Characterization of Acid-Sensing Ion Channel Expression in Oligodendrocyte-Lineage Cells

Keywords: ischemia, inflammation, intracellular calcium

Oligodendrocyte Lineage Cultures

Primary OLC cultures were prepared from 1-day-old postnatal Sprague–Dawley rats by a panning method as previously described (Itoh et al., 2002). Briefly, mixed glial cultures prepared from enzymatically and mechanically dissociated, subcortical, rat brain were grown for 5 days in the presence of growth medium (GM) supplemented with 30% B104 neuroblastoma conditioned medium, l-glutamine (6 mM), biotin (10 ng/mL), insulin (5 μg/mL), transferrin (50 μL/mL), sodium selenite (30 nM), progesterone (20 nM), and putrescine (100 μM). Confluent mixed glial cells were resuspended and purified by two cycles of negative panning with a RAN-2 antibody and one cycle of positive panning with an A2B5 antibody. As previously described (see Itoh et al., 2002), the resulting oligodendrocyte progenitor (OP) cultures were 95% A2B5 positive, and O4- and glial fibrillary acidic protein-negative (see Fig. 1). A2B5 (Eisenbarth et al., 1979) and O4 (Sommer and Schachner, 1982) primary antibodies were collected as supernatants from hybridomas. Anti-myelin basic protein was obtained from Novus (Littleton, CO.) Progenitor cells were expanded by proliferation in GM further supplemented with 30% B104 neuroblastoma conditioned medium, PDGF (1 ng/ mL), and FGF2 (5 ng/mL) up to three passages. Immature oligodendrocytes were produced by substitution of GM with unsupplemented differentiation medium (DM) for 2 days, and mature oligodendrocytes (MOs) by substitution with DM for 4 days.

Reverse Transcriptase Polymerase Chain Reaction and Quantitative PCR

Total RNA was isolated from OLC cultures by a standard Trizol method, followed by repurification with RNeasy kits (Qiagen, Valencia, CA). Passage 3 OP cultures (as earlier) were grown in T-75 flasks with GM supplemented with 30% B104 neuroblastoma conditioned medium, PDGF (1 ng/mL), and FGF2 (5 ng/mL). Substitution of GM with DM, lacking conditioned medium and growth factors, produced IO cultures after 2 days and MO after 4 days. Rat forebrain total RNA was similarly prepared from neonatal whole forebrain samples. White matter (optic nerve and corpus callosum) and cortical grey matter tissues were dissected from adult rat brains, and RNA was extracted from these tissues with RNeasy Lipid Tissue kits (Qiagen). RNA was diluted to 1 μg/mL total RNA, estimated by A₂₆₀. Reverse transcription reactions for conventional RT-PCR were performed with Superscript II reverse transcriptase (Invitrogen, Carlsbad, CA) primed with oligo-dT. Conventional PCR was performed with Platinum TAQ hot-start polymerase (Invitrogen) in 50 μL reactions, using 2 μL (10%) of RT reaction as template. The primers employed are shown in Table 1. The cycling parameters used were as follows: 2 min, 94°; 35 cycles of [30 s, 94°; 30 s, 55°; 1 min, 72°]; 10 min, 72°. Agarose (2%) minigels were loaded with 10 μL PCR product, and stained with SYBR Gold (Invitrogen) and imaged with a CCD camera-based gel documentation system (Kodak, Rochester, NY).

Quantitative RT-PCR was performed using Taqman hydrolysis probes (ABI, Foster City, CA), as specified in Table 1. Random primed cDNA templates were prepared with Stratascript or Accuscript First Strand Synthesis kits (Stratagene, La Jolla, CA). Amplitaq Gold-based chemistry (ABI) was used, together with 0.5 μL cDNA reaction as substrate, in 25 μL reactions, and run on a Stratagene MPC3005 thermocycler. Relative levels of gene expression were estimated by a delta C_T method, where the threshold cycle (C_T) for each ASIC probe, from each tissue/developmental stage cDNA template, was normalized to the C_T for the GAPDH housekeeping probe determined for the same template. Levels were expressed by the formula:

Immunofluorescence

Mice expressing an enhanced green fluorescent protein (EGFP) transgene driven by the 2,3-cyclic nucleotide-3-phosphohydrolase (CNPase) promoter in oligodendroglial progenitors and oligodendroglia (CNPase-EGFP mice) (Yuan et al., 2002) were backcrossed to a C57BL/6 background for at least 10 generations prior to use. Two-month-old CNPase-EGFP mice were deeply anesthetized with ketamine/xylazine, then perfused with saline, followed by phosphate-buffered 4% (w/v) paraformaldehyde. Brains were removed then post-fixed at 25°C with phosphate-buffered 4% paraformaldehyde for 1 h. Brains were transected coronally at the level of the corpus callosum and cryoprotected in 30% sucrose for 2 days prior to embedding in OCT. Ten-micrometer frozen sections were cut and subjected to immunohistochemical analysis as follows. Sections were incubated with affinity purified rabbit anti-ASIC1 polyclonal antibodies (Alpha Diagnostic International, San Antonio, TX, Cat. no. ASIC11-A used at 20 μg/mL; Alomone Labs, Jerusalem, Israel, Cat. no. ASC-014) or purified rabbit immunoglobulins (control) overnight at 4°C. The Alpha Diagnostic antibody is selective for ASIC1a, and the Alomone antibody recognizes either ASIC1a or ASIC1b. Following incubation with biotin conjugated donkey anti-rabbit secondary antibody (species specific, Jackson Immunoresearch), the sections were treated with rhodamine conjugated streptavidin (Jackson, diluted 1:500) for 20 min. The sections were post-fixed with −20°C methanol and counterstained with Hoechst 33258. Between all steps the sections were washed 5× with phosphate buffered saline. Fluorescent images were captured with a 40× objective by confocal microscopy (Nikon C1). Figures represent projections of 15 serial 300-nm optical sections.

Electrophysiology

Whole-cell patch clamp recordings were performed on OLC cultures grown on poly-l-lysine-coated glass coverslips, under observation on an upright Nikon E660FN microscope equipped with a 40× water-dipping objective, and differential interference contrast optics. The coverslip was placed in a perfusion chamber with a coverslip bottom (Warner Instrument, Hamden, CT), which was bulk-perfused (~0.4 mL min⁻¹) with physiological external solution of the following composition (in mM): NaCl, 130; KCl, 5.4; CaCl₂, 1.8; MgCl₂, 1; HEPES, 25; d-glucose, 20; pH = 7.4, adjusted with NaOH. Internal pipette solutions were either Cs⁺ or K⁺ based, and had the following composition (in mM): CsCl or KCl, 130; MgCl₂, 2; EGTA, 10; HEPES, 10; pH = 7.4, adjusted with either CsOH or KOH. The local external solution surrounding recorded cells was rapidly changed to alter the external pH, or apply drugs/toxins, with the aid of a multireservoir, pressurized, solenoid-valve driven microperfusion apparatus (Automate Scientific, Berkeley, CA). A flexible glass-polyimide tip, 100 μm ID, was mounted on the supplied perfusion pencil. The tip of this delivery system was brought to within ~20 μm of the cell under study, producing a local laminar stream 10–20 times the diameter of the cells. A continuous stream of pH = 7.4 HEPES-buffered solution was locally applied for control conditions. This solution was of similar composition to the extracellular solution given earlier, except that the CaCl₂ concentration was reduced to 1 mM to minimize the Ca²⁺ block of ASICs (Immke and McCleskey, 2003), and glucose was replaced by d-mannitol. Acid-shifted solutions had identical composition, except that for solutions with pH < 7, MES buffer (25 mM) was used instead of HEPES. Our standard ASIC activating solution was buffered at pH 5.5, but other pH (5.2–6.0) was occasionally employed when appropriate, as described in the text. Qualitatively similar results were obtained regardless of the specific acidic pH employed. Amiloride was dissolved in these extracellular solutions from a 100 mM stock in DMSO. Psalmopoeus venom, reconstituted from a lyophilized stock (Spider Pharm, Yarnell, AZ) in 0.05% bovine serum albumin, was diluted 1:20,000–1,000,000 in the pH 7.4 buffer for application. Synthetic psalmotoxin peptide (PcTx1) was obtained from either Peptides International (Louisville, KY) or Phoenix Pharmaceuticals (Burlingame, CA) and was prepared as aqueous stock solutions from the lyophilized peptide. PcTx1 was used at 10 or 20 nM in the pH 7.4 application buffer, without added carrier protein. The osmolality of all solutions was measured with an osmometer (Wescor, Logan, UT) and adjusted to ~315 mosm/kg with d-mannitol. The local perfusion system could exchange the extracellular solution within ~50 ms.

Recordings were made with fire-polished, Sylgardcoated capillaries (Clark no. 8520, Warner Instrument) that had resistances of 1.2–2.5 MΩ. The signal was measured with an Axon Axopatch 200B; voltage stimuli, including software control of the Automate microperfusion system, were generated by, and signals digitized, with pClamp 9.2 software (Molecular Devices, Sunnyvale, CA). Membrane capacitances were measured with the membrane test routine of pClamp. When that measurement was unavailable (due to occasional nonconvergence of the routine) capacitance was estimated from the telegraphed readout of the whole-cell capacity compensation dial of the Axopatch 200B. Cells were held nominally at −60 mV in voltage clamp experiments, except for brief clamp to −90 mV during applications of acid-shifted solution. Signals were not adjusted for liquid junction potentials, which were calculated to be 2.5 mV with KCl-based pipette solution, and 3.1 mV with CsCl-based solution, such that true transmembrane potentials would be shifted negative to the nominal command or measured values. Data were analyzed with pClamp, Excel (Microsoft, Redmond, WA) and Origin (OriginLab, Northampton, MA).

Calcium Imaging

OP cultures on poly-L-lysine coated glass coverslips were incubated 40–50 min at 23°C in glucose-containing, pH = 7.4, physiological saline (composition as described earlier) with fluo-4-AM (Invitrogen) at 2 μg/mL, supplemented with Pluronic-F127 (Invitrogen), 0.02%. Cells were mounted in the same chamber as used for electrophysiology, perfused with pH = 7.4 buffered saline for at least 30 min prior to initiating imaging to remove excess fluo-4 and allow for intracellular hydrolysis. Cells were locally perfused with the 100-μm tip, under conditions and with the same solutions as described earlier for electrophysiology experiments. Cells were imaged with the 40× water dipping objective (NA = 0.8) on the Nikon E600FN microscope, and illuminated by a xenon arc lamp (Sutter LS; Novato, CA) through an epifluorescence filter set excited at 500 nm and emitting at 542 nm (filter set YFP-2427A, Semrock, Rochester, NY). Images were detected by a Coolsnap ES (Photometrics, Tucson, AZ) interline CCD camera, and acquired with SimplePCI software (Hamamatsu, Bridgewater, NJ). Images were acquired at ~0.5–1 s⁻¹ with exposure times of 0.2 s, with 3 × 3 binning. The fluorescence intensities of regions of interest (ROI; typically entire cell bodies, or proximal regions of processes) were analyzed with the Dynamic Intensity Analysis module of SimplePCI. Averaged intensity values for each ROI on each image frame so obtained were exported to Excel and/or Origin for further analysis. To normalize for variations in dye loading or cell thickness, intensity data are expressed as ΔF/F_o, where F is the ROI average from each frame and F_o is the “resting” baseline intensity for the same ROI, averaged over five frames immediately prior to the application of lowered extracellular pH. Background fluorescence (from a region within the same field lacking cells or processes) was subtracted from ROI averages.

Materials

Chemicals whose sources were not otherwise mentioned were from Sigma (St. Louis, MO) or Fisher (Pittsburg, PA.).

RT-PCR in OLC

Reverse transcription was performed on total RNA extracted from OP, IO, and MO. Thirty-five cycles of PCR were performed with each cDNA using the primer pairs listed in Table 1. The products of these PCR reactions are shown in Fig. 2A. The appearance of the expected fragments in PCR reactions indicated the presence of ASICs 1a, 2, and 4 at all three developmental stages of cultured OLC. ASIC3 template was weakly detectable with the highly-sensitive dye SYBR Gold only in cDNA prepared from IO and MO; the amount of product produced after 35 cycles of PCR suggested that the ASIC3 substrate was present at low copy number. ASIC4 product appeared most abundant in OP.

Quantitative PCR

To establish mRNA expression levels more definitively, quantitative, real-time RT-PCR (qPCR) was performed on cDNA derived from OP, IO, and MO. The results are summarized in Fig. 2B. Taqman probes were selected to identify ASIC1a, ASIC1b, ASIC2 (nonselective), ASIC3, and ASIC4. A probe for GAPDH was used as a house-keeping probe to normalize for mRNA/cDNA template. qPCR experiments with newborn rat forebrain cDNA were also performed with the same probes, to compare expression levels in OP to a neuron-rich mRNA source. In agreement with standard PCR experiments, ASIC1a, ASIC2, and ASIC4 were all detected at each developmental stage, with maximal levels in OP, and decreased levels in IO and MO. At all stages, ASIC1a and ASIC2 were detected at similar levels and ASIC4 at levels 2–4-fold lower. ASIC 3 was detectable only in late cycles (≥35), if at all. ASIC1b was not detected. In general, the expression levels for each of these genes observed in OP were similar (within 1–2 C_T) to those seen in whole forebrain.

RT-PCR in White Matter

To verify that ASIC transcripts are expressed in white matter tissue in situ, RT-PCR was also performed on RNA isolated from adult rat corpus callosum and optic nerve (see Fig. 3). Adult rat cortical grey matter was used as a positive control. In these experiments additional primer pairs (Table 1) were used to distinguish between ASICs 2a and 2b. ASICs 1a and 2a mRNAs were detected strongly in RNA preparations from each of these sources. Lesser abundances of mRNAs encoding ASICs 3 and 4 were observed, with both being weakly detectable in optic nerve and cortical grey, but only ASIC4 being detected in corpus callosum. ASIC 2b was not detected, suggesting that ASIC2 expression in cultured cells is likely to represent the ASIC2a splice variant. The relative expression levels of ASIC4 in these white matter tissues is apparently lower than in OP, possibly indicating an upregulation of ASIC4 expression in our cultured cells. These experiments demonstrate the expression of several ASIC transcripts in white matter, and show that expression of these transcripts in white matter is by and large similar to that in grey matter.

Immunofluorescent Localization of ASIC1a in White Matter

To verify that ASIC proteins are expressed in cells of the oligodendroglial lineage in situ, we labeled cells in the corpus callosum from a 2-month-old CNPase-EGFP mouse with antibodies against ASIC1a (see Fig. 4). In this transgenic mouse strain, oligodendroglial progenitors and immature and mature oligodendroglia are intrinsically labeled with EGFP driven by the CNPase promoter (Bannerman et al., 2007; Yuan et al., 2002) Our attempts to label cells in white matter of perinatal or adult rats were unsuccessful with available antibodies (four commercial anti-ASIC1 antibodies raised against different immunogenic peptides in all). Two different antibodies, one selective for an epitope in the extracellular loop of ASIC1a, and one that recognizes the C-terminus of both ASIC1a and ASIC1b, successfully labeled mouse white matter and gave similar staining patterns. The results shown in Fig. 4 are with the ASIC1a selective antibody.

As shown in Fig. 4, many, but not all of the EGFP expressing cells in the corpus callosum are co-labeled with the anti-ASIC1a antibody. The cluster of small anti-ASIC1a labeled cells in the upper left corner of Fig. 4A, that are not positive for EGFP, are presumptively neurons of the cingulate cortex. Widespread labeling of neurons in the CNS by anti-ASIC1 antibodies has been previously reported (Alvarez de la Rosa et al., 2003).

Electrophysiological Characterization

Whole-cell voltage clamp recordings from OP (n = 45) detected inward currents representing functional ASIC channels (I_ASIC). Cells were held at −60 mV, and then switched to −90 mV during acidic solution application. In response to brief (1.5–3 s) applications of acidic (pH 5.2–6.0) extracellular solution delivered at 30-s intervals, rapidly desensitizing inward currents in the range of −7 to −325 pA were observed. Qualitatively similar results were obtained regardless of the specific acidic pH applied, and the following values are pooled results from all experiments. Thirty-six of 45 OP cells (80%) responded to acid with detectable inward currents. For those OP cells with non-zero responses, the mean initial current magnitude was −63.5 ± 72 pA, and the mean current density was −7 ± 7 pA/pF. Current densities ranged from −0.7 to −22.6 pA/pF, indicating a highly variable level of ASIC expression across the population.

A small number of high-quality recordings were made from IO cells. Currents were detected in only one of five IO cells; that cell had a current magnitude of −37 pA and a current density of −2.2 pA/pF. Although the number of recorded IO cells is too small to draw statistical conclusions, these results suggest that while IO cells may express functional ASIC currents, they are apparently of lower density than those of OP cells.

The currents were virtually completely blocked (n = 10) by amiloride (100 μM), as shown in Fig. 5. Amiloride block was quickly reversed upon washout, but currents generally did not recover to their original magnitude due to long-lasting desensitization or tachyphylaxis (Chen and Grunder, 2007) which was commonly observed in these cells. Typically, after even one application (3 s, pH = 5.5) of low pH solution, the response magnitude dropped 40–50% on subsequent trials delivered at 30 s or longer intervals. These results are in agreement with the observations of Sontheimer et al. (1989) in a prior study of OLC responses to lowered pH. Tachyphylaxis lasted many minutes (up to 1 h) and was not reversed by application of high calcium (10 mM) pH = 7.4 solution. Similar long-lasting tachyphylaxis was described in oocytes expressing cloned ASIC1a (Chen and Grunder, 2007), but was not observed with other ASIC variants expressed alone or in combination with ASIC1a. This result therefore suggests that homomeric ASIC1a contributes predominantly to the I_ASIC in OP.

A further property that distinguishes homomeric ASIC1a from other functional ASIC variants is the sensitivity of ASIC1a to a toxin (PcTx1) from the venom of the tarantula spider Psalmopoeus christei (Chen et al., 2005; Escoubas et al., 2000). As shown in Fig. 6, I_ASIC was virtually completely blocked (n = 7) by Psalmopoeus christei venom (1:20,000), indicating that the functional I_ASIC was carried predominantly through homomeric ASIC1a (Diochot et al., 2007; Escoubas et al., 2000). The action of the toxin was rapid; with complete block within 30 s. Block was partially reversible with a time course of the order of minutes. The lack of complete recovery was likely due to a combination of slow dissociation/washout of the toxin and tachyphylaxis, as described earlier. Although block by Psalmopoeus venom has only been reported for ASIC1a homomeric channels (Diochot et al., 2007; Escoubas et al., 2000), we cannot rule out the possibility that at 20,000-fold dilution non-specific blocking of other ASIC variants might occur. Therefore, we performed similar experiments with synthetic PcTx1 peptide (10–20 nM) and observed block of most or all current in three OP cells. In one cell current inhibition by the peptide was virtually complete, while in two others, the current was inhibited by nearly 80% (see Fig. 7), suggesting that in at least some cells a fraction of the current may be carried through channels that are not homomeric ASIC1a, which might be expected since transcripts for ASICs 2 and 4 were detected in OP.

Application of acidic extracellular solution causes a rapid depolarization of OP, detectable under whole-cell current clamp conditions (see Fig. 8). The magnitude of depolarization and the resting membrane potential was dependent on the quality of the recording seal, as expected, but in cells with high input resistance (1–2 GΩ) and strongly negative resting potentials (near −60 mV,) the membrane depolarized reversibly to levels of −30 to −20 mV even when the magnitude of I_ASIC was small (10–20 pA). The mean resting potential for four OP cells was −55 mV (±7.6) and mean depolarization was 33.4 mV (±14).

Calcium Imaging

In contrast to the high selectivity for sodium permeability shown by ASICs including 1b or 2a subunits, homomeric ASIC1a channels are reported to have considerable calcium permeability (Gunthorpe et al., 2001; Waldmann et al., 1997). To determine whether intracellular calcium levels rise detectably during ASIC activation by acidic extracellular medium application, we imaged cells loaded with the calcium-sensing dye Fura-4-AM. Image frames were collected every 0.2–0.5 s. After a baseline period for several frames in standard pH 7.4 extracellular solution, pH 5.5 solution was locally applied with the rapid exchange system for 1.5 s before returning to pH 7.4. In some experiments amiloride was co-applied at pH 5.5. In experiments with dilute Psalmopoeus venom (1:100,000; n = 3 different fields) or PcTx1 synthetic peptide (20 nM; three different fields) brief pH drops were delivered after 1–2 min preapplication of toxin, and again after 8–10 min of perfusion with toxin free solution. In some experiments kainic acid (300 μM) and cyclothiazide (50 μM) were applied for 1.5 s as a positive control for calcium perturbation.

Figure 9 shows results of a typical experiment in which dilute Psalmopoeus venom (1:100,000) was used to block the response to low pH. Figure 10 shows an experiment that employed synthetic PcTx1 to inhibit the response. As predicted for homomeric ASIC1a channels, upon application of pH 5.5 extracellular solution (1.5 s application) the fluorescence intensity began to rise within a fraction of 1 s (Figs. 9 and 10) and peaked within 1–4 s. Little or no change in fluorescence was observed when amiloride (100 μM) was co-applied with the pH 5.5 (n = 8; not shown.) Recovery to 1/e of peak value occurred within 10–20 s. Calcium elevation was detected both in the cell body and processes of OP in >20 different experiments. As shown in Figs. 9 and 10, cells varied in the degree of calcium response to an ~1.5 s application of acidic solution. Twenty percent to 40% of ROIs in a given field showed no detectable response to application of pH 5.5. In experiments in which kainic acid was used as a positive control, virtually every imaged cell responded to kainic acid, including the 20–40% that lacked responses to pH. For most cells the kainic acid response was stronger than that for low pH (see Fig. 10). Both Psalmopoeus venom (see Fig. 9) and synthetic PcTx1 (see Fig. 10) completely eliminated the responses to low pH, indicating that calcium elevation requires activation of homomeric ASIC1a channels. In the experiment depicted in Fig. 10 kainic acid elicited strong responses in the presence of PcTx1.

We also examined whether acid application could initiate calcium responses in differentiating OLC. In contrast to the readily observed responses in the majority of OP, calcium responses in IO were very rare. Of more than 200 IO cells, distributed in 12 different fields, only one cell responded unequivocally to application of pH 5.5. Results from that cell are shown in Fig. 11. The cell showed a dramatic increase in fluorescence in both the cell body and in most of its network of processes, starting within 0.5 s after the switch to pH 5.5 for 1.5 s. After a return to pH 7.4 for ~30 s, a second application of pH 5.5 elicited a weaker fluorescence increase. The relative fluorescence increase observed in that IO cell was greater than responses typically observed in seen in OP cells, and took longer to reach a peak. A possible explanation for these differences is that secondary amplification of calcium signaling, perhaps including release from intracellular calcium stores, occurred subsequent to initiation by ASIC1a activation. Considering the rarity of acid-induced responses in IO, it will be challenging to clarify the nature of the response and compare it to the responses of OP. The observation of this single response suggests that although differentiating OLC may occasionally express levels of homomeric ASIC1a sufficient to produce measurable calcium responses, apparently such acid-induced calcium responses are largely restricted to OP cells.

To explore the requirement for extracellular calcium influx via ASICs, we performed experiments comparing responses in oligodendroglial progenitors in reduced calcium (i.e. no calcium added to extracellular solutions, but with no calcium chelators present) to those in our standard 1 mM calcium-containing solutions. Because of the intimate role of calcium ions in ASIC gating and inactivation mechanisms (Chen and Grunder, 2007; Immke and McCleskey, 2003; Paukert et al., 2004), coupled with the uncontrolled release of protons and resulting acidification that would occur upon exchange of calcium containing solutions with chelator-containing solutions, we did not employ chelators to assure very low calcium concentrations. As a result, under our conditions with zero-added calcium solutions, we have by no means completely removed calcium. Nevertheless, despite these concerns, responses to pH 6.0 in zero-added calcium (after a pre-incubation in zero-added calcium, pH 7.4 for 10 s) were highly variable, but on average 30–50% smaller than subsequent responses in the same cells after return to 1 mM calcium (not shown). Although not ideal, these results support the conclusion that acid-induced calcium elevation does involve influx of extracellular calcium.

Expression and Molecular Identity of OLC ASICs

We have shown that OLC, as well as adult CNS white matter tissues, express three of the four genes that comprise the ASIC family, and that the genes expressed and their relative levels resemble those found in total brain RNA. The ASIC1 and ASIC2 genes were expressed at similar levels, and were more abundant than ASIC4. The ASIC1a splice variant was detected, but not the ASIC1b variant, consistent with prior studies localizing ASIC1b exclusively to peripheral neurons (Bassler et al., 2001). Similarly, in standard RT-PCR experiments with white matter- derived cDNA templates, ASIC2a, but not 2b, was detected. ASIC3, which is also largely restricted to peripheral neurons (Chen et al., 1998), was weakly detected in differentiating OLC cDNAs and in adult optic nerve, but not at all in OP cDNA or in corpus callosum.We have additionally detected the presence of immunoreactive ASIC1a protein in mouse oligodendroglial cells. The appearance of ASIC3 transcript only in differentiating OLC, albeit at a very low copy number, suggests a pattern of developmental regulation opposite that of each of the other ASICs genes. ASICs 1a, 2, and 4 all were detected several fold higher in OP than in differentiating OLC. This developmental regulation may reflect some yet unknown role(s) played by ASICs in early OLC development.

Functional Properties of OP ASICs

Although mRNAs for three different ASIC genes are expressed at readily detectable levels in OLC, the ASIC1a subtype appears to contribute a majority of functional ASICs detectable under our whole-cell recording conditions, as shown by our observation that the tarantula toxin blocks most or all of the detected current. The pronounced ASIC tachyphylaxis observed in OP further supports the conclusion that homomeric ASIC1a comprises a substantial fraction of functional channels in OP. In addition, the complete inhibition of detectable acid-induced calcium elevation by psalmotoxin PcTx1 lends further support to the conclusion that toxin-insensitive ASICs are likely present at low density, since depolarization and sodium influx resulting from their activation would be expected to activate voltage gated calcium channels and/or reverse cycling of sodium/calcium exchangers (see discussion below). The apparent low level of functional PcTx1-insensitive ASICs is a surprising finding since coexpression of multiple cloned ASIC subtypes in Xenopus oocytes or transfected cells generally results in the expression of ASICs with altered properties compared with ASIC1a expressed alone, indicating that multiple subtypes readily co-assemble.

Implications for Cellular Signaling

ASIC1a has garnered considerable interest due to its low, yet significant calcium permeability. The many signaling pathways mediated by calcium raise the possibility that activity of ASICs may trigger important physiological processes and may contribute to pathophysiology of OLC. It was recently demonstrated with a mouse ASIC1a knockout model that ASIC1a activity under ischemic conditions accompanied by lowered extracellular pH in the CNS can cause the loss of neurons through a calcium-dependent mechanism analogous to excitotoxic cell death (Xiong et al., 2004, 2006); a neuroprotective effect of psalmotoxin has also been described in a stroke model (Pignataro et al., 2007); those studies considered only neuronal damage and did not discuss white matter damage that accompanies neuronal injury in these models. As we and others have shown, OLC are vulnerable to calcium-mediated excitotoxicity under conditions of inflammation or ischemia due to activation of calciumpermeable AMPA (Deng et al., 2003; Follett et al., 2004; Itoh et al., 2002; McDonald et al., 1998; Sanchez-Gomez and Matute, 1999) or NMDA (Karadottir et al., 2005; Salter and Fern, 2005) receptors. Our observation that ASIC activation can produce elevation of intracellular calcium supports a new independent pathway through which calcium can enter OLC under pathological conditions accompanied by lowered extracellular pH. This calcium-permeable pathway may operate independently from, or in concert with, glutamate-operated channels. A synergy between the actions of NMDA receptors and ASIC1a has been demonstrated in neurons in an ischemia model, involving phosphorylation of the ASIC1a protein through a Ca²⁺/calmodulin-dependent protein kinase II (CaMKII) mediated pathway (Gao et al., 2005), which leads to enhancement of ASIC-mediated currents.

The expression of many types of ligand- and voltage gated channels in OLC at various developmental stages provides a tableau rich with possibilities for crosstalk that could work in synergy with ASICs. In addition to direct calcium influx through ASIC1a, ASIC activation may also affect OLC by membrane depolarization, which as we have shown here, can be considerable (>50 mV depolarization from resting potential). NMDA receptors, which are expressed by OLC in vivo (Karadottir et al., 2005; Salter and Fern, 2005), could be activated by relief from Mg²⁺-block due to depolarization initiated by I_ASIC. Such a mechanism would provide a reciprocally synergistic network, together with the CaMKII-mediated activation of ASIC1a described earlier that could act when glutamate and low pH are present simultaneously, as would be expected under ischemic or inflammatory conditions. The net result would be a stimulation of Ca²⁺ influx greater than either pathway would produce on its own.

Alternatively, ASIC-mediated membrane depolarization could activate voltage-dependent channels, including sodium (Sontheimer et al., 1996; Sontheimer and Waxman, 1993), potassium (Barres et al., 1990; Bevan et al., 1987; Sontheimer and Waxman, 1993), or calcium (Berger et al., 1992; Blankenfeld Gv et al., 1992), all of which are expressed by developing OLC. Both low-voltage and high-voltage activated subtypes of voltage-dependent calcium channels are found in OLC, with a differential pattern of distribution such that low-voltage activated T-type channels are localized at the tips of processes, while high-voltage activated channels are distributed on the soma (Kirischuk et al., 1995) Both types of voltage-gated calcium channels could be activated by depolarizations of the magnitude we report here. Activation of voltage-dependent calcium channels by AMPA/KA receptor-mediated membrane depolarization has been previously reported (Borges et al., 1994; Liu et al., 1997) and therefore seems a likely consequence of ASIC-mediated depolarization. An additional means by which ASIC activation could lead to elevated intracellular calcium would be by reverse action of the Na⁺/Ca²⁺ exchanger, driven by intracellular Na⁺ accumulation and depolarization. Reversed Na⁺/Ca²⁺ exchange has been reported to supplement AMPA receptor-mediated calcium accumulation in OLC (Liu et al., 1997). Furthermore, such a mechanism contributes to axonal damage in demyelinating conditions (Craner et al., 2004; Stys et al., 1992, 1993), and might be especially relevant in the fine processes of OLC where significant Na⁺ accumulation likely occurs during receptor activation. Finally, calcium- induced calcium release from intracellular stores could serve to further amplify the calcium signal (reviewed in Bootman et al., 2002). Inositol triphoshate and/or ryanodine receptors expressed by OLC (Haak et al., 2001) could be triggered to release calcium from intracellular stores after an initial influx of calcium via ASICs. Despite the presence of these various potential pathways for calcium elevation, our finding that psalmotoxin blocks any detectable acid-induced calcium elevation suggests that the responses to acid must be initiated by homomeric ASIC1a activation. This discussion of candidate calcium pathways that could be triggered by, or interact with, ASIC activation is presently speculative. An important next step will be a determination of the relative contributions of calcium influx vs. mobilization of intracellular calcium stores that result from ASIC activation. Further study will be needed to dissect the contributions of the afore-mentioned candidate path-ways to the calcium elevations we describe here.

We have shown that OLC ASICs can respond to acidshifted extracellular environment by both depolarizing the cells and elevating intracellular calcium. It is not yet known what physiological functions are served by ASICs in OLC. Since both normal and pathological physiological processes in the brain can lead to perturbations of pH, it seems likely that ASICs could play roles in oligodendrocyte development, pathology, and perhaps regeneration.

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