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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS
Department of Pathobiology, University of Pennsylvania School of Veterinary Medicine, Philadelphia, Pennsylvania
Submitted 11 May 2005 ; accepted in final form 20 October 2005
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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calcium; arachidonic acid; membrane potential; hypotonicity; cytochrome P-450
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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(for whole cell recording) or 5–10 M (for single-channel recording) tip resistance (Sutter Instrument, Novato, CA) from borosilicate glass and were backfilled with appropriate internal solution. Liquid junction potentials were calculated and corrected manually using the patch-clamp amplifier. Capacitance and access resistance were monitored continuously. Between 50% and 70% of the series resistance was electronically compensated to minimize voltage errors. Command potentials were generated using an EPC-9 patch-clamp amplifier (Heka Elektronik, Lambrecht, Germany), and currents were acquired, stored, and analyzed using PulseFit software (Heka Elektronik). Single-channel amplitude frequency analysis was performed using QuB software (Research Foundation, State University of New York, Buffalo, NY). Whole cell currents were recorded using the standard whole cell mode of the patch-clamp technique. B cells were initially held at 0 mV and then stepped to –80 mV for 100 ms, followed by a 100-ms linear voltage ramp to +80 mV with a sampling interval of 0.2 ms. The entire protocol was repeated every 2 s, and the time course of whole cell currents was obtained by plotting the current amplitude at –80 mV for each cycle. Hypotonicity- and agonist-activated inward currents typically reached a steady-state level within 1–2 min. These steady-state amplitude values were tabulated and reported as mean current amplitudes. Cell membrane capacitance values were used to calculate current densities and expressed as mean current amplitudes per unit of membrane capacitance. All ramp currents shown were leak corrected by subtracting ramp currents obtained after the establishment of a stable whole cell recording from the ramp current obtained after agonist-induced current activation.
The standard extracellular bath solution contained (in mM) 154 Na+-gluconate, 4.5 K+-gluconate, 1 MgSO4, 2 Ca2+-gluconate, 10 HEPES, and 10 D-glucose (pH adjusted to 7.30 using NaOH), and the normal isotonic bath solution contained 90 Na+-gluconate, 4.5 K+-gluconate, 2 Ca2+-gluconate, 1 MgSO4, 10 HEPES, 10 D-glucose, and 100 D-mannitol [100 D-mannitol was omitted to produce hypotonic (200 mosmol/l) bath solution], pH 7.30. The standard pipette solution contained 155 Cs+-methanesulfonate, 6 MgSO4, 1 EGTA, 0.25 Ca2+-gluconate, and 10 HEPES (pH adjusted to 7.3). For measurement of the relative Ca2+ vs. Na+ permeability of channels, the bath solution contained 100 N-methyl-D-glucamine (NMDG)-Cl–, 30 CaCl2, 20 HEPES, 10 D-glucose, and 100 µM 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) (pH 7.3), and the pipette solution contained 145 NMDG-Cl–, 0.2 EGTA, and 10 Na+-HEPES (pH 7.3). Hypotonic bath solution was generated by omitting 50 NMDG-Cl–. The relative ion permeability (PCa/PNa) was calculated from the measured reversal potential using the equation Erev = RT/2F ln {4PCa2+ [Ca2+]o/PNa+ [Na+]i} as described previously (10), with [Ca2+]o representing external Ca2+ concentration and [Na+]i representing intracellular Na+ concentration.
Single-channel recordings were obtained using the cell-attached configuration of the patch clamp at a holding potential (Vmembrane) of –60 mV. For single-channel recordings, the bath solution contained 150 KCl, 5 MgCl2, 10 HEPES, and 10 glucose. Ca2+ was not added to these solutions, because it has been shown to accelerate the inactivation of certain transient receptor potential vanilloid (TRPV) channels (25). The pipette solution contained 150 NaCl, 1 MgCl2, and 10 HEPES, pH 7.3. Both bath and pipette solutions contained 100 µM NPPB to block Cl– channels, 100 nM charybdotoxin (CTX) to block Kv and KCa channels, and 5 mM tetraethylammonium chloride (TEA-Cl–) to block other K+ channels. The single-channel conductance was calculated from the current amplitude histogram. Membrane-permeant drugs were applied by direct addition to the bath; however, membrane-impermeant inhibitors were dialyzed into single cells from the patch-clamp recording microelectrode during whole cell recordings. AA, 4
-phorbol-12,13-didecanoate (4-PDD), and RHC-80267 were obtained from EMD Biosciences (San Diego, CA). 5,6-EET, 20-HETE, 5,8,11,14-eicosatetraynoic acid (ETYA), and 17-octadecynoic acid (17-ODYA) were obtained from Biomol Research Laboratories (Plymouth Meeting, PA), and 5-HPETE and ruthenium red (RR) were obtained from Sigma-Aldrich (St. Louis, MO).
Membrane potential measurements. The B-lymphocyte Vm was measured directly using the current-clamp configuration of the whole cell patch-clamp technique as described previously (10). Cells were bathed in a solution containing (in mM) 155 NaCl, 4.5 KCl, 2 CaCl2, 1 MgCl2, 10 Na+-HEPES, and 10 glucose adjusted to pH 7.3 with NaOH. A low-Na+ Ringer solution used to confirm that changes in Vm were due to Na+-permeant NSCC channels contained 155 NMDG-Cl–, 4.5 KCl, 2 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES. The pipette solution for Vm measurements contained 30 KCl, 125 K+-gluconate, 2 MgSO4, 0.25 CaCl2, 1 EGTA (62 nM free Ca2+), and 10 HEPES.
Cell adhesion assays. Cell integrin avidity changes were assessed using an ICAM-1 and VCAM-1 adhesion assay similar to that described previously (10, 26). Immunolon 4HBX plates (Thermo Labsystems, Franklin, MA) were blocked with 1% BSA in PBS for 1 h at 37°C to prevent nonspecific binding, after which plates were washed twice with PBS. Murine ICAM-1 (human) Fc fusion protein (10 µg/ml; R & D Systems, Minneapolis, MN), murine VCAM-1 (human) Fc fusion protein (20 µg/ml; R&D Systems), or human IgG (10 µg/ml; Jackson ImmunoResearch, West Grove, PA) were added to wells for 1.5 h at 37°C, followed by two PBS washes. Splenocytes (4 x 106/ml, 100 µl) in Complete RPMI 1640 and an additional 100 µl of Complete medium containing the indicated stimuli were added to each well. Cultures were incubated for 0.5 h at 37°C. To remove the unbound cells and media, the wells were washed extensively with PBS. Adherent cells were subsequently removed from wells by incubating them with Complete RPMI 1640 medium containing 5 mM EDTA (200 µl) for 15 min at 4°C, and then they were counted, stained with anti-B220, and analyzed using flow cytometry to determine the absolute number of B cells recovered from each well. Results are expressed as the percentage of total B cells recovered from plates after subtraction of nonspecific binding values obtained from wells treated with human IgG. These studies were reviewed and approved by the Institutional Animal Care and Use Committee of the University of Pennsylvania.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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To further define the mechanism of hypotonicity-induced Ca2+ entry into B cells, we measured membrane currents to characterize hypotonicity-activated channels. We first determined whether DAG lipase activity is required for channel activation. In control cells, hypotonicity typically (>65% of primary B cells tested; n = 47) elicited an inward current with a mean peak steady state amplitude (mean current amplitude) of –59.5 ± 10.1 pA (n = 28) (Fig. 1A). These hypotonicity-induced currents were identical to those described previously (10). Because these recordings were performed with Cs+ in the pipette (to block K+ channels) and Cl–-free pipette or bath solutions (to prevent the development of Cl– conductance), the current reversal potential of 
0 mV (see Figs. 1 and 2, right) indicates that these stimuli activated a channel with equal permeability to intracellular Cs+ cations and extracellular cations Na+ and Ca2+; therefore, by definition, the channel was a NSCC. The frequency of hypotonicity-induced current activation was significantly reduced by treatment with the DAG lipase inhibitor RHC-80267 (–3.5 ± 1.8 pA; n = 12), consistent with a role for AA (derived from DAG) in hypotonicity-induced current activation. Moreover, we found that AA itself directly activated a current (Fig. 1B) (mean current amplitude = –40.7 ± 25.1 pA at Vmembrane = –80 mV, Erev = –4.4 ± 3.4 mV; n = 5), which was indistinguishable from those produced by hypotonicity. However, the DAG lipase inhibitor RHC-80267, which blocked hypotonicity-induced currents (Fig. 1C), did not block the response to subsequent application of AA. Because AA is also produced from membrane lipids by PLA2 isoforms, we examined the effect of the iPLA2 blocker bromoenol lactone (BEL), the sPLA2 blocker 4-bromophenacyl bromide (4-BPB), and the cPLA2 blockers arachidonyltrifluoromethyl ketone (AACOCF3) and chloroquine on hypotonicity-induced currents and found that none of these agents inhibited NSCC activation by hypotonicity (data not shown). These findings are consistent with our assertion that AA derived from DAG is involved in NSCC activation after hypotonic stimulation.
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-PDD (21), we initially focused on TRPV4 as a possible candidate for hypotonicity-induced currents. We found that 2 µM 5,6-EET elicited an inward current (Fig. 1D) (Vmembrane = –80 mV) in primary B cells whose amplitude (–75.4 ± 28.9 pA; n = 7) and reversal potential (–1.2 ± 2.5 mV; n = 7) were similar to those of hypotonicity- and AA-induced NSCCs. 5,6-EET-induced currents such as those elicited from heterologously expressed TRPV4 channels (23–25) also were blocked by 1 µM RR (Fig. 1D). Another unique property of expressed TRPV4 channels is their sensitivity to the non-PKC-activating phorbol ester derivative 4-PDD (24). In B cells, 4-PDD elicited an inward current (Fig. 1E) whose amplitude (–59.8 ± 13.7 pA, Vh = –80 mV; n = 6), reversal potential (–2.6 ± 2.6 mV; n = 11), and RR sensitivity were similar to those of 5,6-EET-activated channels. Multiple eicosanoid-activated NSCCs are expressed in B lymphocytes. If 5,6-EET-activated, RR-sensitive channels (Fig. 1) are the sole pathway for hypotonicity-induced Ca2+ entry into B cells, then RR should block AA- and hypotonicity-induced cation currents completely. In general, we found that this was the case, although we found that AA-activated currents were sometimes incompletely blocked by RR (Fig. 1B), suggesting that distinct NSCCs might also be activated by other products of AA. We focused on oxylipids generated by CYP450 hydroxylase (20-HETE) and by 5-LOX (5-HPETE), which have been reported to elicit RR-insensitive NSCC currents (5, 19, 22) and/or Ca2+ entry into B cells (28). We found that 20-HETE (5 µM) activated an inward NSCC current (Fig. 2A) (–26.4 ± 13.6 pA, Vmembrane = –80 mV; n = 3) (Erev = –10.3 ± 2.5 pA; n = 3) in B cells, as did 2 µM 5-HPETE (Fig. 2B) (–26.5 ± 6.2 pA, Erev = –7.3 ± 2.3 mV; n = 4), although neither was sensitive to RR inhibition (Fig. 2). This difference in RR sensitivity demonstrates that 5,6-EET-activated channels expressed in B cells are distinct from those activated by 20-HETE and 5-HPETE.
To further define these NSCCs in B cells and to determine whether 20-HETE and 5-HPETE activate the same NSCC as hypotonicity, AA, and the other eicosanoids, we defined signature biophysical properties (conductance and Na+ and Ca2+ permeability) of channels activated by each stimulus. For conductance measurements, cells were bathed in 150 mM KCl to clamp the membrane potential at 0 mV and minimize variations in the holding potential of the patch due to passive current flow from adjacent membrane regions. The mean amplitude of agonist-activated single-channel currents was derived by performing amplitude histogram analysis of single-channel recordings (Fig. 3, right) and was used to calculate the conductance. Similar conductance values were obtained for 5,6-EET-activated channels (42.9 ± 3.3 pS; n = 5 records analyzed) (Fig, 3A), 4-PDD-activated channels (42.7 ± 1.9 pS; n = 5) (Fig. 3B), and hypotonicity activated channels (43.8 ± 2.9 pS; n = 7) (Fig. 3C), whereas those activated by 20-HETE (28.1 ± 2.1 pS; n = 5) (Fig. 3D) and 5-HPETE (31.2 ± 1.3 pS; n = 4) (Fig. 3E) had smaller conductances. Because hypotonic solutions are necessarily formulated with lower cation concentrations {in this case, 90 mM external K+ concentration ([K+]o)} for single-channel recordings, we also compared the conductance of 5,6-EET-activated channels in high (150 mM)- and low (90 mM)-[K+]o solutions to determine whether the [K+]o or ionic strength affects this channel property. We found that the conductance of 5,6-EET-activated channels was not different in 90 mM [K+]o solution (41.7 ± 3.4 pS, Vh = –80 mV; n = 4). These measurements confirmed that 5,6-EET-activated channels are distinct from those activated by 20-HETE and 5-HPETE.
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-PDD, and hypotonicity-activated channels in the absence of blockers (Table 1). Finally, when CYP450 epoxygenase was inhibited using miconazole (26.5 ± 1.7 pS; n = 4) (Fig. 3H) and when cells were treated with both miconazole and NDGA (27.2 ± 2.9 pS; n = 6) (Fig. 3I), hypotonicity activated channels whose conductance was similar to that activated by 20-HETE. Thus hypotonicity preferentially activated 5,6-EET-sensitive NSCCs. However, when 5,6-EET production was blocked, other products of AA appeared to be generated in B cells as indicated by the activation of distinct eicosanoid (20-HETE and 5-HPETE)-sensitive NSCCs.
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-PDD-activated channels (0.22 ± 0.1; n = 5) (see Table 2) was similar to previous estimates obtained for hypotonicity (0.20; see Ref. 10). By contrast, 20-HETE- and 5-HPETE-activated channels exhibited Ca2+ and Na+ permeability markedly different from these other channels and from one another (0.95 ± 0.3, n = 4, vs. 0.16 ± 0.1, n = 4, respectively) (Table 2). In summary, measurements of conductance and ion permeability demonstrate that three distinct eicosanoid-activated NSCCs are expressed in primary B cells and that the predominant consequence of hypotonic (and AA) stimulation is activation of TRPV4-like channels.
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-PDD, 20-HETE, and 5-HPETE in this physiological response. We found that each ligand triggered significant plasma membrane depolarization (Fig. 5), which is summarized in Table 3. Both 5,6-EET (+36 mV) (Fig. 5A) and 4-PDD (+38 mV) (Fig. 5B) produced similar changes in Vm. 20-HETE produced the largest change (+47 mV) (Fig. 5C) and 5-HPETE evoked the smallest change in Vm (+31 mV) (Fig. 5D). Thus, for each ligand, the extent of depolarization correlated with the single-channel conductance, with the exception of 20-HETE. This bigger depolarization reflects its greater (5-fold) permeability to divalent Ca2+ (see above). However, for each channel agonist, depolarization was reversed rapidly by superfusion of cells with Na+-free, Cl–-containing solution. Finally, membrane depolarization induced by 5,6-EET and 4-PDD, like respective currents (Fig. 1, D and E) and Ca2+ signals (28), was blocked by RR. Vm changes (Fig. 5, C and D), currents (Fig. 2, A and B), and [Ca2+]i elevation (28) induced by 20-HETE and 5-HPETE were insensitive to this drug. Altogether, these results implicate directly each of the distinct NSCCs we have identified in eicosanoid-induced changes in Vm.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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2 is activated by both; see Refs. 10, 28), NSCC activation by osmotic stress does not involve inositol 1,4,5-trisphosphate but rather AA (and its eicosanoid products) derived from DAG. Our findings therefore point to a new general mechanism by which eicosanoid-inflammatory mediators can activate NSCCs, increase [Ca2+]i, depolarize the plasma membrane, and regulate B-cell adhesion and trafficking independently of BCR and CRAC channels.
Although the goal of these studies was to define the biophysical properties of eicosanoid- and/or AA-activated NSCCs in primary B cells, our results offer some insight into their molecular identity. Previously, we hypothesized that hypotonicity-operated NSCCs in B cells are transient receptor potential (TRP) family members (10). Biophysical and pharmacological similarities between currents in primary B cells and those elicited by heterologously expressed TRPV4 point to TRPV4 as a likely candidate responsible for hypotonicity-induced responses. For example, TRPV4 is activated by changes in osmolarity such as the changes used in our studies. TRPV4 also is activated directly by 5,6-EET and 4-PDD and is inhibited by RR (14, 25). The conductance that we calculated for hypotonicity-activated currents in B cells is also comparable to that reported for TRPV4 (45 pS vs. 50–60 pS). Although our estimates of Ca2+ vs. Na+ permeability are different from those reported for heterologously expressed TRPV4 (14), we think that this variation may reflect the different methodologies used to determine this permeability. Specifically, we used bionic conditions with a single permeant extracellular cation (30 mM Ca2+) and intracellular cation (10 mM Na+) and no Cs+ in the recording solutions, whereas Watanabe et al. (24) determined the difference in reversal potential of currents recorded under multiple sets of recording solutions in different cells. Nonetheless, the degree of difference between these two approaches is somewhat surprising, given the comparable permeability of NSCCs to Na+ and Cs+. To understand this discrepancy better, we also performed a limited set of permeability measurements using conditions and calculations identical to those used by Watanabe et al. (24) and obtained comparable estimates of the relative Cs+-Na+ (1.3 vs. 1.1) and Ca2+-Na+ (8.0 vs. 5.8) permeability of 4-PDD-activated channels in B cells as reported for heterologously expressed TRPV4 channels. Thus the methodology appears to be a critical determinant of the permeability values obtained. However, regardless of which approach provides a better estimate of this property, it is significant that our measurements were internally consistent (Table 2) and enabled us to distinguish between the NSCCs studied. Thus stimuli that are expected to activate only TRPV4 (4-PDD and 5,6-EET) activated a channel with the same permeability and conductance. By contrast, similar conductance values were obtained for 20-HETE- and 5-HPETE-activated channels, but permeability measurements enabled us to distinguish between them.
One important question raised by our findings is the extent to which each of the eicosanoid-activated NSCCs we identified is also activated by hypotonicity. Our single-channel data and permeability measurements indicated that the same 43- to 46-pS channel is activated by hypotonicity, AA, 5,6-EET, and 4-PDD, whereas 20-HETE and 5-HPETE activated distinct channels with smaller conductances. However, hypotonicity- and AA-induced currents sometimes exhibited partial insensitivity to the TRPV4 blocker RR, suggesting that additional NSCCs might also contribute to these responses. It was surprising, therefore, that none of the single-channel recordings exhibited multiple current levels. In fact, the only instance in which hypotonicity was found to activate a smaller conductance channel was when CYP450 epoxygenase and/or hydroxylase was blocked with 17-ODYA (see Fig. 3F and Table 1) or when LOX was blocked alone with NDGA or together with the CYP450 epoxygenase inhibitor miconazole (Fig. 3, G and I; see also Fig. 4). These results suggest that LOX and CYP450 hydroxylase activity are not normally induced by hypotonicity and that TRPV4-like channel activation is the primary consequence of osmotic stimulation.
Our findings also raise important questions concerning the physiological and immunological consequences of hypotonicity- and/or 5,6-EET-induced NSCC activation. Although the extremes in osmolarity used to activate NSCCs in these studies may exceed changes B cells typically experience in vivo, we showed previously (10) that other forms of mechanical stress, such as fluid shear forces found in the vasculature and lymphatics, elicit similar responses. Importantly, shear force has been documented to induce integrin activation in other cell types (1, 17). Of interest in the context of this model are our observations that eicosanoid mediators and the NSCCs that they activate directly regulate the avidity of B-cell binding to ECM integrin ligands, which regulate lymphocyte adhesion to vascular endothelium, extravascular trafficking, and physical interactions between B and T cells in secondary lymphoid organs. These same eicosanoids are produced by the vascular endothelium (2), the pathogens themselves (15), and other leukocytes (20, 27). Therefore, NSCCs might also regulate a much broader range of B-cell immunological functions resulting from interactions with antigen or physical contact with T cells, including the expression of costimulatory molecules or the production of cytokines and antibodies.
Altogether, our results point to a novel general mechanism by which proximal signals that control the production of AA or its metabolic products could have an impact on the proinflammatory functions of B cells. Studies utilizing genetic strategies to alter the expression of NSCCs in lymphocytes are under way, and progress toward the identification of selective and potent antagonists are critical steps needed to understand the full complement of immunological functions for these channels and their utility as targets for modulating inflammatory responses in vivo.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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