Department of Physiology and Neurobiology, University of Connecticut, Storrs, Connecticut 06269
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ABSTRACT |
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Kiss, Laszlo and Stephen J. Korn. Modulation of N-type Ca2+ channels by intracellular pH in chick sensory neurons. Both physiological and pathological neuronal events, many of which elevate intracellular [Ca2+], can produce changes in intracellular pH of between 0.15 and 0.5 U, between pH 7.4 and 6.8. N-type Ca2+ channels, which are intimately involved in exocytosis and other excitable cell processes, are sensitive to intracellular pH changes. However, the pH range over which N-type Ca2+ channels are sensitive, and the sensitivity of N-type Ca2+ channels to small changes in intracellular pH, are unknown. We studied the influence of intracellular pH changes on N-type calcium channel currents in dorsal root ganglion neurons, acutely isolated from 14-day-old chick embryos. Intracellular pH was monitored in patch-clamp recordings with the fluorescent dye, BCECF, and manipulated in both the acidic and basic direction by extracellular application of NH4+ in the presence and absence of intracellular NH4+. Changes in intracellular pH between 6.6 and 7.5 produced a graded change in Ca2+ current magnitude with no apparent shift in activation potential. Intracellular acidification from pH 7.3 to 7.0 reversibly inhibited Ca2+ currents by 40%. Acidification from pH 7.3 to pH 6.6 reversibly inhibited Ca2+ currents by 65%. Alkalinization from pH 7.3 to 7.5 potentiated Ca2+ currents by approximately 40%. Channels were sensitive to pHi changes with high intracellular concentrations of the Ca2+ chelator, bis-(o-aminophenoxy)-N,N,N',N'-tetraacetic acid, which indicates that the effects of pHi did not involve a Ca2+-dependent mechanism. These data indicate that N-type Ca2+ channel currents are extremely sensitive to small changes in pHi in the range produced by both physiological and pathological events. Furthermore, these data suggest that modulation of N-type Ca2+ channels by pHi may play an important role in physiological processes that produce small changes in pHi and a protective role in pathological mechanisms that produce larger changes in pHi.
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INTRODUCTION |
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Both excitatory and inhibitory stimuli can acidify
neurons. Elevation of intracellular Ca2+ (Ahmed and
Connor 1980; Meech and Thomas 1977
; Werth
and Thayer 1994
), application of glutamate (Canzoniero
et al. 1996
; Dixon et al. 1993
; Hartley
and Dubinsky 1993
; Irwin et al. 1994
;
Wang et al. 1994
), and application of gamma-amino
butyric acid (GABA) (Kaila et al. 1993
;
Luckermann et al. 1997
) produce transient intracellular
acidification by as much as 0.15-0.5 pH units in mammalian and
nonmammalian neurons. In addition, Ca2+ influx associated
with action potentials acidifies neurons by ~0.2 U (Ahmed and
Connor 1980
; Trapp et al. 1996
), experimentally induced hypoxia acidifies neurons by as much as 0.5 U
(O'Donnell and Bickler 1994
), and reuptake of glutamate
into rat hippocampal neurons is accompanied by intracellular
acidification of ~0.2 pH units (Amato et al. 1994
).
Intracellular acidification is associated with changes in membrane
excitability (Church 1992
), neurotoxicity (Nedergaard et al. 1991
), and possibly protection from
pathological events such as reperfusion injury after ischemia
(Bond et al. 1993
; Scholz et al. 1992
;
Tombaugh and Sapolsky 1990
; Vornov et al.
1996
). However, the potential importance of small changes in
intracellular pH (pHi), and the mechanisms by which small
or large changes in pHi can influence physiological and/or
pathological events, is poorly understood.
Both N- and L-type Ca2+ channels in mammalian and
nonmammalian tissues are sensitive to changes in pHi
(Kaibara and Kameyama 1988; Klockner and Isenberg
1994
; Mirinov and Lux 1991
; Takahashi et
al. 1993
; Tombaugh and Somjen 1997
). In catfish
horizontal cells, L-type Ca2+ channels are quite sensitive
to pHi changes between 6.6 and 7.6; acidification from pH
7.3 to 7.0 produced a 30-40% reduction in current amplitude, and
alkalinization from pH 7.3 to 7.6 potentiated Ca2+ currents
by ~40% (Dixon et al. 1993
). Although not as
precisely quantified, L-type Ca2+ channels from mammalian
vascular smooth muscle were also sensitive to pHi changes
between 6 and 8.4 (Klockner and Isenberg 1994
). In
contrast to these results, the pHi range over which N-type Ca2+ channels are sensitive is not known.
In hippocampal neurons, N-type Ca2+ channels appear to be
more sensitive than L-type Ca2+ channels to changes in
pHi (Tombaugh and Somjen 1997). This
difference was reflected as a larger change in amplitude of N- than
L-type currents on application of NH4+. However,
pHi was not measured, and pHi among cells and
over time can vary by as much as 0.4-0.5 U under identical patch clamp recording conditions (unpublished data). Consequently, it is not known
whether this difference reflected a difference in the pHi ranges over which these two channel types were sensitive or different channel responses to identical absolute changes in pHi.
The observations described above raise two important questions: are the
small (0.15-0.5 U) changes in pHi that can be produced by
a variety of physiological and pathological stimuli relevant to the
physiology of N-type Ca2+ channels and is the
pHi sensitivity of N-type Ca2+ channels similar
to or different from that reported for L-type Ca2+
channels? To address these questions, we examined the pHi
dependence of N-type Ca2+ channel currents in acutely
isolated chick dorsal root ganglion neurons (DRGs), which contained a
pharmacologically pure population of N-type Ca2+ channels.
To obtain consistent, quantitative data regarding pH sensitivity below
pH 7.3, we controlled pHi with both intracellular and
extracellular NH4+ (Grinstein et al.
1994). Our results indicate that N-type Ca2+
channels are quite sensitive to small changes in pHi
between 6.6 and 7.5, and that the pH sensitivity is quantitatively
similar to that reported for L-type Ca2+ channels in
catfish horizontal cells (Dixon et al. 1993
). These data
suggest that acidification-induced inhibition of N-type
Ca2+ channel activity may be important during both
physiological and pathological events. Some of these data were
presented in abstract form (Callahan et al. 1995
).
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METHODS |
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Cells
Dorsal root ganglion neurons were acutely isolated from lumbar
level ganglia of 14-day white leghorn chick embryos (UCONN Poultry
Farm, Storrs, CT). Cells were prepared as described previously (Polo-Parada and Korn 1997). In photometry experiments,
cells were plated on polyornithine-coated glass cover slips glued to the drilled-out bottom of 35-mm culture dishes. Otherwise, cells were
plated on polyornithine-coated plastic culture dishes. Cells were used
in experiments 1-8 h after plating.
Patch clamp recording
Recordings were made with both the standard whole cell
patch-clamp configuration (Hamill et al. 1981) and
perforated patch technique (Korn and Horn 1989
). Patch
pipettes were fabricated from N51A glass (Garner Glass, Claremont, CA),
coated with silicone elastomer (Sylgard 184, Dow Corning, Midland, MI)
and fire-polished. Pipette resistance varied from 0.5 to 2.0 M
.
Capacitive transients were neutralized electronically, and series
resistance compensation was used at 80-90% (Dagan 3911A patch-clamp
amplifier, Dagan, Minneapolis, MN, or Axopatch 1D, Axon Instruments,
Foster City, CA). In whole cell recordings, series resistance ranged
from 1-4 M
(2.3 ± 0.1, mean ± SE; n = 82). Series resistance in perforated patch experiments ranged from 8.8 to 19.8 M
(13.5 ± 1.2, n = 8). Membrane
currents were filtered at 2 kHz (internal patch-clamp filter) and
digitized at sample intervals of 100-400 µs. Unless otherwise
stated, the holding potential was
80 mV, and Ca2+
currents were evoked by a 100-ms depolarizing stimulus once every 6-10
s. Experiments were performed at room temperature (20-24°C). Data
were acquired and measured with pClamp 6 (Axon Instruments).
Photometry
Intracellular pH was measured in cells loaded with BCECF. To load cells with dye, cells were incubated at 37°C for 10 min with 2.5 µM of the membrane permeant, BCECF-AM (Molecular Probes, Eugene OR, or Texas Fluorescence Laboratories, Austin, TX). Cells then were washed with media and used immediately. When whole cell patch-clamp experiments were combined with photometry, 10 µM of the acid form of BCECF was added to the pipette solution. This prevented the loss of intracellular BCECF during the course of a patch clamp experiment. Electrophysiological experiments were begun after the fluorescence intensity stabilized (stabilization took 1-2 min).
Cells were excited with light from a 150 W Xenon lamp (cut by 60-90% with a neutral density filter), alternately passed through a 450- or 490-nm filter (Omega Optical, Brattleboro, VT) and then through a liquid light guide with quartz-collecting lenses on either end. The light then passed through a 515-nm dicroic mirror. Emitted light passed through a 535-nm barrier filter through an aperture to a photometer. The aperture was set to be as close to the diameter of the cell as possible. The analog voltage output from the photometer was routed into an A/D converter for acquisition by pClamp. In this way, electrophysiological and photometric signals were collected simultaneously.
Photometry and electrophysiology were under the control of a user-written program. The protocols used for simultaneous recording of photometry and electrophysiology data are illustrated in Figs. 5 and 6. Briefly, every trace measured a 450- and a 490-nm signal before the voltage-clamp stimulus and a 490-nm signal during and after the voltage-clamp stimulus. There was no change in 490-nm signal during the depolarizing command. Calibration of the 490/450 ratio for pH is described later. In several experiments, the order of the 450- and 490-nm excitation was alternated so that the 450 nm measurement was made during the stimulus. The results obtained were identical.
For photometry experiments on intact cells (Fig.
1), cells were exposed to excitation
light for 60 ms at each wavelength with a delay of 60 ms between each
excitation. A ratio measurement was collected every 5 s for 15
min. Cells that showed measurable photobleaching were rare and were
discarded.
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BCECF calibration
Intracellular pH was calibrated for BCECF with the nigericin
method (Boyarski et al. 1988). Briefly, cells loaded
with BCECF were exposed to high-potassium bathing solutions (which
contained, in mM, 150 KCl, 15 N-methylglucamine Cl, 5 MgCl2, 10 glucose, and 10 HEPES or MES buffer; pH 7.3, osmolality = 325 ± 5 mosm/kg) of different pH in the
presence of 10 µM nigericin. In the presence of nigericin,
intracellular and extracellular pH equilibrated within 1-2 min. After
equilibration, emission intensity was measured after excitation at 450 and 490 nm in five cells at each pH, and the average plotted as a
function of pH (Fig. 1A). The ratio was normalized according
to Eq. 1 (Boyarski et al. 1988
)
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(1) |
Whereas BCECF was calibrated in "intact" cells, pH measurements usually were recorded in cells loaded with 10 mM HEPES and, in some cases, high concentrations of bis-(o-aminophenoxy)-N,N,N',N'-tetraacetic acid (BAPTA). With internal and external solutions set to pH 7.30, the following pHi was calculated from BCECF measurements in resting patch-clamped cells: 7.30 ± 0.03 (n = 8) and 7.30 ± 0.03 (n = 14) in perforated patch experiments with and without intracellular BAPTA, respectively; 7.32 ± 0.04 (n = 6) and 7.20 ± 0.02 (n = 30) in whole cell experiments with and without intracellular NH4+. Thus neither HEPES nor BAPTA appeared to influence the accuracy of the calibration.
Solutions
Recordings were made from cells plated in 35-mm Nunc
tissue culture dishes containing 1.5-2.0 ml of either static or
flowing bathing solution. In static bath solutions (Figs. 3 and 4), the solution bathing the cells was changed by manually lowering a large-bore pipette that contained the desired test solution near the
cell under study. Application of test solution was terminated by
removal of the large-bore pipette from the bath. In continuous flow
experiments (Figs. 2, 5-8), six separate
solution wells were fed via tygon tubing into a single quartz tip (100 µM diam) that was placed near the recorded cell at the start of the
experiment. One well was filled with control bath solution, and flow
was started immediately after break-in. The solution bathing the cell
was changed by manually turning valves to start and stop flow in each of the six tubes. Solutions bathing the cells were exchanged within 5-10 s using this technique. In most experiments, the control external
solution bathing the cell contained (in mM): 110 NaCl, 30 tetraethylammonium (TEA)-Cl, 20 N-methylglucamine (NMG)-Cl, 2 CaCl2 or BaCl2, 1 MgCl2, 20 glucose, 10 HEPES, and 0.001 tetrodotoxin, pH 7.3 (NaOH),
osmolality = 320 ± 5 mosm/kg. In the standard whole cell
configuration, the pipette solution usually contained (in mM): 150 CsCl, 10 EGTA-Cs, 10 HEPES, 4 MgCl2, 4 creatine phosphate, 4 ATP-Na, and 0.2 mM GTP-Na, leupeptin and creatine kinase, pH 7.3 (CsOH), osmolality = 305 ± 5 mosm/kg. In perforated-patch recordings, the pipette solution contained (in mM): 55 CsCl, 75 Cs2SO4, 8 MgCl2, and 10 HEPES, pH
7.3 (CsOH), osmolality 285 ± 5 mosm/kg. Substitutions are listed
in the figure legends. In some perforated patch experiments, cells were
loaded with BAPTA. To accomplish this, 40 min before recording, cells
were incubated for 30 min with media containing 100 µM BAPTA-AM
(Molecular Probes) in the 37°C incubator. The media containing BAPTA
was replaced with BAPTA-free media and cells incubated for an
additional 10 min. This procedure completely eliminated both
intracellular Ca2+ elevation as measured by Fura-2, and
activation of Ca2+-dependent Cl currents,
after Ca2+ channel activation (data not shown). Except for
the solution in the tip of the electrode, the pipette solution in
perforated-patch experiments contained 16 µg/ml nystatin.
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Two different methods were used to manipulate intracellular pH. In some
experiments, intracellular pH was alkalinized by equimolar substitution
of 20 mM NH4+ (Cl salt) for extracellular
NMG+ (Figs. 3-6)
(Boron and De Weer 1976
). In other experiments,
intracellular pH was controlled by establishing a dual
NH4+ equilibrium, with 20 mM intracellular
NH4+ and varying concentrations of extracellular
NH4+ (Figs. 7 and 8) (Grinstein et al.
1994
). This technique is described in more detail in
RESULTS section.
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Data analysis
All curve fitting and statistics were done with SigmaPlot 2.0 for Windows (Jandel Scientific, Corte Madera, CA). Error bars in plots represent the standard error of the mean. Statistical significance was tested by unpaired Student's t-test except in Fig. 8D, which used a paired Student's t-test.
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RESULTS |
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Ca2+ channel subtype
Although chick DRGs contain predominantly N-type Ca2+
channels (Cox and Dunlap 1994), they can express L-type
channels depending on age and time in culture (Cox and Dunlap
1992
, 1994
). Because L-type Ca2+ channels are
sensitive to changes in pHi, it was important to determine
whether our cell population contained a fraction of L-type
Ca2+ channels.
We characterized the subtype of Ca2+ channel(s) in our
cells by voltage-dependent inactivation and pharmacological
sensitivity. To examine inactivation, cells were held at progressively
more depolarized potentials for 10 s followed by a test stimulus
to 0 mV (Fig. 2A). In all cells tested, Ca2+
currents completely inactivated between 10 and 0 mV, with the half-maximal inactivation at
63.7 ± 1.7 mV (Fig. 2B;
n = 8).
We then tested the -conotoxin GVIA sensitivity and dihydropyridine
sensitivity. In 14 cells tested, 10 µM conotoxin irreversibly blocked
the Ca2+ current by 94.4 ± 2.7%. Ten of these 14 cells were blocked by 100% by 10 µM conotoxin (Fig. 2, C
and D). The mean block in the other four cells was 80.6 ± 4.3%. Importantly, 100% current block was achieved in cells of all
sizes (Fig. 2D). One µM conotoxin also irreversibly
blocked Ca2+ currents by 100% in four of four cells
tested. Consistent with these results, the dihydropyridine nimodipine
(1 µM) had no effect on Ca2+ currents in six cells
tested. Thus as previously described for acutely isolated chick DRGs
taken from 11- to 12-day-old embryos (Cox and Dunlap
1994
), cells of all sizes contained either entirely or almost
entirely
-conotoxin GVIA-sensitive Ca2+ channels.
Potentiation of N-type Ca2+ currents by extracellular NH4+
Application of 20 mM NH4+ reversibly potentiated voltage-activated Ca2+ currents (Fig. 3A). There was no shift in the voltage dependence of activation and little or no change in reversal potential (Fig. 3B).
Extracellular application of NH4+ is a standard technique for elevation of intracellular pH. Because NH4+ permeates through K+ channels, however, it was possible that this increase in voltage-activated inward current reflected the addition of an inward current carried by NH4+ to the Ca2+ current. The experiment in Fig. 4 demonstrates that this was not the case. The data in Fig. 4 were collected under identical conditions as that in Fig. 3, except that TEA was eliminated from the bath solution (equimolar replacement with Na+). As in Fig. 3, application of NH4+ resulted in an increase in the inward current (Fig. 3A). Also evident in this figure is the presence of a slow inward tail current (Fig. 4A, arrow) during NH4+ application. Immediately after removal of NH4+, the slow tail current disappeared and the inward current during the depolarization was reduced slightly. This tail current was blocked by extracellular application of 30 mM TEA, consistent with the tail current representing NH4+ flux through K+ channels. We interpret these data to mean that, in the absence of TEA, the slow tail and perhaps a fraction of the step current during NH4+ application was carried by NH4+. However, the complete disappearance of tail current at a time when inward step current still was potentiated markedly indicates that the dominant effect of NH4+ application was potentiation of the voltage-activated Ca2+ current. This interpretation is supported by the plots in Fig. 4B. The top plot shows that the inward tail current was present during application of NH4+ and disappeared immediately on termination of NH4+ application. This is consistent with what is expected with application and removal of an ion that permeates a voltage-gated channel. In contrast, Fig. 4B, bottom, shows that the inward current during the step grew slowly on NH4+ application and decayed back to the control level slowly after removal of NH4+. In all other experiments, the bathing solution contained 30 mM TEA to minimize the contribution of an NH4+ current through K+ channels to measured currents.
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pH dependence of change in Ca2+ current magnitude
We used two approaches to examine the pHi dependence and sensitivity of Ca2+ channel current magnitude. In the first set of experiments (Figs. 5 and 6), we simultaneously measured pHi and Ca2+ current magnitude from cells loaded with the pH-sensitive dye, BCECF. In these experiments, 20 mM NH4+ was applied externally to alkalinize the cells.
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Figure 5 shows results from a cell also preloaded with BAPTA, recorded with the perforated patch configuration. Figure 5A illustrates digitized light intensity measurements obtained at 450 and 490 nm before, during, and after application of NH4+. The internal pH in these three conditions, calculated from the calibration in Fig. 1, was 7.02, 7.78, and 7.23, respectively. Figure 5B illustrates Ca2+ currents recorded simultaneously with the photometric output in Fig. 5A. As observed previously, Ca2+ currents were potentiated reversibly by NH4+ application. The association of changes in pHi and Ca2+ current magnitude can be compared in Fig. 5, C and D. On application of NH4+, intracellular pH rose from 7.0 and 7.7, and there was a corresponding increase in Ca2+ current magnitude. On removal of external NH4+, pHi fell and Ca2+ current magnitude was reduced. The change in Ca2+ current magnitude routinely lagged behind the measured change in pHi during the on-phase of NH4+ application, making precise correlation of pHi and Ca2+ current magnitude difficult (this lag will be addressed in DISCUSSION). Note, however, that pHi stabilized at four different plateau levels, and with each increment of alkalinization, which was as little as 0.2 pH units, Ca2+ current magnitude was increased. More detailed analysis of these data are presented in the following text.
Figure 6A illustrates
simultaneous recordings of pHi (top) and
Ca2+ channel currents (bottom) from a standard
(ruptured patch) whole cell recording. As described earlier,
application of NH4+ rapidly alkalinized the cell
interior to pH 7.5, and the Ca2+ current was potentiated.
Similar to the results observed in Fig. 5, C and
D, the change in Ca2+ current magnitude more
closely paralleled the change in pHi during the off-phase
of NH4+ application. Consequently, we plotted
Ca2+ current magnitude as a function of pHi,
measured after removal of NH4+ (Fig. 6B).
illustrate data from the cell in Fig. 6A;
plot the
pH dependence of Ca2+ currents from the cell described in
Fig. 5. Data were recorded once every 6 s, so the decline from pH
7.5 to 7.3 took ~90 s. These data indicate that Ca2+
current magnitude is responsive to pHi changes between pH
7.0 and 7.5 and that prevention of intracellular Ca2+
elevation with BAPTA had no apparent effect on pH sensitivity (NH4+-induced potentiation also was observed in 6 whole
cell recordings that included 5 mM intracellular BAPTA; data not
shown). Of particular interest from these experiments though, is the
observation that Ca2+ currents were sensitive to very small
changes between pH 7.3 and 7.5.
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Control of intracellular pH by a dual-NH4+ equilibrium technique
Manipulation of pHi solely by extracellular
NH4+ posed three problems. First, the possibility of a
long-lasting influence of NH4+ itself on
Ca2+ current magnitude could not be ruled out from these
experiments. Second, as described in Figs. 4-6, the change in
Ca2+ current magnitude lagged behind the change in
pHi, especially during the on-phase of NH4+
application. This effect, which may have been due at least partially to
the pHi buffering effect of BCECF, precluded an accurate
quantification of pH dependence of Ca2+ current magnitude.
Third, pHi could be measured and manipulated, but not
quantitatively controlled, with this technique. To ameliorate these
problems, we used a dual NH4+ equilibrium technique, in
which NH4+ is included in the intracellular solution
and varied in the external solution (Fig.
7) (Grinstein et al.
1994). With NH4+ in the pipette solution and no
external NH4+, cells are acidified by the exit of
NH3 across the cell membrane (Fig. 7A, top).
With NH4+ in both the internal and external solution,
the internal proton concentration is determined by the relative
concentrations of internal and external NH4+ (Fig.
7A, bottom). In contrast to techniques that use just
internal or external NH4+, this allows for precise,
reproducible control of pHi. This is illustrated in Fig.
7B. In patch-clamp recordings that included 20 mM
NH4+ in the pipette solution, application of different
external [NH4+] resulted in rapid, controlled changes
in pHi (Fig. 7B). Note that application of 40 mM
NH4+ did not produce the expected alkalinization of
pHi and produced a significant reduction in
Ca2+ current magnitude (not shown), so this technique was
not used to alkalinize cells in the experiments below. The plot in Fig. 7C illustrates the results of NH4+
application to six cells recorded in the whole cell patch-clamp configuration. Cells were held at
80 mV throughout, and no
depolarizing stimuli were applied. At external [NH4+]
of 0, 5, and 20 mM, pHi attained values of 6.6, 7.0, and
7.3, respectively.
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Figure 8 illustrates the pH sensitivity of Ba2+ currents using the dual NH4+ equilibrium technique. The intracellular solution contained 20 mM NH4+. Figure 8A illustrates currents activated by depolarization to 0 mV in the presence of 20 mM external NH4+ (control and recovery traces are shown) and in the presence of 5 mM NH4+. The complete current-voltage relationship from the same cell (Fig. 8B) illustrates that Ba2+ current was reduced by acidification with no accompanying change in reversal potential or voltage sensitivity. At 0 mV, Ba2+ current magnitude was reduced by 40% on switching from 20 to 5 mM NH4+ (Fig. 8D). Figure 8C illustrates four superimposed currents, recorded alternately, in the presence of 20 or 0 mM external NH4+. Currents recorded in the presence of 0 NH4+ were reduced in magnitude by 65% compared with those recorded in the presence of 20 mM NH4+ (Fig. 8D). These results demonstrate that Ca2+ channel currents are extremely sensitive to pHi changes between 7.0 and 7.3 (the physiological range) and that acidification below pH 7.0 results in further reduction in Ba2+ current magnitude. Ba2+ current responses to changes in pHi were readily reversible, which suggests that acidification to values as low as pH 6.6 did not damage the Ca2+ channel, but rather, that changes in pHi modified channel function. Finally, because NH4+ always was present in these experiments, these results demonstrate that it was the change in pHi, not the presence of NH4+, that modulated Ca2+ current magnitude.
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DISCUSSION |
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Ca2+ influx via voltage-gated Ca2+
channels, glutamate application, GABA application, and hypoxia all
produce intracellular acidification by 0.15-0.5 U (Ahmed and
Connor 1980; Canzoniero et al. 1996
; Dixon et al. 1993
; Hartley and Dubinsky
1993
; Kaila et al. 1990
, 1993
; Luckermann
et al. 1997
; Trapp et al. 1996
; Wang et
al. 1994
; Werth and Thayer 1994
). Subsequent
recovery from acidification is relatively slow and depends largely on
the ability of H+ transporters to remove H+
from the cytoplasm, either by extrusion or sequestration into mitochondria. Consequently, sensitivity of physiological processes to
variation in pHi between pH 6.8 and 7.4 may provide an
important mechanism for modulation of neuronal activity. The main
finding of our experiments is that N-type Ca2+ channels are
extremely sensitive to small changes in pHi near physiological pH values. Acidification from pH 7.3 to 7.0 resulted in a
40% decrease in Ca2+ current magnitude, and acidification
from pH 7.3 to 6.6 resulted in greater than a 65% decrease in
Ca2+ current magnitude. Conversely, alkalinization from pH
7.3 to 7.5 potentiated Ca2+ currents by ~40%.
Comparison of pH sensitivity of L- and N-type Ca2+ channels
The pHi-sensitivity of N-type Ca2+
channels in chick DRGs is nearly identical to that of L-type
Ca2+ channels in catfish horizontal cells (Dixon et
al. 1993). Both L-type Ca2+ currents (Dixon
et al. 1993
) and N-type Ca2+ currents (Fig. 8) were
reduced by ~40-45% when pHi was reduced from pH 7.3 to
7.0 and potentiated by 40-45% with alkalinization from pH 7.3 to 7.5 (Fig. 6). In rat hippocampal neurons, N-type Ca2+ channel
currents appeared to be more sensitive to application of
NH4+ than L-type Ca2+ channel currents
(Tombaugh and Somjen 1997
). In these studies, however,
pHi was not measured. In whole cell patch-clamp
experiments, with the pipette solution buffered to pH 7.3 with 10 mM
HEPES, pHi can vary between 6.9 and 7.3 from cell to cell
and over time (unpublished data). Consequently, it is possible that
under the different conditions necessary to isolate N- and L-type
currents in hippocampal neurons (Tombaugh and Somjen
1997
), resting pHi varied and/or the
pHi values traversed on application of NH4+
differed. Alternatively, the apparently different channel sensitivities to pHi in hippocampal neurons indeed may reflect a
difference between mammalian and nonmammalian channel sensitivity to
pHi or different mechanisms of pH sensitivity in different
cell types. Resolution of this discrepancy awaits precise quantitative
examination of the pH dependence of different Ca2+ channel
subtypes from different cells.
Time lag between pH change and Ca2+ current response
In all of our experiments, there was a delay between the measured
pHi change and Ca2+ current response (see Fig.
5 for example). This delay was not entirely due to the presence of
intracellular BCECF or HEPES, both H+ buffers, because the
lag occurred in the absence of these chemicals (Figs. 4 and 5). We
imagine one of three possible reasons for the delay. First, the
pHi-induced modulation may have been produced indirectly by
protonation of another Ca2+ channel modulator rather than
by a direct protonation of the Ca2+ channel. Second, the
pHi-induced modulation may have resulted from protonation
of a site in the Ca2+ channel pore near the outer vestibule
(Chen et al. 1996). Whereas H+-induced
modulation of this externally located site usually is studied via
extracellular pH changes (cf. Chen et al. 1996
;
Prod'hom et al. 1989
), it is possible that
intracellular protons could make their way to this site on repetitive
Ca2+ channel activation. Finally, this delay may have
represented a lag in pHi change adjacent to the membrane
relative to that measured in the entire cell by BCECF. Elucidation of
the mechanism of pHi-induced modulation of Ca2+
currents must await future studies.
Functional significance
The variety of physiological and pathological stimuli that alter pHi and the observation that Ca2+ channel currents are quite sensitive to small changes in pHi in the physiological range suggest that changes in pHi may play an important role in both physiological and pathological events. Some potential roles are described here.
FEEDBACK INHIBITORY MECHANISM.
Mitochondrial uptake of Ca2+ is an important
Ca2+ removal mechanism under conditions of both moderate
and high intracellular Ca2+ load (Herrington et al.
1996; Meech and Thomas 1980
; Park et al.
1996
; Thayer and Miller 1990
; Wang et al.
1994
). After Ca2+ influx, the H+ pumped
out of the mitochondria during respiration can become a significant
source of intracellular acidification and can produce changes in
pHi of
0.2 U (Werth and Thayer 1994
). Thus
Ca2+-induced acidification after Ca2+ influx
might form a feedback inhibitory control on Ca2+ channel
activity. Given the prominent role of N-type Ca2+ channels
in the secretory process in both neurons and endocrine cells, such a
feedback mechanism may be particularly important in the secretion process.
GABA AS A POTENTIAL MODULATOR OF CA2+
CHANNEL FUNCTION.
Activation of GABAA receptors results in changes in
pHi by flux of HCO3 through
GABA-activated Cl
channels (Kaila et al. 1990
,
1993
; Luckermann et al. 1997
; Voipio et
al. 1991
). Under physiological conditions, GABA can acidify the
cell interior, on average, by as much as 0.25-0.4 U. Thus inhibition
of cell function by GABA may be accomplished, at least in part, by
acidification-induced inhibition of Ca2+ channel function.
Interestingly, this effect may depend on the physiological state of the
cell or organism because membrane potential and pCO2 will
influence the magnitude and even direction of GABA-induced pHi changes without influencing GABA-activated conductance
changes (Luckermann et al. 1997
).
NEURONAL PROTECTION DURING ISCHEMIC CONDITIONS.
Cellular damage from ischemia and reperfusion injury after ischemia are
associated with large increases in intracellular Ca2+ (cf.
Bickler and Hansen 1998; Kristian and Siesjo
1998
; Meissner and Morgan 1995
; Shimazaki
et al. 1998
; Vornov 1998
). In both neurons and
cardiac tissue, maintenance of acidic pHi during recovery from ischemia protects cells from cell death (Bond et al.
1993
; Scholz et al. 1992
; Tombaugh and
Sapolsky 1990
; Vornov et al. 1996
).
Acidification-induced inhibition of Ca2+ channels, under
conditions that might otherwise serve to activate Ca2+
channels, may be one mechanism by which cells are protected from neurotoxic damage.
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ACKNOWLEDGMENTS |
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We thank L. Polo-Parada for writing the software used to control the photometry experiments and for the calibration in Fig. 1.
This work was supported in part by the National Science Foundation, The Whitaker Foundation, and the University of Connecticut Research Foundation.
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FOOTNOTES |
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Address for reprint requests: S. Korn, Dept. of Physiology and Neurobiology, University of Connecticut, Box U-156, 3107 Horsebarn Hill Rd., Storrs, CT 06269.
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.
Received 20 August 1998; accepted in final form 2 December 1998.
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REFERENCES |
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