Department of Biology, Ripon College, Ripon, WI 54971, USA
* Author for correspondence (e-mail: lightd{at}ripon.edu)
Accepted 27 September 2002
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Summary |
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Key words: Fluo-4, EGTA, BAPTA, Cell volume regulation, Hexokinase, P2 receptor
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Introduction |
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Exposure of vertebrate cells to a hypotonic solution results in an initial
increase in cell volume owing to the relatively rapid influx of water. During
continuous hypotonic stress, increases in cell volume are followed by a
slower, spontaneous recovery towards the pre-shock level, a process known as
regulatory volume decrease (RVD). This recovery is accomplished by selectively
increasing the permeability of the plasma membrane during cell swelling to
allow for efflux of specific intracellular osmolytes, thereby reversing the
driving force for water influx (Grinstein
and Foskett, 1990; Hoffman,
2000
; Lewis and Donaldson,
1990
; McCarty and O'Neil,
1992
; Strange,
1994
). Most vertebrate cells lose K+ and Cl-
during RVD (Grinstein and Foskett,
1990
; Hazama and Okada,
1990
; Hoffman,
2000
; Holtzman,
1991
; Lewis and Donaldson,
1990
; McCarty and O'Neil,
1992
; Strange,
1994
). This may occur by electroneutral ion transport pathways
(Lewis and Donaldson, 1990
) or
by the separate activation of K+ and Cl- channels
(Bergeron et al., 1996
;
Grinstein and Foskett, 1990
;
Hoffman et al., 1986
;
Lewis and Donaldson, 1990
;
Rubera et al., 1997
). Loss of
organic anions and osmolytes also may occur during RVD
(Kirk and Strange, 1998
).
The cellular mechanisms that activate and regulate permeability pathways
during RVD are not completely understood and appear to differ between cell
types (Land et al., 1998;
Lewis and Donaldson, 1990
;
McCarty and O'Neil, 1992
;
Strange, 1994
). Calcium,
however, is a pivotal signaling agent for a wide variety of physiological
processes. An elevation in the concentration of intracellular free
Ca2+ from a resting level of approximately 50-100 nM to a
stimulated level of 1-10 µM occurs as a rapid response to a number of
cellular stimuli, including growth factors, neurotransmitters, hormones,
peptides, toxins and cell swelling (Cheek
et al., 1993
).
Calcium has been shown to play a key role during cell volume regulation in
a number of cell types (Foskett,
1994; Hoffman,
2000
; McCarty and O'Neil,
1992
; Tinel et al.,
2000
). In some instances, RVD strictly depends on Ca2+
influx across the plasma membrane
(Montrose-Rafizadeh and Guggino,
1991
; Wong et al.,
1990
), whereas in other cell types the Ca2+ response is
mediated by Ca2+ release from intracellular stores
(Negulescu et al., 1992
;
Terreros and Kanli, 1992
). In
addition, although it has been suggested that Ca2+ can directly
activate ion channels during RVD (Hazama
and Okada, 1990
; McCarty and
O'Neil, 1992
), there also is evidence that several
Ca2+-dependent intracellular messengers and enzymes (e.g.,
calmodulin, phospholipase A2, eicosanoids and protein kinases) are
involved with cell volume regulation
(Hoffman, 2000
;
McCarty and O'Neil, 1992
;
Rubera et al., 1997
;
Strange, 1994
;
Tinel et al., 2000
).
Furthermore, the cytoskeleton is thought to play a role in volume regulation
and actin polymerization is dependent on intracellular Ca2+ levels
(Hoffman, 2000
;
Mills et al., 1994
).
Recent studies in our laboratory indicate that RVD in Necturus
erythrocytes depends on a quinine-inhibitable K+ conductance that
is regulated during cell swelling by a calmodulin-dependent mechanism
(Bergeron et al., 1996) and by
a 5-lipoxygenase metabolite of arachidonic acid
(Light et al., 1997
), as well
as extracellular ATP activation of P2 receptors
(Light et al., 1999
;
Light et al., 2001
). Because
calmodulin, phospholipase A2 and 5-lipoxygenase are Ca2+
sensitive (Holtzman, 1991
) and
because P2X receptors are ligand-gated, Ca2+-permeable cation
channels (Fredholm et al.,
1994
), the purpose of this study was to investigate a more
definitive role for Ca2+ in the regulation of cell volume following
hypotonic challenge. To this end, we used four different approaches: (1)
hemolysis studies to examine osmotic fragility; (2) fluorescence microscopy to
detect changes in intracellular Ca2+ levels; (3) a Coulter counter
to measure the volume of osmotically stressed cells; and (4) the whole-cell
patch clamp technique to study membrane currents.
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Materials and Methods |
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Osmotic fragility
Osmotic fragility was examined by determining the degree of cell lysis for
a suspension of RBCs in hypotonic Ringer. The level of hemolysis was
determined via a turbidity shift (cloudy to clear) that occurs when the
integrity of the plasma membrane is compromised. This was detected with a
spectrophotometer (Spectronic 20D, Milton Roy Co.) 10, 15 or 20 minutes after
blood (30-50 µl) was added to saline solutions (3 ml) of different
osmolalities and compositions. Spectrophotometric experiments were conducted
at 625 nm because this wavelength provided the greatest difference in optical
density (OD) between intact and lysed cells
(Bergeron et al., 1996).
A percent hemolytic index (HI) was determined using the formula: HI(%)=(OD of test compoundOD of negative control)/(OD of positive controlOD of negative control) x 100, where OD of test compound refers to the OD of a cell suspension in diluted Ringer to which a test compound was added, OD of negative control refers to the OD of a cell suspension in diluted Ringer and OD of positive control refers to the OD of a cell suspension in distilled water. All reported hemolytic indices were calculated using a concentration of Ringer that gave an OD reading between 0.025 and 0.040. We chose this concentration range because it was sufficiently dilute to lyse approximately half the cells in suspension. Consequently, we could assess whether a test compound increased osmotic fragility by a subsequent reduction in OD compared to the negative control solution. Conversely, a rise in OD indicated that a test compound decreased osmotic fragility.
Fluorescence microscopy
Intracellular free Ca2+ levels were monitored using a Nikon
diaphot microscope, equipped with Hoffman DIC optics (400x) and
epifluorescence (mercury lamp and FITC filter cube), and the fluorescent
Ca2+ indicator fluo-4-AM (10 µM, Molecular Probes, Eugene, OR).
This indicator has a high-affinity binding for Ca2+
(Kd=345 nM) and a very large fluorescence intensity increase in
response to Ca2+ binding (>100 fold). The acetoxymethyl (AM)
ester derivative permeates cell membranes and, once inside a cell, the
lipophilic blocking groups are cleaved by non-specific esterases. This results
in a charged form that is relatively impermeable.
Aliquots of fluo-4-AM were mixed with DMSO and diluted to give a final concentration of 10 µM. The non-ionic detergent Pluronic F-127 was used to assist in dispersion of the non-polar AM ester in aqueous media. This was accomplished by mixing an aliquot of AM ester stock solution in DMSO with an equal volume of 20% (w/v) Pluronic in DMSO before dilution into the loading medium. Cells were incubated with the AM ester for 60-90 minutes at room temperature [incubation at this temperature helps reduce potential compartmentalization of dye within cells (Molecular Probes). However, we did not test for compartmentalization and therefore cannot rule out this possibility]. Cells were then washed in indicator-free medium to remove any dye that was not specifically associated with the cell surface and then incubated for another 30-60 minutes to allow for complete de-esterification of intracellular AM esters.
Our experimental approach was designed to provide a qualitative assessment of fluorescence intensity while still maintaining identical imaging parameters for all conditions. This was accomplished by taking photographs with a Nikon camera (N2000) mounted directly to the microscope and using the same exposure time (3 seconds) and film speed (ASA 400) for all pictures. All photographs of swollen cells were taken 5 minutes after hypotonic shock, a point at which cells were maximally swollen.
Coulter counter
Cell volume distribution curves were obtained by electronic sizing using a
Coulter counter model Z2 with channelyzer (Coulter Electronics, Hialeah, FL).
Mean cell volume was taken as the mean volume of the distribution curves. The
diameter of the aperture tube orifice was 200 µm and the metered volume was
0.5 ml. Absolute cell volumes were obtained using polystyrene latex beads
(20.13 µM diameter or 4.271x103 fl volume) as standards
(Coulter). Experiments with the latex beads showed that measured volumes were
unaffected by changes in osmolality and ionic composition within the ranges
used for this study. Cell suspensions were diluted to give a final cell
density of approximately 5,000-7,000 cells per ml.
Relative cell volume is defined as the average volume of cells compared to
that in an isotonic medium. As described by others
(Jorgensen et al., 1997;
Wang et al., 1996
), a percent
volume recovery at X minutes after hypotonic exposure was calculated as
[(VmaxVX
min)/(VmaxV0)]x100, where
Vmax is the peak relative cell volume, V0 is the initial
relative volume (or one) and VX min is the relative cell volume
measured X minutes after hypotonic exposure. A percent volume decrease was
calculated as [(percent recoveryexperimental)/(percent
recoverycontrol)]x100, where maximal recovery in hypotonic
Ringer is 100%.
Patch clamp
Patch pipettes were fabricated from Kovar sealing glass (Corning model
7052, 1.50 mm OD, 1.10 mm ID; Garner Glass, Claremont, CA) using a two-pull
method (Narishige PP-7). Pipette tips were fire polished (Narishige MF-9) to
give a direct current resistance of approximately 5-8 M in symmetrical
100 mM KCl solutions (the large size of Necturus RBCs allows for the
use of relatively wide tip pipettes). All pipette solutions were filtered
immediately before use with a 0.22 µm membrane filter (Millex-GS, Bedford,
MA), and the pipettes were held in a polycarbonate holder (E. W. Wright,
Guilford, CT). Membrane currents were measured with a 1010
feedback resistor in a headstage (CV-201A, Axon Instruments, Foster City, CA)
with a variable gain amplifier set at 1 mV/pA (Axopatch 200A, Axon
Instruments, Foster City, CA). The current signals were filtered at 1 kHz
through a four-pole low-pass Bessel filter and digitized at 5 kHz with an
IBM-486 computer.
Acquisition and analysis of data were conducted with P-Clamp® (version 6, Axon Instruments, Foster City, CA). Data were acquired during 100 msecond voltage pulses, and the command potential was set to -15 mV (close to the resting potential for RBCs) for 100 msec between each pulse. All voltage measurements refer to the cell interior.
RBCs, attached to glass coverslips (5 mm diameter, Bellco Biotech.,
Vineland, NJ) with poly-D-lysine (150,000-300,000; 1 mg/ml), were placed in a
specially designed open-style chamber (250 µl volume, Warner Instruments
Corp., Hamden, CT). The bath solution could be changed by a six-way rotary
valve (Rheodyne Inc., Cotati, CA). The whole-cell configuration was achieved
following formation of a gigaohm seal (cell-attached configuration) by
applying suction to disrupt the patch of membrane beneath the pipette or by
applying a large voltage (>200 mV) to the patch. A sudden increase in the
capacitance current transient accompanied disruption of the membrane. (N.B.,
although the perforated patch method is preferable when trying to maintain
normal intracellular Ca2+ levels, we were not successful at
obtaining stable G seals with either nystatin or amphotericin B in
patch pipettes. Nonetheless, we were able detect a response that was
internally consistent with our other three experimental protocols when we
manipulated extracellular calcium levels using the standard whole-cell
technique.)
Solutions
Amphibian Ringer solution consisted of (in mM) 110 NaCl, 2.5 KCl, 1.8
CaCl2, 0.5 MgCl2, 5 glucose and 10 HEPES (titrated to pH
7.4 with NaOH, 235 mosm/kg H2O). A `low Na+ Ringer' was
prepared by substituting choline chloride for NaCl (used for all experiments
with gramicidin), and a 0.5x Ringer was obtained by reducing NaCl
accordingly. A `low Ca2+ Ringer' was obtained by adding 5 mM
ethylene glycol-bis(2-aminoethylether)-N,N,N',N'-tetraacetic acid
(EGTA) to amphibian Ringer, which reduced the free Ca2+
concentration to a calculated level of 35 nM (MaxChelator:
http://www.standford.edu/cpatton/).
Cells were loaded with
1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid
tetra(acetoxymethyl) ester (BAPTA-AM) in a manner similar to that described
above for fluo-4-AM. A stock solution of gramicidin was dissolved in methanol,
and a stock solution of A23187 was prepared in DMSO, both at 1000x the
final concentration and then diluted to give an appropriate working
concentration. All stock aqueous solutions were diluted 100x to give an
appropriate final concentration.
Patch pipettes were filled with an intracellular Ringer solution containing (in mM) 100 KCl, 3.5 NaCl, 1.0 MgCl2, 1.0 CaCl2, 2.0 EGTA, 5 glucose, 1.0 Mg-ATP, 0.5 GTP and 5.0 HEPES (pH 7.4 with KOH, 235 mosm/kg/H2O, calculated free Ca2+ level of 60 nM). During seal formation, the extracellular solution contained (in mM) 110 NaCl, 2.5 KCl, 1.8 CaCl2, 0.5 MgCl2, 5 glucose and 10.0 HEPES (pH 7.4). A hypotonic (0.5x) high K+ bath contained (in mM) 2.5 NaCl, 50 KCl, 1.8 CaCl2, 0.5 MgCl2, 5 glucose and 10.0 HEPES (pH 7.4, 120 mosm/kg/H2O).
For hemolysis experiments, pharmacological agents or their vehicle were present prior to the addition of cells. For cell volume studies, pharmacological agents were added with hypotonic exposure (0 minutes) or at peak cell volume (5 minutes after hypotonic challenge). Osmolality of solutions was measured with a vapor pressure osmometer (#5500, Wescor, Logan, UT). Chemicals were purchased from Sigma Chemical Co. (St. Louis, MO), Alexis Biochemicals (San Diego, CA), and ICN Pharmaceuticals Inc. (Costa Mesa, CA). All experiments were conducted at room temperature (21-23°C).
Statistics
Data are reported as means±s.e.m. The statistical significance of an
experimental procedure was determined by a paired Student's t-test or
least significant difference test with paired design of analysis of variance
(ANOVA)/multivariate ANOVA (MANOVA), as appropriate (Data Desk® software,
Ithaca, NY). A P<0.05 was considered significant. Each animal
served as its own control, and cell volumes at specific times were tested
against each other. For patch clamp studies, each cell served as its own
control.
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Results |
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To determine whether osmotic fragility depended on extracellular Ca2+, we repeated this assay with a `low Ca2+ Ringer' (amphibian Ringer solution containing 5 mM EGTA). In this case, the OD measured at the same level of dilution as the control (resulting in an extracellular Ca2+ concentration of <10 nM) was 0.011±0.002 (n=10, P<0.001, Fig. 1), which gave a hemolytic index of 86%. By contrast, addition of the cationophore gramicidin (5 µM) to a diluted `low Ca2+ choline Ringer solution' blocked the inhibitory effect of bathing cells in low Ca2+ (n=10, Fig. 1; gramicidin is a cationophore that was used to maintain a high K+ permeability).
By contrast, we also conducted experiments that raised intracellular Ca2+ levels. Addition of the Ca2+ ionophore A23187 (0.5 µM) to the original diluted amphibian Ringer (230 µM extracellular Ca2+) increased the OD from 0.062±0.006 to 0.083±0.006, indicating a 34% decrease in lysed cells compared to the control (n=10, P<0.01, Fig. 1).
We next examined the effect of blocking P2 receptor activation. This was
accomplished by hydrolyzing extracellular ATP with hexokinase (2.5 U/ml), an
enzyme that traps ATP by transferring its -phosphoryl group to a
variety of C6 sugars
(Schwiebert et al., 1995
). In
this case, the OD decreased from 0.071±0.007 to 0.042±0.006
(n=10, P<0.01, Fig.
2), giving a hemolytic index of 41%. As illustrated in
Fig. 2, the presence of
gramicidin (5 µM) prevented the inhibitory effect of hexokinase
(n=10). In addition, the general P2 receptor antagonist suramin [100
µM, (Fredholm et al., 1994
)]
decreased OD from 0.075±0.002 to 0.047±0.003 (n=10,
P<0.001, Fig. 2),
giving a hemolytic index of 37%. As with hexokinase, the presence of
gramicidin (5 µM) prevented the inhibitory effect of suramin
(n=10). In addition, A23187 (0.5 µM) also prevented suramin from
increasing osmotic fragility (n=10). (N.B., variation in the average
OD between control groups resulted from adding different volumes of blood for
some experiments; volumes ranged from 30-50 µl). However, the same volume
of blood per volume of extracellular medium was used within an experimental
group.
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Fluorescence microscopy studies
Having established that osmotic fragility was dependent on extracellular
Ca2+ and activation of P2 receptors, we next determined whether
intracellular Ca2+ levels changed during hypotonic challenge. As
shown in Fig. 3A, cells loaded
with fluo-4-AM (10 µM) did not display fluoresce under isosmotic conditions
(n=6). By contrast, addition of A23187 (0.5 µM) stimulated
fluorescence under isosmotic conditions, indicating cells were properly loaded
with dye and we could detect qualitative changes in the level of intracellular
free Ca2+ (n=6, Fig.
3B).
|
We next examined the effect of hypotonic shock on the level of cytosolic
Ca2+. Exposure of cells to a hypotonic (0.5x) Ringer
increased the level of fluorescence, indicating a rise in intracellular
Ca2+ in swollen cells (n=6,
Fig. 3A versus 3C). By
contrast, exposing cells to a `low Ca2+ hypotonic Ringer' did not
result in an increase in fluorescence associated with hypotonic challenge
(n=4, Fig. 3C versus
3D). Additionally, the presence of hexokinase (2.5 U/ml) or
suramin (100 µM) in hypotonic Ringer prevented an increase in fluorescence
in swollen cells (n=4, Fig.
4). This inhibitory effect was blocked with the addition of A23187
(0.5 µM, n=4, Fig.
4). Further, the stretch-activated channel antagonist gadolinium
(10 µM) (Yang and Sachs,
1989) inhibited the fluorescence associated with hypotonic
swelling (n=4; data not shown). Finally, the calcium channel blockers
verapamil (10 µM, n=4) and nifedipine (10 µM, n=4) had
no effect on fluorescence following hypotonic challenge (data not shown).
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Cell volume studies
When RBCs were placed in a hypotonic amphibian Ringer, they quickly swelled
and then slowly and spontaneously decreased in volume
(Fig. 5). When repeated with
the `low Ca2+ hypotonic Ringer', the rate of volume recovery was
reduced (n=8, P<0.05 after 40 minutes compared to control
values), lowering the percent volume decrease to 57% of control values at 120
minutes. Similarly, when intracellular Ca2+ was buffered with
BAPTA-AM (100 µM), volume recovery also was inhibited (n=8,
P<0.05 after 40 minutes compared to control), reducing the percent
volume decrease to 51% of control values at 120 minutes
(Fig. 5). There was no
significant difference at any time between the `low Ca2+ hypotonic
Ringer' and the BAPTA-AM groups. Further, the inhibitory affect of BAPTA was
prevented by adding gramicidin (5 µM, n=8, ionophore was added 5
minutes after hypotonic shock when cells were maximally swollen,
Fig. 5, P<0.05
within 5 minutes following addition of ionophore compared to the other three
groups).
|
We previously demonstrated that addition of A23187 to the extracellular
solution potentiates volume recovery following hypotonic challenge
(Light et al., 1999). In this
study we assessed whether the effectiveness of pharmacologically increasing
intracellular Ca2+ has a time-dependent nature. This was
accomplished by adding A23187 (0.5 µM) 5, 40 and 70 minutes after hypotonic
shock. In all three cases, this agent quickly reduced cell volume to near
basal levels (n=8, Fig.
6, P<0.05 compared to control values immediately
following addition of ionophore, regardless of when it was applied). In
addition, although gadolinium (10 µM, n=6) inhibits volume
recovery (Light et al., 1999
),
in this study we found both nifedipine (10 µM, n=6) and verapamil
(10 µM, n=6) had no affect.
|
We also have shown in a previous study that the ATP scavenger hexokinase
blocks cell volume recovery following hypotonic shock, and both A23187 and
gramicidin prevent this inhibitory response
(Light et al., 1999). Here we
examined whether inhibition of RVD with hexokinase (2.5 U/ml) has a
time-dependent response by adding it 5, 40 and 70 minutes after cell swelling.
In all three instances, volume recovery was inhibited to virtually the same
extent (n=8, Fig. 7,
P<0.05 compared to the control immediately following addition of
hexokinase at the 5 minute mark, and P<0.05 after 10 and 5 minutes
when hexokinase was added at the 70 and 40 minute marks, respectively). There
was no significant difference between the three hexokinase groups once this
enzyme exhibited an inhibitory effect that was significantly different from
control values.
|
Patch clamp studies
We previously reported that the K+ channel antagonist quinine,
the calmodulin inhibitors pimozide and W-7, the stretch-activated channel
blocker gadolinium and the P2 receptor antagonist suramin each inhibit
whole-cell currents that are activated under hypotonic conditions
(Bergeron et al., 1996;
Light et al., 2001
). Here we
determined the effect of exposing swollen cells to a low Ca2+
hypotonic Ringer solution. For these experiments we used a high KCl Ringer in
the pipette and a 0.5x KCl Ringer in the bath. The only major ions of
significance with these solutions were K+ and Cl-, and
the equilibrium potentials for perfect cation- and anion-selective
conductances were -16.2 mV and +14.7 mV, respectively. Addition of 5 mM EGTA
to the extracellular medium reduced whole-cell conductance by 72%, from
20.1±2.4 nS to 5.6±1.4 nS (n=5, P<0.01,
Fig. 8). However this maneuver
did not significantly shift the reversal potential.
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Discussion |
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Our finding that hypotonic shock increased cytosolic Ca2+ is
consistent with reports for several other cell types. For instance, reducing
the osmolarity of the bathing medium causes intracellular Ca2+ to
increase in cultured toad bladder cells
(Wong et al., 1990), rabbit
medullary thick ascending limb cells
(Montrose-Rafizadeh and Guggino,
1991
), gastric parietal cells
(Negulescu et al., 1992
) and
Intestine 407 cells (Hazama and Okada,
1990
). Recent reviews provide multiple examples of different cell
types that elevate intracellular Ca2+ in response to acute osmotic
swelling (Foskett, 1994
;
McCarty and O'Neil, 1992
).
Our findings also show that extracellular Ca2+ was necessary for
the swelling-induced rise in cytosolic free Ca2+. This was
demonstrated by a lack of fluorescence associated with hypotonic shock when
cells were exposed to a `low Ca2+ medium'. It should be noted,
however, that once internalized, fluo-4 is diluted as cells swell in response
to hypotonic shock. This in turn decreases fluorescence intensity in a
corresponding manner. Because we did not measure relative fluorescence
intensity concomitant with cell volume changes, it is possible there was a
small rise in intracellular Ca2+ in swollen cells exposed to the
`low Ca2+ Ringer' that we could not detect. However, we do not
believe this was a significant problem in our study. For example, cells in a
normal Ca2+ hypotonic Ringer displayed bright fluorescence even
though they initially swelled to the same extent as cells in a low
Ca2+ hypotonic Ringer. Conversely, although cells in a low
Ca2+ hypotonic Ringer had an inhibited regulated volume decrease,
they still initially swelled to approximately the same extent as cells in
normal Ca2+ hypotonic Ringer, yet displayed virtually no
fluorescence. Taken together, these observations support our conclusion that
extracellular Ca2+ is necessary for an increase in fluorescence
associated with cell swelling. We cannot, however, rule out the presence of a
Ca2+-induced calcium release from intracellular stores. The
importance of extracellular Ca2+ also has been shown for other cell
types, including kidney cells
(Montrose-Rafizadeh and Guggino,
1991) and toad bladder cells
(Wong et al., 1990
), which do
not display a rise in cytosolic Ca2+ following hypotonic shock in
the presence of low extracellular Ca2+.
Additionally, we found the rise in intracellular Ca2+ that
accompanied hypotonic challenge also was correlated with subsequent volume
decrease. For example, application of Ca2+ ionophore caused a
decrease in osmotic fragility and also potentiated the rate of volume recovery
following hypotonic challenge. In addition, we previously demonstrated that
ionophore causes isosmotic cells to shrink
(Light et al., 1999). It is
possible, however, that the rise in Ca2+ resulting from A23187 may
have stimulated solute efflux unrelated to a normal RVD process. However, our
results are more consistent with a rise in cytosolic Ca2+ being
necessary, at least in part, for regulated volume decrease. We make this claim
because a low Ca2+ Ringer solution increased osmotic fragility and
also reduced the rate of volume decrease in swollen cells, demonstrating the
importance of extracellular calcium. Furthermore, buffering intracellular
Ca2+ with BAPTA had a similar response to chelating extracellular
Ca2+. Thus, if a rise in Ca2+ during cell swelling was
an epiphenomenon accompanying RVD, as described by others
(Pasantes-Morales and Morales-Mulia,
2000
), then buffering cytosolic Ca2+ should not have
inhibited volume decrease. Finally, a low Ca2+ hypotonic Ringer
also reduced whole-cell currents that are normally activated during cell
swelling.
We conducted experiments to assess the specific nature of a Ca2+
entry pathway during cell swelling. Interestingly, we found conventional
Ca2+ channel blockers, such as verapamil and nifedipine, had no
effect on fluo-4 fluorescence nor on cell volume recovery and whole-cell
conductance. This is in contrast to several other reports that show these
antagonists inhibit a rise in intracellular Ca2+ following
hypotonic challenge (Montrose-Rafizadeh
and Guggino, 1991; Wong et
al., 1990
). On the other hand, in this study gadolinium, a
stretch-activated channel antagonist (Yang
and Sachs, 1989
), blocked the increase in fluorescence associated
with cell swelling. We previously reported that this lanthanide increases
osmotic fragility and inhibits volume recovery and whole-cell currents
activated by hypotonic challenge (Bergeron
et al., 1996
; Light et al.,
1999
). Further, all the inhibitory actions of gadolinium were
prevented with the Ca2+ ionophore, indicating that the
Ca2+ entry and Ca2+-dependent steps are `downstream' to
the gadolinium-sensitive site. On the basis of this information, a
stretch-activated channel is a reasonable candidate for a permeability pathway
for Ca2+ influx during cell swelling.
However, gadolinium has also been shown to inhibit P2X receptors
(Nakazawa et al., 1997), which
are ATP-gated, Ca2+-permeable, non-selective cation channels
(Fredholm et al., 1994
).
Because we have shown that Necturus erythrocytes express P2X
receptors and activation of these receptors play a role in cell volume
decrease (Light et al., 1999
;
Light et al., 2001
), it is
possible the affect of gadolinium in this study was due to antagonism of P2X
receptors. In fact, this supposition was supported by our studies using
hexokinase and suramin. That is, both of these agents inhibited the
fluorescence increase associated with cell swelling, indicating a lack of
Ca2+ influx in the absence of P2 receptor activation.
Further, our previous findings have shown that hexokinase and suramin
increase osmotic fragility, inhibit regulated volume decrease and block
whole-cell currents associated with hypotonic swelling
(Light et al., 1999;
Light et al., 2001
). Finally,
in all cases the inhibitory nature of hexokinase and suramin was prevented
with Ca2+ ionophore, indicating that Ca2+-dependent
processes are `downstream' of the site of action of hexokinase and suramin,
similar to observations made with cultured neurons
(Garcia-Lecea et al., 1999
).
Taken together, our results are most consistent with extracellular ATP
activation of P2X receptors leading to a rise in cytosolic Ca2+. It
also seems likely that the receptor acts as a permeability pathway for
Ca2+ influx during cell swelling.
It is interesting to note that the effect of a low Ca2+ Ringer
on osmotic fragility was greater than the effect of blocking P2 receptors with
suramin. We can only speculate as to why this might be so. For example,
although suramin is considered a general P2 receptor antagonist
(Fredholm et al., 1994), it may
not completely block all P2 receptors. However, the observation that
hexokinase has virtually the same effect as suramin, and a lesser effect than
a low Ca2+ Ringer, suggests another explanation. There may have
been a suramin- and hexokinase-insensitive Ca2+ permeability
pathway that was open during cell swelling, independent of P2 receptors.
Alternatively, a low Ca2+ Ringer may have additional effects on
osmotic fragility, independent from regulation of permeability pathways, such
as altering membrane integrity.
Some reports indicate that the increase in cytosolic free Ca2+
following cell swelling is transient
(Negulescu et al., 1992;
Wong et al., 1990
). In
addition, others have reported that there is a time-dependence concerning the
activation of transport pathways used for RVD. For example, Ehrlich ascites
tumor cells show activation of a Cl- conductance during cell
swelling that becomes inactivated within the next 10 minutes
(Hoffman et al., 1986
). With
this in mind, we examined whether there was a time dependence for
Ca2+-stimulated volume decrease in Necturus erythrocytes.
This was accomplished by adding Ca2+ ionophore at different points
in time following hypotonic challenge. In all cases, this procedure
potentiated volume recovery. In addition, we also examined the converse; that
is whether lack of P2 receptor activation, and therefore Ca2+
influx, has a time-dependent nature. Application of hexokinase at different
points in time following hypotonic shock also did not display a time-dependent
affect, at least not during the time course of our experiments. Therefore,
Necturus erythrocytes do not display a narrow window of time
regarding activation of RVD mechanisms, at least as they relate to
Ca2+.
Our finding that intracellular Ca2+ levels increase during cell
swelling, which thereby stimulates volume decrease, is consistent with our
previous observations that volume regulation is inhibited by agents that
antagonize calmodulin (Bergeron et al.,
1996). Additionally, we reported that volume regulation depends on
phospholipase A2 and 5-lipoxygenase activity
(Light et al., 1997
), two
Ca2+-dependent enzymes involved with arachidonic acid metabolism
and the generation of leukotrienes
(Holtzman, 1991
). Thus, on the
basis of our previous work, it is not surprising that we found buffering
Ca2+ with BAPTA or EGTA inhibited volume decrease.
Our study also provides evidence that a rise in intracellular
Ca2+ leads to an increase in K+ efflux. This was shown
pharmacologically using the cationophore gramicidin with a choline Ringer.
With this solution, K+ and Cl- were the only two
permeable ions of significance, and addition of gramicidin ensured a continual
high K+ permeability. Gramicidin consistently prevented the
inhibitory effect of hexokinase and suramin, as well as that of BAPTA and
EGTA. The reason for examining the effect of gramicidin 5 minutes after
hypotonic shock is because that point in time corresponded with maximum cell
swelling, indicating it took several minutes for endogenous K+
channels to activate. We also previously reported that addition of gramicidin
causes cells to shrink under isosmotic conditions
(Light et al., 1999). Taken
together, our observations are consistent with this cell type having a low
K+ permeability under isotonic conditions and an elevated
K+ permeability during hypotonic stress in response to a rise in
cytosolic Ca2+.
Moreover, our electrophysiological studies demonstrated that cell swelling
leads to an increase in K+ permeability, via a conductive pathway,
in response to a rise in intracellular Ca2+. We have previously
shown that hypotonic shock activates a K+ conductance that is
necessary for volume recovery (Bergeron et
al., 1996), and this conductance is inhibited by agents that
antagonize calmodulin and phospholipase A2 (Light et al., 1996;
Light et al., 1997
). In this
study, we show that bathing cells in a low Ca2+ solution
significantly decreased the swelling-activated whole-cell conductance.
However, because the reversal potential did not change, we cannot rule out the
possibility that lowering Ca2+ also inhibited a Cl-
conductance concomitantly with a K+ channel. Further, it also is
possible that an increase in intracellular Ca2+ may have led to
membrane depolarization. This in turn could have resulted in activation of ion
channels required for volume recovery.
In conclusion, hypotonic challenge activates a P2 receptor, which leads to a rise in cytosolic Ca2+, thereby stimulating volume decrease by activating a K+ conductance. The coupling of a P2 receptor to Ca2+ and cell volume decrease represent a novel mechanism for osmotic regulation of cell function.
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Acknowledgments |
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