Astrocyte swelling leads to membrane unfolding, not membrane insertion

Astrocyte swelling leads to membrane unfolding, not membraneinsertion

Tina Pangrsic,*,� Maja Potokar,*,� Philip G. Haydon,� Robert Zorec*,� and Marko Kreft*,�

*Laboratory of Neuroendocrinology-Molecular Cell Physiology, Institute of Pathophysiology, Medical Faculty, University of

Ljubljana, Ljubljana, Slovenia

�Celica Biomedical Sciences Center, Ljubljana, Slovenia

�Department of Neuroscience, School of Medicine, University of Pennsylvania, Philadelphia, PA, USA

Abstract

The mechanisms mediating the release of chemical trans-

mitters from astrocytes are the subject of intense research.

Recent experiments have shown that hypotonic conditions

stimulate the release of glutamate and ATP from astrocytes,

but a mechanistic understanding of this process is not avail-

able. To determine whether hypotonicity activates the process

of regulated exocytosis, we monitored membrane capacitance

by the whole-cell patch-clamp technique whilst a hypotonic

medium was applied to cultured astrocytes. If exocytosis is

triggered under hypotonic conditions, as it is following

increases in cytosolic calcium, a net increase in membrane

surface area, monitored by measuring the whole-cell mem-

brane capacitance, is expected. Simultaneous measurements

of cell size and whole-cell membrane conductance and sur-

face area demonstrated that hypotonic medium (210 mOsm

for 200 s) resulted in an increase in membrane conductance

and in the swelling of cultured astrocytes by an average of

40%, as monitored by cell cross-sectional area, but without

any corresponding change in membrane surface area. As we

have demonstrated that capacitance measurements have the

sensitivity to detect increases in cell surface area as small as

0.5%, we conclude that cell swelling occurs via an exocytosis-

independent mechanism, probably involving the unfolding of

the plasma membrane.

Keywords: ATP, calcium, exocytosis, hypotonicity, mem-

brane capacitance.

J. Neurochem. (2006) 99, 514–523.

Although astrocytes are known to provide important support-ive functions to neurons, more recent studies have demon-strated that astrocytes release chemical messengers, includingglutamate, ATP and D-serine, that affect the function ofneuronal networks (Parpura et al. 1994; Innocenti et al. 2000;Haydon 2001; Nedergaard et al. 2002; Newman 2003;Volterra and Meldolesi 2005). In addition to the recentlyappreciated role of astrocytes releasing chemical transmittersby exocytosis in response to receptor-mediated elevations ininternal Ca2+, glutamate and ATP may also be released inresponse to swelling (Mongin et al. 1999; Mongin andKimelberg 2002; Darby et al. 2003). As ischemia can inducethe swelling of astrocytes, this pathway of chemical messen-ger release may play a deleterious role following trauma,stroke and other pathologic conditions.

Many mechanisms have been proposed to mediate therelease of glutamate from astrocytes, but which mechanismplays a major role under specific conditions has not yet beenfirmly established. Several release pathways have been

considered, both calcium-dependent and calcium-independ-ent: regulated exocytosis (Evanko et al. 2004), cystine–glutamate exchangers (Warr et al. 1999), hemichannels(Ye et al. 2003), P2X7 receptors (Duan et al. 2003), thereversal of plasma membrane glutamate transporters (Szat-kowski et al. 1990; Rossi et al. 2000) and anion channel-mediated mechanisms (Mongin et al. 1999). Although thework of several laboratories supports a role for exocytosis inmediating the Ca2+-regulated release of glutamate during

Received March 15, 2006; revised manuscript received June 2, 2006;accepted June 7, 2006.Address correspondence and reprint requests to Robert Zorec,

Laboratory of Neuroendocrinology-Molecular Cell Physiology, Instituteof Pathophysiology, Medical Faculty, University of Ljubljana, Zaloska 4,1000 Ljubljana, Slovenia. E-mail: [email protected] used: [Ca2+]i, intracellular free calcium concentration;

Cm, membrane capacitance; CSA, cross-sectional area; Idc, d.c. current;FM 1-43, N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium dibromide.

Journal of Neurochemistry, 2006, 99, 514–523 doi:10.1111/j.1471-4159.2006.04042.x

514 Journal Compilation � 2006 International Society for Neurochemistry, J. Neurochem. (2006) 99, 514–523� 2006 The Authors

physiologic activation, there is still debate over themechanisms mediating the release of this transmitter underother conditions.

Astrocytes are known to release glutamate as part of cellvolume regulation (Mongin et al. 1999). Hypotonic swell-ing-induced glutamate release has been associated with theactivation of volume-sensitive organic anion channels (Mon-gin et al. 1999). In contrast, it has been proposed that theglutamate release during astrocyte swelling is mediated byexocytosis (Pasantes-Morales et al. 2002), and that astro-cytes swell in a Ca2+-dependent manner (Hansson 1994;Nedergaard et al. 2002). Therefore, an increase in intracel-lular free calcium concentration ([Ca2+]i) may cause anincrease in astrocyte cell volume by an unknown mechanism,leading to the subsequent release of glutamate throughvolume-sensitive organic anion channels (Nedergaard et al.2002; Takano et al. 2005). To address these possibilities, weinvestigated whether regulated exocytosis is associated withhypotonically induced swelling.

In our experiments, we measured the swelling of astro-cytes by monitoring the morphologic appearance of singlecells, whilst simultaneously measuring membrane capaci-tance (Cm) by patch-clamp recording to detect surface areachanges, which reflect exocytotic activity.

The results showed that, during hypotonically inducedastrocyte swelling, Cm remained unchanged, but a significantincrease in membrane conductance was recorded. Therefore,our data support the view that channel activation, rather thanregulated exocytosis, underlies the astrocyte response tohypotonic stimuli.

Materials and methods

Primary astrocyte cultures

Astrocyte cultures were prepared from the cerebral cortices of 2–3-

day-old rats as described previously (Schwartz and Wilson 1992).

Cells were grown in high-glucose Dulbecco’s modified Eagle’s

medium containing 10% fetal bovine serum, 1 mM pyruvate, 2 mM

glutamine and 25 lg/mL penicillin/streptomycin in 92% air/8%

CO2. Confluent cultures were shaken at 1 g overnight and the

medium was changed the next morning; this was repeated a total of

three times. After the third shaking, the cells were dissociated using

cell dissociation solution (Sigma, St. Louis, MO, USA; C-5789) and

cultured for 24 h in 10 lM cytosine arabinoside. After reaching

confluence again, the cells were subcultured onto 22-mm-diameter

poly-L-lysine-coated coverslips and used within 4 days after plating.

If not mentioned otherwise, all chemicals were obtained from

Sigma.

Cell surface cross-sectional area (CSA) measurements

Transmitted light microscopyExperiments were performed using an imaging system (Till

Photonics, Grafelfing, Germany). Coverslips with astrocytes were

placed in a 250-lL recording chamber on an inverted microscope

(Zeiss Axiovert 135, Jena, Germany). The cells were imaged using a

charge coupled device camera (Imago, Till Photonics). Images were

recorded with Vision software (Till Photonics) every 5 s and the

sequence was stored on the hard disk of a computer. The image

sequence was subsequently read into Matlab (MathWorks Inc.,

Natick, MA, USA), and the surface area of the soma cross-section

was estimated using a custom-written program. Briefly, the outline

of the soma was traced interactively and marked manually in

consecutive digitized images with a mouse-generated cursor. The

surface area of the cross-section was then calculated and stored. In

all CSA measurements, focal planes of maximum cell diameter were

used.

Confocal microscopyAstrocyte-loaded coverslips were transferred into the recording

chamber of a confocal microscope (Zeiss LSM 510-META)

and supplied with 250 lL of extracellular solution containing

N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium

dibromide (FM 1-43) (4 lM; Molecular Probes, Leiden, the

Netherlands). Fluorescent and transmitted light images were

acquired by a · 63 plan-apochromatic oil-immersion objective

(NA 1.4) using 477 nm argon ion laser excitation and an LP

505 nm emission filter. Images were stored on a computer where

the z-axis diameter (fluorescent images) and CSA (fluorescent and

transmitted light images) changes were analyzed using LSM

510-META software (Zeiss) and software subroutines written in

Matlab. During medium osmolarity reduction, the concentration

of FM 1-43 did not change.

Electrophysiological recordings

Membrane capacitanceSimultaneously with the cell surface CSA measurements, we

performed electrophysiological patch-clamp recordings in the

whole-cell configuration. Astrocytes on coverslips were bathed in

an extracellular medium containing (in mM): NaCl, 140; KCl, 5;

MgCl2, 2.5; CaCl2, 2.5; Na HEPES, 10; D-glucose, 12.5; adjusted to

pH 7.3 with NaOH [osmolarity ¼ 305 ± 1.5 mOsm (n ¼ 5), meas-

ured with freezing point osmometer Osmomat030, Gonotec GmbH,

Berlin, Germany]. The 30% reduction in osmolarity of the

extracellular medium (210 mOsm) was achieved by diluting the

medium with distilled water. Patch-clamp pipettes had a resistance of

approximately 3 MW when filled with electrode solution containing

(in mM): K gluconate, 140; tetraethylammonium chloride (TEACl),

10; MgCl2, 2; HEPES, 10; Na2ATP, 2; adjusted to pH 7.3 with KOH

[osmolarity ¼ 307 ± 2.5 mOsm (n ¼ 6)]. [Ca2+]i was set to 100 nM

using Ca2EGTA/EGTA buffer. All recordings were made at room

temperature (20�C).Uncompensated Cm measurements were performed (Zorec et al.

1991) using a SWAM IIB amplifier (Celica, Ljubljana, Slovenia),

operating at 800 Hz lock-in frequency (sine wave of 1.1 mV rms).

Cells were voltage clamped at a holding potential of ) 70 mV. The

d.c. current (Idc), holding potential (V), real (YRe) and imaginary

(YIm) components of the admittance signals were acquired every

10 ms, digitized and stored on a second computer by CELL

software (Celica, Ljubljana, Slovenia). The time-courses of ATP-

stimulated Cm and Idc were independent. Signals were subsequentlyread into Matlab where all further analysis was performed.

Swelling of astrocytes 515

� 2006 The AuthorsJournal Compilation � 2006 International Society for Neurochemistry, J. Neurochem. (2006) 99, 514–523

Recordings with an access conductance of less than 50 nS were

discarded from the analysis.

All data are given as the mean ± SEM; n denotes the number

of individual cells assessed in the simultaneous patch-clamp

and imaging studies. Statistical differences were determined by

two-tailed unpaired Student’s t-test and considered to be significant

at p < 0.05.

Membrane conductanceTo determine membrane conductance changes, current–voltage

relations were measured in response to 30-ms voltage steps from

a holding potential of ) 100 mV to final values ranging between

) 150 mV and + 10 mV in 20-mV increments before and after the

decrease in the extracellular medium osmolarity. Currents were

digitized on the same computer by WinWCP software (Strathclyde

University, Glasgow, UK). Further analysis was performed in

Matlab. Briefly, membrane currents 3 ms before the end of the

voltage pulses were measured and plotted against the voltages to

obtain I/V curves, taking into account liquid junction potentials due

to solution composition differences () 30 ± 5 mV, n ¼ 7).

Calcium measurements

[Ca2+]i was measured with Fura-2 (applied as Fura-2/AM; Molecu-

lar Probes Inc., Eugene, OR, USA). The cells were incubated for

30 min at 37�C in medium containing 4 lM Fura-2/AM. The

loading solution was then washed out and replaced by extracellular

solution. The fluorescence of Fura-2 was measured using a Till

Photonics System equipped with a · 40 fluar oil-immersion

objective (NA 1.3) and a charge coupled device camera. Ratio

measurements were performed every 5 s by collecting image pairs

exciting the cells at 340 and 380 nm, respectively. The emission

light was passed through an LP 440 nm filter. For calcium

calibration (Fmin and Fmax), a Ca2+-free solution containing

10 mM EGTA with 10 lM ionomycin and a solution containing

10 mM Ca2+ with 10 lM ionomycin were used. [Ca2+]i was

calculated using standard equations as described by Grynkiewicz

et al. (1985). In all experiments, a constant Kd value of 224 nM

was used.

Results

Quantification of hypotonic astrocyte swelling by

monitoring time-dependent cell CSA changes

In our experiments, we used electrically separated, non-confluent process-bearing cells from rat cortical astrocytecultures. Their identity was determined in separate experi-ments using the astrocyte-specific markers GFAP and S100b(data not shown). We estimated cell swelling in hypotonicconditions by carefully examining changes in cell size underthe microscope. We first tested whether cell swellingmeasurements in transmitted light images were comparablewith measurements in fluorescent images, where membraneswere marked with fluorescent membrane dye FM 1-43. Theastrocytes were bathed in medium containing FM 1-43 andconfocal microscopy was performed. To quantify swellingduring a 30% decrease in bathing medium osmolarity,

changes in the z-axis diameter of cells were measured first(Figs 1a and b, top panels). The diameter of the cell shown inFig. 1 transiently increased and then declined to a level equalto around 8% above the resting diameter measured before theintroduction of the hypotonic stimulus (Fig. 1c); 35 s afterthe decrease in extracellular osmolarity, an average increasein the z-axis diameter of 13 ± 4% (n ¼ 9, Fig. 1d) wasdetected. After the initial peak, the diameter of the cellsremained approximately 8% above resting values.

Next, changes in cell size in the x,y plane were analyzed(Figs 1a and b, middle panels). With fluorescently labelledplasma membrane, the position of the cell limits in theconfocal x,y plane was easily detected (white dots in themiddle panels of Fig. 1) in consecutive images and the CSAwas calculated (Fig. 2). On average, the CSA of nine cellsincreased in 35 s by 9 ± 3% (n ¼ 9, Fig. 2b, gray line). Thetime course of CSA increase was similar to the time course ofthe z-axis diameter change (Fig. 1). Separately, cell borderson transmitted light images were observed (Figs 1a and b,

(a) (b)

(c) (d)

Fig. 1 The effect of hypotonic medium (+ hypo) on a single, FM 1-43-

stained astrocyte. Confocal z-plane (a and b, top panels) and x,y-

plane (a and b, middle panels) images and transmitted light images (a

and b, bottom panels) of a cell bathed in isotonic (a, 305 mOsm) and

hypotonic (b, 210 mOsm) medium. The image in (b) was taken 36 s

after the osmotic change. White lines in the top panels depict the

z-axis diameter of the cell. White dots in the middle and bottom panels

represent the off-line detection of cell borders used to calculate

changes in the cross-sectional area. (c) After reducing the extracel-

lular osmolarity to 210 mOsm, cell swelling in the z-axis direction was

observed. The z-axis diameter changes were normalized to the resting

diameter immediately prior to the osmotic change. (d) On average, a

maximal increase in the relative z-axis diameter of 13 ± 4% (n ¼ 9)

was observed 35 s following the hypotonic stimulus. Error bars rep-

resent the standard error of the mean. The medium osmolarity was

reduced by adding distilled water supplemented by FM 1-43 to the

bathing chamber.

516 T. Pangrsic et al.

Journal Compilation � 2006 International Society for Neurochemistry, J. Neurochem. (2006) 99, 514–523� 2006 The Authors

bottom panels, white dots). The amplitude and time course ofthe calculated CSA changes obtained from the analysis oftransmitted light images corresponded well with the CSAchanges calculated from fluorescent images (Fig. 2). Onaverage, the CSA changes in transmitted light images yieldeda maximum of 7 ± 3% in 30 s (n ¼ 9, Fig. 2b, black line).Linear regression analysis revealed a strong correlationbetween the results obtained by the two imaging methodsused (Fig. 2c, r ¼ 0.96, slope of the regression line ¼1.3 ± 0.0, n ¼ 9). Therefore, to avoid optical interference ofFM 1-43 staining the patch-pipette, we used transmitted lightmicroscopy to monitor cell swelling in electrophysiologyexperiments.

Hypotonic swelling accompanied by the activation of an

outward plasma membrane current does not lead to

changes in Cm in astrocytes

The circumference of single, patch-clamped cells wasmeasured and the CSA was calculated. As shown inFigs 3(a) and (b), the exposure of the patch-clamped cellto a hypotonic medium resulted in an increased CSA from101 to 159 lm2. Simultaneously, Idc, V, YRe and YIm wererecorded. Assuming a single time-constant cell model,depicted in Fig. 3(c), Cm was estimated. Figure 3(d) showsa representative real-time patch-clamp Cm measurement ofa cortical astrocyte dialysed with a 307 mOsm pipette-filling solution and exposed to reduced osmolarity bathingmedium. As Cm is linearly proportional to the membranesurface area, changes in Cm are interpreted to reflect a neteffect of exocytotic and endocytotic activity: vesiclemembrane fusion with and fission from the plasmamembrane, respectively (Neher and Marty 1982). It shouldbe noted that the Cm trace was stable over 8 min, whereasthe middle trace, which represents the change in Idc,

(a) (b)

(c)

Fig. 2 Comparison between the analysis of confocal (gray lines) and

transmitted light (black lines) images. In response to hypotonic shock

(+ hypo), time-dependent changes in cross-sectional area (CSA) of

the cell shown in Fig. 1 (a) and the average of nine cells (b) were

observed. Maximal CSA changes, calculated by outlining the cell

borders in consecutive fluorescent or transmitted light images, yielded

values of 9 ± 3% and 7 ± 3%, 35 and 30 s after medium osmolarity

reduction, respectively. CSA changes were normalized to the resting

CSAs of each cell immediately prior to the osmotic change. Error bars

represent the standard error of the mean. Note that CSA changes

calculated from fluorescence and transmitted light data have similar

amplitudes and time courses. (c) Linear regression (gray line) calcu-

lated for the relative CSA changes obtained by analysis of transmitted

light and fluorescent images. A high correlation was observed between

the two measurements (r ¼ 0.96, n ¼ 85, slope of the regression line

1.3 ± 0.0). White and gray circles represent CSA changes before and

during the hypotonic stimulus, respectively.

(a) (b)

(c) (d)

Fig. 3 The effect of hypotonic medium (+ hypo) on single, patch-

clamped astrocyte. (a) The cell was bathed in isotonic medium

(305 mOsm); white dots represent off-line analysis with outlined cell

shape (bar, 10 lm). (b) After reducing the extracellular osmolarity to

210 mOsm, cell swelling was observed. The image was taken 200 s

after the osmotic change. (c) An electrical model of the patch-clamped

astrocyte comprises membrane capacitance Cm, membrane conduct-

ance Gm and access conductance Ga. (d) A typical recording of Cm,

membrane current (Idc) and Ga time-dependent changes in a single

astrocyte during hypotonic stimulation (bar). The medium osmolarity

was reduced by adding distilled water to the bathing chamber.



showed a hypotonicity-induced change in membraneconductance.

In isotonic medium, the CSA of astrocytes yielded anaverage value of 113 ± 12 lm2 (n ¼ 7). Hypotonicity-induced changes in CSA are expressed as the relativechange (in percentage) in CSA with respect to the image inwhich cells last experienced isotonic medium. Figure 4(a)shows a representative response of CSA of a single patch-clamped astrocyte to hypotonic medium, which increasedby 40% following the addition of the hypotonic medium(Fig. 4a, top panel). The increase in CSA, however, was notaccompanied by a change in Cm (Fig. 4a, bottom panel),measured simultaneously from the same cell. The applica-tion of hypotonic medium (see the artefact in Cm recording)did not affect the time course of Cm. To obtain the timedependence of CSA changes, we measured them in 25-sintervals and calculated the linear regression (Fig. 4b, toppanel, see line). The regression was significantly differentfrom zero (p < 0.05, n ¼ 63, where n represents thenumber of individual observations in all cells tested). TheCSA in the hypotonic medium increased on average by38 ± 12% at 200 s after the application of hypotonicsaline. In contrast, when the Cm changes of the same cellswere plotted against time (Fig. 4b, bottom panel), nosignificant time dependence was found. The slope of theregression line was not significantly different from zero(p > 0.05, n ¼ 63). Changes in Cm measured 200 sfollowing the application of hypotonicity were on average37 ± 153 fF, equivalent to a change of 0.6 ± 1.5% rela-tive to the resting Cm of each cell determined prior tothe application of the hypotonic medium. The absence ofa significant change in Cm in cells relative to the restingCm (on average 10.5 ± 1.8 pF, n ¼ 7) indicates that thereis no net change in membrane surface area associatedwith the significant increase in CSA. These results indicatethat hypotonicity evokes an unfolding of the plasmamembrane without a significant change in the plasmamembrane area.

To further test whether the hypotonic challenge affects theplasma membrane properties in astrocytes, we monitoredmembrane conductance. A common observation in experi-ments investigating the effect of osmotic shock on mamma-lian cells is the activation of regulatory mechanisms, whichoften involve the activation of electrically non-neutralmembrane currents, opposing cell volume changes (Wehneret al. 2003). To test whether these mechanisms are present inastrocytes in our experimental conditions, we measuredmembrane currents in conditions of normal and reducedosmolarity. In isotonic medium, square-voltage stimulationprotocols were applied (Fig. 4c), revealing that the cells havetypical characteristics of non-rectifying astrocytes (Matthiaset al. 2003), with resting membrane potentials (zero-currentpotential) of around ) 80 to ) 70 mV (Fig. 4d). When theosmolarity of the extracellular medium was reduced, we

observed an increase in membrane conductance (Fig. 4d),which is in agreement with previous reports (Olson and Li1997; Darby et al. 2003), with no accompanying netexocytosis.

(a) (b)

(c) (d)

Fig. 4 Simultaneous time-dependent changes in cross-sectional area

(CSA) and membrane capacitance (Cm) in response to hypotonic

shock (bar, 210 mOsm), and the effect of hypotonicity on membrane

current in cultured rat astrocytes. (a) The response of a single cell with

the initial Cm of 10.4 pF. (b) Linear regression (see lines in top and

bottom panels) calculated for the average relative CSA (CSA [%] ¼(0.19 ± 0.04 [%/s]) · t [s] ) (0.23 ± 4.91 [%]), n ¼ 63) and Cm (Cm

[%] ¼ (0.00 ± 0.01 [%/s]) · t [s] ) (0.26 ± 0.67 [%]), n ¼ 63) chan-

ges. Capacitance records were normalized to the resting capacitance

immediately prior to the osmotic change. The slope of the CSA

changes (11.7 ± 2.5%/min) is significantly different from zero. Error

bars represent the standard error of the mean. (c) Representative

traces of membrane currents of astrocytes measured in isotonic (left

panel) and hypotonic (right panel) medium. The cells were voltage-

clamped at ) 100 mV and stimulated with a train of 30-ms square-

voltage pulses ranging from ) 150 to + 10 mV in 20-mV increments.

(d) The I/V curve obtained in isotonic (filled circles) and hypotonic

(open circles) conditions. The asterisks represent statistically signifi-

cant differences between the two curves (p < 0.05, Student’s t-test,

n ¼ 9).



Hypotonic swelling of astrocytes is accompanied by only

a moderate increase in [Ca2+]i, whereas ATP induces a

micromolar increase in [Ca2+]i

To understand why there was no net increase in Cm whenastrocytes were exposed to hypotonic medium (Fig. 4), weinvestigated whether hypotonic conditions led to changes in[Ca2+]i. As published previously (McCarty and O’Neil 1992;Altamirano et al. 1998; Morales-Mulia et al. 1998; Koyamaet al. 2001), we found only a moderate increase in [Ca2+]i onhypotonic stimulation of astrocytes (Fig. 5a). On average,[Ca2+]i increased from the resting level of 60 ± 8 nM to amaximum of 116 ± 13 nM (Fig. 5a, iv; maximal valuesmeasured at 95 s; n ¼ 15). These values are far below therequired [Ca2+]i to trigger Ca2+-dependent exocytosis inastrocytes, which occurs at 10 lM [Ca2+]i (Kreft et al. 2004).Therefore, the absence of a Cm increase in cells exposed tohypotonicity is consistent with the small increase in [Ca2+]iduring hypotonic challenge.

To test whether astrocytes may swell in a Ca2+-dependentmanner (Hansson 1994; Nedergaard et al. 2002), we usedATP, a potent stimulator of elevations of internal [Ca2+]i(Salter and Hicks 1994), and which may also be releasedduring hypotonic swelling (Wang et al. 1996; Mongin andKimelberg 2002). The application of ATP (1 mM) elicited amicromolar increase in [Ca2+]i as measured by Fura-2/AM(Fig. 5b; maximal values measured at 10 s; 2.9 ± 1.3 lM,n ¼ 7). This response is probably due to activated purinergicreceptors, as 100 nM Brilliant Blue (Jiang et al. 2000) causeda 91% inhibition of [Ca2+]i (maximal increase measured at20 s; 0.3 ± 0.3 lM, n ¼ 13). A micromolar increase in[Ca2+]i was also observed with 10 lM ATP (maximalincrease measured at 20 s; 1.1 ± 0.9 lM, n ¼ 9). Themaximal responses to 1 mM and 10 lM ATP were notsignificantly different (p > 0.05, t-test).

ATP-induced increase in Cm and inward current does not

lead to significant swelling of astrocytes

As ATP addition increases [Ca2+]i, ATP-dependent swellingof astrocytes would be expected if previous proposals werecorrect (Nedergaard et al. 2002). Moreover, if ATP elicits amicromolar increase in [Ca2+]i, a process of regulated Ca2+-dependent exocytosis is likely to be detected in these cells(Kreft et al. 2004). Therefore, we applied ATP to the bathingmedium whilst imaging and Cm measurements were per-formed simultaneously. Six of the 11 cells tested exhibited anincrease in Cm induced by either 100 lM or 1 mM ATP(Fig. 6). On average, the Cm of responding cells in 200 sincreased by 334 ± 130 fF (with maxima near 1000 fF) or3.5 ± 1.1% (n ¼ 6, p < 0.05; Fig. 6b, bottom panel) relativeto the resting Cm of each cell (on average 9.8 ± 1.3 pF, n ¼6) determined prior to the application of ATP (Fig. 6a and b,bottom panels). Linear regression was calculated for the firstfour time points (Fig. 6b, bottom panel, dotted line). Theslope of the regression line was significantly different from

zero (p < 0.05, n ¼ 24). Further, the ATP stimulationinduced a slight increase in the CSA of astrocytes(15 ± 6% in 200 s, linear regression, p < 0.05; Fig. 6, toppanels). The slope of the time dependence of CSA after ATPaddition was significantly different from zero; however, itwas not significantly different from the slope in controlexperiments, where an isotonic vehicle without ATP wasapplied (Fig. 6b, top panel, gray line). Isotonic solution

(a) (b)

Fig. 5 Increase in intracellular free calcium concentration ([Ca2+]i) on

hypotonic and ATP stimulation. Astrocytes loaded with Fura-2/AM and

bathed in isotonic medium (ai and bi); (aii) 80 s after the exposure to

hypotonic medium; (bii) 80 s after the exposure to 1 mM ATP. After

exposure to hypotonic medium (aiii and aiv, n ¼ 15) or 1 mM ATP (biii

and biv, n ¼ 7), a significant increase in [Ca2+]i was observed, which,

after 80 s, yielded values of 109 ± 12 and 368 ± 54 nM, respectively.

Note that the maximal increase in [Ca2+]i in response to ATP stimu-

lation exceeds values of 2 lM, which can be measured by Fura-2/AM.

(aiv) Inset with smaller ordinate scale.



without the addition of ATP also induced no increase in Cm

(data not shown). From these data, we conclude that the Cm

increase is clearly attributable to ATP stimulation. We cannotentirely rule out the possibility of a small increase in volumefollowing ATP application to astrocytes (Takano et al. 2005),which, in our experiments, could not be distinguished fromthe control.

ATP addition activated inward membrane currents in allcells tested (Fig. 6c). The increase in current was significantat voltages below ) 90 mV (Fig. 6d, p < 0.05, n ¼ 4). Thenature of the ATP-induced inward current was not studiedfurther in our experiments. However, strong inhibition of[Ca2+]i responses by 100 nM Brilliant Blue G implies apossible involvement of purinergic receptors (Jiang et al.2000).

Discussion

ATP and glutamate have both been shown to be releasedduring hypotonic cell swelling, and both can act as autocrineor paracrine factors (Wang et al. 1996; Mongin and Kimel-berg 2002; Nedergaard et al. 2002; Darby et al. 2003;Takano et al. 2005). Currently, however, the nature of theirrelease from cells during swelling is not entirely clear. ATPrelease is thought to involve ATP transporters and/or ATP-permeable channels, whereas, in some epithelial and clonalcells, exocytosis has been proposed as the pathway of ATPrelease (Wang et al. 1996; Coco et al. 2003; van der Wijket al. 2003; Fabbro et al. 2004). For glutamate release, atleast two possible mechanisms have been considered. First,the release of the anionic form of glutamate through Cl–

channels has been proposed, although glutamate is present inthe cytosol mostly in its zwitterionic form (Pasantes-Moraleset al. 2002). Second, in hypotonically swollen astrocytes,exocytosis has been proposed as the mechanism of glutamaterelease (Pasantes-Morales et al. 2002).

A key issue to be addressed in this study was whether ornot exocytosis (membrane area changes) accompaniedastrocyte cell swelling. In other cell types, the results arenot uniform. Although some studies have shown an increasein surface area due to exocytosis in hypotonic conditions inbiliary epithelial cells (Gatof et al. 2004), Jurkat T lympho-cytes (Ross et al. 1994), human intestine 407 cells (Okadaet al. 1992; van der Wijk et al. 2003) and plant cells(Kubitscheck et al. 2000), results in rat hepatocytes andbovine chromaffin cells show no activation of exocytosis byhypotonic saline (Graf et al. 1995; Kilic 2002).

To detect changes in cell size and morphology, twoapproaches were used. Cells were bathed in mediumcontaining the fluorescent membrane marker FM 1-43,which stains plasma membrane. Confocal three-dimensionalfluorescent images of cells, and transmitted light images,were taken. The analysis of the CSA measurements of cellscalculated from transmitted light images corresponded well

(a) (b)

(c) (d)

Fig. 6 Simultaneous changes in cross-sectional area (CSA) and

membrane capacitance (Cm) in response to 1 mM ATP (bar), and

the effect of ATP application on membrane current in cultured rat

astrocytes. (a) The activation of exocytosis in a single astrocyte with

the initial Cm of 9.8 pF. (b) The average change in CSA and Cm

following the application of ATP. In six of 11 cells, a significant

increase in Cm was observed (mean ± SEM, asterisks). Linear

regression was calculated for the first four time points (bottom pa-

nel, dotted line, Cm [%] ¼ (0.05 ± 0.02 [%/s]) · t [s] ) (0.02 ± 0.76

[%]), n ¼ 24). CSA also increased slightly (top panel, CSA [%] ¼(0.07 ± 0.02 [%/s]) · t [s] ) (0.54 ± 1.79 [%]), n ¼ 54). The slope of

the linear regression line (4.1 ± 0.9%/min) was significantly different

from zero (p < 0.05), but this increase was not significantly different

from controls (top panel, gray line, CSA [%] ¼ (0.03 ± 0.02 [%/

s]) · t [s] + (1.05 ± 1.90 [%]), n ¼ 27, slope ¼ 1.6 ± 1.0%/min). Er-

ror bars represent standard error of the mean. (c) Representative

traces of membrane currents of voltage-clamped astrocytes meas-

ured in isotonic medium before (left panel) and after (right panel)

ATP stimulation. The same voltage pulse protocols as for hypotonic

stimulation were used. (d) The I/V curve obtained in non-stimulated

cells (filled circles) and ATP-stimulated cells (open circles). The

asterisks represent statistically significant differences between the

two curves (p < 0.05, Student’s t-test, n ¼ 4).



with the CSA measurements in fluorescent images. Theswelling of patch-clamped cells (Fig. 3) in hypotonicmedium was comparable with the initial swelling of non-patch-clamped cells (Fig. 2), where the swelling is probablypartially compensated.

An ideal parameter by which plasma membrane areachanges may be monitored is Cm, which is dynamicallymeasured by the patch-clamp technique (Lindau and Neher1988; Zorec et al. 1991). We therefore measured Cm inastrocytes to determine whether the effect of osmotic shockon CSA and cell volume was associated with changes inmembrane surface area, reflecting a net activation ofexocytotic activity (Fig. 3).

The results showed that the surface area was not alteredunder conditions of hypotonic stress in cultured astrocytes,but the cell volume, monitored as CSA, consistent withTakano et al. (2005), increased significantly (Fig. 4). There-fore, we concluded that the increase in CSA duringhypotonically induced astrocyte swelling was not associatedwith a significant activation of exocytosis, but was probablya result of plasma membrane unfolding (Figs 4 and 7).Similar results with cultured astrocytes have been obtainedby Olson and Li (1997), who measured time constants ofcapacitive transients in response to repeated square-voltagepulses, although these are not necessarily proportional to Cm

and membrane surface area changes only.We compared the surface area of cells, obtained from Cm

measurements, with the surface area calculated from CSAs.For the latter, the cell was modelled by a cylinder with aheight of 5 lm, which was the measured thickness ofastrocytes by confocal microscopy (Potokar et al. 2005;Takano et al. 2005), and with the measured CSAs repre-senting the base of the cylinder. From this, an averageestimate of the plasma membrane area of 457 ± 51 lm2 wasobtained, which is significantly smaller than the measuredarea of 1318 ± 226 lm2 (n ¼ 7) from Cm measurements,

taking into account the specific Cm of 0.008 pF/lm2 (White1986). The discrepancy between the values determined bythe two approaches suggests that the plasma membrane ofastrocytes is folded (Fig. 7). When the same calculationswere performed for cells in hypotonic medium, the discrep-ancy between the cell areas, calculated from morphologicaland capacitance data (578 ± 68 and 1322 ± 224 lm2,respectively, n ¼ 7), was decreased. This confirms that theexposure to hypotonic medium leads to the straightening outof the folds to account for the observed CSA increase in theabsence of a significant increase in Cm (Figs 3 and 4).

Our findings show that regulated exocytosis is notinvolved in astrocyte swelling and is therefore unlikely tobe the mechanism of autocrine or paracrine release ofneuroligands (such as ATP and glutamate) during astrocyteswelling. This is consistent with previous results (Monginand Kimelberg 2002; Takano et al. 2005) showing thattetanus toxin does not affect hypotonically induced aminoacid release. However, it should be noted that, in thoseexperiments, the cleavage of the soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) pro-tein, synaptobrevin, by tetanus toxin was not confirmed,casting some doubt on the interpretation of the results.

Several studies have shown an increase in [Ca2+]i duringhypotonic stimulation (McCarty and O’Neil 1992; O’Connorand Kimelberg 1993; Morales-Mulia et al. 1998; Koyamaet al. 2001), but this increase rarely exceeded 400 nM. In ourprevious study on astrocytes (Kreft et al. 2004), we demon-strated that the increase in [Ca2+]i, induced by flashphotolysis of Ca2+-o-nitrophenyl-EGTA (Ca2+-NP-EGTA)and measured by Furaptra, should be in the micromolar rangeto activate the process of regulated exocytosis. In the presentstudy, hypotonic shock increased [Ca2+]i moderately (Fig. 5),but below the threshold of activation of regulated exocytosis(Kreft et al. 2004). This is in agreement with the resultsshowing no Cm increase after hypotonic stimulation (Fig. 4).

Nedergaard et al. (2002) proposed a Ca2+-dependentastrocyte swelling mechanism, whereas Jeremic et al.(2001) proposed that ATP may cause cell swelling, which,in turn, stimulates glutamate release from astrocytes. It hasbeen shown that this release may involve the loss ofcytoplasmic glutamate through volume-sensitive ion chan-nels (Takano et al. 2005). We therefore tested whether ATPapplication caused Ca2+-dependent astrocyte swelling and,consequently, the activation of regulated exocytosis. Inter-estingly, although ATP, which, in our experiments, evoked amicromolar increase in [Ca2+]i, triggered an increase in Cm inmore than 50% of the cells tested, we could not detectadditional changes in CSA that could be accounted for by aprocess of membrane unfolding (Fig. 6). These results are inagreement with our previous report of the stimulation ofexocytosis with an agonist of metabotropic glutamatereceptor ((1S, 3R)-1-aminocyclopentane-1,3-dicarboxylicacid; Zhang et al. 2004).

Fig. 7 A model of the astrocyte cell surface before and after exposure

to a hypotonic medium. Astrocytes with initial diameters of 12 ± 1 lm

and cross-sectional areas (CSAs) of 113 ± 12 lm2 (n ¼ 7) swell in

response to hypotonic shock. In hypotonic conditions, the diameter

increases to 14 ± 1 lm (n ¼ 7, by a factor of 1.17), and the CSA

increases to 156 ± 21 lm2. Note that the number of vesicles during

cell swelling is depicted unchanged before and after the hypotonic

application.



In summary, our data show that Cm is unchanged inresponse to hypotonicity, supporting the view that regula-ted exocytosis is not involved in the release of glutamateand/or ATP during hypotonically induced swelling inastrocytes.

Acknowledgements

This work was supported by grants #P3521 and #P3310 of The

Ministry of Education, Sciences and Sports of The Republic of

Slovenia, EC Contract DECG QLG3-CT-2001-02004, Fogarty

International Research Collaboration Award grant R03-TW01293

to RZ and grants from the National Institutes of Health (NIH) (R37

NS37585, RO1 NS43142) to PGH.

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Astrocyte swelling leads to membrane unfolding, not membrane insertion