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Endocrinology Vol. 144, No. 11 4916-4922
Copyright © 2003 by The Endocrine Society

Role of Cell Volume in K+-Induced Ca2+ Signaling by Rat Adrenal Glomerulosa Cells

Judit K. Makara, Péter Koncz, Gábor L. Petheö and András Spät

Department of Physiology and Laboratory of Cellular and Molecular Physiology, Semmelweis University Medical School and Hungarian Academy of Sciences, H-1444 Budapest, Hungary

Address all correspondence and requests for reprints to: Prof. A. Spät, Department of Physiology, Semmelweis University Medical School, P.O. Box 259, H-1444 Budapest, Hungary. E-mail: Spat{at}Puskin.SOTE.Hu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The involvement of cell volume in the K+-evoked Ca2+ signaling was studied in cultured rat glomerulosa cells. Previously we reported that hyposmosis (250 mOsm) increased the amplitude of T-type Ca2+ current and, accordingly, enhanced the Ca2+ response of cultured rat glomerulosa cells to K+. In the present study we found that this enhancement is not influenced by the cytoskeleton-disrupting drugs cytochalasin-D (20 µM) and colchicine (100 µM). Elevation of extracellular potassium concentration ([K+]e) from 3.6 to 4.6–8.6 mM induced cell swelling, which had slower kinetics than the Ca2+ signal. Cytoplasmic Ca2+ signal measured in single glomerulosa cells in response to stimulation with 5 mM K+ for 2 min showed two phases: after a rapid rise reaching a plateau within 20–30 sec, [Ca2+]c increased further slowly by approximately one third. When 5 mM K+ was coapplied with elevation of extracellular osmolarity from 290 to 320 mOsm, the second phase was prevented. These results indicate that cell swelling evoked by physiological elevation of [K+]e may contribute to the generation of sustained Ca2+ signals by enhancing voltage-activated Ca2+ influx.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
ALDOSTERONE SECRETION BY adrenal glomerulosa cells is regulated mainly by plasma K+ concentration ([K+]e) and angiotensin II. Whereas angiotensin II activates G protein-coupled plasma membrane receptor to stimulate phospholipase C and Ca2+ release from the endoplasmic reticulum, K+ exerts its effect by depolarization of the plasma membrane, evoking Ca2+ influx through activation of T- and L-type voltage-gated Ca2+ channels (1). Aldosterone secretion in vivo (2) and production in vitro (3, 4, 5) are stimulated by increases in K+ concentration as small as a few tenths of millimole. Rat glomerulosa cells may respond to an elevation of [K+]e as small as 0.5 mM with Ca2+ signal (6) and increased activity of mitochondrial nicotinamide adenine dinucleotide phosphate (reduced) [NAD(P)H] formation (7).

One of the possible factors modifying voltage-gated Ca2+ channels involved in K+-induced Ca2+ signaling is cell volume. In a previous paper, we reported that treatment of cultured rat glomerulosa cells with hyposmotic (250 mOsm) extracellular solution augmented the Ca2+ response of single cells to 5 mM K+ (8). This may be well explained by the increased amplitude of the T-type current under hyposmotic conditions, because generation of the Ca2+ influx during small, physiological increases of [K+]e was attributed to this channel type (9, 10). Similarly, Wang et al. (11) observed that basal and K+-evoked aldosterone production measured in bovine glomerulosa cell suspension was dependent on the osmolarity of the incubating solution in the 233- to 303-mOsm range. Cell volume changes were shown to alter the function of a number of ion channels (12, 13). The two most likely mechanisms that can provide a simple and online transfer of volume changes to plasma membrane elements are 1) alteration of the concentration of cytosolic proteins or ions that regulate directly or indirectly the function of transport elements in the membrane and 2) mechanical stress induced in the cytoskeleton network or the plasma membrane (12, 14, 15, 16). Several observations support the role of the actin cytoskeleton in modifying the function of Na+, Cl-, and Ca2+ channels (17, 18, 19, 20, 21, 22).

Besides changes in osmolarity of the extracellular milieu, water influx and concomitant cell swelling can be induced also by elevation of the concentration of osmotically active molecules in the cytosol. Altered activity of transport mechanisms that result in net movement of ions through the plasma membrane may accordingly change cell volume. Isosmotic elevation of [K+]e higher than 25 mM was shown to induce cell swelling in some cell types, e.g. astrocytes (23) and epithelial cells (24, 25, 26, 27, 28, 29). Although the applied K+ concentration in these studies was well out of the physiological range, it raises the possibility that a similar mechanism could work in glomerulosa cells. Considering the stimulatory effect of hyposmotic swelling on the function of T-type Ca2+ channels, an eventual K+-evoked cell volume increase could be an essential element of the high sensitivity of glomerulosa cells to this ion. In the present study, we examined the effect of physiological changes in [K+]e on the volume and cytoplasmic Ca2+ concentration ([Ca2+]c) of rat glomerulosa cells.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell preparation and culture
Animals were used with the approval (No. 17/1998) of the Animal Care and Ethics Committee of the Semmelweis University. All procedures followed legal and institutional guidelines of animal care. Male Wistar rats (200–250 g) were kept on a standard semisynthetic diet. Glomerulosa cells were prepared from the adrenal capsular tissue with collagenase digestion, as described (30). Freshly digested cells were plated onto polylysine-coated glass coverslips and incubated in 5% CO2 at 37 C in a mixture of modified Krebs-Ringer-bicarbonate solution and M199 (38:62, vol/vol, final concentrations: 146 mM Na+, 3.6 mM K+, 0.5 mM Mg2+, 1.2 mM Ca2+). Cells were used after 1 ([Ca2+]c measurements) or 2 d (cell area measurements).

Ratiometric measurement of cytoplasmic [Ca2+] with indo-1
Experiments were performed exactly as described previously (8). Resting [Ca2+]c, measured with indo-1 in this study was lower than that reported in our previous paper (8). At that time, an incorrect calibration equation, provided by the PClamp software (6.0), was used.

For kinetic analysis of the K+-induced Ca2+ signal, the indo-1 fluorescence ratio curves were low-pass filtered offline with 20 kHz (Bessel filter) and derivated using the software PClamp 8.0 (Axon Instruments, Foster City, CA). The number of positive deflections (indicating an increase in the slope of the elevation of [Ca2+]c) during the 2-min stimulation of glomerulosa cells with K+ was determined using a single-channel analysis module of PClamp (Fetchan 6.0). Events (positive deflections) shorter than 10 sec or smaller than 1.5 times the amplitude of spontaneous positive peaks before K+ stimulation were ignored to exclude artifacts.

Simultaneous measurement of [Ca2+]c and cell area with fura-2 and calcein
In simultaneous [Ca2+]c and cell area measurements, [Ca2+]c was monitored by fluorescence intensity of fura-PE3 at either 380- or 340-nm excitation wavelength, whereas calcein-AM was used for determination of cell area as described below. Cells were simultaneously loaded with fura-PE3-AM (2 µM, Teflabs, Austin, TX) and calcein-AM (10 µM, Sigma, St. Louis, MO) in cell culture medium for 30–60 min in a CO2 incubator at 37 C. After loading, cells were incubated further in dye-free extracellular solution at 30 C for 10–20 min. Excitation wavelength was altered every 1.5 sec between 494 nm (for calcein) and 380 or 340 nm (for fura). Fluorescence intensity was measured at 517 nm using an intensified charge-coupled device camera (IC-200, Photon Technology International, Lawrenceville, NJ) connected to an inverted microscope (Axiovert, Zeiss, Jena, Germany). In every 1.5 sec, eight images were acquired. These were averaged pairwise and the averages were summed (ImageMaster, Photon Technology International). Cell area was determined by calcein fluorescence using ImageJ (a Java-based version of NIH Image, http://rsb.info.nih.gov/ij/). A threshold level above background fluorescence was determined that corresponded to the phase-contrast image of the cell (Fig. 1AGo), and cell area was expressed as a number of pixels exhibiting fluorescence intensity above the threshold. Responsiveness of the method was confirmed by the rapid and sustained increase of pixel number by reducing the extracellular osmotic concentration from 290 to 250 mOsm (Fig. 1BGo).



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  		FIG. 1. Measurement of cell area with calcein using digital image technique. A, Fluorescence profile along the diameter of a single glomerulosa cell loaded with calcein. Threshold fluorescence level of cell area was determined by correspondence with the phase-contrast picture of the cell. Pixels with fluorescence below the threshold were regarded as background whereas pixels with fluorescence above the threshold were regarded as those belonging to the cell area. B, Changes in the pixel number corresponding to cell area in three cells in the same vision field. Osmolarity was reduced from 290 to 250 mOsm for 2 min. Pixel number was normalized to the average of the first minute of the recording.

 
In all experiments, the test solutions were applied by a gravity-driven perfusion system located at approximately 50 µm from the cells. Measurements were performed at 30 C. Results for each coverslip mean the average of two to four cells measured in one field of view. In some measurements, the fluorescence of both dyes changed slowly under control conditions. These recordings were fitted to the control 5-min period of the experiment.

Measurement of cell volume changes by calcein dequench technique
In some experiments, cell volume changes were monitored without measuring [Ca2+]c using the calcein dequench technique (31). This method is based upon the strong self-quenching ability of calcein when loaded in cells at high concentration. Increase of cell volume lowers the concentration of the dye resulting in weaker quenching and increase of the fluorescence. We found that this method was simpler and gave a better signal-to-noise ratio than the simultaneous [Ca2+]c-area measurement. In these measurements, the same experimental setup was used as for [Ca2+]c measurements performed with indo-1. Cells were loaded with 10 µM calcein-AM as described above. Excitation was performed at 494 nm, and the emitted fluorescence was measured at 517 nm.

Solutions
In [Ca2+]c measurements alone or combined with cell area determination, the control extracellular solution was (in mM): 87 NaCl, 3.6 KCl, 2 CaCl2, 0.5 MgCl2, 85 sucrose, 10 HEPES, and 11 glucose (pH 7.4, 295 mOsm). In those [Ca2+]c measurements where the effect of the cytoskeleton was tested, 50 mM sucrose was replaced by 25 mM NaCl. When [K+]e was increased by adding KCl, osmolarity of the control solution was also increased by adding N-methyl-D-glucamine-Cl at the same concentration.

Osmolarity of the extracellular solutions was modified by addition or removal of an appropriate amount of sucrose and checked by a freezing-point osmometer (MicroOsmometer 3MO, Advanced Instruments, Norwood, MA).

Cytochalasin-D (Sigma) was dissolved in dimethylsulfoxide, and colchicine (Sigma) was dissolved in ethanol. Cells were pretreated with 20 µM cytochalasin-D or 100 µM colchicine for 14–18 h. The concentration of dimethylsulfoxide or ethanol was 0.1%.

Statistical analysis
Data are expressed as means ± SEM. Statistical significance was estimated by Student’s paired t test, ANOVA, or ANOVA for repeated measures, using a statistical software (Statistica 4.5 or 5.1, Statsoft). Post hoc comparisons were performed by the Tukey‘s honest significant difference test. P < 0.05 was considered significant for all tests.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effect of [K+]e on [Ca2+]c and cell area
To address the question of whether cell volume changes could play a role in the action of physiological stimuli, we measured cell area as an indicator of cell volume simultaneously with [Ca2+]c during elevation of [K+]e. The effect of K+ on cell area was initially tested by elevation of [K+]e from the control 3.6 to 8.6 mM for 2 min. To maintain extracellular osmolarity constant, N-methyl-D-glucamine-Cl was added to the control solution at the same concentration. This increase in [K+]e evoked a slow and continuous increase in cell area in seven of eight experiments, which was usually detectable approximately 30 sec after starting the stimulation (Fig. 2AGo). Upon restoring the control K+ concentration, the increased cell area was maintained for an additional 1–3 min and thereafter returned to control level within approximately 5 min. Area changes exhibited longer lag times and smaller rates than changes in [Ca2+]c. Physiological elevation of [K+]e from 3.6 to 4.6 mM also resulted in an increase of cell volume in four of five experiments (Fig. 2BGo). Cell area changes by 4.6 mM K+ also followed the gradual development of the Ca2+ signal; however, the resolution of individual measurements did not allow reliable time-related separation of the two responses.



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  		FIG. 2. Effect of [K+]e on [Ca2+]c and cell area. Simultaneous measurement of [Ca2+]c and cell area was performed as described in Materials and Methods. Upper panels show normalized Ca2+ signals; lower panels represent relative cell area. [K+]e was elevated isosmotically from the control 3.6 to 8.6 mM (A) or 4.6 mM (B) (representative traces). Similar results were obtained in seven of eight and in four of five experiments for 8.6 and 4.6 mM K+, respectively. Fluorescence (F) indicating [Ca2+]c and pixel number indicating cell area (A) were normalized to the control period (F0 and A0, respectively).

 
Kinetics of the Ca2+ signal evoked by 5 mM K+ in single glomerulosa cells
In view of the potentiating effect of hyposmosis-induced swelling on T-type Ca2+ channels, we assumed that the K+-induced swelling plays a role in the high K+ sensitivity of glomerulosa cells. We therefore analyzed the kinetics of the Ca2+ signal elicited by a 2-min isosmotic elevation of [K+]e from 3.6 to 5 mM. The calcein dequench technique detected the expected increase in cell volume (Fig. 3AGo). [Ca2+]c increased from the resting level of 70 ± 5 to 184 ± 23 nM in 20–30 sec (n = 29). This steep initial rise was followed by an additional increase of [Ca2+]c. For kinetic analysis, the Ca2+ signals in each cell were normalized as follows. The value of [Ca2+]c was averaged for the period between the 20th and 30th sec after the onset of stimulation. The difference between this value and control [Ca2+]c was termed as first response (1 U). By the end of the 2-min stimulation [Ca2+]c further increased and reached 1.34-fold (SEM = ±0.08) of the first response (P < 0.05; Fig. 3BGo).



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  		FIG. 3. A, Effect of 5 mM K+ on cell volume measured by the calcein dequench technique (representative recording). Trace was normalized to the slow continuous decrease in calcein fluorescence. Isosmotic elevation of [K+]e from 3.6 to 5 mM increased fluorescence (indicating cell volume increase). The dequench technique was tested by increasing osmolarity to 320 mOsm (hyper). Similar results were obtained in eight of nine experiments performed on two preparations. B, Kinetics of the Ca2+ response to K+ in a population of glomerulosa cells (n = 29). [K+]e was increased from the control 3.6 to 5 mM for 2 min. Ca2+ response was normalized as described in Results.

 
The Ca2+ response of the single cells displayed various kinetic patterns (Fig. 4Go). On top of the rapid initial [Ca2+]c rise, a second rise began 30–60 sec after the onset of stimulation in several cells. This second rise was either persistent (cell 2, upper panel) or transient (cell 3, upper panel). In other cells [Ca2+]c either remained unaltered or increased slowly but continuously (cell 1, upper panel). For evaluation of the number of rising phases during the 2-min stimulation, the derivative of the [Ca2+]c curves was calculated and analyzed by single-channel analyzing software (see Materials and Methods). Of the 29 examined cells, 16 (55%) exhibited more than one phase of [Ca2+]c rise.



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  		FIG. 4. Kinetic pattern of the Ca2+ response in three single glomerulosa cells. [K+]e was elevated isosmotically from the control 3.6 to 5 mM for 2 min. Upper panels show representative [Ca2+]c traces. Lower panels demonstrate derivatives of the [Ca2+]c curves shown in the upper panels (generation of derivatives is described in Materials and Methods). Deflections larger than the threshold (see Materials and Methods) were regarded as events (one, two, and three events in cells 1, 2, and 3, respectively). Threshold is indicated by vertical line on the right side of each trace.

 
Prevention of swelling attenuates the second phase of the rise in [Ca2+]c
To test whether the second, slow phase of increase in [Ca2+]c was indeed a consequence of cell swelling evoked by K+, cells were stimulated with 5 mM K+ under hyperosmotic conditions. In preliminary experiments, applying an iterating approach, we found that raising extracellular osmolarity to 320 mOsm largely compensates cell swelling caused by 5 mM K+ (Fig. 5AGo). In accordance with previous observation (8), hyperosmotic [K+]e elevation did not affect the first phase of Ca2+ response. In isosmotic control experiments, [Ca2+]c increased from 67 ± 7 to 151 ± 21 nM (n = 13). When cell swelling was opposed with hyperosmosis, [Ca2+]c increased from 75 ± 7 to 154 ± 19 nM (n = 11). However, the second, slow rise of [Ca2+]c was markedly inhibited under hyperosmotic conditions. In control cells, the Ca2+ response at the end of the 2-min stimulation attained 1.44-fold (±0.10) of the first response (n = 14, data not shown). In contrast, in cells stimulated under hyperosmotic conditions, the [Ca2+]c measured at the end of the 2-min stimulation did not differ from that measured at the first phase (i.e. 20–30 sec after the onset of stimulation) (1.01 ± 0.07-fold change; n = 11; Fig 5BGo).



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  		FIG. 5. Effect of hyperosmotic [K+]e elevation on the kinetics of the Ca2+ response. A, Simultaneous elevation of [K+]e (from 3.6 to 5 mM) and osmolarity (from 290 to 320 mOsm) largely prevented cell volume increase compared with the response evoked by isosmotic elevation of [K+]e. B, Kinetics of the Ca2+ response to hyperosmotic elevation of [K+]e in a population of glomerulosa cells (n = 11). [K+]e was increased from 3.6 to 5 mM for 2 min simultaneously with elevation of osmolarity from 290 to 320 mOsm. Ca2+ response was normalized as described in Results.

 
Effect of cytoskeleton-disrupting drugs on the swelling-induced increase of the K+-evoked Ca2+ response
Because elements of the cytoskeleton network have been shown to modulate the function of several voltage-gated ion currents (21), we examined the involvement of the cytoskeleton in the transmission of cell volume changes to the voltage-gated Ca2+ channels in rat glomerulosa cells. The cells were stimulated repeatedly for 20 sec with 6 mM K+. After recording a Ca2+ signal in the control isosmotic (290 mOsm) solution, the stimulation was repeated in hyposmotic (250 mOsm) solution. Finally, the stimulation was applied under restored isosmotic conditions. Cells were either untreated or preincubated for 14–18 h with 20 µM cytochalasin-D (to disrupt actin filaments) or with 100 µM colchicine (to disrupt the microtubule network). In untreated cells, elevation of [K+]e from 3.6 to 6 mM raised [Ca2+]c from the control 43 ± 9 nM by 89 ± 25 nM (Fig. 6AGo; n = 9). Similarly to that reported previously (8), the Ca2+ signal evoked subsequently by 6 mM K+ under hyposmotic conditions was augmented (174 ± 44 nM; 2.25 ± 0.42-fold increase of Ca2+ response), and the effect was reversible upon restoration of osmolarity. The effect of hyposmosis on the amplitude of the K+-evoked Ca2+ signals was significant (P < 0.01, Figs. 6AGo and 7Go). Although control [Ca2+]c in cytochalasin-D- or colchicine-treated cells did not differ from untreated cells, both drugs reduced slightly (but not significantly) the K+-induced Ca2+ signal [cytochalasin-D, 47 ± 10 nM (n = 6); colchicine, 37 ± 8 nM (n = 4)]. However, the amplifying effect of hyposmosis on the K+-evoked Ca2+ signal was unaffected by the cytoskeletal drugs; in 250 mOsm solution the K+-evoked Ca2+ signal increased to 3.02 ± 0.65-fold of the control (isosmotic) response in cytochalasin-treated cells (n = 6; Figs. 6BGo and 7Go) and 3.36 ± 1.41-fold in colchicine-treated cells (n = 4; Figs. 6CGo and 7Go).



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  		FIG. 6. Effect of cytoskeletal agents on the osmosensitivity of the K+-evoked Ca2+ response in single glomerulosa cells. Elevations of [K+]e from the control 3.6 to 6 mM and reduction of osmolarity from the control 290 to 250 mOsm are represented by horizontal bars on each panel. Cells pretreated with 20 µM cytochalasin-D (B) or 100 µM colchicine (C) were compared with control cells (A) from the same preparations. Representative for nine (control), six (cytochalasin-D), and four (colchicine) experiments measured in three preparations.

 


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  		FIG. 7. Effect of cytoskeletal agents on the osmosensitivity of the K+-evoked Ca2+ signal. Ca2+ responses were related to the first isosmotic response. The columns and bars show the means +SEM of the experiments, which were representatively shown in Fig. 6Go.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study we report that elevation of [K+]e in the 4.6–8.6 mM range induces cell swelling in rat adrenal glomerulosa cells. The volume increase augments the Ca2+ response to K+ possibly by enhancing Ca2+ influx through T (and L) type voltage-gated Ca2+ channels and therefore contributes to the high K+ sensitivity of the glomerulosa cell.

Alterations of cell volume affect a number of cellular functions. Cells swollen or shrunken in anisosmotic media often respond with activation of appropriate transport mechanisms to normalize their volume by uptake or release of net osmotic active molecules (mostly ions) and water (12, 13). This process may intervene disadvantageously with cell function by distorting normal intracellular ion concentrations; however, alterations of cell volume were also proposed to act as signal-transmitting mechanisms in several cell types. For example, cell volume changes brought about by absorption or secretion of osmotically active molecules in epithelial cells may couple transport activity of the apical and basolateral membranes (13); swelling induced by insulin plays a role in the metabolic response of hepatocytes to this hormone (16, 32), and anion channels regulated by volume changes of astrocytes in the hypothalamus may contribute to the accommodation of vasopressin release to changes of plasma osmolarity (33). Vesicular peptide hormone secretion by several endocrine cell types is also stimulated by hyposmotic swelling (34).

The effect of cell volume on steroid hormone secretion is less well explored. In a previous study we investigated the effect of extracellular osmolarity on Ca2+ homeostasis and aldosterone production in cultured rat glomerulosa cells, and we showed that cell swelling in hyposmotic solution (250 mOsm) potentiates the Ca2+ response and aldosterone production evoked by elevation of [K+]e from 3.6 to 5 mM. This result was similar to that measured by Wang et al. (11) in bovine glomerulosa cell suspension using supraphysiological (7–13 mM) K+ concentrations. Nevertheless, there was a difference in the two studies; whereas we found no effect of osmolarity on basal [Ca2+]c and aldosterone production, Wang et al. reported an increase in both parameters by hyposmosis. Although this could be explained by differences in species and cell preparations, we found that hyposmosis was capable of evoking a small and slow increase in [Ca2+]c in rat glomerulosa cells as well, when the cells were incubated in bicarbonate- and not in HEPES-buffered solution (Makara, J. K., G. L. Petheo, and A. Spät, unpublished observation). Nevertheless, in the present experimental approach the application of HEPES rather than bicarbonate was advantageous for analysis of the Ca2+ response to small increases in [K+]e under different osmotic conditions, because basal [Ca2+]c remained unaffected.

The stimulatory effect of hyposmotic swelling on the Ca2+ signal evoked by 5 mM K+ could well be explained by the observed increase of the T-type current amplitude (8). This increase in current intensity may be due to a shift of activation threshold to more negative Em values, a shift of steady-state inactivation curve to more positive membrane potential (Em) values, a change in activation and inactivation kinetics, etc. Although presently we cannot offer any well-established mechanism for the phenomenon, a hyposmosis-induced (250 mOsm) shift of the current-voltage curve (activated by depolarizing ramps from -100 to + 40 mV) to more negative potentials supports our previous data on the swelling-evoked increase in T-type current amplitude (Makara, J. K., G. L. Petheo, and A. Spät, unpublished observation). Hormonal regulation of the T-type Ca2+ channels has been studied extensively in several cell types (for review see Ref.35). A number of G protein-coupled receptors were shown to influence the current by shifting the activation curve to positive or negative directions. In bovine glomerulosa cells Ca2+ signal evokes a calmodulin-dependent protein kinase II (CaMKII)-mediated shift of the voltage dependence of the T-type Ca2+ current to more negative values (36, 37). The present observation showing that prevention of cell swelling during exposure to K+ prevents the development of the second phase of Ca2+ signal suggests that cell swelling rather than Ca2+-induced kinase action is required for sensitizing rat glomerulosa cells to physiological increases in extracellular [K+]e.

Elements of the cytoskeleton were shown to interact with the function of several channel types, including voltage-gated Ca2+ channels (21). Disruption or stabilization of the actin filament network (17, 18) and the microtubular system (38, 39) influenced the kinetic properties of high-voltage-activated Ca2+ channels, but we are not aware of such a role of the cytoskeleton in the regulation of T-type channel. In the present study cytochalasin-D and colchicine treatments evoked only a slight but not significant reduction of the Ca2+ response to 6 mM K+ under isosmotic conditions, and they exerted no effect on the hyposmosis-induced potentiation of the K+-evoked Ca2+ signals. This suggests that in glomerulosa cells swelling influences the T-type channel through a mechanism other than the cytoskeleton. A possible small contribution of L-type channels to the Ca2+ influx evoked by 6 mM K+ in rat glomerulosa cells (10) does not invalidate this conclusion.

Elevation of [K+]e was shown to induce cell swelling in a few cell types (23, 25, 27); however, the applied K+ concentration was very high (several tens of a millimole), which made the physiological relevance of this phenomenon questionable. In early studies moderate yet still supraphysiological K+ concentrations (8.4 mM) did not increase water content of freshly isolated rat glomerulosa cells (40). Similar K+ concentrations (7–10 mM) induced sustained swelling in bovine glomerulosa cells, and this swelling was dependent on [Cl-]e (11, 41). The present report is the first to show that even small physiological elevation of [K+]e is sufficient to cause cell swelling in glomerulosa cells and that this volume increase has considerable impact on Ca2+ signaling. Several pathways were proposed to mediate K+-induced swelling in other cell types, including K+ and Cl- conductances as well as the Na+-K+-2Cl-, Na+-Cl-, and K+-Cl- cotransporters (24, 26, 27, 28, 29). Apart from K+ channels, the presence of such transport systems in glomerulosa cells is almost completely unexplored. Nevertheless, it should be recalled that irrespective of their uptake mechanism, solutes accumulated by the cell induce cell swelling even under hyperosmotic conditions. Elevation of extracellular [K+] results in a net K+ gain of the cell, which may be followed additionally by a secondary accumulation of anions. Chloride, as the most abundant extracellular anion, is a likely candidate to be involved in the K+-induced swelling. However, very little is known about the Cl- homeostasis of glomerulosa cells. Recently, an ACTH-activated outwardly rectifying Cl- current was described in rat glomerulosa cells (42), but its relation to volume regulation has not yet been investigated.

When we measured cell volume together with [Ca2+]c, the Ca2+ signal evoked by elevation of [K+]e always preceded the volume increase. This is not surprising, as swelling is presumably a slower process than depolarization. In this case the volume change is expected to influence the Ca2+ signal with latency. Analyzing the Ca2+ response of single glomerulosa cells to 5 mM K+, we found that the first, rapid peak (achieved in 20–30 sec after the onset of stimulation) was followed by an additional slow increase of [Ca2+]c. The second rise in [Ca2+]c could be prevented by applying appropriate hyperosmosis together with K+, suggesting that the second rise was a consequence of swelling. Derivation of the [Ca2+]c curves revealed a significant cell-to-cell heterogeneity, in harmony with previous observations on rat glomerulosa cells exposed to slightly elevated [K+]e (43, 44). It should, however, be emphasized that half of the cells exhibited more than a single [Ca2+]c rising phase during the 2-min elevation of [K+]e.

Increased secretion of aldosterone in response to a small elevation of [K+]e is a vitally significant function of glomerulosa cells. Here we presented data showing that even physiological elevations of [K+]e induce an increase in cell volume that, in turn, may increase the intensity of voltage-activated Ca2+ influx and further augment the K+-evoked [Ca2+]c increase. We propose that this cell volume-mediated potentiation of Ca2+ signaling contributes to the aldosterone secretory effect of physiological K+ stimuli.


   Acknowledgments
 
The excellent technical assistance of Ms. Anikó Rajki, Ms. Erzsébet Horváth, and medical student Kata Wágner is highly appreciated.


   Footnotes
 
This work was supported by grants from the Hungarian Science Foundation (OTKA TS 040865), the Hungarian Council for Medical Research (ETT 28/2000), and the National Research and Development Program (NKFP-1/044/2001).

Received March 26, 2003.

Accepted for publication July 21, 2003.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Spät A, Enyedi P, Hajnóczky Gy, Hunyady L 1991 Generation and role of calcium signal in adrenal glomerulosa cells. Exp Physiol 76:859–885[Medline]
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A. SPAT and L. HUNYADY
Control of Aldosterone Secretion: A Model for Convergence in Cellular Signaling Pathways
Physiol Rev, April 1, 2004; 84(2): 489 - 539.
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