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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS
Department of Anatomy and Cell Biology, Hebrew University, Hadassah Medical School, Jerusalem, Israel
Submitted 14 March 2005 ; accepted in final form 31 August 2005
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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L-type Ca2+ channels; osmosensitivity; mechanosensitivity; osmolarity; hyperosmolarity
-cells (12), and hippocampal neurons (54). Therefore, it appears that voltage-gated Ca2+ channels in pituitary cells and other excitable cells are osmosensitive. This osmosensitivity of Ca2+ channels might reflect sensitivity to mechanical stress that is produced by cell shrinkage or swelling. Indeed, it was suggested that Ca2+ channels in pituitary cells (2, 44), in smooth muscle cells (30), and in cardiac myocytes (34), are mechanosensitive. In addition, it has been demonstrated that recombinant L-type (32) and N-type (6) Ca2+ channels are sensitive to membrane stretch, and that both native and recombinant glutamate receptors are mechanosensitive (8). Because voltage-gated Ca2+ channels play a key role in regulating the secretion of pituitary hormones (26, 59) it was of interest to investigate whether or not hyperosmotic induced cell shrinkage modulates directly voltage-gated Ca2+ currents in pituitary cells. Both L-type and T-type Ca2+ currents (IL and IT, respectively) were observed in anterior pituitary cells (2, 9, 11, 40, 59). In a previous study (35), we have demonstrated that hypertonicity decreases Ca2+ influx through L-type and T-type channels in anterior pituitary cells. In this study, we further characterized the hyperosmotic effects on IL and IT. We show that the hyperosmotic suppression of IL and IT is differential (IL is more sensitive than IT), that the hyperosmotic suppression of IL is correlated with pituitary cell shrinkage and that the hyperosmotic suppression of IL and IT is not dependent on the integrity of the actin cytoskeleton. Furthermore, we show that the hyperosmotic suppression of IL stems from reduction in the activity of Ca2+ channels rather than from a decrease in single channel conductance. This hyperosmotic suppression of Ca2+ currents may contribute to the previously reported hyperosmotic suppression of hormone secretion from pituitary cells. The possible cellular mechanisms underlying these effects and their possible functional significance are discussed.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Actin cytoskeleton imaging. The actin cytoskeleton in pituitary cells was labeled with fluorescein isothiocyanate (FITC)-phalloidin as previously described (28). The staining procedure was performed at room temperature, in petri dishes containing pituitary cells attached to glass coverslips (see above). Shortly thereafter, pituitary cells were fixed with 4% paraformaldehyde (Fluka) in PBS for 40 min, and then permeabilized with 0.1% Triton X-100 (Baker) in PBS for 5 min. Nonspecific binding was then blocked by the addition of PBS containing 1% BSA (10 min) to the cells. Afterward, the pituitary cells were incubated with 13 µM FITC-phalloidin (Sigma) for 40 min, washed extensively (4x) with PBS, mounted with an antifade medium (Agar Scientific) to prevent rapid photobleaching, and then examined with an Olympus BX51 fluorescence microscope (x100 oil-immersion objective). Color images were captured using an Olympus DP70 digital camera. The involvement of the actin cytoskeleton in the hyperosmotic effects was examined with cytochalasin D (Cyto D; Sigma) and phalloidin (Sigma). Stocks were prepared in DMSO and kept at –20°C. Aliquots containing Cyto D or phalloidin were dissolved in the physiological solution before the start of experiments. In several control experiments, we found that final DMSO concentrations (0.001%-0.1%) were not affecting our results (see Fig. 6C).
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, when filled with the "patch pipette solutions" (see below). Membrane currents were recorded at room temperature (20–24oC), sampled with an analog-to-digital converter (TL-1 DMA interface or Digidata 1320A, Axon Instruments) at 5 kHz, filtered with a four-pole low-pass Bessel filter with a cut-off frequency (–3 dB) of 1 kHz and stored in the hard drive of an IBM-based computer. Capacitive currents and access resistances (Ra) were electronically compensated with the potentiometers provided with the amplifier. Final access resistance was usually <15 M. Linear leak currents (and residual capacitive currents) were digitally subtracted after extrapolation of averaged leak currents that were obtained in response to P/2 or P/4 pulse protocols. The pCLAMP6 or pCLAMP8 programs (Axon Instruments) were used for on-line acquisition and for off-line analysis of the membrane currents. IBa through L-type and T-type channels were usually activated with 200-ms voltage steps (interval 10–15 s) from a holding potential (Vh) of –80 mV to various test potentials (Vt). In some of the experiments double-pulse protocols were used to activate simultaneously T-type currents (Vt = –30 mV) and L-type currents (Vt = 0 mV). To obtain instantaneous I-V relationships of Ca2+ channel currents we have used 600-ms voltage ramps ranging from –100 to +80 mV.
For simultaneous monitoring of IBa, Cm, and Ra we used double-pulse protocols. The first subthreshold pulse, a 10-ms depolarization from –80 to –70 mV, activated capacitive currents. The second pulse, a 20-ms depolarization from –80 to 0 mV, activated IBa. In these experiments, currents were sampled at a rate of 50 or 100 kHz (filter 10 kHz and cell membrane capacitive currents were not compensated electronically). The uncompensated membrane capacitive currents were used to calculate the values of Cm and Ra by using the relation 
= Ra·Cm, where is the time constant of the decay of the capacitive currents (31). The was calculated by fitting a monoexponential function to the decay of the capacitive currents. Cm was calculated from the relation C = Q/V, where Q is the amount of charge needed to discharge the cell membrane (obtained by integrating the capacitive current) and V is the amplitude of the subthreshold pulse. Ra was calculated from the relation Ra = /Cm. Calculated Cm values may be affected by alterations in electrode capacitance due to changes in bath level. However, electrode capacitive currents were consistently compensated in all the experiments. In addition, Cm values stayed constant throughout long perfusion experiments, suggesting that changes in electrode capacitance were not a major factor affecting Cm values.
Single channel Ca2+ currents were recorded with the cell-attached mode of the patch-clamp technique. Single channel currents were activated by voltage steps from a Vh of –80 mV to various test potentials (pulse duration 200 or 340 ms, interval 15 s), sampled at a rate of 5 kHz, and filtered at 0.5–1 kHz. To clamp the membrane patch at –80 mV in cell-attached recording, the patch pipette was held at +80 mV. Recordings were always obtained from multi-Ca2+ channel patches. To examine the hyperosmotic effects on these multichannel patches single traces were leak subtracted and analyzed (pCLAMP 6 or 8) as follows: 1) individual current traces in control and during hyperosmotic stimuli were averaged (ensemble currents) and compared. 2) NPo was used as a measure of Ca2+ channel activity (where N is the number of channels in the patch and Po is the probability of Ca2+ channels to be in the open state). The value of NPo for each current trace in the experiment was calculated from the ratio I/i (where I is the average current of this trace and i is the corresponding single channel current amplitude). 3) All point amplitude histograms (APAH). These histograms were constructed from data points in control, hyperosmotic stimulus, and fitted with a Gaussian fit. From the peaks of these histograms, representing closed times and the number of channels in the patch, we usually monitored changes in i and changes in closed probability (Pc).
Statistical differences among groups of tested parameters were examined either by paired or by unpaired t-tests. Multiple-comparison tests were performed by one-way ANOVA. When significant differences were indicated in the F-test (P < 0.001), the significance of differences between the means of any of these groups was determined by the Tukey method for multiple comparisons with 
= 0.05 (see Fig. 4). Results are always reported as means ± SE.
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For cell-attached recordings of single channel currents the extracellular solution contained (in mM) 140 K-aspartate, 2 MgCl2, 5 EGTA, 10 glucose, and 10 HEPES [adjusted to pH 7.3 with K(OH)]. The patch-pipette solution contained (in mM) 100 BaCl2, 15 TEA-Cl, and 10 HEPES. BAY K 8644 or FPL 64176 (3–10 µM) were added to the bath and patch pipette solutions in all the single channel experiments. [Os]e in these single channel experiments was increased as described above, by adding mannitol to the bath solution. All chemicals for these extracellular and intracellular solutions were purchased from Sigma, except QX-314 (Alomone Labs, Jerusalem, Israel), which was used to block Na+ currents.
Before each experiment coverslips containing pituitary cells were placed in a perfusion chamber (RC-16, Warner Instruments). Cells were exposed to control (isosmotic) or hyperosmotic solutions by perfusing the chamber at a rate of 
1 ml/min. The volume of solution in the perfusion chamber was 0.4 ml and the cells were exposed to the different experimental solutions for 2–4 min.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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40% and 20%, respectively. Similar results were obtained in additional double pulse experiments. In these experiments, 63% increase in [Os]e suppressed IL by 49 ± 3% and IT by 30 ± 5% (n = 4) (P < 0.02, paired t-test). This differential sensitivity was also observed when we compared IL and IT that were recorded from different cells, in response to different hyperosmotic stimuli, as illustrated in Fig. 1C.
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60 mV that was not changed during the hyperosmotic stimulus. Similar results were obtained in additional experiments, in response to 30% increase in [Os]e. V0.5 negatively shifted by 1.6 ± 0.3 mV (P < 0.0001, paired t-test, n = 17) and Vpeak negatively shifted by 2.2 ± 0.4 mV (P < 0.0001, paired t-test, n = 17). However, Er in control (67 ± 2 mV) was not different from Er during hyperosmotic challenges (66 ± 5 mV), (P = 0.64, paired t-test, n = 8). Thus our results demonstrate a small, but significant, negative voltage shift during hyperosmotic challenges. Thus it cannot be argued that the hyperosmotic suppression of IL results from a depolarizing shift in the activation of the IL, or from changes in the reversal potential of the currents.
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40%, without any significant effects on the capacitive current, suggesting that changes in Ra and Cm are not responsible for the decrease in IL. Similar experiments were performed in additional cells. Figure 4, B and C, summarizes 14 experiments in which pituitary cells were challenged with 30% increase [Os]e for 150–225 s. The hyperosmotic suppression of IL in these experiments (by 35 ± 3%, n = 14) was not accompanied with significant changes in Cm (Fig. 4B), but was associated with 30% increases in Ra (Fig. 4C), from 14 ± 2 to 20 ± 4 M (P < 0.004, paired t-test, n = 14). This raises the possibility that increase in Ra underlies (or contributes) to the hyperosmotic suppression of IBa. However, closer inspection of Fig. 4C, right panel, reveals that this is not the case. First, there is no correlation between the hyperosmotic suppressions of IL and their corresponding increases in Ra. Second, in at least 4 out of these 14 experiments the hyperosmotic suppression of IL was not accompanied with a significant increase in Ra.
Hyperosmotic suppression of IL and IT: independence on the actin cytoskeleton.
The hyperosmotic suppression of IBa may have resulted from decrease in membrane tension that is transferred to the channel proteins via the cytoskeleton. The involvement of the actin cytoskeleton in the hyperosmotic effects on IL and IT was examined by incubating pituitary cells either with the actin cytoskeletal disrupter Cyto D or with the actin cytoskeletal stabilizer phalloidin. The submembranal actin cytoskeleton (cortical actin) in pituitary cells is demonstrated in Fig. 5A with the use of phalloidin-FITC staining. Figure 5B shows that incubation of pituitary cells with 10 µM Cyto D (for 15 min) resulted with disruption of cortical actin and with typical appearance of actin clusters inside the cell (verified with confocal microscopy, not shown). Figure 5C shows that this pattern of actin filament disruption persisted 60 min after the end of incubation with 20 µM Cyto D (for 30 min). Similar patterns of actin disruption were observed in nine different pituitary cell preparations. We therefore examined hyperosmotic effects on IL and IT shortly (15 min) after preincubations with 20 µM Cyto D (for 30 or 60 min). One of these experiments is illustrated in Fig. 6, A and B. The hyperosmotic suppression of IL and IT persisted after preincubation with 20 µM Cyto D (for 30 min). An increase in [Os]e by 37% suppressed IL and IT by 42% and 16%, respectively. Similar effects were observed after longer preincubations with 20 µM Cyto D (for 60 min). Figure 6C compares similar preincubation experiments with 20 µM Cyto D, 12.7 µM phalloidin, and 0.1% DMSO. Increase in [Os]e by 37% suppressed IL by 34 ± 2% (n = 9), 40 ± 3% (n = 7), and 38 ± 6% (n = 6), for Cyto D, phalloidin, and DMSO, respectively. The persistence of hyperosmotic effects and the lack of difference between the effects, after preincubation with these three chemical agents, suggest that the actin cytoskeleton is not involved in hyperosmotic suppression of Ca2+ currents.
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). These possibilities were investigated by recoding the activity of L-type channels in the cell-attached mode, while exposing pituitary cells, outside the patch pipette, to hyperosmotic media.
Hyperosmotic effects on activity of L-type Ca2+ channels.
Figure 7 shows that 63% increase in [Os]e results with reversible suppression of Ca2+ channel activity in a multichannel patch. This reduction in L-type Ca2+ channel activity is also manifested as a reduction in the amplitude of ensemble Ca2+ channel currents, as illustrated in Fig. 7B. These effects of hyperosmolarity on Ca2+ channel activity were quantitatively estimated by using two additional methods of analysis; by calculating values of NPo, as a measure of L-type Ca2+ channel activity, and by constructing APAH (see MATERIALS AND METHODS). The results of this analysis, for the experiment shown in Fig. 7, are illustrated in Fig. 8. Figure 8A illustrates that 63% increase in [Os]e reversibly decreased NPo values by 50%. This decrease in the activity of L-type channels was also manifested as an increase in the proportion of Pc (by 50%) in the APAH of the same experiment, as illustrated in Fig. 8B. Similar results (63% increase in [Os]e), were obtained in additional cells. NPo values were reduced by 45% from 0.82 ± 0.10 to 0.44 ± 0.09 (P < 0.001, paired t-test, n = 9), and Pc values were increased by 60% from 0.43 ± 0.06 to 0.69 ± 0.07 (P < 0.001, paired t-test, n = 9). Hence, these experiments demonstrate that exposing pituitary cells to hyperosmotic media reduced the activity of L-type Ca2+ channels inside the cell-attached membrane patch.
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), as shown in Fig. 9A. in control (23.8 pS) was not different from during hyperosmotic challenges (63% increase in [Os]e, 25 pS). The small increase in the amplitude of single channel currents at each one of the voltage steps, observed during exposure to the hyperosmotic challenge, may stem from a 4 mV hyperpolarizing shift across the membrane patch. This hyperpolarization by itself may reduce the activity of L-type Ca2+ channels. However, from the activity curve (NPo voltage relationship) of this experiment (Fig. 9B), it is clear that a hyperpolarization of 4 mV cannot explain the strong hyperosmotic suppression of Ca2+ channel activity. Similar results were obtained from averaged I-V curves of single channel currents (n = 16; Fig. 10A); in control (27.3 pS) was not different from during hyperosmotic challenges (26.8 pS). A small positive voltage shift was also observed in these averaged I-V (2 mV). However, from the activity curve that was constructed from the same experiments (Fig. 10B), it is clear that this hyperpolarizing shift cannot explain the hyperosmotic suppression of L-type channels. The reason for this small voltage shift is not clear, however, it may stem from hyperosmotic effects on membrane potential outside the membrane patch. Thus it appears that the hyperosmotic suppression of whole cell Ca2+ channel currents results from reduction in NPo and not from reduction in single channel conductance.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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To assess the biophysical basis for these hyperosmotic effects on whole cell Ca2+ channel currents we recorded the activity of L-type Ca2+ channels from cell-attached patches, while exposing the cell, outside the patch pipette, to hyperosmotic media. Our results show that increase in [Os]e suppressed NPo and increased Pc (Figs. 7 and 8) without having any significant changes in single channel conductance (Figs. 9 and 10). The suppression in NPo was similar in magnitude to the suppression of whole cell currents; 63% increase in [Os]e suppressed both IL and NPo by 40–50%. This similarity suggests that the mechanisms underlying hyperosmotic suppression of whole cell and cell-attached currents are similar, despite the different configurations of recordings and despite the different modes of exposure to hyperosmotic media. Moreover, the similarity in responses may suggest that our cell-attached patches were similar in morphology to the whole cell membranes, and that our cell-attached membrane patches were not lipid blebs devoid of cortical cytoskeleton (15, 36, 64). It has been shown that uncoupling of cortical actin from the plasma membrane, and the formation of lipid blebs during tight seal cell-attached recordings, underlies the discrepancy between the activities of mechano-gated ion channels under cell-attached and whole cell recordings (16, 37, 60, 65).
The results of this study show that decrease in Ca2+ channel activity underlies the hyperosmotic suppression of whole cell Ca2+ currents. This is the first study, to the best of our knowledge, which compares hyperosmotic effects both at the whole cell and single channel level. The decrease in Ca2+ channel activity may result either from a decrease in N or in Po. The prominent suppression of maximal IBa in contrast to the lack of prominent effects on V0.5 (Fig. 3) might suggest that N rather than Po is affected by hyperosmotic stress, as proposed for the effects of membrane stretch on N-type channels (6). On the other hand, the clear increase in Pc (Fig. 6) might suggest that Po rather than N is affected by hyperosmotic stress. This increase in Pc cannot be attributed to changes in the efficacy of BAY K 8644, due to osmotic alterations in intracellular ionic strength. BAY K 8644 was added extracellularly and it was shown that dihydropyridine (DHP) agonists gain access to their receptor site from the outer surface of the cell membrane (18, 57). Additional experiments are needed to resolve this question as to whether N or Po is affected by osmotic stress.
Sensitivity of Ca2+ channels to osmotic cell shrinkage: possible cellular mechanisms.
The present study shows that the hyperosmotic suppression of IL depends on the percent increase in [Os]e (Fig. 2A) and is highly correlated with the percent decrease in pituitary cell volume (Fig. 2B). Hence voltage-gated Ca2+ influx in pituitary cells can be suppressed by pituitary cell shrinkage. This osmosensitivity may result either from alterations in mechanical membrane tension or from alterations in ionic strength underneath the plasma membrane. Changes in membrane tension may be conveyed to the channel protein either through the cytoskeleton of the cell or through the phospholipid bilayer (14, 46). An increasing number of studies point to a functional link between the actin cytoskeleton and L-type Ca2+ channels (28, 38, 45, 47, 52). Moreover, it was shown that hyposmotic-induced increase in IL in smooth muscle cells was abolished by the actin microfilament disrupter Cyto D, and augmented by the actin microfilament stabilizer phalloidin (63). Similar effects were reported for T-type Ca2+ currents in cardiac myocytes (42). However, in contrast to these findings, we demonstrate here that the hyperosmotic suppression of IL and IT persisted after disruption of the actin cytoskeleton with Cyto D or stabilizing it with phalloidin (Figs. 5 and 6). Hence, our results suggest that the actin cytoskeleton does not play a significant role in the hyperosmotic suppression of Ca2+ currents in pituitary cells. This conclusion is based on the assumption that cortical actin is not washed away under conditions of whole cell recordings (60, 65). In this context, it is interesting that shear-stress effects on L-type Ca2+ currents (but not on sodium currents) persisted after disruption of the actin cytoskeleton in smooth muscle cells (56). In addition, shear-stress effects on recombinant L-type channels persisted after truncation of the 1c COOH terminus, a putative site for interaction of Ca2+ channels with the cytoskeleton (32). These findings are in support to the notion that cortical actin cytoskeleton is not the key player in mechanical modulation of L-type channels. It is therefore tempting to speculate that decrease in membrane tension is conveyed directly to the Ca2+ channel proteins through the phospholipid bilayer. It is well established that increased tension in the lipid bilayer activates mechano-gated ion channels in both prokaryotic and eukaryotic cells (14). Previous studies (8) have shown by using amphiphilic compounds that tension in the phospholipid bilayer underlies the mechanosensitivity of glutamate receptor in mouse central neurons. Similarly, it has been shown that tension in the phospholipid bilayer underlies the mechanosensitivity of two-pore domain K channels, TREK and TRAAK (25, 43).
It is also possible that the osmosensitivity of whole cell Ca2+ currents results from local alterations in ionic strength underneath the plasma membrane. Hyperosmotic shrinkage may be associated with efflux of water molecules and, as a result, with local increases in ion concentration underneath the plasma membrane, despite the constant ionic composition of the intracellular solution. It is possible that these local increases in ionic strength underneath the plasma membrane suppress in some way the activity of Ca2+ channels and as a result reduce whole cell Ca2+ influx. This ionic-strength hypothesis is also supported by our single channel results. Hyperosmotic induced cell shrinkage, during cell-attached recordings, is expected to increase ionic strength all over the cytoplasm of the intact cell, including the cytoplasm underneath the cell-attached membrane patch. Some indirect support for this ionic-strength hypothesis comes also from studies that demonstrated that decrease in extracellular ionic strength facilitates Ca2+ influx in hippocampal neurons (5, 54). In addition, it was shown that decrease in intracellular ionic strength triggers the activation of volume regulated anion channels (7, 39), and it was proposed that dehydration of the open-channel volume underlies the hyperosmotic suppression of potassium channels in squid giant axons (66).
Sensitivity of Ca2+ channels to osmotic cell shrinkage: functional significance. The findings of this study demonstrating hyperosmotic suppression of IL and IT are in agreement with previous studies demonstrating hyperosmotic suppression of stimulated hormone secretion from pituitary cells (49, 61). The relevance of these findings to hormone secretion is not clear because pituitary cells, like most mammalian cells, experience very small alterations in [Os]e under normal physiological conditions. However, alterations in intracellular osmolarity ([Os]i), due to changes in the metabolic state of pituitary cells, may occur under normal physiological conditions (29). It is plausible that such putative alterations in [Os]i will cause continuous alterations in pituitary cell volume under normal physiological conditions, thereby affecting Ca2+ influx and hormone secretion.
Another interesting possibility is that Ca2+ channels sense alterations in membrane tension during the process of exocytosis. A substantial decrease in membrane tension during stimulated secretion occurs in rat basophilic leukemia cells (10) and possibly occurs also in bovine chromaffin cells (24). This decrease in membrane tension, similar to hyperosmotic-induced pituitary cell shrinkage, may reduce voltage-sensitive Ca2+ influx and thereby hormone secretion. Hence, it is possible that the hyperosmotic suppression of voltage-gated Ca2+ influx reflects an autoregulatory mechanism for hormone secretion. Similarly, it has been suggested that the sensitivity of L-type channels in smooth muscle cells to shear stress and membrane stretch plays an autoregulatory role in intestinal contractility (19).
Acute increases in plasma osmolarity may occur under certain pathophysiological conditions such as severe dehydration or hyperglycemia in diabetes mellitus. For example, in nonketotic hyperosmolar coma, a severe form of hyperglycemia in diabetic patients, plasma osmolarity can be elevated to 350–450 osmol/l (1, 55), similar to the hyperosmotic stimuli used in this study. It has been demonstrated that similar increases in [Os]e result with suppression of excitability and synaptic transmission in central nervous system neuron (21, 48). It is therefore possible that hyperosmotic suppression of voltage-gated Ca2+ influx plays a role in the neurological symptoms caused by hyperosmolarity (1, 55).
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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) and during (
) exposure to 30% increase in [Os]e. B: I-V curves in A were normalized with maximal current (Imax). Note that the hyperosmotic effect was accompanied with a small negative shift in V0.5 and 1 Vpeak by –2.3 and –4.5 mV, respectively. C: I-V curves that were generated by voltage ramps before and during exposure to 30% increase in [Os]e. D: I-V curves in C were normalized with Imax. Note that the hyperosmotic effect was accompanied with a small negative shift in V0.5 and Vpeak by –4 and –3 mV, respectively. Note also that the hyperosmotic effects were not accompanied with changes in reversal potential (C and D).
