January 2009
Volume 50, Issue 1
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Glaucoma  |   January 2009
Activation of Stretch-Activated Channels and Maxi-K+ Channels by Membrane Stress of Human Lamina Cribrosa Cells
Author Affiliations
  • Mustapha Irnaten
    From the Molecular Medicine Laboratories, RCSI Education and Research Centre, Beaumont Hospital, Dublin, Ireland;
  • Richard C. Barry
    From the Molecular Medicine Laboratories, RCSI Education and Research Centre, Beaumont Hospital, Dublin, Ireland;
    Ophthalmology, Mater Misericordiae University Hospital and Conway Institute, University College Dublin, Dublin, Ireland; and
  • Barry Quill
    Ophthalmology, Mater Misericordiae University Hospital and Conway Institute, University College Dublin, Dublin, Ireland; and
  • Abbot F. Clark
    Glaucoma Research, Alcon Research, Ltd., Forth Worth, Texas.
  • Brian J. P. Harvey
    From the Molecular Medicine Laboratories, RCSI Education and Research Centre, Beaumont Hospital, Dublin, Ireland;
  • Colm J. O'Brien
    Ophthalmology, Mater Misericordiae University Hospital and Conway Institute, University College Dublin, Dublin, Ireland; and
Investigative Ophthalmology & Visual Science January 2009, Vol.50, 194-202. doi:10.1167/iovs.08-1937
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      Mustapha Irnaten, Richard C. Barry, Barry Quill, Abbot F. Clark, Brian J. P. Harvey, Colm J. O'Brien; Activation of Stretch-Activated Channels and Maxi-K+ Channels by Membrane Stress of Human Lamina Cribrosa Cells. Invest. Ophthalmol. Vis. Sci. 2009;50(1):194-202. doi: 10.1167/iovs.08-1937.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

purpose. The lamina cribrosa (LC) region of the optic nerve head is considered the primary site of damage in glaucomatous optic neuropathy. Resident LC cells have a profibrotic potential when exposed to cyclical stretch. However, the mechanosensitive mechanisms of these cells remain unknown. Here the authors investigated the effects of membrane stretch on cell volume change and ion channel activity and examined the associated changes in intracellular calcium ([Ca2+]i).

methods. The authors used primary LC cells obtained from normal human donor eyes. Confocal microscopy was used to investigate the effect of hypotonic cell membrane stretch on cell volume changes. Whole-cell patch-clamp and calcium imaging techniques were used to investigate the effect of hypotonicity on ion channel(s) activity and [Ca2+]i changes, respectively. RT-PCR was used to examine for the maxi-K+ signature in LC cells.

results. In this study, LC cells showed significant volume changes in response to hypotonic cell swelling. The authors characterized a large conductance K+ channel (maxi-K+) in LC cells and demonstrated its increased activity during cell membrane hypotonic stretch. RT-PCR revealed the presence of maxi-K+ signature in LC cells. The authors showed the [Ca2+]i and maxi-K+ channels to be dependent on extracellular Ca2+ and inhibited by gadolinium, which blocks stretch-activated channels (SACs). Pretreatment with thapsigargin, which blocks the release of Ca2+ from endoplasmic reticulum stores, showed no significant difference in [Ca2+]i concentration on hypotonic swelling.

conclusions. The results show that hypotonic stress of human LC cells activates SAC and Ca2+-dependent maxi-K+ channels and that the increase in [Ca2+]i during cell swelling was predominantly from extracellular sources (or intracellular stores other than the endoplasmic reticulum). These findings improve the understanding of how LC cells respond to cell membrane stretch. Further experiments in this area may reveal future targets for novel therapeutic intervention in the management of glaucoma.

Glaucomatous optic neuropathy has an estimated worldwide prevalence of 67 million, making it the second most common form of blindness after cataract. 1 Primary open-angle glaucoma (POAG), the most common form of glaucoma, is characterized by irreversible and progressive loss of axons of the retinal ganglion cells (RGCs), usually in response to abnormally elevated intraocular pressure (IOP). 2 The clinical hallmarks of glaucomatous optic neuropathy are excavation of the tissues of the optic nerve head and visual field loss. 3  
Much work has focused on the lamina cribrosa (LC) of the optic nerve head (ONH), and there is substantial evidence that damage to the RGC axons occurs at this region. 4 5 6 In glaucoma, cupping of the optic disc and stretching, compression, and rearrangement of the collagenous cribriform plates occur in response to an increase in IOP. 7 Resident glial cells of the optic nerve head, namely astrocytes and LC cells, are likely to play a role in this remodeling of the extracellular matrix (ECM). 8 9 10 These two cell types differ in that the LC cell, unlike the astrocyte, does not express glial fibrillary acid protein (GFAP). 11 In addition, their pattern of ECM and cell surface molecule expression suggest that they are a unique ONH cell type. 2 Morphologically, the LC cells are broad, flat, and polygonal, whereas the astrocytes are star shaped and have longer, thinner processes. 12  
Previous work by Kirwan et al. 10 13 has demonstrated the profibrotic nature of the LC cells when exposed to stress in the form of cyclical stretch or TGF-β1. Microarray analysis demonstrated the upregulation of TGF-β2, BMP-7, elastin, collagen VI α1, biglycan, versican, EMMPRIN, VEGF, and thrombomodulin in response to cyclical stretch. 10 Because the LC is a compliant tissue in normal human eyes, an alteration of the composition of the ECM caused by this overexpression of profibrotic modulators may lead to eventual reduction in LC compliance. 
Kirwan et al., 10 using microarray experiments on glaucoma cells, revealed upregulation in certain cell membrane channels and ECM genes that led us to investigate the fundamental components involved in the process by which these LC cells sense mechanical force and convert it to a biochemical response, 14 that is, LC cell mechanotransduction. Among the major cellular components of the mechanotransduction process are ion channels, including the stretch-activated ion channel (SAC). 15 These SACs open in response to mechanical stimuli and allow the movement of cations, including Ca2+ entry and K+ extrusion across the cell membrane. 16 Thus any enhancement in cytosolic concentration of Ca2+ may result in the activation of many physiological or pathophysiological processes that include activation of second-messenger systems, initiation of gene transcription, release of calcium from intracellular stores, opening of calcium-dependent ion channels, cell volume regulation, apoptosis, contraction, and differentiation. 17 18 19 Among the most studied of these events is the activation of calcium-dependent potassium channels (maxi-K+). 
Intracellular calcium is a well-characterized modulator of maxi-K+ channels and is thus intimately involved in volume regulation in a variety of cells. 20 21 For example, in vascular smooth muscle, calcium sparks (local increases in Ca2+) generate spontaneous transient outward currents that are produced by maxi-K+ channels (K+ efflux). This hyperpolarizes the membrane and inhibits calcium entry through voltage-gated channels, thus causing muscle relaxation. 22 In many other type of cells, the maxi-K+ channel is central to cell volume regulation. 17 23 24 Cell swelling (caused by hypotonic shock) has a variety of consequences, including alterations in morphology, membrane tension, ion content, and metabolic state. 25 Maxi-K+ channels are regulated by [Ca2+]i, voltage and membrane tension, thus making them important cellular components in the limitation of Ca2+ entry. 26 Maxi-K channels have demonstrated mechanosensitive properties in skeletal muscle, smooth muscle, and many other human tissues, including the myometrium, trabecular meshwork cells, 27 renal tubular epithelium, and endothelial cells. 28 29 30 31 32  
Indeed, no whole-cell patch-clamp studies have been performed in freshly obtained or cultured human LC cells. Thus, the goal of the present study was to identify and characterize ion channels that could be involved in the response to LC cell membrane stretch and volume change, with particular emphasis on their regulation by increases in intracellular Ca2+. Because we believe these cells respond in a profibrotic manner to mechanical stress, an understanding of the upstream regulatory mechanism(s) of this response is important to further our understanding of the pathogenesis of glaucomatous optic neuropathy. 
Materials and Methods
Culture of Human LC Cells
LC cells were derived from two male donors with no evidence of glaucoma. LC cells used were previously characterized by immunofluorescent staining for various markers, including glial fibrillary acidic protein (GFAP), NCAM, α-smooth muscle actin, and a variety of ECM proteins, including collagen types I, III, and IV, elastin, laminin, and fibronectin. 12 The LC cells used in this study were from passages 4 to 8. The cells were maintained at 37°C and 5% CO2 in Dulbecco modified Eagle medium (DMEM; Sigma Chemical, Poole, UK), supplemented with 10% (vol/vol) fetal calf serum (Gibco, Paisley, UK), 2 mM l-glutamine (Gibco), 2 U/mL penicillin, and 2 mg/mL streptomycin (Gibco). When confluent, cells were subcultured onto glass coverslips or plastic tissue culture dishes (Sarstedt, Newton, NC). For Fura-2 AM fluorescence measurements, confluent cells were used 48 to 72 hours after plating. For electrophysiological recordings, cells grown on six-well plates were trypsinized (0.25% trypsin and 1% EDTA) and were replated on 12-mm round glass coverslips before the experiment began. Cells were then used for patch-clamp recording within 30 minutes or were maintained at room temperature up to 3 hours before recording. 
Solutions
The isotonic solution contained 120 mM NaCl, 6 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 5.4 mM HEPES, and 80 mM D-mannitol, pH 7.4 adjusted with NaOH (osmolality, 323 ± 6 mOsm). The hypotonic solution (osmolarity, 232 ± 8 mOsm) was prepared by omitting D-mannitol from the isotonic solution. For patch-clamp experiments, the patch pipette solution contained 120 mM KCl, 1 mM MgCl2, 5 mM EGTA, and 5.4 mM HEPES, pH 7.2 adjusted with KOH (osmolality, 292 ± 5 mOsm). In Ca2+-free conditions, Ca2+ was replaced by equimolar Mg2+
Volume Change Analysis
LC cells were cultured in imaging dishes and grown to greater than 90% confluence. Cells were preloaded for 15 minutes with 5 μM calcein (Sigma), dissolved in dimethyl sulfoxide, and washed with the isotonic solution. The imaging dishes were then mounted on the confocal microscope (LSM 510 Meta; Zeiss, Jena, Germany). Images were acquired every 5 seconds over a period of 4 minutes and analyzed using an image examiner software program (LSM 5; Zeiss). Calcein was excited at 488 nm with the argon-ion laser, whereas the emitted fluorescence was recorded at a wavelength of 530 nm. Hypotonic solution was added after the third image in each experiment, and the subsequent volume change was calculated using the formula:  
\[\mathrm{V}_{\mathrm{c}1}\mathrm{C}_{\mathrm{c}1}{=}\mathrm{V}_{\mathrm{c}2}\mathrm{C}_{\mathrm{c}2},\]
where Vc1 and Cc1 are the volume and calcein concentration under resting (isotonic conditions), respectively. Vc2 and Cc2 are changed cell volume and calcein concentration, respectively. The concentration of calcein is proportional to its fluorescence intensity; thus, the ratio of concentrations is equal to the ratio of intensities. Assuming that Vc1 is 1, the change volume can be calculated from the changed calcein fluorescence intensity (Fc2) and the resting intensity (Fc1) using the formula 33 :  
\[\mathrm{V}_{\mathrm{c}2}{=}\mathrm{F}_{\mathrm{c}1}/\mathrm{F}_{\mathrm{c}2}.\]
 
Patch Clamp
Whole-cell currents were measured in ruptured patches, as described previously. 34 All experiments were performed at room temperature (20°-22°C). Patch pipettes were prepared from capillary glass (GC150F-10; Harvard Apparatus Ltd., Edenbridge, UK), pulled using a programmable puller (DMZ-Universal; Zeitz-Instruments GmbH, Munich, Germany). Patch electrodes had an electrical resistance of 2 to 5 MΩ when filled with pipette solution. An Ag-AgCl wire was used as reference electrode. The patch-clamp apparatus consisted of a head stage (CV-203BU; Axon Instruments, Union City, CA) connected to a series amplifier (Axopatch 200B; Axon Instruments). Recorded membrane currents were filtered at 1 kHz and digitized at 5 kHz. In brief, freshly prepared LC cells were allowed to attach to the bottom of the cell chamber. Whole-cell access to the inside of the cell was obtained by rupturing the membrane under the pipette tip. Cells were voltage clamped at a holding potential of 0 mV, and membrane currents were recorded in response to voltage steps (from −120 mV to +100 mV, with steps of 20 mV). Average membrane capacitance of the cell was approximately 28.5 ± 6.2 pF. During experiments, whole-cell patched cells were allowed to stabilize and dialyze for at least 5 minutes. Currents were recorded over this time period in isotonic bathing solution (323 ± 6 mOsm) to ensure stability of the current recording. The experiments consisted of exposing cells to hypotonic solution (232 ± 8 mOsm) and measuring the whole-cell current amplitude at different voltage-clamp steps. 
RNA Extraction and RT-PCR
Total RNA was extracted from LC cells (Tri-Reagent Kit; Molecular Research Center, Cincinnati, OH). Total RNA (1–2 μg) was reverse transcribed to obtain cDNA using a reverse transcriptase kit (ImProm II; Promega, Southampton, UK). cDNA samples were amplified with three primers corresponding to three different regions of maxi-K using Taq-polymerase in a DNA thermal cycler (MJ Research, South San Francisco, CA). Control RT-PCR reactions without reverse transcriptase or cDNA served as negative controls and did not result in amplification products. RT-PCR was performed with three sets of primers. Two primer sets (accession numbers U11717 and U11058) have been previously published, 23 35 and the third primer set was designed from its cDNA sequence using a software tool (GeneFisher). 36 BLASTN search was performed on primers to confirm that the sequences were not shared with other known genes. PCR primers are listed in Table 1 . The RT-PCR product was analyzed on a 1% 1× Tris-borate-EDTA (TBE) agarose gel and was imaged using a UV light source. 
Calcium Imaging
[Ca2+]i was measured using the Ca2+-sensitive dye Fura-2 AM (Bioscience, Molecular Probes, Dublin, Ireland). In brief, cells were preloaded with Fura 2-AM in a final concentration of 5 μM for 45 minutes, rinsed twice with 1 mL isotonic solution, mounted on the stage of an inverted epifluorescence microscope (Diaphot 200; Nikon, Toyko, Japan) and treated with the various conditions/inhibitors (hypotonic solution, gadolinium, or thapsigargin or maxi-K+ blockers). Cells were excited alternately at wavelengths of 340 and 380 nm. The resultant fluorescence at each excitation wavelength was measured at 510 nm collected using a charge-coupled device camera system (Hamamatsu, Japan). For Ca2+ calibration, values for [Ca2+]i were obtained from the following equation:  
\[{[}\mathrm{Ca}^{2{+}}{]}_{\mathrm{i}}{=}\mathrm{K}_{\mathrm{d}}\ \frac{\mathrm{F}_{380\ \mathrm{Ca\ free}}(R_{0}{-}R_{\mathrm{min}})}{\mathrm{F}_{380\ \mathrm{Ca\ max}}(R_{\mathrm{max}}{-}R_{0})}\]
where the dissociation constant (Kd) was assumed to be 225 nM based on the work of Grynkyiewicz et al. 37 R min is the ratio of fluorescence measured at 340 nm (F340) over 380 nm (F380) in a nominally Ca2+-free solution. R max is the ratio of fluorescence measured at 340 nm over 380 nm in the presence of saturating amounts of Ca2+ (10 mM) and ionomycin (10 μM) in the bathing solution. Drug actions were measured only after steady state conditions were reached. All calcium-imaging experiments were performed in dark at room temperature (20–22°C) to minimize dye leakage. Images were digitized and analyzed (Openlab2 software; Improvision, Coventry, UK). In all the experiments performed using the calcium imaging technique, representative time course experiments are in [Ca2+]i fluorescence ratio (340/380), and the histograms (average data) are presented as the absolute changes in [Ca2+]i
Statistical Analysis
Data are presented as mean ± SEM for a series of the indicated number of experiments. Statistical analysis of the data was performed using t-tests and one-way ANOVA followed by Tukey post hoc test to compare multiple groups, with P ≤ 0.05 considered significant. The use of a paired test reflected that control and experimental measurements were obtained in the same cell. 
Patch-clamp data analysis (Clampfit software of the p-clamp suite version 9.2; Molecular Devices, Eugene, OR) and data analysis and graphing (Origin 7.5; OriginLab, Northampton, MA) were performed. 
Results
Volume Change in LC Cells
The relative cell volume changes of LC cells in response to hypotonic stretch were measured as described in Materials and Methods. Exposure of LC cells to hypotonic solution resulted in a marked increase in cell volume (maximum mean increase of 30.4% ± 0.4% [n = 5 experiments; 31 cells; P < 0.05]; Fig. 1 ) and then gradually recovered to nearly the original volume after the return of LC cells to an isotonic solution. Alterations of cell volume require the participation of ion transport across the cell membrane, including appropriate activity of Ca2+ and K+ channels. K+ channel activity further maintains the cell membrane potential that is a key determinant of Ca2+ entry into the cell through Ca2+ channels. 27 Ca2+ may, in addition, enter through stretch-activated channels when hypotonic shock is applied. 
Characterization of the Outward Currents in Human Lamina Cribrosa Cells
Hypotonic stretch has been reported in other cell types to stretch the cell membrane. 28 29 38 This procedure increases cell membrane tension caused by cell swelling. Whole-cell experiments were performed to evaluate outward currents in response to hypotonic shock in LC cells. Figure 2Ashows typical traces of whole-cell currents recorded from an LC cell under isotonic stretch (left panel), hypotonic stretch (middle panel), and after washout (right panel). Cells displayed outward current rectification at positive membrane potentials. The summary of whole-cell outward currents obtained from 23 cells is represented as the current-voltage (I–V) relationship seen in Figure 2B . The reversal potential was approximately 0 mV, which is close to the theoretical Nernst potential for the K+-selective channel in our experimental conditions. Whole-cell mean conductance measured at +100 mV was increased from 114 ± 5 pS under isotonic (control) conditions to 193 ± 6 pS under hypotonic conditions (n = 23; P < 0.05). Mean maximal current density measured at Vp = +100 mV was 13.3 ± 11 pA/pF before and 99 ± 8.5 pA/pF (at Vp = +100 mV) after hypotonic stretch, and the difference was statistically different (n = 23; P < 0.05). The current increase was reversed on return to isotonic conditions (washout) 28.6 ± 8.7 pA/pF (at Vp = +100 mV; n = 23). These current profiles resembled maxi-K+ currents 23 26 ; therefore, we used the potassium channel blocker BaCl2 to test whether the outward currents observed in LC cells were K+ currents and found that they were. Tetraethylammonium (TEA) was then used to test whether the K+ currents were Ca2+-dependent, and iberiotoxin (Ibtx) was used to test the maxi-K+ signature. Comparable studies have been reported by Fernández-Fernández et al. 23 in human bronchial epithelial cells. Adding K+ channel blockers, TEA (5 mM; n = 8; P < 0.02) and Ba2+(5 mM; n = 8; P < 0.02) or Ibtx (100 nM; n = 12; P < 0.02) resulted in a large inhibition of the outward current (Fig. 3) , indicating that most of the outward current in LC cells occurs through K+ channels. 
Effect of Iberiotoxin on Hypotonic-Induced K+ Currents in Human LC Cells
The identity of the hypotonic-induced K+ channels was investigated by treatment of LC cells with Ibtx, a well-known specific blocker of Ca2+-dependent maxi-K+ channels. Figure 4Ashows that Ibtx blocked the outward current in all tested cells. On average (Fig. 4B) , the mean peak current density measured at +100 mV was 112 ± 2.5 pA/pF before and 14.0 ± 2.5 pA/pF after 100 nM Ibtx (n = 9; P < 0.05). Thus, most of the outward current (approximately 90%) activated in response to hypotonic stretch in LC cells was carried through Ca2+-dependent maxi-K+ channels. 
We examined the presence of maxi-K+ mRNA in LC cells by RT-PCR analysis. Figure 4Cshows that a single band of 309, 500, and 479 bp was obtained for each amplification product, respectively. All the bands corresponded to the predicted size of maxi-K+ mRNA amplimers. Subsequent sequencing of the bands confirmed that they indeed corresponded to the large conductance potassium (maxi-K+) sequence. 
Effect of Extracellular Ca2+ on Maxi-K+ Channels and [Ca2+]i Increase in LC Cells during Membrane Stretch
Because hypotonic shock increases [Ca2+]i and subsequently activates maxi-K+ currents, whole-cell maxi-K+ currents were recorded under conditions of low extracellular Ca2+, in which Ca2+ was replaced by the same concentration of Mg2+. In the presence of 2 mM extracellular Ca2+ (control), perfusion of hypotonic solution induced a substantial increase of maxi-K+ currents from 26.3 ± 5.2 pA/pF to 63.2 ± 10.5 pA/pF (n = 12 cells; P < 0.02; Fig. 5A ). This increase was nearly completely prevented on removal of extracellular Ca2+ (Fig. 5B) . These results indicate that extracellular Ca2+ is required for the activation of maxi-K+ channels during membrane stretch. 
Parallel studies have been performed in Fura-2 AM–loaded LC cells to examine the requirement of extracellular Ca2+ for the increase of [Ca2+]i in response to hypotonic shock. Figure 5Cshows a representative experiment illustrating the time course of the [Ca2+]i increases in response to hypotonic shock measured as the 340/380 fluorescence ratio. In the presence of 2 mM extracellular Ca2+ (control), the hypotonic-induced increase in [Ca2+]i levels was a transient peak followed by a gradually sustained increase in [Ca2+]i with levels above the baseline. Under the nominally extracellular Ca2+-free conditions (0 mM Ca2+), in the presence of 0.5 mM EGTA in the bath solution, this Ca2+ response was almost completely blocked (7.06 ± 1.12 nM; n = 11; 91 cells; P < 0.05; Figs. 5C 5D ). Taken together, these results indicate that Ca2+ influx from the extracellular environment plays an important role in hypotonic-induced [Ca2+]i increases. 
Effect of Thapsigargin on [Ca2+]i in Human LC Cells
To examine whether intracellular stores also contribute to the increase of [Ca2+]i in response to hypotonic cell membrane stretch, we tested the effect of hypotonic stretch on [Ca2+]i after treatment with 1 μM thapsigargin (TG), a well-known inhibitor of the Ca2+-ATPase pump in endoplasmic reticulum (Fig. 6) . Treatment of LC cells with TG in Ca2+-free medium produced a transient large peak increase of Ca2+. Readdition of Ca2+ to the medium under hypotonic conditions resulted in a normal Ca2+ response. The effect of TG on [Ca2+]i is summarized in Figure 6B . No significant difference in the hypotonic stimulation of the [Ca2+]i was observed between control (untreated) and thapsigargin-treated LC cells, suggesting the source of Ca2+ mobilization is mainly extracellular calcium entry into the cell. 
Effect of Gd3+ on Basal and Swelling Activated Channels and [Ca2+]i in Human LC Cells
Gadolinium is known to block stretch-activated ion channels. 39 Whole-cell currents were recorded in LC cells under hypotonic conditions in the presence or absence of Gd3+. Gd3+ did not affect basal whole-cell current densities under isotonic conditions, but it did prevent the increase in whole-cell currents in response to hypotonic cell membrane stretch (Fig. 7A)
The effect of Gd3+ was also tested on [Ca2+]i in Fura-2 AM–loaded LC cells (Figs. 7B 7C) . In the absence of Gd3+, hypotonic cell membrane stretch induced a significant transient increase in [Ca2+]i (113 ± 8.23 nM; n = 15; 125 cells; P < 0.05) compared with normalized basal levels (the basal level of [Ca2+]i is normalized to 0 nM). However, when cells were preincubated with Gd3+, the [Ca2+]i increase in response to hypotonic stress was nearly completely prevented (8.5 ± 3 nM; n = 9; 84 cells; P < 0.05). The effect of Gd3+ on [Ca2+]i is summarized in Figure 7C . Taken together, these results suggest that the hypotonic cell membrane stretch-induced [Ca2+]i elevation occurs through Gd3+-sensitive SAC. 
Discussion
Increased IOP is a well-recognized risk factor for the development of glaucomatous optic neuropathy, and the LC region of the ONH is believed to be the primary site of glaucomatous damage in POAG. 4 5 The pathogenesis of POAG is still unknown but is likely to be multifactorial, with mechanical, vascular, and other factors influencing individual susceptibility to optic nerve damage. 40 Increased IOP induces significant deformation of the level of the LC 3 and consequently the LC cells. Among the different types of stresses experienced by these cells are shear, compressive, and stretch. 41 Different models have been used to investigate the mechanosensitive mechanisms in different cell types, including cyclic stretch, hydrostatic pressure, and hypotonic stress. The hypotonic stress model has been used in many different cell types to stretch the cell membrane. 42 Hypotonic cell swelling has also been shown to activate a variety of ion channels, ion transporters, and their regulatory proteins. 27 In addition, this model activates various second-messenger systems, such as increases in the intracellular levels of Ca2+, cAMP inositol triphosphate, and arachidonic acid metabolites. 43 We previously demonstrated the profibrotic nature of human LC cells when exposed to stress in the form of cyclical stretch or TGF-β. 10 13 This led us to believe that LC cells play an important role in the pathogenesis of glaucoma. Thus, to further our understanding of this LC mechanotransduction process, our goals were to investigate the mechanism(s) by which LC cells respond to a stretch paradigm and to identify the mechanosensitive pathway in the LC cell. 
In this study, we choose the hypotonic cell membrane rather than the physical cell membrane as a model to stretch the LC cell membrane because the hypotonic cell membrane stretch model allowed us to use the same experimental conditions for the Fura-2 AM experiments (calcium imaging) and the patch-clamp recordings. The use of different stretch models (i.e., hypotonic stretch and mechanical cyclic stretch) would give us data that might not be entirely compatible and might lead to some misinterpretations. 
With the use of hypotonic shock as a model to stretch the cell membrane, we first examined the changes in cell volume. The results showed that exposure of LC cells to hypotonic solution resulted in a marked increase in cell volume, followed by the return to near-original volume when the cells were returned to isotonic conditions. 
Given that it is well known that changes of cell volume require the participation of ion transport across the cell membrane, we examined the ion channels involved in response to hypotonic stretch in LC cells. In this study, we provide evidence that a channel exhibiting [Ca2+]i dependence and other properties, similar to those of the Ca2+-activated maxi-K+ channel, is present in LC cells. To our knowledge, the presence of maxi-K+ in LC cells has not previously been investigated, and our experiments show a significant decrease in cell membrane currents when cells were treated with BaCl2, TEA, or Ibtx, indicating that K+ channels play an important role in generating cell membrane potential in LC cells. Ibtx blocked the hypotonic-induced membrane current, indicating that most of the outward currents activated in response to membrane stretch are carried out by maxi-K+ channels. These results are consistent with those from previous studies on pituitary tumor cells, trabecular meshwork cells, and parotid acinar cells. 21 44 45 In addition, RT-PCR product sequencing revealed the presence of mRNA for maxi-K+ channel in these LC cells, supporting the electrophysiological data. 
Although the characteristics of maxi-K+ currents reported in this study (dependence on the extracellular Ca2+, sensitivity to Ibtx) compare well with those reported in a number of other cell types, 28 29 38 the mechanism that mediates hypotonic-induced, [Ca2+]i-dependent, maxi-K+ activation is less defined. Our results show that in LC cells, the activity of [Ca2+]i and maxi-K+ were increased after hypotonic challenge. These increases were inhibited by the removal of extracellular Ca2+ but not by the depletion of internal [Ca2+]i stores (using thapsigargin), indicating that Ca2+ influx through the cell membrane plays an important role in hypotonic-induced [Ca2+]i increases, though internal releases (other than the endoplasmic reticulum Ca2+-ATPase pump) may also contribute at some minor level. The well-known phenomenon of calcium-induced calcium release (CICR) through ryanodine receptors on the endoplasmic reticulum (ER) is stimulated by Ca2+ influx. 46 The use of thapsigargin empties the ER, thus preventing CICR. Hypotonic stretch in a calcium-free bath obviously prevents calcium influx and any possible CICR. Stretching the cells in a 2-mM Ca2+ bath solution, after emptying the ER stores, showed an increase in [Ca2+]i similar to stretch in 2 mM Ca2+ without thapsigargin treatment, indicating the increase is likely to be from the extracellular environment (with a possible minor contribution from calcium-regulated store release from other intracellular stores such as mitochondria). 
It has been shown that the [Ca2+]i response to extracellular stimulus is biphasic, consisting of an initial rapid, transient increase and a slower sustained rise. 24 The initial peak increase in [Ca2+]i mobilization has been attributed to the release of calcium from cytosolic stores, predominantly in the ER. The later sustained increase in [Ca2+]i, however, has been attributed to the influx of extracellular calcium through the activation of Ca2+ channels. 24 44 47 Our results are consistent with this concept because we showed both the initial transient spike and the subsequent slow sustained rise in [Ca2+]i during membrane stretch. 
We showed that the hypotonic stretch-induced [Ca2+]i increase and the subsequent maxi-K+ activation were significantly inhibited by Gd3+. As Gd3+ blocks the Ca2+-permeable SAC channels, we hypothesized that Ca2+ influx might be ascribed to the activation of SACs by hypotonic shock. The primarily distinctive property of SACs is that their gating is dependent on membrane tension. SACs are selective and permeable to various cations, 48 particularly divalent cations, allowing Ca2+ influx during stretch. It has been proposed that Ca2+ may function as a second messenger for translating mechanical perturbation to regulation of ion transport, 49 which may provide an important role in cell volume regulation. SACs thus seem capable of mechanotransduction, transferring mechanical signals to elevations in cytosolic calcium, thereby activating membrane kinases to specifically phosphorylate other signaling molecules. The activation of SACs followed by Ca2+ entry is the primary signal transduction for the activation of maxi-K+, and elevation of maxi-K+ channel activity in turn enhances further Ca2+ entry. In summary, these results suggest that (i) extracellular Ca2+ is required for the observed [Ca2+]i response and activation of maxi-K+ channels in LC cells and (ii) the maxi-K+ channels are not directly activated by cell swelling but are secondary to Ca2+ influx through SACs. Comparable results have been obtained in the human bronchial epithelial cell line. 23  
Maxi-K+ channels were first identified and classified in chromaffin cell membranes in 1981 by Marty et al. 50 Maxi-K+ channels are important components of cellular systems limiting Ca2+ entry and cell membrane excitability. They play a key role in the maintenance of vascular smooth muscle tone through cell membrane potential and Ca2+ entry regulation, 51 Maxi-K+ channels are believed to be sensors of intracellular Ca2+ and are found to regulate membrane potential in an intracellular Ca2+-dependent manner. 52 The role of the maxi-K+ channels described here in the extracellular Ca2+-mediated regulation of membrane current activated by hypotonic stretch is not entirely clear. Activation of this channel under hypotonic conditions would tend to hyperpolarize the cell membrane. In addition to these channels, there may be other types of channels that determine the integrated response of membrane current to hypotonic stretch in LC cells. Further studies will be needed to clarify more fully the relationship between changes in Ca2+ entry through SAC and the regulation of maxi-K+ channels in LC cells. 
We hypothesize that the increased IOP in ocular hypertension and glaucoma results in LC cell membrane stretch, causing activation of the maxi-K+ channels because of increases in intracellular Ca2+ entering by way of SACs. This alteration in [Ca2+]i could influence the regulation of ECM gene transcription, as has been described, 10 and subsequently the matrix structure of the LC region of the ONH. Any change in the compliance of the LC region caused by altered ECM deposition in glaucoma could contribute to the vulnerability of the RGC axons under the border of increased IOP associated with glaucoma. 
 
Table 1.
 
Oligonucleotide Sequences of Maxi-K Primers Used for RT-PCR
Table 1.
 
Oligonucleotide Sequences of Maxi-K Primers Used for RT-PCR
Accession No. Forward Primers (5′-3′) Reverse Primers (5′-3′) Position Length (bp)
ACAACATCTCCCCCAACC TCATCACCTTCTTTCCAATTC 1222–1531 309
U11058 ATCTCCCCCAACCTGGA ACAGTAGGGAAGGACAGA 979–1479 500
U11717 ACCAAGACGATGATGACC AGCAGAAGATCAGGTCCGTC 2675–3154 479
Figure 1.
 
Regulatory volume changes in response to hypotonic shock in LC cells. Time course of percentage of volume changes after exposure of LC cells to a hypotonic bath solution (n = 5 experiments; 31 cells). LC cells were allowed to stabilize in isotonic solution for at least 5 minutes before the application of hypotonic solution at room temperature; mean cell volume promptly increased and then gradually recovered after return to the initial isotonic solution. Arrows: time at which osmolarity was changed to hypotonic and then back to isotonic condition, respectively.
Figure 1.
 
Regulatory volume changes in response to hypotonic shock in LC cells. Time course of percentage of volume changes after exposure of LC cells to a hypotonic bath solution (n = 5 experiments; 31 cells). LC cells were allowed to stabilize in isotonic solution for at least 5 minutes before the application of hypotonic solution at room temperature; mean cell volume promptly increased and then gradually recovered after return to the initial isotonic solution. Arrows: time at which osmolarity was changed to hypotonic and then back to isotonic condition, respectively.
Figure 2.
 
Hypotonic-induced whole-cell outward currents in LC cells. (A) Original traces of whole-cell outward currents elicited by step voltage increments (inset), recorded under isotonic (iso) and hypotonic (hypo) conditions and after a return to initial conditions (washout). (B) Current-voltage relationship of whole-cell outward currents recorded under isotonic, hypotonic, and after washout conditions. Note the hypotonically induced activation of whole-cell, outwardly rectifying currents at positive potentials from +20 mV to +100 mV.
Figure 2.
 
Hypotonic-induced whole-cell outward currents in LC cells. (A) Original traces of whole-cell outward currents elicited by step voltage increments (inset), recorded under isotonic (iso) and hypotonic (hypo) conditions and after a return to initial conditions (washout). (B) Current-voltage relationship of whole-cell outward currents recorded under isotonic, hypotonic, and after washout conditions. Note the hypotonically induced activation of whole-cell, outwardly rectifying currents at positive potentials from +20 mV to +100 mV.
Figure 3.
 
Effect of K+ channel blockers on whole-cell basal current in LC cells. (AC; left, middle) Typical traces of whole-cell K+ currents obtained as described in Figure 2under (control) isotonic conditions in the presence or absence of (A) 100 nM Ibtx, (B) 5 mM Ba2+, and (C) 5 mM TEA in the bath solution. (AC; right) Measured K+ current densities depicted in pA/pF (recorded at +100 mV) in the absence of (Cont) or after treatment by (A) 100 nM Ibtx (n = 12), (B) 5 mM Ba2+ (n = 8), or (C) 5 mM TEA (n = 8). *Values statistically different from untreated (cont) cells (P < 0.02). Cont, control.
Figure 3.
 
Effect of K+ channel blockers on whole-cell basal current in LC cells. (AC; left, middle) Typical traces of whole-cell K+ currents obtained as described in Figure 2under (control) isotonic conditions in the presence or absence of (A) 100 nM Ibtx, (B) 5 mM Ba2+, and (C) 5 mM TEA in the bath solution. (AC; right) Measured K+ current densities depicted in pA/pF (recorded at +100 mV) in the absence of (Cont) or after treatment by (A) 100 nM Ibtx (n = 12), (B) 5 mM Ba2+ (n = 8), or (C) 5 mM TEA (n = 8). *Values statistically different from untreated (cont) cells (P < 0.02). Cont, control.
Figure 4.
 
Effect of iberiotoxin on hypotonic-induced maxi-K+ currents in LC cells. Addition of 100 nM Ibtx blocked the hypotonic-induced maxi-K+ current. (A) Representative maxi-K+ current traces recorded under isotonic conditions (Iso), 5 minutes after the cell was bathed in hypotonic solution (5 minutes Hypo), and after 3 minutes under hypotonic conditions in the presence of Ibtx (3 minutes Hypo + 100 nM Ibtx). (B) Mean ± SEM of K+ current densities obtained at +100 mV under isotonic (Iso) and hypotonic (Hypo) conditions and after the addition of 100 nM Ibtx (Hypo + Ibtx) in LC cells (n = 9). *Results are statistically different in the presence or absence of Ibtx under hypotonic conditions (P < 0.05; Hypo vs. Hypo + Ibtx). (C) RT-PCR product of three different primers coding for three distinct regions of maxi-K+ channel protein. A single band of 309, 500, and 479 bp (lanes 1–3) was obtained for each RT-PCR product, respectively. M represents a 100-bp molecular marker. All the bands corresponded to the predicted size of maxi-K+ mRNA amplimers and were sequence verified.
Figure 4.
 
Effect of iberiotoxin on hypotonic-induced maxi-K+ currents in LC cells. Addition of 100 nM Ibtx blocked the hypotonic-induced maxi-K+ current. (A) Representative maxi-K+ current traces recorded under isotonic conditions (Iso), 5 minutes after the cell was bathed in hypotonic solution (5 minutes Hypo), and after 3 minutes under hypotonic conditions in the presence of Ibtx (3 minutes Hypo + 100 nM Ibtx). (B) Mean ± SEM of K+ current densities obtained at +100 mV under isotonic (Iso) and hypotonic (Hypo) conditions and after the addition of 100 nM Ibtx (Hypo + Ibtx) in LC cells (n = 9). *Results are statistically different in the presence or absence of Ibtx under hypotonic conditions (P < 0.05; Hypo vs. Hypo + Ibtx). (C) RT-PCR product of three different primers coding for three distinct regions of maxi-K+ channel protein. A single band of 309, 500, and 479 bp (lanes 1–3) was obtained for each RT-PCR product, respectively. M represents a 100-bp molecular marker. All the bands corresponded to the predicted size of maxi-K+ mRNA amplimers and were sequence verified.
Figure 5.
 
Effect of extracellular Ca2+ concentration on cell swelling–activated maxi-K+ currents in LC cells. (A) Current-voltage relationship of maxi-K+ current densities recorded with the voltage protocol in the presence of extracellular Ca2+ under isotonic (Iso) or hypotonic (Hypo) conditions (n =12 for each condition). (B) Maxi-K+ current densities recorded under isotonic (Iso) or hypotonic (Hypo) conditions in the absence of extracellular Ca2+ (n = 12 for each condition). (C) Time course of changes in [Ca2+]i in a representative human LC cell measured as the ratio of emitted fluorescence (340/380) in response to normal Ca2+-containing (2 mM Ca2+) and Ca2+-free (0 mM Ca2+) hypotonic solutions. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed. (D) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from multiple experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence (2 mM) or absence (0 mM) of [Ca2+]o (n = 5; 45 cells). We compared the results in the presence or absence of 2 mM [Ca2+]o. *P < 0.05.
Figure 5.
 
Effect of extracellular Ca2+ concentration on cell swelling–activated maxi-K+ currents in LC cells. (A) Current-voltage relationship of maxi-K+ current densities recorded with the voltage protocol in the presence of extracellular Ca2+ under isotonic (Iso) or hypotonic (Hypo) conditions (n =12 for each condition). (B) Maxi-K+ current densities recorded under isotonic (Iso) or hypotonic (Hypo) conditions in the absence of extracellular Ca2+ (n = 12 for each condition). (C) Time course of changes in [Ca2+]i in a representative human LC cell measured as the ratio of emitted fluorescence (340/380) in response to normal Ca2+-containing (2 mM Ca2+) and Ca2+-free (0 mM Ca2+) hypotonic solutions. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed. (D) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from multiple experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence (2 mM) or absence (0 mM) of [Ca2+]o (n = 5; 45 cells). We compared the results in the presence or absence of 2 mM [Ca2+]o. *P < 0.05.
Figure 6.
 
Effect of thapsigargin on [Ca2+]i in LC cells. (A) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded cells resuspended in isotonic solution under Ca2+-free conditions in the presence of thapsigargin (TG; 1 μM), followed by the removal of TG and the readdition of Ca2+ to the medium under hypotonic conditions. First arrow: start of the experiment (under isotonic conditions). Second arrow: time at which the bath was changed to hypotonic solution. (B) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the absence or presence of thapsigargin (n = 9; 72 cells). Note that there was no significant difference between untreated (Cont) and thapsigargin-treated LC cells.
Figure 6.
 
Effect of thapsigargin on [Ca2+]i in LC cells. (A) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded cells resuspended in isotonic solution under Ca2+-free conditions in the presence of thapsigargin (TG; 1 μM), followed by the removal of TG and the readdition of Ca2+ to the medium under hypotonic conditions. First arrow: start of the experiment (under isotonic conditions). Second arrow: time at which the bath was changed to hypotonic solution. (B) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the absence or presence of thapsigargin (n = 9; 72 cells). Note that there was no significant difference between untreated (Cont) and thapsigargin-treated LC cells.
Figure 7.
 
Effect of gadolinium on hypotonic-induced whole-cell currents and [Ca2+]i increases in LC cells. (A) Current-voltage relationship in LC cells showing whole-cell K+ currents with the voltage protocol, under isotonic (Iso) solution and during exposure to hypotonic (Hypo; n = 10 for each condition) solution in the presence (Iso + Gd3+, Hypo + Gd3+) or absence (Iso, Hypo) of 100 μM Gd3+ (n = 10 for each condition). (B) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded LC cells resuspended in isotonic solution and during exposure to hypotonic solution in the presence (○) or absence (•) of 100 μM Gd3+. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed.. (C) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence or absence of Gd. 3 We have compared the results in the presence or absence of Gd3+. *P < 0.05 (n = 9; 72 cells).
Figure 7.
 
Effect of gadolinium on hypotonic-induced whole-cell currents and [Ca2+]i increases in LC cells. (A) Current-voltage relationship in LC cells showing whole-cell K+ currents with the voltage protocol, under isotonic (Iso) solution and during exposure to hypotonic (Hypo; n = 10 for each condition) solution in the presence (Iso + Gd3+, Hypo + Gd3+) or absence (Iso, Hypo) of 100 μM Gd3+ (n = 10 for each condition). (B) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded LC cells resuspended in isotonic solution and during exposure to hypotonic solution in the presence (○) or absence (•) of 100 μM Gd3+. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed.. (C) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence or absence of Gd. 3 We have compared the results in the presence or absence of Gd3+. *P < 0.05 (n = 9; 72 cells).
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Figure 1.
 
Regulatory volume changes in response to hypotonic shock in LC cells. Time course of percentage of volume changes after exposure of LC cells to a hypotonic bath solution (n = 5 experiments; 31 cells). LC cells were allowed to stabilize in isotonic solution for at least 5 minutes before the application of hypotonic solution at room temperature; mean cell volume promptly increased and then gradually recovered after return to the initial isotonic solution. Arrows: time at which osmolarity was changed to hypotonic and then back to isotonic condition, respectively.
Figure 1.
 
Regulatory volume changes in response to hypotonic shock in LC cells. Time course of percentage of volume changes after exposure of LC cells to a hypotonic bath solution (n = 5 experiments; 31 cells). LC cells were allowed to stabilize in isotonic solution for at least 5 minutes before the application of hypotonic solution at room temperature; mean cell volume promptly increased and then gradually recovered after return to the initial isotonic solution. Arrows: time at which osmolarity was changed to hypotonic and then back to isotonic condition, respectively.
Figure 2.
 
Hypotonic-induced whole-cell outward currents in LC cells. (A) Original traces of whole-cell outward currents elicited by step voltage increments (inset), recorded under isotonic (iso) and hypotonic (hypo) conditions and after a return to initial conditions (washout). (B) Current-voltage relationship of whole-cell outward currents recorded under isotonic, hypotonic, and after washout conditions. Note the hypotonically induced activation of whole-cell, outwardly rectifying currents at positive potentials from +20 mV to +100 mV.
Figure 2.
 
Hypotonic-induced whole-cell outward currents in LC cells. (A) Original traces of whole-cell outward currents elicited by step voltage increments (inset), recorded under isotonic (iso) and hypotonic (hypo) conditions and after a return to initial conditions (washout). (B) Current-voltage relationship of whole-cell outward currents recorded under isotonic, hypotonic, and after washout conditions. Note the hypotonically induced activation of whole-cell, outwardly rectifying currents at positive potentials from +20 mV to +100 mV.
Figure 3.
 
Effect of K+ channel blockers on whole-cell basal current in LC cells. (AC; left, middle) Typical traces of whole-cell K+ currents obtained as described in Figure 2under (control) isotonic conditions in the presence or absence of (A) 100 nM Ibtx, (B) 5 mM Ba2+, and (C) 5 mM TEA in the bath solution. (AC; right) Measured K+ current densities depicted in pA/pF (recorded at +100 mV) in the absence of (Cont) or after treatment by (A) 100 nM Ibtx (n = 12), (B) 5 mM Ba2+ (n = 8), or (C) 5 mM TEA (n = 8). *Values statistically different from untreated (cont) cells (P < 0.02). Cont, control.
Figure 3.
 
Effect of K+ channel blockers on whole-cell basal current in LC cells. (AC; left, middle) Typical traces of whole-cell K+ currents obtained as described in Figure 2under (control) isotonic conditions in the presence or absence of (A) 100 nM Ibtx, (B) 5 mM Ba2+, and (C) 5 mM TEA in the bath solution. (AC; right) Measured K+ current densities depicted in pA/pF (recorded at +100 mV) in the absence of (Cont) or after treatment by (A) 100 nM Ibtx (n = 12), (B) 5 mM Ba2+ (n = 8), or (C) 5 mM TEA (n = 8). *Values statistically different from untreated (cont) cells (P < 0.02). Cont, control.
Figure 4.
 
Effect of iberiotoxin on hypotonic-induced maxi-K+ currents in LC cells. Addition of 100 nM Ibtx blocked the hypotonic-induced maxi-K+ current. (A) Representative maxi-K+ current traces recorded under isotonic conditions (Iso), 5 minutes after the cell was bathed in hypotonic solution (5 minutes Hypo), and after 3 minutes under hypotonic conditions in the presence of Ibtx (3 minutes Hypo + 100 nM Ibtx). (B) Mean ± SEM of K+ current densities obtained at +100 mV under isotonic (Iso) and hypotonic (Hypo) conditions and after the addition of 100 nM Ibtx (Hypo + Ibtx) in LC cells (n = 9). *Results are statistically different in the presence or absence of Ibtx under hypotonic conditions (P < 0.05; Hypo vs. Hypo + Ibtx). (C) RT-PCR product of three different primers coding for three distinct regions of maxi-K+ channel protein. A single band of 309, 500, and 479 bp (lanes 1–3) was obtained for each RT-PCR product, respectively. M represents a 100-bp molecular marker. All the bands corresponded to the predicted size of maxi-K+ mRNA amplimers and were sequence verified.
Figure 4.
 
Effect of iberiotoxin on hypotonic-induced maxi-K+ currents in LC cells. Addition of 100 nM Ibtx blocked the hypotonic-induced maxi-K+ current. (A) Representative maxi-K+ current traces recorded under isotonic conditions (Iso), 5 minutes after the cell was bathed in hypotonic solution (5 minutes Hypo), and after 3 minutes under hypotonic conditions in the presence of Ibtx (3 minutes Hypo + 100 nM Ibtx). (B) Mean ± SEM of K+ current densities obtained at +100 mV under isotonic (Iso) and hypotonic (Hypo) conditions and after the addition of 100 nM Ibtx (Hypo + Ibtx) in LC cells (n = 9). *Results are statistically different in the presence or absence of Ibtx under hypotonic conditions (P < 0.05; Hypo vs. Hypo + Ibtx). (C) RT-PCR product of three different primers coding for three distinct regions of maxi-K+ channel protein. A single band of 309, 500, and 479 bp (lanes 1–3) was obtained for each RT-PCR product, respectively. M represents a 100-bp molecular marker. All the bands corresponded to the predicted size of maxi-K+ mRNA amplimers and were sequence verified.
Figure 5.
 
Effect of extracellular Ca2+ concentration on cell swelling–activated maxi-K+ currents in LC cells. (A) Current-voltage relationship of maxi-K+ current densities recorded with the voltage protocol in the presence of extracellular Ca2+ under isotonic (Iso) or hypotonic (Hypo) conditions (n =12 for each condition). (B) Maxi-K+ current densities recorded under isotonic (Iso) or hypotonic (Hypo) conditions in the absence of extracellular Ca2+ (n = 12 for each condition). (C) Time course of changes in [Ca2+]i in a representative human LC cell measured as the ratio of emitted fluorescence (340/380) in response to normal Ca2+-containing (2 mM Ca2+) and Ca2+-free (0 mM Ca2+) hypotonic solutions. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed. (D) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from multiple experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence (2 mM) or absence (0 mM) of [Ca2+]o (n = 5; 45 cells). We compared the results in the presence or absence of 2 mM [Ca2+]o. *P < 0.05.
Figure 5.
 
Effect of extracellular Ca2+ concentration on cell swelling–activated maxi-K+ currents in LC cells. (A) Current-voltage relationship of maxi-K+ current densities recorded with the voltage protocol in the presence of extracellular Ca2+ under isotonic (Iso) or hypotonic (Hypo) conditions (n =12 for each condition). (B) Maxi-K+ current densities recorded under isotonic (Iso) or hypotonic (Hypo) conditions in the absence of extracellular Ca2+ (n = 12 for each condition). (C) Time course of changes in [Ca2+]i in a representative human LC cell measured as the ratio of emitted fluorescence (340/380) in response to normal Ca2+-containing (2 mM Ca2+) and Ca2+-free (0 mM Ca2+) hypotonic solutions. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed. (D) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from multiple experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence (2 mM) or absence (0 mM) of [Ca2+]o (n = 5; 45 cells). We compared the results in the presence or absence of 2 mM [Ca2+]o. *P < 0.05.
Figure 6.
 
Effect of thapsigargin on [Ca2+]i in LC cells. (A) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded cells resuspended in isotonic solution under Ca2+-free conditions in the presence of thapsigargin (TG; 1 μM), followed by the removal of TG and the readdition of Ca2+ to the medium under hypotonic conditions. First arrow: start of the experiment (under isotonic conditions). Second arrow: time at which the bath was changed to hypotonic solution. (B) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the absence or presence of thapsigargin (n = 9; 72 cells). Note that there was no significant difference between untreated (Cont) and thapsigargin-treated LC cells.
Figure 6.
 
Effect of thapsigargin on [Ca2+]i in LC cells. (A) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded cells resuspended in isotonic solution under Ca2+-free conditions in the presence of thapsigargin (TG; 1 μM), followed by the removal of TG and the readdition of Ca2+ to the medium under hypotonic conditions. First arrow: start of the experiment (under isotonic conditions). Second arrow: time at which the bath was changed to hypotonic solution. (B) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the absence or presence of thapsigargin (n = 9; 72 cells). Note that there was no significant difference between untreated (Cont) and thapsigargin-treated LC cells.
Figure 7.
 
Effect of gadolinium on hypotonic-induced whole-cell currents and [Ca2+]i increases in LC cells. (A) Current-voltage relationship in LC cells showing whole-cell K+ currents with the voltage protocol, under isotonic (Iso) solution and during exposure to hypotonic (Hypo; n = 10 for each condition) solution in the presence (Iso + Gd3+, Hypo + Gd3+) or absence (Iso, Hypo) of 100 μM Gd3+ (n = 10 for each condition). (B) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded LC cells resuspended in isotonic solution and during exposure to hypotonic solution in the presence (○) or absence (•) of 100 μM Gd3+. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed.. (C) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence or absence of Gd. 3 We have compared the results in the presence or absence of Gd3+. *P < 0.05 (n = 9; 72 cells).
Figure 7.
 
Effect of gadolinium on hypotonic-induced whole-cell currents and [Ca2+]i increases in LC cells. (A) Current-voltage relationship in LC cells showing whole-cell K+ currents with the voltage protocol, under isotonic (Iso) solution and during exposure to hypotonic (Hypo; n = 10 for each condition) solution in the presence (Iso + Gd3+, Hypo + Gd3+) or absence (Iso, Hypo) of 100 μM Gd3+ (n = 10 for each condition). (B) Representative time tracings of [Ca2+]i in Fura-2 AM–loaded LC cells resuspended in isotonic solution and during exposure to hypotonic solution in the presence (○) or absence (•) of 100 μM Gd3+. First arrow: start of the experiment. Second arrow: time at which osmolarity was changed.. (C) Mean ± SEM of the [Ca2+]i (in nM) in Fura-2 AM–loaded LC cells from different experiments calculated as the maximum increase in [Ca2+]i in response to hypotonic shock in the presence or absence of Gd. 3 We have compared the results in the presence or absence of Gd3+. *P < 0.05 (n = 9; 72 cells).
Table 1.
 
Oligonucleotide Sequences of Maxi-K Primers Used for RT-PCR
Table 1.
 
Oligonucleotide Sequences of Maxi-K Primers Used for RT-PCR
Accession No. Forward Primers (5′-3′) Reverse Primers (5′-3′) Position Length (bp)
ACAACATCTCCCCCAACC TCATCACCTTCTTTCCAATTC 1222–1531 309
U11058 ATCTCCCCCAACCTGGA ACAGTAGGGAAGGACAGA 979–1479 500
U11717 ACCAAGACGATGATGACC AGCAGAAGATCAGGTCCGTC 2675–3154 479
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