Department of Biology, University of North Carolina, Charlotte, North Carolina 28223
Submitted 5 May 2003 ; accepted in final form 22 November 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
water movement; major intrinsic protein; channel; enzyme
In response to an osmotic gradient, water will cross a membrane in two ways: simple diffusion across the hydrophobic phospholipid bilayer or through specific proteinaceous channels called aquaporins (AQPs). Simple diffusion is slow, unregulated, and dictated primarily by the lipid composition of the plasma membrane. In contrast, water movement through AQPs is fast, selective, and subject to rapid cellular regulation (for a review of AQPs, see Refs. 10, 16, 17, 34). Because cell shrinkage also appears to be a rapid phenomenon (8, 14, 28, 30), we hypothesize that AQPs play a role in the movement of water across the plasma membrane in dying cells during the AVD.
AQPs are a subset of the major intrinsic protein (MIP) family of which >80 have been identified in many different organisms ranging from bacteria to humans (44, 47). All members of this family appear to act as transmembrane channel proteins. The conservation of these proteins across evolutionary lines suggests that they play important and very basic roles in cell biology. Currently, 11 mammalian AQPs have been identified, AQP-0 through -10 (1, 26, 44, 47), and we have recently published studies describing a role for AQP-7, -8, and -9 in rat granulosa cells (38).
All members of the AQP family contain six transmembrane segments and two "hemi-channels" that fold together into an hourglass conformation to mediate water movement (for review of AQP structure, see Refs. 10 and 53). Most AQPs possess a cysteine residue on the extracellular side of the membrane between the fifth transmembrane segment and the second hemi-channel. This cysteine residue has been shown to bind Hg2+, which sterically blocks the flow of water through the channel. Thus Hg2+ acts as an effective and general inhibitor of most AQPs. In the present study, we utilized both Hg2+ and a cell line that overexpresses AQP-1 to explore a role for these water channels in the AVD of rat granulosa cells and thymocytes and to assess the importance of AQP-mediated water loss in the regulation of the AVD and, therefore, the beginning of the process of cell death.
Activation of caspases and apoptotic nucleases are dependent on a total decrease in ionic strength of the cytoplasm that is brought about primarily by a loss of intracellular K+ concentration ([K+]i). Logically, for this to occur, ion efflux must be significantly greater than water loss. Thus we hypothesize that after the AVD, AQPs are inactivated, which makes the plasma membrane significantly less permeable to water. Inactivation of these water channels coupled with the continued efflux of ions would reduce the ionic strength of the cytoplasm to levels conducive to the activity of apoptotic enzymes. Accordingly, we have also examined the water permeability characteristics of normal and shrunken (apoptotic) cells within the same population as well as the regulation of AQP-1 in thymocytes after the AVD.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Chinese hamster ovary K1 cells stably transfected with vector alone (CHO-vector) or AQP-1 (CHO-AQP-1) under a constitutive promoter as previously described (36) were kindly donated by Dr. Alan S. Verkman (University of California at San Francisco). These cells were grown in Ham's F-12K nutrient medium that contained 10% fetal calf serum, 50 U/ml penicillin, 50 µg/ml streptomyosin sulfate at 37°C in 5% CO2-95% air. At the beginning of each experiment, cells were seeded at 0.5 x 106 cells in a T-25 flask that contained 5 ml of medium and were cultured for 3 days to confluency. After this period, cells were washed and either harvested immediately (t = 0) or cultured for the indicated time before harvest.
Cell size and DNA content. Cell size and DNA degradation were analyzed on a Becton Dickinson (San Jose, CA) FACSCalibur flow cytometer as previously described (25). Briefly, granulosa cells (0.5 x 106 cells) and thymocytes (2.5 x 106 cells) were cultured for 24 or 48 h in 12 x 75-mm polystyrene flow tubes in 0.5 ml McCoy's medium alone, with 2 µM thapsigargin (Sigma Chemical, St. Louis, MO), or with 50 µM C6-ceramide (Sigma). At the end of the culture period, cells were vortexed vigorously to detach them, pelleted, resuspended in ice-cold 70% ethanol, and stored at 4°C until analyzed (>1 h). To harvest CHO cells, medium was removed and the cells were lifted by treatment with trypsin-EDTA. These cells were then recombined with the original medium that contained the nonadherent cells and were pelleted and resuspended in ice-cold 70% ethanol. For analysis, all cells were washed in ice-cold PBS and resuspended in PBS that contained 20 µg/ml propidium iodide and 1 mg/ml RNase A. After a 10-min incubation at room temperature, cells were analyzed for cell size on a dot plot of forward vs. side scatter. Gating of the normal-size population was set on a freshly isolated cell population that was fixed and processed as described above. DNA content was measured as FL-1 fluorescence (at 530 nm). Cells for analysis included normal and shrunken cells from the size analysis. The normal DNA-content histogram was set using a freshly isolated cell population. Cells with reduced FL-1 fluorescence were considered hypodiploid and thus apoptotic.
Plasmid degradation assays. Granulosa cells were harvested and incubated as described above. After incubation, cells were pelleted and resuspended in ice-cold 10 mM MgCl2 with 0.25% Nonidet P-40. Debris, chromatin, and membranes were pelleted by ultracentrifugation (100,000 g) for 30 min at 4°C, and the resultant supernatant was collected and analyzed for protein content via Bradford assay (15). To assess nuclease activity (45), 5 µg of lysate protein was incubated with 1 µg of linearized pUC18 plasmid (Stratagene, La Jolla, CA; plasmid was linearized with the SmaI restriction enzyme) without or with 50 µM HgCl2 or 50 mM EDTA in a total volume of 10 µl that contained 50 mM Tris·HCl, 1 mM MgCl2, and 1 mM CaCl2. Samples were incubated at 37°C for 1.5 h and then at 55°C for 1 h with 20 µg of proteinase K. Samples were then electrophoresed on a 1% agarose gel (1.5 h at 80 V), stained with ethidium bromide, and visualized by UV transillumination.
Caspase assay. Caspase-3-like protease activity was measured using a fluorometric assay as previously described (14, 28). Briefly, cytoplasmic extracts were prepared by resuspending granulosa cells in 10 mM MgCl2 with 0.25% Nonidet P-40. Debris was pelleted at 100,000 g for 30 min and supernatants were placed on ice. Next, 1050 µg of extract (measured by Bradford assay; Ref. 15) was preincubated in solution that contained 50 mM HEPES (pH 7.5), 10 mM dithiothreitol, 10% sucrose, and 0.1% 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) ± treatments with 200 µM of the noncompetitive inhibitor DEVD-fmk (Asp-Glu-Val-Asp-fluoromethylketone; Kamiya Biomedical, Seattle, WA). Parallel samples were prepared without inhibitor. DEVD-afc substrate (Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin; Kamiya Biomedical) was then added to all tubes (200 µM final concentration). Samples were incubated for 5 min at 30°C, and fluorescence at 505 nm was measured (excitation, 400 nm) on a fluorescence spectrophotometer (Hitachi, Tokyo, Japan). Samples were incubated an additional 1 h and fluorescence was again measured. A standard curve of fluorescence vs. free 7-amino-4-trifluromethylcoumarin (afc) was then used to calculate the specific activity of caspase-3-like enzymes in each sample.
Mitochondrial membrane potential analysis. Granulosa cells obtained from day-2 PMSG-treated rats were cultured at 106 cells/0.5 ml of serum-free McCoy's 5A medium in 5-ml polystyrene round-bottom tubes with or without 50 µM HgCl2 for 48 h. After this incubation, the mitochondrial membrane-potential-sensitive dye 5,5',6,6'-tetrachloro-1,1'3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1; Molecular Probes, Eugene, OR) was added to a final concentration of 10 µM. Cells were incubated for 30 min in the dark and then analyzed by flow cytometry. Cells were first analyzed on a dot plot of forward vs. side scatter and gates were drawn to exclude cell aggregates and debris (data not shown). Gated cell populations were then compared on a dot plot of FL-1 vs. FL-2 (585 nm) fluorescence. In cells with normal mitochondrial membrane potential, JC-1 forms aggregates and displays a high FL-2 fluorescence. Loss of mitochondrial membrane potential causes the aggregates to dissociate into monomers and results in a decreased FL-2 fluorescence. Previous studies have shown that this dye is specific for mitochondrial membrane potential and does not detect changes in plasma membrane potential (11).
Cell swelling assay: flow cytometer. Cells were harvested as described above for DNA and cell size analysis, washed, and resuspended in medium at a concentration of 1 x 106 cells/ml. The cell size distribution was then analyzed by flow cytometry. For granulosa and CHO cells, water was then added to adjust the osmolarity to 170 mosM. For thymocytes, the osmolarity was adjusted to 210 mosM, because thymocytes were found to be susceptible to lysis at 170 mosM during the course of the experiment. After 2 min of incubation, the cell size distribution was again analyzed by flow cytometry.
Flow cytometry. Thymocytes were harvested as described, exposed to growth factor withdrawal for 24 h, and labeled with anti-AQP-1 antibodies using a Cytofix/Cytoperm kit purchased from Pharmingen (Franklin Lakes, NJ). Briefly, harvested cells were pelleted, resuspended in 150 µl of Cytofix/Cytoperm solution, and incubated for 20 min on ice. Cells were then washed twice with 200 µl of 1x Perm/Wash solution. Cells were resuspended in 0.5 ml of Perm/Wash solution and antiserum was added to a final dilution of 1:500. For control (no antibody) samples, antibodies were not added. After a 1-h incubation on ice, cells were washed twice as described above and all samples were resuspended in 1 ml of Perm/Wash solution that contained a 1:1,000 dilution of phycoerythrin-conjugated goat anti-rabbit IgG (Sigma). No-antibody samples were also treated with this secondary antibody. Samples were then incubated for 1 h on iceinthe dark. After this incubation, cells were washed twice more in Perm/Wash solution before being resuspended in 1x PBS. Fluorescence was analyzed on a Becton Dickinson FACSCalibur flow cytometer.
Immunofluorescence and colocalization. Thymocytes were harvested as described above and cultured in serum-free media for 48 h (37°C in 5% CO2). Cells were then fixed in 4% paraformaldehyde for 1 h at 4°C, and 200 µl of the cell suspension was dried onto positively charged slides at 37°C overnight. Sections were rehydrated in 1x PBS for 10 min and then incubated in blocking solution (10% normal goat serum) for 30 min before incubation with primary antibodies. Rabbit anti-rat-AQP-1 (Alpha Diagnostics International, San Antonio, TX) and mouse anti-rat-tumor necrosis factor receptor-1 (TNF-R1; Santa Cruz Biotechnology, Santa Cruz, CA; Ref. 50) were diluted 1:100 with PBS, applied to sections, and allowed to incubate for 12 h at 4°C in a humidified chamber. The slides were then washed in 3x PBS for 10 min each. A Texas red-conjugated goat anti-rabbit IgG secondary antibody (1:500 dilution; Sigma Aldrich, St. Louis, MO) and a FITC-labeled goat anti-mouse IgG secondary antibody (1:100 dilution; Sigma Aldrich) were added to sections and allowed to incubate at room temperature for 2 h. The slides were again washed in 3x PBS for 10 min each, coverslips were mounted, and the slides were stored at 4°C until analysis. Slides were examined using confocal microscopy to obtain fluorescent pictures (both AQP-1 and TNF-R1) and Normaski (light-contrast) images. AQP-1 fluorescent images were overlaid with Normaski images as well as TNF-R1 fluorescence to show membrane-specific localization of this protein. Negative-control samples were incubated in an identical manner with the substitution of PBS for primary antibody.
Statistics. Data presented in Figs. 3 and 7 were analyzed by Student's t-test. All others were analyzed by one-way ANOVA on ranks and Student-Newman-Keuls test. All statistical analyses were calculated with SigmaStat software version 2.0.
|
|
|
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
To address the significance of AQPs to the overall death process, we have analyzed three common characteristics that occur in most models of cell death: DNA degradation, caspase-3 activation, and mitochondrial membrane permeability change (). DNA degradation was assessed by flow cytometry. Figure 2A illustrates typical DNA histograms obtained with granulosa cells incubated in the presence or absence of HgCl2 (50 µM) and defines the gates used to score cells with degraded (hypodiploid; M1) or normal (diploid to tetraploid; M2) amounts of DNA. As shown quantitatively in Fig. 2B, in the absence of HgCl2 there was a significant increase in the proportion of the population that displayed a hypodiploid amount of DNA over the control (t = 0). The decreased water movement blocks the appearance of this apoptotic population in a dose-dependent manner down to levels seen in the nonapoptotic cells (t = 0). This suggests that the inhibition of water movement through AQPs can block downstream activation of DNA degradation.
|
Analysis of the biochemical properties of apoptotic nucleases has revealed them to be sensitive to the ionic composition of the buffer in which they are analyzed (29, 31). To control for the possibility that HgCl2 may directly inhibit the apoptotic nuclease activity in dying granulosa cells, nucleases were extracted from granulosa cells incubated as above (in 0 µM HgCl2), and the extracts were analyzed for the ability to degrade an isolated pUC18 plasmid in the presence or absence of 50 µM HgCl2. As shown in Fig. 2C, incubation with extract from dying granulosa cells degraded the plasmid into smaller fragments that generated a smear on an ethidium bromide-stained agarose gel (control lane). Inclusion of 50 µM HgCl2 in the assay had no effect on the nuclease activity of the extracts, which suggests that the effects seen in Fig. 2B were mediated by inhibition of AQPs and not by direct effects of HgCl2 on nucleases. As a control, the right lane in Fig. 2C demonstrates that this nuclease activity, which is Ca2+/Mg2+ dependent, is inhibited by EDTA.
Caspase-3 is a central effector caspase that is implicated in many different models of cell death. As shown in Fig. 3, caspase-3-like enzyme activity was significantly enhanced in dying granulosa cells (control) compared with a freshly isolated population (t = 0). The appearance of caspase-3-like activity in these cells could be completely blocked by inclusion of 50 µM HgCl2 in the culture medium (in vivo samples). To assess the possibility that HgCl2 was exerting a direct inhibitory effect on this enzyme, preparations of active caspase-3-like enzymes were prepared from dying granulosa cells and assayed in the presence or absence of 50 µM HgCl2 (in vitro samples). As shown in Fig. 3, HgCl2 had no direct effect on the activity of caspase-3-like enzymes, which suggests that the in vivo data resulted from blockage of AQPs.
Growth factor withdrawal-induced apoptosis, such as that utilized in this study, is known to engage the intrinsic pathway of apoptosis. One prominent feature of the intrinsic pathway is that the mitochondrial membranes undergo that leads to release of cytochrome c and eventual activation of caspase-3. We next analyzed the effect inhibition of AQPs had on the
values in dying granulosa cells using the mitochondrial membrane potential dye JC-1. As shown in Fig. 4, there is a significant increase in the percentage of the population that has undergone
after 48 h in culture relative to the freshly isolated population (t = 0). In contrast, 50 µM HgCl2 completely blocked this depolarization event. This suggests that AQP-mediated water loss is required for apoptotic mitochondrial changes.
|
Inhibition of AVD and apoptosis is not specific to cell type or signal. The results thus far have been obtained with granulosa cells undergoing growth factor withdrawal. To ensure that these effects were not specific to this cell type or stimulus, we expanded these studies to examine apoptosis of thymocytes and in response to thapsigargin and C6-ceramide. To better detect the enhancement of apoptosis in response to these agents, the incubation times were reduced to 24 h for both granulosa cells and thymocytes. As shown in Fig. 5, thapsigargin and C6-ceramide stimulated cell shrinkage and DNA degradation in both granulosa cells and thymocytes above growth factor withdrawal. HgCl2 attenuated the effects of these agents with the same efficiency as growth factor withdrawal. Thus the ability to block apoptosis through the inhibition of AQPs is neither cell type or signal specific.
|
HgCl2 was used in this study to inhibit the water movement through AQPs. Previous studies have suggested that HgCl2 may directly influence signaling pathways including the MAPK pathway (35, 59). To ensure that HgCl2 was blocking apoptosis only by inhibition of AQPs and not inadvertently by activating MAPK or through the well-known phosphatidylinositol 3-kinase (PI3-K) survival pathway, we analyzed the ability of inhibitors of these pathways to attenuate the effects of HgCl2. As shown in Fig. 6, inhibition of MAPK kinase with PD-098,059 (3, 20, 43) or PI3-K with wortmannin (4, 46) or LY-294002 (54) in granulosa cells undergoing growth factor withdrawal enhanced cell death slightly, which indicates a nominal level of activity in the cells. Thymocyte apoptosis was not significantly affected by these agents, which indicates very low basal activity levels of these survival pathways in these cells. Importantly, these compounds did not influence the ability of HgCl2 to block cell shrinkage or DNA degradation in either cell type. This suggests that HgCl2 is not significantly activating these pathways, and thus inhibition of apoptosis seen with the use of HgCl2 is most likely through the ability of this compound to inhibit AQP function.
|
Overexpression of AQP-1 enhances AVD and apoptosis. The results from Figs. 1, 2, 3, 4, 5, 6 suggest that inhibition of plasma membrane permeability may influence the ability of a cell to undergo the AVD and initiate the apoptotic cascade. To determine whether increasing the plasma membrane water permeability may enhance the rate of apoptosis in an appropriately stimulated cell, we examined CHO cells transfected with an AQP-1 expression vector (CHO-AQP-1) or an empty vector (CHO-vector). Expression levels of AQP-1 in these cells have been previously characterized by Ma et al. (36), and our lab has confirmed these results by Western blot analysis (data not shown). To assess plasma membrane water permeability, cell size was measured by flow cytometry before and after 2-min exposure to medium diluted to 170 mosM. Figure 7A depicts overlay histograms of the relative cell size (forward scatter) from these two measurements in these two cell lines (0 and 2 min are shown). CHO-vector cells displayed distinct swelling in this time frame, although the magnitude of this swelling was greatly enhanced in the CHO-AQP-1 cells, which indicates (as expected) increased water permeability in the CHO-AQP-1 cells. We next determined whether this increased water permeability in CHO-AQP-1 cells would increase the number of cells in the population that undergo apoptosis (apoptosis detected through cell shrinkage and DNA degradation). As shown in Fig. 7B, at the beginning of the experiment (t = 0), CHO-AQP-1 cells displayed a slightly higher although not significantly different level of apoptosis (both cell shrinkage and DNA degradation) than CHO-vector cells. This difference was greatly magnified after 48 h of growth factor withdrawal. This indicates that increasing the permeability of the plasma membrane enhances the rate of cell death.
Water permeability is significantly decreased in apoptotic cells after AVD. Water loss after the AVD must be attenuated to allow the reduction of [K+]i, which is required for activation of apoptotic enzyme activity. Thus we hypothesized that the plasma membrane of the shrunken cells undergoes a significant decrease in water permeability. We investigated this hypothesis by analyzing the cell swelling properties of both normal and shrunken (apoptotic) cells. Granulosa cells, thymocytes, and CHO-AQP-1 cells were induced to undergo apoptosis by growth factor withdrawal for 48 h, and the ability of normal and shrunken (apoptotic) subpopulations to swell in response to a hypotonic insult was measured using the flow cytometer (as described in MATERIALS AND METHODS). Depicted in Fig. 8, A, C, and E are the dot plots from each of these populations of cells at 48 h and the gates used to differentiate normal (R2) from shrunken (R1) subpopulations. The distribution of each subpopulation was then separately plotted on an overlay (Fig. 8, B, D, and F), and peak heights were normalized for each histogram. The population was then subjected to hypotonic challenge and the results were overlaid on the initial measurements. In all three cell types, the shrunken subpopulation (R1) displayed virtually no swelling in response to the hypotonic insult, whereas the normal cells (R2) displayed a measurable increase in size. This response was exaggerated in the highly permeable CHO-AQP-1 cells (Fig. 8, E and F), where the normal cells increased in size to an amount equivalent to that seen in Fig. 7, whereas the shrunken cells showed no increase in size, which suggests very low plasma membrane permeability in this apoptotic population. Figure 9 displays the dot plots from these cells to illustrate the dramatic disjuncture in the permeability rates of these two subpopulations.
|
These results support our hypothesis that water movement through AQPs is vital to the cell death process, and this water movement may be inactivated after the AVD to provide ionic conditions in the cell that are suitable to activation of caspases and nucleases required for proper progression of the cell death repertoire.
AQP-1 is present on plasma membrane of thymocytes and remains after AVD. We have shown that plasma membrane water permeability is significantly inhibited in cells after the AVD. It is possible that this decrease is mediated by AQP degradation and/or removal from the plasma membrane. Thus we next determined AQP-1 expression, levels, and subcellular localization in thymocytes before and after the AVD. For these studies, thymocytes were used because they are nonadherent and relatively uniform and are therefore easy to differentiate between normal and apoptotic cells based on size.
As shown in Fig. 10A, AQP-1 is expressed in thymocytes, and there is not a significant decrease in the percentage of the population staining for this homolog after 24 h of growth factor withdrawal-induced apoptosis. Thus degradation of AQP-1 seems an unlikely mechanism by which the plasma membrane permeability is decreased after the AVD. We next examined the subcellular location of AQP-1 in shrunken and normal thymocytes exposed to growth factor withdrawal for 24 h to determine whether this protein remained on the plasma membrane. In this population, we were clearly able to distinguish between pre- and post-AVD cells simply by size. The location of the plasma membrane was first determined from Normaski optics. As seen in Fig. 10B, the majority of AQP-1 localized to the plasma membrane of the normal-sized cells (arrowhead) and did not change location after the AVD (open arrows). To further ensure that this AQP was indeed on the membrane, a colocalization experiment was performed with AQP-1 and TNF-R1, a cell surface receptor previously shown to be in the thymocyte plasma membrane (50). In Fig. 10C, AQP-1 localization is indicated by red fluorescence whereas TNF-R1 is indicated by green. By overlaying these images, colocalization is indicated by yellow fluorescence. This experiment clearly shows that AQP-1 and TNF-R1 colocalize both before and after the AVD, which suggests that AQP-1 remains on the plasma membrane throughout the apoptotic process.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
A reduction in [K+]i is essential for activation of apoptotic enzymes (7, 8, 14, 19, 28, 42, 60), but a loss of ions does not necessarily imply a reduction in concentration if the loss is matched by an efflux in water and a reduction in intracellular size. We have shown within the same population of cells that the subpopulations that have undergone the AVD have very low water permeability, whereas the nonapoptotic cells have normal permeability. This change in water permeability during the AVD would thus allow the loss of K+ to be unbalanced from the loss of water and facilitate a decrease in [K+]i. We have also shown that AQP-1 in thymocytes is not proteolytically cleaved or removed from the cell membrane after apoptosis, which suggests that this water channel is inactivated by a posttranslational nondegradative mechanism.
We have shown that thymocytes express AQP-1, which is known to be Hg2+ sensitive (52). In addition, we have shown in previous studies that granulosa cells express AQP-7, -8, and -9 (38). Interestingly, AQP-7 is Hg2+ insensitive because it does not contain the Hg2+-binding cysteine in its water pore, whereas AQP-8 and -9 are Hg2+ sensitive (52). Moreover, we have shown that granulosa cell apoptosis is in fact inhibited by Hg2+, and therefore we believe that the AVD during granulosa cell apoptosis is predominantly mediated through AQP-8 and -9. It is important to note that the AVD is a critical event during apoptosis. Thus it is reasonable to speculate that most cell types that are highly sensitive to apoptosis would express more than one AQP family member to ensure cell death.
A particularly intriguing finding in this study is that HgCl2 blockage of water movement inhibits a number of apoptotic effects downstream of the AVD. In this regard, it is interesting to note that several studies have blocked the AVD by a variety of methods including the use of K+ and Cl channel inhibitors (8, 27, 41, 49, 56, 62), caspase inhibitors (30, 55), and high-K+ medium (14, 28). In those studies, downstream apoptotic events were also abrogated, further suggesting an essential role for the AVD in the death process. In one study (27), some apoptotic characteristics (such as caspase activity) were detected in the absence of an AVD, although this effect was restricted to nitric oxide-induced apoptosis in a macrophage cell line (RAW 264.7). AQP expression has never been studied in these cells, and a deficiency of water channels may be one explanation for the apparent lack of an AVD and ability to die independent of it. All other inducing agents used with these cells or any other cell types displayed a strict correlation between the AVD and activation of apoptotic enzymes. Because the AVD and the subsequent decrease in [K+]i are absolutely critical for the activation of apoptotic enzymes, it is reasonable to infer that all cells that express AQPs will use one or more of these water channels to mediate water loss during apoptosis. One interesting scenario is cells that do not appear to express AQPs. As more family members become known, a comprehensive analysis will be needed to ensure that these cells do not express any water channels. Because the AVD is a nearly universal characteristic during apoptosis, it is also possible that in cells that do not normally express AQPs, one or more of the AQP family members may be upregulated in response to an apoptotic stimulus. Conversely, they may rely exclusively on simple diffusion of water across the plasma membranes to facilitate the AVD. However, based on the rapid and regulated nature of the AVD, simple diffusion through the lipid bilayer seems unlikely.
Potential caveats exist in the literature concerning the use of Hg2+. Whitekus et al. (58) showed that Hg2+ blocked cell death induced by FasL, an effect that is hypothesized to be caused by receptor clustering. Thus in these studies, we have intentionally avoided the use of cell surface death receptors to obviate any concerns with receptors clustering. In addition, there are reports that both inorganic and organic Hg2+ compounds induce apoptosis in lymphoid and nonlymphoid cells (2, 21, 32, 48). Based on the argument put forth by Whitekus et al. (58), this variation may be accounted for by differences in both concentration and distribution of Hg2+. More membrane-permeable organomercurials such as methylmercury display higher toxicity (18), which suggests a mechanism of action independent of the death pathways initiated here. Studies have also shown that HgCl2 may have direct effects on the MAPK pathway; this signaling pathway has clearly been implicated in survival of apoptotic insult (35, 59). In this study, we addressed the possibility that HgCl2 may suppress cell death by activating this survival pathway or the well-characterized PI3-K-mediated survival pathway. Inhibition of either pathway had no effect on the ability of HgCl2 to block apoptosis. Our studies used very low concentrations of HgCl2, and we have never detected toxicity and/or apoptosis in our controls that contained HgCl2 alone.
Presently there is debate in the literature concerning the effects of Hg2+ on K+ channels. For example, Ballatori et al. found that the plasma membrane K+ permeability was enhanced by treatment with Hg2+ (5, 6), whereas several other studies detected Hg2+-activated K+ currents across the membrane (22, 33, 40). In contrast, Gallagher et al. detected a K+ current in B lymphocytes that was blocked by Hg2+ (23). A single channel mediating K+ loss during apoptosis in all cells has not been identified, and it is possible that different channels may be involved in a cell or signal-specific manner (8, 9, 41, 57, 61, 62). Using a model of AQP-1 overexpression (CHO-AQP-1 cells), we demonstrated that increased water permeability of the plasma membrane greatly enhanced the rate of cell death, which provides strong evidence independent of this heavy metal that membrane permeability to water can influence the susceptibility of a cell to apoptosis when the cell is appropriately stimulated.
Not only is water movement during the AVD critical to the progression of the cell death cascade, but interestingly, the permeability of the apoptotic cell after the AVD is significantly suppressed. This result can be seen most clearly in the exaggerated response of the CHO-AQP-1 cells. Inhibition of water movement after the AVD would allow [K+]i to decrease to levels necessary to facilitate the activation of downstream apoptotic enzymes. The results with AQP-1 in thymocytes suggest that this inhibition occurs through a posttranslational nondegradative mechanism while these channels remain on the cell surface. Ongoing studies in our laboratory are focused on the mechanisms of inhibition of these water channels after the AVD.
The results of this study significantly strengthen the importance of the AVD to apoptosis and suggest for the first time that the water loss occurs through specific proteinaceous water channels. In addition, we have shown that AQP inhibition suppresses the appearance of apoptotic characteristics, whereas overexpression of AQPs enhances the rate of intrinsically induced cell death. This suggests that the water permeability of the plasma membrane may alter the ability of the cell to die. Furthermore, although functional AQPs may be necessary for a cell to shrink and die, they are also inactivated after the AVD, presumably to allow the [K+]i to be reduced to levels conducive to activation of the apoptotic program. This inactivation of AQPs after the AVD seems to be accomplished through means other than degradation of these water channels.
![]() |
ACKNOWLEDGMENTS |
---|
GRANTS
This study was supported by National Institute of Child Health and Human Development Grants HD-39683 and HD-39234 (to F. M. Hughes, Jr.) and internal research funds from the University of North Carolina at Charlotte.
![]() |
FOOTNOTES |
---|
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.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
2. Akhand AA, Kato M, Suzuki H, Miyata T, and Nakashima I. Level of HgCl2-mediated phosphorylation of intracellular proteins determines death of thymic T-lymphocytes with or without DNA fragmentation. J Cell Biochem 71: 243253, 1998.[CrossRef][ISI][Medline]
3. Alessi DR, Cuenda A, Cohen P, Dudley DT, and Saltiel AR. PD 098059 is a specific inhibitor of the activation of mitogen-activated protein kinase kinase in vitro and in vivo. J Biol Chem 270: 2748927494, 1995.
4. Arcaro A and Wymann MP. Wortmannin is a potent phosphatidylinositol 3-kinase inhibitor: the role of phosphatidylinositol 3,4,5-trisphosphate in neutrophil responses. Biochem J 296: 297301, 1993.[ISI][Medline]
5. Ballatori N and Boyer JL. Disruption of cell volume regulation by mercuric chloride is mediated by an increase in sodium permeability and inhibition of an osmolyte channel in skate hepatocytes. Toxicol Appl Pharmacol 140: 404410, 1996.[CrossRef][ISI][Medline]
6. Ballatori N, Shi C, and Boyer JL. Altered plasma membrane ion permeability in mercury-induced cell injury: studies in hepatocytes of elasmobranch Raja erinacea. Toxicol Appl Pharmacol 95: 279291, 1988.[ISI][Medline]
7. Barbiero G, Duranti F, Bonelli G, Amenta JS, and Baccino FM. Intracellular ionic variations in the apoptotic death of L cells by inhibitors of cell cycle progression. Exp Cell Res 217: 410418, 1995.[CrossRef][ISI][Medline]
8. Beauvais F, Michel L, and Dubertret L. Human eosinophils in culture undergo a striking and rapid shrinkage during apoptosis. Role of K+ channels. J Leukoc Biol 57: 851855, 1995.[Abstract]
9. Benson RSP, Heer S, Dive C, and Watson AJM. Characteristics of cell volume loss in CEM-C7A cells during dexamethasone-induced apoptosis. Am J Physiol Cell Physiol 270: C1190C1203, 1996.
10. Borgnia M, Nielsen S, Engel A, and Agre P. Cellular and molecular biology of the aquaporin water channels. Annu Rev Biochem 68: 425458, 1999.[CrossRef][ISI][Medline]
11. Bortner CD and Cidlowski JA. Caspase independent/dependent regulation of K+, cell shrinkage, and mitochondrial membrane potential during lymphocyte apoptosis. J Biol Chem 274: 2195321962, 1999.
12. Bortner CD and Cidlowski JA. A necessary role for cell shrinkage in apoptosis. Biochem Pharmacol 56: 15491559, 1998.[CrossRef][ISI][Medline]
13. Bortner CD, Gomez-Angelats M, and Cidlowski JA. Plasma membrane depolarization without repolarization is an early molecular event in anti-Fas induced apoptosis. J Biol Chem 276: 43044314, 2001.
14. Bortner CD, Hughes FM Jr, and Cidlowski JA. A primary role for K+ and Na+ efflux in the activation of apoptosis. J Biol Chem 272: 3243632442, 1997.
15. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248254, 1976.[CrossRef][ISI][Medline]
16. Brown D. The ins and outs of aquaporin-2 trafficking. Am J Physiol Renal Physiol 284: F893F901, 2003.
17. Brown D, Katsura T, and Gustafson CE. Cellular mechanisms of aquaporin trafficking. Am J Physiol Renal Physiol 275: F328F331, 1998.
18. Clarkson TW. The toxicology of mercury. Crit Rev Clin Lab Sci 34: 369403, 1997.[ISI][Medline]
19. Dallaporta B, Hirsch T, Susin SA, Zamzami N, Larochette N, Brenner C, Marzo I, and Kroemer G. Potassium leakage during the apoptotic degradation phase. J Immunol 160: 56055615, 1998.
20. Dudley DT, Pang L, Decker SJ, Bridges AJ, and Saltiel AR. A synthetic inhibitor of the mitogen-activated protein kinase cascade. Proc Natl Acad Sci USA 92: 76867689, 1995.[Abstract]
21. Duncan-Achanzar KB, Jones JT, Burke MF, Carter DE, and Laird HE 2nd. Inorganic mercury chloride-induced apoptosis in the cultured porcine renal cell line LLC-PK1. J Pharmacol Exp Ther 277: 17261732, 1996.[Abstract]
22. Dyatlov VA, Platoshin AV, Lawrence DA, and Carpenter DO. Mercury (Hg2+) enhances the depressant effect of kainate on Ca-inactivated potassium current in telencephalic cells derived from chick embryos. Toxicol Appl Pharmacol 138: 285297, 1996.[CrossRef][ISI][Medline]
23. Gallagher JD, Noelle RJ, and McCann FV. Mercury suppression of a potassium current in human B lymphocytes. Cell Signal 7: 3138, 1995.[CrossRef][ISI][Medline]
24. Gomez-Angelats M, Bortner CD, and Cidlowski JA. Protein kinase C (PKC) inhibits fas receptor-induced apoptosis through modulation of the loss of K+ and cell shrinkage. A role for PKC upstream of caspases. J Biol Chem 275: 1960919619, 2000.
25. Gross SA, Newton JM, and Hughes FM Jr. Decreased intracellular potassium levels underlie increased progesterone synthesis during ovarian follicular atresia. Biol Reprod 64: 17551760, 2001.
26. Hatakeyama S, Yoshida Y, Tani T, Koyama Y, Nihei K, Ohshiro K, Kamiie JI, Yaoita E, Suda T, Hatakeyama K, and Yamamoto T. Cloning of a new aquaporin (AQP10) abundantly expressed in duodenum and jejunum. Biochem Biophys Res Commun 287: 814819, 2001.[CrossRef][ISI][Medline]
27. Hortelano S, Zeini M, Castrillo A, Alvarez AM, and Bosca L. Induction of apoptosis by nitric oxide in macrophages is independent of apoptotic volume decrease. Cell Death Differ 9: 643650, 2002.[CrossRef][ISI][Medline]
28. Hughes FM Jr, Bortner CD, Purdy GP, and Cidlowski JA. Intracellular K+ suppresses the activation of apoptosis in lymphocytes. J Biol Chem 272: 3056730576, 1997.
29. Hughes FM Jr and Cidlowski JA. Apoptotic DNA degradation: evidence for novel enzymes. Cell Death Differ 1: 1117, 1994.
30. Hughes FM Jr and Cidlowski JA. Glucocorticoid-induced thymocyte apoptosis: protease-dependent activation of cell shrinkage and DNA degradation. J Steroid Biochem Mol Biol 65: 207217, 1998.[CrossRef][ISI][Medline]
31. Hughes FM Jr and Cidlowski JA. Utilization of an in vitro apoptosis assay to evaluate chromatin degradation by candidate apoptotic nucleases. Cell Death Differ 4: 200208, 1997.[CrossRef][ISI]
32. InSug O, Datar S, Koch CJ, Shapiro IM, and Shenker BJ. Mercuric compounds inhibit human monocyte function by inducing apoptosis: evidence for formation of reactive oxygen species, development of mitochondrial membrane permeability transition and loss of reductive reserve. Toxicology 124: 211224, 1997.[CrossRef][ISI][Medline]
33. Jungwirth A, Ritter M, Paulmichl M, and Lang F. Activation of cell membrane potassium conductance by mercury in cultured renal epithelioid (MDCK) cells. J Cell Physiol 146: 2533, 1991.[ISI][Medline]
34. King LS, Yasui M, and Agre P. Aquaporins in health and disease. Mol Med Today 6: 6065, 2000.[CrossRef][ISI][Medline]
35. Kong AN, Yu R, Chen C, Mandlekar S, and Primiano T. Signal transduction events elicited by natural products: role of MAPK and caspase pathways in homeostatic response and induction of apoptosis. Arch Pharmacol Res (Seoul) 23: 116, 2000.
36. Ma T, Yang B, and Verkman AS. Cloning of a novel water and urea-permeable aquaporin from mouse expressed strongly in colon, placenta, liver, and heart. Biochem Biophys Res Commun 240: 324328, 1997.[CrossRef][ISI][Medline]
37. Maeno E, Ishizaki Y, Kanaseki T, Hazama A, and Okada Y. Normotonic cell shrinkage because of disordered volume regulation is an early prerequisite to apoptosis. Proc Natl Acad Sci USA 97: 94879492, 2000.
38. McConnell NA, Yunus RS, Gross SA, Bost KL, Clemens MG, and Hughes FM Jr. Water permeability of an antral ovarian follicle is predominantly transcellular and mediated by aquaporins. Endocrinology 143: 29052912, 2002.
40. Muller TH, Swandulla D, and Lux HD. Activation of three types of membrane currents by various divalent cations in identified molluscan pacemaker neurons. J Gen Physiol 94: 9971014, 1989.[Abstract]
41. Nietsch HH, Roe MW, Fiekers JF, Moore AL, and Lidofsky SD. Activation of potassium and chloride channels by tumor necrosis factor alpha. Role in liver cell death. J Biol Chem 275: 2055620561, 2000.
42. Nobel CS, Aronson JK, van den Dobbelsteen DJ, and Slater AF. Inhibition of Na+/K+-ATPase may be one mechanism contributing to potassium efflux and cell shrinkage in CD95-induced apoptosis. Apoptosis 5: 153163, 2000.[CrossRef][ISI][Medline]
43. Pang L, Sawada T, Decker SJ, and Saltiel AR. Inhibition of MAP kinase kinase blocks the differentiation of PC-12 cells induced by nerve growth factor. J Biol Chem 270: 1358513588, 1995.
44. Park JH and Saier MH Jr. Phylogenetic characterization of the MIP family of transmembrane channel proteins. J Membr Biol 153: 171180, 1996.[CrossRef][ISI][Medline]
45. Perez GI, Maravei DV, Trbovich AM, Cidlowski JA, Tilly JL, and Hughes FM Jr. Identification of potassium-dependent and -independent components of the apoptotic machinery in ovarian germ cells and granulosa cells. Biol Reprod 63: 13581369, 2000.
46. Reinhold SL, Prescott SM, Zimmerman GA, and McIntyre TM. Activation of human neutrophil phospholipase D by three separable mechanisms. FASEB J 4: 208214, 1990.
47. Reizer J, Reizer A, and Saier MH Jr. The MIP family of integral membrane channel proteins: sequence comparisons, evolutionary relationships, reconstructed pathway of evolution, and proposed functional differentiation of the two repeated halves of the proteins. Crit Rev Biochem Mol Biol 28: 235257, 1993.[Abstract]
48. Shenker BJ, Datar S, Mansfield K, and Shapiro IM. Induction of apoptosis in human T-cells by organomercuric compounds: a flow cytometric analysis. Toxicol Appl Pharmacol 143: 397406, 1997.[CrossRef][ISI][Medline]
49. Szabo I, Lepple-Wienhues A, Kaba KN, Zoratti M, Gulbins E, and Lang F. Tyrosine kinase-dependent activation of a chloride channel in CD95-induced apoptosis in T lymphocytes. Proc Natl Acad Sci USA 95: 61696174, 1998.
50. Tartaglia L, Weber R, Figari I, and Reynolds C. The two different receptors for tumor necrosis factor mediate distinct cellular responses. Proc Natl Acad Sci USA 88: 92929296, 1991.[Abstract]
51. Thompson GJ, Langlais C, Cain K, Conley EC, and Cohen GM. Elevated extracellular [K+] inhibits death-receptor and chemical-mediated apoptosis prior to caspase activation and cytochrome c release. Biochem J 357: 137145, 2001.[CrossRef][ISI][Medline]
52. Verkman AS. Applications of aquaporin inhibitors. Drug News Perspect 14: 412420, 2001.[ISI][Medline]
53. Verkman AS and Mitra AK. Structure and function of aquaporin water channels. Am J Physiol Renal Physiol 278: F13F28, 2000.
54. Vlahos CJ, Matter WF, Hui KY, and Brown RF. A specific inhibitor of phosphatidylinositol 3-kinase, 2-(4-morpholinyl)-8-phenyl-4H-1-benzopyran-4-one (LY294002). J Biol Chem 269: 52415248, 1994.
55. Vu CC, Bortner CD, and Cidlowski JA. Differential involvement of initiator caspases in apoptotic volume decrease and potassium efflux during Fas- and UV-induced cell death. J Biol Chem 276: 3760237611, 2001.
56. Wang X, Xiao AY, Ichinose T, and Yu SP. Effects of tetraethylammonium analogs on apoptosis and membrane currents in cultured cortical neurons. J Pharmacol Exp Ther 295: 524530, 2000.
57. Wang L, Xu D, Dai W, and Lu L. An ultraviolet-activated K+ channel mediates apoptosis of myeloblastic leukemia cells. J Biol Chem 274: 36783685, 1999.
58. Whitekus MJ, Santini RP, Rosenspire AJ, and McCabe MJ Jr. Protection against CD95-mediated apoptosis by inorganic mercury in Jurkat T cells. J Immunol 162: 71627170, 1999.
59. Yu R, Chen C, Mo YY, Hebbar V, Owuor ED, Tan TH, and Kong AN. Activation of mitogen-activated protein kinase pathways induces antioxidant response element-mediated gene expression via a Nrf2-dependent mechanism. J Biol Chem 275: 3990739913, 2000.
60. Yu SP and Choi DW. Ions, cell volume, and apoptosis. Proc Natl Acad Sci USA 97: 93609362, 2000.
61. Yu SP, Yeh C, Strasser U, Tian M, and Choi DW. NMDA receptor-mediated K+ efflux and neuronal apoptosis. Science 284: 336339, 1999.
62. Yu SP, Yeh CH, Sensi SL, Gwag BJ, Canzoniero LMT, Farhangrazi ZS, Ying HS, Tian M, Dugan LL, and Choi DW. Mediation of neuronal apoptosis by enhancement of outward potassium current. Science 278: 114117, 1997.