Sustained hyposmotic stress induces cell death: apoptosis by defeat

Tina Jäckle1,*, Cornelia Hasel1,*, Ingo Melzner1, S. Brüderlein1, Peter M. Jehle2, and Peter Möller1

1 Institute of Pathology and 2 Second Department of Internal Medicine, University of Ulm, D-89081 Ulm, Germany


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We describe sustained hyposmotic stress as a novel type of environmental condition enforcing apoptosis. In a dose- and time-dependent fashion, hyposmotic stress leads to a delayed type of apoptosis with considerable variations in constitutive sensitivity among different cell types. For example, after 48 h at 84 mosmol/l, the death rate ranged from 10.8 ± 0.7% in AsPc1 human pancreatic carcinoma cells to 72.0 ± 1.6% in HK-2 human kidney tubule cells. Caspase inhibitors rendered cells more resistant to hyposmolar stress; the caspase 3 inhibitor Ac-Asp-Glu-Val-aspartic acid aldehyde was the most efficient. After 24 h of stress, HT-29 colon carcinoma and HK-2 cells had increased their mitochondrial mass. This went along with an increase in mitochondrial membrane potential in HT-29 cells but with a decrease in HK-2 cells. Starting at 2 h of stress, we detected transient CD95L transcription followed by surface expression of CD95L in HT-29 but not in HK-2 cells. Inhibitory CD95L antibody partially inhibited specific death in HT-29 but not in HK-2 cells. Thus, as in other types of stress-induced apoptosis, the CD95/CD95L system is one of the different routes to suicide optionally used by hyposmotically stressed cells. Our findings may have clinical implications for the prevention and treatment of tissue damage caused by severe hyposmolar states.

hyposmolarity; cell stress; mitochondrial membrane potential; caspase 3; CD95; CD95 ligand


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

STRESS-INDUCED APOPTOSIS is the terminal response of mammalian cells to a variety of environmental physical stimuli that critically destabilize the cell's integrity and functional homeostasis. Osmotic stress is a critical alteration of the ionic strength of the extracellular medium in the direction of hyper- or hyposmolarity. Both conditions have been applied to cells as osmotic shock, meaning that cells were exposed to hyper- or hyposmolar medium for a few minutes to a few hours, followed by a recovery time, after which cells were analyzed for their response. The first visible alteration is cell shrinkage under hyperosmolar conditions and cell swelling under hyposmolar conditions. Both states induce volume-regulatory mechanisms (25). Hyperosmotic stress rapidly (within minutes) triggers the mitogen-activated protein kinases (MAPKs), extracellular signal-regulated kinase (ERK) 1/2, c-Jun NH2-terminal kinase (JNK) 1/2, and p38 (37, 11), and activates heat shock transcription factor 1 (HSF1) (10). Astonishingly enough, the adverse stress, hyposmotic shock, has very similar effects. Hyposmotic shock of 5-min duration leads to activation of HSF1 (10), while after 10-min duration ERK1/2, the p38 MAPK cascade, and stress-activated protein kinase/JNK are phosphorylated (23, 41). These effects may not be ubiquitous but might be cell-type associated because MAPKs were not found to be phosphorylated by hyposmotically shocked human fibroblasts (30). By a pathway not entirely understood, hyposmotic stress later leads to nuclear factor (NF)-kappa B activation, prolonged NF-kappa B binding activity, and Ikappa B-alpha degradation (23, 33). The effects of NF-kappa B-activated gene transcription are multiple and very complex (21). In most cells, NF-kappa B activation protects the cell from apoptosis, through induction of survival genes. It may, however, also promote apoptosis, e.g., in glutamate-induced toxicity in neuronal cells, where NF-kappa B causes cell death (3, 11, 39).

Osmotic stress in a natural context is likely to last longer than the exposure time of osmotic shock. It is therefore of interest to study the effects of long-term exposure to hyper- and hyposmotic stress. A recent study reports that in human fibroblasts, 4 h of exposure to hyperosmotic (800 mosmol/l) and hyposmotic stress (100 mosmol/l) leads to apoptosis (30). We undertook this study to examine this late effect of hyposmolarity in more general terms. We report here that sustained hyposmotic stress is an apoptosis inducer with dose- and time-dependent dynamics operative in every cell type examined. The various cell types differ over a considerable range in their susceptibility/resistance and use different apoptotic pathways.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Cell lines and culture conditions. A panel of different lines was chosen to examine a broad spectrum of cell types. Colon carcinoma cell lines HT-29, COLO-205, SW-1116, the pancreas carcinoma cell line AsPc1, the ALL cell line Reh, as well as the SV40-transfected human kidney cell line HK-2, were obtained from American Type Culture Collection (ATCC, Rockville, MD). The B-cell lymphoma line MedB-1 was established in our laboratory (4). Cells were seeded in six-well plates and grown in IMDM (PAA, Linz, Austria)/RPMI (Life Technologies, Paisley, UK) 4:1 supplemented with 10% fetal calf serum (PAA), 5 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin (Biowhittaker, Heidelberg, Germany) until confluence. Human coronary artery smooth muscle cells (SMC) were obtained from Biowhittaker and also seeded in six-well plates and maintained in RPMI supplemented with 5% fetal calf serum, 5 mM glutamine, 0.5 ng/ml epidermal growth factor, 2 ng/ml fibroblast growth factor, 5 mg/ml insulin, 60 µg/ml gentamycin, and 1.5 µg/ml amphotericin (Biochrom, Berlin, Germany) until confluence. All cells were maintained at 37°C in 5% CO2 atmosphere.

Hyposmotic stress. The above medium was removed from the confluent cultures, and cells were incubated with IMDM/RPMI diluted with distilled water to obtain hyposmolar conditions ranging from 264 to 84 mosmol/l. To exclude possible side effects caused by changes in tonicity, HT-29 cells were incubated for 48 h with either isotonic saline or 84 mosmol/l diluted media supplemented with raffinose to a physiological level (294 mosmol/l) as controls. In one set of experiments, cells were incubated with 84 mosmol/l hyposmolar solution containing either 50 µM Z-Val-Ala-Asp-fluoromethylketone (ZVAD-fmk), a broad spectrum caspase inhibitor, 100 µM Ac-Val-Glu-Ile-L-aspartic acid aldehyde (Ac-VEID-CHO), a caspase 6 inhibitor, or 100 µM Ac-Asp-Glu-Val-aspartic acid aldehyde (Ac-DEVD-CHO), a caspase 3 inhibitor (12), all purchased from Bachem (Heidelberg, Germany). Hyposmolar solution and caspase inhibitors were changed every 48 h. In blocking experiments, the CD95L-neutralizing monoclonal antibody NOK-1 (PharMingen, San Diego, CA) was added every 24 h at a concentration of 1 µg/ml. Cells were harvested every 24 h until a death rate of at least 80%. To detect early responses at the mRNA level, cell samples were taken at 2, 4, 6, 8, and 10 h after the onset of hyposmotic stress. Cells were washed once with PBS without magnesium, calcium, and sodium bicarbonate (Life Technologies), then harvested with 1:250 trypsin (500 mg/l):EDTA (200 mg/l) (Biowhittaker). Cells were resuspended in the supernatant for cytospin preparations, centrifuged at 2,000 g for 5 min at room temperature, and further prepared for flow cytometry analysis of DNA content.

Cytospin preparations. An aliquot of complete cell suspension was filled into cytospin chambers and pelleted at 650 rpm for 3 min to place cells on slides. The slides were air-dried, fixed in methanol, and then stained in May-Grünwald (Merck, Darmstadt, Germany) for 5 min and 1.5% Giemsa (Merck) in H2O for 15 min. The slides were rinsed in H2O and air-dried before examination.

DNA fragmentation. Cell pellets were incubated with lysis buffer as described by Walker et al. (44). Cell lysates were directly loaded on a 1.5% agarose gel, then electrophoretically separated, ethidium-bromide stained, and visualized by an ImageMaster VDS (Pharmacia Biotec, San Francisco, CA).

Confocal laser scanning microscopy. To examine the localization of CD95L, the cell lines HT-29 and HK-2 were cultured on Lab-Tek II borosilicate-coated chamber slides (Nalge Nunc, Naperville, IL) under the same conditions as described above, then treated with 84 mosmolar solution for 30 h. Cells were incubated with the monoclonal CD95L antibody G247-4 (IgG1 isotype; PharMingen) and monoclonal antibody CD95(DX2) (IgG1 isotype; Dako, Copenhagen, Denmark) after washing twice with Dulbecco's modified Eagle's medium (Life Technologies) for 30 min at 37°C. After three washing steps, FITC-conjugated anti-mouse F(ab)2 (Dako) and HOE-33256 (Sigma-Aldrich, Deisenhofen, Germany) for nuclear staining were added for another 30 min. Cells were washed again four times, then kept in DMEM with low-level phenol red to quench autofluorescence. Cells were examined using an inverted Leica TCS SP/UV confocal laser scanning microscope and the Scan system software (Leica Microsystems, Heidelberg, Germany).

Flow cytometric analysis of DNA fragmentation. To quantify cells with advanced DNA degradation, we used a procedure described by Nicoletti et al. (34). In short, ~106 cells per sample were gently resuspended in 500 µl of hypotonic fluorochrome solution containing 0.1% Triton X, 0.1% sodium citrate, and 50 µg/ml propidium iodide (Sigma-Aldrich). The cell suspensions were placed at 4°C in the dark overnight before flow cytometric analysis; 104 events were examined for each determination. Percentage-specific death is defined as percentage DNA fragmentation in the presence of hyposmotic medium (84 mosmol/l) compared with percentage DNA fragmentation of the control kept in isosmolar medium. Flow cytometry was performed on a FACScalibur and with CellQuest software (Becton Dickinson, Mountainview, CA).

Flow cytometric analysis of annexin V binding. To detect annexin V binding as a marker of early apoptosis, cells exposed to hyposmolar medium (264, 213, 154, and 84 mosmol/l) for 48 h were harvested as described above. Cells kept in isosmolar medium served as controls. After washing twice in PBS, cells were incubated with 5 µl of a FITC-labeled mouse anti-human annexin V monoclonal antibody (IgG1) (PharMingen) and 2 µl propidium iodide (50 mg/ml) for 15 min in the dark. After gating for propidium iodide negativity to exclude dead cells, 105 events were examined for each determination. Flow cytometry was performed on a FACScalibur and with CellQuest software.

Flow cytometry of surface CD95/CD95L expression. The following primary mouse anti-human monoclonal antibodies (MAb) were used: CD3(Leu4) (IgG1 isotype; Dako) as an isotype-matched negative control, CD95L antibody NOK-1 (IgG1 isotype; PharMingen), and CD95(DX2). Immunofluorescent staining was performed in polystyrene round-bottom tubes (Falcon, San Jose, CA). For dilutions and washings, Hanks' balanced salt solution containing 2% bovine serum albumin and 0.1% sodium azide was used, referred to as fluorescence-activated cell sorter (FACS) medium. After treatment with hyposmolar solution for 30 h, ~106 cells/sample were resuspended in FACS medium and incubated on ice with the appropriate volume of each MAb. After 1 h, cells were washed twice in FACS medium, and 2 µg of FITC-labeled F(ab')2 goat anti-mouse immunoglobulins (Dako) was added for another 30 min. Cells were washed twice and resuspended in 300 µl FACS medium containing 1 µg/ml propidium iodide (Sigma-Aldrich) to allow selective gating of dead cells showing propidium iodide positivity.

Preparation of RNA and cDNA synthesis. Total RNA was extracted with TRIzol reagent (Life Technologies) according to the manufacturer's instructions, precipitated with 1 vol of 2-propanol, and rinsed with 70% ethanol. Total RNA was digested with RNase-free DNase I (Boehringer-Mannheim, Mannheim, Germany) for 30 min at 37°C and precipitated with 3 vol of ethanol at -20°C for 1 h. The RNA pellet was air-dried and dissolved in diethyl pyrocarbonate (DEPC)-water. Optical density (OD) was measured at 260 and 280 nm. RNA was quantified by the equation OD260 = 1 = 40 µg/ml, where OD260 is the OD measured at 260 nm. The OD260/280 ratio showed values between 1.6 and 1.8, as required for pure RNA content. To assure the purity of the RNA, a PCR as described below was performed with an SP1-primer and a RNA template and yielded no amplification product (data not shown). Five micrograms of total RNA were incubated with 1 µl of poly(dT)15 (500 µg/ml), denatured at 80°C for 5 min to guarantee linear cords, followed by a brief centrifugation and quick chill on ice. First-strand buffer (5×), 0.1 M dithiothreitol, 10 mM dNTP-mix, 1 unit of Superscript reverse transcriptase (Life Technologies), 40 units of RNasin (Promega, Madison, WI), and DEPC-water were added. cDNA synthesis was performed with a DNA thermal cycler (Perkin Elmer, Norwalk, CT).

PCR. Semiquantitative triplex PCR was performed on cDNA using the Taq PCR core kit (Qiagen, Hilden, Germany) and the following sense and antisense primers: human transcription factor SP1, 5'-ACT ACC AGT GGA TCA TCA GGG-3' and 5'-CTG ACA ATG GTG CTG CTT GGA-3'; CD95L, 5'-GGA TTG GGC CTG GGG ATG TTT CA-3' and 5'-TTG TGG CTC AGG GGC AGG TTG TTG-3'. The length of amplicons are 241 bp and 344 bp. Thermocycling included the following steps: denaturation at 95°C for 5 min, 25 cycles of 95°C for 1 min, 56°C for 1 min, 72°C for 1 min, and prolongation for 10 min at 72°C. Primers were used at a final concentration of 0.5 µM each, dNTPs at 10 µM, 2.5 units of Taq DNA polymerase in a total of 50 µl, 10× buffer, and 5× Q-Solution according to the manufacturer's instructions. Omitting reverse transcriptase reaction resulted in no detectable PCR products. Electrophoretically separated PCR products were ethidium bromide stained, and the fluorescence images of the amplicons were analyzed by ImageMaster VDS (Pharmacia Biotec, San Francisco, CA). The ratios of the background-corrected integrated optical densities of the DNA bands related to SP1 expression were calculated.

Flow cytometric analysis of mitochondrial mass and membrane potential. To measure both mitochondrial mass and differences in the mitochondrial membrane potential (Delta psi m), cells treated with hyposmolar (84 mosmol/l) solution for 24 h were incubated with 10 µg/ml 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1) (Molecular Probes, Leiden, Netherlands). This dye accumulates in the mitochondrial matrix under the influence of the mitochondrial membrane potential difference and forms J aggregates that have characteristic absorption and emission spectra (Poot and Pierce, 36). After incubation for 10 min at room temperature in the dark, cells were washed once with PBS and immediately subjected to flow cytometric analysis. The excitation wavelength was 488 nm, and observation wavelengths were 530 nm for green and 585 nm for red fluorescence. After gating out small-sized debris, 2 × 105 events were collected and measured for red and green fluorescence.


    RESULTS
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INTRODUCTION
MATERIALS AND METHODS
RESULTS
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Hyposmotic stress leads to apoptotic cell death. Hyposmotic cell swelling was induced and evidenced by FACS analysis. Cells kept under hyposmolar conditions for a prolonged time period started to change their morphology. We observed membrane blebbing, nuclear fragmentation, and chromatin condensation, suggesting apoptosis (Fig. 1), and this proved to be the case. Apoptotic death was evidenced by annexin V binding studies, flow cytometry, and gel electrophoretical DNA fragmentation assays (see Figs. 2, 3, and 5). Compared with the effects of hyposmotic shock that were observed within a few minutes up to 2.5-3 h (5, 33, 41), apoptosis turned out to be a delayed event, directly related to the extent of hyposmolarity. Compared with the isosmolar situation of controls (294 mosmol/l), we observed increasing amounts of annexin V binding 48 h after transfer to the hyposmolar medium, ranging from 10.4 ± 2.0% (P <=  0.010) for cells exposed to 264 mosmol/l to 50.8 ± 3.2% (P <=  0.010) for cells maintained at 84 mosmol/l (controls kept in isosmolar medium, 2.2 ± 0.1%) (Fig. 2A). Consistently postponed, DNA fragmentation was detected in 6.1 ± 0.4% (P <=  0.063; 264 mosmol/l) to 73.2 ± 0.7% (P <=  0.010; 84 mosmol/l) after 48 h compared with controls (294 mosmol/l; 6.0 ± 0.5%). The percentage of cells with fragmented DNA increased over time (Fig. 2B). In principle, this was true for all cell lines tested. To exclude possible side effects caused by changes in tonicity, HT-29 cells were incubated for 48 h with either isotonic saline or 84 mosmol/l diluted media supplemented with raffinose to a physiological level (294 mosmol/l). For cells exposed to isotonic saline for 48 h, the percentage of DNA fragmentation was 14.9 ± 0.8% (P <=  0.010). For cells exposed to 84 mosmol/l medium supplemented with raffinose, the percentage of fragmented DNA was 6.7 ± 0.9% (P <=  0.344). HT-29 cells kept in isosmolar medium for 48 h showed a DNA fragmentation of 6.0 ± 0.5% (Fig. 2C). Figure 3 paradigmatically depicts DNA ladders of vascular SMCs, COLO-205 and SW-1116 colon carcinoma cells, and AsPc1 pancreas carcinoma cells. However, substantial differences in susceptibility to hyposmotic stress were observed. As shown in Table 1, under identical conditions, death rates of our panel of cells differed within one order of magnitude. Another way to calculate this vulnerability/relative resistance is the half-life at a given hyposmotic condition, which ranged from 20 h (HK-2) to 211 h (AsPc1) (data not shown). Thus hyposmotic stress led to apoptosis in all tested cell lines, with individual variations in velocity and extent. To compare the vulnerability of different cell lines when exposed to several hyposmolar conditions, the percentage of cells with fragmented DNA after 48 h of incubation was detected (Table 1). HT-29 and HK-2 cell lines showed a similar and substantial effect after 48 h of exposure to 84 mosmol/l medium. For HT-29, the percentage of DNA fragmentation was 73.2 ± 0.7% (P <=  0.010) compared with controls kept in isosmolar medium (6.0 ± 0.5%), whereas for HK-2 it was 72.0 ± 1.6% (P <=  0.006; controls 10.3 ± 1.1%). The weakest effect of hyposmotic stress (84 mosmol/l for 48 h) was found in AsPc1 cells. DNA fragmentation was detected in 10.8 ± 0.7% for this cell line (P <=  0.010) compared with controls (294 mosmol/l; 4.1 ± 0.2%).


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Fig. 1.   Exposure to hyposmolar conditions led to an increase in cell size and induced morphological features of apoptosis. HT-29 cells were treated with hyposmolar culture medium (84 mosmol/l) for 48 h and morphologically examined as cytospin preparations (left, May-Grünwald/Giemsa staining) and by flow cytometry measuring forward (FSC) and side scatter (SSC) heights (right, demonstrated as contour plots). A: control cells in isosmolar medium. B: cells in hyposmolar medium.



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Fig. 2.   Hyposmotic stress induced apoptosis in a dose-dependent and time-dependent manner. A: relation between degree of hyposmolarity and percentage of annexin V binding (mean ± SE) of HT-29 cells as revealed by flow cytometry. B: corresponding percentage of DNA fragmentation (mean ± SE) under different degrees of hyposmolarity. The degree of DNA fragmentation was determined by flow cytometrically measuring hypodiploid events after propidium iodide staining. C: to exclude possible side effects such as alterations in tonicity and ion concentration, HT-29 cells were exposed to isotonic saline and 84 mosmol/l diluted medium whose tonicity was corrected to a physiological value (294 mosmol/l) by adding raffinose for 48 h. Values of DNA fragmentation are given as means ± SE.



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Fig. 3.   Hyposmotic stress caused DNA fragmentation in all cell lines tested. Cell lines were exposed to hyposmolar (84 mosmol/l) medium until a flow-cytometrically determined apoptotic death rate of 50% was reached. This was achieved in smooth muscle cells (SMC) at 40 h, in HT-29 at 30 h, in AsPc1 at 144 h, in COLO-205 at 96 h, and in SW-1116 at 72 h. DNAs were run in an ethidium bromide-stained 1.5% agarose gel.


                              
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Table 1.   Specific apoptotic death rates induced by hypoosmotic stress

We restricted the subsequent experiments to two cell lines, a neoplastic intestinal epithelial cell, HT-29, and a nonneoplastic renal tubulus cell, hence an epithelial cell of mesonephric origin, HK-2, which we regarded as suitable since they both had relatively high and comparable death rates (Table 1).

Caspases are critically involved in apoptosis by hyposmotic stress. To further substantiate our observation, we performed caspase inhibition assays. HT-29 and HK-2 cells were incubated in 84 mosmol/l medium with or without the caspase inhibitors Ac-DEVD-CHO, Ac-VEID-CHO, and ZVAD-fmk, respectively. After 24, 48, and 96 h, cells were harvested, and the sub-G1 peak was flow cytophotometrically determined. In HT-29 cells, each of these inhibitors extensively, although not completely, blocked hyposmotically induced cell death at each time point, whereas hyposmotically stressed control cells showed a steady decrease in viability (Fig. 4A). The broadly reacting caspase inhibitor ZVAD-fmk inhibited specific death in the range of 26.9 ± 0.3% (P <=  0.017) at 24 h to 74.9 ± 2.5% (P <=  0.010) at 96 h compared with control cells with ZVAD-fmk (5.4 ± 0.3%) and untreated controls (6.0 ± 0.5%). Caspase 3 inhibitor Ac-DEVD-CHO was most effective and inhibited specific death in the range of 31.4 ± 0.7% (P <=  0.025) at 24 h to 78.9 ± 3.1% (P <=  0.017) at 96 h (controls with Ac-DEVD-CHO 7.1 ± 0.4%). The caspase 6 inhibitor Ac-VEID-CHO was the drug least efficient in rescuing 24.7 ± 1.2% (P <=  0.017) of cells at 24 h and 52.0 ± 4.9% (P <=  0.010) at 96 h (controls with Ac-VEID-CHO 5.0 ± 0.7%). The situation was rather different in HK-2 cells; although the death rate of hyposmotically stressed control cells was comparable to that in HT-29 cells, caspase inhibitors were clearly less effective: Ac-VEID-CHO marginally increased the survival rate by a maximum of 12.1 ± 2.7% (P <=  0.018) at 96 h. ZVAD-fmk was slightly superior in rescuing 39.5 ± 2.8% (P <=  0.024) of cells at 24 h and 44.7 ± 2.9% (P <=  0.018) at 96 h (controls with ZVAD-fmk 7.4 ± 1.1%). Ac-DEVD-CHO rescued 45 ± 2.5% (P <=  0.024) at 24 h and 48.3 ± 2.0% (P <=  0.018) at 96 h (controls with Ac-DEVD-CHO 10.0 ± 1.2%) (Fig. 4B). In conclusion, the efficacy of caspase inhibitors was cell type dependent. This suggests that different signaling pathways exist for hyposmotically induced apoptosis.


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Fig. 4.   Caspase inhibition reduced the apoptotic rate. HT-29 and HK-2 cells were exposed to hyposmolar (84 mosmol/l) medium, supplemented with different caspase inhibitors (ZVAD-fmk, Ac-VEID-CHO, Ac-DEVD-CHO), for 24, 48, and 96 h, respectively. The degree of DNA fragmentation (mean ± SE) was determined by flow cytometrically measuring hypodiploid events after propidium iodide staining. A: HT-29 cells. B: HK-2 cells.

Hyposmotic stress leads to changes in relative mitochondrial mass and mitochondrial membrane potential Delta psi m. Because caspase inhibitors did not completely prevent apoptosis by hyposmotic stress in HT-29 cells and, even less so, in HK-2 cells, we examined by JC-1 staining (36) whether and to what extent the mitochondrial mass and membrane potential are altered under this condition. After 12 h of exposure to hyposmolar (84 mosmol/l) medium, both cell lines showed a shift in JC-1 green fluorescence, indicating an increase in relative mitochondrial mass as a function of green fluorescence intensity (bold lines in Fig. 5, A and A') (31). This effect persisted at 24 h of exposure. Furthermore, both cell lines featured changes in mitochondrial membrane potential by their JC-1 red fluorescence. Bivariate analysis of mitochondrial membrane potential (Fig. 5, B and B') further revealed that, after 12 h of stress, the mitochondria of both HT-29 and HK-2 had decreased their membrane potential. This decrease was more pronounced in HK-2 cells. After 24 h, the HT-29 line had regained the initial level of membrane potential in the majority of cells, although there were clearly more cells with high levels of green fluorescence compared with the control. In contrast, HK-2 cells continuously shifted from red to green fluorescence. Because the transit from red to green fluorescence is regarded as a measure for loss of mitochondrial membrane potential, HK-2 cells, while dying, showed a higher degree and extent of mitochondrial depolarization than HT-29 cells. This suggests that the mitochondrial apoptotic pathway was more extensively involved in HK-2 than in HT-29 cells.


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Fig. 5.   Hyposmotic stress alters the mitochondrial mass and mitochondrial membrane potential (Delta psi m). The intensity of green fluorescence is a function of the mitochondrial mass, whereas the ratio of green to red fluorescence of JC-1 depends on the mitochondrial membrane potential Delta psi m (see MATERIALS AND METHODS). HT-29 (A and B) and HK-2 cells (A' and B') were exposed to hyposmolar medium for 12 and 24 h. Subsequent staining with JC-1 for green fluorescence intensity detected by flow cytometry is shown in histograms at 12 and 24 h (thin line, control; bold line, 84 mosmol/l) (A and A'). Bivariate cytograms display the JC-1 green and red fluorescence intensities of control (0 h) and hyposmotically stressed (12, 24 h) HT-29 and HK-2 cells (B and B').

Involvement of CD95/CD95L in apoptosis by hyposmotic stress. HT-29 and HK-2 cells showed comparable death rates (Table 1) but reacted differently on caspase inhibitors. This prompted us to examine the potential role of the CD95/CD95L system in this experimental setting. Constitutively, HT-29 and HK-2 cell surfaces express CD95 at low levels (data not shown). We have previously published evidence showing that even the low CD95 levels expressed on HT-29 cells may be sufficient to transmit the signal, provided that the cell is sensitive to CD95-mediated apoptosis (43). Maintenance of both HT-29 and HK-2 cells in 84 mosmol/l for up to 30 h did not change either the amounts of CD95 transcripts or the levels of CD95 surface expression (data not shown). However, hyposmotic stress induced CD95L transcripts in both cell lines which, constitutively, are devoid of CD95L mRNA (Fig. 6). At 84 mosmol/l CD95L mRNA was detected as early as 2 h after exposure in both cell lines. mRNA levels rose only slightly over the next 8 h, to subsequently decline and drop below the detection level at later time points (Fig. 6). Next, we examined living HT-29 and HK-2 cells for surface expression of CD95L by laser scanning microscopy. As shown in Fig. 7, after 30 h of exposure to 84 mosmol/l, HT-29 but not HK-2 cells expressed detectable amounts of CD95L protein. CD95L surface exposure was evidenced by FACS analysis (Fig. 7). Next, we applied the NOK-1 antibody, which neutralizes CD95L (20). As shown in Fig. 7, NOK-1 antibody, when added to the hyposmolar medium, substantially reduced the apoptotic rate in HT-29 cells while the antibody had no effect on HK-2 cells. In conclusion, in HT-29 cells hyposmotic stress induced the protein expression and cell surface exposure of CD95L which, under this condition, acted not as the unique factor but as a cofactor in apoptosis induction. This was not seen in HK-2 cells, suggesting that these cells use a different apoptosis pathway(s).


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Fig. 6.   Hyposmotic stress-induced CD95L transcripts. Cell were exposed to 84 mosmolar medium for 2, 4, 6, 8, and 10 h. CD95L mRNA was detected after RT-PCR amplification. As an intrinsic standard, SP1 mRNA was coamplified. A: HT-29 cells. B: HK-2 cells. Top lanes show CD95L and SP1 transcripts detected in an ethidium bromide-stained agarose gel after RT-PCR. Bottom lane shows ratio profiles of integrated optical densities.



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Fig. 7.   Hyposmotic stress-induced CD95L protein is surface expressed in HT-29 (A) but not in HK-2 cells (B). After 30 h of exposure to hyposmolar (84 mosmol/l) medium, living cells were stained by FITC-labeled CD95L monoclonal antibody (MAb) G247-4 and by Hoechst-33256 dye for nuclear staining and subjected to confocal laser microscopy. Left micrographs, control cells; right micrographs, cells after hyposmotic stress. Correspondingly, control cells and stressed cells were flow-cytometrically analyzed for surface expression of CD95L (left histograms). In parallel, cells were exposed to hyposmolar medium in the presence and in the absence of the CD95L inhibitor MAb NOK-1. Subsequently, DNA content of cells with and without antibody treatment was compared after propidium iodide (PI) staining (right histograms).

In conclusion, sustained hyposmotic stress induces apoptosis. Cells respond in a time-dependent and dose-dependent fashion while showing substantial differences in constitutional sensitivity/resistance to this kind of stress. Furthermore, different apoptotic mechanisms are operative, involving the mitochondrial pathway to different extents. As in other types of stress-induced apoptosis, the CD95/CD95L system is one of the triggers optionally used.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Hyposmolarity after hyponatremia is the core symptom of the syndrome of inappropriate antidiuretic hormone (ADH) secretion, which is associated with malignant neoplasia, infectious diseases, and central nervous system and psychiatric disorders. Hyponatremic hyposmolarity also occurs as an undesired therapeutic side effect (35, 40) in glucocorticoid deficiency, hypothyroidism (32), and, by accident, as a consequence of polydipsia (water-intoxication; Ref. 15) or as an iatrogenic complication (32). Moreover, salt-losing nephropathy occurs in some patients with advanced chronic renal disease who are unable to conserve sodium. This disorder can be associated with medullary cystic disease, polycystic kidney disease, analgesic nephropathy, chronic pyelonephritis, and obstructive nephropathy (24). Severe hyponatremic hyposmolarity causes significant neurological damage (cerebral edema, osmotic demyelinating syndrome) and is associated with an increased mortality rate (24, 35). In the past, hyposmolar cell damage was mainly regarded as mechanical lysis resulting from progressive cell swelling. This may indeed be true for erythrocytes and extreme hyposmolar conditions. In the more natural context, however, things are not that simple.

Here we have shown that sustained hyposmotic stress leads in a dose-dependent and time-dependent manner to late-onset apoptosis. All cell types tested underwent apoptosis but varied within one order of magnitude in sensitivity/reactivity. This suggests that different cell types use different means to compensate hyposmotic stress and survive before surrendering to this adverse environmental condition by committing suicide. This hypothesis will have to be tested in further studies. Apart from these alleged differences, we have shown that there is more than one route to hyposmotically enforced cell death. HT-29 cells and HK-2 cells, although featuring very similar death kinetics, differed considerably in the death programs they activated.

We have shown that HT-29 cells and HK-2 cells make different use of the mitochondrial pathway toward apoptosis in sustained hyposmotic stress. In murine cytochrome c knockout cells growing in cell culture conditions that compensated for oxidative phosphorylation, stress-induced apoptosis was attenuated (28). This is evidence that release of cytochrome c is an important cofactor in the stress-induced suicide program. Part of the caspase 3 activation we found in both HT-29 and HK-2 cells might be the indirect consequence of cytochrome c release (6), associated with the mitochondrial permeability shift we observed.

In HT-29 cells, hyposmotic stress-induced apoptosis was almost completely blocked by caspase inhibitors and, to a lesser extent, by the addition of CD95L neutralizing antibody. HT-29 cells featured a slight decrease in mitochondrial membrane potential at 12 h of exposure followed by a regaining of mitochondrial membrane potential 12 h later. This effect may perhaps be due to the observable augmentation of the mitochondrial mass and/or to CD95 signaling itself (2). Altogether, this setting suggests that HT-29 cells behave like type 1 cells in CD95-mediated cell death (38). Type 1 cells execute apoptosis mainly by death-inducing signaling complex (DISC) formation and caspase 8 activation followed by activation of the effector caspase 3. The slight decrease in mitochondrial membrane potential after 12 h may be due to a massive activation of caspase 8 known to initiate the mitochondrial pathway by BID cleavage (27). In HK-2 cells, caspase inhibitors were less protective, CD95L antibody showed no effect, and the decrease in mitochondrial membrane potential was considerably higher compared with HT-29. Although not formally excluded, it appears nonetheless unlikely that other members of the tumor necrosis factor receptor superfamily play a major role in triggering the suicide of HK-2 cells. Upon various stimuli (cellular stress), mitochondria alone are able to release apoptogenic factors like apoptosis-inducing factor (AIF) or TR3 (16). The mitochondrial AIF is reported to induce peripheral chromatin condensation and DNA fragmentation, independent of caspase activation (29). TR3, an orphan member of the steroid-thyroid hormone-retinoid receptor superfamily, was recently described as a proapoptotic transcription factor, causing mitochondrial membrane depolarization and cytochrome c release (7). The trigger molecules and their signaling partners upstream of the mitochondrion, which are involved in hyposmotic stress-enforced apoptosis, are still elusive.

We have shown that hyposmotic stress leads to a transient expression of CD95L transcripts beginning after 2 h. It was detectable for ~10 h and vanished thereafter. This was followed by cytoplasmic CD95L protein synthesis in HT-29 but, at least at 30 h, not in HK-2 cells. At this time point, HT-29 cell surface expressed CD95L, and the addition of inhibitory CD95L antibody specifically reduced the apoptotic rate. This is evidence that, as one of the major death receptors, CD95 is optionally involved in hyposmotic stress-induced apoptosis. Stress-induced induction or upregulation of CD95L expression has already been observed for gamma irradiation and UV exposure (9, 15), oxidative stress (8, 42), and survival factor withdrawal (26). Hyposmolarity adds to this list of adverse environmental conditions, triggering the stress kinase pathway. In different cell systems NF-kappa B activation was followed by upregulation of CD95L (17, 18, 22). It will have to be clarified whether the transient CD95L mRNA expression is induced by NF-kappa B activation, which is known to be one of the consequences of hyposmotic shock (23, 33). CD95L gene expression might also/alternatively be initiated by MAPK kinase (MEK) kinase 1 (MEKK1)-mediated CD95L promoter activation (14). Because in our experiments caspase 3 inhibition was very efficient in reducing the apoptotic rate, MEKK1, known to be cleaved and activated by caspase 3 (Ref. 7), might be critically involved at this point.

In conclusion, transient hyposmotic shock activates antiapoptotic programs, probably supporting counterregulatory mechanisms. Sustained hyposmotic pressure leads to a delayed type of enforced cell death, apoptosis by defeat. Blocking the apoptotic machinery strikingly augments the resistance to this stress and makes the cells stay alive. Transported into the clinical context, our findings might form a rational basis for novel strategies for therapeutic intervention, e.g., when cells with vital functions and/or restricted regenerative capacity like neurons are severely endangered by hyposmolarity.


    ACKNOWLEDGEMENTS

We thank A. Gruber and S. Röderer for excellent technical assistance and C. Higginson for editorial help.


    FOOTNOTES

* T. Jäckle and C. Hasel contributed equally to this work.

This work was supported by Deutsche Forschungsgemeinschaft Grant SFB-518/A6 to P. Möller.

Address for reprint requests and other correspondence: P. Möller, Institute of Pathology, Univ. of Ulm, Albert-Einstein-Allee 11, D-89081 Ulm, Germany (E-mail: peter.moeller{at}medizin.uni-ulm.de).

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.

Received 21 December 2000; accepted in final form 5 July 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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