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Molecular and Cellular Biology, July 2005, p. 6259-6266, Vol. 25, No. 14
0270-7306/05/$08.00+0     doi:10.1128/MCB.25.14.6259-6266.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Inhibiting Myosin Light Chain Kinase Induces Apoptosis In Vitro and In Vivo

Fabeha Fazal,¹ Lianzhi Gu,¹ Ivanna Ihnatovych,¹ YooJeong Han,¹ WenYang Hu,¹ Nenad Antic,¹ Fernando Carreira,¹ James F. Blomquist,² Thomas J. Hope,² David S. Ucker,² and Primal de Lanerolle^1*

Departments of Physiology and Biophysics,¹ Microbiology and Immunology, College of Medicine, University of Illinois at Chicago, Chicago, Illinois 60612²

Received 6 February 2004/ Returned for modification 19 March 2004/ Accepted 13 April 2005

Top Abstract Introduction Materials and Methods Results Discussion References

 
Previous short-term studies have correlated an increase in the phosphorylation of the 20-kDa light chain of myosin II (MLC₂₀) with blebbing in apoptotic cells. We have found that this increase in MLC₂₀ phosphorylation is rapidly followed by MLC₂₀ dephosphorylation when cells are stimulated with various apoptotic agents. MLC₂₀ dephosphorylation is not a consequence of apoptosis because MLC₂₀ dephosphorylation precedes caspase activation when cells are stimulated with a proapoptotic agent or when myosin light chain kinase (MLCK) is inhibited pharmacologically or by microinjecting an inhibitory antibody to MLCK. Moreover, blocking caspase activation increased cell survival when MLCK is inhibited or when cells are treated with tumor necrosis factor alpha. Depolymerizing actin filaments or detaching cells, processes that destabilize the cytoskeleton, or inhibiting myosin ATPase activity also resulted in MLC₂₀ dephosphorylation and cell death. In vivo experiments showed that inhibiting MLCK increased the number of apoptotic cells and retarded the growth of mammary cancer cells in mice. Thus, MLC₂₀ dephosphorylation occurs during physiological cell death and prolonged MLC₂₀ dephosphorylation can trigger apoptosis.

Top Abstract Introduction Materials and Methods Results Discussion References

 
The ability of the cytoskeleton to deform and reform is a crucial aspect of many cellular responses (5). This is especially true of motile and dividing cells where the cytoskeleton must deform and reform on demand. Interactions between cells and the extracellular matrix also appear to be important in cell survival (22). Integrin ligation by the extracellular matrix plays a crucial role in organizing the cytoskeleton (25), and the loss of substrate attachment is known to induce apoptosis (anoikis) (14). On the other hand, studies on epithelial cells grown in three-dimensional culture have shown that integrin-extracellular matrix interactions promote the organization of the cytoskeleton and resistance to apoptotic stimuli (42).

The organization and stiffness of the cytoskeleton are determined in large part by the forces generated by actin and myosin II (12). The actin-myosin II interaction in smooth muscle and nonmuscle cells is regulated by the phosphorylation of serine 19 of the 20-kDa light chain of myosin II (1, 11, 37, 39, 44). This reaction, which is catalyzed by myosin light chain kinase (MLCK), stimulates the actin-activated, Mg²⁺-dependent ATPase activity of myosin II (1). Work from many laboratories has shown that MLC₂₀ phosphorylation and dephosphorylation are required for smooth muscle contraction and relaxation (for reviews, see references 11, 37, and 39). Other experiments have shown that MLC₂₀ phosphorylation/dephosphorylation plays a central role in cell motility (25, 33, 43, 45), endothelial (41, 46) and epithelial (3, 15, 19) barrier function, and cell division (13, 34, 47).

Apoptosis is a carefully regulated cellular process that is important in developing and maintaining tissue homeostasis (40). Dysregulation of the apoptotic process underlies pathologies including cancer, autoimmune diseases, and neurodegenerative disorders. Biochemical events associated with apoptosis include caspase activation, mitochondrial disruption, and genome digestion (20, 24). Another hallmark of apoptosis is a profound change in cell shape that is apparently mediated by restructuring the cytoskeleton. While actin (4) and actin-binding proteins (26) have been implicated in mediating these cytoskeletal changes, the role of myosin II in apoptosis is poorly understood. Because actin and myosin II work together to stabilize the cytoskeleton and to define cell shape, we investigated how MLCK and the phosphorylation/dephosphorylation of the 20-kDa light chain of myosin II (MLC₂₀) are involved in apoptosis. In the present study we show that MLC₂₀ is dephosphorylated during apoptosis and that the dephosphorylation of MLC₂₀, effected by destabilizing the cytoskeleton or by direct inhibition of MLCK, triggers cell death. We also show that targeted inhibition of MLCK induced apoptosis in vivo.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Cell culture. Smooth muscle cells (SMC) were isolated from porcine pulmonary artery by enzymatic digestion as described previously (7). Cells were grown in culture dishes in Dulbecco's modified Eagle medium (DMEM; Gibco BRL, Gaithersburg, MD) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin and streptomycin. Cells were not used beyond seven passages. All drug treatments were performed in DMEM containing 0.5% FBS without antibiotics.

Measurement of MLC phosphorylation. Changes in MLC₂₀ phosphorylation in NIH 3T3 cells, HeLa cells, or SMC were quantified essentially as described by Chew et al. (8). Briefly, floating and adherent cells were collected and washed with phosphate-buffed saline (PBS) and the cellular proteins were precipitated with ice-cold 10% trichloroacetic acid and 10 mM dithiothreitol (DTT). The pellets were washed with acetone; dissolved in 9 M urea, 10 mM DTT, and 20 mM Tris, pH 7.5; and separated using glycerol-urea polyacrylamide gel electrophoresis. The proteins were transferred to nitrocellulose, and the un-, mono-, and diphosphorylated forms of MLC₂₀ were identified using an affinity-purified antibody to MLC₂₀ (30) and horseradish peroxidase-linked secondary antibody (Jackson ImmunoResearch, West Grove, PA). Protein bands were visualized with enhanced chemiluminescence reagent, and the stoichiometry of phosphorylation (mol PO₄/mol MLC₂₀) was calculated as described previously (30).

Fluorescence-activated cell sorter analysis. Cells were trypsinized; washed twice with cold PBS; resuspended in 100 µl of 10 mM HEPES, pH 7.4, 140 mM NaCl, and 2.5 mM CaCl₂ (binding buffer); and incubated with 5 µl of fluorescein isothiocyanate (FITC)-conjugated annexin V (Pharmingen, San Diego, CA) and 10 µl of propidium iodide (PI; 50 µg/ml) for 15 min in the dark at 25°C. After incubation, 400 µl of binding buffer was added per sample and cells were analyzed cytofluorimetrically using a Coulter Epics Elite ESP flow cytometer (excitation, 488 nm; emission, 585 nm). At least 10,000 cells were counted per analysis, and cells that stained positive for annexin V and PI were judged to be apoptotic.

Caspase assays. Two types of assays were performed to detect caspase activation. In one, NIH 3T3 cells were extracted in HKEB (100 mM HEPES, pH 7.4, 10 mM MgCl₂, 5 mM EGTA, 100 µM phenylmethylsulfonyl fluoride, 1 mM DTT) containing 50 µg/ml digitonin and assayed for DEVD-specific caspase activity using fluorogenic acetyl-Asp-Glu-Val-Asp-4-methyl-coumaryl-7-amide (DEVD-MCA; Peptides International, Louisville, KY) as previously described (17, 18). In the second, SMC grown in six-well plate were lysed in 200 µl ice-cold 50 mM Tris-HCl, 150 mM NaCl, 1% NP-40, 1 mM EGTA, pH 7.4, and protease inhibitors. The soluble fraction was collected by centrifugation (20 min at 12,000 x g at 4°C), and protein concentrations were determined using the Bradford assay. Equal amounts of protein were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to a nitrocellulose membrane. Following transfer, the blot was cut into two parts at about 25 kDa. The top was probed with a polyclonal procaspase-3/CPP32 antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) and the bottom with a monoclonal anti-caspase-3 antibody that recognizes the 17-kDa cleaved form (Cell Signaling Technology, Beverly, MA). The blots were incubated with horseradish peroxidase-conjugated secondary antibodies, and immunoreactive bands were detected with enhanced chemiluminescence.

Microinjection experiments. SMC were grown in DMEM containing 10% FBS plus 1% penicillin/streptomycin on fibronectin-coated delta T dishes (Fisher Scientific, Chicago, IL) at a confluency of 50 to 70%. Live cells were injected with affinity-purified goat anti-human immunoglobulin G (IgG) antibodies (BioSource International, Camarillo, CA) or with affinity-purified MLCK antibodies. The antibodies (7 mg/ml) were mixed with either FITC or rhodamine-labeled dextran (Molecular Probes, Eugene, OR) (5 mg/ml) in order to identify injected cells. Microinjections were performed for 30 min because we did not want to incubate the cells injected with the first antibody for a significantly longer period than the second antibody. About 25 cells were injected per coverslip, and this experiment was repeated three times. Following injection, the cells were incubated for 3 to 4 h at 37°C and 5% CO₂ and examined using an Olympus IX70 equipped with a Cooke Sensicam. Because many of the cells injected with the MLCK antibody died and detached from the coverslips, we counted a total of 60 cells per group.

In vivo assays. Mm5MT mouse mammary tumor cells grown in culture were harvested immediately before injection into syngeneic MMTV-C3H/HeN mice. Cells (10⁶) were washed and resuspended in the serum-free DMEM. Healthy, mouse mammary tumor virus-free female mice (14 to 20 weeks old) were anesthetized and the cells injected subcutaneously into the right flank. Following injection, a small horizontal incision was made in the interscapular area and a 100-µl osmotic pump (Alzet, Cupertino, CA) with a 0.25-µl/hour release rate filled with either 27 mM ML-7 in dimethyl sulfoxide (DMSO) (experimental group) or DMSO (control group) was implanted and the wound closed. All procedures were performed according to University of Illinois at Chicago Animal Care Committee guidelines. Animals were monitored daily for the signs of distress and tumor growth. After 14 days, the old osmotic pump was replaced with a new pump filled with 27 mM ML-7 or DMSO. Animals were sacrificed 28 days after the initial surgery, and the tumors were excised, weighed, fixed in the buffered formaldehyde, embedded in paraffin blocks, cut into 5-µm sections, and processed using standard histological methods.

TUNEL staining. Cells grown on coverslips were fixed in 4% paraformaldehyde/PBS and permeabilized in a buffer containing 0.1% sodium citrate and 0.1% Triton X-100 in PBS for 2 min on ice. Permeabilized cells and tissue sections that were prepared as described above were stained with FITC-labeled terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) enzyme reagent using the In Situ Cell Death Detection kit (Roche Molecular Biochemicals, Indianapolis, IN) as described by the manufacturer. Coverslips and tissue sections were mounted using Vectashield containing DAPI (4',6'-diamidino-2-phenylindole) and examined using a Zeiss LSM 510 laser confocal microscope.

Statistical analyses. Results are expressed as means ± standard errors. The data were analyzed using an unmatched Student t test or one-way analysis of variance (ANOVA) (SigmaStat; Systat, Point Richmond, CA). The type of analysis and significance are stated in the figure legends.

Top Abstract Introduction Materials and Methods Results Discussion References

 
MLC₂₀ dephosphorylation during apoptosis. MLC₂₀ is phosphorylated on Thr 18 and Ser 19 by MLCK (39). When NIH 3T3 cells (Fig. 1A) or HeLa cells (not shown) were treated with actinomycin D, there was an initial increase in MLC₂₀ phosphorylation that was due largely to an increase in diphosphorylation. Following this initial increase, MLC₂₀ was rapidly dephosphorylated. The band representing diphosphorylated MLC₂₀ is completely lost and the band representing monophosphorylated MLC₂₀ is significantly decreased by 8 h. Furthermore, MLC₂₀ dephosphorylation precedes caspase activation (Fig. 1A), suggesting that it is a relatively early event in apoptosis. The decrease in MLC₂₀ phosphorylation was not due to proteolysis of MLC₂₀, myosin II, or MLCK because Western blot analyses of NIH 3T3 cells showed the presence of similar amounts of these proteins at all time points (data not shown).

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FIG. 1. MLC20 dephosphorylation during apoptosis. (A) NIH 3T3 cells were treated with 500 nM actinomycin D for the indicated times. MLC20 phosphorylation, caspase activation, and cell death were quantified as described in Materials and Methods. The inset shows the migration of the MLC20 species (un-, mono-, and diphosphorylated MLC20 from top to bottom). Note the initial increase followed by a decrease in MLC20 phosphorylation. *, P value <0.001, ANOVA (n = 3). (B) The stoichiometry of MLC20 phosphorylation (see inset) and cell viability were quantified in untreated SMC (Un) and SMC treated with 10 µM dexamethasone (Dex), 10 µM actinomycin D (ActD), 100 µM cycloheximide (Chx), or 2 µM camptothecin (Cam) for 24 h. *, P value <0.001, t test (n = 3).

To determine the generality of this observation, we treated primary cultures of SMC with various agents that induce apoptosis. Actinomycin D, cycloheximide, or camptothecin resulted in MLC₂₀ dephosphorylation and apoptosis (Fig. 1B). Dexamethasone, which induces apoptosis in other cell types (17), neither dephosphorylated MLC₂₀ nor induced smooth muscle cell death. These data suggest a specific link between prolonged and extensive MLC₂₀ dephosphorylation and cell death.

Inhibiting MLCK leads to MLC₂₀ dephosphorylation, caspase 3 cleavage, and apoptosis, in vitro. We next asked if MLC₂₀ dephosphorylation triggers cell death. SMC were treated with ML-7 or KT5926, two inhibitors of MLCK (2, 16, 29). ML-7 resulted in a dose-dependent decrease in MLC₂₀ phosphorylation and a corresponding increase in cell death (Fig. 2A and B). Similarly, KT5926 resulted in cell death, including genome digestion, in NIH 3T3 cells (data not shown). As in the case with nonmuscle cells treated with actinomycin D, there was a significant decrease in MLC₂₀ phosphorylation that preceded cell death in SMC treated with 20 µM ML-7 (Fig. 2C). Moreover, the cleaved, active form of caspase 3 and a coincident decrease in the holoenzyme were first detected at 8 h, after the decrease in MLC₂₀ phosphorylation and before the onset of apoptosis. Thus, MLC₂₀ dephosphorylation is an early event in apoptosis that precedes caspase activation and the onset of apoptosis and inhibiting MLCK appears to induce cell death.

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FIG. 2. Inhibiting MLCK leads to MLC20 dephosphorylation and apoptosis. SMC were treated with the indicated concentrations of ML-7 for 16 h (A and B) or with 20 µM ML-7 for different times (C). *, P value <0.001, t test (n = 3). (C, inset) Western blots of cell extracts were probed with antibodies against the inactive 32-kDa caspase-3 precursor or the larger fragment (17 kDa) of activated caspase-3 (top and bottom rows, respectively). Note that the cleaved caspase-3 appears at 8 h following the addition of ML-7. It is preceded by MLC20 dephosphorylation, which takes place within two hours of treatment and before the cells become annexin V positive. *, P value <0.001, ANOVA (n = 3). Panel D shows confocal images of cells microinjected with a control antibody or an affinity-purified inhibitory antibody to MLCK immediately following microinjection (top two rows) and 4 h later (bottom two rows). In this experiment, the control antibody (affinity-purified goat anti-human IgG) was mixed with FITC-labeled dextran prior to injection and the MLCK antibody was mixed with rhodamine-labeled dextran. Although the control antibodies (green cells) had no effect, the cells injected with the MLCK antibody (red cells) displayed morphological characteristics typical of apoptotic cells. These are representative micrographs of cells from three separate experiments.

The role of MLC₂₀ dephosphorylation in apoptosis was investigated further by microinjecting SMC with an affinity-purified, inhibitory antibody to MLCK (10). An affinity-purified, goat anti-human IgG was used as a control. Each antibody was mixed separately with either rhodamine-labeled dextran or fluorescein-labeled dextran as an injection marker. Three to four hours after injection, the cells were scored for morphological markers of cell death without prior knowledge of which antibody had been mixed with which marker (Fig. 2D). While 77% of the cells (48 out of 62) injected with the MLCK antibody had morphological changes typical of apoptotic cells, only 13% of the cells (8 out of 60) injected with the control antibody were apoptotic. A t test showed that these differences were statistically significant (P value < 0.001). Thus, inhibiting MLCK by a variety of means induces apoptosis.

Destabilizing the cytoskeleton induces MLC₂₀ dephosphorylation and apoptosis. The stiffness of the cytoskeleton is determined largely by interactions between actin and myosin filaments (12). Increasing MLC₂₀ phosphorylation stabilizes actin filaments (35) and increases cytoskeletal stiffness (6) whereas cytochalasin D decreases cytoskeletal stiffness (6) and KT5926 leads to the loss of actin filaments (P. de Lanerolle, unpublished data). Moreover, destabilizing actin filaments with cytochalasin D (4) or by expressing the actin-severing protein gelsolin (26) or by interfering with integrin signaling by preventing substrate attachment (14) also led to apoptosis.

Because attachment increases MLC₂₀ phosphorylation and stabilizes actin filaments through integrin signaling (9, 35), we postulated that the loss of substrate attachment would induce apoptosis by abrogating integrin signaling and decreasing MLC₂₀ phosphorylation and cytoskeletal stiffness. We investigated this possibility by growing cells on polyhydroxyethylmethacrylate (polyHEMA)-coated dishes to prevent attachment. Growing cells on polyHEMA resulted in significant MLC₂₀ dephosphorylation and an increase in the number of apoptotic cells (Fig. 3). Similarly, destabilizing the cytoskeleton by treating cells with 10 µM cytochalasin D also resulted in significant MLC₂₀ dephosphorylation and an increase in cell death (Fig. 3).

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FIG. 3. Destabilizing the cytoskeleton leads to MLC20 dephosphorylation and apoptosis. Treating SMC with 10 µM cytochalasin D (CytD) and growing cells on polyHEMA (poly) overnight resulted in significant decreases in MLC20 phosphorylation (A) and significant increases in cell death (B) compared to the untreated controls (Un). Treating cells with various concentrations of blebbistatin also induced cell death (C) and MLC20 dephosphorylation (inset). *, P value <0.05, t test (n = 3).

To further characterize the importance of the actin-myosin II interaction in cell death, we treated smooth muscle cells with blebbistatin. Blebbistatin is a specific inhibitor of myosin II ATPase activity (38). Diminishing MLC₂₀ phosphorylation decreases ATP hydrolysis by actin and myosin II (1, 11, 37, 39) and also induces apoptosis (see above). Therefore, inhibiting myosin ATPase activity with blebbistatin should also induce apoptosis. We found dose-dependent increases in annexin V-positive cells following blebbistatin treatment (Fig. 3C). Interestingly, MLC₂₀ phosphorylation was also decreased in these cells. These data suggest that actomyosin II ATPase activity is distal to MLC₂₀ dephosphorylation in the cell death pathway.

Caspase inhibition blocks apoptosis induced by cytoskeletal destabilization. Because cytochalasin D and loss of adhesion lead to caspase activation, we investigated whether caspases also are involved in cell death induced by inhibiting MLCK. Pretreating with 50 µM z-VAD-fluoromethyl ketone (z-VAD-fmk), a cell-permeable caspase inhibitor (18), protected cells subsequently treated with 20 µM ML-7 from cell death as judged by fluorescence-activated cell sorter analysis (Fig. 4A) and FITC-TUNEL staining (Fig. 4B). Interestingly, while z-VAD-fmk protected cells treated with ML-7 from cell death, MLC-P remained low in these cells, further supporting the idea that caspase activation is downstream of MLC₂₀ dephosphorylation. Other experiments showed that overexpressing Bcl-2 protected against apoptosis induced by ML-7 (not shown). These data are consistent with MLC₂₀ dephosphorylation being an early event in apoptosis that is upstream of caspase activation.

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FIG. 4. Caspase inhibition protects against apoptosis but not MLC20 dephosphorylation. (A) SMC were left untreated or treated with 20 µM ML-7 or 10 ng/ml TNF- plus 5 µM cycloheximide (CHX) for 24 h. Other SMC were treated with 50 µM z-VAD-fmk for 1 h followed by 20 µM ML-7 or 10 ng/ml TNF- plus 5 µM cycloheximide for 24 h. z-VAD-fmk significantly increased cell viability compared to cells treated with ML-7 alone, or with TNF- plus cycloheximide. *, P value <0.01, t test (n = 3). (B) Cells were treated with 50 µM z-VAD-fmk for 1 h, 20 µM ML-7 for 4 h, or 50 µM z-VAD-fmk for 1 h followed by 20 µM ML-7 for 4 h. The cells were then fixed and stained with TUNEL reagent and DAPI. There were very few TUNEL-positive nuclei (blue) in untreated cells or cells treated with z-VAD-fmk. Most nuclei in cells treated with ML-7 were TUNEL positive as indicated by the cyan color from the colocalization of the blue (DAPI) and green (FITC-TUNEL) stains. z-VAD-fmk decreased the number of TUNEL-positive nuclei in ML-7-treated cells.

We also tested the effect of tumor necrosis factor alpha (TNF-) on SMC. TNF- by itself does not induce apoptosis in many cells and is usually used in combination with cycloheximide (18, 36). Preliminary experiments on SMC showed that 10 ng/ml TNF- or 5 µM cycloheximide (in contrast to 100 µM cycloheximide used in Fig. 1B), individually had no effect on cell death or MLC₂₀ phosphorylation (not shown). However, when used in combination, 10 ng/ml TNF- and 5 µM cycloheximide resulted in MLC₂₀ dephosphorylation and cell death (Fig. 4A). Moreover, as with ML-7, zVAD-fmk also protected against cell death induced by TNF- and cycloheximide without affecting the level of MLC₂₀ phosphorylation (Fig. 4A).

Inhibiting MLCK induces apoptosis in vivo. We then asked if MLC₂₀ dephosphorylation is part of the cell death process in vivo. We addressed this question using a mouse mammary cancer model. Mice were inoculated with equal numbers of tumor cells and allowed to form tumors in the continuous presence or absence of ML-7. After 4 weeks, tumors harvested from mice treated with ML-7 were significantly smaller in size relative to tumors from untreated mice. The mean weights ± standard errors were 1.23 ± 0.30 g and 0.56 ± 0.20 g for the tumors removed from the control mice (n = 13) and the mice receiving ML-7 (n = 14), respectively. Furthermore, TUNEL staining showed more apoptotic cells in the sections from the mice receiving ML-7 (Fig. 5). Quantification of the TUNEL-positive nuclei in 500 cells from randomly chosen fields in each group showed that 2.5% and 15.6% of the nuclei were TUNEL positive in tumors removed from control mice and mice receiving ML-7, respectively.

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FIG. 5. Inhibition of MLCK induces apoptosis in vivo. Paraffin-embedded tissue sections of tumors removed from control mice (A to D) and from mice receiving ML-7 (E to H) were stained with FITC-TUNEL (B and F) and DAPI (C and G). Phase-contrast (A and E) and merged TUNEL/DAPI (D and H) images are also shown. Tumors from mice receiving ML-7 (F) showed significantly higher numbers of TUNEL-positive cells than tumors removed from control (B) mice. The cyan color in panels D and H indicates the colocalization of the TUNEL and DAPI staining. Bar = 25 µm.

Top Abstract Introduction Materials and Methods Results Discussion References

 
The cytoskeleton undergoes dramatic changes during apoptosis. Despite the fact that myosin II plays a central role in organizing the cytoskeleton, how myosin II and its regulation by MLC₂₀ phosphorylation are involved in apoptosis is largely unknown. In the present study, we report that inactivating myosin II by MLC₂₀ dephosphorylation is an important part of the mechanism of apoptosis. We have previously correlated increases in MLC₂₀ phosphorylation and cytoskeletal stiffness with slower progression through the cell cycle (6). Thus, the tension generated in the cytoskeleton by actomyosin interactions appears to play an important role in determining cell fate. Equally important is the observation that inhibiting MLCK decreases MLC₂₀ phosphorylation and induces apoptosis. These observations suggest that MLC₂₀ dephosphorylation is part of the cell death process. The generality of this effect, demonstrated in vitro using transformed and nontransformed cells and in vivo using a mouse model of breast cancer, suggests that MLC₂₀ dephosphorylation may be an important step in apoptosis during development, organogenesis, and other physiologically important processes.

Our data demonstrate that two different MLCK inhibitors, ML-7 and KT 5926, induce apoptosis. Although no pharmacological agent is totally specific, the inhibitory effect of ML-7 on MLCK is highly selective. The K_i of ML-7 for MLCK is 0.3 µM, while its K_i for protein kinase A is 21 µM and for protein kinase C is 42 µM (27). Thus, at 20 µM, ML-7 is predicted to inhibit MLCK by about 90% and protein kinase A by less than 50%. In addition, microinjecting affinity-purified inhibitory antibodies to MLCK also resulted in cell death. Although we cannot exclude the possibility that ML-7 is affecting another target besides MLCK, taken together the data strongly suggest an important role for MLC₂₀ dephosphorylation in cell death. It is important to note as well that cell death associated with actinomycin D, cycloheximide, camptothecin, TNF-, cytochalasin D, blebbistatin, and growth on polyHEMA was accompanied by MLC₂₀ dephosphorylation. Thus, MLC₂₀ dephosphorylation is an important step in cell death induced by a broad range of apoptotic stimuli.

We also investigated the ability of Y27632, which inhibits the Rho effector protein ROCK, to induce apoptosis in SMC. ROCK increases MLC₂₀ phosphorylation, either directly by phosphorylating MLC₂₀ or by inhibiting myosin phosphatase 1 (36). Consequently, inhibiting ROCK is predicted to decrease MLC₂₀ phosphorylation and induce apoptosis. However, we found that 20 mM Y27632 resulted in only partial dephosphorylation of MLC₂₀ without significantly increasing the number of apoptotic cells (not shown). These data support the idea that prolonged, almost-complete MLC₂₀ dephosphorylation is required to induce apoptosis.

It has been reported that MLC₂₀ phosphorylation increases during the early stages of the cell death process. Short-term studies that quantified MLC₂₀ phosphorylation up to 4 h after cells were treated with an apoptotic agent have suggested that this increase in MLC₂₀ phosphorylation correlates with blebbing (28, 36). Another study reported that MLC₂₀ phosphorylation peaked at 30 min in MDCK cells treated with TNF- (23). We also found an initial increase in MLC₂₀ phosphorylation in nonmuscle cells (Fig. 1A) but not in smooth muscle cells (Fig. 2C), suggesting some subtle cell specific differences in MLC₂₀ dephosphorylation during apoptosis.

Importantly, our data demonstrate a profound decrease in MLC₂₀ phosphorylation that temporally precedes caspase activation and the onset of cell death in both nonmuscle (Fig. 1A) and smooth muscle (Fig. 2C) cells. The data in Fig. 2C show that significant MLC₂₀ dephosphorylation precedes caspase 3 activation which, in turn, temporally precedes cell death. We also found that zVAD-fmk, a cell-permeable caspase inhibitor (18), protected against cell death without affecting MLC₂₀ phosphorylation (Fig. 4A). These data suggest that MLC₂₀ dephosphorylation is upstream of caspase activation. Consistent with the notion that MLC₂₀ dephosphorylation is part of the cell death process, we found that overexpressing Bcl-2 protected cells against apoptosis induced by ML-7 (not shown). Thus, prolonged and extensive dephosphorylation of MLC₂₀ is apparently an early event in the apoptotic pathway. While not negating a biphasic pattern of change, our data strongly support a central role for MLC₂₀ dephosphorylation in apoptosis.

Interestingly, Petrache et al. (31, 32) reported that caspase-dependent cleavage of MLCK plays a role in apoptosis in endothelial cells. It is important to emphasize that there are two forms of MLCK, with predicted molecular weights of 150,000 and 108,000 according to the amino acid sequences, in endothelial cells. Petrache et al. found that they could inhibit apoptosis induced by TNF- by knocking out only the large form of MLCK, the predominant form of MLCK found in endothelial cells (31). However, MLC₂₀ phosphorylation was not quantified in this paper, and one cannot be entirely sure that the effects described are due solely to changes in MLC₂₀ phosphorylation. Nevertheless, the data in this paper suggest that there is something unique about the large form of MLC and its role in apoptosis in endothelial cells.

That inhibiting MLCK induces cell death suggests that MLC₂₀ dephosphorylation may be part of a broader mechanism involved in determining cell fate. Filaments composed of actin and myosin II determine the physical characteristics of the cytoskeleton (5, 21). These filaments insert into focal adhesions, and actomyosin-based contractility generates tension within a cell by tugging on focal adhesions attached to the extracellular matrix (35). Previous experiments have shown that destabilizing actin filaments with cytochalasin D decreases cytoskeletal tension or stiffness while increasing MLC₂₀ phosphorylation increases cytoskeletal stiffness (6). The loss of focal adhesions also leads to the disruption of the actin cytoskeleton, cell rounding, and, presumably, a decrease in cytoskeletal stiffness. Cytochalasin D, blebbistatin, and the loss of attachment, all of which destabilize the cytoskeleton, led to MLC₂₀ dephosphorylation and cell death, further supporting a connection between the two processes.

It is worth noting that blebbistatin, which inhibits myosin ATPase activity (38), results in apoptosis. It is striking that blebbistatin also causes MLC₂₀ dephosphorylation. That treating cells with actinomycin D, cycloheximide, camptothecin, TNF plus cycloheximide, and cytochalasin D, agents that do not inhibit MLCK activity, also results in MLC₂₀ dephosphorylation and cell death (Fig. 1B and 3A and B) underscores the importance of MLC₂₀ phosphorylation as an important signal of cell survival. Therefore, our working hypothesis is that apoptotic stimuli destabilize the cytoskeleton by an unknown mechanism, which, in turn, results in MLC₂₀ dephosphorylation. Simultaneously, actin-myosin II interactions determine the stiffness of the cytoskeleton (12) and inhibiting MLCK decreases MLC₂₀ phosphorylation and destabilizes the cytoskeleton. Thus, we believe that there is an important reciprocal relationship between MLC₂₀ phosphorylation and cytoskeletal stiffness and that cytoskeletal stiffness plays an important role in determining cell fate.

In summary, the data presented here demonstrate that myosin II is dephosphorylated during apoptosis triggered by a variety of suicidal stimuli and that inhibiting MLCK leads to apoptosis in vitro and in vivo. Moreover, MLC₂₀ dephosphorylation is an early event in cell death because it precedes caspase 3 activation. Other experiments showed that inhibiting MLCK or myosin II ATPase activity, cytochalasin D treatment, and the loss of attachment trigger a common cell death response that involves MLC₂₀ dephosphorylation and the destabilization of the cytoskeleton. One implication of these data is that destabilizing the cytoskeleton, by inhibiting MLCK or other means, may be effective in retarding the growth of transformed cells in vivo.

 
We thank Mei Ling Chen, Research Resources Center, UIC, for assistance with the confocal microscopy.

F.F. was supported by a NRSA from NIH (HL68458). This work was supported, in part, by grants from the U.S. Public Health Service to T.T.H. (AI AI47770), D.U. (GM 38800), and P.D.L. (HL 64702 and NIH HL 59618).

 
* Corresponding author. Mailing address: Department of Physiology and Biophysics, University of Illinois at Chicago, 835 South Wolcott Avenue, Chicago, IL 60612. Phone: (312) 996-6430. Fax: (312) 996-1414. E-mail: Primal{at}UIC.edu.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Molecular and Cellular Biology, July 2005, p. 6259-6266, Vol. 25, No. 14
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