Phorbol ester-mediated pulmonary artery endothelial barrier dysfunction through regulation of actin cytoskeletal mechanics

Alan B. Moy,1,2 Ken Blackwell,1 Ning Wang,3 Kari Haxhinasto,1 Mary K. Kasiske,1 James Bodmer,1 Gina Reyes,1 and Anthony English4

1Department of Internal Medicine, 2Biomedical Engineering, University of Iowa College of Medicine, Iowa City, Iowa 52242; 3Physiology Program, Harvard School of Public Health, Boston, Massachusetts 02115; and 4Department of Mechanical, Aerospace, and Biomedical Engineering, University of Tennessee, Knoxville, Tennessee 37996

Submitted 27 August 2003 ; accepted in final form 28 February 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The mechanisms of phorbol ester- and thrombin-mediated pulmonary artery endothelial barrier dysfunction were compared. Phorbol ester dibutyrate (PDBU) mediated slow force velocity and less force than thrombin. Taxol did not attenuate PDBU-mediated tension, while it reversed nocodazole-mediated tension. PDBU-mediated tension was not affected by acrylamide; PDBU increased cell stiffness and produced greater declines in transendothelial resistance (TER) than acrylamide. Thus PDBU caused a net increase in tension and did not unload microtubule or intermediate filaments. Microfilament remodeling, determined on the basis of immunocytochemistry and actin solubility, lacked the sensitivity and specificity to predict actin-dependent mechanical properties. Thrombin increased myosin light chain (MLC) kinase site-specific MLC phosphorylation, according to peptide map analysis, whereas PDBU did not increase PKC-specific MLC phosphorylation. The initial PDBU-mediated tension development temporally correlated with PDBU-mediated decline in TER and increased low-molecular-weight caldesmon (l-CaD) phosphorylation. PDBU-mediated tension development and decreases in TER were associated with a temporal loss of endothelial cell-matrix adhesion, based on a numerical model of TER. Although, on the basis of immunocytochemistry, thrombin-mediated tension was associated with actin insolubility, actin reorganization, and gap formation, these changes did not predict thrombin-mediated gap formation, based on TER and time-lapse differential interference contrast microscopy. These data suggest that PDBU may disrupt endothelial barrier function through loss of cell-matrix adhesion through l-CaD-dependent actin contraction.

phorbol ester dibutyrate; porcine pulmonary artery endothelial cells; myosin light chain; low-molecular-weight caldesmon


IT HAS BEEN WELL DOCUMENTED that phorbol esters directly increase endothelial permeability through loss of cell adhesion in association with remodeling of the actin cytoskeleton (2, 27). Phorbol esters represent unique edemagenic molecules that allow greater insight into how signal transduction pathways remodel endothelial cell adhesion through actin-dependent mechanical forces (2, 17, 27, 30). The precise mechanism by which phorbol esters regulate endothelial cell adhesion through remodeling of actin-dependent mechanical forces remains poorly understood.

Because cell adhesion sites are mechanically coupled to the actin cytoskeleton, there are at least two potential mechanisms by which phorbol esters could decrease endothelial barrier function. Phorbol esters could increase centripetally directed actin tension. Because the actin cytoskeleton is mechanically coupled to adhesion sites, an increase in centripetal tension could, in turn, strain or separate adhesion sites with its extracellular ligand (21). Alternatively, phorbol esters could disrupt cell adhesion through a decrease in local tethering. Loss of tethering could decrease cell adhesion through expression of less-opposed constitutive, centripetal tension (21, 22).

Consistent with this first paradigm, we previously reported increased isometric tension in cultured porcine pulmonary artery endothelial cells (PPAEC) exposed to phorbol esters (2). In this report we document that phorbol ester dibutyrate (PDBU) generated a slow force velocity in PPAEC, which is unlike the rapid force velocity observed in thrombin-treated cells. Thrombin-mediated tension development is dependent on myosin light chain (MLC) phosphorylation at the Ser19 and Thr18 sites, which is regulated, in part, by myosin light chain kinase (MLCK) and inhibition of myosin-associated dephosphorylation (2, 11, 21, 26).

On the basis of a tensegrity paradigm, several mechanisms could account for the observed unique increase in isometric tension in phorbol ester-treated endothelial cells. In tensegrity, cell shape and prestress are dependent on a counterbalance between tensile actin-myosin forces and compressive-resistive struts composed of actin cables, microtubules, and intermediate filaments (21, 36). According to this paradigm, phorbol esters could increase force by activating signal transduction pathways that mediate a net increase in actin-myosin tension.

PKC mediates differential effects on MLC. PKC inhibits MLCK activity under in vitro conditions in smooth muscle (23). In contrast, PKC directly increases MLC phosphorylation at the Ser1/2 site in smooth muscle and in platelet cells and is associated with low force velocity (13, 29). Thus total MLC phosphorylation may underestimate myosin ATPase activity because of these offsetting effects on MLC phosphorylation in the endothelium.

Alternatively, phorbol esters could increase isometric tension through phosphorylation of low-molecular-weight caldesmon (l-CaD). Phosphorylation of l-CaD could release the constitutive inhibition on myosin ATPase (32), which would then mediate a net increase in actin-myosin tension. Stasek et al. (30) reported increased l-CaD phosphorylation in phorbol ester-treated bovine pulmonary artery endothelial cells (BPAEC), whereas phorbol esters did not increase MLC phosphorylation (10). However, these same authors did not report the stoichiometric changes in l-CaD phosphorylation, which would provide greater insight into the potential function of such phosphorylation on myosin ATPase activity. Also, phorbol ester-mediated PKC-specific phosphorylation of MLC and the quantitative physical properties of myosin ATPase activity were not measured in these reports nor in a more recent report that evaluated the effect of phorbol ester on actin and myosin physical properties in BPAEC (3).

Alternatively, phorbol esters could increase isometric tension by unloading intermediate filaments or microtubule compressive-resistive forces, which would mediate less opposition to expressed constitutive actin-myosin forces. However, this measured increase in isometric tension would reflect a shift or redistribution of load-bearing forces rather than a net increase in force generation. Under such conditions, unloading of intermediate filament or microtubule compressive-resistive struts would produce a decrease in cell stiffness as previously reported by Wang and colleagues (36, 38, 39). In contrast, a net increase in actin-myosin contraction would produce an increase in cell stiffness (4, 40).

Stasek et al. (30) reported increased vimentin phosphorylation in response to phorbol ester stimulation, suggesting that phorbol esters may remodel endothelial cell mechanics through effects on intermediate filaments. However, the functional relevance of vimentin-mediated phosphorylation or vimentin-regulated mechanics on endothelial barrier function has not been elucidated.

Elucidating the precise mechanism by which inflammatory stimuli regulate endothelial barrier function through cytoskeletal remodeling is a complex task requiring the integration of mechanical, microscopic, biochemical, and numerical approaches. This report represents the first comprehensive evaluation in which such approaches are used in a complementary fashion to understand the complex mechanisms by which phorbol esters regulate endothelial barrier function. A series of hypotheses was tested in this study. First, do phorbol esters mediate a net increase in actin-myosin tension, and is there evidence to suggest that there is a mechanical coupling between force generation and loss of barrier function? Second, is phorbol ester-mediated force generation dependent on MLC and/or l-CaD phosphorylation? Third, what are the specific cytoskeletal-membrane sites that are altered by phorbol ester stimulation (18, 22)? Lastly, do conventional assays like immunocytochemistry, actin solubility, and time-lapse microscopy have the sensitivity and predictability to evaluate endothelial, actin-dependent mechanical properties and gap formation in response to edemagenic stimuli like thrombin and phorbol esters?


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. PDBU and cytochalasin D were obtained from Sigma (St. Louis, MO). Human {alpha}-thrombin was obtained from Enzyme Research Laboratories (South Bend, IN). Vitrogen collagen (type 1, bovine dermal collagen) was obtained from Celtrix Pharmaceuticals (Santa Clara, CA). Cultured PPAEC were prepared according to previously described techniques (25).

Measurement of isometric tension. Isometric tension of cultured endothelial cells was measured by a modification of a technique we previously described (2). One to two milliliters of a previously defined, cold, unpolymerized collagen mixture (21) were poured between two 10-mm polyethylene bars and allowed to polymerize at 37°C. The collagen matrix was inoculated with 1.0 x 106 cells after the membrane surfaces were coated with 100 µg/ml of fibronectin for 15 min. Data postprocessing was accomplished with a custom-written program in LabView, which subjected the data to a digital low-pass filter, allowing the user to choose data subsets of interest (i.e., sham, baseline, or drug response) for each transducer and calculating the maximum change in tension (mg) and force velocity (mg/min) for each data subset.

Measurement of MLC phosphorylation. Identification and quantification of the phosphorylation of the 20-kDa MLC were accomplished by laser densitometry of two-dimensional gels of MLC isoforms immunoprecipitated from [35S]methionine-labeled cells according to a previously described procedure (20). We calculated stoichiometry by determining the relative fraction of phosphorylated isoforms to all isoforms according to the expression: stoichiometry = fraction of monophosphorylated isoform + 2 x fraction of diphosphorylated isoforms.

Two-dimensional gel separation and quantification of l-CaD phosphorylation. Cultured cells were plated on six-well tissue culture plates and grown to 2-day postconfluence. Cells were stimulated with test agents for specified time periods and lysed with 150 µl of a 3-([3-cholamidopropyl]dimethylammonio)-1-propanesulfonate(CHAPS) solubilization buffer (8 M urea, 4% CHAPS, 100 mM DTT, 40 mM Tris base, 2% pH 3–10 pharmalytes, 2% pH 5–8 pharmalytes, 50 mM NaFl, 100 µg/ml leupeptin, and 1.25 mM PMSF). Samples were snap-frozen in a dry-ice, methanol bath, scraped into microfuge tubes, sheared through a 22-gauge needle, and centrifuged at 100,000 g for 15 min. Isoelectric focusing separated ~100 µl of each sample. To prepare 12 gels (10 ml final volume), the following were mixed: 0.1 ml of 40% pharmalyte stock (Pharmacia) pH 3–10 and 0.4 ml of 40% pharmalyte at pH 5–8 to give a final vol/vol concentration of 0.4% pH 3–10 and 1.6% pH 5–8 pharmalyte. The final composition of each gel also contained 4.5% acrylamide, 9.2 M urea, 1.5% CHAPS, and 0.5% Nonidet P-40. Tube gels were polymerized in 2.4 x 4.0 x 160-mm glass tubes. Tube gels were first subjected to prefocusing at 200 V for 1 h, 400 V for 1 h, and 800 V for 1 h. Samples were loaded and then subjected to 200 V for 2 h, 500 V for 2 h, 800 V for 16 h, and 2,000 V for 6–9 h. Isoelectric focusing was conducted with an upper buffer containing 0.01 M histadine and 0.01 M glutamic acid. l-CaD was further separated in the second dimension by 13% acrylamide SDS-PAGE at 40 mA/gel for 4.5 h at 6°C. The 2.4-mm tube gels were overlaid with an agarose mixture (0.5% agarose, 398 mM Tris, pH 8.8, 0.1% SDS, and 50 mM DTT) on the top of the 1.5-mm slab gel. Samples were then subjected to Western blot procedures in which proteins were transferred to nitrocellulose overnight at 13 V, 4°C, washed briefly in TBS (20 mM Tris base, pH 7.6, 137 mM NaCl), incubated in TBS and 5% dry skim milk for 1 h, and then probed with a mouse anti-CaD antibody (Transduction Labs) at a working dilution of 1:250 for a period of 2.5 h. The membranes were then washed in TBS and 5% milk and then probed with an anti-mouse IgG, horseradish peroxidase-linked secondary antibody (Amersham catalog no. NA931V) at a working dilution of 1:1,200 in TBS and 5% milk for a period of 1 h. The membranes were washed in TBS and 5% milk and then in TBS and then were exposed to X-ray film after being developed with standard ECL (Amersham) procedures and reagents. We calculated stoichiometry by determining the relative fraction of phosphorylated proteins among the regulatory phosphorylated isoforms according to the expression: stoichiometry = fraction of monophosphorylated isoform + 2 x fraction of diphosphorylated isoforms + 3 x fraction triphosphorylated.

Radiolabeling of cultured cells and immunoprecipitation of l-CaD. Confluent cells were inoculated on six-well tissue culture dishes. Cells were radiolabeled with [35S]methionine at 555 µCi/ml in methionine-deficient DMEM (21013-024, GIBCO) supplemented with l-cysteine at 48 mg/ml and 10% FBS for 48 h at 37°C and 5% CO2. The medium was changed to serum-free M-199 (11150-059, GIBCO) supplemented with basal medium Eagle vitamins and amino acids, 2 mM glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10 mg BSA, and then cells were exposed to 10–7 M PDBU for 1 h.

Two-day postconfluent monolayers were alternatively radiolabeled with [32P]orthophosphate in six-well tissue culture plates. Cells were washed in a phosphate-free phosphorylation buffer consisting of 119 mM NaCl, 5 mM KCl, 5.6 mM glucose, 0.4 mM MgCl2, 1 mM CaCl2, and 25 mM PIPES at pH 7.2. Cells were then labeled with 400 µCi/ml of [32P]orthophosphate in the same phosphorylation buffer for 2 h at 37°C with no CO2. The cells were then treated in the same buffer with 10–7 M PDBU for 1 h. At the end of the time periods, the cells were washed in the phosphorylation buffer. Cells were lysed with 200 µl of Nonidet lysing buffer as previously described (20), and the cells were snap-frozen on a dry-ice, methanol bath, thawed, and scraped into 2-ml polycarbonate ultracentrifuge tubes. The lysate was then centrifuged at 100,000 g, 10 min at 4°C. The supernatants were then transferred to 1.5-ml microfuge tubes, and the volume was brought to a total of 400 µl with buffer A (150 mM NaCl, 10 mM Tris·HCl pH 8.0, 0.1% Triton X-100, and 5 mM EDTA). We performed immunoprecipitation overnight at 4°C with rocking by adding anti-l-CaD (C56520 [GenBank] , Transduction Labs) at 1:50 dilution. ImmunoPure (A/G) beads (60 µl; 20422, Pierce), which were equilibrated by being washed in buffer A, were then added to each sample and incubated with rocking for at least 1 h at 4°C. The samples were then centrifuged at 4°C, 14,000 rpm for 1–2 min. The pellets were washed in buffer A, washed in TBS (10 mM Tris·HCl pH 7.6, 150 mM NaCl), and then resuspended in 100 µl of CHAPS solubilization lysis buffer. Samples were heated at 37°C for 15 min and centrifuged for 15 min at 4°C at 14,000 rpm. The supernatants were transferred to new tubes, and the entire sample was either loaded onto the first-dimension tube gels or stored at –80°C until used.

Peptide mapping of tryptic peptide fragments of phosphorylated MLC. One-dimensional peptide maps were constructed from MLC immunoprecipitated from 32P-labeled PPAEC cells as previously described (21). In situ peptide maps were compared with standards of MLC phosphorylated in vitro with MLCK or PKC.

Measurement of actin assembly. Actin assembly was quantified by the measured fraction of insoluble actin in a Triton X-100-based buffer as previously described by Carson et al. (5). Equivalent volume fractions of the soluble and insoluble fractions were subjected to electrophoresis on an 8–15% SDS-polyacrylamide gradient gel, along with known concentrations of purified nonmuscle actin (Cytoskeleton Incorporated), stained with Coomassie blue, and dried, and actin content was quantitated by laser densitometry. The concentration of actin in each cell fraction was quantitated from an in vitro standard calibration curve of the measured radiance volume of known amounts of purified nonmuscle actin separated on the same gel.

Measurement of endothelial barrier function by transendothelial resistance. Barrier function was measured by the technique previously reported (21). We cultured cells on a small gold electrode (5 x 10–4 cm2) using culture medium as the electrolyte and measured barrier function dynamically by determining the electrical impedance of a cell-covered electrode. The in-phase voltage (proportional to the resistance) and the out-of-phase voltage (proportional to the capacitive reactance) were measured. Barrier integrity was expressed as a fractional change in the transendothelial resistance (TER).

Breakdown of cytoskeletal-membrane properties by numerical modeling of experimental transcellular impedance. A numerical analysis was used to calculate specific cell-cell and cell-substrate adhesion and membrane capacitance (Cm) based on the measured transcellular impedance. The procedure is described in more detail elsewhere (18, 22). The total impedance across a cell-covered electrode is composed of the impedance created between the ventral surface of the cell and the electrode (related to {alpha} and due to cell-matrix adhesion), the impedance created between cells (indicated by Rb and due to cell-cell adhesion), the transcellular impedance created from transcellular current conduction (Zm), and the impedance of a naked electrode. For these calculations, Zm is inversely related to Cm, which is dependent on membrane convolution, which, in turn, is dependent on the cortical cytoskeleton. The real and imaginary data of the cell-covered electrode and naked electrode were measured at frequencies between 25–60,000 Hz and at multiple time points so that Cm could be treated as a freestanding variable. A calculated real and imaginary value was generated from the solutions of {alpha}, Rb, and Cm obtained from a multiresponse Levenberg-Marquardt nonlinear optimization model of the real only, imaginary only, real and imaginary (in complex form), and real and imaginary (in magnitude form) data. An error evaluation was calculated by a {chi}2 analysis of the least-squared sum of the calculated and the experimental residuals as a function of current frequency. Solutions for {alpha}, Rb, and Cm were typically obtained by the Levenberg-Marquardt approach from the real data subset, and the analysis was confirmed by also modeling both the real and imaginary to ensure reproducibility.

Measurement of endothelial stiffness by magnetic twisting cytometry. All cells were serum deprived for 48 h before being plated on 100 µg/ml of fibronectin-coated Immulon II dishes (Dynatech) at 30,000 per well in serum-free medium. After the cells were plated overnight, Arg-Gly-Asp-containing peptide (Peptite 2000, Telios)-coated ferromagnetic beads (4 µm diameter) were added to the cells at 20 µg/ml (about one to two beads per cell) for 20 min, and then unbound beads were washed off with serum-free medium. A twisting stress of 40 dyn/cm2 was applied to a well as previously described (36). Briefly, a strong magnetic pulse (1,000 gauss, 10 µs) was applied to magnetize all the beads in one direction, and then a homogeneous twisting field was applied in the vertical direction to rotate the beads. The angular strain was measured with an in-line magnetometer. The applied stress was calibrated in a viscous standard (37). The apparent stiffness was defined as the ratio of stress to strain. After the cells in a well were twisted, 10–7 M PDBU was added to the well, and the same stress was applied again at different time points.

Microscopy. Cultured monolayers were fixed with 3.75% formaldehyde for 10 min and then permeabilized for 10 min with a 0.1% Triton X-100 solution in PBS. Nonspecific binding sites were blocked for 1 h with 1% BSA in PBS (solution A), rinsed with solution A, and then probed for 1 h at room temperature with a mouse IgG antivimentin (1:1,000 dilution) or {alpha}-tubulin (1:500 dilution) (Sigma). The monolayer was then secondarily probed for 1 h with an Oregon green-conjugated, goat anti-mouse antibody (10 µg/ml). Alternatively, permeabilized cell monolayers were rinsed with PBS and probed with 1 unit/monolayer of Texas red phalloidin for 45 min. Images were randomly selected and examined under a Zeiss Axiovert 135 microscope and a x63 Antiflex objective equipped with epifluorescence with fluorescein and rhodamine excitation filters. Images were captured with a 10-bit Hamamatsu ORCA camera and Snapper frame grabber controlled by custom-written software for a Macintosh computer.

For interference reflection microscopy (IRM), we fixed confluent cells in PBS containing 3.7% formaldehyde for 10 min. A Zeiss Axiovert microscope with a x63 Antiflex objective was used to capture IRM images. The images yield varying intensities of light depending on the contact distance between the cell and the glass coverslip.

For time-lapse microscopy, cultured monolayers were inoculated on Delta-T dishes (Bioptechs) and grown to confluence. Environmental temperature was controlled with a Delta-T temperature controller and objective heater (Bioptechs). Time-lapse images were collected at 60-s intervals by differential interference contrast (DIC) microscopy at x630 magnification and subsequently digitized with the ORCA camera and stored on a Macintosh computer. As a positive control to induce cell retraction, cells were continuously perfused with a solution of 0.05% trypsin and 0.53 mM EGTA by a peristaltic pump (Bioptechs) at a flow rate of 1 ml/min.

Statistical analysis. Data are reported as means ± SE. Comparisons between groups were made by a paired or unpaired Student t-test depending on whether the samples were independent. Comparisons between more than two groups were made by analysis of variance. Individual group comparisons were done by Tukey's honestly significant difference test for post hoc comparison of means. Differences were considered significant at the P <= 0.05 level.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Phorbol esters do not mediate force generation through unloading microtubule struts. The change in the centripetal force was measured in cultured PPAEC inoculated on a polymerized collagen membrane by a modification of the technique we previously reported (2). PDBU (10–7 M), at a dose that increases endothelial permeability (27), mediated a slow increase in isometric tension (Fig. 1). Subsequent exposure to 3 µM cytochalasin D abolished tension development, demonstrating that PDBU-mediated tension development was dependent on the actin cytoskeleton.



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Fig. 1. Representative effect of 10–7 M phorbol ester dibutyrate (PDBU) on isometric tension in a confluent monolayer of cultured porcine pulmonary artery endothelial cells (PPAEC) inoculated on the surface of polymerized collagen. PDBU-mediated tension generation is abolished by 3 µM cytochalasin D. See text for explanation. Tracing is representative of >25 experiments.

 
The next-tested hypothesis was whether PDBU increased tension through unloading microtubule struts. Under this paradigm, phorbol esters would induce microtubule disassembly, which, in turn, would oppose less constitutive centripetal actin-myosin tension. If this notion is valid, then taxol should reverse force generation mediated by microtubule disassembly. However, PDBU-mediated tension development was not reversed by the subsequent exposure to 2 µM taxol (Fig. 2A). In contrast, taxol predictably reversed tension development in monolayers exposed to 2 µM nocodazole (Fig. 2B). PDBU-mediated tension generation was unaffected in the presence of taxol. There was no statistical difference in force generation in PDBU-stimulated cells that received taxol (under pretreatment or posttreatment conditions) compared with cells exposed to PDBU alone (Fig. 2C). In contrast, pretreatment of cells with nocodazole attenuated PDBU-mediated tension generation. These data indicate that microtubule disassembly, not microtubule assembly, affects PDBU-mediated tension development.



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Fig. 2. Comparison of the effect of PDBU-mediated tension development in the presence or absence of taxol or nocodazole. A: representative example of cultured PPAEC exposed to 10–7 M PDBU while isometric tension is monitored. Cells were subsequently exposed to 2 µM taxol. B: representative example of 2 µM taxol reversing tension development in cultured cells exposed to 2 µM nocodazole. C: comparison of the mean (± SE) of maximal force amplitude (mg) among sham-controlled nocodazole-designated cells [S (N)], cells exposed to 2 µM nocodazole (N) for 30 min, cells exposed to 2 µM taxol for 30 min (T), cells exposed to 2 µM nocodazole for 30 min followed by exposure to 2 µM taxol (N+T) for 30 min, sham-controlled PDBU-designated cells [S (P)], cells exposed to 0.1 µM PDBU (P) for 60 min, cells exposed to 0.1 µM PDBU for 30 min followed by exposure to 2 µM taxol (P+T) and tension monitored for additional 30 min, cells exposed to 2 µM taxol for 30 min followed by exposure to 0.1 µM PDBU for 60 min (T+P), and cells exposed to 2 µM nocodazole for 30 min followed by exposure to 0.1 µM PDBU for 60 min (N+P). Each data category represents >15 measurements. *Statistical significance compared with sham-treated cells; **nonsignificant (NS) difference compared with cells exposed to PDBU alone; ***significant difference (P < 0.05) compared with cells exposed to PDBU alone. D: immunocytochemistry of tubulin organization in sham-treated cells. Cells were treated with 2 µM nocodazole for 30 min and with 0.1 µM PDBU for 60 min (magnification x630).

 
PDBU did not induce shortening of microtubule filaments, as was documented by immunocytochemistry approaches, although shortening of microtubule filaments, in contrast, was observed in nocodazole-treated cells (Fig. 2D). Together, these data are consistent with a model in which PDBU increased centripetal actin force independently of disrupting microtubule struts.

Nocodazole did not alter actin organization, based on immunocytochemistry (data not shown), and nocodazole did not induce any significant change in the fraction of insoluble actin extracted by Triton X-100 in cultured monolayers. The insoluble actin fraction was 0.75 ± 0.018 in sham-treated cells, whereas the insoluble actin fraction was 0.75 ± 0.02 in nocodazole-treated monolayers. These data demonstrate that nocodazole induced unopposed expression of actin-myosin forces through unloading of microtubule forces, whereas PDBU altered actin mechanics independently of this mechanism.

Phorbol esters do not mediate force generation through unloading intermediate filaments. PDBU also did not increase tension by unloading intermediate filament forces (Fig. 3A). If PDBU increased force through unloading intermediate filament struts, then pretreatment with an agent like acrylamide that perturbs intermediate filaments should decrease the amplitude of force generation induced by PDBU. Exposure of monolayers to 4 mM acrylamide, a dose previously reported to disturb intermediate filament stability (8, 9, 36), increased tension as predicted from a tensegrity paradigm. Subsequent exposure to PDBU further increased tension. Force generation in PDBU-stimulated cells that were pretreated with acrylamide was not statistically greater than that measured in cells exposed to PDBU alone (Fig. 3B). Also, posttreatment of acrylamide did not cause a significant change in PDBU-mediated force generation compared with cells exposed to PDBU alone (Fig. 3B). These data are consistent with a mechanical model in which phorbol esters increased tension independently of unloading of intermediate filament struts.



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Fig. 3. Effect of PDBU on isometric tension in cultured PPAEC in which intermediate filaments struts have been perturbed. A: representative example of cultured PPAEC exposed to 4 mM acrylamide while tension was continuously monitored. Cells were subsequently exposed to 10 –7 M PDBU. B: comparison of the means (± SE) of the maximal change in isometric tension in cultured cells exposed to sham buffer. Cells were exposed to 4 mM acrylamide for 30 min (A), 0.1 µM PDBU for 60 min (P), 0.1 µM PDBU for 60 min after pretreatment with 4 mM acrylamide (A+P) for 30 min, and 0.1 µM PDBU for 30 min followed by exposure to 4 mM acrylamide and tension monitored for additional 30 min (P+A). Each data point represents a sample of >=12 measurements. *Significant difference compared with sham-treated group (P < 0.05). NS, no significant difference in force generation compared with cells treated with PDBU alone.

 
The notion that phorbol esters remodel endothelial mechanics independently of unloading intermediate filament or microtubule struts was further validated through several other mechanical assays. First, cell stiffness was measured in PDBU-treated cells by magnetic twisting cytometry (Fig. 4). If PDBU disrupted intermediate filaments or microtubule struts, then PDBU should decrease cell stiffness. Conversely, if PDBU mediated a net increase in contraction, then PDBU would increase cell stiffness. Cell stiffness was measured in cultured cells that had adhered on substrate for 10 or 16 h. A twisting stress of 40 dyn/cm2 was applied under unstimulated conditions. Cells were subsequently exposed to 10–7 M PDBU, and the same load was applied at different time points. As shown in Fig. 4, cells plated on substrate for 10 h displayed lower basal stiffness than cells plated for 16 h. However, PDBU mediated a slow increase in stiffness under both plating conditions (Fig. 4). PDBU increased stiffness to levels by 30 and 20% by 60 min in cells plated for 16 and 10 h, respectively.



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Fig. 4. Effect of phorbol esters on cell stiffness in cultured PPAEC measured by magnetic twisting cytometry. All cells were plated on 100 µg/ml of fibronectin-coated dishes for either 10 or 16 h (n = 5 and 4, respectively) in serum-free medium. Arg-Gly-Asp-containing, peptide-coated ferromagnetic beads were added to the cells. After a strong magnetic pulse was applied to magnetize all the beads in one direction, a homogeneous twisting field of 40 dyn/cm2 was applied in the vertical direction to rotate the beads. The apparent stiffness was defined as the ratio of stress to strain (dyn/cm2). After the cells in a well were twisted, 10–7 M PDBU was added to the well, and the same stress was applied again at different time points. The mean (± SE) stiffness (dyn/cm2) is reported. The sham responses were only 5% of the PDBU responses (data not shown).

 
Acrylamide mediated a very modest decline in TER (~8%, Fig. 5A). In contrast, PDBU mediated a twofold greater decline in TER (15–20%). Together, these data support the notion that phorbol esters increased tension independently of unloading compressive-resistive, intermediate filament struts.



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Fig. 5. A: comparison of the mean (± SE) fractional change in resistance over time (min) in cells treated with 4 mM acrylamide or 10–7 M PDBU in cultured PPAEC. See text for explanation. Each data point represents a sample size of >=5. B: immunocytochemistry of vimentin (top) and actin (bottom) organization in sham-treated (left) cells and those treated with 4 mM acrylamide for 30 min (middle) or 0.l µM PDBU for 60 min (right). The magnification is x630. See text for explanation.

 
Mechanical unloading between actin-myosin and intermediate filaments as measured by the measured isometric tension, cell stiffness, and TER could not be predicted by changes in actin and intermediate filament organization or mass, based on immunocytochemistry approaches. Few cells displayed juxtanuclear aggregation of intermediate filaments upon exposure to 4 mM acrylamide at 30 min of exposure (Fig. 5B) or even at 4 h of exposure (data not shown). Those few cells that did exhibit juxtanuclear aggregation displayed a less cuboidal cell shape, but this was a subtle feature (8) (Fig. 5B). Juxtanuclear aggregation of intermediate filaments was not observed in cells treated with PDBU for 60 min (Fig. 5B). Remodeling of microtubule organization in cells exposed to 4 mM acrylamide for 30 min was also not observed (data not shown). Together these data demonstrate that an evaluation of actin and intermediate filament cytoskeleton organization based on immunocytochemical approaches lacked the same level of sensitivity as mechanical assays to detect perturbation of load-bearing forces between intermediate filaments and microfilaments in the setting of phorbol ester stimulation.

Differential effects of phorbol ester and thrombin on actin assembly. PDBU-mediated increases in cell stiffness were consistent with actin-myosin contraction. However, we also tested the alternate hypothesis that increased cell stiffness could be due to increased actin assembly, which was evaluated by the measured fractional actin insolubility. The insoluble fraction of detergent-extractable actin did not statistically differ between control cells and cells exposed to PDBU for 60 min (Fig. 6A). In contrast, thrombin mediated a statistically significant increase in insoluble actin at 30 min, whereas 3 µM cytochalasin D mediated a predictable and statistically significant decline in insoluble actin at 30 min. These additional controls confirm that the failure to detect a change in actin assembly in PDBU-treated cells was not due to methodological concerns of the assay. Consistent with this notion, PDBU did not alter microfilament organization according to immunocytochemistry, whereas thrombin increased the number of cortical and stress fiber filaments at 30 min (Fig. 6B). Together these data support the notion that PDBU had a direct and measurable effect on actin-dependent cell mechanics that could not be detected by changes in actin organization or mass, as determined by immunocytochemistry or through changes in actin solubility.



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Fig. 6. A: comparison of the mean (± SE) difference in actin assembly based on the measured fraction of insoluble actin extracted by Triton X-100 in cultured monolayers exposed to PDBU for 60 min, cells exposed to 7 U/ml of thrombin for 30 min, and cells exposed to 3 µM cytochalasin D for 30 min. *Significant difference compared with controlled cells (P < 0.05). B: actin organization based on immunocytochemistry of controlled cells, cells exposed to 0.1 µM PDBU for 60 min, and cells exposed to 7 U/ml of thrombin for 30 min (magnification is x630).

 
Phorbol ester stimulation contracts the actin cytoskeleton through an MLC-independent pathway. Because PDBU mediated a net increase in actin-myosin tension, we next performed a series of experiments to evaluate whether PDBU contracted microfilaments in an MLC-dependent fashion. To address this issue we compared the differences in force amplitude, force velocity, and MLC phosphorylation between cells exposed to PDBU and thrombin, a molecule that induces MLC-dependent force generation (19, 21). Exposure to 7 U/ml of thrombin mediated additional levels of force in cells pretreated with PDBU (Fig. 7).



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Fig. 7. Representative effect of 10–7 M PDBU followed by exposure to 7 U/ml of thrombin on isometric tension in a confluent monolayer of cultured PPAEC inoculated on the surface of polymerized collagen.

 
PDBU mediated less force amplitude and a lower force velocity than that observed in thrombin-treated cells (Table 1). Also, the force velocity was 6–12 times greater in thrombin-stimulated cells than that observed in cells exposed to PDBU pretreated in the absence or presence of acrylamide. Although nocodazole mediated a similar force amplitude as thrombin in cultured cells, the measured force velocity was less than that in thrombin-stimulated cells. These data are consistent with the notion that the signaling mechanisms that govern PDBU-mediated cell mechanics are quite different from those governing thrombin and nocodazole-mediated cell mechanics, in which the former is a model of MLC-dependent force generation.


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Table 1. Effect of treatments on cells' maximal force amplitude and force velocity

 
Consistent with the mechanical data, PDBU did not significantly increase MLC phosphorylation (Fig. 8A). Compared with controlled monolayers, 10–7 M PDBU modestly increased MLC phosphorylation by 0.022 ± 0.03 mol of phosphate per mol of light chain (mP/mMLC) at 10 min, by 0.015 ± 0.015 mP/mMLC at 20 min, and by 0.072 ± 0.01 mP/mMLC at 40 min. In contrast, 7 U/ml of thrombin increased phosphorylation by 1.015 ± 0.030 mP/mMLC at 5 min and by 0.61 ± 0.045 mP/mMLC at 10 min. Only changes in phosphorylation observed in thrombin-treated cells achieved statistical significance.



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Fig. 8. A: stoichiometry of myosin light chain (MLC) phosphorylation [in mol of phosphate per mol of MLC (mP/mMLC)] measured by 2-dimensional gel electrophoresis of MLC immunoprecipitated from [35S]methionine-labeled cells. Comparison of the mean (± SE) stoichiometry among controlled cells (C), cells exposed to 0.1 µM PDBU for 10 (P-10), 20 (P-20), or 40 (P-40) min, and cells exposed to 7 U/ml of thrombin for 5 (T-5) and 10 (T-10) min. *Significant difference from control levels (P < 0.05). Each data point represents an n > 6. See text for explanation. B: composite illustration of peptide maps of tryptic digests from MLC immunoprecipitated in 32P-labeled control cells, cells exposed to 7 U/ml of thrombin for 30 min, cells exposed to 10–7 M PDBU for 40 min, and a standard of gizzard MLC phosphorylated in vitro with myosin light chain kinase (MLCK) or PKC. See text for explanation.

 
We also evaluated a PKC-specific effect on MLC phosphorylation in response to PDBU by comparing the peptide map patterns of tryptic fragments of in vitro standards of PKC and MLCK with peptide maps of MLC in 32P-labeled cells under control, thrombin-, and PDBU-stimulated conditions (Fig. 8B). Tryptic digests were separated by a one-dimensional isoelectric focusing technique. The peptide map for the PKC standard revealed a monophosphorylated Ser1/2 and a monophosphorylated Thr9 peptide fragment, which has been previously described (7, 11, 20). In contrast, the in vitro standard for MLCK revealed a monophosphorylated Ser19 peptide fragment and a Thr18Ser19 diphosphorylated peptide fragment (7, 20).

The peptide map pattern in control and thrombin-stimulated cells was the same as the in vitro standard for MLCK, indicating that MLCK mediated phosphorylation of MLC in cultured PPAEC. Thrombin increased phosphorylation at Ser19 and increased diphosphorylation at the Thr18Ser19 site. The peptide map pattern for cells exposed to PDBU for 30 and 60 min was the same as that of the in vitro standard for MLCK, indicating that phorbol ester treatment did not increase MLC phosphorylation at PKC-specific sites.

Phorbol ester stimulation increases l-CaD phosphorylation. Because phorbol esters contract the actin cytoskeleton independently of MLC phosphorylation, we next evaluated whether there was biochemical evidence to support the hypothesis that phorbol esters contract the actin cytoskeleton through increased l-CaD phosphorylation. l-CaD was immunoprecipitated from lysates of [35S]methionine- or [32P]orthophosphate-labeled cells and separated by two-dimensional protein gel electrophoresis. We identified four 80-kDa nonphosphorylated isoforms in [35S]methionine-labeled cells (Fig. 9A). However, only two of these isoforms were phosphorylated, which was confirmed from immunoprecipitated lysates of 32P-labeled cells. Western blot procedures of l-CaD isolated from unlabeled whole cell lysates and subsequently separated by two-dimensional gel electrophoresis using a commercial antibody against l-CaD identified only the two regulatory isoforms. Stoichiometry of l-CaD phosphorylation was calculated from the two phosphorylated isoforms (Fig. 9B). PDBU increased phosphorylation from 0.91 ± 0.05 mol of phosphate per mol of l-CaD (mP/mCaD) in unstimulated cells to 1.15 ± 0.07 mP/mCaD at 20 min, 1.29 ± 0.04 mP/mCaD at 40 min, and 1.26 ± 0.04 mP/mCaD at 60 min. These stoichiometric levels achieved statistical significance with P values of <0.05 at the 40- and 60-min time points, whereas it achieved borderline statistical significance at 20 min (P = 0.054). These biochemical data support the notion that there is a potential coupling between PDBU-mediated tension development and l-CaD phosphorylation.



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Fig. 9. A: 2-dimensional gel separation of phosphorylated low-molecular-weight caldesmon (l-CaD). l-CaD was immunoprecipitated from lysates of [35S]methionine-labeled cells (35S) and [32P]orthophosphate-labeled cells (32P) and separated by 2-dimensional protein gel electrophoresis. ECL, a Western blot of l-CaD separated by 2-dimensional gel electrophoresis from whole cell lysates using a commercial antibody against l-CaD. The autoradiogram was developed with an ECL reagent. Proteins labeled as A, B, C, and D represent 4 isoforms (only isoforms B and D are phosphorylated). B1 and D1, monophosphorylated proteins of isoforms B and D, respectively. B2 and D2, diphosphorylated proteins of isoforms B and D, respectively. B3 and D3, triphosphorylated proteins of isoforms B and D, respectively. See text for explanation. IEF, isoelectric focusing. B: stoichiometry of l-CaD phosphorylation (mP/mCaD) measured by Western blot of 2-dimensional gel separation of l-CaD from cell lysates. Shown is a comparison of the mean (± SE) stoichiometry between cells exposed to 0.1 µM PDBU for 20, 40, and 60 min. *Significant difference from controlled levels (P < 0.05). Each data point represents a sample size of >=7. See text for explanation.

 
Differential temporal correlation between tension development and TER in PDBU- and thrombin-treated cells. Because thrombin and PDBU increased tension through different mechanisms, we investigated whether there was evidence to suggest a mechanical coupling between tension generation and gap formation for each stimulus. PDBU mediated a dose-dependent decrease in TER with a maximal decrease in resistance of 18 ± 1% at 10–7 M (data not shown). There was a temporal correlation between PDBU-mediated tension development and an initial loss in TER (Fig. 10A). PDBU mediated a slow decline in TER that temporally correlated with slow force generation. In contrast, TER plateaus while the force continues to increase during later time periods upon exposure to PDBU.



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Fig. 10. Comparison of the temporal relationship between isometric tension, expressed as the fractional maximal force, and transendothelial resistance (TER), expressed as the mean fractional change, in cells exposed to 0.1 µM PDBU (A) and cells exposed to 7 U/ml of thrombin (B). Data represent >=6 experiments. See text for explanation.

 
In contrast, force generation was out-of-phase with changes in TER in thrombin-treated cells (Fig. 10B). Thrombin caused an immediate but transient decline in TER. The TER decreased immediately upon exposure to a level of 35 ± 5% within 1 min. Subsequently, the TER increased and returned to its original basal level within 10 min, and the resistance subsequently increased by 23 ± 4% to above initial basal levels. At time points at which force generation was maximally expressed, TER had already recovered to its initial basal state. Also, at time points at which TER exceeded the initial basal resistance, there was still >50% expression of maximal force. Together, thrombin-mediated tension development was not sufficient to account for disruption in barrier function, nor was there a temporal correlation between force generation and barrier recovery. In contrast, PDBU-mediated tension development exhibited a better temporal correlation with loss of barrier function.

Gap formation detected by immunocytochemistry in fixed permeabilized confluent monolayers did not correlate with functional gap formation in living cells. Immunocytochemistry detected gap formation at 30 min of thrombin exposure (Fig. 11A). Yet these apparent gap regions still had intact cell membrane-substrate integrity, which was confirmed by IRM microscopy, which indicated that membrane still completely covered these gap regions (Fig. 11B). In contrast, gap formation was not observed under time-lapse DIC microscopy in response to thrombin in living cells over a 30-min period (Fig. 11, C–E). Cells subsequently exposure to 0.05% trypsin and 0.53 mM EDTA solution resulted in gap formation and cell retraction (Fig. 11, F–H). At time points (30 min) at which immunocytochemistry assays detected thrombin-induced gap formation, TER achieved a level that exceeded initial basal levels by 20%. Together, immunocytochemistry in fixed cell preparations misrepresented the impact of thrombin on functional gap formation in living cells.



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Fig. 11. A: actin organization in thrombin-treated confluent monolayers taken in fixed and permeabilized cells after exposure to thrombin for 30 min. Arrowhead points to apparent gap formation. The controlled sham response is shown in Fig. 6B. B: image of same region captured by interference reflection microscopy; arrowhead points to the same apparent gap formation as shown in A. The figure demonstrates intact membrane integrity (magnification is x630). C: time-lapse image taken under differential interference contrast microscopy at baseline. D: time-lapse image taken at 1 min of exposure to 7 U/ml of thrombin in same cells, which demonstrate no gap formation. E: time-lapse image after 30-min exposure to thrombin. F: same cells were challenged with 0.53 mM EDTA and 0.05% trypsin for 1 min to induce cell retraction. Arrowheads indicate early regions of cell retraction. G and H: subsequent images of same cells as E taken at subsequent 5-s intervals.

 
Localizing the cytoskeletal-membrane regions that are targeted by phorbol esters. To localize the regions at which phorbol ester stimulation disrupts endothelial barrier function, we used a numerical model we previously reported (18, 22) that identifies the contribution of cell-cell adhesion, cell-matrix adhesion, and Cm on transendothelial impedance. Repeated measurements of transendothelial impedance were made at 23 alternating-current frequencies between 25 and 60,000 Hz in PDBU-stimulated cells, so that Cm could be treated as a freestanding variable. Under this arrangement, there was no change in Cm in response to PDBU (Fig. 12). The numerical model revealed that the PDBU-mediated decline in TER was best predicted by a primary decrease in {alpha}. This indicates that PDBU initiated the decrease in barrier function through a primary decrease in cell-matrix adhesion. There was a subsequent further but modest decline in TER that correlated with decreases in Rb at later time points (post-30 min), indicating that PDBU further decreased barrier function during this time period through loss of cell-cell adhesion. PDBU-mediated tension generation temporally correlated with the initial loss in cell-matrix adhesion, suggesting that there may be a mechanical coupling between tension generation and the loss in cell-matrix adhesion. There was also a temporal correlation between tension development and loss in cell-cell adhesion but only at later time points.



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Fig. 12. Mean fractional change over time in impedance between cells (Rb), {alpha}, TER, membrane capacitance (Cm), and isometric tension in cultured PPAEC exposed to 10–7 M PDBU based on a numerical model of transendothelial impedance. Dynamic impedance measurements were obtained at multiple frequencies (25–60,000 Hz) so that Cm could be treated as a freestanding variable. Each data point represents an n of >=10. See text for explanation.

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
Phorbol esters represent unique edemagenic molecules from a mechanical standpoint that provide insight into how signal transduction pathways regulate endothelial barrier function through actin-dependent mechanical forces. We conducted the first comprehensive evaluation of how an edemagenic agent regulates endothelial barrier function through cytoskeleton-based mechanical forces. We demonstrate that phorbol esters increase endothelial isometric tension and stiffness through a net increase in actin-myosin contraction. These mechanical behaviors are associated with increased l-CaD phosphorylation and are mediated in the absence of increased total MLC phosphorylation, PKC-specific MLC phosphorylation, and changes in actin assembly. Phorbol ester-mediated force generation temporally correlated with an initial loss in barrier function, suggesting a potential mechanical coupling between these two measurements, and this initial loss in barrier function is due to disruption in endothelial cell-matrix adhesion based on a numerical model of endothelial barrier function.

PDBU-mediated force generation was abolished by cytochalasin D, indicating that force generation was dependent on intact microfilaments. There are two general mechanisms by which PDBU increases actin-dependent isometric tension. PDBU could activate signal transduction pathways that induce a net increase in tension of actin-myosin cross bridges. Under this paradigm, there would be an increase in the measured isometric tension and cell stiffness. Alternatively, PDBU could activate signal transduction pathways that unload compressive-resistive struts from microtubules or intermediate filaments. Under these latter conditions, decreased counterbalance forces from microtubules and intermediate filaments would oppose constitutive centripetal actin-myosin prestress less, which results in a redistribution of prestress and not a net increase in cell tension. Under this paradigm, activation of signal transduction pathways would result in a measured increased isometric tension but a decrease in cell stiffness. In support of a counterbalance mechanical paradigm, Kolodney and Elson (15) reported an increase in isometric tension in cultured fibroblasts in response to nocodazole, which was inhibited or reversed by taxol. Although the authors reported an increase in MLC phosphorylation, taxol did not reverse or prevent tension development in response to thrombin, which is dependent on MLC phosphorylation (14).

Consistent with a counterbalance paradigm, Wang and colleagues (35, 36) reported a decrease in cell stiffness in endothelial cells exposed to nocodazole or acrylamide. Additionally, Cai et al. (4) reported significantly greater levels of MLC phosphorylation and cell stiffness in cultured fibroblasts expressing a constitutively active MLCK. Wang and Stamenovic (39) reported decreased cell stiffness in cultured wild-type fibroblasts exposed to acrylamide or fibroblasts derived from transgenic mice that were null for expressing vimentin. Together, these data demonstrate that a net increase in actin-myosin cross-bridge formation results in increased centripetal tension and cell stiffness. However, a shift in load-bearing forces from compressive-resistive struts of microtubules or intermediate filaments results in decreased cell stiffness but increased isometric tension.

Our data do not support the hypothesis that phorbol esters increase isometric tension by unloading compressive-resistive microtubule struts. PDBU-mediated tension development was not reversed by the addition of taxol. Pretreatment or posttreatment of taxol did not attenuate PDBU-mediated tension generation. In contrast, pretreatment of cells with nocodazole attenuated PDBU-mediated force generation. These data demonstrate that microtubule disassembly, and not microtubule assembly, modulates the amplitude of tension generation of PDBU-mediated actin-myosin contraction.

In contrast, nocodazole-mediated tension development was reversed by the addition of taxol. PDBU did not mediate microtubule shortening, whereas nocodazole induced microtubule shortening. Nocodazole also did not mediate any nonspecific effects on microfilaments or intermediate filaments. Nocodazole did not disturb actin and vimentin organization, and nocodazole did not alter actin solubility. Together, these data are consistent with a mechanical model in which PDBU does not increase tension through a shift in load-bearing microtubule forces. Uncovering the precise mechanisms by which microtubule drugs modulate actin-myosin tension still needs to be accomplished. Microtubule drugs may modulate centripetal actin-myosin tension by a counterbalance tensegrity model (36); others have purported that agents like nocodazole induce cross talk with actin-myosin by increasing MLC phosphorylation (15, 33). Nonetheless, the data are quite convincing that phorbol ester stimulation increases isometric tension independently of unloading microtubule struts.

The data also do not support the hypothesis that phorbol esters increase tension by unloading compressive-resistive intermediate filaments. PDBU mediated greater increases in isometric tension in cultured PPAEC that were first exposed to acrylamide than in cells exposed to PDBU alone. If phorbol esters activate signal transduction pathways that perturb intermediate filament struts, then force generation due to PDBU should be attenuated by acrylamide. However, perturbation of intermediate filaments by acrylamide stimulation did not statistically alter PDBU-mediated force generation, which is consistent with a model that PDBU increases isometric tension independently of unloading intermediate filament struts.

The difference in the magnitude of the response between acylamide and PDBU on TER is also consistent with the notion that these agents regulate endothelial cell mechanics through different mechanisms. Acrylamide mediated very little loss in barrier function. In contrast, PDBU mediated a twofold greater loss in TER. Together, these data reinforce the notion that perturbations of intermediate filament struts were not sufficient to account for the effects of PDBU stimulation on isometric tension and TER.

The effects of PDBU on cell stiffness measured by magnetic twisting cytometry also validate the notion that phorbol esters mediate a net increase in force generation rather than unloading compressive-resistive intermediate filament or microtubule struts. PDBU slowly increased stiffness in a manner similar to measured changes in isometric tension independent of actin assembly. The increase in stiffness is consistent with a model of actin-myosin cross-bridge formation, not through effects on actin assembly or through shifts in load-bearing forces between microtubules or intermediate filaments and actin-myosin tensile elements.

Immunocytochemistry did not have the resolution to detect shifts in counterbalance forces between intermediate filaments and actin-myosin cross bridges. Eckert (8) reported juxtanuclear aggregation of intermediate filaments in subconfluent Ptk1 cells at 4 h of exposure to low concentrations of acrylamide, although the author did not observe disturbances in microfilaments or microtubules. In contrast, very little juxtanuclear aggregation of intermediate filaments in confluent endothelial monolayers was observed at 30 min or 4 h of exposure to low-dose acrylamide, whereas mechanical approaches had the resolution to evaluate vimentin-dependent mechanics in intact cells. Together, immunocytochemistry represents a poor approach to evaluate shifts in load-bearing forces between actin-myosin tensile elements and intermediate filaments in confluent endothelial cells.

The impact of phorbol esters on the actin cytoskeleton is regulated independently of MLC phosphorylation. Maximal physiological doses of PDBU produced less maximal force generation and force velocity compared with cells exposed to thrombin or nocodazole. The differential response in force generation and force velocity between PDBU, thrombin, and nocodazole supports a paradigm in which thrombin and nocodazole contract through signal transduction pathways that are distinct from PDBU stimulation. Consistent with this notion, PDBU did not significantly increase MLC phosphorylation or increase PKC-specific phosphorylation at the Ser1/2 site on MLC.

These results differ from the responses of phorbol ester-treated BPAEC reported by Garcia and colleagues (3, 10). In a recent report, Bogatcheva et al. (3) reported increased MLC phosphorylation and actin remodeling without increased contraction in response to phorbol ester stimulation. Although it remains to be validated whether this can be attributed to differences in cell-specific responses, there are several important criticisms of this report. First, the authors reported a 40% increase in densitometry of MLC phosphorylation in 32P-labeled cells, which conflicts with the unchanged MLC phosphorylation that was measured by urea gel electrophoresis in the same report. Second, these results conflict with the observed Thr18Ser19 dephosphorylation in response to PMA. Third, these observations also conflict with an earlier report (10) in which it was reported that phorbol esters, both in the presence and absence of A-23187, decreased the stoichiometry at 2 min of exposure, based on urea gel electrophoresis. It was not documented whether phorbol esters had a direct effect on Ser1/2 phosphorylation in BPAEC.

It has been well documented that there is a nonlinear relationship between stoichiometry and myosin ATPase activity (24). At least half of the MLC molecules need to be phosphorylated before measurable levels of myosin ATPase activity can be detected under in vitro conditions, which is consistent with a model in which there is a cooperative interaction between the myosin heads. This notion is empirically consistent in intact endothelial cells, which are stimulated with edemagenic stimuli. Histamine, which does not increase isometric tension in the endothelium (21), does not increase the stoichiometry to measurable levels >0.5 mP/mMLC (20, 28). In contrast, thrombin, which does increase isometric tension, increases the stoichiometry to measurable levels >0.5 mP/mMLC in cultured human endothelial cells (11, 21). In our present report, the measured stoichiometry of MLC phosphorylation determined by two-dimensional gel electrophoresis in thrombin-stimulated PPAEC is also consistent with this principle. The absence of any statistical increase in MLC phosphorylation and the absolute measured stoichiometry in phorbol ester-stimulated cells support the notion that phorbol ester-mediated tension development is governed independently of MLC phosphorylation in cultured PPAEC.

Myosin ATPase activity was qualitatively measured by a silicone wrinkling methodology in subconfluent cells at 10 min of exposure in the Bogatcheva et al. report (3). On the basis of the force velocity of phorbol esters that was measured in our report, 10 min of stimulation with phorbol esters would generate only 1–2 mg of force, which could very conceivably not be detected in subconfluent cells by a silicone wrinkling approach. Thus force generation needs to be dynamically quantified by measuring isometric tension in a confluent monolayer.

Actin remodeling was determined by immunocytochemistry in the Bogatcheva report (3) based on selected single cell images often captured from nonconfluent monolayers. There is potential bias from morphological approaches and statistical intrasubject variance in actin organization within a cultured monolayer, which makes it challenging to interpret intersubject cytoskeletal organization differences from different treatment interventions. These statistical sampling obstacles are resolved by measuring actin solubility from detergent-extracted cell lysates. In our current report, phorbol ester had no effect on actin remodeling on the basis of the measured actin solubility and immunofluorescent microscopy in confluent monolayers.

The notion that phorbol esters mediate force generation through actin-myosin contraction is supported by the measured increase in stoichiometry of l-CaD phosphorylation in response to phorbol ester treatment. Our results are in agreement with those of Stasek et al. (30) in PMA-treated BPAEC. However, these authors did not measure myosin ATPase activity and l-CaD stoichiometry, which is critical to interpret the potential functional impact of l-CaD phosphorylation in intact cells. The stoichiometry increased to levels similar to those that Adam et al. (1) reported with high-molecular-weight caldesmon (h-CaD) phosphorylation in phorbol ester-stimulated smooth muscle cells. Tanaka et al. (31) observed an increase in tropomyosin-stimulated myosin ATPase activity under in vitro conditions when PKC phosphorylates caldesmon at 1 mP/mol h-CaD. Thus the stoichiometric changes in l-CaD phosphorylation measured in phorbol ester-stimulated PPAEC were comparable to those reported under in vitro and in vivo conditions in smooth muscle.

The signaling pathways by which phorbol esters regulate l-CaD phosphorylation in the endothelium have yet to be elucidated. Phorbol esters could activate PKC, which directly phosphorylates l-CaD in intact smooth muscle and other nonmuscle cells at PKC-specific putative sites (12, 16). Phorbol esters also could activate ERK, which phosphorylates l-CaD in intact mammalian smooth muscle and other nonmuscle cells at ERK-specific putative sites (6). However, it is not known how phosphorylation of these sites dysinhibits myosin ATPase activity, and it is not known what impact these phosphorylation sites have on endothelial barrier function. Verin et al. (34) reported ERK activation in PMA-treated BPAEC. However, these authors reported only partial abrogation of PMA-mediated loss in BPAEC barrier function upon exposure to antisense oligonucleotides and pharmacological inhibitors of ERK (34).

The initial loss in phorbol ester-mediated barrier disruption function was temporally correlated with tension development, suggesting a potential mechanical coupling between these two mechanical properties. In contrast, thrombin-mediated tension development was completely out-of-phase with changes in TER. At time points of maximal tension expression, TER was restored to its initial basal state. At time points at which the restoration of TER exceeded initial basal levels, thrombin still expressed considerable force. This shows that tension development is not closely linked to the disruption or restoration of barrier function in thrombin-stimulated PPAEC.

Our report also demonstrates that gap formation measured by immunocytochemistry in fixed permeabilized cultured monolayers does not correlate with functional gap formation in living cells. Gap formation measured by immunocytochemistry did not correlate with TER in thrombin-treated cells. In fact, IRM demonstrated enhanced cell-substrate contact and membrane integrity at apparent gap regions that were documented by immunocytochemistry. In contrast, thrombin did not induce discernible gap formation in living cells captured under time-lapse microscopy on a scale observed by immunocytochemistry in fixed, permeabilized specimens. These results are not unexpected considering that we previously reported cell-cell gap formation on a scale of 70–100 nm in histamine-stimulated cells (22). Thus our findings are consistent with the notion that immunocytochemistry is not sensitive and predictive of membrane-cytoskeletal properties of living cells in the setting of edemagenic stimuli.

Additionally, PDBU induced changes in actin-dependent mechanical properties that could not be detected by immunocytochemistry or by the measured actin insolubility. Together, these data reinforce the notion of the limited sensitivity and specificity of immunocytochemistry in fixed cells to extrapolate actin-dependent cell mechanics in living confluent endothelial cells in response to edemagenic stimuli.

Phorbol esters initiated a disruption in barrier function by activating signal transduction pathways that initially disrupt cell-matrix adhesion sites. Based on a numerical model that characterizes transendothelial impedance (22) into indexes of cell-cell adhesion, cell-matrix adhesion, and Cm, phorbol esters initiate a disruption in barrier function through loss in cell-matrix adhesion. The kinetic changes in cell-matrix adhesion correlate with temporal changes in tension development, suggesting a potential mechanical coupling between force generation and loss of cell-matrix adhesion. Changes in tension development were temporally correlated with modest losses in cell-cell adhesion but only at later time points. Phorbol esters had no apparent effect on Cm, indicating that changes in barrier function were entirely due to changes in adhesive forces. These data suggest two potential mechanisms by which phorbol esters disrupt barrier function. Activation of signal transduction pathways contracts the actin cytoskeleton, which, in turn, strains cell-matrix adhesion sites and possibly later cell-cell adhesion sites. Alternatively, phorbol esters decrease cell-matrix adhesion independently of tension development; the rate constants that regulate resistance and tension are similar, but the reactions are not interdependent. Under these conditions, phorbol esters activate signal transduction pathways that disrupt cell-matrix tethering, which, in turn, induces cell retraction by less-opposed expression of the constitutive centripetal actin-myosin tension. We previously reported that histamine and thrombin transiently disrupt human endothelial barrier function through a sequential loss in cell-cell adhesion and a restoration in cell-matrix adhesion (18, 22). Thus phorbol esters represent an edemagenic stimulus that alters endothelial barrier function through a unique mechanical mechanism that is unlike agents like histamine and thrombin.

In summary, phorbol esters mediate a net increase in microfilament force, which temporally correlates with an initial loss in endothelial barrier function. Force generation is regulated independently of MLC phosphorylation and is associated with increased l-CaD phosphorylation at stoichiometric levels consistent with in vitro dysinhibition of myosin ATPase activity. Phorbol esters initiate a disruption in barrier function by decreasing cell-matrix adhesion based on a numerical analysis of TER. It remains to be determined whether phorbol ester-mediated force generation is mechanically coupled to a loss in endothelial cell-matrix adhesion. Furthermore, the identity of the kinase that mediates l-CaD phosphorylation and the functional impact of these phosphorylation sites on cell mechanics in response to phorbol esters still need to be resolved.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by National Institutes of Health (NIH) Grant GM-61732 (A. B. Moy), American Heart Association Grants 0256019Z (A. B. Moy) and 02650298 (A. English), and NIH Grant HL-33009 (N. Wang).


    ACKNOWLEDGMENTS
 
We thank Sandy Shasby and Dana Ries for support on the MLC experiments. We thank Jeff Van Engelenhoven for support on the tension and impedance experiments. We thank Jianxin Chen for performing the magnetic twisting cytometry studies. We thank D. Michael Shasby for feedback on this manuscript.


    FOOTNOTES
 

Address for reprint requests and other correspondence: A. B. Moy, Dept. of Internal Medicine, C33 GH, Univ. of Iowa College of Medicine, Iowa City, IA 52242 (E-mail: alan-moy{at}uiowa.edu).

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
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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