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Regular Article |
a Department of Anesthesiology, College of Physicians & Surgeons, Columbia University, New York, New York 10032
b Westfälische Wilhelms-Universität, Münster 4400, Germany
c The Johns Hopkins Institutions, Baltimore, Maryland 21287-7294
ABSTRACT
The actin cytoskeleton is important for stress fiber formation and contributes to the initiation and maintenance of smooth muscle contraction. To determine if oxytocin and lysophosphatidic acid (LPA) induce stress fiber formation, cultured human myometrial cells were exposed to oxytocin (10-5 M) or LPA (10-6 M), and filamentous (F) and globular (G) actin pools were stained with fluorescein isothiocyanate-phalloidin and Texas red DNase I, respectively. The F- to G-actin fluorescent-staining ratio was measured by fluorescence microscopy. Oxytocin and LPA increased stress fiber formation, as indicated by an increase in the F- to G-actin fluorescent-staining ratio. The Rho-kinase inhibitor Y-27632 markedly attenuated this increase. Oxytocin-induced stress fiber formation was completely inhibited in the presence of the oxytocin antagonist compound VI. Tyrosine kinase inhibition with tyrphostin A23 partially blocked the increase induced by oxytocin but had no effect on LPA-induced stress fiber formation. Stress fiber formation was not blocked by pertussis toxin, mitogen-activated protein kinase, or protein kinase C inhibitors. Our results show that human myometrial cells respond to oxytocin and LPA with the formation of stress fibers that may be involved in the maintenance of uterine contractions. Rho-kinase appears to be a key signaling factor in this pathway.
growth factors, kinases, oxytocin, signal transduction, uterus
INTRODUCTION
The mechanisms leading to uterine quiescence during pregnancy and the initiation of uterine contraction during labor are incompletely understood. The actin cytoskeleton of smooth muscle cells is dynamically regulated by contractile agonists [1] and plays a major role in the formation of stress fibers and focal adhesions, both of which have been shown to contribute to smooth muscle contraction [2]. RhoA, a member of the Ras superfamily of monomeric G proteins, and its downstream effector Rho-kinase are necessary for agonist-induced stress fiber formation [1, 3]. The implicated mechanisms include inhibition of myosin light chain (MLC) phosphatase [4], resulting in an increase of phosphorylated MLCs. So far, two isoforms of Rho-kinase, ROK
and ROKß, have been identified [3], and both isoforms are upregulated in pregnant myometrium [5]. The signaling pathways by which RhoA and Rho-kinase induce stress fiber formation are both agonist- and cell type-specific [6, 7], and little information exists in myometrial cells.
Oxytocin is a well-known contractile agonist of uterine smooth muscle, and its signaling pathways have been extensively studied. In uterine smooth muscle oxytocin couples to the heterotrimeric G-protein Gq to generate inositol 1,4,5-trisphosphate, 1,2-diacylglycerol, and activate protein kinase C [8]. An additional coupling to the pertussis toxin-sensitive G protein Gi to inhibit adenylyl cyclase has been suggested [8]. More recently, oxytocin exposure has been shown to induce shortening of the actin cytoskeleton in cultured myometrial cells [9], but the pathways involved were not identified.
Lysophosphatidic acid (LPA) is a naturally occurring phospholipid that is released by activated platelets during inflammatory processes and is present in serum in micromolar concentrations. It stimulates cell proliferation, platelet aggregation, cell adhesion, and stress fiber formation via coupling to heterotrimeric G proteins in many cell types [10] and is produced by fetal membranes [11]. LPA induces uterine contraction [12], raises intracellular Ca2+ concentrations in myometrial cells [13], and increases MLC phosphorylation in human myometrial tissues [5], suggesting a role in parturition. Both pertussis toxin-sensitive and -insensitive signaling pathways leading to stress fiber formation have been described [14].
The intracellular signaling pathways that couple either LPA or oxytocin to stress fiber formation in myometrial cells are not known. Therefore, using cultured human myometrial cells that express oxytocin and LPA receptors, the present study was performed to determine whether LPA and oxytocin induce stress fiber formation in myometrial cells and if so, to elucidate the signaling pathways involved.
MATERIALS AND METHODS
Cell Culture
After Institutional Review Board approval from the Johns Hopkins Institutions and written informed consent, biopsies of the lower uterine segment were obtained from a pregnant woman undergoing elective cesarean delivery at term. Primary myometrial cultures were established from enzymatically dispersed cells and were maintained in culture over several passages. Cells were grown in high-glucose (4500 mg/L) Dulbecco modified Eagle medium, containing 10% fetal calf serum without the addition of antibiotics or antimycotics, and were tested for mycoplasma. Cells were kept in an atmosphere of 5% CO2 in air at 37°C, and passages 3–8 were used for the experiments. Cells were plated on eight-well microscope slides (Nunc Chambers, Naperville, IL) and grown until almost confluent for each experiment. They were extensively washed and kept in serum-free media for 96 h. Quiescent serum-deprived cells were either exposed to the vehicle alone (controls) or treated with oxytocin 10-5 M or LPA 10-6 M for 5 min in separate wells of the same slide. Agonists and/or inhibitors (see below) were added directly to the well without prior manipulation to avoid nonspecific activation.
Western Blotting
The smooth muscle phenotype of the cells was confirmed by detecting smooth muscle -actin, calponin and h-caldesmon using Western blotting. Previously characterized human airway smooth muscle cells and homogenized smooth muscle from porcine trachea from the authors' laboratory served as positive controls; human fibroblasts served as a negative control. After serum starvation for 4 days, cells were harvested from culture flasks, membranes were disrupted with a cell scraper, and cells were dissolved in an extraction buffer (62.5 mM Tris, pH 6.8; 2% SDS; 10% glycerol; 5% ß-mercaptoethanol). Protein lysates (20 µg/lane) were separated by SDS-PAGE (80 V) in a buffer containing 25 mM Tris, 0.1% SDS, 192 mM glycine, and transferred to polyvinylidene difluoride membranes (PVDF) in transfer buffer (25 mM Tris, 192 mM glycine, 10% methanol) at a constant voltage of 20 V overnight. Blots were incubated in Tris-buffered saline (20 mM Tris, pH 7.5; 500 mM NaCl) with 0.1% Tween-20 (TBS-T) and 5% nonfat milk for 90 min. PVDF membranes were incubated with monoclonal antibodies against smooth muscle -actin (clone 1A4, dilution 1:500), smooth muscle calponin (dilution 1:2000), or smooth muscle h-caldesmon (dilution 1:1000) for 4 h under constant rotation at room temperature. After washing membranes in TBS-T twice for 15 min, membranes were incubated with sheep anti-mouse Ig secondary antibody conjugated with horseradish peroxidase for 1 h. Immunoreactive bands were detected by chemiluminescence according to the manufacturer's protocol (ECL-plus; Amersham Pharmacia Biotech, Piscataway, NJ).
Pretreatment with Inhibitors
To determine the role of Rho-kinase in agonist-induced stress fiber formation, human myometrial cells were pretreated with 10 µM of the specific Rho-kinase inhibitor Y-27632 [15] for 60 min (n = 9 experiments) prior to treatment with oxytocin 10-5 M or LPA 10-6 M. To exclude a nonspecific effect of oxytocin on stress fiber formation, cells were exposed to the selective oxytocin antagonist compound VI [d(CH2)5, Tyr(Me)2, Thr4, Tyr-NH29]-OVT (10-6 M) 10 min before the addition of oxytocin (10-5 M) or LPA (10-6 M) [16]. The involvement of tyrosine kinases was tested by treating cells with the tyrosine kinase inhibitor tyrphostin A23 (150 µM) or the vehicle (DMSO 0.05%) for 30 min before the addition of oxytocin or LPA in 10 experiments [17]. Further experiments included the use of PD98059 (10 µM, 30 min), a specific inhibitor of the activation and phosphorylation of mitogen-activated protein (MAP) kinase kinase 1, in a dose previously shown to inhibit oxytocin-induced ERK-phosphorylation [18]. GF109203X (100 nM, 20 min) was employed to elucidate the involvement of protein kinase C (PKC). GF109203X is a highly selective inhibitor acting competitively at the ATP-binding site of PKC with a 50% inhibitory concentration (IC50) of 10–20 nM for the various PKC isoforms [19]. The involvement of heterotrimeric Gi/o proteins was tested by exposing cells to pertussis toxin (100 ng/ml, 24 h, dissolved in H2O) [20], or no treatment prior to agonist stimulation in five separate experiments.
Fluorescence Staining Protocol
Fluorescence microscopy was performed by methods described previously with some minor modifications [20, 21]. In brief, cells were fixed and agonist action terminated after 5 min by adding equal volumes of freshly made paraformaldehyde 7.4% into the media-containing wells, resulting in a final paraformaldehyde concentration of 3.7%. After a fixation period of 15 min, cells were washed with phosphate-buffered saline (PBS) three times and permeabilized with 0.2% Triton-X 100 in PBS for 5 min. Cells were then blocked with 1% BSA in PBS for 15 min. Filamentous actin pools (F-actin) were stained with fluorescein isothiocyanate (FITC)-labeled phalloidin (1 µg/ml) and globular actin pools (G-actin) with Texas red DNase I (10 µg/ml) in 1% BSA in PBS simultaneously. Staining was performed for 20 min at room temperature protected from light. The wells were washed three times with PBS and a coverslip was mounted with Vectashield to prevent rapid fading. Stimulation, fixation, and staining were always performed in parallel in all wells of the same slide.
Fluorescence Microscopy
Actin pools were visualized with an inverted fluorescence microscope (Olympus IX70, Tokyo, Japan), using a 20x short-working distance objective, and the images obtained were stored with Metamorph software (Universal Imaging Corporation, West Chester, PA) on a personal computer. A 100-W mercury arc lamp served as the excitation light source. Fluorescence intensities of FITC-phalloidin and Texas red DNase I were calculated simultaneously from a view containing >15 cells. The excitation and emission wavelengths were 490 and 525 nm for FITC-phalloidin, and 596 and 615 nm for Texas red DNase I, respectively. Image capturing times, image intensity gain, image enhancement, and black levels were optimized before each experiment and were kept constant throughout the experiments to standardize fluorescent intensities. A control group was included in each slide to account for differences in fluorescence intensities between different experiments. Images were captured in triplicate from each well and were digitized (640 x 480 pixels) with a color resolution from 0 (minimum) to 256 (maximum) intensity.
Background and total fluorescence intensities for FITC-phalloidin and Texas red DNase I were calculated from each image. The filamentous-to-globular actin fluorescent-staining ratios (F/G-actin ratios) were calculated after background fluorescent intensities were subtracted from total fluorescence intensities and the triplicates averaged to obtain a single value from each well.
Materials
The monoclonal antibodies anti-smooth muscle -actin, anti-calponin, and anti-caldesmon, oxytocin, pertussis toxin, and FITC-labeled phalloidin were obtained from Sigma (St. Louis, MO). Anti-mouse IgG was obtained from Amersham Pharmacia Biotech (Piscataway, NJ), and Texas red DNase I was purchased from Molecular Probes (Eugene, OR). Vectashield H-1000 was obtained from Vector Laboratories (Burlingame, CA). Tyrphostin A23, PD98059, and GF109203X were purchased from Calbiochem (La Jolla, CA). Compound VI ([d(CH2)5, Tyr(Me)2,Thr4,Tyr-NH29]-OVT) was purchased from Peninsula Laboratories, Belmont, CA. Y-27632 was a kind gift from the Welfide Corporation (Osaka, Japan).
Statistical Analysis
Statistical analysis was performed using repeated measures of ANOVA, followed by Bonferroni posthoc comparison, if indicated, using Instat software (Graph Pad, San Diego, CA). Data are presented as mean ± SEM, a P value <0.05 was considered significant.
RESULTS
Characterization of Human Myometrial Cells
Nonconfluent human myometrial cells displayed a flattened shape with the ends fanned out for broad attachment. At confluence the cells assumed an elongated shape and were tightly packed and aligned in parallel bundles as is typical for smooth muscle cells (Fig. 1). This morphology was maintained over several passages, but cell growth slowed down after 10 passages and cells senesced after 12–13 passages. Western blots demonstrated expression of -actin, calponin, and h-caldesmon in cultured myometrial cells, airway smooth muscle cells, and homogenized tissue from porcine trachea that were not detected in human fibroblasts, confirming the smooth muscle phenotype of the cells (Fig. 2).
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Stress Fiber Formation in Myometrial Cells
Unlabeled myometrial cells displayed no autofluorescence. In the absence of agonists, cells labeled with FITC-phalloidin showed stress fibers traversing each cell in parallel arrays (Fig. 3a). The intensity of F-actin staining increased markedly after stimulation with oxytocin and stress fibers appeared more pronounced (Fig. 3b). Staining of the G-actin pool with Texas red DNAse I displayed a diffuse distribution throughout the cytoplasm with higher fluorescence intensities perinuclear (Fig. 3c). The intensity of G-actin staining decreased in response to stimulation with oxytocin, suggesting a conversion of G-actin into F-actin (Fig. 3d).
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Effect of Rho-Kinase Inhibition on Stress Fiber Formation
Stimulation of myometrial cells with oxytocin (10-5 M) or LPA (10-6 M) significantly enhanced the F-actin fluorescence intensity as an indicator of myometrial stress fiber formation (Fig. 4, a–c). The importance of Rho-kinase in stress fiber formation was determined by treating half of the wells with the selective Rho-kinase inhibitor Y-27632 (10 µM, 60 min) or the vehicle (water) in nine separate experiments. Although Y-27632 appeared to decrease the F- to G-actin fluorescent-staining ratio compared to controls treated with the vehicle, these data did not reach statistical significance (P > 0.05, n = 9 experiments) (Fig. 4d). However, Y-27632 markedly attenuated stress fiber formation in cells exposed to either oxytocin or LPA (Fig. 4, e and f). In oxytocin- and LPA-stimulated cells, the F- to G-actin fluorescent-staining ratios increased by 40.2 ± 10.62% and 53.3 ± 8.74%, respectively, whereas no increase was observed in the presence of Rho-kinase inhibition (P < 0.001). The results are summarized in Table 1.
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Inhibition of Oxytocin-Induced Stress Fiber Formation by an Oxytocin Antagonist
To demonstrate that oxytocin-induced stress fiber formation was mediated through its receptor, five separate experiments were performed in which half of the wells were pretreated with the selective oxytocin antagonist compound VI (10-6 M) added 10 min prior to agonist stimulation. In the presence of compound VI, oxytocin-induced stress fiber formation was completely abolished (Fig. 4h), whereas LPA-induced increases in the F/G-actin fluorescent-staining ratio remained unchanged (Fig. 4i). Oxytocin-induced stress fiber formation was 65.1 ± 7.31% in the absence of and 9.1 ± 4.90 in the presence of compound VI. LPA-induced increases in the F/G-actin fluorescent-staining ratio were 89.6 ± 9.49% in the absence of and 93.2 ± 6.1% in the presence of compound VI (Table 1).
Effect of Tyrosine Kinase Inhibition on Stress Fiber Formation
To determine whether tyrosine kinases were involved in the signaling pathway leading to agonist-induced stress fiber formation in myometrial cells, cells were pretreated with 150 µM tyrphostin A23 for 30 min prior to oxytocin or LPA exposure (n = 10 experiments). This dose of tyrphostin A23 was previously shown to inhibit neurite outgrowth completely [17]. The results are summarized in Table 1. Tyrosine kinase inhibition had no significant effect on the basal F- to G-actin fluorescent-staining ratio. Tyrphostin A23 inhibited stress fiber formation induced by oxytocin but had no effect on LPA-induced stress fiber formation (Fig. 4, l and m). In the absence of tyrphostin A23 oxytocin increased the F/G-actin fluorescent-staining ratio by 62.4 ± 8.12%, whereas in the presence of tyrphostin A23 only a 31.5 ± 7.59% increase was observed (P < 0.05). LPA-induced increases in the F/G-actin fluorescent-staining ratio were 56.8 ± 6.39% in the absence of and 61.8 ± 10.31% in the presence of tyrphostin A23.
Effect of PKC Inhibition on Stress Fiber Formation
A potential role of PKC in the signaling pathway was determined by exposing cells to the PKC inhibitor GF109203X (100 nM, 20 min). Pretreatment of cells with GF109203X did not significantly inhibit basal, oxytocin- or LPA-induced stress fiber formation in myometrial cells at the dose studied (P > 0.05, n = 6 experiments) (Table 1), suggesting that PKC was not involved in agonist-stimulated stress fiber formation.
Effect of MAP Kinase Inhibition on Stress Fiber Formation
To investigate whether MAP kinase is an intermediate in the signaling pathway leading to stress fiber formation in human myometrial cells, cells were treated with the MAP kinase inhibitor PD98059 (10 µM, 30 min) (n = 6 experiments) prior to exposure to oxytocin or LPA. PD98059 did not significantly inhibit basal, oxytocin- or LPA-induced stress fiber formation (Table 1).
Pretreatment with Pertussis Toxin
The involvement of pertussis toxin-sensitive G-proteins was studied by inhibiting Gi/o proteins with pertussis toxin (100 ng/ml for 24 h) and subsequently stimulating the cells with oxytocin or LPA. In a dose previously shown to inhibit carbachol-induced stress fiber formation in airway smooth muscle cells [20], pertussis toxin (n = 5 experiments) did not significantly inhibit basal or agonist-induced stress fiber formation in human myometrial cells (Table 1). These results indicate that pertussis toxin-sensitive G-proteins are either not involved or do not represent the only pathway leading to stress fiber formation.
DISCUSSION
This study demonstrates for the first time that oxytocin and LPA induce stress fiber formation in human myometrial cells via a signaling pathway requiring Rho-kinase. The cultured myometrial cells used resemble previously described human myometrial cells with respect to functional oxytocin receptors that are maintained over several passages [8]. The smooth muscle phenotype was confirmed by the presence of the smooth muscle-specific proteins -actin, calponin and h-caldesmon. In addition, the cells contained large numbers of actin fibers traversing the whole cell, a pattern characteristic of smooth muscle cells and different from the diffuse pattern described in myometrial fibroblasts [22].
Staining of filamentous actin with quantification of fluorescence intensity is frequently performed to describe changes in stress fiber formation and reorganization of the actin cytoskeleton [20, 23-25]. Dual labeling techniques with additional staining of the total protein content [24] or concurrent labeling of F- and G-actin pools [21, 26, 27] are employed to correct for differences in cell size and cell number per image and reduce their influence on fluorescence intensities. The simultaneous labeling of F actin with FITC-phalloidin and G actin with Texas red DNase I is thought to be separate spatially and specific for the respective actin pool, and there is little interference due to the differences in emission and absorption spectra [21]. Although variations of staining occur between different experiments, the influence is diminished by the inclusion of treated groups and control groups on the same slide that undergo identical culture, staining, and microscopy conditions. Using this technique, we demonstrated that both oxytocin and LPA increased stress fiber formation in myometrial cells (Fig. 4). The results extend the work of Yu et al. [9] who demonstrated a shortening of stress fibers in myometrial cells exposed to oxytocin with immunocytochemical techniques. Although the exact role of stress fiber formation in uterine contraction is not known, Yu et al. [9] suggested that an interaction of the cytoskeleton with focal adhesion proteins is essential for uterine activity.
In the present study, pretreatment with Y-27632 led to a profound inhibition of stress fiber formation after oxytocin and LPA exposure, indicating that the activation of Rho kinase is essential. These data agree with the results of Leung et al. [3], who showed that the expression of constitutively active Rho-kinase induced stress fiber formation in HeLa cells. Furthermore, the Rho-kinase inhibitor reversed oxytocin-induced Ca2+ sensitization in rat uterus [28] and GTP
S-induced Ca2+ sensitization in rabbit mesenteric arteries, and inhibited stress fiber formation in fibroblasts [15]. The mechanism is thought to be the inhibition of MLC phosphatase by Rho-kinase [4] with a resultant increase in phosphorylated MLC. A mutant of Rho-kinase lacking kinase activity dissolves stress fibers [3], further suggesting that stress fiber formation and smooth muscle Ca2+ sensitization share a common pathway.
The involvement of tyrosine kinases upstream and downstream of RhoA has been implicated in stress fiber formation in several cell types [20, 29]. Although tyrphostins inhibit a broad range of tyrosine kinases with little effect on other protein kinases, their potencies against different tyrosine kinases vary [30]. It is likely that differences in tyrosine kinase expression between various tissues exist and that tyrosine kinases expressed in myometrial cells may only be incompletely inhibited by tyrphostin A23. This is supported by the observation that tyrosine kinase inhibitors profoundly inhibit vascular smooth muscle contraction [31], whereas the inhibition of myometrial contraction is incomplete [32]. In addition, Gohla et al. [33] recently showed that stress fiber formation after expression of G12 was independent of tyrosine kinases, suggesting that tyrosine kinase-independent pathways of stress fiber formation also exist.
The lack of effect of a PKC inhibitor to prevent stress fiber formation confirms previous studies showing that agonist-induced stress fiber formation occurs independent of PKC, even if higher concentrations of GF109203X are used [34]. Activation of PKC and its downstream effectors results in phosphorylation of MLC phosphatase at a different site than Rho-kinase-induced phosphorylation [35], suggesting that parallel signaling pathways for PKC and RhoA exist [36]. Different signaling pathways have also been suggested for MAP kinase, as inhibition of MAP kinase fails to inhibit stress fiber formation [20] and Ca2+ sensitization in smooth muscle [37].
Although Gi proteins mediate some myometrial responses to oxytocin [8] as well as to LPA as evidenced by the ability of pertussis toxin to prevent LPA-induced myometrial cell proliferation [13], the lack of effect of pertussis toxin is not surprising. LPA induces stress fiber formation by coupling to the heterotrimeric G proteins Gi and Gq [7] as well as to G12 and G13 [6], indicating that several G proteins may be involved.
In summary, findings of the present study demonstrate that Rho-kinase plays a central role in the signaling pathways linking activation by oxytocin or LPA to the increased formation of stress fibers. In light of recently demonstrated changes in RhoA/Rho-kinase expression during pregnancy [5, 38], changes in the activity of this pathway with resultant increases in stress fiber formation are likely to enhance the efficiency of uterine contraction at the time of delivery. Upregulation of Rho-kinase may also contribute to the enhancement of myometrial oxytocin sensitivity that occurs at term gestation.
FOOTNOTES
First decision: 27 November 2000.
1 The National Institutes of Health NHLBI RO1 HL 62340 in part supported this study. W.G. is a postdoctoral fellow supported by the Innovative Medizinische Forschung, Germany. 
2 Correspondence: Carol A. Hirshman, Department of Anesthesiology, College
of Physicians & Surgeons of Columbia University, P & S Box 46, 630
West 168th Street, New York, NY 10032. FAX: 212 305 8980; cah63{at}columbia.edu
Accepted: February 16, 2001.
Received: November 2, 2000.
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