1Department of Molecular and Cellular Physiology, College of Medicine, University of Cincinnati, Cincinnati, Ohio 45267-0576; and 2Department of Pharmacology, School of Pharmaceutical Sciences, Showa University, Tokyo 142-8555, Japan
Submitted 19 February 2003 ; accepted in final form 28 July 2003
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ABSTRACT |
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collagen matrix; signal transduction; wound repair
In 1995, we initially reported that fibroblast contraction was induced by 30% calf serum (CS) in NIH 3T3 fibroblast cells reconstituted into fibers by growing them in a three-dimensional collagen matrix (21). Use of the fibroblast fiber enabled precise mechanical measurements of force, stiffness, and shortening velocity. CS elicited a maintained force that was readily reversed and reproducible on washout of CS. Additionally, the response was time and dose dependent (19, 21). Cytochalasin D, a cytoskeleton inhibitor, abolished the contraction and confirmed that force was derived from the fibroblast cells (22).
We characterized the intracellular signaling pathways underlying the CS-induced fibroblast contraction. However, CS is a mixture of physiological activating factors and additional factors, such as newly synthesized provisional matrix, that might localize at wound sites. Several contractile factors have also been suggested from experiments conducted with collagen gels, as described above. The nature of these factors and their mechanisms are not known with certainty. At wound repair sites, it is widely accepted that thromboxanes, including thromboxane A2 (TxA2), play central roles in the activation of blood coagulation and vascular smooth muscle contraction (1, 4, 10). TxA2 is one of the major products of arachidonic acid (AA) metabolism in platelets and activated macrophages. It is thought that it plays a key role in vascular homeostasis at the site of inflammation and injury (1, 16, 18). These functions are reasonable and effective in the initial stage of wound repair. It is also reported that TxA2 receptor stimulation activated PKC and/or Rho kinase mediated by inositol trisphosphate-associated intracellular Ca2+ increases (33). However, the critical effect of these kinases on the TxA2-induced fibroblast contraction has not been understood. Because TxA2 occurs at the wound site after the initial steps, we hypothesized that TxA2 might play a role in later stages.
We found that U-46619, a TxA2 analog, could elicit a significant contraction and investigated the intracellular signaling pathways involved. Interestingly, TxA2 does not appear to be one of the activating factors in CS but works in a parallel and additive fashion.
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METHODS |
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Measurement of isometric force development. NIH 3T3 fibroblast fibers were cut into 5-mm pieces and mounted between glass posts with cyanoacrylate glue. One post was fixed, and the opposite side was connected to a silicone strain gauge force transducer (model AME801, SensoNor, Horten, Norway). The fibers were bathed in MOPS-buffered physiological salt solution (MOPS-PSS) containing (mM) 140 NaCl, 4.7 KCl, 1.2 NaH2PO4, 0.02 EDTA, 1.2 MgSO4, 2.5 CaCl2, 11.1 glucose, and 20 MOPS, pH 7.4 at 37°C.
Measurement of intracellular Ca2+ concentration. Intracellular Ca2+ concentration ([Ca2+]i) was measured in NIH 3T3 fibroblast cells in fibers loaded with the Ca2+-sensitive fluorescent dye fura 2 on the basis of the techniques of Grynkiewicz et al. (8). Fura 2-AM was prepared as a stock solution of 1 mM dye in DMSO. The fura 2 loading solution consisted of 3 µM fura 2-AM, 0.015% Pluronic F-127, and 0.5% DMSO in MOPS-PSS; additionally, the solution was sonicated for 5 min to facilitate dispersion of the fura 2-AM. Fibroblast fibers were incubated in this solution at room temperature for 3 h. Subsequently, fibroblast fibers were rinsed in MOPS-PSS for 15 min to remove extracellular and nonhydrolyzed fura 2-AM. A segment (
2 mm) of the fura 2-loaded fibroblast fiber was placed in a glass-bottom culture dish and covered with nylon mesh, which maintained the fiber in an isometric manner. The fiber was placed in a chamber with a total volume of 500 µl and perfused (5 ml/min) with MOPS-PSS, while a temperature of 37°C was maintained. [Ca2+]i was measured with an Intracellular Imaging (INCA, Cincinnati, OH) microscope-based system. The chamber containing the fiber was positioned in a Nikon Diaphot inverted microscope with fluorphase objectives, permitting illumination at 340 nm. Fluorescent images of cells excited at 340 and 380 nm and emitting at 510 nm were obtained with a Dage silicone-intensified target camera. After subtraction of the background fluorescence, ratios of the images obtained at 340 nm to those obtained at 380 nm (R340/380) were acquired on a pixel-by-pixel basis at a frequency of 1 Hz. A previously generated standard curve (see Standard curve for [Ca2+]i calibration) was employed to convert R340/380 to [Ca2+]i. Quantitative analysis of the average [Ca2+]i was achieved by defining the outline of the cell, summing the signal in all the pixels within the defined area, and dividing by the number of pixels.
Standard curve for [Ca2+]i calibration. Solutions containing known concentrations of free Ca2+ for standard curves were obtained from Molecular Probes (Eugene, OR). Fluorescence intensity was measured in 150 µl of each standard solution (0, 0.065, 0.100, 0.225, 0.351, and 0.602 µM free Ca2+) containing 13.3 µg/ml fura 2 pentapotassium salt. The R340/380 values were used by the INCA system software for Ca2+ calibration of experimental data.
Materials. NIH 3T3 fibroblast cells were purchased from American Type Culture Collection (Manassas, VA). DMEM and CS were obtained from Life Technologies (Grand Island, NY), rat tail collagen type I from Upstate Biotechnology (Lake Placid, NY), 1-(2-(5'-carboxyoxazol-2'-yl)-6-aminobenzofuran-5-oxy)-2-(2'-amino-5'-methyl-phenoxy)-ethane-N,N,N',N'-tetraacetic acid (fura 2) and fura 2 penta-AM (fura 2-AM) from Molecular Probes, phosphatidic acid (PA), lyso-PA, platelet-activating factor (PAF), lyso-PAF, sphingosin-1-phosphate (sphingosine), AA, histamine, U-46619, cytochalasin D, cyclopiazonic acid (CPA), calphostin C, ML-7, W-7, Gö-6976, and rottlerin from Sigma Chemical (St. Louis, MO), SQ-29548 from Cayman Chemical (Ann Arbor, MI), and KN-62 from Calbiochem (San Diego, CA). (R)-(+)-trans-N-(4-pyridyl)-4-(1-aminoethyl)-cyclohexanecarboxamide (Y-27632) was kindly provided by Mitsubishi Pharma (Osaka, Japan). Calphostin C, KN-62, and ML-7 were dissolved in DMSO as stock solutions. U-46619 was dissolved in ethanol as a stock solution. Each DMSO and ethanol concentration in the bathing medium was <0.05% and had no effect on mechanical responses. The remaining agents were dissolved in deionized water. CaCl2 was replaced with 10 mM EGTA (pH 7.4) in Ca2+-free MOPS-PSS.
Data analysis. Values are means ± SE. Group data were compared with a one-way analysis of variance. Bonferroni's method was employed to determine the level of significance of differences between groups. P < 0.05 was considered statistically significant.
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RESULTS |
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Several known contractile agonists were tested, and force and [Ca2+]i responses were measured. Treatment for 15 min with the phospholipids 10 µM PA, 10 µM lyso-PA, 3 µM PAF, 3 µM lyso-PAF, and 10 µM sphingosine did not induce significant increases (<20% of the CS-induced response). These concentrations were chosen to be in the near-maximal range based on the literature. Resting levels of [Ca2+]i were enhanced by lyso-PA and PAF; however, other agonists did not affect [Ca2+]i. Signal transduction molecules, 10 µM AA and 3 µM histamine, also did not affect force. However, the stable TxA2 analog U-46619 (100 nM) induced significant increases in force and [Ca2+]i: maximal responses were 76.8 ± 5.5 µN and 156.2 ± 7.3 nM (n = 5), respectively. On the other hand, high KCl (50 mM) increased [Ca2+]i but not force, as described previously (20). A Ca2+ ionophore, ionomycin (10 µM), also elicited results similar to KCl (data not shown).
U-46619-induced contractile responses in NIH 3T3 fibroblast fibers. Within 2 min after it was added, 100 nM U-46619 transiently increased [Ca2+]i (Fig. 2B). The maximal response was 165.3 ± 7.6 nM (n = 7). After the peak, [Ca2+]i declined to the resting level (97.0 ± 4.6 nM, n = 7) at 7 min after U-46619 addition. During the transient increase in [Ca2+]i, isometric force increased (Fig. 2A) but was maintained. The maximal level was detected 12-15 min after U-46619 treatment. The time courses of [Ca2+]i and force development were similar to those observed with 30% CS stimulation (20).
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The increases in force and [Ca2+]i were dependent on U-46619 concentration (Fig. 3A). Significant increases were detected at 3 nM U-46619, and a maximal response was evident at 100 nM U-46619. EC50 values were 10.0 and 11.0 nM, respectively. U-46619-induced force and [Ca2+]i responses were suppressed by pretreatment with 1 µM SQ-29548 (4), a specific TxA2 receptor antagonist (Fig. 3B). In the presence of SQ-29548, 100 nM U-46619-induced responses were 8.4 ± 1.3 µN and 81.6 ± 2.2 nM (n = 5), respectively. To confirm that the U-46619-induced force was derived from fibroblast cell contraction, the cytoskeleton inhibitor cytochalasin D was used. Treatment with 10 µM cytochalasin D for 10 min reduced resting force from 52.0 ± 0.9 to 45.6 ± 0.8 µN(n = 5). Addition of 100 nM U-46619 induced transient increases in [Ca2+]i identical to the normal response; however, the force response disappeared. The value was 46.2 ± 0.7 µN (n = 5) 15 min after introduction of U-46619.
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Intracellular Ca2+ dependence of NIH 3T3 fibroblast fiber contraction. To delimit the sources of Ca2+, we first inhibited Ca2+ influx from the extracellular medium using Ca2+-free MOPS-PSS (Fig. 4). Pretreatment with Ca2+-free MOPS-PSS for 5 min did not influence resting levels of [Ca2+]i. However, the transient increase in [Ca2+]i in response to 100 nM U-46619 was suppressed. Ca2+-free MOPS-PSS slightly reduced the resting force levels, and the sustained increase in force in response to U-46619 was inhibited (13.0 ± 2.0 µN, n = 5); furthermore, the rate of force development was reduced. Similar results were detected in the presence of the Ca2+ channel inhibitor nifedipine (3 µM, 10-min preincubation; data not shown).
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The sarco- and endoplasmic reticulum Ca2+-ATPase (SERCA) inhibitor CPA was used to assess the role of intracellular Ca2+ stores (Fig. 5). CPA (1 µM) induced a small transient increase in [Ca2+]i (112.6 ± 5.8 nM, n = 5); subsequently [Ca2+]i returned to basal levels. During the [Ca2+]i response, isometric force slightly increased. In the presence of CPA, U-46619-induced increases in [Ca2+]i and force development were significantly inhibited: 27.2 ± 4.6% and 23.4 ± 2.9% (n = 5) of the normal response. Similar results were obtained with 10 min of preincubation with 3 µM thapsigargin (data not shown).
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Intracellular signaling pathways for the U-46619-induced contracture in NIH 3T3 fibroblast fibers. Using a pharmacological approach, we investigated the role of Ca2+-dependent signaling pathways in the U-46619-induced increases in isometric force and [Ca2+]i (Fig. 6). The known regulatory pathways in most types of contractile cells (24, 28) involve calmodulin (CaM), Ca2+-dependent CaM kinase (Ca2+/CaM kinase), and/or myosin light chain kinase (MLCK). Pretreatment for 10 min with a CaM inhibitor, W-7 (30), a Ca2+/CaM kinase inhibitor, KN-62 (31), or an MLCK inhibitor, ML-7 (26), had no effects on basal or U-46619-induced force or the [Ca2+]i responses. On the basis of their effectiveness in a wide variety of cells and tissues, >85% of the normal response was maintained after pretreatment with the maximal concentration of these inhibitors.
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TxA2 receptor is known to enhance the activity of PKC. However, the isoform(s) of PKC involved has not been characterized. In this study, we used three types of PKC inhibitors (general, Ca2+-dependent, and Ca2+-independent) to narrow the possible isoforms. As shown in Fig. 7, 1 µM calphostin C (11) did not affect basal levels of force or [Ca2+]i. However, the U-46619-induced increase in force development was significantly inhibited without affecting the transient [Ca2+]i response. The responses to 100 nM U-46619 in the presence of calphostin C were 78.2 ± 2.6 µN and 164.0 ± 6.0 nM (n = 5), which were 46.8 ± 3.1% and 96.3 ± 5.4%, respectively, of the control responses in the absence of calphostin C. As in our previous study, calphostin C had little effect on the CS-induced contraction. Neither Gö-6976 (15), a Ca2+-dependent PKC inhibitor, nor rottlerin (9), a Ca2+-independent PKC inhibitor, had any effects on CS-induced contractures, similar to calphostin C. Importantly, Gö-6976, but not rottlerin, inhibited the U-46619-induced force. These data are summarized in Fig. 8.
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To assess the potential involvement of Rho kinase signaling pathways in the U-46619 contracture, we used the Rho kinase inhibitor Y-27632 (Fig. 9). Treatment with 1 µM Y-27632 for 10 min slightly reduced resting levels from 51.2 ± 1.9 to 43.4 ± 3.9 µN (n = 5). During the treatment, [Ca2+]i was unaltered. Stimulation with 100 nM U-46619 induced a transient increase in [Ca2+]i; however, the sustained increase in force development was suppressed. With Y-27632 pretreatment, the U-46619-induced responses were 170.8 ± 6.0 nM and 40.8 ± 4.5 µN (n = 5), respectively.
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Relation between CS- and U-46619-induced responses in NIH 3T3 fibroblast fibers. In light of our previous results, we investigated the degree to which the responses to CS could be attributable to U-46619-like activation of TxA2 receptors. From these results, it appeared that the intracellular signaling pathway in U-46619 stimulation was different from that in CS stimulation. To confirm that the signaling pathways for U-46619 and CS were different, we measured effects on contractile responses using combinations of both stimulators. We first measured the sustained force elicited by 100 nM U-46619 or 30% CS (Fig. 10A). After the fibers were rinsed, the 30% CS treatment was repeated (143.4 ± 4.0 µN, n = 5). Then 100 nM U-46619 was added, and force was significantly increased to 168.2 ± 3.9 µN (n = 5). The identical strategy was used to measure the [Ca2+]i response (Fig. 10B). Because treatment with U-46619 or CS elicited only transient increases in [Ca2+]i, 100 nM U-46619 and 30% CS were introduced simultaneously. In this case, the peak of the transient increase was significantly elevated (228.8 ± 4.6 nM, n = 5) from each respective U-46619 peak level (168.8 ± 5.2 and 196.8 ± 3.8 nM, respectively, n = 5).
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To confirm that the CS-induced response was not mediated by the TxA2 receptor, we pretreated fibers with the TxA2 receptor inhibitor SQ-29548 (1 µM; Fig. 11). CS-induced increases in [Ca2+]i and force were not influenced by this treatment. After pretreatment with SQ-29548, CS-induced force and [Ca2+]i were 100.8 ± 11.6% and 96.5 ± 7.1%, respectively, of a single treatment of CS (n = 5).
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DISCUSSION |
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Phospholipids and several molecules related to the signal transduction were reported to exert contractile effects measured by a morphological observation using fibroblast cells in collagen gels (7, 13). Furthermore, Ridley and Hall (25) reported that lyso-PA induced stress fiber formation in the fibroblast cells. However, in our NIH 3T3 fibroblast fibers, these substances, with the exception of U-46619, did not increase isometric force. It is possible that the receptor phenotype or cellular enzymes may differ under different culture conditions. Alternatively, the change in cell shape observed in the collagen gels may be caused by small and chronic forces below our level of detectability and may not reflect the isometric force development observed in the fibroblast fibers.
Most reagents had no influence on the basal tension of the fibroblast fiber; however, a TxA2 analog, U-46619, significantly increased force to 65% of the CS-induced response (Fig. 1). To our knowledge, this is the first report of isometric force development in response to a single physiological molecule in fibroblast fibers. It is widely accepted that TxA2 is localized in wound areas at initial stages and functions as an activator of the blood coagulation system and smooth muscle contraction (2, 27). TxA2 released by macrophages and leukocytes during inflammation is also an important factor associated with wound repair. Inasmuch as a low concentration (100 nM) of U-46619 induced maximal force development, we hypothesized that TxA2 may play a specific role in fibroblast contraction in wound repair.
We confirmed that treatment with Ca2+-free MOPS-PSS or the Ca2+ channel inhibitor nifedipine inhibited the CS-induced [Ca2+]i response, but not force development (20). In contrast, U-46619-induced increases in [Ca2+]i and force were inhibited under these conditions (Fig. 4). SERCA inhibition with CPA or thapsigargin also significantly inhibited the U-46619-induced (Fig. 5) and CS-induced responses. Thus Ca2+ mobilization for the CS- and U-46619-induced fibroblast contractions was not identical.
In smooth muscle cells, it is widely accepted that [Ca2+]i regulates contractile responses associated with CaM-, Ca2+/CaM kinase-, and MLCK-dependent pathways (28). Inhibitors of these pathways did not affect the U-46619-induced responses, similar to our previously reported CS-induced responses. However, inhibition by Y-27632 (32) suggested that the Rho kinase pathway was essential for CS- and U-46619-induced contraction.
Both types of stimulation were also characterized by similar time courses. However, the intracellular signaling pathways did not completely overlap. The TxA2 receptor antagonist SQ-29548 (4) blocked the U-46619 contraction but did not affect CS-induced responses. Moreover, addition of 100 nM U-46619 enhanced the maximal force of a 30% CS contraction (Fig. 11). The additivity of U-46619 and CS is of interest, suggesting the existence of multiple, independent pathways. In addition, inhibition of another key enzyme of the Ca2+-signaling pathway, PKC, led to a decrease in the U-46619-induced contraction without affecting the [Ca2+]i response (Fig. 7). These findings suggested that the U-46619-induced contraction may involve PKC downstream from [Ca2+]i signals. Moreover, our studies showed that Gö-6976, but not rottlerin, blunted the U-46619-induced contraction, suggesting that a Ca2+-dependent PKC was involved. In contrast, our previous study indicated that the CS-induced contraction does not involve PKC (20). These data indicate that TxA2-induced contraction does not correspond with CS-induced contraction.
Using Ca2+-free PSS and/or CPA pretreatments, we previously showed that an increase in Ca2+ derived from intracellular Ca2+ stores was essential in the initial phase of the CS-induced contraction, although not for maintenance. This appears similar for the U-46619-induced contraction. Our present results differ, in that Ca2+ influx and Ca2+ release from intracellular Ca2+ stores have important roles in the U-46619-induced contraction. In addition, the initial contraction rate was significantly reduced by Ca2+-free PSS and CPA. From these results, it appears that the initial contraction phase is dependent on Ca2+. On the other hand, PKC inhibition did not change the Ca2+ response but did inhibit contractile responses. Thus the link between tension development and [Ca2+]i is far from clear and remains the subject of ongoing research.
In summary, TxA2-induced contractile response in fibroblast fiber involves [Ca2+]i-signaling pathways that are mediated by PKC and Rho kinase. Ca2+ influx from extracellular medium and Ca2+ release from intracellular Ca2+ stores are required. Inasmuch as TxA2-induced responses do not appear to be involved in the CS-induced contraction, we hypothesize that TxA2 functions not only as a blood coagulation factor at the initial stage of wound repair but also as a contractile factor for fibroblasts in later repair processes.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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