Am J Physiol Cell Physiol Watch the video to see how APS reaches out to developing nations.
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Cell Physiol 293: C1049-C1058, 2007. First published July 5, 2007; doi:10.1152/ajpcell.00027.2007
0363-6143/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/3/C1049    most recent
00027.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Chan, M. W. C.
Right arrow Articles by McCulloch, C. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Chan, M. W. C.
Right arrow Articles by McCulloch, C. A.

EXTRACELLULAR MATRIX, CELL INTERACTIONS

Cyclosporin inhibition of collagen remodeling is mediated by gelsolin

Matthew W. C. Chan, Pamela D. Arora, and Christopher A. McCulloch

Canadian Institutes of Health Research Group in Matrix Dynamics, Faculty of Dentistry, University of Toronto, Toronto, Ontario, Canada

Submitted 19 January 2007 ; accepted in final form 2 July 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cyclosporin A (CsA) inhibits collagen remodeling by interfering with the collagen-binding step of phagocytosis. In rapidly remodeling connective tissues such as human periodontium this interference manifests as marked tissue overgrowth and loss of function. Previous data have shown that CsA inhibits integrin-induced release of Ca2+ from internal stores, which is required for the binding step of collagen phagocytosis. Because gelsolin is a Ca2+-dependent actin-severing protein that mediates collagen phagocytosis, we determined whether gelsolin is a CsA target. Compared with vehicle controls, CsA treatment of wild-type mice increased collagen accumulation by 60% in periodontal tissues; equivalent increases were seen in vehicle-treated gelsolin-null mice. Collagen degradation by phagocytosis in cultured gelsolin wild-type fibroblasts was blocked by CsA, comparable to levels of vehicle-treated gelsolin-null fibroblasts. In wild-type cells treated with CsA, collagen binding was similar to that of gelsolin-null fibroblasts transfected with a gelsolin-severing mutant and treated with vehicle. CsA blocked collagen-induced Ca2+ fluxes subjacent to bound collagen beads, gelsolin recruitment, and actin assembly at bead sites. CsA reduced gelsolin-dependent severing of actin in wild-type cells to levels similar to those in gelsolin-null fibroblasts. We conclude that CsA-induced accumulation of collagen in the extracellular matrix involves disruption of the actin-severing properties of gelsolin, thereby inhibiting the binding step of collagen phagocytosis.

adhesion molecules; fibroblasts; knockout mice; actin

CYCLOSPORIN A (CsA), an immunosuppressant that is used for the prevention of organ transplant rejection, causes numerous side effects including nephrotoxicity, chronic infections, and gingival overgrowth (27, 47, 54, 55). The immunosuppressive properties of CsA derive from its inhibition of the calcineurin pathway by binding to cyclophilin in the cytoplasm (38, 51), which subsequently leads to the formation of a complex that inhibits IL-2 activation (14, 48). In contrast, the impact of CsA on gingival overgrowth is thought to be caused by perturbation of intracellular Ca2+ signaling (9) and, consequently, interference with collagen remodeling (19). While it is not clear whether CsA upregulates collagen synthesis (20, 26, 43, 50), collagen degradation in CsA-induced gingival overgrowth is clearly inhibited (32), thereby leading to a net accumulation of collagen in tissues with rapid matrix turnover such as gingiva.

The relative abundance of collagen in tissues is determined by the functional balance between collagen synthesis and degradation (45). The maintenance of this balance depends on collagen phagocytosis by fibroblasts, a critical degradative process that is required for normal remodeling and the structural integrity of collagen-rich extracellular matrixes (22). Collagen phagocytosis is a multistep, integrin-dependent process involving the initial recognition and binding of collagen fibrils by the {alpha}2beta1-integrin (6, 36) and the subsequent engulfment of fibrils by plasma membrane extensions (8, 40). For efficient engulfment, actin filament length and branching are actively modified to facilitate extension of the plasma membrane around collagen fibers. Based on their ability to sever actin filaments and promote actin assembly at free barbed ends, Ca2+-dependent actin-binding proteins are likely involved in enhancing extension of lamellipodia (12) that are essential for collagen phagocytosis (40).

We previously (7) identified gelsolin as a critical actin-binding protein in the {alpha}2beta1-integrin-mediated collagen phagocytosis pathway. Gelsolin has been extensively characterized in studies of cell motility and phagocytosis (34, 56, 59) and is a multifunctional, six-subunit, Ca2+-activated actin-severing protein that caps the barbed ends of actin filaments (35). On release by phosphoinositides, actin assembly occurs at free barbed ends (35). Our previous work (5) showed that the Ca2+-dependent actin-severing activity of gelsolin facilitates the collagen-binding step of collagen phagocytosis.

Previous studies have shown that CsA blocks Ca2+ release from endoplasmic reticulum (ER) stores (9) by inhibiting conductance of the mitochondrial inner membrane pore (4) that is involved in ER Ca2+ regulation. Because increased free intracellular Ca2+ is an important determinant of the severing activity of gelsolin (30), we assessed the impact of CsA on gelsolin function in collagen phagocytosis. The data indicate that disruption of gelsolin's severing activity in fibroblasts may contribute to the matrix overgrowth following CsA treatment.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents. Latex (2-µm diameter) beads were purchased from Polysciences (Warrington, PA). Antibodies to beta-actin (clone AC-15) and to bovine type I collagen (clone COL-1), as well as fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse antibody, tetramethylrhodamine B isothiocyanate-phalloidin, and CsA were from Sigma-Aldrich (St. Louis, MO). FITC-goat anti-rabbit and anti-mouse {alpha}2beta1-integrin antibodies were purchased from Cedarlane (Hornby, ON, Canada). The affinity-purified polyclonal antibody to recombinant gelsolin was described previously (10).

Animal and histological studies. Wild-type and background-matched gelsolin-null (Gsn–) mice (58) (6–8 wk of age) were given daily intraperitoneal injections of 40 mg/kg of CsA in saline or saline alone (41, 53). Six male mice were used in each experimental group and were killed after 4 wk of treatment. Mandibles and maxillae were prepared for histology and stained with picrosirius red for collagen. The gingival connective tissue lamina propria of the molar regions were analyzed by computer-assisted morphometry as described previously (41). All animal studies were conducted according to animal care protocols reviewed and approved by the Animal Care Committee of the University of Toronto.

Cells. Fibroblasts were obtained from either wild-type or Gsn– 12-day mouse fetuses as described previously (58). Genotyping by polymerase chain reaction of tail snips was done to confirm deletion of gelsolin. The fibroblastic identity of the cells was verified by staining for vimentin and collagen as well as the absence of desmin. Cells were grown at 37°C and were used at passage 4.

Flow cytometry. Unfixed cells were stained for surface {alpha}2beta1-integrins and assessed by flow cytometry as described previously (36). Nonspecific staining was estimated from samples incubated with an irrelevant isotype control, and these fluorescence channel numbers were subtracted from samples.

Collagen bead binding. Collagen-coated latex beads (2 µm) were applied to microbiological (i.e., non-tissue culture plastic) dishes and attached as described previously (6). The number of beads plated per dish was adjusted to produce specific bead-to-cell ratios. Detached cells were removed by repeated washes. Attached cells spread and rapidly internalized the collagen beads (8). To evaluate collagen bead internalization, FITC-collagen-coated beads were incubated with cells for timed incubation periods. Internalization was stopped by cooling on ice. Fluorescence from extracellular beads was quenched by Trypan blue, whereas internalized beads retained their bead-associated fluorescence (8). In some experiments, because the severing activity of gelsolin enhances collagen bead binding (7), we expressed bead internalization data as the percentage of beads internalized, thereby normalizing for the reduced numbers of beads bound to Gsn– cells.

Actin monomer incorporation. In permeabilized cells incubated with rhodamine actin monomers we measured increases of rhodamine fluorescence due to incorporation into nascent actin filaments (10, 16, 28, 29). Cells were permeabilized with octyl glucoside. Freshly sedimented rhodamine actin monomer was added to the samples, followed by fixation with formaldehyde. Rhodamine fluorescence in single cells was quantified by microscope fluorimetry. For estimation of background correction, detergent treatments were omitted, fluorescence was quantified, and this background signal was subtracted from experimental samples.

Immunofluorescence and confocal microscopy. Cells plated on beads were allowed to spread and bind to collagen beads for 15 min. Cells were fixed, permeabilized, stained with polyclonal gelsolin antibody and FITC-tagged second antibody, and imaged by confocal microscopy.

Collagen bead-associated proteins. Collagen- or BSA-coated magnetic beads (Spherotech) were attached to non-tissue culture plastic dishes. Cell suspensions were allowed to attach to beads for 20 min. Detached, floating cells were aspirated and replaced with medium warmed to 37°C to synchronize phagocytosis. Cells and collagen-coated magnetic beads were collected by scraping into ice-cold extraction buffer containing protease inhibitors. Beads were pelleted with a side-pull magnet. Isolated beads were resuspended, sonicated, and centrifuged to remove unbroken cells. After clarification, equal amounts of proteins were incubated with antibodies to gelsolin or beta-actin to form immunocomplexes that were captured on Sepharose-G beads. The samples were boiled, separated on SDS-PAGE gels, and immunoblotted.

Collagen degradation. Glass coverslips were coated with FITC-collagen, followed by incubation with cell suspensions. Areas of depletion of the FITC-collagen signal from the glass coverslip were measured by morphometry and used as an indication of collagen uptake by cells. To restrict measurements of collagen to a narrow zone above the glass slide, images were captured with total internal reflection fluorescence (TIRF) microscopy, a method that generates fluorescence excitation within a narrow zone (100–150 nm) above the coverslip. Only fluorescent collagen at or below the ventral cell surface was imaged.

Calcium. Cells on glass coverslips were incubated with fura-2 AM as described previously (9) and then with collagen beads. TIRF imaging was performed in regions of interest near the collagen beads or in non-bead-associated parts of the cell so that submembrane intracellular Ca2+ concentration ([Ca2+]i) could be estimated in the vicinity of these beads as we described previously (57). Cells were analyzed with alternating excitation wavelengths of 340 and 380 nm and an emission wavelength of 520 nm. Fluorescence intensities from bead and nonbead regions of interest were plotted as fold increase from baseline level.

Transfections. Gsn– cells were transfected with a DsRed2-tagged wild-type gelsolin expression vector or gelsolin-severing double mutant in the G1 and G2 domains (RLK-210-AAA/AAA-100-DDD) as described previously (5). Cells were transfected with FuGENE6 (Roche, Indianapolis, IN). For study of phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2] regulation of gelsolin in CsA treatment, Gsn– cells were transfected with a DsRed2-tagged wild-type gelsolin and pretreated with the peptides PBP-10 and RhB-QRL (17) for 30 min before bead incubations.

Severing assays. Gelsolin severing of actin filaments was measured as described previously (3). Briefly, rhodamine-phalloidin was added to actin filaments, and the rate of fluorescence loss at 570 nm was measured fluorimetrically. Reduction of fluorescence is caused by the ability of gelsolin to sever actin and displace phalloidin after the addition of CaCl2. Dialyzed cell lysates from wild-type and Gsn– cells treated with either saline or CsA (10 µM) were prepared with detergent plus protease inhibitors. Severing assays were performed with equivalent amounts of protein from cell lysates. Controls included lysates prepared from ionomycin (2 µM) in calcium buffer and ionomycin in 1 mM EGTA-treated cells.

Gelsolin:actin complex fractionation. To examine whether CsA influences the relative abundance of gelsolin:actin complexes, we quantified gelsolin that was associated with actin in pellets prepared by ultracentrifugation. Cells treated with saline or CsA (10 µM) were collected and suspended in a nondetergent buffer (in mM: 1 EGTA, 2 Tris, pH 7.4, 0.2 ATP, 0.2 MgCl2, 0.2 dithiothreitol, 2 phenylmethylsulfonyl fluoride) containing 1 µM phalloidin to stabilize actin filaments. Freshly prepared cell lysates were pipetted several times with a narrow-gauge pipette tip and ultracentrifuged (100,000 g for 1 h at 4°C). The pellet contained gelsolin complexed with actin filaments, and this pellet was resuspended in buffer (in mM: 1 MgCl2, 0.1 CaCl2, 0.2 ATP, 1 NaHCO3, and 1 NaN3, with 0.1 M KCl). Protein concentrations in the fractions were equalibrated, separated by SDS-PAGE, and immunoblotted for detection of actin and gelsolin.

Statistical analyses. For continuous variables, means and SE were computed. Differences between groups were evaluated by Student's unpaired t-test or analysis of variance for multiple comparisons. Statistical significance was set at P < 0.05. Post hoc comparisons were performed with Tukey's test. For all experiments, at least three independent experiments were evaluated, each performed in triplicate.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
CsA-induced collagen accumulation. Compared with vehicle control-treated wild-type mice, the areas of gingival connective tissues of vehicle control-treated Gsn– mice and CsA-treated wild-type and Gsn– mice were apparently enlarged (Fig. 1A). Morphometric assessment of picrosirius red-stained tissue showed ~60% greater amounts of collagen in these same groups of mice compared with vehicle-treated wild-type mice (P < 0.05; Fig. 1B), but there were no differences of collagen-stained tissue areas between vehicle control-treated Gsn– mice and CsA-treated wild-type and Gsn– mice.


Figure 1
View larger version (33K):
[in this window]
[in a new window]

  		Fig. 1. Histological studies of cyclosporin A (CsA)-induced accumulation of collagen and analysis of collagen bead binding and internalization by fibroblasts. A: wild-type (WT) or gelsolin-null (Gsn–) mice were treated daily with intraperitoneal injections of saline (control) or CsA for 4 wk. Gingiva surrounding molar regions stained with hematoxylin and eosin (H&E) is shown. Note 60% increased area of gingival connective tissue (indicated by arrows) in WT (+/+) mice treated with CsA or Gsn– (–/–) mice treated with vehicle or CsA compared with vehicle-treated WT mice. B: quantification of collagen area in gingival connective tissues stained with picrosirius red. WT mice treated with CsA and vehicle and CsA-treated Gsn– mice exhibit similar collagen content. CsA-treated WT mice, Gsn– mice, and vehicle-treated Gsn– mice showed ~60% more collagen area than controls (P < 0.02). Data (mean ± SE) were computed from 6 mice in each experimental group. C: Fluorescein isothiocyanate (FITC)-collagen-coated beads were loaded onto a monolayer culture of fibroblasts treated with vehicle (control) or 10 µM CsA at a bead-to-cell ratio of 8:1 at 37°C for indicated time points. Bead binding (i) and internalization (ii) were determined by microscopy. The number of beads bound per cell was significantly lower in the CsA-treated group (P < 0.01 for all time points). There were no differences in % of internalized beads between control and CsA-treated groups (P > 0.05). iii and iv: BSA beads incubated with cells by the approaches described in i and ii.

 
We characterized the effect of CsA on collagen phagocytosis by determining whether CsA treatment affects surface expression levels of collagen receptors. Cells were treated with CsA (10 µM) or vehicle for 30 min, immunostained for {alpha}2- and beta1-integrins, and analyzed by flow cytometry. There was no statistically significant difference in the surface expression of {alpha}2beta1-integrins between control and CsA-treated cells (data are mean background-corrected fluorescence channel numbers from 3 separate experiments: control = 1.61 ± 0.04, CsA = 1.50 ± 0.05; P > 0.05).

CsA inhibits collagen phagocytosis of cultured fibroblasts in a dose-dependent manner (9), but it is not known whether CsA affects the binding or the internalization steps of collagen phagocytosis. A time-course study was performed with FITC-collagen-coated beads (bead-to-cell ratio = 8:1). Collagen internalization was confirmed by Trypan blue quenching of the fluorescence of external beads. Surface-bound beads exhibited fluorescence quenching with Trypan blue. The numbers of collagen beads bound per cell were significantly greater in the control group compared with the CsA-treated group at 15–90 min (Fig. 1Ci; P < 0.05). To examine the effect of CsA on collagen internalization, the number of internalized beads was calculated as a percentage of bound beads, thereby adjusting for the lower numbers of beads that bound to CsA-treated cells. For this analysis there was no difference between the control and CsA treatments at any of the time points (P > 0.2; Fig. 1Cii), indicating that CsA did not affect the collagen internalization step of phagocytosis once beads were bound. Collagen-coated beads that were incubated with CsA overnight and washed with PBS before a 90-min time-course study of incubation with cells showed no reduction in binding compared with vehicle-treated collagen-coated beads (data not shown), indicating that CsA did not inhibit collagen binding because of a direct effect of CsA on collagen itself. We also examined the effect of CsA on the binding and internalization of beads that were coated with BSA (Fig. 1C, iii and iv). In these experiments, there were no differences in binding or internalization between CsA and control vehicle for any of the time points and, as expected, binding and internalization of BSA-coated beads were only a small fraction of those for collagen-coated beads.

Actin monomer addition. Because actin assembly is required for collagen phagocytosis (21), we examined the effect of CsA on de novo actin incorporation into filaments at the collagen bead-cell interface (30 min after incubation of collagen beads with cells). We examined cells with beads on the cell surface that had not become internalized as demonstrated by Trypan blue quenching (Fig. 1Civ). Although controls and CsA-treated cells exhibited similar cytoplasmic staining intensity for actin filaments (Fig. 2A, c and f), CsA-treated cells showed greatly reduced fluorescence intensity of actin filaments and reduced de novo actin incorporation around collagen beads compared with vehicle-treated controls (Fig. 2A, a and d). Quantification of rhodamine fluorescence around collagen beads showed no increase between 2 and 30 min in the CsA-treated samples (P > 0.2) compared with vehicle-treated controls, which exhibited greater than twofold increases between 2 and 15 min after incubation (P < 0.02; Fig. 2B). Furthermore, rhodamine actin fluorescence was more than twofold lower in the CsA-treated cells than in the controls. BSA-coated beads showed no accumulation of rhodamine actin fluorescence in either CsA- or vehicle-treated cells (data not shown). As a control to determine whether rhodamine actin monomers were preferentially added at the barbed ends of actin filaments, we preincubated cells with cytochalasin D (1 µM, 15 min) before incubation with collagen beads for 30 min. Cytochalasin D reduced the fluorescence intensity in control cells incubated with collagen beads from 18 ± 2.5 to 8 ± 1.5 fluorescence units (P < 0.02).


Figure 2
View larger version (35K):
[in this window]
[in a new window]

  		Fig. 2. Effect of CsA on actin monomer assembly at collagen bead-cell interface. A: cells treated with CsA were incubated with collagen-coated beads. Optical sections of the bead-cell interface were acquired by confocal microscopy. a and d: Actin monomer incorporation in permeabilized cells was observed with rhodamine (Rd)-labeled monomers (red) after 15 min of bead incubation with cells. b and e: Differential interference contrast (DIC). c and f: Staining of actin filaments with Alexa {lambda}488-phalloidin (green). Amount of de novo actin assembly was minimal in CsA-treated samples compared with control treatment. B: rhodamine fluorescence around beads was quantified from 25 cells for each time point in defined 4-µm2 zones. For background correction, detergent treatments were omitted and fluorescence was quantified. Background signals were subtracted from the experimental groups. CsA-treated cells show lower de novo actin monomer addition at collagen bead sites from 2 to 30 min (P < 0.01). Histogram shows mean ± SE rhodamine actin fluorescence.

 
Gelsolin recruitment. By confocal microscopy to examine fluorescence of bead-associated proteins, there was abundant enrichment of gelsolin subjacent to collagen beads 15 min after incubation in vehicle-treated cells; this recruitment was absent in the CsA-treated group (Fig. 3Ai). Because inhibition of gelsolin recruitment, and, as a result, actin severing, could affect actin filament length and actin assembly near collagen bead binding sites, we incubated equal numbers of collagen-coated magnetite beads with control or CsA-treated cells. Subsequently, the beads were removed from the cell surface and bead-associated proteins were immunoblotted. There were marked reductions in the amount of gelsolin and beta-actin that associated with collagen beads after CsA treatment, but the bead-associated amounts of {alpha}2- and beta1-integrin were not detectably different (Fig. 3Aii).


Figure 3
View larger version (17K):
[in this window]
[in a new window]

  		Fig. 3. Gelsolin recruitment and collagen degradation. A, i: confocal microscopy at 15 min after bead incubation with cells shows that CsA-treated cells abolished enrichment of gelsolin around collagen beads. DIC images of same cell immunostained for gelsolin show location of beads. ii: Immunoblot analysis of proteins eluted from collagen-coated magnetite beads. Equal numbers of beads were loaded onto cells, and total protein from all isolated magnetic beads was used for blotting in each lane. Compared with vehicle controls, minimal amounts of gelsolin and beta-actin were detected in CsA-treated samples. There were no obvious differences in the amounts of {alpha}2-and beta1-integrin in the bead-associated proteins. B: time course of collagen degradation. Cells were plated on FITC-collagen-coated glass coverslips and imaged at the cell-substrate interface by total internal reflection fluorescence (TIRF) microscopy. WT fibroblasts treated with vehicle showed loss of FITC-collagen fluorescence over time, while cells treated with CsA showed minimal collagen degradation, similar to Gsn– cells treated with vehicle or CsA at 30–90 min after incubation. Histogram shows mean ± SE FITC-collagen fluorescence.

 
Collagen degradation. Collagen degradation at the cell-collagen interface was evaluated by TIRF microscopy of FITC-collagen-coated coverslips. After 30–90 min of incubation, only wild-type cells treated with vehicle mediated statistically significant reduction of FITC-collagen from glass coverslips (P < 0.02 for 60 and 90 min compared with 0 min; Fig. 3B). In contrast, there were no statistically significant reductions of collagen fluorescence in the three other groups over time (P > 0.5).

Intracellular Ca2+. [Ca2+]i was measured by TIRF in regions of interest at peripheral contact sites with beads because of the ability of TIRF to analyze a narrow zone of fluorescence 100–150 nm above the glass coverslips (57). Peak [Ca2+]i as measured in single fura-2-loaded cells occurred ~3 min after collagen bead incubation in vehicle controls (Fig. 4A), while no obvious increases of signal were evident during the first 5 min in cells treated with CsA (Fig. 4B). Quantification of TIRF images from five different cells showed a 52% increase of fluorescence intensity of control-treated samples compared with a 10% increase for CsA-treated samples at 3 min after incubation. In cells incubated with BSA beads that visually showed attachment to these beads, we found no change in [Ca2+]i in vehicle-treated cells (Fig. 4C) or in cells treated with CsA (data not shown).


Figure 4
View larger version (35K):
[in this window]
[in a new window]

  		Fig. 4. Collagen-coated beads induce intracellular Ca2+ flux. Cells were loaded with fura-2 AM and analyzed after 5-min incubations with collagen-coated beads (CCB). Fluorescence images were taken by TIRF microscopy at 1-min intervals. Phase-contrast images show location of beads at cell periphery (shown by white lines). Region of interest (ROI; red lines) near collagen-coated beads shows area of Ca2+ concentration measurement after CsA treatment (B) compared with vehicle (A). Fluorescence intensity in ROI showed insignificant change after CsA treatment compared with 62% increase of signal from baseline (0 min to a peak at 3 min). Region of cell with no collagen-coated beads was measured and showed no significant change of fluorescence from 0 to 5 min. C: ROI analysis of a vehicle-treated cell incubated with BSA beads.

 
Gelsolin-severing activities. Cell lysates were prepared from wild-type or Gsn– fibroblasts previously treated with vehicle or CsA and then incubated with collagen beads to stimulate (or not, as is the case with CsA) calcium entry (9). Gsn– cells with or without CsA treatment showed ~40% of the actin filament-severing activity of control-treated wild-type cells. CsA almost completely abolished the severing activity of wild-type cells (Fig. 5A). Because CsA reduced the severing activity of gelsolin, we determined whether CsA treatment would mediate an increase in the relative abundance of the gelsolin:actin complexes. Immunoblots showed that in cells treated with CsA there was relatively more gelsolin:actin complex compared with controls (Fig. 5A).


Figure 5
View larger version (27K):
[in this window]
[in a new window]

  		Fig. 5. Effect of CsA on gelsolin-mediated actin filament severing and capping activities. A: actin filament-severing activities analyzed in cell lysates from WT and Gsn– cells treated with CsA or vehicle and then with collagen beads. CsA inhibits severing activity of WT cells. Note that in Gsn– cells, CsA did not affect severing activity, which is ~40% of that in WT cells. Right: ultracentrifugation was used to prepare cell pellets, which were immunoblotted for gelsolin and beta-actin. In CsA-treated WT cells, there was more gelsolin in the pellets, presumably complexed to actin filaments. B: Gsn– cells transfected with WT gelsolin construct (i) or actin-severing mutant AAA-100-DDD/RLK-210-AAA construct (ii) were treated with vehicle or CsA and incubated with FITC-collagen-coated beads. At each time point, extracellular fluorescence was quenched with 0.2% Trypan blue for 5 min to distinguish internalized beads from surface-bound beads. In contrast to WT gelsolin constructs, CsA had no effect on bead binding in the experimental group transfected with the actin-severing mutant construct. C: WT cells incubated with rhodamine B-labeled PBP-10 or control peptide QRL for 10 min after vehicle (i) or CsA (ii) treatment. PBP-10 inhibited collagen internalization, but CsA had no further effect. Bead binding was quantified by fluorescence microscopy. Data (means ± SE) were computed from 3 independent experiments. Beads were only counted in positively transfected cells.

 
To characterize the impact of CsA on gelsolin's severing and capping activities separately we generated double mutations of putative F-actin side binding residues in gelsolin domains G1 (AAA-100-DDD) and G2 (RLK-210-AAA) that cause reduced actin-severing activity as described previously (5). The wild-type or severing mutant constructs were transfected into Gsn– fibroblasts, treated with control or 10 µM CsA, incubated with FITC-collagen-coated beads, and followed in a time-course phagocytosis experiment. Cells transfected with the gelsolin wild-type construct exhibited reductions in collagen binding but not collagen internalization after CsA treatment (Fig. 5Bi; P < 0.05 at t = 90 min). When cells transfected with severing mutant constructs were compared, CsA had no impact on collagen binding or internalization (Fig. 5Bii).

After collagen-receptor association, PI(4,5)P2 binds to gelsolin and promotes uncapping of gelsolin from actin filament and subsequent collagen internalization (5). To study the collagen internalization step of collagen phagocytosis, we utilized a cell-permeant peptide (PBP-10: rhodamine B-QRLFQVKGRR) to disrupt the gelsolin-PI(4,5)P2 interactions by mimicking the PI(4,5)P2 binding site of gelsolin. Wild-type cells pretreated for 10 min with 30 µM PBP-10 or the control peptide, 10 µM QRL, showed no difference in collagen binding, and, as anticipated, the percentage of internalized beads with PBP-10 treatment was significantly less (Fig. 5Ci; P < 0.01 at t = 90 min). When we included 10 µM CsA in the PBP-10 or the control peptide regimen, there was no further reduction in collagen binding or internalization (Fig. 5Cii) compared with the QRL or the PBP-10 peptide.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
CsA is a widely used immunosuppressant that downregulates IL-2 expression and T-cell activation (14) through calcineurin (51). Currently the mechanisms by which CsA blocks extracellular matrix remodeling through the collagen phagocytic pathway are not defined. Collagen phagocytosis is a critical mechanism for extracellular matrix homeostasis (22) and is facilitated by the actin-severing protein gelsolin (5, 7). Impaired collagen phagocytosis by CsA is thought to contribute to collagen accumulation in gingival connective tissues leading to gingival overgrowth and loss of function (39). With a short-term in vivo model of CsA-induced overgrowth (32) and a well-characterized collagen bead phagocytosis model (5–8), we examined one mechanism by which CsA may interfere with collagen remodeling. Our principal finding is that CsA inhibits the initial binding step of collagen phagocytosis by preventing the recruitment and actin filament-severing activity of gelsolin, in part through disruption of calcium signaling at the collagen-receptor interface.

Collagen phagocytosis. Previous studies of CsA-induced gingival overgrowth showed disturbances in collagen turnover (18, 25). Although there is a potential role for increased collagen production in gingival overgrowth (26), we focused on the intracellular collagen degradation pathway by fibroblasts since this pathway is impaired in vivo (31, 32, 44). Collagen phagocytosis involves extension of fibroblast lamellipodia that engulf collagen fibrils (40), a process that requires extensive remodeling of the actin cytoskeleton. Actin remodeling and assembly at the leading edge require actin-severing proteins such as gelsolin, cofilin, and twinfilin (23, 42). The reduced but still measurable severing activity of Gsn– cells is almost certainly attributable to some of these other severing proteins, and in this context there is abundant cofilin in fibroblasts that can organize actin filaments (60). We studied the role of gelsolin in mediating the effect of CsA on collagen phagocytosis because disruption of the intracellular calcium signaling system by CsA could potentially affect the calcium-dependent severing activity of gelsolin. The area of collagen accumulation in the gingival connective tissue of CsA-treated wild-type mice was similar to that of vehicle-treated Gsn– animals, indicating that CsA treatment is phenocopied by the absence of gelsolin in altering collagen turnover. Accordingly, we considered that gelsolin may be an important target in CsA-induced disruption of collagen homeostasis.

Collagen phagocytosis is a multistep process that begins with collagen binding and is followed by internalization and intracellular degradation of collagen in phagolysosomes (22). The effectiveness of the collagen-binding step is dependent in part on cell surface expression levels of {alpha}2beta1-integrins (36). Although there are wide, cell type-dependent variations of surface expression levels of {alpha}2- and beta1-integrins after CsA treatment (13, 31, 49), we found that short-term CsA treatment did not affect surface expression levels of {alpha}2- or beta1-integrins but specifically inhibited the collagen-binding step (9).

Cytoskeletal remodeling. Actin assembly is required for phagocytosis in fibroblasts (21, 33) and in professional phagocytes such as macrophages (2). Regulation of local actin polymerization at the collagen binding site is critical for plasma membrane extension during phagocytosis (1, 37). Previous work has shown that gelsolin is important in IgG-dependent phagocytosis by neutrophils (52) and in collagen phagocytosis by fibroblasts (7). Our present data show that CsA affects actin filament assembly by inhibiting de novo actin monomer addition at the collagen binding site. Blocked actin assembly at collagen binding sites can impair the mobility and clustering of integrin surface receptors (7), a process that is required for full receptor activity (15).

Actin monomer addition at barbed ends is an important process for lamellipodial extension (46). Creation of new barbed ends is mediated by actin-severing proteins such as cofilin or gelsolin (24). Gelsolin is recruited to collagen binding sites, which is followed by actin filament severing and later by uncapping of barbed ends (7, 8). Gelsolin recruitment and localized activation at bead sites are therefore important for efficient actin assembly and for the activity of actin nucleators such as the Arp2/3 complex in platelets and fibroblasts (23). We found that gelsolin recruitment to collagen binding sites and subsequent collagen internalization were impaired by CsA. Since gelsolin's severing activity is regulated by calcium and because CsA treatment can disrupt intracellular calcium signaling (9), we determined whether alterations in calcium signaling can account in part for the lack of gelsolin recruitment and activity. We found reduced calcium flux around collagen-coated beads after CsA treatment, consistent with a lack of gelsolin activation and also with the ability of CsA to block gelsolin's severing activity. Our previous data (9) showed that collagen-induced increases of whole cell free [Ca2+]i are also inhibited by CsA. The basis for the ability of CsA to block the in vitro severing activity of gelsolin as measured here (3) is likely the ability of CsA to block collagen-induced calcium entry (9), which was modeled here by pretreatment of cells with collagen beads before production of cell lysates. Notably, the data from Fig. 5A indicate that Gsn– cells retain 40% of the severing activity of wild-type cells, probably reflecting the presence of other actin-severing proteins such as cofilin. In contrast to Gsn– cells, the almost complete loss of actin-severing activity in CsA-treated wild-type cells suggests that CsA may affect other actin-related functions that are not dependent on gelsolin. Indeed, the actin-dependent extensions of invadopodia are strongly inhibited by CsA, and this inhibition is thought to be mediated by dynamin through the calcineurin pathway (11).

We previously (5) dissected the dual functions of gelsolin in collagen phagocytosis, using mutagenesis techniques to examine the role of isolated gelsolin domains and their impact on actin severing and uncapping. Gelsolin is a six-domain actin-binding protein. The actin-severing properties are attributable in part to domains 1 and 2. A PI(4,5)P2-regulated binding site in domain 2 mediates uncapping of gelsolin from the actin barbed end, thereby enabling filament growth. Disruption of the structure of actin-severing domains in gelsolin reduces collagen binding but not capping, whereas interfering with PI(4,5)P2 regulation inhibits collagen internalization (5). We found that CsA did not further inhibit gelsolin-mediated collagen internalization if cells were previously incubated with PIP2 binding peptides that mimic the phosphoinositide binding region of gelsolin (17). However, CsA did substantially reduce actin severing by gelsolin. Collectively, these data implicate gelsolin and its actin-severing activity as important CsA targets of the binding step of collagen phagocytosis.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by Canadian Institutes of Health Research (CIHR) Group Major Equipment and Operating grants to C. A. McCulloch and a CIHR Network for Oral Research Training and Health (NORTH) Studentship.


   ACKNOWLEDGMENTS
 
We thank Paul Janmey (University of Pennsylvania) for discussions and suggestions in regard to the use of the gelsolin peptides.


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. A. McCulloch, Rm. 244, Fitzgerald Bldg., 150 College St., Toronto, ON, Canada M5S 3E2 (e-mail: christopher.mcculloch{at}utoronto.ca)

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
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Aderem A. Phagocytosis and the inflammatory response. J Infect Dis 187, Suppl 2: S340–S345, 2003.[CrossRef][Web of Science][Medline]

2. Allen LA, Aderem A. Molecular definition of distinct cytoskeletal structures involved in complement- and Fc receptor-mediated phagocytosis in macrophages. J Exp Med 184: 627–637, 1996.[Abstract/Free Full Text]

3. Allen PG, Janmey PA. Gelsolin displaces phalloidin from actin filaments. A new fluorescence method shows that both Ca2+ and Mg2+ affect the rate at which gelsolin severs F-actin. J Biol Chem 269: 32916–32923, 1994.[Abstract/Free Full Text]

4. Altschuld RA, Hohl CM, Castillo LC, Garleb AA, Starling RC, Brierley GP. Cyclosporin inhibits mitochondrial calcium efflux in isolated adult rat ventricular cardiomyocytes. Am J Physiol Heart Circ Physiol 262: H1699–H1704, 1992.[Abstract/Free Full Text]

5. Arora PD, Chan MW, Anderson RA, Janmey PA, McCulloch CA. Separate functions of gelsolin mediate sequential steps of collagen phagocytosis. Mol Biol Cell 16: 5175–5190, 2005.[Abstract/Free Full Text]

6. Arora PD, Fan L, Sodek J, Kapus A, McCulloch CA. Differential binding to dorsal and ventral cell surfaces of fibroblasts: effect on collagen phagocytosis. Exp Cell Res 286: 366–380, 2003.[CrossRef][Web of Science][Medline]

7. Arora PD, Glogauer M, Kapus A, Kwiatkowski DJ, McCulloch CA. Gelsolin mediates collagen phagocytosis through a rac-dependent step. Mol Biol Cell 15: 588–599, 2004.[Abstract/Free Full Text]

8. Arora PD, Manolson MF, Downey GP, Sodek J, McCulloch CA. A novel model system for characterization of phagosomal maturation, acidification, and intracellular collagen degradation in fibroblasts. J Biol Chem 275: 35432–35441, 2000.[Abstract/Free Full Text]

9. Arora PD, Silvestri L, Ganss B, Sodek J, McCulloch CA. Mechanism of cyclosporin-induced inhibition of intracellular collagen degradation. J Biol Chem 276: 14100–14109, 2001.[Abstract/Free Full Text]

10. Azuma T, Witke W, Stossel TP, Hartwig JH, Kwiatkowski DJ. Gelsolin is a downstream effector of rac for fibroblast motility. EMBO J 17: 1362–1370, 1998.[CrossRef][Web of Science][Medline]

11. Baldassarre M, Pompeo A, Beznoussenko G, Castaldi C, Cortellino S, McNiven MA, Luini A, Buccione R. Dynamin participates in focal extracellular matrix degradation by invasive cells. Mol Biol Cell 14: 1074–1084, 2003.[Abstract/Free Full Text]

12. Bearer EL, Prakash JM, Manchester RD, Allen PG. VASP protects actin filaments from gelsolin: an in vitro study with implications for platelet actin reorganizations. Cell Motil Cytoskeleton 47: 351–364, 2000.[CrossRef][Web of Science][Medline]

13. Bolcato-Bellemin AL, Elkaim R, Tenenbaum H. Expression of RNAs encoding for {alpha} and beta integrin subunits in periodontitis and in cyclosporin A gingival overgrowth. J Clin Periodontol 30: 937–943, 2003.[CrossRef][Web of Science][Medline]

14. Bunjes D, Hardt C, Rollinghoff M, Wagner H. Cyclosporin A mediates immunosuppression of primary cytotoxic T cell responses by impairing the release of interleukin 1 and interleukin 2. Eur J Immunol 11: 657–661, 1981.[Web of Science][Medline]

15. Carman CV, Springer TA. Integrin avidity regulation: are changes in affinity and conformation underemphasized? Curr Opin Cell Biol 15: 547–556, 2003.[CrossRef][Web of Science][Medline]

16. Chan AY, Raft S, Bailly M, Wyckoff JB, Segall JE, Condeelis JS. EGF stimulates an increase in actin nucleation and filament number at the leading edge of the lamellipod in mammary adenocarcinoma cells. J Cell Sci 111: 199–211, 1998.[Abstract]

17. Cunningham CC, Vegners R, Bucki R, Funaki M, Korde N, Hartwig JH, Stossel TP, Janmey PA. Cell permeant polyphosphoinositide-binding peptides that block cell motility and actin assembly. J Biol Chem 276: 43390–43399, 2001.[Abstract/Free Full Text]

18. Dannewitz B, Edrich C, Tomakidi P, Kohl A, Gabbert O, Eickholz P, Steinberg T. Elevated gene expression of MMP-1, MMP-10, and TIMP-1 reveal changes of molecules involved in turn-over of extracellular matrix in cyclosporine-induced gingival overgrowth. Cell Tissue Res 325: 513–522, 2006.[CrossRef][Web of Science][Medline]

19. Deliliers GL, Santoro F, Polli N, Bruno E, Fumagalli L, Risciotti E. Light and electron microscopic study of cyclosporin A-induced gingival hyperplasia. J Periodontol 57: 771–775, 1986.[Web of Science][Medline]

20. Eleftheriades EG, Ferguson AG, Samarel AM. Cyclosporine A has no direct effect on collagen metabolism by cardiac fibroblasts in vitro. Circulation 87: 1368–1377, 1993.[Abstract/Free Full Text]

21. Everts V, Beertsen W, Tigchelaar-Gutter W. The digestion of phagocytosed collagen is inhibited by the proteinase inhibitors leupeptin and E-64. Coll Relat Res 5: 315–336, 1985.[Web of Science][Medline]

22. Everts V, van der Zee E, Creemers L, Beertsen W. Phagocytosis and intracellular digestion of collagen, its role in turnover and remodelling. Histochem J 28: 229–245, 1996.[CrossRef][Web of Science][Medline]

23. Falet H, Hoffmeister KM, Neujahr R, Italiano JE Jr, Stossel TP, Southwick FS, Hartwig JH. Importance of free actin filament barbed ends for Arp2/3 complex function in platelets and fibroblasts. Proc Natl Acad Sci USA 99: 16782–16787, 2002.[Abstract/Free Full Text]

24. Frank DJ, Hopmann R, Lenartowska M, Miller KG. Capping protein and the Arp2/3 complex regulate non-bundle actin filament assembly to indirectly control actin bundle positioning during Drosophila melanogaster bristle development. Mol Biol Cell 22: 222–223, 2006.

25. Gagliano N, Moscheni C, Dellavia C, Torri C, Stabellini G, Ferrario VF, Gioia M. Effect of cyclosporin A on human gingival fibroblast collagen turnover in relation to the development of gingival overgrowth: an in vitro study. Biomed Pharmacother 58: 231–238, 2004.[CrossRef][Medline]

26. Ghiggeri GM, Altieri P, Oleggini R, Valenti F, Ginevri F, Perfumo F, Gusmano R. Cyclosporine enhances the synthesis of selected extracellular matrix proteins by renal cells "in culture." Different cell responses and phenotype characterization. Transplantation 57: 1382–1388, 1994.[Web of Science][Medline]

27. Gremese E, Ferraccioli GF. Benefit/risk of cyclosporine in rheumatoid arthritis. Clin Exp Rheumatol 22: S101–S107, 2004.[Web of Science][Medline]

28. Hartwig JH. Mechanisms of actin rearrangements mediating platelet activation. J Cell Biol 118: 1421–1442, 1992.[Abstract/Free Full Text]

29. Hartwig JH, Bokoch GM, Carpenter CL, Janmey PA, Taylor LA, Toker A, Stossel TP. Thrombin receptor ligation and activated Rac uncap actin filament barbed ends through phosphoinositide synthesis in permeabilized human platelets. Cell 82: 643–653, 1995.[CrossRef][Web of Science][Medline]

30. Janmey PA, Chaponnier C, Lind SE, Zaner KS, Stossel TP, Yin HL. Interactions of gelsolin and gelsolin-actin complexes with actin. Effects of calcium on actin nucleation, filament severing, and end blocking. Biochemistry 24: 3714–3723, 1985.[CrossRef][Medline]

31. Kataoka M, Seto H, Wada C, Kido J, Nagata T. Decreased expression of {alpha}2 integrin in fibroblasts isolated from cyclosporin A-induced gingival overgrowth in rats. J Periodontal Res 38: 533–537, 2003.[CrossRef][Web of Science][Medline]

32. Kataoka M, Shimizu Y, Kunikiyo K, Asahara Y, Yamashita K, Ninomiya M, Morisaki I, Ohsaki Y, Kido JI, Nagata T. Cyclosporin A decreases the degradation of type I collagen in rat gingival overgrowth. J Cell Physiol 182: 351–358, 2000.[CrossRef][Web of Science][Medline]

33. Knowles GC, McKeown M, Sodek J, McCulloch CA. Mechanism of collagen phagocytosis by human gingival fibroblasts: importance of collagen structure in cell recognition and internalization. J Cell Sci 98: 551–558, 1991.[Abstract/Free Full Text]

34. Kwiatkowski DJ. Functions of gelsolin: motility, signaling, apoptosis, cancer. Curr Opin Cell Biol 11: 103–108, 1999.[CrossRef][Web of Science][Medline]

35. Kwiatkowski DJ, Janmey PA, Yin HL. Identification of critical functional and regulatory domains in gelsolin. J Cell Biol 108: 1717–1726, 1989.[Abstract/Free Full Text]

36. Lee W, Sodek J, McCulloch CA. Role of integrins in regulation of collagen phagocytosis by human fibroblasts. J Cell Physiol 168: 695–704, 1996.[CrossRef][Web of Science][Medline]

37. Lefkir Y, Malbouyres M, Gotthardt D, Ozinsky A, Cornillon S, Bruckert F, Aderem AA, Soldati T, Cosson P, Letourneur F. Involvement of the AP-1 adaptor complex in early steps of phagocytosis and macropinocytosis. Mol Biol Cell 15: 861–869, 2004.[Abstract/Free Full Text]

38. Liu J, Farmer JD Jr, Lane WS, Friedman J, Weissman I, Schreiber SL. Calcineurin is a common target of cyclophilin-cyclosporin A and FKBP-FK506 complexes. Cell 66: 807–815, 1991.[CrossRef][Web of Science][Medline]

39. McCulloch CA. Drug-induced fibrosis: interference with the intracellular collagen degradation pathway. Curr Opin Drug Discov Devel 7: 720–724, 2004.[Web of Science][Medline]

40. Melcher AH, Chan J. Phagocytosis and digestion of collagen by gingival fibroblasts in vivo: a study of serial sections. J Ultrastruct Res 77: 1–36, 1981.[CrossRef][Web of Science][Medline]

41. Meller AT, Rumjanek VM, Sansone C, Allodi S. Oral mucosa alterations induced by cyclosporin in mice: morphological features. J Periodontal Res 37: 412–415, 2002.[CrossRef][Web of Science][Medline]

42. Moseley JB, Okada K, Balcer HI, Kovar DR, Pollard TD, Goode BL. Twinfilin is an actin-filament-severing protein and promotes rapid turnover of actin structures in vivo. J Cell Sci 119: 1547–1557, 2006.[Abstract/Free Full Text]

43. Ono M, Hatamochi A, Arakawa M, Ueki H. Effects of cyclosporin A on cell proliferation and collagen production by human skin fibroblasts. J Dermatol Sci 2: 274–280, 1991.[CrossRef][Medline]

44. Paik JW, Kim CS, Cho KS, Chai JK, Kim CK, Choi SH. Inhibition of cyclosporin A-induced gingival overgrowth by azithromycin through phagocytosis: an in vivo and in vitro study. J Periodont 75: 380–387, 2004.[CrossRef][Medline]

45. Perez-Tamayo R. Pathology of collagen degradation. A review. Am J Pathol 92: 508–566, 1978.[Medline]

46. Pollard TD, Borisy GG. Cellular motility driven by assembly and disassembly of actin filaments. Cell 112: 453–465, 2003.[CrossRef][Web of Science][Medline]

47. Ringden O, Bergstrand A, Collste H, Grimfors G, Lundgren G, Groth CG. Immunosuppression by Cyclosporin A in human renal transplant recipients. Scand J Urol Nephrol 64: 220–229, 1981.

48. Russell RG, Graveley R, Coxon F, Skjodt H, Del Pozo E, Elford P, Mackenzie A. Cyclosporin A: mode of action and effects on bone and joint tissues. Scand J Rheumatol 95: 9–18, 1992.

49. Scavuzzo M, Sagripanti A, Mosca F, Ambrogi F. Modulation of beta2 integrin phenotype, adhesion, chemotaxis, and oxidative burst of neutrophils by cyclosporine. Biomed Pharmacother 55: 61–69, 2001.[CrossRef][Medline]

50. Schincaglia GP, Forniti F, Cavallini R, Piva R, Calura G, del Senno L. Cyclosporin-A increases type I procollagen production and mRNA level in human gingival fibroblasts in vitro. J Oral Pathol Med 21: 181–185, 1992.[CrossRef][Web of Science][Medline]

51. Schreiber SL, Crabtree GR. The mechanism of action of cyclosporin A and FK506. Immunol Today 13: 136–142, 1992.[CrossRef][Web of Science][Medline]

52. Serrander L, Skarman P, Rasmussen B, Witke W, Lew DP, Krause KH, Stendahl O, Nusse O. Selective inhibition of IgG-mediated phagocytosis in gelsolin-deficient murine neutrophils. J Immunol 165: 2451–2457, 2000.[Abstract/Free Full Text]

53. Spolidorio LC, Merzel J, Villalba H, Vargas PA, Coletta RD, Almeida OP. Morphometric evaluation of gingival overgrowth and regression caused by cyclosporin in rats. J Periodontal Res 36: 384–389, 2001.[CrossRef][Web of Science][Medline]

54. Spolidorio LC, Spolidorio DM, Massucato EM, Neppelenbroek KH, Campanha NH, Sanches MH. Oral health in renal transplant recipients administered cyclosporin A or tacrolimus. Oral Dis 12: 309–314, 2006.[CrossRef][Web of Science][Medline]

55. Tonshoff B, Hocker B. Treatment strategies in pediatric solid organ transplant recipients with calcineurin inhibitor-induced nephrotoxicity. Pediatr Transplant 10: 721–729, 2006.[CrossRef][Web of Science][Medline]

56. Valerius NH, Stendahl O, Hartwig JH, Stossel TP. Distribution of actin-binding protein and myosin in polymorphonuclear leukocytes during locomotion and phagocytosis. Cell 24: 195–202, 1981.[CrossRef][Web of Science][Medline]

57. Wang Q, Herrera Abreu MT, Siminovitch K, Downey GP, McCulloch CA. Phosphorylation of SHP-2 regulates interactions between the endoplasmic reticulum and focal adhesions to restrict interleukin-1-induced Ca2+ signaling. J Biol Chem 281: 31093–31105, 2006.[Abstract/Free Full Text]

58. Witke W, Sharpe AH, Hartwig JH, Azuma T, Stossel TP, Kwiatkowski DJ. Hemostatic, inflammatory, and fibroblast responses are blunted in mice lacking gelsolin. Cell 81: 41–51, 1995.[CrossRef][Web of Science][Medline]

59. Yin HL, Albrecht JH, Fattoum A. Identification of gelsolin, a Ca2+-dependent regulatory protein of actin gel-sol transformation, and its intracellular distribution in a variety of cells and tissues. J Cell Biol 91: 901–906, 1981.[Abstract/Free Full Text]

60. Zhao XH, Laschinger C, Arora P, Szaszi K, Kapus A, McCulloch CA. Force activates smooth muscle {alpha}-actin promoter activity through the Rho signaling pathway. J Cell Sci 22: 222–223, 2007.





This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/3/C1049    most recent
00027.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Chan, M. W. C.
Right arrow Articles by McCulloch, C. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Chan, M. W. C.
Right arrow Articles by McCulloch, C. A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.