help button home button Endocrine Society Endocrinology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Purchase Article
Right arrow View Shopping Cart
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 PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pedrozo, H. A.
Right arrow Articles by Boyan, B. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pedrozo, H. A.
Right arrow Articles by Boyan, B. D.
Endocrinology Vol. 140, No. 12 5806-5816
Copyright © 1999 by The Endocrine Society

ARTICLES

Potential Mechanisms for the Plasmin-Mediated Release and Activation of Latent Transforming Growth Factor-ß1 from the Extracellular Matrix of Growth Plate Chondrocytes1

H. A. Pedrozo, Z. Schwartz, M. Robinson, R. Gomez, D. D. Dean, L. F. Bonewald and B. D. Boyan

Departments of Orthopaedics (H.A.P., Z.S., M.R., R.G., D.D.D., B.D.B.), Periodontics (Z.S., B.D.B.), Biochemistry (L.F.B., B.D.B.), and Medicine (L.F.B.), The University of Texas Health Science Center, San Antonio, Texas 78229-3900; and Department of Periodontics (Z.S.), Hebrew University, Hadassah Faculty of Dental Medicine, Jerusalem, Israel 91010

Address all correspondence and requests for reprints to: Barbara D. Boyan, Ph.D., Department of Orthopaedics (7774), The University of Texas Health Science Center at San Antonio, 7703 Floyd Curl Drive, San Antonio, Texas 78229-3900. E-mail: BoyanB{at}uthscsa.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Chondrocytes produce latent transforming growth factor-ß1 (TGF-ß1) in a small, circulating form of 100 kDa and also store latent TGF-ß1 in their matrix in a large form of 290 kDa containing the latent TGF-ß1 binding protein 1. As growth plate cartilage cells are exceptionally sensitive to TGF-ß1 and are known to produce plasminogen activator, the role of plasmin in the activation of soluble and matrix-bound latent TGF-ß1 was examined. As is true for other cell types, low-dose plasmin (0.01 U/ml) was found to release both active and latent TGF-ß1 from chondrocyte matrix in a time-dependent manner over 3 h. However, high-dose plasmin (1.0 U/ml) was found to release active TGF-ß1 more rapidly than low-dose plasmin, and this release ceased within 30 min; latent complex continued to be released over time (3 h). When high-dose plasmin was titrated against the serine protease inhibitors, aprotinin and {alpha}-(2-aminoethyl)benzenesulfonyl fluoride, results similar to low-dose plasmin were obtained, indicating that the effects of high-dose plasmin could be altered to mimic those of low-dose plasmin. No differences were observed on the effects of plasmin on the release of TGF-ß1 from the matrices of either growth zone or resting zone chondrocytes.

We examined whether plasmin could further activate the truncated large latent TGF-ß1 complex of 230 kDa that was released into the media by plasmin. It is known that plasmin will activate the small latent complex, so this was compared with the truncated form. Plasmin completely activated the small latent complex, whereas a smaller, but significant, activation of the truncated form of latent TGF-ß1 also occurred. These studies may have relevance to normal physiological conditions, where plasminogen and/or plasmin is present in very small amounts in the cartilage and, therefore, small amounts of active TGF-ß1 would be present, and to pathological conditions such as fractures, where chondroprogenitor cells would be exposed to high concentrations of plasmin and, therefore, to short-term high concentrations of this potent chondrogenic growth factor.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GROWTH PLATE chondrocytes produce and release transforming growth factor-ß1 (TGF-ß1) in latent form (1, 2). These cells are particularly sensitive to TGF-ß1, exhibiting phenotypic responses to the growth factor at concentrations 10- to 100-fold lower than has been reported for osteoblasts (3, 4, 5). Thus, the mechanisms by which cartilage cells store and activate TGF-ß1 are of interest.

Like latent TGF-ß1 produced by osteoblasts (3), latent TGF-ß1 produced by chondrocytes consists of a 100-kDa complex of mature TGF-ß1 homodimer (25 kDa) noncovalently associated with a latency-associated peptide (LAP) homodimer (75 kDa) (6, 7), termed small latent TGF-ß1. In addition to the small 100-kDa latent TGF-ß1, chondrocytes produce a large latent complex composed of the small 100-kDa form covalently bound to a 190-kDa protein, latent TGF-ß binding protein 1 (LTBP1). The relative amounts of small and large latent TGF-ß1 appear to be tissue specific. Whereas chondrocytes produce free small and large latent TGF-ß1 in similar proportion to those produced by osteoblasts (8, 9), fibroblasts and liver cells only produce large latent TGF-ß1 (10, 11). Platelets produce a large latent TGF-ß1 complex containing a truncated form of LTBP1 with a molecular mass of 130 kDa (12). Despite these differences, in all latent complexes, dissociation of mature TGF-ß1 homodimer from the LAP is necessary for biological activity (12).

LTBP1 has been shown to play a role in directing the latent complex to the extracellular matrix for storage in a number of cell types (8, 13, 14). In growth plate chondrocytes, LTBP1 expression is regulated by 1,25-(OH)2D3 in a cell maturation-dependent manner, resulting in a decrease in production of small latent TGF-ß1 into the medium of growth zone cell cultures and an increase in the incorporation of large latent TGF-ß1 into the extracellular matrix (6). LTBP1 appears to be bound to the extracellular matrix via cross-links catalyzed by transglutaminase (13). The primary structure of LTBP1 supports the hypothesis that it is involved in matrix structure and function, at least in bone (14), since LTBP1 shares some characteristics in common with structural proteins, especially with the fibrillin family of extracellular proteins (15, 16, 17). Although it binds small latent TGF-ß1, the LTBP1 molecule does not confer latency to TGF-ß1 (7, 18).

Little is known about the activation of the large latent complex after it has been stored in the extracellular matrix. It has been proposed that certain serine proteases, such as plasmin, may play a role in this activation process by releasing the latent complex after cleavage of the LTBP1 molecule at its plasmin-sensitive hinge (9, 13, 14). This truncated form of the large latent complex is similar to the truncated complex released by platelets (12). A mechanism for the activation of stored latent TGF-ß1 has been proposed by Nunes et al. (13). According to this mechanism, plasmin-mediated release of the matrix-associated large latent TGF-ß1 complex exposes the mannose-6-phosphate residues in the LAP that interact with the mannose-6-phosphate/insulin-like growth factor II receptors on the cell surface. The LAP molecule is then cleaved by membrane-associated plasmin to liberate mature TGF-ß1 from the complex. However, there is no evidence to date that membrane-associated plasmin acts directly on the latent complex to release the active homodimer.

Recent studies have demonstrated that recombinant latent TGF-ß1 can also be activated by discrete regions of thrombospondin in vitro (19, 20) and in vivo (21). Thrombospondin is an extracellular matrix protein and appears to activate latent TGF-ß1 by inducing conformational changes in the LAP (19). More recently, it has been shown that binding of {alpha}vß6 integrin to the RGD sequences in the LAP induces activation of the small latent TGF-ß1 (22). These observations suggest that multiple mechanisms exist for the release of the large latent complex from the extracellular matrix and for mediating the release of the active homodimer.

We have shown that plasmin releases the large latent TGF-ß1 complex from the extracellular matrix of growth plate chondrocytes and leads to its activation (8). Whether this is due to a direct action of plasmin on the large complex or part of a cascade in which activation follows release is not known. Plasminogen activator is present in the chondrocyte cultures, and its activity is enriched in extracellular matrix vesicles (23), which have been shown to activate small latent TGF-ß1 in vitro (1). Moreover, plasminogen activator activity is higher in matrix vesicles from growth zone chondrocytes, indicating that the enzyme may function in a cell maturation-dependent manner in the release and activation of the large latent TGF-ß1 complex. To better understand the role of plasmin in this process, we compared the ability of plasmin to release and activate TGF-ß1 associated with the extracellular matrix of chondrocytes at two distinct states of endochondral development.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Chondrocyte culture model
The chondrocyte culture system used in this study has been described in detail previously (24, 25). Chondrocytes were isolated from the resting zone (RC) and growth zone (GC) of the costochondral cartilages from 125 g Sprague Dawley rats. The cells were cultured at 37 C in DMEM containing 10% FBS, 1% penicillin/streptomycin, and 50 µg/ml vitamin C in an atmosphere of 5% CO2 and 100% humidity. Fourth passage cells were used for all experiments. Previous studies have shown that these cells retain their chondrocytic phenotype and differential responsiveness to 1,25-dihydroxyvitamin D3 (1,25-(OH)2D3) and 24,25-dihydroxyvitamin D3 (24,25-(OH)2D3), as well as to TGF-ß1, through four passages in culture (5, 26, 27).

Release of LTBP1 and TGF-ß1 from the matrix
Extracellular matrix preparation. Fourth passage RC and GC cells were cultured to confluence in 24-well plates. At harvest, the media were removed, the cell layers washed three times with PBS, and the cells lysed by three successive 10-min washes with RIPA buffer containing 50 mM Tris, 150 mM NaCl, 1% NP40, and 0.5% deoxycholate. The remaining cell-free nonsolubilized matrix was washed three times with PBS and digested with 0.01 or 1.0 U/ml of plasmin in DMEM for 3 h at 37 C to release large latent TGF-ß1 complexes through cleavage of LTBP1 from the matrix. The reaction was stopped by addition of aprotinin (Sigma, St. Louis, MO) to a final concentration of 5 µg/ml and immediately assayed for active and latent TGF-ß1 by enzyme-linked immunosorbent assay (ELISA) as described below. For each experiment, an equal number of cells was seeded into each well. Because the entire resulting matrix was used to determine the amount of TGF-ß1 released, we did not determine either the protein content of the culture or the DNA content of the lysed cells. Thus, these data are expressed as picograms/well on the assumption that any differences are a direct consequence of treatment of the cultures. Wells containing only media but no matrices served as controls and were treated and assayed as those containing matrix. The inclusion of the "no matrix" groups enabled us to control for the potential interference of media components. In addition, wells containing no plasmin were included to control for the spontaneous release of TGF-ß1 from the matrix.

Measurement of TGF-ß1. The ELISA for measuring TGF-ß1 levels was performed according to the manufacturer’s instructions (catalog no. G1230, Promega Corp., Madison, WI). The plates were coated overnight at 4 C and incubated with blocking buffer for 35 min at 37 C, and the samples and standard were added to the wells for 1.5 h at room temperature. Plates were washed and incubated with anti-TGF-ß1 antibody for 2 h at room temperature, followed by a wash and incubation with conjugated antibody for 2 h at room temperature. Color development was achieved by addition of the substrate provided in the kit, and the reaction was allowed to proceed for 4 min. When color development was complete, the reaction was stopped by addition of 1 M phosphoric acid and the absorbance at 450 nm was measured.

Since the immunoassay is designed to detect the active TGF-ß1 homodimer alone, quantitation of latent TGF-ß1 was performed using acid activation. The samples were prepared in the following fashion. To test for active TGF-ß1, 100 µl of sample were added directly to each well. To determine total TGF-ß1, 50 µl of sample were brought to a final volume of 90 µl with buffer and then acidified by addition of 10 µl of 1 M HCl. After 15 min, the samples were neutralized with 1 M NaOH. One hundred microliters of each sample were tested in the ELISA immediately after acid activation. The amount of latent TGF-ß1 was determined by subtracting the amount of active TGF-ß1 from total TGF-ß1 in each sample.

Matrix digestion with plasmin
Dose response. Fourth passage RC and GC cells were plated on 24-well culture plates, and at confluency, the extracellular matrices were isolated by sequential cell lysis with RIPA buffer as described above. Matrices were digested with 0.0005, 0.001, 0.005, 0.01, 0.05, 0.5, and 1 U/ml of plasmin (Sigma) for 3 h at 37 C. Active and latent TGF-ß1 was measured by ELISA.

Time course. Fourth passage RC and GC cells were plated on 24-well culture plates, and at confluency, the extracellular matrices were isolated by sequential cell lysis with RIPA buffer as described above. Matrices were digested with either 0.01 or 1 U of plasmin/ml DMEM for 1, 5, 15, 30, 60, 120, and 180 min at 37 C. Active and latent TGF-ß1 was measured by ELISA.

Effect of serine protease inhibition. To determine whether plasmin contributes to the release of active TGF-ß1 from the chondrocyte matrix, we incubated RC matrices with plasmin in the presence of aprotinin, which is a specific inhibitor of serine proteases such as trypsin, chymotrypsin, kallikrein, and plasmin (28). These studies also provided a control on the dose-dependent effects of plasmin, since a constant concentration of plasmin was inhibited by various concentrations of aprotinin, creating a dose-response experimental design. To verify that the effects measured resulted from inhibition of plasmin, another inhibitor, {alpha}-(2-amino- ethyl)benzenesulfonyl fluoride (AEBSF) (Sigma) (29) was also used.

To assess the effects of aprotinin on plasmin activity, we used a plasmin assay based on modifications of the plasminogen activator assay developed by Coleman and Green (30). The reaction mixture contained 0.1% Triton-X100, 22 mM 5'5-dithiobis(2-nitrobenzoic acid), 50 mM Na2HPO4, and 20 mM thiobenzyl benzyloxycarbonyl-L-lysinate (Z-Lys-SBzl), 200 mM NaPO4, and 200 mM NaCl. Two-fold serial dilutions of plasmin were prepared in the reaction mixture, starting with 1 U/ml DMEM. To start the reaction, 50 µl of sample were added to 950 µl of plasmin solution and incubated at room temperature for 60 min. The reaction was terminated by the addition of 100 µl of 1 mg/ml of soybean trypsin inhibitor-dissolved 1.0 mM HCl. The assay was run in the presence or absence of aprotinin at final concentrations of 0.5, 5.0, and 50 µg/ml. All reagents were enzyme grade and were purchased from Sigma.

Aprotinin inhibited plasmin activity in a dose-dependent manner. At low concentrations of the inhibitor (0.5 µg/ml), there was no effect on plasmin activity. At low concentrations of plasmin, 5 µg/ml aprotinin blocked 90% of the enzyme activity, but only 50% of plasmin activity at high enzyme concentrations. Aprotinin (50 µg/ml) blocked 90% of the activity of 1 U/ml plasmin.

Fourth passage RC cells were plated on 24-well culture plates, and at confluency, the extracellular matrices were isolated by sequential cell lysis with RIPA buffer, as explained earlier. Matrices were digested with either 0.01 or 1 U/ml of plasmin for 15, 90, 120, and 180 min at 37 C. Five minutes after digestion started, aprotinin was added to a final concentration of 5 µg/ml of DMEM, and the samples were again incubated at 37 C for the time remaining. Active and latent TGF-ß1 were measured by ELISA as described above.

Matrices were also incubated with 1.0 U/ml plasmin in DMEM ± AEBSF at 0.1, 0.5, or 1.0 mM concentrations. AEBSF has been shown to inhibit thrombin and plasmin (29). Control samples received 1.0 mM AEBSF alone. As before, digestion took place for 3 h at 37 C, after which active TGF-ß1 was measured by ELISA.

Characterization of the large latent TGF-ß1 released by plasmin from the matrix
Preparation of soluble complexes. RC cells were plated in T-75 flasks, and at confluency, the matrices were prepared as described above by lysing the cells with 3 ml RIPA buffer and washing with excess PBS. The isolated matrices were digested with 2 ml 0.5 U plasmin/ml DMEM for 3 h at 37 C, immediately after which the digests were collected into a 50-ml tube and the high molecular mass proteins (>100 kDa) were isolated using an Ultrafree centrifugal filter device with a Biomax 100-kDa cut-off membrane (Millipore Corp., Bedford, MA) to remove any active TGF-ß1. Samples were spun at 2,000 x g for 20 min to force the lower molecular mass proteins down into the collecting tube. This step was repeated three times, and the contents in the filter device were mixed with a pipettor between spins. The protein concentration of each fraction was determined by a macro BCA protein assay (Pierce Chemical Co., Rockford, IL). Aliquots of both fractions were taken and the proteins were concentrated by ethanol precipitation. Ice-cold ethanol (100%) was added at 2.5-fold the volume of the sample; the samples were incubated in crushed, dry ice and centrifuged at 16,000 x g for 20 sec using an Eppendorf microcentrifuge (Brinkmann Instruments, Inc. Westbury, NY). The pellets were resuspended in PBS for protein determination.

Western blot analysis. To determine the nature of the latent TGF-ß1 complex, the pellets were resuspended in 10 µl of 2x nonreducing sample buffer, boiled for 5 min, run on a 4–20% SDS-polyacrylamide gel, and transferred overnight to a nitrocellulose membrane. The membrane was blocked with 5% Blotto for 1 h, washed in Tween-Tris buffered saline (T-TBS) containing 20 mM Tris base, 0.1% Tween 20, pH 7.6, and probed with rabbit anti-LTBP1 antibody (Ab39, a generous gift of Dr. Kohei Miyazono) (9) (1:1000) for 1 h at room temperature. After three 20-min washes in T-TBS, the membrane was incubated in a 1:1000 vol/vol dilution of horseradish peroxidase-labeled antirabbit IgG, for 1 h at room temperature. To visualize the bands, the enhanced chemiluminescence (ECL) Western blot analysis system (Amersham Pharmacia Biotech, Buckinghamshire, UK) was used according to the manufacturer’s instructions. To demonstrate association of LTBP1 with the latent TGF-ß1 molecule, the membrane was reprobed with anti-LAP antibody (1:1000 vol/vol), which specifically recognizes the latent TGF-ß1 homodimer (R & D Systems, Minneapolis, MN). Between Western blots, the membrane was stripped by incubation in a small volume of stripping buffer containing 100 mM 2-mercaptoethanol, 2% SDS, and 62.5 mM Tris-HCl, pH 6.7, for 30 min at 60 C. All antibody dilutions were prepared in T-TBS.

Effect of plasmin on large latent TGF-ß1 released from the matrix. To determine whether plasmin could directly activate the 230-kDa large latent TGF-ß1 released from matrix, we incubated plasmin-cleaved large latent TGF-ß1 with plasmin. Aliquots were prepared from the higher molecular mass fraction containing 0.5, 0.25, 0.1, 0.05, and 0.025 µg of total protein and incubated with either 1.0 or 0.01 U/ml of plasmin for 3 h at 37 C in a reaction volume of 150 µl. Active TGF-ß1 present in a 100 µl aliquot of each sample was measured by TGF-ß1 ELISA, as described above. Total TGF-ß1 present in the reaction volume was calculated from ELISA measurements in 50 µl aliquots after acid activation. Control samples containing 1 µg protein were subjected to the same conditions, but received no plasmin and were acid activated before the 3-h incubation period.

For comparison to the truncated plasmin-generated 230 kDa complex, recombinant simian latent TGF-ß1 was used to represent the 100-kDa small latent TGF-ß1 also produced by chondrocytes. Recombinant simian TGF-ß1 was treated with plasmin, and the production of active TGF-ß1 was measured as described above. This small complex lacking LTBP1 has been shown to be activated by plasmin (31). The recombinant simian small latent TGF-ß1 was a generous gift of Dr. Dan Twardzik. Recombinant simian TGF-ß1 (50 µl containing 200 ng/ml) was acid activated by the addition of 10% (vol/vol) of 1 M HCl for 15 min followed by neutralization with equimolar amounts of NaOH as a positive control. Alternatively, samples were incubated with 0.01 or 1 U/ml of plasmin. All reactions were performed in a total volume of 100 µl for 3 h at 37 C. At the termination of the reaction, the entire reaction volume (100 µl) was added to the ELISA plate.

Statistical analysis
All experiments described in this study were performed at least twice to ensure validity of the results. Data presented are from representative experiments and are the mean ± SEM for six separate cultures. Data were analyzed by ANOVA, and post hoc testing was performed using Student’s t test with Bonferroni’s modification.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effect of plasmin on the release of TGF-ß1 from chondrocyte matrices
Dose response. Plasmin released both active and latent TGF-ß1 from the extracellular matrix of RC and GC chondrocytes (Fig. 1Go). However, there was a differential effect of plasmin dose on release vs. activation of latent TGF-ß1. In matrices prepared from RC cells, the release of active growth factor peaked at 0.01 U/ml of plasmin (Fig. 1AGo), while release of the latent complex continued to increase with increasing plasmin concentration (Fig. 1BGo). At 0.01 U/ml plasmin, 380 pg of active TGF-ß1 and 214 pg of latent TGF-ß1 were released per culture well, whereas at 1 U/ml plasmin, 104 pg active TGF-ß1 and 2,326 pg latent TGF-ß1 per well were released. The extracellular matrix of GC chondrocytes responded in a similar manner. The release of active TGF-ß1 peaked at 0.005 U/ml of plasmin (Fig. 1CGo), while the release of latent TGF-ß1 continued to increase with increasing enzyme concentrations (Fig. 1DGo). At 0.005 U/ml plasmin, 270 pg active TGF-ß1 and 482 pg latent TGF-ß1 per well were released, while at 1 U/ml plasmin, 104 pg active TGF-ß1 and 2,902 pg latent TGF-ß1 per well were released.



View larger version (26K):
[in this window]
[in a new window]
  		Figure 1. Dose-dependent release of active and latent TGF-ß1 from RC and GC extracellular matrix. Extracellular matrix was isolated from RC (panels A and B) and GC (panels C and D) cultures as described in Materials and Methods, digested with various concentrations of plasmin for 3 h at 37 C, and then assayed for release of active and total TGF-ß1 by ELISA. Latent TGF-ß1 (bottom panels) was determined by subtracting active (top panels) from total TGF-ß1. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (plasmin = 0 U/ml).

 
Time course. The differential effects of plasmin dose on latent TGF-ß1 release and activation were time dependent (Fig. 2Go). Treatment of RC matrices with the high concentration of plasmin resulted in release of active TGF-ß1 as early as 1 min after addition of the enzyme; peak release occurred by 5 min (Fig. 2AGo). No active TGF-ß1 could be detected after 60 or more minutes of incubation. In contrast, the release of latent TGF-ß1 increased with increasing time (Fig. 2BGo). After 5 min of treatment with plasmin, 29 pg of active TGF-ß1 were released per well, whereas 988 pg latent TGF-ß1 per well were released. After the 3-h incubation, 2,506 pg per well of the latent growth factor were released from the matrix and reached levels that were 100-fold greater than that seen for maximal release of active TGF-ß1. Treatment of RC matrices with the low plasmin concentration resulted in release of active TGF-ß1 that peaked at 90 min (Fig. 2CGo). The amount released after 5 min with low plasmin treatment was comparable to that released by high plasmin (Fig. 2Go, A vs. C). However, at 90 min, low plasmin treatment caused a 5.6-fold greater increase in active TGF-ß1 release. The release of latent TGF-ß1 also increased with time, reaching maximal amounts at 90–120 min (Fig. 2DGo); however, maximal release was only 20% of that seen with the high plasmin treatment.



View larger version (30K):
[in this window]
[in a new window]
  		Figure 2. Time-dependent release of active and latent TGF-ß1 from RC extracellular matrix by plasmin. Extracellular matrix was isolated from RC cultures as described in Materials and Methods, digested with 1.0 (panels A and B) or 0.01 (panels C and D) U/ml plasmin for 0–180 min at 37 C, and then assayed for release of active and total TGF-ß1 by ELISA. Latent TGF-ß1 (bottom panels) was determined by subtracting active (top panels) from total TGF-ß1. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (plasmin = 0 U/ml).

 
In GC matrices, release of active TGF-ß1 by 1 U/ml of plasmin was rapid; maximal release occurred within 1 min of enzyme addition and then decreased with time (Fig. 3AGo). Release of latent TGF-ß1 was also significant within 1 min, but it continued to increase with time (Fig. 3BGo).



View larger version (25K):
[in this window]
[in a new window]
  		Figure 3. Time-dependent release of active and latent TGF-ß1 from GC extracellular matrix by 1 U/ml plasmin. Extracellular matrix was isolated from GC cultures as described in Materials and Methods, digested with 1.0 U/ml plasmin for 0–180 min, and then assayed for active (panel A) and latent (panel B) TGF-ß1 by ELISA. Latent TGF-ß1 was determined by subtracting active from total TGF-ß1. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (no matrix).

 
At the low plasmin concentration (0.01 U/ml), no latent TGF-ß1 was released from GC matrices (Fig. 4Go), although latent TGF-ß1 was present. Active TGF-ß1 was released after 30–60 min, and release continued over time, resulting in a peak of 124 pg per well at 180 min.



View larger version (25K):
[in this window]
[in a new window]
  		Figure 4. Time-dependent release of active and latent TGF-ß1 by 0.01 U/ml plasmin from GC extracellular matrix. Extracellular matrix was isolated from GC cultures as described in Materials and Methods, digested with 0.01 U/ml plasmin for 0–180 min, and then assayed for active and latent TGF-ß1 by ELISA. Latent TGF-ß1 was determined by subtracting active from total TGF-ß1. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (no matrix).

 
Aprotinin (5 µg/ml) inhibited the release of active TGF-ß1 by 0.01 U/ml plasmin from RC matrices by 57–85% at all times examined (Fig. 5AGo). At 15 min, exogenous plasmin had a modest effect on basal release of latent TGF-ß1 when compared with the no-plasmin control (Fig. 5BGo). At later incubation times, aprotinin reduced the plasmin-dependent release of latent TGF-ß1 to levels comparable to the no-plasmin control.



View larger version (36K):
[in this window]
[in a new window]
  		Figure 5. Time-dependent release of active and latent TGF-ß1 from RC extracellular matrix by 0.01 and 1.0 U/ml plasmin in the presence of aprotinin. Isolated RC extracellular matrix was digested with 0.01 U/ml (panels A and B) or 1.0 U/ml (panels C and D) plasmin for 5 min, followed by addition of the plasmin inhibitor, aprotinin (5 µg/ml), or DMEM as control. The samples were then incubated for an additional 10–175 min and assayed for active (panels A and C) and latent (panels B and D) TGF-ß1 by ELISA. Latent TGF-ß1 was determined by subtracting active from total TGF-ß1. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (no matrix); #, P < 0.05, aprotinin vs. no aprotinin.

 
When RC matrices were incubated with 1 U/ml plasmin, aprotinin had no effect on release of active TGF-ß1 at 15 min, but at later time points, release was increased by 135–360% (Fig. 5CGo). In contrast, aprotinin reduced release of latent TGF-ß1 by 66.7% at 15 min and by 36% at 90, 120, and 180 min (Fig. 5DGo). Treatment of RC matrices with 1 U/ml of plasmin in the presence of the serine protease inhibitor AEBSF caused a dose-dependent increase in TGF-ß1 release (Fig. 6Go).



View larger version (18K):
[in this window]
[in a new window]
  		Figure 6. Activation of latent TGF-ß1 from RC extracellular matrix by 1.0 U/ml plasmin in the presence of AEBSF. Isolated RC extracellular matrices were digested with 1.0 U/ml plasmin in the presence or absence of AEBSF for 3 h. The digests were assayed for active TGF-ß1 by ELISA. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (0); #,P < 0.01, 1.0 mM or 0.5 mM vs. 0.1 mM AEBSF; •, P < 0.01, 1.0 mM vs. 0.5 mM AEBSF.

 
Effect of plasmin on soluble large latent TGF-ß1 complexes
Western blot analysis. Western blots using anti-LTBP1 antibody and anti-LAP antibody showed the presence of cleaved large latent complex of 230 kDa and free LTBP of 130 kDa in plasmin digests of RC matrices. Immunoblotting with anti-LAP antibody recognized the 230-kDa band (Fig. 7AGo). Other bands were also recognized by the antibody, notably a band at 205 kDa. This band corresponds to the large latent TGF-ß1 complex lacking the 60-kDa LTBP1 fragment and the 25-kDa TGF-ß1 homodimer and is believed to result from activation of the complex by SDS in the sample buffer (12, 32). Immunoblotting using anti-LTBP1 antibody demonstrated specific recognition of two bands, one at approximately 230 kDa and one at 130 kDa (Fig. 7BGo). Western blots of the low molecular mass fraction of the plasmin digests did not contain immunoreactive complex.



View larger version (43K):
[in this window]
[in a new window]
  		Figure 7. Western blot analysis of latent TGF-ß1 complexes released from RC extracellular matrix and fractionated based on molecular size. Isolated RC matrices were digested with 0.5 U/ml plasmin for 3 h at 37 C, and proteins less than 100 kDa were separated from those more than 100 kDa by an Ultrafree centrifugal filter device. The effective separation of proteins in this digest was demonstrated by Western blot analysis of 1- and 2-µg aliquots of the two fractions with antibodies specific for the LAP (panel A) and LTBP1 (panel B).

 
Release of active TGF-ß1 from soluble truncated large latent TGF-ß1 generated by plasmin digestion of RC matrices. Plasmin further activated TGF-ß1 in the 230-kDa released large latent TGF-ß1 complex. Plasmin at 1 U/ml caused a modest activation of soluble, latent TGF-ß1, which increased with increasing amounts of soluble plasmin-released complex (Fig. 8AGo). At 0.125 µg of soluble complex, the enzyme activated 34% of the total latent TGF-ß1 that could be activated with acid activation. However, at higher concentrations of the soluble complex, activation was reduced to 22%. Plasmin at 0.01 U/ml activated only 5.6% of the latent TGF-ß1 present in solution (Fig. 8BGo).



View larger version (36K):
[in this window]
[in a new window]
  		Figure 8. Dose-dependent activation of soluble latent TGF-ß1 complexes. Varying concentrations of soluble complex isolated from RC extracellular matrix were treated with 1 U/ml (panel A) or 0.01 U/ml (panel B) plasmin for 3 h at 37 C or treated with plasmin at the same concentration (i.e. 1.0 or 0.01 U/ml) for 3 h, acid activated, and then assayed for active TGF-ß1 by ELISA. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. control (0 µg soluble complex); #,P < 0.05, plasmin vs. plasmin + acid activated (AA).

 
In comparison with the effects of plasmin on the truncated, plasmin-generated complex, plasmin activated recombinant simian latent TGF-ß1 in a concentration-dependent manner (Fig. 9Go). Plasmin (0.01 U/ml) activated approximately 21% of the latent TGF-ß1 activated by 1 U/ml plasmin. Plasmin appeared to be more effective than acid activation in generating TGF-ß1. Whereas acid activation resulted in a 6.7-fold increase in active TGF-ß1, treatment with 1 U/ml plasmin resulted in a 9.5-fold increase.



View larger version (17K):
[in this window]
[in a new window]
  		Figure 9. Activation of purified recombinant latent TGF-ß1. CHO-expressed recombinant simian latent TGF-ß1 was incubated in the presence of 0.01 or 1 U/ml plasmin for 3 h at 37 C. For controls, one group was left untreated, and the other was acid activated (AA) with 1 M HCl for 15 min and neutralized with the same milliequivalents of NaOH. Values shown are the mean ± SEM, n = 6 per group. *, P < 0.05, treatment vs. no AA; #, P < 0.05, plasmin vs. AA.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Activation of even minute amounts of latent TGF-ß1 can have profound effects on cartilage. Chondrocytes have been shown to respond to active TGF-ß1 with greater sensitivity than any other cell type. Chondrocytes exhibit maximal responses at 0.1 to 0.2 ng/ml and can respond at concentrations as low as 0.01 ng/ml (33, 34). These concentrations are 10- to 100-fold below plasma levels of TGF-ß, which range between 1–50 ng/ml (35, 36). In this study, we show that latent TGF-ß1 found in the extracellular matrix of both resting zone and growth zone cells can be released both in a soluble, inactive form and as active TGF-ß1 through the enzymatic activities of plasmin. The ratio of latent and active TGF-ß1 and the dynamics of activation of latent TGF-ß1 were different depending on whether high- or low-dose plasmin was used. This has implications for both physiological and pathological conditions in which plasmin enzyme activity and, therefore, active TGF-ß1 levels may be dramatically different (37, 38).

In previous studies, we have shown that resting zone and growth zone chondrocytes respond to factors and function in a maturation-specific manner (26). However, in the present study, no significant differences were observed between the extracellular matrices of these cells with respect to release of latent TGF-ß1 complex or the generation of active TGF-ß1. With both matrices, both high- and low-dose plasmin resulted in the continuous release of latent TGF-ß1 over time. With both matrices, low-dose plasmin resulted in continuous release of active TGF-ß1 over time, but high-dose plasmin resulted in a burst of active TGF-ß1, followed by reduced or nondetectable growth factor. It is more likely that maturation-specific differences in availability of active TGF-ß1 may be due to variations in the amount of latent TGF-ß1 present in the matrix (8), the activity of membrane-associated plasminogen activator (23), or the local plasminogen concentration.

Activation of matrix-bound latent TGF-ß1 by plasmin was not dependent on prior release of latent complex from the matrix. High-dose plasmin resulted in an early burst of active TGF-ß1 when latent TGF-ß1 was slowly being released. Therefore, high-dose plasmin may be activating latent TGF-ß1 before it is cleaved from the matrix. This same dose of plasmin was also more efficient in activating the truncated, plasmin-released latent complex (230 kDa) compared with low-dose plasmin. However, the soluble truncated latent form is still much less susceptible to activation than the small, 100-kDa latent form. These data suggest that the LTBP1 molecule may be partially protecting the latent complex from activation. Therefore, the primary target of low-dose plasmin is LTBP1, causing the release of latent complex from the matrix, and a secondary, weaker effect is the activation process. The reverse may be true for high-dose plasmin.

LTBP1 does not confer latency to the complex (7, 18), but does appear to retard or prevent protease-mediated activation. LTBP1 has been shown to contain a plasmin-sensitive hinge region from amino acids 413 to 506, cleavage of which results in the truncated soluble form of latent TGF-ß1 (10). Antibody to LTBP1 or free excess LTBP1 inhibits the activation of latent TGF-ß1 in the endothelial cell coculture system (39). Other data using the same culture system show that the large latent TGF-ß1 complex cannot be activated unless it is cross-linked to the extracellular matrix and that treatment with an antibody specific for the carboxy terminus of the LTBP1 molecule abrogates activation without interfering with the cross-linking of the LTBP1 molecule to the extracellular matrix (13). The present data suggest that, in addition to its role as an extracellular matrix protein (14) and in mediating the storage and release of latent TGF-ß1 from the matrix (3, 8, 10), the LTBP1 molecule protects the latent complex from activation. It has been proposed that the truncated form of latent TGF-ß1 released from the matrix through plasmin proteolysis has exposed carbohydrate that can then bind to mannose-6-phosphate receptors for cell surface activation, suggesting that intact LTBP1 masks these binding sites and, therefore, prevents activation (13). This suggests that the truncated large latent complex is protected from activation until it can associate with the cell surface.

However, plasmin has also been shown to activate latent TGF-ß1 in the conditioned media of fibroblasts (40), neural crest cells (41), and Chinese hamster ovary (CHO) cells producing small latent TGF-ß1 (31). Most cell types, including fibroblasts, mainly produce the large latent complex containing intact LTBP1 of 290 kDa. Therefore, these data show that plasmin can also activate large complexes not associated with the matrix. Our studies show that plasmin can activate the truncated, soluble large latent complex. This activation was considerably less efficient than the activation of the small latent complex, but still significant. Considering the sensitivity of chondrocytes to TGF-ß1, this activation mechanism is relevant for chondrocyte function.

Serine protease inhibitors altered the effects of high-dose plasmin to resemble those of low-dose plasmin. For example, at the time point when little or no active TGF-ß1 is observed with high-dose plasmin, AEBSF reversed this effect dose dependently, resulting in the production of active TGF-ß1. Aprotinin efficiently inhibited low-dose plasmin, but only partially inhibited high-dose plasmin, except at 15 min, where it had no effect. The inhibitor may not have completely saturated enzyme active sites by that time point. These results indicate that the underlying activation process is highly sensitive to plasmin dose and time of exposure.

Chondrocytes, therefore, have separate and distinct mechanisms for generating reservoirs of latent TGF-ß1 and for controlling the activation of these reservoirs. In a previous study, we showed that production and activation of latent TGF-ß1 are regulated by 1,25-(OH)2D3 and that matrix vesicles, which are extracellular organelles rich in neutral metalloproteinases and plasminogen activator, activate latent TGF-ß1 upon treatment with 1,25-(OH)2D3 (1). While stored in the matrix, latent TGF-ß1 is available to matrix vesicles for activation. This mechanism is under genomic and nongenomic regulation, allowing the cells to regulate the temporal and spatial activation of latent TGF-ß1 at sites remote from the cells. The results presented here further demonstrate that the extracellular matrix is an important site for the regulation of TGF-ß1.

The story becomes more complex when we consider the fact that the costochondral chondrocytes produce at least two isoforms of latent TGF-ß, TGF-ß1 and TGF-ß2 (1), which may exhibit overlapping functions. Although we only characterized the forms of latent TGF-ß1, it is likely that TGF-ß2 is also produced in two molecular forms, i.e. small latent and large latent TGF-ß2. The relative distribution of various latent forms of TGF-ß1 in vivo is also not known to date.

The results presented here are summarized in Fig. 10Go. The LTBP1 hinge region appears to play a crucial role in the production of active TGF-ß1 from latent matrix complexes. This region is highly susceptible to cleavage by plasmin, as is the small latent TGF-ß1 complex. The truncated, soluble large latent complex is less susceptible. Under conditions of high enzyme activity, such as may occur during trauma or inflammation, all three latent complexes become targets of plasmin, the matrix-bound latent TGF-ß1, the truncated, large latent complex, and the small latent complex. Under normal physiological conditions, matrix vesicles may be involved in the activation of local latent TGF-ß1 present in the extracellular matrix, resulting in primarily the release of latent complex for activation on the surfaces of cells distant from the matrix and secondarily in the release of smaller, yet biologically potent, amounts of active TGF-ß1.



View larger version (39K):
[in this window]
[in a new window]
  		Figure 10. Hypothetical model comparing the effect of low- (0.01 U/ml) and high-dose (1 U/ml) plasmin treatment on the release of active and latent TGF-ß1 over time. In panel A, low levels of plasmin are shown to release active TGF-ß1 directly from the large TGF-ß1 complex anchored in the matrix (1 ). Plasmin is also shown to cleave small quantities of the large complex in the plasmin-sensitive hinge region, releasing large latent TGF-ß1 complexes (2 ), and small amounts of active TGF-ß1 from the soluble large complex (3 ). In panel B, high levels of plasmin are shown to predominantly release large latent TGF-ß1 complexes by cleavage at the plasmin-sensitive hinge region (1 ). In addition, plasmin also releases a small amount of active TGF-ß1 (2 ) from matrix-associated large latent TGF-ß1 and a small amount from the soluble large complex (3 ). With short digestion times, only the high dose of plasmin releases active TGF-ß1.

 


   Acknowledgments
 
The authors acknowledge the technical contributions of Mr. Javier Chapa to this work and the assistance of Ms. Sandra Messier in the preparation of the manuscript.


   Footnotes
 
1 This work was supported by Public Health Service Grants DE-08603, DE-05937, and AR-43775, the Center for the Enhancement of the Biology/Biomaterials Interface at University of Texas Health Science Center, San Antonio, and the American Association for Dental Research. Back

Received April 19, 1999.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Boyan BD, Schwartz Z, Park-Snyder S, Dean DD, Yang F, Twardzik D, Bonewald LF 1994 Latent transforming growth factor-ß is produced by chondrocytes and activated by extracellular matrix vesicles upon exposure to 1,25-(OH)2D3. J Biol Chem 269:28374–28381[Abstract/Free Full Text]
  2. Gelb DE, Rosier RN, Puzas JE 1990 The production of transforming growth factor-ß by chick growth plate chondrocytes in short term monolayer culture. Endocrinology 127:1941–1947[Abstract]
  3. Bonewald LF 1996 Transforming growth factor-ß. In: Bilezikian JP, Raisz LG, Rodan GA (eds) Principles of Bone Biology. Academic Press, San Diego, CA, pp 647–659
  4. Schwartz Z, Bonewald LF, Caulfield K, Brooks BP, Boyan BD 1993 Direct effects of transforming growth factor-ß on chondrocytes are modulated by vitamin D metabolites in a cell maturation-specific manner. Endocrinology 132:1544–1552[Abstract]
  5. Schwartz Z, Sylvia VL, Liu Y, Dean DD, Boyan BD 1998 Treatment of resting zone chondrocytes with transforming growth factor-ß1 induces differentiation into a phenotype characteristic of growth zone chondrocytes by downregulating responsiveness to 24,25-(OH)2D3 and upregulating responsiveness to 1,25-(OH)2D3. Bone 23:465–470[Medline]
  6. Pedrozo HA, Schwartz Z, Mokeyev T, Ornoy A, Xin-Sheng W, Bonewald LF, Dean DD, Boyan BD 1999 Vitamin D3 metabolites regulate LTBP1 and latent TGF-ß1 expression and latent TGF-ß1 incorporation in the extracellular matrix of chondrocytes. J Cell Biochem 72:151–165[CrossRef][Medline]
  7. Bonewald LF, Wakefield LM, Oreffo RO, Escobedo A, Twardzik DR, Mundy GR 1991 Latent forms of transforming growth factor-ß (TGFß) derived from bone cultures: identification of a naturally occurring 100 kDa complex with similarity to recombinant latent TGFß. Mol Endocrinol 5:741–751[CrossRef][Medline]
  8. Pedrozo HA, Schwartz Z, Gomez R, Ornoy A, Xin-Sheng W, Dallas SL, Bonewald LF, Dean DD, Boyan BD 1998 Growth plate chondrocytes store latent TGF-ß1 in their matrix through latent TGFß binding protein. J Cell Physiol 177:343–354[CrossRef][Medline]
  9. Dallas SL, Park-Snyder S, Miyazono K, Twardzik D, Mundy GR, Bonewald LF 1994 Characterization and autoregulation of latent transforming growth factor ß (TGFß) complexes in osteoblast-like cell lines. J Biol Chem 269:6815–6822[Abstract/Free Full Text]
  10. Taipale J, Miyazono K, Heldin C-H, Keski-Oja J 1994 Latent transforming growth factor-ß1 associates to fibroblast extracellular matrix via latent TGFß binding protein. J Cell Biol 124:171–181[Abstract/Free Full Text]
  11. Marra F, Bonewald LF, Park-Snyder S, Park IS, Woodruff KA, Abboud HE 1996 Characterization and regulation of the latent transforming growth factor-ß (TGF-ß) complex secreted by vascular pericytes. J Cell Physiol 166:537–546[CrossRef][Medline]
  12. Miyazono K, Hellman U, Wernstedt C, Heldin CH 1988 Latent high molecular weight complex of transforming growth factor ß1. Purification from human platelets and structural characterization. J Biol Chem 263:6407–6415[Abstract/Free Full Text]
  13. Nunes I, Gleizes P-E, Metz CN, Rifkin DB 1997 Latent transforming growth factor-ß binding protein domains involved in activation and transglutaminase-dependent cross-linking of latent transforming growth factor-ß. J Cell Biol 136:1151–1163[Abstract/Free Full Text]
  14. Dallas SL, Miyazono K, Skerry TM, Mundy GR, Bonewald LF 1995 Dual role for the latent transforming growth factor-ß binding protein in storage of latent TGF-ß in the extracellular matrix and as a structural matrix protein. J Cell Biol 131:539–549[Abstract/Free Full Text]
  15. Sakai LY, Keene DR, Engvall E 1986 Fibrillin, a new 350 kD glycoprotein, is a component of extracellular microfibrils. J Cell Biol 103:2499–2509[Abstract/Free Full Text]
  16. Sakai LY, Keene DR, Glanville RW, Bachinger HP 1991 Purification and partial characterization of fibrillin, a cysteine-rich structural component of connective tissue microfibrils. J Biol Chem 266:14763–14770[Abstract/Free Full Text]
  17. Zhang H, Apfelroth SD, Hu W, Davis EC, Sanguineti C, Bonadio J, Mecham RP, Ramirez F 1994 Structure and expression of fibrillin-2, a novel microfibrillar component preferentially located in elastic matrices. J Cell Biol 124:855–863[Abstract/Free Full Text]
  18. Gentry LE, Webb NR, Lim GJ, Brunner AM, Ranchalis JE, Twardzik DR, Lioubin MN, Marquardt H, Purchio AF 1987 Type 1 transforming growth factor ß: amplified expression and secretion of mature and precursor polypeptides in Chinese hamster ovary cells. Mol Cell Biol 7:3418–3427[Abstract/Free Full Text]
  19. Schultz-Cherry S, Ribeiro S, Gentry L, Murphy-Ullrich JE 1994 Thrombospondin binds and activates the small and large forms of latent transforming growth factor-ß in a chemically defined system. J Biol Chem 269:26775–26782[Abstract/Free Full Text]
  20. Schultz-Cherry S, Chen H, Moshers DF, Misenheimer TM, Krutzsch HC, Roberts DD, Murphy-Ullrich JE 1995 Regulation of transforming growth factor-ß activation by discrete sequences of thrombospondin 1. J Biol Chem 270:7304–7310[Abstract/Free Full Text]
  21. Crawford SE, Stellmach V, Murphy-Ullrich JE, Ribeiro SMF, Lawler J, Hynes RO, Boivin GP, Bouck N 1998 Thrombospondin-1 is a major activator of TGF-ß1 in vivo. Cell 93:1159–1170[CrossRef][Medline]
  22. Munger JS, Huang X, Kawakatsu H, Griffiths MJD, Dalton SL, Wu J, Pittet J-F, Kaminski N, Garat C, Matthay MA, Rifkin DB, Sheppard D 1999 The integrin {alpha}vß6 binds and activates latent TGFß1: a mechanism for regulating pulmonary inflammation and fibrosis. Cell 96:319–328[CrossRef][Medline]
  23. Dean DD, Boyan BD, Muniz OE, Howell DS, Schwartz Z 1996 Vitamin D metabolites regulate matrix vesicle metalloproteinase content in a cell maturation-dependent manner. Calcif Tissue Int 59:109–116[CrossRef][Medline]
  24. Boyan BD, Schwartz Z, Swain LD, Carnes Jr DL, Zislis T 1988 Differential expression of phenotype by resting zone and growth region costochondral chondrocytes in vitro. Bone 9:185–194[Medline]
  25. Boyan BD, Schwartz Z, Swain LD 1992 In vitro studies on the regulation of endochondral ossification by vitamin D. Crit Rev Oral Biol Med 3:15–30[Abstract/Free Full Text]
  26. Boyan BD, Dean DD, Sylvia VL, Schwartz Z 1997 Cartilage and vitamin D: genomic and nongenomic regulation by 1,25-(OH)2D3 and 24,25-(OH)2D3. In: Feldman D, Glorieux FH, Pike JW (eds) Vitamin D. Academic Press, San Diego, CA, pp 395–421
  27. Schwartz Z, Sylvia VL, Dean DD, Boyan BD 1998 The synergistic effects of vitamin D metabolites and TGFß on costochondral chondrocytes are mediated by increases in protein kinase C activity involving two separate pathways. Endocrinology 139:534–545[Abstract/Free Full Text]
  28. Trautschold I, Werle E, Zickgraf-Rüdel G 1967 Trasylol. Biochem Pharmacol 16:59–72[CrossRef]
  29. Markwardt F, Hoffmann J, Körbs E 1973 The influence of synthetic thrombin inhibitors on the thrombin-antithrombin reaction. Thromb Res 2:343–348
  30. Coleman PL, Green GDJ 1981 A coupled protometric assay for plasminogen activator. In: Lorand L (ed) Methods in Enzymology: Proteolitic Enzymes, Part C. Academic Press, New York, pp 408–414
  31. Lyons RM, Gentry LE, Purchio AF, Moses HL 1990 Mechanisms of activation of latent recombinant transforming growth factor-ß by plasmin. J Cell Biol 110:1361–1367[Abstract/Free Full Text]
  32. Taipale J, Koli K, Keski-Oja J 1992 Release of transforming growth factor-ß1 from the pericellular matrix of cultured fibroblasts and fibrosarcoma cells by plasmin and thrombin. J Biol Chem 267:25378–25384[Abstract/Free Full Text]
  33. Schwartz Z, Dean DD, Boyan BD 1994 Interdependent regulation of chondrocyte differentiation by TGF-ß and vitamin D3 metabolites. In: Suzuki F, Boyan BD (eds) Proceedings of the International Symposium on Cartilage Metabolism. Osaka, Japan, pp 73–75
  34. Schwartz Z, Sylvia VL, Dean DD, Boyan BD 1996 The synergistic effect of TGFß and 24,25-(OH)2D3 on resting zone chondrocytes is metabolite-specific and mediated by PKC. Connect Tissue Res 35:101–106[Medline]
  35. Kyrtsonis MC, Repa C, Dedoussis GV, Mouzaki A, Simeonidis A, Stamatelou M, Maniatis A 1998 Serum transforming growth factor-ß 1 is related to the degree of immunoparesis in patients with multiple myeloma. Med Oncol 15:124–128[Medline]
  36. Rosensweig JN, Omori M, Page K, Potter CJ, Perlman EJ, Thorgeirsson SS, Schwarz KB 1998 Transforming growth factor-ß1 in plasma and liver of children with liver disease. Pediatr Res 44:402–409[Medline]
  37. Williams DM, Grubbs BG, Park-Snyder S, Rank RG, Bonewald LF 1996 Activation of latent transforming growth factor ß during chlamydia trachomatis-induced murine pneumonia. Res Microbiol 147:251–262[Medline]
  38. Hamilton JA, Piccoli DS, Leizer T, Butler DM, Croatto M, Royston AKM 1991 Transforming growth factor ß stimulates urokinase-type plasminogen activator and DNA synthesis, but not prostaglandin E2 production, in human synovial fibroblasts. Proc Natl Acad Sci USA 88:7180–7184[Abstract/Free Full Text]
  39. Flaumenhaft R, Abe M, Sato Y, Miyazono K, Heldin CH, Rifkin DB 1992 Role of latent TGFß1 binding protein in the activation of latent TGFß in co-cultures of endothelial and smooth muscle cells. J Cell Biol 120:995–1002[Abstract/Free Full Text]
  40. Lyons RM, Keski-Oja J, Moses HL 1988 Proteolytic activation of latent transforming growth factor-ß from fibroblast-conditioned media. J Cell Biol 106:1659–1665[Abstract/Free Full Text]
  41. Brauer PR, Yee JA 1993 Cranial neural crest cells synthesize and secrete a latent form of transforming growth factor ß that can be activated by neural crest cell proteolysis. Dev Biol 155:281–285[CrossRef][Medline]



This article has been cited by other articles:


Home page
 Reproductive SciencesHome page
Yan Wen, Y.-Y. Zhao, M. L. Polan, and B. Chen
Effect of Relaxin on TGF-{beta}1 Expression in Cultured Vaginal Fibroblasts From Women With Stress Urinary Incontinence
Reproductive Sciences, March 1, 2008; 15(3): 312 - 320.
[Abstract] [PDF]

Home page
 Stem CellsHome page
J. K. Mouw, J. T. Connelly, C. G. Wilson, K. E. Michael, and M. E. Levenston
Dynamic Compression Regulates the Expression and Synthesis of Chondrocyte-Specific Matrix Molecules in Bone Marrow Stromal Cells
Stem Cells, March 1, 2007; 25(3): 655 - 663.
[Abstract] [Full Text] [PDF]

Home page
 Endocr. Rev.Home page
K. Janssens, P. ten Dijke, S. Janssens, and W. Van Hul
Transforming Growth Factor-{beta}1 to the Bone
Endocr. Rev., October 1, 2005; 26(6): 743 - 774.
[Abstract] [Full Text] [PDF]

Home page
 JBJSHome page
R. T. Ballock and R. J. O'Keefe
The Biology of the Growth Plate
J. Bone Joint Surg. Am., March 31, 2003; 85(4): 715 - 726.
[Full Text] [PDF]

Home page
 Exp. Biol. Med.Home page
S. Tsutsumi, W. Tomisato, T. Hoshino, T. Tsuchiya, and T. Mizushima
Transforming Growth Factor-{beta}1 Is Responsible for Maturation-Dependent Spontaneous Apoptosis of Cultured Gastric Pit Cells
Experimental Biology and Medicine, June 1, 2002; 227(6): 402 - 411.
[Abstract] [Full Text] [PDF]

Home page
 DevelopmentHome page
S. Plisov, K Yoshino, L. Dove, K. Higinbotham, J. Rubin, and A. Perantoni
TGF beta 2, LIF and FGF2 cooperate to induce nephrogenesis
Development, January 4, 2001; 128(7): 1045 - 1057.
[Abstract] [PDF]

Home page
 Circ. Res.Home page
H. Chen, D. Li, T. Saldeen, and J. L. Mehta
Transforming Growth Factor-{beta}1 Modulates Oxidatively Modified LDL-Induced Expression of Adhesion Molecules: Role of LOX-1
Circ. Res., December 7, 2001; 89(12): 1155 - 1160.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Purchase Article
Right arrow View Shopping Cart
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 PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow