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Molecular and Cellular Biology, September 2007, p. 6309-6322, Vol. 27, No. 18
0270-7306/07/$08.00+0 doi:10.1128/MCB.00291-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
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Oral and Pharyngeal Cancer Branch,1 Craniofacial and Skeletal Diseases Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, 30 Convent Drive, Bethesda, Maryland 20892,3 Finsen Laboratory, Rigshospitalet, Copenhagen Biocenter, Ole Maalø Vej 5, DK-2200 Copenhagen N, Denmark2
Received 16 February 2007/ Returned for modification 10 April 2007/ Accepted 27 June 2007
| ABSTRACT |
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| INTRODUCTION |
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Extensive turnover of collagen takes place during a variety of physiological and pathological tissue-remodeling processes, including development, tissue repair, degenerative connective tissue diseases, and cancer. In this context, collagen turnover serves at least five different functions. It facilitates the physical expansion of a tissue (normal or aberrant), liberates latent growth factors embedded within the extracellular matrix, enables vascular development, subverts the proliferative restrictions imposed on cells by the extracellular matrix, and directly regulates cellular differentiation (5, 22, 30). Inhibition of extracellular matrix degradation, therefore, has long been recognized as an attractive target for therapeutic intervention in a variety of human diseases (1, 6).
Three molecular pathways are known for the turnover of collagen in physiological and pathological tissue-remodeling processes. The best-studied pathway involves a group of secreted or membrane-associated matrix metalloproteinases, the collagenases, that directly cleave collagens within the pericellular or extracellular environment (4, 44). A second, cathepsin K-mediated pathway is specific for osteoclast-mediated bone resorption and takes place in the acidic microenvironment that is created between the ruffled border of the osteoclast and the bone interface (16, 33). The third pathway is intracellular and involves the binding of collagen fibrils to specific cell surface receptors (ß1 integrins or urokinase plasminogen activator receptor-associated protein [uPARAP]/Endo180), followed by the cellular uptake, lysosomal delivery, and proteolytic degradation of the acid-denatured collagen by cathepsins (10, 11, 14, 29).
The functional relationship between the intracellular and pericellular collagen degradation pathways is poorly understood and controversial. The cellular uptake of collagen has been reported to be insensitive to metalloproteinase inhibitors (13). However, a recent study proposed that cleavage of collagen by membrane type 1-matrix metalloproteinase (MT1-MMP), the principal mesenchymal cell collagenase (19, 25, 45), was essential for the cellular uptake and lysosomal degradation of collagen by cultured gingival fibroblasts, based on the colocalization of MT1-MMP and actin during collagen phagocytosis (25). This finding contrasts with previous reports of a 40- to 60-fold increase in intracellular collagen in collagen-rich tissues of MT1-MMP-deficient (M–) mice (2, 19). The elucidation of the functional relationship between intracellular and pericellular pathways is important for evaluating the prospects for successfully using specific inhibitors of pericellular collagenases to prevent extracellular matrix destruction in a variety of diseases (32).
Here we asked whether intracellular and pericellular collagen turnover pathways could act in a complementary manner and, thus, could functionally compensate for each other during the remodeling of a collagen-rich tissue. We focused on bone development because of the high collagen content in cartilage and bone and the well-documented involvement of the pericellular collagenases (19, 23, 38, 45). Specifically, we examined the effect of single and combined null mutations in uPARAP/Endo180, a recently identified cell surface receptor that targets collagen for lysosomal degradation and is highly expressed in developing bone (3, 8, 10-12, 24, 36, 41, 43) (see below), and MT1-MMP, which has a well-established role in collagen turnover during bone formation (19-21, 34, 45). We show that combined deficits in these two collagen turnover pathways have strong synergistic effects on bone formation by directly impairing the survival and proliferation of cells embedded within the collagen-rich bone microenvironment. These data show that the intracellular and pericellular collagen degradation pathways defined by uPARAP/Endo180 and MT1-MMP, respectively, are complementary in vivo, and they identify a novel role for uPARAP/Endo180 in collagen turnover associated with bone formation.
| MATERIALS AND METHODS |
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Immunohistochemistry. Slides were deparaffinized in xylene (Mallinckrodt Baker, Phillipsburg, NJ), and the tissue was rehydrated in graded ethanol (EtOH) (100%, 95%, 70%, and H2O). Antigen retrieval was performed using proteinase K (Fisher Scientific, Pittsburgh, PA) treatment (5 mg/ml in 50 mM Tris-HCl [pH 8.0] [Quality Biologicals, Gaithersburg, MD] and 5 mM EDTA [Quality Biologicals]) at 37°C for 10 min and washing in tap water. Endogenous peroxidase activity was quenched by treatment of slides with 1% hydrogen peroxide (Fisher Scientific) in H2O for 15 min, which were then washed with tap water and transferred to 1x Tris-buffered saline (TBS) (Quality Biologicals) buffer with 0.5% Triton X-100 (Sigma-Aldrich, St. Louis, MO). Slides were incubated overnight at 4°C with purified rabbit polyclonal antibody (2.3 µg/ml) against uPARAP/Endo180, which was generated by immunization with the synthetic peptide N-IPRGVDVREPDIGRQGRLEWV-C as described previously (35), in 1x TBS with 0.25% bovine serum albumin (Sigma-Aldrich). The following day, slides were removed from the cold and incubated for an additional 30 min at room temperature. The slides were washed twice with 0.5% Tween 20 (Bio-Rad Laboratories, Hercules, CA) in 1x TBS. EnVision+ peroxidase anti-mouse antibody (DakoCytomation, Carpinteria, CA) was applied to the slides and incubated for 45 min at room temperature, followed by three washes with TBS-Tween 20 buffer. Dako Nova Red chromogenic substrate (DakoCytomation) was used to visualize expression of uPARAP/Endo180 according to the manufacturer's instructions and incubated for 7 min. The tissue was counterstained in Mayer's hematoxylin (Sigma-Aldrich), blued using 1x TBS, and rinsed in tap water. The slides were mounted with aqueous mounting medium (DakoCytomation) and photographed. For determination of proliferation, mice were injected with 200 µg bromodeoxyuridine (BrdU)/g body weight (Sigma-Aldrich) 2 h prior to euthanasia. Tissues were fixed in 4% paraformaldehyde (PFA) in 1x PBS overnight and embedded in paraffin. Tissue sections (5 µm) were dewaxed in xylene and rehydrated in graded EtOH. Staining for BrdU was performed using a BrdU staining kit (Zymed, South San Francisco, CA) according to the manufacturer's instructions. Quantification was performed by counting the number of positive chondrocytes as a fraction of the total number of chondrocytes in the growth plate, counting a minimum of 500 cells per sample.
Apoptosis analysis was performed by staining with the ApopTag Plus peroxidase in situ apoptosis detection kit (Chemicon International, Temecula, CA) according to the manufacturer's instructions. Slides were counterstained with contrast green (KPL, Gaithersburg, MD). Quantification was performed by counting the number of positive cells per high-power field, with a minimum of three high-power fields analyzed for each sample. Analysis was carried out by an investigator blinded to the genotype of mice.
Collagen type I telopeptides were detected with LF-67 rabbit polyclonal antiserum used at 1:500 (kindly provided by L. Fisher, NIDCR, NIH, Bethesda, MD). Serum from rabbits injected with adjuvant alone was used as a negative control at 1:500. Collagen type II was detected with monoclonal antibody II-II6B3 (1 µg/ml), developed by Thomas F. Linsenmayer and obtained from the Developmental Studies Hybridoma Bank, which was developed under the auspices of the NICHD and is maintained by the University of Iowa, Department of Biological Sciences (Iowa City, IA). Isotype control antibody (EBiosciences, San Diego, CA) (clone P3, immunoglobulin G1 kappa) was used at 1 µg/ml. Immunohistochemistry was performed using the Histomouse-Max kit (Zymed) according to the manufacturer's instructions, and visualization was with diaminobenzidine (Sigma-Aldrich).
In situ hybridization. In situ hybridization for collagen types I and II was performed exactly as described previously (20) by using sense and antisense murine collagen type I and II cDNA probes (28, 40). In brief, paraffin sections prepared as described above were dewaxed in xylene, rehydrated through a series of EtOH solutions of decreasing concentrations, treated with 5 µg/ml proteinase K, postfixed in 4% PFA in phosphate-buffered saline (PBS), acetylated in triethanolamine hydrochloride-acetic anhydride, washed in PBS, dehydrated, and air dried. The sections were incubated overnight at 50°C in hybridization buffer containing a single-stranded riboprobe that was radiolabeled with [33P]UTP (Perkin-Elmer, Waltham, MA) and complementary sense probes. Collagen type I mRNA was detected using a 2-kb rat cDNA probe, and collagen type II mRNA was detected using a 1.1-kb cDNA probe (both kindly provided by Y. Yamada, NIDCR, NIH, Bethesda, MD). After hybridization, the sections were washed extensively, dehydrated, and air dried. The slides were then dipped in photographic emulsion (Hypercoat LM-1; Amersham Biosciences, Piscataway, NJ) and were exposed for 3 to 5 days at 4°C. After exposure, the slides were developed, counterstained with Mayer's hematoxylin, and photographed using bright-field illumination.
Hematoxylin and eosin staining. Tissue was dewaxed in xylene and rehydrated in graded EtOH. Slides were stained with hematoxylin 2 (Richard-Allan Scientific, Kalamazoo, MI) for 3 min, followed by two incubations in tap water. Slides were then incubated in clarifier (Richard-Allan Scientific) for 1 min and rinsed twice in tap water. The slides were blued using bluing reagent (Richard-Allan Scientific) for 1 min and rinsed once in tap water. The slides were then incubated in 95% EtOH and then incubated in eosin (Richard-Allan Scientific) for 40 seconds. The slides were dehydrated through graded EtOH and xylene and mounted with Permount.
Primary osteoblast and chondrocyte cultures. Calvarial osteoblasts were isolated from 3-day-old uPARAP/Endo180-sufficient (U+) and uPARAP/Endo180-deficient (U–) littermates. The calvariae were dissected out and placed into alpha-minimal essential medium (Gibco, Carlsbad, CA) containing L-glutamine (Bio-Whittaker, East Rutherford, NJ), 5% fetal bovine serum (FBS) (Gemini Bioproducts, Woodland, CA), and penicillin-streptomycin (Gibco). Calvariae from three mice were pooled for each genotype. Soft tissue from the outside of the calvariae was carefully removed using fine forceps. The calvariae were then cut in half along the midline suture and were placed into 10 ml of 4 mM EDTA (Quality Biological) in a 50-ml glass Erlenmeyer flask in a shaking water bath at 37°C for 10 min. The EDTA was aspirated off, and a second incubation with fresh EDTA was performed. The EDTA was aspirated, and the calvariae were rinsed with 1x PBS. The PBS was aspirated, and the calvariae were incubated at 37°C for 10 min with a digestion solution of 200-U/ml collagenase 2 (Worthington Biochemicals, Lakewood, NJ) in 1x PBS with 1.6 µg/ml tosyl-lysyl-chloromethylketone (Calbiochem, La Jolla, CA). The digestion solution was aspirated, and fresh digestion solution was added as before. The second digestion solution was aspirated, and 10 ml of fresh digestion solution was added to the calvariae in the flask and incubated with shaking at 37°C for 15 min. The second digestion solution was passed through a 70-µm nylon cell strainer (BD Falcon, San Jose, CA) into a 50-ml conical tube containing 1x PBS with 5% FBS and placed on ice. This step was repeated twice. After the final digestion, the solution was removed, the calvariae were washed with 1x PBS and the wash was added to the previous digestion solutions. The digestion solution was then spun at 1,000 rpm for 5 min. Cells were seeded onto 10-cm dishes (BD Falcon) at a density of 2 x 104 cells/ml and grown in alpha-minimal essential medium, L-glutamine, 10% FBS, penicillin-streptomycin, and 55 µM 2-mercaptoethanol (Gibco). For chondrocyte isolation, the cartilaginous part of the rib cage was removed under sterile conditions and incubated at 37°C for 70 min in 1 ml of a solution of 3-mg/ml collagenase 2 in Dulbecco modified Eagle medium (DMEM) (Gibco) in a 50-ml conical tube. After incubation, the tube was vortexed, 1x PBS was added to a total volume of 35 ml, and the tube was shaken vigorously. The bones and cartilage were allowed to settle at the bottom of the tube, and the PBS was aspirated. This step was repeated twice. The cartilage and bone were then incubated overnight at 37°C in 3 ml 0.5-mg/ml collagenase 2 in DMEM with 10% FBS in a six-well plate (Corning Incorporated, Corning, NY). After the overnight digest, the cell solution was transferred to a 50-ml conical tube by filtering through a 70-µm nylon cell strainer. PBS was added, and the suspension was centrifuged at 1,000 rpm for 8 min. This wash step was repeated twice, and the final cell pellet was resuspended in 10% FBS in DMEM with penicillin-streptomycin and L-glutamine and cultured at 37°C at a cell density of 5 x 103 cells/cm2 for 5 days.
Collagen degradation assays. For analysis of pericellular collagenase activity, dishes were coated with a film of reconstituted dried fibrils of either type I collagen, obtained from rat tail tendons as described previously (42), or type II collagen (bovine articular cartilage; US Biological, Swampscott, MA). Type II collagen was dissolved in 0.05 M acetic acid at 4 mg/ml. The solution was brought to neutral pH at a final concentration of 3 mg/ml collagen, and the solution was transferred sequentially through the wells on a 24-well plate, coating the bottom of each well with approximately 30 µl of collagen solution. The collagen was gelled by incubating at 37°C for 1 h and allowed to dry in a laminar flow hood. The dried gel was washed with three changes of water and allowed to dry again. For both collagen types, 25,000 cells were seeded in a 25-µl droplet in the center of each well and allowed to attach for 5 h. The cells were washed and incubated in DMEM with 10–9 M interleukin-1ß (Peprotech, Rocky Hill, NJ) and 10–8 M tumor necrosis factor alpha (Peprotech) for 1 week. The cells were removed with 1% Triton X-100 (Sigma-Aldrich) in trypsin-EDTA (Invitrogen), and the collagen was stained with Coomassie blue (Sigma-Aldrich). The collagen film was photographed on an inverted Nikon Axiophot microscope with a 10x objective. For intracellular collagen degradation, acid-extracted rat tail tendon collagen was conjugated to fluorescein as described previously (18). The gel was then washed extensively with borate buffer at room temperature, the borate was washed out with water, and the labeled collagen was dissolved in 20 mM HCl at 4°C. Chondrocytes or osteoblasts isolated from uPARAP/Endo180-sufficient and -deficient mice were seeded onto glass coverslips coated with polylysine (Sigma-Aldrich) in 24-well tissue culture plates (Corning Incorporated) at a density of 1 x 104 cells/well in their appropriate growth media (see above). Cells were grown for 48 h and then washed in serum-free medium. E-64d (Calbiochem) was added to a final concentration of 20 µM and left for 1 h in serum-free conditions. The medium was changed to fresh serum-free medium with E-64d after 1 h, and collagen type I fibrils were added to a final concentration of 25 µg/ml and left for 8 h. LysoTracker-red (Molecular Probes, Eugene, OR) (50 nM final concentration) was added to cells and left for 90 min. Cells were washed with 1x PBS and fixed in 4% PFA, 1x PBS, and 0.05% Triton X-100 (Sigma-Aldrich) for 20 min in the dark. The cells were washed with PBS, incubated for 20 min at room temperature in 100 mM glycine (Invitrogen) in 1x PBS, and washed in PBS and then in water. Nuclei were stained with Hoechst (1/10,000) (Molecular Probes) for 5 min and washed with PBS. Coverslips were mounted onto glass slides using Vectashield (Vector Laboratories, Burlingame, CA) to prevent fading, according to the manufacturer's instructions. Confocal images were collected on a Leica TCS SP2 confocal system using a DM-RE-7 upright microscope and a 63x, 1.32-NA objective in the NIDCR Imaging Facility. Projection images from 0.15-µm Z-stacks were made using the Leica LCS software.
Cartilage and bone staining. Mice were euthanized. The skin was removed, the abdominal cavity was opened, and the pups were fixed in 95% EtOH for a minimum of 4 days. Any remaining skin and fat from mice were carefully dissected off of the carcasses, and the internal organs were removed. The remaining adipose tissue was dissolved by incubation in acetone (Mallinckrodt) for 48 h. Cartilage and bone were stained for 1 week in 1 volume 0.3% alcian blue in H2O, 1 volume 0.1% alizarin red in H2O (both Sigma-Aldrich), 1 volume glacial acetic, acid and 17 volumes of 75% EtOH. The samples were rinsed in water, and tissue was cleared in a solution of 1% KOH with frequent changes of solution, for approximately 1 to 2 weeks depending on age of mice. The KOH solution was removed, and samples were incubated with increasing concentrations of glycerol (10% to 50%) in water over several days. Samples were photographed using a Kodak DCS 760 camera.
X-ray analysis. Freshly euthanized pups were positioned on a Plexiglas sheet in a Faxitron MX-20 (X-ray Corporation, Wheeling, IL), and Kodak X-OMAT V film was used to image mice by exposure for 900 s at 30 kV at a magnification of either 1.5x, 3x, or 4x, depending on the age of the mice. Autoradiographs were scanned with an Epson expression 1680 at a setting of 900 dpi using Adobe Photoshop. Bone length was measured using the measurement tool in ImageJ by an investigator blinded to mouse genotype.
Determination of bone mineral density by computer-assisted tomography. Pups were sacrificed using a lethal intraperitoneal injection of xylazine, ketamine, and azepromazine. The skin was removed, and the body cavity was exposed. Pups were fixed and stored in 70% EtOH. To image the calvarium, the top of the skull was cut off, the brain was removed, and bone tissue was trimmed. To image femurs, the legs were cut off at the hip and femur, and surrounding soft tissue was carefully dissected. The day prior to imaging, tissue was transferred to 1x PBS. For imaging, tissues were wrapped in a sponge soaked in 1x PBS and placed in the specimen holder. Tissues were imaged at 14-µm resolution (80 kV, 80 µA) using an Explore Locus SP CT-scanner (General Electric Healthcare, Fairfield, CT). Image analysis and determination of bone mineral density and cortical bone thickness were performed using MicroView 2.1 software (General Electric Healthcare).
Determination of osteoblast activity. Pups were injected intraperitoneally with calcein (Sigma-Aldrich) at a dose of 25 mg/kg in 2% NaHCO3 at 4 days postbirth. The pups were sacrificed 2 days later, and heads were removed and fixed in 70% EtOH overnight at room temperature in the dark. The calvarium was dissected from the remainder of the skull, the brain was removed, and the calvarium was visualized under UV light. Digital images were obtained using the AlphaImager system (Alpha Innotech, San Leandro, CA). The digital images were then analyzed using ImageJ software. For presentation, the images were pseudocolored in ImageJ using the "fire/ice" setting. Quantitation was done using the "ROI manager and measurement" tool in ImageJ.
| RESULTS |
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We next performed calcein incorporation assays to determine if the synergistic reduction in ossification of bones in combined U–/M– pups was caused by a diminution of the rate of bone deposition by osteoblasts. U+/M+, U–, M–, and U–/M– littermate pups were injected with calcein, and the incorporation of this calcium fluorophor into calvarial bones was quantified after 48 h (Fig. 8). In accordance with the effect on bone mineral density, a single deficiency in either uPARAP/Endo180 or MT1-MMP had a more modest effect on osteoblast activity (4 and 20% reduction, respectively) than their combined deficiencies (50% reduction of osteoblast activity).
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| DISCUSSION |
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The findings presented here demonstrate that a molecular plasticity exists in the turnover of collagen associated with bone formation and likely in other physiological and pathological tissue-remodeling processes. The data provide direct experimental support for our previous speculation that the two pathways for the turnover of collagen (intracellular and pericellular) can act in a complementary manner during physiological and pathological tissue remodeling (11, 19). By inference, this implies that pharmacological prevention of the pathological destruction of collagen during the progression of degenerative diseases may be most efficiently achieved through the inhibition of both intracellular and pericellular pathways, although this also could increase the risk of undesired side effects. In this respect, it is important to note that intracellular inclusions of phagocytosed collagen have been observed in connective tissue cells associated with cancer, rheumatoid arthritis, emphysema, and periodontal disease, suggesting that the intracellular collagen degradation pathway is active in these human diseases (7, 8, 17, 27, 31, 37). Our data also lend support to a recent proposal that matrix metalloproteinase inhibitors may have been ineffective in blocking tumor extracellular matrix and basement membrane degradation, at least partially due to the unimpeded functionality of an intracellular, non-MMP-dependent pathway (32).
The specific reasons for the strong synergistic effect of combined uPARAP/Endo180 and MT1-MMP deficiency on cell proliferation and survival in developing bones remain to be determined. uPARAP/Endo180 expression is strictly confined to cells of the osteoblast/osteocyte lineage and chondrocytes of developing bone, and the receptor was not expressed by endothelial cells, either in the developing bones or at other anatomical sites (data not shown). The inability of resident cells with combined uPARAP/Endo180 and MT1-MMP deficiencies (chondrocyte/osteoblasts) to remodel collagen may lead to a failure to create a microenvironment that is conducive to their survival and growth, either due to physical constraints imposed by the collagen, failure to liberate latent growth factors and bioactive extracellular matrix fragments, or combinations thereof.
In summary, our studies demonstrate that both intracellular and pericellular collagen turnover pathways participate in physiological tissue remodeling, that both promote the survival and proliferation of cells located in a collagen-rich microenvironment, and that these two pathways can act in a complementary manner.
| ACKNOWLEDGMENTS |
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This work was supported by the NIDCR Intramural Research Program and by a grant from the Department of Defense (DAMD-17-02-0693) to Thomas H. Bugge. Lars H. Engelholm and Niels Behrendt were supported by EU contract LSHC-CT-2003-503297 and by grants from the Danish Cancer Society, the Danish Cancer Research Foundation, and the Danish Medical Research Council.
| FOOTNOTES |
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Published ahead of print on 9 July 2007. ![]()
Supplemental material for this article may be found at http://mcb.asm.org/. ![]()
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