Introduction Morphogenesis, tissue remodeling, and tissue repair all require the targeted remodeling of interstitial and basement membrane collagen to allow for organ growth, cell migration, and translation of contextual cues that are embedded within the extracellular matrix. Perturbed collagen homeostasis underlies a remarkable array of important human diseases, including fibrosis, that are attributable to excess interstitial deposition of collagen and degenerative diseases, such as osteoporosis, osteoarthritis, and rheumatoid arthritis, which are characterized by a loss of collagen from tissues. Finally, the ability of tumor cells to leave their site of origin and seed at novel locations is dependent on their ability to orchestrate the local degradation of collagen (Joyce and Pollard, 2009; Rowe and Weiss, 2009; Kessenbrock et al., 2010). Understanding the precise mechanisms underlying normal and abnormal collagen homeostasis is, therefore, of significant basic biological and clinical importance. The mechanisms of collagen turnover have been the subject of extensive studies over the past several decades, and two mechanistically different collagen catabolic pathways have been proposed: an extracellular pathway, in which collagen is degraded by membrane-bound and soluble proteases, and an endocytic pathway, in which collagen is first internalized by cells and then degraded by lysosomal cysteine proteases (Aimes and Quigley, 1995; Everts et al., 1996; Lauer-Fields et al., 2002; Hotary et al., 2006; Lee et al., 2006; Madsen et al., 2007; Krane and Inada, 2008; Lecaille et al., 2008; Messaritou et al., 2009; Segovia-Silvestre et al., 2009; Bugge and Behrendt, 2011). The existence of the latter pathway for collagen turnover in vivo is primarily supported by electron microscopy studies showing an abundance of collagen fibers in the endosomal and lysosomal compartments of mesenchymal cells presumed to be engaged in connective tissue degradation during cancer invasion, articular cartilage destruction in rheumatoid arthritis, bone resorption in periodontal disease, periodontal ligament turnover, and uterine involution (Cullen, 1972; Beertsen and Everts, 1977; Harris et al., 1977; Soames and Davies, 1977; Jurukova and Milenkov, 1981; Neurath, 1993; Lucattelli et al., 2003; Ryvnyak and Dulgieru, 2003). However, other studies have suggested that the observed intracellular collagen does not reflect the actual degradation of collagen derived from the extracellular matrix but rather consists of misfolded collagen that has been rerouted for immediate lysosomal degradation during the synthesis process (Bienkowski et al., 1978; Berg et al., 1980). Furthermore, considerable gaps in knowledge also exist as to the specific mechanisms by which collagen would be endocytosed and routed for lysosomal degradation. This particularly regards the requirement for collagenase cleavage before cellular uptake, the identity of cells endowed with the capacity to internalize and degrade collagen intracellularly, and the cellular receptors that mediate the initial collagen uptake. With respect to the latter, the collagen binding α1β1 and α2β1 integrins were the first proposed collagen internalization receptors, based on the observation that integrin-blocking antibodies impair the uptake of collagen-coated beads by cultured fibroblasts (Arora et al., 2000; Segal et al., 2001). A role of milk fat globule epidermal growth factor 8 in opsonizing collagen for specific uptake by macrophages was also recently proposed (Atabai et al., 2009). Furthermore, two members of the mannose receptor (MR) family of endocytic recycling receptors, the MR (MRC1 and CD206) and the urokinase plasminogen activator receptor–associated protein (uPARAP; endo180, MRC2, and CD280), were shown to aggressively internalize and target an assortment of collagen types for lysosomal degradation by cultured cells (Engelholm et al., 2003; Wienke et al., 2003; Martinez-Pomares et al., 2006; Madsen et al., 2011). Cognizant of these considerable mechanistic gaps in the knowledge of collagen turnover in vivo, we took advantage of recent advances in optical microscopy to develop a novel assay to directly visualize the fate of exogenous collagen introduced to the mouse dermis. By using this assay, we were able to characterize the cell types and molecules engaged in collagen degradation in vivo. Most importantly, we identify MR-dependent intracellular collagen degradation by M2-like macrophages as a novel and dominant collagen turnover pathway. Results Generation of an assay to visualize intracellular collagen degradation in vivo Acid-extracted rat tail tendon type I collagen fibrils self-assemble within minutes to form native, trypsin-resistant collagen fiber networks when incubated at physiological pH in vitro. Collagen networks formed this way have been used extensively as a model system to study cellular collagen turnover by cells explanted ex vivo (Birkedal-Hansen et al., 2003). To determine whether similar collagen networks could be reconstituted in vivo and then be visualized in situ, we fluorescently labeled acid-extracted rat tail tendon type collagen I with either Alexa Fluor 488, 594, or 647, which are highly photostable and pH-insensitive fluorescent dyes (see Materials and methods for details on this and other procedures; Panchuk-Voloshina et al., 1999). These fluorescently labeled collagen fibrils preserved their ability to rapidly form trypsin-resistant, collagenase-sensitive collagen networks ex vivo (Fig. 1 A and not depicted). Importantly, when neutralized from an acidic solution and then immediately injected into the subcutaneous space of mice, the fluorescently labeled collagen fibrils self-assembled to form similar collagen networks within the mouse dermis. These networks either formed around preexisting endogenous collagen/elastin bundles, or they formed de novo, independent of existing matrix scaffolds. In both cases, they easily could be visualized by either two-photon microscopy (not depicted) or confocal fluorescence microscopy of the exposed ventral dermis (Fig. 1, B–D). Fortuitously, the collagen injection procedure disrupted dermal homeostasis, as revealed by a substantial influx of cells into the injection field (Fig. 1 E) and by a large increase in the expression of genes associated with matrix turnover 24 h after injection, including genes encoding collagenases and gelatinases (Fig. 1 F). Furthermore, the introduced collagen appeared to be continuously degraded with an approximate half-life of 3.4 d, as determined by extraction of fluorescent material from injected skin (Fig. 1 G). Collectively, this suggested that the tissue damage inflicted by the subcutaneous collagen placement procedure creates a matrix catabolic microenvironment, thereby potentially making it possible to study the degradation of the exogenously introduced collagen networks in situ. By coinjecting the collagen with 10-kD dextran fluorescently labeled with Texas red, the lysosomal compartment of cells could be visualized (Fig. 1 H; Sandoval et al., 2004; Masedunskas and Weigert, 2008). Likewise, intravenous injection of the mice with fluorescent Hoechst 33342 dye 4–6 h before imaging allowed for easy visualization of cell nuclei (Fig. 1 H). A representative example of the appearance of an injection field reconstituted from serial z stacks is shown in Video 1. Figure 1. An assay to visualize intracellular collagen degradation in situ. (A–D) Microscopic appearance of collagen fibers formed after incubation of Alexa Fluor 647–labeled acid-extracted rat tail tendon type I collagen at neutral pH in vitro (A) or formed in vivo 24 h after injection into the dermis of live mice (B–D). Dermal-injected fluorescently labeled rat tail collagen fibrils form collagen fiber networks (white fibers in B) that may be associated with existing collagen/elastin scaffolds (blue fibers in B and white fibers in C, visualized by confocal microscopy) or may form ab initio (compare C and D). (E–G) Dermal collagen injection disrupts tissue homeostasis and generates a matrix catabolic environment. (E) Total cell counts in noninjected (left bar), vehicle-injected (middle bar), or collagen-injected (inj.; right bar) dermis, as determined by the counting of nuclei in four to six serial z stacks from the injection field. n = 3 for each treatment. *, P 90% of the dextran-uptaking cells also contained internalized collagen (see following paragraph). Importantly, these collagen-containing endocytic vesicles included lysosomes, as shown by the frequent colocalization of collagen and dextran (Fig. 2, B–D), demonstrating that internalized collagen is routed to the lysosomes. Imaging of dermis distant from the injection site revealed that the injected collagen remained localized to the area of injection, whereas trace amounts of dextran occasionally were detected (Fig. 2 E). Fluorescently labeled collagen injected in the absence of dextran displayed a similar vesicular distribution as observed when coinjected with the dextran, demonstrating that the dextran is not affecting the uptake of collagen (Fig. 2 F). The routing of internalized collagen to the lysosomes is consistent with previous data showing that fluorescently labeled collagen internalized by cultured cells is directed to the lysosomes (Kjøller et al., 2004). In previous studies, we have shown that inhibition of lysosomal cysteine proteases leads to increased levels of internalized collagen in both macrophages and fibroblasts, providing evidence that the internalized collagen is lysosomally degraded (Kjøller et al., 2004; Madsen et al., 2011). To directly examine the fate of internalized collagen, we allowed cultured fibroblast to internalize radiolabeled collagen and subsequently analyzed the physical state and the location of the collagen as a function of time by TCA precipitation of cell lysates and culture supernatants (Fig. 2 G). 5 h after collagen exposure, 59% of the radioactivity was in a cell-associated and already non-TCA–precipitable form, showing vigorous degradation of internalized collagen. After 9 h, most of this degraded collagen was released from cells and accumulating in the medium, with only 9% remaining cell associated in a TCA-precipitable form. A similar or even more efficient lysosomal collagen degradation process by professional phagocytes such as macrophages would be expected. Figure 2. Cellular uptake and lysosomal routing of collagen in vivo. (A) Representative image of mouse dermis 24 h after injection shows vigorous cellular uptake of collagen, as revealed by multiple cells presenting with fluorescently labeled collagen located within perinuclear endocytic vesicles. (B–D) High magnification of a single collagen-uptaking cell (box in A), illustrating the frequent colocalization of collagen and Texas red dextran in perinuclear vesicles (yellow; examples with arrows), indicative of lysosomal routing of endocytosed collagen. (E) Representative image of mouse dermis distant to the site of collagen and dextran injection. (F) Representative image of mouse dermis 24 h after injection of fluorescently labeled collagen. (G) Primary fibroblasts from wild-type or littermate uPARAP-deficient mice (hatched bars) were allowed to internalize 125I-labeled collagen for 5 h, after which the cells were trypsinized, washed, and reseeded. At 5 and 9 h after collagen exposure (0 and 4 h after reseeding), the cells (left) and the supernatant (right) were separated, and radioactivity in the TCA-precipitable fraction and the non-TCA–precipitable fraction was measured. Error bars indicate SDs. (H and I) Effect of systemic metalloproteinase inhibition on intracellular collagen degradation. (H) GM6001 treatment of mice inhibits MMP activity in vivo. Mice were treated with GM6001 or vehicle 24 and 0.5 h before i.p. injection with either the MMP-activated toxin PrAg-L1 or the nonactivatable control toxin PrAg-U7 (circles, vehicle + PrAg-L1 [n = 8]; squares, GM6001 + PrAg-L1 [n = 5]; triangles, vehicle + PrAg-U7 [n = 5]). *, P = 0.041, log-rank test. (I) Percentage of collagen-internalizing cells in the dermis of vehicle-treated or GM6001-treated mice. n = 3 for each group of collagen-injected mice treated with vehicle or with GM6001. *, P 95% of the GFP-positive cells internalizing collagen in both uPARAP-deficient and -sufficient mice (Fig. 8, A–C), suggesting that other receptors govern cellular collagen uptake in this context. Figure 8. Fibroblasts internalize collagen independently of uPARAP in vivo. (A) Percentage of collagen-internalizing GFP-positive and GFP-negative cells in the dermis of collagen- and dextran-injected Mrc2 +/+ (wild type; n = 4) and Mrc2 −/− (n = 6) mice. *, P 15 collagen-positive vesicles). To determine the level of collagen uptake by the high-level internalizing cells, a comparison to Cx3cr1-GFP–positive or Col1a1-GFP–positive cells was made. Based on z stacks from each of these transgenic mice, the high-level internalizing cells were identified as described in the previous paragraph, and an outline of the high-level internalizing cells and of the GFP-positive cells was drawn using ImageJ software (National Institutes of Health) to generate the regions of interest. The threshold for the fluorescent collagen was selected to allow maximum detection of collagen-positive vesicles while minimizing the contribution from extracellular fibrillar collagen. For each region of interest, the raw integrated density values were used for calculating the relative collagen uptake by high-level internalizing cells compared with GFP-positive cells in the same plane. From each z stack, only slices with both GFP-positive cells and high-level internalizing cells present were used for the analysis, and a mean fold difference between the two cell populations was calculated. The total relative fold difference in collagen uptake was calculated based on the analysis of 42 GFP-positive cells and 45 GFP-negative high-level internalizing cells in four Col1a1-GFP transgenic mice and 56 GFP-positive cells and 43 GFP-negative high-level internalizing cells in four Cx3cr1-GFP transgenic mice (in both cases, three to four z stacks per mouse were analyzed). To calculate the amount of internalized collagen by Cx3cr-GFP–positive cells, Col1a1-GFP–positive cells, and high-level internalizing GFP-negative cells, the fold differences in collagen internalization were multiplied to the fraction of collagen-internalizing Cx3cr1-GFP–positive cells (48.6%), Col1a1-GFP–positive cells (36.8%), or high-level internalizing Cx3cr1-GFP–negative, Col1a1-GFP–negative, cells (14.9%). Western blotting Approximately 1 cm2 of mouse skin was excised and homogenized in lysis buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1% Triton X-100, 0.5% Nonidet P-40, and 0.1% SDS with protease inhibitor cocktail [EMD Millipore]). Protein concentrations were determined using a bicinchoninic acid assay (Thermo Fisher Scientific), and 20 µg protein was used for Western blotting under nonreducing conditions. Western blotting against MR was performed using 2 µg/ml goat anti-MR antibody (clone MR5D3; AbD Serotec). Western blotting against uPARAP was performed with a solution containing 0.5 µg/ml monoclonal mouse anti-uPARAP antibody (clone 2h9F12; Sulek et al., 2007) mixed with alkaline phosphatase–conjugated secondary antibody. Anti-uPARAP antibody and the secondary antibody were mixed at a molar ratio of 3:1 and preincubated for 20 min to avoid the signal from endogenous mouse IgG. Protein lysates from M1 macrophages, M2 macrophages, and resting macrophages (generated as described in the section Ex vivo collagen internalization assay) were analyzed by Western blotting for the detection of MR and tubulin (loading control). Protein concentrations were determined, and equal protein amounts were loaded. The following primary antibodies were used: 2 µg/ml monoclonal mouse anti-MR antibody (BD) and monoclonal mouse antitubulin antibody (Santa Cruz Biotechnology, Inc.). Gene expression analysis by real-time PCR Gene expression was performed by real-time PCR in three tissue samples from collagen-injected or noninjected skin. In brief, 1 µg of total RNA was used for first-strand cDNA synthesis (Superscript III First-Strand Synthesis SuperMix; Invitrogen). The real-time PCR detection system (iCycler iQ; Bio-Rad Laboratories) and iQ SYBR Green Supermix (Bio-Rad Laboratories) were used for analysis. In brief, the reaction mixture contained 500 ng cDNA, housekeeping gene primers, and gene-specific forward and reverse primers for each gene at a final concentration of 1 µM. The real-time cycler conditions were as follows: PCR initial activation step at 95°C for 3 min, 40 cycles each of denaturing at 95°C for 15 s, and annealing/extension at 60°C for 1 min followed by a melting curve analysis of 55–95°C with 0.5°C increment, 5 s per step. A negative control without a template was included to assess the overall specificity of the reaction. The comparative cycle threshold (ΔΔCT) method was used to achieve the relative fold changes in gene expression between noninjected and collagen-injected skin. Primers are shown in Table S1. Whole-mount staining Whole-mount staining was performed 24 h after collagen and dextran injection. Skin was fixed for 4 h (MR and Fizz1 staining) or 16 h (F4/80 staining), and for the MR and Fizz1 staining, the skin was subsequently permeabilized by incubation in 1% Triton X-100 in PBS for 45 min. The skin was washed in PBS + 0.5% BSA + 0.2% Triton X-100 (PBS-BT), blocked in PBS-BT + 2% goat serum for ≥1 h, and incubated overnight at 4°C with primary antibody diluted in PBS-BT + 2% goat serum at the following ratios: rat anti-F4/80 (clone BM8, eBioscience) diluted 1:200, rat anti-MR (AbD Serotec) diluted 1:200, and rabbit anti-Fizz1 (PeproTech) diluted 1:400. Tissues were washed thoroughly in PBS-BT and incubated overnight at 4°C with Alexa Fluor 488–conjugated secondary antibodies diluted 1:500 in PBS-BT. After washing in PBS-BT, the tissue was incubated for 1 h with Hoechst diluted 1:10,000 in PBS-BT, and after subsequent washing, the skin was imaged as described in the “Confocal microscopy” section. Ex vivo collagen internalization assay Human monocytes were isolated from healthy donors as previously described (Jürgensen et al., 2011). In brief, mononuclear cells were isolated from whole blood using density gradient centrifugation (Lymphoprep; Axis-Shield). After this, monocytes were isolated using an automated cell isolator (RoboSep; StemCell Technologies, Inc.) and CD14 selection kit (EasySep Human Buffy Coat; StemCell Technologies, Inc.). The monocytes were differentiated in AIM-V media (Invitrogen) supplemented with the following specific cytokines: M1 macrophages, 5 ng/ml recombinant human GM-CSF (Sigma-Aldrich) for 7 d followed by 5 ng/ml recombinant human GM-CSF, 5 ng/ml recombinant human IFN-γ (R&D Systems), and 100 ng/ml lipopolysaccharide (Sigma-Aldrich) for 2 d; M2 macrophages, 20 ng/ml recombinant human M-CSF and 12.5 ng/ml recombinant human IL-4 (both obtained from R&D systems) for 9 d; and resting macrophages were generated by culturing for 9 d without cytokines. Collagen internalization assay was performed essentially as previously described (Jürgensen et al., 2011). In brief, macrophages were incubated in AIM-V medium supplemented with 1.5% BSA, 20 mM Hepes, and 10 µM of the cysteine protease inhibitor E64d (Merck Biosciences) for 30 min. 125I-labeled rat tail collagen I was added to a concentration of 150 ng/ml, and after a 4-h incubation at 37°C, the intracellular fraction was isolated and measured using a γ counter. M1 and M2 macrophages were evaluated for their ability to internalize collagen in the presence or absence of a function-blocking anti-MR antibody (10 µg/ml; R&D Systems). All samples were analyzed in triplicates. Fibroblasts from the skin of newborn Mrc2 −/− mice or wild-type littermates were isolated by trypsinization overnight at 4°C as previously described (Engelholm et al., 2003). One million cells were seeded in 6-well plates in duplicates in DMEM with 10% FCS and allowed to adhere overnight. The following day, the medium was replaced with fresh medium containing 133 ng/ml of 125I-labeled rat tail collagen I, and the cells were incubated for 5 h. Cells were washed three times in ice-cold PBS, trypsinized briefly, transferred to an Eppendorf tube, and centrifuged for 2 min at 200 g. Cells were then resuspended in DMEM with 10% FCS, reseeded in 6-well plates, and allowed to adhere for 0, 4, or 16 h. After this incubation, the culture supernatant was collected, and the cells were washed twice in PBS, briefly trypsinized, centrifuged at 200 g for 2 min, and resuspended in 1 ml of medium to neutralize the remaining trypsin. Cells were then centrifuged at 1,000 g for 1 min and lysed in lysis buffer (1% Triton X-100, 10 mM Tris, and 140 mM NaCl, pH 7.4). Subsequently, fresh medium was added to a total volume of 1 ml. Total protein content in either culture supernatant or lysates was precipitated by the addition of 90 µl of ice-cold 100% (wt/vol) TCA per milliliter and incubating the samples on ice for 10 min. Precipitated protein was separated from nonprecipitated material by centrifugation for 5 min at 12,000 g, and the amount of radioactivity in the fraction of precipitated protein (including intact nondegraded collagen) and nonprecipitated material (including degraded collagen) was determined using a γ counter. Online supplemental material Video 1 shows a reconstituted field of collagen formed in vivo. Table S1 shows the primers used for real-time PCR. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.201301081/DC1. Additional data are available in the JCB DataViewer at http://dx.doi.org/10.1083/jcb.201301081.dv. Supplementary Material Supplemental Material
