ABSTRACT
Membrane-type 1 matrix metalloproteinase (MT1-MMP, MMP-14), a transmembrane proteinase with a short cytoplasmic tail, is a major effector of extracellular matrix (ECM) remodeling. Genetic silencing of MT1-MMP in mouse (Mmp14-/-) and man causes dwarfism, osteopenia, generalized arthritis and lipodystrophy. These abnormalities have been ascribed to defective collagen turnover. We have previously shown non-proteolytic functions of MT1-MMP mediated by its cytoplasmic tail, where the unique tyrosine (Y573) controls intracellular signaling. Mutation of Y573 (Y573D) blocks signaling without affecting proteolytic activity. Here we report that a mouse with the MT1-MMP Y573D mutation (Mmp14Y573D/Y573D) shows abnormalities similar to those of Mmp14-/- mice. Skeletal stem cells (SSC) of Mmp14Y573D/Y573D mice show defective differentiation consistent with the mouse phenotype, which is rescued by wild-type SSC transplant. These results provide the first in vivo demonstration that MT1-MMP, an ECM-degrading proteinase, modulates bone, cartilage and fat homeostasis by controlling SSC differentiation through a mechanism independent of proteolysis.
INTRODUCTION
Membrane-type 1 matrix metalloproteinase (MT1-MMP, or MMP-14), the product of the gene MMP14, is a cell-membrane-bound proteinase with an extracellular catalytic site and a 20-amino acid cytoplasmic tail (Itoh, 2015; Sato et al., 1994). It degrades a variety of ECM components and is expressed by a wide array of normal and tumor cells. Notably, MT1-MMP is the only MMP whose genetic deficiency in the mouse results in severe phenotypes and early death. These features have implicated MT1-MMP as an important component of the proteolytic mechanisms of physiological and pathological processes including bone, cartilage and adipose tissue homeostasis, as well as tumor invasion, angiogenesis and metastasis. The analysis of the phenotype of mice genetically deficient in MT1-MMP (Mmp14-/-) has shown key roles of this proteinase in the postnatal development and growth of cartilage, bone and adipose tissue. Mmp14 deficiency in the mouse results in dwarfism, severe osteopenia, generalized arthritis and lipodystrophy, among other abnormalities (Chun et al., 2006; Chun & Inoue, 2014; Holmbeck et al., 1999; Zhou et al., 2000). In humans a mutation of MMP14 causes multicentric osteolysis and arthritis disease, or Winchester syndrome, which recapitulates much of the phenotype of the Mmp14-/- mouse (Evans et al., 2012). Conditional Mmp14 knockout in uncommitted skeletal stem cells (SSC, also referred to as mesenchymal stem cells), the common progenitors of osteoblasts, chondrocytes and adipocytes, recapitulates the skeletal phenotype of the global Mmp14 knockout mouse (Y. Tang et al., 2013). Conversely, unlike global Mmp14 deficiency conditional Mmp14 knockout in SSC results in thickening of articular cartilage and increased bone marrow (BM)-associated fat but decreased subcutaneous fat, which derives from distinct progenitor cells (Chun et al., 2006; Gupta et al., 2012; Tran et al., 2012). In light of the fundamental role of MT1-MMP in ECM degradation, it has been proposed that the phenotypes of Mmp14-/- mice result from defective collagen turnover (Chun et al., 2006; Chun & Inoue, 2014; Holmbeck et al., 1999; Zhou et al., 2000).
However, a number of in vitro studies have provided evidence for a variety of non-proteolytic roles of MT1-MMP. We have previously shown that the MT1-MMP cytoplasmic tail activates Ras-ERK1/2 and Akt signaling by a non-proteolytic mechanism that controls cell proliferation, migration and apoptosis in vitro, as well as tumor growth in vivo (D’Alessio et al., 2008; Valacca et al., 2015). Signaling is activated in a dose- and time-dependent manner by MT1-MMP binding of low nanomolar concentrations of tissue inhibitor of metalloproteinases-2 (TIMP-2), a physiological protein inhibitor of MT1-MMP. Signaling activation is also mediated by mutant TIMP-2 lacking MMP inhibitory activity (Ala+ TIMP-2) as well as by mutant MT1-MMP devoid of proteolytic activity (MT1-MMP E240A), showing that the signaling mechanism is proteolysis-independent. We also showed that MT1-MMP signaling requires the unique tyrosine (Y573) in the cytoplasmic tail, which is phosphorylated by Src and LIM1 kinases (Lagoutte et al., 2016; Nyalendo et al., 2007), and that Y573 substitution with aspartic acid (Y573D), a negatively charged amino acid like phosphotyrosine, abrogates MT1-MMP-mediated activation of Ras-ERK1/2 (D’Alessio et al., 2008). Other groups have subsequently found that Y573 controls a variety of signaling pathways and cell functions in vitro (Annabi et al., 2009; Belkaid et al., 2007; Gonzalo et al., 2010; Proulx-Bonneau et al., 2011; Sakamoto & Seiki, 2009; Sina et al., 2010). Y573 is required for activation of the small GTPase Rac1 (Gonzalo et al., 2010), controls macrophage migration and infiltration at sites of inflammation (Gonzalo et al., 2010; Sakamoto & Seiki, 2009), as well as MT1-MMP interaction with Src and focal adhesion kinase (FAK) (Y. Wang & McNiven, 2012).
Based on these observations, we generated a mutant mouse with the Y573D substitution in the cytoplasmic tail of MT1-MMP (MT1-MMP Y573D). Here we report that this mouse shows abnormalities in postnatal bone, cartilage and adipose tissue development and growth that partly recapitulate the phenotype of the MMP14-/- mouse. These phenotypes derive from dysregulation of BM SSC differentiation, and are rescued by wild-type (wt) BM transplant.
RESULTS
Generation and macroscopic characterization of MT1-MMP Y573D mice
Mice heterozygous (MmpY573D/wt) or homozygous (MmpY573D/Y573D) for the MT1-MMP Y573D mutation, generated as described in Materials and Methods (Fig. 1 A and B), were macroscopically indistinguishable from Mmp14wt/wt mice at birth. However, after the first 2 months Mmp14Y573D/wt and Mmp14Y573D/Y573D mice showed a slightly lower growth rate than their Mmp14wt/wt littermates (Fig. 1 C). In mice older than 3 months the weight of Mmp14Y573D/Y573D male and female mice was 15 % lower than that of age- and sex-matched Mmp14wt/wt littermates (Fig. 1 D). Both male and female Mmp14Y573D/Y573D mice were fertile, bred and lived up to > 2 years of age similarly to Mmp14wt/wt mice.
The Y573D mutation does not affect the proteolytic activity of MT1-MMP
A major consequence of the genetic deficiency of MT1-MMP collagenolytic activity in Mmp14-/- mice is the progressive development of fibrosis of soft tissues, including fibrosis of the dermis and hair follicles, with subsequent hair loss (Holmbeck et al, 1999). Mmp14Y573D/Y573D mice up to 2 years of age showed no hair loss, and histological analysis of their skin showed no fibrosis of the dermis or hair follicles (Fig. 2 A), suggesting that the collagenolytic activity of MT1-MMP Y573D is comparable to that of wt MT1-MMP. Consistent with this finding, primary fibroblasts from 3-weeks old Mmp14Y573D/Y573D and Mmp14wt/wt mice showed a similar capacity to degrade collagen and activate pro-MMP-2 in vitro (Fig. 2 B – D), as well as comparable levels of MT1-MMP (Fig. 2 F).
In addition to degrading collagen and other ECM components, MT1-MMP cleaves a variety of cell membrane proteins including the collagen receptors CD44 and discoidin domain receptor 1 (DDR1), as well as the metalloproteinase ADAM9 (Chan et al., 2012; Fu et al., 2013; Kajita et al., 2001). We therefore characterized these proteins by Western blotting analysis of tissues and cells from Mmp14Y573D/Y573D and Mmp14wt/wt mice. Fat, cartilage and bone (Fig. 2 E), the tissues affected by the MT1-MMP Y573D (as described under the following subheading), as well as other tissues (Figure S1A), showed minor variations in the levels of MT1-MMP expression in mice of the two genotypes. In fat MT1-MMP was expressed almost exclusively in the proenzyme form (62 kDa), whereas in cartilage and bone the 42-44 kDa autocatalytic degradation product was prevalent, indicating high levels of proteolytic activity (Rozanov et al., 2001). We could not detect CD44 in the fat of our mice, and qPCR analysis showed extremely low levels of CD44 mRNA in both Mmp14Y573D/Y573D and Mmp14wt/wt mice (Ct values 3.21 and 3,79, respectively). However, the 80 kDa native form of CD44was present in cartilage and bone in comparable amounts and no degradation products were detected in mice of either genotype. DDR1 could not be detected in the bone of Mmp14Y573D/Y573D or Mmp14wt/wt mice, a result consistent with qPCR analysis (Ct values 31.89 and 31.76, respectively). Conversely, DDR1 was detected in fat and cartilage extracts exclusively as a ∼ 62 - 65 kDa form consistent with the degradation product resulting from MT1-MMP proteolysis (Fu et al., 2013). Comparable levels of ∼ 62 - 65 kDa DDR1 were present in cartilage, whereas higher levels were expressed in Mmp14Y573D/Y573D than in Mmp14wt/wt fat. No other degradation products were detected, showing that wt MT1-MMP and MT1-MMP Y573D have comparable capacity to cleave CD44 and DDR1 in vivo.
To corroborate these findings we analyzed CD44, DDR1, ADAM9 and FGFR2 in primary fibroblasts from adult Mmp14Y573D/Y573D and Mmp14wt/wt (Fig. 2 F). By Western blotting analysis of cell extracts we could not positively identify bands consistent with native or cleaved CD44 (Figure S1B). However, analysis of cell-conditioned medium showed comparable levels of the ∼ 70 kDa degradation product derived from MT1-MMP cleavage in cells of the two genotypes (Kajita et al., 2001). ADAM9 degrades FGFR2, and MT1-MMP cleavage of ADAM9 prevents FGFR2 degradation (Chan, et al., 2012). By Western blotting (Fig. 2 F) extracts of Mmp14Y573D/Y573D and Mmp14wt/wt cells showed comparable levels of native, ∼ 90 kDa ADAM9 and lower-Mr bands with Mrs ranging ∼ 40 – 60 consistent with ADAM-9 degradation products generated by MT1-MMP (Chan, et al., 2012), as well as similar levels of native FGFR2. Similarly, like in the tissues, comparable levels of the ∼ 62 - 65 kDa DDR1 cleavage product were detected in cells from both genotypes.
In addition, in agreement with our previous findings (D’Alessio et al., 2008; Valacca et al., 2015), the primary fibroblasts from Mmp14Y573D/Y573D mice showed strongly reduced Erk1/2 and Akt activation in response to exogenous TIMP-2 relative to Mmp14wt/wt cells (Fig. 2 G). Thus, these results showed that the Y573D mutation does not alter MT1-MMP expression or its capacity to cleave interstitial collagen or membrane-bound protein substrates; however, the mutation abrogates MT1-MMP-mediated activation of intracellular signaling.
MT1-MMP Y573D mice show structural and gene expression abnormalities in bone, articular cartilage and adipose tissue
A striking phenotype of Mmp14-/- mice is the dramatically decreased length of long bones, with severe osteopenia and arthritis (Holmbeck et al., 1999; Zhou et al., 2000). At 5 months of age the femurs of Mmp14Y573D/Y573D mice were only ∼ 5% shorter than those of Mmp14wt/wt mice (14.81 ± 0.1519 mm vs. 15.61 ± 0.0619 mm; n=10; p = 0.0001). However, cortical bone thickness at the femur mid-diaphysis was markedly decreased (20-25%) in Mmp14Y573D/Y573D vs. Mmp14wt/wt mice (Fig. 3 A), a finding consistent with the osteopenia of the Mmp14-/- mouse (Holmbeck et al., 1999; Zhou et al., 2000). Conversely, unlike Mmp14-/- mice, which have reduced trabecular bone, and in the homozygous but not in the heterozygous state (Holmbeck et al., 1999; Zhou et al., 2000), femurs and tibias showed significantly increased (40-50%) trabecular bone in both Mmp14Y573D/Y573D and Mmp14Y573D/wt mice relative to age- and sex-matched Mmp14wt/wt mice (Fig. 3 B - D). This effect was accompanied by increased TRAP-positive osteoclasts (Figure S2), indicating enhanced bone remodeling, a feature also observed in Mmp14-/- mice (Holmbeck et al., 1999).
Gene expression profiling of bone from Mmp14Y573D/Y573D mice (Fig. 3 E) showed 76 genes significantly (p ≤ 0.05) up- or downregulated relative to Mmp14wt/wt littermates. A subset of 17 of these transcripts were upregulated 2-fold or more, and 26 were downregulated 2-fold or more. Consistent with the morphometric analyses, gene ontology analysis showed highly significant enrichment for biological processes related to osteoblast differentiation, bone remodeling, ossification and bone growth (Fig. 3 F).
The knee joints of 2-month old mice showed several abnormalities. Both Mmp14Y573D/wt and Mmp14Y573D/Y573D mice displayed marked thinning of articular cartilage (Fig. 4 A top panels, and B) with loss of proteoglycans (Fig. 4 C and D), clustering and cloning of chondrocytes (Fig. 4 A lower panels), classic histologic features of the articular cartilage degeneration associated with human osteoarthritis (OA) and surgical models of OA in mice. The knee cartilage of 2-year old mice also showed fissures, chondrocyte clustering and cloning, abnormalities typical of ageing-associated cartilage degeneration that were not observed in age- and sex-matched Mmp14wt/wt mice (Fig. 4 E).
RNA-Seq analysis of the transcriptome of cartilage from Mmp14Y573D/Y573D mice (Fig. 5 A) showed significant dysregulation of the expression of 1,549 genes relative to Mmp14wt/wt mice (p ≤ 0.05). Of these genes, 694 were upregulated 2-fold or more, and 92 were downregulated 2-fold or more. Gene ontology analysis (Fig. 5 B) showed highly significant enrichment for biological processes related to extracellular matrix homeostasis, cartilage development and chondrocyte differentiation. As these biological processes are strongly dysregulated in human OA, we analyzed the transcriptome of Mmp14Y573D/Y573D cartilage for expression of genes involved in human OA (Fig. 5 C). Gene expression profiling of human OA cartilage has revealed 1,423 genes significantly (p ≤ 0.05) up- or downregulated relative to normal cartilage, 111 of which are strongly up- or downregulated (≥ 2-fold or ≤ 2-fold) (Geyer et al., 2009; Karlsson et al., 2010). We found that 48 of these 111 OA-associated genes are also strongly regulated in Mmp14Y573D/Y573D mouse cartilage, including all the genes for collagens and other ECM proteins upregulated in human OA, ECM-degrading proteinases, as well as genes involved in cell metabolism (Figure 5 D). Thus, consistent with its histological features, Mmp14Y573D/Y573D joint cartilage showed significant gene expression similarity to human OA.
Sections of the long bones of adult Mmp14Y573D/wt and Mmp14Y573D/Y573D mice also showed marked decrease in BM-associated fat relative to Mmp14wt/wt littermates (Fig. 6 A and B). A similar reduction was apparent in all other fat pads, as evidenced by ∼ 50% decrease in body adiposity by DEXA analysis and in the gonadal fat of Mmp14Y573D/Y573D mice relative to age- and sex-matched Mmp14wt/wt littermates (Fig. 6 C and D). This effect was observed in mice of both sexes and ages ranging 3 months to 2 years (Figures S3 and S4). Histological analysis of abdominal and subcutaneous WAT from Mmp14Y573D/Y573D mice revealed marked decrease in the size of adipocytes (Fig. 7 A, C and S5). These findings are consistent with the lipodystrophy of Mmp14-/- mice (Chun et al., 2006; Chun et al., 2010). In contrast, the brown adipose tissue (BAT) of Mmp14Y573D/Y573D mice showed pronounced adipocyte hypertrophy, with decreased expression of the characteristic marker of BAT, uncoupling protein-1 (UCP-1; Fig. 7 B and S6).
The transcriptome of abdominal WAT from Mmp14Y573D/Y573D mice (Fig. 7 D) showed significant (p ≤ 0.05) up- or downregulation of the expression of 151 genes, relative to Mmp14wt/wt mice. Gene ontology analysis (Fig. 7 E) identified highly significant enrichment for biological processes including control of small molecule synthesis, lipid and fatty acid metabolism, response to corticosteroids, and extracellular matrix homeostasis. Some of these pathways are also overrepresented in Mmp14+/- mice on a high-fat diet (Chun et al., 2010). Moreover, similar to Mmp14+/- mice, Mmp14Y573D/Y573D mice were protected from body weight gain induced by high-fat diet (Figure S7).
In addition to reduced WAT, Mmp14Y573D/Y573D mice showed strongly decreased fasting levels of plasma insulin relative to age- and sex-matched Mmp14wt/wt littermates, with normoglycemia and normal food consumption (Fig. 8 and S8). Mmp14Y573D/Y573D and Mmp14wt/wt mice also had comparable levels of ACTH, which controls adipose tissue metabolism, and leptin, a hormone secreted predominantly by adipose tissue. Conversely, Mmp14Y573D/Y573D mice had extremely low serum levels of the inflammatory cytokines interleukin-6 (IL-6), monocyte chemoattractant protein 1/ chemokine (C-C motif) ligand 2 (MCP-1/CCL2), and IL-10 relative to age- and sex-matched Mmp14wt/wt mice (Fig. 8).
MT1-MMP Y573D expression alters skeletal stem cell proliferation and apoptosis, and skews differentiation from chondro- and adipogenesis towards osteogenesis
The observation that the phenotype of Mmp14Y573D/Y573D mice involves abnormalities of bone, cartilage and adipose tissue indicated that the MT1-MMP Y573D mutation might affect the differentiation of SSC, the common progenitor cells of these tissues. Therefore, we isolated SSC from the BM of Mmp14wt/wt and Mmp14Y573D/Y573D littermates, and characterized them in vitro (Fig. 9). Mmp14Y573D/Y573D SSC contained fewer colony-forming units-fibroblasts (CFU-F) than Mmp14wt/wt SSC (1.5×10-6 vs. 4.5×10-6, respectively), formed much smaller colonies (Fig. 9 A), and showed remarkably lower proliferation rate and higher apoptosis index (Fig. 9 B and C). RNA-Seq analysis (Fig. 9D) showed 486 genes significantly dysregulated (p ≤ 0.05) in Mmp14Y573D/Y573D vs. Mmp14wt/wt SSC. Gene ontology analysis (Fig. 9 E) revealed highly significant enrichment of genes involved in DNA synthesis, cell cycle regulation and response to DNA damage, indicating dysregulation of cell proliferation and survival.
We then induced BM-derived SSC from Mmp14Y573D/Y573D and Mmp14wt/wt littermates to differentiate in vitro into the osteoblast, chondrocyte and adipocyte lineages. qPCR analysis of the expression of lineage-specific markers showed dramatic increase in osteogenesis (Fig. 9 F, top panel), and marked decrease in chondrocyte and adipocyte differentiation (Fig. 9 F, middle and bottom panels, respectively) in Mmp14Y573D/Y573D vs. Mmp14wt/wt SSC. The expression levels of wt MT1-MMP and MT1-MMP Y573D did not change during osteoblast differentiation, and on day 17 comparable levels of MT1-MMP mRNA were expressed by Mmp14wt/wt and Mmp14Y573D/Y573D SSC (Fig. 9 F; top panel). Conversely, in agreement with previous reports (Y. Tang et al., 2013), MT1-MMP expression decreased by ∼ 80% during chondrocyte differentiation in both Mmp14wt/wt and Mmp14Y573D/Y573D SSC, and on day 17 Mmp14Y573D/Y573D cells expressed significantly lower MT1-MMP levels than Mmp14wt/wt SSC (Fig. 9 F, middle panel).
To confirm that the differences in BM-SSC differentiation between Mmp14wt/wt and Mmp14Y573D/Y573D mice are mediated by the MT1-MMP Y573D mutation, we transfected wt MT1-MMP or MT1-MMP Y573D cDNA into mouse C3H10T1/2 cells, which are functionally similar to SSC (Q. Q. Tang et al., 2004), and analyzed their capacity to differentiate into osteoblasts, chondrocytes and adipocytes (Fig. 9 G) by lineage-specific staining. Consistent with the results obtained with primary BM-derived SSC, C3H10T1/2 cells transfected with MT1-MMP Y573D showed markedly increased osteogenesis, and decreased chondro- and adipogenesis relative to wt MT1-MMP transfectants (Fig. 9 G). Therefore, in both BM-derived SSC and C3H10T1/2 cells MT1-MMP Y573D expression skewed differentiation towards the osteogenic lineage.
The bone and fat phenotypes of Mmp14Y573D/Y573D mice are rescued by wt BM transplant
These results indicated that the bone, cartilage and fat phenotypes of Mmp14Y573D/Y573D mice result from a defect in SSC. Therefore, we hypothesized that the phenotypes of these mice could be rescued by transplantation of Mmp14wt/wt BM; and, vice versa, transplantation of Mmp14Y573D/Y573D BM could transfer the phenotype to Mmp14wt/wt mice. We therefore transplanted 3- and 5-month old mice with BM from mice of the same age and the opposite genotype or, as controls, from mice of the same age and genotype, and analyzed their phenotypes two months later.
We characterized the trabecular and cortical bone of the femurs of the transplanted mice by microCT analysis (Fig. 10). The results showed that transplantation of Mmp14wt/wt BM had no effect on the cortical bone of Mmp14Y573D/Y573D mice transplanted at 3 months or 5 months of age (Fig. 10 A and S9); however, it completely rescued their trabecular bone phenotype (Fig. 10 B and S9). Conversely, transplantation of Mmp14Y573D/Y573D BM did not transfer the trabecular or cortical bone phenotype to wt mice.
We also found that BM transplant induced no significant changes in articular cartilage thickness relative to the original phenotype of the transplant recipients in this short-term study (data not shown).
Similarly to the bone phenotype, Mmp14wt/wt BM transplant completely rescued the decrease in fat mass of Mmp14Y573D/Y573D mice, but Mmp14Y573D/Y573D BM did not transfer the adipose tissue phenotype to Mmp14wt/wt mice (Fig. 11 A). However, consistent with the existence of BM-derived adipocyte precursors able to colonize peripheral WAT and BAT (Crossno et al., 2006), histological analysis of WAT and BAT from transplanted mice (Fig. 11 B and C) showed mixtures of normal and hypotrophic white adipocytes in both Mmp14wt/wt mice transplanted with Mmp14Y573D/Y573D BM and Mmp14Y573D/Y573D mice transplanted with Mmp14wt/wt BM. Conversely, BM recipients acquired the BAT phenotype of donor mice (Fig. 11 C).
The analysis of BM engraftment, described in Materials and Methods, showed virtually complete replacement of the recipient’s BM with the donors’ BM (Fig. 11 and S10). The circulating blood cells of Mmp14Y573D/Y573D mice displayed no significant abnormalities and numbers comparable to those of Mmp14wt/wt mice, showing that the MT1-MMP Y573D mutation does not affect hematopoietic cells. Therefore, these results showed that the bone and fat phenotype of transplanted Mmp14Y573D/Y573D mice resulted from the transfer of SSC, and not from other BM cells.
DISCUSSION
MT1-MMP plays an important role in the postnatal development of a variety of tissues including bone, cartilage and fat. The striking abnormalities of Mmp14-/- mice – dwarfism, severe osteopenia, generalized arthritis, fibrosis and lipodystrophy - have been ascribed to impaired collagen turnover, which also plays a fundamental role in SSC differentiation and affects adipocyte development (Chun et al., 2006; Chun & Inoue, 2014; Holmbeck et al., 1999; Zhou et al., 2000). The data presented in this paper show that, in addition to its proteolytic activity, MT1-MMP contributes to bone, cartilage and adipose tissue homeostasis through a proteolysis-independent mechanism mediated by its cytoplasmic domain. This conclusion is based on the following observations.
We have previously shown that MT1-MMP activates ERK1/2 and Akt signaling upon binding of physiological concentrations of TIMP-2 (D’Alessio et al., 2008; Valacca et al., 2015). Signaling is activated by mutant MT1-MMP devoid of proteolytic activity (MT1-MMP E240A) or TIMP-2 lacking MMP inhibitory activity (Ala+ TIMP-2), and is blocked by the Y573D substitution in the MT1-MMP cytoplasmic tail (D’Alessio et al., 2008; Fig. 2 G). Here we showed that the Y573 mutation does not significantly alter the collagenolytic activity of MT1-MMP, or its cleavage of cell membrane proteins in vitro and in vivo (Fig. 2 A - F). It should also be noted that changes in the levels of membrane proteins cleaved by MT1-MMP result in mouse phenotypes different from those of Mmp14Y573D/Y573 mice. CD44-/- or ADAM9-/-mice have no phenotype (Protin et al., 1999; Weskamp et al., 2002); whereas the genetic deficiency of FGFR2 or Notch1 results in embryonic lethality, and Notch1 haploinsufficiency causes virtually no phenotype (Conlon et al., 1995; Xu et al., 1998). DDR1 deficiency in the mouse results in female infertility and defective lactaction, which we did not observe in Mmp14-/- mice; conversely, DDR1 haploinsufficiency causes no phenotype (Vogel et al., 2001). Therefore, these findings show that MT1-MMP activation of ERK1/2 and Akt signaling is independent of its proteolytic activity and mediated by its cytoplasmic domain.
To investigate the physiological significance of our in vitro findings, we generated a mouse in which MT1-MMP activation of ERK1/2 and Akt is abrogated by the Y573D mutation. The data presented in this paper show that mice bearing this mutation present phenotypes that recapitulate those caused by Mmp14 deficiency. The phenotypes of Mmp14Y573D/Y573 mice appear to be milder than those of Mmp14-/- mice, indicating that both the proteolytic and signaling functions of MT1-MMP are required for normal postnatal development and homeostasis. However, while comparing the phenotypes of Mmp14Y573D/Y573 and Mmp14-/- mice provides information about the relative contribution of the proteolytic and non-proteolytic functions of MT1-MMP to postnatal development and tissue homeostasis, significant limitations must be considered. Mmp14-/- mice die within 2 months of age (Holmbeck & al., 1999), whereas Mmp14Y573D/Y573 mice have a normal life span and their phenotypes become apparent in animals older than 2 months. The global deficiency of Mmp14 causes severe runting and wasting that ultimately lead to death. It is possible that the dramatic abnormalities of some tissues including cartilage and fat result from indirect effects. For instance, the generalized inflammatory state of Mmp14-/- mice (Shimizu-Hirota et al., 2012) may be the cause of, or contribute to their severe arthritis. Indeed, conditional Mmp14 knockout in uncommitted SSC results in increased thickness of the articular cartilage in 3-month old mice (Y. Tang et al., 2013), in contrast with the reduced thickness and degeneration of Mmp14-/- and Mmp14Y573D/Y573D cartilage (Holmbeck & al., 1999).
Furthermore, the relative contribution of MT1-MMP proteolytic and signaling functions to postnatal development and homeostasis may vary at different stages of postnatal life. The lethality of Mmp14 deficiency limits our understanding of the role of MT1-MMP to relatively early stages of postnatal development; in contrast, the normal life span of Mmp14Y573D/Y573 mice affords studying the role of MT1-MMP in adult animals. Our finding that the phenotype of the Mmp14Y573D/Y573 mouse becomes apparent in adult mice indicates that the proteolytic and the signaling function of MT1-MMP have predominant roles at different stages of postnatal development. The severe phenotype and early mortality of the Mmp14-/- mouse shows that MT1-MMP proteolytic activity has a fundamental role at early stages. Conversely, silencing MT1-MMP signaling is compatible with normal development but results in altered tissue homeostasis in the adult animal.
Our analysis of the articular cartilage of Mmp14Y573D/Y573D mice showed signs of tissue degeneration similar to, but milder than that of Mmp14-/- mice. However, whereas Mmp14-/- mice have severe, acute arthritis (Holmbeck & al., 1999), the articular cartilage of Mmp14Y573D/Y573D mice shows histological signs and gene expression profile comparable to human OA, a chronic degenerative condition. The striking similarity of the gene expression profile of the articular cartilage of Mmp14Y573D/Y573D mice to that of human OA raises an interesting point about the role of MT1-MMP in articular cartilage homeostasis and the pathogenesis of OA. MT1-MMP expression decreases during chondrocyte differentiation, and in differentiated chondrocytes is ∼ 80% lower than in undifferentiated SSC (Y. Tang et al., 2013). Consistent with this finding, Mmp14 deficiency in uncommitted SSC results in increased differentiation into chondrocytes and thickening of articular cartilage (Y. Tang et al., 2013), suggesting that MT1-MMP expression contrasts normal chondrocyte development and articular cartilage homeostasis. Indeed, MT1-MMP is upregulated in OA cartilage relative to normal cartilage (Dreier et al., 2004; Kevorkian et al., 2004; Tchetina et al., 2005), indicating that MT1-MMP expression must be strictly controlled for normal cartilage homeostasis. In Mmp14Y573D/Y573D SSC-derived chondrocyte, expression of MT1-MMP is lower than in Mmp14wt/wt mice (Fig. 9 F); however, their cartilage is thinner and presents signs of degeneration, showing that altering MT1-MMP signaling has a pathological effect even in the presence of low levels of MT1-MMP proteolytic activity. Thus, MT1-MMP signaling is required for articular cartilage homeostasis.
Our phenotypic analysis of Mmp14Y573D/Y573D mice showed significant abnormalities in WAT, consistent with the phenotype of Mmp14-/- mice and of mice with conditional Mmp14 knockout in uncommitted SSC (Y. Tang et al., 2013). However, some differences are noteworthy. In contrast to WAT hypotrophy, we surprisingly found BAT hypertrophy in Mmp14Y573D/Y573 mice, a finding at variance with the normal BAT of Mmp14-/- mice (Chun et al., 2006). Similarly, BM-associated WAT is decreased in Mmp14Y573D/Y573 mice but increased in mice with conditional knockout of Mmp14 in uncommitted SSC (Y. Tang et al., 2013) (unfortunately, the severe wasting and early death of Mmp14-/- mice preclude a reliable assessment of BM-associated fat, which typically develops with aging). However, while the SSC of conditional Mmp14-/- mice show increased adipocyte differentiation in vitro (Y. Tang et al., 2013), the SSC of Mmp14Y573D/Y573D mice show decreased adipocyte differentiation. MT1-MMP is a fundamental effector of adipocyte growth through collagen degradation, a process required for adipocyte increase in size (Chun et al., 2006; Chun & Inoue, 2014). However, Mmp14-/- mice have multiple, severe developmental and metabolic defects that can affect adipose tissue development indirectly. Our data show that in vivo MT1-MMP contributes to both WAT and BAT homeostasis by a proteolysis-independent mechanism mediated by its cytoplasmic tail. Several studies have shown the involvement of MT1-MMP in adipose tissue homeostasis (Chun et al., 2006; Feinberg et al., 2016; Fenech, Gavrilovic, Malcolm, et al., 2015; Fenech, Gavrilovic, & Turner, 2015), and genetic associations between MT1-MMP and obesity in humans have been reported (Chun et al., 2010). MT1-MMP has also been proposed to control metabolic balance (Mori et al., 2016), a function that could explain the WAT and BAT abnormalities of MT1-MMP Y573D mice. Understanding the proteolysis-independent mechanism of MT1-MMP control of adipose tissue homeostasis can therefore have significant clinical and pharmacological implications.
The cortical bone of Mmp14Y573D/Y573 mice shows a significant decrease in thickness, a phenotype similar to – if milder than - that caused by Mmp14 deficiency. In contrast, the increased trabecular bone of Mmp14Y573D/Y573 mice contrasts with the severe osteopenia of Mmp14-/- mice and mice with Mmp14 knockout in uncommitted SSC (Holmbeck & al., 1999; Y. Tang et al., 2013). This discrepancy between the phenotypic effects of the MT1-MMP Y573D mutation and Mmp14 deficiency suggests that the proteolytic and signaling functions of MT1-MMP can have opposing roles in bone physiology, and that a balance between the two functions is required for tissue homeostasis. Deletion of the gene abrogates both functions, whereas the MT1-MMP Y573D mutation only affects signaling, altering the balance between ECM proteolysis and intracellular signaling. Moreover, it should be noted again that the bone phenotype of the global or conditional Mmp14 knockout in SSC can only be observed within the first two-three months of age, whereas the bone abnormalities of Mmp14Y573D/Y573 mice become apparent in animals older than two months. It is possible that the proteolytic and signaling functions of MT1-MMP play different roles in bone modeling (postnatal development) and remodeling (adult life), respectively.
Consistent with their respective phenotypes, the BM-SSC of Mmp14Y573D/Y573D mice show increased osteoblast differentiation, and decreased chondrocyte and adipocyte differentiation. Our finding that the bone and adipose tissue phenotypes can be rescued by Mmp14wt/wt BM transplant shows that the in vivo effects of the MT1-MMP Y573D mutation result from dysregulation of SSC differentiation. Several considerations can explain the failure of Mmp14Y573D/Y573D BM to transfer the mutant phenotypes to Mmp14wt/wt mice, as well as the incapacity of Mmp14wt/wt BM to rescue the cartilage phenotype of Mmp14Y573D/Y573D mice. Mmp14Y573D/Y573D SSC have significantly reduced proliferation and increased apoptosis relative to Mmp14wt/wt SSC (Fig. 9 A - C). We examined the phenotypes of the transplanted mice 2 months after the transplant. While hematopoietic cells from Mmp14Y573D/Y573D mice were able to efficiently repopulate the BM of wt mice – indeed, we found no peripheral blood abnormalities in Mmp14Y573D/Y573D mice – Mmp14Y573D/Y573D SSC might have required a longer time than wt cells for the phenotypic effects to become apparent. Similarly, BM transplant did not affect the cortical bone phenotype of Mmp14Y573D/Y573D mice. Cortical bone has a much slower turnover than trabecular bone (Clarke, 2008); therefore a longer time is required for its homeostasis to be altered. The failure of our BM transplant experiments to affect the articular cartilage phenotype is consistent with the absence of vascularization of this tissue. Indeed, no attempts at treating joint cartilage diseases by systemic stem cell administration have thus far been effective (Jevotovsky et al., 2018).
MT1-MMP is constitutively expressed in SSC and its levels are differentially modulated during osteogenic vs. chondrogenic/adipogenic differentiation (Y. Tang et al., 2013). The MT1-MMP Y573D mutation and Mmp14 deficiency have opposing effects on SSC differentiation in vitro. Mmp14 knockout in uncommitted SSC has no effect on their differentiation in 2D culture; however, it causes decreased osteogenesis and increased chondro- and adipogenesis in 3D collagen gel (Y. Tang et al., 2013). In contrast, in 2D culture MT1-MMP Y573D expression upregulates SSC differentiation into osteoblasts and downregulates chondrocyte and adipocyte differentiation (Fig. 9 F and G). While the in vitro differentiation of Mmp14Y573D/Y573D SSC and SSC with conditional Mmp14 knockout is consistent with the phenotypes of the respective mice, both mutations fail to fully recapitulate the bone, cartilage and fat abnormalities of the global Mmp14-/- mouse, which has osteopenia, lipodystrophy and arthritis. These discrepancies indicate that MT1-MMP controls SSC differentiation by both proteolytic and non-proteolytic mechanisms, and the balance of these two functions is required for normal differentiation. MT1-MMP-mediated ECM degradation modulates mechanosignaling that controls gene expression during SSC differentiation into osteoblasts, and conditional Mmp14 deficiency in SSC blocks osteogenesis and causes severe osteopenia (Y. Tang et al., 2013). Normal SSC differentiation requires the concerted action of ECM remodeling and intracellular signaling, cell functions modulated by the extracellular environment. In vitro, in the presence of abundant ECM - such as in collagen gel culture (Y. Tang et al., 2013) – the proteolytic activity of MT1-MMP is indispensable. Conversely, in the presence of relatively low amounts of ECM – such as in 2D culture – the role of intracellular signaling becomes prevalent. In vivo, cell differentiation in the stem cell niche, tissue/organ development and remodeling have different proteolytic and signaling requirements, which are spatially and temporally modulated.
The relative contribution of proteolysis and signaling to MT1-MMP function is also coordinated by extracellular ligands that can inhibit extracellular MT1-MMP proteolytic activity and activate intracellular signaling. This hypothesis is supported by our previous finding that TIMP-2 binding to MT1-MMP activates ERK1/2 and Akt signaling (D’Alessio et al., 2008; Valacca et al., 2015), as well as by the observation that in the mouse embryo MT1-MMP is temporally and spatially co-expressed with TIMP-2 in the developing skeleton (Apte et al., 1997; Kinoh et al., 1996). TIMP-2-/- mice do not display the severe phenotype of Mmp14-/- mice (Z. Wang et al., 2000). However, in these mice signaling can be activated by MT1-MMP binding of TIMP-3 or TIMP-4, as well as a variety of extracellular and transmembrane proteins, including integrins and CD44, that physiologically interact with the MT1-MMP ectodomains (Mori et al., 2002; Zhao et al., 2004).
Thus, the loss of signaling function caused by the Y573D substitution (D’Alessio et al., 2008; Valacca et al., 2015) (Fig. 2 E) can be at the basis of the defects in SSC differentiation and the consequent phenotypes of the Mmp14Y573D/Y573D mouse. Studies by other groups, as well as our own (unpublished), have shown that, although ERK1/2 signaling is important for osteoblast differentiation (Miraoui et al., 2009; Wu et al., 2015; Xiao et al., 2002), chronic inhibition of ERK1/2 activation results in increased SSC differentiation into osteoblasts (Ge et al., 2007; Higuchi et al., 2002; Nakayama et al., 2003; Schindeler & Little, 2006; Zhang et al., 2012); conversely, inhibition of PI3K/Akt signaling blocks adipo- and chondrogenesis (Kim & Chen, 2004; Lee et al., 2015; Li & Dong, 2016; Yang et al., 2011).
The molecular mechanism that relays signaling from the Y573 residue of the MT1-MMP cytoplasmic tail to the Ras-ERK1/2 and Ras-Akt pathways (D’Alessio et al., 2008; Valacca et al., 2015), remains to be investigated. The adaptor protein p130Cas, which binds to the cytoplasmic tail of MT1-MMP by a mechanism involving Y573 (Y. Wang & McNiven, 2012), recruits Src and/or focal adhesion kinase (FAK), which activate Ras (Bunda et al., 2014; Schlaepfer & Hunter, 1997). We speculate that TIMP-2 binding to MT1-MMP triggers the assembly of the p130Cas/Src/FAK complex at the MT1-MMP tail, and that this effect is abrogated by the Y573D substitution, which thus prevents the downstream activation of ERK1/2 and Akt signaling. In addition, Y573 could control intracellular signaling by modulating cytoplasmic tail interactions with a variety of transmembrane or membrane-bound proteins including caveolin-1 (Annabi et al., 2001; Galvez et al., 2002; Galvez et al., 2004; Labrecque et al., 2004) and ß1 integrins or growth factor receptors (Langlois et al., 2007).
In conclusion, our findings provide the first in vivo evidence for an important role of MT1-MMP mediated, proteolysis-independent signaling in postnatal development and tissue homeostasis. Understanding the relative contribution of the proteolytic and signaling functions of MT1-MMP to the control of metabolic processes that affect a variety of tissues and organs will require the development of additional genetically engineered mouse models, as well as the molecular dissection of extracellular and intracellular components of the MT1-MMP signaling mechanism. The knowledge obtained from these studies can increase our understanding of the pathogenesis of important diseases that affect bone, cartilage and adipose tissue homeostasis, such as osteopenia, osteoarthritis and obesity, and potentially direct the design of novel pharmacological tools for their treatment.
MATERIALS AND METHODS
Generation of MT1-MMP Y573D mice
To generate a mouse with the Y573D mutation in the MT1-MMP cytoplasmic tail we constructed a targeting vector as described in Figure 1 A. The construct was electroporated into W4 embryonic stem (ES) cells, followed by G418 and gancyclovir double selection. The W4 cells were provided to us by the Rodent Genetic Engineering of NYU School of Medicine, who carried out the blastocysts injections and subsequent procedures. Homologous recombination targeting events were identified in 8 ES cell colonies. Two of these, devoid of chromosome abnormalities by cytogenetic analysis, were used for injection into blastocysts of C57BL/6 mice (C57BL/6NTac; Taconic), and one gave germline transmission. Mice heterozygote for the mutation (Mmp14Y573/wt), identified by PCR analysis with primers flanking the LoxP sites (Fig. 1 A), were crossed with Rosa26Cre Deleter mice (C57BL/6NTac-Gt(ROSA)26Sortm16(cre)Arte; Taconic) to excise the neo cassette. Homozygote MmpY573D/Y573D mice were generated by heterozygote mating, and crossed back to wt C57BL/6 mice to remove the Cre recombinase gene. Sequencing of MT1-MMP cDNA from MmpY573D/Y573D mouse tissue showed identity with wt MT1-MMP except for the Y573D mutation. Heterozygote mating produced wt, heterozygote and homozygote mice in the expected Mendelian ratios. The mice were genotyped by PCR using the same primers flanking the loxP sites, which afford identification of the three genotypes (Fig. 1 B). The sequences of these primers are:
Forward: GCT TGG CAG AGT GGA AAG AC
Reverse: GGG CAG TGA TGA AGG TGA GT
All the animal studies were carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of New York University School of Medicine.
Preparation of bone sections for histological analysis
Mice were euthanized by carbon dioxide narcosis, and the entire limb was removed by excising the femur at the joint in the upper extremity of the hip socket and the tibia at the ankle. The remaining skin was peeled off and the connective tissue removed. Limbs were placed in cold 4% paraformaldehyde for 24 h, and decalcified by incubation with Immunocal tm (Decal Corporation, Tallman, NY) or 10% EDTA for 72 h, with the decalcifying solution replaced with fresh one daily. Following decalcification, the limbs were rinsed 4 times for 30 min in PBS, 30 min in 0.85% NaCl and dehydrated for 30 min in 50% and 70% ethanol. Samples were embedded in paraffin blocks and sequentially cut into 5-µm sections, mounted onto slides and stained with the indicated reagents.
Micro-computed tomography (CT)
Micro-CT was performed according to published guidelines (Bouxsein et al., 2010). Bones were scanned using a high-resolution SkyScan micro-CT system (SkyScan 1172, Kontich, Belgium). Images were acquired using a 10 MP digital detector, 10W energy (70kV and 142 mA), and a 0.5-mm aluminum filter with a 9.7-μm image voxel size. A fixed global threshold method was used based on the manufacturer’s recommendations and preliminary studies, which showed that mineral variation between groups was not high enough to warrant adaptive thresholds. The cortical region of interest was selected as the 2.0-mm mid-diaphyseal region directly below the third trochanter. The trabecular bone was assessed as the 2.0-mm region at the distal femur metaphysis; measurements included the bone volume relative to the total volume (BV/TV), bone mineral density (BMD), trabecular number (Tb.N), trabecular spacing (Tb.Sp), and trabecular thickness (Tb.Th).
Immunohistochemistry
Paraffin-embedded specimens of brown adipose tissue (BAT) were cut and immunostained with antibody to uncoupling protein-1 UCP-1 (R&D Systems, cat. # MAB6158, Minneapolis, MN) at a concentration of 10 µg/mL by personnel of the Experimental Pathology Core of NYU School of Medicine. The antibody was validated using, as a negative control, sections of subcutaneous WAT devoid of BAT (Fig. 7 – figure supplement 2).
Western blotting
Western blotting analysis was done as described (D’Alessio et al., 2008; Valacca et al., 2015) using the following antibodies and predetermined dilutions/concentrations. MT1-MMP antibody: Anti-MMP14 antibody [EP1264Y] rabbit mAb (Abcam, cat. # ab51074; Cambridge, MA), dilution: 1:5,000; phospho-ERK1/2 antibody: Phospho-p44/42 MAPK (Erk1/2) (Thr202/Tyr204) (197G2) Rabbit mAb (Cell Signaling Technology, cat. # 4377; Danvers, MA), dilution: 1:1,000; phospho-Akt antibody: Phospho-Akt (Ser473) (D9E) XP® Rabbit mAb (Cell Signaling Technology, cat. # 4060), dilution: 1:2,000; total ERK1/2 antibody: p44/42 MAPK (Erk1/2) (137F5) Rabbit mAb (Cell Signaling Technology, cat. # 4695), dilution: 1:1,000; total Akt antibody: Akt (pan) (C67E7) Rabbit mAb (Cell Signaling Technologies, cat. # 4691), dilution: 1:1,000; CD44 antibody: mouse monoclonal antibody to mouse CD44 (DSHB; cat. # 5D2-27-s), dilution: 0.2 – 0.5 µg/ml; ADAM9 antibody: mouse anti-ADAM-9 monoclonal antibody (R&D Systems; cat. # MAB939); dilution: 1 µg /ml; DDR1 antibody: rabbit anti-DDR1 monoclonal antibody D1G6 (Cell Signaling Technology; cat. # 5583, dilution: 1:1,000; FGFR2 antibody: rabbit anti-FGFR2 antibody (Abcam; cat. # ab125496); dilution 1:1,000; β-tubulin: rabbit anti-tubulin monoclonal antibody 11H10 (Cell Signaling Technology; cat. #2125), dilution: 1:1,000; lamin B antibody: goat anti-lamin B antibody B-20 (Santa Cruz; cat. # sc-6217); dilution: 1:1,000; rabbit IgG antibody: anti-rabbit IgG, HRP-linked antibody (Jackson ImmunoResearch; cat. # AB_10015289), dilution 1:10,000, or (Cell Signaling Technologies, cat. # 7074), dilution: 1:10,000; mouse IgG antibody: anti-mouse HRP-linked antibody (Jackson ImmunoResearch; cat. # AB_2307391), dilution 1:10,000.
Analysis of proMMP-2 activation by gelatin zymography
The analysis of pro-MMP-2 activation by primary fibroblasts was performed as described (D’Alessio et al., 2008), using human recombinant proMMP-2 (Millipore Sigma).
Collagen degradation assay
Single cell degradation of fibrillar collagen was analyzed as described (Sakr et al., 2018), using rat tendon or bovine collagen labeled with Alexa Fluor 488 (ThermoFisher Scientific). In a modification of this assay confluent cells were seeded onto Alexa Fuor 488 labeled fibrillar collagen in Dulbecco’s Modified Minimal Essential Medium (DMEM) supplemented with 10% fetal calf serum to permit cell attachment. After 24 h incubation at 37o C, the cell layer was washed three times with serum-free DMEM and incubation was continued in serum-free phenol red-free to preclude MMP inhibition by serum inhibitors and interference with fluorescence measurement by phenol red. The medium was removed at 24 h intervals, immediately analyzed for Alexa Fluor 488 fluorescence with a fluorescence microplate reader, and replaced with the same volume of medium. Medium incubated with the labeled collagen in the absence of cells was used as a blanc whose fluorescence readings were subtracted from the samples’ readings at each time point. The time of the first addition of serum-free, phenol red-free DMEM was considered as time 0.
Isolation, culture and differentiation of BM SSC
Cultures of BM SSC were established by a modification of the method described (Soleimani & Nadri, 2009). BM was flushed from cut femurs and tibiae using 3-ml syringes with Minimum Essential Medium Alpha (αMEM without ribonucleosides or deoxyribonucleosides; Gibco, Life Technologies cat. # 12561-056, Grand Island, NY) supplemented with 15% fetal calf serum (FBS; Atlanta Biologicals, cat. # S11150, Flowery Branch, GA) and antibiotics (Penicillin-Streptomycin). After separating the BM into a single-cell suspension, the cells were passed through a 70-μm cell strainer, incubated for 2 min in NH4Cl red blood cell lysis solution (StemCell Technologies, cat. # 07800, Vancouver, Canada), and seeded into culture plates in αMEM supplemented with 15% FBS and antibiotics. Three hours later non-adherent cells were removed by replacing the medium with fresh complete medium. This procedure was repeated twice a day for the first 72 h. Subsequently, the adherent cells were washed with phosphate buffer saline (PBS) and incubated with fresh medium every 3-4 days until the cultures became subconfluent. To induce differentiation the cells were seeded into 24-well plates (5 × 104 cells/well). After 24 h, the culture medium was substituted with osteoblast or adipocyte differentiation medium, and incubation was continued for the indicated times, changing the medium with freshly prepared medium twice a week. For chondrocyte differentiation the cells were seeded into 48-well plates as a pellet (2.5 × 105 cells/well). The medium was substituted with freshly prepared differentiation medium 2 h after seeding, and subsequently three times a week. Differentiation medium consisted of αMEM supplemented with 10% FBS, penicillin/streptomycin and, for osteoblast differentiation: dexamethasone 1 µM (Millipore Sigma, cat. # D4209; St. Louis, MO), beta-glycerolphosphate 20 mM (Millipore Sigma, cat. # 500200), L-ascorbic acid 50 µM (Millipore Sigma, cat. # A5960); for adipocyte differentiation: dexamethasone 1 µM, indomethacin 50 µM (Millipore Sigma, cat. # I7378), 3-Isobutyl-1-methylxanthine (IBMX) 500 nM (Millipore Sigma, cat. # I5879), insulin 5 µg/ml (Millipore Sigma, cat. # I9278); and for chondrocyte differentiation: sodium pyruvate 100 µg/ml, ascorbic acid 50 µg/ml, dexamethasone 0.1 µM, insulin, transferrin, selenium (ITS) mix 1X (Millipore Sigma, cat. # I3146) and bone morphogenetic protein-2 (BMP2) 300 ng/ml (Peprotech, cat. # 120-02-2UG, Rocky Hill, NJ). The SSC used for differentiation experiments were no older than passage 3 in culture.
CH310T1/2 cell culture, transfection and differentiation
C3H10T1/2 cells obtained from the American Type Culture Collection (ATCC CLL226, Gaithersburg, MD) were grown in Dulbecco’s MEM (DMEM; Gibco, Life Technologies, cat. # 10566-016) supplemented with 10% FCS (Atlanta Biologicals). Cells at 50-70% confluency were transfected by overnight incubation with the indicated cDNAs in complete growth medium using TransIT®-LT1 - Transfection Reagent according to the manufacturer’s instructions (Mirus Bio, LLC, cat. # MIR2300; Madison, WI). Stable transfectants were selected in medium containing 500 μg/ml hygromycin B (InvivoGen, cat. # ant-hg-1; San Diego, CA). Pools of resistant cell colonies were characterized for MT1-MMP expression by reverse transcription-PCR and Western blotting, and used for differentiation assays. To induce differentiation the cells were seeded into 24-well plates at a density of 2.5 × 104 cells/well and then shifted to DMEM with low glucose (1 g/L; Gibco, Life Technologies, cat. # 11885076) with the same FCS concentration and supplements used for BM-derived SSC differentiation.
Quantitative PCR analysis of gene expression in differentiating cells
Cells were lysed in Trizol Reagent (Invitrogen, Waltham, MA) and total RNA was isolated with RNeasy (Qiagen, Valencia, CA), according to the manufacturer’s instructions. RNA was quantified using a Nanodrop 2000 spectrophotometer (ThermoFisher Scientific, Waltham, MA), and 1 µg of total RNA was used for cDNA preparation using SuperScript III (Invitrogen, Life Technologies). Quantitative real-time PCR reactions were performed with SYBR green PCR reagents using the ABI Prism 7300 sequence detection system (Applied Biosystems-Life Technologies, Waltham, MA). Relative gene expression levels were calculated using the 2 delta Ct method (Livak & Schmittgen, 2001). Target mRNA levels were normalized to the geomean of Supplementary File 1.
Analysis of circulating hormones and cytokines
Serum and/or plasma levels of the indicated hormones and cytokines were measured by multiplex ELISA kits (Millipore, Burlington, MA) at the High-Throughput Biology Laboratory of NYU School of Medicine.
RNA-Seq and data analysis
The indicated tissues and SSC were isolated from Mmp14wt/wt and Mmp14Y573D/Y573 mice (3 mice/tissue/genotype) and total RNA was extracted using the RNeasy kit (Qiagen). The subsequent steps were carried out by personnel of the Genome Technology Center of NYU School of Medicine. RNA-Seq libraries were prepared with the TruSeq sample preparation kit (Illumina, San Diego, CA). Sequencing reads were mapped to the mouse genome using the STAR aligner (v2.5.0c) (Dobin et al., 2013). Alignments were guided by a Gene Transfer Format file (Ensembl GTF version GRCh37.70). The mean read insert sizes and their standard deviations were calculated using Picard tools (v.1.126). Read count tables were generated using HTSeq (v0.6.0) (Anders et al., 2015), normalized to their library size factors with DESeq (v3.7) (Anders & Huber, 2010), and differential expression analysis was performed. Read Per Million (RPM) normalized BigWig files were generated using BEDTools (v2.17.0) (Quinlan & Hall, 2010) and bedGraphToBigWig tool (v4). Statistical analyses were performed with R (v3.1.1), GO analysis with David Bioinformatics Resources 6.8 (Huang da et al., 2009a, 2009b) and GSEA with the hallmarks (h.all.v6.1.symbols.gmt) gene sets of the Molecular Signature Database (MSigDB) v6.1 (Mootha et al., 2003; Subramanian et al., 2005).
BM transplant
Mice of 3 and 5 months of age were used for BM transplant experiments. The BM of female mice was transplanted into age-matched male mice. The recipient mice were lethally irradiated with a single dose of 9 Gy. Twenty-four hours later BM cells (BMCs) were collected from the tibias and femurs of donor mice, suspended in PBS containing 2% heat-inactivated FBS, and 5 × 106 BMCs in 200 µl were injected into the recipient mice via the tail vein. Eight weeks later the mice were subjected to DEXA scanning analysis and then sacrificed. To verify BM engraftment we transplanted female mice with male BM, in order to characterize the recipients’ BM by immunohistochemistry and/or flow cytometry with antibody to UTY, a Y chromosome marker (H. Wang et al., 2013). However, several commercially available antibodies failed to recognize UTY in murine BM cells, probably because this marker is not expressed in these cells. For the same reasons we were not able to identify the genotype of transplanted animals’ WAT or BAT by immunohistochemistry with UTY antibody, or RNA in situ hybridization. Therefore, we genotyped the BM of recipient mice for the Mmp14Y573D/Y573D mutation using the PCR primers designed for genotyping our mice. We reasoned that partial engraftment of donor’s BM would result in both the Mmp14wt/wt and Mmp14Y573D/Y573D genotypes, with a pattern of PCR products identical to that of Mmp14Y573D/wt mice (Fig. 1 B). In addition, running the PCR reaction for a high number of cycles would afford detection of small residual amounts of BM from recipient mice. By this method, the results (Fig. 10 – figure supplement 1) showed virtually complete replacement of the recipient’s BM with the donors’ BM. However, we could not use this method to identify donor-derived adipocytes in the transplanted animals’ WAT or BAT as these tissues contain circulating cells – e.g. macrophages – from the BM.
Statistical analysis
For the BM transplant experiments, to calculate the number of mice required in each group to achieve statistical significance for the expected differences between controls and experimental groups, we used the analysis of Power of Test based on the variance observed in our previous analyses of our mice. In our characterization of the bone, cartilage and fat phenotypes we found differences between MT1-MMP Y573D wt mice ranging 25% - 75% in the mean (m) values of the various parameters, with standard deviations (SD) ranging 25% - 50%. Therefore, we assumed m = 50% and SD = 35%. To calculate the number of animals needed we used the formula: n = (Za +Zb)2 × 2 (%SD)2/(%D)2, where: n is number of animals per group, (Za + Zb)2 for a 2-Tailed test with power of 90% at p=0.05 is 10.5, %SD is the percent standard deviation, and %D is the assumed percent difference between control and experimental group(s). Thus, n = 10.5 × 2(35)2 / 502 = 10.29. Therefore, we planned to use 11 mice for each experimental and control group. The statistical analysis of the results of all the experiments was performed with the two-tailed Student’s t-test (unless otherwise indicated) using GraphPad Prism, Version 7.05.
COMPETING INTERESTS
The authors declare that they have no financial or non-financial competing interests.
SUPPLEMENTAL INFORMATION
ACKNOWLEDGMENTS
We gratefully acknowledge the precious collaboration of the Rodent Genetic Engineering, High-Throughput Biology Laboratory, Experimental Pathology Research Laboratory, and Genome Technology Center of NYU School of Medicine, which are partially supported by NIH grant P30CA016087 to the Laura and Isaac Perlmutter Cancer Center and the Shared Instrumentation Grant S10 OD021747, and the MicroCT Core of NYU College of Dentistry, supported by NIH grant S10 OD010751 to Dr. Nicola C. Partridge.
Footnotes
The summary remains the same.