Stromal regulation of vessel stability by MMP14 and TGFβ

When tissues are injured, vasodilation of capillaries and extravasation of plasma proteins into the interstitial tissue mark the onset of vascular remodeling following tissue assault (Bhushan et al., 2002). These processes are crucial not only for initiating a healing response, but also for enabling re-establishment of tissue homeostasis. Although molecules that regulate aspects of vascular stability and/or leakage have been identified, the molecular mechanisms controlling transport of macromolecules across the endothelium have only recently begun to be defined. Extravasation of plasma proteins is subject to regulation by many factors – some affect vessel leakiness by regulating the formation of openings in venular endothelium, resulting in exposure of subendothelial basement membranes to capillary lumens (Feng et al., 1997; Hashizume et al., 2000; Feng et al., 2002; McDonald and Baluk, 2002), whereas others regulate the diffusion of macromolecules into interstitium (McKee et al., 2001; Pluen et al., 2001; Brown et al., 2003). In the resting state, large plasma proteins such as albumin are transported across the endothelial body through a series of vesicles that may or may not fuse to form transcellular channels (Mehta and Malik, 2006), underscoring the fundamental importance of transcellular pathways in maintaining the semi-permeable nature of continuous endothelium (Drab et al., 2001). In contrast to this transcellular-type transport, the majority of plasma protein leakage in response to inflammatory stimuli occurs through the formation of gaps between cells, i.e. ‘paracellular’ leakage (Predescu et al., 2002; Mehta and Malik, 2006).

Vascular responses to tissue damage are accompanied by type I collagen remodeling in perivascular stroma (Page and Schroeder, 1982). The extracellular matrix (ECM), including fibrillar type I collagen, is rapidly remodeled around blood vessels following the acute inflammatory processes that accompany tissue damage, as well as during chronic vascular pathologies, e.g. atherosclerosis, hypertension, varicosis, restenosis, etc. (Jacob et al., 2001). Matrix metalloproteinases (MMPs) that cleave interstitial collagens also play a crucial role in regulating perivascular matrix remodeling. Indeed, sustained MMP activity is associated with some vascular pathologies, including atherosclerosis, hypertension restenosis and aneurysm (Mott and Werb, 2004; Page-McCaw et al., 2007). MMPs can further contribute to vascular remodeling by liberating vasoactive cytokines from stromal matrices, including the angiogenic/permeability factor vascular endothelial growth factor (VEGF) (Bergers et al., 2000; Sounni et al., 2002), as well as activating latent growth factors such as transforming growth factor β (TGFβ) (Yu and Stamenkovic, 2000; Mu et al., 2002; Wang et al., 2006). Despite extensive investigations into the roles of MMPs as mediators of chronic vascular pathologies, surprisingly little is known regarding their role in acute vascular responses, or how they contribute to vascular homeostasis. Accordingly, we investigated whether MMPs participate in the early phases of acute tissue repair, and whether they contribute to the appropriate vascular responses to tissue damage. In this study, we identified a post-translational pathway whereby type I collagen fibrils regulate perivascular MMP activity and TGFβ bioavailability, which in turn regulate vascular homeostasis by altering vessel stability and leakage.

Previous studies have reported that ectopically applied collagenase, or a reduced accumulation of collagen fibrils in tissue, correlates with enhanced ‘drug’ delivery to tumors (McKee et al., 2001; Brown et al., 2003; Loeffler et al., 2006; Gade et al., 2009), thus indicating that the organization and structure of perivascular collagen fibrils regulate vascular leakage. To directly assess whether inhibition of collagenolytic MMP activity impacted vascular leakage, we used the Miles assay (Miles and Miles, 1952), which is an in vivo assay of vascular leakage, to analyze the acute cutaneous vascular response to mustard oil (MO) in mice that were pretreated systemically with the broad-spectrum metalloproteinase (MP) inhibitor GM6001. Wild-type (wt) mice were administered with GM6001 (or vehicle) for 5 days, followed by acute challenge with MO versus vehicle (mineral oil, MnO) (Inoue et al., 1997), one minute after receiving an intravenous (i.v.) injection of Evans Blue (EB) dye, which binds to and marks serum albumin. As expected, following MO exposure, EB extravasation rapidly increased in ear tissue interstitium, indicative of plasma protein leakage (Fig. 1A). Surprisingly, however, prior treatment with GM6001 significantly increased EB leakage (Fig. 1A). This indicated that the inhibition of steady-state MP activity rendered vessels more susceptible to MO-induced acute leakage, and implied a link between steady-state MP activity and vascular leakage.

To identify the MP associated with this response, we assessed steady-state and MO-induced EB leakage, using the Miles assay, in control and homozygous null mice lacking genes that encode collagenolytic MMPs, i.e. MMP2, MMP8, MMP13 and MMP14 (Fig. 1B,C). Steady-state and MO-induced vascular leakage characteristics in the ear skin were similar in MMP2, MMP8 and MMP13 null skin as compared with the skin from littermate control mice (Fig. 1B, and data not shown). However, in mice lacking MMP14, the steady-state vascular leakage of EB from ear skin was elevated compared with aged-matched controls (Fig. 1C), and similar to the leakage following MO exposure. The steady-state leakage of capillaries in back skin was also higher in MMP14 null mice versus age-matched heterozygous and wild-type littermates (Fig. 1D).Thus, the enhanced leakage of cutaneous vessels in mice lacking MMP14 indicates that MMP14 participates in regulating steady-state vascular leakage.

To further explore how collagen fibril organization and MMP14 activity impacted vascular leakage, we utilized a mouse model of age-related dermal fibrosis, Colα1(I)^r/r mice (Liu et al., 1995), which exhibit threefold higher tissue collagenolytic activity in skin (Fig. 2A). The Colα1(I)^r/r mice also exhibited a ~50% reduction in leakage following MO exposure (Fig. 2B), which could be rescued fully by prior treatment of mice with GM6001 (Fig. 2C). Although the levels of MMP14 protein are similar in wt and Colα1(I)^r/r mice (Fig. 2D), we observed that MMP14 activity, as evidenced by activation of proMMP2, is markedly enhanced in an ex vivo bioassay using collagen from Colα1(I)^r/r mice (Fig. 2E), and that this enhanced activity is reflected by constitutive activation of MMP2 in Colα1(I)^r/r mice in vivo (Fig. 2F). To re-affirm that MMP2 was not directly responsible for the restricted vascular leakage in Colα1(I)^r/r) mice, selective loss of MMP2 in Colα1(I)^r/rMMP2^−/− mice (Egeblad et al., 2007) did not alter Colα1(I)^r/r susceptibility to MO-induced capillary leakage (Fig. 2G).

Colα1(I)^r/r mice were also found to be resistant to acute vascular leakage induced by either serotonin or VEGF (Fig. 3A). Moreover, the resistance to VEGF leakage in Colα1(I)^r/r mice was not linked to changes in the expression or activation of VEGF receptor 2 (VEGFR2) (Fig. 3B), and together suggest that the elevated collagenolytic activity in Colα1(I)^r/r mice reduces the acute vascular leakage induced by a variety of stimulants.

To further evaluate the pathologic basis underlying reduced vascular leakage in Colα1(I)^r/r mice, we assessed the organization, architecture and diameter of cutaneous capillary networks in ears from Colα1(I)^r/r mice and their control littermates following MO exposure. We assessed capillary beds by fluorescent angiography using fluorescein-labeled Lycopersicon esculentum (tomato) lectin that specifically binds to the luminal surfaces of vascular endothelium (Thurston et al., 1996). The organization and diameter of capillaries in quiescent vascular beds of control and Colα1(I)^r/r mice were similar, both qualitatively (Fig. 3C) and quantitatively (Fig. 3D). However, following MO treatment, the total vessel area, and in particular the vessel diameter, increased significantly in control mice but remained unchanged in the skin of Colα1(I)^r/r mice (Fig. 3C,D).

Since mural cell coverage of blood vessels contributes to vasodilation and extravasation of plasma proteins, we examined the presence of perivascular cells on venules in the skin from wt and Colα1(I)^r/r mice by whole-mount immunodetection of α-smooth muscle actin (αSMA), a marker for mature perivascular cells (Verbeek et al., 1994). Interestingly, the position and morphology of αSMA-positive perivascular cells in vehicle-treated control versus MO-treated Colα1(I)^r/r skin were indistinguishable (Fig. 3E, panels a–c).

Extravasation of plasma proteins into tissue interstitia is also regulated by the frequency of openings in venular endothelium (McDonald and Baluk, 2002); accordingly, we injected control and Colα1(I)^r/r mice with fluorescein-labeled Lycopersicon esculentum lectin (to visualize vasculature), in combination with rhodamine-labeled Ricinus communis agglutinin I (to visualize luminal openings). Following MO exposure, numerous leakage sites appeared in capillary beds of tissue from control mice (Fig. 3E, panel e), specifically in capillaries where perivascular cell coverage was prominent (Fig. 3E, panel b). However, the leakage sites were less abundant in MO-treated Colα1(I)^r/r capillaries (Fig. 3E, panel f), indicating that the enhanced stability of vessels in Colα1(I)^r/r mice correlated with reduced vasodilation following MO challenge and a diminished frequency of venular openings, independent of similar perivascular cell coverage.

In addition to activating proMMP2, MMP14 also regulates the bioavailability of several chemokines and growth factors, most notably latent TGFβ (Werb, 1997; Mu et al., 2002; Alfranca et al., 2008), which can profoundly impact vascular integrity. Given that MMP2 deficiency failed to alter vascular stability alone (Fig. 1B), or in combination with the Colα1(I)^r/r mutation (Fig. 2G), we evaluated whether TGFβ₁ might mediate a MMP14- or collagen-regulated vascular response. We assessed whole-skin lysates by ELISA and found decreased levels of total TGFβ₁ in hyperleaky MMP14-deficient mice (Fig. 4A). We also found, in leakage-resistant Colα1(I)^r/r mice, an elevated level of total TGFβ₁ that was independent of MMP2 (Fig. 4B and supplementary material Fig. S1A). Moreover, although Tgfb1 mRNA expression (Fig. 4C) and the synthesis of latent TGFβ [the TGFβ latency-associated peptide (LAP)] (Fig. 4D) were similar in both wild-type and Colα1(I)^r/r mice, we observed an increase in the mature, bioactive form of TGFβ₁ in Colα1(I)^r/r tissue lysates (Fig. 4E,F), which was sensitive to GM6001 inhibition (Fig. 4G). Using an organotypic cell-based assay with cells that constitutively express MMP14, which were cultured on collagen derived from Colα1(I)^r/r mice, we confirmed that the enhanced bioavailability of TGFβ₁ (and MMP2 activity) was indeed regulated directly by MMP14, and enhanced in the presence of mutant collagen fibrils (Fig. 4H and supplementary material Fig. S1B).

To directly demonstrate the role of TGFβ in regulating vascular stability and response, we administered a pan-TGFβ-neutralizing antibody, which neutralizes the bioactivity of all TGFβ isoforms in vivo (Yamamoto et al., 1999; Neptune et al., 2003), to 6-week-old control and Colα1(I)^r/r mice prior to MO challenge. Similar to our results following the administration of GM6001 (Fig. 2C), neutralization of TGFβ in Colα1(I)^r/r mice following MO stimulation resulted in appropriate acute vascular remodeling and reinstated the characteristic EB leakage to a level comparable to untreated controls (Fig. 4I).

The TGFβ type I receptor ALK5 (activin-like kinase 5) is expressed in both CD31⁺ endothelial cells and αSMA-positive perivascular cells (data not shown). Similar to treatment with the pan-TGFβ-neutralizing antibody, responsiveness to MO-induced vascular leakage was restored following administration of an inhibitor of ALK5 (ALK5-I) (Fig. 4J); it also significantly enhanced the steady-state vascular leakage in unstimulated skin (Fig. 4K). Thus, consistent with the hyperleaky phenotype of MMP14 null mice, which have decreased steady-state levels of TGFβ, vessel stability and vascular leakage in Colα1(I)^r/r mice are susceptible to regulation by modulation of metalloproteinase activity, TGFβ bioavailability and signaling downstream of ALK5.

We reasoned that if neutralization of TGFβ or blockade of ALK5 signaling enhanced steady-state and acute leakiness, then sustained, ectopic expression of TGFβ₁ would phenocopy the elevated MMP14 activity that we observed in Colα1(I)^r/r mice, and result in resistance to leakage. To test this hypothesis, we utilized an inducible TGFβ₁ bigenic mouse model (Cao et al., 2002) that expresses TGFβ₁ in basal keratinocytes of skin following induction by RU486 (Lu et al., 2004). Because exposure to RU486 for 6 days induced a psoriatic phenotype (Fig. 5A, panels a,b), we performed the EB leakage assay after 3 days of exposure to RU486, prior to the onset of psoriasis, when the skin of the bigenic mice appeared ‘normal’ (Fig. 5A, panel a) even though RU486-induced expression of TGFβ₁ was prominent in keratinocytes (Fig. 5A, panel c). Significantly, the bigenic mice treated with RU486 for 3 days were resistant to MO-induced EB leakage (Fig. 5B), confirming that sustained ectopic expression of TGFβ phenocopied the vascular effects linked to elevated MMP14 activity in Colα1(I)^r/r mice.

The data above indicate that MMP14-activated TGFβ₁ regulates vascular homeostasis and the vascular response to acute tissue damage via signaling through ALK5. To determine whether this signaling pathway was susceptible to regulation in aged tissues where fibrosis and the cross-linking of collagen fibrils can compromise tissue hemodynamics, we utilized aged C57BL/6 mice. Similar to young animals, transient blockade of ALK5 in aged mice led to enhanced vessel leakiness following MO stimulation (Fig. 6A).

To determine whether the ALK5 signaling pathway was also active in chronically leaky angiogenic vessels, such as those found in neoplastic tissues, we utilized two transgenic models of epithelial carcinogenesis (Guy et al., 1992; Coussens et al., 1996). Angiogenic blood vessels in the premalignant skin of K14-HPV16 transgenic mice exhibit increased steady-state leakiness (Eichten et al., 2007) in capillaries that are largely devoid of αSMA-positive perivascular cells (Fig. 6B,C). Transient ALK5 blockade in K14-HPV16 transgenic mice further increased vessel leakage and extravasation of EB dye (Fig. 6C). Together, these data indicate that the ALK5 signaling pathway remains susceptible to therapeutic modulation in both aged and angiogenic vessels.

Based on this, we hypothesized that antagonizing ALK5-mediated signaling in tumor vessels would enhance leakage and potentially improve the delivery of high molecular weight compounds, such as antibodies, liposomes or contrast agents, into the tumor interstitium. Moreover, to ensure that inhibition of TGFβ was influencing vascular permeability and not simply modulating bulk flow, we performed live imaging of mice to dynamically follow the leakage of high and low molecular weight dextrans into mammary carcinomas in MMTV-PyMT transgenic mice (Guy et al., 1992). ALK5 inhibition conferred a significant increase in the leakage of 70 kDa dextran into late-stage carcinomas (Fig. 6D,E). Moreover, although ALK5 blockade had no effect on the initial leakage of 10 kDa dextran, it significantly increased the retention of dextran in the tumor. Thus, transient blockade of ALK5 in live mice bearing large invasive tumors enhanced the leakiness of angiogenic vessels, enabling increased delivery of high molecular weight compounds and increased retention of low molecular weight compounds to tumor tissue. Together, these data indicate that TGFβ-mediated signaling through the ALK5 receptor in vascular cells regulates vascular stability and leakage in homeostatic, acutely damaged, aged, and angiogenic tissue, and thus reveals a therapeutic opportunity to enhance the delivery of drugs to tissue where hemodynamics limit that possibility.

Herein we describe a novel mechanism by which TGFβ regulates vessel stability and leakage in homeostatic, acutely damaged and angiogenic blood vessels. Our findings demonstrate that, in cutaneous vasculature, steady-state levels of MMP14 and TGFβ regulate homeostatic vascular leakage via ALK5 signaling, and that, together, they regulate extravasation of plasma proteins into the tissue interstitium. Moreover, we provide further evidence that a post-translational pathway, mediated by perivascular type I collagen and MMP14 activity, controls the bioavailability of TGFβ. Taken together, these data indicate a central role for stromal metalloproteinase-activated TGFβ in mediating vascular homeostasis and remodeling, and further indicate that TGFβ and/or MMP14-selective antagonists may enhance vascular leakage and the delivery of therapeutics to tissues where hemodynamics limit efficient drug delivery (Fig. 7).

TGFβ has long been implicated as a regulator of vascular integrity (Pepper, 1997; Gleizes and Rifkin, 1999; Tuxhorn et al., 2002); vasculogenic and angiogenic processes (Dickson et al., 1995; Pepper, 1997); and endothelial and perivascular cell proliferation and/or differentiation (Sato, 1995; Yan and Sage, 1998; Vinals and Pouyssegur, 2001). Vascular abnormalities have also been described in TGFβ null mice, as well as in mice lacking various TGFβ type I and II receptors (Dickson et al., 1995; Seki et al., 2003; Seki et al., 2006). Despite extensive research on the impact of TGFβ on vasculature, the mechanisms by which TGFβ regulates vascular development and function are complex and remain controversial.

TGFβ differentially impacts endothelial and vascular smooth muscle cells in a concentration-, substrate- and tissue-specific context (Madri et al., 1992; Yan and Sage, 1998). For example, in bleomycin-induced pulmonary edema, TGFβ promotes vascular leakage by acting directly on pulmonary endothelial cells in concert with integrin α5β1 (Pittet et al., 2001; Su et al., 2007). TGFβ also induces the expression of angiopoietin-like protein 4 (Angpt-4) that, in turn, disrupts endothelial cell junctions, increases leakage, and promotes tumor cell emigration and metastasis to the lungs (Padua et al., 2008).

By contrast, in pancreatic tumors, vascular leakage is induced when TGFβ type I receptors are inhibited, accompanied by decreased Smad2-regulated signaling cascades, specifically in endothelia (Kano et al., 2007). In the brain, the inhibition of TGFβ type I ALK5 receptors also increases leakage (of the blood brain barrier), accompanied by changes in the expression of tight junction adhesion proteins (Ronaldson et al., 2009).

Our data indicate that, in cutaneous and mammary vessels, TGFβ acts primarily through the ALK5 receptor to restrict vessel leakage. Indeed, our localization data showing ALK5 expression in perivascular support cells (data not shown) concur with recent in vivo studies showing that the deletion of ALK5 in mice results in the loss of mural cells, whereas vessel lumens remain intact (Seki et al., 2006). Although we cannot rule out the possibility that TGFβ may also directly or indirectly influence the endothelial component of the vascular wall in vivo (Cambier et al., 2005; Roviezzo et al., 2005; Parikh et al., 2006), recent studies with targeted, endothelial cell-selective deletion of ALK5 reveal normal vascular development and morphology, supporting previous observations that ALK5 is more crucial for smooth muscle rather than endothelial cell function (Park et al., 2008). Moreover, despite the fact that either MMP14 (Lehti et al., 2005) or TGFβ can promote vessel investment by perivascular cells, and that TGFβ alters the accumulation of macromolecules in pancreatic tumors treated with TGFβ inhibitors and has been linked to reduced pericyte coverage (Hirschi et al., 1998; Kano et al., 2007), we did not observe any differences in perivascular cell coverage in Colα1(I)^r/r tissues, indicating that TGFβ may activate multiple pathways in perivascular cells leading to stabilization of vasculature.

In vascular smooth muscle cells, TGFβ can bind to ALK5 and activate Smad2 and Smad3 to promote differentiation, and increase contractility and ECM synthesis (Dumont and Arteaga, 2003). Although studies in cultured endothelial cells also described ALK5-mediated increases in Smad3 through an MMP14-dependent activation of TGFβ (Alfranca et al., 2008), we did not observe any changes in vascular leakage in Smad3-deficient mice (N.E.S. and L.M.C., unpublished observations), indicating that ALK5-mediated changes in vascular leakage in vivo probably impact the Smad2 pathway in smooth muscle cells.

Importantly, several studies have found that ALK5 inhibition effectively reduces matrix synthesis and impairs fibrotic reactions in organs including the skin (scleroderma) and kidney (Ishida et al., 2006). Moreover, TGFβ-induced fibroblast or smooth muscle cell contractility can increase the tension on extracellular matrices leading to increased interstitial fluid pressure, which further restricts capillary outflow (Heldin et al., 2004), whereas ALK5 inhibition reduces dermal fibroblast activation in vivo (Ishida et al., 2006). In tumors, drug delivery is influenced considerably by the increased interstitial fluid pressure caused by the increased permeability of tumor vasculature and the absence of functional lymphatics (Minchinton and Tannock, 2006), and is exacerbated further by increased collagen deposition and fibrosis in activated tumor stroma Heldin et al., 2004). Reduced type I collagen accumulation enhances drug penetration and drug responses (McKee et al., 2001; Brown et al., 2003; Loeffler et al., 2006; McKee et al., 2006), as does ectopic delivery of interstitial collagenases (Gade et al., 2009). Thus, transient blockade of TGFβ would be expected to not only reduce matrix synthesis, but also to reduce perivascular and stromal cell contractility, leading to reduced interstitial fluid pressure, and improved tissue perfusion and drug delivery.

Indeed, in the MMTV-PyMT transgenic mouse model of mammary carcinogenesis, ALK5 inhibition increased the leakage of high molecular weight dextran into tumor tissue, and increased the retention of low molecular weight dextran in late-stage tumors, indicating that TGFβ and ALK5 regulate vascular paracellular transport mechanisms responsible for the transport of high molecular weight molecules. Although tissue specificity may dictate which intracellular signaling pathway regulates the TGFβ effects on vascular homeostasis, it is clear that multiple tissues, in addition to skin and mammary tissue, respond similarly to TGFβ blockade. In thyroid tumors, inhibition of the TGFβ type II receptor (TGFβRII) leads to reduced interstitial fluid pressure and outward flow of serum proteins from capillaries (Lammerts et al., 2002), and in pancreatic tumors, this leads to the increased accumulation of macromolecules in stroma (Kano et al., 2007).

In Colα1(I)^r/r mice, where steady-state levels of bioavailable TGFβ are maintained at elevated levels, we observed a reduction in luminal openings in vessels with a diameter greater than 5 μm. However, in both Colα1(I)^r/r mice and aged wild-type mice that accumulate non-enzymatically cross-linked collagen and exhibit vascular dysfunction (Susic, 2007), inhibiting signaling by the ALK5 receptor restores the appropriate vascular response. Thus, the ability of TGFβ to impact matrix synthesis and organization, as well as the contractility of perivascular cells, and to reduce vascular openings, may collectively contribute to restricted vascular leakage in Colα1(I)^r/r mice.

Our data indicate the existence of a novel feed-forward mechanism by which type I collagen fibrils regulate the activity of MMP14 and the bioavailability of TGFβ. Previous studies have reported that type I collagen upregulates MMP14 mRNA expression (Haas et al., 1999), and increases cell surface stabilization of the MMP14 protein (Haas et al., 1999), consistent with our data (supplementary material Fig. S1B). Moreover, collagen interactions with the MMP14 hemopexin C domain enhance the formation of multimeric MMP14 complexes, which, in turn, activate proMMP2 (Tam et al., 2002). Typically, MMP14 protein is degraded autolytically following MMP2 activation, and subsequent degradation of collagen further downregulates MMP14 mRNA expression, whereas addition of metalloproteinase inhibitors leads to prolonged stabilization of MMP14 on the cell surface in cell-based assays (Bernardo and Fridman, 2003). Thus, in Colα1(I)^r/r mice, the inability to cleave collagen probably results in surface clustering of MMP14, and stabilized MMP14 enables localized cleavage of the TGFβ LAP to liberate active TGFβ, as well as the enhanced presence of the active form of MMP2.

LAP-TGFβ₁ can localize to cell surfaces via integrins (αvβ1, αvβ6 and αvβ5) (Annes et al., 2003) to facilitate TGFβ activation (Cambier et al., 2005). In airway epithelia, LAP-TGFβ also binds to αvβ8, and cleavage of LAP from the TGFβ-αvβ8 complex is mediated by MMP14 (Mu et al., 2002; Araya et al., 2006). As αvβ8 is not expressed by vascular smooth muscle cells (Mu et al., 2002), stabilized collagen fibrils in Colα1(I)^r/r mice may directly activate and stabilize MMP14, which then subsequently activates latent TGFβ. Moreover, although MMP2 can also induce maturation of TGFβ₁ or degrade fibrillin, a microfibril-anchoring protein whose loss is linked to activation of TGFβ (Dallas et al., 2002; Mu et al., 2002; Annes et al., 2003), our data demonstrate that MMP14 null mice, but not MMP2 null mice, display vascular phenotypes similar to TGFβ inhibition. Evaluation of steady-state and MO-induced vascular leakage in MMP2-deficient Colα1(I)^r/r mice was similar to that found in Colα1(I)^r/r mice. Thus, by interfering with the dynamic interplay between collagen, MMP14 and TGFβ, we have identified a novel post-translational pathway that regulates vascular homeostasis.

Findings from this study support several important new concepts (Fig. 7). First, steady-state levels of TGFβ in cutaneous tissue restrict vascular leakage and confer vascular stability. Second, ECM proteins that provide structural integrity also play crucial roles in regulating TGFβ activation, as well as MMP activity (Tam et al., 2002). In a fibrillar state, pericellular type I collagen regulates the expression and activity of the collagenolytic proteases that degrade it (i.e. MMP14 and MMP2) (Overall, 2001), as well as the bioavailability of TGFβ, to directly regulate cell and tissue status. Thus, fibrillar collagen represents an abundant sensor-type molecule endowed with an inherent self-regulating ‘switch’ that is engaged upon rapid collagen degradation following acute trauma and that, in turn, suppresses a post-translational pathway maintaining vascular stability. Third, these data provide a novel link between type I collagen, MMP14 and TGFβ activation and the regulation of vasodilation and extravasation of plasma proteins – perturbations of which manifest in the pathogenesis of several cutaneous diseases. The present study demonstrates that, in microenvironments where type I collagen fibrils remain stable following acute tissue stimulation, sustained steady-state metalloproteinase activity and TGFβ bioavailability exert dominant control over vascular cells and limit their ability to mount an acute response. Together, these observations support the hypothesis that antagonizing MMP14 or TGFβ activity could repress vascular quiescence in fibrotic tissues, increase plasma protein extravasation in hyperactive tissues, and normalize infiltrating responses to resolve fibrotic disorders, whereas enhanced MMP or TGFβ activity may encourage vascular quiescence. Clinically, these data imply that pharmacologic agents that modulate type I collagen fibril status, MMP14 activity and/or TGFβ bioavailability, and/or downstream ALK5 signaling, could profoundly alter tissue hemodynamics that could be therapeutically exploited to improve drug delivery or molecular imaging.

All mice were maintained within the UCSF Laboratory for Animal Care barrier facility, and all experiments involving animals were approved by the Institutional Animal Care and Use Committee (IACUC) of UCSF. Colα1(I)^r/r mice were derived from the colony at Massachusetts General Hospital, Boston, where the mutation was targeted to the J1/129 embryonic stem cells and introduced into the C57BL/6 strain, as described previously (Liu et al., 1995). Backcrosses to FVB/n (N5) mice were performed to create an inbred line of Colα1(I)^r/+ mice, and a breeding colony of homozygous mutant Colα1(I)^r/r mice was established (UCSF). Controls were the progeny of Colα1(I)^+/+ breeding pairs that did not possess the mutated gene but were on the same genetic background. The presence of the mutant Col1a1 allele was assessed by PCR genotyping of tail DNA using oligonucleotide primers that discriminated between the wt allele (5′-TGGACAACGTGGTGTGGTC-3′ and 5′-TTGAACTCAGGAATTTACCTGC-3′) and the mutant allele (5′-TGGACAACGTGGTGTGGTC-3′ and 5′-TGGACAACGTGGTGCCGCG-3′) when DNA was successively amplified for 30 cycles at 95°C for 60 seconds, 59°C for 30 seconds, and 72°C for 120 seconds. Mice carrying targeted null mutations in the Mmp2, Mmp8, Mmp13 or Mmp14 genes (Itoh et al., 1997; Zhou et al., 2000; Balbin et al., 2003; Stickens et al., 2004) were individually backcrossed into the FVB/n background for five generations, before they were intercrossed, and homozygous null genotypes were then generated and compared with littermate controls. Gene-switch K14-GLp65/tata.TGFβ1 mice were generated and maintained as described previously (Cao et al., 2002; Lu et al., 2004). Expression of the TGFβ₁ transgene (Tgfb1) was induced in 4-week-old bigenic mice by topical application of RU486 on ear skin, once a day for 3 days, at a dose of 20 μg/mouse (dissolved in 50 μl of sesame oil); mice were then analyzed on day 4. K14-HPV16 and MMTV-PyMT mice were maintained in the FVB/n strain and analyzed as described previously (Guy et al., 1992; Coussens et al., 1996).

Neutralization of TGFβ activity in vivo was accomplished by intraperitoneal (i.p.) injections of pan-specific TGFβ antibody (R&D Systems, MN; #AB-100; 1.0 mg/ml in sterile PBS, pH 7.4), at 5.0 mg/kg body weight, at 120, 96 and 24 hours before MO challenge. Control animals received normal rabbit IgG (R&D Systems; #AB-105-C). Five animals per cohort were injected and the experiment was repeated three times. GM6001 {N-[(2R)-2-hydroxyamidocarbonylmethyl-0-4-methylpentanoyl]-L-tryptophan ethylamide} (Galardy et al., 1994), a broad, class-specific MP inhibitor (Chemicon International, CA), was administered i.p., at 100 mg/kg body weight, as a 20 mg/ml slurry in 4.0% carboxymethylcellulose (CMC) in 0.9% PBS, daily, for 3 days prior to cutaneous challenge. Controls were treated with a daily injection of 4% CMC in PBS. Four animals per cohort were injected and the experiment was repeated four times. This concentration of GM6001 has been demonstrated to inhibit in vivo MP activity (Zheng et al., 2000). Pharmacological inhibition of ALK5 was achieved using the ALK5 kinase inhibitor [3-(pyridin-2-yl)-4-(4-quinonyl)]-1H-pyrazole (Calbiochem, San Diego, CA), which was administered i.p., at 1.0 mg/kg, in 2% DMSO in sterile PBS every other day for 6 days. Control mice received equivalent i.p. injections of the solvent vehicle DMSO. For biodistribution studies with dextran, mice were intravenously given 100 μl of lysine-fixable dextran 10,000 kDa Alexa Fluor 647 at 2.0 mg/ml (Molecular Probes) and 50 μl dextran 70,000 kDa tetramethylrhodamine at 4.0 mg/ml (Molecular Probes), and were studied 3 hours later. All analyses used age-matched mice and five mice per treatment group, and all experiments were repeated a minimum of three separate times. For all other experiments, analyses were conducted in triplicate on cohorts containing at least three mice and P values of <0.05 were considered significant. All animal experiments were conducted in accordance with procedures approved by the IACUC, UCSF.

Evans Blue (EB) dye (30 mg/kg in 100 μl PBS; Sigma-Aldrich, St Louis, MO) was injected into the tail vein of 7- to 8-week-old mice in all analyses, except for MMP14 null animals where, owing to early neonatal lethality, 17-day-old mice were injected i.v. with EB dye retro-orbitally, and assayed as described below. In some experiments, after 1 minute, 30 μl of 5% MO (phenyl isothiocyanate, 98%, Sigma-Aldrich) diluted in MnO (Sigma-Aldrich) was applied to the dorsal and ventral surfaces of the ear; the application process was repeated 15 minutes later. Isoflurane-anesthetized mice were photographed 30 minutes after the injection of EB dye. Anesthetized mice were then cardiac-perfused (see below), their ears removed, and blotted dry and weighed. EB dye was extracted from ears in 1 ml of formamide, for 12-48 hours at 60°C, and measured spectrophotometrically at 610 nm in a SpectraMax 340 (Molecular Devices, Sunnyvale, CA). Data are expressed as mean ± S.E.M. Comparisons of the amounts of dye extravasation were evaluated by Mann-Whitney statistical test with P values less than 0.05 considered significant. In some experiments, 5 minutes prior to the infusion of EB dye, shaved 5- to 7-week-old mice were injected intradermally (10 μl) with one of the following agents at the concentrations shown: VEGF₁₂₀ (R&D Systems), VEGF₁₆₄ (Chemicon International), histamine (Calbiochem) or serotonin (Sigma-Aldrich). The appearance of a blue spot was monitored for 30 minutes, at which time mice were euthanized, cardiac-perfused, photographed, and the area of skin surrounding the site of injection was excised (~5 mm²), photographed, and the EB dye extracted as above.

Isoflurane-anesthetized mice were injected with fluorescein-labeled Lycopersicon esculentum lectin (100 μl, 2 mg/ml; Vector Laboratories, Burlingame, CA) or rhodamine-labeled Ricinus communis agglutinin I (50 μl, 5.0 mg/ml; Vector Laboratories) into the femoral vein, as described (Thurston et al., 1996; Thurston et al., 1998). Two minutes after lectin injection, mice were perfused with fixative (1% paraformaldehyde plus 0.5% glutaraldehyde in PBS, pH 7.4, at 37°C) through the ascending aorta for 2 minutes to fix the vasculature and flush out non-adherent leucocytes. Confocal images were acquired on a Zeiss LSM 510 META NLO with an ultrafast, tunable Coherent Ti:Sa MIRA laser with Verdi pump for multi-photon excitation.

Immunodetection of α-smooth muscle actin was performed on tissue pieces following injection of Ricinus communis lectin and cardiac perfusion, as described above. Tissue pieces were fixed in 4% paraformaldehyde for 4 hours in the dark at 4°C, with gentle agitation, followed by several washes in 4°C PBS, before being permeabilized in 0.3% Triton X-100 overnight with gentle agitation at 4°C. Tissue pieces were then incubated with a Cy3-labeled anti-α-smooth muscle actin monoclonal antibody (mAb) (Sigma-Aldrich; Clone 1A4 #C6198, 1:500) diluted in 5% normal goat serum, 2.5% BSA, 0.3% Triton X-100 in PBS, overnight at 4°C on a rotating platform, followed by extensive washing in 4°C PBS and mounting with Vectashield (Vector Laboratories) mounting medium. Images were acquired on a Zeiss LSM 510 META NLO with an ultrafast, tunable Coherent Ti:Sa MIRA laser with Verdi pump for multi-photon excitation. Immunodetection of TGFβ in ear skin was performed on formalin-fixed, 5-μm, paraffin-embedded sections incubated with a mouse monoclonal anti-TGFβ antibody (R&D Systems; clone: 1D11) at a 1:50 dilution, revealed with biotinylated goat anti-mouse secondary antibody, at a 1:200 dilution, and counterstained in 1.0% methyl green and mounted with Cytoseal 60.

Immunofluorescence (IF) for αSMA and ALK5 was performed on 10-μm cryo-frozen tissue sections of ear skin that were fixed for 10 minutes with cold acetone, blocked for 30 minutes, and incubated with primary antibodies (ALK5, 1:50 dilution; Abcam #31013; αSMA clone 1A4, 1:100 dilution; Sigma) overnight at 4°C, followed by successive PBS washes and the application of secondary antibodies (1:500). Following further PBS washes, the sections were mounted with Vectashield media containing DAPI (Vector) and analyzed using a confocal microscope LSM 510 with a 63× oil objective, and images were then analyzed with a Zeiss LSM image examiner.

Immunodetection of MMP14 protein in collagen matrices was performed after incubation of cells in three-dimensional (3D) collagen gels for 18 hours without serum. The medium was then aspirated from each well and the cells were immediately fixed with 2.0% paraformaldehyde (2% PFA in PBS, pH 7.4, freshly prepared) for 20 minutes at room temperature (RT), then permeabilized with PBS containing 0.5% Triton X-100 for 10 minutes at 4 C, washed three times with PBS-glycine (130 mM NaCl, 7.0 mM Na₂HPO₄, 3.5 mM NaH₂PO₄, 10 mM glycine), and then blocked with 200 μl/well of IF buffer (130 mM NaCl, 7.0 mM Na₂HPO₄, 3.5 mM NaH₂PO₄, 0.1% BSA, 0.2% Triton X-100, 0.05% Tween-20) with 10% goat serum for 1 hour at RT. The primary rabbit polyclonal anti-MT1-MMP (hinge region) antibody (AB815, Chemicon), which was diluted at 1:250 in IF Buffer with 10% goat serum, was incubated overnight at 4°C. Samples were washed three times with IF buffer for 20 minutes each, followed by incubation with Alexa Fluor 488 goat anti-rabbit antibody, diluted 1:200 in IF buffer with 10% goat serum, for 45 minutes. Cells were then washed three times with IF buffer, cover slipped with the gel, and mounted in Vectashield with DAPI mounting medium, and allowed to air dry. Once dry, slides were observed by laser confocal microscopy using a Zeiss LSM META.

Tissue pieces (5 mm²) were collected from ears, or shaved back skin, following the injection (i.d.) of 10 μl of 10 ng VEGF₁₆₄ or 0.1% BSA in PBS. Tissues were pulverized in liquid nitrogen (N₂), followed by lysis in ice-cold buffer containing 20 mM Tris, pH 7.6, 150 mM NaCl, 1.0 mM EDTA, 50 mM NaF, 1.0% Triton X-100, 0.5% Na-deoxycholate, 0.1% SDS, 2.0 mM Na₂VO₄, 10 μg/ml aprotinin and 1.0 mM phenylmethylsulfonylfluoride, and centrifuged at 10,000 rpm (9503 × g) for 30 minutes at 4°C. The supernatants were recentrifuged at 10,000 rpm for 30 minutes at 4°C. Lysates were then incubated in a slurry of heparin-sepharose CL-6B (Pharmacia, Peapack, NJ) and incubated overnight, rocking at 4°C, and then centrifuged and equilibrated to 150 mM NaCl. Protein was dialyzed against PBS and quantified using the Bio-Rad protein assay system (Bio-Rad, Hercules, CA). Before immunoprecipitation, BSA was added to the pre-cleared lysates at 0.5%. Equal amounts of protein (1.0 mg) from lysates were used for immunoprecipitations and western blotting. Incubation of tissue lysate with goat anti-Flk-1 (Santa Cruz Biotechnology, Santa Cruz, CA) followed by protein-G sepharose beads was performed for 2 hours at 4°C. Immunoprecipitates were washed three times with 20 mM Tris (pH 7.6), 150 mM NaCl and 0.1% Triton X-100, and bound proteins were eluted by boiling in 1× SDS-PAGE sample buffer for 5 minutes, followed by electrophoresis on 10% SDS-PAGE under reducing conditions. The resolved proteins were transferred to a nitrocellulose membrane (BA-S85; Schleicher & Schuell BioScience, Inc., Keene, NH). Anti-phosphotyrosine PY-20 (Upstate Biotechnology, Lake Placid, NY) and anti-Flk-1 (Santa Cruz Biotechnology) antibodies were used on western blots. Immunodetection was performed by incubation with specific peroxidase-conjugated secondary antibodies followed by enhanced chemiluminescence (ECL) (Amersham Biosciences, Piscataway, NJ).

Tissues were pulverized in liquid N₂ and solubilized in lysis buffer containing 50 mM Tris, 75 mM NaCl, 10 mM EDTA, protease inhibitor cocktail mix without EDTA (Roche, Indianapolis, IN), 0.01 mg/ml aprotinin (Sigma-Aldrich), 0.1 mg/ml leupeptin (Sigma-Aldrich) and 10 mM phenylmethylsulfonyl fluoride (PMSF) (Sigma-Aldrich) using a 2.0 ml tissue grinder (Fisher Scientific, Pittsburgh, PA), followed by centrifugation at 10,000 × g for 30 minutes at 4°C. Protein concentration was determined using the Bio-Rad DC Protein assay reagent according to the manufacturer’s instructions (Bio-Rad). To determine total levels of TGFβ₁ in tissue lysates, a standard protocol for quantitative sandwich enzyme immunoassay was used. For ELISA analysis, a monoclonal antibody specific for active TGFβ₁, β₂ and β₃ (1.0 μg/ml in PBS; R&D Systems; MAB1835) was used to pre-coat maxisorb immunoplates (Nalge Nunc International, Rochester, NY) overnight at RT. Lysates (100 μg) were activated by adding 1.0 N HCl (1:25) and incubated for 1 hour at 4°C with gentle agitation. Acidified samples were neutralized by adding 1.0 N NaOH (1:25) and diluted with ELISA sample buffer (1× PBS, 0.05% Tween-20, 1.4% fatty acid-free BSA). Samples were then added to pre-coated maxisorb immunoplates and incubated for 3 hours at RT, which was followed by the addition of biotinylated anti-TGFβ₁ antibody (200 ng/ml; R&D Systems; BAF240) and incubation overnight at 4°C. Avidin-peroxidase conjugate (1:1000; Sigma-Aldrich) was added for 1 hour at RT followed by a 20-minute incubation with o-phenylenediamine (OPD) substrate at RT in the dark (Sigma-Aldrich). The reaction was stopped by addition of 1.0 M H₂SO₄ and absorbance was measured at 450 nm (570 nm for background corrections) on a Molecular Device Spectra Max 340. Recombinant human TGFβ₁ (R&D Systems) was used as a standard control. Six tissues samples per genotype were analyzed and all samples were analyzed in duplicate.

For immunoprecipitation of TGFβ, 4.2 mg of protein lysates were pre-cleared with protein A-agarose beads (Roche) for 1 hour at 4°C, followed by centrifugation at 3000 rpm (5 minutes). The supernatant was then incubated with 2.0 μg of antibody for TGFβ₁, β₂ and β₃ (R&D Systems; MAB1835) or MMP14 (Chemicon International; AB8102, catalytic domain; MAB3317, hemopexin domain) for 3 hours at 4°C in HNTG buffer (20 mM HEPES, pH 7.5, 150 mM NaCl, 0.1% Triton X-100, 10% glycerin, 10 mM Na pyrophosphate, 10 mM NaF, 1 mM Na-o-vanadate, 1 mM PMSF and 10 μg/ml aprotinin). After incubation with protein agarose G or A beads (Roche) for an additional hour at 4°C, the lysates bound to agarose beads were washed three times with HNTG buffer, and bound proteins were eluted by boiling in 1× reduced SDS-PAGE sample buffer for 5 minutes and then centrifuged at 13,000 rpm for 10 minutes. Tissue lysates (20 μg for LAP) or eluted immunoprecipitated complexes were separated by electrophoresis on 10% SDS-polyacrylamide gels, and transferred to nitrocellulose membranes overnight at 4°C. Membranes were blocked, incubated with primary antibodies for 1–2 hours at RT, washed, and then incubated with secondary antibodies (Bio-Rad; goat anti-rabbit- or goat anti-mouse-HRP conjugate 1:2000) or strepavidin-HRP conjugate (Sigma-Aldrich; 1:20,000) for 1 hour at RT. Membranes were then washed and developed by using an ECL kit (Amersham Biosciences). Biotinylated LAP antibodies (R&D Systems; BAF246, 1:1000), biotinylated anti-TGFβ₁ antibodies (R&D Systems; BAF240, 1:1000) and antibodies to MMP14 (Oncogene Science, Cambridge, MA, 1M397, 1:1000; Chemicon International, AB8104, 1:1000) were used for detection on membranes. For the loading control in LAP western analysis, a rat monoclonal antibody (AbCam, Cambridge, MA; YL1/2, 1:5000) against α-tubulin and a goat anti-rat-HRP (Pierce, Rockford, IL; 1:2000) were used.

TGFβ bioavailabilty in tissue lysates from mice, and in conditioned medium from MDA-MB-MMP14 cells grown in either +/+ or r/r collagen matrices, was assayed using mink lung epithelial cells (MLEC) transfected with a luciferase reporter gene under the control of the TGFβ-sensitive promoter (PAI-1), and maintained as described previously (Abe et al., 1994). PAI-1-MLEC cells (2.5×10⁴) were cultured in 96-well culture plates in 500 μl DMEM containing 10% FBS, and allowed to attach for 6 hours at 37°C. After two PBS washes, the medium was replaced with tissue lysate or conditioned medium. Tissue lysates were prepared as described above for ELISA. 50 μg of total proteins was diluted in incubation medium [high-glucose DMEM, 0.2% BSA (Fraction V, fatty acid-free; Calbiochem), penicillin/streptomycin, 2.0 mM L-glutamine, 250 μg/ml G-418 sulfate] and incubated with PAI-1-MLEC cells for 20 hours in the presence or absence of anti-TGFβ antibody (15 μg/ml; R&D Systems; clone 1D11). For conditioned medium analyses, MDA-MB-231 cells were grown in either +/+ or r/r collagen, as described above, and incubated in the presence or absence of GM6001 (10 μM; Chemicon International) or anti-TGFβ antibody (15 μg/ml). Conditioned medium (50 μl) was diluted in 50 μl of incubation medium and incubated with PAI-1-MLEC cells for 20 hours. Cells were washed twice with PBS, incubated with lysis buffer (Promega, Madison, WI), and luciferase activity was then measured using a Luciferase Assay System kit (Promega) following the manufacturer’s instructions. Relative luminescence units (RLU) were calculated as an average from triplicate wells per culture condition and repeated in triplicate.

Total RNA was extracted from shaved back skin or ear pieces with TRIzol reagent (Invitrogen, New York, NY), according to the manufacturer’s recommendations, by powdering fresh-frozen tissue samples in liquid N₂, homogenizing with a microtube pestle (USA Scientific, Ocala, FL), shearing by multiple passages through a syringe and 21-gauge needle (Becton Dickinson, Franklin Lakes, NJ), followed by chloroform extraction, isopropanol precipitation, and an ethanol wash. Northern blot analysis was performed using standard methods with 10 μg of total cellular RNA. Probes were generated by random primed labeling of DNA isolated from plasmids using standard methodology (Maniatis et al., 1982). Northern blots were hybridized at 65°C overnight in Church buffer (0.5 M sodium phosphate, pH 7.2, 1.0 mM EDTA, 7.0% w/vol SDS, 250 μg/ml tRNA), and subsequently washed at 62°C in 2× SSC containing 1.0% SDS. The probes used for hybridization were: a 335 bp fragment of Mmp2 (EMBL: M84324; position: 2053–2387 bp), a 335 bp fragment of Mmp14 (EMBL: NM_008608; position: 54–388 bp), a 669 bp fragment of Timp2 (EMBL: X62622; position: 2–670 bp), a 974 bp fragment of Tgfb1 (EMBL: M13177; position: 421–1395 bp) and a 207 bp fragment for 18S RNA as a loading control (EMBL: J00623; position: 13–219 bp). Hybridized filters were exposed overnight on phosphor screens, analyzed in a Phosphoimager (Molecular Dynamics, Sunnyvale, CA; Storm 860, ImageQuant 5.2 software), and then additionally exposed for 1–3 days on Kodak Biomax MS films (Kodak, Rochester, NY) with an intensifier screen at −80°C.

Shaved dorsal skin pieces isolated from 5–8-week-old mice were pulverized in liquid N₂ and solubilized in 500 μl buffer [0.25 M sucrose, 5 mM Tris, pH 7.5, protease inhibitor cocktail mix without EDTA (Roche), 0.25 mg/ml Pefablock (Roche), 0.01 mg/ml aprotinin (Sigma-Aldrich)] and centrifuged at 800 × g for 15 minutes at 4°C. Supernatants were then centrifuged for 1 hour at 100,000 × g at 4°C, and pellets were resuspended in 100 μl solubilization buffer and homogenized by sonication at 4°C. Protein concentration was determined using the Bio-Rad DC Protein assay reagent according to the manufacturer’s instructions (Bio-Rad). Lysate buffers were then exchanged with 10 mM Tris, pH 7.5, using Micro Bio-Spin chromatography columns (Bio-gelP-6; Bio-Rad). To determine gelatinolytic activity in tissue lysates, supernatant fractions (50 μg) were incubated with 400 ng DQ-gelatin (Molecular Probes) in reaction buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 5.0 mM CaCl₂, 0.2 mM sodium azide and 0.05% BrJ35) at 37°C. Reactions were analyzed for 5 hours at 37°C and fluorescence measured (excitation 485 nm, emission 530 nm) every 3 minutes using a microplate spectrofluorometer (SpectraMax Gemini EM, Molecular Devices) operated by SoftMax Pro 4.1 software. Values shown represent the mean ± S.E.M. from three tissue pieces and are representative of analyses performed in triplicate. Each experiment was repeated three independent times.

Tissue samples (ear) from 5–8-week-old mice were weighed and homogenized (1:8 weight to volume) in lysis buffer containing 50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 0.1% NP-40, 0.5% deoxycholate and 0.1% SDS. Soluble and insoluble extracts were separated by centrifugation (10,000 × g) and subsequently stored at −80°C. Equivalent amounts of soluble extract were analyzed by gelatin zymography (Herron et al., 1986a) on 10% SDS-polyacrylamide gels co-polymerized with substrate (1.0 mg/ml of gelatin) in sample buffer (2% SDS, 50 mM Tris-HCl, 10% glycerol, 0.1% Bromophenol Blue, pH 6.8). After electrophoresis, gels were washed three times for 30 minutes in 2.5% Triton X-100, then washed three times for 15 minutes in ddH₂O, incubated overnight at 37°C in 50 mM Tris-HCl and 10 mM CaCl₂ (pH 8.2), and then stained in 0.5% Coomassie Blue and destained in 20% methanol, 10% acetic acid. Negative staining indicates the location of active protease bands. Exposure of pro-enzymes within tissue extracts to SDS during the gel separation procedure leads to activation without proteolytic cleavage (Herron et al., 1986b).

To prepare collagen gels for culture experimentation, mouse tail collagen was purified as described previously (Miller and Rhodes, 1982) and quantified by determination of hydroxyproline content (Woessner, 1961; Bergman and Loxley, 1963). Subsequently, eight volumes of +/+ and r/r collagen (4.4 mg/ml) were neutralized by the addition of one volume of 10× PBS containing 0.005% Phenol Red and one volume of NaOH. 50 μl of MDA-MB-231 breast carcinoma cells expressing a full-length human MMP14 cDNA (Tam et al., 2004), at 5×10⁶ cells/ml in serum-free DMEM, were added to 200 μl of neutralized +/+ and r/r collagen. The collagen/cell suspensions were mixed thoroughly and then four 50 μl aliquots were added per well into a 96-well culture dish (Corning, Corning, NY) and incubated at 37°C for 1 hour to allow collagen polymerization. 100 μl of DMEM containing 10% fetal bovine serum was then added to cells and incubated at 37°C for 18 hours. Collagen gels were washed with 200 μl serum-free DMEM, and cells were then incubated in 100 μl serum-free DMEM containing human proMMP2, since the MDA-MB-231 cells essentially express no MMP2. Conditioned medium was harvested after 48 hours and collagen gels were washed in 200 μl of PBS. 50 μl of non-reducing SDS-PAGE sample buffer was then added to collagen gels to extract the collagen-bound MMP2 and then, after collection, the sample buffer was used to bring the total volume to 200 μl. Equivalent amounts of supernatants and collagen-bound MMP2 extracts were analyzed by gelatin zymography and incubated for 4 hours at 37°C.

MMTV-PyMT mice (Guy et al., 1992) (approximately 100 days old) were treated with the ALK5 inhibitor [3-(Pyridin-2-yl)-4-(4-quinonyl)]-1H-pyrazole (Calbiochem), which was given i.p. every other day at 1.0 mg/kg in 2% DMSO in sterile PBS. Control mice received vehicle alone. On day 6 of treatment, tumors were imaged in live mice as described previously (Egeblad et al., 2008). Briefly, the mice were anesthetized and the inguinal mammary gland containing a tumor was surgically exposed. Images from live mice were acquired using a Fluor 10×/0.5 NA lens and a micro-lensed spinning disk confocal microscope (Solamere Technology Group, Salt Lake City, UT) equipped with argon and krypton lasers (Dynamic Lasers, Salt Lake City, UT). Images were collected with an ICCD camera (XR-Mega-10EX S-30, Stanford Photonics, Palo Alto, CA). The anesthetized mice were injected i.v., into the tail vein, with 100 μl sterile PBS containing 1.0 mg/ml 10 kDa Alexa Fluor 647-conjugated dextran, 2.0 mg/ml 70 kDa rhodamine-conjugated dextran and 0.4 μM Qtracker 705 non-targeted quantum dots (all from Invitrogen). Five animals in each group were imaged and the tumors were between 6–11 mm in diameter.

From the acquired image sets, maximum intensity projections of 16-μm optical sections were generated using Bitplane Imaris (version 5.7 for Windows X64). To determine dextran leakage, these sets were analyzed further using Improvision Volocity (version 4.1.0 for Windows). Pixel intensities in the quantum dot channel that were below 125% of the mean background intensity were used to define the extravascular space (negative for quantum dots). Leakage of 10 kDa or 70 kDa dextran to the extravascular space was defined as the percentage of the extravascular space with pixel intensities above 150% of the mean background intensity levels. From each mouse, 3–5 fields of view (imaged in parallel) were analyzed. The average percentage of extravascular leakage for the five mice in each group is reported. Differences in dextran leakage between controls and ALK5 inhibitor-treated mice were compared using Student’s t-test (two-tailed, unequal variance) using GraphPad Prism 4 software.

Statistical analyses were performed using GraphPad Prism and/or InStat software. The specific tests used were Student’s t-test, Mann-Whitney (unpaired, nonparametric, two-tailed) test, and unpaired t-test with Welch’s correction. P values <0.05 were considered statistically significant.

The authors extend thanks to Dr Rosemary Akhurst for providing MLE-PAI-1 cells and Smad3 null mice; Dr Xiao-Jing Wang for providing bigenic TGFβ transgenic mice; Dr Suneel S. Apte for MMP14^ko/+ (FVB/n) mice; Dr Carlos Lopez-Otin for MMP8^ko/ko mice; Dr Mitsunobu R. Kano for information on ALK5 inhibitor dosing; the UCSF Helen Diller Family Comprehensive Cancer Center Mouse Pathology and Laboratory for Cell Analysis cores for technical expertise; and Drs Thea Tlsty, David Bates, Douglas Hanahan, Donald McDonald, Bonnie Sloane and Gabriele Bergers, and members of the Coussens laboratory for valuable comments and critical discussions. We also thank Drs David DeNardo and Pauline Andreu for critical reading of the manuscript. The authors acknowledge the NIH National Institute of Aging for allowing the purchase of aged C57BL/6 mice, and support from the Centre Anticancéreux près l’Université de Liège-Belgium (N.E.S.); the Dutch Cancer Society (L.v.K.); AR44815 (S.M.K.); The Canadian Institutes in Health Research and the National Cancer Institute of Canada (C.M.O.); NIH grants CA72006, AI053194 and CA105379 (Z.W. and M.E.); and CA140943, CA72006, CA98075, CA94168, a National Technology Center for Networks and Pathways (U54 RR020843), and a Department of Defense Breast Cancer Research Program Era of Hope Scholar Award (W81XWH-06-1-0416) to L.M.C. C.M.O. is supported by a Canada Research Chair in Metalloproteinase Biology. N.B. is supported by NIH grants U54CA126552 and NS044155. Deposited in PMC for release after 12 months.