Live-cell imaging demonstrates extracellular matrix degradation in

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Research Article

Live-cell imaging demonstrates extracellular matrix degradationin association with active cathepsin B in caveolae of endothelialcells during tube formation

Dora Cavallo-Medveda,b,⁎ , Deborah Rudya, Galia Blumc, Matthew Bogyoc,d,Dejan Caglice, Bonnie F. Sloanea,b

aDepartment of Pharmacology, Wayne State University, School of Medicine, 540 East Canfield, Detroit, MI 48201, USAbBarbara Ann Karmanos Cancer Institute, Wayne State University, School of Medicine, Detroit, MI 48201, USAcDepartment of Pathology, Stanford University, School of Medicine, Stanford, CA 94305, USAdDepartment of Microbiology and Immunology, Stanford University, School of Medicine, Stanford, CA 94305, USAeDepartment of Biochemistry, and Molecular and Structural Biology, Jozef Stefan Institute, Jamova 39, 1000 Ljubljana, Slovenia

A R T I C L E I N F O R M A T I O N

⁎ Corresponding author. Department of PharmFax: +1 313 577 6739.

E-mail address: [email protected] (D

0014-4827/$ – see front matter © 2009 Elseviedoi:10.1016/j.yexcr.2009.01.021

A B S T R A C T

Article Chronology:

Received 24 September 2008Revised version received11 January 2009

Accepted 22 January 2009Available online 3 February 2009

Localization of proteases to the surface of endothelial cells and remodeling of the extracellularmatrix (ECM) are essential to endothelial cell tube formation and angiogenesis. Here, wepartially localized active cathepsin B and its cell surface binding partners, S100A/p11 (p11) ofthe annexin II heterotetramer (AIIt), to caveolae of human umbilical vein endothelial cells

(HUVEC). Via a live-cell proteolysis assay, we observed that degradation products of quenched-fluorescent (DQ)-proteins (i.e. gelatin and collagen IV) colocalized intracellularly with caveolin-1(cav-1) of HUVEC grown in either monolayer cultures or in vitro tube formation assays. Activity-based probes that bind covalently to active cysteine cathepsins and degradation products of DQ-collagen IV partially localized to intracellular vesicles that contained cav-1 and active cysteinecathepsins. Biochemical analyses revealed that the distribution of active cathepsin B in caveolarfractions increased during in vitro tube formation. Pro-uPA, uPAR, MMP-2 and MMP-14, whichhave been linked with cathepsin B to ECM degradation pathways, were also found to increase incaveolar fractions during in vitro tube formation. Our findings are the first to demonstratethrough live-cell imaging ECM degradation in association with active cathepsin B in caveolae ofendothelial cells during tube formation.

© 2009 Elsevier Inc. All rights reserved.

Keywords:

Endothelial cellsCaveolaeCathepsin BProteasesECM degradationAngiogenesisFunctional imaging

Introduction

Angiogenesis, the formation of new blood vessels from the pre-existing vasculature, is a process in which stimulated endothelialcells remodel extracellular matrix (ECM), migrate through theECM, proliferate, differentiate and eventually form endothelial

acology, Wayne State Univ

. Cavallo-Medved).

r Inc. All rights reserved.

tubules capable of blood transport [1]. Proteases of at least threeclasses (serine, cysteine, and metallo-) [including matrix metallo-proteinases (MMPs), members of the urokinase plasmin(ogen)system and cysteine cathepsins] play crucial roles in angiogenesis(for review see [2]). Proteases participate in the angiogenic processby generating both pro- and anti-angiogenic factors from ECM

ersity, School of Medicine, 540 East Canfield, Detroit, MI 48201, USA.

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http://dx.doi.org/10.1016/j.yexcr.2009.01.021

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proteins [3–5] and by processing growth factors and receptors,including integrins [6,7]. For example, cleavage of type IV collagenby endothelial cell MMPs results in the exposure of cryptic αvβ3-integrin binding sites, thus promoting angiogenesis [7]. Urokinaseplasminogen activator (uPA) converts plasminogen to plasminwhich can be further processed to generate angiostatin, an anti-angiogenic compound [8]. Cathepsin B, a lysosomal cysteineprotease, regulates the “angiogenic switch” [9] by initiatingproteolytic cascades involved in ECM degradation that promoteangiogenesis [10], or conversely, by cleaving collagen XVIII andgenerating endostatin, an anti-angiogenic factor [11]. Overall, thebalance of protease activity and the interaction of proteases withthe ECM contribute to regulation of the angiogenic process.

ECMremodeling anddegradationby tumor cells are facilitatedbythe translocation of proteases to cell surfaces and their secretion intothe extracellular milieu [12–15]. Many proteases implicated in ECMremodeling and degradation are also associatedwith caveolae [12], alipid-rich region of the plasma membrane involved in endocytosis,cholesterol transport and cell signaling events (for reviews, see[16,17]). These proteases include uPA and its receptor uPAR,cathepsin B and its cell surface binding protein S100A10/p11 (p11)[the light chain of the annexin II heterotetramer (AIIt)], MMP-2 andMMP-14 [12]. In colorectal carcinoma cells, downregulation ofcaveolin-1 (cav-1), themain structural protein of caveolae, decreasesdistribution of cathepsin B anduPA to caveolae and ECMdegradationby these cells [18]. The association of proteases with caveolae ofendothelial cells is intriguing since caveolae are involved inangiogenesis (for review, see [19]).Duringendothelial cellmigration,cav-1 localizes to the rear of migrating cells [20–22]. Moreover,downregulation of cav-1 impedes endothelial cell polarization anddirectional movement [20]. Indeed, knockdown of cav-1 suppressestube formationbyendothelial cells andalso reduces vessel formationin the chicken chorioallantoic membrane assay [23]. Thus, wehypothesize that compartmentalization of proteases to caveolae inendothelial cells modulates cell migration, proteolysis of ECMproteins and tube formation.

In the present study, we identified active cathepsin B as wellas its cell surface-binding partner, S100A10/p11 (p11), incaveolae of human umbilical vein endothelial cells (HUVEC):an association that is augmented during in vitro endothelial tubecell formation. Using a live-cell proteolysis assay in combinationwith in vitro tube formation assays and cysteine cathepsinactivity-based probes, we observed extracellular degradation ofECM proteins (i.e., type IV collagen) by migrating HUVEC andintracellular colocalization of ECM degradation products withcav-1 in vesicles containing active cysteine cathepsins. Thesedata suggest an association of active cathepsin B with caveolaeand degradation and remodeling of the ECM during endothelialcell migration and tube formation.

Materials and methods

Materials

M199 medium, heparin, N-Octyl β-D-glucopyranoside, 2-[N-morpholino]ethanesulfonic acid (MES), methyl-β-cyclodextrin(MβCD), and all other chemicals unless otherwise stated werefrom Sigma (St. Louis, MO); fetal bovine serum (FBS), Lipofectinreagent, dye-quenched fluorescent (DQ)-gelatin and DQ-collagen

IV were from Invitrogen (Carlsbad, CA); bovine endothelial cellgrowth factor (bECGF) was from Roche Applied Science (India-napolis, IN); polyclonal anti-caveolin (610059), monoclonal anti-annexin II (mAb 5) and monoclonal anti-p11 (mAb 148)antibodies were purchased from BD Biosciences (Bedford, MA);polyclonal anti-cathepsin B [24] antibodies were produced,affinity-purified and characterized in our laboratory; polyclonalanti-β1 integrin antibodies were a kind gift from Dr. KennethYamada (National Institutes of Health/National Institute of Dentaland Craniofacial Research, Bethesda, MD); polyclonal anti-uPA(ab20046) and polyclonal anti-uPAR (ab27423) antibodies werefrom Abcam (Cambridge, MA); polyclonal anti-MMP-14 (AB815)antibodies were from Chemicon (Temecula, CA); horseradishperoxidase-labeled goat anti-rabbit and goat anti-mouse IgG werefrom Pierce (Rockford, IL); purified MMP-2 and MMP-9 enzymeswere a kind gift from Dr. Rafael Fridman (Wayne State University,Detroit, MI); cysteine cathepsin activity-based probes, GB-111-FLand GB-123, were recently described in references [25] and [26],respectively; Cultrex was from Trevigen (Gaithersberg, MD);acrylamide and nitrocellulose membranes were from BioRad(Hercules, CA); protein markers and chemiluminescent westernblotting detection kits were from Amersham Pharmacia Biotech(Piscataway, NJ); benzyloxycarbonyl-L-arginyl-L-arginine-4-methyl-7-coumarylamide (Z-Arg-Arg-NHMec) was from Bachem(Torrance, CA); and Amicon Ultra-4 10 K centrifugal filters werefrom Millipore (Bedford, CA).

Cell culture

HUVEC from the American Type Culture Collection (ATCC) (Rock-ville, MD) were grown on 1.5% (w/v) gelatin-coated tissue culturelab-ware in endothelial cell M199 media supplemented with40 μg/ml bECGF, 100 μg/ml heparin, antibiotics (penicillin/streptomycin) and 10% (v/v) FBS (as recommended by the ATCC)in 5% CO2/humidified atmosphere at 37 °C unless otherwise stated.

Transient transfection of HUVEC with cav-1-mRFP expressionvector (a kind gift from Drs. Radu V. Stan and Steve F. Dowdy,University of California at San Diego, San Diego, CA) wasperformed using Lipofectin reagent according to the manufac-turer's instructions.

In vitro tube formation assay

In vitro tube formation assays were performed in 35 mm tissueculture dishes coated with 200 μl of Cultrex [i.e., reconstitutedbasement membrane (rBM)] that was allowed to solidify at 37 °Cfor 10 min. Thereafter, 2×105 cells were seeded onto the Cultrexin M199 media containing 2% (v/v) FBS and incubated at 37 °C for16–18 h to allow formation of tubular structures [27,28].

Preparation of caveolae-enriched fractions

Caveolae-enriched fractions were isolated by both non-detergentand detergent based methods. The non-detergent method wasperformed as previously described [18,29]. Briefly, cells from four100 mm dishes were washed twice with PBS at 4 °C and lysed in2 ml 500 mM sodium carbonate buffer, pH 11.0. Cell lysates werehomogenized on ice by 10 strokes in a loose-fitting Douncehomogenizer and then sonicated on ice in a 50 W ultrasonicatorthree times for 20 s each. Cell homogenates were adjusted to 45%

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(w/v) sucrose by mixing them with 2 ml of 90% (w/v) sucroseprepared inMES-buffered saline (25mMMES, pH 6.5, 0.15mNaCl)and then placed on the bottom of a 12 ml ultracentrifuge tube. Thehomogenate was then overlaid with a 5–35% sucrose gradientcomprised of 4 ml of 35% (w/v) sucrose (in MES-buffered salinecontaining 250 mM sodium carbonate) and 4 ml of 5% (w/v)sucrose (in MES-buffered saline containing 250 mM sodiumcarbonate). Gradients were subjected to ultracentrifugation at185,000 ×g in a SW41 rotor (Beckman Instruments) for 18–20 h at4 °C. One ml fractions were collected from the top of the gradientand equal volume aliquots were analyzed by SDS-PAGE andimmunoblotting. Where indicated, cells were treated overnightwith 1 μM GB111-FL, a quenched activity-based probe for cysteinecathepsins [25], prior to caveolae isolation. SDS-PAGE gels wereimaged on a Typhoon Imaging Scanner (Amersham Biosciences,Pittsburgh, PA) to detect bound probe.

Caveolae-enriched fractions could not be isolated from HUVECgrown on rBM for in vitro tube-formation assays using the non-detergent based method described above because the amount ofrBM (131.2 mg in 8 ml) required for sufficient cells, i.e., four100mmdishes, prevented proper preparation of sucrose gradients.Therefore, we employed a successive detergent based method thatseparates Triton X-100-soluble from -insoluble membrane com-ponents [30]. Cells were incubated in lysis buffer containing 1%Triton X-100 in the absence of octylglucoside for 30 min on ice,collected and centrifuged at 14,000 ×g for 15 min at 4 °C. Thesupernatant, hereafter termed the Triton-soluble fraction, wasrecovered. The insoluble pellet was resuspended in an equalvolume of lysis buffer containing both 1% Triton X-100 and 60 mMoctylglucoside, incubated on ice for 30 min, passed 10 timesthrough a syringe with a 20-gauge needle and centrifuged for10 min at 14,000 ×g at 4 °C. The supernatant, hereafter termed theTriton-insoluble fraction, was recovered and the final volumeadjusted to that of the Triton-soluble fraction. Caveolae wereenriched in the Triton-insoluble fraction as was verified by treatingcells with 10 mM MβCD, a cholesterol sequestering drug thatdisrupts caveolae, at 37 °C for 1 h prior to successive detergentextraction of membranes.

Preparation of conditioned media

Equal numbers of cells were serum-starved overnight, media werecollected, centrifuged at 800 ×g to remove whole cells, re-centrifuged at 2000 ×g to remove cell debris, and concentratedto equal final volumes.

SDS-PAGE and immunoblot analysis

Sampleswere equally loaded and separated by SDS-PAGE (10%,12%or 15% acrylamide) and transferred onto nitrocellulosemembranesunder either reducing or non-reducing conditions. Immunoblot-ting was performed with primary antibodies against cathepsin B(1:4000), caveolin (1:5000), annexin II (1:5000), p11 (1:5000),uPA (5 μg/ml), uPAR (5 μg/ml), β1-integrin (1:3000) and MMP-14(1:1000) and secondary antibodies conjugated with horseradishperoxidase (1:10,000) in Tris-buffered saline wash buffer (20 mMTris, pH 7.5, 0.5 M NaCl) containing 0.5% Tween 20 and 5% (w/v)non-fat dry milk. After washing, bound antibodies were detectedby enhanced chemiluminescence according to the manufacturer'sinstructions.

Gelatin zymography

Subcellular fractions and conditioned media samples were equallyloaded and applied without heating or reduction to a 10% SDS-polyacrylamide gel containing 1 mg/ml of gelatin [31]. Afterelectrophoresis, the gel was washed twice for 15 min each timewith 2.5% Triton X-100, followed by a 20 min wash with distilledwater, and incubated overnight at 37 °C in 50 mM Tris buffer, pH7.8, containing 5 mM CaCl2 and 0.05% Brij35. The gel was stainedwith a solution of 0.5% Coomassie brilliant blue R-250 in 50%methanol and 10% acetic acid and then de-stained in a 50%methanol and 10% acetic acid solution. Clear bands represent areasof proteolytic activity. Purified MMP-2 and MMP-9 were loadedseparately as positive controls.

Cathepsin B activity assay

Cathepsin B activity was measured in subcellular fractions andconditionedmedia as previously described [32]. Briefly, to measureintracellular cathepsin B activity 50 μl of sample was incubatedwith 300 μl of activator buffer (5mMEDTA,10mMDTT, pH 5.2) for15 min at 37 °C. In a 96 well flat-bottom black microtiter plate,100 μl of cell lysate/activator buffer mixture was added to 200 μl ofassay buffer [Hank's balanced-salt solution lacking sodiumbicarbonate and phenol red and containing 0.6 mM CaCl2,0.6 mM MgCl2 and 25 mM piperazin-N-N′-bis[2-ethanesulfonicacid] (disodium salt), pH 7.3] containing 150 μM Z-Arg-Arg-NHMec substrate. The enzymatic assay was carried out at pH 6.0.Fluorescence was measured in triplicate at one-minute intervalsfor 30 min using a Tecan SpectraFluor Plus plate-reader at anexcitation of 360 nm and an emission of 465 nm. Tomeasure latentor procathepsin B activity in media samples, autoactivation ofprocathepsin B was accelerated in the presence of dextran sulfate[33]. Briefly, 50 μl of conditioned media was mixed with 300 μl ofcitrate buffer (20 mM citrate buffer, pH 4.6, 10 mM DTT and 25 μg/ml dextran sulfate) and incubated for 45 min at 37 °C. Then 100 μlof media-citrate buffer solutionwas mixed in a 96 well flat-bottomblack microtiter plate with 200 μl of 150 μM Z-Arg-Arg-NHMecsubstrate prepared in 200 mM sodium phosphate buffer, pH 6.8.Fluorescence was read in triplicate at one-minute intervals over a30 min period at an excitation of 360 nm and an emission of465 nm. Cathepsin B activity was recorded as pmol/min/cellnumber. Cells were counted using a hemocytometer prior tosetting up overnight experiments. Statistical significance wasdetermined by a two-tailed t-test with assumed equal variance.In the figures, ⁎ represents a p value less than 0.05 and ⁎⁎

represents a p value less than 0.01.

Immunocytochemistry

Cells were grown on coverslips for 16–24 h, permeabilized withsaponin, fixed with cold methanol for 5 min and then blocked for45min by incubatingwith PBS containing 2mg/ml BSA. Cells werethen washed with PBS and incubated with primary antibody(rabbit anti-human caveolin or preimmune rabbit IgG) for 2 h atroom temperature according to our published procedures [34].After washing with PBS, the cells were incubated for 1 h with Texasred-conjugated affinity-purified donkey anti-rabbit IgG containing5% normal donkey serum (Jackson ImmunoResearch). Cells werethen washed, mounted upside-down with Slow Fade anti-fade


reagent (Invitrogen Life Technologies) on glass slides and observedwith a Zeiss 510 LSM confocal microscope.

Live cell proteolysis assay

Proteolysis by live cells was assessed as previously described[35,36]. Briefly, glass coverslips were coated with either 50 μl of1.5% (w/v) gelatin containing 25 μg/ml DQ-gelatin and incubatedon ice for 15 min to solidify or 50 μl of 16.4 mg/ml Cultrexcontaining 25 μg/ml DQ-collagen IV and incubated for 10 min at37 °C to solidify. Subsequent steps were carried out at 37 °C. Cells(2.5×104) were seeded onto coated coverslips and incubated for60 min. Medium containing 2% FBS was added to the cells andincubated for 16–18 h. Proteolysis of DQ-gelatin and DQ-collagenIV (green fluorescence) was observed in live cells with a Zeiss LSM510 META NLO microscope with 10× and 40× water immersionobjectives. Time series analyses of DQ-collagen IV degradationwere performed with a Zeiss LSM 510 META NLO microscopeequippedwith a controlled environmental chamber thatmaintainsa 5% CO2/humidified atmosphere at 37 °C. Where indicated, assayswere conducted in the presence of 1 μM GB-123, a non-quenchedcysteine cathepsin activity-based probe, for 16–18 h. Cells werewashed 3 times with PBS and incubated in complete growthmedium in the absence of the probe immediately prior to imaging.

Results

Gelatin and collagen IV degradation products localize tovesicles containing cav-1 in HUVEC

Remodeling of the ECM is a process critical to the formation oftubular structures by endothelial cells. To analyze this process, weemployed a well-established live-cell proteolysis assay [35] andcompared HUVEC grown on coverslips coated with gelatincontaining DQ-gelatin with HUVEC grown on coverslips coatedwith rBM containing DQ-collagen IV. Following 16 h of incubation,live cells were imaged by confocal microscopy for degradationproducts (green fluorescence) of the two DQ-protein substrates.On gelatin, HUVEC grew as a monolayer of individual cells (Fig. 1A)and degradation products of DQ-gelatin were observed primarilyintracellularly in the perinuclear region (Fig. 1B). In comparison,HUVEC grown on rBM were stimulated to migrate and form largemulticellular structures often referred to as “cord-like”, “tube-like”or “tubular” structures (Fig. 1C), thus mimicking angiogenesis invitro, a process hereafter referred to as tube formation [27,28].Degradation products of DQ-collagen IV were localized bothintracellularly and pericellularly (i.e., on the cell membrane andimmediately outside the cell) (Fig. 1D).

To assess whether cav-1, the main structural protein forcaveolae, is associated with ECM degradation, we first immunos-tained HUVEC grown in monolayer versus tube-like structures forendogenous cav-1. Under both conditions, endogenous cav-1(green fluorescence) was localized both within intracellularperinuclear vesicles and along the surface of HUVEC cells (Figs.1E–H). We were unable to determine whether endogenous cav-1colocalized with degradation products of DQ-substrates as thefixation required for immunostaining interferes with the fluores-cence emitted by the DQ-substrate degradation products. There-fore, we performed a live-cell proteolysis assay with cav-1-mRFP

transfected HUVEC cells. The images in Figs. 2A and B illustrate thatcav-1-mRFP (red fluorescence) and DQ-substrate degradationproducts (green fluorescence) colocalized in a perinuclear punc-tate pattern, a pattern similar to that observed for the DQ-substratedegradation products in Fig. 1, suggesting that ECM degradation inHUVEC grown as a 2D monolayer and as 3D tube-like structuresoccurs within vesicles containing caveolin-1, or that ECM degrada-tion products are internalized via caveolae.

Cathepsin B and its cell surface binding partner partiallyco-localize to caveolae of HUVEC

We have previously established that cathepsin B is present incaveolae of human colorectal carcinoma cells [34]. Here, weexamined the colocalization of cathepsin B with cav-1 in caveolaeof HUVEC. Isolation of caveolae from HUVEC in monolayer culturewas performed using a well-established protocol that is based onthe insolubility of caveolae to carbonate extraction and theirspecific buoyant density in equilibrium sucrose gradients [29].Cav-1, a marker protein for caveolae, was distributed in fractions4–5, hereafter denoted as the caveolae fractions (Fig. 3A). Matureand thus active forms of cathepsin B were also present in thecaveolae fractions: 31 kDa single chain cathepsin B and 25/26 kDaheavy chain of double chain cathepsin B. The majority ofcathepsin B was isolated in dense fractions 8–11, which containlysosomes as identified using the lysosomal marker proteinsLAMP-1 and β-galactosidase [34] (data not shown). To confirmthat cathepsin B in the caveolae fractions was active, prior tosubcellular fractionation, we pretreated HUVEC with a FITC-labeled activity-based probe (GB111-FL) that selectively andcovalently binds the active sites of two cysteine cathepsins,cathepsin B and cathepsin L [25]. SDS-PAGE analysis and laserscanning imaging of subcellular fractions revealed only activecathepsin B (31 kDa) in the caveolae fractions (fractions 4 and 5)of HUVEC (Fig. 3B). Active cathepsin B [25] and active cathepsin L(28 kDa) were present in the dense or lysosomal fractions(fractions 9 and 10). The presence of active cathepsin B incaveolae of HUVEC would be consistent with a functionalsignificance for cathepsin B in these membrane microdomainsas has been shown in human colon carcinoma cells [34]. We havepreviously identified the light chain of AIIt, p11, as a cell surfacebinding partner for procathepsin B in human breast carcinomaand glioma cells [37] and determined that cathepsin B and p11cosediment in caveolae fractions of HCT116 human coloncarcinoma cells [34]. Here, we found p11 along with annexin II,the heavy chain of AIIt, localized to the caveolae of HUVEC (Fig.3A). This suggests that the association of cathepsin B withcaveolae may be mediated by its binding partner p11.

Distribution of cathepsin B to caveolae of HUVEC increasesduring endothelial cell tube formation

We further examined the translocation of cathepsin B to caveolaein HUVEC cells during endothelial cell tube formation. Due to thelarge volume of rBM required to generate sufficient HUVEC tube-like structures for isolation of caveolae, we were not able to usesodium carbonate extraction and subcellular fractionation on asucrose gradient to isolate caveolae. Therefore, we used analternative approach of successive detergent extractions [30] thatis based on the insolubility of caveolae in 1% Triton X-100. Cells

Fig.1 – Gelatin and collagen IV degradation byand localization of endogenous cav-1 inHUVEC grown as amonolayer versus tube-likestructures. Equal numbers of HUVECwere grown for 16 h on glass coverslips coated with gelatin alone (A), gelatin containing 25 μg/ml DQ-gelatin (B), rBM (Cultrex) alone (C), or rBM containing 25 μg/ml DQ-collagen IV (D). After 16 h, confocal images were taken oflive cells [differential interference contrast (DIC)] and DQ-substrate degradation products (green), which are present intracellularly(arrow) in HUVEC grown on gelatin (B) and both intracellularly (arrow) and pericellularly (arrowhead) in HUVEC grown on rBM (D).Immunostaining for endogenous cav-1 using rabbit anti-human caveolin antibodies was performed on fixed and permeabilizedHUVEC cells grown on gelatin (E and F; two different fields of view) or rBM (G andH; two different fields of view). Themorphology ofHUVEC inmonolayer culture is apparent from the immunostaining for tubulin (red) usingmouse anti-human tubulin antibodies (Eand F). Staining for caveolin is observed in the perinuclear region (arrow) and at the cell surface (arrowhead). N, nucleus. Bar, 20 μm.


grown as 2D monolayers and 3D tube-like structures werefractionated into Triton-soluble (TS) and Triton-insoluble (TI)fractions; lipid rich microdomains including caveolae are localized

to TI fractions whereas proteins resident to other cellularorganelles including lysosomes are found in the TS fraction [30].The advantage of this method of caveolae isolation over the

Fig. 2 – Intracellular colocalization of cav-1 with gelatin and collagen IV degradation products in HUVEC. Equal numbers of HUVECtransfected with cav-1-mRFP construct were grown for 16 h on glass coverslips coated with either gelatin containing 25 μg/mlDQ-gelatin (A) or rBM (Cultrex) containing 25 μg/ml DQ-collagen IV (B). After 16 h, confocal images were taken of live cells (DIC),cav-1-mRFP (red) and DQ-substrate degradation products (green). DQ-substrate degradation products (green) are presentintracellularly inHUVECgrownongelatinandbothintracellularlyandpericellularly inHUVECgrownonrBM(see inserts). Colocalizationof cav-1-mRFP and DQ-substrate degradation products appears yellow in the merged images. N, nucleus. Bar, 20 μm.


carbonate extraction method described above is that it preservesthe stability and thus activity of proteases allowing us to measureenzymatic activity following fractionation. Cav-1 served as apositive control for the presence of caveolae in the TI-fraction(Fig. 3C): however, it was also present in the TS-fraction since it is

not exclusive to caveolae, but is found in the ER and Golgi prior toits association with lipids and formation of caveolae at the cellsurface [38]. To confirm that caveolae were present in the TI-fraction, we treated HUVEC for 1 h prior to successive detergentextractionwith 10 mMMβCD, a cholesterol sequestering drug that

Fig. 3 – Increased distribution of active cathepsin B and p11 to caveolae of HUVEC cells during tube formation. HUVEC grown as amonolayer were subjected to subcellular fractionation on a sucrose gradient following homogenization in sodium carbonate buffer,pH 11.0. One ml fractions were collected from the top of the gradient and equal volume aliquots of fractions 2–11 were analyzed bySDS-PAGE and immunoblotted with anti-caveolin, anti-cathepsin B, anti-annexin II and anti-p11 antibodies (A). Aliquots fromcaveolae fractions 4 and 5 (Cav) andnon-caveolae fractions 9 and 10 (non-Cav) isolated fromHUVEC treatedwith 1 μMGB111-FLwereanalyzed by SDS-PAGE and imaged using a Typhoon laser scanner. Cathepsin B (31 kDa) and cathepsin L (28 kDa) were labeled (B).Equal numbers of HUVEC were grown on gelatin (mono) or rBM (tube) for 18 h and caveolae were isolated by successive detergentextraction as described in Materials and methods. Triton-soluble (TS) represents non-caveolae fractions and Triton-insoluble (TI)represents caveolae fractions. Equal volume aliquots of each fraction were analyzed by SDS-PAGE and immunoblotted with anti-caveolin, anti-cathepsin B, anti-annexin II and anti-p11 antibodies (C). To verify distribution of caveolae toTI fractions, HUVEC grownfor 18 h on rBM were treated with 10 mM MβCD for 1 h at 37 °C prior to caveolae isolation (D). TS (closed bars) and TI (open bars)fractions of HUVEC monolayers (mono) and tube-like structures (tube) were assayed for cathepsin B (CTSB) activity against Z-Arg-Arg-NHMec substrate and activity was recorded as pmol/min/cell number. The specific activity for caveolae-associated cathepsin Bactivity is expressed as a percentage of cathepsin B activity in the non-caveolar fraction (TS) (E). Overnight conditioned media ofHUVEC grown as a monolayer on gelatin (mono) and during tube formation (tube) were also analyzed for cathepsin B activity andexpressed as a percentage of secreted activatable cathepsin B activity from monolayer HUVEC (F). Immunoblots and graphs,presented as mean±standard deviation, are representative of at least three experiments. ⁎P<0.05 and ⁎⁎P<0.01.



disrupts the structure of caveolae. As a result, there was a loss incav-1 distribution to the TI-fraction of treated cells (Fig. 3D).Analysis of cathepsin B revealed an increase in the distribution ofthe mature 31 kDa and 25/26 kDa forms and the 38 kDaintermediate form to caveolae of HUVEC tube-like structures ascompared to caveolae of monolayer HUVEC (Fig. 3C); thisassociation is lost following 10 mM MβCD treatment (Fig. 3D),thus verifying that distribution of cathepsin B to caveolae is not anartifact of the fractionation procedure. Quantitation of theimmunoblotting data in Fig. 3C revealed a 15% increase in thedistribution of cathepsin B to the TI fraction as compared to the TSfraction during HUVEC tube formation (data not shown). Similarly,the distribution of p11 to the TI fractions was increased during tubeformation [90% increase (data not shown)] while the distributionof annexin II remained unchanged (Fig. 3C). Enzymatic assays alsodetected increased cathepsin B activity in the caveolae fraction ofHUVEC tube-like structures as compared to HUVEC monolayer,consistent with a functional role for this protease within thesemembrane domains (Fig. 3E). Cathepsin B activity was alsodetected in the media conditioned during HUVEC tube formationat levels greater than that observed in the media conditioned byHUVEC grown as a monolayer on gelatin (Fig. 3F). These data alongwith the pericellular degradation observed during tube formationsuggest that association of cathepsin B with caveolae may increasecathepsin B secretion and ECM degradation during endothelial celltube formation.

Fig. 4 – Active cathepsin B partially localizes to caveolae of HUVEC ducav-1-mRFP construct and grown on glass coverslips coated with rBGB-123 for 18 h. Thereafter, unbound GB-123 was washed away andwere then taken of live cells (DIC), cav-1-mRFP (red), DQ-collagencathepsins (pseudocolored blue). Colocalization of cav-1-mRFP andcolocalization of DQ-collagen IV degradation products and GB-123 aIV degradation products and GB-123 appears white (arrows; see ins

Collagen IV degradation products colocalize intracellularlywith active cysteine cathepsins and cav-1

We performed live cell proteolysis assays with HUVEC transfectedwith cav-1-mRFP in the presence of GB-123, a non-quenchedactivity-based probe for cysteine cathepsins B and L [26]. Note thatwe used GB-123 (a Cy5 labeled probe) rather than GB111-FL inthese live-cell proteolysis assays because the FITC-labeledGB111-FLcannot be differentiated from the degradation products of DQ-collagen IV. Cells grown in rBM containing DQ-collagen IV for 18 hin the presence of GB-123werewashed to removeunboundGB-123and incubated 1–2 h inprobe-freemediumprior to imaging for DQ-collagen IV degradation, cav-1 and cysteine cathepsin activity (Fig.4). Our results revealed that DQ-collagen IV degradation products(green) partially colocalized intracellularly with cav-1-mRFP (red)and GB-123 (blue), thus providing further support for thefunctional significance of cathepsin B localizing to caveolae.

Redistribution of other ECM degrading proteases to HUVECcaveolae during tube formation

Cathepsin B has been previously described as a potential initiatorof ECM proteolytic pathways that include proteases of the plasmin(ogen) cascade and MMPs. For that reason, we examined thedistribution of pro-uPA, a cathepsin B substrate [10], and itsreceptor uPAR to caveolae of HUVEC during tube formation and

ring tube formation. HUVECwere transiently transfectedwith aM containing 25 μg/ml DQ-collagen IV in the presence of 1 μMcells incubated 1–2 h in probe-free medium. Confocal images

IV degradation products (green) and GB-123 bound to cysteineDQ-collagen IV degradation products appears yellow,

ppears pale blue, and colocalization of cav-1-mRFP, DQ-collagenert) in the merged image. Bar, 10 μm.


found an increased association of both pro-uPA and uPAR to the TIfractions of HUVEC tube-like structures as compared to TI fractionsof HUVEC monolayers (Fig. 5A). β1-integrin, a protein that forms atrimeric complex with uPAR and cav-1 on cell surfaces, also co-colocalized to these caveolae fractions during tube formation (Fig.5A). These results are consistent with caveolae being the site forthe initiation of a network of proteases involved in ECMdegradation on the surface of endothelial cells.

MMP-2 and MMP-14 have both been identified in caveolae ofglioblastomas, fibrosarcomas and endothelial cells [39,40]. Wetherefore examined the association of these MMPs with caveolaeof HUVEC. MMP-14, also known as MT1-MMP, had previously beenshown to localize to caveolae and to internalize and traffic throughendothelial cells via a caveolae-mediated pathway [39–41]. In ourstudies, we detected an increased distribution of MMP-14 tocaveolae (TI-fraction) of HUVEC during tube formation (Fig. 5A).

We performed gelatin-zymography to compare the intracel-lular distribution of gelatinases MMP-2 and -9 in HUVEC grown asa monolayer versus tube-like structures. Our results revealed anassociation of MMP-2 with caveolae (TI) fractions of HUVECfollowing tube formation (Fig. 5B). Although MMP-9 was detected

Fig. 5 – Increased distribution of pro-uPA, uPAR, β1-integrin,MMP-2 and MMP-14 to HUVEC caveolae during tube formation.Equal numbers of HUVECwere grown on gelatin (mono) or rBM(tubes) for 18 h and caveolae were isolated by successivedetergent extraction as described in Materials and methods.Triton-soluble (TS) represents non-caveolae fractions andTriton-insoluble (TI) represents caveolae fractions. Equalvolume aliquots of each fraction were analyzed by SDS-PAGEand immunoblotted with anti-uPA, anti-uPAR, anti-β1-integrinand anti-MMP-14 antibodies (A). Equal volume aliquots of eachfraction were also analyzed by gelatin zymography for MMP-2and MMP-9 activities (B). Purified MMP-2 (10 ng) and MMP-9(5 ng) were used as positive controls. Immunoblots andzymograms are representative of at least three experiments.

in both the TS- and TI-fractions of HUVEC tube-like structures, wedetected MMP-9 in the rBM (i.e., in the Cultrex used in the in vitrotube formation assays) in the absence of cells as well as in mediaconditioned by Cultrex in the absence of cells (data not shown).MMP-2, in contrast, was not detected in Cultrex in the absence ofcells or in media conditioned by Cultrex in the absence of cells(data not shown).

Pericellular proteolysis of collagen IV by migrating HUVECduring tube formation

We performed live cell proteolysis assays in 4D (i.e., three-dimensional space+time) during endothelial tube formation. Toconduct these assays, we seeded HUVEC onto rBM containing DQ-collagen IV and imaged the cells every 10 min over 16 h with aZeiss LSM 510 META NLO microscope equipped with a controlledenvironmental chamber. Images taken at six time points over thecourse of the 16 h incubation period are illustrated in Fig. 6A. Amovie of proteolysis by the migrating and differentiating HUVEC isavailable online (Supplementary materials, movie SM1). After 2 hof incubation, HUVEC were still single cells with some cellsexhibiting sprouting (arrowheads). By 4 h the cells had started toalign and form small tubular networks with degradation productsof DQ-collagen IV (green) present pericellularly. At 6 h and 12 h,the smaller networks had begun to coalesce into larger networks.By 14 h and 16 h, larger tubular structures had formed from thesmaller tubes produced earlier during the assay (4 h) anddegradation products were observed immediately adjacent to thetubes. We also observed pericellular degradation products of DQ-collagen IV at the rear of one cell (arrow) that was migratingtowards the developing tubular structure. When we performedlive cell proteolysis assays using HUVEC transfected with cav-1-mRFP, we found that DQ-collagen IV degradation productscolocalized intracellularly with cav-1-mRFP in the perinuclearregion. Degradation products were also found pericellularly atboth the front and at the rear of a migrating endothelial cells(arrows) [Fig. 6B and movie available online (Supplementarymaterials, movie SM2)]. These data are quite intriguing sincepolarization of cav-1 to either the front or the rear of migratingcells including endothelial cells has been reported [20,22,42–44]and we detected cav-1 colocalized with DQ-collagen IV degrada-tion products at the rear of migrating endothelial cells (seearrowhead in insert). These data are consistent with a role forcaveolae-associated proteases in endothelial cell migrationthrough ECM during tube formation.

Discussion

Differentiation of endothelial cells during angiogenesis is a processthat involves proteolytic enzymes, which degrade basementmembrane and thereby facilitate cell migration, invasion andcapillary tube formation. The accessibility of proteases to the ECMis enhanced by their localization on the cell membrane. Here, weexplored one possible mechanism for cell surface association, i.e.,the distribution of cathepsin B and other proteases associated withECM degradation to caveolae on the cell surface of endothelialcells. Our results are the first to show through live-cell imagingECM degradation in associationwith active cathepsin B in caveolaeof endothelial cells during tube formation.

Fig. 6 – Time courses for collagen IV proteolysis and colocalization of collagen IV degradation products with cav-1 during HUVECtube formation. HUVEC were grown on glass coverslips coated with rBM containing 25 μg/ml DQ-collagen IV. (A) Confocal imageswere taken of live cells between 2 and 16 h. DQ-collagen IV degradation products (green) are seen surrounding tubular structuresand at the rear of a migrating endothelial cell (arrow). Cell sprouting is observed at 2 h (arrowheads). Bar, 100 μm. (B) Confocalimages were taken between 1 and 4 h of live HUVEC transfected with cav-1-mRFP. DQ-collagen IV degradation products (green) andcav-1 (red) are seen colocalized (yellow) intracellularly (arrowheads) and pericellular degradation of DQ-collagen IV is observed atthe rear of a migrating endothelial cell (arrows). Black arrows represent the original location of a migrating HUVEC cell (in box) at1 h. Bar, 20 μm.


Fig. 7 – Potential protease network in endothelial cell caveolaeduring ECM degradation. In endothelial cells, proteases(i.e., cathepsin B, pro-uPA and MMPs) and their associatedproteins/receptors (i.e., AIIt, uPAR and β1-integrin) aretranslocated to cell surface caveolae via intracellularvesicles (i.e., multivesicular bodies and exosomes) [14].Caveolae-associated proteases are activated and degrade ECMproteins both pericellularly and intracellularly viacaveolae-mediated endocytosis.


Caveolae are abundant in endothelial cells [45] and during anangiogenic response caveolae appear to coalesce and form larger,vesicular structures [46]. Previous studies have shown a correlationbetween cav-1 expression and angiogenesis. Overexpression ofcav-1 in endothelial cells enhances caveolae formation andaccelerates capillary tube development [47] and, conversely,downregulation of cav-1 reduces the number of capillary tubes[23,47]. Moreover, caveolae are hypothesized to be sites for thecompartmentalization of cell surface proteases [12], includingproteases linked to angiogenesis. In the present study,we identifiedactive cathepsin B and its cell surface receptor, p11, alongwith otherECM degrading proteases (i.e., pro-uPA, MMP-2 and -14), incaveolar fractions of human endothelial cells, an association thatwas enhanced during tube formation.

Cathepsin B regulates both pro- and anti-angiogenic factorssuggesting that this enzyme may be a contributor to the“angiogenic switch” of endothelial cells [9]. Lack of cathepsin Breduces angiogenesis in a mouse model of pancreatic islet cellcarcinogenesis (RIP1-Tag2) crossed with cathepsin B-null mice; incontrast, a lack of cathepsin L does not affect angiogenesis in thismouse model [48]. Cathepsin B also has been shown to exert anti-angiogenic affects through its ability to cleave collagen XVIII togenerate endostatin, an anti-angiogenic factor that has beenidentified in caveolae of endothelial cells [11,49].

uPA also has variable roles in angiogenesis. Binding of uPA to itsreceptor, uPAR, enhances organization of endothelial cells intotubular structures [50]. Conversely, uPA, along with tissueplasminogen activator (tPA), mediates the generation of angiosta-tin, an anti-angiogenic factor, from plasminogen [51]. Theincreased levels of β1-integrin in caveolae of HUVEC may reflectintersections between proteolytic and signaling pathwaysobserved in human chondrocytes in which uPAR complexes withcav-1 via β1-integrin [52] and disruption of this complex results inthe loss of uPAR-dependent adhesion and β1-integrin-inducedsignal transduction [53]. Our findings of increased localization ofp11, pro-uPA, uPAR and β1-integrin to caveolae of endothelial cellsduring tube formation would be consistent with a functional rolefor caveolae as a site for the compartmentalization and regulationof proteolytic events associated with angiogenesis.

Colocalization of cathepsin B and pro-uPA to caveolae ofendothelial cells is interesting since cathepsin B can activate pro-uPA to uPA, thereby initiating a network of cell surface proteolyticevents that includes activation of plasminogen and MMPs [10,54].The associations of MMP-2 and -14 with endothelial caveolae,when the cells formed tubular structures, are in concert with anincrease in secretion of MMP-2. MMP-14 and MMP-2 havepreviously been localized to caveolae of endothelial cells alongwith αvβ3 and TIMP-2, a receptor and an inhibitor of MMP-2,respectively [39,40]. The colocalization of these proteins tocaveolae may be of functional significance since MMP-14 activatesMMP-2, a process that also involves TIMP-2 [55]. In endothelialcells, the cytoplasmic tail of MMP-14 binds phosphorylated cav-1[56], and the enzyme is internalized and trafficked by caveolae-mediated endocytosis [41].

Although the mechanism(s) for translocation of proteases tocaveolae remains unclear, we hypothesize that their localization tothese specialized membrane domains influences their activities atthe cell surface. Our previous studies in human colorectalcarcinoma cells revealed that downregulation of cav-1 decreasesdistribution of cathepsin B and uPA to caveolae and secretion of

these enzymes [18]. Consequentially, these cav-1 knockdown cellsare less invasive and degrade less ECM than their parentalcounterparts [18]. In the present study, we show intracellularcolocalization of cav-1 with ECM degradation products inendothelial cells. In particular, intracellular colocalization of ECMdegradation products and cav-1 was observed at the rear of someendothelial cells migrating on rBM. Other studies have shown thatcav-1 translocates to the rear of migrating endothelial cells [22].Here we propose that a possible function for caveolae and theirassociated proteases at the rear of migrating endothelial cells is inthe uptake and intracellular degradation of ECM proteins, aprerequisite for cell detachment and one that also has been observedfor cathepsin B in migrating tumor cells [14]. An involvement ofproteases in the intracellular degradation of ECM proteins has beenreported [35,57–59]. In breast carcinoma cells, intracellular cysteinecathepsins degrade collagen IV [59], a process suppressed byCA074Me, a cell-permeable, selective inhibitor of cathepsins B andL [60]. Degradation of collagen IV by fibroblasts is also linked tocysteine cathepsins following endocytosis of collagen IV via amechanism mediated by the uPAR-associated protein (uPARAP)/Endo180 [61], a member of the macrophage mannose receptorfamily of endocytic transmembrane glycoproteins [62]. uPARAP


forms a trimeric complex with uPA and uPAR on the cell surface andbinds several types of collagens including collagen I, IV and V [63]. Inmice deficient in uPARAP, fibroblasts show an almost completeabrogation of collagen endocytosis and diminished cell adhesion andmigration on various collagens [61]. Although there are no knownreports on the expression of uPARAP in endothelial cells, itsassociation with uPA/uPAR in other cell types leads us to speculatethat uPARAP associates with caveolae and participates in ECMendocytosis and degradation. In Fig. 7, we have illustrated a potentialpathway for the association of proteases (i.e., cathepsin B, uPA andMMPs) with caveolae and caveolae-associated ECM degradation inendothelial cells. Further dissection of the mechanisms of ECMendocytosis and degradation via caveolae and their associatedproteases in endothelial cell differentiation remains for futurestudies.

Acknowledgments

This work was supported by a National Institutes of Health (NIH)National Technology Center for Networks and Pathways Grant(U54-RR020843) and a DOD Breast Cancer Center of Excellence(DAMD17-02-1-0693). The Microscopy and Imaging ResourcesLaboratory is supported by the NIH National Technology CenterGrant and NIH Center Grants P30-ES06639 and P30-CA22453.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.yexcr.2009.01.021.

R E F E R E N C E S

[1] W. Risau, Mechanisms of angiogenesis, Nature 386 (1997)671–674.

[2] V.W. van Hinsbergh, M.A. Engelse, P.H. Quax, Pericellularproteases in angiogenesis and vasculogenesis, Arterioscler.Thromb. Vasc. Biol. 26 (2006) 716–728.

[3] G. Bergers, R. Brekken, G. McMahon, T.H. Vu, T. Itoh, K. Tamaki, K.Tanzawa, P. Thorpe, S. Itohara, Z. Werb, D. Hanahan, Matrixmetalloproteinase-9 triggers the angiogenic switch duringcarcinogenesis, Nat. Cell Biol. 2 (2000) 737–744.

[4] M.S. Pepper, Role of the matrix metalloproteinase and plasminogenactivator-plasmin systems in angiogenesis, Arterioscler. Thromb.Vasc. Biol. 21 (2001) 1104–1117.

[5] J.M. Rakic, C. Maillard, M. Jost, K. Bajou, V. Masson, L. Devy, V.Lambert, J.M. Foidart, A. Noel, Role of plasminogenactivator-plasmin system in tumor angiogenesis, Cell. Mol. LifeSci. 60 (2003) 463–473.

[6] K.T. Preissner, A.E. May, K.D. Wohn, M. Germer, S.M. Kanse,Molecular crosstalk between adhesion receptors and proteolyticcascades in vascular remodelling, Thromb. Haemost. 78 (1997)88–95.

[7] J. Xu, D. Rodriguez, E. Petitclerc, J.J. Kim, M. Hangai, Y.S. Moon, G.E.Davis, P.C. Brooks, Proteolytic exposure of a cryptic site withincollagen type IV is required for angiogenesis and tumor growth invivo, J. Cell Biol. 154 (2001) 1069–1079.

[8] B.C. Patterson, Q.A. Sang, Angiostatin-converting enzymeactivities of human matrilysin (MMP-7) and gelatinase B/typeIV collagenase (MMP-9), J. Biol. Chem. 272 (1997)28823–28825.

[9] E. Im, A. Venkatakrishnan, A. Kazlauskas, Cathepsin B regulatesthe intrinsic angiogenic threshold of endothelial cells, Mol. Biol.Cell 16 (2005) 3488–3500.

[10] H. Kobayashi, M. Schmitt, L. Goretzki, N. Chucholowski, J. Calvete,M. Kramer, W.A. Gunzler, F. Janicke, H. Graeff, Cathepsin Befficiently activates the soluble and the tumor cell receptor-boundform of the proenzyme urokinase-type plasminogen activator(Pro-uPA), J. Biol. Chem. 266 (1991) 5147–5152.

[11] M. Ferreras, U. Felbor, T. Lenhard, B.R. Olsen, J. Delaisse,Generation and degradation of human endostatin proteins byvarious proteinases, FEBS Lett. 486 (2000) 247–251.

[12] D. Cavallo-Medved, B.F. Sloane, Cell-surface cathepsin B:understanding its functional significance, Curr. Top. Dev. Biol. 54(2003) 313–341.

[13] R. Fridman, M. Toth, I. Chvyrkova, S.O. Meroueh, S. Mobashery,Cell surface association of matrix metalloproteinase-9 (gelatinaseB), Cancer Metastasis Rev. 22 (2003) 153–166.

[14] M.M. Mohamed, B.F. Sloane, Cysteine cathepsins: multifunctionalenzymes in cancer, Nat. Rev. Cancer 6 (2006) 764–775.

[15] M. Seiki, N. Koshikawa, I. Yana, Role of pericellular proteolysisby membrane-type 1 matrix metalloproteinase in cancerinvasion and angiogenesis, Cancer Metastasis Rev. 22 (2003)129–143.

[16] R.G. Anderson, The caveolae membrane system, Annu. Rev.Biochem. 67 (1998) 199–225.

[17] T.M. Williams, M.P. Lisanti, The caveolin proteins, Genome Biol. 5(2004) 214.

[18] D. Cavallo-Medved, J. Mai, J. Dosescu, M. Sameni, B.F. Sloane,Caveolin-1 mediates the expression and localization of cathepsinB, pro-urokinase plasminogen activator and their cell-surfacereceptors in human colorectal carcinoma cells, J. Cell Sci. 118(2005) 1493–1503.

[19] M.L. Massimino, C. Griffoni, E. Spisni, M. Toni, V. Tomasi,Involvement of caveolae and caveolae-like domains in signalling,cell survival and angiogenesis, Cell Signal. 14 (2002) 93–98.

[20] A. Beardsley, K. Fang, H. Mertz, V. Castranova, S. Friend, J. Liu, Lossof caveolin-1 polarity impedes endothelial cell polarization anddirectional movement, J. Biol. Chem. 280 (2005) 3541–3547.

[21] M. Isshiki, J. Ando, K. Yamamoto, T. Fujita, Y. Ying, R.G. Anderson,Sites of Ca(2+) wave initiation move with caveolae to the trailingedge of migrating cells, J. Cell Sci. 115 (2002) 475–484.

[22] M.O. Parat, B. Anand-Apte, P.L. Fox, Differential caveolin-1polarization in endothelial cells during migration in two andthree dimensions, Mol. Biol. Cell 14 (2003) 3156–3168.

[23] C. Griffoni, E. Spisni, S. Santi, M. Riccio, T. Guarnieri, V. Tomasi,Knockdown of caveolin-1 by antisense oligonucleotides impairsangiogenesis in vitro and in vivo, Biochem. Biophys. Res.Commun. 276 (2000) 756–761.

[24] K. Moin, N.A. Day, M. Sameni, S. Hasnain, T. Hirama, B.F. Sloane,Human tumour cathepsin B. Comparison with normal livercathepsin B, Biochem J 285 (1992) 427–434.

[25] G. Blum, S.R. Mullins, K. Keren, M. Fonovic, C. Jedeszko, M.J. Rice,B.F. Sloane, M. Bogyo, Dynamic imaging of protease activity withfluorescently quenched activity-based probes, Nat. Chem. Biol. 1(2005) 203–209.

[26] G. Blum, G. von Degenfeld, M.J. Merchant, H.M. Blau, M. Bogyo,Noninvasive optical imaging of cysteine protease activity usingfluorescently quenched activity-based probes, Nat. Chem. Biol. 3(2007) 668–677.

[27] D.S. Grant, P.I. Lelkes, K. Fukuda, H.K. Kleinman, Intracellularmechanisms involved in basement membrane induced bloodvessel differentiation in vitro, In Vitro Cell. Dev. Biol. 27A (1991)327–336.

[28] Y. Kubota, H.K. Kleinman, G.R. Martin, T.J. Lawley, Role of lamininand basement membrane in the morphological differentiation ofhuman endothelial cells into capillary-like structures, J. Cell Biol.107 (1988) 1589–1598.

[29] K.S. Song, S. Li, T. Okamoto, L.A. Quilliam, M. Sargiacomo, M.P.Lisanti, Co-purification and direct interaction of Raswith caveolin,

http://dx.doi.org/doi:10.1016/j.yexcr.2009.01.021


an integral membrane protein of caveolae microdomains.Detergent-free purification of caveolae microdomains, J. Biol.Chem. 271 (1996) 9690–9697.

[30] K.R. Solomon, M.A. Mallory, R.W. Finberg, Determination of thenon-ionic detergent insolubility and phosphoprotein associationsof glycosylphosphatidylinositol-anchored proteins expressed onT cells, Biochem. J. 334 (1998) 325–333.

[31] M. Toth, D.C. Gervasi, R. Fridman, Phorbol ester-induced cellsurface association of matrix metalloproteinase-9 in humanMCF1OA breast epithelial cells, Cancer Res. 57 (1997) 3159–3167.

[32] B.E. Linebaugh, M. Sameni, N.A. Day, B.F. Sloane, D. Keppler,Exocytosis of active cathepsin B enzyme activity at pH 7.0,inhibition and molecular mass, Eur. J. Biochem. 264 (1999)100–109.

[33] D. Caglic, J.R. Pungercar, G. Pejler, V. Turk, B. Turk,Glycosaminoglycans facilitate procathepsin B activation throughdisruption of propeptide-mature enzyme interactions, J. Biol.Chem. 282 (2007) 33076–33085.

[34] D. Cavallo-Medved, J. Dosescu, B. Linebaugh, M. Sameni, D. Rudy,B.F. Sloane, Mutant K-ras regulates cathepsin B localization on thesurface of human colorectal carcinoma cells. Neoplasia 5 (2003)507–519.

[35] M. Sameni, J. Dosescu, K. Moin, B.F. Sloane, Functional imaging ofproteolysis: stromal and inflammatory cells increase tumorproteolysis, Mol. Imaging 2 (2003) 159–175.

[36] B.F. Sloane, M. Sameni, I. Podgorski, D. Cavallo-Medved, K. Moin,Functional imaging of tumor proteolysis, Annu. Rev. Pharmcol.Toxicol. 46 (2006) 301–315.

[37] J. Mai, D.M. Waisman, B.F. Sloane, Cell surface complex ofcathepsin B/annexin II tetramer in malignant progression,Biochim. Biophys. Acta 1477 (2000) 215–230.

[38] P. Liu, M. Rudick, R.G. Anderson, Multiple functions of caveolin-1,J. Biol. Chem. 277 (2002) 41295–41298.

[39] B. Annabi, M. Lachambre, N. Bousquet-Gagnon, M. Page, D.Gingras, R. Beliveau, Localization of membrane-type 1 matrixmetalloproteinase in caveolae membrane domains, Biochem. J.353 (2001) 547–553.

[40] A. Puyraimond, R. Fridman, M. Lemesle, B. Arbeille, S. Menashi,MMP-2 colocalizes with caveolae on the surface of endothelialcells, Exp. Cell Res. 262 (2001) 28–36.

[41] B.G. Galvez, S. Matias-Roman, M. Yanez-Mo, M. Vicente-Manzanares,F. Sanchez-Madrid, A.G. Arroyo, Caveolae are a novel pathway formembrane-type 1 matrix metalloproteinase traffic in humanendothelial cells, Mol. Biol. Cell 15 (2004) 678–687.

[42] V. Santilman, J. Baran, B. Anand-Apte, P.L. Fox, M.O. Parat,Caveolin-1 polarization in migrating endothelial cells is directedby substrate topology not chemoattractant gradient, Cell Motil.Cytoskelet. 63 (2006) 673–680.

[43] D. Lentini, F. Guzzi, F. Pimpinelli, R. Zaninetti, A. Cassetti, S. Coco,R. Maggi, M. Parenti, Polarization of caveolins and caveolae duringmigration of immortalized neurons, J. Neurochem. 104 (2008)514–523.

[44] X.H. Sun, D.C. Flynn, V. Castranova, L.L. Millecchia, A.R. Beardsley,J. Liu, Identification of a novel domain at the N terminus ofcaveolin-1 that controls rear polarization of the protein andcaveolae formation, J. Biol. Chem. 282 (2007) 7232–7241.

[45] M.P. Lisanti, P.E. Scherer, J. Vidugiriene, Z. Tang,A. Hermanowski-Vosatka, Y.H. Tu, R.F. Cook, M. Sargiacomo,Characterization of caveolin-rich membrane domains isolatedfrom an endothelial-rich source: implications for human disease,J. Cell Biol. 126 (1994) 111–126.

[46] S. Esser, K. Wolburg, H. Wolburg, G. Breier, T. Kurzchalia, W.Risau, Vascular endothelial growth factor induces endothelialfenestrations in vitro, J. Cell Biol. 140 (1998) 947–959.

[47] J. Liu, X.B. Wang, D.S. Park, M.P. Lisanti, Caveolin-1 expressionenhances endothelial capillary tubule formation, J. Biol. Chem.277 (2002) 10661–10668.

[48] V. Gocheva, W. Zeng, D. Ke, D. Klimstra, T. Reinheckel, C. Peters,D. Hanahan, J.A. Joyce, Distinct roles for cysteine cathepsin genesin multistage tumorigenesis, Genes Dev. 20 (2006) 543–556.

[49] S.A. Wickstrom, K. Alitalo, J. Keski-Oja, Endostatin associates withlipid rafts and induces reorganization of the actin cytoskeleton viadown-regulation of RhoA activity, J. Biol. Chem. 278 (2003)37895–37901.

[50] H.W. Schnaper, E.S. Barnathan, A. Mazar, S. Maheshwari, S. Ellis,S.L. Cortez, W.H. Baricos, H.K. Kleinman, Plasminogen activatorsaugment endothelial cell organization in vitro by two distinctpathways, J. Cell. Physiol. 165 (1995) 107–118.

[51] S. Gately, P. Twardowski, M.S. Stack, D.L. Cundiff, D. Grella, F.J.Castellino, J. Enghild, H.C. Kwaan, F. Lee, R.A. Kramer, O. Volpert,N. Bouck, G.A. Soff, Themechanism of cancer-mediated conversionof plasminogen to the angiogenesis inhibitor angiostatin, Proc. Natl.Acad. Sci. U. S. A. 94 (1997) 10868–10872.

[52] W. Schwab, J.M. Gavlik, T. Beichler, R.H. Funk, S. Albrecht, V.Magdolen, T. Luther, M. Kasper, M. Shakibaei, Expression of theurokinase-type plasminogen activator receptor in human articularchondrocytes: association with caveolin and beta 1-integrin,Histochem. Cell Biol. 115 (2001) 317–323.

[53] Y. Wei, X. Yang, Q. Liu, J.A. Wilkins, H.A. Chapman, A role forcaveolin and the urokinase receptor in integrin-mediated adhesionand signaling, J. Cell Biol. 144 (1999) 1285–1294.

[54] Y. Ikeda, T. Ikata, T. Mishiro, S. Nakano, M. Ikebe, S. Yasuoka,Cathepsins B and L in synovial fluids from patients withrheumatoid arthritis and the effect of cathepsin B on theactivation of pro-urokinase, J. Med. Investig. 47 (2000) 61–75.

[55] M.M. Bernardo, R. Fridman, TIMP-2 (tissue inhibitor ofmetalloproteinase-2) regulates MMP-2 (matrixmetalloproteinase-2) activity in the extracellular environmentafter pro-MMP-2 activation by MT1 (membrane type 1)-MMP,Biochem. J. 374 (2003) 739–745.

[56] L. Labrecque, C. Nyalendo, S. Langlois, Y. Durocher, C. Roghi,G. Murphy, D. Gingras, R. Beliveau, Src-mediated tyrosinephosphorylation of caveolin-1 induces its association withmembrane type 1 matrix metalloproteinase, J. Biol. Chem. 279(2004) 52132–52140.

[57] V. Everts, E. van der Zee, L. Creemers, W. Beertsen, Phagocytosisand intracellular digestion of collagen, its role in turnover andremodelling, Histochem. J. 28 (1996) 229–245.

[58] L. Kjoller, L.H. Engelholm, M. Hoyer-Hansen, K. Dano, T.H. Bugge,N. Behrendt, uPARAP/endo180 directs lysosomal delivery anddegradation of collagen IV, Exp. Cell Res. 293 (2004) 106–116.

[59] M. Sameni, K. Moin, B.F. Sloane, Imaging proteolysis by livinghuman breast cancer cells, Neoplasia 2 (2000) 496–504.

[60] D.J. Buttle, M. Murata, C.G. Knight, A.J. Barrett, CA074 methylester: a proinhibitor for intracellular cathepsin B, Arch. Biochem.Biophys. 299 (1992) 377–380.

[61] L.H. Engelholm, K. List, S. Netzel-Arnett, E. Cukierman, D.J. Mitola,H. Aaronson, L. Kjoller, J.K. Larsen, K.M. Yamada, D.K. Strickland,K. Holmbeck, K. Dano, H. Birkedal-Hansen, N. Behrendt, T.H.Bugge, uPARAP/Endo180 is essential for cellular uptake ofcollagen and promotes fibroblast collagen adhesion, J. Cell Biol.160 (2003) 1009–1015.

[62] L. East, C.M. Isacke, The mannose receptor family, Biochim.Biophys. Acta 1572 (2002) 364–386.

[63] N. Behrendt, The urokinase receptor (uPAR) and theuPAR-associated protein (uPARAP/Endo180): membraneproteins engaged in matrix turnover during tissue remodeling,Biol. Chem. 385 (2004) 103–136.

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Live-cell imaging demonstrates extracellular matrix degradation in