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Protein Kinase C-ε Promotes Survival of Lung Cancer Cells by Suppressing Apoptosis through Dysregulation of the Mitochondrial Caspase Pathway*

The serine/threonine protein kinase C (PKC) has been implicated in the regulation of drug resistance and cell survival in many types of cancer cells. However, the one or more precise mechanisms remain elusive. In this study, we have identified and determined the mechanism by which PKC-ε, a novel PKC isoform, modulates drug resistance in lung cancer cells. Western blot analysis demonstrates that expression of PKC-ε, but not other PKC isoforms, is associated with the chemo-resistant phenotype of non-small cell lung cancer (NSCLC) cell lines. Northern blotting and nuclear run-on transcription analysis further reveals that the failure of expression of PKC-ε in the chemo-sensitive phenotype of small cell lung cancer (SCLC) cells results from transcriptional inactivation of the gene. Importantly, forced expression of PKC-ε in NCI-H82 human SCLC cells confers a significant resistance to the chemotherapeutic drugs, etoposide and doxorubicin. Resistance is characterized by a significant reduction in apoptosis in PKC-ε-expressing cells. Treatment of NCI-H82 cells with etoposide induces a series of time-dependent events, including the release of cytochromec from the mitochondria to the cytosol, activation of caspase-9 and caspase-3, and cleavage of poly(ADP-ribose) polymerase (PARP). All of these events are blocked by PKC-ε expression. Furthermore, caspase-specific inhibitors, z-VAD-fmk and z-DEVD-fmk, significantly attenuate the accumulation of sub-G1population and block the PARP cleavage in response to etoposide. These results suggest that PKC-ε prevents cells from undergoing apoptosis through inhibition of the mitochondrial-dependent caspase activation, thereby leading to cell survival. Finally, down-regulation of PKC-ε expression by the antisense cDNA in NSCLC cells results in increased sensitivity to etoposide. Taken together, our findings suggest an important role for PKC-ε in regulating survival of lung cancer cells.

Lung cancer is the leading cause of cancer-related deaths in the United States. Human lung cancer is a disease of heterogeneous histology which can be divided into two major catego-ries; small cell lung cancer (SCLC) 1 and non-small cell lung cancer (NSCLC). SCLC represents ϳ25% of all lung cancer and is characterized by a marked initial response to chemotherapy and radiation. However, nearly all patients with SCLC relapse and develop resistance to cytotoxic therapy and eventually die from the disease. Overall, the 5-year survival rate is only 3-8% (1). The underlying mechanism of developing drug resistance in relapsed SCLC patients is unclear. Studies have suggested that tumor cells that survive chemotherapy display phenotypes distinctly different from those expressed by the primary tumor prior to chemotherapy (2). It is clear that understanding the molecular determinants of chemo-resistance in SCLC upon relapse will lead to more efficient therapy.
Apoptosis, or programmed cell death, is a highly specific and regulated process that plays an important role in the development and homeostasis of multicellular organisms as well as tumorigenesis (3,4). There is an increasing body of evidence that the control of apoptosis is disrupted in many tumor cells. The dysregulated apoptosis is thought to contribute to enhanced tumor progression and metastasis. Furthermore, it has become evident that resistance to apoptosis is one potential mechanism whereby tumor cells escape from chemotherapyinduced cytotoxicity, leading to cell survival (5). Development of chemo-resistance in tumor cells may, therefore, be due to their selective defects in the intracellular signaling proteins central to apoptotic pathways. These defects may provide a selective advantage for the tumor cells thereby rendering them resistant to chemotherapy.
Recent studies have defined a number of key molecules that are critical in apoptotic process. These include the Bcl-2 family of proteins and a family of aspartate-specific cysteine proteases named caspases. The Bcl-2 family proteins play a critical role in regulation of mitochondria function during apoptosis (6,7). Members of the Bcl-2 family can have both pro-apoptotic and anti-apoptotic activities. The interactions between the different Bcl-2 family members play an important role in determining cell fate during apoptosis. Caspases are the central engines of the apoptotic process, which are synthesized as latent zymogens and are activated by proteolytic cleavage (8). There are currently two well characterized caspase-activating cascades that regulate apoptosis: one is mediated by cell surface death receptors, and the other is regulated by the mitochondria. In the death receptor pathway, death receptor ligation promotes clustering and activation of caspase-8 and -10, which in turn activate downstream caspases. By contrast, caspase activation via the mitochondrial pathway involves mitochondrial integration of apoptotic signals and subsequent release of cytochrome c into the cytosol. Once released from the mitochondria, cytochrome c initiates formation of a complex (apoptosome) with procaspase-9 and an adaptor molecule, apoptotic protease-activating factor-1 (Apaf-1) thereby activating caspase-9, which in turn activates downstream caspases such as caspase-3, -6, and -7.
Protein kinase C (PKC) is a family of at least 11 structurally related serine/threonine protein kinases that play crucial roles in transducing signals that regulate diversified biological functions, including proliferation, differentiation, and apoptosis (9,10). The PKC family can be divided into three subgroups based on differences in their structures and biochemical properties: the classic isoforms (PKC-␣, ␤I, ␤II, and ␥), which are Ca 2ϩ and phorbol ester/diacylglycerol (DAG)-dependent; the novel isoforms (PKC-␦, ⑀, , , and ), which are phorbol ester/DAG-dependent but Ca 2ϩ -independent; and the atypical isoforms (PKC-and /), which are Ca 2ϩ -and phorbol ester/DAG-independent. The heterogeneous expression profiles of each PKC isoform in tissues strongly suggest that each isoform has a unique individual role. Their different subcellular localization and isoform-specific cofactor and activator requirements indicate each isoform is regulated differently (10,11). Recent studies using strategies to explore the isoform-specific functions of PKC suggest that specific PKC isoforms may be either antiapoptotic or pro-apoptotic, which is likely dependent on the nature of the apoptotic stimuli and the specific cell types involved (12). PKC isoforms that appear to be anti-apoptotic include PKC-␣, PKC-␤II, PKC-⑀, and the atypical isoforms PKC-/ and PKC- (13)(14)(15)(16). In contrast, PKC-␦ has been shown as a common intermediate in the apoptotic pathway (17,18). However, the precise biochemical mechanisms by which PKC regulates anti-apoptotic effects remain elusive.
Human lung cancer cells express multiple PKC isoforms. It has been reported that the drug-resistant phenotype is associated with expression and/or activity of PKCs in lung cancer cell lines and lung carcinomas (19,20). However, it is unclear what role(s), if any, a specific PKC isoform may play in this process. In this study, we identify and determine the function of the PKC-⑀ isoform in lung cancer cells. Using isoform-specific antibodies, we demonstrate that PKC-⑀, but not other PKC isoforms, is specifically linked to the chemo-resistant phenotype of the NSCLC cell lines and its induction in SCLC cells confers resistance to chemotherapy. Evidence presented in this study suggests that the chemo-protective effect of PKC-⑀ is mediated by the inhibition of drug-induced caspase activation through dysregulation of one or more of the mitochondrial apoptotic pathways, thereby leading to cell survival. These observations demonstrate a functional role for PKC-⑀ in the regulation of survival of lung cancer cells and provide potential novel targets for therapeutic intervention to overcome drug resistance.

EXPERIMENTAL PROCEDURES
Materials-Human PKC-⑀ cDNA was obtained from Dr. A. P. Fields (University of Texas Medical Branch, Galveston, TX). The human PKC-⑀ cDNA was tagged with a FLAG epitope at its N terminus and subcloned into expression vector pCI-neo (Promega, Madison, WI). The plasmid that expresses the antisense PKC-⑀ cDNA was constructed by subcloning the human full-length PKC-⑀ cDNA into pRc/CMV vector (Invitrogen, Carlsbad, CA) in an antisense orientation. Mouse monoclonal antibodies to specific PKC isoforms were purchased from BD Transduction Laboratories (Lexington, KY). Antibodies against cleaved and full-length versions of caspase-3 and caspase-9 were from Cell Signaling (Beverly, MA). Rabbit anti-PARP and goat anti-actin polyclonal antibodies were purchased from Upstate Biotechnology (Lake Placid, NY) and Santa Cruz Biotechnology (Santa Cruz, CA), respectively. Anti-cytochrome c (clone 7H8.2C12) and anti-cytochrome c oxidase (COX) subunit IV (clone 20E8) were from BD PharMingen and Molecular Probes (Eugene, OR), respectively. Caspase inhibitors (z-VAD-fmk and z-DEVD-fmk) were purchased from Enzyme System Products (Livemore, CA). Etoposide and doxorubicin were from Calbiochem (San Diego, CA). G418 sulfate (Geneticin) and LipofectAMINE were from Invitrogen (Rockville, MD). Hoechst 33342 and propidium iodide were obtained from Sigma (St. Louis, MO). [␣-32 P]dCTP and [␣-32 P]CTP were from Amersham Biosciences (Piscataway, NJ).
Cell Culture and DNA Transfection-Human lung cancer cell lines were obtained from ATCC (Rockville, MD) and maintained in RPMI 1640 medium supplemented with 9% (v/v) calf serum, 100 units/ml penicillin, and 100 units/ml streptomycin. Stable transfection of NCI-H82 cells was carried out using an electroporation method. Transfected cells were selected for G418 resistance. Clonal populations that express the FLAG-tagged PKC-⑀ were generated and maintained in the culture medium containing 500 g/ml G418. Stable transfection of NCI-H157 with an antisense PKC-⑀ cDNA was carried out using LipofectAMINE according to the manufacturer's recommendation. 48-h post-transfection, cells were selected for G418-resistance. A pool of G418-resistant clones was used for clonogenic survival assays.
Northern Blotting and Nuclear Run-on Transcription Analysis-Total cellular RNA was isolated using the acid phenol-guanidine isothiocyanate method (21). 20 g of RNA was fractionated on a denaturing 1% agarose gel containing formaldehyde (2.1 M), transferred to a Gene-Screen membrane (PerkinElmer Life Sciences, Boston, MA), and hybridized with 32 P-labeled human PKC-⑀ cDNA probe in ULTRAhyb hybridization buffer (Ambion, Austin, TX). The same membrane was then stripped and re-probed with human ␤-actin cDNA for loading control.
Nuclear run-on transcription assays were carried out as described previously (22). The 32 P-labeled nascent RNA transcripts purified from nuclei of representative cell lines were used to hybridize with the immobilized cDNA on Zeta-Probe membrane (Bio-Rad, Hercules, CA) according to the manufacturer's recommendations. After high stringency washes, membranes were subjected to autoradiography.
Soft Agar Assay and Growth Studies-Soft agar assays were carried out in six-well plates as described by Clark et al. (23). Briefly, cells were seeded in 0.3% agarose in growth medium overlaid on a base of 0.6% agarose. Cultures were fed weekly. Colonies were scored 21 days later after plating. For growth curves, 5 ϫ 10 4 cells per well were plated in six-well plates in complete medium. Media were changed every 2 days. Cell numbers were determined by trypan blue exclusion. Triplicate cultures of each cell clone were prepared and processed in all experiments.
Cell Viability and Clonogenic Survival Assays-Cells (1 ϫ 10 5 cells/ well) were cultured in six-well plates in the presence of either etoposide or doxorubicin at indicated concentrations for 96 h. Control cells were treated with the vehicle, dimethyl sulfoxide (Me 2 SO). Cell viability was determined by trypan blue dye exclusion and presented as the ratio of viable cell numbers in drug-treated cultures relative to that in Me 2 SOtreated controls times 100. Clonogenic assays were performed by plating cells (5000 cells/well) in six-well plates, treating with etoposide or vehicle (0.1% Me 2 SO) for 48 h. Cells were then cultured in drug-free media for 12 days and stained with 0.5 mg/ml crystal violet in 20% methanol. Drug-resistant colonies were scored, and cloning efficiency was expressed as a ratio of the number of colonies formed in the antisense-transfected cells to that in the vector-transfected cells times 100. All treatments were performed in triplicate.
Measurement of Apoptosis-Control or treated cells were collected and stained by incubating in a staining solution of 100 g/ml Hoechst 33342 and 20 g/ml propidium iodide in phosphate-buffered saline (PBS) at 37°C for 15 min in dark. Stained cells were viewed under the Zeiss Axioplan-2 fluorescence microscope with the appropriate filters to simultaneously visualize the blue fluorescence from Hoechst 33342 and the red fluorescence from propidium iodide. Apoptotic cells were defined by condensed and/or fragmented chromatin of blue fluorescence-emitting nuclei. To quantify apoptotic cells, a least 500 cells per preparation were scored for the incidence of apoptotic chromatin changes. Results are expressed as a ratio of apoptotic cells to the total number of blue nuclei examined times 100.
Flow cytometry analysis was used to determine the sub-G 1 populations of etoposide-treated cells. Cells were fixed in 90% methanol, treated with RNase A (100 g/ml) at 4°C for at least 4 h, and stained with 50 g/ml propidium iodide in PBS on ice for 15 min. DNA contents were assessed by fluorescence-activated cell sorting (FACS) using a FACSort (Becton Dickinson Immunocytometry Systems, San Jose, CA). A total of 50,000 cells was analyzed in every event. Data were evaluated using CellQuest software (Becton Dickinson) and represent the average of three measurements.
Detection of Internucleosomal DNA Fragmentation-Cells (1 ϫ 10 6 ) were harvested and lysed in a buffer of 500 mM Tris⅐Cl (pH 9.0), 20 mM EDTA, 10 mM NaCl supplemented with proteinase K (1 mg/ml) and 1% SDS at 37°C overnight. DNA was extracted with phenol/chloroform/ isoamyl alcohol (25:24:1, v/v), followed by ethanol precipitation. DNA was recovered by centrifugation at 13,000 ϫ g for 30 min and dissolved in LTE buffer (3 mM Tris⅐Cl (pH 8.0) and 0.3 mM EDTA) containing 500 g/ml RNase A. 20 g of DNA per sample was digested with RNase A at 37°C for 1 h and separated on a 1.8% agarose gel. DNA fragmentation was examined under UV lights after ethidium bromide staining and photographed.
Colorimetric Caspase-3 Activity Assay-Caspase-3 activity was assessed using a Caspase-3 Cellular Activity Assay Kit (Calbiochem) according to the protocol suggested by the manufacturer with minor modifications. Briefly, cells were resuspended and incubated at 5 ϫ 10 7 cells/ml in a lysis buffer of 50 mM HEPES (pH 7.4), 1 mM dithiothreitol, 0.1 mM EDTA, 0.1% CHAPS, and 0.1% Nonidet P-40 for 30 min at 4°C. Cell lysates were cleared at 10,000 ϫ g for 10 min at 4°C. Caspase-3 activity was measured in the supernatant using colorimetric Ac-DEVD-pNA as a substrate. The cleavage of the substrate was determined using a kinetic microplate reader (Molecular Devices Corp., Sunnyvale, CA) at a wavelength of 405 nm. Caspase-3 activity was calculated according to the slope of the A 405 versus time plot from linear regression analysis. Data were expressed as picomoles of the colorimetric substrate pNA liberated per minute and microgram of protein.
Subcellular Fractionation-Cells were treated with etoposide or Me 2 SO for 96 h and harvested by centrifugation at 900 ϫ g for 5 min at 4°C. Cell pellets were washed twice with ice-cold PBS and resuspended in buffer A (20 mM HEPES (pH 7.5), 250 mM sucrose, 1.5 mM MgCl 2 , 10 mM KCl, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol) containing 1 mM phenylmethylsulfonyl fluoride, 10 g/ml aprotinin, and 10 g/ml leupeptin. After 10-min incubation on ice, cells were homogenized using a Dounce tissue grinder. Cell homogenates were centrifuged at 1,000 ϫ g for 10 min at 4°C to discard unbroken cells and nuclei. Supernatants were subjected to additional centrifugation at 10,000 ϫ g for 25 min at 4°C. The resulting pellets (rich in mitochondria) were resuspended in buffer A and designated as the heavy-membrane fraction. The supernatants were further centrifuged at 150,000 ϫ g for 90 min at 4°C and collected as the cytosolic fraction.
Western Blot Analysis-Whole cell lysates were prepared in a modified radioimmune precipitation assay buffer as described previously (24). Equal amounts of lysates were resolved on a SDS-polyacrylamide gel and then transferred to a nitrocellulose membrane. Protein expression was determined by immunoblotting with appropriate antibodies as described previously (24). The protein was visualized by using the enhanced chemiluminescence (ECL) detection system (Amersham Biosciences).
Statistics-All data are presented as means Ϯ S.D. Differences in anchorage-independent colony formation and effects of caspase inhibitors on the sub-G 1 accumulation were examined using Student's t test. SigmaPlot (SPSS Inc., Chicago, IL) was used to perform statistical analyses. The difference was considered to be statistically significant if the p value was Ͻ0.05.

Expression of PKC-⑀ Isoform Is Phenotype-specific and Regulated Primarily at the Level of Gene Transcription-
The underlying mechanism of developing drug resistance in relapsed SCLC patients is unclear. This resistance may be due, in part, to a transition of the SCLC toward the NSCLC histology (2,25), which shows poor response rates to chemotherapy. Furthermore, it has been reported that the drug-resistant phenotype is associated with expression and/or activity of PKCs in lung cancer cell lines and lung carcinomas (19,20). Therefore, expression profiling of PKC isoforms was determined by Western blot analysis using isoform-specific antibodies in a panel of human lung cancer cell lines, which includes eight NSCLC lines and four SCLC lines. As shown in Fig. 1A, a striking differential expression pattern for the PKC-⑀ isoform was observed between the NSCLC and SCLC phenotypes: its expression was detected in all of the NSCLC lines but none of the SCLC lines. Expression of other PKC isoforms (␣, ␤I, ␤II, ␥, ␦, , and ) shows no specific association to either phenotype, and expression of PKC-and -isoforms was undetectable ( Fig. 1A and data not shown). Furthermore, NCI-H82 SCLC cells, when phenotypically transformed to the NSCLC-like phenotype by oncogenic Ras, express PKC-⑀ (Fig. 1A, H82ϩRasV12 lanes) (24), suggesting that PKC-⑀ expression is associated with the NSCLC-phenotype in lung cancer cells. Expression of PKC-⑀ was further examined at levels of steady-state mRNA and gene transcription to determine the molecular basis of the phenotype-specific expression. Northern Blot analysis (Fig. 1B or data not shown) demonstrates that the lack of PKC-⑀ protein FIG. 1. Phenotype-specific expression of the PKC-⑀ isoform in human lung cancer cell lines. A, differential expression of PKC isoforms. Equal amounts of whole-cell lysates prepared from log-phase growing cells were subjected to Western blot analysis with antibodies against specific isoforms, including ␣, ⑀, ␤I, ␤II, and ␦. B, Northern blot analysis. Total cellular RNAs (20 g) were separated on a denaturing agarose gel, transferred to nylon membrane, and hybridized to 32 Plabeled PKC-⑀ cDNA probe. The membrane was stripped and re-probed with the ␤-actin cDNA (middle panel). Lower panel, photography of 18 S RNA on 1% formaldehyde-agarose gel in the presence of ethidium bromide. C, nuclear run-on transcription assay. The 32 P-labeled RNA transcripts isolated from nuclei of representative cell lines were hybridized to the immobilized cDNAs as indicated. ␤-Actin and 18 S ribosomal cDNA were used as positive controls. H82ϩRasV12, two clonal lines of c-Ha-Ras(V12)-transfectants of NCI-H82 (H82) cells (24). expression in SCLC cell lines is associated with a deficiency in the expression of steady-state PKC-⑀ mRNA in these cells. Undetectable expression of PKC-⑀ mRNA in SCLC cells may be due to changes in the message half-life or transcription of the gene. To distinguish between these two possibilities, nuclear run-on transcription experiments were performed to determine the transcription rate for the PKC-⑀ gene. As shown in Fig. 1C, PKC-⑀ gene transcription was undetectable in SCLC lines (H82 and H69), whereas NSCLC lines (H23 and H157) were capable of transcribing the gene. It appears that in SCLC cells the PKC-⑀ gene is transcriptionally inactive. Together, these data suggest that the phenotype-specific expression of PKC-⑀ is controlled primarily at the level of gene transcription.
Expression of PKC-⑀ Enhances Cell Survival to Chemotherapeutic Drugs-It has been reported that the Myc/Ras-mediated phenotypic transition of SCLC toward NSCLC-like phenotype is associated with an acquired resistance to 2-difluoromethylornithine (25). Additionally, we have found that induction of PKC-⑀ expression in Ras-transformed NCI-H82 SCLC cells is correlated with altered drug sensitivity to N 1 ,N 12 -bis(ethyl)spermine, an anti-cancer polyamine analogue. 2 These data suggest that phenotype-specific expression of PKC-⑀ may be important for cellular response to drug treatment. To explore the functional significance of PKC-⑀ expression on chemo-sensitivity, NCI-H82 (H82) human SCLC cells that do not express endogenous PKC-⑀ ( Fig. 1) were stably transfected with an expression plasmid encoding the FLAG epitope-tagged full-length human PKC-⑀. Several G418-resistant clones were isolated and expanded, in which PKC-⑀ expression was confirmed by Western blot analysis using anti-PKC-⑀ antibody ( Fig. 2A). Three PKC-⑀-expressing clonal cell lines (1, 7, and 12) were used in this study. As a control, a pool of G418-resistant clones from the empty vector-transfected NCI-H82 cells was generated and designated as H82-Vector. Compared with the control (H82-Vector) cells, PKC-⑀-expressing clonal cell lines displayed very similar growth kinetics in complete medium (Fig. 2B) but showed an increased ability to grow on soft agar (Table I). Differences in the anchorage-independent colony formation between PKC-⑀-expressing cells and H82-Vector cells were statistically significant (p values were 0.0034, 0.0004, and 0.014 for PKC-⑀ clones #1, #7, and #12, respectively, compared with H82-Vector). It should be noted that forced expression of PKC-⑀ did not result in an alteration of expression levels of other PKC isoforms and had no significant effect on cell cycle progression ( Fig. 2A and data not shown).
Strikingly, significantly increased cell survival was observed in PKC-⑀-expressing cells relative to control cells (H82-Vector) in response to the chemotherapeutic drugs, etoposide and doxorubicin, at all concentrations tested (Fig. 2, C and D). Cell viability studies indicate that induction of PKC-⑀ expression enhanced cell survival in a dose-and time-dependent manner (data not shown). The IC 50 values were increased by 4-to 5-fold for etoposide and 2-fold for doxorubicin in PKC-⑀-expressing cells, compared with H82-Vector cells (Table I) of PKC-⑀-expressing cells remained viable. At higher concentrations of the drug, i.e. at 1 M etoposide, although the viability of PKC-⑀-expressing cells decreased to 28 -35%, they maintained the survival advantage over H82-Vector cells, compared with 2% viability (Fig. 2C). Although the PKC-⑀ expression level varied in three clonal cell lines ( Fig. 2A), no significant differences were observed with respect to chemo-resistance. These data demonstrate that PKC-⑀ has protective effects on drug-induced cytotoxicity and that its induction contributes to the development of drug resistance in NCI-H82 SCLC cells.
Expression of PKC-⑀ Suppresses Etoposide-induced Apoptosis-Etoposide has been shown to induce both necrosis and apoptosis in cancer cells. To determine the nature of etoposideinduced cell death in H82 cells, we assessed morphological changes in the nuclear/chromatin structure of cells in response to etoposide by double-staining cells with Hoechst 33342 and propidium iodide, followed by fluorescence microscopy. It was found that, in H82-Vector cells, etoposide induced morphological alterations typical for apoptosis (26), including chromatin condensation and fragmentation, nuclear swelling, and the appearance of apoptotic bodies (Fig. 3A, upper panel). However, etoposide-treated PKC-⑀-expressing cells retained much of their original morphological features with few condensed nuclei (Fig. 3A, lower panel). Furthermore, time course studies indicate that expression of PKC-⑀ profoundly delayed the onset of apoptosis induced by etoposide (Fig. 3B). Consistent with this result, internucleosomal DNA fragmentation was detected at early time points between 48 and 72 h in H82-Vector cellstreated with etoposide but was not detected until later points (Ͼ96 h) in PKC-⑀-expressing cells (Fig. 3C). Importantly, the integrity of the genomic DNA from PKC-⑀-expressing cells was largely retained, whereas that of H82-Vector cells was significantly affected in response to etoposide, showing a clear sign of degradation in the large-size genomic DNA in addition to the internucleosomal DNA fragmentation. These data indicate that PKC-⑀-mediated cell survival is associated with the inhibition of apoptosis.
It was reported that exposure to topoisomerase II inhibitors, such as etoposide, induce S phase delay in SCLC cells that may contribute to differential sensitivity to the drug (27). The protective effect of PKC-⑀ against etoposide-induced apoptosis may reflect the ability of PKC-⑀ to disrupt cell cycle control mechanism. Effects of PKC-⑀ expression on cell cycle progression were determined by flow cytometry analysis to examine DNA contents of etoposide-treated and untreated cells. As shown in Fig.  3D, untreated H82-Vector and H82-PKC-⑀-expressing cells were distributed very similar among different phases of cell cycle. However, marked differences were observed in etoposidetreated cells. The sub-G 1 population (DNA content Ͻ2N) was increased in H82-Vector cells from 6.7% to 20.4%, reflecting increases in the proportion of apoptotic cells, whereas that of PKC-⑀-expressing cells remained basically unchanged (4.6% versus 7.1%). Time course studies indicate that increases in sub-G 1 population was time-dependent, which occurred at 24 h of post-treatment and progressively increased over the 96-h treatment period (data not shown). It appears that in response to etoposide, the increased sub-G 1 population was accompanied by a decrease in the G 1 population in H82-Vector cells, whereas populations in the S and G 2 /M phases were very similar among PKC-⑀-expressing cells and H82-Vector cells. These results further confirm a role of PKC-⑀ on the inhibition of the etoposideinduced apoptosis, suggesting that the influence of PKC-⑀ expression on cell survival may affect events that control the G 1 /S phase checkpoint function in response to DNA damage.

Expression of PKC-⑀ Results in Inhibition of Etoposide-induced
Caspase Activation-Caspase activation plays a central role in the execution of apoptosis. Studies have suggested that genotoxic agents such as etoposide induce apoptosis via the mitochondrial/caspase-9-mediated pathway, which involves sequential activation of caspase-9 and caspase-3 (8,28). It is possible that the protective effect of PKC-⑀ on etoposide-induced apoptosis may result from blocking mitochondria-dependent caspase activation. Therefore, activation of procaspase-9 was evaluated in response to etoposide treatment by Western blot analysis using antibodies that specifically detect the 37-kDa cleaved caspase-9 (the active form). As shown in Fig. 4A, treatment of H82-Vector cells with etoposide induced the proteolytic cleavage of procaspase-9 into a 37-kDa fragment, whereas the etoposide-induced procaspase-9 cleavage was blocked in PKC-⑀-expressing cells. Consistent with caspase-9 activation, etoposide treatment in H82-Vector cells led to caspase-3 activation, evident by the appearance of the cleaved caspase-3 product (19-kDa). By contrast, the proteolytic cleavage of caspase-3 was abrogated in PKC-⑀-expressing cells. Consistently, we found that the cellular activity of caspase-3 was attenuated in PKC-⑀-expressing cells in response to treatment with etoposide (Fig. 4C). Together, these findings suggest that PKC-⑀ is involved in the regulation of etoposide-induced activation of caspase-9 and caspase-3. It should be noted that a faint band corresponding to the 37-kDa cleaved caspase-9 product was detected in both H82-Vectorand PKC-⑀-expressing cells in the absence of etoposide, suggesting that there is a basal level of caspase-9 activation. This may be due, in part, to the basal level of apoptosis seen in these cells (Fig. 3D, sub-G 1

populations in untreated cells).
Caspase-3 is one of the key executioners of apoptosis, being responsible either partially or totally for the proteolytic cleavage of many key proteins, such as poly(ADP-ribose) polymerase (PARP). Intact PARP (116 kDa) is cleaved into 24-and 89-kDa fragments representing the N-terminal DNA binding domain and the C-terminal catalytic domain of the enzyme. To further validate the induction of apoptosis by etoposide, Western blot analysis was performed to examine the cleavage of PARP in response to drug treatment. Consistent with caspase-3 activa- tion, etoposide treatment resulted in the cleavage of PARP only in H82-Vector cells, as shown by the appearance of an 89-kDa fragment (Fig. 4B). To determine if the cleavage of PARP is caspase-dependent, the cell-permeable, irreversible, caspase inhibitors z-VAD-fmk and z-DEVD-fmk were used. H82-Vector cells were pretreated with the inhibitors for 1 h prior to the addition of etoposide. As seen in Fig. 5A, inclusion of either caspase inhibitor significantly reduced the PARP cleavage in response to etoposide, although z-VAD-fmk was more potent. z-VAD-fmk shows a broad range of specificity against multiple caspases and is thought to inhibit apoptosis at an early stage, prior to activation of caspase-2, -3, -6, or -7, which may explain its higher potency (29). Consistently, we found that z-VAD-fmk was more efficient in decreasing the accumulation of the sub-G 1 population in response to etoposide in H82-Vector cells (Fig. 5B). At the concentration of 20 M, treatment with z-VADfmk resulted in a significant decrease (50% of reductions, p Ͻ 0.02) in the sub-G 1 population as compared with that of the etoposide treatment alone, whereas z-DEVD-fmk had less effect (35% of reduction, p Ͻ 0.05). Together, these observations suggest that PKC-⑀-mediated cell survival to etoposide is due, at least in part, to the inhibition of drug-induced activation of caspase-dependent apoptotic signaling pathway(s).

PKC-⑀ Blocks Etoposide-induced
Release of Cytochrome c-Apoptosis induced by chemotherapeutic agents or ␥-irradiation is likely mediated by the mitochondria pathway (28). Recent studies have demonstrated that the primary regulatory step for mitochondria-mediated caspase activation might be at the level of cytochrome c release from the mitochondrial intermembrane space to the cytosol (8). Therefore, levels of cytochrome c in the cytosol were determined by subcellular fractionation followed by Western blot analysis. As shown in Fig. 6A etoposide treatment in H82-Vector cells resulted in an increased level of cytochrome c in the cytosolic fraction, whereas levels of cytochrome c in PKC-⑀-expressing cells were not affected by etoposide. Interestingly, low levels of cytochrome c were readily detected in the cytosolic fraction of PKC-expressing cells in the absence of etoposide. It is unlikely that this was due to mitochondrial contaminations, because the expression of cytochrome c oxidase subunit IV (COX IV), an integral mitochondrial membrane protein, can not be detected in the cytosolic fraction. It may, therefore, reflect a basal level of cytochrome c release in cells that is independent of etoposide treatment. These data suggest that one possible mechanism underlying PKC-mediated cell survival is to prevent cytochrome c translocation to the cytosol, thereby inhibiting etoposide-induced caspase activation.
Down-regulation of PKC-⑀ Expression with Antisense cDNA Sensitizes NSCLC Cells to Etoposide-The effect of PKC-⑀ induction on survival of SCLC cells suggests that the status of PKC-⑀ expression may correlate to how well the cells respond to chemotherapeutic drugs. This hypothesis prompted us to investigate the effect of specific down-regulation of PKC-⑀ expression on drug sensitivity in NSCLC cells. NCI-H157 human NSCLC cells, expressing a high level of PKC-⑀ (Fig. 1A, H157 lane), were transfected with an empty vector or a plasmid expressing the human PKC-⑀ antisense cDNA. Transfected cells were selected for G418 resistance and down-regulation of the PKC-⑀ protein level in the antisense-expressing cells was confirmed by Western blotting (Fig. 7, inset). Clonogenic cell survival assays were performed to determine the etoposide sensitivity. Results (Fig. 7) demonstrate that PKC-⑀ downregulation (depletion) led to a profound change in sensitivity to etoposide. The cloning efficiency of the antisense-expressing cells was about 80% of that of vector-transfected cells in the absence of etoposide. After exposure to etoposide, the cloning efficiency of antisense-expressing cells was reduced to 64 and 38% at 100 and 500 nM of etoposide, respectively. These data support the notion that the expression level of PKC-⑀ is a key determinant of cellular susceptibility to etoposide in lung cancer cells. DISCUSSION In this study, we have revealed a phenotype-specific expression pattern for the PKC-⑀ isoform in human lung cancer cells.
Our studies indicate that the transcriptional control of the PKC-⑀ gene is a critical step for its phenotype-specific expression. Furthermore, induction of PKC-⑀ expression renders cells resistant to chemotherapy, whereas its down-regulation (depletion) by antisense cDNA results in an enhanced chemo-sensitivity. These results suggest that regulation of the PKC-⑀ gene expression is a key event involved in cellular response to chemotherapeutic drugs. It is predicable that changes in PKC-⑀ gene expression would lead to alternations of the expression of many cellular genes, which may be the genetic determinants of chemo-sensitivity and chemo-resistance. Therefore, transcriptional activation of PKC-⑀ gene provides an intriguing mechanism by which drug resistance might be regulated.
PKC-⑀ is a member of the novel PKC subgroup, which appears to play an important role in growth regulation. Among PKC isoforms, PKC-⑀ is the only isoform that has been demonstrated to promote proliferation and transformation in both fibroblast and epithelial cells upon overexpression (30,31). However, we find that expression of PKC-⑀ in NCI-H82 SCLC cells does not lead to profound changes in cell proliferation and serum dependence. Consistently, PKC-⑀-expressing cells display very similar patterns of cell cycle progression as that of H82-Vector cells (Fig. 3D and data not shown). One explana- FIG. 4. Expression of PKC-⑀ blocks etoposide-induced caspase activation and PARP cleavage. Cells were treated with either vehicle or 200 nM etoposide for 96 h. Whole-cell lysates were prepared and subjected to immunoblotting with indicated antibodies. A, activation of caspase-3 and caspase-9 were detected using antibodies that specifically recognizes the cleaved (active caspase) products. The membranes were stripped and re-blotted with antibodies to the corresponding fulllength caspase. Actin expression was detected for loading controls. B, cleavage of PARP was detected using an antibody that recognizes both full-length (116 kDa) and the cleaved (89 kDa) PARP. C, in vitro caspase-3 activity. Cell lysates prepared from etoposide-treated cells were measured for caspase-3-like activity using colorimetric Ac-DEVD-pNA as a substrate. To determine the specificity of caspase activity, cell lysates were incubated with the caspase-3-specific inhibitor, z-DEVDfmk, prior to measuring the activity. Data are means Ϯ S.D. of three independent experiments. tion for seeing no effect of PKC-⑀ on cell proliferation in our system might be due to the difference in the genetic background of the cells studied. Furthermore, the activation of the mitogen-activated protein kinase (MAPK) pathway, the Raf-ERK pathway, is critical for PKC-⑀-mediated cell proliferation and transformation in fibroblasts and epithelial cells (30,32). However, we do not observe the activation of the ERK pathway in response to PKC-⑀ expression (data not shown), suggesting that the inability of PKC-⑀ to activate the ERK pathway might account for its little effect on proliferation. However, PKC-⑀ expression results in increased anchorage-independent growth ( Table I), suggesting that PKC-⑀ may promote cell survival.
Resistance to apoptosis has been implicated as one potential mechanism whereby tumor cells escape death induced by chemotherapy. In this study, we show that induction of PKC-⑀ expression renders cells resistant to the chemotherapeutic drug etoposide, which is accompanied with a significantly reduction in apoptosis. This suggests that the chemo-protective effect of PKC-⑀ is mediated, at least in part, by suppression of drug-induced apoptosis. Etoposide induces the activation of caspase-9 and caspase-3, along with evidence of PARP cleavage. These events are correlated with the release of cytochrome c from the mitochondria to the cytosol, suggesting that mitochondrial pathways are involved in apoptosis induced by etoposide. In contrast, the death receptor-mediated pathway is unlikely to play a role in this process, because etoposide does not induce caspase-8 activation. Moreover, the caspase-8-specific inhibitor, z-IETD-fmk, has no effect on PARP cleavage and the accumulation of the sub-G 1 population in response to etoposide (data not shown). That the pan-caspase inhibitor, z-VAD-fmk, blocks the cleavage of PARP and attenuates the accumulation of the sub-G 1 population suggests that etoposideinduced apoptosis is caspase-dependent. In any detectable etoposide-induced events, including changes in nuclear/chromatin morphology, accumulation of the sub-G 1 population, activation of caspase-9 and -3, and the mitochondrial release of cytochrome c, were blocked or significantly reduced by the expression of PKC-⑀. These findings suggest that PKC-⑀ inhibits etoposide-induced apoptosis at the level of, or upstream from, the mitochondria by preventing the release of apoptosis-promoting factors, such as cytochrome c.
In H82 cells, PKC-⑀ effectively blocks the etoposide-induced release of cytochrome c, thereby resulting in the inhibition of caspase-9 activation. However, PKC-⑀ seems to have little effect on the basal level of release of cytochrome c, which was detected in PKC-⑀-expressing cells as well as H82-Vector cells in the absence of etoposide ( Fig. 6A and data not shown). This is likely due to basal levels of apoptosis occurring independently of PKC-⑀ expression. It was observed that log phase growing cells display a basal level of caspase-9 activation, and a small portion of cells (ϳ5-7%) were accumulated in the sub-G 1 fraction (Fig. 3D). This suggests that the anti-apoptotic function of PKC-⑀ with etoposide may require additional steps such as activation of PKC-⑀ and/or other signaling molecules. In support of this notion, a recent study (33) reported that the ⑀V1-2 peptide derived from the V1 region of PKC-⑀ blocked the protective effect of TPA in tumor necrosis factor ␣ and calphostin C-induced apoptosis in U937 cells. The ⑀V1-2 peptide has been shown to specifically inhibit PKC-⑀ translocation by interfering with its anchoring (34). A plasma membrane translocation mechanism for PKC activation is well described (10), suggesting TPA-induced activation of PKC-⑀ is required for the suppression of apoptosis. It should be noted that, in the absence of etoposide, there is a higher level of cytosolic cytochrome c in PKC-⑀-expressing cells than in H82-Vector cells. However, PKC-⑀-expressing cells do not show an increased level of apoptosis (Fig. 3, A and D), suggesting expression of PKC-⑀ may raise the apoptotic threshold of a cell. It is possible that PKC-⑀ may employ one or more different mechanisms to promote the basal level of survival from that in response to etoposide.
Interestingly, a significant fraction of exogenously expressed PKC-⑀ is localized to the mitochondria-rich heavy membrane (Fig. 6A). It is unclear how PKC-⑀ is localized to the mitochondria. This may result from cytosolic to mitochondrial translocation due to the mitochondrial generation of diacylglycerol as reported previously (35). Several recent studies have suggested that the mitochondrial recruitment of PKC-⑀ may be associated with its activation. In PKC-⑀ transgenic mice, it was found that PKC-⑀ was colocalized with the mitogen-activated protein kinase (MAPK) in the murine heart mitochondria, which appears to be responsible for the increased PKC-⑀-MAPK signaling (36). Ohnuma et al. (37) reported that PKC-⑀ activation by ischemic preconditioning in rabbit hearts is involved in the direct translocation of PKC-⑀ to the mitochondria. Additionally, we have observed that the PKC activator TPA can induce PKC-⑀ translocation from the cytosol to the heavy membrane in lung cancer cells expressing endogenous PKC-⑀. 2 Furthermore, Ruvolo et al. (38) reported that either the treatment of human pre-B REH cells with the PKC agonist Bryostatin-1 or forced expression of exogenous PKC-␣ induces mitochondrial localization of PKC-␣, which is tightly associated with increased Bcl-2 phosphorylation and enhanced resistance to chemotherapy. This study has defined a functional role for mitochondrial associated PKC-␣ in Bcl-2 phosphorylation and in suppressing drug-induced apoptosis (38). Therefore, the mitochondrial associated PKC-⑀ may have a functional significance in mediating its anti-apoptotic function.
A growing body of evidence has suggested a general antiapoptotic role for PKC-⑀ in different cell types in response to a variety of apoptotic stimuli. For examples, PKC-⑀ has been implicated in the protection of cardiac myocytes from hypoxiainduced cell death (34). Overexpression of PKC-⑀ in human TF-1 cells suppressed apoptosis upon IL-3 withdrawal and increased Bcl-2 expression (40). In particular, enhanced PKC-⑀ expression has been linked to a decreased cellular sensitivity to chemotherapeutic drugs, possibly through inhibition of druginduced apoptosis. It has been reported that PKC-⑀ overexpression in R6 rat embryo fibroblasts prevented cisplatin-induced apoptosis and protected cells against cisplatin cytotoxicity (41). Additionally, compared with the cisplatin-sensitive ovarian carcinoma 2008 cells, the resistant-variant 2008/C13*5.25 cells displayed increased PKC-⑀ expression and a significant reduction in PKC-␣ expression (42). Consistent with our findings, Wu et al. (43) recently demonstrated that overexpression of PKC-⑀ rendered LNCaP human prostate cancer cells resistant to apoptosis, which was accompanied with a lack of stimulation of caspase-3 proteolytic activity in response to apoptotic stimuli. In addition, we find that expression of a dominant negative mutant of PKC-⑀ in NSCLC cells results in apoptosis instead of a G 2 /M cell cycle arrest in response to paclitaxel. Apoptosis is accompanied with the activation of caspase-9 and the cleavage of PARP. 2 These data suggest that inhibition of caspase activation may be a common mechanism by which PKC-⑀ protects cells from death. Interestingly, proteolytic activation of PKC-⑀ during chemotherapeutic agent-induced apoptosis by caspase-3 has been reported in U937 cells (44). However, the functional significance of this proteolytic activation of PKC-⑀ is unclear.
In summary, we have defined an anti-apoptotic mechanism by which PKC-⑀ promotes survival of human lung cancer cells to chemotherapeutic drug etoposide. PKC-⑀ induction in H82 SCLC cells confers drug resistance without affecting cell proliferation. The influence of PKC-⑀ on the sensitivity to etoposide is mediated by the inhibition of caspase activation through one or more of the mitochondrial pathways. Although other mechanisms potentially involved in drug resistance (e.g. modulation of P-glycoprotein expression) might also be affected by PKC-⑀ expression (39), our findings suggest that the abnormal expression of PKC-⑀ could raise the apoptotic threshold of cells, thereby leading to resistance to chemotherapy. Elucidating the precise mechanism(s) of drug-induced apoptosis and how PKC-⑀ may affect this process in mediating lung cancer cell survival will likely lead to better therapeutic regimens that target the apoptosis machinery to overcome drug resistance.


extfieldsofstudy: Biology Medicine

text:
ECM regulates MT1-MMP localization with β1 or αvβ3 integrins at distinct cell compartments modulating its internalization and activity on human endothelial cells

Regulation of membrane-type 1 matrix metalloproteinase (MT1-MMP) by different extracellular matrices (ECMs) on human endothelial cells (ECs) has been investigated. First, MT1-MMP is found at the intercellular contacts of confluent ECs grown on β1 integrin–dependent matrix such as type 1 collagen (COL I), fibronectin (FN), or fibrinogen (FG), but not on gelatin (GEL) or vitronectin (VN). The novel localization of MT1-MMP at cell–cell contacts is assessed by confocal videomicroscopy of MT1-MMP-GFP–transfected ECs. Moreover, MT1-MMP colocalizes with β1 integrins at the intercellular contacts, whereas it is preferentially found with αvβ3 integrin at motility-associated structures on migrating ECs. In addition, clustered integrins recruit MT1-MMP and neutralizing anti-β1 or anti-αv integrin mAb displace MT1-MMP from its specific sites, pointing to a biochemical association that is finally demonstrated by coimmunoprecipitation assays. On the other hand, COL I, FN, or FG up-regulate cell surface MT1-MMP on confluent ECs by an impairment of its internalization, whereas expression and internalization are not modified on GEL or VN. In addition, MT1-MMP activity is diminished in confluent ECs on COL I, FN, or FG. Finally, MT1-MMP participates and cooperates with β1 and αvβ3 integrins in the migration of ECs on different ECM. These data show a novel mechanism by which ECM regulates MT1-MMP association with β1 or αvβ3 integrins at distinct cellular compartments, thus modulating its internalization, activity, and function on human ECs.

Introduction
During angiogenesis, ECM changes dynamically to provide new binding sites to endothelial cell (EC)* receptors involved in adhesion and migration (Stupack and Cheresh, 2002). ECM supports endothelial cell attachment by binding to distinct adhesion receptors being the integrins ␣ 2 ␤ 1, ␣ 5 ␤ 1, and ␣ v ␤ 3, the principal endothelial receptors for type I collagen (COL I), fibronectin (FN), and vitronectin (VN), respectively (Bazzoni et al., 1999). Integrin expression is modulated during angiogenesis by different factors and plays important roles during endothelial migration and the formation of new vessels as shown by the blocking effects of ␣ v ␤ 3 antagonists (Stupack and Cheresh, 2002).
The dynamics of ECM during the angiogenic response is accomplished by the deposition of provisional components as well as by its remodeling by proteases (Werb, 1997). The major enzymes involved in degrading ECM belong to the matrix metalloproteinase (MMP) family (Nagase and Woessner, 1999). Membrane-type 1 matrix metalloproteinase (MT1-MMP), one of the membrane-anchored metalloproteinases, degrades several ECM components including type I, II, and III collagens, gelatin (GEL), VN, FN, fibrinogen (FG), and laminins 1 and 5 apart from activating pro-MMP-2 and pro-MMP-13 (Werb, 1997). MT1-MMP is also the best candidate for focused proteolysis because it is recruited to the invadopodia and to the leading edge of human melanoma cells (Nakahara et al., 1997;Lehti et al., 2000). Proteolytic ECM remodeling not only removes ECM, but also provides a promigratory environment. Thus, cleavage of laminin-5 by MT1-MMP induces epithelial cell migration (Koshikawa et al., 2000). In turn, integrins, by interaction with ECM, regulate pro-MMP-2 activation and MT1-MMP expression in human capillary ECs and in tumor cells (Stanton et al., 1998;Yan et al., 2000;Ellerbroek et al., 2001). Recent studies have demonstrated a functional interaction between MT1-MMP and integrins. Thus, it is known that ␣ v ␤ 3, induced on vascular sprouts during angiogenesis, interacts with MMP-2, a substrate for MT1-MMP (Brooks et al., 1996). It has also been reported the cooperative role of MT1-MMP and ␣ v ␤ 3 in activating pro-MMP-2 (Hofmann et al., 2000;Deryugina et al., 2001). Finally, MT1-MMP has been shown to directly participate in ␣ v processing during its maturation, thus modulating the adhesive and migratory behavior of tumor cells Ratnikov et al., 2002). However, the mechanisms by which ECM and integrins might regulate MMP functionality in human ECs remain unexplored.
Here, we report that ECM modulates MT1-MMP association with ␤ 1 and ␣ v ␤ 3 integrins at specific cell sites on ECs, thus regulating MT1-MMP internalization, expression, and activation. Moreover, MT1-MMP and integrins are shown to cooperate during the migration of human ECs, a critical step in the angiogenic response.

MT1-MMP localizes at the intercellular contacts of ECs on COL I, FN, or FG
The effect of different ECM in regulating MT1-MMP on human ECs was investigated. First, the localization of MT1-MMP was found to be different depending on the matrix. As shown in Fig. 1 A, MT1-MMP decorated intercellular contacts of ECs adhered to COL I, FN, or FG, in contrast to the diffuse staining observed on GEL and VN. To elucidate the adhesion receptors contributing to attachment of ECs to the different ECM, adhesion assays in the presence of neutralizing anti-integrin antibodies were performed. As shown in Fig. 1 B, ECs mainly adhered to COL I or FN in a ␤ 1 integrin-dependent manner, whereas contribution of ␣ v ␤ 3 integrins was lower (90 and 60% inhibition, respectively). In contrast, EC adhesion to GEL or VN was greatly reduced by anti-␣ v or anti-␣ v ␤ 3, and at a lower extent by anti-␤ 1 integrin mAb (80 and 50% inhibition, respectively). Both ␤ 1 and ␣ v integrins similarly contributed to EC adhesion to FG ( Fig.  1 B). Thus, distinct recognition of ECM by integrins seems to regulate MT1-MMP subcellular localization on human ECs.
The novel MT1-MMP localization at cell-cell contacts was also assessed through an independent approach. Fulllength MT1-MMP was fused with GFP, and the construct was transfected into ECs. MT1-MMP-GFP protein had an expected molecular mass of ف 90 kD and was active as analyzed by Western blot and zymography (Fig. 2, A and B, respectively). The dynamics of MT1-MMP subcellular localization was observed by time-lapse confocal videomicroscopy. Transiently transfected ECs displayed an enrichment of MT1-MMP-GFP at cell sites contacting other cells (Fig. 2 C; Video 1). In addition, an active traffic of MT1-MMP-GFP-positive intracellular vesicles to the membrane, forming clusters at motility-associated structures, was also observed (Fig. 2 C; Video 1).

MT1-MMP distinctly colocalizes with ␤ 1 or ␣ v ␤ 3 integrins on ECs depending on the ECM and the migratory state
Because MT1-MMP could be found at interendothelial contacts and motility-associated structures of ECs, its putative association with integrins also present at those sites was investigated. Interestingly, MT1-MMP colocalized with ␤ 1 integrins at the intercellular junctions of confluent ECs on COL I, but not at the motility structures of migrating cells (Fig. 3 A). ␤ 1 integrins were occasionally observed at the leading edge and lamellipodia of ECs together with MT1-MMP (unpublished data). Upon wound-induced migration, MT1-MMP was recruited to discrete regions of motile structures independently of the matrix (GEL or COL I), where it colocalized with ␣ v ␤ 3 integrin (Fig. 3 B). However, ␣ v ␤ 3 integrin had a diffuse pattern on confluent ECs and was not observed at cell-cell contacts on COL I (Fig. 3 B).
To assess the putative role of the MT1-MMP cytoplasmic domain in its matrix-regulated localization, ECs were transiently transfected with either MT1-MMP-GFP or MT1-MMP ⌬ cyt-GFP constructs. As shown for the full-length protein, the MT1-MMP ⌬ cyt-GFP fusion protein had an expected molecular mass of ف 90 kD and proteolytic activity in both FG and GEL zymography; however, the expression of mature MT1-MMP ⌬ cyt-GFP was lower than the full-length protein as demonstrated by the Western blot analysis (Fig. 2  A). Although MT1-MMP-GFP was found colocalizing with ␤ 1 or ␣ v ␤ 3 integrins at some intercellular contacts as well as at motility structures of migrating ECs on COL I, respectively ( Fig. 3 C), MT1-MMP ⌬ cyt-GFP could rarely be observed at those cell sites colocalizing with integrins ( Fig. 3 C). The GFP protein transfected into ECs did not display any specific localization (Fig. 3 C). These data suggest that the cy- ECs were grown to confluency on coverslips coated with 1% GEL or 10 g/ml VN, COL I, FN, or FG for 24 h and then stained for immunofluorescence analysis with the anti-MT1-MMP mAb LEM-2/15. Negative control of ECs on COL I without primary antibody is also included. Bar, 20 m. (B) Adhesion assays of ECs to plates coated with 1% GEL or 10 g/ml VN, COL I, FN, or FG were performed for 30 min at 37ЊC in the absence or presence of 10 g/ml neutralizing anti-integrin mAb (␤1, ␣v, or ␣v␤3). The arithmetic mean Ϯ SD of three independent experiments run in triplicate is shown.
tosolic domain of MT1-MMP might participate in its localization with either ␤ 1 or ␣ v ␤ 3 integrins at specific cell sites.

MT1-MMP can associate with ␤ 1 and ␣ v ␤ 3 integrins on human ECs
Because MT1-MMP was observed to colocalize with ␤ 1 and ␣ v ␤ 3 integrins at cell-cell contacts and motility structures, respectively, the association of these proteins was further investigated. As shown in Fig. 4 A, ␤ 1 integrin clustering induced by COL I-coated beads resulted in an efficient redistribution of MT1-MMP around the clustered integrins. Similarly, ␣ v ␤ 3 integrin clustering also promoted rearrangement of MT1-MMP around the COL I-coated beads (Fig.  4 A). However, no MT1-MMP staining was observed around occasional CD31 clustering, and also, CD31 did not redistribute around ␤ 1 or ␣ v ␤ 3 integrins (Fig. 4 A). Similar results were obtained with GEL, FN, or FG-coated latex beads, although the efficiency in recruiting ␤ 1 and ␣ v ␤ 3 integrins as well as in the recruitment of MT1-MMP by these integrins was slightly different depending on the ECM used as shown in Fig. 4 A.
Furthermore, treatment of ECs grown on COL I with the neutralizing anti-␤ 1 integrin mAb LIA1/2 impaired MT1-MMP localization at the cell-cell contacts, although it preserved ␤ 1 integrin staining at these sites as well as MT1-MMP localization at motility structures ( Fig. 4 B). On the contrary, treatment of ECs with the neutralizing anti-␣ v integrin mAb ABA 6D1 blocked MT1-MMP staining at motility-associated clusters, but neither affect ␣ v integrin localization at these sites nor MT1-MMP localization at cell-cell contacts (Fig. 4 B).
Finally, the biochemical association of MT1-MMP with ␤ 1 or ␣ v ␤ 3 integrins was directly assessed by coimmunoprecipitation assays. Immunoprecipitation of ␤ 1 integrins from ECs lysed in mild conditions showed a faint but consistent band of 60 kD, corresponding to mature MT1-MMP ( Fig.  4 C). However, ␣ v integrins coprecipitated the immature and mature forms of MT1-MMP as a double band of 63 and 60 kD, with 63 kD being the main one ( Fig. 4 C). Anti-MT1-MMP immunoprecipitates also contained traces of ␤ 1 and ␣ v integrins under these conditions (Fig. 4 C).

MT1-MMP surface expression is distinctly modulated by integrin-ECM interactions on human ECs
The intensity of MT1-MMP staining seemed to be higher when localized at the cell-cell contacts of ECs on COL I, FN, or FG compared with GEL or VN ( Fig. 1 A). To investigate this point further, the expression of MT1-MMP on different matrices was quantitated. The expression of MT1-MMP on confluent ECs increased after 6 or 24 h of attachment to COL I, FN, or FG in contrast to GEL or VN as assessed by flow cytometry; this ECM-mediated MT1-MMP up-regulation diminished in the presence of cycloheximide (Fig. 5, A and B). Because migration induced an increase of MT1-MMP expression on ECs on GEL (Gálvez et al., 2001), the effect of migration on different matrix in modulating MT1-MMP expression was also analyzed. Woundinduced migration further enhanced MT1-MMP surface expression on VN, COL I, FN, and FG, suggesting a matrix-independent effect; moreover, the migration-induced increase was inhibited in the presence of cycloheximide (Fig.  5, A and B).  Next, we analyzed whether ECM was mediating the effects observed in MT1-MMP expression by distinct clustering of integrins. First, clustering of ␤1 integrins was independently achieved by coating plates with either the activating TS2/16 or the neutralizing LIA1/2 anti-␤1 integrin mAb. Both mAb promoted a consistent accumulation of cell surface MT1-MMP on ECs, pointing to clustering rather than activation as the mechanism responsible for integrinmediated effects (Fig. 5 C). Additionally, clustering ␣v␤3 integrins by coating plates with LM609 mAb increased MT1-MMP expression at a lower extent (Fig. 5 C). Anti-␣3 and anti-␣5 integrin mAb also up-regulated MT1-MMP surface expression in contrast to anti-␣2 integrin and to the control anti-CD31 mAb (Fig. 5 C).
MT1-MMP level was also analyzed by Western blot of total cellular lysates from ECs grown on different ECM. No major changes in the whole amount of MT1-MMP were detected on lysates of ECs grown on GEL and VN versus COL I, FN, or FG in contrast to wound-healing stimulation that induced a similar MT1-MMP up-regulation on all substrates ( Fig. 5 D). These results suggest that other mechanisms apart from increased expression are likely involved in the up-regulation of MT1-MMP on the surface of confluent ECs by distinct ECM.

Integrin-ECM interactions regulate MT1-MMP internalization on human ECs
MT1-MMP internalization has been proposed as a new mechanism for regulating MT1-MMP balance on the cell surface (Jiang et al., 2001;Uekita et al., 2001). Because the ECM-mediated increase in MT1-MMP endothelial surface expression did not correlate with changes in the total MT1- (A) Double staining of COL I-coated beads added to migratory ECs with biotinylated anti-MT1-MMP and anti-␤1 or anti-␣v␤3 integrin mAb was performed. MT1-MMP (green) is recruited around beads where ␤1 or ␣v␤3 integrin (red) are clustered. CD31 staining (red) was also included as negative control. DIC images are also shown. Bar, 20 m. The percentage of stained beads with the different combination of mAb and the efficiency of beads in clustering ␤1 or ␣v␤3 integrins are shown in the right panel. The arithmetic mean of three independent experiments is represented. (B) MT1-MMP (green) and integrin (red) staining was performed on confluent or migratory ECs grown on COL I and preincubated or not with the neutralizing anti-␤1 LIA1/2 or anti-␣v integrin ABA 6D1 mAb. Note in merged images (yellow) that MT1-MMP colocalization at cell-cell contacts and motile structures (arrowheads) is impaired by pretreatment with anti-␤1 or anti-␣v integrin mAb, respectively. Bar, 20 m. (C) Immunoprecipitation with anti-␤1 TS2/16 or anti-␣v␤3 integrin LM609 mAb from ECs on COL I lysed in mild conditions was performed. Western blot of these immunoprecipates with anti-MT1-MMP LEM-2/15 mAb shows bands of 60 kD (asterisk) and 63/60 kD (arrowhead/asterisk), respectively, that correspond to the mature (60 kD) and immature (63 kD) forms of MT1-MMP, as well as other nonspecific lower bands. Anti-MT1-MMP LEM-2/63 mAb also pulls down faint bands corresponding to ␤1 and ␣v integrins. One out of three independent experiments is shown.
MMP amount analyzed by Western blot, the putative regulatory role of ECM on MT1-MMP internalization was investigated. Transferrin receptor was included in the analysis as a well-characterized marker of endothelial endocytosis (Moos and Morgan, 2000).
The internalization kinetics of MT1-MMP on GEL was rapid, being complete at 6 h; however, no MT1-MMP internalization was observed on ECs grown on COL I, and after 6 h all the MT1-MMP still remained at the cell surface (Fig. 6, A and C). VN behaved similarly to GEL and FN or FG to COL I in this respect (unpublished data). The effect of COL I on impairing MT1-MMP internalization was mimicked by clustering ␤1 integrins; in contrast, clustering ␣v␤3 integrins resulted in a similar MT1-MMP internalization to that obtained on GEL (Fig. 6 B). Besides, MT1-MMP internalization on COL I was completely restored upon wound-induced migration of ECs (Fig. 6, A and C). The association of MT1-MMP with ␤1 or ␣v␤3 integrins might be related to the regulation of its internalization because ␤1 integrins were not internalized compared with ␣v␤3 integrin that was completely internalized after 6 h independently of matrix or migration (unpublished data).
The initially labeled MT1-MMP at the cell surface was almost completely recovered in permeabilized cells after 6 h of receptor internalization, ruling out shedding as an important mechanism for the decrease of the protease at the cell surface (Fig. 6, A and B). To confirm this point, MT1-MMP internalization was also visualized by immunofluorescence microscopy. As shown in Fig. 6 D, after 6 h at 37ЊC, MT1-MMP was internalized in a vesicle-like pattern similar to the transferrin receptor on ECs grown on GEL in contrast to COL I on which MT1-MMP staining remained mostly at the cell-cell contacts.

MT1-MMP activity is diminished on human ECs on ␤1 integrin-dependent substrates
Because ECM modulates MT1-MMP localization and internalization, the effect of distinct ECM on MT1-MMP activity was also assessed by FG and GEL zymography. Despite higher levels of MT1-MMP on the cell membrane, no detectable FG degradation was observed on confluent ECs grown on COL I, FN, or FG compared with GEL or VN, in which a basal fibrinolytic activity was present (Fig. 7 A). Moreover, wound-induced migration stimulated FG degradation in ECs cultured on COL I, FN, or FG, but much less efficiently than on GEL (Fig. 7 A). Similar results were obtained when assessing pro-MMP-2 processing by GEL zymography of lysates and supernatants (Fig. 7 A). Moreover, the fraction of pro-MMP-2 associated to the cell lysate was reduced in resting ECs grown on COL I, FN, or FG in contrast to the increase of this pro-form observed in the corresponding supernatant (Fig. 7 A). No significant changes in TIMP-2 levels were detected under the different conditions (unpublished data). These data show a down-regulation of MT1-MMP activity on ␤1 integrin-dependent substrates.
ECM effects on MT1-MMP activity were likely mediated by integrin interactions. Thus, clustering ␤1 integrins with the two different mAbs, activating TS2/16 and neutralizing LIA1/2, mimicked the effects seen on COL I, FN, or FG with no MT1-MMP activation on confluent cells and reduced MT1-MMP activation on migratory cells (Fig. 7 B). In contrast, clustering ␣v integrins with ABA 6D1 mAb elicited an activation of MT1-MMP comparable to that observed on GEL or VN (Fig. 7 B). Therefore, MT1-MMP activity is down-regulated by ␤1 integrin-mediated interactions with COL I, FN, and FG, whereas ␣v integrin-mediated interactions with GEL or VN, like wound-induced migration, enable MT1-MMP activation.
To analyze the relation between regulation of MT1-MMP internalization and activity, okadaic acid (a modulator of receptor internalization) was used (Parton et al., 1994). Interestingly, upon treatment with okadaic acid, MT1-MMP internalization increased up to 50% in confluent ECs grown on COL I. As shown in Fig. 7 C, treatment with okadaic acid also restored the activity of MT1-MMP on COL I, pointing to a link between MT1-MMP internalization and its activation state.

MT1-MMP participates and cooperates with integrins during EC migration on ECM
During EC migration, MT1-MMP localizes at migratory structures on different ECMs, its expression increases, and its activity is up-regulated. Therefore, the role of MT1-MMP during EC migration on ECM was investigated. First, it was observed that anti-MT1-MMP inhibitory mAb LEM-2/15, when used at saturating doses of 10 g/ml, prevented EC migration on COL I, FN, FG, or VN by more than 50% on wound-healing and transmigration assays (Fig. 8 A). Interestingly, the Fab monovalent fragment of anti-MT1-MMP LEM-2/15 mAb inhibited EC migration on COL I at a similar extent to the complete IgG, suggesting that clustering of MT1-MMP is not necessary for the impairment of migration. Anti-VE cadherin and anti-␤1 integrin TS2/16 mAb or anti-␣v integrin ABA 6D1 mAb were also included as negative and positive controls of inhibition, respectively (Fig. 8 A). The role of MT1-MMP in EC migration was also explored by transiently transfecting ECs with GFP, MT1-MMP-GFP, or MT1-MMP⌬cyt-GFP constructs and assessing their ability to migrate across COL I-coated filters. Interestingly, ECs transfected with the truncated mutant showed an impaired migration when compared with ECs transfected with the full-length protein, suggesting that intact MT1-MMP is required for migration of ECs on COL I (Fig. 8 B). Moreover, MT1-MMP-dependent migration was significantly inhibited by anti-MT1-MMP mAb on ECs transfected with MT1-MMP-GFP (Fig. 8 B).
Finally, we wanted to analyze whether the association of MT1-MMP with ␤1 and ␣v integrins might also have a functional relevance during EC migration. The combination of the inhibitory anti-MT1-MMP mAb LEM-2/15 at suboptimal doses with anti-␤1 and anti-␣v integrin mAb resulted in an additive inhibitory effect on EC migration on COL I in wound-healing assays (Fig. 8 C). Furthermore, pretreatment of ECs with suboptimal doses of anti-MT1-MMP and either anti-␤1 or anti-␣v integrin mAb had an additive inhibitory effect on EC transmigration on COL I or VN, i.e., the migration was completely abolished when the three mAbs were used in combination (Fig. 8 C). Similar results were obtained when cells were migrating on FN or FG (unpublished data). These findings suggest a functional cooperation of MT1-MMP with ␤1 and ␣v integrins in modulating the migratory potential of human ECs.

Discussion
We have reported a novel role of the ECM in regulating MT1-MMP subcellular localization, internalization, and activity in human ECs through its differential association with ␤1 or ␣v␤3 integrins at cell-cell contacts and motility structures, respectively (Fig. 9).
MT1-MMP has been involved in pericellular proteolysis by focusing ECM degradation to specific cell sites such as invadopodia or the cellular leading edge (Nakahara et al., 1997;Lehti et al., 2000). However, we have demonstrated a novel localization of MT1-MMP at endothelial cell-cell contacts on COL I, FN, and FG, where it colocalizes with ␤1 integrins. Dynamic assessment of MT1-MMP localization by fluorescence microscopy in live cells during EC mi- Lysates from resting or woundhealing-stimulated ECs grown on plates coated with 10 g/ml of anti-␤1 activ TS2/16, anti-␤1 neutr LIA1/2, anti-␣v integrin ABA 6D1, or anti-CD31 TP1/15 mAb included as control were analyzed by FG and GEL zymography. Lysates of ECs grown on GEL or COL I are also included for comparison. A representative out of three experiments is shown. (C) Lysates from confluent ECs grown on COL I and pretreated or not overnight with 10 M okadaic acid were analyzed by FG and GEL zymography. A representative out of two experiments is shown.
gration confirmed an enrichment of MT1-MMP at areas where ECs contact each other. The direct association of MT1-MMP with ␤1 integrins and/or with other receptors present at endothelial junctions (such as claudins) might be involved in the recruitment to these sites as recently shown in 293T-transfected cells (Miyamori et al., 2001). Conceivably, MT1-MMP at EC contacts might constitute a reservoir ready to be mobilized to other cell sites or to be activated in situ. Moreover, emerging new functions for MT1-MMP, such as cleavage of endothelial intercellular re-ceptors for detachment of ECs from each other during migration or for leukocyte transmigration might rely on that new localization. However, under migratory conditions and independently of the ECM, MT1-MMP is recruited to regions on ECs likely involved in ECM degradation colocalizing with ␣v␤3 similarly to tumoral cells (Hofmann et al., 2000;Deryugina et al., 2001). Thus, integrins could form complexes with proteases to focus EC proteolytic activity more efficiently. In this regard, it has been proposed that ␣v␤3/MT1-MMP cooperation would be required to fully assessed by wound-healing and/or transmigration assays at 8 and 5 h, respectively. Migration was analyzed in the absence or presence of saturating doses (10 g/ml) of the inhibitory anti-MT1-MMP mAb LEM-2/15. The inhibitory effects obtained were statistically significant for wound healing (*, P Ͻ 0.04; **, P Ͻ 0.01) and for transmigration assays (ϩ, P Ͻ 0.02; ϩϩ, P Ͻ 0.015). Fab monovalent fragments of anti-MT1-MMP LEM-2/15 mAb used at 10 g/ml also had an inhibitory effect on COL I (ϩϩ, P Ͻ 0.015). Anti-␤1 TS2/16 and anti-␣v integrin ABA 6D1 mAb were also included. Anti-VE-cadherin TEA1/31 mAb was used as negative control. The arithmetic mean Ϯ SD of a representative out of four independent experiments run in duplicate is shown. (B) ECs were transiently transfected with GFP, MT1-MMP-GFP, or MT1-MMP⌬cyt-GFP constructs and then assayed for migration across COL I-coated filters for 5 h. The number of transfected cells counted in eight independent fields and normalized respect the efficiency of transfection is represented. The impaired migration observed with the mutant as well as the inhibition of migration of MT1-MMP-GFP transfected cells by anti-MT1-MMP mAb were statistically significant (*, P Ͻ 0.03). The arithmetic mean Ϯ SD of two independent experiments run in duplicate is shown. (C) EC migration on COL I or transmigration across COL I or VN-coated filters were analyzed at 8 and 5 h, respectively, in the absence or presence of suboptimal doses (5 g/ml) of one, two, or three of anti-MT1-MMP LEM-2/15, anti-␤1 TS2/16, or anti-␣v integrin ABA 6D1 mAb. Anti-CD31 mAb TP1/15 was included as negative control. The inhibitory effects obtained were statistically significant for wound-healing (*, P Ͻ 0.04; **, P Ͻ 0.02) and transmigration assays (ϩ, P Ͻ 0.01; ϩϩ, P Ͻ 0.005). The arithmetic mean Ϯ SD of a representative out of three independent experiments run in duplicate is shown.
activate pro-MMP-2 in human breast carcinoma cells . Also, a protease-docking function of ␣3␤1 integrin at invadopodia by forming a functional complex with seprase has been shown in melanoma cells (Mueller et al., 1999). Interestingly, MT1-MMP cytosolic domain might be participating in the localization with ␤1 and ␣v␤3 integrins at cell-cell contacts and motility structures, respectively, as suggested by immunofluorescence staining of transfected ECs.
The colocalization of MT1-MMP with ␤1 or ␣v␤3 integrins pointed to a biochemical association of these receptors on ECs. MT1-MMP association with ␣v␤3 or ␤1 integrins had been suggested by chemical cross-linking on breast carcinoma cells and by studies with anti-␤1 integrin mAbcoated beads in ovarian carcinoma cells, respectively Ellerbroek et al., 2001). We have demonstrated by coimmunoprecipitation assays that MT1-MMP can associate with ␤1 and ␣v␤3 integrins on primary ECs. This interaction might be important for MT1-MMP localization because anti-␤1 or anti-␣v␤3 integrin neutralizing mAb impaired MT1-MMP localization at cell-cell contacts and motile structures, respectively. Interestingly, ␤1 integrins are mainly associated to the 60-kD mature form of MT1-MMP, pointing to an association with MT1-MMP at the cell membrane. On the other hand, ␣v integrins associate to both the 63-kD immature and the 60-kD mature forms of MT1-MMP, suggesting that both proteins might travel together from intracellular synthesis toward the cell membrane in similar compartments, such as caveolae (Puyraimond et al., 2001). In this regard, the processing of the immature pro-␣v chain by MT1-MMP has recently been demonstrated .
Regulation of cellular proteolysis by ECM-induced increase of protease expression was first reported in fibroblasts interacting with FN (Werb et al., 1989). We have demonstrated two different mechanisms for regulating MT1-MMP expression by ECM interactions on ECs. First, MT1-MMP can be up-regulated by an increase in its synthesis de novo, as seems to be the case upon migration-inducing conditions independent of the ECM type. Accordingly, it has previously been reported that an ECM-mediated signaling increased Egr-1-dependent transcriptional expression of MT1-MMP in rat ECs (Haas et al., 1999). Interestingly, our data reveal an alternative mechanism by which ECM modulates MT1-MMP expression by regulating its internalization. Internalization of MT1-MMP through a dileucine cytoplasmic motif has recently been reported on MT1-MMP-transfected cells, but its endogenous regulation has not been characterized (Jiang et al., 2001;Uekita et al., 2001). We demonstrate that MT1-MMP internalization is abrogated on confluent ECs grown on ␤1 integrin-dependent ECM (COL I, FN, or FG) compared with the constitutive endocytosis on ␣v␤3 integrin-dependent substrates (GEL or VN). This mechanism might explain the ECM-mediated MT1-MMP increase in the absence of mRNA changes reported by other groups (Stanton et al., 1998;Zhuge and Xu, 2001). The increase in MT1-MMP surface expression is also achieved by ␤1 integrin clustering with mAb anti-␤1, anti-␣3, or anti-␣5 integrins, but not by anti-␣2 integrin mAb, pointing to specific ␤1 integrins as responsible for the effect. Possibly, clustering ␤1 integrins might induce phosphorylation or recruit cytoskeletal/signaling machinery that could interfere with the endocytosis motif, thus blocking MT1-MMP internalization. Additionally, physical retention of MT1-MMP through distinct interactions with ␣v␤3 or ␤1 integrins at the cell membrane might enable or block MT1-MMP endocytosis. Interestingly, upon migration-inducing conditions, MT1-MMP internalization is restored in accordance with the requirement of MT1-MMP internalization for its function in cell migration and invasion recently reported (Jiang et al., 2001;Uekita et al., 2001).
Surprisingly, MT1-MMP proteolytic activity is reduced on COL I, FN, or FG despite its increased surface expression. Previous reports had shown an increase of MT1-MMP/MMP-2 activation by distinct ECM such as FN or COL I in tumoral or transformed cells (Stanton et al., 1998;Ellerbroek et al., 2001). However, on human capillary cells or fibroblasts, ECM induced a down-regulation of MMP-2 activation in accordance with our data (Werb et al., 1989;Yan et al., 2000). This suggests that primary and tumor cells might have different mechanisms for regulating MT1-MMP activity through ECM interactions. The decrease in MT1-MMP activity observed in confluent ECs on COL I correlated with the blockade of its internalization. Interestingly, treatment with okadaic acid that enforces MT1-MMP internalization on COL I restores MT1-MMP activity, pointing to a causal relation between the blockade of MT1-MMP internalization and the decrease of its activity. ␤1 integrin association with MT1-MMP might directly interfere with its catalytic domain or induce modifications around its internalization motif also affecting the oligomerization site (Cys 574 ) required for its function (Lehti et al., 2002). Alternatively, ␤1 or ␣v␤3 integrin-mediated signals might distinctly modulate MT1-MMP activity in accordance with previous differences in signaling triggered by COL I and VN reported during EC migration (Leavesley et al., 1993).
Independent of ECM, migration induces MT1-MMP localization at EC motility structures, an increase of its expres- Figure 9. MT1-MMP is regulated by ECM and migration on human ECs. MT1-MMP in confluent ECs on COL I displays a localization at cell-cell contacts with ␤1 integrins, an impaired internalization rate, and a low enzymatic activity. MT1-MMP under migratory conditions exhibits a localization at motility structures with ␣v␤3 integrin, a high rate of internalization, and an induction of enzymatic activity. Blockade of MT1-MMP activity leads to an inhibition of EC migration that is larger when combined with neutralizing ␤1 and/or ␣v␤3 integrin function.
sion without affecting internalization, and an up-regulation of its activity pointing to an active role of this protease during EC migration. Here, it is demonstrated that MT1-MMP is required for EC migration on several ECM because both the whole IgG as well as the Fab monovalent fragment of inhibitory anti-MT1-MMP mAb decreased EC migration; in addition, the cytoplasmic deletion mutant of MT1-MMP had impaired migratory ability on COL I, showing that intact MT1-MMP is required for EC migration in accordance with previous reports in other cell types (Lehti et al., 2000;Uekita et al., 2001). Moreover, MT1-MMP cooperates with ␤1 and ␣v␤3 integrins in this respect because inhibition of the three pathways in combination abrogates EC migration on ECM. This effect may be due to overlapping or independent mechanisms acting in concert. Thus, integrin-mediated effects on migration are likely dependent on the regulation of cell attachment/detachment as well as on signaling through small GTPases (Schwartz and Shattil, 2000;Stupack and Cheresh, 2002). However, the mechanisms by which proteases play a role in cellular migration remain largely undefined (Murphy and Gavrilovic, 1999). Interestingly, no effect of the inhibitory anti-MT1-MMP mAb on EC adhesion to ECM was observed, pointing to other mechanisms involved (unpublished data). MT1-MMP might regulate migration by unmasking cryptic migratory sites, as shown for epithelial cells on laminin 5 (Koshikawa et al., 2000), or as it might happen to ␣v␤3 integrin that can bind cleaved collagen (Messent et al., 1998); in this regard, proteolysis of COL I was observed during EC migration (unpublished data). MT1-MMP-mediated processing of cellcell contact receptors might also favor detachment and migration of ECs. In this regard, the ability of MT1-MMP to process the transmembrane receptors CD44 and tissue transglutaminase involved in tumor cell migration has been shown (Belkin et al., 2001;Kajita et al., 2001). Direct modifications of integrin receptors by MT1-MMP might also be relevant because MT1-MMP can process pro-␣v, -␣5, and -␣3 integrins, and this processing seems to improve ␣v␤3mediated signaling and migration Ratnikov et al., 2002). Therefore, there is a dual contribution of MT1-MMP and integrins in regulating EC migration on ECM, although more efforts are required to elucidate if there is a functional crosstalk between them.
In summary, the regulation of MT1-MMP depends on integrin interactions with the ECM found by ECs during their migration. This might correlate with the sequential events of detachment of basal membrane, remodeling of subendothelial tissue and provisional matrix with appearance of new integrin ligands, and finally matrix deposition with arrest of migration that takes place in vivo during angiogenesis, vascular injury, wound healing, or tissue remodeling.
Type IV GEL, FN, and okadaic acid were purchased from Sigma-Aldrich. COL I from ICN Biomedicals, and human plasma, plasminogen-depleted, FG from Calbiochem-Novabiochem were also used. VN was a gift from Dr. E. Dejana (FIRC Institute of Molecular Oncology, Milan, Italy;Dejana et al., 1988).

Cells and cell cultures
Human ECs from umbilical vein were obtained and cultured as described previously (Arroyo et al., 1992). Cells up to the third passage were used in all the assays. ECs were changed to serum-free medium HE-SFM (Life Technologies) and seeded on dishes coated with 1% GEL or VN, COL I, FN, or FG at 10 g/ml before the functional assays were performed.

Immunofluorescence microscopy
ECs were grown on coverslips coated with different ECM until confluence. Next, they were stimulated by disrupting the monolayer for 24 h, fixed, and blocked as described previously (Gálvez et al., 2001). Coverslips were incubated with the primary antibody (anti-␣v␤3 LM609, anti-␤1 TS2/16, or anti-CD31 TP1/15 mAb) and labeled with a secondary Rhodamine X-conjugated goat anti-mouse antibody. After saturating coverslips with mouse serum, cells were labeled with the anti-MT1-MMP LEM-2/15 biotinylated mAb and then incubated with Streptavidin-Alexa 488. Samples were examined in a photomicroscope (DMR; Leica) with a 63ϫ oil immersion objective, and images were recorded using a CCD camera from Leica.
ECs grown to confluence on COL I-coated coverslips were incubated for 30 min at RT with 10 g/ml of the inhibitory anti-␤1 (LIA1/2) or anti-␣v integrin (ABA 6D1) mAb, and after 24 h at 37ЊC they were fixed and processed for immunofluorescence assays as described above. Coverslips were, in this case, incubated with the secondary Rhodamine X-conjugated goat anti-mouse antibody, and after blocking with mouse serum, incubated with the biotinylated anti-MT1-MMP LEM-2/15 mAb.

Adhesion assays
96-well plates (Costar) were coated with different ECM for 1 h at 37ЊC, washed with PBS twice, saturated with PBSϩ1% BSA for 1 h at 37ЊC, and again washed with PBS twice. ECs were resuspended in 1 ml HNSS (Bio-Whittaker) with BCFECS at 1 M (Molecular Probes, Inc.) and incubated at 37ЊC for 15 min. Next, they were washed twice with HNSS and resuspended in HE-SFM. Cells were preincubated with blocking antibodies for 20 min at RT, seeded at 3 ϫ 10 4 cells per well, and then incubated at 37ЊC for 30 min on the plates. Finally, plates were washed with HNSS twice and 100 l per well of lysis buffer (PBS ϩ 0.1% SDS) was added. Absorbance was measured at 480 nm with a microplate fluorescence reader (model FL500; Bio-Tek Instruments, Inc.). Experiments were performed in triplicate.

Generation of MT1-MMP-GFP fusion protein and time-lapse confocal videomicroscopy analysis
MT1-MMP-GFP and MT1-MMP⌬cyt-GFP were obtained using human MT1-MMP cDNA cloned into pCDNA3.1 as template to amplify by PCR the complete encoding region or the cytosolic domain truncated region (⌬Arg 563 ϪVal 582 ), respectively, of this molecule without the stop codon. A HindIII site was added to the 5Ј end and a SacII site at the 3Ј end. The PCR products were cloned into pcDNA3.1/V5-HIS-TOPO from Invitrogen. PCR products were then subcloned into pEGFP-N1. Sequence and orientation was confirmed by sequence analysis. ECs were transiently transfected by either calcium phosphate procedure or electroporation. In brief, 1.5 ϫ 10 6 cells were transfected with 20 g DNA, and after 15 min, 2% FBS was added; cells were plated in the presence of the DNA during 4 h. Alternatively, 20 g DNA was transfected into 2 ϫ 10 6 cells by electroporation at 200 V and 975 F using a Gene Pulser (Bio-Rad Laboratories). ECs transfected with MT1-MMP-GFP constructs were used 24 h after transfection for different assays (immunofluorescence microscopy and Transwell assays as described). For videomicroscopy analysis, transfected ECs were grown to confluence on COL I-coated glass bottom dishes (WillCo Wells B.V.) and then placed on the microscope stage. Plates were maintained at 37ЊC in a 5% CO 2 atmosphere using an incubation system (PeCon). Confocal series of fluorescence and differential interference contrast (DIC) images were simultaneously obtained at 3-min intervals during 2 h with a 40ϫ oil immersion objective. Images were processed and assembled into movies using Leica confocal software.

Clustering integrin receptors with latex beads
2.97-m-diam latex beads (Sigma-Aldrich) were incubated at RT with 1% GEL or 10 g/ml COL I, FN, or FG in 1 ml buffer Tris-HCl 0.1 M, pH 9.0, overnight under agitation and then blocked with PBS plus 1% BSA for 2 h at RT. Blocked beads were pelleted and resuspended in 1 ml HNSS plus 0.1% azide. Matrix-adsorbed latex beads (2 ϫ 10 4 ) were added to EC plated on COL I-coated coverslips, incubated for 24 h, washed, and fixed. Then, double-immunofluorescence staining was performed. 100 latex beads per field with a total of 4 fields per coverslip were counted.

Coimmunoprecipitation
Coimmunoprecipitation assays were essentially performed as described previously (Yáñez-Mó et al., 1998). In brief, ECs were grown to confluence for 24 h on COL I-coated plates and lysed under mild conditions (TBS, 1 mM CaCl 2 , 1 mM MgCl 2 , 1% Brij-96, 1% hemoglobin, and 1 mM PMSF). Lysates were incubated for 2 h at 4ЊC with 40 l of the primary mAb or glycine-Sepharose. Immunoprecipitates were washed twice with 1:10 dilution of the lysis buffer, boiled 5 min at 96ЊC in Laemmli buffer, and separated by 12% SDS-PAGE under nonreducing conditions. After transferring to a nitrocellulose membrane, Western blot was performed as described above.

Flow cytometry analysis
ECs grown on plates coated with different ECM or 10 g/ml of different purified mAb were detached with PBS plus 5 mM EDTA on ice and washed twice with PBS. In some cases, endothelial monolayer was disrupted to induce migration as described previously (Gálvez et al., 2001). About 2 ϫ 10 5 cells were incubated with 100 l hybridoma culture supernatant or biotinylated primary mAb for 20 min at 4ЊC. Cells were then washed with PBS and incubated with 100 l of the proper dilution of a FITC-conjugated anti-mouse Ig or Streptavidin-Alexa 488. Finally, fluorescence of samples was acquired in logarithmic scale using a FACSCalibur™ flow cytometer (Becton Dickinson).

Analysis of receptor internalization
ECs seeded on 6-well plates coated with different ECM were incubated with the primary mAb at RT for 20 min, exhaustively washed with PBS, and transferred to 37ЊC for 6 h to allow labeled-receptor internalization. Half of the cells were then fixed and permeabilized for 10 min at 4ЊC with lysis buffer (Becton Dickinson) and all processed for flow cytometry. The fluorescence of permeabilized and nonpermeabilized samples was acquired in linear scale for optimizing quantitation and analyzed at time points 0 and 6 h. Anti-intermediate filament CHELO-3 mAb was used to test efficiency of permeabilization that was always close to 100%. A similar approach was used to visualize receptor internalization under the fluorescence microscope. Internalization percentages at 6 h were estimated with Eq. 1: (1)

Western blot assays
ECs were grown on different ECM and stimulated to migrate or not by disrupting the monolayer. Resting and stimulated cells were washed twice with PBS and directly lysed in Laemmli buffer on ice. Lysates were analyzed by Western blot as described previously (Gálvez et al., 2001).

Zymography assays
ECs were changed to serum-free medium 24 h before the assay. ECs were grown on different ECM or on mAb-coated plates, and were then stimulated to migrate or not by disrupting the monolayer. Culture supernatants and cell lysates were processed by zymography as described previously (Gálvez et al., 2001).

Wound-healing assays
ECs were grown to confluence on COL I, FN, or FG-coated 24-well plates. Cells were preincubated with different purified mAb at 10 g/ml or at 5 g/ml, when used in combination with other mAb, 30 min before the injury. Next, wound-healing assays were performed as described previously (Gálvez et al., 2001). Experiments were done in duplicate and four fields of each well were recorded.

Cell transmigration assay
EC transmigration assays were performed in 8-m pore Transwell chambers (Costar). Cells were resuspended in serum-free medium and seeded at 2 ϫ 10 4 cells/well on GEL, VN, COL I, FN, or FG-coated filters in the absence or presence of 5-10 g/ml different purified mAb on the upper chamber. Transmigrated cells were stained and counted after 5 h of migration. Experiments were done in duplicate and four fields of each Transwell were counted with a 40ϫ objective in a microscope (Eclipse E400; Nikon). In transmigration assays with transfected ECs, transmigrated cells in eight independent fields were counted under the fluorescence microscope.

Statistical analysis
Tested and control samples in the functional assays were compared for statistical significance by t test.