Targeting of Tumor Cells by Cell Surface Urokinase Plasminogen Activator-dependent Anthrax Toxin*

Shihui LiuDagger , Thomas H. Bugge§, and Stephen H. LepplaDagger

From the Dagger  Oral Infection and Immunity Branch and § Oral and Pharyngeal Cancer Branch, NIDCR, National Institutes of Health, Bethesda, Maryland 20892

Received for publication, December 11, 2000, and in revised form, March 9, 2001


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Urokinase plasminogen activator receptor (uPAR) binds pro-urokinase plasminogen activator (pro-uPA) and thereby localizes it near plasminogen, causing the generation of active uPA and plasmin on the cell surface. uPAR and uPA are overexpressed in a variety of human tumors and tumor cell lines, and expression of uPAR and uPA is highly correlated to tumor invasion and metastasis. To exploit these characteristics in the design of tumor cell-selective cytotoxins, we constructed mutated anthrax toxin-protective antigen (PrAg) proteins in which the furin cleavage site is replaced by sequences cleaved specifically by uPA. These uPA-targeted PrAg proteins were activated selectively on the surface of uPAR-expressing tumor cells in the presence of pro-uPA and plasminogen. The activated PrAg proteins caused internalization of a recombinant cytotoxin, FP59, consisting of anthrax toxin lethal factor residues 1-254 fused to the ADP-ribosylation domain of Pseudomonas exotoxin A, thereby killing the uPAR-expressing tumor cells. The activation and cytotoxicity of these uPA-targeted PrAg proteins were strictly dependent on the integrity of the tumor cell surface-associated plasminogen activation system. We also constructed a mutated PrAg protein that selectively killed tissue plasminogen activator-expressing cells. These mutated PrAg proteins may be useful as new therapeutic agents for cancer treatment.


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Dissolution of the extracellular matrix is a prerequisite for invasive growth and metastatic spread of tumors as well as for physiological tissue remodeling and tissue repair. Matrix dissolution is accomplished by the concerted effort of a number of extracellular proteolytic systems, including serine, metallo-, and cysteine proteases (1-3). A particularly well studied proteolytic system implicated in tumor progression is the plasminogen activation system, a complex system of serine proteases, protease inhibitors, and protease receptors, that governs the conversion of the abundant plasma protease zymogen, plasminogen, to the active protease, plasmin (1, 2).

Plasmin is formed by the proteolytic cleavage of plasminogen by either of two plasminogen activators, the urokinase plasminogen activator (uPA)1 and the tissue plasminogen activator (tPA). uPA is a 52-kDa serine protease that is secreted as an inactive single chain proenzyme (pro-uPA) that is efficiently converted to active two-chain uPA by plasmin (4). Two-chain uPA, in turn, is a potent activator of plasminogen, leading to a powerful feedback loop that results in productive plasmin formation. However, both pro-uPA and plasminogen are catalytically inactive pro-enzymes, and the mechanism of initiation of uPA-mediated plasminogen activation is not fully understood. Pro-uPA binds with high affinity (Kd = 0.5 nM) to a specific glycosylphosphatidylinositol-linked cell surface receptor, the uPA receptor (uPAR), via an epidermal growth factor-like amino-terminal fragment (ATF; amino acids 1-135, 15 kDa) (5). uPAR is a 60-kDa, three-domain glycoprotein whose first and third domains constitute a composite high affinity binding site for the ATF of pro-uPA (5-8). The concomitant binding of pro-uPA to uPAR, and of plasminogen to as yet uncharacterized cell surface receptors, strongly potentiates uPA-mediated plasminogen activation (9-12), possibly due to the formation of ternary complexes, aligning the two proenzymes in a way that exploits their low intrinsic activity and thereby favors a mutual activation process (13). The net result of this process is the efficient and localized generation of active uPA and plasmin on the cell surface.

Although many studies have documented the central role of uPA-mediated cell-surface plasminogen activation requiring uPAR, recent studies in uPAR-deficient mice have demonstrated the existence of additional, uPAR-independent pathways of uPA-mediated plasminogen activation, in the context of both physiological cell migration and fibrin dissolution (14, 15).

uPAR and uPA are overexpressed with remarkable consistency in malignant human tumors, including monocytic and myelogenous leukemias (16, 17) and cancers of the colon (18), breast (19), bladder (20), thyroid (21), liver (22), pleura (23), lung (24), pancreas (25), ovaries (26), and the head and neck (27). Extensive in situ hybridization and immunohistochemical studies of various human tumor types have demonstrated that cancer cells typically express uPAR, whereas pro-uPA may be expressed by either the cancer cells or by adjacent stromal cells (18, 28, 29).

Plasminogen activation by uPA is regulated by two physiological inhibitors, plasminogen activator inhibitors-1 and -2 (PAI-1 and PAI-2) (30-32), each forming a 1:1 complex with uPA. Plasmin generated by the cell surface plasminogen activation system is relatively protected from its primary physiological inhibitor alpha 2-antiplasmin (11, 33, 34). Unlike uPA, plasmin is a relatively nonspecific protease, capable of degrading fibrin and several other glycoproteins and proteoglycans of the extracellular matrix (35). Therefore, cell surface plasminogen activation facilitates invasion and metastasis of tumor cells by dissolution of restraining tissue barriers. In addition, cell surface plasminogen activation may facilitate matrix degradation through the activation of latent matrix metalloproteinases (MMP) (36). Plasmin can also activate growth factors, such as transforming growth factor-beta , which may further modulate stromal interactions in the expression of enzymes and tumor neo-angiogenesis (37).

Another protein that requires cell surface proteolytic activation is anthrax toxin. This three-component toxin consists of protective antigen (PrAg, 83 kDa), lethal factor (LF, 90 kDa), and edema factor (EF, 90 kDa) (38-40). PrAg binds to an unidentified cell surface receptor and is cleaved at the sequence, 164RKKR167, by a cell-surface, furin-like protease (41, 42). This cleavage is absolutely required for the subsequent steps in toxin action. The carboxyl-terminal 63-kDa fragment (PrAg63) remains bound to receptor, associates to form a heptamer, and binds and internalizes LF and EF (40, 43-45). LF kills animals (46, 47) and lyses mouse macrophages (48, 49), probably due to the proteolytic cleavage of mitogen-activated protein kinase kinases (50, 51). EF damages cells due to its intracellular adenylate cyclase activity (52). A potent PrAgdependent cytotoxin, FP59, created by fusing LF amino acids 1-254 to the ADP-ribosylation domain of Pseudomonas exotoxin A can kill any cell having receptors for PrAg and the ability to activate PrAg by cleavage at amino acids 164-167 (53, 54).

The unique requirement that PrAg be activated on the target cell surface provides an opportunity to re-engineer this protein to make its activation dependent on the tumor cell surface urokinase plasminogen activation system. Our previous work showed that PrAg can be made specific for MMP-expressing cells by replacing the 164RKKR167 furin site with sequences preferentially cleaved by MMPs (55). In this report we extended this approach to exploit the localized activity of the uPA protease on tumor cells. uPA and tPA possess an extremely high degree of structural similarity (56, 57), share the same primary physiological substrate (plasminogen) and inhibitors (PAI-1 and PAI-2) (58), and exhibit restricted substrate specificity. Recent elegant genetic studies using substrate phage display and substrate subtraction phage display identified peptide substrates that are cleaved with high efficiency as well as high selectivity by either uPA or tPA (59, 60). We used the amino acid sequences defined in that work to replace the furin cleavage site in PrAg to produce several mutated PrAg proteins susceptible to cleavage by uPA and tPA. These uPA- and tPA-targeted PrAg proteins were activated selectively on the surface of tumor cells and caused their killing by the recombinant cytotoxin FP59, as described below.

    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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REFERENCES

Reagents-- FP59 and a soluble form of furin were prepared as described previously (61). Rabbit anti-PrAg polyclonal antiserum (serum no. 5308) was made in our laboratory. Reagents obtained from American Diagnostica Inc. (Greenwich, CT) included pro-uPA (single-chain uPA, no. 107), uPA (no. 124), tPA (no. 116), human urokinase amino-terminal fragment (ATF, no. 146), human Glu-plasminogen (no. 410), human PAI-1 (no. 1094), alpha 2-antiplasmin (no. 4030), monoclonal antibody against human uPA B-chain (no. 394), and goat polyclonal antibody against human t-PA (no. 387). tPA not containing protein stabilizer was purchased from Calbiochem (San Diego, CA). Aprotinin and tranexamic acid were purchased from Sigma Chemical Co. (St. Louis, MO). The uPAR monoclonal antibody R3 was a gift from Dr. Gunilla Høyer Hansen (Finsen Laboratory, Copenhagen, Denmark).

Construction of Mutated PrAg Proteins-- A modified overlap PCR method was used to construct the mutated PrAg proteins in which the furin site is replaced by: 1) the plasminogen-derived sequence PCPGRVVGG in PrAg-U1; 2) the preferred uPA substrate sequences PGSGRSA and PGSGKSA in PrAg-U2 and PrAg-U3, respectively; and 3) the preferred tPA sequence PQRGRSA in PrAg-U4 (Table I). Plasmid pYS5 (62) was used as both PCR template and expression vector. The native Pfu DNA polymerase (Stratagene, La Jolla, CA) was used in the PCR reactions. We used 5'-primer F, AAAGGAGAACGTATATGA (Shine-Dalgarno and start codons are underlined), and the phosphorylated reverse primer R1, pTGGTGAGTTCGAAGATTTTTGTTTTAATTCTGG (the first three nucleotides encodes P, the others anneal to the sequence corresponding to P154 to S163), to amplify a fragment designated "N." We used the mutagenic phosphorylated primer H1, pTGTCCAGGAAGAGTAGTTGGAGGAAGTACAAGTGCTGGACCTACGGTTCCAG, encoding CPGRVVGG and S168 to P176, and reverse primer R2, ACGTTTATCTCTTATTAAAAT, annealing to the sequence encoding I589 to R595, to amplify a mutagenic fragment "M1." We used a phosphorylated mutagenic primer H2, pGGAAGTGGAAGATCAGCAAGTACAAGTGCTGGACCTACGGTTCCAG, encoding GSGRSA and S168 to P176, and reverse primer R2, to amplify a mutagenic fragment "M2." We used a phosphorylated mutagenic primer H3, pGGAAGTGGAAAATCAGCAAGTACAAGTGCTGGACCTACGGTTCCAG, encoding GSGKSA and S168 to P176, and reverse primer R2, to amplify a mutagenic fragment "M3." We used a phosphorylated mutagenic primer H4, pCAGAGAGGAAGATCAGCAAGTACAAGTGCTGGACCTACGGTTCCAG, encoding QRGRSA and S168 to P176, and reverse primer R2, to amplify a mutagenic fragment "M4." Primers F and R2 were used to amplify the ligated products of N + M1, N + M2, N + M3, and N + M4, respectively, resulting in the mutagenized fragments U1, U2, U3, and U4 in which the coding sequence for the furin site (164RKKR167) is replaced by uPA or tPA substrate sequence. The 670-bp HindIII/PstI fragments from the digests of U1, U2, U3, and U4 were cloned between the HindIII and PstI sites of pYS5. The resulting mutated PrAg proteins were accordingly named PrAg-U1, PrAg-U2, PrAg-U3, and PrAg-U4. We also constructed a mutated PrAg protein, PrAg-U7, in which 164RKKR167 is replaced by the sequence PGG. This protein is expected to be resistant to all cell surface proteases. DNA sequencing analyses confirmed the sequences of the mutated PrAg constructs.

Expression and Purification of PrAg Proteins-- Plasmids encoding the constructs described above were transformed into the non-virulent strain Bacillus anthracis UM23C1-1, and transformants were grown in FA medium (62) with 20 µg/ml kanamycin for 16 h at 37 °C. The mutated PrAg proteins were concentrated from the culture supernatants and purified by chromatography on a MonoQ column (Amersham Pharmacia Biotech, Piscataway, NJ) by the methods described previously (63).

In Vitro Cleavage of PrAg Proteins by uPA, tPA, and Furin-- Reaction mixtures of 50 µl containing 5 µg of the PrAg proteins were incubated at 37 °C with 5 µl of soluble furin or 0.5 µg of uPA or tPA. Furin cleavage was done as described previously (55). Cleavage with uPA or tPA was done in 150 mM NaCl, 10 mM Tris-HCl (pH 7.5). Aliquots withdrawn at intervals were analyzed by SDS-polyacrylamide gel electrophoresis (PAGE) using 4-20% gradient Tris-glycine gels (Novex, San Diego, CA), and proteins were either visualized by Coomassie Blue staining or were electroblotted to a nitrocellulose membrane (Novex). Membranes were blocked with 5% (w/v) non-fat milk, incubated sequentially with rabbit anti-PrAg polyclonal antibody (no. 5308) and horseradish peroxidase-conjugated goat anti-rabbit antibody (sc-2004, Santa Cruz Biotechnology, Inc., Santa Cruz, CA), and visualized by detection of horseradish peroxidase by SuperSignal West Pico chemiluminescent substrate (Pierce, Rockford, IL). To verify the cleavage sites, digestions of native PrAg by furin, PrAg-U2 and-U3 by uPA, and PrAg-U4 by tPA (Calbiochem) were performed for 3 h at 37 °C as described above. Then the resulting PrAg63s were separated by SDS-NuPAGE electrophoresis (Novex), and the proteins were transferred onto Immobilon-P polyvinylidene difluoride membranes (Millipore, Bedford, MA) and visualized by Coomassie Blue staining. The protein bands were cut out and sequenced by the Protein and Nucleic Acid Laboratory, Center for Biologics Evaluation and Research, FDA using an ABI model 494A protein sequencer.

Cells and Culture Medium-- Human 293 kidney cells, human cervix adenocarcinoma HeLa cells, human melanoma A2058 cells, and human melanoma Bowes cells were obtained from American Type Culture Collection (Manassas, VA). Mouse Lewis lung carcinoma cell line LL3 was kindly provided by Dr. Michael S. O'Reilly (Boston, MA). These cells were grown in Dulbecco's modified Eagle's medium (DMEM) with 0.45% glucose, 10% fetal bovine serum (FCS), 2 mM glutamine, and 50 µg/ml gentamicin. Human umbilical vein endothelial cells (HUVEC) were obtained from Clonetics Corp. (Walkersville, MD) and were grown in RPMI 1640 containing 20% defined and supplemented bovine calf serum (HyClone Laboratories, Inc, Logan, UT), 5 units/ml heparin (Fisher Scientific, Pittsburgh, PA), 100 units/ml penicillin, and 0.2 mg/ml endothelial cell growth supplement (Collaborative Research), 100 µg/ml streptomycin, 50 µg/ml gentamicin, and 2.5 µg/ml amphotericin B (Life Technologies, Rockville, MD). Cells were maintained at 37 °C in a 5% CO2 environment.

Cytotoxicity Assays with MTT-- Cells were cultured in 96-well plates to ~50% confluence and washed twice with serum-free DMEM to remove residual serum. Then the cells were preincubated for 30 min with serum-free DMEM containing 100 ng/ml pro-uPA and 1 µg/ml Glu-plasminogen with or without PAI-1, aprotinin, alpha 2-antiplasmin, ATF, or the uPAR blocking antibody R3. PrAg proteins (0-1000 ng/ml) combined with FP59 (50 ng/ml) were added to the cells to give a total volume of 200 µl/well. Cells were incubated with the toxins for 6 h, after which the medium was replaced with fresh DMEM supplemented with 10% fetal calf serum. Cell viability was assayed by adding 50 µl of 2.5 mg/ml MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) at 48 h. The cells were incubated with MTT for 45 min at 37 °C, the medium was removed, and the blue pigment produced by viable cells was dissolved in 100 µl/well of 0.5% (w/v) SDS, 25 mM HCl, in 90% (v/v) isopropanol. The plates were vortexed and the oxidized MTT was measured as A570 using a microplate reader.

Binding and Processing of Pro-PA and PrAg-U2 by Cultured Cells-- Cells were cultured in 24-well plates to confluence, washed, and incubated in serum-free DMEM with 1 µg/ml pro-uPA, 1 µg/ml PrAg-U2, and 1 µg/ml Glu-plasminogen, and 2 mg/ml bovine serum albumin (BSA) at 37 °C for various lengths of times. The cells were washed five times to remove unbound pro-uPA and PrAg-U2. When PAI-1 was tested, it was incubated with cells for 30 min prior to the addition of pro-uPA and PrAg-U2. When tranexamic acid was tested, cells were preincubated with serum-free DMEM containing 2 mg/ml BSA, 1 mM tranexamic acid, without plasminogen, for 30 min before the addition of pro-uPA and PrAg-U2. Cells were lysed in 100 µl/well of modified radioimmune precipitation lysis buffer (50 mM Tris-HCl, pH 7.4, 1% Nonidet P-40, 0.25% sodium deoxycholate, 150 mM NaCl, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 1 µg/ml each of aprotinin, leupeptin, and pepstatin) on ice for 10 min. Equal amounts of protein from cell lysates and equal volumes of the conditioned media were separated by PAGE using 4-20% gradient Tris-glycine gels (Novex). Western blotting was performed as described above to detect pro-uPA and PrAg-U2 and their cleavage products by using the monoclonal antibody against human uPA B-chain (no. 394) and anti-PrAg polyclonal antibody (no. 5308).

Cytotoxicity Assay in a Co-culture System-- A co-culture model like that described previously (55) was employed to determine whether PrAg-U2 killed uPAR-overexpressing tumor cells without affecting bystander, uPAR non-expressing cells. Briefly, HeLa and 293 cells were co-cultured in separate compartments of eight-chamber slides. With the partitions removed, the culture slides were incubated for 6 h with native PrAg or PrAg-U2 (each 300 ng/ml) combined with FP59 (50 ng/ml) in serum-free DMEM containing 100 ng/ml pro-uPA and 1 µg/ml Glu-plasminogen. After 48 h, the partitions were replaced and MTT-containing medium was added to each chamber to assess cell viability, as described above.

    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Directing uPA- and tPA-specific Proteolysis to Anthrax PrAg-- The furin cleavage site, 164RKKR167, is located in a surface-exposed, flexible loop of PrAg composed of residues 162-175 (64). We constructed mutated PrAg proteins in which this sequence is replaced by sequences that are preferred uPA or tPA substrates (Table I). The mutated PrAg protein PrAg-U1 contains the sequence PCPGRVVGG, corresponding to positions P5 to P4' in the physiological substrate plasminogen. Protein PrAg-U2 contains the sequence PGSGRSA, which includes the consensus sequence SGRSA, recently identified as the minimized optimum substrate for uPA (59). Because the sequence SGRSA is cleaved by uPA 1363-fold times more efficiently than the physiological cleavage site present in plasminogen, and because it exhibits a uPA/tPA selectivity of 20 (59), the PrAg-U2 protein is expected to be a specific substrate of uPA. uPA/tPA selectivity of the sequence SGRSA can be further enhanced by placing lysine in the P1 position (59). Thus, the sequence PGSGKSA, which exhibits a uPA/tPA selectivity of 121 (59), was selected for insertion into the mutated PrAg protein PrAg-U3, which was expected to have an even higher uPA selectivity than PrAg-U2. Ke et al. (59) further showed that the P3 and P4 residues were the primary determinants of the ability of a substrate to discriminate between tPA and uPA. Thus, substitution of both the P4 glycine and the P3 serine of the most labile uPA substrate (GSGRSA) with glutamine and arginine, respectively, decreased the uPA/tPA selectivity by a factor of 1200 and yielded a tPA-selective substrate (59). Based on that result, we constructed the mutated PrAg protein PrAg-U4 containing the sequence PQRGRSA, so as to produce a tPA-specific substrate. We also constructed a mutated PrAg protein PrAg-U7, in which 164RKKR167 was replaced by the sequence PGG. PrAg-U7 is not expected to be cleaved by any known protease and was used as a control protein in this study. The designations of the mutated PrAg proteins along with the expected properties based on the study of Ke et al. (59) are summarized in Table I.

                              
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Table I
Predicted and observed properties of mutated PA proteins

Plasmids encoding these mutated PrAg proteins were constructed by a modified overlap PCR method, cloned into the Escherichia coli-Bacillus shuttle vector pYS5, and expressed in B. anthracis UM23C1-1. The proteins were secreted into the culture supernatants at 20-50 mg/liter. The mutated PrAg proteins were concentrated and purified by MonoQ chromatography to one prominent band at the expected molecular mass of 83 kDa, which co-migrated with native PrAg in SDS-PAGE. Thus, using a production protocol that is now standard in this laboratory, these mutated PrAg proteins could be expressed and purified easily in high yield and purity.

To verify that the mutated PrAg proteins had the expected susceptibility to cleavage by proteases, they were incubated separately with uPA, tPA, and a soluble form of furin. As expected, these mutated PrAg proteins were not cleaved by furin, whereas the native PrAg was cleaved by furin to produce the active PrAg63 product (Fig. 1A). The cleavage by furin after the 164RKKR167 sequence was confirmed by amino-terminal sequencing of the resulting PrAg63. The relative susceptibilities of the mutated PrAg proteins to cleavage by uPA and tPA agreed closely with what was predicted from the phage display data used in their design (Fig. 1, B and C, Table I). In particular, uPA cleaved PrAg-U2 very efficiently but was less active on PrAg-U3. Moreover, PrAg-U2 was quite resistant to tPA, with just trace amounts being cleaved even with a 3-h incubation period (Fig. 1C). PrAg-U3 was even more resistant to tPA, in that no cleavage could be detected at any time point (Fig. 1C). These results showed the high uPA specificity for these two mutated PrAg proteins. In contrast, PrAg-U4 was a very weak substrate for uPA, but a good substrate for tPA (Fig. 1, B and C). The cleavage of PrAg-U2 and PrAg-U3 at the predicted peptide bonds by uPA and that of PrAg-U4 by tPA was confirmed by amino-terminal sequencing of the resulting PrAg63s. PrAg-U7 and PrAg-U1 were both completely resistant to uPA and tPA (Fig. 1, B and C). Native PrAg was completely resistant to tPA (Fig. 1C) but was slightly cleaved by uPA at the furin recognition site (Fig. 1B). When we replaced the furin site with the sequence PGG to produce PrAg-U7, the protein was completely resistant to uPA (Fig. 1B).


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Fig. 1.   Protease sensitivity of the mutated PrAg proteins. PrAg proteins were incubated with furin (A), uPA (B), or tPA (C), for the times indicated and then analyzed by SDS-PAGE and Coomassie Blue staining (0.5 µg of PrAg per lane in A), or by Western blotting with rabbit polyclonal antibody against PrAg (0.1 ng of PrAg per lane, B and C).

PrAg-U2 and PrAg-U3 Selectively Kill uPAR-expressing Tumor Cells-- To test the hypothesis that PrAg-U2 and PrAg-U3 would selectively kill uPAR-overexpressing tumor cells, cytotoxicity assays were performed with two human tumor cell lines, cervix adenocarcinoma HeLa and melanoma A2058. The non-tumor human kidney cell line 293 was used as a control. Expression of uPAR by these two tumor cell lines but not by 293 cells was reported previously (65) and was confirmed in this study by performing a pro-uPA binding and processing assay (Fig. 2A). In the presence of plasminogen, both HeLa and A2058 cells bound pro-uPA and processed it to the active, two-chain form, as identified by the uPA B-chain antibody. In contrast, the uPAR non-expressing 293 cells showed only a weak binding and failed to convert pro-uPA to two-chain uPA (Fig. 2A).


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Fig. 2.   Interaction of mutated PrAg proteins with uPAR-expressing tumor cells. A, binding and processing of pro-uPA by cultured cell lines. HeLa cells, A2058 cells, and human kidney 293 cells were cultured to confluence and incubated in serum-free media containing 2 mg/ml BSA and 1 µg/ml each of pro-uPA and Glu-plasminogen for the times indicated. The cells were then washed, and lysates were prepared for Western blotting analysis with a monoclonal antibody against uPA B-chain (no. 394). B, C, D, cytotoxicity of the mutated PrAg proteins to 293 (B), HeLa (C), and A2058 (D) cells. Cells were cultured to 50% confluence and incubated with PrAg proteins together with 50 ng/ml FP59 for 6 h in serum-free DMEM containing 100 ng/ml of pro-uPA and 1 µg/ml of Glu-plasminogen. Then the toxins were removed and replaced with fresh serum-containing DMEM. MTT was added to determine cell viability at 48 h. The analysis was performed two additional times with similar results. (Mean ± S.D., n = 4.)

Cytotoxicity of native PrAg and the mutated PrAg proteins to these cells was measured in 96-well plates. In tumor tissues, cancer cells typically overexpress uPAR, whereas either the cancer cells or the adjacent stromal cells express pro-uPA, which is activated on the cancer cell surface after binding to uPAR. We showed that HeLa and A2058 cells did not express pro-uPA under the current culture condition ("0lanes in Fig. 2A). Therefore, in the cytotoxicity assay, 100 ng/ml pro-uPA was added to the tumor cells to mimic the role of pro-uPA secreted in tumor tissues in vivo. We also added 1 µg/ml Glu-plasminogen, because plasminogen is present in high concentration (1.5-2.0 µM) in plasma and interstitial fluids and is required for uPAR-dependent conversion of pro-uPA to active uPA. The cells were then incubated with the native or the mutated PrAg proteins combined with FP59 for 6 h, and cell viability was measured after 48 h. The results showed that the uPAR non-expressing 293 cells were sensitive to native PrAg in a dose-dependent manner but were completely resistant to killing by all the mutated PrAg proteins (Fig. 2B). In contrast, the uPAR-expressing HeLa and A2058 cells were highly susceptible to killing by native PrAg, PrAg-U2, and PrAg-U3, were less susceptible to PrAg-U4, and were completely resistant to PrAg-U1 and PrAg-U7 (Fig. 2, C and D). The EC50 values (concentrations needed to kill half of the cells) for native PrAg and the mutated PrAg proteins are summarized in Table I. The rank order of the cytotoxicities of these PrAg proteins correlated well with the uPA cleavage profiles shown in Fig. 1B, strongly suggesting that the cytotoxicity observed was dependent on the uPA activity generated by the uPAR-expressing tumor cells. The selective cytotoxicity of the mutated PrAg proteins for the tumor cells was retained when the experiments were repeated in medium containing 10% fetal calf serum (data not shown). This indicates that serum proteases do not activate the PrAg proteins, nor do serum protease inhibitors block proteolytic cleavage of mutated PrAg proteins by the cell surface proteases. To simplify further analysis, all subsequent experiments were performed in serum-free medium.

Killing of uPAR-expressing Tumor Cells by the Mutated PrAg Proteins Is Strictly Dependent on the Integrity of the Cell Surface-associated Plasminogen System-- To verify that the cytotoxicity of the mutated PrAg proteins was dependent on the cell surface-associated plasminogen activation system, we first tested the role of pro-uPA in the action of the mutated PrAg proteins. When the cytotoxicity experiments shown in Fig. 2, C and D, were repeated without addition of pro-PA, the mutated PrAg proteins PrAg-U2, PrAg-U3, and PrAg-U4 were not toxic to HeLa and A2058 cells, whereas native PrAg retained the same cytotoxicity (data not shown). Furthermore, the killing of HeLa cells by PrAg-U2 was directly dependent on the concentration of pro-uPA added (Fig. 3). No cytotoxicity was detected in the absence of pro-uPA, whereas substantial killing occurred at a pro-uPA concentration of only 12.5 ng/ml (Fig. 3). These data prove that the toxicity of these mutated PrAg proteins to the tumor cells is absolutely dependent on the presence and activation of pro-uPA. Within tissues, the pro-uPA bound to cell surface uPAR is usually produced by neighboring cells or adsorbed from plasma. Few types of cultured cells produce both cell surface uPAR and secreted pro-uPA. One example is the Lewis lung carcinoma cell line LL3, which produces both proteins (66-69). Therefore, it was expected that the LL3 line would be susceptible to PrAg-U2 even in the absence of added pro-uPA, and this was confirmed in the experiment shown in Fig. 4. Killing was especially pronounced when exposure to toxin was extended to 48 h.


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Fig. 3.   Cytotoxicity of PrAg-U2 to HeLa cells requires addition of pro-uPA. HeLa cells were cultured and treated with toxin for 6 h as in Fig. 2, except that various concentrations of pro-uPA were added. The analysis was performed one additional time with results similar to those presented here.


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Fig. 4.   PrAg-U2 kills Lewis lung carcinoma cell line LL3 without addition of pro-uPA. The cells were cultured to 50% confluence and washed, and the medium was replaced with serum-free DMEM containing 1 µg/ml Glu-plasminogen. Then PrAg-U2 together with 50 ng/ml FP59 was incubated with the cells for 48 h or removed after 6 h and replaced with serum-containing DMEM. MTT was added to determine cell viability at 48 h. The analysis was performed one additional time with similar results. (Mean ± S.D., n = 2.)

We next assessed the binding and proteolytic activation of pro-uPA and PrAg-U2 on uPAR-expressing and uPAR non-expressing cells. uPAR-expressing HeLa cells and non-expressing 293 cells were incubated with 1 µg/ml each of pro-uPA and PrAg-U2 in the absence or presence of plasminogen, PAI-1, and tranexamic acid for various durations of time. Thereafter, cell lysates and conditioned media were examined by Western blotting to detect the binding and processing of pro-uPA and PrAg-U2. uPAR-expressing HeLa cells proteolytically activated pro-uPA, with active uPA accumulating both on the cell surface and in the medium (Fig. 5A). In contrast, the uPAR non-expressing 293 cells bound weakly but could not cleave pro-uPA, and only trace amounts of active uPA accumulated in the medium (Fig. 5A). The activation of pro-uPA by HeLa cells was completely blocked by PAI-1 (Fig. 5A), providing further evidence that uPA is activated on the cell surface through a reciprocal activation loop involving pro-uPA and plasminogen. Activation of PrAg-U2 on the HeLa cell surface, determined by the production of the processed form PrAg 63 and the formation of SDS-stable PrAg 63 oligomer (Fig. 5B), exactly matched the activation profile of pro-uPA on the cell surface (Fig. 5A). In particular, when the activation of pro-uPA was blocked by PAI-1 (Fig. 5A), PrAg-U2 activation was blocked in parallel (Fig. 5B), demonstrating that the activation of PrAg-U2 on the HeLa cell surface required the activation of pro-uPA. As expected, the uPAR non-expressing 293 cells could process neither pro-uPA nor PrAg-U2 (Fig. 5, A and B). As a control experiment, we showed that HeLa and 293 cells could process native PrAg (by furin), and this could not be inhibited by PAI-1 (Fig. 5C). The effect of PAI-1 on cytotoxicity of native PrAg and PrAg-U2 was also assessed. As expected, PAI-1 conferred strong protection to HeLa cells from PrAg-U2 plus FP59, but not from native PrAg plus FP59 (Fig. 6).


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Fig. 5.   Binding and processing of pro-uPA and PrAg-U2 by HeLa and 293 cells. HeLa and 293 cells were cultured to confluence in 24-well plates and preincubated with serum-free DMEM containing 2 mg/ml BSA, 1 µg/ml Glu-plasminogen, with or without 10 µg/ml PAI-1 for 30 min. Some cells were preincubated with serum-free DMEM containing 2 mg/ml BSA, 1 mM tranexamic acid, without plasminogen. Then 1 µg/ml each of pro-uPA and PrAg-U2 were added to the cells and incubated for the times indicated. The cells were thoroughly washed, and the cell-conditioned media and the cell lysates were analyzed by Western blotting using a monoclonal antibody against the uPA B-chain (no. 394) (A), or by using a rabbit anti-PrAg polyclonal antibody (no. 5308) (B) to determine the binding and processing status of pro-uPA and PrAg-U2. As a control experiment (C), HeLa cells and 293 cells were preincubated with or without PAI-1 for 30 min and 1 µg/ml PrAg was added for the indicated times. The cells were washed, and the cell lysates were analyzed by Western blotting by using a rabbit anti-PrAg polyclonal antibody (no. 5308).


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Fig. 6.   The cytotoxicity of PrAg-U2 to HeLa cells is blocked by PAI-1. HeLa cells were cultured to 50% confluence, preincubated with serum-free DMEM containing 100 ng/ml pro-uPA and 1 µg/ml Glu-plasminogen with or without 2 µg/ml PAI-1 for 30 min. Then native PrAg and PrAg-U2 combined with FP59 (50 ng/ml) were added to the cells and incubated for 6 h. The toxins were removed and replaced with fresh serum-containing DMEM. MTT was added to determine cell viability at 48 h. The analysis was performed two additional times with similar results. (Mean ± S.D., n = 2.)

Although active uPA could also be detected in the conditioned medium of HeLa cells (Fig. 5A), just a trace amount of PrAg-U2 was activated in the medium (Fig. 5B), indicating that the coincident binding of PrAg-U2 and uPA on the cell surface facilitated the activation of PrAg-U2 by uPA. To further support this, we also assessed the effects of PrAg-U7, the uncleavable PrAg variant, on the binding and processing of PrAg-U2 by HeLa cells. We showed that PrAg-U2 binding and processing on the HeLa cell surface was completely blocked by the excess amount (200-fold) of PrAg-U7, and the cytotoxicity of PrAg-U2 to HeLa cells was blocked in parallel (Fig. 7). In agreement with this, the selective cytotoxicity of PrAg-U2 to uPAR-expressing HeLa cells was retained even in a co-culture with the uPAR non-expressing 293 cells, whereas native PrAg killed both cell types (Fig. 8). The fact that PrAg-U2 activated on HeLa cells did not spill over and cause killing of the bystander 293 cells suggests that the specificity toward tumor cells may be retained in vivo.


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Fig. 7.   The effects of PrAg-U7 on the binding and processing of PrAg-U2 by HeLa cells. HeLa cells were cultured to confluence in 24-well plates, then incubated with serum-free DMEM containing 2 mg/ml BSA, 1 µg/ml Glu-plasminogen, 1 µg/ml pro-uPA, with 1 µg/ml PrAg-U2, and 50 ng/ml FP59 or 1 µg/ml PrAg-U2, 200 µg/ml PrAg-U7, and 50 ng/ml FP59 at 37 °C. After 2-h incubation, the cells were washed, and the cell lysates were analyzed by Western blotting by using a rabbit anti-PrAg polyclonal antibody (no. 5308) (in A). For the cytotoxicity assay (in B), the toxins were removed and replaced with fresh serum-containing DMEM after 6 h. MTT was added to determine cell viability at 48 h.


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Fig. 8.   PrAg-U2 selectively kills HeLa cells in a co-culture model. HeLa and 293 cells were cultured to confluence in the separate compartments of eight-chamber slides. With the partitions removed, the slides were placed in 100-mm culture dishes with serum-free medium containing 100 ng/ml pro-uPA and 1 µg/ml glu-plasminogen, so that the different cells were in the same culture environment. Native PrAg and PrAg-U2 (300 ng/ml), each combined with FP59 (50 ng/ml), or FP59 alone were added to the cells, and incubated for 48 h. Partitions were replaced at 48 h, and MTT was added to determine the viability of the cell type present in each chamber.

The involvement of cell surface-bound plasminogen in the activation of pro-uPA and PrAg-U2 was investigated by the use of tranexamic acid, which inhibits the binding of plasminogen to the cell surface (11, 70). Pretreatment of cells with 1 mM tranexamic acid strongly inhibited the activation of pro-uPA and PrAg-U2 (Fig. 5, A and B) but not the activation of native PrAg (data not shown). The involvement of cell surface-bound plasminogen in the cascade activation of pro-uPA and PrAg-U2 was further demonstrated by comparing the effects of two plasmin inhibitors, alpha 2-antiplasmin and aprotinin. Aprotinin, which can inhibit the activity of plasmin both on the cell surface and in solution (10, 11), protected HeLa cells from killing by PrAg-U2 plus FP59. In contrast, alpha 2-antiplasmin, which is an inefficient inhibitor of cell surface-bound plasmin (10, 11), could not protect the cells (Fig. 9). Aprotinin and alpha 2-antiplasmin had no effect on the killing of cells by native PrAg plus FP59 (Fig. 9).


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Fig. 9.   Cell surface plasmin activity is required for the cytotoxicity of PrAg-U2 to HeLa cells. HeLa cells were cultured to 50% confluence and preincubated with serum-free DMEM containing 100 ng/ml pro-uPA and 1 µg/ml Glu-plasminogen with or without 40 µg/ml alpha 2-antiplasmin or 100 µg/ml aprotinin for 30 min. Then 300 ng/ml PrAg or PrAg-U2 combined with 50 ng/ml FP59 were added to the cells for 6 h. The toxins were removed and replaced with fresh serum-containing DMEM. MTT was added to determine cell viability at 48 h. The analysis was performed two additional times with similar results. (Mean ± S.D., n = 2.)

We next addressed the role of uPAR in the cytotoxicity of PrAg-U2 to the uPAR-expressing HeLa cells by preincubating cells with two reagents that specifically block the binding of uPA to its receptor. ATF, the amino-terminal fragment of uPA, competes with pro-uPA for binding to uPAR. It protected the tumor cells from killing by PrAg-U2 plus FP59 in a dose-dependent manner but had no effect on killing by native PrAg plus FP59 (Fig. 10A). Similarly, the monoclonal uPAR antibody R3 that specifically blocks the binding of pro-uPA to uPAR also protected the tumor cells from killing by the uPA-activated cytotoxin (Fig. 10B) but had no effect on the killing of cells by native PrAg. These results demonstrate that the activation of PrAg-U2 and the tumor cell killing was absolutely dependent on the binding of pro-uPA to uPAR. Taken together, we conclude that the cytotoxicity of the uPA-activated PrAg proteins to uPAR-expressing tumor cells was strictly dependent on the integrity of the cell surface-associated plasminogen activation system.


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Fig. 10.   Binding of pro-uPA to uPAR is required for the cytotoxicity of PrAg-U2 to HeLa cells. HeLa cells cultured to 50% confluence were preincubated for 30 min with serum-free DMEM containing 100 ng/ml pro-uPA and 1 µg/ml Glu-plasminogen, and different concentrations of ATF (A) or uPAR-blocking antibody R3 (B). Then 300 ng/ml each of PrAg and PrAg-U2 combined with 50 ng/ml FP59 were added to the cells for 6 h. Then toxins were removed and replaced with fresh serum-containing DMEM. MTT was added to determine cell viability at 48 h. (Mean ± S.D., n = 2.)

PrAg-U4 Is Toxic to tPA-expressing Cells Whereas PrAg-U2 and PrAg-U3 Are Only Weakly Toxic-- Because PrAg-U4 is the mutated PrAg that is most susceptible to cleavage by tPA (Fig. 1C), we expected it to be toxic to tPA-expressing cells. To test this hypothesis, cytotoxicity assays were performed on two tPA-expressing cells, human Bowes melanoma cells and primary human umbilical vein endothelial cells (HUVEC). The expression of tPA by these cells was demonstrated by Western blotting of culture supernatants using a polyclonal antibody against human tPA (Fig. 11A). The cytotoxicity assay was done in serum-free DMEM without addition of pro-uPA and Glu-plasminogen. Different concentrations of native PrAg, PrAg-U2, PrAg-U3, and PrAg-U4 combined with 50 ng/ml FP59 were incubated with cells for 12 h, and cell viability was measured at 48 h. PrAg-U4 was toxic to the two tPA-expressing cells, whereas PrAg-U2 and PrAg-U3 showed very low toxicity (Fig. 11, B and C). The EC50 values of the PrAg proteins to these tPA-expressing cells are summarized in Table I. These and the above results clearly show that these mutated PrAg proteins, PrAg-U2, PrAg-U3, and PrAg-U4, have differential cytotoxicity to the uPA/uPAR and tPA-expressing cells.


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Fig. 11.   PrAg-U4 efficiently kills tPA-expressing cells whereas PrAg-U2 and PrAg-U3 have low toxicity. A, Bowes cells and HUVEC cells express tPA. Serum-free conditioned media from Bowes cells and HUVEC cells were collected 24 h after addition to confluent cells and were analyzed by Western blotting using a polyclonal goat anti-tPA antibody. B and C: Bowes cells (B) and HUVEC cells (C) cultured to 50% confluence were treated with native PrAg, PrAg-U2, PrAg-U3, and PrAg-U4 together with 50 ng/ml FP59 for 12 h in serum-free DMEM without pro-uPA and Glu-plasminogen. Then toxins were removed and replaced with fresh serum-containing DMEM. MTT was added to determine cell viability at 48 h. The analysis was performed two additional times with similar results. (Mean ± S.D., n = 2.)


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Since the discovery in 1976 that uPA is produced and released from cancer cells (71), convincing evidence has accumulated that the urokinase plasminogen activation system is involved in the proliferation, invasion, and metastasis of human tumors (1, 2). Recent data suggests that tumor invasion factors can serve as targets for novel therapies to prevent cancer invasion and metastasis (2, 72). Strategies for interfering with the expression or the activity of uPA, uPAR, and PAI-1 have involved antisense oligonucleotides, antibodies, inhibitors, and recombinant or synthetic uPA and uPAR analogues (72). However, these approaches would only be expected to slow the progression of tumors without having a direct cytotoxic action that could eradicate the malignant cells. Several studies have targeted protein toxins to uPAR, in several cases by fusion of toxin catalytic domains to ATF (73-75). However, the present study is the first attempt of which we are aware to exploit the substrate specific protease activity of the plasminogen activators to target cytotoxic bacterial toxin fusion proteins to tumor cells. We constructed mutated anthrax toxin PrAg proteins in which the furin activation site is replaced by either the amino acid sequence present at the cleavage site in plasminogen (PrAg-U1) or by sequences deduced by a substrate phage display methods as being highly susceptible to cleavage by uPA (PrAg-U2 and PrAg-U3) or tPA (PrAg-U4) (59, 60). The uPA and tPA cleavage profiles of these mutated PrAg proteins matched very well with those of the isolated peptides used to replace the furin activation site in these proteins. Thus, PrAg-U2 and PrAg-U3 are efficiently and preferentially activated by uPA, whereas PrAg-U4 is preferentially activated by tPA. The above results demonstrate that new activator specificities can be conferred on PrAg by replacement of the furin site with appropriate substrate sequences. When combined with FP59, PrAg-U2 and PrAg-U3 selectively killed uPAR-overexpressing tumor cells in the presence of pro-uPA whereas showing very low toxicity to tPA-expressing cells such as human vascular endothelial cells. In contrast to PrAg-U2 and PrAg-U3, PrAg-U4 showed less uPA-dependent cytotoxicity but efficiently killed tPA-expressing cells. The ability of these mutated PrAg proteins to differentiate between uPA/uPAR- and tPA-expressing cells suggests that they may have potential use for targeting tumors in vivo.

uPAR has been proposed to play an important role in pro-uPA activation, serving as a template for the binding and localization of pro-uPA near to its substrate plasminogen on the plasma membrane. The concomitant binding of pro-uPA to uPAR and of plasminogen to as yet uncharacterized cell surface receptors was suggested to strongly potentiate uPA-mediated plasminogen activation possibly due to the formation of complexes aligning the two proenzymes in a way that compensates for their low intrinsic activity and favors a mutual activation process. The results reported in this study agree well with this "two proenzyme reciprocal activation model." In the presence of plasminogen, pro-uPA could be activated on uPAR-expressing cells but not on the surface of uPAR non-expressing cells. Blocking the binding of pro-uPA to uPAR protected cells from PrAg-U2 (plus FP59). We further showed that cell surface-bound plasminogen is required for the cascade activation of pro-uPA and PrAg-U2. Thus, pro-uPA and PrAg-U2 activation were significantly decreased by reducing the amount of cell surface-bound plasminogen by treatment of the cells with tranexamic acid. Moreover, the ability of plasmin inhibitors to decrease the cytotoxicity of PrAg-U2 toward HeLa cells further demonstrated that surface-bound plasmin plays a crucial role in the pro-uPA and PrAg-U2 activation cascade. For example, aprotinin, which inhibits plasmin activity both on the cell surface and in solution, protected HeLa cells from PrAg-U2 plus FP59, whereas alpha 2-antiplasmin, which can only inhibit plasmin in solution (10, 11), did not provide such protection. This set of data shows that uPAR expressed on the tumor cell surface, serving as a template to place pro-uPA near its substrate plasminogen, is essential for initiation of the pro-uPA activation cascade and therefore for the subsequent activation of the uPA-activated, mutated PrAg proteins.

We further showed that pro-uPA activated on the uPAR-expressing cell plasma membrane led also to activation of pro-uPA in the supernatant. However, PrAg-U2 was preferentially activated on the cell surface, with only a trace amount being activated in the supernatant. This can be explained as being due to the high affinity binding of PrAg-U2 and uPA to the cell surface, which effectively concentrates them there and results in high local concentrations. Both these receptor-binding events have nanomolar dissociation constants, 0.5 nM for pro-uPA binding to uPAR and 1 nM for PrAg binding to its as yet unidentified cell surface receptor (76, 77). These results suggested that the mutated PrAg proteins would be selectively cytotoxic to cells presenting activated uPA on their cell surfaces. This was confirmed by the co-culture experiment in which PrAg-U2 killed only uPAR-expressing HeLa cells while sparing uPAR non-expressing cells.

The results reported here clearly demonstrate that the cytotoxicity of these mutated PrAg proteins is strictly dependent on the tumor cell surface-associated plasminogen activation system, and in particular requires the presence of pro-uPA and its receptor uPAR on the tumor cell surface. Thus, these mutated PrAg proteins can be expected to target tumor tissues that overexpress both these factors. These results encourage the further testing of PrAg-U2 and PrAg-U3 in animal tumor models. The tPA-specific mutated PrAg protein, PrAg-U4, may be useful for targeting of tumors overexpressing tPA such as melanomas (78, 79), although the activation of PrAg-U4 by vascular endothelial cells warrants caution.

Many tumor-cell-selective cytotoxins have been created by replacing the receptor recognition domains of bacterial and plant protein toxins with cytokines, growth factors, and antibodies (80). Some of these "immunotoxins" derived from diphtheria toxin, Pseudomonas exotoxin A, and ricin have shown efficacy and have been approved for clinical use. However, a recurrent problem with these materials is nonspecific toxicity, due to uptake of trace amounts into normal, bystander cells. Because these toxins act catalytically, even a small amount of internalized toxin can seriously damage normal tissue. For this reason, it is important to increase the specificity of these recombinant fusion proteins for tumor cells.

Previous efforts to develop anthrax toxin fusion proteins as therapeutic agents have been modeled on the work described above and have focused on modification or replacement of domain 4, the receptor-binding domain of PrAg. Thus, work is ongoing to create cell-type-specific cytotoxic agents by modifying or replacing domain 4 with new targeting ligands (63, 81). However, we suggest that an optimum strategy for improving specificity is to combine two conceptually distinct targeting strategies in a single PrAg protein. Thus, a PrAg protein that is both retargeted to a tumor cell surface protein and dependent on the cell surface plasminogen activation system may achieve therapeutic effects while being free of the side effects observed with many of the existing immunotoxins.

    FOOTNOTES

* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Oral Infection and Immunity Branch, NIDCR, National Institutes of Health, 30 Convent Dr., MSC 4350, Bldg. 30, Rm. 303, Bethesda, MD 20892-4350. Tel.: 301-594-2865; Fax: 301-402-0396; E-mail: Leppla@nih.gov.

Published, JBC Papers in Press, March 12, 2001, DOI 10.1074/jbc.M011085200

    ABBREVIATIONS

The abbreviations used are: ATF, amino-terminal fragment of urokinase plasminogen activator; DMEM, Dulbecco's modified Eagle's medium; EF, edema factor; FP59, fusion protein of LF amino acids 1-254 and Pseudomonas exotoxin A domain III; HUVEC, human umbilical vein endothelial cells; LF, lethal factor; MMP, matrix metalloproteinase; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; PrAg, anthrax toxin-protective antigen; PrAg20, amino-terminal 20-kDa fragment of PrAg; PrAg63, carboxyl-terminal 63-kDa fragment of PrAg; PAGE, polyacrylamide gel electrophoresis; PAI-1, plasminogen activator inhibitor-1; PAI-2, plasminogen activator inhibitor-2; tPA, tissue plasminogen activator; uPA, urokinase plasminogen activator; uPAR, urokinase plasminogen activator receptor.

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DISCUSSION
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