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INTRODUCTION |
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
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-
, 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.
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MATERIALS AND METHODS |
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),
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,
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.
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RESULTS |
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.
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).
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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.)
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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
("0 " lanes 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.)
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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.)
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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.
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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,
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,
2-antiplasmin, which is an
inefficient inhibitor of cell surface-bound plasmin (10, 11), could not
protect the cells (Fig. 9). Aprotinin and
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 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.)
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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.)
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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.)
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DISCUSSION |
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
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.