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INTRODUCTION |
Tissue plasminogen activator
(tPA)1 is a multidomain
serine protease responsible for converting the zymogen plasminogen into the broad specificity serine protease plasmin. Plasmin is involved in a
range of biological processes, including fibrinolysis, tissue development, and tumor invasion and metastasis (1). Fibrin binding is a
critical step in the expression of significant tPA activity because
this enzyme is not produced as a zymogen but has only low levels of
activity in free solution (2). Although tPA is mainly associated with
hemostasis, in contrast to urokinase-type plasminogen activator (uPA),
which is considered to be more important in pericellular proteolysis,
several groups have identified or isolated a number of different cell
surface tPA receptors on endothelial cells and shown that binding of
tPA and plasminogen to the same cell can lead to accelerated plasmin
generation (3-5). Published dissociation constants for cell-tPA
interactions vary over a wide range,
10
11-10
7 M, although it now
appears that the highest affinity interactions may be due to cell
surface-associated plasminogen activator inhibitor type 1 (6). Binding
of tPA to plasminogen activator inhibitor type 1 requires the enzyme
active site and results in an inactive complex. The physiological
significance of putative tPA receptors of lower affinity is unclear
considering the low plasma level of tPA (70 pM). For
example, the tPA receptor annexin II, which binds tPA with a
Kd of 48 nM, is capable of producing modest stimulation of enzyme activity (<10-fold) in vitro
(7). Monocytes and monocytoid cells have been studied and found to bind
tPA with a low affinity, but high capacity, and are able to stimulate
tPA enzyme activity up to around 20-fold (8, 9). Other receptors on
liver cells have also been investigated in an attempt to understand the
structural features of tPA, both protein and carbohydrate, involved in
receptor binding and responsible for its short in vivo
half-life and rapid clearance from the circulation when tPA is
administered as a thrombolytic agent (10).
It is known that tPA-surface interactions are critical for the
regulation of plasminogenolytic activity, and this has been investigated extensively where fibrin is the biological surface (11,
12). Detailed structure/function studies on a large number of mutant
enzymes indicate that many areas of the protein are involved in
modulating the interaction with fibrin and hence regulating tPA
activity (13). The purpose of the present study was to look at the
regulation of tPA-catalyzed plasminogen activation on cell surfaces
using a number of cultured cell lines and to identify structural
features involved in tPA-cell interactions leading to stimulation of
plasminogen activation. Five cell types have been investigated in
conjunction with five tPA variants lacking various domains and/or
carbohydrate. To understand the factors regulating the kinetics of
plasminogen activation, our studies included ranges of cell (receptor)
concentration, as well as tPA and plasminogen concentration. Using this
approach, it has been possible to develop a model that explains how
cell interactions with tPA and plasminogen regulate the generation of
plasmin activity and changes in apparent Km and
kcat.
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EXPERIMENTAL PROCEDURES |
Proteins--
tPA is a multidomain protein comprising
finger-epidermal growth factor-kringle 1-kringle 2-protease domains
(F-E-K1-K2-P) and is glycosylated. In this study, full-length
glycosylated tPA was compared with nonglycosylated enzyme and deletion
mutants as follows (structure and alternative names): F-E-K1*-K2*-P*, full-length glycosylated tPA (asterisk indicates glycosylated domain);
F-E-K1-K2-P, full-length nonglycosylated tPA; E-K1-K2-P,
FtPA; K2-P,
kringle 2 and protease, K2P, BM 06.022, Reteplase; P, protease domain.
Full-length glycosylated tPA was International Standard melanoma tPA
(code 86/670, National Institute for Biological Standards and Control,
South Mimms, United Kingdom). Full-length nonglycosylated tPA was a
gift from Rhone-Poulenc Rorer (Vitry sur Seine, France) expressed and
purified from Escherichia coli (14). Both full-length enzymes had similar specific activities in fibrinolytic assays.
FtPA
and protease domain were generated as described below using the same
plasmid as for the full-length protein (pXL130), provided by
Rhone-Poulenc Rorer and expressed and renatured from E. coli inclusion bodies in the same way as full-length nonglycosylated tPA
(14). Expression of tPA was induced by addition of 40 mg/ml naladixic
acid (15) to cultures of E. coli strain N99cI+
(Pharmacia, Uppsala, Sweden) grown at 37 °C with aeration up to an
A600 nm of approximately 1.0. Cultures were
grown at 37 °C for a further 90-120 min before harvesting and
processing as described previously for the full-length enzyme (14) with the following modifications. After concentrating renatured protein using SP Trisacryl (IBF Biotechnics, Villeneuve-la-Garenne, France). The final lysine-Sepharose column step was replaced by fast protein liquid chromatography using a Mono S column (Pharmacia) run in 25 mM sodium acetate buffer, pH 4.3, containing 0.01% Tween
80, and purified tPA was eluted with a gradient of 0-1 M
NaCl. The serine protease domain purification was further modified by
omitting the pH 5.5 wash step on the SP Trisacryl column.
FtPA was produced by deleting residues Ser-1-Lys-49 and serine
protease domain by deleting residues Ser-1-Arg-275 using the polymerase chain reaction technique of deletion by overlap extension (16, 17) using the plasmid pXL130. Primers A and D were used for
both mutant proteins: primer A, 5'-ACCTGCAGCCAAGCTT-3', and primer D,
ATCCTGAAATCAGACCAAGTCCTG. For
FtPA, additional primers were B,
TCGCTGCAACTCATATGTAAGTAT, and C, ACTTACATATGAGTTGCAGCGAGC. For serine
protease domain generation, primers were B, CCTCCTTTGATCATATGTAAGTAT, and C, ATACTTACATATGATCAAAGGAGGGCTC. The final polymerase chain reaction product, ABCD, was digested with AvaI (New England
Biolabs, Beverly, MA) for insertion into pXL130. Re-ligated plasmids
were checked for correct size on agarose gels, and proteins were
examined by SDS-polyacrylamide gel electrophoresis to ensure correct
molecular weights. The final step in protein purification was fast
protein liquid chromatography and variants were checked for purity and correct molecular weight by SDS-polyacrylamide gel electrophoresis.
K2P (also known as BM 06.022 or Reteplase) was a gift from Roche
Molecular Biochemicals. This protein was expressed, renatured and
purified in a nonglycosylated form from E. coli (18). All tPA variants were measured in direct assays against chromogenic substrate S-2288 (Chromogenix, Mölndal, Sweden), and activity was
quantitated against the 2nd International Standard for tPA (code
86/670, National Institute for Biological Standards and Control, South
Mimms, United Kingdom) in a plasminogen activation assay in the absence
of stimulator as outlined below.
Cell Culture--
THP1, U937, Nalm6, Molt4, and K562 cells were
cultured in RPMI 1640 medium with 2-4 mM glutamine,
7-10% fetal bovine calf serum, and 1 mM sodium pyruvate
(pyruvate excluded for U937 cells) (Sigma or Life Technologies Ltd.).
Cells were grown to a density of 1.5 × 106 to
2.5 × 107 cells/ml in the presence of 5%
CO2. Cells were harvested by centrifugation and washed
three times in serum-free RPMI 1640 medium at 0-4 °C. The cell
pellet was drained and resuspended with gentle vortexing and washed
with 0.05 M glycine buffer, pH 3.5, containing 0.1 M NaCl for 2-3 min on ice (19). Cells were washed twice
with RPMI medium for final counting, resuspension and dilution in assay buffer (below) containing albumin at 1 mg/ml.
Plasminogen Activation Kinetics--
Plasminogen activation
reactions were carried out in microtitre plates in reaction volumes of
100 µl. In initial studies using full-length tPA enzymes, reaction
mixtures were 20 µl of tPA (final concentration 5 ng/ml,
70
pM), 40 µl of cells (final concentration, between
103 and 108 cells/ml) and 40 µl of substrate
mix containing Glu-plasminogen (Enzyme Research Laboratories, Swansea,
United Kingdom) at a final concentration of 100 nM and
chromogenic substrate S-2251 (Val-Leu-Lys-p-nitroanilide, Chromogenix, Mölndal, Sweden) at a final concentration of 0.15 mM. Reactions were performed in assay buffer consisting of
Tris-HCl buffer, pH 7.4, at 37 °C and a final ionic strength of
0.12, containing 1 mg/ml human serum albumin. In experiments with
higher concentrations of tPA (up to 1.5 nM), reaction
volumes remained the same. In experiments with tPA variants having
different molecular weights, starting activities of tPA had equivalent
activation rates with Glu-plasminogen in the absence of cells and
stimulation was calculated as activation rate with cells/activation
rate without cells under identical conditions. In studies using a range
of plasminogen concentrations reaction mixtures were made up of 20 µl
of tPA, 20 µl of cells (to give a range of cell densities as above),
20 µl of plasminogen (to give a range of concentrations up to a final concentration of 520 nM), and 40 µl of substrate solution
containing S-2251 as above. Cells were incubated for 15 min with tPA to
equilibrate to 37 °C before substrates were added to begin the
reaction. Absorbance was monitored at 405 nm, using a Thermomax
thermostatted plate reader (Molecular Devices Corporation, Stanford,
CA), producing the expected exponential increase for
p-nitroanilide resulting from plasmin production. Rates of
plasmin production were calculated from slopes of plots of A
versus s2 generated automatically from Thermomax
data by a program specifically written for this purpose (J
Waterman-Smith, Molecular Devices, Crawley, United Kingdom). These
slopes are proportional to plasmin generation and were calculated using
Enzfitter (Elsevier, Cambridge, United Kingdom), as outlined previously
(20, 21). Plasmin generation could be calculated from rates of
A/s2 by dividing by 22 310 A
M
1 s
1 from previously
determined values for the Km and
kcat of plasmin on S-2251 and the extinction
coefficient of p-nitroanilide under these conditions (21).
Simultaneous kinetic experiments were carried out with and without tPA
to control for intrinsic activator (most probably uPA) synthesized by
the cells and bound to the surface, which was not removed in the low pH
wash. Different cell lines had reproducible levels of associated
intrinsic activator giving background rates in the order THP1 > U937 > K562 > Molt4 > Nalm6 (essentially zero
background rate). Where background rates were high, i.e.
>10% of maximum rate achieved with added tPA (found with THP1 and
U937 cells), these were subtracted from observed rates with added tPA
in order to calculate rates due to exogenous activator only.
Plasminogen activation kinetics in the presence of varying
concentrations of 6-aminohexanoic acid (6-AHA) (Sigma) to investigate conformational changes and enzyme resulting from lysine analogue binding were conducted in a similar manner. Concentrations used for
these experiments were 50 pM tPA, 1 pM uPA, and
450 nM plasminogen.
To investigate the involvement of oligosaccharide moieties in binding
to cells, kinetic experiments were performed in the presence of 20 mM monosaccharides, mannose, fucose,
N-acetylglucosamine, N-acetylgalactosamine,
N-acetylneuraminic acid, and ovalbumin (10 mg/ml) (all from Sigma).
tPA Binding Studies--
Nalm6 and U937 cells were washed as
described above and finally resuspended in Hepes-buffered saline, pH
7.4. Ligand binding assays were performed by incubating full-length
125I-tPA (0.25-8.0 nM in a total reaction
volume of 100 µl) or K2P (0.5-16 nM in a total reaction
volume of 200 µl) with the washed cells (107 cells/ml)
for 2 h at 4 °C. Cells were then separated from the whole
reaction mixture by centrifugation of aliquots, in triplicate, in 20%
sucrose solution. A parallel set of reactions were incubated in the
presence of an excess of cold, unlabeled full-length tPA (glycosylated)
or K2P to determine nonspecifically bound radioactive counts.
Low affinity binding sites were investigated using higher
concentrations of tPA (up to 12 µM), and cell-bound tPA
was measured by enzyme activity as described previously (22). In these
experiments 200 µl of 107 cells/ml were incubated with a
range of tPA concentrations for 30 min at room temperature before bound
and free tPA were separated by centrifugation through 20% sucrose, as
above. Bound enzyme was determined in plasminogen activation assays, as
described above. Parallel experiments were performed without cells to
determine the amount of free tPA carryover in kinetic assays. These
activities were subtracted from specifically bound rates prior to data
analysis. Each tPA concentration was incubated and activity measured in triplicate.
Models--
The usual way to analyze tPA data in the presence of
a stimulator is to use a simple Michaelis-Menten equation,
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(Eq. 1)
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where [tPA] and [Pgn] are the total added concentrations of
enzyme and substrate, respectively. Km' and
kcat' refer to the kinetic parameters of the
bound enzyme. The contribution of the solution reaction is ignored.
Model 2 is based on the concentration effect of cells on tPA and
plasminogen, which interact with cell surface binding sites leading to
a localized raised reactant concentration in a cell-associated compartment. The principle of the model is as previously developed by
Nesheim et al. (23) to study regulation of the
prothrombinase complex on phospholipid vesicles and can be understood
in our system with reference to Scheme
1.
The abbreviations used in Scheme 1 are listed below, and the prime
notation denotes cell-associated protein or parameter. Thus, Pgn' and
tPA' are bound plasminogen and tPA, respectively; vc is the cell volume;
vca is the volume of the cell-associated compartment. Vc and Vca
are the corresponding volumes for the whole reaction mixture.
Vt is the total reaction volume, and
Vbulk = Vt
(Vc + Vca). The intrinsic
catalytic parameters and binding constants applied in the present study
are shown in Table I. Molar concentrations of bound enzyme and
substrate, relative to the whole reaction mixture, can be calculated
from simple equilibrium considerations knowing Kd,
the number of binding sites/cell, and cell concentration (hence
receptor concentration). However, rateca will depend not
only on the equilibrium between free and bound reactants but also on
the local concentration, which will depend on the size of
Vca. Thus it is necessary to calculate
[tPA'l]and [Pgn'l] (local concentration of
tPA' and Pgn', respectively) in order to calculate the local rate using
Km' and kcat'.
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(Eq. 2)
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(Eq. 3)
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Thus, it can be seen that the concentration factor
(R) for reagents in the cell-associated compartment relative
to the concentration for the whole reaction mixture is a ratio of 2 volumes and will vary with the cell concentration,
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(Eq. 4)
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(Eq. 5)
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or for a fixed reaction volume (e.g. 1 ml), as
follows.
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(Eq. 6)
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Because we are assuming that vca is a
constant, we define
as a constant equivalent to
1/vca in ml. Thus, under these conditions, the
following equation is used.
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(Eq. 7)
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can be used to calculate the volume of
vca for one cell and also h in Scheme
1. To calculate local concentrations, a concentration relative to the
whole solution is multiplied by R. That is,
[tPA'l ] = [tPA']·R and
[Pgn'l ] = [Pgn']·R, etc.
An equation for the reaction rate in the cell-associated compartment
can be derived using the Michaelis-Menten approach for the reaction A + S
AS
A + X, where A is enzyme (tPA) and S is substrate
(plasminogen).
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(Eq. 8)
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The measured rate, relative to the whole reaction mixture, is as
follows.
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(Eq. 9)
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This is a result of the following equation,
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(Eq. 10)
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(Eq. 11)
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and therefore the following equations are used.
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(Eq. 12)
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Applying the enzyme conservation equation,
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(Eq. 13)
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(Eq. 14)
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where [B] is the molar concentration of enzyme receptors and
Kd is the dissociation constant for enzyme and
receptor. Rearranging results in the following equation.
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(Eq. 15)
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Then, dividing Equation 12 by Equation 15 results in the
following equation.
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(Eq. 16)
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This can be rearranged as follows and used as model 2, with
respect to tPA and plasminogen.
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(Eq. 17)
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Bound plasminogen [Pgn'] is calculated in the usual
way,
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(Eq. 18)
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where [Pgno] is added plasminogen,
KD is the dissociation constant for
plasminogen-receptor binding, and [D] is the molar concentration of
plasminogen receptors. Because [tPA]
[Pgn] in the system,
there is no need to account for enzyme-substrate complex in the
distribution of bound and free plasminogen.
This equation is closely related to the relationship for specific
activation of an enzyme-catalyzed reaction (24) but with the
modification of Km' to
Km'/R. This model was used to derive
values for kcat' and
(and hence
vca and h) by nonlinear regression
analysis using Grafit version 4 (25). Model data were generated using
Mathcad, version 7 (Mathsoft Inc., Cambridge, MA).
Model 3 was also considered; it could account for the observed data and
is related to the model of Lu and Nelsestuen (26) for prothrombinase
activity on vesicles. This model was similar to model 2, being a
version of the specific activation equation, although in this case it
was assumed that binding (giving tPAb*) modified the
intrinsic enzyme parameters Km' and
kcat' directly. To account for the profile of
rateca with cell concentration (the "template effect"),
it was necessary to assume that bound enzyme could only react with
nonbound plasminogen in free solution (pgnf). This is shown
in Scheme 2.
Model 2 can then be modified to give model 3.
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(Eq. 19)
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RESULTS |
Full-length tPA and Deletion Variants--
All of the cell types
tested in these studies, U937, THP1, K562, Molt4, and Nalm6, were able
to act as promoters in kinetic assays measuring plasminogen activation
by full-length tPA. These cell types gave similar results in terms of
level of stimulation and cells/ml required for peak stimulation.
Results are presented in detail for U937 cells, a widely used
monocytoid line, and Nalm6 cells, a pre-B leukemic line that expresses
negligible levels of intrinsic plasminogen activators. Fig.
1 shows the relationship between
activation rate and cell density for full-length glycosylated tPA,
nonglycosylated tPA, and nonglycosylated deletion variants. To ensure
good comparability between variants, all data for one cell type were
collected simultaneously on one microtitre plate using one batch of
cells. Here, initial enzyme activities in the absence of cells gave
similar rates of plasminogen activation and the effectiveness of cells
in promoting activation can be seen in changes in rate of plasmin
production. Degree of stimulation was also calculated, as rate with
cells/rate without cells. Over a range of cell densities, a peak of
stimulation was seen between 106 and 107
cells/ml for all tPA variants. The bell shaped pattern for stimulation versus log of cell density is in accord with a template
mechanism dependent on ternary complex formation of cell-bound
activator and substrate. This is supported by radioligand binding
studies that demonstrate that both these cell types are able to bind
tPA and plasminogen to the cell surface (8). All cell types used in
these studies showed this relationship between stimulation and cell
density, although peak activity could fall between
106-108 cells/ml, and stimulation could
approach 80-fold, with some variation between batches of cells. The
ranking order of variants shown in Fig. 1 was constant. For the data
presented in Fig. 1, maximum levels of stimulation were observed with
full-length glycosylated tPA and were 51-fold with U937 cells and
24-fold with Nalm6 cells.
FtPA and K2P showed a reduced level of
stimulation with both cell types examined, 12- and 21-fold,
respectively, with U937 cells and 9- and 12-fold, respectively, with
Nalm6 cells in the experiments shown. There was a small but measurable
stimulation for the protease domain of <2-fold with Nalm6 cells. This
was difficult to measure in the presence of U937 cells due to the higher background rates (from intrinsic activator) associated with
these cells. Unexpectedly, K2P stimulation was always greater than
FtPA even though
FtPA lacks only one domain and is structurally much closer to full-length tPA than K2P. However, it is clear that loss
of finger domain in either mutant enzyme results in a marked drop in
stimulation by cells, but there was still a significant level of
stimulation, apparently due to sites on kringle 2. The lower activity
of nonglycosylated versus glycosylated full-length tPA was
studied in more detail.

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Fig. 1.
Activation of Glu-plasminogen by tPA variants
in the presence of U937 and Nalm6 cells. Rates of plasmin
generation were measured in the absence of cells using concentrations
of each variant titrated to give similar rates and the effects of cells
over a range 0 to 4 × 107 cells/ml. Rates with U937
cells were corrected for background activation rate due to intrinsic
cell-produced activator determined simultaneously in reactions without
added tPA. tPA variants used were full-length glycosylated tPA ( ),
full-length nonglycosylated tPA ( ), FtPA ( ), K2-P tPA ( ),
protease domain (×), and no tPA (+).
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Glycosylated and Nonglycosylated tPA--
Fig.
2 shows the degree of stimulation for
glycosylated (A and C) and nonglycosylated
(B and D) full-length tPA with U937 and Nalm6
cells. Over a range of cell densities, there was a peak of stimulation
that was independent of the tPA concentration from 0.075 to 1.5 nM. Over this range of added tPA, there was no evidence of
any saturation of binding sites, indicating that binding is not
restricted to low numbers of receptors per cell. In the experiments shown in Fig. 2, comparative stimulation for U937 and Nalm6 were 78.9 ± 2.8 (mean stimulation ± S.D.) and 60.0 ± 4.8, respectively, for full-length glycosylated tPA and 34.9 ± 5.8 and
30.5 ± 3.3, respectively, for full-length nonglycosylated tPA.
Lack of glycosylation resulted in a drop in peak stimulation of around
50%. Thus, it is possible that glycosylation sites are directly
involved in cell interactions, or alternatively, these could be
indirect effects such that glycosylation of kringles affects lysine
binding. In competition experiments including monosaccharides up to 20 mM or 10 mg/ml ovalbumin in kinetic assays, normal patterns
of stimulation were observed with no detectable inhibition over a range
of cell densities from 0 to 3 × 107 cells/ml. These
levels of monosaccharides have previously been shown to inhibit binding
of tPA to glycosylation site-specific cell surface receptors (27). Our
results indicate that lack of glycosylation has an indirect effect on
stimulation of tPA activity by cells.

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Fig. 2.
Stimulation of Glu-plasminogen activation by
tPA in the presence of increasing concentrations of cells.
A and B are results using U937 cells corrected
for background activation in the absence of tPA due to intrinsic
cell-produced activator. C and D are results with
Nalm6 cells (no correction for negligible background rates).
A and C show stimulation of activation using
full-length glycosylated tPA; B and D are with
full-length nonglycosylated tPA. Concentrations of tPA were 0.075 nM ( ), 0.3 nM ( ), and 1.5 nM
( ).
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Plasminogen Concentration--
The experiments shown in Figs. 1
and 2 were conducted at a fixed plasminogen concentration of 100 nM, and further experiments were performed to investigate
the effects of varying substrate concentration. If the interaction
between tPA and cell receptors is affecting only the
Km of the plasminogen activation reaction it should
be possible to overcome the lower relative stimulation of the
nonglycosylated tPA variants using higher substrate concentrations.
Fig. 3A shows similar
Michaelis-Menten curves for the tPA variants at the optimal cell
density of 107 cells/ml. Curve fitting was by nonlinear
regression to the standard Michaelis-Menten equation, although there is
some suggestion of apparent substrate inhibition with the full-length
enzymes (see under "Discussion"). The apparent
Km values were similar for the full-length enzymes
(20 ± 4 and 12 ± 5 nM for glycosylated and
nonglycosylated, respectively) and somewhat higher for the deletion
variants (46 ± 11 and 33 ± 8 nM for K2P and
FtPA, respectively). The maximum rate of activation was 0.22 pM/s for full-length tPA, which is lower than for the free
solution reaction (compare Fig. 3, A and B).
Further reductions in Vmax were apparent for the deletion variants. Clearly, cell binding affects both apparent Km and Vmax. The activity of
two variants, full-length tPA and
FtPA, in the absence of cells is
shown for comparison, and these variants have similar activity over
this range of plasminogen, where the slope of the line can be used to
estimate kcat/Km. The
literature values for full-length tPA applied in these studies were
Km = 65 µM and
kcat-0.06 s
1,
kcat/Km = 923 M
1 s
1 (11). This is close to
the value derived from Fig. 3A of 786 M
1 s
1. Fig. 3B shows
the change in plasminogen activation in the absence of cells for
full-length tPA up to 16 µM plasminogen. Rates did not
approach saturation, and hence it was not possible to calculate Km and Vmax or
kcat.

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Fig. 3.
Relationship between plasminogen activation
rate and plasminogen concentration for tPA variants at fixed cell
density. A shows plasminogen activation rates,
determined at 1 × 107 cells/ml, over plasminogen
concentration range of 1-520 nM for each tPA variant,
showing how cells affect the apparent Vmax and
Km for the activation reaction with each tPA
variant. Curves shown are full-length glycosylated tPA ( ),
full-length nonglycosylated tPA ( ), FtPA ( ), K2P tPA ( ),
full-length glycosylated tPA with no cells (×), and FtPA with no
cells (+). The curves are fitted to a normal Michaelis-Menten equation
by nonlinear regression analysis. B shows data for
full-length glycosylated tPA activation of plasminogen (up to 16 µM) in the absence of cells.
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tPA-Cell Binding Studies--
To investigate the levels of tPA
binding under the conditions used in these kinetic experiments,
radiolabeled tPA was incubated with cells over a low concentration
range, up to 16 nM. Under these conditions, very little
specific binding of 125I-tPA could be observed, indicating
the absence of significant levels of a high affinity binding site with
a Kd in this range of tPA concentrations (results
not shown). In order to investigate lower affinity binding sites, cells
were incubated with higher concentrations of tPA and detection was by
enzyme activity rather than radioactivity. This approach allows higher
concentrations of tPA to be used as no excess of unlabeled tPA is
required, and only bound, active enzyme is detected, which is the
species of interest in these studies. Results from these studies are
shown in Fig. 4 using full-length tPA and
K2P. Values for maximum binding and Kd were
determined from direct fitting to these data using a single site
binding isotherm model. The Kd values were 487 ± 117 nM for the full-length enzyme and 1122 ± 320 nM for K2P. There was also an approximately 2-fold
difference in the maximum level of bound activity of these two enzymes
(1659 ± 250 for full-length tPA and 835 ± 166 A/s2 for K2P). This could be due to a decreased
number of binding sites for K2P compared with full-length tPA, or
alternatively a reduction in the activity of K2P compared with
full-length tPA. In either case, it is possible in theory to use these
data to calculate the number of binding sites per cell. However, in
practice, this is likely to be inaccurate, as weak interactions such as this will undergo some dissociation during processing and determination of activity (28).

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Fig. 4.
Ligand binding data for the interaction of
full-length tPA and K2P with Nalm6 cells. Nalm6 cells
(107 cells/ml) were incubated with varying concentrations
of tPA, either full-length glycosylated ( ) or K2P ( ), and bound
enzyme was determined, after separation of nonbound, in plasminogen
activation assays. Parallel experiments without cells were conducted to
determine nonspecific enzyme carryover (open symbols).
Nonspecific binding was subtracted from bound activity (× for K2P),
and nonlinear regression analysis was used to determine
Kd and maximum level of activation by fitting to a
single binding isotherm using corrected data.
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Comparison of Model and Experimental Data--
Model 1 was not
considered in detail as it cannot account for the rate
versus cell concentration profile observed in experimental data (Figs. 1 and 2). The values used with models 2 and 3 to simulate cell-associated plasminogen activation rates are shown in Table I. The initial assumptions of the model
outlined above stated that the effect of cells in this system is to
concentrate reactants and intrinsic enzyme parameters,
kcat and Km, remain the same
whether bound or free. However, it became clear that concentration of
reactants alone was insufficient to explain the results as no
satisfactory fitting of the data could be achieved. It was necessary to
allow for a reduction in kcat' for the
cell-bound reaction. This requirement is apparent from the equation for
model 2 and Fig. 3. Because model 2 is a related to competitive
inhibition, Vmax' (cell-associated) should
approach Vmax (in solution) at high [Pgn'].
This is not the case from the data shown in Fig. 3, A and
B, indicating that kcat' is lower
than kcat.
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Table I
Model parameters for the cell associated plasminogen activation used in
conjunction with model 2
Km and kcat are the enzyme
parameters for the activation of plasminogen by tPA in free solution.
Km' and kcat' are the parameters
for the cell associated reaction. is a measure of the concentration
effect of cells for plasminogen activation.
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Using the values for Kd, binding sites, and
Km shown in Table I, data were fitted by nonlinear
regression analysis to determine values for
kcat' and
to a set of data for which the
activation rate was measured with varying cell and plasminogen
concentrations. Because there are two independent variables in this
system (plasminogen and cells), multiple regression analysis was used
to derive values for the unknown parameters, using Grafit, version 4.0, software (25). The results from fitting in this way gave the results
shown in Table I for kcat' and
. The
kcat' of 0.004 s
1 represents a
15-fold decrease relative to the free solution reaction. The value of
is a measure of the concentration of reactants around a cell. This
estimate of 3.65 × 1011 can be thought of as the
reciprocal of the reaction volume (in ml) around 1 cell. Model data and
experimental data are shown in Fig. 5 for
comparison and are in reasonable agreement as can be seen from the
pattern of stimulation, the absolute rates, and the optimum cell
concentration. Both sets of data have the same trends for increasing
apparent Km with increasing cell concentration and
increasing optimum cell concentration with increasing plasminogen
concentration. The level of stimulation of bound enzyme activation rate
was also in good agreement with experimental results over this range of
cell and plasminogen concentrations.

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Fig. 5.
Model results and experimental data showing
the dependence of activation rate on cell and plasminogen
concentration. Experimental data were collected using Nalm6 cells
and Glu-plasminogen over the ranges shown in the presence of 70 pM full-length glycosylated tPA. Model data were generated
for full-length tPA from model 2 and the parameters in Table I using
kcat' and from nonlinear regression
analysis. Model results using model 3 and values for
Km' = 7 nM and
kcat' = 0.004 s 1 from nonlinear
regression analysis are not shown, but were very similar to those
presented for model 2.
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Model 3 was also applied to the data shown in Fig. 5. Fitting by
multiple regression provided estimates for cell-bound enzyme parameters
of Km' = 7 ± 2 nM and
kcat' = 0.0039 ± 0.0003 s
1.
Thus, the same value for kcat' as with model 2, but with a decrease of almost 10,000-fold in Km from
65 µM to 7 nM on cell binding. The
three-dimensional plot was essentially superimposable on the plot shown
in Fig. 5 (model), with only minor differences at the extremes.
Further exploration of model 2 was performed using constants and
variables for full-length tPA and the deletion variant K2P given in
Table I, in an attempt to replicate the results shown in Figs. 1 and 3.
Model results are shown in Fig. 6 for
fixed plasminogen (100 nM) and varying cells (Fig.
6A) and fixed cells (107 cells/ml) with varying
plasminogen (Fig. 6B). Data were generated applying the two
possible alternative explanations for the results in Fig. 4. That is,
the reduced maximum binding to cells of K2P relative to full-length tPA
was due to a 2-fold reduction in binding sites, or to a 2-fold
reduction in the cell-bound kcat'. By comparing Fig. 6 with the experimental data shown in Figs. 1 and 3, it is clear
that the best explanation is a reduction in
kcat', which is 0.004 s
1 for
full-length tPA and 0.002 s
1 for K2P at the cell surface.
This provides levels of stimulation in Fig. 6B of 36.1- and
16.7-fold for full-length tPA and K2P, respectively, in the same range
as the data from Fig. 1. Model 2 also explains the approximate 2-fold
increase in apparent Km' noted above for deletion
variants of tPA relative to full-length enzyme (Fig. 3) as being due to
the 2-fold increase in Kd (from Fig. 4 and Equation 17).

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Fig. 6.
Model results showing how rate of plasminogen
activation depends on cell concentration or plasminogen
concentration. A is a classic template profile of cells
versus activation rate generated using model 2 and the
values in Table I in the presence of 100 nM plasminogen for
full-length tPA ( ) and K2P (diamonds) for comparison with
data in Fig. 1. B shows the effect of varying plasminogen at
1 × 107 cells/ml for the same enzymes and tPA with nl
cells (×) for comparison with Fig. 3. Open diamonds show
the assumption that K2P binds to Nalm6 cells with a
Kd = 1122 nM and has half the binding
sites/cell as full-length tPA and kcat'=0.004
s 1, as for full-length tPA, to account for the data in
Fig. 4. Closed diamonds are for Kd = 1122 nM with the same number of binding sites per cell as
full-length tPA (1.6 × 107/cell), but
kcat' for K2P is now 0.002 s 1.
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Plasminogen and tPA Conformational Changes--
To investigate
possible involvement of lysine binding sites on tPA or plasminogen
inducing conformational changes leading to stimulation of enzyme
activity, kinetic experiments were performed in the presence of a range
of 6-AHA concentrations instead of cells. Fig.
7 shows the effect of 0.005-10
mM 6-AHA on activation rate of Glu-plasminogen by
full-length glycosylated tPA, tPA protease domain, and an alternative
activator, uPA. Clearly, there was little effect of 6-AHA binding to
tPA as full-length tPA and protease were identical (overall, a slight
inhibition of 14% was observed in both cases). A different mechanism
operates in the case of uPA where the conformational change in
Glu-plasminogen at around 2 mM 6-AHA, previously found
(29), did produce an enhanced activation rate (3.2-fold stimulation in
this case), in agreement with earlier findings (30). Activation by tPA
was much less sensitive to this conformational change. Similar
experiments using Lys-plasminogen showed only inhibition with
increasing 6-AHA such that over the same concentration range,
activation rates were inhibited by 69.5, 60.9, and 35.1% for tPA,
protease domain, and uPA, respectively.

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Fig. 7.
The effect of 6-AHA on the activation rate of
Glu-plasminogen by tPA and uPA. Activation kinetics were
determined in parallel for the activation of 450 nM
Glu-plasminogen in the presence of 0-10 mM 6-AHA for 1 pM uPA ( ) and approximately 50 pM tPA
full-length ( ) or protease domain ( ).
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All of the studies shown above using Glu-plasminogen as substrate were
repeated using Lys-plasminogen and gave similar results for these cell
types and tPA variants, although activation rates were higher with and
without cells resulting in lower levels of stimulation (data not shown).
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DISCUSSION |
From the data presented here, it is clear that the cells used are
able to act as efficient promoters of plasminogen activation by tPA.
These cells have previously been characterized for binding of
plasminogen, uPA and tPA (8, 9, 31, 32). In the present detailed study,
the levels of stimulation with full-length glycosylated tPA were up to
30-80-fold, which is as good as or better than observed in previous
studies with cells or isolated receptors or heparin or fibrinogen
fragments (5, 21). Furthermore, the present study was performed using
physiological tPA concentrations. The two cell types studied in detail
here were of different origins (U937, a monocytoid cell line, and
Nalm6, a pre-B leukemic cell line) but were very similar in their
behavior as promoters of tPA activity. In fact, all cells investigated
showed similar levels and patterns of stimulation. The pattern of
stimulation of the different tPA variants, the ubiquitous binding
sites, and the results of binding studies all suggest that relatively
weak nonspecific interactions regulate the activity of cell-associated
tPA. It is also interesting that stimulation of kinetics by cells or by fibrin are similar in so far as they both depend on multiple sites of
interactions. Kringle 2 and finger-growth factor domains have been
identified as important in fibrin binding and activity (33, 34), but it
is now apparent that multiple interactions with fibrin throughout the
tPA molecule may play a role in regulating activity (13), and the
mechanism of binding may be complex (35). The situation appears to be
similar when cells are promoters, although in our studies finger domain
is very important, as suggested in previous studies using endothelial
cells (36, 37).
The effects of domain deletion on stimulation seen here with tPA
variants was best explained by lower affinities for binding and a
reduction in activity of cell-bound enzyme. It is interesting in this
regard that a recent study using vascular smooth muscle cells found a
3-fold lower maximal cell-bound activity for K2P relative to
full-length tPA and concluded this was due to the loss of a higher
affinity site available only to the full-length molecule (22). In the
present study, data fitting was not improved using a two site model.
The observation of Kohnert et al. (18) that fibrin
stimulation of K2P was only 35% the stimulation observed for
full-length tPA is also very similar to our results with cells. Furthermore, optimal heparin concentrations have been found to stimulate full-length tPA more than K2P, by 22- and 13-fold,
respectively (38). All these observations would be expected for a
mechanism in which bound K2P has a lower kcat'
than bound full-length tPA.
Mechanism of Stimulation--
From a practical viewpoint,
reporting observed kcat and
Km values is useful as a measure of degree of
stimulation by comparing
kcat/Km in the presence and
absence of cells. However, caution is necessary not to over-interpret
these parameters in proposing direct effects on bound proteins.
Furthermore, the apparent Km and
kcat values from model 1 are related to total
substrate and enzyme added, irrespective of whether it is cell-bound or
in solution.
Model 2 has been applied in the present study as an alternative
approach. Here, stimulation is primarily a result of the concentration of reactants by the cell. The initial assumptions in model 2 were that
Km and kcat are the same
whether reactions are in free solution or cell-associated. The
experimental data in Fig. 3 showed that this simple assumption could
not be applied and there was a need to reduce the
kcat' for the cell-bound enzyme. Curve fitting
could then be used to generate values for
and kcat'. When this was done, model and
experimental results were in good agreement, within the variation
observed from batch to batch of cells. The reason for the reduction in
cell kcat' relative to solution reaction is not
known but was necessary in all models. The lower activity could result
from different conditions close to the cell surface due to different
physical conditions (pH, ionic strength, etc.) due to the effect of
cell membrane and components (39) or factors relating to replenishment
of substrate and removal of product in this compartment such that the
maximum kcat' is not observed (40).
Alternatively, this could be a real regulatory effect on the cell
surface. For example, under conditions of high local plasminogen
concentration close to the cell, there may be allosteric regulation by
substrate, which has been noted in some previous studies (41, 42).
Model 2 is also able to account for the template profile seen in Figs.
1 and 2 for varying cell concentration, which is explained by two
competing effects. As cell concentration increases, mass action causes
the equilibrium to shift toward bound reactants up to a plateau, with
concomitant increase in observed rate. Conversely, as cell
concentration increases, the volume of Vca also
increases, and hence the concentration of reactants in
Vca decreases.
The value of
derived from multiple regression fitting can be used
to calculate the volume of the cell-associated compartment and from
this the height of the layer (Scheme 1, h) can be
determined. A
value of 3.65 ± 0.97 × 1011
translates into a value of h = 11 nm (range, 8-15 nm)
using a uniform cell diameter for Nalm6 cells of 9 µm. This is a
significant value from several points of view. The previous study by
Nesheim et al. (23), using the same principles to develop
their model to study prothrombinase activation in the presence of
phospholipid vesicles, used previously published physical studies to
estimate the height of their "interface shell." The height of this
shell was initially set at 12 nm but was adjusted upwards to 39 nm to fit the data more closely. Furthermore, light scattering studies on
plasminogen have provided information on dimensions of the molecule
(29). Native Glu-plasminogen is a prolate elipsoid of dimensions
14.7 × 5.7 × 5.7 nm, which extends to a longer open form in
the presence of 6-AHA and presumably as Lys-plasminogen and plasmin.
The longest dimension of these forms is then increased to 24 nm. From
these values, it would appear that the cell-associated compartment is a
single layer of molecules around the cell. Interestingly, prothrombin
is also a prolate elipsoid of similar dimensions to plasminogen and has
been postulated to bind and extend from the vesicle membrane to a
distance of 11 nm, the longest dimension (43). The significance of the
height of interface shell has also been discussed by Nesheim et
al. (23) and clearly does not represent a distinct isolated
compartment. What these observations most likely mean is that the
probability of plasminogen activation taking place is high at distances
very close to, or in contact with, the cell surface. Another
observation arising from the model developed by Nesheim et
al. (23) for prothrombin activation was apparent substrate
inhibition resulting from competition between enzyme and substrate for
the same binding sites. The consequences were similar in the tPA and
prothrombin systems.
Model 3 has also been included in the analysis and, like model 2, can
explain the experimental data well. The crucial difference between
models 2 and 3 is that the latter requires a huge change in intrinsic
enzyme Km of almost 10,000-fold on cell binding.
There is no evidence that this magnitude of change can happen, for
example, as a result of conformational changes. Indeed, attempts to
induce conformational changes using lysine analogues that are known to
act through kringle domains did not produce any significant stimulation
of activation rate with tPA, as seen in Fig. 7. Previous studies have
also suggested that the protease domain acts independently of the
N-terminal, A-chain domains (34, 44). It is interesting that the uPA
system is stimulated in this way and is more sensitive to the
conformation of plasminogen as affected by 6-AHA, fibrinogen fragments,
or monoclonal antibodies to the 6-AHA binding site (30, 45). Model 3 also assumes that only free plasminogen is available for reaction with
cell-bound tPA if the correct "template" profile is to be obtained.
Again, there is no evidence to support this. On the contrary, lysine analogues that are able to block fully plasminogen binding, but only
partially block tPA binding to cells, are completely effective at
inhibiting cellular stimulation of plasminogen activation in our
studies (data not shown) and elsewhere (22).
Although there is reasonably good agreement between the data and model
2, there are a number of sources of error in the parameters we have
applied. A simplification that was made in multiple regression analysis
was to ignore any competition between plasminogen and tPA for the same
binding sites although this has been demonstrated. It was not possible
to include this feature in the data fitting. However, it could be
included in simulated data using model 2 and had a noticeable effect at
high plasminogen concentrations (>500 nM) at which rates
began to fall. This effect would lead to an underestimation of the true
cell-bound kcat'. In vivo, this phenomenon may play a major role in inhibiting excess plasminogen activation as the concentration of plasminogen is around 2 µM. In the present study any effects of single chain to
two chain tPA conversion or generation of Lys-plasminogen have also
been disregarded. This is justified because of the conditions chosen for the study in which reactants were maintained at low concentrations and plasmin generation monitored at early times to gather initial rates
of plasmin generation. Furthermore, previous work has shown that single
chain and two chain tPA behave similarly in the presence of cells (21).
Other inherent problems are the difficulties in accurately measuring
dissociation constants for weak complexes, as here for both tPA and
plasminogen. The cell lines used also showed some inevitable batch to
batch variation, although Nalm6 cells were ideal, being exceptionally
homogeneous in terms of size and shape, and were very robust in our
assays. tPA is not an easy enzyme to work with due to low solubility
and difficulties in quantitation: for example, active site titration
and artifacts arising from allosteric interactions with chromogenic
substrates (46). However, the approach used in the present study to
standardize free solution activities of plasminogen activation of all
tPA variants against full-length tPA would minimize these difficulties and afford the most direct way of studying stimulation. Another potential problem with model 2 is that it ignores the contribution of
the free solution reaction. Although this is insignificant at low
substrate concentrations, it cannot be disregarded as substrate is
increased. Model 2 is closely related to the equation for specific activation of an enzyme, which makes the simplifying assumption that
the unstimulated enzyme is not active (24). More complex equations have
been developed when this simplification does not hold (47). As apparent
from the equation for model 2, the value for
is closely related to
Km'. There is no way of measuring Km' directly, which adds uncertainty to the value
for
and h. However, what can be said from the results using model 2 is that large changes in Km on cell binding are not essential to explain stimulation of activity.
Despite these problems, model 2 did provide a good explanation for the
experimental data obtained. Nevertheless, it is undoubtedly a
simplification, and as with the work of Nesheim et al. (23), it is open to criticisms. A number of alternative theoretical treatments dealing with surface activation kinetics have been proposed,
some of which are also concerned with coagulation enzymology in the
presence of phospholipid vesicles (26, 48-50) or on platelets (51) or
involved other systems (40). These approaches deal with problems
related to heterogeneous catalysis in a variety of ways, placing more
emphasis on the effect of co-localization of reactants, kinetics of
interactions with and on the surface, orientational effects,
conformational changes, receptor occupancy, and the effects of
transition from three-dimensional to two-dimensional geometry. Model 2 does have the advantage of being intuitively simple and relies
primarily on the concentration effect of cells with only minor
modifications to cell-associated enzyme behavior, which appear
reasonable. The model can simulate observed data over tPA, plasminogen,
and cell concentration ranges, with different tPA variants. The data
support the idea that low affinity, high capacity interactions can
regulate tPA activity by cells, even at physiological tPA
concentrations. This approach may be useful in understanding tPA
regulation by fibrin and in the regulation of other, surface-bound,
enzyme activities.