(Received for publication, July 27, 1995; and in revised form, October 17, 1995)
From the
Fluorescent analogs of the proteinase zymogen, plasminogen (Pg),
which are specifically inactivated and labeled at the catalytic site
have been prepared and characterized as probes of the mechanisms of Pg
activation. The active site induced non-proteolytically in Pg by
streptokinase (SK) was inactivated stoichiometrically with the
thioester peptide chloromethyl ketone, N-[(acetylthio)acetyl]-(D-Phe)-Phe-Arg-CH
Cl;
the thiol group generated subsequently on the incorporated inhibitor
with NH
OH was quantitatively labeled with the fluorescence
probe, 2-((4`-iodoacetamido)anilino)naphthalene-6-sulfonic acid; and
the labeled Pg was separated from SK. Cleavage of labeled
[Glu]Pg
by urokinase-type plasminogen activator
(uPA) was accompanied by a fluorescence enhancement
(
F
/F
) of 2.0,
and formation of 1% plasmin (Pm) activity. Comparison of labeled and
native [Glu]Pg
as uPA substrates showed that
activation of labeled [Glu]Pg
generated
[Glu]Pm
as the major product, while native
[Glu]Pg
was activated at a faster rate and
produced [Lys]Pm
because of concurrent
proteolysis by plasmin. When a mixture of labeled and native Pg was
activated, to include plasmin-feedback reactions, the zymogens were
activated at equivalent rates. The lack of potential proteolytic
activity of the Pg derivatives allowed their interactions with SK to be
studied under equilibrium binding conditions. SK bound to labeled
[Glu]Pg
and [Lys]Pg
with
dissociation constants of 590 ± 110 and 11 ± 7
nM, and fluorescence enhancements of 3.1 ± 0.1 and 1.6
± 0.1, respectively. Characterization of the interaction of SK
with native [Glu]Pg
by the use of labeled
[Glu]Pg
as a probe indicated a
6-fold higher
affinity of SK for the native Pg zymogen compared to the labeled Pg
analog. Saturating levels of
-aminocaproic acid reduced the
affinity of SK for labeled [Glu]Pg
by
2-fold
and lowered the fluorescence enhancement to 1.8 ± 0.1, whereas
the affinity of SK for labeled [Lys]Pg
was
reduced by
98-fold with little effect on the enhancement. These
results demonstrate that occupation of lysine binding sites modulates
the affinity of SK for Pg and the changes in the environment of the
catalytic site associated with SK-induced conformational activation.
Together, these studies show that the labeled Pg derivatives behave as
analogs of native Pg which report functionally significant changes in
the environment of the catalytic site of the zymogen.
Activation of the serine proteinase zymogen, plasminogen (Pg) ()to form plasmin is the central event in the dissolution of
blood clots by the fibrinolytic system. The physiological serine
proteinases, urokinase-type plasminogen activator (uPA) and tissue-type
plasminogen activator (tPA), activate Pg by cleavage of the
Arg
-Val
bond in the catalytic domain of the
zymogen, thereby initiating degradation of fibrin by plasmin (reviewed
in Henkin et al.(1991) and Ponting et al. (1992a)).
The rate of Pg activation and its physiological localization are
regulated by assembly of activator-Pg, enzyme-substrate complexes bound
to the fibrin surface. The rate of plasmin formation is increased as a
result of coordinated interactions among the activating enzymes, Pg,
and fibrin, with fibrin binding mediated substantially by lysine
binding sites on tPA and Pg (Hoylaerts et al., 1982; Nesheim et al., 1990; Liu and Gurewich, 1992; Fleury et al.,
1993). The rate of Pg activation is also controlled by conformational
equilibria between compact and extended forms of [Glu]Pg
which are shifted to the more rapidly activated, extended forms by
binding of lysine analogs (Violand et al., 1978; Urano et
al., 1987, 1988; Mangel et al., 1990; Christensen and
Molgaard, 1992; Marshall et al., 1994). Reactions catalyzed by
plasmin play a complex role in accelerating the rate and localizing Pg
activation. Plasmin converts the activating enzymes from the
single-chain to the two-chain forms (Rijken et al., 1982;
Collen et al., 1986; Longstaff et al., 1992); it
cleaves a 77-residue peptide from the amino terminus of
[Glu]Pg to produce the more reactive, [Lys]Pg form
of the zymogen (Violand and Castellino, 1976; Christensen, 1977; Lucas et al., 1983; Fredenburgh and Nesheim, 1992); and it
transforms fibrin into a more effective surface and cofactor of the
reactions by generating new sites for productive complex assembly
(Norrman et al., 1985; Bok and Mangel, 1985; Higgins and
Vehar, 1987; Fleury et al., 1993).
The bacterial protein,
streptokinase (SK) activates Pg by a different mechanism than that of
uPA and tPA. In this mechanism, specific binding of SK to Pg induces
formation of an active catalytic site in the zymogen through a
conformational change (McClintock and Bell, 1971; Reddy and Markus,
1972; Schick and Castellino, 1974). Non-proteolytic activation of Pg by
SK is closely coupled with enzymatic activation of Pg by Pg-SK and
plasmin-SK complexes, by their cleavage of the
Arg-Val
bond (Bajaj and Castellino, 1977;
Gonzalez-Gronow et al., 1978; Davidson et al., 1990).
Similar to activation by the physiological proteinases, the rate of
SK-initiated plasmin formation is regulated by interactions with fibrin
(Strickland et al., 1982; Chibber et al., 1985;
Cassels et al., 1987), Pg conformational equilibria (Chibber
and Castellino, 1986), and plasmin proteolytic reactions (Strickland et al., 1982). Studies aimed at defining the molecular
mechanisms of Pg activation have been complicated by the concurrent
plasmin-catalyzed reactions. This problem has been addressed recently
by the use of engineered variants of Pg, uPA, and tPA, in which plasmin
cleavage sites have been mutated to prevent the reactions, or the
catalytic site serine residue of Pg has been replaced to render the
plasmin formed inactive (Boose et al., 1989; Lijnen et
al., 1990; Davidson et al., 1990; Liu and Gurewich, 1992;
Fleury et al., 1993).
The present studies were undertaken to develop a new approach for investigating fibrinolytic reaction mechanisms, based on derivatives of Pg in which the catalytic site of the zymogen has been inactivated and labeled with an extrinsic fluorescence probe. To prepare these analogs, the active site generated non-proteolytically in Pg by SK binding is irreversibly inactivated by site-directed alkylation with a thioester peptide chloromethyl ketone (Bock, 1988, 1992a). The thiol group generated subsequently at the amino terminus of the covalently incorporated inhibitor provides a unique site for selective labeling with a variety of fluorescence probes (Bock, 1992b). Dissociation of the labeled Pg-SK complex allows purification of the fluorescent derivatives of Pg. The first Pg derivatives of this type, labeled with the fluorescence probe, 2-anilinonaphthalene-6-sulfonic acid, were prepared and characterized in the present studies. The labeled [Glu]Pg derivative was found to have properties similar to native Pg and to report changes in the catalytic site of the zymogen through changes in the probe fluorescence accompanying proteolytic activation by uPA and non-proteolytic activation by SK. The absence of potential proteolytic activity of the Pg analogs allowed SK-Pg interactions to be studied under equilibrium binding conditions for the first time. The approach developed in these studies is expected to enable further definition of binding interactions and elementary reaction steps of Pg activation which are coupled to conformational changes affecting the catalytic site of the zymogen.
Least-squares fitting was performed with the computer program SCIENTIST (MicroMath Software). All reported estimates of error represent ± 2 S.E.
The kinetics of the fluorescence changes
following addition of uPA to labeled Pg were measured under the same
conditions described above. Measurements were corrected for background
by use of a blank lacking the labeled protein, and for long reactions,
corrected for changes (10%) in fluorescence of an identical
reference mixture containing labeled Pg and 10 µM
FFR-CH
Cl that was incubated in parallel. The effect of
native Pg on SK binding to labeled Pg was measured from the
fluorescence changes recorded with time following addition of SK to
pre-equilibrated mixtures of [Glu]Pg
and
[AANS]FFR-[Glu]Pg
. Two traces from
identical reactions were averaged, and the amplitudes of the rapidly
established, initial fluorescence changes were obtained from the
blank-corrected data collected within the first 18-50 s of the
lag phase. The fluorescence amplitudes as a function of native Pg
concentration were analyzed as competitive binding of labeled and
native Pg to SK to obtain the dissociation constant for SK binding to
[Glu]Pg
by least-squares fitting of the cubic
binding equation described previously (Olson et al., 1991;
Lindahl et al., 1991). For this analysis, the dissociation
constant (590 ± 100 nM) and maximum fluorescence change
(2.8 ± 0.1) for SK binding to labeled Pg were fixed at the
values independently determined for the same protein preparations.
In
the second step of labeling, generation of the inhibitor thiol group in
ATA-FFR-(Pg-SK) with NHOH in the presence of 5-IAF resulted
in covalent incorporation of the probe selectively into the zymogen, as
shown by the SDS gel results in Fig. 1. SK or Pg treated
separately with 5-IAF were not significantly labeled (Fig. 1).
An additional experiment (not shown) in which Pg alone was subjected to
incubations with the inhibitor and probe demonstrated that the presence
of SK was required for labeling. Labeling was also prevented in other
control reactions in which NH
OH was omitted, or when the
catalytic site was blocked with FFR-CH
Cl prior to
incubation with ATA-FFR-CH
Cl and 5-IAF in the presence of
NH
OH (Fig. 1). These results indicated that
ATA-FFR-CH
Cl was covalently incorporated specifically into
the active catalytic site produced in Pg by SK binding, and that the
fluorescence probe was subsequently incorporated by selective
modification of the inhibitor thiol generated with NH
OH.
Figure 1:
Specificity of labeling
of Pg-SK complex with ATA-FFR-CHCl and 5-IAF. The
fluorescence (A) and protein-stained bands (B) on a
10% SDS gel are shown for reduced samples (5-15 µg) of Pg,
SK, and the products of labeling reactions performed as described under
``Experimental Procedures'' at final concentrations of 10
µM Pg or Pg-SK complex, 125 µM 5-IAF, and 0.1 M NH
OH where indicated, pH 7.0, and 25 °C. Lane 1, untreated Pg; lane 2, untreated SK; lane
3, Pg-SK complex inactivated with ATA-FFR-CH
Cl; lane 4, ATA-FFR-(Pg-SK) labeled with 5-IAF in the presence of
NH
OH; lane 5, Pg incubated with 5-IAF in the
presence of NH
OH; lane 6, ATA-FFR-(Pg-SK)
incubated with 5-IAF in the absence of NH
OH; lane
7, Pg-SK complex active site-blocked with FFR-CH
Cl
before incubation with ATA-FFR-CH
Cl and subsequently, with
5-IAF in the presence of NH
OH; lane 8, isolated
[5-AF]FFR-[Glu]Pg; lane 9,
[5-AF]FFR-[Glu]Pg after incubation at 9 µM with 0.1 µM uPA for 3 h at I = 0.15 M, pH 7.4, and 25 °C.
Figure 2:
Effect of cleavage by uPA and SK binding
on the fluorescence emission spectrum of
[AANS]FFR-[Glu]Pg. A,
fluorescence emission spectra are shown of 760 nM
[AANS]FFR-[Glu]Pg
(lower curve,
Pg) and the same concentration of labeled Pg after incubation for
1.5 h with 75 nM uPA (middle curve, +uPA) in I = 0.15 M buffer, pH 7.4, at 25 °C, or
after incubation with 5 µM SK for 50 min in the same
buffer containing 10 µM FFR-CH
Cl (upper
curve, +SK). Spectra were collected as described under
``Experimental Procedures.'' The inset shows results
of SDS-gel electrophoresis of a reduced sample (10 µg) of the
labeled plasminogen preparation visualized by fluorescence (right) and protein stain (left). B, SDS gels are shown for
samples from the incubations in A run under nonreducing
conditions (lanes 1-6) and reducing conditions (lanes 7-12). Samples contained 1.3 µg of Pg or
plasmin and 4.7 µg of SK, as follows: lanes 1 and 7, [AANS]FFR-[Glu]Pg
; lanes 2 and 8,
[AANS]FFR-[Glu]Pg
incubated with SK; lanes 3 and 9, SK; lanes 4 and 10,
[AANS]FFR-[Glu]Pg
incubated with uPA; lanes 5 and 11, native [Lys]Pm
; lanes 6 and 12, equal volume mixture of samples of
uPA-activated [AANS]FFR-[Glu]Pg
and
native [Lys]Pm
. The migration positions of
molecular weight markers are indicated on the right by their
molecular weights in thousands.
Figure 3:
Comparison of the time courses of the
activity and fluorescence increases accompanying cleavage of
[AANS]FFR-[Glu]Pg and native
[Glu]Pg
by uPA. Reactions of 750 nM native [Glu]Pg
or
[AANS]FFR-[Glu]Pg
in I = 0.15 M, pH 7.4, buffer were initiated at 25
°C by addition of 75 nM uPA. A, chromogenic
substrate activity measured on samples removed from the reaction
mixtures containing native Pg (
) or
[AANS]FFR-[Glu]Pg
(
) expressed as
a percent of the maximum measured for activation of native Pg. Inset, SDS gel of reduced samples (1.4 µg) removed from
the native Pg reaction mixture at the indicated times in minutes. Bands
corresponding to Pg, plasmin heavy chain (Pmh), and light
chain (Pml) are indicated. B, the increase in
fluorescence (
,
F/F
) with time
for the [AANS]FFR-[Glu]Pg
reaction. Inset, SDS gel of samples from the labeled Pg reaction, as
described in A. C, percent of maximum activity
(
) and fluorescence (
,
F/F
) increases in a reaction
containing 750 nM [Glu]Pg
and 75 nM [AANS]FFR]-[Glu]Pg
in the
above buffer plus 10 mM
-ACA, initiated with 25 nM uPA. Electrophoresis, activity, and fluorescence measurements were
performed as described under ``Experimental
Procedures.''
Figure 4:
Fluorescence titrations of
[AANS]FFR-[Glu]Pg with SK. The
fractional increase in fluorescence (
F/F
) of 0.21 µM
[AANS] FFR-[Glu]Pg
is plotted as a
function of the total SK concentration in the absence (
) and
presence of 10 mM (
) or 100 mM (
)
-ACA in pH 7.4 buffer at 25 °C. The lines represent
the nonlinear least-squares fits to the data with the parameters given
in the text. Titrations were performed and analyzed as described under
``Experimental Procedures.''
The possible role of
lysine binding sites in the interaction of SK with Pg was examined from
the effect of -ACA on SK binding. Analysis of fluorescence
titrations of [AANS]FFR-[Glu]Pg
in the
presence of 10 or 100 mM
-ACA showed a decrease in the
amplitude of the fluorescence change from a maximum of 3.1 ± 0.1
in the absence of
-ACA to 1.6 ± 0.1 at 10 mM and
1.8 ± 0.1 at 100 mM
-ACA. Corresponding
dissociation constants of 820 ± 160 and 1160 ± 150 nM were obtained, showing saturation of the effect with a modest,
2-fold decrease in affinity (Fig. 4).
Results of similar
experiments with [AANS]FFR-[Lys]Pg revealed significant differences in the binding of SK to labeled
[Glu]Pg
and [Lys]Pg
. In the
absence of
-ACA, binding of SK to
[AANS]FFR-[Lys]Pg
resulted in a lower
maximum fluorescence enhancement of 1.6 ± 0.1 compared to 3.1
± 0.1 seen with labeled [Glu]Pg
(Fig. 5). The dissociation constant for SK binding to
labeled [Lys]Pg
was 11 ± 7 nM,
showing a
54-fold higher affinity than for
[Glu]Pg
. Because of the higher affinity, analysis
of this data also allowed estimation of a stoichiometry for this
interaction of 1.0 ± 0.2 mol of SK/mol of Pg. In further
contrast to the results for [Glu]Pg
,
-ACA
greatly decreased the affinity of SK binding to
[AANS]FFR-[Lys]Pg
, while having little
effect on the amplitude of the fluorescence change. As shown by
analysis of the data in Fig. 5, the presence of 10 or 100 mM
-ACA resulted in SK binding to labeled
[Lys]Pg
with dissociation constants of 960
± 140 and 1080 ± 100 nM, respectively, and
maximum fluorescence changes of 1.9 ± 0.1. These results showed
saturation of the effect of
-ACA with an overall
98-fold
decrease in affinity of SK for [Lys]Pg
, yielding
a dissociation constant indistinguishable from that for
[Glu]Pg
. Results obtained with other preparations
of labeled [Glu]Pg and [Lys]Pg that were mixtures
of forms 1 and 2, or which contained significant levels of labeled
plasmin showed similarly large differences in affinity for SK and the
differential effect of
-ACA on the interactions.
Figure 5:
Fluorescence titrations of
[AANS]FFR-[Lys]Pg with SK. The
fractional increase in fluorescence (
F/F
) of 0.21 µM
[AANS] FFR-[Lys]Pg
is plotted as a
function of the total SK concentration in the absence (
) and
presence of 10 mM (
) or 100 mM (
)
-ACA in pH 7.4 buffer at 25 °C. The lines represent
the nonlinear least-squares fits to the data with the parameters given
in the text. Titrations were performed and analyzed as described under
``Experimental Procedures.''
Figure 6:
Effect of native [Glu]Pg on SK binding to [AANS]FFR-[Glu]Pg. The right panel shows the fractional increase in fluorescence (
F/F
) recorded with time for
reactions initiated by addition of 1.0 µM SK to mixtures
of 0.20 µM [AANS]FFR-[Glu]Pg
and 0, 0.20,
0.50, 1.1, 2.0, or 2.9 µM native [Glu]Pg
in pH 7.4 buffer at 25 °C. The amplitudes of the initial
fluorescence increases (
), indicated by the dashed
lines, are plotted in the left panel as a function of the
corresponding concentrations of [Glu]Pg
([Pg]). The solid line represents the
nonlinear least-squares fit of the equation for competitive binding to
the data with the parameters given in the text. Reactions were
performed and the data were analyzed as described under
``Experimental Procedures.''
These studies were undertaken to develop fluorescent derivatives of the plasminogen zymogen of a new type, to be used as probes for investigation of fibrinolysis mechanisms. The feasibility of the labeling strategy for preparing a family of probe-labeled derivatives of the zymogen was demonstrated in the preparation and characterization of the [AANS]FFR-Pg derivatives. These derivatives were found to have properties functionally analogous to native Pg, and to report conformational changes associated with activation of the catalytic site resulting from specific proteolytic cleavage by uPA, or induced nonproteolytically by binding of SK. Irreversible inactivation of the catalytic site in the labeled Pg analogs allowed activation by uPA and equilibrium binding of SK to be studied without significant complications from plasmin-catalyzed feedback reactions.
Evaluation of the specificity of labeling of the Pg-SK complex and characterization of the isolated Pg derivatives supports the conclusion that the Pg analogs were specifically labeled at the active catalytic site induced in Pg by SK binding. The results indicated a high degree of specificity of the labeling reactions, similar to that seen with other proteinases by the same method (Bock, 1992a, 1992b). These results and the mechanism of peptide chloromethyl ketone inhibition support the idea that the inhibitor is attached to the imidazole group of the catalytic site histidine residue and the probe is attached to the amino-terminal thiol of the tripeptide inhibitor (Powers and Harper, 1986; Bock, 1992a). The isolation and characterization of the affinity-labeled active species generated by SK provided direct evidence that the active catalytic site in the Pg-SK complex is the same as the proteinase active site of plasmin, in agreement with the conclusions of previous studies (Schick and Castellino, 1974). Results of labeling experiments with the carbohydrate variants of [Glu]Pg and [Lys]Pg indicate that all of these forms can be similarly labeled with comparable specificity.
[AANS]FFR-[Glu]Pg was compared to native [Glu]Pg
as a uPA
substrate to assess its properties as an analog of the native zymogen.
To our knowledge, no catalytic site-labeled serine proteinase zymogen
derivatives have been previously described. Because these derivatives
contain the tripeptide affinity label covalently bound at the catalytic
site, it was of interest to determine whether they exhibited properties
analogous to the native zymogen or to the activated proteinase.
Activation of native [Glu]Pg by uPA occurs by cleavage of the
Arg
-Val
bond in Pg, transiently generating
[Glu]plasmin, with the final product, [Lys]plasmin,
being formed by additional plasmin cleavage of [Glu]Pg and
[Glu]plasmin (Violand and Castellino, 1976). Cleavage of
[AANS]FFR-[Glu]Pg
by uPA produced
[AANS]FFR-[Glu]Pm
as the major initial
product, correlated with a large enhancement in the probe fluorescence
that signaled activation of the catalytic site. Comparison of the rates
of the uPA activation reactions for native and labeled Pg showed that
cleavage of [AANS]FFR-[Glu]Pg
was
slower than generation of [Lys]Pm
from native Pg.
These results were consistent with the established role of
plasmin-catalyzed feedback reactions in the process (Violand and
Castellino, 1976; Christensen, 1977; Violand et al., 1978;
Lucas et al., 1983) and the low level (1%) of active plasmin
formed from the labeled zymogen. The absence of active plasmin
formation, however, prevented a direct assessment of the properties of
the labeled Pg as a uPA substrate analog. To address this question, uPA
activation of native and labeled Pg were compared in a mixture of the
zymogens and in the presence of
-ACA. Under these conditions, the
rates of activation of [AANS]FFR-[Glu]Pg
and native [Glu]Pg
and the resulting time
courses of their conversion to the [Lys]Pm
forms
were indistinguishable. The observation that labeled Pg and the native
zymogen behave similarly in the overall activation process, under
conditions where the rate is dependent on interactions of
-ACA and
plasmin-catalyzed reactions, in addition to cleavage by uPA, indicates
that the labeled derivatives can be employed as reporting substrate
analogs of plasminogen activation.
The fluorescent Pg analogs
allowed quantitative equilibrium binding studies of the SK-Pg
interaction to be done for the first time. The mechanism of Pg
activation by SK involves SK-induced conformational activation of the
Pg catalytic site, closely coupled with irreversible proteolytic
conversion of Pg to plasmin (McClintock and Bell, 1971; Reddy and
Markus, 1972; Schick and Castellino, 1974; Bajaj and Castellino; 1977;
Gonzalez-Gronow et al., 1978; Davidson et al., 1990).
Inactivation of the catalytic site in the Pg analogs allowed the
binding interactions to be studied without the complications caused by
subsequent proteolysis. The results of these studies provided two types
of information: the affinities of binding and the amplitudes of the
fluorescence changes reporting perturbations of the catalytic
site-bound probe. Comparison of SK binding to labeled
[Glu]Pg and [Lys]Pg
revealed a
54-fold higher affinity for
[Lys]Pg
. Although the affinities of these
interactions have not been previously quantitated, the dissociation
constants of 590 ± 110 nM obtained for
[Glu]Pg
and 11 ± 7 nM for
[Lys]Pg
represented lower affinities than were
anticipated on the basis of previous studies. High affinity binding was
previously inferred mainly from: (i) active-site titrations of Pg
activation under experimental conditions where estimation of affinity
below the micromolar range would have been uncertain (McClintock and
Bell, 1971; Reddy and Markus, 1972), and (ii) indirectly from the
kinetic behavior of mixtures of nanomolar concentrations of SK and Pg
(Wohl et al., 1980, 1983; Chibber and Castellino, 1986;
Davidson et al., 1990). It should be noted, however, that the
affinity determined for labeled [Lys]Pg
in the
present studies may be underestimated because the relatively low
fluorescence yield of the probe limited the titrations to a
concentration range in excess of the dissociation constant. The
properties of the labeled derivatives as Pg analogs and reporters of SK
binding were evaluated further by quantitatively characterizing the
interaction between native Pg and SK using
[AANS]FFR-[Glu]Pg
as a binding probe.
The resulting estimate of 90 ± 60 nM for the
dissociation constant for SK binding to native [Glu]Pg
indicated that the presence of the label in the catalytic site
resulted in a
6-fold reduced affinity of SK for the Pg derivative.
Such an effect is not surprising because it is thought to reflect the
influence of occupation of the catalytic site by the probe-tripeptide
label on the thermodynamically linked binding and conformational change
which result in Pg activation and accompanying formation of the
substrate binding subsites. On this basis, the affinity of SK may also
be expected to vary for other active-site-liganded Pg species,
depending on the structures of the bound fluorescent labels,
active-site-titrants, or substrates employed to report the interaction.
In this respect, it is interesting that the presence of the label
decreased rather than increased SK affinity and thus did not apparently
stabilize a more active enzyme-like conformation of the labeled
zymogen. These observations support the conclusion that the active-site
labeled Pg derivatives should be considered analogs of the native
zymogen, which can be expected to exhibit functionally similar but not
quantitatively identical properties. The magnitude of the effect of the
presence of the label on the properties of the derivative studied here
indicates that this will not compromise the use of the fluorescent Pg
analogs as probes of Pg interactions and activation mechanisms.
Examination of the influence of -ACA on the affinity of SK for
the fluorescent Pg derivatives revealed additional differences between
[Glu]Pg
and [Lys]Pg
. The
affinity of SK for [Glu]Pg
was reduced only
2-fold by saturating levels of
-ACA, while the higher
affinity for [Lys]Pg
was reduced
98-fold.
This differential effect resulted in indistinguishable affinities of
the Pg species for SK at saturating concentrations of
-ACA. These
effects are presumably due to interactions of
-ACA with lysine
binding sites on Pg, because there is presently no evidence for such
sites on SK. It has been previously shown that [Glu]Pg is in
a compact conformation, which is shifted to an extended form by
-ACA binding (Violand et al., 1978; Mangel et
al., 1990; Ponting et al., 1992b; Christensen and
Molgaard, 1992; Marshall et al., 1994). [Lys]Pg is
in a more extended conformation than that of [Glu]Pg in the
absence of
-ACA, which is shifted to a fully extended form similar
to that of [Glu]Pg in the presence of
-ACA (Ramakrishnan et al., 1991; Marshall et al., 1994). These and other
studies have demonstrated differences between
-ACA binding to
[Glu]Pg and [Lys]Pg (Markus et al., 1978;
Christensen, 1984). It has been proposed that the compact conformation
of [Glu]Pg is stabilized by an intramolecular interaction
between the amino-terminal 77-residue sequence and lysine binding sites
in kringles 4 and 5 (Wiman and Wallen, 1975; Christensen, 1984;
Ramakrishnan et al., 1991; Christensen and Molgaard, 1992;
Marshall et al., 1994). This interaction is lost when the
peptide is removed in [Lys]Pg, exposing these lysine binding
sites. Thus, the high affinity of SK for [Lys]Pg and its
dependence on
-ACA could reflect preferential binding of SK to the
partially extended conformation of [Lys]Pg and/or interaction
of SK with the lysine binding sites exposed in kringles 4 and/or 5. In
the latter case, the lower affinity of SK for [Glu]Pg may be
due to competition with the amino-terminal peptide for these sites.
Saturation of these sites by
-ACA would be expected to lower
selectively the affinity of SK for [Lys]Pg to that of
[Glu]Pg, as is observed. Additional studies will be required
to determine whether this is the basis for the differences in SK
affinity.
The large enhancements in the fluorescence of the
catalytic site-bound probe in [AANS]FFR-Pg that accompanied
SK binding are thought to report conformational changes associated with
SK-induced activation of the catalytic site and formation of the
substrate-binding specificity subsites. The changes in the fluorescence
spectral properties of the active-site-bound probe were similar but not
the same as those accompanying proteolytic activation by uPA.
Fluorescence enhancements of 0.5-1.0 have been observed for SK
binding to labeled plasmin, suggesting that the difference in the
amplitudes of the fluorescence changes may be accounted for by an
additional perturbation of the environment of the activated catalytic
site due to SK binding. The larger difference observed in the magnitude
of the fluorescence changes accompanying SK binding to
[Glu]Pg and [Lys]Pg
suggests that there are also significant differences in the
environments of the catalytic sites in these complexes. These
differences were modulated by
-ACA, as shown by the reduction of
the amplitude of the fluorescence change for [Glu]Pg
while that for [Lys]Pg
was not similarly
affected. Thus, although further studies will be needed to determine
precisely the sources of these effects, it appears that occupation of
lysine-binding sites by
-ACA, Pg conformational equilibria, as
well as direct effects of SK binding are linked to perturbations in the
environment of the Pg catalytic site in complexes with SK that may be
expected to have functional significance.
In summary, derivatives of Pg that are specifically labeled with fluorescence probes attached to the catalytic site via a tripeptide chloromethyl ketone have properties analogous to those of native Pg. The fluorescent Pg analogs provide unique opportunities to observe events at the catalytic site of the zymogen that accompany its activation. Results obtained with the [AANS]FFR-Pg analogs demonstrate the utility of the derivatives for investigation of the mechanisms of conformational and proteolytic Pg activation, under conditions where principal activation reaction steps can be separated from concurrent plasmin-catalyzed reactions. Although the present studies have focused on the Pg derivatives prepared with one fluorescence probe, the capabilities of the labeling approach indicate that this can be extended to include a family of Pg derivatives labeled with different fluorescence probes, as well as other types of labels.