Serpins are thought to inhibit proteinases by
first forming a Michaelis-type complex that later converts into a
stable inhibitory species. However, there is only circumstantial
evidence for such a two-step reaction pathway. Here we directly observe
the sequential appearance of two complexes by measuring the
time-dependent change in fluorescence resonance energy
transfer between fluorescein-elastase and
rhodamine-
1-protease inhibitor. A moderately tight
initial Michaelis-type complex EI1
(Ki = 0.38-0.52 µM) forms and
dissociates rapidly (k1 = 1.5 × 106 M
1 s
1,
k
1 = 0.58 s
1).
EI1 then slowly converts into
EI2 (k2 = 0.13 s
1), the fluorescence intensity of which is stable for at
least 50 s. The two species differ by their donor-acceptor energy
transfer efficiency (0.41 and 0.26, respectively).
EI2 might be the final product of the elastase + inhibitor association because its transfer efficiency is the same as
that of a complex incubated for 30 min. The timedependent
change in fluorescence resonance energy transfer between
fluorescein-elastase and rhodamine-eglin c, a canonical inhibitor,
again allows the fast formation of a complex to be observed. However,
this complex does not undergo any fluorescently detectable
transformation.
 |
INTRODUCTION |
Proteolysis, a biologically important event, is regulated by
protein proteinase inhibitors that belong to two major classes: the
so-called "canonical" inhibitors and the serine proteinase inhibitors, called "serpins." The former are relatively small proteins (29-190 amino acid residues) that belong to numerous structural families (1). The latter are larger proteins (400-450 residues) that form a single family with highly conserved secondary structural elements (nine
-helices and three
-sheets) (2, 3).
Canonical inhibitors form tight reversible complexes with their cognate
enzymes. X-ray crystallography shows that they have an exposed peptidic
sequence, the reactive site loop, that forms a "lock and key"
complex with the substrate-binding crevice of the proteinase.
This Michaelian-like complex is stabilized by a large number of
noncovalent bonds which account for the high enzyme-inhibitor binding
energy. Its hydrolysis is probably prevented by a small distortion of
the P1-P1' linkage of the inhibitor (1).
Unlike canonical inhibitors, serpins form denaturant-stable complexes
with their target proteinases and behave kinetically like irreversible
inhibitors (3, 4). Their mechanism of action is not well understood
mainly because the tertiary structure of their complexes with
proteinases has not yet been solved. A significant number of x-ray
structures of active and inactive serpins are, however, available
(5-11). These structures reveal that the reactive site loop of serpins
is much longer and much more flexible than that of canonical
inhibitors. It may easily insert into
-sheet A to form the central
strand of this unusually malleable (12) sheet. Full insertion of the
P1 to P14 sequence of the reactive site loop
may occur spontaneously (9) or following cleavage of the
P1-P1' bond (5). Serpins that have undergone such a
-sheet rearrangement are inactive. On the other hand, introduction into
-sheet A of a tetradecapeptide structurally related to P1-P14 inactivates serpins because
it renders them unable to insert their own
P1-P14 sequence (13). Thus, full loop-sheet
insertion and lack of insertion both yield inactive serpins. This led
to the suggestion that loop-sheet insertion modulates the inhibitory
activity of serpins (14) and to the proposal of a number of models for
the serpin-proteinase interaction (15-18). All these models assume
that the inhibition reaction takes place in at least two steps: an
initial binding followed by a structural rearrangement that stabilizes
the complex. However, these steps have not been observed directly and
it is not known at which rate the proteinase moves from its initial to
its final position.
Just as chemical kinetics helps elucidating reaction mechanisms (19),
inhibition kinetics affords insight into inhibition mechanisms. For
instance, several years before publication of the first
three-dimensional structure of a canonical inhibitor-proteinase complex, an inhibition mechanism for a canonical inhibitor has been
proposed on the basis of a kinetic analysis (20). Most kinetic studies
on serpins have reported measurements of kass, the second-order inhibition rate constant, which is useful for delineating inhibitor efficiency and physiological function (21) but of
poor mechanistic significance because it is a combination of rate
constants for the individual reaction steps (22). On the other hand,
two-step serpin-proteinase interactions
have been demonstrated by a limited number of investigators. The
measurements were based on competition experiments in which the serpins
displaced a substrate or another ligand from the active center of the
proteinase (23-26). The time course of the reaction was thus followed
indirectly. In addition, these experimental approaches did not provide
access to k
1 and k1 but
only to Ki, the ratio of these two constants.
We hypothesized that labeling of one reaction partner with a
fluorescence donor and of the other with a fluorescence acceptor and
measuring the time dependence of the fluorescence resonance energy
transfer with a fast kinetic apparatus would allow us to directly
observe the individual reaction species as well as the dynamics of
their interconversion. The present work checks this hypothesis by
investigating the interaction of elastase with
1PI, a serpin, and comparing it with binding of elastase with eglin c, a
canonical inhibitor.
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MATERIALS AND METHODS |
Porcine pancreatic elastase
(elastase)1 was isolated and
active site titrated as described previously (27). Recombinant
1PI and eglin c expressed in Escherichia coli
were obtained from Novartis (Basel) and were titrated with human
neutrophil elastase (27, 28).
Fluorescent Labeling of
1PI, Eglin c, and
Elastase--
1PI was specifically labeled on Cys-232,
its single free cysteinyl residue using
tetramethylrhodamine-5-maleimide (Molecular Probes). A 140 µM solution of
1PI was made up in 50 mM TES, 150 mM NaCl, pH 7.0, and reacted with a
1.5 molar excess of the labeling reagent dissolved in
dimethylsulfoxide. After 1 h at room temperature, excess reagent
was removed by two gel filtrations on a PD-10 column (Amersham
Pharmacia Biotech) equilibrated with 50 mM Hepes, 150 mM NaCl, pH 7.4. Active site titration with human
neutrophil elastase showed that
1PI was fully active
after labeling. Yields varied from 0.7 to 0.9 label/molecule of
1PI as determined spectrophotometrically using
555 nm = 75,000 M
1
cm
1 for protein-bound label. Prior reaction of
1PI with
4-N,N-dimethylaminoazobenzene-4'-iodoacetamido-2'-sulfonic acid (Protein Institute, Philadelphia, PA), a specific reagent of free
thiols (29), yielded a protein that did not further bind
tetramethylrhodamine-5-maleimide, indicating that the latter specifically labels Cys-232.
Eglin c (1.2 mM) was dissolved in the above TES buffer and
reacted with a 3-fold molar excess of the succinimidyl derivative of
tetramethylrhodamine dissolved in dimethylsulfoxide. After 1 h at
room temperature, excess reagent was removed as described above. Active
site titration with human neutrophil elastase showed that the labeled
inhibitor retained full activity. There was 0.96 molecule of
label/molecule of eglin c as determined spectrophotometrically using
the above
555 nm.
Elastase (0.8 mM) was dissolved in 0.1 M
carbonate buffer, pH 9.0, and reacted with a 20-fold molar excess of
FITC (Molecular Probes) dissolved in methanol. After 1 h at room
temperature, excess reagent was removed by gel filtration on two PD-10
columns equilibrated and developed successively with the above
carbonate buffer and with 1 mM HCl. Active site titration
of FITC·elastase with nonlabeled
1PI showed that the
enzyme did not loose activity during labeling. There was 1.1 label/molecule of elastase as determined spectrophotometrically using
495 nm = 66,800 M
1
cm
1 for protein-bound FITC (30). A higher degree of
labeling could not be achieved.
Absorption and Fluorescence Measurements--
Absorption and
emission spectra were recorded on a Cary 4 spectrophotometer and a
SLM-8000 spectrofluorometer, respectively. Fluorescence quantum yields
of free and
1PI-bound FITC elastase were measured using
acryflavin as a reference (
= 0.45). Emission spectra were corrected
for screening effects at both excitation and emission wavelengths
(31).
Kinetics of interaction of FITC·elastase with
TMR·
1PI or TMR·eglin c was monitored by fluorescence
resonance energy transfer from FITC to TMR using a Bio-Logic SFM-3
stopped flow apparatus with a dead time of 1.7 ms (Bio-Logic, Claix,
France). The reaction was done in 100 mM Hepes, 150 mM NaCl, pH 7.4, 25 °C. The excitation and emission
wavelengths were 450 and 514 nm (Melles-Griot interferential filter),
respectively.
 |
RESULTS |
Steady-state Fluorescence of Free and Bound FITC·Elastase and
TMR·
1PI--
Fig. 1
shows the fluorescence spectra of the free and bound species. It can be
seen that reaction of FITC·elastase with TMR·
1PI significantly quenches the emission of FITC and enhances that of TMR,
strongly suggesting fluorescence resonance energy transfer between the
donor and the acceptor. To see whether part of the fluorescence
intensity variation is due to a change in the environment of the
label(s) following complex formation, we have recorded the spectrum of
the complex formed of FITC·elastase and unlabeled
1PI
as well as that formed of TMR·
1PI and unlabeled
elastase (Fig. 1). In both curves, the binding of the unlabeled protein only marginally affected the fluorescence of the labeled one. This
clearly indicates that all of the fluorescence intensity variation is
associated with a nonradiative energy transfer process between FITC and
TMR.

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Fig. 1.
Fluorescence resonance energy transfer
between FITC·elastase and TMR· 1PI. The
excitation wavelength was 450 nm. Spectra a-c correspond to
2.6 µM FITC·elastase in the absence (a) or
presence of 5.2 µM unlabeled 1PI
(b) or TMR· 1PI (c). Normalizing
spectrum a on spectrum c at 515 nm where
FITC·elastase is the only emitting species yields spectrum
d, which represents the contribution of FITC·elastase to
spectrum c. Likewise, subtraction of d from
c yields spectrum e, which represents the
contribution of TMR· 1PI. Spectra f and
g represent the fluorescence of 5.2 µM
TMR· 1PI in the absence or presence of 2.6 µM unlabeled elastase, respectively. Proteins were
dissolved in 50 mM Hepes, 150 mM NaCl, pH 7.4. Elastase and 1PI were reacted for 30 min before
recording the spectra.
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Kinetics of the Interaction of FITC·Elastase with
TMR·
1PI--
Fig. 2
shows a typical biphasic stopped flow trace observed upon mixing
FITC·elastase with a 10-fold molar excess of
TMR·
1PI. The fluorescence intensity rapidly decreases
(t1/2
60 ms), falls to a minimum and then slowly
recovers (t1/2
5 s) without reaching its
initial value. We have tentatively assumed that the initial fluorescence quenching describes the formation of an initial complex EI1, which slowly converts into a second complex
EI2 (Fig. 2). The fluorescence intensity was
stable for at least 50 s (i.e. 10 t1/2 of EI2 formation)
whether the reaction was run under pseudo-first-order conditions
([I]o = 10 [E]o = 7 µM) or
under second-order-conditions ([E]o = [I]o = 7 µM). This indicates that within this interval of
time, EI2 neither releases free FITC·elastase,
which would enhance the fluorescence intensity, nor converts into a
further complex with a different donor-acceptor transfer efficiency.
The fluorescence intensity could not be recorded beyond 50 s
because of photodecomposition of the label(s).

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Fig. 2.
Kinetics of the fluorescence resonance energy
transfer between FITC·elastase and TMR· 1PI at pH 7.4 and 25 °C. The excitation and emission wavelengths were 450 and
514 nm, respectively. The elastase and 1PI
concentrations were 0.7 and 7 µM, respectively.
Curve a, FITC·elastase + unlabeled 1PI;
curve b, FITC·elastase + TMR· 1PI. The
data-recording frequency of the stopped flow apparatus was
103 s 1 for observation times 1 s
and 10 s 1 for observation times > 1 s, which
explains the apparent heterogeneity in the noise level.
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A control where FITC·elastase was mixed with unlabeled
1PI showed a fluorescence decay of about 4% over the
observation time of 50 s. This small decrease was considered to be
negligible and was not taken into account for further calculations.
Data analysis showed that the formation of the two complexes could be
described by simple exponentials: the pseudo-first-order rate constant
of EI1 formation (kobs)
was 11.8 ± 0.6 s
1 for [I]o = 10 [E]o = 7 µM, whereas the first-order rate
constant for the conversion of E11 into
EI2 (k2) was 0.130 ± 0.004 s
1.
To see whether the EI1 complex is reversible or
irreversible, we have recorded stopped flow traces using constant
concentrations of FITC·elastase and variable concentrations of
TMR·
1PI. We have found that the minimum fluorescence
intensity (Fig. 2), a measure of the concentration of
EI1, varies hyperbolically with the inhibitor concentration (Fig. 3). This rules out
irreversible binding because reaction of constant concentrations of
enzyme with increasing concentrations of inhibitor should lead to
constant concentrations of complex if binding were irreversible. The
data of Fig. 3 were thus analyzed assuming that FITC·elastase (E) and
TMR·
1PI (I) form a reversible complex
EI1 whose equilibrium dissociation constant Ki may be calculated by fitting the
fluorescence data to Equation 1.
|
(Eq. 1)
|
where
F is the difference between the minimum
fluorescence intensity of EI1 and the
fluorescence intensity at t = 0 while
Fmax is the asymptotic value of
F for infinite concentrations of inhibitor. Nonlinear
regression analysis yielded Ki = 0.52 ± 0.08 µM. The curve calculated using this value fairly well
fits the experimental points (Fig. 3).

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Fig. 3.
Determination of the equilibrium dissociation
constant Ki of the initial complex formed between
FITC·elastase and TMR· 1PI, i.e. the
EI1 complex shown in Fig. 2. Constant
concentrations of elastase (0.7 µM) were mixed with
increasing concentrations of 1PI and the minimal
fluorescence intensities (see Fig. 2) were measured. F
and Fmax are defined in the text. The
curve is theoretical and has been constructed using Equation 1 and the best estimate of Ki (0.52 µM).
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The above equilibrium may also be described by the rate constant of
complex formation and dissociation by the following interaction.
which may also be written as
if pseudo-first-order conditions prevail. Hence,
kobs, the pseudo-first-order rate constant for
the EI1 formation is given by
|
(Eq. 2)
|
To determine k1 and
k
1 we have measured
kobs under pseudo-first-order conditions. Fig.
4 shows that kobs
is linearly related to [I]o as predicted by Equation 2.
The individual rate constants were calculated by linear regression
analysis k1 = (1.5 ± 0.02) × 106 M
1 s
1,
k
1 = 0.58 ± 0.08 s
1. The
equilibrium dissociation constant K1 of the
EI1 complex, calculated from
k1 and k
1 was 0.38 ± 0.06 µM. This value is in good agreement with that
measured under equilibrium conditions and therefore provides good
internal consistency to our data.

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Fig. 4.
Measurement of the rate constants for the
association and the dissociation of the initial complex
EI1 between FITC·elastase and
TMR· 1PI at pH 7.4 and 25 °C. The apparent
first-order rate constant of complex formation
kobs was determined by fluorescence energy
transfer (Fig. 2) using [TMR· 1PI] = 0.7-7
µM and [FITC·elastase] = 0.1 [TMR· 1PI].
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On the other hand, the fluorescence intensity corresponding to
EI2 did not change with the inhibitor
concentration. In addition, k2, the first-order
rate constant for the conversion of EI1 into EI2 was also inhibitor-independent (data not
shown). These two observations are interpreted to mean that
k2 is a true first-order rate constant,
i.e. that the transformation of EI1
into EI2 is an irreversible process. Our data
therefore provide a complete kinetic description of the minimum steps
of the interaction of elastase with
1PI:
Energy Transfer Efficiency within the
FITC·Elastase-TMR·
1PI Complexes--
The transfer
efficiency, ED, within the complex prepared by
manually mixing labeled elastase with labeled
1PI (see
legend to Fig. 1) was derived from steady-state fluorescence
measurements and calculated as follows:
|
(Eq. 3)
|
where
D,A and
D are the quantum
yields of the FITC·elastase-TMR·
1PI complex and the
FITC·elastase-
1PI complex, respectively.
D,A was corrected to take into account the degree of
labeling, f, of
1PI by TMR.
|
(Eq. 4)
|
Following corrections for screening effects,
ED was found to be 0.26 ± 0.02.
The transfer efficiencies, ED, within the
EI1 and EI2 complexes
detected by stopped flow (Fig. 2) were also calculated from the
fluorescence intensities at 515 nm, a wavelength at which FITC is the
only fluorescent chromophore:
|
(Eq. 5)
|
where ID,A and ID
are the fluorescence intensities of the
FITC·elastase-TMR·
1PI complex and the
FITC·elastase-
1PI complex, respectively.
ID,A was again corrected for the degree of
labeling. The screening effect was negligible in the stopped flow
experiments. For the EI2 complex,
ED was 0.26 ± 0.03, a value identical to that corresponding to the elastase-
1PI complex prepared
by manual mixing.
The value of ID,A used to calculate
ED for the EI1 complex
was taken from the minimum fluorescence intensity (Fig. 2) of stopped flow mixtures containing 0.35 µM FITC·elastase and 11.4 µM TMR·
1PI, which yield 97%
EI1 complex, i.e. in which the
fluorescence contribution of free FITC·elastase is negligible. After
correcting for the degree of labeling, ED was
found to be 0.41 ± 0.04. Possible random labeling of the lysine
residues of elastase precluded interchromophore distance calculations
using the ED values.
Kinetics of the Interaction of FITC·Elastase with TMR·Eglin
c--
Eglin c is a reversible proteinase inhibitor that belongs to
the class of canonical inhibitors (1). Mixing FITC·elastase with a
10-fold molar excess of TMR·eglin c yields an exponential fluorescence quenching whose amplitude does not change during the 50-s
observation time (Fig. 5). This behavior
sharply contrasts with that of
1PI (Fig. 2). The
reversible binding of elastase with eglin c (22) therefore takes place
in only one fluorescently detectable step. We have measured the rate
constant kobs of fluorescence energy transfer as
a function of inhibitor concentration under pseudo-first-order
conditions ([I]o = 10 [E]o). Fig.
6 shows that kobs
increases linearly with the inhibitor concentration as already observed
with
1PI and described by Equation 2. The association rate constant k1 calculated from this plot was
found to be 1.8 × 106 M
1
s
1, in good agreement with the value of 106
M
1 s
1, measured by enzymatic
means (22). The k
1 value could not be
determined accurately because the eglin c concentrations used in these
experiments were at least 86-fold higher than the Ki
of the elastase-eglin c complex (Ki = 10
8 M; Ref. 22) so that
k1 [I]o
k-1 and kobs
k1 [I]o in Equation 2.

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Fig. 5.
Kinetics of the fluorescence resonance energy
transfer between FITC·elastase and TMR·eglin c at pH 7.4 and
25 °C. The elastase and eglin c concentrations were 0.7 and 7 µM, respectively. For all other conditions see Fig. 2.
Curve a, FITC·elastase + unlabeled eglin c; curve
b, FITC·elastase + TMR·eglin c.
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Fig. 6.
Effect of TMR·eglin c concentration on the
apparent first-order rate constant for the formation of the
FITC·elastase-TMR·eglin c complex at pH 7.4 and 25 °C. The
rate constant kobs was determined by
fluorescence energy transfer as shown in Fig. 5 using [TMR·eglin c] = 0.87-14 µM and [FITC·elastase] = 0.1 [TMR·eglin
c].
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 |
DISCUSSION |
Stopped flow recording of the changes in fluorescence resonance
energy transfer observed upon mixing FITC·elastase with
TMR·
1PI has enabled us to directly observe the
sequential appearance of two enzyme-inhibitor complexes with
significantly different donor-acceptor energy transfer efficiencies. It
is commonly assumed but rarely demonstrated that the first step of the
reaction of a proteinase with a serpin is the formation of a reversible
Michaelis-type complex involving the substrate-binding site of the
enzyme and the reactive site loop of the inhibitor. Serpin-induced
displacement of ligands from the active center of proteinases (23-26)
or dissociation of serpin-proteinase complexes by
2-macroglobulin (32, 33) have provided indirect evidence
for such a reversible binding step. In the present work an initial
complex referred to as EI1 forms in less than
1 s after mixing. Rate and equilibrium measurements of the
building up of this species clearly demonstrate that it is a fast
equilibrating reversible complex. We believe this is the first direct
evidence for a Michaelis-type complex between a proteinase and a
serpin. On the other hand, the conversion of EI1
into EI2 is a true first-order process because
its rate and amplitude do not depend upon the inhibitor concentration.
This indicates that the formation of EI2 and
hence the overall elastase-
1PI interaction is an
irreversible process, a view supported by enzymatic data showing that
1PI and other serpins behave like irreversible proteinase inhibitors (4). The two-step model presented here is
probably a minimal one because fluorescence detects only those species
that significantly differ in energy transfer efficiency, whereas it is
not unlikely that undetectable reaction intermediates also appear
during the elastase +
1PI reaction.
Molecular modeling studies (15, 16, 18) and biochemical investigations
(17, 34) indicate that in the course of its reaction with a serpin, the
proteinase moves more or less far away from its initial position,
whereas the serpin's reactive site loop inserts more or less deeply
into
-sheet A. It has also been demonstrated that proteinases cleave
the P1-P'1 bond of serpins and form acyl-enzyme
intermediates between the serine residue of their catalytic site and
the P1 residue of the inhibitors. This cleavage is believed
to trigger the above conformational changes and, as a consequence, to
stabilize the final complex by preventing the hydrolysis of the
acyl-enzyme (35, 36). These findings may help discuss the nature of the
EI2 complex and the significance of
k2, the rate constant of its formation. The
stopped flow detected complex is probably structurally identical to the
stable complex prepared by manual mixing and incubated for 30 min
because the two species have identical donor-acceptor transfer
efficiencies. As a consequence, EI2 is likely to
be identical with the elastase-
1PI complex, which
Lawrence et al. (35) and Wilczynska et al. (36)
have shown to be a translocated acyl-enzyme. Although
k2 describes the first-order fluorescence
increase that accompanies the conversion of EI1
into EI2, it is not necessarily the rate
constant for the translocation of elastase. The enzyme's acylation and
translocation are separate events characterized by two individual rate
constants, kacylation and
ktranslocation. Because the
EI1 to EI2 conversion is
a simple exponential process, k2 might have
either one of the following two meanings: (i) k2
kacylation, i.e.
kacylation
ktranslocation
or (ii) k2
ktranslocation, i.e.
kacylation
ktranslocation. The second assumption implies that the acyl-enzyme would be exposed to water during the long lasting
translocation process (t1/2 = 5.3 s). Because
enzyme translocation is required to prevent hydrolysis of the
acyl-enzyme, active enzyme would be released during the translocation
process. This is not the case because the 1:1
elastase·
1PI complex is enzymatically inactive (4). We
may, therefore, tentatively conclude that k2 is
the rate constant for the acylation of elastase. We are not unaware
that the above reasoning is somewhat speculative. For instance, despite
its having an energy transfer efficiency identical to that of the
stable complex prepared by manual mixing, EI2
might not be the final inhibitory complex because it might still
undergo fluorescently silent structural changes. On the other hand, the
acylation and the translocation rates might not be as different from
each other as we have hypothesized. Clearly, the only way to solve
these uncertainties would be to directly measure the rate of
acylation.
Like the serpin
1PI, the canonical inhibitor eglin c (1)
rapidly forms a Michaelis-type complex with elastase, but this complex
does not undergo the characteristic conversion seen with the
elastase-
1PI complex. This is in accord with
crystallographic data showing that the P1-P'1
bond of eglin c is not cleaved within the enzyme-inhibitor complex (37)
and that there are no gross structural differences between free and
proteinase-bound eglin c (38). Thus, kinetics of fluorescence resonance
energy transfer unambiguously discriminates the serpin
1PI from the canonical inhibitor eglin c.
Most extracellular serpins play a proteolysis-preventing function. The
data collected using our new approach may help to better delineate this
function. In vivo, a proteinase is usually released into a
milieu containing both serpin and substrate. Knowledge of
Ki and [I]o, the in vivo
inhibitor concentration, will allow prediction of whether the substrate
may effectively compete with the serpin for the binding of the
proteinase; competition is possible if
[I]o/Ki
[S]o/Km. Besides, the magnitude of
k
1 will predict the rate at which a preformed EI1 complex may be dissociated by substrate.
Last, the magnitude of k2 will predict the
apparent reversible or irreversible nature of the inhibition in
vivo. If k2 is fast, for example say
k2 > 10
2 s
1
(t1/2 < 1 min), the inhibition will be irreversible
within a few minutes. However, if k2 < 10
2 s
1, the inhibition may be reversible
for a prolonged time, a view that is generally overlooked.
We thank Dr. Hans-Peter Schnebli for the gift
of recombinant
1-proteinase inhibitor.