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INTRODUCTION |
Nitric oxide (NO)1 helps
mediate a large number of physiologic and pathophysiologic processes
(1-6). NO is generated by the NO synthases (NOSs), which oxidize
L-arginine (Arg) in a two-step process that generates
N
-hydroxy-L-arginine (NOHA) as an
intermediate (7-9). All NOSs are homodimers with each subunit being
composed of a reductase domain that contains FAD and FMN, and an
oxygenase domain that contains 6(R)-tetrahydrobiopterin
(H4B) and iron protoporphyrin IX (heme) (10-11).
Ca2+-dependent calmodulin (CaM) binding enables
NADPH-derived electrons to transfer from the flavins to the heme and
initiate NO synthesis (12-13).
Three main NOSs are expressed in mammals. Neuronal NOS (nNOS or NOS-1)
and endothelial NOS (eNOS or NOS-3) are constitutively expressed and
largely participate in signal cascades. Inducible NOS (iNOS or NOS-2)
is primarily regulated by transcriptional mechanisms and functions as a
regulator and effector of the immune response. The counterpart to this
diversity of localization and function is a specific regulation of each
isoform. In this regard, nNOS appears unique because it is the only
isoform for which a rapid (within seconds) and stable buildup of a
ferrous heme-NO complex has been observed (14, 15). This differs from
that reported for iNOS and eNOS (16-19), where partial buildup of a ferric heme-NO complex, if it occurs at all, takes minutes. Moreover, ferric heme-NO complex formation in eNOS and iNOS is directly linked to
the exogenous NO concentration and can be suppressed by NO scavengers
like oxyhemoglobin (18, 19), whereas ferrous heme-NO complex formation
in nNOS is independent of external NO (15, 21).
The kinetics of heme-NO complex formation in nNOS and its effect on
steady-state activity has been extensively studied (14, 15, 22, 23).
During steady-state catalysis, a majority of nNOS (up to 80%) is
present as its ferrous heme-NO complex. Buildup of this complex is
linked to a decrease in catalytic activity. Moreover, it induces a
significant increase in the apparent
Km,O2 of the enzyme (14),
which has important implications regarding function in cells or tissues
(24, 25). Different mechanisms have been suggested to explain this
phenomenon (14, 23). However, simulation of these early models (14, 15)
failed to reproduce precisely the pre-steady and steady-state behavior
of nNOS.
Here we describe a kinetic model for nNOS catalysis that is built from
rate constants derived from the literature or completed here. The model
incorporates our recent finding that a ferric heme-NO complex is
generated prior to release of free NO (26). This ferric heme-NO product
partitions between two pathways to regenerate ferric enzyme, but only
one of these pathways efficiently releases NO. The kinetic model is
shown to simulate correctly behaviors of wild-type nNOS or nNOS mutants
that display greater activity in the steady state (22, 23). The general
utility of the model in visualizing the effect of enzyme modifications and in understanding the different catalytic behaviors of the three NOS
isoforms is discussed.
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EXPERIMENTAL PROCEDURES |
Materials--
All reagents and materials were obtained from
Sigma or sources as previously reported (22, 25).
Protein Expression and Purification--
Wild-type nNOS protein
was expressed in Escherichia coli and purified as described
previously (23, 27). UV-visible spectra were recorded on a Hitachi
U3110 Spectrophotometer in the absence or presence of 20 µM H4B and 1 mM Arg. Enzyme
concentration was quantified using the absorption of the ferrous-CO
adduct at 444 nm and an extinction coefficient of 74 mM
1 cm
1
(A444-A500).
Citrulline Binding Assay--
nNOS was incubated in a solution
containing 40 mM EPPS, pH 7.6, 300 µM DTT,
10% glycerol, and 4 µM H4B plus different
L-citrulline concentrations ranging from 100 µM to 166 mM for 10 min at room temperature.
Spectra were recorded, and the absorption increase at 420 nM was monitored to follow citrulline binding. Citrulline release was monitored in a stopped-flow instrument equipped with a
rapid-scanning diode array detector (HI-Tech MG 6000). Solutions containing 2 µM nNOS in 40 mM EPPS, pH 7.6, 300 µM DTT, 10% glycerol, 4 µM
H4B, and 100 mM citrulline were rapidly mixed
with the same buffer containing 100 mM Arg. Two sets of 10 mixings were run. In each run 96 spectra were recorded within 144 or
288 ms. The decrease in absorption at 417 nm and increase at 387 nm
were fit to a mono-exponential curve using software provided by the
instrument manufacturer for the first data set. The second set was
simultaneously fitted to a mono-exponential curve using the multifit
program of Origin 5.0.
nNOS Activity Assays--
NADPH oxidation and NO formation at
steady-state were measured at room temperature by spectroscopic assays
as described previously (15).
Ferrous-Nitrosyl Complex Formation during NO Synthesis--
For
experiments done without an NADPH-regenerating system, a cuvette
containing 1 µM nNOS was incubated for 10 min at room temperature in an air-saturated solution containing 40 mM
EPPS, pH 7.6, 0.5 mM EDTA, 2 µM CaM, 2.5 mM Ca2+, 400 µM DTT, 20 µM H4B, and 1 mM Arg. NO
synthesis was triggered by NADPH addition (40 µM final
concentration) at 10 °C. In the presence of an NADPH-regenerating
system (9 units of glucose-6-phosphate dehydrogenase and 1 mM glucose 6-phosphate), NADPH was added prior to the
reaction, and NO synthesis was triggered by addition of excess
Ca2+. Spectra were recorded before, during, and after NO
synthesis was initiated. The same reactions were also done using a
stopped-flow instrument and monitored by a rapid-scanning diode array
detector. The buffer containing the enzyme was rapidly mixed at
10 °C with the same buffer containing the triggering agent (NADPH or
Ca2+). Absorbance change at 436, 395, and 340 nm were used
to follow the formation of ferrous heme-NO complex, the decay of ferric enzyme, and the oxidation of NADPH, respectively. Spectra were recorded
in a time scale ranging from 144 ms to 7.22 s. A set of eight
representative experiments (with and without the NADPH-regenerating system) was simultaneously fitted using the multifit option of Origin
5.0. The buildup of Fe(II) heme-NO complex and the decay of Fe(III)
enzyme were fit to a double exponential curve. NADPH oxidation rates
were fit to a linear function in both a 0-500-ms and 2-8-s range.
Model Simulations--
The simulation was based on rate
equations derived from the kinetic model in
Scheme 1. The equations are defined under
the "Appendix." See text for details about the rate constants and
their dimension. For simulation the rate equations were treated by
simultaneous iterative calculation. The iteration step was typically 1 ms and up to 100,000 points were calculated. Percentages of each
intermediate were determined at steady state. Initial rates of NADPH
oxidation and citrulline production were calculated between 0 and
0.1 s. The same iterative treatment was applied for the
abbreviated kinetic model in the inset of Fig. 3.

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Scheme 1.
Kinetic model of nNOS catalysis.
Catalysis is approximated by 10 steps. Each number refers to
rate constants displayed in Table I. Single arrows indicate
that each step was assumed to be irreversible. The values of rate
constants for wild-type or mutant nNOS are detailed in the legends of
Tables I and II. See text for details.
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RESULTS |
General Considerations of the Model--
We chose the simplified
sequence "Fe(III) to Fe(II) to Fe(II)O2 to Product" to
describe heme transitions in both steps or NO synthesis (Arg
hydroxylation to NOHA, NOHA oxidation to NO plus citrulline). As
recently shown with the nNOS oxygenase domain in single turnover
experiments by Boggs et al. (26), the Fe(III) heme-NO
complex forms as the first observable product of NOHA oxidation, with
subsequent NO release from the enzyme occurring through its
dissociation from the heme. A comparable buildup of Fe(II) heme-NO
complex has been observed by Abu-Soud et al. (15) within the
first second of initiating a multiple turnover reaction in nNOS. In
both cases, heme-NO complex buildup is not linked to enzyme binding
solution NO because there is no effect of added NO scavengers. It also
differs from NO inhibition as originally described by Ignarro and
colleagues (20), which occurs in nNOS only after a high number of
catalytic turnovers (15 min at 37 °C) and at high concentrations of
exogenous NO (complete inhibition required 100 µM of NO).
Rather, heme-NO complex formation during the initial phase of NO
synthesis is probably linked to a geminate-type combination of Fe(III)
heme and NO generated in the active site. This process was proposed by
Kominami et al. (28), who found through EPR experiments that
NO rebinds quickly to nNOS after photo-dissociation from the ferric
heme even at 5 K. A complete study of NO geminate recombination in eNOS
has been achieved by Negrerie et al. (29); geminate
rebinding of NO to heme after photo-dissociation consists in fact of
both picosecond and nanosecond phases. This phenomenon was also
described by Scheele and colleagues (30) through photo-dissociation
experiments with nNOS. Their kinetic characterization showed a low
dissociation efficiency of NO after photolysis, suggesting its escape
from the heme pocket is relatively slow compared with direct rebinding
(30). The kon derived from their experiments
(1.2 × 107 M
1
s
1 at 23 °C) is close to the values
observed by Abu-Soud et al. (31) and Huang et al.
(32) when iNOS or nNOS was mixed with NO in a stopped-flow instrument
(kon around 106
M
1 s
1
at 10 °C). These values are consistent with a slow step being NO
entry into the heme pocket and a subsequent very fast binding (30).
This was confirmed by the absence of effects on rates of NO binding to
ferric or ferrous NOS by bound substrate or H4B, as is also
the case with O2 binding (27). Although it has not been
directly observed, most current mechanisms for NO synthesis predict
that the NOS heme will end up in a low spin, six-coordinate aqua form
immediately after generating NO from NOHA. Thus, NO binding has to
occur after water dissociation. During the single turnover stopped-flow
experiments of Boggs et al. (26), this water-bound
intermediate is not seen. Instead, there is a quantitative buildup of
the ferric-NO complex. Therefore, it appears that the speed of water
release and NO binding is such that essentially all of the NO molecules
bind before they leave the heme pocket. Thus, a nearly instantaneous
and quantitative binding of NO probably occurs in all NOS to generate a
Fe(III) heme-NO complex as the first observable product.
After forming the ferric heme-NO complex, NO can be released at a
relatively high dissociation rate (the koff for
NO from ferric nNOS varies between 3 and 5 s
1 depending
on the reports) (22, 26, 30). This represents a "productive"
pathway to release NO and regenerate ferric enzyme for a second
catalytic cycle. Nevertheless, during the steady-state nNOS exists
mainly as a ferrous heme-NO complex (14, 15). This was confirmed
recently by Adak et al. (22) and by experiments we will
report here. This implies that reduction of the heme-NO complex occurs,
which is likely because the rate of ferric heme reduction in nNOS (3-4
s
1 at 10 °C) is within the same range as
NO dissociation from the ferric heme. In contrast, NO dissociation from
the ferrous heme-NO complex is quite slow (29, 30), and ferric enzyme
is mainly regenerated by direct reaction with O2 (14).
However, because NO is not released in this reaction (the product is
nitrate), the ferrous heme-NO species represents an intermediate in a
"futile" pathway that regenerates ferric enzyme for the next
catalytic cycle. Thus, NO binding to the ferric heme results in a
circumstance where the enzyme can partition between NO dissociation
versus reduction of the heme-NO complex.
This concept allows us to propose a simplified kinetic model for nNOS
catalysis (Scheme 1). Our model contains 10 steps and is characterized
by a forward pathway that leads to the production of citrulline and NO
(steps 1-6) and two regenerating pathways as follows: the first
involves direct release of NO from ferric heme (step F), and the second
involves reduction of the ferric heme-NO complex (step G) followed
either by a slow NO release (step 10) or oxidation of the ferrous
heme-NO complex (step 9). Our purpose was to simulate both the initial
and steady-state behaviors of nNOS and to understand how NO regulates
enzyme activity and oxygen response. We did not try to model precisely
the forward pathway and assumed some simplifications concerning this
sequence. The validity of these simplifications will be judged on the
ability of the model to reproduce the principal parameters and
particularly NADPH oxidation, NO synthesis, and oxygen response.
Considering the concentration of oxygen and NO, oxygen binding and NO
release steps are assumed to be irreversible in the model, and we
considered only forward rate constants. The constants we used to build
this model were all determined at 10 °C and were obtained from the literature or determined here when necessary.
Substrate Binding and Citrulline Release--
At the Arg
concentrations typically used in our experiments, the association rate
is sufficiently high to neglect Arg binding in the kinetic analysis
(apparent binding rate constant around 300 s
1
considering a kon between 2 and 8 × 105 M
1
s
1 (33, 34)). The Arg
koff is also slow enough to assume that no
substrate molecule is likely to leave the binding site once it is bound
(koff = between 0.8 and 1.6 s
1 (33-35)). Similar arguments can be made
for NOHA after it forms in the active site (34, 35). However, release
of citrulline could affect the overall kinetics if it is slow relative
to kcat. The only published value for citrulline
koff was indirectly derived from single turnover
and steady-state data (36) and led to an estimate of 0.32 s
1, which would be rate-limiting. To check
the reliability of this value, we studied citrulline binding by
equilibrium and stopped-flow methods. Based on the changes observed in
the UV and resonance Raman spectra for citrulline (37, 39) or
thiocitrulline binding (34, 38, 39), we followed citrulline binding to
nNOS as a shift in Soret absorbance from 393 to 417 nm (Fig.
1A). The Kd
derived from the equilibrium titration is 7 ± 2.5 mM.
We then measured the rate of citrulline release in a stopped-flow chase
experiment using excess Arg. An nNOS sample containing 100 mM citrulline was rapidly mixed with a solution containing
100 mM Arg, and 96 spectra were recorded during the first
144 or 288 ms. As shown in Fig. 1B, there was a progressive
decrease in absorption at 417 and an increase at 393, which reflects
dissociation of the ferric-citrulline complex and buildup of the
ferric-Arg complex. The kinetics of spectral change at 387 and 417 nm
are displayed in Fig. 1C. The traces fit well to single
exponential curves and give an apparent off rate of 17 ± 2 s
1. At the Arg concentration used in this
experiment the binding rate of Arg is maximum. Therefore, we can assume
that the koff of citrulline is no slower than
17 ± 2 s
1. By using our experimentally
determined koff and Kd values, the citrulline kon is calculated to be
2 × 103 M
1
s
1, which is significantly slower than the
Arg kon. Thus, citrulline in the range of
concentrations typically generated during NOS activity assays (0-20
µM) will not compete with Arg binding. This is confirmed
by Frey et al. (38), Rogers and Ignarro (20), and
Sennequier and Stuehr (39), who did not see a change in steady-state
activity with added citrulline up to concentrations of 1 mM. We conclude that binding and release of amino acid
substrates or products will not impact the kinetic characteristics of
nNOS catalysis under our experimental conditions.

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Fig. 1.
Kinetics of citrulline binding to ferric
nNOS. A, change in absorbance at 420 nM for
1 µM nNOS as a function of citrulline concentration.
Conditions are described under "Experimental Procedures." The curve
was fit to a one binding site model with Origin©. B,
rapid-scanning spectra recorded after mixing citrulline-bound nNOS with
excess Arg. Conditions are described under "Experimental
Procedures." Traces shown were collected between 0.00135 and 0.135 s
after mixing. These curves are representative of more than 20 mixing
experiments. C, absorbance at 417 nm (solid line)
and 387 nm (dashed line) versus time derived from
mixing experiments depicted in B. The change at 417 nm
represents loss of the citrulline-bound nNOS, whereas the change at 387 nm represents gain in the Arg-bound nNOS. These curves are
representative of more than 20 mixing experiments.
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Flavin and Heme Reduction--
The rate of flavin reduction in
CaM-bound nNOS is relatively high (bimolecular reduction constants
ranged from 0.3 to 2 × 106
M
1 s
1
according to Gachhui et al. (40) and Abu-Soud et
al. (41)) and leads to apparent monomolecular constants higher
than 70 s
1 in the range of NADPH
concentrations that we used. Thus, we assume that flavin reduction is
sufficiently fast to neglect this step, and we only consider the rate
of electron transfer between the reductase domain and oxygenase domain
heme, which is a monomolecular process. Because flavin reduction is
fast, the rate of heme reduction under aerobic conditions can be
inferred from the initial rate of NADPH consumption. We obtain then a
pseudo-first order rate constant independent of the NADPH concentration
that characterizes heme reduction. We checked this hypothesis with
stopped-flow experiments in which nNOS catalysis was triggered by NADPH
addition to CaM-bound enzyme or by adding excess Ca2+ to an
NADPH-reduced enzyme. NADPH consumption was followed as a decrease in
absorption at 340 nm. The initial slope was ~2.4 s
1 in the NADPH-triggered reaction (Fig.
2, Table
I). In experiments where the initial rate
of heme reduction was followed at 395 nm after initiating by
Ca2+ addition, absorption loss followed a double
exponential decay, with a slow phase characterized by a rate constant
of about 2 s
1. This value, which represents
heme reduction (and is the rate-limiting step of the process, as seen
later), coincides well with the initial rate of NADPH consumption.
Moreover, both values are close to rates obtained for nNOS heme
reduction by Adak et al. (3.6 s
1
(22)) and Gachhui et al. (3.6 s
1
(40)) through anaerobic experiments at similar temperature. This
validates our simplification and allows us to derive the rate of heme
reduction from the initial NADPH oxidation rate, whose signal remains
largely unaffected by changes in heme or flavin absorption. Similar
results were obtained in experiments with NOHA (data not shown), where
the initial slope of NADPH consumption gave a pseudo-first order rate
constant of 2.5 s
1. Thus, we assume in our
model that the heme reduction rate is the same for enzyme containing
Arg or NOHA. In addition, based on our unpublished
results2 (data not shown), we
assumed that reduction of the ferric heme proceeds at the same rate
whether NO is bound or not.

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Fig. 2.
Experimental and simulated characterization
of nNOS. A, spectra of 1 µM nNOS before
(solid line) and after (dashed-dotted line)
initiating NO synthesis by Ca2+ addition to the cuvette in
the presence of an NADPH-regenerating system. Spectra were recorded at
23 °C. Other conditions are described under "Experimental
Procedures." Inset, difference spectrum between active and
resting enzyme. B, stopped-flow analysis of Fe(II) heme-NO
complex formation (436 nm) and loss of ferric nNOS (395 nm) after
mixing excess Ca2+ with NADPH-reduced nNOS (1 µM) at 10 °C. Other conditions are described under
"Experimental Procedures." Each trace is representative of eight
mixing experiments. C, stopped-flow analysis of NADPH
consumption at 340 nm after mixing excess Ca2+ with
NADPH-reduced nNOS (1 µM) at 10 °C. Other conditions
are described under "Experimental Procedures." D,
simulated concentration change for the Fe(III) nNOS (solid
line) and Fe(II) heme-NO species (dashed-dotted line)
as a function of time after initiating NO synthesis by 1 µM nNOS. The simulation was based on the kinetic model of
Scheme 1. Data were obtained by iterative calculation using Mathcad©
and assuming constant concentrations of O2 (140 µM) and NADPH (40 µM). The curves were fit
to a two-exponential equation. Further details are under
"Experimental Procedures" and the "Appendix." E,
simulated concentration change for NADPH (dotted line),
citrulline (solid line), and NO (dashed line) as
a function of time in the same reaction as described in
D.
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Table I
Comparison of pre-steady-state and steady-state values from in vitro
experiments and theoretical simulation with wild-type nNOS
Constants and values were derived from the literature or from reported
experiments. See text for details. Values were determined at 10 °C
unless specified. The rate constants used in the simulation are as
follows: k1, 2.6 s 1;
k2, 9 × 105 M 1
s 1; k3, 26 s 1;
k4, 2.6 s 1; k5, 9 × 105 M 1 s 1;
k6, 26 s 1; kF, 5 s 1; kG, 2.6 s 1;
k9, 1 × 10 4 s 1;
k10, 1.3 × 103
M 1 s 1. References for the literature
values are in parentheses.
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O2 Binding to Heme--
The O2
concentration in our experiments is relatively high (~150
µM in half-air-saturated solution at 10 °C), and
little O2 is consumed during the rate measurement. Thus, we
assumed a constant oxygen concentration and binding rate in the model.
We used a pseudo-first order rate based on results obtained by Boggs
and colleagues (26) at the same oxygen concentration and temperature. Their pseudo-first order binding rate
(pseudo-kon) varies between 8 and 10 × 105 M
1
s
1, consistent with more extensive
O2 binding studies by Abu Soud et al. (27) who
for nNOS determined a kon for O2 of
9 × 105 M
1
s
1. The high association rate and similarity
between these estimates validates our considering O2
binding as an irreversible step in the model.
We neglected superoxide or H2O2 release due to
autooxidation of Fe(II)-O2. No direct kinetic data are
available concerning the release of superoxide in the presence of Arg
and H4B, the only rates reported concern conversion of the
Fe(II)-O2 species to ferric enzyme (27, 43, 44), which may
involve reduction of the complex by H4B. Moreover, Arg
seems to suppress completely superoxide generation (45-48). In the
same manner, H2O2 formation (direct release
from the heme or through superoxide dismutation) is typically absent in
assays run in presence of Arg and H4B (49).
Catalytic Step--
We assumed that both steps of the reaction
(conversion of Arg to NOHA and then NOHA to citrulline) could be
approximated by the same rate constants. This assertion is supported by
the results of Iwanaga et al. (36) who found similar rate
constants for the Arg to NOHA and NOHA to citrulline transitions in
nNOS (7 and 6 s
1, respectively). Although
this simplification may modify the overall rate of catalysis in our
model, it will not affect the characteristics that are linked to
heme-NO binding. Boggs and colleagues (26) have studied the NOHA to
citrulline catalytic step in a single turnover at 10 °C starting
with pre-reduced ferrous nNOS oxygenase domain. The transformation rate
of Fe(II)-O2 into Fe(III)-NO product was 26 s
1. Few other reports are available. Iganawa
et al. (36) found global forward constants between 6 and 7 s
1 for nNOS, but this rate included heme
reduction. However, the weighted reciprocal mean of the three constants
we used for describing the same phenomenon is 7 s
1, which coincides with their value.
NO Release from Heme or Oxidation of the Fe(II) Heme-NO
Complex--
We used a rate constant for NO dissociation from ferrous
nNOS of 1 × 10
4
s
1 based on the measurements of Scheele
et al. (30). For O2 reaction with the Fe(II)
heme-NO complex we used a pseudo-first order rate constant (0.19 s
1) that was reported by Adak et
al. (22) at a single O2 concentration (half-air-saturated) at 10 °C. NO dissociation from ferric nNOS corresponds to rate of 5 s
1 in our model (22,
26, 30). Thus, the Fe(III) heme-NO complex can be reduced or have NO
dissociate at similar rates, which allows near equivalent partitioning
of this species between the two recycling pathways in Scheme 1. In
contrast, NO dissociation from the Fe(II) heme-NO complex or its
reaction with O2 is comparatively slow, which explains why
the Fe(II) heme-NO complex accumulates during the steady state.
Mathematic Simulation of the Kinetic Model--
We used a
simultaneous iterative calculation based on rate equations derived from
the kinetic model in Scheme 1. Rate equations are described under the
"Appendix." These calculations were achieved using Mathcad 7.0 software. A single catalytic turnover takes about 1 s for nNOS at
10 °C. Thus, for the calculations we chose a time step of 1 ms,
which should be small enough given that the fastest step in the process
(O2 binding) is around 100 s
1.
Indeed, no changes were seen if we shortened the time step for analysis
to 0.1 ms (data not shown). As described below, this model can simulate
the concentrations of all the heme intermediates depicted in Scheme 1,
the rates of NADPH consumption and citrulline, NO, or nitrate
production at pre-steady state and steady state, the various ratios of
these rates, and how all these parameters change as a function of time,
O2 concentration, or rate of heme reduction. NADPH
consumption rates were derived from the slope between 0 and 500 ms
(burst phase) and then between 10 and 50 s (steady-state phase).
The latter time scale was used to determine the steady-state formation
rate of citrulline and NO.
Comparing Experimental and Simulated Behavior of nNOS--
We
first confirmed the basic behavior of nNOS regarding its partitioning
during steady-state NO synthesis. In Fig. 2A, NO synthesis
was triggered by adding excess Ca2+ to NADPH-reduced nNOS
in the presence of CaM and an NADPH-regenerating system. As previously
reported (14, 22), the reaction at steady state was characterized by an
absorbance increase at 436 nm and an absorbance decrease at 395 nm, due
to the presence of the Fe(II) heme-NO species and a loss of the ferric
species. The relative proportion of these two species was calculated
based on standard spectra of ferric, ferrous, ferrous heme-NO, and
ferric heme-NO nNOS as published by Wang et al. (50, 51) and
Abu-Soud et al. (14). This led us to redefine our estimated
extinction coefficient for the ferrous heme-NO complex to 60 mM
1 s
1
at 436 nm. In our experiment the percentage of Fe(III) enzyme that
remained during steady state was ~28%, and the percentage of ferrous
heme-NO species present was about 72% (Table I). This falls within
estimates reported in literature (14, 15, 22), which range between 67 and 85% ferrous heme-NO complex.
We also confirmed nNOS behavior during the initial phase of NO
synthesis using rapid-scanning stopped-flow spectroscopy. Here, the
reaction was triggered with NADPH and did not include an
NADPH-regenerating system so we could follow consumption of NADPH. As
shown in Fig. 2B, there was a time-dependent
absorbance increase at 436 nm and decrease at 395 nm to levels that
were similar to those in Fig. 2A and to results reported
previously (14, 15, 22). We also observed an inflection in the NADPH
oxidation rate that corresponded to buildup of the 436 nm species (Fig.
2C). The first rate of NADPH consumption was derived from
the slope between 0 and 0.5 s, and the second rate of NADPH
consumption was calculated between 4 and 14 s. A multifit of eight
similar experiments gave an NADPH oxidation rate of 2.4/s for the first
phase and 0.48/s for the second phase, i.e. a ratio of 5. This is in the same range of values reported by Adak et al.
(22) (Table I). We also determined the rate constants of Fe(II) heme-NO
formation and loss of Fe(III) enzyme through a multifit of traces from
several similar experiments. For Fe(II) heme-NO buildup, the curves
best fit to a double exponential equation, giving rate constants that
vary from 15 to 10 s
1 for the fast phase and
from 2.5 to 0.9 s
1 for the slow phase. Again,
this is close to values obtained for nNOS by Adak et al.
(22) and Abu-Soud et al. (14, 15) (Table I). For loss of
Fe(III) enzyme, the traces also follow a double exponential decay, with
rate constants of 12 s
1 for the fast phase
and from 2 to 0.8 s
1 for the slow phase. This
confirms that loss of the ferric species is kinetically coupled to
heme-NO complex formation after NO synthesis is started.
We next simulated the NO synthesis reaction according to our kinetic
model in Scheme 1, using kinetic values and concentration of enzyme,
O2, Arg, and NADPH derived for wild-type nNOS as discussed above. Traces in Fig. 2, D and E, show that the
simulation faithfully reproduces the behavior of nNOS. Within the
correct time period after catalysis is initiated, the Fe(II) heme-NO
species builds up to represent 67% of the total enzyme, whereas the
Fe(III) species decreases to 25% of total enzyme. The remaining 8% of
nNOS mainly distributes between two other enzyme species
(Fe(II)-O2 and Fe(III) heme-NO; data not shown). The
simulation also gives NADPH consumption rates of 2.35 and 0.55 turnovers/s in the initial and steady-state phases of NO synthesis,
respectively. Thus, the simulated change in NADPH oxidation rate occurs
within the correct time frame, corresponds to buildup of the Fe(II)
heme-NO species, and is of a magnitude (5-fold decrease) that is very
close to our experimental value. As shown in Fig. 2E, the
simulation also predicts a deflection in the rate of citrulline
accumulation and has NO release delayed until heme-NO complex buildup
nears completion.
The simulated rate of Fe(III) loss is biphasic with rate constants of
22.8 and 0.61 s
1 which approximates our
experimental values. However, simulated buildup of the Fe(II) heme-NO
species best fit to a mono-exponential equation with a rate constant
corresponding to the slower rate component that was experimentally
observed (Table I). Thus, our model does not predict the fast phase of
Fe(II) heme-NO buildup. One explanation may be that absorption at 436 does not characterize only the buildup of Fe(II)-NO but can also
reflect absorption changes linked to Fe(III) heme-NO that forms prior
to the Fe(II) heme-NO species (26) and could contribute to the fast
phase. The purpose of our model was not to reproduce precisely the
kinetics within the first 50 ms. Indeed, the simplification of the
forward pathway and the limitation of the mathematical simulation
forbid it. But the resolution of this ultra-rapid phase is of no use, and our model faithfully reproduces changes that occur within the 1st s.
The simulation also predicts an NADPH per citrulline ratio of 1.67, which is greater than the theoretical minimum value (1.5 NADPH oxidized
per citrulline formed) (7, 8, 11) but actually matches several
experimental values (53, 54). It also predicts that nNOS will release
somewhat less NO than citrulline during the steady state. Both of these
effects derive from a portion of the Fe(III) heme-NO species being
reduced to the corresponding ferrous species, which wastes an
NADPH-derived electron because it leads the enzyme to generate
citrulline without releasing free NO (see Scheme 1). Thus, our model
gives insight into the functioning and efficiency of catalysis.
Partitioning of nNOS between productive and futile regenerating cycles
is a key aspect of our model. The relative enzyme flux through the two
different paths can be appreciated through different ratios. For
example, the ratio of nitrate versus citrulline reflects the
flux of nNOS through the futile cycle. The simulation gives a ratio of
0.34, which means about a third of the enzyme cycles through this
futile pathway in unit time. The ratio of NADPH-oxidized versus NO-formed depends also on the partitioning between
both regenerating pathways. The simulation gives a ratio of 2.54, which is a bit high but close to several reported values (from 1.7 to 2.2, see Table I). Because this ratio is quite sensitive to the heme
reduction rate, slight changes can modify it drastically and make it
difficult to obtain a perfect fit between experimental and simulated
substrate/product ratios.
The Effect of NO Synthesis on nNOS Apparent
Km,O2--
One remarkable characteristic of
nNOS is that NO synthesis greatly increases its apparent
Km,O2. A
Km,O2 of 350 µM
was obtained when nNOS was synthesizing NO, whereas a value of about 38 µM was obtained when nNOS oxidized NADPH in the absence
of substrate (15). This shift cannot be derived from a direct
competition between NO and O2, because it would require NO
concentrations (20-40 µM) that are out of reach for steady-state experiments. In fact, this behavior appears to involve formation of the Fe(II) heme-NO complex, which influences the apparent
Km,O2 through its
O2-dependent oxidation reaction (see Scheme 1).
Thus, we tested if our kinetic model could simulate how catalysis in
the presence and absence of NO synthesis affects enzyme
Km,O2. We simplified our model
into a substrate-free one, where one-electron reduction of the heme
leads only to superoxide generation (Fig.
3, inset) (48). The numerical
treatment was achieved by the same iterative method discussed above
(see "Appendix"). The rate constants used here were the same as
noted above except for the release of superoxide. We used the values of
Boggs et al. (26) who reported a general
kcat of 20 s
1 for
ferrous nNOS in the absence of substrate. For
O2-dependent decay of the Fe(II) heme-NO
complex, we varied the single rate constant obtained for this reaction
(see above) in direct proportion to the simulated O2
concentration. We then calculated the rate of catalysis (NADPH
oxidation) as a function of O2 concentration in the
presence or absence of substrate (Fig. 3). The
Km,O2 obtained in the absence
of substrate (no NO synthesis) was 2.5 µM. Although this
is somewhat lower than our experimentally determined value (14), it is
close to values generally reported for O2-binding heme
proteins and primarily reflects ferrous heme affinity for O2 (50). In the presence of substrate (and NO synthesis)
the Km,O2 was 271 µM, which is close to the measured value of Abu-Soud
et al. (15). Thus, our model can accurately simulate how NO
synthesis shifts the Km,O2 of
nNOS.

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Fig. 3.
Simulated curves of activity
versus O2 concentration for nNOS in the
presence or absence of Arg. Data points represent steady-state
rates of NADPH oxidation in the presence (solid square) or
absence (open circle) of Arg. Values were calculated
respectively from the kinetic models depicted in Scheme 1 or the
inset. The simulation process is described under
"Experimental Procedures" and "Appendix." The curves were fit
using a single binding site equation with Origin© software.
Inset, kinetic scheme used to simulate nNOS catalysis in the
absence of substrate. Single arrows indicate assumption that
each step is irreversible. Bold numbers refer to rate
constants listed in Table I. See text for details.
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Simulating Behavior of Hyperactive nNOS Mutants--
We next
tested if our kinetic model could simulate the initial and steady-state
behavior of nNOS W409F and W409Y mutants, which have been characterized
by Adak et al. (22, 23). These mutants have lower levels of
Fe(II) heme-NO complex buildup in the steady-state, a negligible
deflection between their initial and steady-state rates of NADPH
oxidation, and an increased rate of NO synthesis during the steady
state as compared with wild-type nNOS. Only two kinetic steps appear to
be changed in the mutants (22, 23). Their rates of heme reduction are
2-3 times slower compared with wild type, and their rates of Fe(II)
heme-NO complex oxidation are 7-8 times faster.
We performed two different simulations based on W409F mutation. In the
first we increased the rate of Fe(II) heme-NO oxidation and left the
heme reduction rate constant (Fig.
4A and Table
II), whereas in the second we both
increased Fe(II) heme-NO oxidation and slowed the heme reduction rate
(Fig. 4B and Table II). In both cases, we obtain a smaller
percentage of Fe(II) heme-NO complex during steady state (21.5 and
9.5%) compared with wild-type nNOS. These reduced levels are close to
percentages directly estimated from experiments (25% for W409F and
14% W409Y; Ref. 23). Moreover, the simulation correctly predicts that
only a small change occurs between the initial and steady-state rates
of NADPH consumption in W409F and W409Y, and their steady-state rates
are predicted to be faster than wild type (1.3 and 0.9 turnover/s for
the mutants compared with 0.55 for wild type). These simulated rates of
NADPH turnover are almost identical to those measured for W409F and W409Y at 10 °C (1.4 and 0.8 turnover/s, Ref. 22).

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Fig. 4.
Simulated pre- and steady-state behavior of
the nNOS W409F mutant. Both panels show simulated concentration
changes as a function of time for the Fe(III) nNOS, Fe(II) heme-NO
complex, and NADPH after initiating NO synthesis at 10 °C. These
were calculated using the kinetic model in Scheme 1 and the reaction
conditions for wild-type nNOS in Fig. 2. A, results using
one different rate constant compared with wild-type nNOS (a faster
Fe(II) heme-NO oxidation rate of 9 × 103
M 1 s 1).
B, results incorporating two different rate constants (the
faster Fe(II) heme-NO oxidation rate and a slower heme reduction rate
of 1.2 s 1). The buildup of Fe(II) heme-NO
complex and decay of Fe(III) nNOS were fit to single- or
two-exponential equations as described under "Experimental
Procedures."
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Table II
Comparison of the values of some pre-steady-state and steady-state
indicators between in vitro experiments and theoretical simulation,
case of W409 mutant nNOS
Constants were derived from the literature. Constants and values
obtained from our simulations correspond to 10 °C unless otherwise
specified. The rate constants used in the simulation are as follows:
k1, 1.4 s 1; k2, 9 × 105 M 1 s 1;
k3, 26 s 1; k4, 1.4 s 1; k5, 9 × 105
M 1 s 1; k6, 26 s 1; kF, 5 s 1;
kG, 1.4 s 1; k9, 1 × 10 4 s 1; k10, 9 × 10 3 M 1 s 1. Simulation I
incorporates only an increase in Fe(II)-NO oxidation rate
(k10 value); Simulation II incorporates both an
increase in Fe(II)-NO oxidation rate (k10 value) and
a decrease in heme reduction rate (k1,
k4, kG values). Values from
literature are drawn from Adak et al. (22). See text for
details.
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Other parameters derived from the simulation match those determined
experimentally (see Table II for general comparison). In particular,
the model correctly simulates a slower buildup of the Fe(II) heme-NO
complex in the mutants (23). The ratio of NADPH oxidized
versus NO produced is also predicted to be smaller for the
Trp-409 mutants than for wild type, as confirmed experimentally by Adak
et al. (23). The slight overestimation of this parameter in
the simulation highlights the sensitivity of the outcome toward modifications in heme reduction rate. In general, slowing heme reduction favors enzyme partitioning toward the active cycle (NO dissociation from the Fe(III) heme-NO complex) and disfavors
partitioning toward the futile cycle (reduction to form the Fe(II)
heme-NO complex) (see Scheme 1). We also simulated catalysis at
different O2 concentrations to derive
Km,O2 values for the W409F mutant. The simulations were run as described for wild type except we
included appropriate heme reduction and Fe(II) heme-NO oxidation rates
as noted above. We obtained
Km,O2 values of 41 ± 1 and 16 ± 1 µM for W409F depending on whether one or
both parameters were modified (data not shown). These values clearly
differ from the wild-type
Km,O2 (271 µM)
but actually match the values determined experimentally for Trp-409
mutants by Adak et al.
(Km,O2 around 40 µM).3 Together,
this highlights how our model can correctly simulate complex effects of
a mutation on both pre-steady-state and steady-state catalysis.
Influence of Heme Reduction Rate--
Our kinetic model can also
predict how graded changes in any kinetic parameter will affect nNOS
catalysis. To examine the influence of heme reduction rate, we ran
simulations at heme reduction rates that ranged from 0 to 10 s
1. Fig.
5A shows how the distribution
of three nNOS species varies in the steady state as a function of heme
reduction rate. At rates below 1 s
1, most of
the enzyme is present in its Fe(III) form. However, as heme reduction
gets faster, the proportion of Fe(III) species goes down and there is a
concomitant gain in Fe(II) heme-NO species, which predominates at rates
above 2 s
1. Clearly, the relative proportion
of these species is quite sensitive to the heme reduction rate. In
contrast, the simulation predicts that the Fe(II)-O2
species builds up to a relatively minor but stable level. In general,
the steady-state concentrations of Fe(III) and Fe(II) heme-NO species
as predicted by the model match experimental data obtained for nNOS
(this report and Ref. 15).

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Fig. 5.
Catalytic behavior of nNOS as a function of
heme reduction rate. All simulations followed the kinetic model in
Scheme 1 and reaction conditions described for Fig. 2. The values of
k1, k4, and
kG (see Scheme 1) were simultaneously changed for
each iterative simulation. A, change in population of three
heme species at steady state. B, change in rates of NADPH
oxidation, NO, and citrulline synthesis at steady state. C,
change in rates of NO synthesis at steady state for different values of
Fe(III)-NO dissociation rate (1 s 1, ; 10 s 1, ) and Fe(II)-NO oxidation rate (0.05 s 1, ; 1 s 1,
). The dotted line corresponds to a simulation run with
the same values as in B. D, change in various
efficiency ratios at steady state.
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Fig. 5B predicts that a surprising relationship exists
between heme reduction rate and steady-state catalysis. Regardless of
how nNOS activity is measured (rates of NADPH oxidation, citrulline, or
NO synthesis) the simulation shows that there is an optimum rate of
heme reduction beyond which activity falls. Although this seems
counterintuitive, it makes sense when one considers that faster rates
of heme reduction favor greater reduction of the Fe(III) heme-NO
product complex, and thus favor cycling of enzyme through the futile
pathway (see Scheme 1). This eventually decreases the rate of NO
release in the steady state. On the other hand, for slower rates of
heme reduction, product formation (the Fe(III) heme-NO complex) becomes
the rate-limiting step and can result in a decreased rate of NO
synthesis. It appears that the system can set the rate of heme
reduction to optimally counterbalance these two effects.
To ensure the robustness of our model, we checked if this surprising
and unexpected regulation feature is not linked to particular values of
the kinetic parameters. Three main parameters qualitatively and
quantitatively control the outcome of the simulation as follows: (i)
the heme reduction rate, which influences the biosynthetic pathway and
enzyme partitioning between the two regenerating cycles; (ii) the
dissociation rate of Fe(III)-NO, which influences enzyme partitioning
and controls the rate of the productive regenerating cycle; (iii) the
rate of Fe(II)-NO oxidation, which controls the rate of the futile
regenerating cycle. As can be seen in Fig. 5B, a variation
of 4-fold in the simulated rate of heme reduction (between 1 and 4 s
1; experimental values ranged between 2.4 and 3.6 s
1) only moderately alters the
simulated NO synthesis rate (0.225 s
1 ± 20%). The same variation alters rates of citrulline production and
NADPH consumption by 10 and 8%, respectively. We also ran simulations
for extreme changes in parameters ii and iii noted above (Fig.
5C). We chose rates of Fe(III)-NO dissociation of 1 and 10 s
1 (experimental values range between 3.5 and
5 s
1). The optimum rate for heme reduction in
this case shifts between 1 and 3 s
1, along
with an obvious increase in the rate of NO synthesis (4-fold versus a 10-fold change in Fe(III)-NO dissociation rate).
When we varied the simulated rate of Fe(II)-NO oxidation between 0.05 and 1 s
1 (experimental value is 0.19 s
1), the optimum rate for heme reduction
shifts between 0.8 to 2 s
1, and the NO
synthesis activity is multiplied by 2 (for a 20-fold increase of
Fe(II)-NO oxidation rate). Clearly, extreme changes of these essential
parameters (up to 20-fold) do not alter drastically the results of the
simulation. In all cases, NO synthesis versus heme reduction
rate is described by a bell-shaped curve with an optimum between 1 and
3 s
1. Although such changes also altered the
predicted rate of NO synthesis, it is encouraging to note that the
experimental rate values we used for i-iii above in our simulation
actually predict a steady-state rate of NO synthesis that is quite
close to the experimentally observed value. Therefore our kinetic model
appears robust and is not overly sensitive to inaccuracies in any
particular rate measurement.
Fig. 5D simulates how the rate of heme reduction influences
catalytic efficiency in nNOS. The NADPH/NO ratio climbs continuously from an initial value of 1.5 as the heme reduction rate increases. NO
release becomes less probable as the heme reduction rate increases because a greater percentage of the Fe(III) heme-NO complex is channeled into the futile cycle (see Scheme 1). Because of this lowering in catalytic efficiency, citrulline and NO release become increasingly uncoupled from NADPH consumption as the heme reduction rate increases. The uncoupling profile is less accentuated for citrulline because citrulline is made when enzyme partitions into either the active or futile cycle, whereas NO is only released by
enzyme molecules that partition into the active cycle. Because the
futile cycle generates nitrate instead of NO, the nitrate/citrulline ratio also increases with the heme reduction rate (Fig. 5D).
This ratio has a maximum value of 1, and the simulation suggests that a
significant percentage of total enzyme will cycle through the futile
cycle at high heme reduction rates.
 |
DISCUSSION |
We developed a kinetic model to understand the unique behavior of
nNOS. This involves a fast buildup of ferrous-NO complex includes about
80% of the steady-state population, with an associated change between
initial and steady-state activities. In the past we proposed models for
nNOS partitioning which typically invoked direct competition between NO
and O2 binding to ferrous heme (15). Nevertheless, none of
the earlier models when simulated were able to mimic closely the
behavior of nNOS, and the competition binding concept eventually did
not fit well with relative rates of heme reduction and ligand binding
that were determined later. Our current model integrates the latest
results in the literature, particularly the concept of fast
recombination between ferric heme and NO formed in the active site (26,
30). Simulations of our kinetic model closely mimic all catalytic
features of wild-type or mutant nNOS, including their initial and
steady-state behaviors, populations of heme species, and apparent
Km,O2 values. Together, this
strongly argues that the fundamental features of our current model are correct.
In our model catalytic turnover is regulated by the following two
distinct features: 1) partitioning of an immediate Fe(III) heme-NO
product between a futile and productive regenerating pathway, and 2)
the relative rate of the futile regenerating pathway. These two
features share no common influences and thus operate completely independent of one another. Partitioning simply reflects the relative rates of NO dissociation versus reduction of the immediate
Fe(III) heme-NO product. Because these are fundamental characteristics of the enzyme, they are immune to the external environment when the
enzyme operates under Vmax conditions
(i.e. sufficient NADPH, Arg, etc.). However, because they
oppose each other, small changes in these two rates can have a
relatively large effect on enzyme partitioning. For our model we
assumed that reduction of the Fe(III) heme-NO complex was identical to
reduction of the ferric enzyme, whose extent and rate is known (40,
41). Because NO binding to hemeproteins typically raises their midpoint
potential (55), the assumption is probably valid at least from a
thermodynamic standpoint. The accuracy of our model in simulating
several aspects of nNOS catalytic behavior also supports the
assumption. Nevertheless, a direct measure of this rate may enable
finer refinement. In contrast to the partitioning step, the rate of the
futile regenerating cycle is clearly influenced by factors that are
both extrinsic and intrinsic to the enzyme. For example, the rate of
Fe(II) heme-NO oxidation is directly related to the O2
concentration, and enzyme mutations at residue Trp-409 greatly increase
the rate at any given O2 concentration. Given the multiple
opportunities for "fine-tuning" both intrinsic and extrinsic
parameters, our kinetic model should be useful to guide and interpret
further investigation of catalysis and regulation.
One imagines that intrinsic parameters of nNOS have been set for its
proper physiologic function. This may be illustrated by its measured
rate of heme reduction corresponding to the activity optimum predicted
in our model (see Fig. 5B). Intrinsic parameters can also
govern the relative impact of extrinsic factors such as O2
concentration. This is illustrated by the high apparent Km,O2 that is unique to nNOS
(14, 56). Our model reveals that a relatively fast heme reduction,
coupled with a relatively slow cycling through the futile pathway,
creates this special regulation in nNOS. Functionally, this allows the
enzyme to minimize its catalytic sensitivity to O2
concentration changes, while generating NO in a near-linear manner as a
function of O2 concentration. Why this evolved is still
unclear (14). However, it must be physiologically important, because
our model clearly shows that the enzyme sacrifices catalytic efficiency
to create this response.
For iNOS (17, 18, 57), eNOS (16, 19), and nNOS (16, 20), a loss of
activity occurs if heme binds NO that accumulates in solution. However,
this occurs only when relatively high NO concentrations are achieved,
can be prevented by NO scavenging, and involves trapping the heme in a
ferric NO complex. It is important to stress that this phenomenon
corresponds to a simple equilibrium between ferric heme and free NO and
is therefore quite different from the kind of regulation described in
this report. This phenomenon occurs quite late during NO synthesis (15 min at 37 °C) and therefore was not taken into account by our model
that focused on initial and steady-state phases.
Negrerie and colleagues (29), who thoroughly studied the NO
recombination process in eNOS, suggested on the basis of results by
Abu-Soud et al. (14) that the Fe(II) heme-NO species may form via reduction of the Fe(III) heme-NO species. However, the Fe(II)
heme-NO species is not observed during steady-state catalysis in eNOS
or iNOS. At this point, our model can provide an explanation for this
difference. eNOS is distinguished by a low steady-state activity, a
slow heme reduction, and an undetectable level of Fe(II) heme-NO
species (19). Its low catalytic activity is therefore not linked to
eNOS cycling through the futile regenerating pathway. In fact,
according to our simulations, the slow heme reduction rate in eNOS (19,
58, 59) is sufficient to explain its low proportion of Fe(II) heme-NO
species during the steady state (Fig. 5A) and its overall
lower activity (Fig. 5B). This suggests that eNOS follows
the same kinetic model as nNOS, but the slower heme reduction minimizes
partitioning into the futile regenerating cycle and diminishes the
global speed of catalysis. In iNOS, one observes a similar catalytic
activity and a slightly slower heme reduction rate compared with nNOS
(19, 60) but little or no Fe(II) heme-NO complex formation in the
steady state.4 This phenotype
is similar to what is seen for the Trp-409 nNOS mutants. Indeed, our
preliminary data suggest that iNOS exhibits accelerated Fe(II) heme-NO
oxidation,5 as occurs in the
Trp-409 nNOS mutants (23), and this can account for such results. Thus,
although several kinetic parameters remain to be determined for iNOS
and eNOS, our kinetic model seems capable of simulating their behavior
as well.
So far, the different behaviors of the three NOS isoforms appear to be
linked to differences in their heme reduction and Fe(II) heme-NO
oxidation rates. But what structural and physical parameters control
these processes? The Trp-409 mutations achieved by Adak et
al. (22, 23) suggest one controlling factor is the heme reduction
potential. Crystal structures show that a Trp-409 indole nitrogen forms
a strong hydrogen bond with the heme thiolate (61, 62). Suppression of
this hydrogen bond through mutation is expected to lower the reduction
potential of the heme, which we suspect inhibits heme reduction while
speeding oxidation of the Fe(II) heme-NO complex. This hypothesis also
arises from the comparison of the results of Wang et al.
(51), who observed a faster rate of Fe(II) heme-NO complex oxidation in
the absence of Arg, and Presta et al. (63), who showed that
Arg binding increases the reduction potential of the NOS heme.
Consequently, it seems possible that catalytic differences among NOS
isoforms arise in part from their differential control of heme
reduction potential. Our kinetic model provides a basis to investigate
this hypothesis and better understand the structure-function
relationships in the three NOS.