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INTRODUCTION |
Nitric-oxide synthase
(NOS)1 is an uncommon
self-sufficient P450-like enzyme catalyzing nitric oxide (NO)
biosynthesis from L-arginine (1-4). There are three
mammalian NOS isozymes: the constitutive neuronal NOS (nNOS) and
endothelial NOS (eNOS) require calmodulin for enzyme activity, whereas
the inducible NOS (iNOS) contains tightly bound calmodulin (1-4). All
three isozymes have a common bi-domain structure with the reductase
domain containing FAD, FMN, and NADPH binding sites, and the oxygenase
domain harboring the heme center and binding sites for
L-arginine and tetrahydrobiopterin (BH4)
(1-4). The main function of the reductase domain is to provide
reducing equivalents to the heme center in the oxygenase domain where
the key chemistry of L-arginine conversion occurs. Three
substrates and four products are involved in NOS catalysis. The overall
reaction is a complicated five-electron oxidation of the key guanidine
nitrogen plus three additional electrons from NADPH to reduce two
molecules of oxygen to water and form the L-citrulline and
nitric oxide. Several x-ray crystallographic structures for the iNOS
and eNOS oxygenase domains have been reported (5-7). The x-ray
crystallographic data at 1.9-Å resolution of the C-terminal FAD-NADPH
binding domain of the nNOS reductase domain was also published recently
(8). These data reveal a three-domain modular design. The
FAD and NADPH binding subdomains are superimposable on those of
cytochrome P450 reductase (CPR) with a root mean square deviation of
1.3 Å, whereas the more flexible FMN-connecting domain shows a 3.9-Å
root mean square deviation to the
-chain of CPR. The fourth domain
that binds FMN is lost during crystallization, but the structure is
projected to be similar to that of CPR. These crystallographic data
give firm support for a modular design of NOS and thus provide a
basis to prepare subdomains for structure/function and reaction
mechanism studies. Investigation into individual breakdown modules
could simplify the data interpretation for each redox center and should
be a useful approach in elucidation of the complicated reaction
mechanism for NOS.
Overexpression systems for the individual oxygenase and reductase
domain of NOS have been developed in bacterial and baculovirus systems,
including our own group (9-17). Only a few are related to eNOS
(6, 17). Large amounts of eNOSox were usually obtained by
trypsinolysis from intact bovine eNOS (13, 18). Although the
baculovirus system is useful (17), it is both time-consuming and
costly. The bacterial expression system (18), although fast, has
unpredictable sudden debilitating mutations in the expression construct
and, in our hands, has resisted being scaled up to more than a few
liters of culture for unknown reasons. Yeast expression could be an
alternative vehicle to generate large amounts of active mammalian
enzymes (19). Yeast has been shown to be effective in overexpressing
eNOS and the reductase domain of nNOS (14, 20). Here, we report the
overexpression in yeast the oxygenase, eNOSox, and
reductase, eNOSred, domains of human eNOS and the characterization for their oxidation-reduction activities. Both domains
show behaviors very similar to the domains present in the whole eNOS
and should be useful tools for future biophysical and mechanistic investigations.
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EXPERIMENTAL PROCEDURES |
Materials--
BH4 was obtained from Schircks
Laboratories (Jona, Switzerland). Plasmids containing human eNOS
cDNA in pGEM3Z and human eNOS polyclonal antibodies were kindly
provided by Dr. Pei-Feng Chen in our division (21). PCR kits (Expand
High Fidelity PCR system) were the product of Roche Molecular
Biochemicals. Restriction enzyme PmeI was purchased
from New England BioLabs, and the other restriction enzymes were from
Invitrogen. All reagents and devices for DNA extraction and isolation
were products of Qiagen. An Easyselect Pichia Expression
kit, containing the expression vector pPICZB, Pichia strain
GS115, and Escherichia coil strain TOP10F', was purchased
from Invitrogen and used for the expression of both eNOS domains.
Reagents for electrophoresis and Western blotting were from Bio-Rad.
The remaining chemicals were from Sigma.
Expression of Human eNOS Domains--
PCR was used to amplify
the cDNA product. Human eNOS cDNA in pGEM3Z was used as
template, and DNA fragments encoding oxygenase (amino acids 1-491) and
reductase domain (amino acids 482-1204) were amplified with specific
primers. For oxygenase domain, the forward primer was
5'-CGGAATTCAACATGCATCACCATCACCATCACGGCAACTTGAAGAGCGTG-3' (translation start codon is underlined), and the backward primer was
5'-GCTCTAGATCAGGTGATGCCGGTGCCCTTGGC-3' (translation
stop codon is underlined). For reductase domain, the forward primer was
5'-CGGAATTCAACATGCATCACCATCACCATCACGGGAGTGCCGCCAAGGGC-3', and the backward primer was
5'-GCTCTAGATCAGGGGCTGTTGGTGTCTGAGCC-3'. In both
forward primers, the EcoRI site and His6 tag
were added, and in each backward primer an XbaI site was
added. The correct sequences of the PCR products were confirmed by
primer extension sequencing. Both PCR products were double-digested
with EcoRI and XbaI and subcloned separately into
the corresponding sites of an alcohol oxidase promoter-driven
expression vector pPICZB to obtain a 1.5-kb insert of
eNOSox and a 2.1-kb insert of eNOSred. The
constructs were linearized with PmeI, transformed into yeast Pichia pastoris GS115, and selected by growing them on the
YPDS/Zeocin plates containing 1% yeast extract, 2% peptone, 2%
dextrose, 1 M sorbitol, and 100 µg/ml Zeocin. The colony
that grew fastest was inoculated into 25 ml of buffered minimal
glycerol medium (100 mM potassium phosphate, pH 6.0, 1%
yeast extract, 2% peptone, 1.34% yeast nitrogen base with ammonium
sulfate without amino acids, 4 × 10
5% biotin and
1% glycerol) and cultured at 30 °C overnight. This culture
was then transferred to a 250-ml buffered minimal glycerol medium and
grown at 30 °C overnight to A600 = 7-10.
Cells were harvested and resuspended in 250 ml of buffered minimal
methanol medium (100 mM potassium phosphate, pH 6.0, 1%
yeast extract, 2% peptone, 1.34% yeast nitrogen base with ammonium
sulfate without amino acids, 4 × 10
5% biotin, and
0.5% methanol) and cultured for 72 h at 30 °C to induce
protein expression.
Protein Purification--
Yeast cells were harvested and washed
with buffer 1 (50 mM Tris-HCl, pH 8.0) with protease
inhibitors (1 µM leupeptin, 1 µM antipain,
1 µM pepstatin A, and 1 mM
phenylmethylsulfonyl fluoride) and resuspended in an equal volume of
buffer 1 with protease inhibitors. An equal volume of glass beads
(425-600 µm) was added to the suspension. Cells were broken by 10 cycles of 30-s vortexing and brief chilling on ice. Cell debris and
glass beads were removed by centrifugation at 3,400 rpm. The
supernatant, obtained after another centrifugation at 12,000 rpm in a
microcentrifuge, was applied to a 2-ml nickel-nitrilotriacetic acid-agarose column. The column was first washed with a 50-bed volume of buffer 1 plus protease inhibitors, then washed with a
~30-bed volume of buffer 1 plus 0.3 M NaCl and 1 mM L-histidine, then by a ~20-bed volume
buffer 1 plus 0.1 M NaCl and 5 mM
L-histidine. Finally, buffer 1 plus 40 mM
L-histidine was used to elute bound oxygenase domain and
100 mM L-histidine for the reductase domain. The eluate was concentrated by Centriprep-50 then applied to a 10-DG
column (Bio-Rad) and eluted with 50 mM HEPES, pH 7.4, containing 0.1 M NaCl and 10% glycerol, to remove histidine.
Biopterin and Flavin Determination--
The content of
BH4, FAD, and FMN of purified eNOS domains was measured as
described previously (17, 21) and quantified from a standard curve of
authentic BH4, FAD, or FMN, respectively. Biopterin
determination was done on eNOSox with or without
reconstitution with exogenous BH4. BH4
reconstitution was done similarly to procedures that were published
previously (9, 22) under anaerobic condition. The excess amount of
BH4 was removed by gel filtration, and the amount of bound
BH4 was determined using our HPLC quantitation similar to
the published procedure using authentic BH4 to build a
standard curve (23).
Pyridine Hemochromogen Assay--
Heme content was determined by
the formation of pyridine hemochromogen as previously described (24).
The total heme content was determined from difference spectrum of
bis-pyridine heme (reduced minus oxidized) using

556-538 nm = 24 mM
1
cm
1.
Quantification of Thiol Functional Groups--
Surface-exposed
thiol groups were determined by chemical modification using
4,4'-dithiopyridine to form a 4-thiopyridone chromophore with major
absorbance at 343 nm. The 4,4'-dithiopyridine itself has almost no
absorption at that wavelength (25).
eNOSred Activity Assay--
Cytochrome c
reductase activity was measured as the absorbance increase at 550 nm
using 
= 21 mM
1 cm
1
as described previously (17). Ferricyanide or 2,6-dichlorophenol indolphenol oxidation assay was carried out using 
= 1 mM
1 cm
1 at 400 nm and

= 21 mM
1 cm
1 at 600 nm, respectively (14).
eNOSox Activity in Generating Biopterin
Radical--
This activity measurement essentially followed previous
published procedure for iNOSox (26, 27). High concentration
of BH4-reconstituted eNOSox was reduced
anaerobically in a tonometer by dithionite titration in the presence of
1 mM L-arginine. The ferrous eNOSox was then
reacted with oxygenated buffer using a rapid-freeze/EPR technique as we
previously published (28, 34). The rapid-freeze apparatus, System 1000 (Update Instrument, Madison, WI), was placed inside an anaerobic
chamber (Coy Laboratory). The oxygen level was lower than 5 ppm during
the whole experiment procedure and monitored by an oxygen/hydrogen
analyzer (Model 10, Coy Laboratory). One or two push programs were used
to obtain samples freeze-trapped at different reaction times.
Spectrometry--
UV-visible spectra were measured on an HP8453
diode array spectrophotometer with a 1-nm spectral bandwidth. EPR
results were recorded at liquid helium or liquid nitrogen temperature
on a Bruker EMX EPR spectrometer. For liquid helium system, a
GFS600 transfer line and an ITC503 temperature controller were used to maintain the temperature. An Oxford ESR900 cryostat was used to accommodate the sample. For liquid nitrogen transfer, a silver-coated double-jacketed glass transfer line and a BVT3000 temperature controller were used. Data analysis was conducted using WinEPR, and
spectral simulations were done using SimFonia programs provided by
Bruker. Flavin fluorescence was measured using an SLM SPF-500C spectrofluorometer using the ratio mode. About 2 µM
eNOSred in a 1-cm quartz cuvette was excited at 450 nm
(5-nm spectral bandwidth), and the emission spectrum between 450 and
650 nm (7.5-nm spectral bandwidth) was collected at 24 °C.
Stoichiometric Titration--
The redox capacities of
eNOSred and eNOSox were determined by anaerobic
stoichiometric titration using sodium dithionite. Stock solution of
sodium dithionite was freshly prepared by dissolving powdered reagent
in 50 mM, pH 8.2 pyrophosphate buffer pre-saturated with
pure nitrogen gas. The concentration of sodium dithionite was
standardized by titration against a fixed amount of lumiflavin-3-acetic acid (
444 = 1.08 × 104
M
1cm
1) anaerobically before and
after individual real sample titration (29). The average concentration
was used to calculate the number of reducing equivalents consumed in
the titrations. Each protein sample was placed in an anaerobic titrator
and made anaerobic by 5 cycles of evacuation (30 s) and argon
replacement (5 min). Standardized dithionite solution contained in a
gas-tight syringe engaged to the side arm of the titrator was
quantitatively delivered and mixed with the protein sample under argon
atmosphere. The electronic spectrum was recorded on an HP8452
diode array spectrophotometer to confirm that the system was
equilibrated after each addition of dithionite reflected by a static absorbance.
Miscellaneous Methods--
The protein content was determined by
BCA method (30). SDS-PAGE was performed on 10% Ready-Gels in a Bio-Rad
mini-gel apparatus. Gel filtration chromatography was performed on a
Sephacryl 200 HR column (1.5 × 50 cm). A kit for molecular weight
12,000-200,000 (product code: MW-GF-200) was used as the gel
filtration marker.
Computer Modeling--
The SCoP program (Simulation Resources
Inc., Redlands, CA) was used for simulating the data obtained from
stoichiometric titration, mainly the eNOSred similar to the
method used by Iyanagi et al. (31). The absorbance changes
at different monitoring wavelengths during titration were simulated
against accumulated reducing equivalents added,
|
(Eq. 1)
|
where A
is the observed absorbance at
wavelength
, and
1 through
6 are the extinction coefficients
for each flavin redox species (F1, F1H, and F1H2 represent
fully oxidized, semiquinone, and fully reduced forms of the first
flavin, respectively, and F2, F2H, and F2H2 are the
equivalents for the second flavin). The concentrations of each flavin
intermediate during a stoichiometric titration are expressed as
follows,
|
(Eq. 2)
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|
(Eq. 3)
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|
(Eq. 4)
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|
(Eq. 5)
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(Eq. 6)
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|
(Eq. 7)
|
where f1 and f2 are the total amounts of each flavin and,
|
(Eq. 8)
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(Eq. 9)
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(Eq. 10)
|
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(Eq. 11)
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where E1 through E4 are midpoint
potentials of the four half-reactions of the two flavins in
eNOSred, Eh is any measured redox potential
value, F is the Faraday constant (96,485 Coulomb mol
1), R is the gas constant (8.314 J
K
1mol
1), and T is temperature
(298 K). The total reducing equivalents were simply expressed as,
|
(Eq. 12)
|
Where ft is the total flavin,
i.e. the sum of f1 and f2.
Simulation was generated by sweeping the Eh values in any
desired potential range and seeking optimal values for
E1-E4 to achieve the best fit to the observed
data. It is not possible to achieve a set of absolute midpoint
potentials, but the relative midpoint potential values can be
converged. In other words, once one of the E1-E4
values is fixed, the other three can be located by simulation.
Absorbance extinction coefficients for the fully oxidized and fully
reduced flavins are readily available, and those for the flavin
semiquinone can be properly estimated from the spectrum at the stage of
one- and three-electron-reduced states. We also let these two
coefficients be variable in a narrow range and optimized their values
via simulation.
 |
RESULTS |
Expression and Purification of eNOS Subdomains--
The yeast
expression vector containing the AOX promoter is promising in
overexpressing both domains of eNOS in P. pastoris. By
introducing a His6 tag in the N terminus of both domains,
purification of the target protein can be conveniently done by
nickel-nitrilotriacetic acid-agarose column chromatography. The average
yields of the purified oxygenase domain and the reductase domain are
~8 and ~22 mg/liter, respectively. Both purified eNOSox
and eNOSred resulted in a single band on SDS-PAGE with
apparent molecular masses of 54 and 82 kDa, respectively (Fig.
1A) and exhibited
immunoreactivity with polyclonal antibodies against eNOS (Fig.
1B).

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Fig. 1.
Homogeneity of purified eNOSox
and eNOSred. Purified eNOSred (lane
2) and eNOSox (lane 3) after
nickel-chelating column chromatography were analyzed by
SDS-PAGE/Coomassie Blue staining (A) and Western blotting
(B). Molecular weight markers are shown in lane
1. Samples of approximately 1 µg were used for each run.
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The heme content determined by pyridine hemochromogen assay was
0.93 ± 0.04 (n = 7), almost stoichiometric to the
protein subunit (Table I). Replenishing
hemin or
-aminolevulinic acid to the cell medium during yeast growth
did not further increase the heme content in purified
eNOSox. The purified oxygenase domain also contained
endogenous biopterin at a stoichiometry lower than 0.3/monomer.
Because our sample buffers did not include dithiothreitol, most of
these biopterin molecules were present as dihydrobiopterin, BH2, as analyzed by our HPLC method (data not shown). The
functional form of biopterin, BH4, can be reconstituted
back to the purified eNOSox according to the anaerobic
procedure similar to that described by Rusche and Marletta (22). The
reconstituted eNOSox has biopterin content as high as 0.72 per monomer (Table I) and is present as the fully reduced form,
BH4, as analyzed by our HPLC method (data not shown).
The content of both FAD and FMN in purified eNOSred is
essentially stoichiometric based on our HPLC determination against authentic FAD and FMN standards (Table I). The ratios of FAD and FMN to
that of the eNOSred monomer are 1.0 and 1.14, respectively, thus further reconstitution of flavins is unnecessary.
Spectroscopic Characteristics of
eNOSox--
UV-visible spectral analyses from 250 to 700 nm of purified eNOSox showed a Soret peak at 400-404 nm,
81 mM
1cm
1, a broad
/
band
at 518 nm, 15.4 mM
1cm
1, and a
charge-transfer band at 645 nm, 5.8 mM
1cm
1. Treatment with
L-arginine shifted the Soret peak to 396 nm with comparable
amplitude, 82 mM
1cm
1, and only
slight changes in the visible region (Fig.
2). When eNOSox was reduced
by dithionite, the Soret band is red-shifted to 413 nm with a sizable
decrease in intensity, 66.7 mM
1cm
1. The
/
band also
shifted to 552 nm, 13.0 mM
1cm
1,
and the charge-transfer band at about 650 nm was abolished as the lower
lying three metal d-orbitals were completely filled. Further addition
of CO resulted in the hallmark 444-nm Soret band for P450 hemeproteins
with an extinction of 91.3 mM
1cm
1 with the features found
at the visible region very similar to that of ferrous
eNOSox. These spectral behaviors are very similar to our
bacterial-expressed eNOSox and other NOS oxygenase domains (9, 13, 17). The ratio of 280 nm to the Soret peak to either the
resting or L-arginine-treated eNOSox was
~1.5, which is an index of the purity of the hemeprotein and is a
reliable number compared with other NOS preparations (9, 13,
17).

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Fig. 2.
UV-visible spectra of eNOSox
(solid line) and its L-arginine complex
(long dash), ferrous form (medium
dash-dot), and ferrous CO complex (short
dash). The spectra between 500 and 700 nm are enlarged
in the inset. Spectra for a typical preparation were given.
Some eNOSox preparations show a purity index,
i.e. A280/A396
in the L-arginine complex, which is as reliable as
1.3.
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Liquid helium temperature EPR of the resting eNOSox showed
a mixture of high spin and low spin heme structures (Fig.
3, spectrum A). The rhombic
high spin heme has g values of 7.53, 4.23, and 1.83 (the
gmin was only observable at somewhat lower
temperature, ~ 4 K), and the low spin heme show conspicuous rhombic
g values at 2.43, 2.28, and 1.90. Both sets of parameters
are typical for NOS and other P450 type hemeproteins containing a
cysteine thiolate proximal heme ligand (32, 33). Addition of excess
amounts of L-arginine essentially wiped out the low spin
heme signals and substantially increased the high spin heme signals
(Fig. 3, spectrum B). The g values of the high
spin heme shifted to 7.56, 4.17, and 1.82, corresponding to a small
rhombicity shift from 20.6 to 21.1%. On the other hand, imidazole
converted eNOSox to fully low spin heme complex (Fig. 3,
spectrum C). There are two well-resolved rhombic low spin
heme complexes with g values of 2.71/2.29/1.75 and
2.60/2.29/1.81. Resolution in the EPR spectrum of the two imidazole low
spin heme was even better than that found for whole eNOS (33).

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Fig. 3.
EPR spectra of eNOSox
(A), L-arginine complex
(B), and imidazole complex (C).
The concentration for eNOSox was 25 µM in 50 mM HEPES, pH 7.4, with 10% glycerol, 0.1 M
NaCl. L-Arginine was 100 mM and imidazole
was 40 mM. EPR conditions were: microwave
frequency, 9.61 gHz; power, 1 milliwatt; modulation, 10 G; temperature,
11 K; time constant, 0.33 s. Each spectrum is the average of two
scans.
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Stoichiometric Titration of the eNOSox--
To
determine the redox capacity of the purified eNOSox, a
stoichiometric titration was conducted using standardized dithionite solution. L-Arginine was added to make the heme
homogeneously high spin. The spectral conversion from ferric to ferrous
heme during the course of titration appeared to involve only one simple redox reaction as evidenced by the isosbestic points at 410, 489, 532, 615, and 678 nm for both the absolute and difference spectra relative
to the resting eNOSox spectrum. An isosbestic point at 338 nm was slightly perturbed by dithionite whose absorption peaks at 314 nm (Fig. 4, A and
inset). However, the optical amplitude changes at 444-388
nm showed a long lag for 1.5-2.5 (in three separate titrations)
reducing equivalents before a sharp rise. 1-1.5 reducing equivalents
were needed to completely reduce eNOSox (Fig.
4B). The additional 1.5-2.5 reducing equivalents consumed during titration were not due to oxygen contamination as assessed by
lumiflavin titration using the same titration vessel and conditions. Furthermore, similar stoichiometric titrations performed on
eNOSred did not show an initial long lag (see below). The
extra reducing equivalents used to titrate eNOSox could be
due to oxidized biopterin or free sulfhydryl group at the protein
surface. The former may be a consequence of autoxidation of
BH4, and the latter could be due to the loss of the zinc
cluster, which coordinates with two cysteines from each monomer (2, 5,
6). However, the samples used in these titrations are not
BH4-replenished. The content of biopterin was as low as
0.2-0.3/heme and was preset as BH2, which is not reducible
by dithionite. This left the zinc loss as the most possible cause of
the additional consumption of dithionite in the titration. Three
experiments were conducted to assess this hypothesis. Zinc analysis by
ICP-MS analysis was carried out using either eNOSox or
purified whole eNOS. Sufficient amount of zinc was determined by ICP-MS
analysis (data not shown). Gel filtration chromatography was conducted
to determine the population of eNOSox monomer and dimer.
Molecular sieving using five molecular weight standards and purified
eNOSox indicated that the whole population was present as a
dimer with a molecular mass of >100 kDa (Fig.
5). Titration of free thiol by
4,4'-dithiopyridine was also conducted on eNOSox using free
L-cysteine as a positive control. Time-dependent modification of the thiol was monitored in
parallel with urea-treated eNOSox and a bovine
eNOSox predetermined to have zinc and present as a dimer
(Fig. 6) (5). Both yeast-expressed human eNOSox and eNOSox, trypsinolyzed
from bacterial expressed bovine eNOS, exhibited almost identical
kinetics of chemical modification (Fig. 6). Two to three thiol groups
were easily modified in both protein samples at a rate of 0.5 min
1, but the next six residues were modified much slower
at 0.02 min
1. Pre-treatment of 5 M urea
significantly enhanced the extent of chemical modification in the fast
phase as a result of exposure of about additional two to three thiol
groups. The rates of the two phases remain similar, 0.5 and 0.02 min
1, respectively, but the contribution of each phase
shifted from 2:5 to 4:2.5 after urea treatment. Moreover, the overall
extent of modification after a 2-h period remained the same as the
eNOSox sample not treated by urea. In contrast, free
L-cysteine followed simple modification kinetics, and the
rate, 8 min
1, is even faster than the fast phase of that
observed for eNOSox samples.

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Fig. 4.
Stoichiometric titration of
eNOSox by sodium dithionite in the presence of
L-arginine. Purified eNOSox, not
replenished with exogenous BH4, at 18.0 µM
and containing 500 µM L-arginine, was
titrated by 16.2 mM sodium dithionite anaerobically. A
1.8-ml total volume of reaction mixture in 50 mM HEPES, pH
7.5, containing 0.1 M NaCl and 10% glycerol, was used in
the titration. Panel A shows the absorption spectra during
the reductive titration. The inset shows the difference
spectra from panel A (against resting). Panel B
gives the changes of A444 minus
A388 as a function of reducing equivalent per
mol of eNOSox. The solid straight lines indicate
the initial, ending levels and the initial slope of the heme
titration.
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Fig. 5.
Molecular mass estimation by gel
filtration. eNOSox (~1 mg) was gel-filtered through
a Sephacryl 200 HR column (1.5 × 50 cm), the ratios of sample
elution volume (Ve) and column void volume
(Vo) were determined by blue dextran and plotted
with five molecular mass standards: horse heart cytochrome c
(12,400 Da), bovine erythrocytes carbonic anhydrase (29,000 Da), bovine
serum albumin (66,000 Da), yeast alcohol dehydrogenase (150,000 Da),
and sweet potato -amylase (200,000 Da) (solid circles).
The data for eNOSox (solid triangle) was
interpolated into the standard curve to obtain the estimated molecular
mass. There was only one protein peak monitored at
A280 absorbance detected for eNOSox
in the gel filtration profile.
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Fig. 6.
Kinetics of chemical modification for thiols
by 4,4'-dithiopyridine. 50 µM 4,4'-dithiopyridine
was added individually to 4 µM human eNOSox
(solid circles), 4 µM bovine
eNOSox (open circles), 4 µM human
eNOSox pretreated with 5 M urea for 2 h
(cross), and 9 µM free L-cysteine
(open triangles) in 50 mM KPi, pH
7.5. Formation of 4-thiopyridine after reaction with the -SH groups of
each sample was monitored at 324 nm ( = 19.8 mM 1cm 1) on an HP8453
diode-array spectrophotometer for a period of 2 h at room
temperature. The solid lines are one- or two-exponential
fits for each set of kinetic data.
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Tetrahydrobiopterin Radical Formation of
eNOSox--
BH4- reconstituted
eNOSox prepared at a concentration of ~300
µM was premixed with excess L-arginine and
reacted with oxygenated buffer anaerobically at room temperature and
freeze-trapped at several time points. The EPR spectrum corresponding
to a 100-ms reaction time is given in Fig.
7. EPR recorded at 11 K between 200 and
4200 G revealed both the heme component and the radical component (Fig.
7A). The EPR spectrum of a control
L-arginine-treated eNOSox was also recorded
under the same EPR conditions. Two spectra are normalized to the same
concentration of heme. Approximately 50% of the BH4
was converted to the BH4 radical, and other
diamagnetic heme intermediates were estimated from the decrease
of the high spin heme signal amplitude. The radical signal observed at
the g = 2 region was measured again at 115 K (Fig.
7B). The hyperfine features pertaining to nitrogen and
proton splittings are clearly revealed. The biopterin radical was
centered at g = 2.002 and had an overall line width of
39 G. Microwave power dependence indicated a strong magnetic
interaction with the heme center with a P1/2 as high as 14 milliwatts at 120 K (data not shown). Spin concentration was estimated
by double integration of the EPR signal, using a copper standard, to be
~20 µM. After correction for the ~4-fold dilution
factor during rapid freezing, we essentially observed ~80
µM radical, equivalent to about 40% of the total
biopterin because BH4/heme ratio was ~0.7. This radical
EPR could be closely simulated by including one strongly coupled
nitrogen at N5, one alpha proton at N5, and one beta proton at C6
(dashed line in Fig. 7B).

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Fig. 7.
Transient formation of BH4
radical by eNOSox. 280 µM
eNOSox containing 0.7 equivalent of BH4/heme
was first mixed with 1 mM L-arginine then
reduced by dithionite titration in a tonometer with a side-arm
attaching an optical cuvette. This ferrous eNOSox was
reacted with oxygenated buffer at a 1:1 ratio on a rapid-freeze quench
apparatus, the reaction mixture was freeze-quenched in isopentane, and
the ice particles were collected at several reaction times from 20 to
200 ms at a ram velocity of 2.5 cm/s at room temperature by our
specially designed packing device (42). Liquid helium EPR
(A) was recorded for the intermediate trapped at 100 ms
(solid) with a parallel control sample of
L-arginine-treated eNOSox (dash).
EPR conditions were the same as those in Fig. 3. Liquid nitrogen
temperature EPR was recorded in the radical region in A as a
100-G scan. The EPR conditions were: microwave frequency: 9.29 gHz;
power, 1 milliwatt; modulation, 2 G; time constant, 0.33 s; and
temperature, 115 K. The spectrum in B (solid) was
from a single scan. Dashed lines is a computer simulation
using the following parameters: gx = gy = gz = 2.0023; line
width, 12/11/11 G. The
Axx/Ayy/Azz
values: for the nitrogen nucleus, 2/1.5/23 G; for the two hydrogens,
4.6/21.6/11.8 G and 12.4/10.9/14.0 G, respectively.
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Enzyme Activities of eNOSred--
Cytochrome
c reductase activity, DCPIP, and ferricyanide reduction
activities were evaluated for eNOSred expressed in yeast (Table II). Cytochrome c
reductase activity was 70.3 mol/min/mol and was increased about 2- to
3-fold, 194.3 mol/min/mol, in the presence of Ca+2/CaM.
This effect of CaM was less than we previously observed for
baculovirus-expressed eNOSred (17). However, the absolute activity, both in the presence and absence of CaM, are higher than the
values reported for sf9-expressed eNOSred and eNOS,
i.e. 13.8/20.9 and 138/224 without and with
Ca+2/CaM, respectively. Both ferricyanide and DCPIP
reductase activities were greater than that of cytochrome c
reductase. Ferricyanide reductase activity increased from 3220 to 4480 min
1 by Ca+2/CaM, whereas DCPIP reductase
activity was increased from 400 to 800 min
1 by adding
Ca+2/CaM (Table II). Our data are compatible with
literature data for eNOSred or full eNOS (Table II).
Although there are some variations of cytochrome c and
ferricyanide reductase activities among different eNOS or
eNOSred preparations, most of them are within the same order of magnitude (Table II). Differences in assay temperature, expression system, and buffer composition could be the reasons that
resulted in these variations.
Flavin Fluorescence of eNOSred and Its Interaction with
CaM--
The emission spectrum of isolated eNOSred shows a
broad band from 470 to 650 nm peaked at ~528 nm (Fig.
8). The intensity of the fluorescence was
increased by ~30% in the presence of Ca+2/CaM, but there
was no obvious shift of the peak. Excess EDTA could only reverse
~80% of the fluorescence change caused by Ca+2/CaM (Fig.
8). This residual fluorescence increase, which is not reversed by EDTA,
could be simply the slight increase of intensity caused by
Ca+2 alone (data not shown). This fluorescence change
provided a nice index to determine the CaM binding. Titration of
micromolar level of eNOSred with CaM in the presence of
Ca+2 showed a sharp breaking point at 1:1 ratio of CaM to
eNOSred (inset of Fig. 8). This result indicated
that the Kd value of CaM is significantly lower than
micromolar, and each reductase domain binds one CaM.

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Fig. 8.
Calmodulin effect on flavin
fluorescence. Flavin emission spectra between 450 and 650 nm of
1.9 µM eNOSred were recorded before
(solid line) and after addition of 250 µM
Ca2+ and 3.4 µM calmodulin (long
dash) to follow the formation of the CaM·eNOSred
complex and the dissociation of this complex by 3.5 mM EDTA
addition (short dash). A negative control, excluding
eNOSred and CaM was subtracted from the data.
Inset, stoichiometric titration of eNOSred by
CaM monitored by flavin fluorescence change at 530 nm. The intersection
of two straight lines is CaM binding stoichiometry.
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Stoichiometric Titration of eNOSred--
Identical
anaerobic titration procedure as eNOSox titration was
applied here to determine the redox capacity in eNOSred
using dithionite as titrant. A total of four reducing equivalents,
required to fully reduce eNOSred, accounted exactly for the
capacity of two flavin cofactors (Fig.
9). No additional redox centers other than the FAD and FMN were disclosed by this titration. There are three
stages of reduction observed in the dithionite titration. The first
stage took one reducing equivalent, and the electronic spectra showed
isosbestic points (~366 and ~508 nm) in this process (Fig.
9A), indicating a single redox transformation step. The absorbance decrease at 456 nm, and the corresponding increase at 600 nm
are attributed to the formation of a neutral flavin semiquinone. The
second stage took two reducing equivalents. It appeared to show one
isosbestic point (~342 nm) but is not conclusive, indicating that
this stage is likely to be involved in at least two chemical reaction
steps (Fig. 9B). There was a large decrease of 456-nm
absorbance accompanied by a small change at 600 nm. The last stage
involved one-electron reduction to reach the fully reduced state (Fig.
9C). The further bleaching of 456- and 600-nm absorbance
evidenced disappearance of both the oxidized flavins and flavin
semiquinone. The general titration profile is very similar to that
published for microsomal CPR and nNOS reductase domain (43,
44).

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Fig. 9.
Stoichiometric titration of
eNOSred and computer simulations. eNOSred
at 65 µM in 50 mM HEPES, pH 7.5, containing
0.1 M NaCl and 10% glycerol, was titrated with
standardized 30.8 mM sodium dithionite anaerobically.
Panel A shows the absorption spectra (1-5) up to
the addition of 1 reducing equivalent. The spectra of the fully
oxidized flavins (dash-dot-dash) and semiquinone form of FMN
(solid line) are highlighted. Panel B shows the
spectra 5-10 for the titration between 1 and 3 reducing equivalents,
whereas the spectra 10-20 shown in panel C represent the
titration data for addition of the 3-6 reducing equivalents. The
hydroquinone of the second flavin (FADH2) (solid
line) and FADH· semiquinone (dash-dot-dash) are
highlighted in C. Panel D is the plot of
absorbance changes at 456, 508, and 600 nm against the reducing
equivalents consumed per mol of eNOS red. Lines
going through data points of each wavelength are the
simulation obtained as detailed in the main text. The arrows
in A-C indicate the direction of the spectra changes with
increasing dithionite. Another two duplications of titration show very
similar results.
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Computer Simulations for the Reductive Titration Data--
The
data shown in Fig. 9D at three different wavelengths were
simulated by the SCoP program according to Equations 1-12 to obtain three redox potentials gaps,
E1-
E3, between
four half-reactions of two flavins in eNOSred (31).
Computer simulation for the data at 456, 508, and 600 nm was successful
as indicated by the close match of the simulations and the actual data
except the initial <0.3 reducing equivalents (Fig. 9D). The
initial short lag was probably due to the residual amount of oxygen in
the titrator. This simulation process was tested using any arbitrary
absolute midpoint potential value for one of the four half-reactions
and to zoom in the values for
E1-
E3. The
variation for each of the redox potential gaps is not significant as
shown in Table III. The optimal value for
E1 is the largest, 180 mV, and a clean conversion from one oxidized flavin to the semiquinone form was expected in the first stage of titration. In contrast,
E2 was almost zero, indicating that the
second stage of reduction consisted of two almost parallel
half-reactions. The value of
E3, 73 mV, was
in the middle and could be used to estimate the cutoff point for
obtaining the absorbance contribution from only one specific half-reaction. The extinction coefficients at three different wavelengths were also converged by several simulation cycles and are
shown in Table IV. In principal, we could
conduct these simulation cycles on any wavelength between 300 and 700 nm and reconstruct the spectrum for each of the six flavin redox
species.
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Table III
The three redox potential gaps of the four half-reactions of flavins of
eNOSred and related redox systems
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DISCUSSION |
We have successfully prepared both eNOSox and
eNOSred using the yeast expression system. Purified
eNOSox domains appear to have most of the heme
characteristics in intact eNOS. The optical spectra of ferric
and ferrous eNOSox, their ferrous-CO complex, and the spin
state change caused by L-arginine are all typical for eNOS
and eNOSox as we observed previously (17, 21, 24). The
purity index, expressed as the ratio of
A280/A396, was as reliable as 1.3-1.6 compared with 1.5-1.7 found in our previous baculovirus-expressed recombinant protein (17). This ratio is not as
reliable as the recent iNOSox preparation expressed from E. coli (9), but our oxygenase domain does not have the
N-terminal heterogeneity as observed for iNOSox as only one
single peptide band was observed on SDS-PAGE at the ~50-kDa region
(9).
The EPR spectrum of the isolated eNOSox showed a dominant
low spin P450-type heme. Using the low field g = 7.5 signal amplitude to estimate the spin state population against the
L-arginine-treated eNOSox, about 75% of the
heme was present as the low spin form. This high proportion of low spin
heme is at odds with the room temperature optical data (Fig. 2
versus Fig. 3) judging from the similar amplitude of the
charge-transfer band at 650 nm of the resting eNOSox and
eNOSox treated with L-arginine. Optical data implied that the majority of the heme was present as the high spin
form, thus a temperature-dependent heme spin state change likely occurred with the low spin heme electronic configuration as the
ground state. A Truth Diagram analysis for the correlation between heme rhombicity and axial ligand field strength for the low
spin heme component put the eNOSox at the lower right edge of the P zone, indicating a P450-like protein (33). The rhombic distortion (V), axial perturbation (
), and heme
rhombicity (%) obtained were 2.09, 5.75, and 36.4, respectively. There
might be another low spin component present as indicated by the
shoulder at the g = 2.43 signal. However, this
additional component was not as visible as that observed for full eNOS,
and only the low rhombicity low spin heme component appeared to be
present in the isolated eNOSox (33). In contrast, the
imidazole low spin heme complex showed two better-resolved EPR species
than the corresponding imidazole complex of full eNOS (compare Fig. 3,
spectrum C, with Fig. 6a in Ref. 33) with very
similar g values for the two sets of low spin heme
complexes. The calculated values of V,
, and percent
rhombicity for the two low spin species are 2.52/4.53/56% and
2.63/4.05/65%, respectively, and they place these two heme complexes at the center of the P zone. Throughout the entire low spin imidazole·heme complex, there was a reciprocal relationship between the heme rhombicity and the tetragonality, indicating that the
heme rhombic distortion attenuates the axial ligand intensity, possibly
via a lengthening of the Fe-S bond.
The extra 1.5-2.5 reducing equivalents required to initiate the heme
reduction in the stoichiometric titration is puzzling (Fig. 4). These
additional equivalents did not originate from biopterin, because the
amount of pterin was too low to account for this amount of reducing
equivalents and dithionite does not reduce BH2 to
BH4 due to unfavorable redox thermodynamics (35). The zinc
loss is not the reason: there was plenty of zinc present in isolated
eNOSox as assessed by ICP-MS analysis. The isolated eNOSox is a perfect dimer as analyzed by gel filtration.
The monomeric form was not even detected. Furthermore, the titration
for the surface-exposed thiol function group showed identical
modification kinetics as another bovine eNOSox whose
crystallographic data indicate the presence of zinc cluster. Thus the
hypothesis that a zinc loss leading to surface-exposed disulfide
linkage as that found in the iNOSox crystallographic data
does not apply to our isolated eNOSox (7). Furthermore, the
possibility of propensity of zinc loss in recombinant
eNOSox but not in intact eNOS may be unfounded in our yeast
expression system. We do not see additional metal redox centers such as
heme, iron-sulfur cluster, or copper by optical or EPR spectroscopy,
thus leaving us with no explanation for the extra reducing equivalents
that show much higher redox potential than the heme center.
Reduction of the heme center appeared to consume more than one reducing
equivalent (1-1.5 in three experiments). A similar case in
iNOSox was also observed recently (9). The sharp increment of absorbance change at the beginning of the titration and a curvature and even tailing approaching the end of the titration seem to indicate
that the titrant may not have electron-donating power strong enough to
completely reduce all of the heme molecules. Considering the very
negative midpoint potential of thiolate-ligated heme, it is possible in
the later portion of the titration that only part of the reducing
equivalents from dithionite were donated to the heme center dictated by
the midpoint potential difference between the heme and dithionite (36).
We thus put more emphasis in using the initial linear sharp rise to
estimate the end point of titration. By doing this, we get a
stoichiometry closer to 1 rather than 1.5.
The biological activity of our eNOSox was demonstrated by
its capability in forming the biopterin radical (Fig. 7).
We chose this method because it is directly linked to the redox
function of the protein and provides detailed information regarding the reaction mechanism of eNOS. In our study, the radical signal plateaued at ~100 ms at room temperature. The line shape, intensity, and initial kinetics of biopterin radical formation appeared very similar
to those published for iNOSox (26, 27). Computer simulation for the BH4 radical indicates a minimal requirement for one
nitrogen and two proton nuclei to match the EPR data. Because N5 (or
its 4a carbon) is positioned para to the electron-releasing amino group
at C2 and thus has high electronegativity, it is thus more favorable
than N8 to give the first electron. N8 (or its 8a carbon) is positioned
meta to C2 and C4; thus electron withdrawal can only occur via
conjugation with 4-oxo group. The pKa of the N5
proton is much higher than neutral and is not dissociated easily; thus
a hyperfine interaction of this proton with the unpaired electron at N5
is expected. The second proton has to come from the C6 beta proton.
Such initial trial of simulation appears fairly promising. Our observed
biopterin radical is thus likely a BH
·
cation radical (37, 38). Although N8 nuclei and its associated proton
have been proposed to be involve in the unpaired spin system (37), it
remains to be clarified by further spectroscopic studies using isotope
replacement. Nonetheless, we present here the first high quality EPR
spectrum of eNOS biopterin radical and will pursue the mechanistic role
of biopterin using a rapid-freeze EPR approach in parallel with
stopped-flow and other kinetic methods.
eNOSred activity was assessed by three different assays.
The cytochrome c reduction and the DCPIP reduction assays
require the participation from both flavins, and ferricyanide reduction activities were believed to involve only FAD (39). In all cases, CaM
enhanced the activity and the enhancement for cytochrome c reduction and DCPIP reduction activity to a similar extent. Why we only
see a ~3-fold activity increase for the cytochrome
c reductase activity by CaM using our yeast protein and a
10-fold increase in our previous sf9-expressed
eNOSred is unclear (Table II). Nonetheless, CaM appeared to
interact both between FMN and the heme as well as between FAD and FMN
as initially observed in nNOS (40). There are many factors that enhance
the reductase activity of eNOS, including the CaM binding, the removal
of the autoinhibitory peptide, and the phosphorylation of the C
terminus of the reductase (1, 39, 41, 42). Furthermore, the presence of
dithiothreitol, EDTA, and variation in ionic strength during different
stages of the purification also affect the sensitivity of the reductase domain activity to Ca+2/CaM (17, 39, 41, 42). Comparing
eNOSred activity values determined from different
laboratories does not yield straightforward conclusions. The effect of
CaM on nNOSred does not appear to shift the midpoint
potential of either half-reaction of the two flavins (43), because
addition of CaM caused only marginal change for the FMN/FMNH midpoint
potential and essentially no change for the other three
half-reactions.
The only redox centers present in eNOSred are the two
flavins, because exactly 4 reducing equivalents were consumed in the stoichiometric titration (Fig. 9D). Optical changes of the
flavins occurring almost immediately after dithionite addition
contrasts with the data for eNOSox, which required 1.5-2.5
additional reducing equivalents before reduction of heme and supports
that additional redox centers were present in eNOSox (Fig.
4B). In addition to quantifying the redox capacity of
eNOSred, stoichiometric titration also enabled
determination of the relative redox potentials between different
half-reactions as illustrated in this study (Fig. 9D and
Table III). Successful simulation in the data for all three wavelengths, using the same set of difference midpoint potential values, attested to the utility of this approach. The redox
potential gap between the first and second half-reaction was 170-190
mV, thus a complete separation of the first half-reaction from the other three is expected. The optical spectrum at the point of addition
of one reducing equivalent should contain one intact flavin (FAD) and
one flavin semiquinone (FMNH·) (Fig. 9A, heavy
line). The extinction coefficient for the FMNH· semiquinone
could thus be unambiguously determined. The spectral change at 600 nm
is completely due to semiquinone forms of the flavins, because the
fully reduced and fully oxidized samples were silent in this region.
The titration data indicate that the second semiquinone was gradually
reduced to its hydroquinone at the addition of the fourth reducing
equivalent. However,
E3 was only 56-90 mV
and prohibited clean separation of the last half-reaction from the
other three. Because
10
E3/0.059 = (Ox1 · Red2)/(Ox2 · Red1) for the last two overlapping half-reactions,
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we expect that only 70-80% of the reaction after addition of the
fourth electron is only contributed by FADH· semiquinone
reduction to FADH2. Simulation for the 600-nm data here was
very useful to define the extinction coefficient for the second
semiquinone species, because the trapezoidal titration profile is very
sensitive to the difference midpoint potential as well as the
extinction coefficient (31). Thus, simulation greatly assisted
in converging the value of the
E values and the
extinction for the second flavin semiquinone. There was a 20%
difference in the extinction coefficient at 600 nm for these two flavin
semiquinones and almost a 3-fold difference at 456 nm and a difference
of ~50% at 508 nm with the FMNH· having the higher values
(Table IV). The middle two half-reactions, corresponding to
that after addition of the second and third reducing equivalents,
attributable to the formation of FAD semiquinone and FMNH·
transformation into the fully reduced form, were almost equivalent in
redox potential and were titrated together. The absolute values of the
midpoint potential for all four half-reactions will be determined by
potentiometric titration. Once these values are available, they will be
used to validate the
E values obtained in this study and
to refine the accuracy of the extinction coefficient derived herein.
The spectral contribution from each half-reaction at every single
wavelength can be reliably determined and will be useful in future
mechanistic studies using stopped-flow measurements.