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INTRODUCTION |
Soluble guanylate cyclase
(sGC),1 a hemoprotein,
catalyzes the conversion of GTP to cGMP. The enzyme is a major receptor
for NO in cell-cell signal transduction pathways such as neuronal communication and vasodilation (1-7). NO binds to the heme iron of the
enzyme and markedly activates the cyclase reaction (8-12). The
purified enzymes from rat and bovine lung were a heterodimer consisting
of two similar but not identical subunits (12-14). Although the enzyme
has been believed to contain 1 protoheme IX/heterodimer (12, 15-17),
Stone and Marletta found recently the heme stoichiometry of 1.5 hemes/heterodimer (18). Based on the result, they argued that the
stoichiometry was actually 2 hemes/heterodimer, with each subunit
binding one equivalent of heme at a homologous site between two
subunits (18). In this context, it is noted that a site-directed
mutagenesis study raises a new issue for the heme coordination (19).
This mutagenesis study aimed at the identification of a heme axial
ligand strongly suggested that the binding domain of protoheme IX
located only in the smaller subunit, the
-subunit, and that the
histidine residue at the 105-position of the
-subunit was an axial
ligand of the enzyme heme. The ligation of the histidine residue at the
105-position to the heme iron was confirmed by a site-directed
mutagenesis study, in which the
-subunit fragment consisting of
residues 1-385 contained a stoichiometric amount of heme (20), while
the
-subunit fragment with H105A mutation was heme-deficient (21).
Although these results demonstrated that the histidine residue
conserved only among the
-subunits was the heme binding site, there
was no evidence for the heme binding site in the
-subunit.
Stone and Marletta (12, 22) reported that the ferric and the ferrous
enzyme hemes were both in a five-coordinate high spin state.
Furthermore, the ferrous NO complex was demonstrated to be the active
form of the enzyme with five-coordinate NO heme (12, 23, 24). Evidence
for the formation of five-coordinate NO complex was also obtained by
using partially purified, reconstituted enzyme preparations (25, 26).
Hence, the breaking of the heme-proximal ligand bond upon NO binding to
the enzyme heme was proposed to be a trigger for the activation of the
enzyme, as has been hypothesized by Traylor and Sharma (27). The
spectroscopic finding for the ligation of a histidine residue at the
proximal position has been obtained by a resonance Raman study (24).
Deinum et al. (24) assigned the 204-cm
1 Raman
band to the iron-histidine stretching vibration based on analogy with
that of other hemoproteins. This result was a first demonstration for
the weak iron-histidine bond in sGC, but the assignment of the
Fe-histidine vibration was incomplete for lack of an isotope shift
experiment such as 57Fe substitution.
Although a resonance Raman spectroscopy was an important spectroscopic
probe for the analysis of properties of the iron-proximal base bond,
the use was usually limited to ferrous high spin enzyme heme. In
contrast, an EPR technique provided structural information for the
metal-axial base bond of six-coordinate ferrous NO hemoproteins (28-32) and of Co2+ porphyrin-substituted derivatives of
hemoproteins (33-37). The Co2+ derivatives that were
proved to retain close structural homology to the native iron
hemoproteins by x-ray crystallography (38, 39) were frequently used to
provide EPR response instead of EPR-silent ferrous hemoproteins. In the
present work, we aimed to identify a histidine residue as the proximal
ligand of the heme and to examine the nature of the metal-histidine
bond of sGC by an EPR method. To take advantage of a EPR spectroscopy as a structural probe, we have prepared the six-coordinate NO complex
and the Co2+-substituted derivative of sGC. The EPR
findings provided firm spectroscopic evidence for the histidine
ligation at the proximal position of metal. Moreover, we found that the
EPR features of the NO complex and the Co2+ derivative
closely resembled those of the corresponding form of
-subunits of
hemoglobin, showing the presence of tension on the iron-histidine bond.
We also obtained EPR evidence that the N3
binding to the ferric enzyme heme
formed presumably a five-coordinate high spin
N3
heme.
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EXPERIMENTAL PROCEDURES |
Enzyme Purification--
Fresh bovine lung (4 kg) was minced and
homogenized with a Waring blender in 12 liters of 50 mM TEA
buffer, pH 7.6, containing 1 mM phenylmethylsulfonyl
fluoride, 1 mM benzamidine, 1 mM EDTA, and 55 mM mercaptoethanol (buffer A). Throughout the purification, these protease inhibitors and mercaptoethanol were included in the
buffer unless otherwise stated. After the homogenate was centrifuged at
13,500 × g for 20 min, 1.2 kg of DEAE-cellulose A-500
(Seikagaku Kogyo) equilibrated with buffer A was added to the
supernatant. The slurry was stirred for 1 h at 4 °C, and the
resin was collected by sedimentation. Subsequently, the resin was
washed three times with buffer A and was poured into a column. The
enzyme was eluted with a 3.5-liter linear NaCl gradient of 0-0.35
M in buffer A. After active fractions were pooled, solid
ammonium sulfate (0.29 g/ml) was added. The precipitate collected by
centrifugation at 10,000 × g for 15 min was dissolved
in 200 ml of 40 mM potassium phosphate buffer, pH 7.6, containing 55 mM mercaptoethanol. The sample was washed
with the phosphate buffer by using a Minitan system (Millipore Corp.)
to remove ammonium sulfate. Then the enzyme was applied to the column
of Matrex Blue A (Amicon) equilibrated with 40 mM phosphate
buffer described above and eluted with a linear gradient of 0-1
M KCl gradient. The pooled enzyme was applied to a ceramic
hydroxylapatite column (Bio-Rad). The enzyme was eluted by increasing
phosphate concentration from 0 to 0.45 M at pH 7.6 containing 55 mM mercaptoethanol and the protease
inhibitors except for EDTA. The concentrated sample was further
purified with a Superdex 200-pg HPLC column (Amersham Pharmacia
Biotech). Then fractions with the cyclase activity were applied to an
80-ml column of GTP-Sepharose 4B with a 12-atom spacer attached through ribose hydroxyl (40). The column was exhaustively washed to remove
contaminated proteins with 25 mM Tricine-NaOH buffer, pH 7.6. The enzyme was eluted with a 1-liter gradient running from 0 to 0. 15 M NaCl. The fractions with a specific activity over 8000 nmol/min/mg of protein in the presence of NO were pooled. Then the
sample was finally purified to an apparently homogenous state with a
Protein Pak G-DEAE HPLC column (Waters). The overall yield was about
10%. The resultant homogenous enzyme was stored in 50 mM
TEA buffer, pH 7.6, containing 10% glycerol and 5 mM DTT
at
80 °C until use.
Co2+ Protoporphyrin IX Substitution--
Ignarro
et al. (17) have reported a method to prepare the apoenzyme
by lowering pH to 5.7. We attempted to prepare the apoenzyme by this
method, but the recovery of the Co2+-substituted enzyme was
very low at the final purification step. We tested the heme depletion
as a function of pH and found that the heme in sGC was depleted by the
DEAE cellulose chromatography under alkaline conditions. In brief, the
supernatant fraction of homogenized tissue described above was adjusted
to pH 8.5, and DEAE cellulose A-500 equilibrated with 50 mM
TEA buffer at pH 8.5 containing protease inhibitors was poured to the
supernatant. The enzyme was eluted by a linear gradient of 0-0.35
M NaCl. The fractions with cyclase activity that was
assayed in the presence of protoporphyrin IX were further purified by
GTP-agarose and Superdex 200-pg columns under the conditions described
above. The apoenzyme was pooled and reconstituted with Co2+
protoporphyrin IX under anaerobic conditions. The remaining
purification steps were the same as those used for the native enzyme
purification. The enzyme Co2+ porphyrin-substituted by our
method exhibited essentially the same optical and EPR spectral
properties as the Co2+-substituted enzyme, which was
obtained by the method of Ignarro et al. (17).
Spectral Measurements--
Absorption spectra were recorded with
a Shimadzu MPS-2000 or a Perkin-Elmer Lamda 18 spectrophotometer at
room and subzero temperatures. The temperature of the cuvette holder
was controlled with thermomodule elements. The buffer systems used were
50 mM TEA buffer (pH 7.6) containing 5% glycerol and the
same buffer containing 40% ethylene glycol for room and subzero
temperature measurements, respectively. Other details were described in
the figure legends.
EPR spectra were measured on a Varian E-12 X-band EPR spectrometer with
100-kHz field modulation. An Oxford flow cryostat (ESR-900) was used
for liquid helium temperature measurements. The microwave frequency was
calibrated with a microwave frequency counter (Takeda Riken, model TR
5212), and the magnetic field strength was determined by the nuclear
magnetic resonance of water protons. Accuracy of g values was ±0.01 in
the low magnetic field and ±0.005 in the high field. Other details
were as described elsewhere (41).
NO complexes for EPR measurements were prepared in buffer containing
5% glycerol at
5 °C or in buffer containing 40% ethylene glycol
at
24 °C as follows. The enzyme solution was transferred to a
septum-capped EPR tube and flushed with oxygen-free argon gas for 10 min. Then NO gas previously washed with 1 N NaOH or an
aliquot of SNAP solution was introduced to the tube with a gas-tight
syringe. The formations of NO complexes in five- and six-coordinate
states were ensured by directly measuring the optical spectrum of the
sample in the EPR tube at
5 or
24 °C.
The ferric enzyme was prepared by adding a 2-fold excess of
ferricyanide to the DTT-free ferrous enzyme, where DTT in the enzyme
solution was removed by a Superdex 200HR (Amersham Pharmacia Biotech)
HPLC column. For EPR measurements, the residual ferricyanide was freed
of the solution by passing through a Superdex 200HR HPLC column. The
ferric enzyme was converted to N3
complexes by adding a desired amount of NaN3. The EPR
spectra of ferric N3
complex were
measured at 5 or 15 K.
Kinetic Measurements--
The NO binding to the ferrous enzyme
was analyzed by a Photal stopped flow spectrophotometer, model RA-401,
equipped with a photodiode array detector. The buffer solution in
reservoirs was bubbled with oxygen-free argon for 10 min, and then the
catalytic amount of glucose oxidase and catalase and 2 mM
glucose was added to assure anaerobic conditions. The enzyme and an
aliquot of NO-saturated solution were then added to the buffer solution
under a constant stream of argon.
Resonance Raman Measurements--
The resonance Raman spectra
were measured with a JASCO NR-1800 spectrometer equipped with a liquid
nitrogen-cooled CCD detector (Princeton Instruments). Excitation
wavelengths were 413.1- and 406.7-nm lines from a Krypton ion laser
(Coherent, Innova 90). Calibration of the Raman spectrometer was
performed by using indene.
Activity Measurements--
The enzyme activity during the
purification was measured in a reaction mixture containing 2 mM GTP, 5 mM DTT, 3 mM
MgCl2, and an appropriate amount of the enzyme solution in
a total volume of 0.5 ml of 40 mM TEA buffer, pH 7.4. When
desired, 1 mM isobutylmethylxantine was added to inhibit
phosphodiesterase activity. The reaction was started by the addition of
0.2 mM SNAP and conducted at 37 °C for 10 min. The
reaction was terminated by the addition of 20 µl of 30% acetic acid.
The mixture was centrifuged for 10 min at 15,000 rpm, and cGMP was
quantitated with a C18 HPLC column at a constant flow rate of 1 ml/min
of 40 mM potassium phosphate buffer, pH 6.0, containing
10% methanol.
The activity of the homogenous enzyme was assayed under the same
condition as that described above, except that the concentration of GTP
was increased to 4 mM. The activation by NO was performed in a septum-capped sample tube. The assay mixture in the tube was
flushed with a purified argon, and the reaction was started by the
addition of 30 µl of saturated NO solution with a gas-tight syringe.
For the activation by CO, the reaction mixture was saturated with CO
gas prior to the addition of the enzyme.
Electrophoresis--
Reducing SDS-polyacrylamide gel
electrophoresis was carried out by using 9% acrylamide running gel.
Protein was visualized with a silver stain method (Daiichi Chemical
Co.).
Reagents--
GTP and cGMP were purchased from Wako Pure
Chemical Industries. Research grade NO was obtained from Takachiho
Chemical Co. S-Nitroso-N-acetyl-DL-penicillamine
was purchased from DOJINDO or ALEXIS. Other chemicals, purchased from
Nacalai Tesque Co., were of highest commercial grade and were used
without further purification.
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RESULTS |
Properties of Native Enzyme--
The homogenous enzyme exhibited a
basal activity of 98 nmol/min/mg of protein at 37 °C in the presence
of Mg2+. The activity increased to 26,811 nmol/min/mg of
protein upon the addition of NO, while it increased to 650 nmol/min/mg
of protein upon the addition of CO. An addition of protoporphyrin IX
(2.4 µM) slightly increased the activity (580 nmol/min/mg
of protein). The enzyme preparation stimulated by a combined addition
of protoporphyrin and NO exhibited an activity (25,560 nmol/min/mg of
protein) rather lower than the NO-stimulated activity. The
SDS-polyacrylamide gel electrophoresis analyses indicated that the
enzyme was a heterodimeric protein consisting of an
-subunit of 75 kDa and a
-subunit of 71 kDa. The enzyme contained 0.97 ± 0.04 protoheme IX/heterodimer, in which protein was determined by the
modified biuret method of Yonetani (42) using bovine serum albumin as
the standard, and the heme content was determined by the pyridine
hemochromogen method (43). When protein was determined by the Bradford
protein assay, essentially the same heme stoichiometry was obtained
(0.95 ± 0.03). The cyclase activity per heme, defined as turnover
number (µmol of cGMP/min/µmol of heme) was 3800 min
1
at 37 °C in the presence of NO.
Detection and Characterization of Six-coordinate Ferrous NO
Complex--
The ferric enzyme exhibited a Soret band at 390 nm
(
mM 103), which was indicative of the
five-coordinate high spin state (data not shown). The ferrous enzyme,
prepared by adding a slight excess of
Na2S2O4 to the ferric enzyme under
anaerobic conditions, exhibited the Soret maximum at 431 nm
(
mM 105). The optical spectrum of the enzyme
reduced by Na2S2O4 was identical
with that of the ferrous enzyme as isolated in the presence of
mercaptoethanol. An anaerobic addition of SNAP or NO to the ferrous
enzyme yielded the five-coordinate NO complex with the Soret maximum at
400 nm (
mM 75). These optical and EPR spectral properties of the ferric enzyme (see Figs. 6 and 8) agreed with previous results (18, 20) but significantly differed from the
result obtained by using a partially purified reconstituted enzyme
(25). Deinum et al. (24) pointed out that the
partially purified reconstituted enzyme preparation had a different
heme environment from the native enzyme.
When the ferrous enzyme was mixed with SNAP at
24 °C in the
presence of 40% ethylene glycol used as an antifreeze, a new spectral
species with a sharp Soret band at 419 nm and 544- and 579-nm bands in
the visible region was produced (Fig.
1A). The peak positions
closely agreed with those of the six-coordinate NO complex of
hemoglobin (44), suggesting that the new species was six-coordinate
ferrous NO complex. By raising the temperature to
15 °C, the
species fully converted to the five-coordinate ferrous NO complex with
400-nm Soret maximum, giving clear isosbestic points (Fig.
1A, inset).

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Fig. 1.
Optical and EPR spectra of six-coordinate
14NO complex. A, an optical spectrum of the
six-coordinate NO complex of sGC. The six-coordinate NO complex was
prepared by adding 100 µM SNAP to the ferrous enzyme (2.1 µM as heme) at 24 °C in 50 mM TEA, pH
7.6, containing 40% ethylene glycol. Inset, the degradation
of the six-coordinate ferrous NO complex to the five-coordinate ferrous
NO complex after raising the temperature to 15 °C. B,
EPR spectrum of the ferrous NO complex (16 µM as heme).
The EPR spectrum illustrated in trace a was that
of the fully five-coordinate NO complex, which was prepared by adding
NO under anaerobic conditions at 5 °C in 50 mM TEA
buffer, pH 8.6, containing 40% ethylene glycol, and that in
trace b was that of the six-coordinate NO complex
prepared by adding SNAP under anaerobic conditions at 24 °C in 50 mM TEA buffer, pH 8.6, containing 40% ethylene glycol. In
the spectrum of trace b, the presence of the
six-coordinate NO species at g = 1.979 besides that of the
five-coordinate one was noted. The EPR signal of the six-coordinate NO
complex, which was obtained by subtracting the three-line signal of the
five-coordinate NO complex from spectrum b, was shown in
trace c. The EPR measurements were done at 35 K
and at microwave power of 10 milliwatts. NOA is the
coupling constant for hyperfine splitting by 14NO.
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The coordination state of the new species was examined by an EPR method
at 35 K. When the new species, prepared in EPR tube at pH 7.6 and
24 °C, was immediately frozen by immersing into liquid nitrogen,
the EPR signal of the new species was negligibly small in the spectrum,
where the three-line signal of five-coordinate NO complex was
predominant. We thought that the new species was rapidly degraded to
the five-coordinate NO complex upon freezing. The examination of
stability of the new species as a function of pH at
24 °C revealed
that the species was more stable at pH 8.6 than at pH 7.6. As expected,
the EPR spectrum of the species prepared at pH 8.6 displayed a new EPR
signal at g = 1.979 besides the three-line signal of the
five-coordinate NO complex (b in Fig. 1B). The
new EPR species was not a modified form of the five-coordinate NO
complex produced by the effect of pH or the binding of antifreeze, since the changes in pH from 7.6 to 8.6, the changes in the antifreeze from ethylene glycol to glycerol, or changes in the concentration of
antifreeze did not alter the EPR spectrum of the five-coordinate NO
complex. The EPR signal of the new species was obtained by subtracting
the three-line signal of the five-coordinate species (a in
Fig. 1B) from the spectrum of trace b.
The resultant spectrum (c in Fig. 1B) was typical
of a six-coordinate ferrous NO complex as indicated by a triplet
superhyperfine splitting of 14NO in the central resonance
signal around g = 2. This was the first clear identification of a
six-coordinate ferrous NO complex of sGC. Superhyperfine structure of a
triplet of triplets in the gz region, which was indication
of the ligation of axial ligand with 14N nucleus, a
histidine residue (29), was unclear in the spectrum due to the low
signal quality.
The formation of the six-coordinate NO complex was examined at room
temperature by monitoring the absorbance at 400, 419, and 430 nm under
stopped flow conditions (Fig.
2A). It was particularly noted
that the magnitude of the absorbance changes at 419, 400, and 430 nm
was different from that expected. For instance, if the five-coordinate
NO complex is assumed to be directly produced in the reaction between
the ferrous enzyme and NO, the absorbance decrease at 430 nm must be
much larger than the increase at 400 nm, indicative of the formation of
the five-coordinate NO complex. However, the absorbance change at 430 nm was smaller than that at 400 nm (Fig. 2A). Furthermore,
the absorbance decrease at 419 nm was unexpectedly large. The most
reasonable interpretation was that the six-coordinate NO complex was
produced within a dead time of the apparatus (about 2.5 ms), and then
converted to the five-coordinate NO complex. To confirm the formation
of the six-coordinate NO complex within a dead time of the apparatus,
the reaction was analyzed by a rapid scan spectrophotometer (Fig.
2B). The spectrum taken at 4 ms after mixing agreed with
that of the six-coordinate NO complex shown in Fig. 1A and
was converted to that of the five-coordinate NO complex with the Soret
band at 400 nm through one set of isosbestic points.

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Fig. 2.
NO binding analyses by a stopped flow
method. A, time course of NO binding to ferrous sGC
measured by the stopped flow method. In this experiment, 1.8 µM DTT-free ferrous enzyme was mixed with 60 µM NO at 15 °C. Absorbance changes were monitored at
400, 419, and 430 nm. Inset, pH dependence of the conversion
rate from the six-coordinate NO to the five-coordinate NO complex was
illustrated. B, spectral changes in the ferrous sGC followed
by a rapid scan spectrophotometer after mixing with NO. The DTT-free
ferrous enzyme (2.4 µM) was mixed with 60 µM NO, and spectra were recorded with 4-ms gate time at
times indicated in the figure at 15 °C. For comparison,
the spectrum of the ferrous enzyme (Fe2+) is illustrated.
In these experiments, 50 mM TEA buffer, pH 7.6, containing
5% glycerol was used, and DTT was omitted to avoid the reaction
between NO and DTT.
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The time course at 419 nm (Fig. 2A) obeyed that of a first
order reaction with a rate constant of 38 s
1. The
conversion rates did not show significant pH-dependent
changes between pH 7.0 and 8.6 at 15 °C (Fig. 2A,
inset), being different from the above described results at
24 °C in the presence of ethylene glycol. The finding that the
six-coordinate NO complex formation was completed within the dead time
indicated that the binding rate of NO to the ferrous enzyme,
i.e. the formation rate of the six-coordinate NO complex,
was much faster than 1 × 107
M
1 s
1. These results together
were the first clear evidence of the formation of the six-coordinate NO complex.
Co2+ Protoporphyrin IX-substituted
Enzyme--
Apoenzyme used for the reconstitution with
Co2+ protoporphyrin IX exhibited the basal and
NO-stimulated activities of 38 and 165 nmol/min/mg of protein,
respectively. The following titration experiments indicated that the
apoenzyme preparation retained a correct binding site for protoheme IX.
When the apoenzyme was titrated with protoheme under anaerobic
conditions maintained by the addition of a slight excess of
Na2S2O4, the absorbance at 431 nm
as well as the cyclase activity increased with an increased amount of
protoheme, giving a clear inflection point. At the point the cyclase
activity reached a plateau (data not shown), with the NO-stimulated
activity of 2550 nmol/min/mg of protein. The basal activity of the
reconstituted enzyme was 16 nmol/min/mg of protein. The resultant
reconstituted enzyme exhibited an optical spectrum essentially
identical to that of the native enzyme. Similar results were also
obtained when titrated with Co2+ protoporphyrin IX. The
Co2+-substituted enzyme further purified by the method of
the previous section exhibited retention times identical to those of
the native enzyme when analyzed by a Superdex 200HR column and a
Protein Pak G-DEAE HPLC column (data not shown). The results indicated that the Co2+-enzyme had the same metal binding site as
that of native enzyme, and had essentially identical molecular mass and
protein surface charges to those of the native enzyme. The
Co2+-enzyme exhibited a specific activity of 8600 nmol/min/mg of protein in the presence of NO. Since an attempt to
purify it to a homogenous state was unsuccessful, the comparisons of
activity between cobalt- and iron-enzymes were done using a turnover
number defined as µmol of cGMP/min/µmol of heme or cobalt
porphyrin. The turnover number of the partially purified
Co2+-substituted enzyme was 5840 min
1 in the
presence of NO, which was about 1.5-fold higher than that of native
enzyme (3800 min
1). The activation of the
Co2+-substituted enzyme by NO was about 50-fold, which was
significantly low when compared with the native enzyme (270-fold). The
lower activation of the Co2+-substituted enzyme by NO was
attributable to the high basal activity of 115 min
1,
which was about 8-fold higher than that of native enzyme (14 min
1).
The partially purified Co2+-substituted enzyme showed the
Soret band at 405 nm and the visible band at 559 nm (Fig.
3A), which were nearly
identical to those of Co2+-substituted myoglobin (45). The
shoulder absorption around 430 nm marked by an asterisk was
attributed to the contamination of the native iron-enzyme by the
pyridine hemochromogen assay. The content was estimated to be less than
15%. The addition of NO to the Co2+-enzyme slightly
blue-shifted the Soret band to 399 nm and red-shifted the visible band
to 569 nm (Fig. 3B). The spectral pattern of the NO complex
was entirely different from that of Co2+-myoglobin NO
complex in a six-coordinate state, which exhibited the Soret band at
421 nm and 539- and 577-nm bands in the visible region. These results
suggested that the NO complex of Co2+-substituted sGC was
in a five-coordinate state. The coordination state was confirmed by a
resonance Raman spectroscopy as described below.

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Fig. 3.
Optical absorption spectra of
Co2+ protoporphyrin IX-substituted sGC. A,
optical spectrum of Co2+-substituted enzyme under anaerobic
conditions at 5 °C. B, optical spectrum of the NO complex
of Co2+-substituted enzyme at 5 °C. The NO complex was
prepared by adding NO under anaerobic conditions. In these experiments,
50 mM TEA buffer, pH 7.6, containing 5% glycerol and 5 mM DTT was used.
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The resonance Raman spectra of the Co2+-substituted enzyme
and the NO derivatives were summarized in Fig.
4, A and B. The
Co2+-enzyme exhibited the
4 and
3 Raman bands at 1371 cm
1 and at 1504 cm
1, respectively, which closely agreed with those of
Co2+-myoglobin and -hemoglobin in a six-coordinate state
(46). The addition of 14NO to the
Co2+-substituted enzyme shifted the
4 band
to 1376 cm
1 from 1371 cm
1 with an
appearance of the Raman band at 1682 cm
1. The resonance
Raman spectrum was markedly different from those of 14NO
complexes in a six-coordinate state (46). The replacement of
14NO by 15NO eliminated the Raman band at 1682 cm
1 with a concomitant appearance of the
1648-cm
1 band and without detectable shift of other bands
(Fig. 4A, c and d). In the low
frequency region, we detected the shift of 523 cm
1 band
upon the replacement of 14NO with 15NO (Fig.
4B, b and inset). These results
indicated that the 1682- and 523-cm
1 bands were assigned
to the NO stretching vibration (
N-O) and Co-NO vibration
(
Co-NO), respectively. Both
N-O and
Co-NO values agreed with those of the corresponding
vibration of five-coordinate NO complexes of Co2+ model
porphyrins (48) but not of six-coordinate NO complexes (47). These
findings indicated that the NO complex of Co2+-substituted
sGC was in a five-coordinate state.

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Fig. 4.
Resonance Raman spectra of Co2+
protoporphyrin IX-substituted sGC. A, high frequency
resonance Raman spectra of Co2+ porphyrin-substituted
enzyme and its NO complexes. The spectra of the
Co2+-enzyme, Co2+-14NO complex, and
Co2+-15NO complex are illustrated in
a, b, and c, respectively. The
difference spectrum between 14NO and 15NO
complexes (14NO 15NO) is presented in
d. B, low frequency resonance Raman spectra of
Co2+ porphyrin-substituted enzyme (trace
a) and the 14NO complex (trace
b). The difference spectrum between 14NO and
15NO is shown in the inset. These spectra were
taken at 406.7 nm excitation wavelength. The buffer used in these
experiments was 50 mM TEA buffer, pH 7.6, containing 5%
glycerol and 5 mM DTT.
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The EPR spectrum of Co2+-substituted enzyme was shown in
Fig. 5A, with that of
Co2+-myoglobin for comparison. The
Co2+-substituted enzyme exhibited five-coordinate low spin
signals at g
= 2.37 and g
= 2.04 with
poorly resolved eight-line hyperfine splitting due to 59Co
nucleus (CoA
= 7.4 mT). The g
= 2.37 component was significantly broad compared with that of
Co2+-myoglobin. The hyperfine splitting constant
(CoA
= 7.4 mT) agreed with that of other
Co2+-substituted hemoproteins with proximal histidine (34,
37), suggesting the presence of a histidine residue as the proximal ligand in sGC. However, the triplet superhyperfine splitting due to the
14N nucleus of the axial ligand was not well resolved in
Fig. 5A. To gain firm evidence for the histidine ligation,
EPR signals with 20-mT sweep width centered at 300 mT were accumulated
to obtain a high quality spectrum. As shown in Fig. 5B, we
could detect the three-line superhyperfine splitting due to the
14N nucleus (Na = 1.7 mT). Thus, the EPR
signal of Co2+-substituted sGC was characterized by a
poorly resolved triplet splitting of the 14N nucleus and
the relatively broad g
component. These features resembled those of the
-subunit in Co2+-hemoglobin
tetramer in T-state (34) rather than that of the
-subunit of
Co2+-hemoglobin in T-state (34),
Co2+-myoglobin, or Co2+-horseradish peroxidase
(35).

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Fig. 5.
EPR spectra of Co2+
protoporphyrin IX-substituted sGC. A, EPR spectrum of
Co2+ protoporphyrin IX-substituted sGC (Co sGC)
at 35 K (upper trace). The EPR spectrum of
Co2+ protoporphyrin IX substituted myoglobin (Co
Mb) is shown for comparison (lower trace).
B, accumulated EPR spectrum of Co2+
protoporphyrin IX-substituted sGC between 0.29 and 0.31 mT. In this
experiment, 40 scans were averaged. The buffer used in these
experiments was 50 mM TEA buffer, pH 7.6, containing 10%
glycerol and 5 mM DTT. Na and CoA
denote coupling constants for hyperfine splitting by 14N
nucleus of proximal base and 59Co nucleus,
respectively.
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Azide Complex--
As shown in Fig.
6A, the addition of
N3
to the ferric enzyme caused a
decrease in the intensity of the Soret band and a remarkable increase
in the intensity of the 635-nm band at room temperature, confirming the
previous result (22). These spectral changes were unusual, because the
N3
addition to other ferric
hemoproteins such as metmyoglobin produced 420- and 540-nm low spin
bands and reduced the 640-nm band intensity (49). The low spin bands in
the metmyoglobin N3
complex were
intensified by lowering the temperature, showing that the spin state
was in a thermal spin equilibrium between low and high spin states (50,
51). In contrast, the N3
complex of
sGC did not display the spectral change by lowering the temperature to
77 K (data not shown). This indicated that the ferric heme of the
N3
-bound sGC was in a high spin state,
not in a thermal spin equilibrium.

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Fig. 6.
Optical and resonance Raman spectra of the
ferric azide complex. A, the optical absorption
spectrum of ferric sGC in the absence and presence of azide (50 mM). The spectra were taken at 5 °C, and the buffer used
was 50 mM TEA buffer, pH 7.6, containing 5% glycerol.
B, the resonance Raman spectra of ferric sGC with or without
50 mM of azide in 50 mM TEA buffer, pH 7.6, containing 5% glycerol. The spectra were taken at 406.7-nm excitation
wavelength.
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The ferric enzyme exhibited Raman bands assignable to the oxidation
state marker at 1371 cm
1 (
4), the
coordination marker at 1492 cm
1 (
3), and
2 at 1568 cm
1 (Fig. 6B). The
result showed that the coordination and spin states of the enzyme heme
were categorized to ferric five-coordinate high spin heme (52). The
addition of N3
only intensified and
sharpened the coordination marker band (
3) at 1492 cm
1 without detectable shifts of the other bands,
indicating that the coordination state of the ferric enzyme heme
remained unchanged upon the addition of
N3
. Although we tried to detect the
ligation of N3
to the enzyme heme
through the detection of Fe-N3
stretching vibration, it was unsuccessful.
The N3
addition markedly enhanced the
cyclase activity of the ferric enzyme, but not the ferrous enzyme (Fig.
7). The activation reached a maximum at
50 mM N3
and gradually
decreased with an increase in the N3
concentration (Fig. 7). The reason for the activity decrease over 50 mM N3
was unknown. The
maximum activity expressed as turnover number (µmol of
cGMP/min/µmol of heme) was 970 min
1, which corresponded
to the specific activity of 9100 nmol of cGMP formed/min/mg of protein.
This was about one-third of the specific activity of the ferrous NO
complex. The result was a first observation for the activation of the
enzyme in the ferric state. The addition of 10 mM KCN
markedly inhibited the activity in the presence of 50 mM
N3
, but the inhibitory effect was
significantly diminished by increasing the
N3
concentration to 150 mM
(Fig. 7). This strongly suggested that N3
and cyanide combined to the same
site, i.e. the ferric enzyme heme.

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Fig. 7.
Activation of ferric enzyme by azide.
Assays were performed in the reaction mixture containing 2 mM GTP, 3 mM MgCl3 and a desired
amount of N3 in a total volume of 0.5 ml of 40 mM TEA buffer, pH 7.4. The reaction was started by
the addition of an appropriate amount of the ferric enzyme and
conducted at 37 °C for 10 min. The reaction was terminated by the
addition of 20 µl of 30% acetic acid. The amount of cGMP was
determined by the method described under "Experimental Procedures."
The enzyme activity was expressed as turnover number (µmol of cGMP
formed/min/µmol of heme). As shown in the figure, a final
concentration of 10 mM KCN was added to the assay solution
containing 50 or 150 mM
N3 . The activities measured in the
presence of KCN are indicated by closed squares. The
activities of the ferrous enzyme in the presence of
N3 were measured in the above reaction
mixture supplemented with 5 mM DTT. Each data point is the
average value of triplicate determinations.
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The oxidized resting enzyme exhibited only rhombic high spin signals
with g values of g1 = 6.62, g2 = 5.36, and
g3 = 1.98 at 5 K (Fig. 8,
trace A). The EPR spectrum was not altered by raising the temperature to 15 K (not shown), indicating that the ferric
enzyme heme did not contain a low spin component. The result essentially agreed with the previous report measured at 20 K (22). These results with resonance Raman data confirmed that the ferric enzyme heme in sGC was in a single coordination and in pure high spin
state. The addition of 50 mM
N3
produced two types of new high spin
species with EPR signals of g'1 = 6.94 and g"1 = 6.10 at 5 K (Fig. 8, trace B). To estimate the amount of
N3
-bound heme in the presence of 50 mM N3
, the spectrum of
trace A divided by some factors was subtracted from the spectrum of trace B. As illustrated in
the spectrum B'), the subtraction of the spectrum divided by
2 satisfactorily eliminated the residual unreacted ferric enzyme,
indicating that a half of the enzyme combined
N3
ion. The reason why the
N3
addition yielded two types of high
spin species remained unclear. These new high spin signals were
intensified by increasing the N3
concentration to 150 mM (Fig. 8, trace
C). The EPR spectrum at 15 K (Fig. 8, trace
D) was essentially identical with that at 5 K, indicating
that the N3
complex did not contain
low spin components.

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Fig. 8.
EPR spectra of ferric sGC and its ferric
azide complex. Trace A, EPR spectrum of the ferric
enzyme at 5 K; trace B, EPR spectrum of the
ferric N3 (50 mM) complex
at 5 K; trace B', difference spectrum
(trace B 0.5 × trace
A); trace C, spectrum of the ferric
N3 (150 mM) complex at 5 K; trace D, spectrum of the ferric
N3 (150 mM) complex at 15 K. EPR spectra were taken at a microwave power of 5 milliwatts and by
100-kHz modulation with 0.5-mT width, and the enzyme concentrations
were 75 µM as heme. Throughout these EPR measurements, 50 mM TEA buffer, pH 7.6, containing 10% glycerol was
used.
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DISCUSSION |
Stone and Marletta (53) have reported the mechanism for the
five-coordinate NO complex formation. They proposed that NO first
combined with the ferrous heme to produce a six-coordinate NO complex,
which then converted to a five-coordinate NO complex. The conversion
was a complex process, which was interpreted by assuming two
populations of the enzyme with different activities for NO. About 30%
of the population of the heme rapidly converted to the five-coordinate
NO complex from the six-coordinate one via a single exponential pathway
(20 s
1 at pH 7.4 and 10 °C), while the conversion of
the remaining 70% of the population was very slow and depended on NO
concentration. The slow NO-dependent conversion of the
latter major population was explained by assuming an unidentified
nonheme iron binding site of NO on the protein. However, these
experimental findings were inadequate to argue the formation of a
six-coordinate NO complex and inconsistent with our results in some
points. In the present work, we obtained definite evidence to show the
formation of a six-coordinate NO complex in the reaction (Figs. 1 and
2). In contrast to the above findings by Stone and Marletta, our data showed that the entire population of the six-coordinate NO complex rapidly converted to the five-coordinate NO complex through a single
exponential pathway (38 s
1 at pH 7.6 and 15 °C).
Furthermore, the conversion rate was almost unchanged at the lower
concentration of NO (10 µM). Our results did not
contradict an interpretation that the enzyme heme was the sole binding
site for NO. One might claim that the discrepancy for the kinetic
results was due to the difference in the heme content. The 1.5-heme
stoichiometry reported by Stone and Marletta (18) was estimated by the
precise protein determination, in which the protein content obtained by
the Bradford protein assay was corrected by a factor of about 1.6 based
on the quantitative amino acid analyses; i.e. the Bradford
protein assay overestimated the amount of protein. When the correction
was not made, the heme content in their preparations essentially agreed
with the 1-heme stoichiometry obtained by the present and the recent
studies (54). Thus, the difference was considered apparent, although
efforts to obtain the precise amount of protein were not made in this and other studies (54). Whatever the reason for the discrepancy, our
data including the EPR and resonance Raman studies agreed with a view
that the enzyme heme in our preparation was a single population with a
single coordination structure and definitely indicated that the
five-coordinate NO complex was produced via the formation of the
six-coordinate NO complex.
The optical and resonance Raman spectra of the
Co2+-reconstituted enzyme presented here differed
considerably from the results of Dierks et al. (48),
especially for the NO complex. They reported the Soret absorption
maximum at 390 nm of the NO complex, but the corresponding complex in
our preparation exhibited the maximum at 399 nm. The discrepancy might
be attributed to the difference in the heme environment. Indeed, Deinum
et al. (24) pointed out that the apoenzyme obtained by
Dierks et al. had a different environment from the native
enzyme. By using the apoenzyme preparation with a correct heme binding
site (see "Results"), we demonstrated that the
Co2+-substituted enzyme formed a five-coordinate NO complex
with a high cyclase activity. Hence, the coordination structure of the active NO complex was essentially the same irrespective of whether the
central metal of porphyrin was iron or cobalt.
The iron-histidine stretching frequency of sGC essentially agreed with
that of
-hemes of hemoglobin in T-state (55), implying a tension to
pull the proximal histidine from the porphyrin plane in sGC (24). EPR
characterization of native and Co2+-enzymes also revealed
the similarity in metal environment between
-subunits of hemoglobin
and sGC. The EPR spectrum of the six-coordinate NO complex of native
sGC differed from those of horseradish and cytochrome c
peroxidases with an anionic proximal histidine residue but closely
resembled those of hemoglobin and myoglobin with a neutral proximal
histidine residue. Among the latter hemoproteins, the six-coordinate NO
complex of
-subunits of hemoglobin (31, 32) exhibited an EPR
spectrum similar to that of sGC with the paramagnetic center of rhombic
symmetry. The Co2+-substituted enzyme also exhibited an EPR
signal analogous to
(Co) subunits in Co2+-hemoglobin
tetramer in T-state with the broad g
component and the
poorly resolved 14N superhyperfine splitting (33), where
the Co2+-proximal histidine bond in
(Co) subunits was
reported to be more tensioned than that of
(Co) subunits (56). The
present data revealed that the nature of the metal-histidine bond in
sGC was strikingly analogous to that of
-subunits of hemoglobin and provided additional evidence for tension on the metal-proximal histidine bond of sGC proposed by a resonance Raman study (24).
Heme iron was reported to lie about 0.4 Å out of the porphyrin plane
toward the proximal side in a five-coordinate ferrous high spin state
and to move into the porphyrin plane upon NO ligation, yielding an
~0.4-Å movement of the iron atom from the initial position (57-60).
The movement upon NO ligation might impose further tension on the
iron-proximal histidine bond in sGC but did not cause the immediate
cleavage of the proximal bond, as demonstrated by the formation of the
six-coordinate NO complex with the proximal histidine (Figs. 1 and 2).
There were several factors to facilitate the cleavage of the
iron-histidine bond, including a repulsive trans effect of NO (27, 61)
and the protonation of the proximal histidine residue (62, 63). The
latter was proposed for the proximal histidine release of myoglobin and
peroxidases at acidic pH. This possibility might be excluded, because
the rate for the release was almost unchanged between pH 7.0 and 8.6 (Fig. 2A, inset), and the pK value
higher than 8.6 might be unlikely for the imidazole group. The
repulsive trans effect of NO on the proximal histidine, therefore, was
concluded to be an important driving force for the proximal histidine
release, as has been hypothesized by others (24, 27). The resultant
five-coordinate NO complex possibly retained the iron atom displaced
from the porphyrin plane toward NO, as shown for the iron-porphyrin
complexes with five-coordinate NO structure (64). The displacement of
the iron atom toward NO might stabilize the five-coordinate NO complex
by preventing the access and the rebinding of the proximal histidine
residue to the heme iron. Thus, the tension imposed by the in plane
iron movement and the repulsive trans NO effect are crucial for the iron-proximal histidine bond cleavage of sGC upon NO ligation.
The overall movement of metal associated with the formation of
five-coordinate NO complex significantly differed between
Fe2+ and Co2+ porphyrins. The overall movement
from the initial position was ~0.35 Å for cobalt atom and ~0.6 Å for iron atom, which were assessed from the results of model porphyrin
complexes and myoglobins (39, 64-66). If the estimation is valid for
the Co2+-substituted sGC, one may assume that the 0.35-Å
movement is sufficient for breaking the metal-histidine bond in
Co2+-substituted enzyme upon NO binding. Thus, it was
predicted that the proximal base bond of the
Co2+-substituted enzyme was more readily broken than in the
native iron-enzyme, and the instability of the bond correlated with the higher basal activity in the Co2+-enzyme.
It has been known that N3
activated
sGC in the presence of exogenously added catalase and DTT under aerobic
conditions (16). In this case, catalase oxidized
N3
to NO using
H2O2 produced by autoxidation of DTT, resulting
in the NO complex formation of sGC. In the present study, DTT and catalase were not exogenously added. Under these conditions, we found
that the ferric enzyme was capable of catalyzing cGMP production in the
presence of N3
. The optical and EPR
spectral studies revealed that the N3
complex was in the five-coordinate high spin state (Figs. 6 and 8). The
five-coordinate N3
heme has been
proposed for the high spin N3
complex
of carp hemoglobin in T-state with inositol hexaphosphate (67), and
McCoy and Caughey (68) have found the similarity between the infrared
N3
stretching frequency of high spin
N3
complex of hemoglobin and that of
the five-coordinate N3
protoheme
complex. Although we could not obtain a clear indication for the
formation of the six-coordinate N3
heme in the reaction between the ferric enzyme and
N3
, the formation of the
six-coordinate one was assumed important as a trigger for the release
of the proximal histidine in sGC; the formation moved the
Fe3+ atom into the porphyrin plane from the position
initially displaced toward the proximal side as reported for model
compounds (69, 70). The movement probably imposed further tension on
the proximal histidine bond of sGC. Consequently, the six-coordinate
N3
heme might experience the release
of the proximal histidine, yielding a small fraction of a
five-coordinate N3
heme with in plane
configuration of the iron atom. The five-coordinate N3
heme once formed would be
stabilized by the iron displacement from the porphyrin plane toward the
N3
ion. These considerations led to a
reasonable conclusion for why CO and cyanide complexes of sGC were
practically inactive; to our knowledge it has not been known that both
CO and cyanide complexes formed the stable five-coordinate complex with
the iron displacement from the porphyrin plane toward the corresponding ligand.2 It is emphasized
that our results provide the first clear experimental evidence for the
five-coordinate high spin N3
heme, no
matter whether or not the mechanism for the proximal histidine release
is correct.