From the Department of Chemistry, Jackson State
University, Jackson, Mississippi 39217 and the ¶ Department of
Biochemistry, University of Illinois, Urbana, Illinois 61801
Received for publication, September 16, 2002, and in revised form, December 9, 2002
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ABSTRACT |
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The heme active site structure of
chloroperoxidase (CPO), a glycoprotein that displays versatile
catalytic activities isolated from the marine mold Caldariomyces
fumago, has been characterized by two-dimensional NMR
spectroscopic studies. All hyperfine shifted resonances from the heme
pocket as well as resonances from catalytically relevant amino acid
residues including the heme iron ligand (Cys29)
attributable to the unique catalytic properties of CPO have been firmly
assigned through (a) measurement of nuclear Overhauser effect connectivities, (b) prediction of the Curie
intercepts from both one- and two-dimensional variable temperature
studies, (c) comparison with assignments made for cyanide
derivatives of several well characterized heme proteins such as
cytochrome c peroxidase, horseradish peroxidase, and
manganese peroxidase, and (d) examination of the crystal
structural parameters of CPO. The location of protein modification that
differentiates the signatures of the two isozymes of CPO has been
postulated. The function of the distal histidine (His105)
in modulating the catalytic activities of CPO is proposed based on the
unique arrangement of this residue within the heme cavity. Contrary to
the crystal state, the high affinity Mn(II) binding site in CPO (in
solution) is not accessible to externally added Mn(II). The results
presented here provide a reasonable explanation for the discrepancies
in the literature between spectroscopists and crystallographers
concerning the manganese binding site in this unique protein. Our study
indicates that results from NMR investigations of the protein in
solution can complement the results revealed by x-ray diffraction
studies of the crystal form and thus provide a complete and better
understanding of the actual structure of the protein.
Chloroperoxidase is a monomeric glycoprotein (42 kDa) secreted by
the mold Caldariomyces fumago (1). Like most members of the
peroxidase superfamily, CPO1
contains an iron protoporphyrin IX moiety (heme b; Fig.
1) as its prosthetic group and shares
major common reaction intermediates with other heme peroxidases (1).
However, extensive biochemical and biophysical studies carried out on
this enzyme have revealed dramatic structural and catalytic differences
between CPO and traditional heme peroxidases. For example, the axial
ligand to the heme iron in CPO is a cysteine (Cys29)
sulfur atom rather than a histidine nitrogen atom commonly found in
most heme peroxidases (2-5). Furthermore, CPO employs a glutamic acid
(Glu183) as the distal acid-base catalyst, whereas most
other heme peroxidases use a histidine to fulfill the same function
(6).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Schematic presentation of different
conformations of iron protoporphyrin IX in peroxidases (CPO and HRP
(A) and CcP and MnP
(B)).
The unique active site structure of CPO dictates a broad spectrum of catalytic activities such as oxidation of organic substrates (peroxidase activity) (7), dismutation of hydrogen peroxide (catalase activity) (8, 9), and monooxygenation of many organic molecules (monooxygenase activity) (10-13). Furthermore, CPO has a unique ability to utilize halide (except fluoride) ions to halogenate a wide variety of organic acceptor molecules in the presence of hydrogen peroxide or other organic hydroperoxides (14-16). Most importantly, CPO is adept in catalyzing the stereoselective epoxidation of alkenes (10, 17, 18), hydroxylation of alkynes (19, 20), and oxidation of organic sulfides (21-23).
The versatile catalytic activities of CPO have attracted much interest in understanding the structural properties of the enzyme. Especially, the increasing current interest in chiral synthesis has made CPO an attractive candidate for making important chiral synthons that are of both industrial and medicinal significance. Therefore, detailed structural insight into this structurally unique and catalytically diverse heme enzyme would help to further understand the structure-activity relationship of heme proteins in general and the structural basis for the broad range of activities displayed by CPO in particular.
Many chemical and spectroscopic techniques are now available for structural investigations of heme proteins. Among them, NMR spectroscopy and x-ray crystallography represent the most powerful methods for high resolution structural characterization of paramagnetic metalloenzymes. The two methods are complementary in most cases, and it is difficult to determine which one is better. Despite the great success with cytochrome c peroxidase in the early 1980s (24, 25), x-ray crystallography of heme peroxidases has suffered from difficulties in obtaining suitable diffracting protein crystals. Nonetheless, the solid-state structure of CcP has served as a convenient and independent structural basis for evaluating results derived from NMR studies of the protein in solution. Consequently, CcP has served as a prototype model for NMR spectroscopists interested in hyperfine resonance assignment and structural refinement of the protein in solution, a state that is more closely related to the physiological conditions under which the enzyme functions (26-34).
Compared with the extensive and in depth NMR studies on CcP (28-36) and horseradish peroxidase (34, 37-53), relatively few NMR studies of CPO have been reported (54-58). Furthermore, the most powerful NMR approach, the two-dimensional NMR technique that has led to the unambiguous assignment of major hyperfine-shifted signals in a number of heme peroxidases (59), has not been applied to the investigation of CPO. As a result, no extensive and definitive resonance assignments are available for this structurally unique yet functionally diverse enzyme.
Here we report the first application of the two-dimensional NMR method
to the elucidation of the active site structure of CPO in solution. The
observation of both COSY and NOESY connectivities among
paramagnetically shifted signals as well as NOESY connectivities between hyperfine shifted resonances and signals within the crowded diamagnetic envelope, in combination with the Curie intercepts obtained
from variable temperature experiments coupled with previous one-dimensional NOE studies performed on this enzyme (54) have allowed
us to assign most of the signals from the heme group, which in turn
allows the assessment of the conformations of the heme side chains. Of
particular importance, the two-dimensional studies have allowed firm
assignment of the heme iron ligand, Cys29 spin system that
is critical to the unique catalytic properties of CPO. Surprisingly,
the addition of excess Mn(II) to CPO resulted in no detectable effects
on the NMR spectral properties of the protein. This is in sharp
contrast with the results observed for CPO isolated from cultures grown
in the presence of manganese (60) and from manganese (II) binding
variants of CcP (36, 61, 62) and native MnP (63). The
difference between solution and solid-state structural features of the
same protein demonstrates the need for structural characterization
using NMR to complement x-ray structural analysis.
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EXPERIMENTAL PROCEDURES |
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Sample Preparation-- Chloroperoxidase was isolated from the growth medium of C. fumago according to the method established by Morris and Hager (64) with minor modifications using acetone rather than ethanol in the solvent fractionation step. Protein preparations with Rz values of 1.4 or higher were used in all experiments.
Protein samples for NMR experiments were prepared in either
D2O buffer or 90% H2O, 10% D2O
buffer solutions containing 100 mM potassium phosphate at
pH 5.5 (direct meter readings from an Orion 720A pH meter using a
standardized calomel combination microelectrode uncorrected for any
isotope effects). Samples in D2O were prepared by at least
five isotope exchanges of the protein solution in H2O with
D2O buffered at pH 5.5. The isotope exchanges were carried out in either Centricon or Centriprep tubes (both from Amicon, Inc.) at
4 °C. All NMR samples contain ~1.5 mM protein as
determined by electronic absorption spectroscopy for the Soret
absorbance at 398 nm and the reported absorption coefficient of 91,200 M1 cm
1 (65). The cyanide
adducts of the protein were prepared by the addition of a 10-20%
molar excess of cyanide from a freshly made 500 mM stock
solution of KCN in 99.9% D2O.
UV-visible Titrations-- Manganese titration experiments were performed on a Hewlett-Packard 8453 diode array spectrophotometer. CPO was dialyzed against 100 mM KH2PO4, pH 5.5, and 10 mM EDTA twice overnight, followed by three dialyzes against 100 mM KH2PO4, pH 5.5, to remove EDTA from the sample. CPO (16 µM) is placed in both the sample and reference cuvettes. Under gentle stirring, aliquots of 50 mM MnSO4 solution were added to the sample with 2, 3, 4, 5, 10, 30, 40, 50, and 60 equivalents of Mn2+ per enzyme equivalent. At the same time, an equal amount of buffer was added to the reference cuvette to compensate for any dilution effects. UV-visible spectra were obtained in the range of 250-750 nm.
EPR-- EPR experiments were carried out on a 95-GHz (W-band) spectrometer at room temperature. Spectra were obtained on ~2 mM protein samples in 100 mM KH2PO4, pH 5.5. For Mn(II) binding studies, CPO was first dialyzed against buffer containing 10 mM EDTA twice overnight. After removal of excess EDTA, Mn(II) was titrated into CPO under gentle stirring at 4 °C. The samples were then subjected to repeated dilution and concentration in a Centriprep concentrator to remove any free and adventitiously bound Mn(II). Instrument settings used for the experiments were as follows: microwave frequency = 95 GHz, modulation amplitude = 32.4 G, and microwave power = 1.00 milliwatts.
NMR Spectroscopy-- Proton NMR spectra of both native and cyanide-bound forms of CPO were recorded at 25 °C on a Varian Unity 600 FT NMR spectrometer operating at a proton frequency of 599.97 MHz. The residual solvent signal was suppressed with either the super WEFT method (66) or presaturation during relaxation delay. Chemical shift values were referenced to the residual HDO signal at 4.76 ppm.
Variable temperature experiments were carried out on a Varian
Unity-Inova 500 FT NMR spectrometer operating at a proton frequency of
499.77 MHz. The reference chemical shift of the residual HDO signal was
calculated according to the relationship of T =
25
0.012 (T
25), where
T is the chemical shift of HDO at temperature T
in °C, and
25 is the chemical shift of HDO at 25 °C
(67). A value of 4.76 ppm rather than 4.81 ppm (67) was used for
25 to match the previous NMR studies of CPO (54).
Phase-sensitive NOESY spectra for the cyanide-bound derivative of CPO were acquired at 25 °C with mixing times ranging from 1.5 to 35 ms. Typical NOESY spectra were collected with 256 experiments in the F1 dimension using the hypercomplex method of States et al. (68). In general, 400 scans were accumulated for each F1 experiment, which was acquired with 4096 complex points in the F2 dimension over a spectral width of 27 or 60 kHz. The residual solvent signal in all NOESY experiments was suppressed using a 200-ms presaturation with a weak decoupler power. NOESY spectra with mixing times of 3 ms or less were collected with the incorporation of the super WEFT sequence (66) to suppress the intense diamagnetic signals from the protein matrix and the residual signal from the solvent.
Clean TOCSY (69) spectra of CPOCN were recorded at both 500 and 600 MHz over different spectral windows using 4096 F2 points and 256 complex F1 points of 320-400 scans. Solvent suppression was achieved by a 200-ms direct saturation during the relaxation delay period. Various mixing times (2, 10, 30, and 40 ms) were used to allow effective spin lock for protons with different relaxation properties.
All two-dimensional data were processed on a Dell Dimension 8200 PC with a Pentium 4 processor using Felix 2001 (Accelrys, Inc.). Various apodization functions were employed to emphasize protons with different relaxation properties. For example, apodization over 256, 512, and 1024 points was used to emphasize fast relaxing broad cross-peaks at the expense of resolution, whereas apodization over 2048 points is necessary to emphasize slowly relaxing cross-peaks. All two-dimensional data were zero-filled to obtain 2048 × 2048 matrices as required by the large hyperfine shift dispersion exhibited by the paramagnetic nature of this protein.
The structure of CPO was examined on either a Silicon Graphics Indigo 2 Extreme workstation using Quanta (Accelrys) or a Dell Dimension 8200 computer using ViewerLite (Accelrys) to visualize the crystal
coordinates supplied by the Brookhaven Protein Data Bank (6).
Generally, atom separations are reported as distances between protons
of interest with the exception of methyl protons, where methyl carbons
are used for distance measurements.
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RESULTS |
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The proton NMR spectrum of the native ferric high spin CPO (data not shown) is essentially identical to the results reported previously (55, 57). Because of the high spin nature of the native CPO, considerably broad resonances are observed in the NMR spectra that provide only limited information about the structural properties of the enzyme. Therefore, no further efforts were made to analyze the spectra of native CPO in this study. We have focused on the NMR spectral properties of the cyanide-bound, ferric low spin derivative of CPO. This protein form, although not physiologically active, has been the most favorable system on which paramagnetic NMR investigations are carried out (35, 37, 70, 71). The short electronic relaxation times and large magnetic anisotropy of the low spin peroxidase cyanide complexes give much sharper and better resolved signals in their proton NMR spectra, providing much more information about the electronic, magnetic, and molecular structural properties of the heme pocket as compared with the native, high spin resting forms (27, 38, 50, 72). In addition, the cyanide adduct of heme peroxidases has been implicated as an important analogue for the active, oxidized low spin enzyme intermediates for which proton NMR spectroscopy is currently inapplicable due to the large resonance line widths (73).
Although the CPO preparations used in this study were
spectrophotometrically homogeneous, the 1H NMR spectra of
the cyanide-bound CPO complex are NMR spectroscopically inhomogeneous
due to the presence of two isozymes. This is reflected by the pairwise
pattern for most of the hyperfine-shifted signals as shown in Fig.
2. The approximately equal intensities of
the two sets of signals suggest that the ratio of A and B isozymes is
close to 1 in the current enzyme preparation. The spectral features are
equivalent to that reported previously (54, 57). No noticeable solvent
isotope effect on the chemical shifts of heme protons was observed when
spectra were recorded in H2O (Fig. 2, lower
trace). This is anticipated, since the distal acid-base catalyst in CPO is a glutamic acid (6) rather than a histidine as in
other heme peroxidases (24, 74-76). The proton/deuteron exchange on
the N atom of the distal histidine has been shown to be responsible
for the observed solvent effect on heme resonances in several heme
peroxidases (36, 53, 72).
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The spectral features of CPOCN display a high degree of similarity to
that of other heme peroxidase cyanide derivatives (59). The four
intense signals with integrated intensities of three protons each in
the downfield region are typical of heme methyl groups. They have been
tentatively assigned to the two heme methyls (5- and 1- or 8- and
3-CH3) for the two isozymes (54). Other resonances with
intensities of one proton each in the downfield region represent
protons from other heme substituents and those from amino acid residues
in the proximal and distal heme pocket. The resolved upfield spectral
region displays several single-proton resonances and a few multiproton
signals. Previous work on both heme model compounds and a number of
heme peroxidases have firmly concluded that this spectral region
encompasses the resonances from -protons of the heme vinyl and
propionate groups as well as those from some of the amino acid residues
near the heme center (53, 72, 77-79). The chemical shifts and the
corresponding diamagnetic shift values predicted from Curie plot as
well as the spin-lattice relaxation times for the hyperfine shifted
resonances and their assignments in CPOCN are compiled in Table
I, along with the corresponding
parameters in MnPCN (72, 80) and HRPCN (51) reported previously.
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The assignment of the hyperfine shifted signals for the CPOCN complex
was achieved through comparison with the assignments made for cyanide
derivatives of other heme peroxidases (30, 34, 36, 53, 72) and
examination of the active site structure of CPO revealed by its crystal
structure (6) with confirmation through one- and two-dimensional NOE
measurements as well as through bond connectivities (COSY). Shown in
Fig. 3 is the NOESY spectrum of CPOCN
collected in D2O buffer with a mixing time of 35 ms. The
clear NOESY connectivities and the results from scalar (COSY; Supplemental Fig. S1) connectivities prove the validity of previous NOE
connectivities observed in one-dimensional NOE experiments (54) and
lead to the proposed assignments for all but three of the
nonexchangeable hyperfine shifted protons (resonances A, B, and Z) from
the heme active site in CPOCN. It should be noted that the observed
cross-peaks in the COSY experiment are not necessarily true coherence
peaks due to the large molecular weight of CPOCN (81, 82). Therefore,
the suggested assignments are further verified by the Curie intercepts
predicted from the temperature dependence of the resonances
(Supplemental Fig. S2).
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Fig. 4 shows the NOESY spectrum collected
with a mixing time of 1.5 ms. The clear NOE connectivity between the
two fast relaxing, nonexchangeable, strongly hyperfine shifted
resonances firmly establishes the geminal partner relationship of the
two protons. Because of the extremely short T1
values (1.5 ms) of these two protons, previous efforts to correlate the
two signals by one-dimensional NOE method were unsuccessful (54).
Nonetheless, based on the relaxation properties, the large contact
shift, and the expected distance to the heme iron center, these two
resonances were tentatively assigned to the -CH2 protons
of the heme iron ligand, Cys29 (54). The crystal structure
of CPO (6) and the observed NOESY cross-peaks (Fig. 4) between these
signals now confirm the earlier proposals (54). This is another example
demonstrating that NMR spectroscopy can serve as an independent tool in
characterizing the structural properties of large paramagnetic heme
proteins.
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The structure of metal binding sites, such as Ca2+ and
Mn2+ binding sites, within peroxidase molecules has
received increasing attention in recent years due to its potential to
modulate specific activities of this class of enzymes. This has
stimulated great interest in designing metal binding sites in
peroxidases (36, 61, 62). The crystal structure of CPO (6) has revealed a Mn(II) binding site near the heme group similar to the location of
Mn(II) in MnP as shown in Fig. 5 (76).
However, the exact role of Mn(II) in CPO is unclear. A careful NMR
titration of the CPOCN preparation used in these experiments with
Mn(II) failed to find any detectable effect of Mn(II) as shown in Fig.
6. The results are surprisingly different
from that observed for the Mn(II) binding site in native MnP (63) and
an engineered MnP mimic MnCcP (36, 61, 62). The absence of
any detectable effect of the added Mn(II) on the spectral properties of
CPOCN undoubtedly depends on the growth conditions used to produce the enzyme. C. fumago cultures grown in complex media
bind one manganese ion per molecule (60). However, when the fungus is
grown on synthetic media in the absence of manganese, the enzyme
contains little or no manganese.
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To confirm the NMR studies of Mn(II) binding, both UV-visible
spectrophotometric and EPR spectroscopic Mn(II) titrations were performed (data not shown). These measurements supported the conclusion from NMR studies.
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DISCUSSION |
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Presence of Isozymes
Both one- and two-dimensional NMR spectra of CPOCN (Figs. 2 and 3) display pairwise resonances for most of the paramagnetically relaxed protons, indicating the presence of two structurally and magnetically closely related isozymes of CPO in solution. This is in complete agreement with previous one-dimensional NMR study of this protein (54, 57). The two isozymes are not due to heme disorder or chemical equilibrium between different forms of the same protein. Instead, the isozymes are chemically and structurally distinct, as judged by the observed NOE pattern and the lack of chemical equilibrium between them. This is different from the situation for recombinant CcP variants where multiple forms of the same protein were found to exist in equilibrium as reflected by exchange peaks in the NOESY spectrum and temperature-dependent NMR resonance intensity changes among them (28, 29, 36, 83).
It is interesting to note that not all of the hyperfine shifted resonances display corresponding partners in the NMR spectra. In fact, only signals assignable to the heme group and heme iron ligand show an observable doubled peak pattern as shown in Figs. 2 and 3. This fact, combined with the very similar chemical shifts of the two isozymes, suggests that there are no global structural differences between the two distinguishable protein forms. It seems that the magnetic inequivalence is particularly localized to a specific region of the heme active site, as will be further discussed under "Resonance Assignment."
Resonance Assignment
The Heme Substituents-- As shown in Fig. 3, the heme methyl resonances C at 24.0 ppm and D at 23.8 ppm give weak NOEs to peak J at 14.1 ppm, peak K at 12.9 ppm, and peak "O" at 11.6 ppm. Several other NOEs are also observed in the NOESY spectrum in the diamagnetic aliphatic region, attributable to amino acid residues close to this methyl group. Of critical importance to the assignment of these signals is the observation of the strong NOE connectivity between peaks J and O that are indicative of a geminal relationship between these two signals. Such NOE patterns are consistent with the results reported from the earlier one-dimensional work (54) and are typical of a methyl group near a propionate group as observed for CcPCN (31, 35), MnCcPCN (36), and HRPCN (40). Therefore, resonances C and D can be assigned to the 5- or 8-CH3 group of the two isozymes of CPO.
The other set of heme methyl signals E and F at 20.7 and
20.4 ppm produces an extremely weak NOE to resonance I at 14.8 ppm and
weak NOEs to signals at 2.7 ppm (V) and
4.4 ppm (X). Clear NOEs are
also observed between resonance I and the signals V and X. This is
reminiscent of the NOE pattern observed for the engineered MnP model,
MnCcP (36), and several other heme peroxidases (31, 34, 35,
72, 84). The lack of strong NOE between peak I and any other signals
suggests the absence of geminal partners to this proton. Therefore,
resonance I must be assigned to the
H of a vinyl group, and signals
E and F must be assigned to a nearby methyl group of the two isozymes.
Consequently, signals V and X must be assigned to the
-protons of
the same vinyl group. This assignment is further supported by the
strong temperature dependence and the reasonable Curie intercepts of
these resonances (Supplemental Fig. S2a and Table I). Furthermore, the
proposed assignment is consistent with the expected chemical shift
pattern of a heme vinyl group. The
-CH vinyl proton resonance is
normally observed in the downfield region because of
-spin
delocalization of the unpaired spin density, whereas the
-CH2 vinyl proton peaks are usually positioned between 0 and
5.0 ppm (36, 54, 71, 85).
Since signals E and F display NOE connectivity to signal I assignable
to the H of a vinyl group (see above), the methyl peaks E and F at
20.7 and 20.4 ppm must arise from either the 1-CH3 or the
3-CH3 group of the heme unit. The observed resonance E (F)
for CPOCN is assigned to the 3-CH3 group, since peak E (F) also gives NOE connectivity to signal T at
1.4 ppm. The latter signal
also displays a strong NOE to signal Y at
5.2 ppm. Both signals T and
Y display clear NOEs to resonance P at 3.0 ppm. This NOE pattern and
the observed shift positions coupled with the Curie behavior of these
signals suggest the assignment of signals P, T, and Y to the protons of
a vinyl group other than the vinyl group mentioned before. The presence
of dipolar connectivities to two vinyl groups mandatorily assigns
signal E (F) to 3-CH3, since it is the only candidate that
can display such dipolar connectivities (Fig. 1). The observed NOE
pattern between the 2-vinyl group (resonance T) and 3-CH3
(resonance E (F)) indicates that the
H of the 2-vinyl group is
pointing away from the 3-CH3 group, and the
H of the vinyl group is pointing toward this methyl group as shown in Fig. 1A. This is in complete agreement with the results observed
for cyanide-bound low spin lignin peroxidase (84) and HRPCN (50, 51).
This result is also in perfect agreement with the results obtained from
crystallographic analysis of the CPO structure (5, 6). The position of
the 2-vinyl group shown in Fig. 1B would produce observable
NOE between the
H (usually under the crowded diamagnetic envelope
region) and the 3-CH3 group as in the case of
CcPCN (33, 35), MnCcPCN (36), and MnPCN (72).
With resonance E (F) assigned, signals at 14.8 (I),
2.7 (V), and
4.4 (X) ppm observed in the NOESY spectrum can be readily assigned to
4-H
, 4-H
-t, and 4-H
-c, respectively, according to the
magnitude of NOEs displayed (Fig. 3), the intensity of the scalar
cross-peaks observed in the COSY experiment (Supplemental Fig. S1), and
the temperature dependence displayed in the Curie plot (Supplemental Fig. S2a and Table I). Consequently, peak B (C) at 24.0 (23.8) ppm must
originate from the 8-CH3 group, and the NOE observed at
14.1, 12.9, and 11.6 ppm must arise from the 7-propionate H
. The
7-propionate H
signals produce the expected reciprocal NOE to the
assigned 8-CH3 as well as strong NOEs to signals at 2.0 (1.9) and 2.8 (2.7) ppm assignable to signals of the two
-protons of
the 7-propionate group. The temperature dependence behavior of these
signals is strikingly similar to the corresponding protons in MnPCN
(72) and HRPCN (51), further supporting the assignment proposed. This
assignment is also supported by the observation of NOEs between
7-propionate group and H
2 of His105 (see distal heme
cavity assignment).
It is worth mentioning that although most heme resonances display a
doubled-peak pattern due to the existence of two isozymes of this
protein, the chemical shift of most protons differs only slightly from
their counterparts between the two isozymes. For example, the chemical
shift of the two resolved heme methyl resonances display differences of
only 0.2 and 0.3 ppm for the two isozymes. However, the resonance
positions of the 7-propionate -CH protons (signals J and K at 14.1 and 12.9 ppm; signals O and K at 11.6 and 12.9 ppm) in the two isozymes
exhibited a difference of ~1.2 ppm. The large chemical shift
difference of the 7-propionate
-protons suggests that the protein
modification differentiating the two isozymes is probably located in
the vicinity of this propionate group or is communicated to the heme
via this side chain. Further support for this implication comes from
the fact that the two 7
-protons in one isozyme are observed as an
essentially degenerate pair at 12.9 ppm (signal K), whereas the
corresponding protons in the other isozyme are detected as chemically
and magnetically distinctive groups at 14.1 and 11.6 ppm (signals J and
O). It is interesting to note that the position of the degenerate
7
-protons in one isoenzyme (form B) is exactly at the middle of the
distinctive 7
-protons in the other isoenzyme (form A). This could
suggest that the degeneracy of the two 7
-protons is caused by a
factor that averages the otherwise magnetically inequivalent protons.
Previous studies on CPOCN have suggested the assignment of signals G(H) and W to the heme mesoprotons based on their extremely short T1 values of 6-7 ms (54). The predicted distances of these signals to the heme iron center is about 4.5 Å (Table II) according to the relationship between T1 values of inequivalent protons in noncoordinating groups and their distances to the paramagnetic center (50), assuming that heme methyl protons have an average distance of 6.1 Å from the iron. Therefore, it is reasonable, in the absence of the crystal coordinates of CPO, to assign these signals to the heme mesoprotons, since they are the only remaining protons close enough to heme iron to experience such sufficient paramagnetic relaxation (54). Unfortunately, no NOEs are observed between these signals and any of the assigned heme signals in the current study, suggesting different identities for these protons. The relatively small diamagnetic intercepts of these signals as compared with that of the heme mesoprotons in MnPCN and HRPCN (Table I) also argue for different assignments for these protons. Furthermore, the crystal structure of CPO reveals that there are several nonheme protons within the heme cavity that can experience sufficient paramagnetic relaxation due to their close proximity to the iron (Table II).
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Careful examination of the NOESY map shown in Fig. 3 leads to the
possible assignment of two heme mesoprotons in CPOCN. These are the
- and
-mesoprotons that resonate at 5.8 and 8.7 ppm, respectively. However, the essentially non-Curie behavior of the latter
resonance (8.7 ppm) excludes the assignment of this signal to heme
mesoprotons. The strong temperature dependence, the observed shift
value, the extrapolated Curie intercept (8.4 ppm, Table I), and the NOE
between the assigned 4-H
vinyl proton allow us to assign the signal
at 5.8 ppm as the heme
-mesoproton of CPOCN.
The Heme Thiolate Ligand Cys29--
The extremely
broad, fast relaxing, strongly hyperfine-shifted peaks A (B) and Z at
39.0 (38.3) and 20.7 ppm are characteristic features of protons in
close proximity to the heme iron. These peaks have been previously
proposed to arise from the
-CH2 protons of the
coordinated cysteine, Cys29, based on their relaxation
properties and shift positions (54). However, due to the considerably
broad line widths and the extremely short T1
values of these resonances, no observable NOE was detected in the
one-dimensional NOE work that could firmly relate the geminal relationship between the two protons (54). The clear NOE observed in
the current two-dimensional study (Fig. 4) unequivocally confirms the
earlier hypothesis and definitively establishes the geminal partner
relation of these two resonances. The similar Curie intercepts of these
signals as compared with that of the corresponding protons in MnPCN and
HRPCN (Table I) also support the assignment of these signals to the
-CH2 protons of the coordinated iron ligand,
Cys29. The crystal structure of CPO (6) further supports
this assignment. With distances of 3.13 and 4.10 Å from the heme
iron, respectively, the two
-CH2 protons of
Cys29 are expected to display strong contact shift and
effective paramagnetic relaxation as predicted previously (54). The
firm assignment of the Cys29
H is hindered by the
unsymmetrical nature of the NOESY map obtained. Nonetheless, the NOE
displayed between peak A (B) and a signal at 4.4 ppm (Fig. 4) suggests
the assignment of the latter signal to the Cys29
H. This
is expected, since the extremely short mixing time needed to detect the
NOEs between fast relaxing protons would fail to detect NOEs between
nongerminal protons. The detection of NOE between
Cys29
H and signal A (B) but not Z suggests that
resonance A (B) is cis to the
H of Cys29.
Pro28 and the Presence of Isoenzymes--
The
downfield hyperfine shifted resonances that remain unassigned by now
are signals L, M, and N, since they display no NOE connectivity to any
other hyperfine-shifted resonances (Figs. 3 and 4). This situation adds
uncertainties in assigning these signals. However, the extremely long
nonselective T1 (220 ms) of signal L as compared with other
hyperfine-shifted resonances suggests the assignment of signal L to a
proton at a relatively remote distance to the heme iron. Although the
identity of this signal cannot be determined from the current data,
observation of NOEs between this signal and three other signals at
10.2, 8.7, and 7.9 ppm suggests the assignment of this signal to one of
the CH2 protons in Pro28. This is based on the
fact that the signal at 7.9 ppm displays an observed NOE to 7-H
proton (cross-peak 34 in Fig. 3). With a distance of 2.99 Å between
H of Pro28 and 7-H
, it is reasonable to expect
observable NOEs between this proton pair (cross-peak 34 in
Fig. 3). The temperature-independent nature of this signal (Table I)
also suggests the remote position of this proton relative to the iron
center. The distance of the proton (signal M) to heme iron predicted
from the relaxation properties of this signal suggests the assignment
of signal M to another
H of Pro28 (Table II). It is
interesting to note that the proposed Pro28 protons also
display a doubled peak pattern. This suggests that Pro28 is
involved in protein modification that defines the two isozymes. This
hypothesis complies with the conclusion that the protein modification
that differentiates the two isozymes is located in the vicinity of the
heme 7-propionate group or is communicated to the heme via this side
chain. It is thus reasonable to propose that the existence of
isoenzymes of CPO is a result of conformational change of
Pro28. The isomerization of the prolyl imide bond has been
recently attributed to the modulation of ligand recognition of proteins by controlling the relative orientation of protein-binding surfaces (86). However, the failure to observe any TOCSY connectivities among
these signals makes it impossible to confirm the assignments proposed
here. Difficulties for observing TOCSY connectivities have often been
encountered for paramagnetic metalloproteins due to the spread of the
relaxation properties and the chemical shifts of the resonances (87).
Consequently, resonance assignments in paramagnetic metalloproteins are
commonly achieved through analysis of NOE connectivities (35, 72, 87).
Therefore, further studies are necessary to provide definitive
assignment of this residue. This will in turn, allow the elucidation of
the role this residue plays in defining the signature of the two
isozymes of CPO.
Alternatively, the existence of isozymes of CPO may be attributed to post-translational modifications of the protein (88). The secreted form of CPO is processed from a precursor containing a 21-residue-long, moderately hydrophobic signal sequence, at an atypical Gln-Glu peptide bond. Following cleavage, the N-terminal glutamic acid readily cyclizes into pyroglutamic acid. Furthermore, CPO contains two high mannose N-glycosylation sites, identified as asparagine 12 and 213. Other modifications include deamidation of residues asparagine 13, asparagine 198, and glutamine 183 into the corresponding acids (88). Any difference in these modification processes could result in the formation of isozymes with similar global structural but distinct local environments.
The Distal Heme Cavity--
In contrast to most heme peroxidases
that use a His at the distal heme pocket as the acid-base catalyst, CPO
employs a more acidic amino acid, Glu183 for the same
function. This makes it difficult to assign any signals from this
residue, since it does not possess a characteristic resonance as in the
case of a His. Our assignment for the distal heme environment was
primarily achieved by analysis of the NOESY spectra collected in 90%
H2O solution (Supplemental Fig. S3). The
solvent-exchangeable signal "a" displays an observed hyperfine shift of 16.3 ppm, comparable with the position of the solvent exchangeable proton observed at 16.5 ppm in CcPCN and 16.3 ppm in HRPCN (34). In most of the heme peroxidases studied, the downfield shifted solvent-exchangeable signals have all been assigned to the residues at the distal side of the heme cavity (59). Therefore,
we can assign signal "a" to a solvent-exchangeable proton of the
distal residues in CPO. The corresponding proton has been assigned to
the H1 of the distal His in CcP and horseradish peroxidase (34). However, the distance of H
1 in His105
at the distal side of CPO is much further from the heme iron compared
with that in CcP and horseradish peroxidase. On the other hand, the H
2 is at a reasonable distance from heme iron and is hydrogen-bonded to the acid-base catalyst, Glu183. Since
there are no other exchangeable protons at the distal heme cavity of
CPO, signal "a" is assigned to the H
2 of His105 in
CPO. This assignment is further supported by the NOEs observed between
"a" and the firmly assigned 7-propionate
-protons. With distances of 3.22 and 3.45 Å between N
2 and the two 7-propionate
-protons, it is expected to observe NOEs between H
2 and
the 7-propionate protons (cross-peaks 50 and 51 in Supplemental Fig. S3). Observation of cross-peaks (48 and
49 in Supplemental Fig. S3) between signal "a" and two other
protons assignable to the imidazole ring C
and C
protons also
favors the assignment of "a" as H
2 of His105 in CPO.
The long nonselective T1 (200 ms) of signal "a"
predicts that this proton is 8.13 Å from the heme iron (Table II).
This is in close agreement with the distance measured from the crystal structure of CPO (6).
The protonation of His105 might be critical for the
catalytic activity of CPO. In most heme peroxidases, the concerted
interaction of both the distal acid-base catalyst and a second polar
residue is required to cleave the peroxide bond in the formation of
compound I (89). The only polar residue at the heme distal side of CPO is His105, which is 3.5 Å above the 7-propionate group
(6). Therefore, we propose that this His indirectly participates in the
cleavage of the peroxide bond by hydrogen-bonding to and correctly
positioning of the direct acid-base catalyst,
Glu183, in CPO as shown in Fig.
7. This picture is in perfect agreement with the proposed reaction mechanisms of CPO reported previously (6,
90, 91).
|
In this mechanism, the activation of hydrogen peroxide is initiated by
its binding to the distal heme site of CPO that was occupied by a water
molecule in the enzyme's resting state (Fig. 7, 1).
The hydrogen bond between His105 (N2H) and
Glu183 (O
1) facilitates the abstraction of a peroxide
proton by Glu183 through hydrogen bonding between the
peroxide proton and the other carboxylate oxygen (O
2) of
Glu183 (2). The resulting singly ionized
hydrogen peroxide covalently binds to the heme iron, leading to the
formation of a short lived intermediate (3). This
intermediate then brings the terminal oxygen of the activated peroxide
closer to Glu183, which helps the formation of a strong
hydrogen bond between Glu183 (O
2H) and the terminal oxygen of
peroxide (4). Formation of 4 dramatically
promotes the subsequent proton delivery to the terminal oxygen from
Glu183, resulting in the heterolytic cleavage of the O
O
bond and the formation of the compound I oxoferryl center
(Fe4+=O) and porphyrin
cation radical common to most
heme peroxidases (5). Reaction of 5 with organic
substrates regenerates the resting state enzyme (peroxidase activity).
Alternatively, CPO compound I can be transformed into the resting state
via evolution of dioxygen (catalase activity). Most importantly, the
compound I of CPO can activate most halide ions to halogenate a broad
range of organic acceptor molecules (halogenase activity). Although His105 is not directly involved in the reactions of CPO, it
is critical to the normal function of CPO, because it helps to properly
arrange the direct acid-base catalyst (Glu183) in its
correct orientation.
The NOESY results also showed several cross-peaks between the firmly
assigned heme signals and a signal that is assignable to the other
distal residue, Phe186. The crystal coordinate of CPO
indicated that the H of Phe186 is at a position capable
of producing NOEs to four types of heme protons, 3-CH3,
4-H
, 4-H
-t, and 4-H
-c (cross-peaks 17, 26, 43, and 46 in Fig.
3). This is exactly what was observed. Therefore, the signal at 2.4 ppm
(intercept at 2.2 ppm from Curie plot, Table I) could be assigned to
one of the C
protons of Phe186.
Signals from other distal residues, Phe103, a residue thought to be important in controlling substrate access to heme center, and especially Glu183, the residue that is directly involved in the catalytic process of CPO, cannot be clearly accessed from the current data. Therefore, further studies are needed to locate the NMR properties of these important residues. This can be achieved by either replacing these residues with other amino acids or selectively labeling these residues with an NMR active isotope such as 13C or 15N. This work is currently in progress in our laboratories.
Location of the Putative Mn(II) Binding Site-- The above assignments of paramagnetically shifted signals make it possible to locate the proposed Mn(II) binding site in CPO. Since the unpaired electrons of Mn(II) can interact with protons close to Mn(II) and broaden them, observation of broadening of specific signals indicates specific Mn(II) binding and helps to locate the Mn(II) binding site (36, 63). However, as pointed out earlier, essentially no specific spectral changes were observed upon the addition of up to 10 eq of Mn(II) to the CPO preparations (Fig. 6). This is in sharp contrast with the results reported for CPO crystals, native MnP (63), and the engineered Mn(II) binding variants of CcP (36, 62). As discussed earlier, CPO isolated from fungal cultures grown in the absence of manganese neither contains Mn(II) nor binds added Mn(II). Thus, we conclude that binding of Mn(II) to its specific site in CPO requires the presence of Mn(II) during folding of the enzyme into its native conformation. When CPO folds in the absence of Mn(II), the high affinity binding site remains vacant and is inaccessible to added Mn(II). This conclusion is supported by all of our NMR (Fig. 6), UV-visible, and EPR (data not shown) studies carried out on proteins isolated from cultures grown in the absence of Mn(II).
Other Inferred Assignments-- The NOESY data presented in this work contain several cross-peaks involving the hyperfine shifted heme protons. On the basis of the cross-peak intensities, the specific NOE pattern, the observed shift positions, the extrapolated Curie intercept, assignments in other heme peroxidases, and the distance measured from the crystal structure of CPO, most of these cross-peaks can be assigned to the amino acid side chains that are in close proximity to the heme protons in question. These assignments are given in the legend to Fig. 3 with proton distances (Å) given in the parenthesis after the assignments.
In summary, the results presented here allowed the first reasonable
assignments for most of the hyperfine shifted resonances, including
those of the heme iron ligand (Cys29) of CPO. The location
of protein modification that differentiates the signature of the two
isozymes of CPO is near the heme propionate group, possibly due to the
conformational change of Pro28. The possible function of
His105 at the distal heme cavity is to help the cleavage of
the peroxide bond by hydrogen-bonding to and properly positioning the
acid-base catalyst, Glu183, within the heme center. The
high affinity Mn(II) binding site in CPO (in solution) is not
accessible to externally added Mn(II). The results presented here
provide a reasonable explanation for the discrepancies in the
literature between spectroscopists and crystallographers concerning the
Mn(II) binding site in this unique protein. Our study indicates that
results from NMR investigations of the protein in solution can well
complement the results revealed by x-ray diffraction studies in the
crystal form and thus provide a complete and better picture of the
actual structure of the protein.
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ACKNOWLEDGEMENTS |
---|
We are indebted to M. Feng, Dr. J. Chou, and Dr. G. Rai for technical help. We acknowledge the University of Southern Mississippi Polymer Science NMR facility (Varian UNITY-INOVA 500 MHz) and the Varian Oxford Instrument Center for Excellence in NMR laboratory (Varian UNITY 600 MHz) at the University of Illinois for use of the spectrometers.
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FOOTNOTES |
---|
* This research was sponsored by start-up support from the Chemistry Department of Jackson State University (to X. W.) and National Institutes of Health Grant GM07768 (to L. P. H.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The on-line version of this article (available at
http://www.jbc.org) contains three additional figures.
§ To whom correspondence may be addressed: Dept. of Chemistry, Jackson State University, 1400 J. R. Lynch St., Jackson, MS 39217. Tel.: 601-979-3719; Fax: 601-979-3674; E-mail: xwang@stallion.jsums.edu.
To whom correspondence may be addressed: Dept. of
Biochemistry, University of Illinois, 600 S. Mathews Ave., Urbana, IL
61801. Tel.: 217-333-9686; Fax: 217-265-0385; E-mail:
l-hager@uiuc.edu.
Published, JBC Papers in Press, December 16, 2002, DOI 10.1074/jbc.M209462200
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ABBREVIATIONS |
---|
The abbreviations used are:
CPO, chloroperoxidase;
CcP, cytochrome c peroxidase;
CcPCN, cyanide-ligated low spin form of
CcP;
NOE, nuclear Overhauser effect;
NOESY, two-dimensional
nuclear Overhauser enhancement spectroscopy;
TOCSY, two-dimensional
total correlation spectroscopy;
CPOCN, cyanide-ligated low spin form of
CPO;
MnCcP, cytochrome c peroxidase containing
Gly41 Glu, Val45
Glu, and
His181
Asp triple mutations;
MnCcPCN, cyanide-ligated low spin form of MnCcP;
MnP, manganese
peroxidase;
MnPCN, cyanide-bound low spin form of MnP;
HRPCN, cyanide-bound low spin form of horseradish peroxidase.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Everse, J., Everse, K. E., and Grisham, M. B. (1990) Peroxidases in Chemistry and Biology , CRC Press, Inc., Boca Raton, FL |
2. |
Hollenberg, P. F.,
and Hager, L. P.
(1973)
J. Biol. Chem.
248,
2630-2633 |
3. | Sono, M., Dawson, J. H., and Hager, L. P. (1986) Biochemistry 25, 347-356[Medline] [Order article via Infotrieve] |
4. | Dawson, J. H., Kau, L. S., Penner-Hahn, J. E., Sono, M., Eble, K. S., Bruce, G. S., Hager, L. P., and Hodgson, K. O. (1986) J. Am. Chem. Soc. 108, 8114-8116 |
5. | Sundaramoorthy, M., Mauro, J. M., Sullivan, A. M., Terner, J., and Poulos, T. L. (1995) Acta Crystallogr. Sect. D Biol. Crystallogr. 51, 842-844[CrossRef] |
6. | Sundaramoorthy, M., Terner, J., and Poulos, T. L. (1995) Structure (Lond.) 3, 1367-1377[Medline] [Order article via Infotrieve] |
7. |
Thomas, J. A.,
Morris, D. R.,
and Hager, L. P.
(1970)
J. Biol. Chem.
245,
3129-3134 |
8. | Hewson, W. D., and Hager, L. P. (1979) Porphyrins 7, 295-332 |
9. |
Ortiz de Montellano, P. R.,
Choe, Y. S.,
DePillis, G.,
and Catalano, C. E.
(1987)
J. Biol. Chem.
262,
11641-11646 |
10. | Allain, E. J., Hager, L. P., Deng, L., and Jacobsen, E. N. (1993) J. Am. Chem. Soc. 115, 4415-4416 |
11. | Allain, E. J. (1997) Enantioselective Epoxidation of Alkenes by Chloroperoxidase and the Development of a Chloroperoxidase Expression System.Ph.D. thesis , pp. 27-38, University of Illinois, Urbana, IL |
12. | Dawson, J. H. (1988) Science 240, 433-439[Medline] [Order article via Infotrieve] |
13. | Dawson, J. H., and Sono, M. (1987) Chem. Rev. 87, 1255-1276 |
14. |
Libby, R. D.,
Beachy, T. M.,
and Phipps, A. K.
(1996)
J. Biol. Chem.
271,
21820-21827 |
15. |
Libby, R. D.,
Rotberg, N. S.,
Emerson, J. T.,
White, T. C.,
Yen, G. M.,
Friedman, S. H.,
Sun, N. S.,
and Goldowski, R.
(1989)
J. Biol. Chem.
264,
15284-15292 |
16. |
Libby, R. D.,
Shedd, A. L.,
Phipps, A. K.,
Beachy, T. M.,
and Gerstberger, S. M.
(1992)
J. Biol. Chem.
267,
1769-1775 |
17. | Hu, S., and Hager, L. P. (1999) Tetrahedron Lett. 40, 1641-1644[CrossRef] |
18. | Lakner, F. J., and Hager, L. P. (1996) J. Org. Chem. 61, 3923-3925[CrossRef][Medline] [Order article via Infotrieve] |
19. | Hu, S., and Hager, L. P. (1998) Biochem. Biophys. Res. Commun. 253, 544-546[CrossRef][Medline] [Order article via Infotrieve] |
20. | Hu, S., and Hager, L. P. (1999) J. Am. Chem. Soc. 121, 872-873[CrossRef] |
21. | Colonna, S., Gaggero, N., Manfredi, A., Casella, L., Gullotti, M., Carrea, G., and Pasta, P. (1990) Biochemistry 29, 10465-10468[Medline] [Order article via Infotrieve] |
22. | Colonna, S., Gaggero, N., and Pasta, P. (1992) NATO ASI Ser. Ser. C Math. Phys. Sci. 381, 323-331 |
23. | Colonna, S., Gaggero, N., Casella, L., Carrea, G., and Pasta, P. (1992) Tetrahedron Asymmetry 3, 95-106[CrossRef] |
24. |
Finzel, B. C.,
Poulos, T. L.,
and Kraut, J.
(1984)
J. Biol. Chem.
259,
13027-13036 |
25. |
Poulos, T. L.,
and Kraut, J.
(1980)
J. Biol. Chem.
255,
8199-8205 |
26. | Satterlee, J. D., Erman, J. E., LaMar, G. N., Smith, K. M., and Langry, K. C. (1983) Biochim. Biophys. Acta 743, 246-255[Medline] [Order article via Infotrieve] |
27. | Satterlee, J. D., Erman, J. E., LaMar, G. N., Smith, K. M., and Langry, K. C. (1983) J. Am. Chem. Soc. 105, 2099-2104 |
28. | Alam, S. L., Satterlee, J. D., Mauro, J. M., Poulos, T. L., and Erman, J. E. (1995) Biochemistry 34, 15496-15503[Medline] [Order article via Infotrieve] |
29. | Satterlee, J. D., Alam, S. L., Mauro, J. M., Erman, J. E., and Poulos, T. L. (1994) Eur. J. Biochem. 224, 81-87[Abstract] |
30. | Satterlee, J. D., Russell, D. J., and Erman, J. E. (1991) Biochemistry 30, 9072-9077[Medline] [Order article via Infotrieve] |
31. | Satterlee, J. D., and Erman, J. E. (1991) Biochemistry 30, 4398-4405[Medline] [Order article via Infotrieve] |
32. | Satterlee, J. D., Erman, J. E., Mauro, J. M., and Kraut, J. (1990) Biochemistry 29, 8797-8804[Medline] [Order article via Infotrieve] |
33. |
Satterlee, J. D.,
Erman, J. E.,
and DeRopp, J. S.
(1987)
J. Biol. Chem.
262,
11578-11583 |
34. | Banci, L., Bertini, I., Turano, P., Ferrer, J. C., and Mauk, A. G. (1991) Inorg. Chem. 30, 4510-4516 |
35. | Savenkova, M. I., Satterlee, J. D., Erman, J. E., Siems, W. F., and Helms, G. L. (2001) Biochemistry 40, 12123-12131[CrossRef][Medline] [Order article via Infotrieve] |
36. | Wang, X., and Lu, Y. (1999) Biochemistry 38, 9146-9157[CrossRef][Medline] [Order article via Infotrieve] |
37. | de Ropp, J. S., Sham, S., Asokan, A., Newmyer, S., Ortiz de Montellano, P. R., and La Mar, G. N. (2002) J. Am. Chem. Soc. 124, 11029-11037[CrossRef][Medline] [Order article via Infotrieve] |
38. | Asokan, A., de Ropp, J. S., Newmyer, S. L., de Montellano, P. R. O., and La Mar, G. N. (2001) J. Am. Chem. Soc. 123, 4243-4254[CrossRef][Medline] [Order article via Infotrieve] |
39. | Chen, Z., de Ropp, J. S., Hernandez, G., and La Mar, G. N. (1994) J. Am. Chem. Soc. 116, 8772-8783 |
40. | De Ropp, J. S., and La Mar, G. N. (1991) J. Am. Chem. Soc. 113, 4348-4350 |
41. | de Ropp, J. S., Chen, Z., and La Mar, G. N. (1995) Biochemistry 34, 13477-13484[Medline] [Order article via Infotrieve] |
42. | De Ropp, J. S., Mandal, P. K., and La Mar, G. N. (1999) Biochemistry 38, 1077-1086[CrossRef][Medline] [Order article via Infotrieve] |
43. | Gonzalez-Vergara, E., Meyer, M., and Goff, H. M. (1985) Biochemistry 24, 6561-6567[Medline] [Order article via Infotrieve] |
44. | La Mar, G. N., De Ropp, J. S., Smith, K. M., and Langry, K. C. (1980) J. Am. Chem. Soc. 102, 4833-4835 |
45. | La Mar, G. N., and De Ropp, J. S. (1980) J. Am. Chem. Soc. 102, 395-397 |
46. | La Mar, G. N., and De Ropp, J. S. (1982) J. Am. Chem. Soc. 104, 5203-5206 |
47. |
La Mar, G. N.,
Thanabal, V.,
Johnson, R. D.,
Smith, K. M.,
and Parish, D. W.
(1989)
J. Biol. Chem.
264,
5428-5434 |
48. | La Mar, G. N., Chen, Z., Vyas, K., and McPherson, A. D. (1995) J. Am. Chem. Soc. 117, 411-419 |
49. | Thanabal, V., De Ropp, J. S., and La Mar, G. N. (1986) J. Am. Chem. Soc. 108, 4244-4246 |
50. | Thanabal, V., De Ropp, J. S., and La Mar, G. N. (1987) J. Am. Chem. Soc. 109, 7516-7525 |
51. | Thanabal, V., DeRopp, J. S., and La Mar, G. N. (1987) J. Am. Chem. Soc. 109, 265-272 |
52. | Thanabal, V., La Mar, G. N., and De Ropp, J. S. (1988) Biochemistry 27, 5400-5407[Medline] [Order article via Infotrieve] |
53. | Thanabal, V., De Ropp, J. S., and La Mar, G. N. (1988) J. Am. Chem. Soc. 110, 3027-3035 |
54. | Dugad, L. B., Wang, X., Wang, C. C., Lukat, G. S., and Goff, H. M. (1992) Biochemistry 31, 1651-1655[Medline] [Order article via Infotrieve] |
55. | Wang, X., and Goff, H. M. (1997) Biochim. Biophys. Acta 1339, 88-96[Medline] [Order article via Infotrieve] |
56. | Lukat, G. S., and Goff, H. M. (1990) Biochim. Biophys. Acta 1037, 351-359[Medline] [Order article via Infotrieve] |
57. | Goff, H. M., Gonzalez-Vergara, E., and Bird, M. R. (1985) Biochemistry 24, 1007-1013[Medline] [Order article via Infotrieve] |
58. |
Lukat, G. S.,
and Goff, H. M.
(1986)
J. Biol. Chem.
261,
16528-16534 |
59. | Bertini, I., Turano, P., and Vila, A. J. (1993) Chem. Rev. 93, 2833-2932 |
60. |
Hollenberg, P. F.,
Hager, L. P.,
Blumberg, W. E.,
and Peisach, J.
(1980)
J. Biol. Chem.
255,
4801-4807 |
61. | Yeung, B. K. S., Wang, X., Sigman, J. A., Petillo, P. A., and Lu, Y. (1997) Chem. Biol. 4, 215-221[Medline] [Order article via Infotrieve] |
62. | Wilcox, S. K., Putnam, C. D., Sastry, M., Blankenship, J., Chazin, W. J., McRee, D. E., and Goodin, D. B. (1998) Biochemistry 37, 16853-16862[CrossRef][Medline] [Order article via Infotrieve] |
63. | Banci, L., Bertini, I., Bini, T., Tien, M., and Turano, P. (1993) Biochemistry 32, 5825-5831[Medline] [Order article via Infotrieve] |
64. |
Morris, D. R.,
and Hager, L. P.
(1966)
J. Biol. Chem.
241,
1763-1768 |
65. | Gonzalez-Vergara, E., Ales, D. C., and Goff, H. M. (1985) Prep. Biochem. 15, 335-348[Medline] [Order article via Infotrieve] |
66. | Inubushi, T., and Becker, E. D. (1983) J. Magn. Reson. 51, 128 |
67. | Pierattelli, R., Banci, L., and Turner, D. L. (1996) J. Biol. Inorg. Chem. 1, 320-329[CrossRef] |
68. | States, D. J., Haberkorn, R. A., and Ruben, D. J. (1982) J. Magn. Reson. 48, 286-292 |
69. | Griesinger, C., Otting, G., Wuethrich, K., and Ernst, R. R. (1988) J. Am. Chem. Soc. 110, 7870-7872 |
70. | Walker, A. F., and Simonis, U. (1993) in NMR of Paramagnetic Molecules (Berliner, L. J. , and Reuben, J., eds), Vol. 12 , pp. 133-274, Plenum Press, New York |
71. | Dugad, L. B., and Goff, H. M. (1992) Biochim. Biophys. Acta 1122, 63-69[Medline] [Order article via Infotrieve] |
72. | Banci, L., Bertini, I., Pease, E. A., Tien, M., and Turano, P. (1992) Biochemistry 31, 10009-10017[Medline] [Order article via Infotrieve] |
73. |
Satterlee, J. D.,
and Erman, J. E.
(1981)
J. Biol. Chem.
256,
1091-1093 |
74. | Edwards, S. L., Raag, R., Wariishi, H., Gold, M. H., and Poulos, T. L. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 750-754[Abstract] |
75. | Gajhede, M., Schuller, D. J., Henriksen, A., Smith, A. T., and Poulos, T. L. (1997) Nat. Struct. Biol. 4, 1032-1038[Medline] [Order article via Infotrieve] |
76. |
Sundaramoorthy, M.,
Kishi, K.,
Gold, M. H.,
and Poulos, T. L.
(1994)
J. Biol. Chem.
269,
32759-32767 |
77. | La Mar, G. N., and Walker, F. A. (1979) in Porphyrins (Dolphin, D., ed), Vol. 4 , pp. 61-157, Academic Press, New York |
78. | La Mar, G. N., Viscio, D. B., Smith, K. M., Caughey, W. S., and Smith, M. L. (1978) J. Am. Chem. Soc. 100, 8085-8092 |
79. | La Mar, G. N. (1979) in Biological Applications of Magnetic Resonance (Shulman, R. G., ed) , pp. 305-343, Academic Press, New York |
80. | Banci, L., Bertini, I., Kuan, I. C., Tien, M., Turano, P., and Vila, A. J. (1993) Biochemistry 32, 13483-13489[Medline] [Order article via Infotrieve] |
81. | Qin, J., Delaglio, F., La Mar, G. N., and Bax, A. (1993) J. Magn. Reson. Ser. B 102, 332-336 |
82. | Bertini, I., Luchinat, C., and Tarchi, D. (1993) Chem. Phys. Lett. 203, 445-449[CrossRef] |
83. | Satterlee, J. D., Teske, J. G., Erman, J. E., Mauro, J. M., and Poulos, T. L. (2000) J. Protein Chem. 19, 535-542[Medline] [Order article via Infotrieve] |
84. | Banci, L., Bertini, I., Turano, P., Tein, M., and Kirk, T. K. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 6956-6960[Abstract] |
85. | La Mar, G. N., and De Ropp, J. S. (1979) Biochem. Biophys. Res. Commun. 90, 36-41[Medline] [Order article via Infotrieve] |
86. | Mallis, R. J., Brazin, K. N., Fulton, D. B., and Andreotti, A. H. (2002) Nat. Struct. Biol. 9, 900-905[CrossRef][Medline] [Order article via Infotrieve] |
87. | Salgado, J., Kalverda, A. P., Diederix, R. E. M., Canters, G. W., Moratal, J. M., Lawler, A. T., and Dennison, C. (1999) J. Biol. Inorg. Chem. 4, 457-467[CrossRef][Medline] [Order article via Infotrieve] |
88. | Kenigsberg, P., Fang, G. H., and Hager, L. P. (1987) Arch. Biochem. Biophys. 254, 409-415[Medline] [Order article via Infotrieve] |
89. | Poulos, T. L. (1988) Adv. Inorg. Biochem. 7, 1-36 |
90. | Sundaramoorthy, M., Terner, J., and Poulos, T. L. (1998) Chem. Biol. 5, 461-473[Medline] [Order article via Infotrieve] |
91. | Wagenknecht, H.-A., and Woggon, W.-D. (1997) Chem. Biol. 4, 367-372[Medline] [Order article via Infotrieve] |
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