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J. Biol. Chem., Vol. 275, Issue 41, 31891-31895, October 13, 2000
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From the Departments of
Received for publication, May 1, 2000, and in revised form, May 29, 2000
The flavoprotein nitroalkane oxidase catalyzes
the oxidative denitrification of primary or secondary nitroalkanes to
the corresponding aldehydes or ketones with production of hydrogen
peroxide and nitrite. The enzyme is irreversibly inactivated by
treatment with N-ethylmaleimide at pH 7. The inactivation
is time-dependent and shows first-order kinetics for three
half-lives. The second-order rate constant for inactivation is 3.4 ± 0.06 M The flavoprotein nitroalkane oxidase from the fungus
Fusarium oxysporum (ATCC 695) catalyzes the oxidative
denitrification of primary or secondary nitroalkanes to the
corresponding aldehydes or ketones with production of hydrogen peroxide
and nitrite (Scheme 1). The study of an
enzyme capable of oxidizing nitroalkanes is of considerable interest
from both fundamental and applied standpoints. Nitroalkanes are widely
used as industrial solvents, chemical intermediates, explosives, and
fuels (1). Several nitroalkanes are toxic and/or carcinogenic (1).
Thus, an enzymatic activity that converts nitroalkanes into less
harmful species has significant potential for bioremediation. From a
chemical standpoint, the formation of nitronates in solution is a well
characterized chemical reaction (2) that serves as the basis for
understanding the formation of carbanions involving much weaker carbon
acids, such as amino acids and Nitroalkane oxidase is isolated with the flavin cofactor in the form of
an N(5)-[3-nitrobut-2-yl]-1,5-dihydroflavin adenine dinucleotide and is consequently not active (3, 4). This nitrobutyl-flavin adduct can be converted in vitro to
FAD, yielding active enzyme (4, 5). The FAD-containing enzyme is
active on a broad range of primary and secondary nitroalkane substrates (6, 7). Although other flavoprotein oxidases, such as
D-amino acid oxidase (8), glucose oxidase (9), and
2-nitropropane dioxygenase (10, 11), have been shown to be able to
oxidize nitroalkanes, nitroalkane oxidase is unique in that it requires the neutral form of the substrate for catalysis (4, 12). Mechanistic
studies of nitroalkane oxidase support a chemical mechanism in which an
active site base with a pKa value of 7 removes a
proton from the Materials--
Nitroethane and FAD were from Sigma.
N-Ethylmaleimide and valerate were from Aldrich.
N-[ethyl-1-14C]Maleimide was from
NENTM Life Scientific Products, Inc.
L-1-Tosylamido-2-phenylethyl chloromethyl ketone-treated
trypsin was purchased from Worthington. Nitroalkane oxidase was
purified from F. oxysporum (ATCC 695) as described previously (7). The activated FAD-containing form of the enzyme was
prepared according to Gadda et al. (4) and was stored at Methods--
Enzyme activities were measured with 20 mM nitroethane as substrate in air-saturated 0.5 mM FAD, 50 mM potassium phosphate, 16 mM imidazole, pH 7.5, by monitoring the rate of oxygen
consumption with a computer-interfaced Hansatech Clark oxygen electrode
at 30 °C, as described previously (7). Stock solutions of
N-ethylmaleimide in 25 mM potassium phosphate,
pH 7, were prepared just prior to use.
Nitroalkane oxidase (10-20 µM) was incubated with
N-ethylmaleimide (3-15 mM) in 0.5 mM FAD, 25 mM potassium phosphate, pH 7, at 30 °C. All incubations
were carried out in the presence of FAD to avoid the formation of
apoprotein (5). At different times, aliquots were withdrawn and assayed
for enzymatic activity. For experiments in which the effect of valerate
on the rate of inactivation was measured, the enzyme was incubated for
5 min with this compound before the addition of
N-ethylmaleimide. To stop the inactivation, unreacted
N-ethylmaleimide was removed by gel filtration on a Sephadex
G-25 column equilibrated with 25 mM potassium phosphate, pH
7, at room temperature. The irreversibility of
N-ethylmaleimide inactivation of nitroalkane oxidase was
determined by incubating the modified enzyme isolated by gel filtration
for 3 h in 0.5 mM FAD, 25 mM potassium
phosphate, pH 7, at 30 °C. At different times, aliquots were
withdrawn and assayed for enzymatic activity.
To identify the peptide whose modification by
N-ethylmaleimide resulted in enzyme inactivation, 32 µM enzyme was incubated with 10 mM
N-[ethyl-1-14C]maleimide (1.8 × 103 cpm/nmol) in the presence and absence of 25 mM valerate in a total volume of 0.5 ml. After 25 min, a
second 10 mM aliquot of 10 mM
N-[ethyl-1-14C]maleimide was added,
and the incubation continued for an additional 25 min. The reaction was
stopped by gel filtration using a Sephadex G-25 column equilibrated
with 4 mM calcium chloride, 0.4 M ammonium bicarbonate, pH 8. After the addition of solid urea to a final concentration of 8 M, the samples were incubated for 1 h at 37 °C. The solutions were then diluted with three volumes of
water followed by the addition of trypsin to a final concentration of 3% (w/w, trypsin/nitroalkane oxidase). After a 4-h incubation at
37 °C, a second aliquot of trypsin (1% (w/w) final concentration) was added, and the digestion continued for a further 16 h at
37 °C. The reaction was stopped by adding freshly prepared
phenylmethanesulfonyl fluoride at a final concentration of 1 mg/ml.
Purification of peptides was carried out by
HPLC2 using a Waters
instrument equipped with a model 996 photodiode array detector and a
Vydac 218TP54 (4.6 × 250-mm) reverse-phase column at a flow rate
of 1 ml min Data Analysis--
The time course of inactivation of
nitroalkane oxidase by N-ethylmaleimide was analyzed by
fitting the residual activity (A) at a given time
(t) to Equation 1,
Inactivation of Nitroalkane Oxidase by
N-Ethylmaleimide--
Treatment of nitroalkane oxidase with
N-ethylmaleimide at pH 7 and 30 °C results in a
time-dependent loss of enzymatic activity, as shown in Fig.
1. The inactivation is first-order for
about 3 half-lives, but the rates decrease at longer times (data not shown). This could be due to hydrolysis of N-ethylmaleimide
to form N-ethylmaleamate, a process previously reported to
occur at pH 7 and above (16). Consistent with this hypothesis, the addition of a second aliquot of N-ethylmaleimide after three
half-lives results in a further time-dependent loss of
enzymatic activity. The initial rate of inactivation is dependent on
the concentration of N-ethylmaleimide (Fig. 1). A plot of
the rate of inactivation versus the concentration of the
reagent is linear up to 15 mM N-ethylmaleimide
(Fig. 1B), indicating that there is no significant formation
of a reversible complex between the reagent and the enzyme prior to
inactivation. The second-order rate constant for inactivation
determined from this plot is 3.4 ± 0.06 M
Valerate is a competitive inhibitor of nitroalkane oxidase with a
Ki value of 0.6 mM at pH 7 and 30 °C
(14). In the presence of 25 mM valerate the rate of
inactivation of nitroalkane oxidase by 10 mM
N-ethylmaleimide decreases from 96 min
To determine if the reaction between N-ethylmaleimide and
nitroalkane oxidase is irreversible, the enzyme was separated from the
remaining reagent by gel filtration when the residual activity had
decreased to 6% of the initial value. The inactivated enzyme was then
incubated in the absence of N-ethylmaleimide at pH 7 and
30 °C. No recovery of activity was observed after 3 h,
consistent with the modification being irreversible.
Identification of the Residue Modified by N-Ethylmaleimide--
To
identify the amino acid residue whose modification by
N-ethylmaleimide resulted in the inactivation of the enzyme,
nitroalkane oxidase was incubated with
N-[ethyl-1-14C]maleimide in the
presence and absence of valerate. The reactions were stopped when the
residual activity of the enzyme incubated in the absence of valerate
was 10%. At that time, the valerate-protected enzyme retains about
80% of the initial activity. Both samples were digested with trypsin,
and the resulting tryptic digests were separated by reverse-phase HPLC.
In our initial attempts the reaction was stopped by the addition of
10% trichloroacetic acid. When this was done, the tryptic maps of the
two samples were identical (data not shown), suggesting that the
modification is acid-labile. In contrast, when the reaction was
quenched by using gel filtration to remove the unreacted reagent, the
tryptic map of the sample lacking valerate showed an extra peak eluting at 61.4 min (Fig. 2). This was the only
HPLC fraction showing significant incorporation of radioactivity in
either sample. The N-terminal amino acid sequence of the alkylated
peptide was determined by automated Edman degradation to be
LLNEVMXYPL (Table I), where X indicates the absence of any phenylthiohydantoin
derivative in the chromatogram. This sequence corresponds to that of a
peptide in nitroalkane oxidase previously identified using
tetranitromethane as an active site-directed reagent,
LLNEVMXYPLFDGGNIGLR (15).
Mass spectrometry was used to definitively identify the residue at
position 7 and the site of modification. Positive ion MALDI-TOF mass
spectrometry of the modified peptide yielded a predominant peak with
m/z+ value of 2248.8 (Fig.
3). A cysteine residue at position 7 and incorporation of a single N-ethylmaleimide moiety into the
peptide would yield a m/z+ value of
2249, establishing cysteine as amino acid residue X. A
second peak with m/z+ value of 2264.7 was observed in the mass spectrum (Fig. 3), consistent with an oxidized
form of the same peptide.
The specific residue modified by N-ethylmaleimide was
identified by nanospray mass spectrometry on an ion trap mass
spectrometer in the positive ion mode. Species with mass/charge values
of 1126 and 1132 were seen in the mass spectrum of the same peptide.
The mass/charge value of the smaller species is consistent with that expected for the doubly charged ion of the alkylated peptide, while the
higher mass/charge ratio species can be attributed to an oxidized form
of the same peptide. These two parental ions were further fragmented
while in the trap in order to obtain the sequence of the peptide, as
illustrated in Fig. 4 for the
m/z2+ species of 1132. The sequence
of the alkylated peptide could be determined by comparing the
m/z+ values of the daughter ions to
the calculated values expected for the peptides produced in the
fragmentation process (Table II). The
sequences of both peptides determined from the ion trap spectrometric
analysis were in agreement with that from the Edman analysis. The data
in Table II are consistent with the cysteine residue at position 7 being the site of alkylation by N-ethylmaleimide and the
methionine residue at position 6 being oxidized in the species with
m/z2+ value of 1132. Since no
methionine sulfoxide was seen in the automated sequence analysis of the
same peptide, the oxidized methionine probably formed during the mass
spectrometric analysis.
Mechanistic studies of nitroalkane oxidase are consistent with a
mechanism for catalysis in which a base on the enzyme removes a proton
from the
Identification of a Cysteine Residue in the Active Site of
Nitroalkane Oxidase by Modification with
N-Ethylmaleimide*
,,, and§¶
Biochemistry and Biophysics
and § Chemistry, Texas A & M University,
College Station, Texas 77843-2128
ABSTRACT
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1
min1. The competitive inhibitor valerate
protects the enzyme from inactivation, indicating an active
site-directed modification. Comparison of tryptic maps of enzyme
treated with
N-[ethyl-1-14C]maleimide in the
absence and presence of valerate shows a single radioactive peptide
differentially labeled in the unprotected enzyme. The sequence of this
peptide was determined to be LLNEVMCYPLFDGGNIGLR using Edman
degradation and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. The cysteine residue was
identified as the site of alkylation by ion trap mass spectrometry.
INTRODUCTION
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EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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-hydroxy acids. Therefore, the study
of an enzymatic activity that carries out the oxidation of nitroalkanes provides the unique opportunity to compare the enzyme-catalyzed formation of nitronates with the reaction in solution.
View larger version (4K):
[in a new window]
Scheme 1.
-carbon of the substrate to initiate catalysis (13,
14). The resulting carbanion attacks the N-5 position of FAD to
form a covalent adduct that subsequently decays to form nitrite and the
aldehyde or ketone product. This covalent adduct can be trapped by
nitroethane anion during turnover of the enzyme with nitroethane,
yielding 5-[3-nitrobut-2-yl]-1,5-dihydroflavin adenine dinucleotide
(4). The steady state kinetic mechanism of nitroalkane oxidase has been
determined with nitroethane (13, 14). The data are consistent with a
ping-pong isomechanism in which the release of the aldehyde product
from the reduced enzyme is followed by an irreversible isomerization to
yield the enzyme species that reacts with oxygen. Despite the
considerable advances in the biochemical and mechanistic
characterization of the enzyme, no structural information is available
beyond the sequence of part of the gene encoding for the N-terminal
half of nitroalkane oxidase.1
In the absence of crystallographic data, an effective strategy to
identify active site residues has been the use of irreversible inhibitors. This approach has been recently used to identify an essential tyrosine residue in the active site of nitroalkane oxidase (15). In the present report, we describe a kinetic and structural characterization of the inactivation of the FAD-containing form of
nitroalkane oxidase by the cysteine-directed reagent
N-ethylmaleimide (16).
EXPERIMENTAL PROCEDURES
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70 °C in the presence of 0.5 mM FAD to prevent
formation of the less stable apoprotein. The concentration of
nitroalkane oxidase was determined by the method of Bradford (17) using
bovine serum albumin as standard. All other reagents were of the
highest purity commercially available.
1. Eluent A was 0.05% aqueous
trifluoroacetic acid, and eluent B was 0.04% trifluoroacetic acid in
acetonitrile. The chromatography was carried out with a linear gradient
from 5 to 50% B over 90 min. Peptides were collected manually, and the
amount of radioactivity incorporated in each was determined by
scintillation counting. Automated Edman degradation of the purified
peptide was carried out on a Hewlett-Packard G1000A protein sequencer
at the Protein Chemistry Laboratory of Texas A & M University.
MALDI-TOF mass spectroscopy of the purified peptide was carried out
using a Voyager Elite XL mass spectrometer (PerSeptive Biosystems,
Framingham, MA) at the Laboratory for Biological Mass Spectrometry at
Texas A & M University. MALDI-TOF spectra were acquired in the positive ion mode, using -cyano-4-hydroxysuccinamic acid as matrix. Samples were prepared for MALDI-TOF using the overlayer method of sample preparation previously described (18). Mass spectrometry of the
purified peptide was performed using an Esquire LC Ion Trap Mass
Spectrometer equipped with a nanospray source (Bruker Daltonics, Inc.,
Billerica, MA).
where A0 is the initial activity and
kobs is the observed rate of inactivation.
(Eq. 1)
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1
min1.
View larger version (18K):
[in a new window]
Fig. 1.
Time-dependent inactivation of
nitroalkane oxidase by N-ethylmaleimide.
Nitroalkane oxidase (10 µM) was incubated with different
concentrations of N-ethylmaleimide in 0.5 mM
FAD, 25 mM potassium phosphate, pH 7, and 30 °C. At
different times, aliquots were withdrawn and assayed for enzymatic
activity as described under "Experimental Procedures."
A, time course of inactivation. N-Ethylmaleimide
concentrations were 3 ( 
), 5 (
), 10 (
), 15 (
), and 10 mM (
) in the presence of 25 mM valerate. The
lines are fits of the data to Equation 1. B,
secondary plot of the observed rate of inactivation as a function of
the concentration of N-ethylmaleimide.
1 to 0.1 min1
(Fig. 1), consistent with the inactivation being active
site-directed.
View larger version (26K):
[in a new window]
Fig. 2.
HPLC of tryptic digests of nitroalkane
oxidase treated with N-ethylmaleimide in the presence
and absence of valerate. Nitroalkane oxidase was incubated with
N-[ethyl-1-14C]maleimide in the
presence and absence of valerate as described under "Experimental
Procedures." Tryptic digests of each sample were separated by
reverse-phase HPLC as described under "Experimental Procedures."
Peptide elution was monitored at 214 nm (A). The resulting
peaks were analyzed for radioactive incorporation using a scintillation
counter (B). The radioactive peptide eluting at 61.4 min
found only in the inactivated sample is indicated by an
arrow. The upper line in each
panel is the sample treated with N-ethylmaleimide
alone; the bottom line is the sample treated with
N-ethylmaleimide in the presence of valerate.
N-terminal amino acid sequence of the peptide eluting at 61.4 min in
the tryptic digest of nitroalkane oxidase treated with
N-ethylmalcimide
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[in a new window]
Fig. 3.
MALDI-TOF mass spectrometric analysis of the
peptide modified by N-ethylmaleimide. The
N-ethylmaleimide-modified peptide isolated by reverse-phase
HPLC of a tryptic digest of nitroalkane oxidase inactivated by
N-ethylmaleimide was analyzed by MALDI-TOF mass spectrometry
as described under "Experimental Procedures." Inset,
expansion of the m/z+ region
2245-2256 of the mass spectrum.
View larger version (18K):
[in a new window]
Fig. 4.
Nanospray mass spectrometric analysis of the
peptide modified by N-ethylmaleimide. The
parental ion with an m/z2+ value of
1132 was fragmented in the ion trap mass spectrometer as described
under "Experimental Procedures."
Ion trap mass spectrometric analysis of the N-ethylmaleimide-containing
peptide
DISCUSSION
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-carbon of the substrate to form a carbanion (Scheme 2) (4, 13, 14). During the normal course of
catalysis, the carbanion can attack the N-5 position of the
cofactor to form a covalent adduct. This adduct can lose nitrite to
form a highly reactive cationic imine that can then react with
hydroxide to form the aldehyde or ketone product and reduced flavin.
Alternatively, the cationic imine can react with nitroethane anion in
solution to form the stable and inactive 5-nitrobutyl-flavin. Despite
the advances in the mechanistic and biochemical characterization of the
enzyme, no structural information is available beyond the sequence of
part of the gene encoding for nitroalkane oxidase1. The use
of irreversible chemical modification reagents has long been employed
to study the active site of enzymes whose structures are not available.
By using this approach, we have recently identified a tyrosine residue
in the active site of nitroalkane oxidase (15). The results presented
here identify a cysteine residue as also being in the active site
of the enzyme.
View larger version (22K):
[in a new window]
Scheme 2.
Nitroalkane oxidase is irreversibly inactivated in a time- and concentration-dependent fashion by treatment with N-ethylmaleimide. The inactivation is active site-directed, based on the protection from inactivation afforded by the competitive inhibitor valerate. The tryptic digests of nitroalkane oxidase treated with N-[ethyl-1-14C]maleimide both in the presence and absence of valerate show that a single peptide is differentially labeled by N-ethylmaleimide in the inactivated enzyme. This peptide is the same as the one previously identified using tetranitromethane as an active site probe (15). The cysteine residue identified here is adjacent to the active site tyrosine nitrated by tetranitromethane (15), strengthening the conclusion that both of these residues are in the active site of the enzyme. As previously reported, no direct match of the sequence of the alkylated peptide could be found with the available partial gene sequence of nitroalkane oxidase,1 indicating that the cysteine and tyrosine residues are located in the C-terminal half of the protein.
In terms of the catalytic roles of these two residues, we have
previously proposed that the tyrosine residue participates in substrate
binding by forming a hydrogen bond to the nitro group of the
nitroalkane substrate (15). A possible role for a cysteine residue is
as the active site base that abstracts a proton from the -carbon of
the substrate, as is the case with the flavoprotein dihydroorotate
dehydrogenase (19, 20). The involvement of a cysteine residue in
catalysis is consistent with solvent kinetic isotope effect studies of
nitroalkane oxidase, specifically an inverse effect on the
V/K value of about 0.5.1 Previous pH
dependence studies on the enzyme showed the presence of a base with a
pKa value of 7 in the free enzyme (14). However, the
effects of D2O on the pH dependence do not support the
assignment of the cysteine as the residue responsible for this
pKa. The pKa value of a cysteine
residue should shift less than 0.4 units when changing from water to
D2O (21). Instead, the pKa value
increases to 7.7 in D2O,1 strongly suggesting
that the catalytic base in nitroalkane oxidase is not a cysteine
residue. In the catalytic mechanism of Scheme 2, another active site
base is required to deprotonate a water molecule to form the hydroxide
that reacts with the cationic imine formed during catalysis. Although a
serine residue would seem more appropriate for this reaction due to the
pKa value of about 15 of its functional group (22),
a cysteine residue with an unusually high pKa value
could act as this base. An alternative explanation is that the active
site cysteine is not directly involved in catalysis but that enzyme
inactivation stems from steric hindrance due to the presence of the
N-ethylmaleimide moiety in the active site preventing
substrate binding.
In summary, the chemical modification studies with
N-ethylmaleimide presented here show that a cysteine residue
is present in the active site of the flavoprotein nitroalkane oxidase.
This amino acid residue is located next to a tyrosine residue
previously identified in the active site of nitroalkane oxidase. These
results are a prerequisite to future mutagenesis studies aimed at a
better understanding of the catalytic mechanism of this enzyme.
| |
ACKNOWLEDGEMENTS |
|---|
We thank Dr. William K. Russell at the Laboratory for Biological Mass Spectrometry of Texas A & M University for the MALDI-TOF mass spectrometric analyses and Dr. Catherine Stacey at Bruker Daltonics, Inc. (Billerica, MA) for the ion trap mass spectrometric analyses.
| |
FOOTNOTES |
|---|
* This work was supported in part by National Institutes of Health Grant GM 58698.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.
¶ To whom correspondence should be addressed: Dept. of Biochemistry and Biophysics, 2128 TAMU, College Station, TX 77843-2128. Tel.: 979-845-5487; Fax: 979-845-4946; E-mail: fitzpat@tamu.edu.
Published, JBC Papers in Press, July 25, 2000, DOI 10.1074/jbc.M003679200
1 G. Gadda and P. F. Fitzpatrick, unpublished results.
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ABBREVIATIONS |
|---|
The abbreviations used are: HPLC, high pressure liquid chromatography; MALDI-TOF, matrix-assisted laser desorption/ionization time-of-flight.
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EXPERIMENTAL PROCEDURES
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DISCUSSION
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| 2. | Bernasconi, C. F. (1992) Adv. Phys. Org. Chem. 27, 119-238 |
| 3. | Edmondson, R. D., Gadda, G., Fitzpatrick, P. F., and Russell, D. H. (1997) Anal. Chem. 69, 2862-2865 |
| 4. | Gadda, G., Edmondson, R. D., Russell, D. H., and Fitzpatrick, P. F. (1997) J. Biol. Chem. 272, 5563-5570 |
| 5. | Gadda, G., and Fitzpatrick, P. F. (1998) Biochemistry 37, 6154-6164 |
| 6. | Kido, T., Hashizume, K., and Soda, K. (1978) J. Bacteriol. 133, 53-58 |
| 7. | Gadda, G., and Fitzpatrick, P. F. (1999) Arch. Biochem. Biophys. 36, 309-313 |
| 8. | Porter, D. J. T., Voet, J. G., and Bright, H. J. (1973) Z. Naturforsch 27, 1052-1053 |
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| 10. | Kido, T., Tanizawa, K., Inagaki, K., Yoshimura, T., Ishida, M., Hashizume, K., and Soda, K. (1984) Agric. Biol. Chem. 48, 2549-2554 |
| 11. | Gorlatova, N., Tchorzewski, M., Kurihara, T., Soda, K., and Esaki, N. (1998) Appl. Environ. Microbiol. 64, 1029-1033 |
| 12. | Heasley, C. J., and Fitzpatrick, P. F. (1996) Biochem. Biophys. Res. Commun. 225, 6-10 |
| 13. | Gadda, G., and Fitzpatrick, P. F. (2000) Biochemistry 39, 1400-1405 |
| 14. | Gadda, G., and Fitzpatrick, P. F. (2000) Biochemistry 39, 1406-1410 |
| 15. | Gadda, G., Banerjee, A., and Fitzpatrick, P. F. (2000) Biochemistry 39, 1162-1168 |
| 16. | Riordan, J. F., and Vallee, B. L. (1972) Methods Enzymol. 11, 541-548 |
| 17. | Bradford, M. M. (1976) Anal. Biochem. 72, 248-254 |
| 18. | Edmondson, R. D., and Russell, D. H. (1996) J. Am. Soc. Mass Spectrom. 7, 995-1001 |
| 19. | Bjornberg, O., Rowland, P., Larsen, S., and Jensen, K. F. (1997) Biochemistry 36, 16197-16205 |
| 20. | Rowland, P., Bjornberg, O., Nielsen, F. S., Jensen, K. F., and Larsen, S. (1998) Protein Sci. 7, 1269-1279 |
| 21. | Quinn, D. M., and Sutton, L. D. (1991) in Enzyme Mechanisms from Isotope Effects (Cook, P. F., ed) , pp. 73-126, CRC Press, Inc., Boca Raton, FL |
| 22. | Jencks, W. P., and Regenstein, J. (1976) in Handbook of Biochemistry and Molecular Biology (Fasman, G. D., ed), 3rd Ed., Vol. I , pp. 305-351, CRC Press, Inc., Boca Raton, FL |
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