Characterization of active site residues of nitroalkane oxidase

Article abstract of DOI:10.1016/j.bioorg.2009.12.004

The flavoenzyme nitroalkane oxidase catalyzes the oxidation of primary and secondary nitroalkanes to the corresponding aldehydes and ketones plus nitrite. The structure of the enzyme shows that Ser171 forms a hydrogen bond to the flavin N5, suggesting that it plays a role in catalysis. Cys397 and Tyr398 were previously identified by chemical modification as potential active site residues. To more directly probe the roles of these residues, the S171A, S171V, S171T, C397S, and Y398F enzymes have been characterized with nitroethane as substrate. The C397S and Y398 enzymes were less stable than the wild-type enzyme, and the C397S enzyme routinely contained a substoichiometric amount of FAD. Analysis of the steady-state kinetic parameters for the mutant enzymes, including deuterium isotope effects, establishes that all of the mutations result in decreases in the rate constants for removal of the substrate proton by ～5-fold and decreases in the rate constant for product release of ～2-fold. Only the S171V and S171T mutations alter the rate constant for flavin oxidation. These results establish that these residues are not involved in catalysis, but rather are required for maintaining the protein structure.

Full text of DOI:10.1016/j.bioorg.2009.12.004

Bioorganic Chemistry 38 (2010) 115–119
Contents lists available at ScienceDirect
Bioorganic Chemistry
journal homepage: www.elsevier.com/locate/bioorg
Characterization of active site residues of nitroalkane oxidase
Michael P. Valley a, Nana S. Fenny a, Shah R. Ali a, Paul F. Fitzpatrick b,
*
a Department of Biochemistry and Biophysics, Texas A&M University, College Station, TX 77843, USA
b Department of Biochemistry, University of Texas Health Science Center, San Antonio, TX 78229, USA
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 17 November 2009
Available online 28 December 2009
The flavoenzyme nitroalkane oxidase catalyzes the oxidation of primary and secondary nitroalkanes to
the corresponding aldehydes and ketones plus nitrite. The structure of the enzyme shows that Ser171
forms a hydrogen bond to the flavin N5, suggesting that it plays a role in catalysis. Cys397 and Tyr398
were previously identified by chemical modification as potential active site residues. To more directly
probe the roles of these residues, the S171A, S171V, S171T, C397S, and Y398F enzymes have been char-
acterized with nitroethane as substrate. The C397S and Y398 enzymes were less stable than the wild-
type enzyme, and the C397S enzyme routinely contained a substoichiometric amount of FAD. Analysis
of the steady-state kinetic parameters for the mutant enzymes, including deuterium isotope effects,
establishes that all of the mutations result in decreases in the rate constants for removal of the substrate
proton by ꢀ5-fold and decreases in the rate constant for product release of ꢀ2-fold. Only the S171V and
S171T mutations alter the rate constant for flavin oxidation. These results establish that these residues
are not involved in catalysis, but rather are required for maintaining the protein structure.
Ó 2009 Elsevier Inc. All rights reserved.
Keywords:
Enzyme kinetics
Flavoprotein
Isotope effects
Mutagenesis
Active site
Nitroalkane oxidase
1. Introduction
structural basis by which a common structure can catalyze diver-
gent reactions, NAO allows comparison of the enzyme-catalyzed
The flavoenzyme nitroalkane oxidase (NAO) catalyzes the oxi-
dation of nitroalkanes to the corresponding aldehydes or ketones
with consumption of molecular oxygen and release of nitrite and
hydrogen peroxide (Scheme 1) [1]. While there is growing evi-
dence for hydride transfer mechanisms for the large family of fla-
voprotein oxidases [2–6], NAO is unusual in that it catalyzes
nitroalkane oxidation through formation of a carbanion intermedi-
ate (Scheme 2) [3]. Based on analysis of the sequence of the cloned
enzyme, NAO was identified as a homolog of the acyl-Co dehydro-
genase (ACAD) family of flavoproteins [7]. This assignment is con-
sistent with the similarities in the initial catalytic reactions of the
two enzymes; in both cases a protein carboxylate abstracts an
acidic proton from the substrate [1]. The subsequent determina-
tion of the structure of NAO [8] confirmed this assignment and pro-
vided insight into the use of this fold to catalyze the different
reactions. Critically, while both the cofactors and the active site
bases in NAO and ACAD occupy similar locations in the different
proteins, the substrates access the active site from opposite sides
of the protein [9], providing a structural basis for the reaction spec-
ificities. In addition to providing an opportunity to understand the
reaction with a well-studied solution reaction. The comparable
non-enzymatic reaction, formation of a nitroalkane anion from a
nitroalkane, has long been studied as a model for proton abstrac-
tion from carbon [10,11]. Recently, study of the oxidation of nitroe-
thane by NAO has allowed the evaluation of the contribution of
quantum mechanical tunneling to the rate increase in the en-
zyme-catalyzed reaction [12–14].
As with many other flavoenzymes, the catalytic mechanism of
NAO can be divided into oxidative and reductive half-reactions.
In the reductive half-reaction, Asp402 abstracts the
a-proton from
the neutral nitroalkane substrate to form a nitroalkane anion
which then attacks the N5 position of the FAD cofactor. The initial
adduct rearranges to release nitrite and generate a reactive cationic
imine species that reacts with hydroxide to eventually form re-
duced FAD and the aldehyde or ketone product. This cationic imine
has been trapped with cyanide and its structure determined
(Fig. 1), firmly establishing it as along the catalytic pathway
[9,15]. In the more typical oxidative half-reaction, molecular oxy-
gen attacks the reduced FAD to form hydrogen peroxide and regen-
erate the oxidized cofactor.
The knowledge of the structure of NAO has implicated several
active site residues as potentially important for catalysis. The ac-
tive site base of NAO, Asp402, is part of a catalytic triad that also
contains Arg409 and Ser276 (Fig. 1). Mutation of any of these three
residues decreases the rate constant for removal of the substrate
proton by 2–3 orders of magnitude, confirming their importance
Abbreviations: NAO, nitroalkane oxidase; ACAD, acyl-CoA dehydrogenase;
Knitroethane, Km value for nitroethane.
* Corresponding author. Address: Department of Biochemistry, UTHSCSA, MC
7760, San Antonio, TX 78229, USA.
E-mail address: fitzpatrick@biochem.uthscsa.edu (P.F. Fitzpatrick).
0045-2068/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved.
doi:10.1016/j.bioorg.2009.12.004

116
M.P. Valley et al. / Bioorganic Chemistry 38 (2010) 115–119
Scheme 1.
[9,16,17]. In addition, the structures of both the resting form of the
enzyme and of the cyanide-trapped intermediate show that the
hydroxyl of Ser171 forms a hydrogen bond with the N5 position
of the flavin, suggesting it may modulate the reactivity of the fla-
vin. A similar interaction is seen in the ACAD family, which con-
tains a conserved threonine residue that aligns with Ser171 in
NAO. Mutagenesis of this threonine to alanine in human med-
ium-chain ACAD significantly decreases the rate constant for flavin
reduction [18]. Finally, chemical modification studies have identi-
fied Cys397 [19] and Tyr398 [20] as putative active site residues in
NAO. We report here the results of site-directed mutagenesis to
investigate the roles these three residues play in catalysis.
2. Experimental procedures
2.1. Materials
Fig. 1. Interactions in the active site of nitroalkane oxidase. The structure is that of
the enzyme trapped with cyanide turning turnover with 1-nitrohexane (pdb code
3D9G) and shows the 5-cyanohexyl-FAD.
All chemicals were purchased from Sigma–Aldrich Chemical
Corp. (Milwaukee, WI). Recombinant nitroalkane oxidase was ex-
pressed and purified as previously described [7]. Mutations were
generated with the QuikChange Site-Directed Mutagenesis Kit
(Stratagene), and the mutant enzymes were expressed and purified
following the protocol for the wild-type enzyme. DNA sequencing of
the entire coding sequence of each mutant plasmid was performed
at the Laboratory for Plant Genome Technologies of Texas A&M Uni-
versity. Protein concentrations were determined by the method of
Bradford [21] with bovine serum albumin as standard; for all kinetic
measurements, enzyme concentrations were determined using an
puter-interfaced Hansatech Clark oxygen electrode (Hansatech
Instruments, Pentney King’s Lynn, UK). When varying the concen-
tration of nitroethane, the steady-state kinetic parameters were
determined at ambient oxygen concentrations. (This concentration
of oxygen is 5–10 times the Km value for oxygen for each mutant.)
To vary the concentration of oxygen at a constant concentration of
nitroethane, oxygen and argon were combined in different ratios
with a MaxBlend low flow air/oxygen blender (Maxtec Inc., Salt
Lake City, Utah), and the assay buffer was equilibrated with the
gas mixture. To prevent the formation of the anionic form of the
substrate, stock solutions of neutral nitroethane were prepared in
dimethyl sulfoxide and assays were initiated by the addition of
substrate.
e
446 value of 14.2 mMꢁ1 cmꢁ1, as previously described [22].
2.2. Methods
Enzyme activity was measured in 200 mM HEPES, 0.1 mM FAD,
pH 8.0, 30 °C, by monitoring oxygen consumption with a com-
Scheme 2.

M.P. Valley et al. / Bioorganic Chemistry 38 (2010) 115–119
117
Table 1
Steady-state kinetic parameters for NAO mutant enzymes.a
c
Enzyme
kcat (sꢁ1
)
kcat/Knitroethane (mMꢁ1 sꢁ1
)
Knitroethane (mM)
Kai (mM)
25
71 14
238 32
72 18
kcat/KO (mMꢁ1 sꢁ1
)
KO (lM)
2
2
Wild-typeb
S171A
S171V
S171T
Y398F
15
1
6.3 0.4
2.6 0.3
3.2 0.1
3.6 0.6
2.2 0.18
3.6 0.4
2.3 0.2
2.1 0.4
1.9 0.1
2.0 0.5
4.8 0.7
2.0 0.4
3
310 30
190 40
82
29
44
11
28
58
9
6
2
4
6
4
5.4 0.4
6.2 0.1
7.2 0.8
10.6 0.8
7.3 0.6
140
7
580 210
390 80
290 20
38
6
C397S
72 16
a
b
c
Conditions: pH 8.0, 30 °C.
From Ref. [17], with the exception of the oxygen kinetics.
Determined with 20 mM nitroethane.
Table 2
2.3. Data analysis
Steady-state kinetic isotope effects for NAO mutant enzymes.a
Data were analyzed using the programs KaleidaGraph (Synergy
Software) and Igor Pro (WaveMetrics, Inc., Lake Oswego, OR). Stea-
dy-state kinetic parameters were determined by fitting the data to
the Michaelis–Menten equation or to Eq. (1), which includes the ef-
Enzyme
Dkcat
D(kcat/Knitroethane
)
Wild-typeb
S171A
S171V
S171T
Y398F
1.4 0.2
1.9 0.2
1.6 0.1
2.4 0.4
1.9 0.3
2.4 0.3
9.2 1.1
11.6 2.3
14.6 1.3
8.5 2.8
10.8 1.7
8.5 2.5
fect of substrate inhibition. The parameter
v is the initial velocity,
kcat is the turnover number, Km is the Michaelis constant, S is the
substrate concentration, and Kai is the substrate inhibition con-
stant. Steady-state kinetic isotope effects were calculated from
Eq. (2), where Fi is the fraction of deuterium in the substrate and
Dkcat and D(kcat/Km) are the isotope effects on kcat and kcat/Km,
respectively.
C397S
a
Conditions: pH 8.0, 30 °C.
From Ref. [17].
b
strate for this mutant are given in Tables 1 and 2. The kinetic
parameters for the Y398F enzyme are all close to the wild-type val-
ues, with the largest difference being a threefold decrease in the
kcat/Km value for nitroethane. The isotope effects are similarly close
v
v
¼ kcatS=ðKm þ S þ S2=KaiÞ
ð1Þ
ð2Þ
kcat
S
¼
S2
Kai
D
D
D
Kmð1 þ Fið ðkcat=KmÞ ꢁ 1ÞÞ þ Sð1 þ Fið kcat ꢁ 1ÞÞ þ
to the wild-type values, with a small increase in the kcat value.
3.3. Mutagenesis of Cys397
3. Results
Cys397 was mutated to serine to determine the role of the
thiol in catalysis. Analyses of different preparations of the C397S
enzyme indicated that it was 50–90% apoenzyme. This implies
that the binding of flavin is perturbed by mutation of this residue.
Consistent with such a structural perturbation, the C397S enzyme
was less stable than the wild-type enzyme in that it more readily
precipitated from solution after multiple freeze/thaw cycles. The
steady-state kinetic parameters with nitroethane as substrate
for C397S enzyme are summarized in Table 1. These are unaf-
fected by the presence of apoenzyme, since all enzyme concentra-
tions were determined from the flavin content and thus the
3.1. Mutagenesis of Ser171
Ser171 was mutated to alanine, valine, and threonine to probe
its role in catalysis. In all cases the mutant enzymes were well-be-
haved and could readily be purified using the protocol developed
for the wild-type enzyme. The concentration of active enzyme
determined from the e446 value was typically equivalent to the to-
tal protein concentration, indicating ꢀ100% flavin occupancy. The
steady-state kinetic parameters of the mutant enzymes with
nitroethane as substrate are summarized in Table 1. For all three
mutant enzymes, the kcat values and kcat/Km values for nitroethane
for the three mutant enzymes are down 2 to 3-fold from the wild-
type values. The kcat/Km values for oxygen are similarly about 2-
fold less for the S171A and S171V enzymes, while the value for
the S171T enzyme is at least as large as the wild-type value. Deu-
terium kinetic isotope effects were also determined for the three
mutant proteins. The D(kcat/Knitroethane) values are large and compa-
values are for the holoenzyme only. The kcat value and the kcat
Km value for nitroethane decreased about 2-fold, while the kcat
/
/
Km value for oxygen is unchanged from the wild-type values. This
D
enzyme also shows the largest kcat value of any of the mutant
enzymes.
4. Discussion
D
rable to that for the wild-type enzyme. The kcat values for the
S171V and S171A enzymes show only a slight increase from the
wild-type value, while this isotope effect for the S171T enzyme
shows a larger increase.
None of the mutations described here had large effects on any
steady-state kinetic parameter. However, large changes in individ-
ual rate constants do not always have commensurate effects on
steady-state kinetic parameters. This is especially the case when
3.2. Mutagenesis of Tyr398
Tyr398 was mutated to phenylalanine to probe the role of the
phenol in catalysis. Flavin binding appeared to be unperturbed in
the Y398F enzyme, in that the purified enzyme contained a stoichi-
ometric amount of FAD. However, this mutant enzyme precipitated
from solution over prolonged periods of time at 30 °C, and thus is
less stable than the wild-type enzyme. The steady-state kinetic
parameters and deuterium isotope effects with nitroethane as sub-
Scheme 3.

118
M.P. Valley et al. / Bioorganic Chemistry 38 (2010) 115–119
Table 3
Intrinsic kinetic parameters for nitroalkane oxidase mutants.
Kinetic parameter
Wild-type NAOa
S171A
S171T
S171V
C397S
Y398F
Kd (mM)
14 1 (48 24)b
247 5 (310 150)
310 30
25
9
11
6
48 13
74 16
11
5
52 21
96 33
390 80
k3 (sꢁ1
)
49 12
190 40
6.1 0.5
42 13
580 210
8.7 1.1
42 10
290 20
8.8 0.8
k5 (mMꢁ1 sꢁ1
k7 (sꢁ1
)
140
7
)
17 (16 1)
5.8 0.4
12
1
a
From Ref. [17].
b
Values in parentheses for the wild-type enzyme were calculated from the data in Tables 1 and 2.
late the values of the individual rate constants in Scheme 3 for each
mutant enzyme (Table 3).1
Kd ¼ KDmðkcat=Km ꢁ 1Þ=ð kcat ꢁ 1Þ
ð3Þ
ð4Þ
D
k3k7
KO
¼
2
ðk3 þ k7Þk5
kcat ¼ k3k7=ðk3 þ k7Þ
ð5Þ
ð6Þ
D
Dkcat ¼ ð k3 þ k3=k7Þ=ð1 þ k3=k7Þ
When any of the three residues is mutated, multiple steps are af-
fected, suggesting that the decreases in activity are due to small
structural changes rather than loss of catalytically important resi-
dues. The largest effects of the mutations are on the rate constant
for proton abstraction from the substrate, k3, with smaller effects
on the rate constants for product release, k7. Given that NAO in-
creases the rate constant for proton abstraction for the substrate
by a billion-fold compared to the uncatalyzed reaction, [12], it is
not surprising that even a small perturbation of the structure would
decrease this rate constant. The active site of NAO is at the bottom of
a long tunnel, placing the FAD N5 ꢀ20 Å from the surface of the pro-
tein. Diffusion of the product out of the enzyme limits turnover for
the wild-typeenzyme; the decreases in k7 seen here with the mutant
enzymes suggests that even small disruptions in the overall protein
structure can slow this movement. The effects of the mutations on
the rate constant for the reaction with oxygen are smaller than those
on the reductive half-reaction, with only the S171A and S171V en-
zymes showing any change. The lack of a significant effect of mutat-
ing Ser171 establishes that the hydrogen bond between this residue
and the flavin N5 is not important for modulating the reactivity of
either the reductive or oxidative half-reactions. Instead, the hydro-
gen bond between Ser171 and the FAD may affect the active site
dynamics. This would be consistent with the introduction of larger
residues, but not replacement of serine with the smaller alanine,
altering the kinetics.
Cys397 and Tyr398 were identified as potential active site res-
idues by chemical modification. The modest effects of the C397A
and Y398F mutations on flavin binding and stability suggest that
these mutations have rather general disrupting effects on the pro-
tein structure. The sulfur of Cys397 is 4 Å from both of the flavin
methyl groups, and both Cys397 and Tyr398 are located in a helix
that terminates at Asp402 (Fig. 2). Chemical modification of either
and the resulting increase in bulk would be expected to signifi-
cantly perturb the positioning of the active site base and the flavin,
providing a reasonable explanation for the loss of activity when
these residues are modified. The much smaller effects seen with
the conservative mutations seen here confirm that the roles of
Cys397 and Tyr398 are structural rather than catalytic.
Fig. 2. Relative positions of Cys397, Tyr398, and FAD in nitroalkane oxidase.
the rate-limiting step for turnover is product release, as with NAO.
A combination of steady-state kinetic parameters and kinetic iso-
tope effects can resolve this problem, allowing one to partition
the effects of the mutations among the individual steps in
Scheme 3. The kinetic mechanism of NAO (Scheme 3) has been
established for the wild-type enzyme using a combination of stea-
dy state and rapid reaction kinetics [17]. This mechanism provides
a framework for analysis of the data for the mutant enzymes. With
nitroethane as substrate, the rate-limiting step in the reductive
half-reaction is cleavage of the substrate CH bond with rate con-
stant k3 [23]. Consequently, the deuterium isotope effect on the
kcat/Km value for nitroethane, D(kcat/Knitroethane), equals the intrinsic
isotope effect for CH bond cleavage, and the kcat/Km value for
nitroethane equals k3/Kd, with Kd the dissociation constant for
nitroethane. The relationship between the Km and the Kd values
is given by Eq. (3) [24]. Oxygen reacts with the reduced enzyme
with second-order kinetics, with no indication of saturation at
accessible levels of oxygen [17]. Such kinetics are typical of the
oxygen reaction of flavoprotein oxidases [25]. As a result, the value
of k5, the rate constant for the reaction of the reduced enzyme with
oxygen, is equivalent to the kcat/Km value for oxygen. The KO value
2
does not reflect a true binding event; rather, it equals kcat/(kcat/KO
)
2
(Eq. (4)). For wild-type NAO, overall turnover at saturating concen-
tration of nitroethane and oxygen is limited by product release
from the oxidized enzyme, with rate constant k7 [17]; because this
step is 19-fold slower than reduction with the wild-type enzyme,
the isotope effect on the kcat value is small. Eq. (5) gives the rela-
tionship between the kcat value and k3 and k7. The relationship be-
The residues in NAO analyzed herein by site-directed mutagen-
esis were selected either because the structure of the protein
1
The values in Table 3 were calculated using 9.2 1.1 as the intrinsic isotope effect
for all the enzymes, since the D(kcat/Knitroethane) values for all four mutant enzymes are
tween the isotope effect on kcat,
Dkcat, and the intrinsic isotope
equivalent to the value for the wild-type enzyme. Use of the individual D(kcat
/
effect on the CH bond cleavage step, Dk3, is given by Eq. (6). These
Knitroethane) values for the mutant enzymes does not have a significant effect on the
calculated values.
relationships allow the data of Tables 1 and 2 to be used to calcu-

M.P. Valley et al. / Bioorganic Chemistry 38 (2010) 115–119
119
[9] A. Héroux, D.M. Bozinovski, M.P. Valley, P.F. Fitzpatrick, A.M. Orville,
Biochemistry 48 (15) (2009) 3407–3416.
[10] C.F. Bernasconi, Accounts Chem. Res. 25 (1992) 9–16.
[11] R.P. Bell, in: The Proton in Chemistry, second ed., Cornell University Press,
Ithaca, 1973, pp. 250–296.
[12] M.P. Valley, P.F. Fitzpatrick, J. Am. Chem. Soc. 126 (20) (2004) 6244–6245.
[13] D.T. Major, D.M. York, J. Gao, J. Am. Chem. Soc. 127 (47) (2005) 16374–16375.
[14] D.T. Major, A. Heroux, A.M. Orville, M.P. Valley, P.F. Fitzpatrick, J. Gao, Proc.
Natl. Acad. Sci. USA 106 (2009) 20734–20739.
suggests they play a catalytic role (Ser171) or because previous
solution studies suggested that they were near the active site
(Cys397 and Tyr398). In all three cases, the effects of the mutations
are too small to consider these residues critical for catalysis.
Acknowledgment
[15] M.P. Valley, S.E. Tichy, P.F. Fitzpatrick, J. Am. Chem. Soc. 127 (7) (2005) 2062–
2066.
This research was supported in part by NIH Grant GM58698.
[16] P.F. Fitzpatrick, M.P. Valley, D.M. Bozinovski, P.G. Shaw, A. Héroux, A.M.
Orville, Biochemistry 46 (2007) 13800–13808.
[17] M.P. Valley, P.F. Fitzpatrick, Biochemistry 42 (19) (2003) 5850–8566.
[18] B. Kuchler, A.G. Abdel-Ghany, P. Bross, A. Nandy, I. Rasched, S. Ghisla, Biochem.
J. 337 (Pt 2) (1999) 225–230.
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