Steady-state substrate specificity and O2-coupling efficiency of mouse cysteine dioxygenase

Article abstract of DOI:10.1016/j.abb.2014.11.004

Cysteine dioxygenase (CDO) is a non-heme mononuclear iron enzyme that catalyzes the oxygen-dependent oxidation of l-cysteine (Cys) to produce l-cysteine sulfinic acid (CSA). Sequence alignment of mammalian CDO with recently discovered thiol dioxygenase enzymes suggests that the mononuclear iron site within all enzymes in this class share a common 3-His first coordination sphere. This implies a similar mechanistic paradigm among thiol dioxygenase enzymes. Although steady-state studies were first reported for mammalian CDO over 45 years ago, detailed analysis of the specificity for alternative thiol-bearing substrates and their oxidative coupling efficiencies have not been reported for this enzyme. Assuming a similar mechanistic theme among this class of enzymes, characterization of the CDO substrate specificity may provide valuable insight into substrate-active site intermolecular during thiol oxidation. In this work, the substrate-specificity for wild-type Mus musculus CDO was investigated using NMR spectroscopy and LC-MS for a variety of thiol-bearing substrates. Tandem mass spectrometry was used to confirm dioxygenase activity for each non-native substrate investigated. Steady-state Michaelis-Menten parameters for sulfinic acid product formation and O₂-consumption were compared to establish the coupling efficiency for each reaction. In light of these results, the minimal substrate requirements for CDO catalysis and O₂-activation are discussed.

Full text of DOI:10.1016/j.abb.2014.11.004

Archives of Biochemistry and Biophysics 565 (2015) 49–56
Contents lists available at ScienceDirect
Archives of Biochemistry and Biophysics
journal homepage: www.elsevier.com/locate/yabbi
Steady-state substrate specificity and O2-coupling efficiency of mouse
cysteine dioxygenase q
⇑
Wei Li, Brad S. Pierce
Department of Chemistry and Biochemistry, College of Sciences, The University of Texas at Arlington, Arlington, TX 76019, United States
a r t i c l e i n f o
a b s t r a c t
Article history:
Cysteine dioxygenase (CDO) is a non-heme mononuclear iron enzyme that catalyzes the oxygen-
dependent oxidation of L-cysteine (Cys) to produce L-cysteine sulfinic acid (CSA). Sequence alignment
Received 3 September 2014
and in revised form 7 November 2014
Available online 20 November 2014
of mammalian CDO with recently discovered thiol dioxygenase enzymes suggests that the mononuclear
iron site within all enzymes in this class share a common 3-His first coordination sphere. This implies a
similar mechanistic paradigm among thiol dioxygenase enzymes. Although steady-state studies were
first reported for mammalian CDO over 45 years ago, detailed analysis of the specificity for alternative
thiol-bearing substrates and their oxidative coupling efficiencies have not been reported for this enzyme.
Assuming a similar mechanistic theme among this class of enzymes, characterization of the CDO
substrate specificity may provide valuable insight into substrate-active site intermolecular during thiol
oxidation. In this work, the substrate-specificity for wild-type Mus musculus CDO was investigated using
NMR spectroscopy and LC–MS for a variety of thiol-bearing substrates. Tandem mass spectrometry was
used to confirm dioxygenase activity for each non-native substrate investigated. Steady-state Michaelis–
Menten parameters for sulfinic acid product formation and O2-consumption were compared to establish
the coupling efficiency for each reaction. In light of these results, the minimal substrate requirements for
CDO catalysis and O2-activation are discussed.
Keywords:
Non-heme iron
Cysteine
Dioxygenase
Specificity
Coupling
Thiols
Ó 2014 Elsevier Inc. All rights reserved.
Introduction
disease [1–4]. Recently, the interplay between dysfunction in sulfur
metabolism and human neurodegenerative disease states (Alzhei-
Cysteine dioxygenase (CDO)1 is a mononuclear non-heme iron
enzyme that catalyzes the first concerted step in the O2-dependent
oxidation of L-cysteine (Cys) to produce cysteine sulfinic acid (CSA)
(Scheme 1). Enzymes involved in sulfur-oxidation and transfer are
increasingly being recognized as potential drug targets for develop-
ment of antimicrobials, therapies for cancer, and inflammatory
mers, autism, and Parkinsons) has been of considerable medical
interest [5–7].
Multiple high resolution crystal structures of the resting and
substrate-bound enzyme have been solved which highlight the
atypical mononuclear iron coordination for the mammalian CDO
active site [6,8–10]. Among the non-heme mononuclear iron oxi-
dase/oxygenase class of enzymes, the typical Fe-coordination
sphere is comprised of two protein-derived neutral His residues
and one monoanionic carboxylate ligand provided by either an
Asp or Glu residue. Unlike most enzymes within this family, one
face of the CDO mononuclear iron active site (Fig. 1) is coordinated
by 3 protein derived histidine residues resulting in a 3-His facial
triad. Another unusual feature observed within the mature eukary-
otic CDO active site is a post-translational modification in which
spatially adjacent Cys93 and Tyr157 residues are covalently
cross-linked to produce a C93-Y157 pair. Among CDO enzymes
identified, Y157 is conserved across phylogenic domains whereas
the C93-Y157 pair is unique to eukaryotes. Several reports have
demonstrated that formation of this cross-link increases the
catalytic activity and coupling efficiency of CDO [11–13]. Regard-
less, C93A CDO variants, which lack the ability to produce the
q
This work was supported by the National Science Foundation (CHE) 1213655
(B.S.P.).
⇑
Corresponding author at: Department of Chemistry and Biochemistry, 700
Planetarium Place, Room 130, The University of Texas at Arlington, Arlington, TX
76019, United States. Fax: +1 (817) 272 3808.
E-mail address: bspierce@uta.edu (B.S. Pierce).
Abbreviations used: TDO, thiol dioxygenase; CDO, cysteine dioxygenase; ADO,
1
cysteamine dioxygenase; MSDO, mercaptosuccinate dioxygenase; MPDO, 3-merca-
ptopropionate dioxygenase; Cys, cysteine; Hcy, homocysteine; CA, 2-aminoethane-
thiol (cysteamine); MS, mercaptosuccinate; PA, L-penicillamine, CME, L-cysteine
methyl ester; DME, 2-(dimethylamino)ethanethiol; SC, S-methyl-L-cysteine; 3MP,
3-mercaptopropionate; 1MP; 1-mercaptopropane; HT, hypotaurine; TMSP, trimeth-
ylsilyl propanoic acid; TCEP, tris(2-carboxyethyl)phosphine hydrochloride; HPLC,
high performance liquid chromatography; NMR, nuclear magnetic resonance; LC–MS,
liquid chromatography–mass spectrometry; MRM, multiple reaction monitoring;
SIM, single ion mode.
http://dx.doi.org/10.1016/j.abb.2014.11.004
0003-9861/Ó 2014 Elsevier Inc. All rights reserved.

50
W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
C93-Y157 pair, retain their catalytic activity. Therefore, this post-
translational modification does not appear to be required for catal-
ysis. The exact mechanism of C93-Y157 formation remains
unresolved.
As shown in Fig. 1, the L-Cys substrate coordinates to the Fe-site
in a bidentate manner via neutral amine and thiolate functional
groups [10]. Like other members of the non-heme mononuclear
iron enzyme family, CDO exhibits an obligate-ordered binding of
the
L-Cys substrate prior to molecular oxygen [14,15].
Cysteine dioxygenase and cysteamine (2-aminoethanethiol)
dioxygenase (ADO) are the only known mammalian thiol dioxy-
genase (TDO) enzymes. Until recently, the catabolic dissimilation
of L-cysteine was believed to be unique within the domain of
eukaryotes [16,17]. However, a number of bacterial TDO enzymes
have now been identified, suggesting that the ability to oxidize
excess thiols is advantageous for survival. For example, two
Fe/O2-dependent TDO enzymes isolated from Variovorax paradoxus
have recently been identified [mercaptosuccinate dioxygenase,
(MSDO) and 3-mercaptopropionate dioxygenase (MPDO)] [18,19].
Sequence comparison suggests these enzymes also contain a
3-His active site motif. The conserved metal binding site among
TDO enzymes suggest that the first-coordination sphere is neces-
sary for thiol oxidation whereas the outer-sphere residues most
likely facilitate binding of their specific substrates. This implies
that TDO enzymes have the potential to catalyze the O2-dependent
oxidation of a variety of thiol-substrates to produce the corre-
sponding sulfinic acid providing they are capable of binding to
the mononuclear active site.
All TDO enzymes identified belong to the cupin superfamily
which is defined on the basis of a characteristic b-barrel tertiary
structure. Despite significant deviations in thiol-substrates and
amino acid sequence homology, several conserved features can
be identified among TDO enzymes. A sequence alignment for
selected TDO enzymes is provided in Supplemental information,
Fig. S1. From this analysis, the 3-His active site motif appears to
be a common feature among TDO enzymes. Beyond the active site
3-His residues, several conserved residues are observed in CDO
enzymes across phylogenic domains (Y58, R60, H155, and Y157).
For example, Y157 and H155 residues are present in both eukary-
otic and prokaryotic enzyme forms. Moreover, both Y157F and
H155A CDO variants exhibit abolished (or minimal) enzymatic
activity [20,21]. Recent spectroscopic experiments on the catalyti-
cally inactive ferric enzyme suggest that these residues are
involved in key outer-sphere interactions with the substrate-
bound active site to facilitate catalysis. By contrast, the C93
involved in C93-Y157 cross-link formation is only observed in
mammalian CDO enzymes. In bacterial enzymes this residue is
replaced by a glycine suggesting that this cross-link is not catalyt-
ically essential. In support of this hypothesis, C93A and C93S
variants of mammalian CDO retain catalytic activity, albeit with
decreased kcat [13,20]. X-ray crystallography and computational
models suggest that R60 is involved in charge stabilizing of the
Fig. 1. 1.60 Å resolution X-ray crystal structure for the substrate-bound CDO active
site at pH 8.0 (pdb code 4IEV) [10].
This seems unlikely as S153 is not universally conserved across
phylogenic domains similar to H155 and Y157. No experiments
to data provide any insight into the catalytic role of these residues.
Previous studies suggest that mammalian CDO exhibits high
substrate and stereoselectivity [23–26]. For example, it has been
reported that
L-homocysteine is an inhibitor of CDO but its
potential as a substrate has not been carefully evaluated [27]. In
fact detailed characterization of any non-natural substrates is
severely underreported. In this work, a variety of commercially
available thiol-substrates were utilized to evaluate the substrate-
specificity of wild-type CDO cloned from Mus musculus. Substrates
were selected to evaluate specific active site interactions and
cross-reactivity among other physiological TDO substrates [ADO
(cysteamine), MPDO (3-mercaptopropionate), and MSDO (mercapto-
succinate)]. The steady-state kinetic parameters and coupling
efficiencies for O2-consumption and sulfinic acid formation are
reported. In all substrates evaluated, NMR spectroscopy, differen-
tial 16O/18O-incorperation, and tandem LC–MS/MS were employed
to confirm formation of the appropriate sulfinic acid product.
Materials and methods
Enzyme purification
Recombinant mouse CDO was expressed in Escherichia coli
BL21(DE3) pLysS competent cells (Novagen) and purified using a
10 L bioreactor (New Brunswick Scientific Bioflo100) as previously
described [14]. The as-isolated CDO enzyme typically contains
ꢀ50% ( 10%) of the C93-Y157 cross-link as observed by SDS PAGE.
Therefore, prior to use, isolated CDO is converted to the fully mod-
ified enzyme as described elsewhere [13]. All preparations were
assayed for ferrous and ferric iron content spectrophotometrically
as previously described [15,28]. Typical ferrous iron incorporation
within purified CDO is ꢀ 70% ( 10%). For clarity, the concentrations
reported in enzymatic assays reflect the concentration of ferrous
iron within samples of CDO (FeII-CDO).
L-Cys carboxylate group. This Arg residue is absent in mammalian
and bacterial ADO enzymes which catalyze the O2-dependent oxi-
dation of 2-aminoethanethiol (cysteamine). It has also been noted
that Ser153, His155 and Tyr157 may form a ‘‘catalytic triad’ similar
to those observed among hydrolase or transferase enzymes [22].
NMR kinetic study
NMR kinetic studies were performed on a 300 JEOL nuclear
magnetic resonance spectrometer (Pleasanton, CA). All measure-
ments were made in Wilmad NMR tubes (standard wall, 5 mm
O.D., precision, 507-PP-7). For each reaction, fully modified CDO
(typically 2–25 lM) was added to a buffered substrate solution
in D2O (sodium phosphate buffer, 50 mM NaCl, pD 7.5) to initiate
the reaction at ambient temperature (20 2 °C). Reaction points
were terminated by heat shock at 95 °C for 2 min followed by
spin-filtration to remove denatured protein. Final concentration
Scheme 1. CDO catalyzed oxidation of L-cysteine to produce cysteine sulfinic acid.

W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
51
of 1 mM trimethylsilyl propanoic acid (TMSP) was added as the
internal standard. NMR spectra were integrated using JOEL USA
Delta NMR data processing software (version 5.0.4). The corrected
value of pD was obtained by adding 0.4 pD units to the value
reported by the pH electrode (Mettler Toledo InLab Expert Pro)
[29].
as previously described [13,14]. Instrumentation: Shimadzu
LCMS-2020; Column, Phenomenex Kinetex C18, 100 Å column
100 Â 4.6 mm, 2.6 m; Mobile phase, 20 mM sodium acetate, 0.6%
methanol, 1% heptafluorobutyric acid, pH 2.0; Injection volume,
50
were detected spectrophotometrically at 218 nm. Each reaction
was initiated by addition of enzyme (1 M) to a buffered solution
lL; Flow rate, 1.0 mL/min. Product CSA and hypotaurine peaks
l
Circular dichroism (CD)
(25 mM phosphate, 50 mM NaCl, pD 7.5) containing substrate at
ambient temperature (20 2 °C). At selected times, aliquots were
CD analysis of CDO in H2O and D2O buffer was performed on
JASCO 715 UV–visible circular dichroism spectrometer with xenon
arc light source. Protein samples analyzed by CD were prepared in
10 mM phosphate buffer, 50 mM NaCl, pL 7.5 filtered through a
collected and quenched by addition of 10 lL of 40 mM hydrochlo-
ric acid. Following addition of HCl, samples were heated to 95 °C
for 3 min to ensure full enzyme denaturation and then spin-filtered
by 0.22 lM cellulose acetate membrane (Corning, Spin-X) prior to
0.22
Equine heart myoglobin (100684-32-0), chicken egg white lyso-
zyme (12650-88-3), and poly- -lysine (25988-63-0) purchased
from Sigma–Aldrich were used as standards for secondary struc-
ture determination as described elsewhere [30]. CD results were
also interpreted using the freely available online software K2D3
(http://k2d3.ogic.ca/). Far-UV (185–260 nm) CD spectra were
recorded in a quartz cuvette of 0.1 cm path length and ꢀ0.45 mL
volume at a scan speed of 40 nm/min at 20 2 °C.
l
m
polypropylene membrane filter (VWR international).
analysis on HPLC. The concentration of CSA and hypotaurine pro-
duced in reactions were determined by comparison to standard
calibration curves (0.1–20 mM). Steady-state kinetic parameters
for CDO were determined by fitting data to the Michaelis–Menten
equation using the program SigmaPlot ver. 11.0 (Systat Software
Inc., Chicago, IL).
L
LC–MS and LC–MS/MS analysis
Detection and verification of enzymatic products were per-
formed on a triple quadrupole LC–MS/MS (Shimadzu Scientific
Instruments, LC–MS 8040) in positive ion mode. Instrumentation:
Oxygen electrode
The rate of dissolved oxygen consumption utilized in CDO reac-
tions was determined using a standard Clark type electrode
(Hansatech Instruments, Norfolk, England). Reaction temperatures
were maintained at 20 2 °C) using a circulating water bath (Ther-
moFlex 900, Thermo Scientific). Calibration of the O2-electrode was
performed as previously described [13]. For each potential CDO
substrate utilized, 1.0 mL of a stock substrate solution was pre-
pared in a buffered solution (25 mM HEPES pH 7.5 or equivalent
phosphate buffer), and incubated at 20 °C for 3–5 min to equili-
brate the reaction solution to the cell temperature and establish
a baseline for O2-electrode. Reactions were initiated by addition
of CDO to obtain a final enzyme concentration within the cell of
Column, Phenomenex Luna 3
(P/N 00D-4449-B0); Mobile phase, 70% ACN, 30% H2O, 30 mM
NH4AC, 0.1% trifluoroacetic acid; Injection volume, 2 L; flow rate,
lm HILIC 200Å, 100 Â 2.00 mm,
l
0.25 mL/min. Confirmation of CDO product was verified multiple
reaction monitoring (MRM) using a triple quadrupole LC–MS/MS
[Shimadzu Scientific Instruments, LCMS 8040] [32,33]. The molecu-
lar ions (M+) of the CDO products (CSA, 154 m/z and hypotaurine,
110 m/z) were selected for secondary fragmentation. MRM optimi-
zation was then employed to maximize transition intensity and sen-
sitivity for each fragment. The optimized MRM method was used to
verify both substrate and product by direct injection of enzymatic
assays. These results were compared to direct injection of standards.
Additional verification of dioxygenase activity was confirmed
by select ion mode (SIM) in LC–MS. In these experiments, the mass
of the molecular ion (M+) was compared following the exchange of
16O2 molecular oxygen for 18O2. Incorporation of both oxygen
atoms into the substrate should result in an increase in the molec-
ular ion of 4 m/z.
1 lM.
18O2 enzymatic reactions
Enzyme and substrate solutions were rigorously degassed on a
Schlenk line prior to transferring into the anaerobic chamber. Ana-
lytical grade argon was passed through a copper catalyst (Kontes,
Vineland, N.J.) to remove atmospheric 16O2 impurities and then
sparged through distilled water to hydrate gas. All anaerobic sam-
ples were prepared within sealed vials in a glove box (Coy Labora-
tory Products Inc., Grass City, MI) with the O2 concentration
maintained below 1 ppm. 18O2 reactions were prepared within
the anaerobic chamber by adding excess substrate (25 mM) to a
Results
Validation of CDO steady-state kinetics utilizing native (
substrate by NMR, O2-electrode, and mass spectrometry
L-Cys)
15 mL tube (VWR Catalog Number 89049-170) containing 5.0
lM
The rate of CDO catalyzed L-cysteine oxidation to produce CSA
CDO. Each vial was sealed using a rubber septum (ChemGlass
P/N CG-3022-93) within the glove box and secured by standard
electrical tape. 18O2-saturated buffer (99%) was prepared by sparg-
ing anaerobic buffer with 18O2 gas (Icon 99% 18O2, P/N 11135). For
all substrates, 500
the septum sealed reaction vial by gas-tight Hamilton syringe
has been well characterized by utilizing both HPLC, O2-electrode,
and LC–MS [8,13,34]. While significantly less sensitive than these
methods, NMR does offer a greater flexibility in monitoring a
potentially broad range of sulfinic acids produced by CDO.
Moreover, re-optimization of mobile phase or assay conditions
for each individual substrate is time consuming and impractical.
Since NMR has not previously been utilized to study CDO activity,
it is important to first validate this method using the native CDO
l
L of the 18O2-satuated buffer was spiked into
resulting in an approximate final O2 concentration of ꢀ226
lM
[31]. Reaction mixtures were mixed by gentle inversion and
allowed to react for >1 h prior to heat denaturation, spin-filtration,
and work up for LC–MS analysis.
substrate (
L
-Cys) prior to proceeding to non-native substrates.
-cysteine
As an initial point of comparison, the NMR spectra for
L
and CSA standards within a sodium phosphate buffer in D2O (pD 7.5)
are shown in Supplemental Information, Fig. S2 (panel A). Within
this spectral window (2.50–4.40 ppm), only the non-exchangeable
HPLC analysis
CDO catalyzed oxidation of
L
/D-cysteine and 2-aminoethane-
protons on the
a- and b-carbons (Scheme 1) are observed. Both have
thiol (cysteamine) was performed by isocratic reverse phase HPLC
clearly resolved resonances which can be utilized to monitor either

52
W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
the formation of CSA or loss of
L
-Cys over the course of a typical reac-
(v0/[E]), such that the values for kcat and KM are easily obtained
from fitting the results to the Michaelis–Menten equation. The
steady-state kinetics for wild-type CDO with -Cys obtained by
tion. The NMR spectra of
observed as a doublet of doublets (dd) centered at 3.02, 3.10, and
3.97 ppm. By comparison, the same /b-protons observed for CSA
are observed at 2.70, 2.84, and 4.16 ppm. Relative to the -Cys
L-Cys exhibits three sets of resonances
L
a
NMR spectroscopy is shown in Fig. 2B. The solid line represents a
best-fit to the initial rate of CSA formation (h) as a function of
substrate concentration. From this analysis, the kcat and KM values
for CSA formation were determined to be 0.74 0.06 sÀ1 and
7.3 1.5 mM, respectively. The steady-state kinetics for CSA for-
mation obtained by HPLC under identical reaction conditions in
D2O is overlaid on the same plot (j) for comparison. Here, the
Michaelis–Menten parameters, kcat and KM, were determined to
be 0.65 0.05 sÀ1 and 7.0 1.3 mM, respectively. Therefore, under
equivalent conditions both HPLC and NMR yield nearly equivalent
steady-state results. It should be noted that deuterium adversely
affects both the maximal velocity of CSA formation and the appar-
ent KM-value obtained in steady-state assays. Michaelis–Menten
parameters (kcat and KM) obtained under identical conditions in
H2O were determined to be 1.3 0.2 sÀ1 and 2.3 0.3 mM,
respectively.
The efficiency at which an oxygenase enzyme incorporates one
mol of O2 into the product is referred to as ‘coupling’. Under
steady-state conditions, the (CSA/O2)-coupling efficiency is defined
as the ratio of the kcat determined for CSA formation divided by the
kcat obtained for the rate of O2-consumption. Interestingly, D2O
also has a significant impact on the (CSA/O2) coupling efficiency
for CDO. The coupling efficiency of fully-modified CDO catalyzed
L
starting material, the diastereotopic protons on the b-carbon exhibit
the greatest overall change in chemical shift due to the close prox-
imity of the adjacent sulfinic acid group. For simplicity, the
for -Cys and CSA (3.97 and 4.16 ppm, respectively) were utilized to
monitor the rate of CDO catalyzed CSA formation.
a-proton
L
To demonstrate the utility of NMR spectroscopy for monitoring
the steady state kinetics for CDO catalyzed reactions, both the
decay of
Each reaction was initiated by addition of CDO (0.5
fered D2O solution (25 mM phosphate, 50 mM NaCl, pD 7.5,
20 °C) containing 4.5 mM -Cys. At selected time points, samples
L
-Cys and formation of CSA were monitored with time.
lM) to a buf-
L
were quenched, spin-filtered, and analyzed by NMR spectroscopy.
Trimethylsilyl propanoic acid (TMSP) was added as an internal
standard to each sample for normalization of signal intensity as
described in Material and Methods. Representative NMR spectra
for the CDO catalyzed formation of CSA are shown in Supporting
Information, Fig. S2 (panel B). Quantitation of each species was
performed by comparison of the integrated peak area to freshly
prepared standards of known concentration. Fig. 2A shows the
time-dependent integrated results for
Data were fit to either a single exponential decay (
L
-Cys and CSA (
a-protons).
L-Cys) or a single
exponential rise to maximum (CSA) using the same observed rate
constant (kobs ꢀ 0.12 0.03 hÀ1). The amplitude (A) of the expo-
nential phase was 4.0 0.15 mM, representing ꢀ88% of theoretical
value (4.5 mM). The initial velocity (v0) is obtained from the prod-
uct of the reaction amplitude and rate constant (v0 = kobsÁA). By
L-Cys oxidation was previously reported as 75–80% [13]. Under
identical conditions within a deuterium buffer, the observed
(CSA/O2)-coupling (pD = 7.5) is significantly lower (41 5%). The
observed kcat, KM, and coupling efficiency all appear to be nega-
tively influenced by substituting D2O for H2O. While beyond the
scope of this study on substrate-specificity, a full report of the
solvent isotope effects on CDO catalysis, oxidative coupling, and
proton-inventory is described elsewhere [35].
It is also possible explanation that deuterium negatively influ-
ences the stability of the protein resulting in decreased activity.
Therefore, CD spectroscopy was used to evaluate the influence of
normalizing this value by the concentration of enzyme (0.5
lM),
the initial velocity for CSA formation at 4.5 mM
L-Cys is obtained
(v0/[E] = 0.27 0.07 sÀ1). These results indicate that over the
course of this reaction, nearly stoichiometric formation of CSA is
observed and its rate of formation is kinetically matched to the loss
of
L
-Cys.
Using this method, steady state kinetics for CDO catalyzed CSA
formation were performed as described above while varying the
concentration of -Cys from 1 to 20 mM. Rather than monitor reac-
D2O on secondary structure. In these experiments, 5 lM samples
of CDO were prepared in 25 mM phosphate, 50 mM NaCl, pL 7.5,
20 °C. As illustrated in Supplemental Information, Fig. S3, the UV
CD spectra [185–260 nm] of fully-modified CDO shows a maxima
at 193 nm and two minima at 208 and 222 nm. Analysis of the
CDO secondary structure was made by comparison to known pro-
L
tions over an extended time as described above, the initial rate (v0)
of each reaction was collected within the linear regime of product
formation (ꢀ45-min). To compensate for shorter reaction times,
the enzyme concentration was increased to 4
lM. As before, all ini-
tein standards (myoglobin, poly-L-lysine, and lysozyme) utilizing
tial rates are normalized for Fe-containing enzyme concentration
the freely available software K2D3 as described in Materials and
Fig. 2. (A) Time course of
black circle) using the same observed rate constant of kobs ꢀ 0.12 0.03 h; amplitude, 4.0 0.15 mM (ꢀ84% of expected). The initial velocity for CSA formation at 4.5 mM
is obtained (v0/[E] = 0.27 0.07 sÀ1) by normalizing the observed rate by the concentration of enzyme (0.5
M). B. Comparative Michaelis–Menten plot for CDO catalyzed
steady-state CSA formation determined by NMR (h) and HPLC (j). Assay conditions: 25 mM sodium phosphate, 50 mM NaCl, pD 7.5, 20 2 °C.
L
-Cys decay and CSA formation with time. Each data set was fit to either a single exponential decay (
L
-Cys, white circle) or rise to maximum (CSA,
L
-Cys
l

W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
53
methods. From this analysis, it was observed that the fraction of
-helical and b-sheet secondary structure in buffer prepared in
H2O is 14 4% and 35 3%, respectively. These values correlate
well with the percent -helical (17%) b-sheet (34%) content deter-
mined by X-ray crystallography [10]. By comparison, no significant
change is observed in the fraction of -helix (15%) or b-sheet (33%)
relative to H2O. With this in mind, NMR can be used to evaluate
the relative substrate specificity for CDO using a wide range of
commercially available thiol-substrates. For each substrate evalu-
ated, confirmation of dioxygenase product was independently ver-
ified by mass spectrometry using the methods described above.
a
a
a
observed for CDO in deuterium buffer. This result indicates that the
perturbations observed in the steady-state kinetic parameters can-
not be attributed to a change in the secondary structure of CDO.
While LC–MS methods for CSA detection are described in the
literature, the use of tandem LC–MS/MS has not previously been
demonstrated. In these experiments, both methods were utilized
to verify the identity of the sulfinic acid product produced in
CDO catalyzed reactions and confirm dioxygenase activity. Multi-
ple reaction monitoring (MRM) was performed to verify CSA
formation as described in Materials and methods. In these experi-
ments, the [M+H]+ molecular ion for CSA (154 m/z) was selected for
secondary fragmentation. MRM optimization was then employed
to maximize transition intensity and sensitivity for each fragment
allowing for quantitation of product ions. The optimized MRM
method was used to verify sulfinic acid product by direct injection
of enzymatic assays. These results were compared to direct injec-
tion of standards. Fig. 3(panel A) shows the MRM spectra for a
standard solution of CSA. In addition to the 154 m/z parent
[M + H]+ ion, two additional ions are observed at 44 and 74 m/z.
CDO catalyzed formation of hypotaurine from 2-aminoethanethiol
(cysteamine)
The NMR spectra for cysteamine and hypotaurine in D2O (pD
7.5) are shown in Supplemental Information, Fig. S4. The non-
exchangeable protons on the a- and b-carbon are observed within
the spectral window (2.50–4.00 ppm). The resonances for the
b-protons of cysteamine (d = 2.84, white square) and hypotaurine
(d = 2.65, black square) are well resolved and thus can be monitored
over the course of a typical reaction. Cysteamine exhibits 2 sets of
resonances observed as triplets (t) centered at 2.84 and 3.21 ppm.
By comparison, the same a/b-protons observed for hypotaurine are
observed at 2.65 and 3.36 ppm. Since the b-protons peaks slightly
overlap for substrate and product (3.21 and 3.36 ppm, respec-
tively), the
a-protons were selected to monitor the progress of
the CDO catalyzed reaction.
For reactions involving non-natural substrates, the concentra-
tion of CDO used in these experiments was increased to compen-
sate for slower reaction rates. As described previously, reactions
Fig. 3B demonstrates that direct injection (2
lL) of the CDO cata-
were initiated by addition of enzyme CDO (50 lM) to a buffered
lyzed reaction with -Cys yields and identical fragmentation pat-
L
D2O solution containing 90 mM cysteamine (20 °C). In these reac-
tions, very low (but detectable) features associated with hypotau-
rine (d = 2.65 ppm) formation could be observed in CDO reactions
(Fig. S4). Steady state kinetics for CDO catalyzed hypotaurine for-
mation was performed at varying cysteamine concentrations rang-
ing from 25 to 100 mM. Under these conditions a linear increase in
the enzyme normalized initial (v0/[E]) rate can be observed with
increasing substrate concentration. However, even at these ele-
vated substrate concentrations, full enzyme saturation kinetics
was never observed and thus a value for KM could not be deter-
mined for cysteamine. Regardless, from the best fit to the linear
portion of the curve, the pseudo-second order rate constant
(approximately kcat/KM) was determined (kobs ꢀ 0.01 MÀ1 sÀ1). For
comparison to the product formation data, the efficiency by which
CDO incorporates one mol of O2 into the hypotaurine product was
also determined for cysteamine reactions. Similar to product for-
mation reactions, saturation kinetics as monitored by the Clarke-
type O2-electrode is also not obtained and thus the pseudo-second
order rate constant for O2-consumption was determined from the
best fit to the linear portion of the curve (kobs ꢀ 0.63 MÀ1 sÀ1).
Therefore, in reactions with cysteamine, CDO exhibits >1% coupling
efficiency.
tern. The matching fragmentation pattern and relative intensities
confirm the native CSA product within CDO reactions.
As final confirmation of dioxygenase activity, selected ion mode
(SIM) was used to observe an appropriate shift in the observed
product [M + H]+ ion upon substitution of 18O2 for the naturally
abundant 16O2. As illustrated in Fig. 3C, CDO reactions performed
in the presence of 18O2 results in a + 4 m/z shift (154 m/z versus
158 m/z) in the observed [M+H]+ parent ion, thus verifying the
inclusion of both oxygen atoms into the
expected.
L-Cys substrate as
Now that all assay procedures and analytical methods have
been validated for the quantitation of native substrate and product
within enzyme catalyzed reactions, it is now possible to proceed to
the evaluation of CDO specificity using non-native substrates.
Determination of steady-state CDO kinetics and coupling utilizing
non-native substrates
As presented above, NMR can be used to monitor the steady-
state kinetics of CDO catalyzed CSA formation with the caveat that
the observed (CSA/O2)-coupling efficiency is decreased in D2O
Fig. 3. LC–MS study on CDO’s natural substrate L-cysteine using MRM (Multiple reaction monitoring) and SIM (select ion mode). (A) Blue lines represent products detected
with CSA standard. (B) Red lines represent products detected with reaction sample catalyzed by CDO. (C) SIM MS illustrating the +4 m/z shift in the CSA product ion peak
(154–158 m/z) upon substitution of 18O2 for 16O2. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

54
W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
Given the slow rate of catalysis and high level of oxidative
spectrometry by MRM could not be utilized for verification of
product. Instead, the LC–MS (selected ion mode) was used to
confirm appropriate parent ion mass and shift in parent ion mass
(+4 m/z) in reactions where 18O2 was substituted for 16O2. Remark-
ably, despite the broad range of substrates employed, all of the
oxidized products generated by CDO catalyzed reactions all exhib-
ited the appropriate +4 m/z shift in 18O2 reactions indicating a 4
electron oxidation of the sulfur atom to produce the appropriate
sulfinic acid. Other than formation of a disulfide-bond, no evidence
for partial thiol-oxidation or formation of multiple products was
observed.
uncoupling, it is not surprising that features attributed to disulfide
cross-link formation can be observed in reactions with cysteamine
at prolonged time-points. The chemical shifts associated with
disulfide-linked cysteamine are indicated by (*) in Fig. S4. These
features are completely removed upon addition of a suitable reduc-
tant such as tris(2-carboxyethyl)phosphine hydrochloride (TCEP).
Therefore, only the rate of hypotaurine formation should be con-
sidered for kinetic measurements as opposed to the decay of the
cysteamine substrate.
Verification of the hypotaurine product was performed by tan-
dem mass spectrometry (MRM) as illustrated in Fig. 4. In these
experiments, the [M+H]+ molecular ion for hypotaurine (110 m/z)
was selected for further fragmentation. MRM optimization was
performed on a standard solution of hypotaurine and then com-
pared to direct injections of the enzymatic reaction. In addition
to the 110 m/z parent [M+H]+ shown Fig. 4(panel A), the MRM
spectra for a standard solution of hypotaurine exhibits three char-
acteristic ions at 30, 45 and 65 m/z. By comparison, direct injection
Discussion
While
a variety of highly sensitive methods have been
described in the literature for the detection of cysteine sulfinic acid
[14,36–39], NMR spectroscopy offers the most versatile means to
study a variety of potential enzyme substrates under identical con-
ditions. The high KM value observed for CDO catalyzed reactions in
D2O (7 mM) allows for collection of data at substrate concentra-
tions appropriate for steady-state kinetic studies. One caveat of
using NMR spectroscopy is the significant decrease in the (CSA/
O2)-coupling in D2O relative to H2O. While beyond the scope of this
manuscript, a detailed accounting of the solvent isotope effects on
CDO catalysis is described elsewhere [35].
As with most oxidase and oxygenase enzymes, O2-activation in
TDO enzymes is gated by substrate-binding. Currently two theories
have been proposed to explain the substrate-gated O2 regulation
exhibited by non-heme mononuclear iron enzymes; (1) thermody-
namic gating of the FeII/FeIII redox couple and (2) Fe-site conforma-
tional changes which facilitate direct O2-coordination [40–42].
Taking the structure of the substrate-bound active site into consid-
eration (Fig. 1), the data provided in Table 1 suggest that simulta-
neous coordination of both the substrate-thiol and amine groups
are necessary for gating O2-binding and subsequent substrate
oxidation. For example, 1-mercpatopropane (1MP) and 3-mercap-
topropionic acid (3MP) lack an amino group and thus neither
O2-consumption or sulfinic acid product formation is observed
upon addition to aerobic solutions of CDO. The fact that CDO is
unable to oxidize 3MP, suggests that CDO may also have a means
by which to discriminate between O- and N-atom Fe-coordination.
While 2-(dimethylamino)ethanethiol (DME) has an available
amino functional group, the increased steric bulk and altered pKa
of the alkylated amine relative to the primary Cys-amine clearly
inhibits direct Fe-coordination. While the activity was too low for
accurate kinetic measurements, LC–MS clearly identifies a sulfinic
(2 lL) of the CDO catalyzed reaction with cysteamine yields an
identical fragmentation pattern with matching intensities, thus
confirming formation of the hypotaurine product within CDO reac-
tions. Selected ion mode (SIM) was used to observe an appropriate
shift in the observed product [M+H]+ ion upon substitution of 18O2
for the naturally abundant 16O2 to confirm the dioxygenase activ-
ity. As illustrated in Fig. 4B, CDO reactions performed in the pres-
ence of 18O2 results in a +4 m/z shift (114 m/z versus 110 m/z) in
the observed [M+H]+ parent ion, thus verifying the inclusion of
both oxygen atoms into the substrate.
CDO catalyzed thiol-oxidations
NMR kinetic studies with the natural L-Cys substrate and the
substrate analogue cysteamine effectively illustrate the dynamic
range of this analysis. While not possible in many enzymatic sys-
tems, the elevated KM observed for CDO (6.9 mM) within D2O
allows for collection of steady-state kinetic results at substrate
concentrations ranging from the NMR detection limit (0.3 mM)
up to, and beyond the apparent KM of CDO. As illustrated in Table 1,
several other commercially available thiol-substrates were ana-
lyzed using the same methodology as described for L-Cys and cys-
teamine. In all instances, the steady-state rate of O2-consumption
and product formation was determined to determine the coupling
efficiency for each substrate. When product sulfinic acid standards
were not commercially available, the decrease in the substrate
peak was monitored by NMR as described for Cys and cysteamine
reactions. In the absence of a product standard, tandem mass
Fig. 4. LC–MS study on CDO’s non-natural substrate cysteamine using MRM (Multiple reaction monitoring) and SIM (select ion mode). (A) Blue lines represent products
detected with hypotaurine standard. (B) Red lines represent products detected with reaction sample catalyzed by CDO. (C) SIM MS illustrating the +4 m/z shift in the
hypotaurine product ion peak (110–114 m/z) upon substitution of 18O2 for 16O2. (For interpretation of the references to colour in this figure legend, the reader is referred to
the web version of this article.)

W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
55
Table 1
Steady-state kinetic parameters determined by NMR and O2-electrode for selected CDO substrates.
Substrate
Oxygen consumption
Product formation
kcat (sÀ1
kcat (sÀ1
)
KM (mM)
V/K (MÀ1 sÀ1
)
)
KM (mM)
V/K (MÀ1 sÀ1
)
Coupling (%)
1.8 0.02
2.4 0.04
1.0 0.27
0.2 0.01
0.70 0.2
3.1 0.2
2570
770
30
0.74 0.06
0.08 0.02
0.07 0.03
N/A
7.0 1.3
6.8 3.2
6.2 2.1
N/A
106 21
41%
L
-Cys
-Cys
9
5
3.3%
4.8%
5.6%
D
34.6 8.0
13.4 1.8
10
5
L
L
-PA
119
0.3 0.1
-Hyc
CA
MS
N/A
À
N/A
N/A
N/A
N/A
N/A
N/A
N/A
0.63
N/A
N/A
N/A
N/A
N/A
N/A
N/A
À
N/A
N/A
N/A
N/A
N/A
N/A
N/A
<0.01
N/A
N/A
N/A
N/A
N/A
N/A
0.9%
N/A
N/A
N/A
N/A
N/A
N/A
CME
SC
DME
3MP
1MP
+
+
À
À
À
À
À
À
À
À
In these experiments, kcat is defined as v0/[E]. Substrate abbreviations: Cys, Cysteine; Hcy, homocysteine; CA, 2-aminoethanethiol (cysteamine); MS, mercaptosuccinate; PA,
-penicillamine, CME, -cysteine methyl ester; DME, 2-(dimethylamino)ethanethiol. SC, S-methyl- -cysteine; 3MP, 3-mecaptopropionate, 1MP; 1-mecaptopropane.
L
L
L
acid product in reactions with
contrast, the methylated thiol group of S-methyl-
prevents S-atom coordination to the Fe-site; thus catalysis is not
possible. As most of these substrates are isosteric with -Cys, it is
L
-cysteine methyl ester (CME). By
oxygen species. This hypothesis is supported by previous EPR stud-
ies using cyanide as a spectroscopic probe for substrate-interactions
within the catalytically inert FeIII-CDO. In these experiments, both
L-cysteine (SC)
L
D- and
L-Cys were capable of binding the active site to facilitate cya-
expected that they would induce a similar conformational changes
upon docking within the active site. Therefore, at least qualita-
tively, the results presented here support the hypothesis that
simultaneous (S/N)-bidentate coordination of the CDO substrates
thermodynamically regulate O2-binding by altering the FeII/III
redox couple.
nide binding, however samples prepared form
D
-Cys exhibited sig-
nificantly greater g-strain suggesting greater conformational
heterogeneity [13]. Reactions with cysteamine further support this
model in that both O2-consumption and hypotaurine formation are
significantly decreased. Moreover, as illustrated in Fig. 5B, under
identical conditions, the specificity (V/K) of CDO for cysteamine is
One feature that is becoming increasing apparent is the role of
the C93-Y157 pair in providing appropriate substrate orientation
and stereoselectivity. The (S/N)-bidentate substrate coordination
provides two points of simultaneous interaction. A third point of
interaction is produced by hydrogen bonding between the Cys-
carboxylate group and the C93-Y157 pair (2.94 Å). This third inter-
action is also stabilized electrostatically by interaction with R60
(separated by 2.91–3.16 Å). Collective, these interactions satisfy
the ‘‘three point interaction rule’’ for chiral selection [43]. Assuming
bidentate coordination of the substrate is the only requirement
four orders of magnitude less than that observed for L-Cys
(<0.01 MÀ1 sÀ1). Thus in the absence of a third point of interaction
within the CDO active site pocket, substrate binding is appears to
be weak resulting in attenuated O2-consumption rates, enzymatic
specificities, and coupling efficiencies.
CDO reactions with homocysteine and L-penicillamine (PA)
appear to also support this model in that both substrates have
the capacity to coordinate via bidentate (S/N)-ligation as well as
satisfy the ‘three point interaction’ with CDO. Therefore it is not sur-
prising that the rate of O2-consumption observed for PA (1.0 sÀ1) is
for gating reactivity with O2, then addition of either
D- or
L-isomers
quite similar to both L/D-Cys. Moreover, the specificity of CDO for
should trigger consumption of O2, which is indeed what is observed
experimentally. Despite this, as shown in Fig. 5A, the ratio of
O2-consumed per CSA generated is vastly attenuated (< 3%) in
these substrates is an order of magnitude greater than cysteamine.
Unlike PA, homocysteine exhibits low O2-consumption rate
(0.2 sÀ1). Presumably, the increased steric bulk of these substrates
distort the geometry of the ternary complex resulting in decreased
reactions involving
vant
been reported in the absence of D2O (81% and 4% respectively).
D-Cys as compared to the physiologically rele-
-Cys substrate (ꢀ40%). Similar uncoupling effects have also
L
productive catalysis (8% and 15% relative to
mental error, CDO specificity for -Cys and PA is essentially equiv-
alent (9 < V/K < 10 MÀ1 sÀ1). This value is an order of magnitude
lower than observed for the native -Cys substrate. By contrast,
CDO specificity for homocysteine (0.3 MÀ1 sÀ1) is vastly lower than
observed for -Cys. In principle, the additional (CH2)-group within
L-Cys). Within experi-
D
Indeed, nearly a 10-fold increase is observed in specificity (V/K) of
CDO for
L
-Cys as compared to
D
-Cys [13]. This suggests that, in addi-
L
tion to the chiral selection provided by three simultaneous points of
interaction, outer-sphere interactions with C93-Y157/R60 residues
may influence the geometry of the (O2/Substrate)-bound ternary
enzyme complex to minimize escape of partially reduced reactive
L
the homocysteine side chain significantly increased the steric bulk
within the active site pocket thus distorting substrate–enzyme
Fig. 5. Values determined for CDO kcat (v0/[E]) and specificity (V/K) for selected substrates. (A) The maximum rate of O2-consumption (black bars) and product formation (gray
bars) reflect the impact of substrate on enzymatic coupling. Enzyme specificity (V/K) for each substrate is indicated by the gray bars in panel B.

56
W. Li, B.S. Pierce / Archives of Biochemistry and Biophysics 565 (2015) 49–56
interactions. Interestingly, the oxidative coupling observed for
homocysteine is quite similar to that observed for -Cys and PA.
References
D
[1] K.G. Reddie, K.S. Carroll, Curr. Opin. Chem. Biol. 12 (2008) 746–754.
[2] P.G. Winyard, C.J. Moody, C. Jacob, Trends Biochem. Sci. 30 (2005) 453–461.
[3] D. Trachootham, J. Alexandre, P. Huang, Nat. Rev. Drug Discov. 8 (2009) 579–
591.
[4] D.P. Behave, W.B. Muse, K.S. Carroll, Infect. Disord. Drug Targets 7 (2007) 140–
158.
[5] C. Gordon, H. Bradley, R.H. Waring, P. Emery, Lancet 339 (1992) 25–26.
[6] C.R. Simmons, L.L. Hirschberger, M.S. Machi, M.H. Stipanuk, Protein Expr. Purif.
47 (2006) 74–81.
[7] M.T. Heafield, S. Fearn, G.B. Steventon, R.H. Waring, A.C. Williams, S.G.
Sturman, Neurosci. Lett. 110 (1990) 216–220.
[8] S.C. Chai, A.A. Jerkins, J.J. Banik, I. Shalev, J.L. Pinkham, P.C. Uden, M.J. Maroney,
J. Biol. Chem. 280 (2005) 9865–9869.
[9] P.J. Bagley, L.L. Hirschberger, M.H. Stipanuk, Anal. Biochem. 227 (1995) 40–48.
[10] C.M. Driggers, R.B. Cooley, B. Sankaran, L.L. Hirschberger, M.H. Stipanuk, P.A.
Karplus, J. Mol. Biol. 425 (2013) 3121–3136.
Given the absence of a carboxylate group on cysteamine, it
seems unlikely that ADO would require a Cys-Tyr cross-link to
orientate substrate binding. Indeed, sequence comparison of mam-
malian ADO and CDO enzymes suggest the absence of a Cys-Tyr
pair at an equivalent position in the CDO sequence. Nevertheless,
it has been reported that SDS PAGE of ADO isolated from specific
mouse tissue extracts exhibited a doublet (or triplet) protein band
similar to what is observed for mammalian CDO [44]. This feature
is absent in homogeneously purified recombinant SUMO- or
Flag-tagged ADO and thus this may simply be the result of post-
translational ubiquitination as suggested by the authors.
In the absence of any structural information for thiol dioxygen-
ase enzymes other than CDO, it is difficult to fully consider the rel-
evance of outer-sphere interactions relevant to thiol dioxygenase
substrate specificity. As a first approximation, the structural
threading model for the M. musculus ADO shown in Supplemental
Information (Fig. S5) was generated using the substrate-bound Rat-
tus norvegicus CDO crystal structure (pdb code 3ELN) as a template
[45,46]. While homology models should be considered with appro-
priate skepticism, they do provide a reasonable framework for the
discussion of conserved residues within the enzyme active site and
potential substrate interactions. Within the ADO homology model,
H155 and Y157 are replaced by D192 and L194 as indicated by
sequence alignment. The increased hydrophobicity of the ADO
active site pocket likely facilitates cysteamine binding to the Fe-
site as no other competing hydrogen-bonding interactions are
present. The only Cys and Tyr residues in close enough proximity
to produce a covalent cross-link are Y162 and C169 but these do
not overlap with the C93-Y157 pair of CDO (3ELN). Furthermore,
these residues appear quite distant from the predicted 3-His active
site (H100, H102, and H179) (ꢀ10 Å). Thus, if C162 and Y169 do
indeed form a covalent cross-link analogous to CDO, it is difficult
to see how they would be catalytically relevant on the basis of this
model.
[11] E. Siakkou, M.T. Rutledge, S.M. Wilbanks, G.N.L. Jameson, Biochim. Biophys.
Acta 1814 (2011) 2003–2009.
[12] J.E. Dominy Jr., J. Hwang, S. Guo, L.L. Hirschberger, S. Zhang, M.H. Stipanuk, J.
Biol. Chem. 283 (2008) 12188–12201.
[13] W. Li, E.J. Blaesi, M.D. Pecore, J.K. Crowell, B.S. Pierce, Biochemistry 52 (2013)
9104–9119.
[14] J.A. Crawford, W. Li, B.S. Pierce, Biochemistry 50 (2011) 10241–10253.
[15] B.S. Pierce, J.D. Gardner, L.J. Bailey, T.C. Brunold, B.G. Fox, Biochemistry 46
(2007) 8569–8578.
[16] J.E. Dominy Jr., C.R. Simmons, P.A. Karplus, A.M. Gehring, M.H. Stipanuk, J.
Bacteriol. 188 (2006) 5561–5569.
[17] M. Stipanuk, C. Simmons, P. Andrew Karplus, J. Dominy, Amino Acids (2010) 1–
12.
[18] N. Bruland, J.H. Wubbeler, A. Steinbuchel, J. Biol. Chem. 284 (2009) 660–672.
[19] U. Brandt, C. Waletzko, B. Voigt, M. Hecker, A. Steinbuchel, Appl. Microbiol.
Biotechnol. 98 (2014) 6039–6050.
[20] S. Ye, X. Wu, L. Wei, D. Tang, P. Sun, M. Bartlam, Z. Rao, J. Biol. Chem. 282
(2007) 3391–3402.
[21] C.W. Njeri, H.R. Ellis, Arch. Biochem. Biophys. 558 (2014) 61–69.
[22] C.R. Simmons, Q. Liu, Q. Huang, Q. Hao, T.P. Begley, P.A. Karplus, M.H. Stipanuk,
J. Biol. Chem. 281 (2006) 18723–18733.
[23] M.H. Stipanuk, Annu. Rev. Nutr. 24 (2004) 539–577.
[24] L. Ewetz, B. Sorbo, Biochim. Biophys. Acta 128 (1966) 296–305.
[25] B. Soerbo, L. Ewetz, Biochem. Biophys. Res. Commun. 18 (1965) 359–363.
[26] J.B. Lombardini, T.P. Singer, P.D. Boyer, J. Biol. Chem. 244 (1969) 1172–1175.
[27] C.H. Misra, Neurochem. Res. 8 (1983) 1497–1508.
[28] J.D. Gardner, B.S. Pierce, B.G. Fox, T.C. Brunold, Biochemistry 49 (2010) 6033–
6041.
[29] P.K. Glasoe, F.A. Long, J. Phys. Chem. 64 (1960) 188–190.
[30] N.J. Greenfield, Nat. Protoc. 1 (2007) 2876–2890.
[31] J.A. Dean (Ed.), Lange’s Handbook of Chemistry, 14 ed., McGraw-Hill Inc, New
York, 1992.
[32] R.A. Yost, D.D. Fetterolf, Mass Spectrom. Rev. 2 (1983) 1–45.
[33] A.L. Corder, B.P. Subedi, S. Zhang, A.M. Dark, F.W. Foss, B.S. Pierce, Biochemistry
52 (2013) 6182–6196.
[34] A. Kasperova, J. Kunert, M. Horynova, E. Weigl, M. Sebela, R. Lenobel, M. Raska,
Mycoses 54 (2011) e456–462.
The work presented here provides a much needed evaluation of
the minimum substrate requirements necessary for CDO catalyzed
O2-activation, sulfinic acid formation, and coupling efficiency. Sub-
strates were selected to evaluate cross-reactivity with other thiol-
dioxygenase enzyme substrates as well as provide insight into
potential substrate-active site interactions for CDO.
[35] J.K. Crowell, W. Li, B.S. Pierce, Biochemistry (2014), http://dx.doi.org/10.1021/
bi501241d (in press).
[36] E. Siakkou, S.M. Wilbanks, G.N. Jameson, Anal. Biochem. 405 (2010) 127–131.
[37] S. Heron, M.G. Maloumbi, M. Dreux, E. Verette, A. Tchapla, J. Chromatogr. A
1161 (2007) 152–156.
[38] T. Bantan-Polak, M. Kassai, K.B. Grant, Anal. Biochem. 297 (2001) 128–136.
[39] P.J. Hillas, P.F. Fitzpatrick, Biochemistry 35 (1996) 6969–6975.
[40] E.I. Solomon, T.C. Brunold, M.I. Davis, J.N. Kemsley, S.-K. Lee, N. Lehnert, F.
Neese, A.J. Skulan, Y.-S. Yang, J. Zhou, Chem. Rev. 100 (2000) 235–349.
[41] E.I. Solomon, A. Decker, N. Lehnert, Proc. Natl. Acad. Sci. U.S.A. 100 (2003)
3589–3594.
[42] M. Costas, M.P. Mehn, M.P. Jensen, L.J. Que, Chem. Rev. 104 (2004) 939–986.
[43] L.H. Easson, E. Stedman, Biochem. J. 27 (1933) 1257–1266.
[44] J.E. Dominy Jr., C.R. Simmons, L.L. Hirschberger, J. Hwang, R.M. Coloso, M.H.
Stipanuk, J. Biol. Chem. 282 (2007) 25189–25198.
Acknowledgments
This work was supported by NSF (CHE) 1213655 to B.S.P. We
would also like to acknowledge NSF financial support (CRIF:MU
CHE-0840509) for the purchase of the JEOL ECA500 500 MHz
FT-NMR spectrometers and the UTA Shimadzu Center for
Advanced Analytical Chemistry for the use of HPLC and LC–MS/
MS instrumentation.
Appendix A. Supplementary data
[45] L.A. Kelley, M.J. Sternberg, Nat. Protoc. 4 (2009) 363–371.
[46] C.R. Simmons, K. Krishnamoorthy, S.L. Granett, D.J. Schuller, J.E. Dominy, T.P.
Begley, M.H. Stipanuk, P.A. Karplus, Biochemistry 47 (2008) 11390–11392.
Supplementary data associated with this article can be found, in
the online version, at http://dx.doi.org/10.1016/j.abb.2014.11.004.