Characterization of unstable products of flavin- and pterin-dependent enzymes by continuous-flow mass spectrometry

Article abstract of DOI:10.1021/bi500267c

Continuous-flow mass spectrometry (CFMS) was used to monitor the products formed during the initial 0.25-20 s of the reactions catalyzed by the flavoprotein N-acetylpolyamine oxidase (PAO) and the pterin-dependent enzymes phenylalanine hydroxylase (PheH) and tyrosine hydroxylase (TyrH). N,N′-Dibenzyl-1,4-diaminobutane (DBDB) is a substrate for PAO for which amine oxidation is rate-limiting. CFMS of the reaction showed formation of an initial imine due to oxidation of an exo-carbon-nitrogen bond. Nonenzymatic hydrolysis of the imine formed benzaldehyde and N-benzyl-1,4-diaminobutane; the subsequent oxidation by PAO of the latter to an additional imine could also be followed. Measurement of the deuterium kinetic isotope effect on DBDB oxidation by CFMS yielded a value of 7.6 ± 0.3, in good agreement with a value of 6.7 ± 0.6 from steady-state kinetic analyses. In the PheH reaction, the transient formation of the 4a-hydroxypterin product was readily detected; tandem mass spectrometry confirmed attachment of the oxygen to C(4a). With wild-type TyrH, the 4a-hydroxypterin was also the product. In contrast, no product other than a dihydropterin could be detected in the reaction of the mutant protein E332A TyrH.

Full text of DOI:10.1021/bi500267c

Article
pubs.acs.org/biochemistry
Open Access on 04/08/2015
Characterization of Unstable Products of Flavin- and Pterin-
Dependent Enzymes by Continuous-Flow Mass Spectrometry
Kenneth M. Roberts,† Jose R. Tormos,‡ and Paul F. Fitzpatrick*,†
́
†Department of Biochemistry, University of Texas Health Science Center, San Antonio, Texas 78229, United States
‡Department of Chemistry, St. Mary’s University, San Antonio, Texas 78228, United States
ABSTRACT: Continuous-flow mass spectrometry (CFMS)
was used to monitor the products formed during the initial
0.25−20 s of the reactions catalyzed by the flavoprotein N-
acetylpolyamine oxidase (PAO) and the pterin-dependent
enzymes phenylalanine hydroxylase (PheH) and tyrosine
hydroxylase (TyrH). N,N′-Dibenzyl-1,4-diaminobutane
(DBDB) is a substrate for PAO for which amine oxidation is
rate-limiting. CFMS of the reaction showed formation of an
initial imine due to oxidation of an exo-carbon−nitrogen bond.
Nonenzymatic hydrolysis of the imine formed benzaldehyde and N-benzyl-1,4-diaminobutane; the subsequent oxidation by PAO
of the latter to an additional imine could also be followed. Measurement of the deuterium kinetic isotope effect on DBDB
oxidation by CFMS yielded a value of 7.6 0.3, in good agreement with a value of 6.7 0.6 from steady-state kinetic analyses. In
the PheH reaction, the transient formation of the 4a-hydroxypterin product was readily detected; tandem mass spectrometry
confirmed attachment of the oxygen to C(4a). With wild-type TyrH, the 4a-hydroxypterin was also the product. In contrast, no
product other than a dihydropterin could be detected in the reaction of the mutant protein E332A TyrH.
dentification of the product(s) of an enzyme-catalyzed
reaction is essential to understanding both the catalytic
atom into the pterin substrate to form a 4a-hydroxypterin (4a-
HO-pterin) (Scheme 1).12 The latter readily dehydrates in
I
mechanism of the enzyme and its role in metabolism. In many
cases, the initial product is unstable, hindering structural
characterization.1−4 Product identification in such cases
requires a combination of chemical logic, characterization of
stable compounds produced upon further reaction, and/or
synthesis of the authentic product or an analogue. Two specific
examples occur in the reactions of flavin amine oxidases and
pterin-dependent hydroxylases. The former is a large group of
enzymes that catalyze the oxidation of a C−N bond in the
substrate to an imine;5 the imine typically hydrolyzes to form
an aldehyde or ketone and a different amine as the stable
products, the products that are routinely detected. The
difficulty in detecting the initial imine product because of its
instability raised the question of whether the hydrolysis is
enzyme-catalyzed. The formation of an L-amino acid during
turnover of D-amino acid oxidase in the presence of sodium
borohydride provided the initial evidence of an imine as the
product released by a flavin amine oxidase;6 this was
subsequently confirmed by the use of a cyclic amine so that
the imine product was stable.7 Direct detection of an unstable
imine product required the use of an organic solvent for the
reaction, thereby preventing hydrolysis of the imine.8 However,
in the absence of evidence that an imine is always the product
of flavoprotein-catalyzed amine oxidation, enzyme-catalyzed
hydrolysis of the imine product is still being proposed for flavin
amine oxidases.9−11
Scheme 1
solution to form a quinonoid dihydropterin. The initial
demonstration that the pterin product of these enzymes is a
hydroxypterin was based on the similarity of the near-UV
absorbance spectrum of the pterin product of the phenylalanine
hydroxylase (PheH) reaction to that of a synthetic and stable
4a-hydroxy-5-deazapterin;13 further support for the structure of
the product came from the 13C nuclear magnetic resonance
(NMR) spectrum of the product formed from a specifically
labeled tetrahydropterin at cryogenic temperatures.14 Species
with similar absorbance spectra were subsequently identified as
products in the reactions catalyzed by tyrosine hydroxylase
The pterin-dependent aromatic amino acid hydroxylases
catalyze the incorporation of one atom of molecular oxygen
into the side chain of the amino acid substrate and the other
Received: March 3, 2014
Revised: April 8, 2014
Published: April 8, 2014
© 2014 American Chemical Society
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(TyrH) and tryptophan hydroxylase.15,16 While effective with
the wild-type enzymes, the use of absorbance spectra to identify
the pterin products of these enzymes is less straightforward
when it is applied to mutant enzymes in which multiple pterin
products are formed. Although deconvolution of product
spectra to quantitate the partitioning among different products
can be successful,16,17 such an approach is problematic when
the partitioning becomes complex. It is also not applicable in
addressing the possibility that novel pterin products are
formed.18
To synthesize ethylenediammonium diacetate (EDDA),
acetic acid (71 mL, 2.4 equiv) was added dropwise to
ethylenediamine (30.5 g) in 500 mL of dichloromethane at 4
°C. After the addition of acetic acid, solvent and excess acetic
acid were removed from the precipitate by rotary evaporation at
30 °C. The off-white solid was washed with dichloromethane
and dried under vacuum at room temperature to yield EDDA
1
(97.6 g) as a fluffy, white solid: H NMR (600 MHz, D2O) δ
1.87 (6H, s), 3.30 (4H, s); 13C NMR (175 MHz, D2O) δ 22.76,
36.36, 180.78.
We describe here the use of continuous-flow mass
spectrometry (CFMS) to characterize the initial products in
the reactions catalyzed by the flavin-dependent amino oxidase
N-acetylpolyamine oxidase (PAO) and by the pterin-dependent
hydroxylases PheH and TyrH. Structurally, PAO is a member
of the monoamine oxidase (MAO) family, a large family of
flavoproteins that includes MAO A, MAO B, LSD1, and the L-
amino acid oxidases.19 PAO catalyzes the oxidative cleavage of
N1-acetylspermine and N1-acetylspermidine, yielding 3-acet-
amidopropanal and spermidine or putrescine.20 In the case of
maize PAO, the reaction has been proposed to involve initial
oxidation to an imine followed by its hydrolysis within the
enzyme active site to form the final products.9 N,N′-Dibenzyl-
1,4-diaminobutane (DBDB) (Scheme 2) was selected as a
Expression and purification of His6-tagged mouse PAO were
performed as described by Royo and Fitzpatrick,23 omitting the
final chromatography on Sephacryl S-200. Rat phenylalanine
hydroxylase lacking the N-terminal regulatory domain (PheH
Δ117) was expressed and purified as described by Roberts et
al.24 Expression and purification of wild-type and E332A rat
TyrH were performed using the procedure of Daubner et al.25
with modifications. Bacterial cells were grown at 37 °C to an
OD600 of 0.3; the temperature was then decreased to 20 °C. At
an OD600 of 0.7−0.8, isopropyl β-D-1-thiogalactopyranoside
was added to induce protein expression. After 21 h at 20 °C,
the cells were harvested by centrifugation at 5000g for 30 min
or 7500g for 15 min. The cell pellets were stored at −80 °C.
Cells were resuspended in buffer containing 50 mM 4-(2-
hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 75
μM diethylenetriaminepentaacetic acid (DTPA), 100 μg/mL
phenylmethanesulfonyl fluoride, 10% glycerol, 1 μM leupeptin,
1 μM pepstatin A, and 10% glycerol (pH 7.0) and sonicated.
Nucleic acids were precipitated with streptomycin at a final
concentration of 1%. The protein was precipitated by addition
of ammonium sulfate to 45% saturation and resuspended in the
same buffer. The purified protein was obtained by separation
on a heparin-Sepharose column using a 0 to 0.8 M sodium
chloride gradient. The purified protein was dialyzed against 50
mM HEPES, 100 mM sodium chloride, 20 mM nitrilotriacetic
acid, 20 mM DTPA, 1 μM leupeptin, and 1 μM pepstatin A
(pH 7.0) to remove bound iron. The chelators were removed
by further dialysis against 50 mM HEPES, 100 mM sodium
chloride, 1 μM leupeptin, and 1 μM pepstatin A (pH 7.0).
Assays. The oxidation of DBDB by PAO was followed
directly in chemical-quench assays by monitoring the
concentration of DBDB using a BioLogic (Claix, France)
QFM-400 quenched-flow apparatus. Assays were performed by
mixing 50 μL of 80 μM DBDB in 10% methanol and 50 mM
ammonium acetate (pH 8.5) with 50 μL of 40 μM PAO in 50
mM ammonium acetate (pH 8.5) at 30 °C. N-Methylphene-
thylamine (MPEA) was included with DBDB at a concen-
tration of 80 μM as an internal standard. Reactions were
quenched with 50 μL of 2 M hydrochloric acid. Samples
representing t0 were obtained by first mixing DBDB with 2 M
hydrochloric acid and then adding PAO to the mixture. The
quenched samples were centrifuged to remove protein; the
resulting supernatants were diluted 10-fold with 0.1% trifluoro-
acetic acid in water before being injected onto a Phenomenex
Gemini-NX C18 high-performance liquid chromatography
(HPLC) column (3.0 μm, 2.0 mm × 150 mm). DBDB was
eluted using an isocratic mobile phase of 0.1% trifluoroacetic
acid and 17.5% acetonitrile in water and detected by
fluorescence, with excitation at 255 nm and emission at 278
nm. The amount of DBDB was determined by comparison with
a standard curve.
Scheme 2
substrate for our studies because oxidation of the amine is the
rate-limiting step with it as the substrate for PAO,21 whereas
product release is rate-limiting with physiological substrates.22
PheH was selected for the characterization of the product of a
pterin-dependent enzyme because it was used in the initial
demonstration of the 4a-HO-pterin; in addition, analysis of the
TyrH reaction products was conducted with both the wild-type
enzyme and a mutant protein in which tetrahydropterin
oxidation and amino acid oxidation are uncoupled, so that
the 4a-HO-pterin is not the exclusive product.
EXPERIMENTAL PROCEDURES
■
Materials. 6-Methyl-5,6,7,8-tetrahydropterin (6MPH4) was
purchased from Schircks Laboratories (Jona, Switzerland).
Isopropyl β-D-1-thiogalactopyranoside was from Research
Products International (Mount Prospect, IL). Diethylenetria-
minepentaacetic acid (DTPA), pepstatin A, L-phenylalanine,
and L-tyrosine were purchased from Sigma-Aldrich (St. Louis,
MO). Leupeptin was from Peptide Institute (Osaka, Japan).
N,N′-Dibenzyl-1,4-diaminobutane (DBDB) was purchased
from Prime Organics, Inc. (Woburn, MA). N,N′-Bis-perdeuter-
obenzyl-1,4-diaminobutane (DBDB-d14) was synthesized by the
procedure of Henderson Pozzi et al.22
Steady-state deuterium isotope effects for the oxidation of
DBDB by PAO were measured by monitoring oxygen
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consumption over a range of substrate concentrations using a
Yellow Springs Inc. (Yellow Springs, OH) 5300A biological
oxygen monitor. Assays contained 2.5−250 μM DBDB or
DBDB-d14 and 1.6 μM PAO in 5% methanol and 50 mM
ammonium acetate (pH 8.5) at 30 °C; the total reaction
volume was 1 mL. Reactions were initiated by the addition of
enzyme.
Explorer was used to estimate the confidence in the best-fit
values by determining the sum square error for all pairs of
parameters. The reported confidence intervals are the ranges of
values for each parameter for which global fitting gives X2
values no more than 40% greater than the X2 value obtained
using the best-fit values; this is termed a X2 threshold of 1.4.
RESULTS AND DISCUSSION
Continuous-Flow Mass Spectrometry. DBDB oxidation
was monitored by high-resolution mass spectrometry using a
Thermo Scientific (Waltham, MA) LTQ OrbiTrap Discovery
mass spectrometer equipped with a custom nanoelectrospray
ionization source incorporating a temperature-controlled nano-
volume continuous-flow mixer (Eksigent, Dublin, CA). To
minimize delay volumes, New Objective (Woburn, MA)
SilicaTip emitters with 20 μm inside diameters and 10 μm
tips were mounted directly onto the mixing chip. Reactions
were performed by mixing the substrate and enzyme solutions
at equal flow rates using the same conditions that were used for
the chemical-quench assays. MPEA was included with the
substrate as an internal standard at a final concentration after
mixing of 40 μM. Flow rates ranging from 0.6 to 10 μL/min
were used to obtain multiple reaction times. A sample
corresponding to a reaction time of 0 s was obtained by
mixing DBDB with buffer only. Product isotope measurements
were taken using a mixture of 6.7 μM DBDB and 33 μM
DBDB-d14. Reaction mixtures were monitored in spectral mode
over the range of m/z 85−400. Substrates and reaction
products were identified by their respective [M + H]+ ions with
a mass error of <2 ppm. Each data point represents an average
of at least 200 scans. Ion intensities were normalized against the
intensity of MPEA at each flow rate.
■
Kinetics of Oxidation of DBDB. To utilize CFMS to
analyze the kinetics of oxidation of DBDB by PAO, it was
necessary to first calibrate the system. For continuous-flow
methods, the reaction time at which the analysis occurs is
determined by both the flow rate of the reactants and the
volume between the mixer and the detector. Either can be
varied to analyze the time course of reactions.28,29 In the
continuous-flow approach used here, different reaction times
were obtained by varying the flow rate. Calculation of the
reaction times thus required knowledge of the volume (Vapp
)
from mixing until the reaction is quenched by desolvation upon
electrospray (ESI). The value of Vapp can be determined by
monitoring the progress of a reaction with established kinetics
at various flow rates (Q), an approach that is commonly used to
calibrate rapid-quench systems and determine dead times for
stopped-flow instruments. Consequently, the kinetics of
oxidation of DBDB by PAO were first analyzed by chemical-
quench methods to determine the rate constant for the
reaction. To do so, the enzyme was mixed with DBDB and the
reaction was quenched at 0.25−8 s. The amount of DBDB
remaining at each time was then determined by HPLC with
fluorescence detection. Because the concentration of DBDB
used was below its KM value under these conditions (50 μM),
the time-dependent loss of DBDB (Figure 1) could be fit
For CFMS of pterin oxidation by PheH Δ117, 3 volumes of
25 μM PheH Δ117 and 2.7 mM L-phenylalanine in 50 mM
ammonium acetate and 1% methanol (pH 7.0) was mixed with
1 volume of 4.0 mM 6MPH4 in 1 mM HCl and 1% methanol at
5 °C at a flow rate of 5.0 μL/min. Mass scanning of the reaction
mixtures was performed over the range of m/z 100−300.
Determination of MS/MS spectra of reaction species was
performed at a rate of 1.0 μL/min.
CFMS assays of pterin oxidation by wild-type and E332A
TyrH were performed by mixing an anaerobic solution of 20
μM enzyme and 500 μM tyrosine in 100 mM EDDA/
ethylenediamine (pH 8.0) with an equal volume of an aerobic
solution of 100 μM 6MPH4 in 25 mM ammonium chloride/
HCl and 10% methanol (pH 3.0) at 5 °C. The enzyme/
substrate mixture was prepared in several steps. The stock
apoenzyme was exchanged into 100 mM EDDA/ethylenedi-
amine (pH 8.0) using a Sephadex G-25 column (1.0 cm × 17.5
cm) and added to a tonometer containing tyrosine in the same
buffer. A side arm containing 1 equiv of ferrous ammonium
sulfate in 2 mM HCl was attached. The tonometer was made
anaerobic by alternating vacuum and argon for 15 cycles. The
contents were then mixed by gently inverting the contents into
the side arm several times. Five additional vacuum−argon
cycles were applied, and the tonometer was immediately
mounted onto the CFMS system. Mass scanning of reactions
was performed over the range of m/z 130−400.
●
Figure 1. Time course for the oxidation of DBDB by PAO: ( )
□
chemical-quench assays and ( ) continuous-flow mass spectrometry.
The reaction conditions were 40 μM DBDB, 20 μM polyamine
oxidase, 50 mM ammonium acetate, 5% methanol, pH 8.5, and 30 °C.
All concentrations are after mixing. The line indicates the best-fit curve
for the chemical-quench data to eq 1.
reasonably well as an exponential decrease (eq 1), yielding a
value for the observed rate constant (kobs) for DBDB oxidation
a
of 0.14 0.01 s−1.
obst
[DBDB]t = [DBDB]0e−k
(1)
Data Analysis. Kinetic data from individual experiments
were fit using KaleidaGraph (Synergy Software, Reading, PA).
KinTek Explorer26 (KinTek Corp., Austin, TX) was used to
perform global analyses of multiple time-dependent data sets
through singular-value decomposition to determine observed
rate values. The FitSpace Explorer27 package of KinTek
Next, DBDB and PAO were reacted in the CFMS system
under conditions identical to those used for the chemical-
quench assays, varying the flow rate to obtain multiple reaction
times. As with other mass spectrometry methods, the
quantification of ions of interest by CFMS requires normal-
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ization against an internal standard. However, quantification
using ESI is further complicated by the observation that the
intensity of an ion signal varies somewhat with the
concentration of the enzyme and the flow rate. In our hands,
high concentrations of enzyme decrease the ion intensity, likely
because of an increase in the viscosity and surface tension of the
reaction mixture, which disrupt the desolvation process of the
electrospray. The flow rate dependence of the ion intensity is
the result of variations in droplet size and the extent of
desolvation.30,31 We have found that this flow-dependent signal
variation is ion-specific, such that the signal ratios for two ions
may vary significantly at different flow rates. To correct for this,
several analogues of DBDB were tested as internal standards,
including N,N′-dibenzyl-1,2-diaminoethane, diphenylamine,
lysine, and N-methylphenethylamine. N-Methylphenethylamine
(MPEA) gave the behavior most consistent with that of DBDB
across a range of flow rates, and it is not a substrate for PAO.
Consequently, the concentration of DBDB remaining upon
detection at each flow rate was determined by normalizing the
ion signal against that for MPEA as an internal standard and the
known concentration of DBDB at t0.
Figure 3. Time dependence for products of the PAO-catalyzed
oxidation of DBDB. The data are reported as the ion intensities for 1
( ), BDB ( ), 2 ( ), and putrescine ( ) relative to that of MPEA.
Assay conditions are identical to those described in the legend of
Figure 1. The lines are from global fits of the model in Scheme 4 with
the values for the rate constants listed in Table 1.
■
□
●
○
hydrogen atoms less than the m/z value for the [M + H]+ ion
of DBDB (m/z 269.2012). The simplest possibility for such a
species is that it is an imine (m/z 267.1856) formed by
abstraction of two protons and two electrons from DBDB
(Scheme 3). The magnitude of the signal for 1 increased over
the first few seconds of the reaction and then slowly decayed
(Figure 3), indicating that 1 is not the final product in the
reaction. Consistent with such a conclusion, the magnitude of
the signal for an additional species with an m/z value of
179.1540 increased more gradually over the first 5 s of the
reaction (Figures 2 and 3). This mass is consistent with the [M
+ H]+ ion of N-benzyl-1,4-diaminobutane (BDB, m/z
179.1543). As BDB is the expected product in the hydrolysis
of an exo-imine formed from DBDB (Scheme 3), 1 can be
identified as N-benzyl-N′-benzylidene-1,4-diaminobutane. No
significant signal was observed for the ion of benzylamine or N-
benzyl-4-aminobutanal, ruling out the endo-imine of DBDB as
the major enzymatic product. The intensity of the signal for
BDB decayed at later times, and that of an additional ion (2) at
m/z 177.1384 increased (Figure 3). The m/z value of the latter
ion is exactly two hydrogen atoms less than the signal for BDB,
indicating that it represents the [M + H]+ ion (m/z 177.1386)
of a dehydrogenation product of BDB. The fact that 2 derives
from BDB and not DBDB is supported by the observation of a
lag phase in its formation. Similar to 1, 2 can be reasonably
assigned as N-benzylidene-1,4-diaminobutane, the exo-imine of
BDB (Scheme 3). This establishes that BDB is also a substrate
for PAO. As with 1, the magnitude of the signal for 2 decreased
with time; this decay was accompanied by the slow increase
after several seconds in the magnitude of a signal at m/z
89.1072 (Figure 3). This matches well the [M + H]+ ion of
putrescine (m/z 89.1073), consistent with hydrolysis of 2
(Scheme 3). Benzaldehyde must be an additional product in
the hydrolyses of 1 and 2 to BDB and putrescine, respectively.
However, benzaldehyde is not readily ionized under ESI
conditions, and no corresponding ion was observed. Overall,
the oxidation of DBDB by PAO can be described by the
pathway in Scheme 3. As ESI-MS is sufficiently gentle to keep
multimeric protein complexes intact,32−35 reaction products
that have not dissociated from the enzyme are not expected to
be detectable as individual ions by CFMS. The direct
observation of both 1 and 2 in these assays indicates that the
imine products are released from the enzyme with their
subsequent hydrolysis occurring in solution.
The reaction time for a continuous-flow analysis is
proportional to the inverse of flow rate Q (eq 2).
t = V /Q
app
(2)
The value of Vapp can be determined by combining eqs 1 and 2
and substituting in the value for kobs from the chemical-quench
experiment (eq 3).
−1)Vapp/Q
[DBDB]t = [DBDB]0e(−0.14 s
(3)
This approach yielded a value for Vapp of 100 20 nL for the
mixer used for the experiments described here. Figure 1 shows
an overlay of the data from the rapid-quench and MS analyses
using this Vapp value to determine the reaction times. A similar
analysis conducted with a second mixer yielded a value of 210
20 nL for Vapp, demonstrating the necessity of determining
Vapp upon any change in the experimental system.
Products of the Oxidation of DBDB by PAO. As noted
above, a major advantage of CFMS in following reactions is the
ability to gain structural insight into transient reactant products.
When the oxidation of DBDB was monitored by scanning the
range of m/z 85−400, several additional ions were seen during
the reaction (Figure 2). The intensities of these ions varied with
reaction time in a manner consistent with them being reaction
products (Figure 3). Ion 1 was seen at the earliest reaction time
(0.6 s) and had an m/z value of 267.1855. This is exactly two
Figure 2. Representative mass spectrum from the experiment in Figure
1 at a reaction time of 4.8 s. The spectrum represents the average of
250 scans at a flow rate of 1250 nL/min. Ion signals are normalized to
the most intense ion ([1 + H]+).
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Scheme 3
To obtain the values of the rate constants for the steps in
Scheme 3, the kinetics of formation and decay of the MS signals
for DBDB and the different reaction products were analyzed
globally using KinTek Explorer.26 A simple four-step model
(Scheme 4) was sufficient to fit the complete reaction (Figure
DBDB may result in better binding of the former in the active
site. As the rates of product release do not contribute to kcat/
KM, differences in the rate constants for product release are not
responsible for the differences.
Kinetic Isotope Effects for the Oxidation of DBDB. The
ability of the CFMS system to directly monitor substrate and
products during the oxidation of DBDB by PAO suggested that
it should be possible to directly measure the isotope effect on
DBDB oxidation. Deuterium isotope effects on steady-state
kinetic parameters were first determined by noncompetitive
initial rate assays. Rates of oxygen consumption were measured
with either DBDB or N,N′-bis-perdeuterobenzyl-1,4-diamino-
butane (DBDB-d14) as the substrate for PAO, using conditions
similar to those used for the MS-based assays. The data were
well fit using eq 4, which applies for an identical isotope effect
(Dk) on the kcat and kcat/KDBDB values, yielding an isotope effect
of 6.7 0.6.
Scheme 4
3). In the first step, DBDB is oxidized to 1 by PAO. The second
step is the nonenzymatic hydrolysis of the free imine 1 to BDB.
The two subsequent steps arise from turnover with BDB as the
substrate. The rate constants for the two hydrolysis steps (kH O
)
2
were assumed to be identical in the analysis, because allowing
their rate constants to vary independently did not significantly
improve the fit and gave similar values for all rate constants.
The values from the analysis are listed in Table 1.
v = kcat[DBDB]/{[KDBDB + [DBDB]][1 + (Dk − 1)F]}
(4)
Table 1. Kinetic Parameters for the Kinetic Mechanism in
Scheme 4
a
Alternative analyses assuming different isotope effects on the
kcat and kcat/KDBDB values did not yield improved fits. This
result is consistent with rate-limiting chemistry with this
substrate, as previously concluded.22
k1
k2
6.0 mM−1 s−1 (4.9−7.1)
33 mM−1 s−1 (26−62)
0.52 s−1 (0.43−0.62)
kH O
The isotope effect was then measured by CFMS. A mixture
of DBDB and DBDB-d14 was used as the substrate for PAO,
again following the products over time. The initial concen-
tration of DBDB-d14 was ∼5-fold greater than that for the
undeuterated substrate to compensate for the decreased rate of
formation of the deuterated product due to the large isotope
effect measured in the steady-state assays. Figure 4 shows the
time dependence for the observed isotope effect on the
formation of 1; the value at time zero yields an intrinsic isotope
2
a
Values for the rate constants were determined globally with KinTek
Explorer and the data from Figures 1 and 3. The values in parentheses
are the confidence intervals reported by FitSpace at a X2 threshold of
1.4, that is, the change in the value that results in an increase in X2 of
40% if all of the other parameters are allowed to vary to improve the
fit.
Because these analyses were conducted at a low concen-
tration of DBDB, rate constants k1 and k2 equal the kcat/KM
values for DBDB and BDB, respectively. The value for BDB is
∼5-fold greater than that for DBDB, consistent with the former
being the better substrate. These results do not allow us to
distinguish whether the differences are due to a difference in
binding or chemistry. For oxidation of a polyamine, PAO
requires that the reacting nitrogen be neutral and that a
different nitrogen be charged.22 The primary nitrogen in BDB
would have a pKa value higher than that of the benzylic
nitrogens in DBDB or BDB, so that more of the latter would be
correctly protonated at the pH of these analyses, 8.5. In
addition, because the active sites of polyamine oxidases are
appropriate for linear saturated substrates,36,37 the absence in
BDB of the bulky and rigid second aromatic ring present in
effect of 7.7
0.3. This value is similar to the value for Dk
determined in the steady-state assays and more precise. Because
of the large isotope effect on imine formation, ion intensities for
2 were too low to perform a similar analysis for the oxidation of
BDB in the second turnover.
Pterin Products of the Aromatic Amino Acid
Hydroxylases. The CFMS system was also used to character-
ize the pterin products produced by wild-type aromatic amino
acid hydroxylases and a mutant enzyme. Initial analyses used
PheH Δ117, a truncated enzyme that lacks the N-terminal
regulatory domain and thus no longer exhibits allostery,
simplifying the kinetics.38,39 In reactions with 6MPH4 and L-
phenylalanine as substrates, the ion for tyrosine (m/z
182.0811), the amino acid product of the reaction, could be
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at ambient temperatures. The MS/MS spectrum of the putative
4a-HO-pterin (Figure 5B) showed three major product ions at
m/z 170.1035, 154.1087, and 86.0348. The first two ions likely
result from a fragmentation pathway wherein the pyrimidine
ring contracts into an imidazole, resulting in the loss of the
carbonyl carbon as CO or CO2 (Scheme 5). The observation
Scheme 5
Figure 4. Measurement of the deuterium isotope effect for oxidation
of DBDB by PAO determined by continuous-flow mass spectrometry.
The relative intensities of the ions for deuterated (d13) and
undeuterated (h13) 1 were determined at different flow rates. Reaction
mixtures contained 6.7 μM DBDB, 33 μM DBDB-d14, and 40 μM
PAO in 5% methanol and 50 mM ammonium acetate (pH 8.5) at 30
°C. All concentrations are after mixing. The line indicates the best fit
to a single exponential.
resolved from that of 6MPH4 (m/z 182.1036), despite their
very similar masses (Figure 5A, inset). In addition, two ions
that this contraction occurs both with and without loss of the
second oxygen atom suggests that the additional oxygen atom is
in the proximity of the carbonyl. The third product ion, [M −
C5H8N2O]+, is generated by the loss of the methylpiperazine
ring and an oxygen atom, indicating that the oxygen atom
incorporated during the reaction is bound to the piperazine
ring. These observations confirm the incorporation of oxygen at
position 4a, as this position is a member of the piperazine ring
and adjacent to the carbonyl. Consistent with this conclusion,
the same product ion at m/z 86.0348 ([M − C5H8N2]+) was
also present in the MS/MS spectrum of 6MPH4 (not shown);
in this case, it would be formed by loss of the methylpiperazine
alone from the substrate lacking the second oxygen atom.
A similar analysis of the reaction of wild-type TyrH with
6MPH4 and L-tyrosine was conducted. Again, two pterin
products with m/z values consistent with the dihydropterin and
the 4a-HO-pterin were readily detectable (Figure 6A),
Figure 5. (A) Mass spectrum for the reaction of 20 μM PheH Δ117,
2.0 mM L-phenylalanine, and 1.0 mM 6MPH4 at a reaction time of 1.2
s. All concentrations are after mixing. The spectrum is the average of
20 scans. Ion signals are normalized to the most intense ion ([6MPH4
+ H]+). The inset is a close-up of the spectrum across the range of m/z
182.07−187.12 showing the resolved peaks for L-tyrosine (Tyr) and
6MPH4. (B) MS/MS spectrum of the 4a-HO-pterin product (m/z
198.0982) from the reaction with PheH Δ117. Reaction conditions
were identical to those used for panel A at a reaction time of 6 s. The
spectrum is the average of 10 scans. Ion signals are normalized to the
most intense ion ([M − CO2]+).
that could be identified as pterin oxidation products were
clearly seen (Figure 5A). One had an m/z value of 180.0877,
exactly two hydrogen atoms less than that for 6MPH4,
indicating that this species is the [M + H]+ ion of a
dihydropterin [6MPH2, m/z 180.0880 (Scheme 1)]. The
second ion had an m/z value of 198.0982, greater than that of
6MPH4 by the mass of a single oxygen atom. The likely
candidate for this product is the unstable 4a-hydroxy-6-
methyltrihydropterin (4a-HO-pterin, m/z 198.0986), formed
during the reaction by the incorporation of one atom of
molecular oxygen at position 4a of 6MPH4 (Scheme 1).
The site of attachment of the hydroxyl moiety in the 4a-
hydroxypterin product of the aromatic amino acid hydroxylases
was previously established by analysis of the NMR spectrum of
the product formed under cryogenic conditions from 6MPH4
that had been specifically labeled with 13C at position 4a.14 The
ability to directly detect the HO-pterin by mass spectrometry
allowed us to confirm this conclusion with unlabeled substrates
Figure 6. Mass spectrum for the reaction of 10 μM wild-type (A) or
E332A (B) TyrH, 250 μM L-tyrosine, and 50 μM 6MPH4 at a reaction
time of 1.5 s. All concentrations are after mixing. The spectra are
averages of 150 scans. Ion signals are normalized to the most intense
ion.
confirming directly the latter as a product of the reaction
catalyzed by this enzyme. The magnitude of the relative signal
for 6MPH2 increased at longer reaction times, quickly
becoming the dominant signal, while the magnitude of the
signal for 6MPH4 decayed as the reaction progressed. In
contrast, the magnitude of the signal for the 4a-HO-pterin
decreased with time, indicating that this product dehydrates to
give the dihydropterin.
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Scheme 6
The mutant enzyme E332A TyrH shows almost complete
uncoupling of tetrahydropterin oxidation from amino acid
hydroxylation.25 Spectroscopic studies of this protein suggested
that both the amino acid and the pterin substrate bind normally
and convert the iron to the same oxygen-reactive form that is
found in the wild-type enzyme18,40 but did not establish the
point in the subsequent reaction at which uncoupling occurred.
Stopped-flow absorbance spectroscopy was previously used to
obtain the absorbance spectrum of the pterin product(s) to
clarify the step at which the reaction became unproductive.
Comparison of the spectrum with those of known pterins
suggested that a pterin distinct from the 4a-HO-pterin was
formed in the reaction, possibly a hydroperoxypterin.18 CFMS
was used to more definitively identify the pterin product(s)
produced by E332A TyrH. The mass spectrum of the reaction
mixture did not show evidence of the formation of the 4a-HO-
pterin (Figure 6B). Instead, the dihydropterin (6MPH2) was
the only pterin product detected. It is not possible to determine
from the mass spectrum which tautomer of 6MPH2 forms, but
the near-UV absorbance spectrum of the reaction exhibits
absorbance between 300 and 400 nm consistent with a
significant amount of quinonoid 6MPH2.18 The lack of a signal
for a hydroperoxypterin intermediate suggests that it did not
form, and that the enzyme oxidizes 6MPH4 directly to 6MPH2.
However, we cannot rule out the possibility that the
hydroperoxypterin is too unstable to detect even at these
short times. The absence of a signal for the 4a-HO-pterin
establishes that the E332A TyrH reaction does not catalyze the
appropriate heterolytic cleavage of molecular oxygen to
generate the Fe(IV)O. Our present understanding of the
mechanism of the pterin-dependent hydroxylases is shown in
Scheme 6. The altered binding of the pterin due to mutagenesis
of Glu332 alters the initial reaction of the enzyme with oxygen,
resulting in direct oxidation of the tetrahydropterin, or the
mutation prevents the heterolytic cleavage of the oxygen−
oxygen bond in the initial intermediate. The previous proposal
of Chow et al.18 that Glu332 is responsible for protonation of
one of the oxygen atoms to facilitate oxygen bond cleavage
would be consistent with the results presented here.
CFMS definitively identified the unstable 4a-HO-pterin
proposed as a product for both PheH and TyrH and ruled
out the 4a-HO-pterin as a significant product in the reaction of
a mutant TyrH.
AUTHOR INFORMATION
■
Corresponding Author
*E-mail: fitzpatrickp@uthscsa.edu. Phone: (210) 567-8264.
Fax: (210) 567-8778.
Funding
This work was supported in part by grants from the National
Institutes of Health (R01 GM058698) and The Welch
Foundation (AQ1245).
Notes
The authors declare no competing financial interest.
ABBREVIATIONS
■
PheH, phenylalanine hydroxylase; PheH Δ117, phenylalanine
hydroxylase lacking the N-terminal regulatory domain; TyrH,
tyrosine hydroxylase; CFMS, continuous-flow mass spectrom-
etry; PAO, N-acetylpolyamine oxidase; MAO, monoamine
oxidase; DBDB, N,N′-dibenzyl-1,4-diaminobutane; DBDB-d14,
N,N′-bis-perdeuterobenzyl-1,4-diaminobutane; 6MPH4, 6-
methyl-5,6,7,8-tetrahydropterin; 6MPH2, 6-methyldihydropter-
in; 4-HO-pterin, 4a-hydroxy-6-methyltrihydropterin; DTPA,
diethylenetriaminepentaacetic acid; EDDA, ethylenediammo-
nium diacetate; HEPES, 4-(2-hydroxyethyl)-1-piperazineetha-
nesulfonic acid; ESI, electrospray ionization; MPEA, N-
methylphenethylamine; BDB, N-benzyl-1,4-diaminobutane.
ADDITIONAL NOTE
■
a
The single-exponential fit is not meant to imply a pseudo-first-
order condition of the assay but simply represents a descriptive
reflection of the data that can be used as a calibration for the
CFMS system.
REFERENCES
■
(1) Stumpe, M., and Feussner, I. (2006) Formation of oxylipins by
CYP74 enzymes. Phytochem. Rev. 5, 347−357.
(2) Fenoll, L. G., Rodríguez-Lop
CONCLUSION
■
The data presented here establish the utility of continuous-flow
mass spectrometry for detecting unstable products of enzyme-
catalyzed reactions. In contrast to previous methods for
isolating imine products of reactions by flavoproteins, which
relied on the formation of stable imine products or anhydrous
conditions, this approach established directly that an unstable
imine is the product of the reaction catalyzed by PAO and
allowed measurement of the deuterium kinetic isotope effect
for the formation of the imine. It also allowed analysis of the
kinetics of further oxidation of the enzymatic products in the
reaction. In the reactions of the pterin-dependent enzymes,
́
ez, J. N., García-Sevilla, F., García-
Ruiz, P. A., Varon, R., García-Canovas, F., and Tudela, J. (2001)
́
́
Analysis and interpretation of the action mechanism of mushroom
tyrosinase on monophenols and diphenols generating highly unstable
o-quinones. Biochim. Biophys. Acta 1548, 1−22.
(3) Deery, E., Schroeder, S., Lawrence, A. D., Taylor, S. L.,
Seyedarabi, A., Waterman, J., Wilson, K. S., Brown, D., Geeves, M. A.,
Howard, M. J., Pickersgill, R. W., and Warren, M. J. (2012) An
enzyme-trap approach allows isolation of intermediates in cobalamin
biosynthesis. Nat. Chem. Biol. 8, 933−940.
(4) Yang, W., Moore, I. F., Koteva, K. P., Bareich, D. C., Hughes, D.
W., and Wright, G. D. (2004) TetX is a flavin-dependent
2678
dx.doi.org/10.1021/bi500267c | Biochemistry 2014, 53, 2672−2679

Biochemistry
Article
monooxygenase conferring resistance to tetracycline antibiotics. J. Biol.
Chem. 279, 52346−52352.
and Rate Constants from Single-Turnover Experiments. Biochemistry
52, 1062−1073.
(5) Fitzpatrick, P. F. (2010) Oxidation of amines by flavoproteins.
Arch. Biochem. Biophys. 493, 13−25.
(25) Daubner, S. C., and Fitzpatrick, P. F. (1999) Site-directed
mutants of charged residues in the active site of tyrosine hydroxylase.
Biochemistry 38, 4448−4454.
(6) Hafner, D. W., and Wellner, D. (1971) Demonstration of imino
acids as products of the reactions catalyzed by D- and L-amino acid
oxidases. Proc. Natl. Acad. Sci. U.S.A. 5, 987−991.
(7) Fitzpatrick, P. F., and Massey, V. (1982) Thiazolidine-2-
carboxylic acid, an adduct of cysteamine and glyoxylate, as a substrate
for D-amino acid oxidase. J. Biol. Chem. 257, 1166−1171.
(8) Woo, J. C. G., and Silverman, R. B. (1995) Monoamine oxidase B
catalysis in low aqueous medium. Direct evidence for an imine
product. J. Am. Chem. Soc. 117, 1663−1664.
(9) Binda, C., Coda, A., Angelini, R., Federico, R., Ascenzi, P., and
Mattevi, A. (1999) A 30 Å long U-shaped catalytic tunnel in the crystal
structure of polyamine oxidase. Structure 7, 265−276.
(10) Pawelek, P. D., Cheah, J., Coulombe, R., Macheroux, P., Ghisla,
S., and Vrielink, A. (2000) The structure of L-amino acid oxidase
reveals the substrate trajectory into an enantiomerically conserved
active site. EMBO J. 19, 4204−4215.
(11) Kachalova, G., Decker, K., Holt, A., and Bartunik, H. D. (2011)
Crystallographic snapshots of the complete reaction cycle of nicotine
degradation by an amine oxidase of the monoamine oxidase (MAO)
family. Proc. Natl. Acad. Sci. U.S.A. 108, 4800−4805.
(12) Fitzpatrick, P. F. (2000) The aromatic amino acid hydroxylases.
In Advances in Enzymology and Related Areas of Molecular Biology
(Purich, D. L., Ed.) pp 235−294, John Wiley & Sons, Inc., New York.
(13) Lazarus, R. A., Dietrich, R. F., Wallick, D. E., and Benkovic, S. J.
(1981) On the mechanism of action of phenylalanine hydroxylase.
Biochemistry 20, 6834−6841.
(14) Lazarus, R. A., DeBrosse, C. W., and Benkovic, S. J. (1982)
Phenylalanine hydroxylase: Structural determination of the tetrahy-
dropterin intermediates by 13C NMR spectroscopy. J. Am. Chem. Soc.
104, 6869−6871.
(15) Dix, T. A., Kuhn, D. M., and Benkovic, S. J. (1987) Mechanism
of oxygen activation by tyrosine hydroxylase. Biochemistry 26, 3354−
3361.
(16) Moran, G. R., Derecskei-Kovacs, A., Hillas, P. J., and Fitzpatrick,
P. F. (2000) On the catalytic mechanism of tryptophan hydroxylase. J.
Am. Chem. Soc. 122, 4535−4541.
(17) Ellis, H. R., Daubner, S. C., and Fitzpatrick, P. F. (2000)
Mutation of serine 395 of tyrosine hydroxylase decouples oxygen-
oxygen bond cleavage and tyrosine hydroxylation. Biochemistry 39,
4174−4181.
(18) Chow, M. S., Eser, B. E., Wilson, S. A., Hodgson, K. O.,
Hedman, B., Fitzpatrick, P. F., and Solomon, E. I. (2009) Spectroscopy
and kinetics of wild-type and mutant tyrosine hydroxylase:
Mechanistic insight into O2 activation. J. Am. Chem. Soc. 131, 7685−
7698.
(19) Gaweska, H., and Fitzpatrick, P. F. (2011) Structures and
mechanism of the monoamine oxidase family. Biomol. Concepts 2,
365−377.
(20) Wu, T., Yankovskaya, V., and McIntire, W. S. (2003) Cloning,
sequencing, and heterologous expression of the murine peroxisomal
flavoprotein, N1-acetylated polyamine oxidase. J. Biol. Chem. 278,
20514−20525.
(21) Henderson Pozzi, M., Gawandi, V., and Fitzpatrick, P. F. (2009)
pH dependence of a mammalian polyamine oxidase: Insights into
substrate specificity and the role of lysine 315. Biochemistry 48, 1508−
1516.
(26) Johnson, K. A., Simpson, Z. B., and Blom, T. (2009) Global
Kinetic Explorer: A new computer program for dynamic simulation
and fitting of kinetic data. Anal. Biochem. 387, 20−29.
(27) Johnson, K. A., Simpson, Z. B., and Blom, T. (2009) FitSpace
Explorer: An algorithm to evaluate multidimensional parameter space
in fitting kinetic data. Anal. Biochem. 387, 30−41.
(28) Zechel, D. L., Konermann, L., Withers, S. G., and Douglas, D. J.
(1998) Pre-steady state kinetic analysis of an enzymatic reaction
monitored by time-resolved electrospray ionization mass spectrometry.
Biochemistry 37, 7664−7669.
(29) Li, Z., Sau, A. K., Furdui, C. M., and Anderson, K. S. (2005)
Probing the role of tightly bound phosphoenolpyruvate in Escherichia
coli 3-deoxy-D-manno-octulosonate 8-phosphate synthase catalysis
using quantitative time-resolved electrospray ionization mass spec-
trometry in the millisecond time range. Anal. Biochem. 343, 35−47.
(30) Hopfgartner, G., Wachs, T., Bean, K., and Henion, J. (1993)
High-flow ion spray liquid chromatography/mass spectrometry. Anal.
Chem. 65, 439−446.
(31) Page, J. S., Kelly, R. T., Tang, K., and Smith, R. D. (2007)
Ionization and transmission efficiency in an electrospray ionization-
mass spectrometry interface. J. Am. Soc. Mass Spectrom. 18, 1582−
1590.
(32) Sobott, F., and Robinson, C. V. (2004) Characterising
electrosprayed biomolecules using tandem-MS: The noncovalent
GroEL chaperonin assembly. Int. J. Mass Spectrom. 236, 25−32.
(33) Blackwell, A. E., Dodds, E. D., Bandarian, V., and Wysocki, V. H.
(2011) Revealing the quaternary structure of a heterogeneous
noncovalent protein complex through surface-induced dissociation.
Anal. Chem. 83, 2862−2865.
(34) Morgner, N., Montenegro, F., Barrera, N. P., and Robinson, C.
V. (2012) Mass spectrometry: From peripheral proteins to membrane
motors. J. Mol. Biol. 423, 1−13.
(35) Rose, R. J., Damoc, E., Denisov, E., Makarov, A., and Heck, A. J.
R. (2012) High-sensitivity Orbitrap mass analysis of intact macro-
molecular assemblies. Nat. Methods 9, 1084−1086.
(36) Binda, C., Angelini, R., Federico, R., Ascenzi, P., and Mattevi, A.
(2001) Structural bases for inhibitor binding and catalysis in
polyamine oxidase. Biochemistry 40, 2766−2776.
(37) Adachi, M. S., Taylor, A. B., Hart, P. J., and Fitzpatrick, P. F.
(2012) Mechanistic and structural analyses of the roles of active site
residues in yeast polyamine oxidase Fms1: Characterization of the
N195A and D94N enzymes. Biochemistry 51, 8690−8697.
(38) Daubner, S. C., Hillas, P. J., and Fitzpatrick, P. F. (1997)
Expression and characterization of the catalytic domain of human
phenylalanine hydroxylase. Arch. Biochem. Biophys. 348, 295−302.
(39) Daubner, S. C., Hillas, P. J., and Fitzpatrick, P. F. (1997)
Characterization of chimeric pterin-dependent hydroxylases: Con-
tributions of the regulatory domains of tyrosine and phenylalanine
hydroxylase to substrate specificity. Biochemistry 36, 11574−11582.
(40) Krzyaniak, M. D., Eser, B. E., Ellis, H. R., Fitzpatrick, P. F., and
McCracken, J. (2013) A pulsed EPR study of amino acid and
tetrahydropterin binding in a tyrosine hydroxylase nitric oxide
complex: Evidence for substrate rearrangements in formation of the
oxygen-reactive complex. Biochemistry 52, 8430−8441.
(22) Henderson Pozzi, M., Gawandi, V., and Fitzpatrick, P. F. (2009)
Mechanistic Studies of para-Substituted N,N′-Dibenzyl-1,4-diamino-
butanes as Substrates for a Mammalian Polyamine Oxidase.
Biochemistry 48, 12305−12313.
(23) Royo, M., and Fitzpatrick, P. F. (2005) Mechanistic studies of
mouse polyamine oxidase with N1,N12-bisethylspermine as a
substrate. Biochemistry 44, 7079−7084.
(24) Roberts, K. M., Pavon, J. A., and Fitzpatrick, P. F. (2013)
Kinetic Mechanism of Phenylalanine Hydroxylase: Intrinsic Binding
2679
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