From the Department of Biochemistry, Wake Forest University School of Medicine, Winston-Salem, North Carolina 27157
Received for publication, September 26, 2002, and in revised form, December 30, 2002
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ABSTRACT |
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Escherichia coli thiol peroxidase
(Tpx, p20, scavengase) is part of an oxidative stress defense system
that uses reducing equivalents from thioredoxin (Trx1) and thioredoxin
reductase to reduce alkyl hydroperoxides. Tpx contains three Cys
residues, Cys95, Cys82, and
Cys61, and the latter residue aligns with the
N-terminal active site Cys of other peroxidases in the peroxiredoxin
family. To identify the catalytically important Cys, we have cloned and
purified Tpx and four mutants (C61S, C82S, C95S, and C82S,C95S). In
rapid reaction kinetic experiments measuring steady-state turnover,
C61S is inactive, C95S retains partial activity, and the C82S mutation
only slightly affects reaction rates. Furthermore, a sulfenic acid
intermediate at Cys61 generated by cumene hydroperoxide
(CHP) treatment was detected in UV-visible spectra of
4-nitrobenzo-2-oxa-1,3-diazole-labeled C82S,C95S, confirming the
identity of Cys61 as the peroxidatic center. In
stopped-flow kinetic studies, Tpx and Trx1 form a Michaelis complex
during turnover with a catalytic efficiency of 3.0 × 106 M Oxidative stress defenses combat reactive oxygen species (1, 2)
such as superoxide (O AhpC, BCP, and Tpx are all members of the ubiquitous peroxiredoxin
(Prx) family within the Trx superfamily of protein folds; however, the
three E. coli Prx members are highly diverged from one
another and are representative of three distinct Prx subfamilies (11-13). Generally, the Prx active site contains a disulfide bond composed of a conserved N-terminal Cys (Cys46 from
Salmonella typhimurium AhpC) and the C-terminal Cys
(Cys165' from S. typhimurium AhpC) from the
other subunit of the antiparallel dimer, resulting in two symmetrical
active sites per dimer (14). Most Prxs contain two conserved Cys (2-Cys
Prxs); however, in some homologues, only the N-terminal Cys is retained
(the 1-Cys Prxs) (15). In many instances, the homodimers assemble into toroid-shaped decamers (15), a redox-dependent process in
S. typhimurium AhpC whereby reduced decamers disassociate
into dimers upon oxidation (16). To detoxify peroxides, the reduced
N-terminal Cys attacks the peroxide -O-O- bond, with concomitant
formation of a Cys sulfenic acid (Cys-SOH) intermediate, which then
condenses with the C-terminal Cys to regenerate the stable disulfide
bond at the active site (14, 17). For most bacterial Prxs, disulfide reduction is achieved by a specialized electron donor, AhpF (18), whereas many other Prx systems (both bacterial and eukaryotic) receive
electrons from a reducing system composed of Trx and Trx reductase
(TrxR) (13, 15).
Homologues of tpx are distributed throughout most or all
eubacterial species, both Gram-negative and Gram-positive, and are found in pathogenic strains such as Haemophilus influenzae,
Streptococcus pneumoniae, and Helicobacter pylori
(19), but biochemical and genetic analyses have been limited primarily
to E. coli Tpx. In response to oxidative stress, E. coli up-regulates Tpx expression through an oxygen-responsive
promoter element that is repressed by the transcriptional regulators
ArcA and Fnr under anaerobic conditions (9, 20, 21). In addition,
tpx deletion mutants, while still viable, were more
susceptible to oxidative stress and displayed diminished colony sizes
and numbers after peroxide exposure (22). In vitro studies
have confirmed that Tpx forms a Trx-linked peroxidase system capable of
reducing H2O2 and ROOH and protecting against
glutamine synthetase inactivation by a mixed function oxidation system
(9).
Tpx contains three Cys residues in its primary sequence,
Cys61, Cys82, and Cys95. Of these,
Cys61 aligns with the peroxidatic, N-terminal Cys of other
Prxs; whereas Cys95 does not align with the conserved
C-terminal Cys of other 2-Cys Prxs, it is conserved among all Tpx
homologues (19). Previous mutagenesis studies have presented
conflicting information about which Cys residues are involved in
peroxide attack and have indicated that both Cys61 and
Cys95 are essential for activity, whereas loss of
Cys82 only slightly attenuates activity (22, 23). In this
report, we identify Cys61 as the peroxidatic Cys forming a
Cys-SOH intermediate and firmly establish its intrasubunit linkage to
Cys95 in the redox-active disulfide. Although Tpx monomers
are not covalently linked, analytical ultracentrifugation studies
reported herein demonstrate that the enzyme is a homodimer in solution.
Materials--
SDS, ultrapure glycine, ultrapure urea, EDTA
disodium salt, dithiothreitol (DTT), ammonium sulfate,
Cloning of tpx into an Expression Vector--
Genomic DNA
prepared from a 400-ml overnight culture of an E. coli K-12
strain (XL-1 Blue) was used as the PCR template (24). The gene of
interest was amplified with the following PCR primers: forward primer,
5'-GCGAATTCAGGAGGAAGAATAGATGTCACAAACCGTACATTTCCAGGGC-3' and reverse primer 5'-GCCTGCAGTTATGCTTTCAGTACAGCC-3'
(engineered restriction sites underlined) synthesized in the DNA
Synthesis Core Laboratory of the Comprehensive Cancer Center of Wake
Forest University. PCR mixtures (50 µl) contained 200 µM dNTPs, 2 units of Vent DNA polymerase, 20 pmol each of
forward and reverse oligonucleotides, 1 mM
MgCl2, and 0.5 µg of genomic DNA. The reaction was
carried out in a Mini Cycler (MJ Research, Waltham, MA) as follows:
95 °C at 30 s, 55 °C for 45 s, and 72 °C for 1.5 min
(35 cycles) and then 72 °C for 15 min. The 507-bp tpx PCR
product purified with the QIAquick PCR Cleanup kit (Qiagen, Studio
City, CA) was then ligated into the pCR2.1 TA cloning vector
(Invitrogen) after Taq DNA polymerase/dNTP treatment.
Plasmid DNA purified from the E. coli host using the Wizard
Miniprep Kit (Promega) was screened for the presence of insert by
digestion with the appropriate restriction enzymes (EcoRI
and PstI). The DNA fragment containing tpx was excised from agarose gels and purified using the Gene Clean II Kit (Bio
101, Inc., Vista, CA). The fragment was ligated using T4 DNA ligase
into the pPROK1 expression vector (Clontech, Palo Alto, CA; expression under control of the tac promoter) that
had been similarly digested, isolated from an agarose gel, and
pretreated with calf intestinal phosphatase to generate
pPROK1/tpx.
Site-directed Mutagenesis of Tpx--
The four mutant E. coli Tpx enzymes, C61S, C82S, C95S, and C82S,C95S were created by
following the protocol outlined in the QuikChange site-directed
mutagenesis kit (Stratagene) using primers complementary to the coding
and noncoding template sequence (pPROK1/tpx) containing a
double-base mismatch. To generate the C61S mutation, the forward primer
5'-GATACCGGTGTTTCGGCCGCATCAGTACG-3' and a
complementary reverse primer were used (underlined letters indicate the
base pair mismatch). To generate the C95S mutation, the forward primer 5'-CGCCCAGTCTCGTTTCTCGGGCGCAGAAGG-3' and a complementary
reverse primer were employed, and C82S was constructed using the
forward primer 5'-CACCGTTGTGCTGTCTATCTCTGCCGATCTGCC-3' and
its complementary reverse primer. C82S,C95S was created using C95S
plasmid as template with the forward and reverse primers for the C82S
mutation. Reaction mixtures (50 µl) contained 10-50 ng of template
DNA (pPROK1/tpx or pPROK1/tpxC95S), 125 ng of
each primer, 200 µM dNTPs, and 1 µl of Pfu
Turbo polymerase. Twelve cycles of 95 °C for 30 s and 55 °C
for 13 min were carried out in a Mini Cycler followed by 55 °C for
10 min to finish extending products. To digest methylated template,
each reaction mixture was treated with 1 µl of DpnI at
37 °C for 1 h.
Bacterial Strains and Culture Procedures--
Ligated DNA
(pPROK1/tpx) and mutagenesis products were transformed into
XL-1 Blue cells. Single colonies were selected on Luria-Bertani (LB)
plates containing ampicillin (50 µg/ml), and those containing the
recombinant DNA were evaluated for protein expression by SDS-PAGE after
induction with 0.4 mM isopropyl
Purification of Recombinant and Mutant Tpx Proteins--
A
modification of a previous Tpx purification protocol was used for this
study (9). All procedures were carried out in a standard buffer (pH
7.0) consisting of 25 mM potassium phosphate with 1.0 mM EDTA. Briefly, 100 ml of XL-1 Blue E. coli
harboring the pPROK1/tpx plasmid were added to 10 liters of
LB medium containing 0.5 g of ampicillin supplemented with
0.2% glucose in a BioFlo 2000 fermentor (New Brunswick Scientific,
Edison, NJ). Isopropyl
Purifications of mutant Tpx proteins were carried out in the same
manner as for wild type Tpx with a few exceptions. The pooled C61S
mutant Tpx protein from the Phenyl-Sepharose column was loaded onto a
24 × 2.5-cm DEAE-cellulose column (DE52; Whatman, Kent, UK)
equilibrated with 30 mM potassium phosphate, and eluted
with a linear gradient from 30 to 80 mM potassium phosphate
(1 liter total volume). To completely purify C61S, fractions containing the mutant protein were reapplied to the Phenyl-Sepharose and DEAE-cellulose columns. Initial attempts at purifying the C95S mutant
under the same conditions as wild type resulted in aggregation of the
mutant protein, as observed by multiple bands during SDS-PAGE of the
subsequent fractions, even after many rounds of purification over the
two columns. DTNB titration of the isolated protein even after DTT
treatment gave less than one thiol (data not shown), suggesting that
Cys61 had become irreversibly overoxidized to a sulfinic
(Cys-SO2H) or sulfonic (Cys-SO3H) acid. The
addition of 2 mM DTT to all buffers prior to bacterial
disruption and over the course of the purification of C95S gave pure
protein after one round of purification on the two columns. C82S was
purified in buffers containing 2 mM DTT according to the
same protocol as C95S; however, pure C82S,C95S was obtained using a
slightly altered protocol. 20% instead of 10%
(NH4)2SO4 was used to treat
C82S,C95S prior to its application to a Phenyl-Sepharose column
equilibrated with 20% (NH4)2SO4, and C82S,C95S was eluted using a gradient of 20%
(NH4)2SO4 in 25 mM
potassium phosphate with 2 mM DTT to 0%
(NH4)2SO4 in deionized water with 2 mM DTT. Elution from the Q Sepharose column was achieved with a gradient of 20-120 mM potassium phosphate in 2 mM DTT. Prior to assay, each mutant was subjected to
ultrafiltration and washed with standard buffer to remove DTT and then
immediately incubated with equal volumes of TCEP gel.
Other Protein Purifications--
Purifications of E. coli TrxR (26) and E. coli Trx1 (27) were carried out
as described previously. S. typhimurium AhpF and S. typhimurium AhpC were purified as reported previously (25).
Fluorescein-5-Maleimide Labeling of Proteins--
Wild type,
C61S, C82S, C95S, and C81S,C95S Tpx (100 µg each), prereduced with
DTT that was removed by ultrafiltration, were reacted with fluorescein-
5-maleimide (10 equivalents) under denaturing (4 M urea)
and reducing (100 eq of TCEP) conditions for 16 h at 4 °C in
standard buffer. Protein samples (5 µg) were separated on 18%,
20-cm-long SDS-polyacrylamide gels. Densitometry to assess the protein
contents of fluorescein-labeled bands was conducted using the Quantity
One quantitation software from Bio-Rad, and image files were generated
with a ChemiImager 5500 digital imaging system from Alpha Innotech
Corp. (San Leandro, CA) and a near UV light source and UV filter. For
quantitation, six amounts of wild type Tpx (0.03-5 µg) were
electrophoresed along with each of the mutant samples. Electrophoresis
samples also included one with 5 µg of C61S, to which 0.25 µg of
wild type Tpx was intentionally added, and one into which 1.7 µg each
of wild type Tpx and C61S and C82S,C95S mutants were loaded to assess
the quality of the separation.
The labeled proteins were also submitted for electrospray mass
spectrometry, as described below, following ultrafiltration into water
to remove buffer using Apollo concentrators (Orbital Biosciences).
Analytical Ultracentrifugation Analyses--
To determine the
oligomeric state of wild type and mutant proteins, samples of different
concentrations were analyzed by sedimentation equilibrium at various
speeds on an Optima XL-A analytical ultracentrifuge (Beckman
Instruments, Palo Alto, CA) outfitted with absorbance optics. Tpx,
C61S, and C95S (purified in DTT) were exchanged into a buffer of 25 mM potassium phosphate, 1 mM EDTA, and 0.15 M NaCl, pH 7.05, via ultrafiltration. Three different
concentrations of each protein (39.5, 184.2, and 342.1 µM) in 110 µl were loaded into three of the six sectors
of each cell, and buffer (125 µl) was loaded into the remaining
sectors as a reference. Data were obtained at 11,000, 16,000, and
20,000 rpm at 20 °C following equilibration for 8, 10, and 12 h
at each speed. Data with absorbances higher than 1.4 were excluded from
data analysis. The partial specific volumes for Tpx and the mutants
were calculated from the amino acid composition to be 0.7433 cm3 g Mass Spectrometric Analyses--
Protein samples were
extensively dialyzed in deionized water (6 liters) in a Slide-A-Lyzer
cassette (Pierce) overnight prior to analysis by electrospray
ionization mass spectrometry (ESI-MS; Micromass, Manchester, UK)
precalibrated with horse heart myoglobin. The protein sample (1 µM) in 1% formic acid was injected at a flow rate of 300 µl/h, and positively charged ions in the m/z range of 800-1800 were analyzed using MassLynx software (version 3.5; Micromass).
Spectral Experiments--
Most spectral assays were conducted on
a single wavelength, thermostatted (25 °C) Gilford 220 updated
recording spectrophotometer (Oberlin, OH) with a Beckman DU
monochrometer (Fullerton, CA) unless otherwise noted. Microbiuret
assays for proteins to determine extinction coefficients, disulfide
assays with 2-nitro-5-thiosulfobenzoate, and thiol quantification with
DTNB were conducted as described previously (25, 30). To further
quantify Tpx proteins by absorbance, the following experimentally
determined extinction coefficients at 280 nm were used: Tpx and
C82S,C95S, 3800 ± 300 M Steady-state Kinetic Analysis of Tpx and Mutants--
Michaelis
constants (Km) and turnover numbers
(kcat) with CHP as the substrate for Tpx were
obtained as described previously for H. pylori TrxR (35).
Briefly, reactions were conducted on an Applied Photophysics DX-17 MV
stopped-flow spectrophotometer (Surrey, UK) at 25 °C, and activity
was followed by monitoring the decrease in fluorescence or absorbance
of NADPH over time. Use of the stopped-flow spectrophotometer for
mixing and data acquisition helped alleviate nonlinearity and
reproducibility problems inherent in the manual mixing methods (14).
Reaction mixtures in one syringe contained NADPH (150 µM), Trx1 (2-80 µM), TrxR (0.25-10
µM, in a molar ratio of 1:8 TrxR to Trx1, so that the
electron transfer to Trx1 is not rate-limiting), and Tpx (0.2 µM) in peroxidase buffer consisting of 50 mM
potassium phosphate (pH 7.0), 0.1 M
(NH4)2SO4, and 0.5 mM
EDTA. CHP (1-50 µM) was prediluted into
Me2SO (in a ratio of 1:50 with the solvent) and mixed with
peroxidase buffer in the other syringe. In all cases, concentrations
given are final concentrations achieved after mixing of the contents of
the two different syringes. Under the conditions given, Tpx was
supplied in rate-limiting concentrations, and due to the saturable
interaction between Tpx and Trx1, the reaction was linearly dependent
on Trx1, provided that concentrations supplied were below the
Km. Assays were designed so that Trx1 levels were at
least 10-fold higher than those of Tpx to maintain the peroxidase as
the limiting enzyme. Flavin-dependent oxidase activity did
not contribute significantly to NADPH oxidation (0.05 s Sulfenic Acid Trapping Experiments--
To observe the formation
of Cys-SOH as a reaction intermediate, experiments with C61S, C95S, and
C82S,C95S mutants labeled with NBD chloride were carried out as
described previously (17, 37), with a few exceptions. Each mutant (171 µM in 100 µl, enough protein to give an absorbance of
0.5 at 280 nm) was preincubated with DTT for 1 h and then desalted
by ultrafiltration. C61S and C95S proteins in standard buffer were made
anaerobic by repeated flushing with argon gas and vacuum in alternating
cycles for 30 min. Anaerobic solutions of NBD chloride (50 mM in Me2SO) and CHP (110 mM in
Me2SO) were prepared by bubbling argon through the
preparations for 15 min. Under anaerobic conditions, the C61S and the
C95S mutants were oxidized by H2O2 (1 eq), and
then the proteins were incubated with NBD chloride (10 eq) for 5 min.
Excess NBD chloride was removed by ultracentrifugation, and the
absorbance properties of the protein samples were analyzed (200-600
nm) on a Beckman DU 7500 diode array spectrophotometer (Fullerton, CA). Labeling of prereduced C165S AhpC from S. typhimurium (48.3 µM) with NBD chloride was used as a positive control. Due
to its sensitivity toward oxidation, C82S,C95S was stored with equal
volumes of TCEP gel after removal of DTT and treated with CHP rather
than H2O2 prior to NBD chloride labeling.
Tryptic Peptide Separations by HPLC--
To prepare proteins for
digestion, prereduced Tpx (10 nmol) was treated with or without
H2O2 (1 eq) and then incubated in MES (125 mM, pH 6.5) buffer with EDTA (1 mM) prior to
reaction with 4-vinylpyridine and subsequent denaturation in urea (1.6 M) as described (30). The addition of 4-vinylpyridine to
block free Cys was necessary to prevent adventitious disulfide bond formation or migration following denaturation. Following dialysis and
solvent removal, exhaustive tryptic digestion of Tpx, C61S, or C95S in
either oxidized or reduced forms was carried out as described (30).
Tryptic maps were generated by injecting samples into a Rainin Dynamax
HPLC system equipped with an AquaPore RP-300 C8 column (4.6 × 100 mm) and were eluted using a 90-min gradient consisting of 5-60%
Solvent B (Solvent A was 0.1% trifluoroacetic acid in deionized,
ultrapure H2O; Solvent B was 70% acetonitrile with 0.08%
trifluoroacetic acid in H2O). Peaks were detected at 215 and 254 nm on a Dynamax UV-DII dual wavelength detector. To isolate the
disulfide-containing fragments, peaks at 68 (P1) and 70 min (P2) from
oxidized Tpx were isolated and then further purified on a shallower
gradient. Acetonitrile and trifluoroacetic acid were then removed by
vacuum centrifugation, and solutions were brought to a final volume of
100 µl with deionized H2O prior to analysis by
ESI-MS.
Characterization of Purified Tpx and Mutants--
Homogeneous,
recombinant wild type Tpx and mutant proteins were isolated using a
modification of the previous purification protocol (9), and all mutants
were purified in the presence of 2 mM DTT to prevent
overoxidation of free Cys residues (14). Mutant enzymes were difficult
to prepare under oxidizing conditions due to irreversible protein
dimerization, and C95S purified without DTT displayed significantly
less activity than C95S purified under reducing conditions. DTNB
titrations of denatured, prereduced wild type Tpx and mutants confirmed
the expected thiol content for each (Table
I). Additionally, ESI-MS of Tpx indicated
a mass of 17,700 atomic mass units, 135 atomic mass units less than the mass of 17,835 atomic mass units predicted by the tpx open
reading frame, indicating the loss of the initiating Met from E. coli Tpx. Each single mutant exhibited a mass of 17,686 atomic
mass units, corresponding to a 16 atomic mass units loss due to the Cys
to Ser mutation, whereas the 17,672-atomic mass units mass for
C82S,C95S confirmed the double Cys to Ser mutation. The Cys to Ser
mutations are conservative, and spectral and circular dichroism scans
conducted in the far ultraviolet region revealed no gross structural
perturbations among the mutants compared with the wild type protein
(data not shown).
As mutant proteins were isolated from E. coli expressing low
amounts of wild type Tpx, labeling experiments were conducted using
fluorescein-5-maleimide to shift protein molecular weights according to
the number of cysteine residues (an increase in 427 g/mol per
fluorescein-labeled cysteine residue) and to enhance quantitation by
densitometry. Using long gels and 18% acrylamide, singly, doubly, and
triply labeled proteins were nicely separated (data not shown).
Including data from kinetic studies described below, which give an
upper limit of 0.4% for the degree of contamination of the C61S mutant
by wild type Tpx (if all activity is due to the presence of wild type),
densitometry of the gel samples taking into account this information
indicates a wild type Tpx contamination level of less than 5%.
Verification of the single (C82S,C95S), double (C61S, C82S, and C95S
mutants), and triple (wild type Tpx) labeling of these proteins by
fluorescein was obtained by mass spectrometry. Furthermore,
purification protocols for C61S and C82S,C95S mutants were
significantly different from that for the wild type enzyme, making
contamination by the wild type enzyme less likely in these cases.
Oligomeric State of Tpx--
Earlier studies using a gel
filtration column standardized with molecular weight markers suggested
that oxidized Tpx was a monomer of 16.8 katomic mass units (9). In
addition, the 2-atomic mass units difference for Tpx ESI-MS masses
obtained under reducing (17,698.4 ± 0.62 atomic mass units) and
nonreducing (17,696.7 ± 0.23 atomic mass units) conditions and
the nonreducing SDS-PAGE analysis (see below) exclude the possibility
of a covalent dimer for oxidized Tpx. Nonetheless, most other 2-Cys Prx
proteins examined thus far, except for human PrxV (38), exist as
covalent dimers when oxidized and noncovalent dimers or higher order
oligomers when reduced (15). The oligomeric state of Tpx in solution
was, therefore, examined by analytical ultracentrifugation
sedimentation equilibrium experiments. Data analyses of three separate
data sets each (40-340 µM) gave weight-average molecular
weights for Tpx of 29,500 and 32,000 for the oxidized and
reduced enzyme, respectively, indicating that two Tpx monomers
self-associate in solution independent of redox state. Tpx dimerization
is not concentration-dependent above 40 µM,
because similar molecular weights are obtained at different speeds and
all three concentrations. Detection of dimeric molecular weights for
both C61S and C95S under the same experimental conditions indicates
that these mutations do not perturb self-association.
Most 2-Cys Prxs are linked by an intersubunit disulfide bond at the
active site in their oxidized state (25, 39). However, previous
SDS-polyacrylamide gel studies of wild type Tpx revealed only a 19-kDa
band under reducing and nonreducing conditions, suggesting the
formation of an intrasubunit, rather than intersubunit, disulfide bond
upon oxidation (9, 19). In the present study, a second species (~36
kDa) was observed on SDS-PAGE conducted under nonreducing conditions
only when a thiol-specific blocking agent, methyl
methanethiolsulfonate, was excluded from sample preparations prior to
denaturation (data not shown), indicating that any covalent
dimerization of Tpx is artifactual. Furthermore, during SDS-PAGE
analysis under nonreducing conditions, only a very small downward shift
in protein migration is observed upon oxidation (from an apparent
molecular mass of 18.5 to 18.2 kDa, data not shown), consistent with
intrasubunit disulfide bond formation as analyzed further using tryptic
mapping techniques.
Identification of the Intrasubunit Disulfide Linkage--
To
assess the redox state of Cys residues and identify the location of the
putative intrasubunit disulfide linkage, a series of biochemical assays
were conducted on oxidized and reduced wild type Tpx. Reduced Tpx
contained no disulfide bonds (as quantified by
2-nitro-5-thiosulfobenzoate assays) and 1.90 ± 0.03 or 2.60 ± 0.09 free thiols/monomer under native or denaturing conditions, respectively (Table I), indicating the presence of one buried, slow
reacting Cys thiol. After the addition of 1 eq of peroxide to
prereduced Tpx, the two accessible thiol groups were lost (Table I),
and one disulfide bond (0.90 ± 0.01) per monomer was gained.
To identify the disulfide-forming Cys residues, tryptic maps of
oxidized and reduced, pyridylethylated Tpx were generated by HPLC and
compared at 215 and 254 nm (Fig. 1).
Instead of two strong peaks at 254 nm in the reduced, pyridylethylated
chromatograms, four strong peaks were observed around 54, 56, 58, and
74 min, which allowed for the identification of Cys-containing
peptides. After comparing the oxidized and reduced tryptic maps at 215 nm, two new peaks representing the disulfide-containing peptides were detected in the oxidized map at 68 (P1) and 70 min (P2), representing the disulfide-containing peaks (Fig. 1). P1 and P2 were reisolated on a
shallower gradient and then reduced and pyridylethylated and reanalyzed
by HPLC. P1 and P2 each gave two new peaks corresponding to two of the
254-nm absorbing peaks observed in the original chromatogram of the
reduced, pyridylethylated protein (data not shown). Masses of P1 (3830 atomic mass units) and P2 (3702 atomic mass units) as determined by
ESI-MS correspond to Cys61- and
Cys95-containing peptides and verify that Cys61
and Cys95 compose the active site intrasubunit disulfide
bond (Fig. 1). The presence of adjacent tryptic digest sites upstream
of Cys61 resulted in two disulfide-containing peaks due to
the partial digestion after Arg47.
Kinetic Characterization of Tpx and Inactivation during
Catalysis--
The Trx1-linked peroxidase activity of Tpx and its
differential activity with ROOH and H2O2 were
reported earlier (9, 19). In this study, the true
kcat and Km values of Tpx for Trx1, CHP, and H2O2 were determined from
Hanes-Woolf plots of the initial rate data (Table
II). All lines in the Hanes-Woolf primary
plots intersect the origin, indicating a bisubstrate, ping-pong
(substituted) reaction mechanism for Tpx, which can be depicted as a
sequence of consecutive reactions,
Tpx steady-state reactions followed for longer than 5 s exhibited
rapidly declining NADPH oxidation rates over time for CHP, but not
H2O2, that were dependent on the concentration
of peroxide. This observed inactivation of Tpx required a threshold
level of CHP, whereby at high levels of CHP (>150 µM),
but not at low CHP levels (<100 µM), activity diminished
rapidly and nonlinearly and resulted in incomplete consumption of
peroxide (Fig. 2). Activity could not be
regained with the addition of new substrates; however, supplementation
of reactions with fresh Tpx restored NADPH oxidation, which again
diminished rapidly due to inactivation of the newly added Tpx (data not
shown). Inactivated reaction mixtures (i.e. 200 µM CHP and levels of NADPH, TrxR, Trx1, and Tpx matching
those used for the steady-state assays) that had been incubated with DTT and subsequently washed could not regenerate activity in the presence of fresh NADPH and CHP. These data suggest that inactivation is due to an irreversible overoxidation process that is most likely occurring at the active site Cys-SOH, a transient species that can be
readily and irreversibly overoxidized by excess peroxide to
Cys-SO2H or Cys-SO3H (14, 41). Prereduced Tpx
incubated overnight in excess CHP retained full activity, indicating
that inactivation requires Tpx turnover. ESI-MS verification of active site Cys overoxidation was obtained by injecting freshly inactivated reaction mixtures and immediately determining Tpx mass after turnover. After subtracting the contribution of TrxR and Trx1 ions from the
reaction mixture spectra, the mass of Tpx was found to be 17,732 atomic
mass units, an increase in 32 atomic mass units, which is consistent
with the addition of two oxygen atoms. Together, these data strongly
suggest that during turnover, Tpx's active site Cys is converted to
Cys-SO2H in an overoxidation process relying on
a threshold amount of peroxide and that the Cys-SOH formed at the
active site follows one of two pathways with each catalytic cycle (Fig.
3).
Bisubstrate kinetic analyses have confirmed that Tpx reacts
differentially with soluble peroxides as evidenced by the large difference in Km for H2O2
and CHP (1750 and 9.1 µM, respectively) (Table II). Upon
examination of the Km(app) values for other soluble peroxides (in reactions conducted at [Trx1] = 10 µM), we find that whereas H2O2
and CHP maintain their status as the worst and best substrates,
respectively (Km(app) values of 648 versus 3.9 µM), Tpx has a lower
Km(app) for ethyl hydroperoxide than for
t-butyl hydroperoxide (30.7 versus 66.6 µM). Examination of Tpx reactivity with a physiological
alkyl hydroperoxide, a relatively insoluble fatty acid hydroperoxide, 15-HPETE, resulted in a complex pattern of
[15-HPETE]-dependent enzyme activity and inactivation. At
low concentrations (2-6 µM), activity increased steadily
with peroxide concentrations, but at higher [15-HPETE] (10-30
µM), activity was significantly decreased (Fig.
4). Incubation of the Tpx reaction
mixture (i.e. NADPH, TrxR, Trx1, and Tpx) with increasing
amounts of 15-HPETE followed by the addition of a noninactivating
amount of CHP (50 µM) resulted in loss of Tpx activity
with higher concentrations of 15-HPETE (10-30 µM) (data
not shown), suggesting that Tpx is also inactivated by this substrate.
The resulting, nonhyperbolic curve representing [15-HPETE]-dependent activity (Fig. 4) is not amenable to
typical Michaelis curve fits and is further complicated by the limited solubility of 15-HPETE in aqueous buffers. Therefore, apparent second
order rate constants were calculated from the slope of a line drawn
through the points for 0-4 µM to give a
kcat(app)/Km(app) of 2.6 × 105 M Tpx Specificity toward Trx1 Reduction--
We examined the
specificity of the interaction between Tpx and Trx1 by testing whether
or not other E. coli enzymatic disulfide/dithiol reductants
could serve as reductase systems. Because Trx2 (trxC) is
up-regulated in response to oxidative stress (43, 44) and exhibits
similar disulfide reductase capabilities as Trx1 (44, 45), we
postulated that Trx2 may be able to substitute for Trx1. However,
replacement of Trx2 in the steady-state Tpx assay did not result in
NADPH oxidation (data not shown). Previous work also indicated that Tpx
was active in the presence of high levels of glutathione (10-20
mM) (9); however, replacing the TrxR/Trx1 reductase system
with glutathione reductase and glutathione (up to 40 mM)
did not support Tpx turnover in our stopped-flow assays (data not
shown). Additionally, neither E. coli glutaredoxin 1 nor
E. coli AhpF were capable of reducing Tpx, indicating that Tpx peroxidase activity specifically requires Trx1.
Cys61 Is the Peroxidatic Center in Tpx--
To address
which of the three Cys residues in Tpx are involved in the direct
reduction of peroxide, Cys to Ser mutants were analyzed under
steady-state assay conditions using a stopped-flow spectrophotometer.
Unlike in previous studies, where both C61S and C95S were inactive (22,
23), we detected activity with C95S (~20% of wild type activity with
10 µM Trx1) but not with C61S in our assays (Fig.
5), indicating that Cys61 is
the essential peroxidatic Cys of the disulfide pair (the activity of
C61S is about 0.4% that of wild type Tpx and may be entirely due to a
very small amount of wild type contamination in the C61S protein). C95S
is also more sensitive to inactivation than wild type Tpx, as
illustrated by less robust reaction rates that are more quickly
diminished even in the presence of low concentrations of peroxide (data
not shown). The loss of Cys82 does not have a large effect
on activity (~72% activity compared with wild type Tpx);
interestingly, removal of both Cys82 and Cys95
in the double mutant, C82S,C95S, dramatically decreases activity below
that of C95S to about 10% of wild type activity (Fig. 5), suggesting a
modest, stabilizing effect of Cys82 in the absence of
Cys95. When the single Cys Tpx mutants were assayed with
varying amounts of Trx1 (0-80 µM) at one concentration
of CHP (50 µM), we found that C82S retained the same low
Km(app) for Trx1 as wild type Tpx (14.8 versus 12.3 µM), whereas C95S had a much
higher Km(app) for Trx1 (50.7 µM). Our labeling and densitometry data verifying less
than 5% contamination of the mutants by wild type Tpx, taken together
with the value of 20% for the rate of C95S compared with wild type Tpx
(at 10 µM Trx1) and the unique kinetic properties of this
mutant (higher Km(app) for Trx1 and more
rapid inactivation during turnover), clearly indicate that C95S does
retain activity, although removal of part of the Tpx redox-active
disulfide center adversely affects its normal catalytic activity with
Trx1 and exacerbates its inactivation by peroxides.
Identification of the Sulfenic Acid Form of
Cys61--
NBD chloride labeling and x-ray crystallography
have directly demonstrated the presence of a R-SOH intermediate on the
peroxidatic Cys of several Prx homologues (17, 46). During catalysis, the labile Cys-SOH is quickly attacked by the other half-cysteine of
2-Cys Prxs to reform the redox active disulfide; therefore, to
stabilize and trap Cys-SOH, the C-terminal Cys of the active site
disulfide pair must be removed. Because C95S and C82S, but not C61S,
display a reduced thiol titer upon oxidation (Table I) and
Cys61 is essential for activity (Fig. 5), SOH formation on
Cys61 was hypothesized. Detection of SOH in C95S was
hampered by artifactual enzyme dimerization and apparent NBD migration
to Cys82, resulting in a low Cys-SOH signal (data not
shown). Therefore, NBD chloride labeling to detect the putative Cys-SOH
intermediate was conducted using a Tpx double mutant, C82S,C95S, which
still maintained slight activity during steady-state assays (Fig. 5). In accordance with SOH formation following peroxide treatment, the
thiol titer of C82S,C95S also decreased in a manner similar to that of
C95S (Table I). In the resulting ultraviolet-visible wavelength
spectral scan, oxidized, NBD-labeled C82S,C95S exhibited an absorbance
maximum at 347 nm (Fig. 6), a signal
characteristic of NBD adducts of R-SOH (R-S(O)-NBD) (17, 37), which
clearly signifies the detection and trapping of approximately
stoichiometric amounts of SOH at Cys61, the only Cys in
this mutant protein. The absorbance maximum at 420 nm for reduced
C82S,C95S (Fig. 6), a signal diagnostic for NBD-modified thiols
(R-S-NBD), was much attenuated in the oxidized spectra, confirming the
loss of the thiolate species (RS Although Tpx was excluded initially from the Prx family (9) due to
its low sequence identity (17% compared with E. coli AhpC),
the high z score (>70) for the alignment of Tpx with 2-Cys mammalian
Prx homologues in fold and function assignment system (FFAS)
analysis (47) confirms homology of Tpx with the Prxs. Not surprisingly
then, Tpx shares many features of the Prx catalytic mechanism,
including a reliance on the conserved N-terminal Cys for peroxide
reduction (15). Of Tpx's three Cys residues (Cys61,
Cys82, and Cys95), Cys61 was shown
in our mutagenesis studies to be essential for attack of the
peroxide's -O-O- bond (Fig. 4) and to form a Cys-SOH intermediate upon oxidation (Fig. 6). Our assay data, collected using stopped-flow spectrophotometry, captured initial reaction rates before overoxidation of Cys61, and it is likely that prior inability to detect
mutant C95S activity (22, 23) was due to the rapid overoxidation of
Cys61 during purification in the absence of DTT and/or
during the manual mixing procedures used in assay methods. Tpx does not
form the hallmark intersubunit disulfide linkage of typical 2-Cys Prxs, and instead Cys61 links with Cys95 in an
unusual intramolecular disulfide bond, a reaction intermediate common
to atypical 2-Cys Prxs such as human PrxV (15). Whereas Cys95 does not align with the C-terminal Cys of other 2-Cys
Prxs, the participation of Cys95 as a disulfide partner
signals that Cys95 is functionally equivalent to the
"resolving" Cys of other 2-Cys Prxs (15).
Investigations of Prx interactions with their reducing systems have
revealed two different kinetic patterns where 1) the formation of
enzyme substrate complexes is an observable phenomenon (25, 26, 48, 49)
or 2) reduction of the Prx is a bimolecular process, whereby infinite
values for Km and Vmax
characterize activity (13, 35). The specific reduction of Tpx by Trx1
occurs with a Km of 25 µM and a
catalytic efficiency of ~3 × 106
M Earlier reports of the greater catalytic efficiency of Tpx with
t-butyl hydroperoxide over H2O2 (9)
were confirmed in our own kinetic studies with CHP and
H2O2 as substrates (giving true Km values of 9.1 and 1750 µM,
respectively) (Table II). As a result, Tpx does not seem to exhibit the
requirements for a minimal peroxide binding site as observed for the
Trx-dependent Prx from H. pylori (35). In light
of this, the nature of the Tpx peroxidatic binding site is likely to be
complex, and due to its high affinity for CHP, hydrophobic interactions
may be the predominating forces that influence substrate interaction with the enzyme.
Previously, it was shown that the primary role of AhpC was to keep
concentrations of H2O2 in exponentially growing
E. coli quite low (10 Inactivation has been observed for many different Prx homologues during
activity assays (14, 55-58), and, when investigated, irreversible
overoxidation at the active site Cys is responsible for the loss of
activity (57), with the terminal species being sulfinic acid in some
cases (15). If Tpx inactivation requires a threshold level of peroxide
that is substantially higher than in vivo concentrations,
then the observed in vitro overoxidation for Tpx is due to
the use of very high substrate concentrations in steady-state assays.
This seems probable, considering that the endogenous in vivo
concentrations of H2O2 are reportedly quite low
in the absence of exogenous sources (59). Exposure to peroxide at
endogenous concentrations of 2 µM or more is
growth-inhibitory (7), and in these cases, peroxidase activity may no
longer prevent oxidative damage to cellular components. Whereas the
in vivo concentrations of other ROOH in bacteria are not
known, it is likely that they would not exceed
H2O2 levels. Therefore, subjecting Tpx during
assays to exceptionally high peroxide levels may artificially promote
overoxidation, a future consideration for all Prx steady-state analyses
done without the benefits of stopped-flow reaction techniques.
To explain the existence of multiple Prxs, the differential cellular
localization of Tpx and AhpC has been cited. Tpx has been characterized
as a periplasmic protein (9, 60), despite the lack of a signal sequence
for periplasmic transport,3
whereas AhpC has been localized to the cytoplasm (60). E. coli proteomics studies by Link et al. (60) showed that
during the growth phase in minimal media and without induction by
H2O2, Tpx (~1.6 µM) is 3.5-fold
less abundant than AhpC and 2.5-fold more abundant than BCP. The
relatively high expression of all three Prxs indicates a sustained
requirement for peroxide detoxification during growth, and in these
cases, a periplasmic peroxidase metabolizing peroxides prior to
cytosolic entry would be quite beneficial. Although Trx1 is relatively
abundant (0.3% of the cellular protein) (61), its cytoplasmic location
(62) would restrict access of Tpx to its reductant and decrease the
catalytic efficiency of the Tpx system. The designation of Tpx as a
periplasmic protein should then be treated with caution until more
careful localization studies are completed.
Detailed kinetic analyses of Tpx reaction rates have allowed us to
clarify the roles of the three Cys in E. coli Tpx herein and
demonstrate the essentiality of Cys61. The shared reliance
on Cys-SOH formation and disulfide bond formation points to mechanistic
similarities between Tpx and other members of the Prx family. Further
kinetic and mechanistic studies on Prx family members may begin to
delineate the functional roles of multiple Prxs in the same organism,
the presence of which most likely signifies considerable selection
pressure from oxidative stress and the need to combat reactive oxygen species.
1 s
1, and the
low Km (9.0 µM) of Tpx for CHP
demonstrates substrate specificity toward alkyl hydroperoxides over
H2O2 (Km > 1.7 mM). Rapid inactivation of Tpx due to Cys61
overoxidation is observed during turnover with CHP and a lipid hydroperoxide, 15-hydroperoxyeicosatetraenoic acid, but not
H2O2. Unlike most other 2-Cys peroxiredoxins,
which operate by an intersubunit disulfide mechanism, Tpx contains a
redox-active intrasubunit disulfide bond yet is homodimeric in solution.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-mercaptoethanol, 5,5'-dithiobis(2-nitrobenzoic acid) (DTNB), Tris
base, and other buffer reagents were purchased from Research Organics
(Cleveland, OH). Bacteriological media components were from Difco.
Ethanol was obtained from Warner Graham Company (Cockeysville, MD).
Isopropyl
-D-thiogalactopyranoside and X-gal
(5-bromo-4-chloro-3-inolyl-
-D-galactopyranoside) were from Inalc (Milan, Italy). Vent DNA polymerase was purchased from New
England Biolabs (Beverly, MA). Restriction enzymes, T4 DNA ligase, calf
intestinal phosphatase, Taq DNA polymerase,
MgCl2 solutions, and restriction buffers were obtained from
Promega (Madison, WI). Agarose medium EEO (electrophoresis grade),
organic solvents (high pressure liquid chromatography (HPLC) grade),
water (optima grade), sodium borate, acetic acid, sodium chloride, and H2O2 were from Fisher. Acrylamide/bis (40%)
solution was purchased from Bio-Rad. Ampicillin powder,
chloramphenicol, streptomycin sulfate, formic acid, calcium chloride,
cumene hydroperoxide (CHP), dimethyl sulfoxide (Me2SO),
protocatechuic acid, protocatechuate-3,4dioxygenase, lipoxygenase,
Saccharomyces cerevisiae glutathione reductase, E. coli glutaredoxin 1, and insulin were from Sigma. Ethyl
hydroperoxide was from Polysciences, Inc. (Warrington, PA).
Glutathione, 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole (NBD chloride),
4-vinylpyridine, and tert-butyl hydroperoxide were from
Aldrich. NADPH and NADH were from Roche Molecular Biochemicals. L-1-(Tosylamino)-2-phenylethyl chloromethyl ketone-treated
trypsin was obtained from Worthington. Pierce supplied the immobilized Tris[2-carboxyethyl]phosphine (TCEP) disulfide-reducing gel, Gel Code
Blue stain, and trifluoroacetic acid ampules. Molecular Probes, Inc. (Eugene, OR) supplied the methyl methanethiolsulfonate and fluorescein-5-maleimide. Arachidonic acid was from NuCheck Prep (Elysian, MN). Stratagene (La Jolla, CA) supplied the Pfu
Turbo polymerase, DpnI, and E. coli XL-1 Blue
competent cells. All ultrafiltration was carried out using Apollo 7-ml
high performance centrifugal concentrators (Orbital Biosciences,
Topsfield, MA) unless otherwise noted.
-D-thiogalactopyranoside. Isolated plasmid DNA for each
construct was sequenced throughout the coding region by automated DNA
sequencing at the Comprehensive Cancer Center of Wake Forest
University. Bacterial stocks containing each plasmid with the subcloned
gene were prepared from a single colony and stored at
80 °C in LB
broth containing 15% (v/v) glycerol. Culture procedures were generally
the same as reported earlier (25).
-D-thiogalactopyranoside (0.4 mM) was added at A600 = 0.9, and bacteria were harvested by centrifugation 16 h after induction. Pelleted bacteria were disrupted with a Bead Beater (BioSpec Products, Bartlesville, OK), and cell extracts treated with streptomycin sulfate
to precipitate nucleic acids were subjected to 30 and 75%
(NH4)2SO4 treatments to precipitate
proteins (25). The protein mixture resuspended in standard buffer
containing 10% (NH4)2SO4 was
applied to a 24 × 2.5-cm Phenyl-Sepharose 6 Fast Flow Column (Amersham Biosciences), washed with 10%
(NH4)2SO4 buffer, and eluted with
deionized H2O. Protein fractions were evaluated for contamination of overexpressed Tpx by SDS-PAGE, and the purest fractions were pooled. After dialysis against 5 mM
potassium phosphate buffer (pH 7.0), the protein was loaded onto a
Q-Sepharose column (Amersham Biosciences) pre-equilibrated in 5 mM potassium phosphate and eluted with a linear gradient
from 5 to 30 mM potassium phosphate (1 liter total volume).
Again, fractions were analyzed for Tpx by SDS-PAGE, and the pure
fractions were pooled, concentrated, and aliquotted for storage at
20 °C.
1 and 0.7434 cm3
g
1, respectively (28). Multiple data sets were globally
fit to the equation for a single ideal species using WinNonLin2 (29). The buffer density of 1.00773 g/cm3 was determined using a
DA-310 M precision density meter (Mettler Toledo, Hightstown, NJ) at
20 °C.
1
cm
1; C95S, 3500 ± 200 M
1
cm
1; C82S, 4200 ± 600 M
1
cm
1; C61S, 5300 ± 700 M
1
cm
1. Other extinction coefficients used were as follows:
E. coli TrxR, 11,300 M
1
cm
1 (454 nm) (31); E. coli Trx1, 13,700 M
1 cm
1 (280 nm) (32); S. typhimurium AhpC, 24,300 M
1
cm
1 (280 nm) (25); S. typhimurium AhpF, 13,100 M
1 cm
1 (450 nm) (25); E. coli glutaredoxin 1, 12,400 M
1
cm
1 (280 nm) (33); NADPH, 6200 M
1 cm
1 (340 nm); NADH, 6220 M
1 cm
1 (340 nm);
2-nitro-5-thiobenzoate (TNB2
), 14,150 M
1 cm
1 (412 nm) (34).
1)
and was not subtracted from initial rates. After conversion to
concentration units/min, the primary rate data were fit to a
rectangular hyperbolic curve function in Sigma Plot (Jandel Scientific,
San Rafael, CA). The data obtained for each substrate concentration
were plotted according to the Hanes-Woolf representation of the
Michaelis-Menten equation to give intersecting lines at the
y axis, indicating a substituted (ping-pong) mechanism and were further evaluated using the Hanes-Woolf equation for a
two-substrate, substituted reaction as previously described (35).
Because of the much larger Km of Tpx for
H2O2, larger amounts of substrates were present
in the reaction mixtures, and the sensitivity required previously in
the fluorescence assay was no longer necessary. Therefore, all
H2O2 data were collected in absorbance mode on the stopped flow spectrophotometer with the same protein assay mixtures
as above, except that the second syringe contained varying H2O2 concentrations (0.25-10 mM).
Synthesis of 15-hydroperoxyeicosatetraenoic acid (15-HPETE) was carried
out as described by O'Flaherty et al. (36).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
DTNB analysis of free thiols on wild type and mutant Tpx
View larger version (36K):
[in a new window]
Fig. 1.
HPLC separation of tryptic peptides from
either reduced or oxidized pyridylethylated Tpx and tryptic digest
sites. Exhaustive digests of reduced (A and
C) or oxidized (B and D) Tpx (5 nmol)
that was pyridylethylated during denaturation prior to incubation with
trypsin were fractionated on an AquaPore RP-300 C8 column. Gradient
conditions are given under "Experimental Procedures," and peptides
were detected at 215 nm (A and B) and 254 nm
(C and D). The closed
arrows in each map indicate the following peaks: a
pyridylethylated fragment containing Cys82 (74 min)
(A and C) and disulfide-containing peaks P1 (68 min) and P2 (70 min) (B and D). The
open arrows indicate fragments resulting from
reduction of P1 or P2. E, a portion of the sequence of Tpx
from E. coli (residues 45-113, accession number AAC74406)
is shown with the tryptic sites (underlined) flanking the
three Cys (boldface type) numbered from the initiating Met.
The two Cys61 fragments resulting from the flanking tryptic
sites are highlighted with different lines for P1
(dotted) and P2 (solid), and the total ESI-MS
mass of each disulfide-containing peak (including the
Cys95-containing fragment indicated by the
dashed line) is given above the site
of initial hydrolysis by trypsin.
Steady-state kinetic parameters of Tpx
where ROH is the corresponding alcohol. These redox reactions are
analogous to those observed for other peroxidases from H. pylori and Crithidia fasciculata (35, 40) that are
recycled by Trx or a Trx homologue, but unlike those systems that
display an infinite Km for Trx, Tpx interacts with
Trx1 in a saturable manner with a Km of 22.5-25.5
µM (Table II).
View larger version (19K):
[in a new window]
Fig. 2.
Steady-state kinetic analysis of Tpx with
various concentrations of CHP. All three protein components (TrxR
(1.5 µM), Trx1 (10 µM), and Tpx (1 µM)) were incubated in 150 µM NADPH for 5 min in one syringe and then were mixed with varying amounts of CHP (0 µM (closed circles), 100 µM
(open triangles), 200 µM (closed
squares), and 400 µM (open diamonds)) in
another syringe (final concentrations after mixing; see "Experimental
Procedures"). Reaction progress was monitored at 340 nm on a stopped
flow spectrophotometer at 25 °C, and rates were extrapolated from
the linear portion of the curve (0-2 s) using linear regression
analysis. At 100 µM CHP, Tpx was not inactivated and gave
linear absorbance changes for the duration of the reaction and full
peroxide consumption (open triangles). At higher CHP
concentrations, Tpx activity diminished rapidly and nonlinearly without
complete consumption of CHP or NADPH (closed squares and
open diamonds). Stopped-flow data were collected every 50 ms, but only data from every 1.5 s are represented by the
symbols.
View larger version (14K):
[in a new window]
Fig. 3.
Scheme of TrxR/Trx1 reduction and electron
transfer pathways for Tpx during catalysis and inactivation.
During Tpx turnover, a small proportion of Tpx becomes overoxidized and
is removed from the reaction cycle (path A). The remaining
Tpx reforms the redox-active disulfide to continue the catalytic cycle
(path B). Eventually, after multiple turnovers in the
presence of 150 µM CHP, the enzyme primarily converts
to the R-SO2H species, few disulfide-containing species
remain, and activity declines, leading to irreversible inactivation.
This scheme depicts the overall flow of electrons but not necessarily
the precise redox forms of TrxR involved in turnover.
1
s
1. These studies verify that while Tpx is sensitive to
overoxidation by fatty acid hydroperoxides (and other soluble alkyl
hydroperoxides such as CHP), significant activity at low concentrations
of these substrates is achieved with rates comparable with those
obtained for other soluble peroxides. Similar trends of inactivation
with increasing concentrations of fatty acid hydroperoxide were
obtained during studies of human PrxII with linoleic acid hydroperoxide (42), although the peroxide levels required for overoxidation in this
case were much higher (250 µM) than those required for Tpx inactivation.
View larger version (14K):
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Fig. 4.
Steady-state assay of Tpx as a function of
15-HPETE concentration. Reaction mixtures in one syringe
containing NADPH (150 µM), Trx1 (10 µM),
TrxR (1.5 µM), and Tpx (1 µM) were mixed
with varying amounts of 15-HPETE (0-30 µM) in another
syringe on the stopped-flow spectrophotometer at 25 °C (final
concentrations after mixing). Each rate is the average of three
experiments and was obtained by linear regression analysis of the
linear portion of the reaction progression (0-1 s) prior to
inactivation. Standard peroxidase buffer was used in all cases as
described under "Experimental Procedures."
View larger version (20K):
[in a new window]
Fig. 5.
Steady-state kinetic analysis of wild type
and mutant Tpx proteins. Tpx reaction mixtures in one syringe
containing NADPH (150 µM), Trx1 (10 µM),
TrxR (1.5 µM), and varying amounts (0-1
µM) of Tpx (closed squares), C82S (open
circles), C95S (closed triangles), C82S,C95S
(open triangles), or C61S (closed circles) were
mixed with CHP (50 µM) on the stopped-flow
spectrophotometer in standard peroxidase buffer (final concentrations
after mixing; see "Experimental Procedures"). Each rate is the
average of three experiments conducted at 25 °C and was obtained by
linear regression analysis of the linear portion of the reaction (0-1
s) prior to inactivation.
) upon reaction with
peroxide. C61S with or without peroxide pretreatment exhibits only the
420-nm peak after reaction with NBD chloride, confirming the lack of
SOH formation on Cys82 or Cys95 (data not
shown).
View larger version (15K):
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Fig. 6.
Spectrophotometric analysis of NBD-labeled
Tpx mutants. Prereduced C82S,C95S treated with 1 eq of cumene
hydroperoxide (solid line) or no peroxide (dashed
line) was modified with NBD chloride (10×) for 5 min. To remove
excess reagent, treated samples were washed with 5 ml of buffer by
ultrafiltration, and then labeled proteins were analyzed from 200 to
600 nm.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1 s
1 (Table II). This rate is
about 10-fold faster than rates of ~105
M
1 s
1 achieved generally by
other Prx systems (13) and much faster than the weak peroxidase
activity displayed by BCP (~104
M
1 s
1, calculated from apparent
maximal velocity (Vmax) and
Km values obtained under non-steady-state conditions
for linoleic acid hydroperoxide (8)). On the basis of catalytic
efficiency, Tpx is the most potent reductant of alkyl hydroperoxides in
E. coli when compared with the other two Prx family members,
AhpC and BCP. Whereas some Prxs interact with Trx in a nonsaturable manner, it is unknown if this type of activity arises as a result of
differential affinity for Trx binding or because the
Km for the reducing substrate is higher than the
concentrations tested.
8 to 10
6
M) (7), and the role of AhpC in organic hydroperoxide
detoxification was questioned because bacteria do not synthesize the
polyunsaturated fatty acids required for lipid peroxidation. However,
the ability of Tpx and other Prxs, including yeast type II thioredoxin
peroxidase (50), to preferentially decompose organic hydroperoxides
over H2O2 suggests that Tpx's central role
in vivo involves the reduction of complex ROOH, whereas AhpC
mainly reduces H2O2. Several lines of evidence
also highlight the importance of Prxs in bacterial ROOH resistance,
including the isolation of mutants with increased resistance to organic
solvents linked directly to a mutation in E. coli AhpC (51).
Other bacterial peroxidases, such as Ohr from Xanthomonas
campestris, are specifically up-regulated by organic
hydroperoxides (52, 53), implying that defense against these peroxides
is requisite for bacterial survival. It is also quite possible that
during pathogenesis, bacteria take up polyunsaturated fatty acids from
the host (54), creating the potential for bacterial lipid hydroperoxide
formation. Whereas the exact nature and concentration of organic
hydroperoxides that Prxs are exposed to in vivo is unknown,
the existence of Prxs with selectivity for these peroxides suggests the
possibility of more complex peroxides as Prx substrates.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Daniel Ritz for the generous gift of E. coli Trx2, Dr. Joe O'Flaherty for assistance with 15-HPETE production, Dr. C. M. Reynolds for the cloning and initial purification of wild type E. coli Tpx, and Mark Morris for HPLC assistance and peptide analysis. We are particularly grateful to Mike Samuel at Wake Forest University Medical Center and Dr. Rodney Baker at the University of Mississippi School of Medicine for assistance with mass spectrometry analyses.
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FOOTNOTES |
---|
* Project support was provided by National Institutes of Health Grant R01 GM50389 and by an Established Investigatorship from the American Heart Association (to L. B. P).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Dept. of Biochemistry,
Wake Forest University School of Medicine, Medical Center Blvd.,
Winston-Salem, NC 27157. Tel.: 336-716-6711; Fax: 336-716-7671; E-mail:
lbpoole@wfubmc.edu.
Published, JBC Papers in Press, January 3, 2003, DOI 10.1074/jbc.M209888200
2 L. M. S. Baker and L. B. Poole, unpublished observations.
3 D. Ritz, personal communication.
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ABBREVIATIONS |
---|
The abbreviations used are: AhpC, alkyl hydroperoxide reductase peroxidase component; AhpF, alkyl hydroperoxide reductase flavoprotein oxidoreductase component; Trx, thioredoxin; BCP, bacterioferritin-comigratory protein; Tpx, thiol peroxidase; Prx, peroxiredoxin; Cys-SOH, cysteine sulfenic acid; TrxR, thioredoxin reductase; DTT, dithiothreitol; DTNB, 5,5'-dithiobis(2-nitrobenzoic acid); CHP, cumene hydroperoxide; NBD chloride, 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole; Cys-SO2H, cysteine sulfinic acid; Cys-SO3H, cysteine sulfonic acid; 15-HPETE, 15-hydroperoxyeicosatetraenoic acid; ESI-MS, electrospray ionization mass spectrometry; HPLC, high pressure liquid chromatography; MES, 2-(N-morpholino)-ethane sulfonic acid; TCEP, Tris[2- carboxyethyl]phosphine; Me2SO, dimethyl sulfoxide.
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