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Journal of Bacteriology, December 2001, p. 7053-7057, Vol. 183, No. 24
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.24.7053-7057.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
The PPP-Family Protein Phosphatases PrpA and PrpB
of Salmonella enterica Serovar Typhimurium Possess Distinct
Biochemical Properties
Liang
Shi,*
David G.
Kehres, and
Michael E.
Maguire
Department of Pharmacology, School of
Medicine, Case Western Reserve University, Cleveland, Ohio 44106
Received 10 August 2001/Accepted 21 September 2001
 |
ABSTRACT |
Salmonella enterica serovar Typhimurium requires
Mn2+, but only a few Mn2+-dependent enzymes
have been identified from bacteria. To characterize Mn2+-dependent enzymes from serovar Typhimurium, two
putative PPP-family protein phosphatase genes were cloned from serovar
Typhimurium and named prpA and prpB. Their
DNA-derived amino acid sequences showed 61% identity to the
corresponding Escherichia coli proteins and 41% identity
to each other. Each phosphatase was expressed in E. coli
and purified to near electrophoretic homogeneity. Both PrpA and PrpB
absolutely required a divalent metal for activity. As with other
phosphatases of this class, Mn2+ had the highest affinity
and stimulated the greatest activity. The apparent
Ka of PrpA for Mn2+ of 65 µM was
comparable to that for other bacterial phosphatases, but PrpB had a
much higher affinity for Mn2+ (1.3 µM). The pH optima
were pH 6.5 for PrpA and pH 8 for PrpB, while the optimal temperatures
were 45 to 55°C for PrpA and 30 to 37°C for PrpB. Each phosphatase
could hydrolyze phosphorylated serine, threonine, or tyrosine residues,
but their relative specific activities varied with the specific
substrate tested. These differences suggest that each phosphatase is
used by serovar Typhimurium under different growth or environmental
conditions such as temperature or acidity.
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INTRODUCTION |
MntH, the Salmonella
enterica serovar Typhimurium ortholog of the eukaryotic NRAMP
proteins (natural resistance-associated macrophage protein), is a
highly selective H+-stimulated Mn2+ transporter
(6). Subsequently, the mammalian NRAMP1 protein, expressed
in monocytes and macrophages, was shown to mediate marked changes in
intracellular Mn2+ in macrophages (5).
Transcription of mntH in serovar Typhimurium is induced by
hydrogen peroxide and cation starvation, and mutation of
mntH makes serovar Typhimurium cells more sensitive to
killing by peroxide. These data suggest that Mn2+ is
required for maintenance of some physiological functions and that
Mn2+ could play a role in virulence, especially in
resistance to reactive oxygen species (6). Any role for
Mn2+ is presumably mediated by its use as an enzyme
cofactor. Surprisingly, very few known enzymes are known to be
Mn2+ dependent. In an effort to understand the role of
Mn2+ in cell function and pathogenesis, we have begun to
characterize each of the few known Mn2+-cofactored enzymes.
One such family is the PPP class of protein phosphatases (2, 8,
17).
PPP phosphatases are metalloenzymes that include the catalytic subunits
of mammalian PP1, PP2A, PP2B, and related protein phosphatases. Two
divalent metal ions in their active sites catalyze a single-step
dephosphorylation reaction. In eukaryotes, PPP phosphatases are
involved in regulating multiple cellular functions such as cell
division, glycogen metabolism, and muscle contraction. Homologs of PPP
phosphatases can be discerned in many bacterial and archaeal genomes
(for reviews, see references 8 and 17). All bacterial PPP
phosphatases characterized to date require Mn2+ for maximal
activation (7, 10, 12-16, 18-20). To investigate the
role of Mn2+ further, we have cloned, purified, and
characterized the two PPP phosphatase homologs evident in the extant
genome of serovar Typhimurium. As predicted each is a
Mn2+-dependent protein phosphatase; however, the two
enzymes have markedly different properties suggestive of distinct
physiological roles.
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MATERIALS AND METHODS |
Standard procedures.
Protein concentrations were measured
with the bicinchoninic acid protein assay kit from Pierce (Rockford,
Ill.). Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
(SDS-PAGE) was performed as described by Laemmli (9), and
the gels were stained as described by Fairbanks et al.
(4).
Cloning and sequencing prpA and prpB from
serovar Typhimurium.
The prpA and prpB genes
were cloned from MM1255 (wild-type serovar Typhimurium LT2) by PCR.
Primers (Genosys, The Woodlands, Tex.) were based on available serovar
Typhimurium genomic sequence fragments
(http://genome.wustl.edu /bacterial.salmonella/shtml). The
upstream and downstream primers for prpA were
5'-GCTTACTAAGCAAATGCATCTGCACACGC-3' and
5'-CCTGCGCAGCAGGATCCCGATTTTTAACCC-3', respectively.
The upstream and downstream primers for prpB were
5'-GGCCTACTTTTTATGCATAATACCAGACTC-3' and
5'-GAAGGCTATATCGGATCCCATAATGCAGGG-3', respectively.
Restriction enzyme digestion of PCR products gave clones for each gene
that began with an NsiI site 30 to 40 bp 5' of the presumed
start codon and ended with a BamHI site 30 to 40 bp 3' of
the presumed stop codon. These were ligated into the NsiI
and BamHI sites of pDGK101 to give pDGK235 (containing
prpA) and pDGK236 (containing prpB). Each plasmid
contained 2,916 bp of pBluescriptII SK(+) from the BamHI
site through the SalI site in the polylinker (Stratagene, La
Jolla, Calif.) followed by 505 bp of the serovar Typhimurium corA promoter on a SalI-NsiI fragment,
followed by the prpA or prpB insert.
Inserts in pDGK235 and pDGK236 were sequenced using the above
oligonucleotides as primers at the Howard Hughes Medical Institute Biopolymer/Keck Foundation Biotechnology Resource Laboratory, Yale
University. Both sequences agree with the serovar Typhimurium LT2 sequence available at the above website. prpA and
prpB map to approximately 40.2 and 62.8 min on the serovar
Typhimurium chromosome, comparable to E. coli prpA and
prpB at 41 and 61.5 min, respectively (3).
Expression of ST-PrpA and ST-PrpB.
The open reading frames
for prpA and prpB were amplified by PCR with
Pfu DNA polymerase (Stratagene). The forward primers were
5'-TGGGGATCCATGAACGACAGGAAAAACATGATG-3' for prpA
and 5'-TGGGAATCCATGGAATTAATTCGTTATGCTFAC-3' for
prpB, while the reverse primers were
5'-TGGAAGCTTGCTATTGTATCCGCGCTAACG-3' for prpA and
5'-TGGAAGCTTTTACTTTATTTTAAAAAAAGACAGATTG-3' for
prpB. To facilitate cloning, a BamHI restriction
was introduced at the 5' end of the forward primers and a
HindIII restriction site was introduced at the 5' end of
the reverse primers. The resulting PCR products were ligated into the
pRSET-A expression vector (Invitrogen, Carlsbad, Calif.) and
transformed into E. coli DH5
cells (Gibco BRL, Rockville,
Md.). After plasmid purification, both strands of the insert were
sequenced to verify the fidelity of the PCR amplification. Competent
E. coli BL21(DE3) cells (Promega, Madison, Wis.) were
transformed with plasmid, grown to an optical density at 600 nm of 0.6 to 1, induced with isopropyl-
-D-thiogalactopyranoside (IPTG) at a final concentration of 0.1 mM for 4 h at 37°C, and harvested by centrifugation. The pellet was resuspended in 40 ml of
buffer A (20 mM Tris [pH 7], 500 mM NaCl, 1 mM phenylmethylsulfonyl fluoride [PMSF], 1 mM imidazole-HCl) with 2 mg of lysozyme per ml,
100 µg of DNase I per ml, and 100 µg of RNase per ml and incubated on ice for 30 min. The cells were lysed by being passed through a
French press three times at 8,000 lb/in2; lysates were
centrifuged at 17,000 × g at 4°C for 30 min. The supernatant was loaded on a 3-by 5-cm column of Ni-nitrilotriacetic acid agarose (Qiagen, Valencia, Calif.) charged with NiCl2
and equilibrated with buffer A. The column was washed with 100 ml of
buffer A and then with 100 ml of buffer B (20 mM Tris [pH 7], 500 mM
NaCl, 1 mM PMSF, 40 mM imidazole-HCl). The fusion proteins were then
eluted from the column with 150 ml of buffer C (20 mM Tris [pH 7],
500 mM NaCl, 1 mM PMSF, 80 mM imidazole-HCl), 10-ml fractions were
collected, and 10 µl of each fraction removed and assayed for
phosphatase activities, and peak fractions were pooled. Samples
containing ST-PrpA were dialyzed against 3 liters of buffer D (10 mM
Tris-HCl [pH 7], 1 mM dithiothreitol, 1 mM PMSF, 1 mM EDTA) plus 10 mM NaCl overnight, and loaded on a 3- by 5-cm column of Macro-Prep High
S Support (Bio-Rad, Hercules, Calif.) that had been equilibrated with
buffer D plus 10 mM NaCl. The column was washed with 100 ml of buffer D
plus 50 mM NaCl and eluted with a salt gradient of 50 to 400 mM NaCl in
buffer D. Fractions (5 ml) were collected and assayed for phosphatase
activity. Active fractions were pooled, dialyzed with 1 liter of buffer
E (50 mM Tris-HCl [pH 7], 1 mM dithiothreitol, 1 mM PMSF, 1 mM EDTA,
10% [vol/vol] glycerol) overnight, and stored at 
20°C.
ST-PrpB samples were concentrated with a Centriprep-10 column and
loaded on a 2- by 65-cm column of Superose 6 (Amersham Pharmacia
Biotech, Piscataway, N.J.) equilibrated with buffer F (10 mM ammonium
formate [pH 6], 1 mM PMSF, 1 mM EDTA). The column was eluted with
180 ml of buffer F; 1-ml fractions were collected and assayed
for
phosphatase activity. Peak fractions were pooled, concentrated
with a
Centriprep-10 column, resuspended with buffer E, and stored
at
20°C. All purification procedures were carried out at 4°C
except
gel filtration chromatography, which was performed at room
temperature.
Expression and purification of ST-PrpA and ST-PrpB
were monitored by
Western hybridization with anti-Xpress antibody
(Invitrogen, Carlsbad,
Calif.).
Preparation of phosphorylated MBP.
Serine/threonine- or
tyrosine-phosphorylated myelin basic protein (MBP) was prepared with
[
-33P]ATP as specified by the manufacturer (New
England BioLabs, Beverly, Mass.). Peptide substrates were purchased
from Promega.
Phosphatase assay.
Purified phosphatase protein (200 ng) was
incubated at 37°C in 30 µl of solution containing 50 mM imidazole
(pH 6.5 for ST-PrpA and pH 8 for ST-PrpB), 1 mM MnCl2, 10 µg of bovine serum albumin per ml, BSA, and 2 µM
[33P]phospho-MyBP. The initial rate of the reaction was
linear for at least 30 min for all substrates. The reaction was
terminated by the addition of 100 µl of 20% (wt/vol) trichloroacetic
acid, mixed, and centrifuged for 3 min in a microcentrifuge at maximum speed. 33P radioactivity was determined using 50 µl of
supernatant liquid. A modified molybdic acid extraction method
(7) was used to verify the reaction product as inorganic
phosphate. The peptide substrates were assayed as specified by the
manufacturer (Promega).
pNPP phosphatase activities were assayed at 37°C in 200 µl of
solution containing 50 mM imidazole (pH 6.5 for ST-PrpA and
pH 8 for
ST-PrpB) 1 mM MnCl
2, and 1 mM pNPP. ST-PrpA or ST-PrpB
was
added to start the reaction. After 30 min, the reaction was
stopped
with 800 µl of 0.5 M sodium borate (pH 9). Released
p-nitrophenol
was measured at 410 nm on a Beckman DU-64
spectrophotometer. Most
phosphatase assays were performed on proteins
carrying the His
6 tag and linking sequence. However,
removal of the fusion domains
of ST-PrpA and ST-PrpB with enterokinase
did not change their
affinity for Mn
2+ or other properties
tested. For pH optimum experiments, controls
indicated that the
composition of the buffer had no significant
influence on phosphatase
activity. Final concentrations of inhibitors
used were 10 mM EDTA, 10 mM sodium pyrophosphate, 50 mM NaF, 10
mM sodium potassium tartrate, 1 mM tetramisole, 1 mM sodium orthovanadate,
150 µM trifluoperazine, 10 µM microcystin-LR, 5 µM okadaic acid,
and 1 mM AppppA (diadenosine
tetraphosphate).
Nucleotide sequence accession numbers.
The prpA
and prpB sequences have been deposited with GenBank under
accession no. AY149950 and AY049951, respectively.
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RESULTS |
Cloning of prpA and prpB.
Genes for
two PPP family protein phosphatases identified in extant serovar
Typhimurium genomic sequence by a BLAST (1) search were
cloned by PCR amplification from serovar Typhimurium genomic DNA.
Derived amino acid sequences were each 61% identical to their
respective homologs in E. coli (Fig.
1); therefore, the serovar Typhimurium
genes were designated prpA and prpB. The
predicted protein product of prpA, ST-PrpA, consisted of 222 amino acids with a predicted molecular mass of 25,879 Da and an
isoelectric point of 8.8. ST-PrpB was predicted to have 218 amino
acids, a molecular mass of 25,024 Da, and a pI of 4.6. The serovar
Typhimurium proteins were each 61% identical to their respective
E. coli homologs and 41% identical to each other (Fig. 1).
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FIG. 1.
Sequence alignment of serovar Typhimurium and E. coli PPP phosphatases. The predicted amino acid sequences of
ST-PrpA, ST-PrpB, and the E. coli phosphatases were aligned
using CLUSTAL W and modified slightly by eye to optimize alignment at
the termini. Dashes indicate sequence gaps introduced to optimize the
alignment of conserved regions; asterisks indicate identical residues
in all four phosphatases; colons indicate either three of four residues
identical or all four residues conserved.
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Expression and purification ST-PrpA and ST-PrpB.
To facilitate
purification by metal-chelate affinity chromatography, an
amino-terminal His6 sequence from the pRSET-A vector was
introduced into the recombinant ST-PrpA and ST-PrpB carried in pRSRT-A
and expressed in E. coli after induction with IPTG. After
metal-chelate affinity chromatography, ST-PrpB was concentrated and
further purified by gel filtration chromatography. ST-PrpA, however,
became aggregated when concentrated. Each was further purified by
cation-exchange chromatography. Both ST-PrpA and ST-PrpB were purified
to near electrophoretic homogeneity (Fig.
2), and their identities were confirmed
by Western blot analysis with anti-Xpress antibody (data not shown).
The yield from 1 liter of cells was 1 mg of ST-PrpA or 3 mg of ST-PrpB.
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FIG. 2.
Analysis of purified ST-PrpA and ST-PrpB by SDS-PAGE.
ST-PrpA and ST-PrpB were expressed in E. coli and purified
by metal-chelate affinity chromatography and ion-exchange or gel
filtration chromatography as described in Materials and Methods.
Portions of 1 µg of ST-PrpA and 2 µg of ST-PrpB were analyzed by
SDS-PAGE on a 15% (wt/vol) acrylamide gel. Lanes: 1, ST-PrpA; 2, ST-PrpB. The positions of protein standard (Stds) are indicated at the
left.
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Properties of ST-PrpA and ST-PrpB.
Both ST-PrpA and ST-PrpB
showed divalent metal ion-dependent protein phosphatase activity toward
[33P]phosphoseryl MBP (Fig.
3). Although a few other cations
supported some phosphatase activity, Mn2+ clearly produced
the greatest stimulation. Surprisingly, ST-PrpA and ST-PrpB
displayed different affinities for Mn2+ (Fig. 3A). The
Mn2+ Ka of ST-PrpB was 1.3 µM,
over 50 times lower than the Mn2+ Ka
of ST-PrpA at 65 µM (Table 1).
Ni2+, Co2+, and Fe2+ could support
the phosphatase activity of ST-PrpB (Fig. 3C) but with
Ka values in the mM range (Fig. 3A; Table 1).
Ni2+ was also a weak activator of PrpA activity. Neither
Mg2+ nor Ca2+ had stimulatory effects on either
phosphatase, while Cd2+, Cu2+, and
Zn2+ inhibited both phosphatases when added concomitantly
with Mn2+. Fe2+ and Co2+ slightly
inhibited Mn2+-stimulated ST-PrpA protein phosphatase
activity (Fig. 3B).
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FIG. 3.
Effects of divalent metal ions on the catalytic activity
of ST-PrpA and ST-PrpB. (A) The activity of 200 ng each of purified
ST-PrpA and ST-PrpB was assayed under standard conditions (see
Materials and Methods) using [33P]phosphoseryl MBP as
substrate with the indicated concentrations of Mn2+ or
Ni2+. Results are given as the percent activity observed
with 1 mM of Mn2+ and the standard error of the mean
(n = 4). For points with no error bar, the error was
smaller than the size of the symbol. For ST-PrpA, 100% activity was
equal to 0.0302 pmol of 33Pi released per min,
while the corresponding value for ST-PrpB was 0.0304 pmol/min. (B and
C) The activity of purified ST-PrpA (B) and ST-PrpB (C) was assayed
under the conditions described for panel A, except that, where
indicated, the compounds listed were substituted for the activating
divalent metal ion, Mn2+. All compounds were present at a
final concentration of 1 mM. Iron was maintained in the ferrous state
by the addition of ascorbic acid to a final concentration of 1 mM. All
results are reported as the percentage of activity relative to that
observed with Mn2+ and standard error of the mean
(n = 4).
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Both ST-PrpA and ST-PrpB were active toward pNPP over a pH spectrum
from 5 to 10. However, optimal ST-PrpA activity occurred
at pH 6 to 7, while the optimal pH for ST-PrpB activity was pH
7.5 to 8.5 (Fig.
4A). Each was active over a temperature
range
from 0 to 65°C. Maximal activity was seen between 45 and 55°C
for ST-PrpA but 30 to 37°C for ST-PrpB. ST-PrpA but not ST-PrpB
was
heat stable, with PrpA maintaining substantial activity at
65°C (Fig.
4B). ST-PrpA and ST-PrpB demonstrated a similar specific
activity
toward [
33P]-phosphoseryl MBP, but the activity of
ST-PrpA toward [
33P]-phosphotyrosyl MBP was less than
10% that of ST-PrpB. In contrast,
activity toward synthetic
phosphopeptide substrates, whether phosphotyrosyl
or phosphothreonyl,
was much greater with ST-PrpA than with ST-PrpB
(Table
2).
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FIG. 4.
pH and temperature dependence of ST-PrpA and ST-PrpB.
(A) The activity of 2 µg of purified ST-PrpA and ST-PrpB was assayed
for phosphatase activity against pNPP under standard conditions (see
Materials and Methods), except that the pH was varied as indicated
using different buffers. Shown is the relative protein phosphatase
activity detected as a function of pH and the standard error of the
mean (n = 3). 100% activity was defined as 0.84 nmol
of p-nitrophenol released per min for ST-PrpA at pH 6.5 and
0.92 nmol/min for ST-PrpB at pH 8. Buffer salts (50 mM) were sodium
acetate (pH 4 to 5), imidazole (pH 5 to 8), Tris-HCl (pH 8 to 9), and
glycine (pH 9 to 11). (B) The activity of purified ST-PrpA and ST-PrpB
was assayed for pNPP phosphatase activity under standard conditions
(see Materials and Methods) while varying the temperature. All results
are reported as the percent activity relative to that observed at
37°C and the standard error of the mean (n = 3). For
points with no error bar, the error was smaller than the size of the
symbol.
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The effect of protein phosphatase and other phosphomonoesterase
inhibitors (
11,
14,
16) on the activity of ST-PrpA and
ST-PrpB was also examined (data not shown). At the concentrations
tested, pyrophosphate, the chelating agent EDTA, the nonspecific
phosphatase inhibitor NaF, and the tyrosine phosphatase inhibitor
vanadate inhibited >90% of activity from either enzyme with
[
33P]-phosphoseryl MBP as substrate. Diadenosine
tetraphosphate (AppppA)
inhibited the activity of both enzymes by 20 to
30%, but the acid
phosphatase inhibitor tartrate, the alkaline
phosphatase inhibitor
tetramisole, the protein phosphatase 2B inhibitor
trifluoperazine,
and the protein phosphatase 1 and 2A inhibitors
microcystin-LR
and okadaic acid did not inhibit ST-PrpA and
ST-PrpB
activity.
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DISCUSSION |
Little is known about the physiological functions of
Mn2+. Mn2+ is often a functional substitute for
Mg2+. Intuitively, one might thus expect to find a fairly
large number of Mn2+-dependent enzymes. However, only a
handful of Mn2+-dependent metalloenzymes are known. Because
of our interest in Mn2+ transport and its possible role in
pathogenesis (6), we are characterizing several of these
Mn2+-dependent enzymes, including the two
Mn2+-dependent protein phosphatases of serovar Typhimurium
described in this report.
prpA and prpB of serovar Typhimurium are clear
homologs of the corresponding genes in E. coli
(13), but their sequences diverge much more than many
homologs between such closely related organisms, at only 61% identity
between species for both PrpA and PrpB. In contrast, the percent
identity between serovar Typhimurium and E. coli for other
Mn2+-dependent enzymes is 91 to 97% for SodA
(Mn2+-superoxide dismutase), GpmM (phosphoglyceromutase),
SpoT (AppppA synthase/hydrolase), and PepP (Aminopeptidase P). This
could be indicative of a different origin for the phosphatases compared to other Mn2+-dependent enzymes.
The Ka of ST-PrpA for Mn2+ (65 µM)
is within a factor of 3 of the Ka for
Mn2+ for the homologous phosphatases PP1-arch1 from the
archaeon Sulfolobus solfataricus and PP1-cyano1 and
PP1-cyano2 from Microcystis sp. (7, 12, 16). In
contrast, the Mn2+ Ka of 1.3 µM
for ST-PrpB is much lower than that for other prokaryotic PPP
phosphatases previously characterized. Although Ni2+,
Co2+, and Fe2+ can support some phosphatase
activity, their relative Ka values compared to
that of Mn2+ indicate that Mn2+ is the
physiologically relevant cation.
PrpA and PrpB of E. coli have also been purified and
partially characterized (13). Unfortunately, divalent
cation selectivity cannot be compared since this was not determined for
the E. coli enzymes. The effects of several phosphatase
inhibitors on ST-PrpA and ST-PrpB were generally similar not only to
each other but also to the effects of the same inhibitors on the
homologous phosphatases PP1-cyano1 from Microcystis
aeruginosa PCC 7820 and PP1-cyano2 from M. aeruginosa
UTEX 2063 (16). In contrast, although fluoride ion could
completely block phosphatase activity in the serovar Typhimurium and
M. aeruginosa enzymes, it had little effect on the two
E. coli phosphatases. Vanadate inhibited both phosphatases from serovar Typhimurium but only PrpA from E. coli
(13). Finally, substrate specificity may differ between
serovar Typhimurium and E. coli. In both species, PrpA and
PrpB have similar activity toward phosphoseryl MBP, but the activities
of the E. coli phosphatases toward phosphotyrosyl MBP were
equivalent whereas ST-PrpB had 10-fold greater activity than ST-PrpA
toward phosphotyrosyl MBP. In contrast, ST-PrpA had high activity
against phosphotyrosyl peptide substrates. Therefore, until
physiological substrates are determined, substrate specificity cannot
be determined. Our preliminary data (L. Shi and M. E. Maguire,
unpublished results) indicate that mutation of prpA or
prpB markedly alters the peroxide and temperature
sensitivity. Proteins whose expression is altered with a change in
temperature or exposure to reactive oxygen species might therefore be
candidates for substrates for these phosphatases.
Not all bacterial genomes have PPP-class phosphatases. For those that
have PPP phosphatases, almost all have only a single gene
(17). Serovar Typhimurium and E. coli are the
only bacteria so far studied to have two PPP phosphatase genes. The
E. coli phosphatases are also the only bacterial PPP
phosphatases for which a function can be assigned. They regulate the
transcription of htrA via the CpxR-CpxA two-component signal
transduction pathway that is responsive to protein misfolding in the
periplasm (13). htrA encodes a protease that
degrades misfolded proteins, thus avoiding potential cell toxicity. The
similarities between the PrpA phosphatase in E. coli and
serovar Typhimurium suggest that they probably have a similar function
in serovar Typhimurium. In contrast, the rather different properties of
the PrpB enzymes from the two species suggest the possibility of
overlapping but not identical physiological roles.
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FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Pharmacology, School of Medicine, Case Western Reserve University,
10900 Euclid Ave., Cleveland, OH 44106-4965. Phone: (216) 368-6187. Fax: (216) 368-3395. E-mail: lxs76{at}po.cwru.edu.
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Journal of Bacteriology, December 2001, p. 7053-7057, Vol. 183, No. 24
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.24.7053-7057.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
