From the Pharmacologie et Physico-Chimie des
Interactions Cellulaires et Moléculaires, UMR CNRS 7034, Université Louis Pasteur de Strasbourg, Faculté de
Pharmacie, 74 route du Rhin, B.P. 24, F-67401 Illkirch, France and
¶ Laboratoire de Spectrométrie de Masse Bio-Organique, UMR
CNRS 7509, ECPM, Université Louis Pasteur de Strasbourg, 25 rue
Becquerel, F-67087 Strasbourg, France
Received for publication, September 4, 2002
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ABSTRACT |
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The bifunctional allosteric enzyme HPr
kinase/phosphatase (HPrK/P) from Bacillus subtilis is a key
enzyme in the main mechanism of carbon catabolite repression/activation
(i.e. a means for the bacteria to adapt rapidly
to environmental changes in carbon sources). In this regulation system,
the enzyme can phosphorylate and dephosphorylate two proteins,
HPr/HPr(Ser(P)) and Crh/Crh(Ser(P)), sensing the metabolic state
of the cell. To acquire further insight into the properties of HPrK/P,
electrospray ionization mass spectrometry, dynamic light scattering,
and BIACORE were used to determine the oligomeric state of the protein
under native conditions, revealing that the enzyme exists as a hexamer
at pH 6.8 and as a monomer and dimer at pH 9.5. Using an in
vitro radioactive assay, the influence of divalent cations, pH,
temperature, and different glycolytic intermediates on the activity as
well as kinetic parameters were investigated. The presence of divalent
cations was found to be essential for both opposing activities of the
enzyme. Furthermore, pH values equal to the internal pH of vegetative
cells seem to favor the kinase activity, whereas lower pH values
increased the phosphatase activity. Among the glycolytic intermediates
evaluated, fructose 1,6-diphosphate and fructose 2,6-diphosphate were
found to be allosteric activators in the kinase assay, whereas high concentrations inhibited the phosphatase activity, except for fructose
1,6-diphosphate in the case of HPr(Ser(P)). Phosphatase activity was
induced by inorganic phosphate as well as acetyl phosphate and
glyceraldehyde 3-phosphate. Kinetic parameters indicate a preference
for binding of HPr compared with Crh to the enzyme and supported a
strong positive cooperativity. This work suggests that the oligomeric
state of the enzyme is influenced by several effectors and is
correlated to the kinase or phosphatase activity. The phosphatase
activity is mainly supported by the hexameric form.
Protein kinases and phosphatases play a decisive role in many
biological processes by phosphorylating and dephosphorylating target
proteins. For humans, it was estimated in the mid-1990s that the genome
may contain as many as 2000 protein kinases and 1000 protein
phosphatases (1). Initial analysis, after decoding of the DNA that
constitutes the human genome, indicates a somewhat lower number of
predicted protein kinases and protein phosphatases (2, 3). Since
protein kinases are not only involved in normal cell growth but also in
malignant transformations, these enzymes have been in focus during
recent years as new drug targets (4-6). Protein kinases and
phosphatases also play a fundamental role in modulating signals in
cellular processes in prokaryotic cells. In the Bacillus
subtilis (B. subtilis strain 168) genome
sequencing project (7) 4106 protein genes were identified encoding 52 known and 46 putative kinases and 26 known and 10 putative phosphatases (on the World Wide Web, see
genolist.pasteur.fr/SubtiList/genome.cgi).
The bifunctional HPr kinase/phosphatase
(HPrK/P)1 in the low guanine
and cytosine (low GC) Gram-positive bacteria, B. subtilis, is involved in the main regulatory mechanism for
carbon catabolite repression/activation (CCR/CCA) (9-12). The enzyme
possesses kinase activity in the presence of ATP and a favorable carbon
source, such as glucose, which generates high concentrations of
glycolytic intermediates (e.g. fructose
1,6-diphosphate (FBP)), and can phosphorylate two protein substrates
(i.e. HPr (histidine-containing protein) and Crh
(catabolite repression HPr)) on Ser-46 (10, 11, 13, 14). The presence
of two substrates for HPrK/P seems to be unique among low GC
Gram-positive bacteria, since Crh has only been detected, thus far, in
species of Bacillus (15). High concentrations of inorganic
phosphate and low concentrations of ATP, reflecting the intracellular
state of cells at starvation (16), have been shown to trigger the
phosphatase activity of the enzyme HPrK/P of B. subtilis and, thus, dephosphorylation of HPr(Ser(P)) and Crh(Ser(P)) (9). Furthermore, it has been demonstrated that HPrK/P is
an allosteric homo-oligomeric enzyme yielding sigmoidal velocity curves
and is strongly regulated by allosteric effector molecules such as FBP
(17). In addition to being involved in the CCR/CCA regulatory
mechanism, HPr is also a part of the bacterial phosphoenolpyruvate-sugar phosphotransferase system (PTS) and becomes
phosphorylated on His-15 by enzyme I (EI) (18). The phosphoryl group
can then be transferred to the sugar-specific enzymes II, which
phosphorylate incoming PTS sugars (18). The other protein, Crh, which
was discovered within the B. subtilis genome
sequencing project (7), has a Gln instead of a His at position 15, and
in vitro phosphorylation by phosphoenolpyruvate and EI could
not be demonstrated (14).
When HPr and Crh are phosphorylated at Ser-46, an interaction has been
demonstrated under in vitro conditions with the
trans-acting protein CcpA (carbon catabolite control protein
A) (13, 19). This complex has been shown to recognize a
cis-acting palindromic sequence called cres
(catabolite response elements), leading to repression or activation of
target gene or operon transcription. There is evidence that CcpA and
cre meditate CCR of many catabolic genes or operons such as
acsA (acetyl coenzyme A synthetase) (20), xyl
(encoding enzymes for xylose utilization) (21, 22), gnt (encoding enzymes for gluconate catabolism) (23, 24), hut (encoding enzymes involved in histidine catabolism) (25, 26), and
lev (encoding a fructose-specific PTS and a levanase) (27). CCA involving CcpA as a positively acting regulator has been
demonstrated for genes involved in the overflow metabolism for
excreting excess glucose, such as ackA encoding acetate
kinase (12, 28), pta encoding phosphotransacetylase (29),
and alsS encoding acetolactate synthase (30). Both CCR and
CCA mediated by CcpA appear to be a general mechanism for Gram-positive
bacteria with low GC content (31, 32).
The aim of this study was to further characterize the bifunctional
enzyme HPrK/P from B. subtilis and to propose a
model for the regulation of the kinase/phosphatase "switch" induced
by the metabolic state of the cell.
Purification of HPr, Crh, and HPrK/P from B. subtilis
In order to purify HPr(His)6, Escherichia
coli strain M15 containing plasmid pREP4 (Qiagen, Courtaboeuf,
France) was transformed with plasmid pAG2 (14). For
Crh(His)6, E. coli strain BL21(DE3) containing plasmid pLysS (Novagen, Madison, WI) was transformed with plasmid pAG1 (14). For the purification of HPrK/P, E. coli strain BL21(DE3) was transformed with plasmid pET32LIC
(Novagen, Madison, WI) encoding HPrK/P with a thioredoxin-, His-,
and S-tag (Trx-His6-S-tag). Transformants were grown
overnight at 37 °C in Bouillon Trypto-caseine soja medium (Bio-Rad)
with 50 µg/ml ampicillin for HPrK/P, 50 µg/ml ampicillin and 25 µg/ml kanamycin for HPr, and 50 µg/ml ampicillin and 170 µg/ml chloramphenicol for Crh. The following morning, the cultures
were transformed to 1 liter of medium containing the desired antibiotic
drugs and were grown until A600 was 0.6. Expression of the genes was induced by adding isopropyl
The sequences of the proteins are as follows: HPr(His)6;
MRGSHHHHHH GSMAQKTFKV TADSGIHARP ATVLVQTASK YDADVNLEYN GKTVNLKSIM GVMSLGIAKG AEITISASGA DENDALNALE ETMKSERLGE; Crh(His)6,
MVQQKVEVRL KTGLQARPAA LFVQEANRFT SDVFLEKDGK KVNAKSIMGL MSLAVSTGTE
VTLIAQGEDE QEALEKLAAY VQEEVLQHHHHHH; and
HPrK/P(Trx-His6-S-tag), MSDKIIHLTD DSFDTDVLKA DGAILVDFWA
EWCGPCKMIA PILDEIADEY QGKLTVAKLN IDQNPGTAPK YGIRGIPTLL LFKNGEVAAT
KVGALSKGQL KEFLDANLAG SGSGHMHHHH HHSSGLVPRG SGMKETAAAK FERQHMDSPD
LGTDDDDKMG NVRTKDVMEQ FNLELISGEE GINRPITMSD LSRPGIEIAG YFTYYPRERV
QLLGKTELSF FEQLPEEEKK QRMDSLCTDV TPAIILSRDM PIPQELIDAS EKNGVPVLRS
PLKTTRLSSR LTNFLESRLA PTTAIHGVLV DIYGVGVLIT GKSGVGKSET ALELVKRGHRL
VADDCVEIRQ EDQDTLVGNA PELIEHLLEI RGLGIINVMT LFGAGAVRSN KRITIVMNLE
LWEQGKQYDR LGLEEETMKI IDTEITKLTI PVRPGRNLAV IIEVAAMNFR LKRMGLNAAE
QFTNKLADVI EDREQEE. The concentration of HPrK/P(Trx-His6-S-tag) was determined
spectrophotometrically using the Bio-Rad protein assay (Bio-Rad) with
Bio-Rad protein assay standard I lyophilized bovine plasma Electrospray Ionization Mass Spectrometry (ESI-MS)
Measurements
Further Treatment of the Proteins--
For ESI-MS analysis of
HPrK/P(Trx-His6-S-tag), further purification was performed
using a 1-ml HiTrap Q Sepharose high performance column (Amersham
Biosciences). The HiTrap Q column was equilibrated with 5 column
volumes of 25 mM Tris buffer (pH 8) followed by 5 column
volumes of 25 mM Tris containing 1 M KCl (pH 8)
and finally 10 column volumes of 25 mM Tris buffer (pH 8).
The protein solution was applied to the column, and fractions were
eluted with 25 mM Tris buffer (pH 8), containing increasing
concentrations of KCl (100, 300, 600, and 1000 mM). The
purity of the fractions, after concentration using an Ultrafree
centrifugal filter unit with a molecular mass cut-off of 10,000 Da
(Millipore), was confirmed with SDS-PAGE separation using PhastGel
(Amersham Biosciences) and Coomassie staining. The pure fractions were
then desalted through a PD-10 column (Amersham Biosciences) as
described above.
Prior to mass spectrometry analysis, an additional desalting procedure
was performed with Centricon (Millipore Corp.) using 10 mM
ammonium acetate (pH 6.8) as reconstitution solution. Ammonium acetate
was used, since this buffer preserves the native structure of proteins
and is compatible with the ESI-MS analysis. Concentrators with a
molecular mass cut-off of 10,000 Da were used for
HPrK/P(Trx-His6-S-tag), and devices with a molecular mass
cut-off of 3000 Da were used for HPr(His)6 and
Crh(His)6. The procedure for desalting was conducted following the protocol provided by the company and contained six dilution/concentration steps performed at 4 °C for 60 min. Finally, the concentration of the proteins was determined spectrophotometrically using the Bio-Rad protein assay. The samples were stored at Instrumentation--
Mass spectrometry experiments were
conducted using a hybrid quadrupole time-of-flight mass spectrometer
(Q-TOF II; Micromass, Altrincham, UK) equipped with a Z-Spray ESI
source. All spectra were recorded in the positive ion mode.
ESI-MS Analysis under Denaturing Conditions--
Purity and
homogeneity of HPr(His)6, Crh(His)6, and
HPrK/P(Trx-His6-S-tag) were verified by mass analysis under
denaturing conditions with the proteins diluted to 10 µM
in a 1:1 water/acetonitrile (v/v) mixture acidified with 1% formic
acid. Under these conditions, noncovalent interactions between subunits
are disrupted, thus allowing for the accurate measurement of the
molecular weight of the constitutive enzyme subunits with a precision
of better than 0.01%. Mass spectra were recorded in the positive ion
mode in the mass range of 500-4000 m/z (mass to
charge) after calibration of the instrument with horse heart myoglobin
diluted to 2 µM in a 1:1 water/acetonitrile (v/v) mixture
acidified with 1% formic acid.
ESI-MS Analysis under Native Conditions--
Under native
conditions, the mass measurements of HPr(His)6,
Crh(His)6, and HPrK/P(Trx-His6-S-tag) were
performed using 10 mM ammonium acetate buffer (pH 6.8).
Samples were diluted to ~20 µM and continuously infused
into the ESI ion source at a flow rate of 5 µl/min. To study the
influence of pH variations on the stability of the oligomeric state of
the enzyme, increasing amounts of NH3 were added, up to pH
9.5, to the 10 mM ammonium acetate buffer.
HPrK/P(Trx-His6-S-tag) was then diluted to 20 µM with the pH-adjusted buffer. All of the measurements
were performed at an accelerating voltage equal to 200 V and with the
pressure in the interface region of the mass spectrometer equal to 6.5 millibars. Mass data were acquired in the positive ion mode in the
2500-12,000 m/z mass range. Clusters of CsI
(separate injections of 1 mg/ml CsI in 50% aqueous isopropyl alcohol)
were used for the calibration of an extended mass range in the high
m/z region. The molecular weight of the
noncovalent complex of the enzyme was calculated as the mean of five
peak maximum values ± S.D.
Dynamic Light Scattering Measurements
Dynamic light scattering data were obtained with the DynaPro-801
instrument (Protein Solutions Inc.) using a 30-milliwatt, 833-nm
wavelength argon laser at 20 °C and equipped with a solid-state avalanche photodiode. During the illumination, the photons scattered by
proteins were collected at 90 °C on a 10-s acquisition time and were
fit with the analysis software, Dynamics. Intensity fluctuations of the
scattered light resulting from Brownian motion of particles were
analyzed with an autocorrelator to fit an exponential decay function
and then measuring a translational diffusion coefficient D. For
polydisperse particles, the autocorrelation function was fit as the sum
of contributions from the various size particles using the
regularization analysis algorithm. D is converted to a hydrodynamic
radius Rh through the Stokes-Einstein equation (Rh = kbT/6 BIAcore Surface Plasmon Resonance Analysis
HPrK/P(Trx-His6-S-tag) oligomerization was examined on a
BIAcore instrument (BIAcore J; Amersham Biosciences). The
principle of this technology relies on a surface plasmon resonance
phenomenon that transforms the specific incident angle of the light
reflected from a metallic surface in response to the solute bound to
the surface. The Biacore measurements are arbitrary expressed in
resonance units that are proportional to the concentration of solute
bound to the surface. The monitoring of the surface plasmon resonance response allows analysis such as kinetics in real time and
stochiometries for the formation of multimolecular complexes. To
determine the effect of pH on the quaternary structure of
HPrK/P(Trx-His6-S-tag), the protein was diluted in a solution
of 40 mM Tris-HCl buffered either at pH 6.8 or at pH 9.5 and loaded at a flow rate of 35 µl/min on an NTA sensor chip in
conditions of low density of nickel (50 resonance units) and saturation
of protein over the nickel (200 nM). Typical experiments
were carried out in the same conditions, at 25 °C, with running
buffer 10 mM HEPES, 0.15 M NaCl, 50 µM EDTA, 0.005% surfactant P20, pH 7.4, and with a
constant flow rate of 5 µl/min. Alternatively, to evaluate the
dissociation constants, a running buffer of 10 mM Tris-HCl,
50 µM EDTA, 0.005% surfactant P20, pH 6.8 or 9.5, was
used. To correct for nonspecific binding to dextran matrix, experiments
were performed with two channels of the sensor chip simultaneously, one
not being regenerated with nickel.
Chemicals
Pyruvic acid, D-fructose 2,6-diphosphate (F2,6BP),
D-fructose 1,6-diphosphate (FBP), acetyl coenzyme A,
Radioactive Kinase and Phosphatase Assay
The assay mixture for in vitro phosphorylation of
HPr(His)6/Crh(His)6 contained 50 mM
Tris buffer (pH 8), 5 mM MgCl2, 0.5 mM ATP, 3.3-5 Bq (leading to 200-300 cpm of
[ For the phosphatase assay, HPr(His)6/Crh(His)6
was first phosphorylated in a kinase assay. The final volume was 20 µl and contained 50 mM Tris buffer (pH 8), 7 mM MgCl2, 2 mM ATP, 3.3-5 Bq
(leading to 200-300 cpm of [ Divalent Cations for HPrK/P Activity--
Different
divalent cations, Mg2+, Mn2+, Co2+,
Ca2+, and Cu2+ as chloride salts, were tested
at a final concentration of 5 mM. To exclude possible
influence from the buffer, 50 mM HEPES (pH 8) was used, since this buffer is known to form only weak complexes with
Mg2+, Mn2+, Ca2+, and
Cu2+ (33, 34).
Effects of pH--
The activity of the enzyme was determined at
11 different pH values, ranging from 5.0 to 8.7, for the different
substrates, HPr/HPr(Ser(P)) and Crh/Crh(Ser(P)). The buffer used, to
cover the pH range, was a mixture of 0.2 M citric acid and
0.2 M Tris (final concentration of 50 mM). The
pH was adjusted at 37 °C. For the kinase stability study, the enzyme
was incubated for 10 min at 37 °C at the different pH values
indicated above in 50 mM citric acid/Tris. After
incubation, the residual activity was measured at pH 8.0 for HPr and pH
9.0 for Crh (final concentration of 250 mM Tris) at
37 °C for 10 min. In the phosphatase stability study, the procedure
was the same except that the activity of the enzyme was measured at pH
6.2 for HPr(Ser(P)) and 7.0 for Crh(Ser(P)). In the former case, MES
was used as a buffer.
Effects of Temperature--
The temperature effects on enzyme
stability were monitored in an assay with 50 mM MOPS (pH
8.0) adjusted at room temperature over a temperature range of
4-70 °C, and the assay mixture was allowed to incubate for 10 min.
MOPS was used as a buffer, since Effects of Different Glycolytic Intermediates--
Glycolytic
intermediates were tested at the following concentrations: 1 mM acetyl phosphate, 2 mM FBP, 1 mM
F2,6BP, 0.5 mM D-glucose 6-phosphate, 1 mM DL-glyceraldehyde 3-phosphate, 1 mM NAD+, 0.1 mM NADH, 0.5 mM acetyl coenzyme A, and 1 mM pyruvate. Up to
40 mM FBP and F2,6BP were tested in a phosphatase assay.
Inorganic phosphate was omitted when evaluating the potential of acetyl phosphate and DL-glyceraldehyde 3-phosphate as possible
candidates to induce phosphatase activity.
Kinetic Parameters for HPr/Crh and ATP--
To
determine the kinetic parameters for HPr/Crh, the concentration of the
protein was varied from 0.1 to 200 µM while keeping the
concentration of the second substrate, ATP, at saturation (2 mM) with 7 mM MgCl2. When the
kinetic parameters for ATP were determined, the concentration of
HPr/Crh was 200 and 169 µM, respectively, and the
concentration of ATP was varied between 1 µM and 1 mM. The concentration of MgCl2 was kept with an
excess of 5 mM over the total ATP concentration to have a
constant proportion of ATP existing as MgATP2 Estimation of Kinetic Parameters--
The program GraphPad Prism
was used to determine K0.5 (the half-saturation
constant in the Hill equation), the Hill coefficient (h),
and Vmax using the following equation:
Y = VmaxSh/(K0.5 + Sh), where S is the substrate concentration.
Characterization of the Oligomeric State of the Purified
Recombinant Proteins
ESI mass spectra were first recorded under denaturing
conditions of the different proteins, HPr(His)6,
Crh(His)6, and HPrK/P(Trx-His6-S tag), to
verify purity and homogeneity, and to determine the molecular weight of
constitutive subunits. The molecular mass determined for
HPr(His)6 was 10,688.5 ± 0.2 Da, which is in
agreement with the theoretical value of 10,687 Da calculated from the
amino acid sequence (data not shown). A minor compound (~10%) with a
mass of 10,768.6 ± 0.2 Da was also detected, corresponding most
probably to the phosphorylated form of the protein
( In addition to the use of ESI-MS for the evaluation of the purity and
homogeneity of proteins, the technique has also become a useful method
to study noncovalent complexes (for recent reviews, see Refs. 36-38).
ESI-MS analysis under native conditions of the proteins revealed only
monomers of HPr(His)6 and Crh(His)6 (10 mM ammonium acetate buffer (pH 6.8), data not shown). Using
other biophysical techniques, circular dichroism and NMR, HPr was
reported to be monomeric, whereas Crh was detected both as monomer and dimer (39). Substantially different experimental conditions presumably
explain the different results obtained by ESI-MS and NMR.
ESI-MS analysis unambiguously revealed that
HPrK/P(Trx-His6-S-tag) from B. subtilis is a specific noncovalent hexamer at pH 6.8 with a
measured molecular mass of 310,337 ± 22 Da (Fig.
1A). The molecular mass
obtained under native conditions is 0.05% higher than the mass
predicted from the denatured analysis of
HPrK/P(Trx-His6-S-tag). This mass difference is very low
compared with other mass measurements analyzing noncovalent
subassemblies (40-42) and is not significant given the uncertainty of
the measurements in native conditions. However, this difference is
presumably due to the inclusion of water molecules or small cations
that are not present in the denatured monomer. Despite the discrepancy,
there is no doubt that HPrK/P(Trx-His6-S-tag) exists as a
hexamer at pH 6.8. The ESI-MS analysis is not in agreement with earlier
reported results performed by size exclusion chromatography and
analytical ultracentrifugation, which suggest that the enzyme is an
octamer or possibly a heptamer (17). However, a precise quantification
and an accurate determination of the molecular weight of oligomeric
proteins are not directly and precisely determined with these latter
techniques. The results from ESI-MS measurements are more precise and
consistent with the x-ray crystallography experiments of HPrK/P from
Lactobacillus casei showing that the catalytic domain of the
enzyme is a hexamer in the crystal, suggesting the same state in
solution (43). The x-ray structure was solved at pH 5.2 with the
truncated HPrK/P with an N-terminal fragment missing. Whereas the
N-terminal fragment is poorly conserved in the enzyme from different
bacteria, the remaining residues, including the putative ATP- and
HPr-binding domains, are highly conserved (43). Recently, full-length
HPrK/P from Staphylococcus xylosus was crystallized (pH
7.6), also supporting a hexameric structure of the enzyme (44).
Furthermore, full-length Mycoplasma pneumoniae HPrK/P(His)6 was also reported to be a hexamer as revealed
by biophysical and crystallographic data (pH 7.5) (45).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
CONCLUSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
CONCLUSION
REFERENCES
-D-thiogalactopyranoside (Sigma) to a final concentration of 1 mM. Further incubation was performed at
37 °C for 2 h. Cells were harvested by centrifugation at 5000 rpm for 25 min at 4 °C. The supernatant was discarded, and the cell pellet was stored at
80 °C for at least 1 h before being
resuspended in 5-10 volumes of 50 mM Tris buffer, pH 8, containing 0.3 M NaCl. A small amount of deoxyribonuclease
I type I (Sigma) was added, and the cell mixture was homogenized using
Ultra-TURRAX for 2-3 min. The cells were then ruptured by two passages
through the French press (American Instrument Co.) at 500 p.s.i.
Whole cells and cell debris were removed by centrifugation at 30,000 rpm for 30 min at 4 °C, and the resultant supernatant was then mixed
during 30 min with Ni2+-NTA resin (Qiagen, Courtaboeuf,
France) (1 ml of resin/g of cells) pre-equilibrated with 10 column
volumes of 100 mM Tris, pH 7.4. For HPrK/P, this was
performed at 4 °C as well as all subsequent purification steps,
whereas for HPr and Crh purification was performed at room temperature.
The Ni2+-NTA resin was then transferred to a column, and
the resin was washed with 50 mM Tris, pH 7.4, containing 50 mM Na2SO4 and 15% glycerol. The
proteins were eluted with imidazole in 50 mM Tris buffer,
pH 7.4, containing 50 mM Na2SO4 and
15% glycerol. HPr was collected from the elution fractions with 30 and
300 mM imidazole. Crh and HPrK/P were eluted with 300 mM imidazole. The proteins were verified with SDS-PAGE
separations using PhastGel (Amersham Biosciences) and Coomassie
staining. After concentrating the fractions by an Ultrafree centrifugal
filter unit with a molecular mass cut-off of 5000 Da, 4 ml
(Millipore Corp., Bedford, MA) at 5000 rpm to about 2.5 ml, the protein
solutions were desalted using a PD-10 column (Amersham Biosciences)
pre-equilibrated with 25 ml of 10 mM Tris buffer, pH 8. The
protein solutions were run into the column and eluted with 3.5 ml of
the buffer solution.
-globulin
(Bio-Rad) as a standard, and the concentrations of Crh and HPr were
determined by UV spectrophotometry using the extinction coefficient for
one and two tyrosine residues, respectively (1500 and 2900 M
1 cm
1). Protein solutions were
stored at
20 °C.
20 °C
until ESI-MS measurements were performed.
D, where
represents the solvent viscosity, kb is the
Boltzmann's constant, and T is the temperature), and then to a molecular weight for a spherical particle. Apparent molecular weights were deduced from histograms of distribution of percentage mass
versus Rh. HPrK/P was diluted in a 10 mM ammonium acetate buffer, similar to that used in mass
spectrometry analysis under native conditions. All solutions were
filtered with 0.22-µm Millex filters prior to dilution of the
protein. Sample preparations were achieved either by diluting the
protein in the pre-equilibrated buffer or by adding increasing amounts
of NH3 to a 10 µM
HPrK/P(Trx-His6-S-tag) solution in ammonium acetate until pH
ranged from 6.8 up to 9.5. An aliquot was removed at the desired pH and
kept at room temperature for at least 2 h before dynamic light
scattering measurement.
-nicotinamide adenine dinucleotide (
-NAD+),
-nicotinamide adenine dinucleotide reduced form (
-NADH), D-glucose 6-phosphate, DL-glyceraldehyde
3-phosphate, CoCl2·6H2O, CuCl2·2H2O,
MnCl2·4H2O, and acetyl phosphate were
purchased from Sigma. MgCl2·6H2O and
CaCl2·2H2O were obtained from Merck.
-32P]ATP/pmol of ATP) (Amersham Biosciences), 2 mM FBP, 0.1% bovine serum albumin, 10 µM
HPr/Crh, and, to initiate the reaction, 100 nM
HPrK/P(Trx-His6-S-tag), in a final volume of 20 µl. The
mixture was incubated at 37 °C for 10 min, and the phosphorylation
reaction was then terminated by spotting samples onto 1 × 1-cm
P81 phosphocellulose paper (Whatman, Maidstone, UK) and dropped
immediately into a beaker containing 75 mM
H3PO4. The total volume of phosphoric acid
solution used was ~10 ml for each paper. Unreacted ATP was removed by
washing three times with 75 mM
H3PO4, for 15 min each, and once with ethanol,
just covering the papers, for 5 min. The papers were dried and
transferred to scintillation vials containing 6 ml of scintillation
solution for water samples, Rotiszint ecoplus (Carl Roth, Karslruhe,
Germany), and the radioactivity was determined in a scintillation
counter, LKB 1211 Rackbeta (PerkinElmer Life Sciences). Typically, each
condition was tested in triplicate.
-32P]ATP/pmol of ATP)
(Amersham Biosciences), 5 mM FBP, 0.1% bovine serum
albumin, 200 µM HPr/Crh, and, to initiate the reaction, 800 nM HPrK/P(Trx-His6-S-tag). After incubation
at 37 °C for 2 h, HPr(Ser(P))/Crh(Ser(P)) was purified on
Ni2+-NTA resin using a column procedure. The resin was
pre-equilibrated with 100 mM Tris buffer (pH 7.4) and then
combined with the assay mixture for at least 30 min at room
temperature. To remove ATP and FBP, the resin was washed extensively
with 50 mM Tris buffer (pH 7.4) containing 50 mM Na2SO4 and 15% glycerol until
no radioactivity was detected in the eluent. The enzyme and the
phosphorylated protein were eluted with 50 mM Tris buffer
(pH 7.4) containing 50 mM Na2SO4,
15% glycerol, and 300 mM imidazole. Desalting was performed using a PD-10 column (Amersham Biosciences). The column was
pre-equilibrated with 25 ml of 10 mM Tris buffer (pH 8).
After adding the sample (not more than 2.5 ml) to the column, the
proteins were eluted with 3.5 ml of 10 mM Tris buffer (pH
8). Fractions containing radioactivity were collected and pooled. In
most of the cases, remaining enzyme from the kinase reaction was
removed by S-protein-agarose (Novagen, Madison, WI). After
equilibration with 20 mM Tris buffer (pH 7.5), 150 mM NaCl and 0.1% Triton X-100 (Bio-Rad) the
S-protein-agarose was gently shaken with the protein solution for 30 min at room temperature. The phosphorylated protein was eluted with 20 mM Tris buffer (pH 7.5), 150 mM NaCl, and 0.1% Triton X-100 as confirmed by radioactivity in the eluent. Desalting was
then performed as described above. An Ultrafree centrifugal filter unit
(molecular mass cut-off of 5000 Da) was used to concentrate the protein
solution (4 °C) according to the operating procedure provided by the
company. The amount of phosphorylated protein, pmol/µl, was
determined on the basis of cpm/pmol ATP. Amounts of
HPr(Ser(P))/Crh(Ser(P)) up to 10 µM were then included in
a phosphatase assay containing, in general, 50 mM Tris
buffer (pH 8), 5 mM MgCl2, 0.1% bovine serum
albumin, 1 mM inorganic phosphate (K2HPO4 and KH2PO4, pH
8.0), and, to initiate the reaction, 100 nM
HPrK/P(Trx-His6-S-tag). The reaction mixture was incubated at 37 °C for 2 h. Termination of the reaction and the following steps were the same as for the phosphorylation assay. Typically, each
condition was tested in triplicate.
pKa/°C is low,
0.006, compared with other buffers (33). For the phosphatase assay,
10 mM inorganic phosphate was used, and the reaction was
allowed to proceed for 10 min before termination.
and a
constant concentration of free Mg2+ (35). The initial
velocity was determined after 10 min of incubation.
RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
CONCLUSION
REFERENCES
M = 80 Da). For Crh(His)6, the main
species detected (~85%) had a mass of 10,392.5 ± 0.2 Da, which
is in agreement with the theoretical value of 10,390 Da (data not
shown). About 15% of the signals were converted to a molecular mass of
10,261.6 ± 0.1 Da, which is probably due to the loss of the
N-terminal Met (
M = 131 Da) from Crh. The different constructions of HPr and Crh with a His tag at the N terminus preceded
by a Met, Arg, Gly, and Ser for HPr and a His tag on the C terminus and
a Met followed by Val on the N terminus for Crh could be a reason for
the differences observed in the mass spectra. A possible explanation
for the findings that a part of HPr was phosphorylated, but not Crh,
may be due to the fact that HPr was phosphorylated at His-15 by the EI
during the purification procedure when the plasmid was expressed in
Escherichia coli. In Crh the His in position 15 is exchanged
by a Gln, and no phosphorylation has been demonstrated in
vitro by phosphoenolpyruvate and EI (14). ESI-MS analysis of
HPrK/P(Trx-His6-S-tag) under denaturing conditions revealed
a highly homogeneous sample with a single species of 51,700.7 ± 1.0 Da, which is in agreement with the theoretical value, 51,699 Da,
taking into account the loss of the N-terminal Met (data not shown).
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Fig. 1.
ESI mass spectra of HPrK/P (20 µM) under native conditions at pH 6.8 (A) and pH 9.5 (B). At pH 6.8, the major signals observed in the mass spectrum are due to multiply
charged ions of the hexamer. Minor signals reveal the coexistence of
small amounts of monomer and dimer. Increasing the pH to 9.5 results in
destabilization of the hexamer, and most of the enzyme was detected as
a monomer. Traces of dimer and hexamer are also observed. The
accelerating voltage was equal to 200 V, and the pressure in the
interface region was 6.5 millibars.
Since the conditions for ESI mass measurements differed from the radioactive kinase assay (see below), due to different buffer solutions and an additional desalting procedure, the kinase activity of HPrK/P was also tested under the same conditions as those used for mass spectrometry. Using 10 mM ammonium acetate (pH 6.8) as buffer, the native enzyme was found to be fully active.
Divalent Cations Are Necessary for Both the Kinase and Phosphatase Activity
In the absence of divalent cations, neither kinase nor phosphatase
activity was detected. The ion required to produce maximal kinase
activity, when HPr was included in the assay, was Mg2+
(Fig. 2A), which is in
agreement with previously published results for HPrK/P and HPr from
B. subtilis (11). When Crh was used as a
substrate, Mg2+ gave the same degree of phosphorylation as
the divalent cation Mn2+. However, for HPr, a somewhat
lower activity was observed with Mn2+ compared with
Mg2+. Some activity was also observed with
Co2+, more for HPr than when Crh was used as a substrate.
For Ca2+ and Cu2+, no activity was observed for
either HPr or Crh under the conditions used. The effect of divalent
cations on HPr phosphorylation by HPr kinase from other Gram-positive
bacteria with low GC content has also been investigated. For HPr kinase
from Streptococcus salivarius, Mg2+ was the
preferred cation, but activity was also observed in the presence of
Mn2+ and Co2+ (46). No effect was seen for
Ca2+ and Cu2+ (46). HPr kinase from
Streptococcus pyogenes was maximally activated by
Mg2+ and Mn2+, and no activity was observed for
the cations Sn2+, Ni2+, and Cu2+
(47). For the different HPr kinases, Mg2+ seems to be the
preferred divalent cation.
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For the phosphatase assay, activity was observed in the presence of Mg2+ and Mn2+ (Fig. 2B). However, in this case, the same activity was obtained with either Mg2+ or Mn2+ when including HPr(Ser(P)) as the substrate, but in the case of Crh(Ser(P)) a more pronounced effect was observed with Mg2+ compared with Mn2+ on dephosphorylation of the proteins. For Co2+, Ca2+, and Cu2+, no major effect on the phosphatase activity was detected.
The HPrK/P was completely dependent on divalent cations for kinase
activity. In general, protein kinases require metal ions, preferentially Mg2+, for catalysis (48). The metal ion
neutralizes charge and orients and polarizes the -phosphoryl
group, which facilitates the phosphorylation reaction (48, 49). Most
protein kinases seem to bind two metal ions that surround the
triphosphate of ATP (48). Although divalent metal ions other than
Mg2+ can be involved, Mg2+ is considered the
physiological activator in, at least, eukaryotes, due to its high
concentration in the cell (48). For protein kinases in general, maximal
activity is usually also observed with Mg2+ except for most
tyrosine kinases, which are maximally activated by Mn2+
(50). The phosphatase activity of HPrK/P also required the addition of
metal cations for catalysis, which is a common contributing factor for
catalysis among Ser(P)/Thr(P) protein phosphatases (49).
Is pH Involved in the Switch between the Kinase and Phosphatase Activity?
Effect of pH on the Oligomeric State of the Enzyme Probed by Mass Spectrometry-- Under carefully controlled operating conditions (accelerating voltage set to 200 V and the pressure in the interface region of the mass spectrometer set to 6.5 millibars), HPrK/P was investigated under native conditions at different pH values. The major ion series observed in the mass spectrum at pH 6.8 was related to multiply charged ions of a hexamer (310,337 ± 22 Da) (Fig. 1a). Minor peaks were attributed to the monomer (51,699 Da) and dimer (103,404 Da) species of the enzyme. An estimation of the different oligomeric forms of the enzyme at pH 6.8 revealed that ~80% of the detected ions correspond to the hexamer, whereas ~10% correspond each to the monomer and dimer. Increasing the pH to 9.5, by the addition of ammonium hydroxide but in otherwise strictly identical conditions as at pH 6.8, resulted in a complete dissociation of the hexamer, and the most intense signals in the spectrum were those of the multiply charged monomer and dimer (Fig. 1b). Approximately 70% of the detected ions corresponded to the monomer, and ~20% of the signals were those of the multiply charged dimer. Only weak signals representative of the hexamer (~10%) were observed.
Effect of pH on Kinase and Phosphatase Activity by
HPrK/P of HPr and Crh--
The effect of pH on substrate
binding and catalysis was studied using a mixed citric acid/Tris buffer
covering the pH range between 5 and 8.7 (Fig.
3). The kinase activity versus
pH shows an optimal activity between pH 6.6 and 8.7 for the enzyme and HPr and a maximal activity at pH 8.7 for the enzyme and Crh. To obtain
further information regarding enzyme characteristics and to assess the
pH stability of HPrK/P, the enzyme was incubated for 10 min at 37 °C
at the indicated pH values in 50 mM citric acid/Tris. After
incubation, the residual activity was measured at pH 8.0 for HPr and pH
9.0 for Crh (final concentration of 250 mM Tris) at
37 °C and after incubation for 10 min. A decline in activity was
observed at pH below 5.8, but above this value the enzyme retained its
activity (data not shown). The decrease in activity between 5.8 and 6.6 when using HPr as substrate and between 5.8 and 8.7 when using Crh as
substrate is probably due to improper ionic forms of the active site
and/or substrate (51). The decline in activity below pH 5.8 can in part
be related to irreversible denaturation of the enzyme (51). Thus, the
activity loss between 6.6 and 8.7 for the enzyme and Crh seems to be
related more to an improper ionic form of the substrate than to the
enzyme, since the enzyme plus HPr does not show this reduction in
activity. Regarding the influence of pH on HPr kinase activity from
other species, the pH optima for HPr kinase from S. salivarius and Streptococcus mutans Ingbritt were
reported to be 7.5 and 7.0, respectively (46, 52). The pH optimum for
the stability of the enzyme from S. salivarius
after preincubation for 30, 60, and 90 min prior to measuring its
activity was 8.0 (46).
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The activity and stability of HPrK/P when acting as a phosphatase were also investigated. Whereas the activity, when investigated with HPr(Ser(P)) as substrate, was optimal at pH 6.2 and then decreased successively above and below this value, a different activity was observed when using Crh(Ser(P)) as substrate (Fig. 3). In the latter case, optimal activity was observed between pH 6.2 and 7.5. Stability experiments measuring the activity at pH 6.2 for HPr(Ser(P)) and pH 7.0 for Crh(Ser(P)) after preincubating the enzyme did not influence the activity (data not shown).
A pH-dependent switch between the kinase and the phosphatase activities of the bivalent enzyme was observed. The kinase activity was predominant at higher pH, and increasing phosphatase activity was recorded at lower pH. The reason for this switch may be connected to the starvation metabolism of the bacterium. In general, the internal pH of neutralophilic bacteria, including B. subtilis, is between 7.5 and 8.0 (53). It has also been demonstrated that B. subtilis possesses a rather high cytoplasmic buffering capacity (54). However, entering the sporulation phase in response to starvation for a variety of nutrients, the internal pH of spores from B. subtilis was found to be ~6 (55). Thus, the kinase activity is high at pH around 8 when growing in the presence of certain nutrients, whereas decreasing pH favors phosphatase activity when the bacteria is about to enter sporulation due to starvation before metabolic dormancy. Favoring the phosphatase activity leads to relief of CCR/CCA. For another bifunctional enzyme, the 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase from bovine liver, the pH optimum for the kinase activity was 8.5, whereas the phosphatase reaction was maximal at pH 6.5 (56). Thus, a similar pH dependence for the two divergent opposing activities was observed, as for HPrK/P.
The ESI-MS results and the activity data indicate that the state of oligomerization may be an important factor in the switch between the kinase and phosphatase activity. Thus, when the enzyme exists as a hexamer, the phosphatase activity is favored.
In order to analyze this pH switch, we followed the oligomerization state of the enzyme as a function of pH using mass spectrometry, dynamic light scattering, and BIAcore techniques. The pH titration by mass spectrometry was done using the previously described conditions.
We used dynamic light scattering to evaluate the changes in the
quaternary structure of the HPrK/P as it has been observed from mass
spectrometry analysis upon changes of pH. Interestingly, the analysis
of histograms of distribution in percentage of mass shows that the
apparent molecular masses were about 2-fold higher at pH 6.8 than at pH
9.5, respectively (343 and 143 kDa (data not shown). These average
molecular weights calculated for a spherical particle and obtained from
eight independent measurements are fully consistent with the existence
of the HPrK/P as a hexamer at pH 6.8, whereas at pH 9.5 it more likely
forms a trimer, although we cannot exclude the possibility that a dimer
would be also compatible with the molecular weight of the particles at
pH 9.5 with regard to the shape of the oligomers. We noted the
existence of a mixture of particles in size as suggested by the high
polydispersity value (around 40%) and the poor fit to a monomodal
distribution. We assume that this polydispersity reflected a dynamic
equilibrium of the HprK/P between different oligomeric states and a
trend to form aggregates as well, since it persisted despite attempts to eliminate high size particles or possible dust particles by centrifugation, filtrating, and increasing the ionic strength of the
protein samples. Therefore, the regularization in bimodal distribution
of radii by percentage of mass as a measurement of lower bound particle
size appears to be a suitable approach for surveying the pH effect on
the structure of the multimeric HprK/P. In summary, our results suggest
that HprK/P undergoes a transition in its oligomeric state upon the
increase of basicity from pH 6.8 to pH 9.5. The trimer (and/or dimer)
appears to be a predominant form of the HprK/P at pH 9.5, and the
hexamer appears to be the main form at pH 6.8. Fig.
4C shows the pH titration
experiments performed by mass spectrometry or by dynamic light
scattering. There is a good agreement between the results obtained by
those two techniques and a good correlation with the pH titration of the phosphatase activity using HPr as a substrate (Fig.
3A).
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We used the BIAcore technology and NTA sensor chip to capture pH-dependent different quaternary structures of the HPrK/P. The experiments were designed to measure a different amount of protein depending on whether the nickel was bound to the hexameric, trimeric, dimeric, or monomeric form of the protein. Thus, we generated NTA chips with low density of nickel so that each nickel was bound to only one hexamer of HprK/P at saturation. Under these conditions, we observed that for the same amounts of HprK/P injected on an NTA chip (regenerated with the same density of nickel), the amount of protein bound was about 2 times less when it was loaded in Tris buffer, pH 9.5, than when it was at pH 6.8. Accordingly to the previous results from mass spectrometry and dynamic light scattering studies, this difference was consistent with the binding of hexamer to the NTA-Ni2+ chip when HprK/P is loaded at pH 6.8 and the binding of trimer when loaded at pH 9.5. Furthermore, the analysis of the sensorgrams obtained from the protein loaded at pH 6.8 showed that the level of resonance units decreased rapidly after it had reached a maximum during the phase of binding. This dissociation phase was initiated at the switch between the loading buffer and the running buffer. We explained this effect as a dissociation of the hexamer at the change of pH from 6.8 to 7.4 between both buffers. In order to investigate more thoroughly the kinetic of dissociation, we performed experiments using running buffers similar to the loading buffers, pH 6.8 and 9.5, following each injection of the protein. The analysis of the sensorgram (data not shown) showed that only the protein loaded at pH 6.8 was subject to a sharp decrease of resonance units following the switch to the running buffer, pH 9.5. The absence of such a fast dissociation with the protein loaded at pH 9.5 indicated that this effect was due to the dissociation of the bound oligomers itself rather than to its dissociation from the Ni2+-NTA chip. Both the stochiometry of the fixation of the HprK/P on the NTA-Ni2+ chip and the kinetics of its dissociation provided further evidence of a transition in the protein oligomeric state dependent on the pH that is consistent with the formation of a hexamer at pH 6.8 and a trimer at pH 9.5.
Altogether, these results strongly suggest that the kinase-phosphatase switch is linked to the oligomeric state of the enzyme. Moreover, BIACORE technology may be used to follow the oligomeric states of the enzyme under different physiological conditions.
The Enzyme Activities for the Two Substrates, HPr and Crh, Are Differentially Affected by Temperature
The temperature effect on enzyme activity was carried out over a
temperature range between 4 and 70 °C using 50 mM MOPS
(pH 8.0), since this buffer is more stable with temperature
(pKa/°C =
0.006) (33). Using an
incubation time of 10 min, the optimum temperature for the kinase
activity was recorded to be between 37 and 45 °C for HPr and
45 °C for Crh (Fig. 4). Furthermore, the enzyme activity with HPr
and Crh, respectively, differs with a higher degree of phosphorylation
for HPr at temperatures below 45 °C than for Crh. The reverse is
observed for temperatures above 45 °C (i.e. a
higher degree of phosphorylation for Crh than for HPr). In the
phosphatase assay, the optimal temperature appears to be 50 °C. No
major differences in the behavior of the activity between the two
different substrates were observed in this case.
Species of Bacillus are the only Gram-positive bacteria with low GC content, which in addition to HPr, is provided with a second protein substrate, Crh (15). The kinase activity was more stimulated at temperatures below 45 °C using HPr as a substrate, whereas temperatures above 45 °C favored phosphorylation of Crh. The proposed background for B. subtilis when provided with two different protein substrates for HPrK/P may deal with CCR/CCA in different ecological environments. In this context, an observation worth noticing is that the gene encoding Crh has been localized in a genome area including operons dealing with catabolic degradation of complex substrates found in roots (7).
The Metabolic State of the Cell, through Different Glycolytic Intermediates, Modulates the Kinase/Phosphatase Balance
Several glycolytic intermediates were tested for potential
allosteric effects of HPrK/P (Fig.
5). The concentrations were chosen according to representative values recorded for B. subtilis as well as other Gram-positive bacteria. Thus, four
intermediates were evaluated at concentrations found in vivo
in Streptococcus mutans, another Gram-positive bacterium
with low GC content: 0.5 mM D-glucose
6-phosphate, 1 mM NAD+, 0.1 mM
NADH, and 1 mM pyruvate (57). FBP was initially tested at 2 mM, a concentration found in B. subtilis when grown on D-glucose (58), and
F2,6BP was also initially tested at a similar concentration, 1 mM. DL-Glyceraldehyde 3-phosphate was included
at a concentration of 1 mM, since intracellular
concentrations of this intermediate were found to be as high as 0.6 mM in Streptoccus bovis (59). Acetyl coenzyme A
and acetyl phosphate were tested at concentrations of 0.5 and 1 mM, respectively. Intracellular concentrations of these two
metabolites have been reported to be ~0.2 mM in the Gram-positive bacteria Corynebacterium glutamicum (60).
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Regarding the kinase activity, FBP and F2,6BP were found to stimulate the phosphorylation reaction (Fig. 5A). FBP stimulated HPr phosphorylation more than F2,6BP, whereas in the case of Crh both intermediates caused the same degree of phosphorylation. On the contrary, inorganic phosphate, acetyl phosphate, and DL-glyceraldehyde 3-phosphate inhibited the kinase activity of HPrK/P. The inhibitory effect of inorganic phosphate on kinase activity has previously been reported for HPr kinases (46, 47, 61-63). To further evaluate the effect of inorganic phosphate, 1 mM inorganic phosphate was included in a kinase assay with increasing concentrations of FBP/F2,6BP. Whereas 10 mM FBP partly restored the kinase activity for both proteins, HPr and Crh, 10 mM F2,6BP had only a small effect regarding the phosphorylation of Crh, and no effect of F2,6BP was observed for the phosphorylation of HPr (data not shown). Similar results with FBP and the capacity of restoring the kinase activity in the presence of inorganic phosphate were reported for HPrK/P from Lactobacillus casei (63). A similar experiment was performed to evaluate the possibility to restore the kinase activity in an assay using ATP with 0.1 mM inorganic phosphate in the absence of effector molecules. Increasing concentrations of ATP up to 3 mM only partly restored the kinase activity (data not shown).
For the phosphatase activity, F2,6BP was found to inhibit the dephosphorylation of both HPr(Ser(P)) and Crh(Ser(P)) as well as high concentrations of FBP in the case of Crh(Ser(P)) (Fig. 5B). FBP alone only inhibited dephosphorylation of Crh(Ser(P)), and 2 mM ATP alone did not affect the dephosphorylation activity of either of the proteins, whereas 20 mM FBP with the addition of 2 mM ATP inhibited dephosphorylation of both HPr(Ser(P)) and Crh(Ser(P)) (data not shown). Similar results have been reported previously for HPrK/P from B. subtilis (9). Since acetyl phosphate and glyceraldehyde 3-phosphate inhibited the kinase activity, their role in the phosphatase assay were investigated (Fig. 5C). Both substances seem to have similar potential as a source of inorganic phosphate, thus shifting the enzyme activity to a phosphatase. As in the case with inorganic phosphate, Mg2+ is required for activity. However, phosphatase activity was also observed in the absence of inorganic phosphate, acetyl phosphate, or glyceraldehyde 3-phosphate but in the presence of Mg2+, although not to the same extent. A similar observation, that the enzyme did not require inorganic phosphate to exhibit phosphatase activity, has also been reported for HPrK/P from S. xylosus (64).
Of the evaluated glycolytic intermediates, FBP as well as inorganic phosphate are known as an allosteric activator and inhibitor, respectively, not only for the kinase activity of HPrK/P from B. subtilis (11) but also for the enzymes from, for example, L. casei (63), Lactobacillus brevis (62), S. mutans (61), and S. pyogenes (47). Furthermore, F2,6BP was also found to be an allosteric activator for the kinase activity. The opposing phosphatase activity was found to be regulated by inorganic phosphate, acetyl phosphate, or glyceraldehyde 3-phosphate. Inorganic phosphate is known as an indicator of the metabolic status of the cell (16). In addition to the role of acetyl phosphate in acetate metabolism and in the overflow mechanism of excess carbohydrates (29), acetyl phosphate has been identified as a regulator of two-component signal transduction systems such as ComA in B. subtilis (65). Acetyl phosphate has been postulated as a good sensor reflecting the intracellular metabolic state of the cell as an indicator of glucose availability, and the synthesis of acetyl phosphate was found to be necessary for glucose-starved cells of E. coli to survive glucose starvation (66). Thus, reflecting the intracellular metabolic status of the cell in response to changing concentrations of PTS sugars, acetyl phosphate could shift the kinase activity of HPrK/P to a phosphatase. Regulatory properties were also found for glyceraldehyde 3-phosphate, which induced the phosphatase activity of the enzyme. Recently presented data suggest multiple levels of control of the gapA operon, including the gene encoding glyceraldehyde 3-phosphate dehydrogenase, gapA, involved in the conversion of glyceraldehyde 3-phosphate to 1,3-bisphosphoglycerate (67). This enzyme catalyzes an irreversible reaction, and enzymes catalyzing irreversible reactions in the glycolysis were found to be inducible by glucose. Furthermore, the gapA operon was found to be regulated by a repressor, CggR, and CcpA (67). In view of the complex regulation of the gapA operon to modulate the needs of glycolysis and the tricarboxylic acid cycle under different conditions, the conversion of glyceraldehyde 3-phosphate to 1,3-bisphosphoglycerate is strongly regulated. However, the exact role of glyceraldehyde 3-phosphate in the CCR/CCA mechanism requires further evaluation.
Taking into account the proposed relationship between the oligomeric state of the enzyme and its activity, FBP and F2,6BP are suggested to promote the dissociation of the hexamer into dimer and monomer. The presence of inorganic phosphate, acetyl phosphate, and/or glyceraldehyde 3-phosphate, on the other hand, are suggested to induce the formation of the hexamer.
The Different Substrate and Effector Sites are Strongly Coupled
The kinetic parameters for the three substrates, HPr/Crh and ATP,
were determined by varying one substrate of interest and keeping the
second substrate at a saturating level. When estimating the kinetic
parameters for HPr and Crh, the concentration of HPr/Crh was varied
between 0.1 and 200 µM, whereas the concentration of ATP,
in general, was kept at 2 mM. Two graphs (Fig.
6) illustrate some of the results, and
all of the kinetic parameters are listed in Table
I.
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Results presented in Fig. 6 (Graph B) are nearly identical to results presented in Fig. 5 of Jault et al. (17). This comparison validates the radioactive assay used in this work, whereas in most of the previous work, the enzymatic activity was measured by a gel assay (17).
The enzyme exhibited a positive homotropic response with respect to substrate binding. For the two substrates, HPr and Crh, the affinity was about 5 times higher for HPr, indicating the preference of HPr as a substrate. The addition of the effector molecule, FBP, resulted in increased values for Vmax, whereas no major effect on K0.5 was observed. F2,6BP, on the other hand, increased both Vmax and K0.5 values. Both activators increased the turnover numbers as well as kcat/K0.5. The degree of cooperativity decreased somewhat or was not affected in the presence of the effectors. The inclusion of the negative effector, inorganic phosphate, lowered all of the kinetic parameters except the Hill coefficient for Crh.
Furthermore, the possibility of an interaction between the HPr/Crh and ATP sites was also investigated. The concentration of ATP was decreased for the examination of whether lowered concentrations of the phosphate donor would alter the observed kinetic behavior of HPr/Crh. When the concentration of ATP was decreased 4-fold, the Hill coefficient decreased in the case with HPr and increased in the case with Crh. Thus, the binding of ATP to the enzyme seems to induce and/or influence the positive cooperative binding of HPr/Crh to HPrK/P.
To estimate the kinetic parameters for ATP, the concentration of ATP was varied between 1 µM and 1 mM, and the concentration of HPr/Crh was kept at 200 and 169 µM, respectively. In the absence of effector molecules, the enzyme exhibited low affinity for ATP. The addition of FBP/F2,6BP lowered both the K0.5 and Vmax when using HPr as the second substrate. However, the opposite effect, with increasing values for Vmax, was observed when including Crh in the assay. Whereas the kcat values decreased in the case with HPr, an increase was observed for Crh. In both cases, however, the quotient of kcat/K0.5 increased. For the Hill coefficient, no obvious explanation for either the increase or decrease in cooperativity after the addition of FBP/F2,6BP can be offered. Experiments to determine the kinetic parameters at pH 7.4 for the different substrates were also performed without effector molecules (data not shown). The results indicate that the positive cooperativity was more pronounced at pH 8.0 than at pH 7.4. In addition, the values for the turnover number and kcat/K0.5 were markedly lowered, except for the turnover number for HPr, which remained the same.
Attempts to determine kinetic parameters in a phosphatase assay failed.
Under the different concentrations of phosphorylated substrates that we
were able to use due to our incapacity to purify high concentrations of
phosphorylated substrate, we were not able to obtain a simple and
complete saturation curve. Therefore, we were not able to obtain
kinetic parameters using the Hill equation (see "Experimental
Procedures").
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CONCLUSION |
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A model for the proposed regulation of HPrK/P with HPr as
substrate is illustrated in Fig. 7. The
switch between kinase and phosphatase activity is probably correlated
to the oligomeric state of the enzyme, where the hexamer favors the
phosphatase activity and the trimeric form of the enzyme favors the
kinase activity. Fieulaine et al. (43) proposed a model
where the shift from kinase to phosphatase activity may be due to a
direct competition between ATP and inorganic phosphate. An observation
supporting an additional mechanism is that phosphatase activity was
also observed even in the absence of inorganic phosphate but in the presence of Mg2+. The same phenomenon, no requirement for
inorganic phosphate to exhibit phosphatase activity, was reported for
HPrK/P from S. xylosus (64). However, a close
relationship seems to exist between the kinase and phosphatase activity
supported by point mutations of Asp-176 and Asp-177 replaced by Ala
residues in HPrK/P from B. subtilis affecting
both the kinase and phosphatase activity (68). In addition, several
mutants of HPrK/P from B. subtilis and
L. casei affect phosphatase activity, but with
almost normal kinase activity, and no mutant specifically affecting
kinase activity has been found (8, 69). Some of the mutations, related
with reduced phosphatase activity, are located on the surface in a direct contact with another subunit, suggesting that the mutation may
influence the oligomeric state of the enzyme (8, 69).
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The internal pH of vegetative cells, ~8, will dislocate the balance to the kinase activity, whereas starvation conditions, when the internal pH is lowered, will favor the phosphatase activity. FBP and F2,6BP, possibly interacting with the same site due to their structural similarities, are probably regulating the kinase activity through allosteric modification of the interaction between the monomeric subunits of the enzyme. The kinetic parameters indicate a stronger regulation by FBP than F2,6BP. Other metabolites that have been reported to stimulate the kinase activity are 6-phosphogluconate and glycerate-2-phosphate, although to a lesser extent than FBP (11).
In addition to inorganic phosphate, the glycolytic intermediates, acetyl phosphate and glyceraldehyde 3-phosphate, were found to induce the phosphatase activity of the enzyme. Whether the latter two regulators bind to the same site as inorganic phosphate is not clear, but for both, divalent cations were necessary for enzyme activity, as in the case with inorganic phosphate. The dephosphorylation reaction does not seem to be a simple reversal of the phosphorylation reaction, since the addition of ADP did not stimulate dephosphorylation of HPr(Ser(P)) in an assay with HPrK/P from L. casei (43). Inhibition of the phosphatase activity was observed in the presence of F2,6BP for HPr(Ser(P)) and Crh(Ser(P)), and also of FBP but only when using Crh(Ser(P)) as a substrate.
The binding of the two substrates, HPr and ATP, yield sigmoidal velocity with positive cooperativity, thus increasing the affinities of the vacant binding sites. Furthermore, the results indicate a kinetic coupling between the ATP and HPr binding sites, since the Hill coefficient was changed when the concentration of ATP decreased. For both the opposing activities of the enzyme, divalent cations were necessary for the reactions not only to neutralize the phosphate group of ATP or HPr(Ser(P))/Crh(Ser(P)) but perhaps also through a specific site modulating the conformational state of the enzyme.
In this study, proteins were expressed in E. coli and fused with different tags (His6 and Trx-His6-S) in order to facilitate their purification and to obtain a high yield of a homogenous population of proteins. The different tags were not removed, assuming that they were not going to interfere with the behavior of the enzyme. Comparison of some specific experiments from our laboratory and other groups using different constructions support this assumption (data not shown) (39). However, it cannot be ruled out that the wild-type enzyme may behave in a somewhat different manner. Nonetheless, the proposed model is assumed to be also applicable for the wild-type enzyme.
In summary, the bifunctional HPrK/P from B. subtilis involved in the main mechanism of CCR/CCA is a
strongly regulated allosteric enzyme. The differences in response
obtained regarding phosphorylation and dephosphorylation of the
proteins under different in vitro conditions
(e.g. regulation by effector molecules and in
cooperativity) could be a means by which bacteria "fine tune" the
regulation of metabolic pathways. Thus, the enzyme HPrK/P is involved
in a sophisticated regulation system, sensing environmental changes in
the availability of carbon sources to adapt the uptake and use of
nutrients in a hierarchical manner in at least two different ecological niches.
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ACKNOWLEDGEMENTS |
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We thank Louis Cuccia for valuable linguistic advice and Mireille Gaire for helpful scientific advice. We thank Virginie Lafont and Danièle Altschuh for technical support with the BIACORE system as well as Philippe Dumas and Bernard Lorber for support with the dynamic light-scattering apparatus.
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FOOTNOTES |
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* 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.
§ Recipient of a grant from the Swedish Academy of Pharmaceutical Sciences.
** To whom correspondence should be addressed. Tel.: 33-3-90-24-42-70; Fax: 33-3-90-24-43-12; E-mail: haiech@pharma.u-strasbg.fr.
Published, JBC Papers in Press, October 30, 2002, DOI 10.1074/jbc.M209052200
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ABBREVIATIONS |
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The abbreviations used are: HPrK/P, HPr kinase/phosphatase; CCR/CCA, carbon catabolite repression/activation; FBP, D-fructose 1,6-diphosphate; PTS, phosphotransferase system; ESI-MS, electrospray ionization mass spectrometry; F2, 6BP, D-fructose 2,6-diphosphate; MES, 4-morpholineethanesulfonic acid; MOPS, 4-morpholinepropanesulfonic acid; NTA, nitrilotriacetic acid.
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REFERENCES |
---|
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---|
1. | Hunter, T. (1995) Cell 80, 225-236[Medline] [Order article via Infotrieve] |
2. | International Human Genome Sequence Consortium. (2001) Nature 409, 860-921[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Venter, J. C.,
Adams, M. D.,
Myers, E. W.,
Li, P. W.,
Mural, R. J.,
Sutton, G. G.,
Smith, H. O.,
Yandell, M.,
Evans, C. A.,
Holt, R. A.,
Gocayne, J. D.,
Amanatides, P.,
Ballew, R. M.,
Huson, D. H.,
Wortman, J. R.,
et al..
(2001)
Science
291,
1304-1351 |
4. | Noonberg, S. B., and Benz, C. C. (2000) Drugs 59, 753-767[Medline] [Order article via Infotrieve] |
5. | Sedlacek, H. H. (2000) Drugs 59, 435-476[Medline] [Order article via Infotrieve] |
6. | Traxler, P., Bold, G., Buchdunger, E., Caravatti, G., Furet, P., Manley, P., O'Reilly, T., Wood, J., and Zimmermann, J. (2001) Med. Res. Rev. 21, 499-512[CrossRef][Medline] [Order article via Infotrieve] |
7. | Kunst, F., Ogasawara, N., Moszer, I., Albertini, A. M., Alloni, G., Azevedo, V., Bertero, M. G., Bessieres, P., Bolotin, A., Borchert, S., Borriss, R., Boursier, L., Brans, A., Braun, M., Brignell, S. C., Bron, S., Brouillet, S., Bruschi, C. V., Caldwell, B., Capuano, V., Carter, N. M., Choi, S. K., Codani, J. J., Connerton, I. F., Danchin, A., et al.. (1997) Nature 390, 249-256[CrossRef][Medline] [Order article via Infotrieve] |
8. |
Hanson, K. G.,
Steinhauer, K.,
Reizer, J.,
Hillen, W.,
and Stülke, J.
(2002)
Microbiology
148,
1805-1811 |
9. | Kravanja, M., Engelmann, R., Dossonnet, V., Blüggel, M., Meyer, H. E., Frank, R., Galinier, A., Deutscher, J., Schnell, N., and Hengstenberg, W. (1999) Mol. Microbiol. 312, 59-66 |
10. |
Galinier, A.,
Kravanja, M.,
Engelmann, R.,
Hengstenberg, W.,
Kilhoffer, M. C.,
Deutscher, J.,
and Haiech, J.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
1823-1828 |
11. | Reizer, J., Hoischen, C., Titgemeyer, F., Rivolta, C., Rabus, R., Stulke, J., Karamata, D., Saier, M. H., Jr., and Hillen, W. (1998) Mol. Microbiol. 27, 1157-1169[CrossRef][Medline] [Order article via Infotrieve] |
12. |
Turinsky, A. J.,
Grundy, F. J.,
Kim, J. H.,
Chambliss, G. H.,
and Henkin, T. M.
(1998)
J. Bacteriol.
180,
5961-5967 |
13. | Galinier, A., Deutscher, J., and Martin-Verstraete, I. (1999) J. Mol. Biol. 286, 307-314[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Galinier, A.,
Haiech, J.,
Kilhoffer, M. C.,
Jaquinod, M.,
Stulke, J.,
Deutscher, J.,
and Martin-Verstraete, I.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
8439-8444 |
15. | Darbon, E., Galinier, A., Le, Coq, D., and Deutscher, J. (2001) J. Mol. Microbiol. Biotechnol. 3, 439-444[Medline] [Order article via Infotrieve] |
16. |
Mason, P. W.,
Carbone, D. P.,
Cushman, R. A.,
and Waggoner, A. S.
(1981)
J. Biol. Chem.
256,
1861-1866 |
17. |
Jault, J. M.,
Fieulaine, S.,
Nessler, S.,
Gonzalo, P., Di,
Pietro, A.,
Deutscher, J.,
and Galinier, A.
(2000)
J. Biol. Chem.
275,
1773-1780 |
18. | Postma, P. W., Lengeler, J. W., and Jacobson, G. R. (1993) Microbiol. Rev. 57, 543-594[Abstract] |
19. | Deutscher, J., Kuster, E., Bergstedt, U., Charrier, V., and Hillen, W. (1995) Mol. Microbiol. 15, 1049-1053[Medline] [Order article via Infotrieve] |
20. | Grundy, F. J., Turinsky, A. J., and Henkin, T. M. (1994) J. Bacteriol. 176, 4527-4533[Abstract] |
21. | Jacob, S., Allmansberger, R., Gartner, D., and Hillen, W. (1991) Mol. Gen. Genet. 229, 189-196[CrossRef][Medline] [Order article via Infotrieve] |
22. | Dahl, M. K., and Hillen, W. (1995) FEMS Microbiol. Lett. 132, 79-83[CrossRef] |
23. | Miwa, Y., and Fujita, Y. (1990) Nucleic Acids Res. 18, 7049-7053[Abstract] |
24. | Fujita, Y., Miwa, Y., Galinier, A., and Deutscher, J. (1995) Mol. Microbiol. 17, 953-960[Medline] [Order article via Infotrieve] |
25. | Oda, M., Katagai, T., Tomura, D., Shoun, H., Hoshino, T., and Furukawa, K. (1992) Mol. Microbiol. 6, 2573-2582[Medline] [Order article via Infotrieve] |
26. | Wray, L. V., Jr., Pettengill, F. K., and Fisher, S. H. (1994) J. Bacteriol. 176, 1894-1902[Abstract] |
27. | Martin-Verstraete, I., Stulke, J., Klier, A., and Rapoport, G. (1995) J. Bacteriol. 177, 6919-6927[Abstract] |
28. | Grundy, F. J., Waters, D. A., Allen, S. H., and Henkin, T. M. (1993) J. Bacteriol. 175, 7348-7355[Abstract] |
29. |
Presecan-Siedel, E.,
Galinier, A.,
Longin, R.,
Deutscher, J.,
Danchin, A.,
Glaser, P.,
and Martin-Verstraete, I.
(1999)
J. Bacteriol.
181,
6889-6897 |
30. |
Turinsky, A. J.,
Moir-Blais, T. R.,
Grundy, F. J.,
and Henkin, T. M.
(2000)
J. Bacteriol.
182,
5611-5614 |
31. | Hueck, C. J., and Hillen, W. (1995) Mol. Microbiol. 15, 395-401[Medline] [Order article via Infotrieve] |
32. | Stulke, J., and Hillen, W. (1999) Curr. Opin. Microbiol. 2, 195-201[CrossRef][Medline] [Order article via Infotrieve] |
33. | Gueffroy, D. E. (1993) Buffers A Guide for the Preparation and Use of Buffers in Biological Systems, 11th printing , pp. 13-16, Calbiochem-Novabiochem Corp., GmbH |
34. | Perrin, D. D., and Dempsey, B. (1974) in Buffers for pH and Metal Ion Control (Albert, A., ed) , pp. 24-54, Chapman and Hall, New York |
35. | Cornish-Bowden, A. (1995) Fundamentals of Enzyme Kinetics , 2nd Ed. , pp. 69-72, Portland Press, Ltd., London |
36. | Loo, J. A. (2000) Int. J. Mass Spectrom. 200, 175-186[CrossRef] |
37. | Last, A. M., and Robinson, C. V. (1999) Curr. Opin. Chem. Biol. 3, 564-570[CrossRef][Medline] [Order article via Infotrieve] |
38. | Loo, J. A. (1997) Mass Spectrom. Rev. 16, 1-23[CrossRef][Medline] [Order article via Infotrieve] |
39. | Penin, F., Favier, A., Montserret, R., Brutscher, B., Deutscher, J., Marion, D., and Galinier, A. (2001) J. Mol. Microbiol. Biotechnol. 3, 429-432[Medline] [Order article via Infotrieve] |
40. | Zal, F., Chausson, F., Leize, E., Van Dorsselaer, A., Lallier, F. H., and Green, B. N. (2002) Biomacromolecules 3, 229-231[CrossRef][Medline] [Order article via Infotrieve] |
41. | Rostom, A. A., and Robinson, C. V. (1999) J. Am. Chem. Soc. 121, 4718-4719[CrossRef] |
42. | Tito, M. A., Tars, K., Valegard, K., Hajdu, J., and Robinson, C. V. (2000) J. Am. Chem. Soc. 122, 3550-3551[CrossRef] |
43. |
Fieulaine, S.,
Morera, S.,
Poncet, S.,
Monedero, V.,
Gueguen-Chaignon, V.,
Galinier, A.,
Janin, J.,
Deutscher, J.,
and Nessler, S.
(2001)
EMBO J.
20,
3917-3927 |
44. |
Marquez, J. A.,
Hasenbein, S.,
Koch, B.,
Fieulaine, S.,
Nessler, S.,
Russell, R. B.,
Hengstenberg, W.,
and Scheffzek, K.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
3458-3463 |
45. | Steinhauer, K., Allen, G. S., Hillen, W., Stulke, J., and Brennan, R. G. (2002) Acta Crystallogr. D Biol. Crystallogr. 58, 515-518[CrossRef][Medline] [Order article via Infotrieve] |
46. |
Brochu, D.,
and Vadeboncoeur, C.
(1999)
J. Bacteriol.
181,
709-717 |
47. | Reizer, J., Novotny, M. J., Hengstenberg, W., and Saier, M. H., Jr. (1984) J. Bacteriol. 160, 333-340[Medline] [Order article via Infotrieve] |
48. | Adams, J. A. (2001) Chem. Rev. 101, 2271-2290[CrossRef][Medline] [Order article via Infotrieve] |
49. | Mildvan, A. S. (1997) Proteins 29, 401-416[CrossRef][Medline] [Order article via Infotrieve] |
50. | Ferrari, S., and Thomas, G. (1991) Methods Enzymol. 200, 159-169[Medline] [Order article via Infotrieve] |
51. | Segel, I. H. (1993) Enzyme Kinetics Behavior and Analysis of Rapid Equilibrium and Steady-state Enzyme Systems , pp. 884-942, John Wiley & Sons, Inc., New York |
52. | Thevenot, T., Brochu, D., Vadeboncoeur, C., and Hamilton, I. R. (1995) J. Bacteriol. 177, 2751-2759[Abstract] |
53. | Booth, I. R. (1985) Microbiol. Rev. 49, 359-378 |
54. | Krulwich, T. A., Agus, R., Schneier, M., and Guffanti, A. A. (1985) J. Bacteriol. 162, 768-772[Medline] [Order article via Infotrieve] |
55. | Barton, J. K., den Hollander, J. A., Lee, T. M., MacLaughlin, A., and Shulman, R. G. (1980) Proc. Natl. Acad. Sci. U. S. A. 77, 2470-2473[Abstract] |
56. | Kountz, P. D., el-Maghrabi, M. R., and Pilkis, S. J. (1985) Arch. Biochem. Biophys. 238, 531-543[Medline] [Order article via Infotrieve] |
57. | Iwami, Y., Yamada, T., and Araya, S. (1975) Arch. Oral Biol. 20, 695-697[Medline] [Order article via Infotrieve] |
58. | Fujita, Y., and Freese, E. (1979) J. Biol. Chem. 254, 5340-5349[Abstract] |
59. |
Asanuma, N.,
and Hino, T.
(2000)
Appl. Environ. Microbiol.
66,
3773-3777 |
60. | Wendisch, V. F., Spies, M., Reinscheid, D. J., Schnicke, S., Sahm, H., and Eikmanns, B. J. (1997) Arch. Microbiol. 168, 262-269[CrossRef][Medline] [Order article via Infotrieve] |
61. | Mimura, C. S., Poy, F., and Jacobson, G. R. (1987) J. Cell. Biochem. 33, 161-171[Medline] [Order article via Infotrieve] |
62. | Reizer, J., Peterkofsky, A., and Romano, A. H. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 2041-2045[Abstract] |
63. |
Dossonnet, V.,
Monedero, V.,
Zagorec, M.,
Galinier, A.,
Perez-Martinez, G.,
and Deutscher, J.
(2000)
J. Bacteriol.
182,
2582-2590 |
64. |
Huynh, P. L.,
Jankovic, I.,
Schnell, N. F.,
and Bruckner, R.
(2000)
J. Bacteriol.
182,
1895-1902 |
65. | Kim, S. B., Shin, B. S., Choi, S. K., Kim, C. K., and Park, S. H. (2001) FEMS Microbiol. Lett. 195, 179-183[CrossRef][Medline] [Order article via Infotrieve] |
66. | Nystrom, T. (1994) Mol. Microbiol. 12, 833-843[Medline] [Order article via Infotrieve] |
67. | Ludwig, H., Homuth, G., Schmalisch, M., Dyka, F. M., Hecker, M., and Stulke, J. (2001) Mol. Microbiol. 41, 409-422[CrossRef][Medline] [Order article via Infotrieve] |
68. |
Galinier, A.,
Lavergne, J. P.,
Geourjon, C.,
Fieulaine, S.,
Nessler, S.,
and Jault, J. M.
(2002)
J. Biol. Chem.
277,
11362-11367 |
69. |
Monedero, V.,
Poncet, S.,
Mijakovic, I.,
Fieulaine, S.,
Dossonnet, V.,
Martin-Verstraete, I.,
Nessler, S.,
and Deutscher, J.
(2001)
EMBO J.
20,
3928-3937 |