Download Regulation of E. coli Glycogen Phosphorylase Activity by HPr

J. Mol. Microbiol. Biotechnol. (2001) 3(3): 385-393.
JMMB Symposium
HPr-Glycogen Phosphorylase
Interaction 385
Regulation of E. coli Glycogen Phosphorylase
Activity by HPr
Yeong-Jae Seok1*, Byoung-Mo Koo1, Melissa Sondej2,
and Alan Peterkofsky2,§
1
School of Biological Sciences, Seoul National University,
Seoul 151-742, Korea
2
Laboratory of Biochemical Genetics, National Heart, Lung
and Blood Institute, National Institutes of Health, Bethesda,
MD 20892-4036, USA
Abstract
Bacteria sense continuous changes in their
environment and adapt metabolically to effectively
compete with other organisms for limiting nutrients.
One system which plays an important part in this
adaptation response is the phosphoenolpyruvate:sugar phosphotransferase system (PTS).
Many proteins interact with and are regulated by PTS
components in bacteria. Here we review the interaction
with and allosteric regulation of Escherichia coli
glycogen phosphorylase (GP) activity by the histidine
phosphocarrier protein HPr, which acts as part of a
phosphoryl shuttle between enzyme I and sugarspecific proteins of the PTS. HPr mediates crosstalk
between PTS sugar uptake and glycogen breakdown.
The evolution of the allosteric regulation of E. coli GP
by HPr is compared to that of other phosphorylases.
Introduction
Sensory transduction in bacteria is an important device for
monitoring the surrounding environment. Bacteria have the
capacity to efficiently adapt to environmental changes by
switching on and off the utilization of a large number of
carbon sources. The PTS plays an important role in such
sensing mechanisms. This system mediates transport of
hexoses or hexitols across the cytoplasmic membrane by
a mechanism that couples translocation to phosphorylation
of the substrate (Postma et al., 1996). Glucose transport
in E. coli involves three soluble PTS components (enzyme
I (EI), histidine phosphocarrier protein (HPr), and enzyme
IIAglc (EIIAglc)) and one membrane-bound protein, enzyme
IIBCglc (EIIBCglc). EI, HPr, and EIIAglc are encoded by a
single operon, ptsHIcrr, while the membrane-bound
*For correspondence. Email [email protected]; Tel. 82-2-880-8827;
Fax. 82-2-888-4911.
§
Jonathan Reizer was a Senior Staff Fellow in the Peterkofsky lab at NIH
from 1985-1988. During this period, Dr. Reizer edited a book on Sugar
Transport and Metabolism in Gram-Positive Bacteria and published several
outstanding papers dealing with sugar transport in Lactobacilli and
Mycoplasma. This was the stimulus for the subsequent work from the
Peterkofsky lab characterizing the genes and proteins of the PTS in
Mycoplasma capricolum.
© 2001 Horizon Scientific Press
glucose-specific transporter EIIBCglc is encoded by the
ptsG gene. Expression of these genes is under dual
regulation, i.e., activation by the cAMP-CRP complex and
repression by Mlc (Plumbridge, 1998 and 1999; Kim et al.,
1999). Glucose uptake requires sequential phosphoryl
transfer from four PTS proteins, as follows:
Phosphoenolpyruvate ⇒ EI ⇒ HPr ⇒ EIIAglc ⇒ EIIBCglc
⇒ glucose. In this pathway, HPr acts as part of a phosphoryl
shuttle between EI and sugar-specific proteins.
EI is a 64 kDa protein consisting of an N-terminal and
C-terminal domain. The N-terminal domain of EI (EIN),
containing the His-189 active site, extends from residues
1 to 259 and can be phosphorylated in a fully reversible
manner by phosphorylated HPr (Seok et al., 1996a). The
C-terminal domain is necessary for PEP binding and
dimerization (Seok et al., 1996b and 1998). Only the dimeric
form of EI is active (Seok et al., 1998). E. coli HPr is a
small, monomeric protein with a Mr, predicted from its
amino acid sequence, of 9119 (De Reuse et al., 1985).
Phosphoryl transfer from P-EI (His-189) to HPr is at His15 (Weigel et al., 1982). EIIAglc of E. coli is phosphorylated
by HPr at His-90. The EIIBCglc subunit of the transporter is
comprised of two domains, the EIIC domain (residues 1386) spanning the membrane eight times and the
cytoplasmic EIIB domain (residues 387-477) containing the
phosphorylation site, Cys-421 (Buhr et al., 1994). EIIC and
EIIB together catalyze translocation and phosphorylation
of glucose. X-ray and NMR structures have been
determined for EIN (Liao et al., 1996; Garrett et al., 1997a),
HPr (Hammen et al., 1991; van Nuland et al., 1994; Jia et
al., 1993), EIIAglc (Worthylake et al., 1991), and the EIIB
domain (Eberstadt et al., 1996). Recently, the solution
structure of the phosphoryl transfer complex between EIN
and HPr was also elucidated by high resolution NMR
spectroscopy (Garrett et al., 1999).
Since the PTS was discovered in E. coli nearly 35 years
ago (Kundig et al., 1964), additional studies have
documented other metabolic roles of this system (Figure
1). PTS secondary functions include the regulation of
adenylate cyclase, chemotaxis, non-PTS transport
systems, and metabolic enzymes. The phosphorylation
state of the proteins in the cascade varies with transport
activity; i.e., it increases in the absence and decreases in
the presence of the corresponding sugar substrate. The
ratio of phosphorylated to dephosphorylated proteins in
turn serves as signal input for the control of metabolic
pathways, of gene transcription, and of chemosensory
behavior. Genetic and biochemical studies implicate the
crr gene product (EIIAglc) as the main mediator of these
regulatory phenomena in Gram-negative bacteria.
The current model suggests that EIIAglc and/or PEIIAglc interact with target proteins and that the ratio of PEIIAglc to EIIAglc is the determining factor in the regulation.
Unphosphorylated EIIAglc inhibits glycerol kinase and
several sugar permeases by a mechanism termed inducer
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386 Seok et al.
Figure 1. Regulatory roles of the phosphoenolpyruvate:glucose phosphotransferase system in Escherichia coli. Phosphoryl groups are sequentially transferred
from PEP to the transported glucose. Soluble proteins encoded by a single operon, ptsHIcrr, are phosphorylated at a histidine residue, while the EIIB domain
of the membrane-bound transporter, encoded by the ptsG gene, is phosphorylated at a cysteine residue. The regulatory functions of the soluble proteins are
also indicated (See text for further details).
exclusion (Postma et al., 1996; Seok et al., 1997b; Sondej
et al., 1999 and 2000). Phosphorylated EIIAglc stimulates
cAMP synthesis necessary for CRP to activate transcription
of cAMP-dependent genes and operons (Peterkofsky et
al., 1993).
In Gram-positive bacteria, HPr is centrally involved in
carbon catabolite repression as is EIIAglc in Gram-negative
bacteria. HPrs in Gram-positive bacteria are
phosphorylated at a second site, Ser-46, by a metaboliteactivated, ATP-dependent protein kinase and
dephosphorylated by a Pi-dependent phosphatase (Reizer
et al., 1993). HPr phosphorylated at Ser-46 plays a role in
catabolite repression as a corepressor of CcpA (Deutscher
et al., 1995). HPr also modulates the activity of
transcriptional regulators of genes involved in carbohydrate
utilization, and glycerol kinase by a phosphorylationdephosphorylation mechanism in low-GC Gram-positive
bacteria (Vadeboncoeur, 1995). In B. subtilis, chemotaxis
is governed by phosphorylated EI which modulates CheA
activity (Garrity et al., 1998).
Even though many regulatory roles of EIIAglc in enteric
bacteria and HPrs in Gram-positive bacteria have been
reported, there have been only a few reports on the
participation of the two general PTS proteins (EI and HPr)
in the regulation of other metabolic pathways in enteric
bacteria. EI and/or HPr were reported to trigger chemotaxis
towards PTS carbohydrates (Postma et al., 1996; Lux et
al., 1995). HPr was shown to regulate the activity of
glycogen phosphorylase (GP) and stimulate adenylate
cyclase in E. coli (Peterkofsky et al., 1995; Seok et al.,
1997a). In this review, we focus on interaction with and
allosteric regulation of GP activity by HPr in E. coli. The
evolutionary significance of the physical interaction
between HPr and GP and the metabolic crosstalk between
glucose uptake and glycogen breakdown is discussed.
Activation of Glycogen Phosphorylase by Interaction
with HPr
While there was no history suggesting a role for HPr in
metabolic regulation in enteric bacteria, there was a
theoretical rationale to explore this possibility. First, HPr is
both a phosphocarrier and a regulator in Gram-positive
bacteria which do not possess a cytoplasmic EIIAglc protein.
Second, HPr acts as part of a phosphoryl shuttle between
EI and sugar-specific proteins; this shuttle is futile if the
sole role of HPr is in sugar transport. It would be simpler
for PEP to pass the phosphoryl group directly to enzymes
II. Third, the cellular concentration of HPr in E. coli is
significantly higher than necessary for its role as a
phosphocarrier, and there are between 10- and 20-fold
more HPr molecules in a cell than enzyme I subunits
(Mattoo and Waygood, 1983). Furthermore, the small size
of HPr (less than 10 kDa) makes it an ideal potential
regulatory factor. Based on this perspective, the technique
of ligand fishing, using surface plasmon resonance (SPR)
spectroscopy in a BIAcore instrument, was employed to
search for proteins that interact with and might be regulated
by HPr in E. coli extracts (Seok et al., 1997a). SPR detects
a change in refractive index resulting from the interaction
of a soluble protein with another protein adsorbed to a
surface. HPr, immobilized on a CM-5 sensor chip, was used
as the bait. When a crude extract of a stationary phase
culture of E. coli strain IF-1 was allowed to flow over the
immobilized HPr for 10 min, there was a detectable
increase in SPR response. The association constant
HPr-Glycogen Phosphorylase Interaction 387
Figure 2. Concentration dependence for the interaction between E. coli
glycogen phosphorylase and immobilized HPr. The indicated concentrations
of GP (in µg/ml), in the running buffer (10 mM Hepes (pH 7.4), 150 mM
NaCl, and 0.005% Tween 20), were allowed to flow over an immobilized
HPr surface for the determination of kinetic parameters for the interaction
between GP and HPr. As a control, 8 µg/ml of rabbit muscle glycogen
phosphorylase (RM) was exposed to the surface. The association constant
was calculated using the BIAevaluation software. The arrow indicates the
starting points of the injections. (From Figure 4 in Seok et al., 1997a).
between EI and HPr was recently reported to be about
105 M-1 by isothermal titration calorimetry and NMR
spectroscopy (Garrett et al., 1997b; Nosworthy et al., 1998).
When purified EI and EIIAglc were exposed to the HPr
surface in the BIAcore, however, no interaction was
detectable regardless of their state of phosphorylation. The
factor interacting with HPr was purified from an E. coli cell
lysate. Purification steps include ultracentrifugation at
100,000 x g , fractionation by ammonium sulfate
precipitation, DE-52 anion exchange chromatography, gel
filtration chromatography using a Superose 12 column, and
Phenyl-Superose column chromatography. Fractions from
each purification step were examined for binding to
immobilized HPr using the BIAcore. At the final step, the
HPr-binding protein was estimated by SDS-PAGE to be
about 100 kDa in size and approximately 90% pure.
Sequence analysis following proteolysis revealed the
purified binding protein to be E. coli GP, whose molecular
weight, predicted from its amino acid sequence, is 93,144.
Supporting the sequence analysis, overproduction of E.
coli GP resulted in increased binding of a crude extract to
immobilized HPr (Seok et al., 1997a).
Kinetic analysis indicated that the dissociation constant
for the interaction between HPr and purified GP was
approximately 1.7 x 10-8 M (Figure 2), and phosphorylated
HPr (P-HPr) has about 4 times higher affinity. The
observation that P-HPr competes with HPr for binding to
GP indicated that HPr and P-HPr bind to the same or an
overlapping site on GP. The high affinity interaction of HPr
and GP was also confirmed by gel shift experiments using
nondenaturing polyacrylamide gel electrophoresis. Based
on the tight HPr-GP interaction, it was not surprising that
some HPr (about 9 kDa) co-eluted with GP during
purification steps through Mono Q and gel filtration
columns. Moreover, cells overproducing GP to a high level
showed no color development on MacConkey plates with
glucose or mannitol as the indicator sugar, presumably
resulting from sequestration of the available HPr.
Figure 3. Phosphorylation state-dependent effects of varying concentrations
of HPr on glycogen phosphorylase activity. Glycogen phosphorylase activity
was measured using a phosphoglucomutase- and glucose-6-phosphate
dehydrogenase-coupled enzyme assay. The reaction mixture (1 ml)
contained 50 mM sodium phosphate, pH 7.1, 1 mM MgCl2, 0.2% glycogen,
50 nM glucose-1,6-diphosphate, 1 mM β-mercaptoethanol, 0.6 mM NADP,
10 µg/ml phosphoglucomutase, 2 µg/ml glucose-6-phosphate
dehydrogenase, and 5 µg/ml glycogen phosphorylase. The amount of HPr
(0-15 µg/ml) was varied in the coupled enzyme assay mixture to determine
the effects on GP activity (+ HPr). To determine if phosphorylation of HPr
would affect its stimulatory activity, phosphoenolpyruvate (1 mM), EI (2 µg/
ml), and HPr were added to the reaction assay (+ P-HPr). The reaction
mixture was preincubated for 10 min before the assay was initiated by the
addition of GP. Two minutes after the initiation of the reaction with GP, the
A340, a measure of the increase in NADPH, was recorded every minute for
a total of 10 min. A unit of glycogen phosphorylase activity is defined as the
reduction of 1 µmol of NADP/min at 25 °C. The activity is expressed as the
increase over the basal activity (0.00346 units). (From Figure 8A in Seok et
al., 1997a).
The effect of HPr on GP activity was determined in
the physiological direction (glycogen degradation)
employing a coupled enzyme assay. E. coli GP activity
was increased by E. coli HPr to 250% of the control value,
and the stimulation of GP activity by HPr was saturable
(Figure 3). Double reciprocal plots show that HPrdependent activation leads to an increase (approximately
5-fold) in the affinity for glycogen. When the concentration
of phosphate is varied, HPr promotes an approximately 5fold increase in the Vmax. P-HPr exhibited approximately
20% as much stimulation of GP activity as dephosphoHPr (Figure 3). Since it is difficult to reproducibly generate
a sample of P-HPr totally devoid of the dephospho-form, it
is likely that the weak stimulatory effect by P-HPr is a
reflection of contamination with the dephospho-form and
that P-HPr itself does not activate GP. Because the cellular
concentration of HPr in E. coli is in great excess over that
of GP (Chen and Segel, 1968; Mattoo and Waygood, 1983),
and both dephospho- and phospho-HPr have high affinity
for GP, it is therefore reasonable to assume that, in vivo,
E. coli GP is always bound to HPr and that physiological
perturbations that shift the ratio of HPr to P-HPr lead to
changes in the activity of GP.
Real time interaction analysis by SPR, gel-shift, and
sedimentation equilibrium studies, and GP enzyme activity
assays showed that the interaction between HPr and GP
was highly specific; only E. coli HPr, not HPrs from Grampositive bacteria or mycoplasma, interacts with and
388 Seok et al.
activates E. coli GP. Although NPr (Powell et al., 1995)
and DTP (Reizer et al., 1994) also accept a phosphoryl
group from EI in E. coli, neither of these proteins stimulates
E. coli GP activity. Likewise, the E. coli HPr-dependent
allosteric regulation of GP was specific for the E. coli
enzyme. Although eukaryotic GPs exhibit a high degree of
sequence identity to E. coli GP (Hudson et al., 1993), rabbit
muscle GP does not interact with nor is its activity
stimulated by E. coli HPr. While both GP and maltodextrin
phosphorylase (MP) are involved in the breakdown of α–
1,4-glucans and there is extensive sequence conservation
in these two proteins in E. coli (Hudson et al., 1993), there
was no interaction between HPr and MP. The absence of
HPr binding to E. coli MP and rabbit muscle GP emphasizes
the high degree of specificity of the HPr-GP interaction.
To learn the importance of individual amino acid
residues in HPr involved in the binding and activation of
GP, 51 mutant HPrs were examined. Those residues where
mutation resulted in a significant change in the gel shift of
the HPr-GP complex and resulted in a stimulation of GP
activity of less than 50% compared with wild-type HPr
included Arg-17, Lys-24, Lys-27, Lys-40, Ser-46, Gln-51,
and Lys-72. Mutation at Lys-24 showed the most dramatic
effect on the binding to and activation of GP. It is worth
noting that the residues Lys-24, Lys-27, and Ser-46 of HPr
appear to be essential both for activation of GP and for the
interaction of HPr with GTP (Peterkofsky et al., 1995).
These three residues occupy a limited surface region on
HPr. Sedimentation equilibrium experiments showed that
binding of HPr to E. coli GP enhances monomer-dimer
association almost 4-fold and dimer-tetramer association
66-fold. Mutant HPrs, defective in interaction with GP, show
significantly different associative behavior with GP. For the
K40A mutant, the monomer-dimer association is about 500fold weaker, while the dimer-tetramer association is 2-fold
stronger than for wild-type HPr. For the K72E mutant, the
association for dimer formation is over 300-fold weaker,
while that for tetramer formation is virtually the same as
for wild-type HPr (Seok et al., 1997a). The studies cited
above constitute the first direct evidence that associates
HPr with regulatory functions in E. coli.
Physiology of HPr-Glycogen Phosphorylase
Interaction
Glycogen and other similar α–1,4-glucans have been
reported in over 40 different bacterial species; the
concentration of glycogen accumulated can be over 50 %
of the cell dry weight (Preiss and Romeo, 1989). In E. coli,
a single operon, glgCAP, encodes genes for ADP-glucose
pyrophosphorylase, glycogen synthase, and GP,
respectively (Yang et al., 1996). Synthesis of glycogen
involves ADP-glucose pyrophosphorylase and glycogen
synthase, while breakdown involves GP. Regulation of
glgCAP expression occurs coordinately. Positive and
negative regulations are exerted by cAMP and the CsrA
gene product, respectively (Romeo and Preiss, 1989; Yang
et al., 1996). GP catalyzes the breakdown of glycogen into
glucose -1-phosphate while the PTS effects the uptake of
sugar substrates, like glucose, and hence both play a
central role in carbohydrate metabolism. As mentioned
above, crosstalk between these two important carbon
Figure 4. Regulation of E. coli glycogen phosphorylase activity by the
phosphorylation state of HPr. The model postulates that, when PTS sugars
are being transported (onset of active growth phase), HPr is primarily in
the dephospho-form. Under these conditions, GP is activated, and glycogen
degradation is enhanced. The degraded glycogen serves as an extra source
of energy during early stages of growth. When PTS sugars are not being
transported (stationary phase), HPr is primarily in the phospho-form. Under
these conditions, GP is in the less active form, and glycogen accumulates.
The reactions stimulated by glucose uptake are indicated as thick arrows.
metabolic pathways is mediated by the phosphorylation
state of HPr of the E. coli PTS. Figure 4 summarizes the
role of the HPr-GP interaction in central carbon metabolism
in E. coli. The phosphocarrier function of HPr in the PTS
allows the protein to exist in both phospho- and dephosphoforms. Substantial activation of GP by HPr requires that it
be in the dephospho-form. If the role of glycogen is just to
store energy, it might be plausible to assume that cellular
glycogen accumulates during growth but not starvation.
Thus, one would not predict that only the dephospho-form,
but not the phospho-form, of HPr activates hydrolysis of
glycogen because the phosphorylation state of the PTS
proteins increases in the absence and decreases in the
presence of the corresponding sugar substrate. However,
in fact, glycogen is accumulated during the transition from
exponential growth into stationary phase in E. coli and some
other microorganisms (Preiss and Romeo, 1989).
Many studies suggest that glycogen provides carbon
and energy to prolong viability during stationary phase by
sparing essential macromolecular components such as
proteins and rRNA from turnover (Strange, 1968; Preiss
and Romeo, 1989; Yang et al., 1996). Many bacteria and
HPr-Glycogen Phosphorylase Interaction 389
yeast accumulate glycogen in either stationary phase or
stress conditions such as nitrogen or sulfur starvation, heat
shock or osmotic stress. Recently, it was shown that the
genes coding for the yeast glycogen synthase complex
are induced during a metabolic shift to the stationary phase
by the exhaustion of glucose, by employing DNA microarray
technology (DeRisi et al., 1997). In E. coli, the rate of growth
and quantity of glycogen accumulated are inversely related
when cells are grown in nitrogen-limited, glucosecontaining media. The ratio of phosphorylated to
dephosphorylated forms of soluble PTS proteins decreases
abruptly during the shift to stationary phase in E. coli, and
phosphorylated forms prevail under starvation conditions
(unpublished data). The phosphorylated form of EIIAglc will
increase cellular cAMP levels. Consistent with the concept
that cAMP levels rise in stationary phase (Peterkofsky et
al., 1993), glg operon expression is increased in stationary
phase (Preiss and Romeo, 1989; Yang et al., 1996).
Because regulation of glgCAP expression occurs
coordinately, there must be other mechanisms for the
regulation of glycogen breakdown distinct from its synthesis
to explain the selective accumulation of glycogen in
stationary phase (Chen and Segel, 1968). This differential
regulation is accomplished by HPr in E. coli.
It is important to note that glycogen is not required for
growth since mutants of E. coli and some other bacteria,
unable to synthesize glycogen, grow as well as their normal
parents (Preiss and Romeo, 1989). Several species of
enteric bacteria containing or not containing glycogen have
been compared with respect to their viability in media
devoid of exogenous carbon source. Prolonged survival
rates were observed only with the strains containing
glycogen. However, in media containing 0.5 – 1 mM MgCl2
glycogen was not required for prolonged survival (Krebs
and Preiss, 1975). The Mg++ is believed to stabilize rRNA
against turnover, thus playing a similar role as the
suggested function of glycogen.
In order to evaluate a link of the PTS to survival of E.
coli through the regulation of glycogen breakdown,
chromosomal PTS deletion mutants were studied for their
ability to maintain survival during prolonged incubation in
stationary phase. Cultures were incubated at 37°C for an
extended period of time (7 days), and the numbers of
culturable cells were determined daily. While the colony
forming units per ml (CFU/ml) of wild-type cells dropped
only slightly, from 109 to 107 after 7 days in stationary phase,
no surviving cells (CFU) were observed for strains deficient
in HPr, EI, and EIIAglc after 4 days in stationary phase and
after 6 days for strains deficient in EI, and EIIAglc, or EIIAglc
alone (unpublished data). Since EIIAglc regulates the
production of cAMP and the CRP-cAMP complex regulates
the expression of many stationary phase genes including
GP, the possibility that these results are associated with
depletion of cAMP was ruled out. Examination of deletion
mutants for adenylate cyclase showed only slightly less
CFU/ml than wild-type (106 after 7 days), and the addition
of cAMP to the medium had no effect on culturability of
PTS mutants. These findings suggest that HPr plays a role
in the ability of E. coli to maintain culturability from nutrient
deprivation by enabling utilization of energy reserves from
glycogen, but not via the regulation of cAMP levels.
During the transition from stationary phase into a new
round of growth, the demand for biosynthetic metabolism
increases and a number of metabolic changes occur. E.
coli as well as many other microorganisms rapidly degrade
available glycogen to accommodate the increased growth
rate. The decrease in cellular cAMP level characteristic of
the entry into logarithmic growth phase (Peterkofsky et al.,
1993) turns off the expression of the glgCAP operon, and
glycogen synthase activity preferentially decays. The
uptake of a PTS sugar should promote a shift in the state
of HPr in the direction of dephospho-HPr, resulting in an
activation of GP (Figure 4). Consequently, effective
recovery from the stationary phase of growth may be
enhanced by the energy derived from the utilization of
stored glycogen. There are several reports supporting this
assumption. One study examined synchronized E. coli B1
cells (Planutis et al., 1982). At the beginning of cell division,
the level of cellular glucose is low while that of glycogen is
high. As cell division progresses, the level of glucose
increases while that of glycogen drops possibly via glucose
uptake and glycogen breakdown. Other studies suggest
that exponential-phase glycogen recycling is essential for
growth in Mycobacterium smegmatis (Belanger and Hatfull,
1999) and that glycogen is required for an increase in the
cell division cycle and cell viability in Saccharomyces
cerevisiae (Silljé et al., 1999). Obviously, further clarification
of the role of bacterial glycogen is important.
The Structural Basis for the Evolution of the HPrGlycogen Phosphorylase Interaction
As mentioned above, the interaction between HPr and GP
is highly specific. E. coli NPr, Bacillus subtilis HPr, and
Mycoplasma capricolum HPr show no interaction with or
stimulatory effect on the activity of E. coli GP. Those
residues where mutation resulted in a significant change
in the gel shift of the HPr-GP complex and resulted in a
decrease of the stimulation of GP activity compared with
wild-type HPr included Arg-17, Lys-24, Lys-27, Lys-40, Ser46, Gln-51, and Lys-72. The X-ray crystallographic structure
(Jia et al., 1993) shows that these residues are on the
surface of HPr and distributed over a large area. Five of
the seven residues have a positive charge, while the other
two amino acids are polar, suggesting that electrostatic
interactions are important for assembly of the active HPrGP complex. A comparison of HPr primary sequences
(Postma et al., 1996; Zhu et al., 1993; Pieper et al., 1995)
shows that enteric HPrs are very similar to each other;
however, they differ considerably from those of Grampositive bacteria and mycoplasma except around the site
of phosphorylation (His-15). Comparison of amino acid
sequences of E. coli HPr with B. subtilis HPr, M. capricolum
HPr, and E. coli NPr shows that only two (Arg-17 and Ser46) of the seven residues are identical and the others are
not conserved. Consequently, it is not surprising that only
E. coli HPr interacts with E. coli GP. This argues that E.
coli HPr evolved in a unique way to regulate glycogen
hydrolysis by sensing the availability of carbon sources in
its environment. It is noteworthy that Ser-46 is the site of a
regulatory phosphorylation in Gram-positive bacteria; thus,
Ser-46 of HPr is important in regulatory functions in both
Gram-positive and Gram-negative bacteria.
The specificity of the interaction between GP and HPr
390 Seok et al.
Figure 5. Alignment of the complete amino acid sequences of E. coli glycogen phosphorylase (EcoGP), E. coli maltodextrin phosphorylase (EcoMP), and
rabbit muscle glycogen phosphorylase (rabitM). Amino acid residues whose side-chains participate in subunit-subunit contact (obtained from Watson et al.,
1997) and/or ligand binding (obtained from Hudson et al., 1993) are indicated in the row labeled role: d, residues for subunit-subunit contact; #, active site; *,
glycogen storage site; •, AMP binding site; -, pyridoxal phosphate binding site; and , glucose-6-phosphate binding site. Residues that are identical in the
listed proteins are shown in reverse shading.
HPr-Glycogen Phosphorylase Interaction 391
was further supported by the findings that E. coli HPr didn’t
bind to and stimulate the activity of E. coli maltodextrin
phosphorylase (MP) or rabbit muscle GP (Seok et al.,
1997a). Alpha-glucan phosphorylase has been examined
from a variety of organisms including bacteria, yeast, slime
mold, plants, insects, and vertebrates including humans.
These proteins share similar enzymatic properties. All of
them catalyze the phosphorylation of the α,1-4 glucosyl
link between glucose residues at the non-reducing end of
the glucosyl chain and utilize the 5'-phosphate group of
the cofactor pyridoxal phosphate (PLP) in catalysis
(Hudson et al., 1993). They exhibit similar pH dependence
and reaction kinetics, and require the dimeric state for
activity (Watson et al., 1997). Furthermore, the amino acid
sequences of phosphorylases are highly conserved with
more than 40% sequence identity regardless of the source
(Preiss and Romeo, 1989), and the overall threedimensional structures show similar folds for rabbit muscle
and yeast GPs and E. coli MP (Rath and Fletterick, 1994;
Watson et al., 1997). The initiating events for activation or
expression of phosphorylases also have a common factor,
cAMP, which in eukaryotes acts to stimulate cAMPdependent protein kinase for phosphorylase kinase
activation and in the bacterial system to relieve catabolite
repression through interaction with CRP. The cAMP
regulatory binding subunit of cAMP-dependent protein
kinase is homologous in sequence and structure to CRP,
indicating a common evolutionary origin in response to
cAMP (Su et al., 1995). In both cases, cAMP is synthesized
in response to external stimuli, hormonal stimulation for
the mammalian system and nutrient deprivation for the
bacteria. Nevertheless, phosphorylases differ in their affinity
and specificity for polysaccharides, their modes of
regulation, and their physiological roles. E. coli MP has a
high affinity for linear oligosaccharides and <1% activity
against glycogen (Watson et al., 1997), while GPs from
several organisms have a poor affinity for linear
oligosaccharides and a high affinity for glycogen (Preiss
and Romeo, 1989). Mechanisms for phosphorylase
regulation also differ from species to species. In the case
of E. coli GP, the high affinity interaction with HPr is the
major basis for activity regulation. E. coli MP and other
bacterial and plant phosphorylases are active in the
absence of phosphorylation and AMP and have, thus far,
shown no regulatory properties (Hudson et al., 1993). The
yeast enzyme requires phosphorylation for maximal activity,
but is insensitive to activation by AMP (Hudson et al., 1993),
while mammalian enzymes are activated by
phosphorylation or AMP. Crystallographic studies of yeast
and rabbit muscle GPs and E. coli MP have elucidated the
structural characteristics and changes undergone by a
protein in response to phosphorylation and to various
allosteric effectors. (Hudson et al., 1993; Rath and
Fletterick, 1994; Watson et al., 1997).
To understand the evolutionary significance of the HPrGP interaction in E. coli and the differences between
regulated and unregulated phosphorylases, sequences and
structures of phosphorylases from different sources are
compared. In Figure 5, the complete amino acid sequences
of E. coli GP and MP are aligned with rabbit muscle GP.
The amino acid residues participating in ligand binding are
marked based on the structure of rabbit muscle GP, while
those for subunit-subunit contact are from the structure of
E. coli MP (Hudson et al., 1993; Watson et al., 1997). E.
coli MP and GP are smaller than is mammalian GP, and
lack some residues both at the N- and C-terminus. There
is about 40% identity over all aligned residues. Overall,
the C-terminal half is more conserved than the N-terminal
region, in which most of the regulatory sites are located.
Despite their regulatory differences, all known
phosphorylases share similar catalytic properties and a
highly conserved PLP site (Hudson et al., 1993). Therefore,
it is no surprise that residues for the active site (glucose
binding site) and surrounding the PLP site are almost 100%
conserved in all three enzymes. The amino acid residues
for glycogen storage sites are highly conserved between
E. coli GP and the rabbit muscle enzyme, but not those in
E. coli MP. This probably accounts for the difference in
substrate specificity of E. coli MP compared to the other
enzymes. The fact that only three of eight residues are
identical for the glucose-6-phosphate binding site explains
the difference in the regulatory properties between rabbit
muscle GP and bacterial phosphorylases by this
metabolite. The majority of sequence differences occur in
those regions that are involved in control. In particular, the
first 120 residues, which in rabbit muscle GP contain the
phosphorylatable Ser-14, exhibit minimal similarity in amino
acid sequence with only eleven identities, a level which is
below the threshold where sequence similarity alone would
dictate structural similarity. In E. coli MP, there are no
corresponding interactions that mimic the Ser-14
phosphate and N-terminal tail interactions of rabbit muscle
GP (Watson et al., 1997). During evolution, selective control
by phosphorylation could have been conferred on
eukaryotic GPs (Hudson et al., 1993), while regulation of
E. coli GP could have been accomplished by
phosphorylation of HPr. Furthermore, while AMP binding
residues are highly conserved in mammalian enzymes,
they are not conserved in yeast and bacterial
phosphorylases, which are not regulated by AMP. Rabbit
muscle GP is activated by AMP which binds at the allosteric
site and promotes changes in the subunit-subunit contacts
similar to those observed for activation by phosphorylation.
Residues involved in the subunit-subunit interfaces of the
dimer differ significantly in the crystal structures of yeast
and rabbit muscle GP (Lin et al., 1997) and E. coli MP
(Watson et al., 1997). This has important consequences
with respect to the conformation of a gate to the catalytic
site of phosphorylases. For example, while access to the
catalytic site is controlled by either covalent phosphorylation
or binding with AMP in rabbit muscle GP, the gate to the
catalytic site is held in an open conformation as a result of
the changes at the subunit interface in E. coli MP. The
open access to the conserved catalytic site provides an
explanation for the activity without control in this basic
archetype of a phosphorylase (Watson et al., 1997).
Sedimentation equilibrium studies show that HPr changes
the state of oligomerization of GP in E. coli (Seok et al.,
1997a). Thus, HPr seems to be involved in the subunitsubunit contacts of GP, and the gate to the catalytic site
might be controlled by the phosphorylation state of the HPr
subunit in the E. coli HPr-GP complex. Accordingly, a
separate pathway for allosteric signal transmission across
subunits may have evolved in this enzyme. Like
392 Seok et al.
mammalian GPs, the E. coli enzyme, complexed with HPr,
seems to exist in two different quaternary and tertiary
structural states, which is determined by the availability of
nutrients.
In summary, HPr mediates crosstalk between two
important pathways for carbon metabolism, i.e., uptake of
sugar substrates and breakdown of the storage compound,
glycogen. The changes that occurred during the separate
evolutionary development of these enzymes have led to
different properties in terms of regulatory patterns and
three-dimensional structures. The regulation of GP activity
by the phosphorylation state of HPr is proposed to be the
bacterial analogy to the covalent phosphorylationdephosphorylation cascade characteristic of eukaryotic
GPs. Clearly, ongoing X-ray crystallographic studies of the
HPr-GP complex and mutagenic studies on GP will
enhance our understanding of the details of the interaction
of GP with HPr. Further clarification of the role of bacterial
glycogen is also needed to understand the physiological
basis for activation of glycogen breakdown by
unphosphorylated HPr in the presence of PTS sugars.
Acknowledgement
Y.-J. Seok obtained financial support from the Korea Research Foundation
in the program year 1998, and B.-M. Koo was supported by BK21 Research
Fellowship from the Korean Ministry of Education
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