Survey
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
P-type ATPase wikipedia , lookup
Signal transduction wikipedia , lookup
G protein–coupled receptor wikipedia , lookup
Protein moonlighting wikipedia , lookup
Histone acetylation and deacetylation wikipedia , lookup
Magnesium transporter wikipedia , lookup
List of types of proteins wikipedia , lookup
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 Further Reading Caister Academic Press is a leading academic publisher of advanced texts in microbiology, molecular biology and medical research. Full details of all our publications at caister.com • MALDI-TOF Mass Spectrometry in Microbiology Edited by: M Kostrzewa, S Schubert (2016) www.caister.com/malditof • Aspergillus and Penicillium in the Post-genomic Era Edited by: RP Vries, IB Gelber, MR Andersen (2016) www.caister.com/aspergillus2 • The Bacteriocins: Current Knowledge and Future Prospects Edited by: RL Dorit, SM Roy, MA Riley (2016) www.caister.com/bacteriocins • Omics in Plant Disease Resistance Edited by: V Bhadauria (2016) www.caister.com/opdr • Acidophiles: Life in Extremely Acidic Environments Edited by: R Quatrini, DB Johnson (2016) www.caister.com/acidophiles • Climate Change and Microbial Ecology: Current Research and Future Trends Edited by: J Marxsen (2016) www.caister.com/climate • Biofilms in Bioremediation: Current Research and Emerging Technologies Edited by: G Lear (2016) www.caister.com/biorem • Flow Cytometry in Microbiology: Technology and Applications Edited by: MG Wilkinson (2015) www.caister.com/flow • Microalgae: Current Research and Applications • Probiotics and Prebiotics: Current Research and Future Trends Edited by: MN Tsaloglou (2016) www.caister.com/microalgae Edited by: K Venema, AP Carmo (2015) www.caister.com/probiotics • Gas Plasma Sterilization in Microbiology: Theory, Applications, Pitfalls and New Perspectives Edited by: H Shintani, A Sakudo (2016) www.caister.com/gasplasma Edited by: BP Chadwick (2015) www.caister.com/epigenetics2015 • Virus Evolution: Current Research and Future Directions Edited by: SC Weaver, M Denison, M Roossinck, et al. (2016) www.caister.com/virusevol • Arboviruses: Molecular Biology, Evolution and Control Edited by: N Vasilakis, DJ Gubler (2016) www.caister.com/arbo Edited by: WD Picking, WL Picking (2016) www.caister.com/shigella Edited by: S Mahalingam, L Herrero, B Herring (2016) www.caister.com/alpha • Thermophilic Microorganisms Edited by: F Li (2015) www.caister.com/thermophile Biotechnological Applications Edited by: A Burkovski (2015) www.caister.com/cory2 • Advanced Vaccine Research Methods for the Decade of Vaccines • Antifungals: From Genomics to Resistance and the Development of Novel • Aquatic Biofilms: Ecology, Water Quality and Wastewater • Alphaviruses: Current Biology • Corynebacterium glutamicum: From Systems Biology to Edited by: F Bagnoli, R Rappuoli (2015) www.caister.com/vaccines • Shigella: Molecular and Cellular Biology Treatment Edited by: AM Romaní, H Guasch, MD Balaguer (2016) www.caister.com/aquaticbiofilms • Epigenetics: Current Research and Emerging Trends Agents Edited by: AT Coste, P Vandeputte (2015) www.caister.com/antifungals • Bacteria-Plant Interactions: Advanced Research and Future Trends Edited by: J Murillo, BA Vinatzer, RW Jackson, et al. (2015) www.caister.com/bacteria-plant • Aeromonas Edited by: J Graf (2015) www.caister.com/aeromonas • Antibiotics: Current Innovations and Future Trends Edited by: S Sánchez, AL Demain (2015) www.caister.com/antibiotics • Leishmania: Current Biology and Control Edited by: S Adak, R Datta (2015) www.caister.com/leish2 • Acanthamoeba: Biology and Pathogenesis (2nd edition) Author: NA Khan (2015) www.caister.com/acanthamoeba2 • Microarrays: Current Technology, Innovations and Applications Edited by: Z He (2014) www.caister.com/microarrays2 • Metagenomics of the Microbial Nitrogen Cycle: Theory, Methods and Applications Edited by: D Marco (2014) www.caister.com/n2 Order from caister.com/order 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 References Belanger, A. E. and Hatfull, G. F. 1999. Exponential-phase glycogen recycling is essential for growth of Mycobacterium smegmatis. J. Bacteriol. 181: 6670-6678. Buhr, A., Flükiger, K., and Erni, B. 1994. The glucose transporter of Escherichia coli. Overexpression, purification, and characterization of functional domains. J. Biol. Chem. 269: 23437-23443. Chen, G. S., and Segel, I. H. 1968. Purification and properties of glycogen phosphorylase from Escherichia coli. Arch. Biochem. Biophys. 127: 175186. De Reuse, H., Roy, A., and Danchin, A. 1985. Analysis of the ptsH-ptsI-crr region in Escherichia coli K-12: nucleotide sequence of the ptsH gene. Gene 35:199-207. DeRisi, J. L., Iyer, V. R., and Brown, P. O. 1997. Exploring the metabolic and genetic control of gene expression on a genomic scale. Science 278: 680-686. Deutscher, J., Kuster, E., Bergstedt, U., Charrier, V., and Hillen, W. 1995. Protein kinase-dependent HPr/CcpA interaction links glycolytic activity to carbon catabolite repression in gram-positive bacteria. Mol. Microbiol. 15: 1049-1053. Eberstadt, M., Grdadolnik, S. G., Gemmecker, G., Kessler, H., Buhr, A., and Erni, B. 1996. Solution structure of the IIB domain of the glucose transporter of Escherichia coli. Biochemistry 35: 11286-11292. Garrett, D., Seok, Y.-J., Liao, D.-I., Peterkofsky, A., Gronenborn, A. M., and Clore, G. M. 1997a. Solution structure of the 30 kDa N-terminal domain of enzyme I of the Escherichia coli phosphoenolpyruvate:sugar phosphotransferase system by multidimensional NMR. Biochemistry 36: 2517-2530. Garrett, D. S., Seok, Y.-J., Peterkofsky, A., Clore, G. M., and Gronenborn, A. M. 1997b. Identification by NMR of the binding surface for the histidinecontaining phosphocarrier protein HPr on the N-terminal domain of enzyme I of the Escherichia coli phosphotransferase system. Biochemistry 36: 4393-4398. Garrett, D. S., Seok, Y.-J., Peterkofsky, A., Gronenborn, A. M., and Clore, G. M. 1999. Solution structure of the 40,000 Mr phosphoryl transfer complex between the N-terminal domain of enzyme I and HPr. Nature Struct. Biol. 6: 166-173. Garrity, L. F., Schiel, S. L., Merrill, R., Reizer, J., Saier, M. H. Jr., and Ordal, G. W. 1998. Unique regulation of carbohydrate chemotaxis in Bacillus subtilis by the phosphoenolpyruvate-dependent phosphotransferase system and the methyl-accepting chemotaxis protein McpC. J. Bacteriol. 180: 4475-4480. Hammen, P. K., Waygood, E. B., and Klevit, R. E. 1991. Reexamination of the secondary and tertiary structure of histidine-containing protein from Escherichia coli by homonuclear and heteronuclear NMR spectroscopy. Biochemistry 30: 11842-11850. Hudson, J. W., Golding, G. B., and Crerar, M. M. 1993. Evolution of allosteric control in glycogen phosphorylase. J. Mol. Biol. 234: 700-721. Jia, Z., Quail, J. W., Waygood, E. B., and Delbaere, L. T. J. 1993. The 2.0Å resolution structure of Escherichia coli histidine-containing phosphocarrier protein HPr. A redetermination. J. Biol. Chem. 268: 2249022501. Kim, S.-Y., Nam, T.-W., Shin, D., Koo, B.-M., Seok, Y.-J., and Ryu, S. 1999. Purification of Mlc and analysis of its effects on the pts expression in Escherichia coli. J. Biol. Chem. 274: 25398-25402. Krebs, E.G. and Preiss, J. 1975. Regulatory mechanisms in glycogen metabolism. MTP Int. Rev. Sci. Carbohydr. Biochem. Ser. I. 5:337-389. Kundig, W., Ghosh, S., and Roseman, S. 1964. Phosphate bound to histidine in a protein as an intermediate in a novel phosphotransferase system. Proc. Natl. Acad. Sci. USA 52, 1067-1074. Liao, D.-I., Silverton, E., Seok, Y.-J., Lee, B. R., Peterkofsky, A., and Davies, D. R. 1996. The first step in sugar transport: crystal structure of the amino terminal domain of enzyme I of the E. coli PEP: sugar phosphotransferase system and a model of the phosphotransfer complex with HPr. Structure 4: 861-872. Lin, K., Hwang, P.K., and Fletterick, R.J. 1997. Distinct phosphorylation signals converge at the catalytic center in glycogen phosphorylases. Structure 5: 1511-1523. Lux, R., Jahreis, K., Bettenbrock, K., Parkinson, J. S., Lengeler, J. W. 1995. Coupling the phosphotransferase system and the methyl-accepting chemotaxis protein-dependent chemotaxis signaling pathways of Escherichia coli. Proc. Natl. Acad. Sci. USA 92: 11583-11587. Mattoo, R. L. and Waygood, E. B. 1983. Determination of the levels of HPr and enzyme I of the phosphoenolpyruvate-sugar phosphotransferase system in Escherichia coli and Salmonella typhimurium. Can. J. Biochem. Cell Biol. 61:29-37. Nosworthy, N. J., Peterkofsky, A., König, S., Seok, Y.-J., Szczepanowski, R. H., and Ginsburg, A. 1998. Phosphorylation destabilizes the aminoterminal domain of enzyme I of the Escherichia coli phosphoenolpyruvate:sugar phosphotransferase system. Biochemistry 37: 6718-6726. Peterkofsky, A., Reizer, A., Reizer, J., Gollop, N., Zhu, P.-P., Amin, N. 1993. Bacterial adenylyl cyclases. Prog. Nucleic Acid Res. Mol. Biol. 44: 31-65. Peterkofsky, A., Seok, Y.-J., Amin, N., Thapar, R., Lee, S. Y., Klevit, R. E., Waygood, E. B., Anderson, J. W., Gruschus, J., Huq, H., and Gollop, N. 1995. The Escherichia coli adenylyl cyclase complex: requirement of PTS proteins for stimulation by nucleotides. Biochemistry 34: 8950-8959. Pieper, U., Kapadia, G., Zhu, P.-P., Peterkofsky, A., and Herzberg, O. 1995. Structural evidence for the evolutionary divergence of mycoplasma from gram-positive bacteria: the histidine-containing phosphocarrier protein. Structure 3: 781-790. Planutis, K.S., Planutiene, M.V., Lazareva, A.V., Sel’kov, E.E., and Evtodienko, Yu. V. 1982. Polyglucose content in the cell and the rate of glucose comsumption during synchronous growth of Escherchia coli. Biochem. Biophys. Res. Commun. 109: 583-587. Plumbridge, J. 1998. Expression of ptsG, the gene for the major glucose PTS transporter in Escherichia coli, is repressed by Mlc and induced by growth on glucose. Mol. Microbiol. 29: 1053-1063. Plumbridge, J. 1999. Expression of the phosphotransferase system both mediates and is mediated by Mlc regulation in Escherichia coli. Mol. Microbiol. 33: 260-273. Postma, P. W., Lengeler, J. W., and Jacobson, G. R. 1996. Phosphoenolpyruvate:carbohydrate phosphotransferase systems. In: Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology. F. C. Neidhardt ed. ASM Press, Washington, D.C. p. 1149-1174. Powell, B. S., Court, D. L., Inada, T., Nakamura, Y., Michotey, V., Cui, X., Reizer, A., Saier, M. H., Jr., and Reizer, J. 1995. Novel proteins of the phosphotransferase system encoded within the rpoN operon of Escherichia coli. Enzyme IIANtr affects growth on organic nitrogen and the conditional lethality of an erats mutant. J. Biol. Chem. 270: 48224839. Preiss, J. and Romeo, T. 1989. Physiology, biochemistry and genetics of bacterial glycogen synthesis. Adv. Microb. Physiol. 30: 183-238. Rath, V. and Fletterick, R. J. 1994. Parallel evolution in two homologues of phosphorylase. Nature Struct. Biol. 1: 681-690. Reizer, J., Reizer, A., Kornberg, H. L., and Saier, M. H., Jr. 1994. Sequence of the fruB gene of Escherichia coli encoding the diphosphoryl transfer protein (DTP) of the phosphoenolpyruvate:sugar phosphotransferase system. FEMS Microbiol. Lett. 118: 159-162. Reizer, J., Romano, A. H., and Deutscher, J. 1993. The role of phosphorylation of HPr, a phosphocarrier protein of the phosphotransferase system, in the regulation of carbon metabolism in gram-positive bacteria. J. Cell. Biochem. 51: 19-24. HPr-Glycogen Phosphorylase Interaction 393 Romeo, T. and Preiss, J. 1989. Genetic regulation of glycogen biosynthesis in Escherichia coli: in vitro effects of cyclic AMP and guanosine 5'diphosphate 3'-diphosphate and analysis of in vivo transcripts. J. Bacteriol. 171: 2773-2782. Seok, Y.-J., Lee, B. R., Zhu, P.-P., and Peterkofsky, A. 1996a Importance of the carboxyl-terminal domain of enzyme I of the Escherichia coli phosphoenolpyruvate: sugar phosphotransferase system for phosphoryl donor specificity. Proc. Natl. Acad. Sci. USA 93: 347-351. Seok, Y.-J., Lee, B. R., Gazdar, C., Svenson, I., Yadla, N., and Peterkofsky, A. 1996b. Importance of the region around glycine-338 for the activity of enzyme I of the Escherichia coli phosphoenolpyruvate:sugar phosphotransferase system. Biochemistry 35: 236-42. Seok, Y.-J., Sondej, M., Badawi, P., Lewis, M. S., Briggs, M. C., Jaffe, H., and Peterkofsky, A. 1997a. High affinity binding and allosteric regulation of Escherichia coli glycogen phosphorylase by the histidine phosphocarrier protein, HPr. J. Biol. Chem. 272: 26511-26521. Seok, Y.-J., Sun, J., Kaback, H. R., and Peterkofsky, A. 1997b. Topology of allosteric regulation of lactose permease. Proc. Natl. Acad. Sci. USA 94: 13515-13519. Seok, Y.-J., Zhu, P.-P., Koo, B.-M., and Peterkofsky, A. 1998. Autophosphorylation of enzyme I of the Escherichia coli phosphoenolpyruvate:sugar phosphotransferase system requires dimerization. Biochem. Biophys. Res. Commun. 250: 381-384. Silljé, H. H. W., Paalman, J. W. G., Ter Schure, E. G., Olsthoorn, S. Q. B., Verkleij, A. J., Boonstra, J., and Verrips, C. T. 1999. Function of trehalose and glycogen in cell cycle progression and cell viability in Saccharomyces cerevisiae. J. Bacteriol. 181: 396-400. Sondej, M., Seok, Y.-J., Badawi, P., Koo, B.-M., Nam, T.-W., and Peterkofsky, A. 2000. Topography of the surface of the Escherichia coli phosphotransferase protein enzyme IIAglc that interacts with lactose permease. Biochemistry 39: 2931-2939. Sondej, M., Sun, J., Seok, Y.-J., Kaback, H. R., and Peterkofsky, A. 1999. Deduction of a consensus binding sequence for IIA glc of the phosphoenolpyruvate:sugar phosphotransferase system by cysteine scanning mutagenesis of Escherichia coli lactose permease. Proc. Natl. Acad. Sci. USA 96: 3525-3530. Strange, R. E. 1968. Bacterial glycogen and survival. Nature 220: 606607. Su, Y., Dostmann, W. R., Herberg, F. W., Durick, K., Xuong, N. H., Ten Eyck, L., Taylor, S. S., Varughese, K. I. 1995. Regulatory subunit of protein kinase A: structure of deletion mutant with cAMP binding domains. Science 269: 807-813. Vadeboncoeur, C. 1995. HPr: heteromorphous protein. Res. Microbiol. 146: 525-530. van Nuland, N. A. J., Hangyi, I. W., van Schaik, R. C., Berendsen, H. J. C., van Gunsteren, W. F., Scheek, R. M., and Robillard, G. T. 1994. The high-resolution structure of the histidine-containing phosphocarrier protein HPr from Escherichia coli determined by restrained molecular dynamics from nuclear magnetic resonance nuclear Overhauser effect data. J. Mol. Biol. 237: 544-559. Watson, K. A., Schinzel, R., Palm, D., and Johnson, L. N. 1997. The crystal structure of Escherichia coli maltodextrin phosphorylase provides an explanation for the activity without control in this basic archetype of a phosphorylase. EMBO J. 16: 1-14. Weigel, N., Powers, D. A., and Roseman, S. 1982. Sugar transport by the bacterial phosphotransferase system. Primary structure and active site of a general phosphocarrier protein (HPr) from Salmonella typhimurium. J. Biol. Chem. 257: 14499-14509. Worthylake, D., Meadow, N. D., Roseman, S., Liao, D.-I., Herzberg, O., and Remington, S. J. 1991. Three-dimensional structure of the Escherichia coli phosphocarrier protein IIIglc. Proc. Natl. Acad. Sci. USA 88: 1038210386. Yang, H., Liu, M. Y., and Romeo, T. 1996. Coordinate genetic regulation of glycogen catabolism and biosynthesis in Escherichia coli via the CsrA gene product. J. Bacteriol. 178: 1012-1017. Zhu, P.-P., Reizer, J., Reizer, A., and Peterkofsky, A. 1993. Unique monocistronic operon (ptsH) in Mycoplasma capricolum encoding the phosphocarrier protein, HPr, of the phosphoenolpyruvate:sugar phosphotransferase system. Cloning, sequencing, and characterization of ptsH. J. Biol. Chem. 268: 26531-26540. 394 Seok et al.