How Phosphotransferase System-Related Protein Phosphorylation Regulates Carbohydrate Metabolism in Bacteria

EI is encoded by the ptsI gene. Sequence comparisons reveal that the EIs (about 570 residues; 63 kDa) from various gram-positive and gram-negative bacteria exhibit significant identity (345, 678). EI is autophosphorylated in the presence of Mg²⁺ (940) at the N-3 position of the imidazole ring of a conserved histidine (His-189 in E. coli EI) (16, 940), which is located on the N terminus of the protein. The C terminus contains the PEP binding site and is necessary for dimerization (114, 254, 798). The two-domain structure of the protein was discovered by high-sensitivity differential scanning calorimetry (478). Limited proteolysis of E. coli EI in vitro results in a specific split and provides a stable peptide representing the N-terminal domain (455). The N-terminal domain of EI (EI-N) (115) as well as the C-terminal domain (EI-C) (234) were cloned and characterized. EI-C was shown to complement EI-N both in vivo and in vitro; i.e., EI-N was phosphorylated in the presence of EI-C, PEP, and Mg²⁺ (234). A truncated protein composed of the first 259 amino acids has been synthesized and purified, and both the crystal (475) and solution (258) structures were resolved. It appears that phosphorylation does not drastically change the conformation of the N-terminal domain per se. Recently, the crystal structure of the C-terminal domain of EI of the thermophile Thermoanaerobacter tengcongensis was also elucidated (619). EI exhibits about 30% sequence similarity with pyruvate phosphate dikinase (PPDK) (328, 605, 670), and the various domains in both proteins have similar structures (619). PPDK, which converts ATP, P_i, and pyruvate into AMP, pyrophosphate (PP_i), and PEP; PEP synthase, an enzyme that catalyzes the conversion of pyruvate and ATP into PEP, AMP, and P_i and that is also phosphorylated on a histidyl residue (590); and EI have related functions. For PPDK, a structural model with three domains has been proposed: a C-terminal PEP/pyruvate binding domain, an N-terminal nucleotide binding domain, and a P∼His domain linked to the other two domains via two flexible polypeptide segments (328). Although the nucleotide and PEP/pyruvate binding sites are approximately 45 Å apart, the P∼His domain is thought to bridge the distance by swiveling between the two other domains to enable the phosphoryl transfer. Recent data obtained from a comparison of several PPDK crystal structures are consistent with the proposed model (587). Similar to PPDK, a large distance (>20 Å) between the PEP/pyruvate binding site and the phosphorylatable histidine was observed in a model of EI in which the N-terminal and C-terminal EI structures were fused (619). Swiveling of the N-terminal domain by changing the position of the domains with respect to each other but retaining the structure of the individual domains reduces the distance significantly (to 7.4 Å). In the case of EI, the N-terminal domain contains the HPr binding site rather than the nucleotide binding site of PPDK and PEP synthase. This model was confirmed by recent studies in which the structure of full-length EI of Staphylococcus carnosus was determined (516). The phosphohistidine and HPr-binding domain are clearly separated from the C-terminal dimerization and PEP binding domain. The structure also revealed the extensive interaction surface related to the dimer. Thermodynamic analyses of the influence of several effectors on the conformational stability of EI of Streptomyces coelicolor showed that at a low pH, EI is partly unfolded and that the presence of PEP causes structural changes (356). Two other recent studies on the structure of EI (639) and its C-terminal domain (638) suggest a relatively large structural variability for monomeric EI, which progressively diminishes when EI dimerizes and binds Mg²⁺ and PEP. The observed effects are explained by a swiveling mechanism, as described for PPDK. The structure thus provides additional evidence for the proposed conformational change necessary to facilitate phosphotransfer.

HPr (about 90 residues; 9 to 10 kDa) is encoded by the ptsH gene and has been purified from a variety of organisms (678). Similarity is most pronounced around the active-site histidyl residue, His-15 in the HPr of most enteric bacteria and firmicutes (183, 681, 941). HPr is phosphorylated at the N-1 position of the imidazole ring of His-15. The crystal and solution structures of HPrs from E. coli and B. subtilis as well as a few other species are known (for reviews, see references 33, 123, 329, and 936). The molecule forms an open-faced β-sandwich consisting of a four-stranded antiparallel β-sheet that is covered at one side by one short and two long α-helices. The active-site His-15 is located in the N-terminal part of the first long α-helix and is exposed to the solvent.

The solution structures of the complex between HPr and the N-terminal domain of EI (260) and between HPr and EIIA^Glc of E. coli (929) have been determined. The absence of large changes of the chemical shift values for the backbone atoms upon complexation indicates that the interactions of HPr with EI and EIIA^Glc do not require considerable conformational changes in any of the domains of the PTS binding partners. The binding interface of HPr and glycogen phosphorylase (see “REGULATION OF CARBON METABOLISM IN GRAM-NEGATIVE ENTERIC BACTERIA”) was also mapped, and the interaction surface of the three structures was compared (930). EI and EIIA^Glc interact with the same narrow region of HPr, whereas the region of interaction with glycogen phosphorylase is somewhat wider (929, 930). This is consistent with the higher binding affinity reported for HPr with glycogen phosphorylase than for HPr with EI or EIIA^Glc (259, 799). A similar interaction surface was reported for E. coli HPr binding either EIIA^Mtl (139, 908) or EIIA^Man (949) and B. subtilis HPr binding EIIA^Glc (123). The key interacting residues are located in helices 1 and 2 and the loops preceding helix 1 and following helix 2. Studies with a large number of ptsH mutants, which show lowered affinity for binding to EI, suggest that the same residues are important for the interaction between EI and HPr (85).

In most low-G+C gram-positive bacteria and a few gram-negative organisms, HPr can also be phosphorylated by an ATP-dependent protein kinase on a seryl residue, e.g., Ser-46 in B. subtilis. We will discuss the importance of the second regulatory phosphorylation in detail below (see REGULATORY FUNCTIONS OF P-Ser-HPr). The regulatory phosphorylation is not part of the phosphoryl transfer to carbohydrates, but phosphorylation of the seryl residue slows the phosphoryl transfer from P∼EI to HPr at least 100-fold. E. coli HPr contains a Ser-46 residue but lacks an HPr kinase. Nevertheless, replacement of Ser-46 in E. coli HPr by an aspartyl residue lowers the affinity of P∼EI for HPr almost 1,000-fold (589). This effect is most likely caused by electrostatic repulsion, as the mutation caused by the replacement of Ser-46 with an aspartyl residue (Ser46Asp) leads to only weak structural changes (952).

The carbohydrate specificity of the PTS resides in the EIIs, which consist of an integral membrane domain facing both the periplasmic space and the cytoplasm and cytoplasmic domains or proteins. All EIIs are built basically in a similar modular manner (764) and can consist of up to four separate proteins (for reviews, see references 464, 678, 730, and 813). The PTSs were classified into four (super)families with distinct evolutionary origins on the basis of the phylogenies of the EIIs (759) as follows: (i) the glucose-fructose-lactose superfamily, comprised of the glucose family (e.g., E. coli EIIA^Glc/EIICB^Glc or B. subtilis EIICBA^Glc [CB, CBA, etc., indicate the domain order in multidomain EIIs]), the fructose-mannitol family (e.g., E. coli EIICBA^Mtl), and the lactose family (e.g., L. casei EIIA^Lac/EIICB^Lac); (ii) the ascorbate-galactitol superfamily, comprised of the ascorbate family (e.g., E. coli SgaA/SgaB/SgaT) (358, 989) and the galactitol family (e.g., E. coli EIIA^Gat/EIIB^Gat/EIIC^Gat) (610); (iii) the mannose family (e.g., E. coli EIIAB^Man/EIIC^Man/EIID^Man or B. subtilis EIIA^Lev/EIIB^Lev/EIIC^Lev/EIID^Lev [a mannose- and fructose-specific PTS]); and (iv) the dihydroxyacetone family (e.g., E. coli EIIA-HPr-EI^Dha [DhaM] [304] or EIIA^Dha in firmicutes).

The presence of a common preserved domain in the latter two families (411) might indicate a similar evolutionary origin of the two families and thus the existence of a third superfamily. The sequence-based classification of the various PTSs is supported by X-ray crystallography and nuclear magnetic resonance (NMR) studies, which clearly show that the structures of the EIIA (474, 617, 818, 858, 906, 955) and EIIB (1, 2, 98, 459, 623, 778, 906) domains/proteins belonging to the various classes are quite different. Information on the structure of the integral membrane domain EIIC (and EIID) is limited. The membrane topologies of these proteins have been studied using Cys replacement mutagenesis and chemical modification by thiol reagents for EIIBCA^Bgl (960) and EIICBA^Mtl (918).

Extensive work on the intricate properties of the EII complexes and the detailed mechanisms by which they transport and phosphorylate carbohydrates has been performed in the laboratories of G. T. Robillard, B. Erni, and G. R. Jacobsen. The results that focus predominantly on EIICBA^Mtl, EIIC^Man/EIID^Man, and EIICB^Glc have been reviewed extensively (369, 730, 813), and we will therefore discuss only a few important aspects. Some more recent papers included work on the properties of a cyclized protein formed by the soluble EIIA^Glc and the membrane-bound EIICB^Glc (812), the dimerization of EIICBA^Mtl (903, 904), sugar recognition by the EIIC^Man/EIID^Man and EIICB^Glc of E. coli (256), the membrane topologies of EIIBCA^Bgl (960) and EIICBA^Mtl (918), and transient-state kinetics of enzyme EIICB^Glc (544).

The glucose-specific EII complex of enteric bacteria consists of two distinct proteins, the cytoplasmic protein EIIA^Glc (gene crr) and the membrane-associated protein EIICB^Glc (gene ptsG), in which the EIIB domain is hydrophilic and in contact with the cytoplasm and the EIIC domain is buried within the membrane. The EIIA and EIIB domains are defined as the domains that receive the phosphoryl group from P∼HPr and P∼EIIA, respectively. Whereas the phosphorylated residue in the EIIA domain/protein is a histidyl residue, that in the EIIB domain/protein can be either a histidyl residue (mannose family) or a cysteyl residue (all other families). The latter was first demonstrated in E. coli EII^Mtl (634), and the specific phospho-cysteyl residue was thereafter identified in a number of cases by analytical methods (547, 633). The phosphoryl group of the EIIB domain is transferred to the carbohydrate after translocation of the carbohydrate by the EIIC domain across the membrane and delivery at the inside face of the cytoplasmic membrane. In the case of the mannitol-specific E. coli EII^Mtl or the glucose-specific B. subtilis EII^Glc complex, the EII is a single polypeptide that contains the two hydrophilic domains EIIA and EIIB as well as the transmembrane domain EIIC. Again, in other EII complexes, such as the mannose-specific E. coli EII^Man, phosphoryl groups are carried by two histidyl residues present in the A and B domains of the cytoplasmic EIIAB^Man, whereas translocation of the carbohydrate requires the two transmembrane subunits EIIC and EIID.

Phosphoryl transfer between the PTS proteins proceeds by an in-line associative mechanism (48). The transfer is accommodated by the formation of heterodimeric transition complexes, and considering the kinetics of the transfer, the heterodimeric association should be transient. The solution structures of different transition complexes, including EI and HPr (260), HPr and EIIA^Glc (124, 929), HPr and EIIA^Mtl (139), HPr and EIIA^Man (949), EIIA^Glc and EIIB^Glc (98, 270), and, finally, P∼EIIA^Chb and Cys10Ser mutant EIIB^Chb (398), could be determined. The formation of the heterodimeric complexes has a significant effect on the flux control properties of the individual components of the PTS (738, 896, 901).

Based on the observation that E. coli EI and HPr are attached to membrane fragments of E. coli (757), it was speculated that the PTS components associate into a metabolon (611), which would improve PTS function. However, it is not clear how phosphotransfer from one component to the next could take place in a fully associated system without linkers to provide room for the domains to move, and furthermore, as it was calculated that diffusion should not limit glucose influx (239, 240), there should be no functional driving force to fully associate into an EI-HPr-EII complex. Formation of larger complexes by EI linked to yellow fluorescent protein has been observed in cells grown on PTS as well as non-PTS carbohydrates when they reached stationary phase and high cell densities (637). The observed distribution of the fluorescence over the cells was punctate and sometimes even bipolar (after induction of yellow fluorescent protein-EI with IPTG [isopropyl-β-d-thiogalactopyranoside]). The aggregation appeared to be reversible, as it disappeared after the addition of a carbohydrate, which led to renewed growth of the cells and dispersion of the fluorescence. Thus, phosphotransfer activity within the PTS, as invoked by the addition of a carbohydrate, in fact seems to promote the overall dissociation of the PTS components rather than their association into a metabolon.

Notwithstanding, several PTS proteins were experimentally proved to associate to functional homodimers. The first PTS protein that was found to functionally dimerize was EI (432, 560; also reviewed in reference 114). Later, it was shown that the N-terminal domain does not dimerize (115) but that the C-terminal domain is responsible for dimerization (86). In fact, the association properties of the isolated C-terminal domain are similar to that of entire EI, although the binding constants are orders of magnitude larger (638). It was proposed that the monomer/dimer transition of EI could potentially regulate uptake flux (940), and as a result, the transition has been extensively studied. The idea was supported by the fact that PEP phosphorylates only dimeric EI, and the rate of association/dissociation is very slow, especially compared to sugar uptake (114, 539). The monomer/dimer transition has been studied by various analytical techniques, and many physiological conditions have been tested. It was observed that P∼EI has a 10-fold increased dimerization constant with respect to unphosphorylated EI and that the dimerization of EI is promoted by PEP and Mg²⁺ and inhibited by pyruvate (190-192). It was concluded that the stimulatory effect of Mg²⁺ and PEP on EI association results from a change in conformation. Association of both normal EI and the nonphosphorylatable mutants H189E, H189A, and H189Q was studied by carrying out sedimentation equilibrium experiments (639). The association constant appears to be very sensitive to pH, temperature, and ionic strength and may vary by as much as 700-fold. It was concluded that the ligands Mg²⁺ and PEP are the major determinants for dimerization and that their binding leads to conformational changes. A model that accommodates the known facts about the dimerization was presented. A potential regulatory function of EI dimerization with regard to the role of PEP is discussed below.

Several membrane-spanning EIIs have also been shown to dimerize, such as EIICBA^Mtl (730) and EIICB^Glc (213, 214, 548). Proteome analysis of stable oligomeric protein complexes confirmed the existence of EIICBA^Mtl and EIICB^Glc oligomers in the membrane of E. coli (826). In addition, oligomers of EII^Nag and EII^Tre were detected, which complies with a generalized picture in which the PTS permeases function as dimers (or oligomers). In contrast, it is generally assumed that HPr and EIIA do not form functional homooligomers. However, several experimental observations suggest the opposite. Biochemical evidence strongly suggested that EIIA^Lac forms homotrimers (176, 319). This suggestion was supported by the crystal structure of EIIA^Lac from Lactococcus lactis, which also revealed a homotrimer (818). In addition, it was reported that four molecules of EIIA^Glc associate with one EIICB^Glc homodimer (213). Moreover, multimeric EIIA^Glc was observed during gel filtration (786) and in crystals (224). Dimeric EIIA^Chb and P∼EIIA^Chb were observed when ultracentrifugation was carried out (398). EIIAB^Man was found in the dimeric form both in the cytoplasm and linked to the membrane (216, 826). Also, EIIA^Ntr, a protein homologous to EIIAs that are functional in sugar transport, crystallizes as a dimer (73), although it seems to be a monomer in solution (927). Crh, an HPr-like protein of B. subtilis, was shown to form homodimers in crystals (644), and F29W HPr from Geobacillus stearothermophilus (previously Bacillus stearothermophilus) forms similar domain-swapped dimers in crystallization experiments (827). On the other hand, several measurements performed with the protein in solution suggested that F29W HPr forms monomers. Recently, in vitro experiments provided kinetic evidence that all components of the E. coli glucose PTS form functional dimers (R. Bader, J. G. Blom, K. J. Hellingwerf, M. A. Pelletier, H. V. Westerhoff, and C. Francke, unpublished results).

Although the supposed primary function of the PTS is the transfer of the phosphoryl group from PEP to carbohydrates, all reactions up to and including the EIIs are reversible. The equilibrium constant, K_eq, for the reaction PEP⇔P∼EI⇔P∼HPr is about 80 for the E. coli enzymes (539), whereas the K_eq for the phosphoryl group transfer between HPr and EIIA^Glc from E. coli has a value of between 1 and 1.5 (540). Thus, the overall K_eq for the phosphoryl group transfer from PEP to EIIA^Glc is in the order of 80 to 120. Furthermore, the K_eq for the reaction P∼EIIA^Glc + EIICB^Glc⇔EIIA^Glc + P∼EIICB^Glc was determined to be 12 (539), very close to the estimated value of 7 (737). The number signifies that the phosphotransfer potential of the thiophospho bond in P∼Cys-EIICB^Glc is very close to that of the phosphoamidate bond in phosphohistidine of the other PTS proteins. Only the final step, P∼EIIB^Glc + glucose⇔EIIB^Glc + glucose-6-phosphate (glucose-6-P), is virtually irreversible, with a measured forward rate of 3.2 × 10⁶ M⁻¹ s⁻¹ (539). These values indicate that under physiological conditions, the phosphoryl transfer reactions catalyzed by the PTS between PEP and P∼EIIB^Glc can run in both directions, from one high-energy P∼enzyme intermediate to the next or back to the previous reactions. This differs from most eukaryotic signal transduction systems in which the seryl, tyrosyl, and threonyl residues that are often phosphorylated represent low-energy phosphoenzyme intermediates (837). The reversibility of the phosphoryl group transfer in the PTS has regulatory consequences because it allows the metabolic network to control the phosphorylation state of the PTS proteins in a number of ways, and we will discuss how numerous cellular processes are regulated by the phosphorylation states of certain PTS proteins below (see Table 1 for a summary).

Most of our current knowledge regarding Crp/cAMP-mediated regulation is derived from the study of cyaA, crp, and/or ptsHI-crr mutants (76, 416, 636, 677, 678, 763, 890). The effect of different carbon sources on growth, gene expression levels, cAMP concentrations, and Crp levels has been tested with these mutants. The addition of exogenous cAMP was used to distinguish between the effects caused by concentration changes of either cAMP or Crp. One class of mutants, designated crp* or crp(In), produced Crp, which is active without cAMP. Unfortunately, an interpretation of the mutant data is not easy, as the factors involved in cAMP-mediated regulation are interdependent, as depicted in Fig. 2.

Inactivation of the cyaA gene disables bacteria like E. coli and S. enterica serovar Typhimurium from growing on a large number of carbon sources (76), while cyaA overexpression appears to be detrimental (699). Inactivation of crp also affects the cAMP concentration by leading to a drastic increase in the rate of cAMP synthesis (75, 170, 241, 363, 679). The increase in the cAMP concentration depends on an intact crr gene. In crp crr double mutants, cAMP overproduction is markedly reduced (from about 60-fold of the wild-type level in the crp strain to only 4-fold of the wild-type level in the double mutant) (143, 170).

The phenomena described above are related to feedback regulation on the level of transcription. Expression of cyaA is negatively regulated by Crp/cAMP in both E. coli (10, 387, 395, 576) and S. enterica serovar Typhimurium (221), whereas expression of crp is both positively (310) and negatively (9, 140, 311) affected by Crp/cAMP. As a result, strong crp expression occurs at low and high Crp/cAMP concentrations (due to reduced inhibition and increased stimulation, respectively), whereas low crp expression is observed at intermediate Crp/cAMP concentrations (310). The concentration of Crp/cAMP in exponentially growing cells is an order of magnitude higher than that allowing minimal crp transcription (310, 365). Consequently, a decrease in the cAMP concentration caused by the addition of glucose to these cells should lower the transcription of crp and hence the concentration of Crp. Glucose-grown E. coli cells indeed contain less Crp, and this was shown not to be due to faster degradation of the crp mRNA (364, 365). The glucose effect on crp expression is absent in mutants lacking either adenylate cyclase or Crp, consistent with the postulated role of Crp/cAMP. The growth stage of E. coli cultures also has an influence on the amount of Crp (365). These variations might explain why some authors reported complete relief from glucose repression after the addition of cAMP (647), whereas others observe only partial relief (932).

Expression of the EIIA^Glc-encoding crr gene is almost completely independent of Crp/cAMP (857). In contrast, expression of the EIICB^Glc-encoding ptsG gene is almost fully repressed in cyaA or crp mutants (727). Owing to the low level of the glucose transporter EIICB^Glc in these mutants, the addition of glucose to cyaA or crp mutants does not lead to the same extent of dephosphorylation of P∼EIIA^Glc as in the wild-type strain. The regulation of crp transcription by factors other than Crp/cAMP adds further complexity to catabolite repression data. crp transcription is negatively regulated by both the DNA binding protein Fis (280), the transcription of which is in turn regulated by Crp/cAMP (591), and SpoT (378), the protein that mediates the stringent response and that catalyzes the synthesis of ppGpp (guanosine tetraphosphate).

Whether P∼EIIA^Glc exerts its regulatory effect on adenylate cyclase directly or indirectly has been an open question for a long time. Early experiments with permeabilized cells containing an intact PTS showed that adenylate cyclase activity is stimulated by potassium and phosphate and inhibited by α-methylglucoside (α-MG) and pyruvate (476, 477, 701), which is in line with a stimulatory effect by P∼EIIA^Glc. However, in vitro experiments with purified adenylate cyclase were not conclusive (963). The effect of EIIA^Glc phosphorylation on the activation of adenylate cyclase was studied in crr mutants producing EIIA^Glc in which the phosphorylatable His-90 (197) and the catalytically important His-75 had been changed. In comparison to wild-type EIIA^Glc, His75Gln EIIA^Glc synthesized in a crr mutant caused an approximately fourfold activation of adenylate cyclase. In contrast, neither His90Gln nor His90Glu EIIA^Glc activated adenylate cyclase (700, 852). The latter two mutant proteins are not phosphorylated by P∼His-HPr, while His75Gln EIIA^Glc is. Takahashi and coworkers also found that His75Glu EIIA^Glc, which is barely phosphorylated in vitro (700), can slightly activate adenylate cyclase (852). In wild-type EIIA^Glc, His-75 and His-90 are in very close proximity, and the replacement of His-75 with a negatively charged Glu possibly mimics phosphorylated His-90. Although these results indicate that phosphorylation of His-90 of EIIA^Glc is important for the activation of adenylate cyclase, they do not prove a direct interaction. Recent results with adenylate cyclase tethered to the membrane showed that although both P∼EIIA^Glc and unphosphorylated EIIA^Glc bind with similar affinity to the protein, it is activated by P∼EIIA^Glc only when an additional, not-yet-identified factor from cell extracts is present (632) (Fig. 2).

From studies with truncated adenylate cyclase, it can be concluded that the catalytic activity resides in the N-terminal domain, with the C-terminal domain required for EIIA^Glc-mediated regulation (632, 699, 746). Starting from an E. coli ptsHI-crr deletion strain containing, in addition, an Asp414Asn mutation in cyaA, which prevents the large increase in adenylate cyclase activity in a crp mutant (143), suppressor mutants affected in the cyaA gene could be isolated. Most suppressor mutations were located in the region around Asp-414 and resulted in a truncated adenylate cyclase that was about half the size of the wild-type enzyme (144). The truncated molecules produce about 10 times more cAMP in a crr strain than the wild-type enzyme. These results suggest that the N-terminal domain of adenylate cyclase is active in the absence of P∼EIIA^Glc and that the C-terminal domain acts as an inhibitor that may be removed/inactivated by P∼EIIA^Glc.

Mutation of crp leads to a drastic increase of extracellular cAMP. This increase is 50-fold diminished when the C-terminal 48 amino acids of adenylate cyclase are cut off. In contrast, the increase is diminished by only fourfold when the CRP/cAMP-mediated transcriptional regulation of the cya gene is prevented by replacing the cya promoter with the constitutive bla promoter (363). Therefore, it was concluded that the regulation of cAMP production occurs mainly posttranslationally (at the level of enzyme activity) and not at the level of transcription. However, because the regulation of cyaA transcription is affected by the Crp/cAMP concentration, which in turn is affected by CyaA activity, that conclusion does not seem to be justified. This can be illustrated by the changes in adenylate cyclase activity caused by a crr null mutation. Adenylate cyclase activity measured in crr mutants amounts to only 5% to 20% of the activity measured in wild-type strains (227, 469, 595), and the reintroduction of plasmid-borne crr increases it four- to fivefold, thus approaching levels of wild-type activity (477, 700). If EIIA^Glc was the only factor interacting with and stimulating adenylate cyclase, crr mutants and strains producing adenylate cyclase missing the last 48 amino acids should exhibit the same phenotype. However, it is evident that the positive effect on adenylate cyclase caused by the expression of plasmid-borne crr in a crr mutant is much smaller than the negative effect (50-fold) caused by the deletion of the 48 C-terminal amino acids of adenylate cyclase in a crp mutant. This difference probably owes to the absence of the Crp-dependent effect of cAMP on transcription in a crp mutant.

Despite the enormous amount of experimental data, the control of cAMP and Crp levels in enteric bacteria is still not completely understood. Regulation occurs not only at the level of transcription but also at the level of enzyme activity. These two processes affect each other, making it difficult to quantify their individual contributions. Nevertheless, the general properties of the regulatory mechanism are clear. The immediate response of cells to the addition of a rapidly metabolizable carbohydrate involves regulation only at the level of enzyme activity. The addition of glucose or another PTS sugar causes the dephosphorylation of P∼EIIA^Glc. This deactivates adenylate cyclase and hence lowers the concentration of cAMP and Crp/cAMP inside the cell. On a longer time scale, regulation on the transcriptional level becomes important. At normal physiological levels of cAMP and Crp, the crp gene is negatively regulated by Crp/cAMP. Thus, a reduced activity of adenylate cyclase leads to lower expression of crp. As a result, the concentration of Crp/cAMP drops even further. On the other hand, a lower Crp/cAMP concentration leads to an increased expression of cyaA, and more adenylate cyclase will accumulate, thereby counteracting the decrease in activity and raising the Crp/cAMP concentration. In principle, the elevated pool of adenylate cyclase provides a higher potential for cAMP production once the glucose is depleted, although the Crp concentration will be lower.

Some results cannot be explained by the proposed mechanism. For instance, in S. enterica serovar Typhimurium crp* (595) and E. coli crp* cya (851) mutant strains, the addition of glucose lowers the concentration of Crp* and hence repression. However, because Crp* is constitutively active, i.e., it does not need cAMP for activity, its concentration should not be affected by the presence or absence of glucose. The occurrence of a component that is able to interact with Crp in the presence of glucose could explain the observed decrease in the Crp concentration. In fact, such a mechanism regulates the activity of the repressor Mlc (588): when glucose is present, EIICB^Glc is dephosphorylated and binds the transcription factor Mlc, thus relieving repression. It is tempting to speculate that unphosphorylated EIICB^Glc might also bind Crp, but as yet, there is no experimental proof to support this assumption.

Factors other than those mentioned above have been reported to influence the activity of adenylate cyclase, but their physiological relevance remains uncertain. A decrease in adenylate cyclase activity, for instance, was observed upon lowering the membrane electrochemical potential (649). Mutation of fruB (formerly fruF) can have a negative (272) or positive (225) effect on adenylate cyclase activity. Also, when GTP or the elongation factor Tu, a GTP-binding protein (702), was added to purified adenylate cyclase, it stimulated its activity almost twofold (829).

Although early studies reported hardly any effect of the growth rate on the cAMP concentration under glucose limitation (529, 957), more recent studies report a clear relationship, with elevated cAMP concentrations occurring at low growth rates (613, 614). During growth on glucose in continuous culture, expression of lacZ appears to be independent of ppGpp but directly related to the cAMP concentration and inversely related to the growth rate (438); i.e., growth rate and cAMP concentration are also inversely related. The effect of the growth rate on the cAMP concentration can be explained in terms of the stimulation of adenylate cyclase activity by P∼EIIA^Glc. The phosphorylation state of the PTS components in these experiments should depend on the growth rate because the cells were grown in a chemostat under carbohydrate-limiting conditions, and the growth rate was varied by changing the rate of carbon source supply. As a result, at high growth rates, less P∼EIIA^Glc and hence less cAMP should be formed than that at slow growth rates.

Makman and Sutherland (508) observed that following the addition of glucose, starved E. coli cells halt cAMP synthesis and excrete cAMP into the medium. In a later study, the presence of other metabolizable sugars was shown to elicit the same effect (758). Interestingly, crp mutants excrete much higher amounts of cAMP than wild-type cells (75, 241, 679). A putative low-affinity cAMP exporter (K_m of approximately 10 mM) was identified (275). However, as the internal cAMP concentration is generally in the micromolar range, excretion is not thought to be relevant to cAMP-mediated regulation.

The breakdown of cAMP by the endogenous cAMP phosphodiesterase (80) provides another possible way to affect the cAMP concentration. However, cAMP phosphodiesterase has a low affinity for cAMP (K_m of between 0.5 and 0.8 mM) (80, 604), making it unlikely that this enzyme exerts much, if any, control over cAMP levels. The findings that, in mutants lacking cAMP phosphodiesterase, glucose still lowers the cAMP level (508) and that CCR exerted by various carbon sources is as strong as that in wild-type cells (337) also argue against a physiological role of the cAMP phosphodiesterase.

During inducer exclusion, unphosphorylated EIIA^Glc binds directly to either the respective permease (lactose or melibiose), the ATP-binding subunit of the ATP binding cassette (ABC) transporter (maltose), or, in the case of glycerol, GlpK, the cytoplasmic enzyme catalyzing the formation of the inducer (inducer exclusion is depicted in Fig. 2). Initial evidence for a direct interaction between EIIA^Glc and the inhibited proteins came from studies showing that the lactose transport activity of LacY in E. coli vesicles is strongly reduced when the vesicles contain partially purified unphosphorylated EIIA^Glc (189, 559). Similar results were obtained with whole cells synthesizing either LacY or the melibiose transporter (MelB) (561). Direct binding of unphosphorylated EIIA^Glc to membrane preparations of an E. coli strain overproducing LacY was demonstrated by Osumi and Saier (624), who also found that the binding of EIIA^Glc correlates with the amount of transporter and is enhanced by the presence of lactose, while the phosphorylation of EIIA^Glc abolishes it. These results were confirmed and quantified by Nelson et al. (593, 597). Direct binding and inhibition by unphosphorylated EIIA^Glc were also established for the MalK component of the maltose transport system (163, 675) and for GlpK (166, 676).

The uptake systems for raffinose (876), galactose (559), and arabinose (338) are also subject to inducer exclusion. Raffinose uptake via the plasmid-encoded raffinose permease RafB is inhibited by α-MG, and the effect is enhanced in “leaky” ptsI mutants (876). It was suggested that galactose transport in E. coli would also be under the control of inducer exclusion (7, 385). Indeed, the uptake of galactose by membrane vesicles containing galactose permease was reported to be diminished by the addition of unphosphorylated EIIA^Glc (559). Nevertheless, other experiments from the same group (562, 767) did not reveal an effect of the PTS on galactose fermentation by intact cells. The authors of those reports argued that the absence of an effect probably results from the full induction of the galactose permease in intact cells, thereby reducing the effectiveness of inhibition by EIIA^Glc, as inducer exclusion is mediated by a 1:1 stoichiometric interaction (559). Nevertheless, E. coli gal operon expression is submitted to CCR, as was demonstrated by the early experiments of Stephenson and Gale (836).

Efficient binding of unphosphorylated EIIA^Glc to the target protein occurs only when the substrate of the target is present. For example, EIIA^Glc binds strongly to the lactose transporter LacY only in the presence of a β-galactoside (597, 624, 800, 825). This mechanism ensures that EIIA^Glc, the concentration of which is fairly constant in E. coli and S. enterica serovar Typhimurium cells, will not be wasted on useless binding, i.e., under conditions in which the substrate is not present in the environment. The necessity of substrate binding indicates that the target proteins probably have to undergo a conformational change before unphosphorylated EIIA^Glc can bind. Evidence for conformational changes associated with substrate binding has been presented for LacY (245, 390), MelB (586), MalK (71, 581), and GlpK (355, 654, 874).

EIIA^Glc directly interacts with many different proteins as it serves in PTS phosphoryl transfer, inducer exclusion, and probably activation of adenylate cyclase (Table 1). Although most interaction partners are structurally unrelated, it appears that the binding surface of EIIA^Glc is always similar. The structure of EIIA^Glc is hardly affected by binding to its target proteins (98, 124, 270, 355, 929) or by phosphorylation (642). E. coli EIIA^Glc forms an antiparallel β-sandwich in which His-90 and His-75 of the active site are located off center on one face of the sandwich (224, 643, 955). Besides His-90, the residue that carries the phosphoryl group in E. coli P∼EIIA^Glc (197), His-75, appears to be very important for the phosphocarrier activity. Replacement of His-75 with Gln results in strongly diminished phosphoryl group transfer (686). A structural study of His75Gln mutant EIIA^Glc indicated no significant structural changes around the active site compared to the wild-type protein (643). Two possible explanations were proposed: (i) His-75 stabilizes the negative charge of P∼His-90, or (ii) the active site contains a proton relay network also involving Thr-73. The latter hypothesis was recently falsified by replacing Thr-73 with Ser, Ala, or Val and measuring the effect on the phosphotransfer rates to and from HPr (545). These rates were barely affected by the mutations.

The active site of E. coli EIIA^Glc is surrounded by a ring of solvent-exposed hydrophobic residues flanked by some polar and charged residues. A similar arrangement was found for the EIIA^Glc of Mycobacterium capricolum (349) and the EIIA domain of B. subtilis EIICBA^Glc (122, 474). The NMR structure of E. coli P∼EIIA^Glc shows that the phosphoryl group bound to His-90 (197) protrudes from the ring of hydrophobic residues that surround the active site (642). Supposedly, the disturbance of the hydrophobic binding surface of EIIA^Glc prevents the interaction between P∼EIIA^Glc and the target proteins. Furthermore, the introduction of a negative charge on the protein surface contributes to the diminished interaction.

The structures of B. subtilis HPr (330) and the EIIA domain of B. subtilis EIICBA^Glc (474) were used to construct a model of the HPr:EIIA^Glc transition complex (327). Herzberg concluded that (i) upon the formation of the transition complex, no major conformational change occurs and (ii) the central part of the binding surface consists mainly of hydrophobic residues. These assessments are sustained by the NMR solution structures of the HPr:EIIA^Glc transition complexes of B. subtilis (124) and E. coli (929). The transition complex perfectly accommodates the binding of a phosphoryl group by both the active-site histidine of EIIA^Glc and that of HPr in either an associative or dissociative mechanism. The active-site histidine of EIIA^Glc (His-90 in E. coli) is located in a shallow depression of a slightly concave surface (224, 643, 955), whereas the active-site histidine of HPr is located at the end of the first α-helix on a convex protrusion of the protein (330). The binding of the phosphoryl group in B. subtilis EIIA^Glc is stabilized by His-68 (His-75 in E. coli) (474) and by Arg-17 in HPr (330). In addition, two aspartyl residues are perfectly positioned to form alternate ion pairs with Arg-17 of HPr. The hydrophobic binding surface containing the active site of EIIA^Glc includes several valine and phenylalanine residues (Phe-41, -71, and -88 and Val-40, -46, and -96 in E. coli) (929).

It appears that the binding surfaces of EIIA^Glc for HPr and EIIB^Glc are almost identical (Table 2). The soluble EIIB domain of E. coli EIICB^Glc was isolated, and the solution structure of the EIIA^Glc:EIIB^Glc transition complex was determined (98, 270). The interaction surface consists of a central hydrophobic patch composed of the phenylalanine and valine residues mentioned above, and the same two aspartyl residues form alternate ion pairs with arginine residues. P∼EIIA^Glc transfers its phosphoryl group to Cys-421 of the glucose permease EIICB^Glc (92, 547, 618). This residue is located at the C terminus of β1 of the EIIB domain on a hydrophobic protrusion (203). Two arginine residues neutralize the negative charge added to Cys-421 upon phosphorylation of EIIB^Glc (98). Like Cys-421 (92), the presence of these residues was reported to be compulsory for effective glucose transport and phosphorylation (449). Although the central interaction parts are very similar, the residues forming the edges of the binding surfaces are different between the transition complexes HPr:EIIA^Glc and EIIA^Glc:EIIB^Glc (98, 929). The edges of the binding surface of HPr contain only positively charged residues, whereas those of EIICB^Glc include both positively and negatively charged amino acids. Consequently, the edges of the binding surface of EIIA^Glc for HPr include many negatively charged and some polar residues, while those of EIIA^Glc for EIIB^Glc include a polar residue and both negatively and positively charged amino acids.

Sequence comparisons of EIIA^Glc and EIIB^Glc from various bacteria reveal that the interfacial residues are strongly conserved (98). The few changes are mostly conservative, thus preserving the central network of intermolecular hydrophobic interactions. A higher variability is observed for the peripheral polar and charged residues. There are, for instance, several positively charged residues in E. coli EIIA^Glc replaced with polar or negatively charged residues in the B. subtilis protein. Although these substitutions abolish some salt bridges, their contribution to the stability of the complex appears to be moderate, as illustrated by the similar apparent K_m values for heterologous (PTS proteins from different organisms) and homologous phosphoryl transfer (98, 723). Point mutations in several well-conserved residues of EIIA^Glc also lower the phosphoryl transfer activity but only by between 20 and 70% (822). It was concluded from these observations (98) that multiple interactions contribute to the affinity between PTS protein partners.

The N terminus of EIIA^Glc is disordered in the structures of both the HPr:EIIA^Glc and EIIA^Glc:EIIB^Glc transition complexes and is therefore probably not involved in the protein/protein interactions. Nevertheless, Meadow et al. (538, 541) established that although the naturally occurring N-terminal truncation (the first seven residues are absent, and the truncated EIIA^Glc therefore migrates faster on nondenaturing gels and was called EIIA_fast^Glc) has no effect on phosphoryl transfer between HPr and EIIA^Glc, it reduces the phosphoryl transfer to EIICB^Glc by about 97%. Recent measurements by the same group confirm these observations with genetically truncated EIIA^Glc (deletion of the first 7 and 16 residues) (545). Simultaneously, they showed that phosphoryl group transfer to the soluble EIIB^Glc domain remains unaffected. Misko et al. (559) found that compared to intact EIIA^Glc, EIIA_fast^Glc has a lower affinity for LacY-containing membranes. These findings suggest that the N terminus plays a role in the recruitment of EIIA^Glc to the E. coli inner membrane. Indeed, Wang et al. (926, 928) showed that the N terminus interacts with anionic lipids isolated from E. coli, suggesting that the N terminus serves as a kind of membrane anchor.

The crystal structure of the E. coli EIIA^Glc:GlpK complex has been determined (355). GlpK with bound glycerol and ADP forms tetramers in which each GlpK subunit interacts with one EIIA^Glc molecule. Indeed, complete inhibition of glycerol kinase activity occurs only when at least four molecules of EIIA^Glc are present per GlpK tetramer (900). The structure of EIIA^Glc in the complex is very similar to that of EIIA_fast^Glc (955), except that residues 1 to 11 are in contact with the surface of GlpK. Like EIIA_fast^Glc, the EIIA domain of B. subtilis EIICBA^Glc lacks the seven N-terminal residues. Nevertheless, the protein complements an E. coli ptsHI-crr mutant defective in GlpK regulation (723). Therefore, the N-terminal residues cannot be very important for the inhibition of glycerol kinase (355). The inhibitory contact between the two proteins is mediated by a 3₁₀ helix of GlpK (residues 472 to 481), which protrudes into the concave surface of the active site of EIIA^Glc. The contact surface of EIIA^Glc is very similar to that in the EIIA^Glc:EIIB^Glc complex (Table 2). The hydrophobic residues surrounding the active-site histidine are important, and binding involves negatively and positively charged residues.

The organization of the residues around the active site of EIIA^Glc in the EIIA^Glc:GlpK complex suggests a possible role for Zn²⁺ in the association. Indeed, crystals soaked in a 20 mM ZnCl₂ solution adopted a Zn²⁺ ion at the expected position (223). Subsequent in vitro studies revealed that the presence of the metal ion lowers the K_i of EIIA^Glc for GlpK by about 60-fold (223, 616). The Zn²⁺ ion is liganded by His-75, His-90, and a water molecule in EIIA^Glc and by Glu-478 in GlpK. The stimulatory effect of Zn²⁺ on EIIA^Glc-mediated GlpK inhibition disappeared when Glu-478 was replaced with Cys, Asp, His, or Gln (657).

It is not known how the interaction with unphosphorylated EIIA^Glc inhibits glycerol phosphorylation. The more than 30-Å distance between the sites of EIIA^Glc binding and glycerol phosphorylation rules out a direct effect. A conformational change of glycerol kinase is part of the catalytic cycle, and so it was proposed that the binding of EIIA^Glc could lock the protein in a closed state (355). The lower inhibition by EIIA^Glc observed for some mutant GlpKs affected in the domain associated with “opening” and “closing” GlpK is consistent with this concept (655, 658).

LacY is an H⁺ symporter that uses electrochemical potential to drive the uptake of several galactosides such as lactose and melibiose (390, 391, 942). The recently resolved LacY structure confirms previous predictions that the protein contains 12 hydrophobic transmembrane domains connected by relatively hydrophilic loops (4). The major determinants of substrate binding are located at the interfaces of helices IV, V, and VIII (300, 444, 754). In contrast, the cytoplasmic loop between helices IV and V together with the flanking helical domains and some residues in the large cytoplasmic loop connecting helices VI and VII are important for the interaction of LacY with EIIA^Glc (338, 800, 823). Based on the analysis of various lacY mutants, a model for LacY function in galactoside transport has been proposed (3, 391, 942). LacY is assumed to have a conformation in which the sugar binding site is facing the periplasm (“outward”). The binding of the sugar induces a structural change, which exposes the substrate binding site to the cytoplasm (“inward”). To explain the observation that EIIA^Glc binds to LacY with higher affinity when its substrate is present (597, 624, 800, 825), the “outward”/“inward” transition is assumed to make the EIIA^Glc interaction site more easily accessible (800, 823). Evidence supporting this concept comes from studies of a Cys154Gly mutant LacY. The mutation abolishes both lactose transport and binding of EIIA^Glc and was originally thought to lock the protein in the “outward” conformation (819, 825, 902). However, the crystal structure of Cys154Gly LacY indicates that the protein is locked in an “inward” conformation (4). It was then proposed that EIIA^Glc binds to the “outward” conformation of LacY, thus preventing the structural change to the “inward” conformation and, consequently, the entry of the bound sugar. Other observations support this interpretation. When galactosides are present at saturating amounts, ¹²⁵I-labeled unphosphorylated EIIA^Glc binds in a 1:6 stoichiometry to wild-type LacY (in inside-out vesicles) (824, 825). The replacement of residues Ile-129 and Lys-131 (these residues are not located in the EIIA^Glc interaction site but are located on central helix IV, which is involved in substrate binding) with Cys allows more EIIA^Glc molecules to bind to LacY but does not affect the apparent equilibrium dissociation constant, K_D, of the EIIA^Glc/LacY complex. In addition, different sugars lead to different binding stoichiometries, with the preferred substrates melibiose and lactose causing maximum binding of EIIA^Glc. The data were explained by assuming that LacY exists in two (or more) populations: one population that binds EIIA^Glc (“outward”) and one that doesn't (“inward”). The observed EIIA^Glc binding stoichiometry was thought to reflect the ratio of these populations, which should be affected by the transported sugar (and in mutant LacYs by the conformational flexibility). According to the model, the binding stoichiometry should not affect the dissociation constant of the EIIA^Glc/LacY complex, which is consistent with the observations. It is noteworthy that in contrast to the data reported by Sondej et al. (824, 825), Nelson et al. (597) reported an EIIA^Glc/LacY binding stoichiometry of 1:1. The reason for the discrepancy is not clear.

The interaction surface of LacY and EIIA^Glc has been probed by analyzing the effect of various mutations. Residues of the central loop of LacY were found to be involved in the interaction (338, 951). Changing Thr-7 or Met-11 to Ile prevents EIIA^Glc from binding (597, 625). However, a deletion of residues 2 to 8 hardly affects the regulation of LacY by EIIA^Glc (338). Other residues of LacY involved in binding EIIA^Glc were identified by Cys-scanning mutagenesis (823). Although some of the LacY residues important for EIIA^Glc binding are conserved in MalK, MelB, and RafB, no consensus EIIA^Glc binding motif could be proposed (823). In contrast to the binding surfaces of other EIIA^Glc target proteins (HPr, EIIB^Glc, and GlpK), the LacY binding surface seems to contain only a small number of hydrophobic residues. Several E. coli EIIA^Glc (crr) mutants defective in inducer exclusion of LacY substrates have been identified (822, 987). Some of these residues are also important for the interaction of EIIA^Glc with HPr, EIIB^Glc, and GlpK (Table 2).

Similar to LacY, the melibiose permease MelB is an ion (Na⁺/H⁺) sugar symporter with 12 membrane-spanning segments (74, 586, 966). S. enterica serovar Typhimurium melB mutants synthesizing an inducer exclusion-resistant melibiose permease have been isolated (439) (Table 2). The mutated residues all lie on one side of an α-helical stretch situated in the cytoplasmic C-terminal region of MelB. Other substitutions located on the same α-helical stretch diminished inhibition of the permease by EIIA^Glc. Cutting off the last 22 amino acids of MelB reduces inducer exclusion by fourfold, and mutants lacking more residues exhibit low melibiose and methyl-α-thiogalactoside transport even in the absence of inducer exclusion.

Maltose uptake in E. coli requires the malE-encoded periplasmic maltose binding protein and the multisubunit ABC transporter MalFGK₂ (72, 226, 447). The soluble MalK subunits are tightly associated with the two permease subunits MalF and MalG (354, 580). Binding of maltose-carrying maltose binding protein to MalFGK₂ stimulates ATP hydrolysis by MalK and therefore maltose transport (118, 161, 187). Stimulation of ATP hydrolysis is cooperative, suggesting that the MalK subunits interact with each other (159, 160, 226). MalK possesses an N-terminal ATPase domain (present in all ATP-binding proteins of ABC transporters) and, in certain bacteria including E. coli and S. enterica serovar Typhimurium, a specific C-terminal regulatory domain (66, 187, 782, 828). In the presence of maltose and glucose, EIIA^Glc is assumed to bind to the regulatory domain, thus preventing maltose import. In addition, if no maltose is present, the C-terminal MalK domain is proposed to bind and inhibit the transcription activator MalT (380, 627, 828), which controls the expression of the mal regulon (65, 70).

Several malK mutants exhibiting weak inducer exclusion of maltose but normal inhibition of MalT by MalK and vice versa have been isolated (163, 431) (Table 2). In addition, the binding of monoclonal antibodies directed against linear epitopes extending from either amino acids 113 to 123 or 352 to 361 in the MalK sequence is diminished by the presence of EIIA^Glc (828). The scattered location of the mutations (Table 2) and the EIIA^Glc/antibody competition sites made it difficult to propose a binding site for EIIA^Glc (66, 828). In contrast to Thermococcus litoralis MalK (187), biochemical studies suggest that in the functional MalFGK₂ complex of E. coli, the C termini of the MalK subunits are in close contact (774). This assumption was confirmed by resolving the structures of three different forms of the MalK dimer (117). Several amino acids, the replacement of which leads to weak maltose exclusion, lie on the same face of the dimer, thus identifying a potential binding site for EIIA^Glc. The binding of EIIA^Glc to the nucleotide binding domain of one subunit and the regulatory domain of the other could well prevent the “tweezers-like motion” necessary to accommodate the cycle of hydrolysis and subsequent transport (117).

Using ligand fishing, yet another interaction partner of unphosphorylated E. coli EIIA^Glc was recovered (419). The protein was called fermentation/respiration switch protein (FrsA; formerly YafA) because frsA disruption causes increased respiration on several sugars including glucose, and overexpression results in increased fermentation. EIIA^Glc forms a 1:1 complex with FrsA. Thus, in E. coli, the balance between fermentation and respiration seems to be regulated by the PTS. Furthermore, the E. coli genome has been screened for proteins containing a potential EIIA^Glc binding site using sequence patterns deduced from mutagenesis studies of LacY (823). The search has yielded over 35 potential binding partners including HPr and MalK but excluding MelB and FsrA. Therefore, whether the recovered list of proteins is significant or not remains an open question.

The utilization of a PTS substrate, such as glucose, by enteric bacteria increases the amount of not only EIICB^Glc and the mannose-specific EIIs (215, 420, 769, 839) but also EI and HPr (172, 727, 771, 839). Unfortunately, a detailed analysis of transcription regulation of glucose-activated genes is complicated by the fact that most of them are also under the control of Crp/cAMP, i.e., partly repressed by the presence of glucose. For instance, cyaA or crp mutants have lower levels of EI, HPr, and several EIIs (727). On the other hand, E. coli wild-type cells grown on glucose exhibit increased levels of these proteins compared to cells grown on lactate. The relatively elevated synthesis in the presence of glucose appeared to require the activity of the repressor Mlc. Overexpression of the E. coli mlc gene leads to an increased colony size during growth on glucose-containing solid medium (hence the name mlc, for making large colonies), which is due to retarded glucose uptake and the reduced formation of acetate (343). This growth behavior led to the discovery of mlc in the first place.

Earlier studies of E. coli mutants that were affected in growth had already suggested that ptsG and ptsM expression might be controlled via a repressor. Suppressor mutants that were able to grow anaerobically on glucose, mannose, and glucosamine were obtained from E. coli ptsG strains, which cannot utilize the above-mentioned carbon sources in the absence of oxygen (732). The suppressor mutants were also 2-deoxy-d-glucose (2DG) sensitive under anaerobic conditions, and hence, the locus/gene was called dgsA. The toxic 2DG is transported mainly via EIIC^Man/EIID^Man, and indeed, after anaerobic growth, dgsA mutants exhibited elevated mannose and 2DG transport activities compared to the parental strain. DgsA was therefore assumed to be a repressor for the man operon. In fact, mlc was later found to be allelic with dgsA (662).

Strains overproducing Mlc (DgsA) were constructed, and the protein was purified (407, 857). It was found that the presence of a Zn²⁺ ion is essential for binding to the DNA (780). The amino acid sequence of the 44-kDa Mlc is about 40% identical to that of NagC (343, 579). NagC is a repressor/activator involved in N-acetylglucosamine uptake and metabolism (646, 667, 669, 922), and its activity is induced by N-acetylglucosamine-6-phosphate (668). Both Mlc and NagC belong to the ROK family, which contains repressors, open reading frames (ORFs), and kinases (314, 877). The DNA binding sequences of Mlc and NagC (663) and the putative sugar binding sites of Mlc and the structurally similar ROK family members glucokinase (GlcK) of E. coli and a putative fructokinase (FrcK) of B. subtilis (780) appear to be very similar. Nevertheless, in contrast to NagC, GlcK, and FrcK, Mlc does not seem to bind glucose or metabolites that are derived from it. Gel mobility shift assays showed that these molecules do not affect the interaction of Mlc with its various DNA target sites (167, 407, 408, 664). In addition, activation/deactivation of Mlc by phosphorylation seems unlikely, as no phosphorylation of Mlc has been detected (407).

While studying the expression of ptsHI-crr and ptsG, which are both regulated by Mlc, it became clear that the PTS controls Mlc activity (171, 172, 664, 665, 857). de Reuse and Danchin (171) concluded that transcriptional regulation of the pts operon is a consequence of an increase in the level of unphosphorylated EIICB^Glc. Plumbridge (665) then suggested that unphosphorylated EIICB^Glc might interact with Mlc (Table 1). Affinity chromatography demonstrated that Mlc indeed binds to unphosphorylated, but not to phosphorylated, EIICB^Glc (856). Mlc was found to be sequestered to the membrane when P∼EIICB^Glc was dephosphorylated during the uptake of glucose (458, 588). In addition, the overproduction of EIICB^Glc resulted in the derepression of the Mlc-controlled operons ptsG and ptsHI-crr (588). Finally, surface plasmon resonance experiments and mobility shift assays confirmed that Mlc binds to EIICB^Glc but not to EI, HPr, or EIIA^Glc (588). The affinity of Mlc for unphosphorylated EIICB^Glc (K_D = 10⁻⁷ M) and for its various DNA target sites (K_D = 10⁻⁸ to 10⁻⁷ M) is similar (588).

Initial studies with mutant EIICB^Glc carrying various deletions suggested that Mlc binds to the EIIB domain only when the EIIC-EIIB linker (residues 320 to 390) is present (458). Certain mutations in the EIIC domain interfere with Mlc-mediated regulation (615, 661, 988). In addition to dgs mutants, another class of mutants that also exhibited a phenotype resembling that of mlc strains was isolated. E. coli manXYZ strains grow only poorly on glucosamine, which is also slowly transported via EIICB^Glc. Starting from a manXYZ strain, several mutants that were able to grow efficiently on glucosamine were obtained. The mutants, which synthesized EIICB^Glc constitutively, were called umgC (uptake of α-methylglucoside control), and the umgC mutations were mapped close to the ptsG gene (383). UmgC was hence originally postulated to be a repressor for ptsG. Investigation of the original mutants and some new umgC mutants revealed that these mutations actually map within ptsG (615, 661, 988). Interestingly, some of these mutations affect the putative cytoplasmic loops in the EIIC^Glc domain and cause an up to fivefold increase in α-MG uptake rates compared to wild-type cells as well as elevated ptsG mRNA levels under noninducing conditions. The effects of umgC mutations resemble those found in mlc null mutants (615), and it was therefore assumed that they increase the affinity of EIIB^Glc for Mlc. The binding of Mlc to EIIB is very sensitive to changes in the surface-exposed residues in the vicinity of the phosphorylatable Cys (795), and specific mutations in the EIIC domain possibly affect the arrangement of these residues in the EIIB domain, thereby affecting the binding of the transcription factor. However, the purified EIIB domain alone was found to be sufficient to bind Mlc, but anchoring to the membrane is necessary to prevent Mlc-mediated repression (795, 855). Nevertheless, other effects resulting from the membrane sequestration, like the induction of conformational changes or the masking of binding domains, might contribute to the regulation of Mlc (855).

Mlc has been crystallized, and its structure has been solved (780). The protein appears in two distinct homodimeric forms in the crystallographic unit, and the difference in the distances of the helix-turn-helix (HTH) DNA binding domains between the two forms suggests that only one should be able to bind to the DNA. The helix-turn-helix DNA binding domain of Mlc is stabilized by the C-terminal helix of the protein, and this structural feature seems to be crucial for Mlc function (780). Although deletion of the first 9 C-terminal amino acids does not affect Mlc activity, deletion of the first 18 C-terminal amino acids prevents repression and EII binding (795). The loss of function is accompanied by a transition from the tetrameric form, which is found in dilute solutions (588), to a dimeric form (795).

DNase I footprinting experiments uncovered Mlc binding sites in the regulatory regions of five operons/genes in E. coli. They include ptsHI (407, 665, 857); ptsG (408, 664); manXYZ (662); malT, the positive regulator of the maltose regulon (167); and mlc itself (167). Transcription of these operons/genes is repressed by the binding of Mlc to a site overlapping the downstream promoter (except for the pts operon, where Mlc binds close to P₀) and is stimulated by Crp/cAMP, which binds upstream from Mlc. To find out to what extent Mlc and Crp/cAMP contribute to the regulation of the operons, expression of lacZ fusions in wild-type strains was compared to that in strains devoid of adenylate cyclase, Crp, Mlc, or EIICB^Glc.

A null mutation of the mlc gene caused a threefold increase in manX expression in glycerol-grown cells, and a similar increase was observed for malT (167). Synthesis of a ManX-LacZ fusion protein is very low in a cyaA mutant and is increased 14-fold by the addition of cAMP. The control exerted by Crp/cAMP appears to be stronger than that of Mlc, as a fivefold-higher amount of ManX-LacZ is present in cells grown on glycerol than in cells grown on glucose (662).

Glucose-grown cells also exhibit expression of mlc that is about twofold lower than that of cells grown on glycerol (167, 808). Whereas transcription regulators are often associated with one of the transcription units that they control, mlc is monocistronic. The coding sequence of the gene is preceded by a Crp/cAMP binding site, two promoters (579), and an Mlc binding site overlapping the second promoter (167). Although both promoters are recognized by RNA polymerase containing the housekeeping sigma factor σ⁷⁰ (Eσ⁷⁰), P₂ is also recognized by the heat shock factor σ³² complex (Eσ³²) (808). In vitro transcription assays corroborated the finding that transcription is initiated from both promoters and that Crp/cAMP has a positive effect on P₂ in the presence of Eσ⁷⁰ but a negligible effect in the presence of Eσ³². Crp/cAMP inhibits transcription from P₁, and Mlc inhibits transcription from P₂. The synthesis of σ³² in response to a heat shock leads to the overexpression of mlc. Nevertheless, the Mlc-to-EIICB^Glc ratio remains fairly constant because ptsG transcription is also induced by Eσ³².

The EIICB^Glc-encoding ptsG gene is expressed from two promoters, which are under the control of Mlc (408, 664). One Mlc binding site overlaps P₁ and precedes a Crp/cAMP target site, and the other is upstream from P₂. Crp/cAMP-dependent expression from P₁ prevails, as P₂ transcripts amount to only 10% of the total ptsG mRNA (664). crp or cyaA null mutants do not express ptsG (408, 409, 727), while ptsG is up to 18-fold overexpressed in mlc null mutants (depending on the growth conditions) (373, 664). In wild-type cells, growth on glucose causes an eightfold induction of ptsG expression compared to growth on non-PTS carbon sources (215, 664). It follows that although affected by Crp/cAMP, the expression of ptsG is regulated mainly by Mlc. In addition, other factors have been implicated. It appears that the repression of ptsG transcription from promoters P₁ and P₂ by Mlc and the activation of transcription from promoter P₁ by Crp-cAMP are enhanced by Fis (807), one of the major histone-like proteins of E. coli (279). In addition, under conditions of nutrient limitation, ptsG expression is repressed by Eσ^s (RpoS), and this repression seems to act in synergy with that of Mlc (792). Furthermore, using ligand fishing with the promoter region of E. coli ptsG, it was found that the two-component response regulator ArcA, when phosphorylated, binds the promoter region at three positions: two between P₂ and P₁ overlapping the Crp/cAMP binding sites and one next to P₁ overlapping the Mlc binding site (373). P∼ArcA also binds to the promoter region of the ptsHI-crr operon but with lower affinity. The binding of P∼ArcA leads to a twofold reduction in ptsG transcription. The following link between the Arc two-component system and the glucose PTS might exist. An elevated glycolytic flux might cause a redox imbalance and hence lead to the activation of the response regulator ArcA, which is phosphorylated when reducing equivalents accumulate under aerobic conditions (271). P∼ArcA then binds to the ptsG promoter region and represses the transcription of ptsG (373), thus reducing the uptake of glucose and the glycolytic flux.

A different mode of regulation of the EIICB^Glc concentration is related to the stability of the RNA message. It was established that ptsG mRNA stability is reduced in an RNase E-dependent manner by metabolic blocks early in glycolysis (accumulation of glucose-6-P or fructose-6-P) (209, 410, 577, 578). Destabilization of the mRNA depends on enolase (578), a major component of the RNase E-containing degradosome, and the RNA chaperone Hfq (394). The chaperonin Hfq promotes the annealing of specific small RNA (sRNA) to target mRNA, thereby affecting the translation of the message (269, 842, 895). The ptsG mRNA is destabilized by an Hfq-binding sRNA called SgrS (898). Furthermore, expression of sgrS (formerly ryaA) is controlled by the transcription activator SgrR (formerly YabN) and becomes activated under conditions of α-MG-6-P (glucose-6-P) accumulation. However, Hfq-catalyzed annealing of the SgrS sRNA to ptsG mRNA alone is not enough to explain the observed instability of the ptsG mRNA. Kawamoto et al. (394) showed that the presence of the first two transmembrane helices of translated PtsG (i.e., localization near the membrane) is essential for an effective ptsG mRNA breakdown under conditions of hexose-6-P accumulation.

Transcription of the ptsHI-crr operon is initiated from three promoters (172, 173, 237, 753). P₀ and P₁ each yield two different transcripts: a short transcript coding for HPr only and a long transcript encompassing all three genes (172). The third promoter is located at the 3′ end of ptsI and allows crr transcription. Transcription from P₂ is independent of Crp/cAMP and Mlc (857) and accounts for about 80% of the total crr-containing mRNA (172). As a result, growth on glucose hardly affects the concentration of EIIA^Glc, whereas it increases the number of ptsH and ptsI transcripts three- to fourfold (171, 172, 727, 857). This induction is due to an increase in transcription from P₀. In the absence of glucose, Mlc binds to the P₀ promoter region and prevents the binding of the RNA polymerase (407, 665, 857). Although the presence of Crp/cAMP is essential for expression from P₀ (173, 665, 750, 857), this promoter is controlled mainly by Mlc (407, 857). The P₁ promoter is preceded by a second Crp/cAMP binding site, and expression from this promoter is weakly controlled by Crp/cAMP (173) but is not affected by Mlc (407, 857). Next to the Crp/cAMP is a FruR binding site (751), the inactivation of which has no effect on ptsHI-crr expression (665, 857). Recently, it was found that crr expression from P₂ is affected by the histone-like nucleoid-structuring protein (427), but the physiological relevance of this finding remains to be established.

Repression by Mlc and activation by Crp/cAMP are directly related to the phosphorylation state of EIICB^Glc and EIIA^Glc, respectively. As explained above, during growth on glucose, unphosphorylated EIIA^Glc and EIICB^Glc prevail, with the latter sequestering the repressor Mlc to the membrane. As a result, EIICB^Glc will be synthesized, whereas the amount of Mlc will decrease slightly due to the low amount of Crp/cAMP, which will further increase EIICB^Glc synthesis. Repression by Mlc is relieved not only by growth on glucose but also by growth on other rapidly metabolized PTS substrates such as N-acetylglucosamine and mannitol and to a lesser extent by growth on fructose and mannose (407, 661). Lactose, melibiose, and sucrose have no effect. Mlc repression is also affected by maltose, although maltose is not a PTS sugar. However, maltose degradation yields intracellular glucose and glucose-1-P. The import of maltose can therefore relieve Mlc repression inasmuch as intracellular glucose causes P∼EIICB^Glc dephosphorylation (678).

Many bacteria contain an EIIA^Glc-like protein or domain, but whether EIIA^Glc plays a regulatory role has been investigated for only a few organisms, such as B. subtilis, M. capricolum, Haemophilus influenzae, S. thermophilus, and S. coelicolor.

Similar to E. coli ptsI mutants, B. subtilis ptsI mutants are not able to grow on the non-PTS carbon source glycerol (265). However, in contrast to E. coli, inactivation of EIIA^Glc does not restore growth of the B. subtilis ptsI strain on glycerol (282). It will be discussed below that in gram-positive bacteria, EIIA^Glc does not interact with GlpK but that this enzyme is regulated via P∼His-HPr-mediated phosphorylation (158). In M. capricolum, glycerol kinase is also not inhibited by EIIA^Glc (992).

Sequencing of the H. influenzae genome revealed genes encoding EI, HPr, and EIIA^Glc (ptsHI-crr operon) as well as genes encoding an EIIBC^Fru-like protein (fruA) and a protein in which two FPr domains, each containing a phosphorylatable histidine (His-300 and His-424), are fused to EIIA^Fru (EIIA^Fru-FPr-FPr; fruB) (500, 717). In H. influenzae, only fructose, but not glucose, is taken up via a PTS, and a ptsI mutant can therefore ferment glucose (499). Interestingly, the development of competence, which in H. influenzae requires cAMP and Crp (105, 196), was lowered about 70-fold in a crr mutant (305) and 250- to 500-fold in a ptsI mutant (305, 499). The addition of extracellular cAMP restored the development of competence. It is likely that H. influenzae P∼EIIA^Glc stimulates adenylate cyclase activity in a manner similar to that observed in enteric bacteria. No EIICB^Glc exists in H. influenzae, and thus, the presence of glucose does not lead to direct dephosphorylation of P∼EIIA^Glc. Nevertheless, the uptake of fructose will lower the amount of P∼EI and P∼HPr and consequently will also lower the amount of P∼EIIA^Glc.

Although the main uptake system for glucose in S. coelicolor is GlcP, a non-PTS permease of the major facilitator superfamily (911), CCR by glucose was observed (21, 217, 530, 912). Glucose kinase (443, 911) and the accumulation of glycolytic intermediates (694) are implicated in mediating the effect. Although S. coelicolor contains EI, HPr, EIIA^Glc (designated EIIA^crr), and EIIBC^Glc homologs as well as other sugar-specific EII complexes, PTS proteins do not seem to be involved in CRR in this high-G+C gram-positive organism (58, 629). In contrast to E. coli, the crr gene of S. coelicolor precedes ptsI, and ptsH is located somewhere else on the chromosome. EIIA^crr restores growth of an E. coli crr strain on glucose (392). HPr from S. coelicolor is efficiently phosphorylated by B. subtilis EI and PEP but is barely modified by B. subtilis HprK/P and ATP (97, 630). In fact, S. coelicolor is missing HPrK/P, and the deletion of ptsH has no effect on the glucose repression of galactokinase (97) or glycerol kinase (612). The absence of fluctuations in the cAMP concentration (112) restricts the potential role of the PTS to inducer exclusion-mediated mechanisms. However, although expression of the S. coelicolor crr gene in an E. coli ptsHI-crr deletion strain significantly reduces maltose uptake and although a specific interaction between E. coli MalK and S. coelicolor EIIA^crr was established by using surface plasmon resonance spectroscopy (392), no evidence showing that the PTS protein exerts a similar function in S. coelicolor has been obtained.

PEP-dependent phosphorylation of EI was originally thought to be the only route to generate P∼His-HPr (435, 436). However, two decades after the discovery of the PTS, PEP-independent EI phosphorylation was discovered (236). E. coli acetate kinase (AckA) and [γ-³²P]ATP phosphorylate EI in a reversible reaction (236). When catalyzing the formation of acetyl∼P from acetate and ATP, AckA is transitorily phosphorylated on a glutamyl residue (238, 881), probably at Glu-387 in E. coli AckA (817). PEP and P∼AckA most likely phosphorylate the same histidyl residue in EI (16), as the phosphoryl group of P∼AckA can be passed to EIIA^Glc via EI and HPr. A second protein kinase that also phosphorylates EI at the expense of ATP was found in E. coli (154, 155). It was partially purified, but its activity vanished during purification due to the loss of a cofactor, which was shown to be NAD(P)⁺. The reaction catalyzed by this EI kinase is reversible; i.e., it can transfer the phosphoryl group of P∼EI back to ADP, and the backward reaction requires the presence of NAD⁺. In its absence, EI kinase acts as a phosphatase and converts P∼EI to EI and P_i. It is not yet clear whether phosphorylated EI kinase is transitorily formed during P∼EI dephosphorylation. Although AckA- and EI kinase-mediated phosphorylation of EI have been demonstrated in vitro, whether they have physiological importance remains to be shown.

Motile bacteria such as E. coli and B. subtilis can be attracted to or repelled from various chemical and physical environmental signals. These signals can be processed via different pathways (see reference 11), but ultimately, they affect the frequency with which the rotation direction of the flagella changes. An increase in counterclockwise rotation results in smooth, straight movement of the bacterium (e.g., towards the attractants), whereas an increase in clockwise rotation leads to “tumbling” and a change in direction (e.g., away from the attractant) (see reference 501). The most important chemotaxis signal processing pathway comprises chemoreception in the cytoplasmic membrane by methyl-accepting chemotaxis proteins (MCPs) (for reviews on chemotaxis, see references 77, 82, 838, and 850). Although the number of proteins involved in the chemotactic signaling pathway varies in different bacteria, the main mechanism follows similar principles (850). The chemotaxis pathway of enteric bacteria is displayed in Fig. 3. An incoming chemotactic signal is relayed to the flagellar motor via the CheA/CheY two-component system. The autophosphorylation activity of the ATP-dependent histidine kinase CheA is stimulated by its CheW-mediated recruitment to the MCPs. P∼CheA transfers its phosphoryl group to an aspartyl residue of CheY, the response regulator. P∼CheY binds to the flagellar motor protein FliM, and as a consequence, the tumbling frequency increases. To prevent the accumulation of P∼CheY, the protein has autophosphatase activity, and in various bacteria (including E. coli), dephosphorylation is stimulated by a protein called CheZ. This signal transduction pathway is controlled by an adaptive feedback mechanism involving MCP methylation by the methyltransferase CheR, which stimulates CheA activity, and demethylation by the methylesterase CheB, which inhibits CheA activity. CheB is activated by phosphoryl transfer from P∼CheA. Upon chemoreception, the MCP lowers the activity of CheA considerably, and as a result, the concentration of P∼CheY decreases. The bacterium therefore tumbles less frequently and thus effectively moves up a concentration gradient of chemoattractant. Simultaneously, receptor demethylation diminishes due to lower levels of P∼CheA/P∼CheB, and CheA will be reactivated, leading finally to an adaptation of the system.

E. coli shows chemotaxis towards many nutrients, including several PTS sugars (8, 462). By using a variety of null mutants, it was established that the chemotrophic response towards PTS sugars was elicited by their transport and not by their binding or metabolism (8, 297, 462, 641, 948). It was established that chemotaxis towards PTS sugars at relatively high concentrations (>1 mM) is also mediated by the chemoreceptors Aer and Tsr (296), which sense redox state and proton motive force, respectively (865). In E. coli, chemotaxis by PTS sugars can be abolished by knocking out CheA, CheY, CheW, EI, HPr, or the corresponding sugar-specific EII complexes, whereas the removal of the MCPs, CheR, or CheB has no significant effect (8, 297, 460, 462, 498, 550, 609, 641, 745, 863, 864; see also reference 678). Despite the fact that PTS-mediated sugar uptake and Che protein-mediated chemotaxis involve similar phosphoryl transfer reactions, no phosphoryl transfer between PTS components and Che proteins has been demonstrated (379). Rather, in vitro experiments established that unphosphorylated EI interacts with CheA and thereby inhibits the autokinase activity of CheA by up to 10-fold (497). In addition, CheA activity drops about threefold upon shifts of the PEP concentration in the physiologically relevant range (from 2 to 0.3 mM). Further evidence for an effect of the phosphorylation state of PTS proteins on CheA activity was provided by kinetic analyses of the chemotactic response in E. coli wild-type cells and several che null mutants (498). The response, which was induced by flash photolysis of caged d-glucose and α-MG, shows rapid kinetics and high signal sensitivity (K_m for d-glucose is about 10 nM). Considering the rapidity and sensitivity of the response, those authors proposed that transport-induced dephosphorylation of P∼EI affects CheA activity but has little influence on the PEP concentration (the intracellular concentration of PEP is about 200-fold higher than that of EI). This hypothesis holds only when phosphorylation of EI by PEP would be a slow process controlling glucose influx. However, because even at high glucose concentrations, nearly all control over the flux resides in the glucose transporter (901), it is highly unlikely that EI phosphorylation exerts control at low glucose concentrations. Rather, PTS model calculations predict that the extent of EI phosphorylation rapidly responds to changes in the PEP concentration (239, 240, 777). At high intracellular PEP concentrations, mainly P∼EI and little EI have been calculated to be present. Thus, even a slight reduction in the PEP concentration (as anticipated for the chemotaxis experiments) (497) could cause a relatively significant increase of unphosphorylated EI, which would lower CheA activity. This could in turn slightly lower the amount of P∼CheY and lead to a strong chemotactic response, as the binding of P∼CheY to the flagellar motor is highly cooperative (133).

In contrast to enteric bacteria, chemotaxis towards PTS carbohydrates in B. subtilis is mediated by the methyl-accepting chemotaxis protein McpC, which is also a sensor for several amino acids (582). Deletion of mcpC abolishes chemotaxis towards the carbohydrates d-glucose, d-mannitol, d-fructose, trehalose, N-acetylglucosamine, and α-methyl-d-glucoside (261, 428). Nevertheless, chemotaxis in B. subtilis is affected by the PTS, and the cytoplasmic domain of McpC is thought to receive a transport-related signal from the PTS (428). As a consequence, EII^Mtl, EII^Fru, and EII^Tre null mutations resulted in a loss of chemotaxis towards the related carbohydrate. Deletion of ptsH also prevents B. subtilis from moving up a sugar gradient of mannitol and fructose (261). The response towards glucose is more complex and involves McpA (313). A ptsH or mcpA deletion strain shows chemotaxis towards glucose (which is taken up via a non-PTS transporter by the ptsH mutant), whereas a double mutant does not (261). Binding studies with purified CheA and EI indicate that P∼EI, but not EI or PEP, binds CheA and thereby inhibits its autophosphorylation. In B. subtilis, P∼CheY exerts an effect opposite that in E. coli (61, 62, 945) (i.e., P∼CheY leads to smooth swimming), and therefore, the inhibition of CheA activity by phosphorylated EI in B. subtilis is consistent with CheA inhibition by unphosphorylated EI in E. coli. However, experimental findings with tethered cells raise doubts about the proposed mechanism. For instance, the addition and removal of mannitol had effects similar to those of the addition of glucose on the frequency of counterclockwise rotation in a ptsH mutant (261), although this mutant does not transport mannitol. It is not clear how the phosphorylation state of the PTS proteins could be affected by a sugar that is not transported and metabolized. In addition, conflicting results were obtained when the effect of glucose on the rotation of tethered cells was studied (261, 428).

Many bacteria, including E. coli and S. enterica serovar Typhimurium, store carbohydrates in the form of glycogen. They produce it from glucose-1-P by the sequential action of ADP-glucose pyrophosphorylase (GlgC) and glycogen synthase (GlcA) (see reference 683). When needed, glycogen is reconverted to glucose-1-phosphate by glycogen phosphorylase (GlgP) (962). The related E. coli genes are located in the glgCAP operon, which is positively regulated by Crp/cAMP (684) and negatively regulated by CsrA (38, 486, 962). B. subtilis possesses a glgBCDAP operon (glgB codes for a glycogen branching enzyme, and glgD codes for a second ADP glucose pyrophosphorylase), which is presumably negatively regulated by CcpA/P-Ser-HPr (178). Surface plasmon resonance-based ligand fishing revealed a strong interaction between E. coli glycogen phosphorylase (GlgP) and HPr; indeed, GlgP was the only protein “fished” out of a cell extract by immobilized HPr (799). The interaction between the two proteins was confirmed by mobility shift assays and sedimentation equilibrium centrifugation. Subsequent competition experiments established that the binding was highly specific. Neither B. subtilis and Mycobacterium capricolum HPrs nor E. coli NPr and diphosphoryl transfer proteins (FPr) bind to E. coli GlgP, and vice versa, E. coli maltodextrin phosphorylase, which is very similar to GlgP (352, 797), does not bind HPr (418, 799). The affinity of GlgP for P∼His-HPr is four times higher than that for unphosphorylated HPr. Binding of HPr stimulates the formation of GlgP dimers and tetramers. Only the binding of HPr, but not of P∼His-HPr, leads to a significant increase in GlgP activity (2.5-fold) (799). The concentration of HPr (531) is much higher than that of GlgP (116), and therefore, a major part of cellular GlgP will be complexed with HPr or P∼His-HPr. As a result, GlgP activity will be determined by the phosphorylation state of HPr. Conversely, overproduction of GlgP should lead to the sequestration of most HPr and thereby to the inhibition of the PTS, as was indeed observed (418).

Based on these results, the following model for the regulation of glycogen breakdown was proposed for E. coli. Cells growing on glucose start to accumulate glycogen during the late exponential growth phase and the onset of stationary phase (418, 683, 684). Under these conditions, the concentration of phosphorylated PTS proteins is relatively high (797). cAMP levels will therefore rise (653), and expression of the glgCAP operon will be stimulated. The activity of GlgP is low, because mainly P∼His-HPr is present, and glycogen will accumulate. Later, when growth of the cells has advanced far into stationary phase, the PEP concentration drops, PTS proteins will become dephosphorylated, and the cAMP concentration and expression of the glgCAP operon will decrease. However, GlgP is now activated by binding unphosphorylated HPr, and glycogen breakdown will therefore commence and dominate over glycogen synthesis.

Isocitrate dehydrogenase from E. coli was the first bacterial protein shown to be phosphorylated by an ATP-dependent seryl protein kinase (257). In the early 1980s, when Deutscher, Reizer, and Saier investigated a possible implication of protein phosphorylation in inducer expulsion (185, 710, 712), a process where certain sugar phosphates that have accumulated in bacteria are dephosphorylated and expelled as soon as the cells are exposed to rapidly metabolizable carbon sources such as glucose or mannose, a second seryl-phosphorylated bacterial protein was detected in Streptococcus pyogenes and was identified as being HPr of the PTS (185). Unlike PEP-dependent EI-catalyzed phosphorylation at the catalytic His-15 of HPr, which occurs in both gram-negative and gram-positive organisms (Fig. 1) (262, 941), phosphorylation of HPr at a seryl residue, which requires a specific protein kinase and ATP (185) or GTP (709), was originally believed to be restricted to gram-positive organisms with low G+C content.

The site of ATP-dependent phosphorylation in HPr of Enterococcus faecalis was identified as being Ser-46 (183). An equivalent of Ser-46 is present in HPr of all low-G+C gram-positive bacteria, and the surrounding sequence is usually well conserved (Fig. 4). However, in the HPr of some organisms, the phosphorylatable serine is in a slightly different position (position 45 in HPr of Bacillus thuringiensis subsp. israelensis [400] and position 47 in HPr of Mycoplasma genitalium [713]). Replacement of Ser-46 in B. subtilis HPr with an alanine, tyrosine, or aspartate prevents the FBP-stimulated ATP-dependent phosphorylation of HPr (206, 722), whereas HPr carrying a threonine at position 46 is still slowly phosphorylated (722). Mutations at Ser-46 also affect the PEP-dependent, EI-catalyzed phosphorylation at His-15. In a mutant complementation assay carried out with rate-limiting amounts of B. subtilis HPr (20 μM), the phosphoryl carrier activity was lowered to 35, 30, and 20% when Ser46Ala, Ser46Tyr, or Ser46Thr HPr, respectively, was used in place of wild-type HPr (206) and was lowered to 10% when Ser46Asp HPr was used (722). Phosphorylation of HPr at Ser-46 lowered the rate of PEP-dependent phosphorylation by 2 orders of magnitude (177). In addition, the apparent K_m value for the PEP-dependent phosphorylation of P-Ser-HPr was 15-fold higher than the apparent K_m value determined with HPr (723). In E. coli HPr, Ser-46 is located within a hydrophobic patch implicated in the interaction with EI (259) and EIIAs (139, 691, 949). The fact that Ser-46 of HPr faces the well-conserved Glu-84 of EI in the HPr:EI complex (260) probably explains why HPrs of gram-positive bacteria carrying a negative charge at Ser-46, due to either phosphorylation or mutation, interact poorly with EI. Vice versa, phosphorylation of HPr by EI at His-15 slowed its phosphorylation at Ser-46 by the ATP-dependent HPr kinase (177). In the HPr:HPr kinase complex, helix α4 of HPr kinase interacts with the region around His-15 (228, 533). Phosphorylation at His-15 probably diminishes the affinity between the kinase and its protein substrate. Indeed, in two-hybrid experiments, an interaction of B. subtilis HPr kinase with Ser46Ala mutant HPr could be detected. This interaction was prevented when EI was additionally produced in the yeast cells, suggesting that EI efficiently competes with HprK/P for binding to HPr and/or that EI phosphorylates HPr at His-15, which lowers its affinity for HprK/P (P. Noirot, S. Poncet, and J. Deutscher, unpublished results).

Gram-negative enteric bacteria do not possess HPr kinase, although their HPrs usually contain Ser-46 and exhibit about 45% overall sequence identity to HPrs from gram-positive organisms. Nevertheless, the region around Ser-46 is quite different. This difference is probably responsible for the failure of HPr kinase from gram-positive bacteria to phosphorylate E. coli HPr (177, 722). HPrs of Mycoplasma capricolum, in which amino acids next to Ser-46 were replaced with the corresponding amino acids of E. coli HPr, were poorly phosphorylated with ATP (991). Based on the solution and crystal structures of HPrs from E. coli, E. faecalis, and B. subtilis, the presence of a lysine in position 49 of E. coli HPr, which is a glycine in HPrs of most gram-positive bacteria (Fig. 4), was assumed to represent a major obstacle for the interaction of HPr kinase with HPr from E. coli and presumably with HPrs from other gram-negative bacteria (374).

Interestingly, several gram-negative nonenteric organisms possess an HPr kinase, and the sequence around Ser-46 of their HPrs resembles that of HPr from low-G+C gram-positive bacteria (64) (Fig. 4). In contrast, Streptomyces coelicolor, a gram-positive organism with high G+C content in its DNA, does not possess a protein exhibiting significant sequence similarity to HPr kinase (55), and the sequence around Ser-47 of its HPr barely resembles the sequence around Ser-46 in HPrs of low-G+C gram-positive bacteria. Nevertheless, HPr from S. coelicolor can be phosphorylated by the HPr kinase from B. subtilis (97, 630), suggesting that despite the sequence differences, the structure around Ser-47 sufficiently resembles the structure around Ser-46 in B. subtilis HPr to allow its phosphorylation.

A soluble enzyme that specifically dephosphorylated P-Ser-HPr was purified to near homogeneity from E. faecalis (181). It had a molecular mass of 30 kDa (on sodium dodecyl sulfate-polyacrylamide gels), but it escaped the authors' attention that the purified enzyme must have also exhibited HPr kinase activity. In fact, it was later discovered that similar to the first described bacterial serine protein kinase, isocitrate dehydrogenase kinase/phosphatase (450), HPr kinase is also bifunctional and possesses P-Ser-HPr dephosphorylation activity (423). HPr kinase of E. faecalis has a molecular mass of 33 kDa, which is close to the molecular mass detected for the major protein in the P-Ser-HPr phosphatase preparation. Inorganic phosphate (P_i) was reported to stimulate P-Ser-HPr dephosphorylation (181). This was surprising, as P_i was assumed to be one of the products of P-Ser-HPr dephosphorylation and was therefore rather expected to inhibit this reaction. The paradox was resolved by demonstrating that P-Ser-HPr dephosphorylation is not a hydrolysis reaction. Dephosphorylation of either ³²P-Ser-HPr in the presence of P_i or P-Ser-HPr in the presence of ³²P_i leads to the formation of radioactive ³²PP_i. P_i thus functions as a substrate, and lowering the P_i concentration below the μM range almost completely prevents the dephosphorylation of P-Ser-HPr (557). In analogy to the usual phosphohydrolysis reaction, this novel protein dephosphorylation mechanism was called phospho-phosphorolysis, and the responsible enzyme was termed HPr kinase/P-Ser-HPr phosphorylase (HprK/P). Because P-Ser-HPr dephosphorylation is reversible, PP_i can replace ATP as the phosphoryl donor for HPr phosphorylation. The kinetic parameters indicate that PP_i-dependent HPr phosphorylation is actually the energetically favored reaction (557). To allow the efficient dephosphorylation of P-Ser-HPr in the cell, the resulting PP_i therefore needs to be hydrolyzed. In B. subtilis and other bacilli, PP_i hydrolysis seems to be achieved by YvoE, which exhibits pyrophosphatase activity (557). The yvoE gene is located in the hprK operon of bacilli (Fig. 5), and since YvoE exhibits sequence similarity to P-glycolate phosphatases and stimulates P-Ser-HPr dephosphorylation, it was mistakenly assumed to be P-Ser-HPr phosphatase (251). It is likely that organisms missing a yvoE gene in the hprK operon use one of usually several YvoE homologs present in bacteria to hydrolyze PP_i formed during P-Ser-HPr dephosphorylation.

Cold-chase ATP labeling of HPr in Listeria monocytogenes crude extracts suggested that HprK/P of this organism cannot dephosphorylate P-Ser-HPr (129). However, later experiments with purified HprK/P established that L. monocytogenes HprK/P functions as P-Ser-HPr phosphorylase, similar to the B. subtilis enzyme (326).

Interestingly, although Mycoplasma pneumoniae hprK mutant cells cannot form P-Ser-HPr, crude extracts prepared from this mutant were still able to dephosphorylate P-Ser-HPr. The dephosphorylation activity could be attributed to a PP2C-type P-protein phosphatase, which, in accordance with its B. subtilis homolog, was called PrpC (306). M. pneumoniae HprK/P is unusual, as it can phosphorylate HPr at relatively low ATP concentrations (831). In addition, in vivo P-Ser-HPr formation is stimulated by the presence of glycerol in the growth medium (307). Although the M. pneumoniae enzyme was reported to dephosphorylate P-Ser-HPr in vitro in a P_i-dependent manner, similar to HprK/Ps from other organisms (831), the P-Ser-HPr dephosphorylation activity exhibited by PrpC seems to be of physiological relevance. While an M. pneumoniae wild-type strain grown in the presence of glucose and glycerol contained only low amounts of P-Ser-HPr and doubly phosphorylated HPr, more than half of the HPr was found to be present as P-Ser-HPr and doubly phosphorylated HPr in the prpC mutant (306). Because a B. subtilis strain synthesizing Val267Phe HprK/P, which still functions as a kinase but has lost its phosphorylase activity (see “P-Ser-HPr Regulates PTS Transport Activity by a Feedback Mechanism”), contains more than 95% of its HPr as P-Ser-HPr (571), it is unlikely that PrpC of this organism plays a major role in P-Ser-HPr dephosphorylation. In B. subtilis, PrpC and the corresponding protein kinase, PrkC, were reported to play important roles in stationary-phase cells and to control the phosphorylation state of the translation elongation factor EF-G (248). It is interesting that HprK/Ps of certain gram-negative bacteria such as Neisseria meningitidis (S. Poncet, M.-K. Taha, M. Larribe, and J. Deutscher, unpublished results) and Brucella melitensis (S. Poncet, M. Dozot, X. de Bolle, J. J. Letesson, and J. Deutscher, unpublished results) exhibit no or very low P-Ser-HPr dephosphorylation activity, and although these organisms do not contain a homolog of PrpC, it is possible that they possess another P-Ser-HPr phosphatase.

The crystal structures of truncated HprK/P from L. casei (missing the N-terminal 120 amino acids) (229) and of full-length HprK/Ps from Staphylococcus xylosus (515) and M. pneumoniae (13) have been determined. The crystallized proteins form hexamers composed of two layers of trimers. Size exclusion chromatography and equilibrium sedimentation showed that L. casei HprK/P also forms hexamers in solution (229). B. subtilis HprK/P was reported to form hexamers at a neutral pH but was reported to form monomers and dimers at pH 9.5 (697). HprK/Ps do not exhibit similarity to eukaryotic protein kinases (312, 866) or P-protein phosphatases (43, 805) but resemble PEP carboxykinase and nucleosidediphosphate kinases (229, 252, 748). A P loop (or Walker motif A [GXXGXGKS]) that is usually located between amino acids 150 and 170 serves as a nucleotide binding site. The crystal structures of L. casei and S. xylosus HprK/Ps revealed that P_i binds to the same position in Walker motif A as the β-phosphate of the nucleotide. This explains the inhibitory effect of P_i on the kinase activity, as ATP and P_i compete for the same binding site. Vice versa, ATP inhibits the phosphorylase activity of HprK/P (181). The inhibitory effect of ATP was stronger at low Mg²⁺ concentrations. Mutant studies with hprK from various organisms suggested that Walker motif A is important not only for the kinase but also for the phosphorylase function (315, 571, 831).

The dual role of Walker motif A was confirmed by solving the crystal structures of HprK/Ps complexed with HPr or P-Ser-HPr, which also allowed the prediction of a detailed mechanism for HPr phosphorylation and P-Ser-HPr dephosphorylation (228). His-140 of L. casei HprK/P forms a hydrogen bond to Asp-179, which in turn acts as a base during HPr phosphorylation by forming a hydrogen bond to the hydroxyl group of Ser-46 in HPr. The structure of the HprK/P:P-Ser-HPr complex, which was obtained by cocrystallizing HPr, HprK/P, and PP_i, allowed the identification of the amino acids cooperating in the nucleophilic attack of P_i on the P-Ser bond and revealed that the binding of HPr and P-Ser-HPr to HprK/P involves nearly identical interfaces. Compared to the HprK/P:HPr complex, an additional contact is made between the phosphoryl group in P-Ser-HPr and Arg-245 of a neighboring HprK/P subunit (228). The loop containing Arg-245 is too flexible to be traced in the HprK/P and HprK/P:HPr structures, but it is stabilized in the HprK/P:P-Ser-HPr complex due to the interaction of Arg-245 with the phosphoryl group (228, 229). During P-Ser-HPr dephosphorylation, Asp-179 (present in protonated form) and His-140 (donates a proton to Ser-46 of HPr) (228) play an inverse role compared to the kinase reaction. The interaction site of S. aureus HPr with S. xylosus HprK/P was also determined by NMR, and the HPr interface appears to be similar in solution and in the crystal (533).

The crystal structure of HprK/P from S. xylosus revealed that a P_i ion binds not only to Walker motif A but also to the N-terminal domain (515). It was fixed by three conserved arginines located in two different subunits, one in the upper and one in the lower HprK/P trimer. The second P_i therefore bridges the two trimers (515), but the P_i bridge is not essential for hexamer formation, as truncated L. casei HprK/P (229) and M. pneumoniae HprK/P without bound P_i (13) also form hexamers. The function of the N-terminal domain, which protrudes from the central part like a propeller, is unknown. Truncated L. casei HprK/P that is missing this domain is active as a kinase and phosphorylase and responds to all known effectors (229). As will be discussed below, the N-terminal domain is naturally truncated in HprK/Ps of α-proteobacteria. Surprisingly, the N-terminal domain of HprK/P exhibits a fold similar to that of the N-terminal part of MurE (13), an enzyme implicated in the formation of cytoplasmic precursors for cell wall synthesis (284).

The structure of HprK/P with bound ATP or GTP has not yet been solved. If the nucleotide would occupy the same position as that in the closely related adenylate kinase, it would clash with the central K3 loop of a neighboring HprK/P subunit. Structural changes of ATP and/or HprK/P must therefore accompany nucleotide binding (13, 229, 515). Likewise, structural changes of HprK/P might be responsible for the observed cooperative binding of ATP (372).

About 15 years after the discovery of the ATP-dependent phosphorylation of HPr at Ser-46 (185), at almost the same time, three laboratories succeeded in identifying the HprK/P-encoding hprK gene (named ptsK in one publication) from three different organisms. In the first report, purification of E. faecalis HprK/P and sequencing of its N terminus and internal peptides allowed the amplification of the 5′ part of the hprK gene by PCR. This information was subsequently used to identify hprK of B. subtilis within the sequenced genome (251) and to obtain the complete hprK gene of E. faecalis (423). In the second report, purification and N-terminal sequencing of B. subtilis HprK/P was used to directly identify hprK of B. subtilis (709). Finally, purification of HprK/P and probing of a DNA library with a degenerate oligonucleotide derived from the N-terminal amino acid sequence allowed the identification of hprK from Streptococcus salivarius (84). More recently, the hprK genes from other low-G+C gram-positive bacteria including S. xylosus (357), L. casei (198), M. pneumoniae (830), and G. stearothermophilus (127) have been cloned and sequenced, and the corresponding HprK/Ps were purified and characterized. Genome sequencing revealed the presence of hprK in most low-G+C gram-positive organisms.

In the firmicutes, hprK is usually the first gene within an operon (Fig. 5), which is composed of five genes in B. subtilis. In most gram-positive bacteria, the second gene is the prolipoprotein diacylglyceryl transferase-encoding lgt gene, which carries out the diacylglyceryl lipidation of lipoproteins at a cysteyl residue. The frequent association of hprK with lgt might indicate that the functions of the two enzymes are somehow related, although no experimental evidence for this assumption has been obtained so far. A possible link between the PTS and Lgt is further suggested by the observation that in Enterobacteriaceae (E. coli, S. enterica serovar Typhimurium, Yersinia pestis, etc.), lgt is preceded by ptsP (T. Doan, personal communication), which encodes an EI with an N-terminal extension resembling the GAF domain in NifA. The hprK operon of G. stearothermophilus, Oceanobacillus iheyensis, L. monocytogenes, Listeria innocua, and the seven bacilli, whose genomes have been sequenced, contains the pyrophosphatase-encoding yvoE gene followed by yvoF downstream of lgt. B. subtilis, Bacillus clausii, Bacillus halodurans, Bacillus licheniformis, and G. stearothermophilus contain an additional gene, yvoD, inserted between lgt and yvoE. In S. aureus and Staphylococcus epidermidis, yvoE is missing, and yvoF directly follows lgt. The yvoDEF genes are absent from most other low-G+C gram-positive organisms. O. iheyensis possesses two genes encoding HprK/Ps, with one followed by lgt. It is not known whether the HprK/P ortholog encoded by the other hprK gene can phosphorylate HPr. In the genus Clostridium, hprK and lgt are located in different regions of the genome. In Clostridium acetobutylicum, hprK is preceded by a gene encoding a protein with significant sequence similarity to the glycerol-inducible E. coli GlpX, which resembles fructose-1,6-bisphosphatases (859).

Quite unique is a presumed hprK gene in Fusobacterium nucleatum ATCC 25586 and F. nucleatum subsp. vincentii, which encodes two complete HprK/P proteins fused together in tandem. The two HprK/P domains exhibit 22% sequence identity to each other and about 30% identity to HprK/Ps from gram-positive bacteria. However, a conserved Walker motif A is present only in the C-terminal domain. The corresponding sequence in the N-terminal domain is strongly altered and does not resemble Walker motif A. A prerequisite for HPr phosphorylation is the deprotonation of the hydroxyl group of Ser-46 of HPr by the concerted action of conserved histidyl and aspartyl residues (His-140 and Asp-179 in L. casei HprK/P). In F. nucleatum HprK/P, these residues are present in the C-terminal domain but are replaced with Glu and Lys, respectively, in the N-terminal domain. In contrast, Arg-245, which interacts with the phosphoryl group of P-Ser-HPr and probably plays a major role in P-Ser-HPr dephosphorylation, is present only in the N-terminal domain. It is therefore tempting to assume that in this duplicated enzyme, the two activities have been separated and that the C-terminal domain carries the kinase activity, whereas the N-terminal domain might function as P-Ser-HPr phosphorylase. The two sequenced F. nucleatum strains also contain HPr (FN1782) with conserved His-15 and Ser-46 regions and EI (FN1793), but although the genes are located close to each other on the chromosome, they do not seem to be organized in an operon.

The organization of the hprK gene in certain gram-negative bacteria will be discussed below in more detail.

In vitro experiments suggested that the two antagonistic activities associated with HprK/P are regulated in the cell in response to changes in the concentrations of FBP, ATP, and P_i (177). FBP clearly stimulates the kinase activity of B. subtilis HprK/P (372), whereas its effect is less pronounced with the L. casei enzyme (198). The effect of FBP is usually stronger when low, nonphysiological concentrations of ATP are used. In addition, the stimulatory effect of FBP on HPr phosphorylation was more evident when the experiments were carried out in the presence of a few mM P_i (177, 198, 423), which lowers the HPr kinase activity (84, 181, 251). The main physiological function of FBP might therefore be to prevent the inhibition of HprK/P by P_i. For the S. xylosus enzyme, activation by FBP was observed only when HprK/P was present at low concentrations (357). Surprisingly, the Streptococcus salivarius enzyme was not at all stimulated by FBP (84). Similarly, the M. pneumoniae enzyme is fully functional at relatively low ATP concentrations, and its activity is also not stimulated by FBP (831). The intracellular concentrations of FBP, ATP, and P_i vary largely depending on whether the cells utilize a favored carbon source or not. During the uptake of a rapidly metabolizable carbon source such as glucose, fructose, or mannose by L. lactis, the concentrations of P_i, ATP, and FBP are about 5, 8, and 30 mM, respectively (528, 598, 873). Under these conditions, HprK/P functions as an HPr kinase and not as P-Ser-HPr phosphorylase. In contrast, in the absence of a rapidly metabolizable carbon source, the concentration of P_i increases to about 50 mM, whereas the concentrations of FBP and ATP drop to 1 to 3 mM (528, 598, 873). The effect of growth on a preferred or less favorable carbon source on P-Ser-HPr formation was confirmed with S. salivarius cells. By carrying out crossed immunoelectrophoresis experiments, the fractions of the various forms of intracellular HPr in S. salivarius cells grown in glucose-containing medium were found to be 45% for P-Ser-HPr and 50% for doubly phosphorylated HPr, whereas only little P∼His-HPr (5%) and almost no dephospho-HPr could be detected (263). Cells grown on galactose, a less efficiently metabolized carbon source, also exhibited a high amount of doubly phosphorylated HPr (44%) and little P∼His-HPr (3%), whereas P-Ser-HPr was lowered to 18%, and dephospho-HPr increased to 35%. A similar observation was made for Streptococcus bovis, where an inverse correlation between P-Ser-HPr formation and the P_i concentration was observed. Cells growing on glucose contained more than half of the HPr as P-Ser-HPr, and the P_i concentration was in the low mM range. When glucose was exhausted, P-Ser-HPr disappeared, and the P_i concentration went up to 30 mM (26). In in vitro experiments, 20 mM P_i completely prevents the phosphorylation of HPr by 5 mM ATP, even when 25 mM FBP is present (177).

The observation that PP_i is formed during P-Ser-HPr dephosphorylation and used as a phosphoryl donor by HprK/P suggested that the PP_i concentration affects the intracellular amount of P-Ser-HPr. B. subtilis cells grown in the presence of glucose contain 10-fold more FBP than cells grown on succinate (from 1.4 to 14 mM). A similar increase was observed for PP_i (from 1.2 to 6 mM) (557). This metabolite distribution is probably the reason why about 65% of the HPr in glucose-grown B. subtilis cells is present as P-Ser-HPr, 35% is present as HPr, and little is present as P∼His-HPr and doubly phosphorylated HPr (571). In contrast, in succinate-grown cells, mainly dephospho-HPr was present, together with a few percent of P∼His-HPr. In in vitro experiments, HPr phosphorylation with B. subtilis HprK/P was maximal above 5 mM PP_i but negligible below 1 mM PP_i. The approximately 6 mM PP_i present in cells growing on glucose is therefore expected to be partly responsible for the increase of P-Ser-HPr, whereas in the absence of a rapidly metabolizable carbon source, the low PP_i and high P_i concentrations probably allow efficient P-Ser-HPr dephosphorylation. ATP-dependent HPr phosphorylation by B. subtilis HprK/P was maximal in the presence of 10 mM FBP, whereas it almost disappeared below 2 mM FBP (372). On the contrary, FBP was not required for efficient PP_i-dependent HPr phosphorylation (557).

An endogenous low-molecular-weight HprK/P inhibitor was detected when soluble extracts of Lactobacillus brevis were separated by size exclusion chromatography (715). However, no further details about the nature of the bacterial HPr kinase inhibitor were reported. Because the HPr kinase activity was detected by autoradiography after phosphorylation with [γ-³²P]ATP, it is possible that the reported HPr kinase inhibitor was PP_i. If the PP_i concentration in the assay mixtures exceeded the employed [γ-³²P]ATP concentration (10 μM) by severalfold, primarily nonradioactive (and therefore undetected) P-Ser-HPr would have been formed. Ironically, in the same publication, it was reported that the presence of PP_i can completely inhibit [γ-³²P]ATP-dependent HPr phosphorylation (715). Inhibition of [γ-³²P]ATP-dependent HPr phosphorylation by PP_i was also reported for HprK/P from Treponema denticola and was suggested to be probably due to the competition of the two phosphoryl donors for binding to HprK/P, as P-Ser-HPr was again detected by autoradiography (278). A synthetic inhibitor of HprK/P has also been identified. It is a heterocyclic bis-cationic compound, which was reported not to bind to the Walker motif (696).

In conclusion, the two opposing activities of HprK/P and consequently the ratio of the various HPr forms seem to be regulated mainly via metabolites. In agreement with this assumption, the amount of HprK/P present in B. subtilis cells was reported to have no influence on the ratio of the different HPr forms (59). The amount of HprK/P probably determines only how fast cells can adapt to changes in carbohydrate availability but apparently has no influence on the equilibrium reached under a certain condition.

Interestingly, in oral streptococci (glucose or lactose grown) (660, 893) and S. thermophilus (lactose grown) (134), a major part (up to 75%) of the HPr is converted into doubly phosphorylated HPr. A possible role of (P∼His,P-Ser)-HPr in the regulation of the lactose transporter LacS of S. thermophilus will be discussed below (see “Regulation of EIIA-Containing Non-PTS Transporters by P∼His-HPr-Mediated Phosphorylation”).

Heterofermentative lactobacilli such as L. brevis and Lactobacillus buchneri were reported to contain HPr and HprK/P but to be devoid of EI and EIIs (715). Owing to the presumed absence of a functional PTS, there seemed to be no obvious regulatory role for the ATP-dependent phosphorylation of HPr at Ser-46 in these organisms. Although the report about the absence of EI and EIIs from heterofermentative lactobacilli later turned out to be wrong (an L. brevis ptsHI operon has been cloned and sequenced) (193) and the presence of a fructose-specific PTS induced under anaerobic conditions has been established (772), it initiated the search for additional functions of P-Ser-HPr, one of them being the regulation of CCR.

A connection between the phosphorylation of HPr at Ser-46 and CCR was suggested by the observation that CCR in B. subtilis requires the metabolism of the repressing sugar (490, 606). In gram-negative bacteria, the transport of a carbohydrate via the PTS is sufficient to elicit CCR. By studying CCR in mutants defective at various steps of glycolysis, the number of glycolytic intermediates potentially serving as a signal for CCR in B. subtilis could be narrowed to four candidates: dihydroxyacetone phosphate, glyceraldehyde-3-phosphate, 1,3-diphosphoglycerate, and FBP (606). As FBP, the main activator of HPr kinase, was among these metabolites, ATP-dependent phosphorylation of HPr was suspected to play a role in CCR (184).

To test the proposed hypothesis that P-Ser-HPr plays a role in CCR, Deutscher and coworkers studied CCR in a B. subtilis ptsH1 mutant (Ser46Ala replacement). Indeed, several enzymes, including gluconate kinase, mannitol-1-P dehydrogenase, inositol dehydrogenase, and glucitol dehydrogenase, which are all sensitive to CCR in a wild-type strain, were not or only partly repressed by glucose and other rapidly metabolizable sugars in the ptsH1 mutant (184). Moreover, while the expression of a translational fusion (integrated at the amyE locus) of the promoter region and the 5′ part of gntK (encodes gluconate kinase) to lacZ was sensitive to CCR in a wild-type strain, it was not repressed by glucose in the ptsH1 mutant (184). In addition, if a B. subtilis strain carrying a deletion of the ptsGHI genes, which encode EIICBA^Glc, HPr, and EI, respectively, was transformed with a plasmid that allowed the production of the P-Ser-HPr-resembling Ser46Asp HPr (952), partial repression of the gnt operon was observed even in the absence of a repressing sugar (184). These results clearly established the crucial role of the phosphorylation of HPr at Ser-46 in CCR of B. subtilis.

The ptsH1 mutation turned out to be an important tool to study the implication of P-Ser-HPr in CCR in B. subtilis in more detail (184, 206). All B. subtilis ptsH1 mutants used in various laboratories are derivatives of ptsH1 strain SA003. They were obtained by either introducing additional mutations into SA003 or transforming specific mutants with chromosomal DNA of ptsH1 strains carrying an antibiotic resistance cassette inserted about 6 kb downstream from ptsH (184). The ptsH-linked antibiotic resistance cassette was recovered from strain MO481 (P. Stragier, unpublished results). The use of the ptsH1 mutant demonstrated that many other B. subtilis genes and operons are partly or completely relieved from CCR, such as the acetyl coenzyme A (acetyl-CoA) synthetase-encoding acsA gene (984) as well as the lev (523, 526), xyl (148), xyn (249, 250), and glp (158) operons (for a review, see reference 178). In a few cases, the effect of the ptsH1 mutation on CCR depends on the growth conditions. CCR of the B. subtilis hut operon, for example, is not altered in a ptsH1 mutant grown in minimal medium, whereas CCR in this strain completely disappears when it is grown in complex medium (985). A similar but less pronounced growth medium-dependent effect on CCR was observed with the iol operon (184). The mechanisms underlying the growth-dependent differences in CCR in ptsH1 mutants are not understood but might be related to the different nitrogen sources present in the media (220).

Synthesis of the plasmid-encoded B. thuringiensis larvicidal protoxin Cry4A by stationary-phase cells is also sensitive to CCR. The repressive effect of glucose on Cry4A production disappeared in a ptsH1 mutant (Ser45Ala replacement in B. thuringiensis HPr) (399). In L. casei, the ptsH1 mutation causes a relief from CCR of 6-P-β-galactosidase and N-acetyl glucosaminidase and from diauxic growth in media containing glucose and either lactose, maltose, or ribose (919). An L. lactis ptsH1 mutant is relieved from CCR of the ptbA-bglH operon (569). These results confirm that P-Ser-HPr plays an important role in CCR of not only bacilli but also other gram-positive organisms.

A chromosomal B. subtilis ptsH2 mutant (Ser46Thr replacement) has also been constructed. Although the purified mutant HPr is slowly phosphorylated in ATP-dependent phosphorylation assays (722), the B. subtilis ptsH2 mutant exhibits a relief from CCR of the gnt operon identical to that of the ptsH1 mutant (M. Steinmetz and J. Deutscher, unpublished results). In contrast, an L. casei ptsH2 mutant shows only partial relief from CCR and diauxic growth (919). It is therefore likely that Ser46Thr HPr from L. casei is more efficiently phosphorylated than the B. subtilis mutant protein.

Some genes and operons are more strongly expressed in the presence of glucose or other rapidly metabolizable carbohydrates. This phenomenon is known as CCA. In B. subtilis, the expression of several genes such as alsS (α-acetolactate synthase) (726, 983), ackA (acetate kinase) (886), and pta (phosphotransacetylase) (685) is stimulated by the presence of glucose. CCA of these B. subtilis genes is absent from ptsH1 mutants. Expression of the L. lactis las operon containing the genes encoding the glycolytic enzymes phosphofructokinase, pyruvate kinase, and lactate dehydrogenase is also activated by the presence of glucose in the growth medium. The stimulatory effect of glucose on las operon expression disappeared in an L. lactis ptsH-disrupted strain transformed with a plasmid allowing the synthesis of L. lactis Ser46Ala mutant HPr (493), whereas transformants containing a plasmid carrying L. lactis wild-type ptsH exhibited normal glucose-mediated las activation.

The involvement of P-Ser-HPr in CCR and CCA was further supported by studying mutants that were unable to form P-Ser-HPr due to hprK inactivation. Such mutants have been obtained, for example, with B. subtilis (251, 315, 709), S. xylosus (357), and L. casei (198). In all three organisms, the inactivation of hprK causes a relief from CCR, but for several B. subtilis catabolic operons, the releasing effect in hprK mutants is markedly stronger than that in ptsH1 mutants (249, 523, 685). This phenomenon was observed for many other genes when transcriptome analysis was carried out with B. subtilis hprK and ptsH1 mutants (491). As will be explained below, the difference between ptsH1 and hprK mutants is due to the presence of Crh, a B. subtilis HPr paralog, which is also phosphorylated by HprK/P (250) and which can partly substitute for P-Ser-HPr in CCR and CCA.

Groundbreaking work in the laboratory of Glenn Chambliss led to the discovery of two other components involved in CCR in B. subtilis. Both were identified by isolating mutants in which the α-amylase-encoding amyE gene was relieved from CCR. Two independently isolated mutants were affected in a 14-bp imperfect palindromic sequence called amyR1, which is located just downstream from the amyE promoter and which exhibits similarity to the lac operator site of E. coli (601, 603). In both strains, replacement of the G:C base pair in position 8 of amyR1 with an A:T base pair was responsible for the observed CCR resistance (603, 939). Expression of a plasmid-borne cat gene under the control of the amyE promoter followed by mutated amyR1 was insensitive to CCR, while the simultaneous expression of the chromosomal amyE (promoter followed by wild-type amyR1) was repressed by glucose (602). These findings suggested that amyR1 is a cis-acting element. A series of mutants in which one or two positions of amyR1 were altered was constructed, and most of these mutants were partly or fully relieved from CCR of amyE. Based on mutational analysis and on the occurrence of similar imperfect palindromic sequences in the promoter regions of many other catabolite-repressed B. subtilis genes, the following consensus sequence was proposed for the catabolite response operator: TGWNANCGNTNWCA (939). Slightly different consensus sequences have been proposed by others (353, 563). The general term catabolite response element (cre) has been introduced for these operator sites (421).

Imperfect palindromic sequences resembling the proposed consensus cre sequence are located in the promoter region or at the 5′ ends of many catabolite-repressed transcription units. For some of these cre's, their implication in CCR or CCA has been experimentally confirmed (178). By searching the B. subtilis genome with a degenerate cre sequence as a query, Miwa and coworkers detected a total of 126 tentative cre's (565). Out of these 126 putative cre sites, 32 were inserted into a plasmid between the spac promoter and the lacZ gene, and the effect of these sequences on the glucose repression of β-galactosidase synthesis was tested. In 22 constructs, expression of the lacZ gene was repressed by glucose (565). By using a slightly different search method, 61 additional potential B. subtilis cis elements for CCR could be detected, including several cre's preceding operons known to be submitted to CCR. A list of the about 160 identified or presumed B. subtilis cre's was presented by Deutscher et al. (178). As the transcription units regulated by these cre's contain, on average, two to three cistrons, about 400 genes, which represent 10% of the B. subtilis genome, are estimated to be regulated by CCR or CCA. A similar number was obtained when transcriptome analyses were carried out using a ccpA mutant (see below for ccpA) (575, 980). The number of genes presumed to be regulated by cAMP/Crp-mediated CCR in E. coli is in the same order (34).

A unique potential cre precedes the B. subtilis uxaC gene from the yjm operon, which codes for the enzymes necessary for glucuronide/galacturonate transport and metabolism (549, 729). While most cre's are composed of an imperfect palindromic sequence, the putative cre of the yjm operon is a perfect palindrome. Moreover, the 14-bp-long putative yjm cre is part of a longer perfect palindrome composed of a total of 26 bp (TCAAAATGTTAACGTTAACATTTTGA [the 14-bp consensus cre is in boldface type]). To date, it is not known whether this extended palindrome plays a special role in CCR. The β-galactosidase-encoding mbgA gene of Bacillus megaterium contains two cre's overlapping by 1 bp. Mutations in either one affect CCR of the mbgA gene (803). Two potential cre's overlapping to a much larger extent (by 4 bp) begin at positions 278 and 287 within the B. subtilis gudB (former ypcA) gene, which encodes one of two glutamate dehydrogenases present in this organism and the expression of which is sensitive to the presence of glucose. The second B. subtilis glutamate dehydrogenase, RocG, is also submitted to CCR (53). The first 8 bp in the two presumed gudB cre's are identical (GTGAAGGCGgtgaaggcgCTTTCA [the first potential cre is in boldface type, the second is in italics, and the repeated sequences are underlined and in lowercase letters]). Interestingly, a rocG mutant is not able to utilize proline, ornithine, or arginine as the sole carbon source (growth on these carbon sources requires an active glutamate dehydrogenase), establishing that wild-type B. subtilis contains only low GudB activity. However, spontaneous revertants that were able to grow on these compounds appeared with high frequency. Surprisingly, in four independently isolated revertants, the GTGAAGGCG direct repeat was deleted, providing a gudB1 allele with only a single cre (TGAAGGCGCTTTCA) (54). It is not clear whether the elevated GudB activity in the spontaneous mutants is due to the change in the two overlapping cre's or to the deletion of the -V-K-A- repeat in the protein. The B. subtilis iol (563) and ara (361) operons also contain two functional cre's, which, however, are not located next to each other. One cre precedes the first gene (mmsA and araA, respectively); the other is located within the second gene (iolB and araB). Several other catabolite-repressed or -activated genes or operons contain more than one potential cre in the promoter region or within the 5′ end (93, 290, 564, 886), but in several cases, only one of the cre's is operative in CCR or CCA.

Two divergently transcribed genes or operons can potentially be regulated by a single cre. This seems to be the case for the putative L. casei lev cre, which overlaps the transcription start site of the levABCDX operon and the −10 box of the levR gene, which encodes the transcription activator for the lev operon. Simultaneous repression of levABCDX and the transcription activator gene levR presumably leads to very efficient CCR (534).

The L. casei lev operon is only one of many examples establishing the presence of functional cre sites in bacteria other than B. subtilis (Table 3). In S. xylosus, deletion of the cre located in front of the malRA operon (for maltose/maltotriose utilization) causes a complete relief from CCR for α-glucosidase (204). Similarly, deletion of a cre located right behind the transcription initiation site of the fructan hydrolase-encoding fruA gene of Streptococcus mutans causes a relief from glucose repression (93, 946). CCR of the β-amylase-encoding bamM gene from B. megaterium is mediated by a cre located in the DNA region encoding the signal peptide of this extracellular enzyme (457). A cre site is also present in front of the P-β-glucosidase-encoding bglH gene of Lactobacillus plantarum (511, 512) and the pepQ gene of Lactobacillus delbrueckii (779). These examples confirm that the cre-dependent CCR/CCA mechanism is widely distributed within gram-positive bacteria with low G+C content. Interestingly, the HPr-encoding ptsH gene of several streptococci is also preceded by a potential cre site, and expression of the ptsHI operon in S. salivarius and Streptococcus bovis is up-regulated in the presence of glucose (CCA) (894).

cre's can either overlap the promoter, overlap the transcription start site, or be located between the transcription start site and the initiation codon or within the 5′ part of catabolite-repressed genes (Table 3). Binding of a target protein to cre sites located within the promoter region is thought to interfere with the binding of the RNA polymerase holoenzyme, while cre's situated downstream from the promoter are assumed to be regulated via a road block. As will be discussed below, most transcription units submitted to CCA contain a cre located in front of the promoter region.

Since cre's exhibit similarity to the E. coli lac and gal operators, it was tempting to assume that a LacI/GalR-type repressor might be involved in CCR of B. subtilis and other gram-positive bacteria (603). By carrying out transposon mutagenesis, a B. subtilis strain in which amyE expression was resistant to catabolite repression could be isolated (322). Indeed, this strain carried the transposon inserted into a gene encoding a LacI/GalR-type repressor, which was called CcpA (catabolite control protein A) (938). Interestingly, genetic mapping revealed that the ccpA gene is located close to the previously described B. subtilis alsA1 mutation, which prevented the production of acetoin in glucose-grown cells (983). The failure of the alsA1 mutant to produce acetoin during growth on glucose is due to the almost complete absence of the alsS-encoded acetolactate synthase. The alsS gene forms an operon together with the acetolactate decarboxylase-encoding alsD (726) and is induced by a CCA mechanism. Similar to the alsA1 mutation, the ccpA mutation abolishes acetoin production. Introduction of an SPβ prophage carrying intact ccpA restored acetoin production in the ccpA and alsA1 mutants, strongly suggesting that ccpA and alsA are identical loci (322). Indeed, the alsA1 mutation was identified as a G-to-A exchange within the Shine-Dalgarno box of the ccpA gene (at position −14 with respect to the ccpA start codon) (566). This mutation lowers the synthesis of CcpA about 10-fold. The concentration of CcpA in wild-type B. subtilis was calculated to be 0.6 μM (566).

Disruption of ccpA not only prevented the inhibitory effect of glucose on α-amylase activity and the induction of acetolactate synthase but also affected CCR of many other B. subtilis enzymes (178, 184, 299, 566), thus establishing the pleiotropic character of the ccpA mutation. Whole-genome analysis turned out to be a powerful tool for detecting CcpA-regulated transcription units (575, 980). Interestingly, three B. subtilis ccpA mutants that specifically affected CCA of the alsSD operon, while CCA or CCR of other transcription units was not or only slightly altered, were isolated (887). Two of the mutations affect conserved regions in the DNA binding domain of CcpA (A18V and R48C replacements), whereas the third mutation was in a less conserved region in the core domain (Pro286Leu replacement).

The effect of ccpA inactivation on the synthesis of catabolic enzymes could also be demonstrated by comparing the protein patterns of two-dimensional gels obtained with crude extracts from B. subtilis (880) or E. faecalis (453) wild-type and ccpA strains. E. faecalis proteins isolated from a few spots exhibiting an altered intensity in the ccpA mutant compared to the wild-type strain were identified by microsequencing. They include a protein with strong similarity to the α subunit of l-serine dehydratase, an enzyme catalyzing the conversion of serine to pyruvate, as well as glycerol dehydrogenase and the two subunits of dihydroxyacetone kinase (DhaKL). The genes encoding the three latter proteins are present in the dha operon (453), which also contains the gene for an EIIA of the mannose class PTS (see “SOME UNUSUAL PTS PATHWAYS AND PROTEINS”). In addition, galKETR operon expression as well as the synthesis of 13 proteins identified as glucose starvation proteins were no longer sensitive to CCR in the E. faecalis ccpA mutant. Two-dimensional gel electrophoresis with B. subtilis crude extracts showed that in a ccpA mutant, seven tricarboxylic acid cycle enzymes (CitC, CitG, CitH, CitZ, OdhA, SdhA, and SucC/SucD) were relieved from CCR (880). The citZ (403) and citG genes and the odhAB, sdhCAB, and sucCD operons are preceded by potential cre's (178, 565). Interestingly, E. faecalis and B. subtilis ccpA mutants contain a much larger amount of P-Ser-HPr than wild-type strains (453, 492). The increased percentage of P-Ser-HPr slows glucose uptake by the B. subtilis ccpA mutant when grown on synthetic medium (492), which in turn seems to prevent CCA of the gapA operon, which encodes the enzymes of the lower part of glycolysis (492, 880). When a ptsH1 mutation, which prevents the formation of P-Ser-HPr, was introduced into the ccpA strain, the activating effect of glucose on gapA operon expression (230) was restored (492). Expression of gapA is controlled by the repressor CggR, which is inactivated by elevated amounts of FBP (194). A similar repressive effect of the elevated amount of P-Ser-HPr in a ccpA mutant was observed for the B. subtilis glutamate synthase-encoding gltAB operon. Introduction of a ptsH1 mutation restored gltAB expression in a ccpA background (925). The mechanism leading to enhanced P-Ser-HPr formation in ccpA mutants is not known. In addition, when grown in complex medium, ccpA mutants seem to efficiently transport PTS sugars, because glucose utilization by a B. subtilis ccpA mutant represses the synthesis of several enzymes submitted to a second, CcpA-independent CCR mechanism. For example, CCR of the α-glucosidase- and glycerol kinase-encoding genes is not diminished in a B. subtilis ccpA mutant (184).

Genes regulated by CcpA include not only carbon catabolic genes but also genes related to carbon metabolism in a larger sense. The L. delbrueckii subsp. lactis CcpA ortholog PepR1 stimulates the expression of the prolidase-encoding pepQ gene and seems to affect the expression of pepX, pepI, and brnQ, which encode two other peptidases and a branched-chain amino acid transporter, respectively (779). A connection between CCR and amino acid metabolism also exists in Streptococcus gordonii and Streptococcus rattus, where the arginine deiminase operon is repressed by CcpA (195, 298), and in B. subtilis, where CcpA stimulates the expression of the ilv-leu operon, which encodes the enzymes for the synthesis of branched-chain amino acids (810, 882). A connection between CCR and the utilization of nitrogen sources was provided by the finding that the expression of rocG, which encodes the main glutamate dehydrogenase of B. subtilis, is subject to CcpA-mediated CCR (53). A connection between carbon and nitrogen metabolism in B. subtilis was further supported by the finding that the σ⁵⁴-encoding sigL gene is regulated by CcpA (128). Transcriptome analysis with B. subtilis ccpA, hprK, and ptsH1 mutants revealed that numerous nitrogen and phosphorus metabolic genes are submitted to CCR or CCA (491). CcpA of Lactobacillus pentosus represses the activity of xylulose 5-phosphate phosphoketolase, the central enzyme of the phosphoketolase pathway (674). In Clostridium perfringens, inactivation of ccpA diminishes the expression of the enterotoxin-encoding cpe gene during entry into the stationary growth phase (913). S. mutans ccpA mutants exhibit diminished fructosyltransferase and glucosyltransferase gene expression (88) and biofilm formation (947).

CcpA belongs to the family of LacI/GalR-type repressors but contains characteristic sequences located at the end of the DNA binding domain and within the core protein and can therefore easily be distinguished from other members of this family. These regions are important for CcpA function, because mutations affecting amino acids in the CcpA-specific sequences prevent or lower CCR of a xylA-lacZ fusion in B. megaterium (422). Interestingly, polyclonal antibodies directed against CcpA from B. megaterium do not recognize E. coli LacI (441). Phylogenetic analysis shows that all low-G+C gram-positive bacteria for which the genomes have been sequenced contain a ccpA ortholog (J. Deutscher and C. Francke, unpublished observation). The gene is probably always expressed, as a protein exhibiting the approximate size of CcpA and cross-reacting with the B. megaterium CcpA antibody was present in the crude extracts of all gram-positive bacteria tested (441). For several of these bacteria, including E. faecalis, L. casei, Lactobacillus pentosus, L. lactis, L. monocytogenes, Clostridium saccharobutylicum (formerly C. acetobutylicum P262) (162), and S. xylosus, a role of their CcpAs in CCR was demonstrated experimentally. Either their ccpA genes complement a B. subtilis ccpA mutant or B. subtilis ccpA complements ccpA mutants of these organisms. Genes affected by ccpA mutations in bacteria other than B. subtilis are listed in Table 3.

Interestingly, overexpression of the C. saccharobutylicum ccpA ortholog regA in an E. coli strain harboring the C. saccharobutylicum staA gene inhibited the expression of staA, which encodes a starch-degrading enzyme. As the corepressor of CcpA is absent from E. coli (see the next section), these results imply that, if overproduced, CcpA can bind to cre's without a corepressor. Therefore, regulation of ccpA expression represents one way to control the binding of CcpA to cre's. Indeed, in a few organisms (S. xylosus, L. pentosus, and L. delbrueckii), ccpA is preceded by a cre, and its expression is submitted to autoregulation (204, 507, 574). In L. monocytogenes, the synthesis of CcpA was elevated when the cells were exposed to prolonged salt stress (200). By contrast, expression of B. subtilis ccpA is constitutive and is not influenced by the presence or absence of glucose (566). CCR in B. subtilis is therefore not regulated via ccpA expression. Instead, additional factors must be involved in the CCR signal transduction pathway. Because CcpA at physiological concentrations was not able to bind to the cre of the catabolite-repressed B. subtilis gnt operon (566), it was likely that these additional factors would function as corepressors of CcpA.

When the effects of the B. subtilis ptsH1 mutation on CCR were compared to those of a ccpA mutation, it was found that all enzymes that are partly or completely relieved from CCR in the ptsH1 mutant are also relieved from CCR in the ccpA mutant (184). In addition, two of the enzymes that remained repressed by glucose in the ptsH1 mutant (GlpK and α-glucosidase) remained repressed in the ccpA mutant (184). The similar pleiotropic phenotypes observed for ptsH1 and ccpA mutants suggested that P-Ser-HPr and CcpA participate in the same CCR mechanism and might interact. Binding of P-Ser-HPr to CcpA was indeed demonstrated by elution retardation experiments (182). In the presence of FBP, elution of P-Ser-HPr from a Ni-nitrilotriacetic acid column saturated with His-tagged CcpA was clearly retarded compared to the elution of HPr or P∼His-HPr, while doubly phosphorylated (P∼His,P-Ser)-HPr was only slightly retarded.

The specific interaction between P-Ser-HPr and CcpA was confirmed by NMR measurements (381). The formation of the P-Ser-HPr:CcpA complex could be observed in the absence of FBP, probably owing to the high protein concentrations used in the NMR studies. The NMR measurements also allowed the determination of the residues of P-Ser-HPr interacting with CcpA. In addition to the regions around the active-center His-15 and the regulatory Ser-46 (amino acids 43, 44, and 46 to 56), the interface includes amino acids 21 to 27. The regions at amino acids 21 to 27 and 46 to 56 are well conserved within the HPrs of gram-positive bacteria but are quite different in the HPrs of most gram-negative organisms (Fig. 4).

The size of CcpA (37 kDa) is too large to allow a determination of its interface in the P-Ser-HPr:CcpA complex by NMR measurements. However, based on the crystal structure of PurR (788), a LacI/GalR-type repressor missing the CcpA-specific sequences, and the failure of various mutant CcpAs to form a complex with P-Ser-HPr, a presumed surface-exposed area including Tyr-89, Tyr-295, Ala-299, and Arg-303 was proposed to be part of the interface in the P-Ser-HPr:CcpA complex (422). The four amino acids presumed to participate in P-Ser-HPr binding are located in CcpA-specific regions that are absent from other members of the LacI/GalR repressor family. The crystal structure of the B. megaterium P-Ser-HPr:CcpA complex revealed that residues Tyr-295, Ala-299, and Arg-303 are located on helix IX of the CcpA N subdomain (which directly follows the DNA binding domain) and indeed interact with P-Ser HPr (787). Val-300 and Leu-304 of CcpA are also part of this tight interface, which includes Ile-47, Met-48, and Met-51 of P-Ser-HPr. Replacement of Ile-47 with Thr in S. salivarius HPr leads to the disappearance of diauxic growth and the CCR observed with the wild-type strain during lactose and maltose utilization (263). However, crossed immunoelectrophoresis with HPr antibodies revealed that the amount of P-Ser-HPr in the ptsH(Ile47Thr) mutant was similar to that observed in the wild-type strain. The ptsH(Ile47Thr) mutation was therefore assumed to diminish the affinity of the mutant P-Ser-HPr for CcpA (263). The surface-exposed hydrophobic patch formed by Ile-47, Met-48, and Met-51 is also involved in the interaction of HPr with EI (260) and EIIAs (139, 691, 949).

NMR studies revealed an affinity of CcpA for P-Ser-HPr that was about 50-fold higher than that for HPr (381). This effect must be due mainly to electrostatic repulsion, as no significant structural changes accompany the phosphorylation of HPr at Ser-46 (29, 381). The electrostatic effects were assumed to involve the negatively charged phosphorylated Ser-46 of HPr and a positively charged counterpart in CcpA. According to the crystal structure, Arg-303 and Lys-307 indeed form hydrogen bonds with the phosphoryl group in P-Ser-HPr. Tyr-89, the fourth CcpA residue predicted to be involved in the interaction with P-Ser-HPr, is located on helix I of CcpA. Its close contact to helix IX probably explains why a mutation affecting Tyr-89 prevents the interaction with P-Ser-HPr (422). The P-Ser-HPr:CcpA example therefore proves that based on genetic and biochemical studies, reliable predictions about protein-protein interfaces can be made.

Similar to gram-negative bacteria, where PEP-dependent phosphorylation of EIIA^Glc is important for CCR (see “REGULATION OF CARBON METABOLISM IN GRAM-NEGATIVE ENTERIC BACTERIA”), phosphorylation of HPr at His-15 affects CCR in gram-positive bacteria, probably by weakening the P-Ser-HPr:CcpA interaction. Indeed, the region around His-15 of P-Ser-HPr makes contacts to helix IX of subunit 1 and helices I and II of subunit 2 in the regulatory domains of the CcpA dimer (787). One P-Ser-HPr molecule therefore interacts with both subunits of the CcpA dimer. Asp-69 and Asp-99 on helices I and II in the regulatory domain of subunit 2 of the CcpA dimer form hydrogen bonds with Arg-17 in P-Ser-HPr. The crystal structure provides an explanation as to why CcpA exhibits only a weak affinity for (P-Ser,P∼His)-HPr (184) and why replacing His-15 with Glu or Ala caused an almost complete relief from CCR (707). When a ptsH1 mutant strain was transformed with a plasmid carrying the His15Ala ptsH allele, the Ser46Ala mutant HPr synthesized from the chromosomal ptsH1 allele allowed the uptake of glucose via the PTS, while the His15Ala mutant HPr synthesized from the plasmid-located ptsH allele could be phosphorylated at Ser-46 and was therefore expected to be active in CCR. However, only a weak repressive effect of glucose on gluconate kinase activity was observed in the ptsH1 transformant (707), suggesting that seryl-phosphorylated His15Ala mutant HPr has only a low affinity for CcpA (707). Indeed, in the P-Ser-HPr:CcpA complex, Asp-296 forms hydrogen bonds with the Nδ1 of His-15 and the backbone amide nitrogen of Ala-16, which enhances the affinity between the two proteins. Phosphorylation at Nδ1 of His-15 in P-Ser-HPr by PEP and EI or replacing His-15 with Glu or Ala probably prevents the interaction of Asp-296 with HPr and, in the case of (P-Ser,P∼His)-HPr and His15Glu mutant P-Ser-HPr, probably even leads to electrostatic repulsion. The participation of His-15 in the interaction with CcpA provides a link between CCR and PEP-dependent phosphorylation of PTS proteins (182).

Several mutations leading to “permanent” CCR have been described. Replacing Ser-46 of HPr with a negatively charged Asp provides a mutant protein resembling P-Ser-HPr (952), which was therefore able to bind to CcpA immobilized on a Ni-nitrilotriacetic acid column (707). In agreement with this finding, overexpression of the ptsH(Ser46Asp) allele from a plasmid led to partial CCR of the B. subtilis gnt operon even in the absence of a repressing sugar (184). Two other classes of mutations that caused similar “permanent” CCR have been described. Several B. megaterium ccpA mutants that exhibit partial CCR of the xylose utilization genes in the absence of a repressing sugar have been isolated (442). In contrast to wild-type CcpA, the mutant CcpAs are thought to interact with the xyl cre even in the absence of a corepressor (442). The mutations affected the DNA binding domain (Glu77Leu replacement) or the core domain of CcpA. Permanent CCR was also observed with a B. subtilis mutant producing Val265Phe mutant HprK/P, which exhibits normal kinase but strongly reduced P-Ser-HPr phosphorylase activity, which leads to the accumulation of P-Ser-HPr (571). Similar to the HPr(Ser46Asp)-producing strain, CCR in the hprK(Val265Phe) mutant occurs even in the absence of a repressing sugar. Expression of a xynB′-lacZ fusion in the hprK(Val265Phe) mutant grown in a medium containing xylose and succinate as the sole carbon sources is 100 times lower than that in a wild-type strain (571). This permanent repressive effect is responsible for the failure of the hprK(Val265Phe) mutant to grow on CCR-sensitive sugars such as gluconate, glucitol, and ribose. Inactivation of ccpA or the insertion of a ptsH1 mutation in the hprK(Val265Phe) strain restored growth on the above-mentioned sugars (571). These results confirm that P-Ser-HPr formation represents the main signal for CCR in gram-positive bacteria. When present at elevated concentrations in the cell, P-Ser-HPr is sufficient to lead to strong CCR, while glycolytic intermediates do not seem to be essential.

The above-described results indicate that in gram-positive bacteria with low G+C content, P-Ser-HPr functions as a corepressor by allowing the repressor CcpA to bind to cre sites. This concept was first tested with the B. subtilis gnt operon, which possesses a cre located within the 5′ end of gntR, the first gene of this operon (247). Footprint experiments revealed that CcpA or P-Ser-HPr alone does not bind to the gnt cre. By contrast, if P-Ser-HPr and CcpA were present together in the reaction mixture, a specific protection of a region corresponding to the gnt cre was observed, while no protection occurred with CcpA and dephospho-HPr (247). Mutations affecting the gnt cre and causing a relief from CCR prevented binding of the P-Ser-HPr:CcpA complex. Similar results were obtained with cre's from many other catabolite-repressed or -activated transcription units. In addition, by isolating the DNA:protein complex and separating its components on a denaturing gel, it was demonstrated that P-Ser-HPr and CcpA bind together to cre sites (30).

The binding affinity of the P-Ser-HPr:CcpA complex for a cre site was first studied with circular dichroism spectroscopy. Based on association/dissociation-related changes in the spectrum, an apparent K_D of 4.5 μM was determined for the formation of the ternary complex P-Ser-HPr:CcpA:optimized B. megaterium xyl cre (A:T in position 10 replaced with T:A) (381). A 1:1:2 (cre:CcpA dimer:P-Ser-HPr) binding stoichiometry was proposed. Because no interaction with the cre could be observed when 0.5 mM HPr and CcpA was used, binding of P-Ser-HPr to CcpA was estimated to increase the affinity of the repressor for its DNA targets by more than 2 orders of magnitude (381). Besides, equilibrium constants for the binding of the P-Ser-HPr:CcpA complex to the B. megaterium xyl cre and the B. subtilis amyE cre were determined by electrophoretic mobility shift experiments and were in the orders of 5 × 10⁶ and 3.5 × 10⁸ M⁻¹, respectively (30, 404). A K_D (0.6 nM) that was much lower than the one determined by circular dichroism spectroscopy (4.5 μM) was reported for the formation of the ternary complex of B. subtilis CcpA, P-Ser-HPr, and the xylA cre (794).

The crystal structure of the triple complex composed of the CcpA dimer and two P-Ser-HPr molecules from B. megaterium and a cre has been solved (787). It revealed that the N-terminal HTH motifs and the hinge helices of the CcpA dimer make 32 phosphate contacts to the DNA. They interact with 10 bp out of the 16 bp forming the pseudopalindromic cre, an artificial operator site not present in B. megaterium. However, a reverse complementary sequence identical to bp 2 to 15 of the employed cre is present in Streptococcus iniae and probably controls expression of the lactose permease- and lactose oxidase-encoding lctPO operon (cre located 52 bp upstream from the start codon) (J. Deutscher, unpublished observation). In the ternary complex, important contacts are made with the central C:G and G:C base pairs via Leu-55, the “leucine lever.” These contacts are responsible for DNA kinking and minor groove expansion. DNA bending by the P-Ser-HPr:CcpA complex (35°) is smaller than what was observed for LacI (40°) and PurR (50°). The HTH motif exhibits sufficient flexibility to allow CcpA binding to the variable half sites of pseudopalindromic cre's. The plasticity of the HTH motif also explains the capacity of CcpA to interact with the various cre's of an organism, which generally exhibit significant sequence differences.

A comparison of the structure of the triple complex with that of unliganded CcpA (apoCcpA) revealed that the interaction of P-Ser-HPr with CcpA causes significant structural changes, primarily in the regulatory N subdomain, which allow CcpA to interact with cre's (787). The N subdomain in both subunits of “activated” CcpA is rotated on average by 6° compared to apoCcpA. In addition, the position of helix IX (contains Arg-303) in “activated” CcpA is different from its location in apoCcpA. The interaction of Arg-303 of CcpA with the phosphoryl group in P-Ser-HPr requires the rotation of the side chain of Arg-303, which initiates a series of structural changes culminating at Thr-61, which is located at the core/DNA binding junction. The rotation of Tyr-91, which in apoCcpA interacts with Thr-306 via a hydrogen bond, towards the CcpA dimer interface plays an important role in this process, which ultimately allows high-affinity binding of CcpA to cre sites (787).

The P-Ser-HPr:CcpA interaction mode seems to be independent of the absence or presence of a cre site. The crystal structure of a complex formed between B. subtilis P-Ser-HPr and CcpA missing the first 57 amino acids was very similar to the corresponding structure in the B. megaterium P-Ser-HPr:CcpA complex (106). Interestingly, the B. subtilis protein complex contains two sulfate ions located close to each other. Both sulfate ions interact with Asp-276, and because an FBP molecule could easily be modeled into the structure with its two phosphoryl groups superimposed on the sulfate ions, it was predicted that the region around Asp-276 represents the FBP binding site.

Based on the above-described results, a general CCR and CCA mechanism, as outlined in Fig. 6 can be proposed for low-G+C gram-positive bacteria. The increase in FBP in cells growing on a rapidly metabolizable carbohydrate such as glucose stimulates the HprK/P-catalyzed formation of P-Ser-HPr, which forms a complex with CcpA and allows the LacI/GalR-type repressor to bind to the cre's. For all genes of gram-positive organisms known to be submitted to CCA, the cre is located upstream from the promoter, and binding of the P-Ser-HPr:CcpA complex stimulates transcription, probably by a mechanism similar to that reported for Crp- or Cra-mediated activation of gene expression (142, 645). cre's located upstream from the promoter are found in the catabolite-activated B. subtilis ackA (886), glg (178), pta (685), and ilv-leu (810, 882) transcription units and the pepQ and las genes of L. delbrueckii and L. lactis, respectively (494) (Table 3). CcpA-mediated CCA of the B. subtilis als operon occurs probably via an indirect mechanism, as no obvious cre is located in front of the als promoter (882). The cre of the CCR-sensitive S. xylosus malRA operon is also located in front of the promoter (205). However, the mal cre and the mal promoter are separated by only 8 bp. This might be close enough to enable the P-Ser-HPr:CcpA complex bound to the cre to prevent binding of the RNA polymerase holoenzyme. In carbon catabolite-activated genes (ackA, pta, ilv-leu, and pepQ), the cre and the corresponding promoter are separated by at least 12 bp.

When Ramseier et al. studied the in vitro binding of B. megaterium CcpA to an “optimized” B. subtilis xyl cre, in which the G:C and A:T base pairs in positions 3 and 10 were replaced with T:A, an interaction was observed in the absence of P-Ser-HPr (695). Moreover, the presence of P-Ser-HPr or the structurally related Ser46Asp mutant HPr (952) inhibited the interaction of CcpA with the modified xyl cre. In addition to the two mutations introduced intentionally in the B. subtilis xyl cre, there is also an ambiguity for position 9, which was previously reported to be a T (421) but is a C in the published genome sequence (437). In the nearly identical B. megaterium xyl cre (only one base pair difference) (290), there is a C at position 9. As the experiments reported by Ramseier et al. (695) were carried out with the sequence carrying a T in position 9, the unexpected results obtained in the CcpA binding studies might be due to this sequence difference. Indeed, when footprint experiments were carried out with B. megaterium CcpA and an optimized B. megaterium xyl cre, which differs from the optimized B. subtilis xyl cre used previously (695) only at position 9 (C:G instead of T:A), results similar to those reported for the B. subtilis gnt cre were obtained; i.e., binding of B. megaterium CcpA to the xyl cre was observed only when P-Ser-HPr was present (290).

CcpA has been reported to bind in vitro to the B. subtilis ccpC (401) and ackA cre's (886) in the absence of a corepressor. Nevertheless, the stimulating effect of glucose on ackA expression disappeared in a ptsH1 crh double mutant (for crh, see the next section) (886). The discrepancy between in vivo and in vitro results could not be explained.

Glucose-6-P allowed P-Ser-HPr-independent binding of CcpA to the B. subtilis gnt cre (564) and the optimized B. megaterium xyl cre (290), albeit only under nonphysiological conditions. For the B. subtilis gnt cre, an effect could be detected only when glucose-6-P was used at more than 30 mM, which is far above the 4 mM glucose-6-P detected in glucose-grown B. subtilis cells (233). For the B. megaterium xyl cre, an effect of glucose-6-P on the binding of CcpA could be observed only at pH values below 5.4 (290), i.e., far below the cytoplasmic pH of 7 to 8 reported for B. megaterium cells (503). Glucose-6-P was therefore not considered to play a role in regulating binding of CcpA to the cre's but was suggested to mimic a possibly physiologically relevant unknown effector (290).

In footprint experiments, CcpA was found to bind weakly to the B. subtilis amyE cre in the absence of a corepressor (405). CcpA binding was stimulated by P-Ser-HPr and further enhanced by FBP (404). In in vitro transcription studies, CcpA in the 500 nM range was able to specifically inhibit amyE expression. In the presence of 2.7 μM P-Ser-HPr, inhibition of amyE expression occurred at slightly lower CcpA concentrations (around 100 nM) and was more efficient (90% inhibition). However, the P-Ser-HPr effect was considered to be nonspecific, as expression from control promoters was also inhibited. In addition, the ptsH1 mutation had no influence on CCR of the amyE gene (924). Surprisingly, when NADP or NADPH was present, specific inhibition of in vitro transcription from the amyE promoter was observed at very low CcpA concentrations (in the 5 nM range), but NADP(H) was less efficient (70% inhibition) than P-Ser-HPr. Moreover, in footprint experiments, the addition of NADP(H) had only a weak effect on the protection of the amyE cre by CcpA. It was therefore concluded that NADP(H) does not function as a corepressor by enhancing the binding of CcpA to the cre but, rather, that it allows CcpA to inhibit the transcription machinery, thus leading to reduced amyE expression (404). CcpA has been proposed to directly interact with and to inhibit RNA polymerase, and NADP/NADPH was thought to stimulate the interaction of CcpA with RNA polymerase (406). The suggested role of NADP and NADPH cannot be related to the redox state of the cells, as both molecules were similarly efficient in the stimulation of CcpA-mediated inhibition of amyE expression, while NAD and NADH had no effect. With the exception of dormant spores, the concentration of the NADP(H) pool was reported to remain fairly constant in B. megaterium cells under various growth conditions, including growth in the absence and presence of glucose (801). It is therefore not clear to which signal the interaction of NADP(H) with CcpA responds.

In several instances, CCR of catabolic genes is mediated by an indirect mechanism. For example, cre sites can be located in front of genes encoding transcription regulators of catabolic genes. This is the case for B. subtilis CcpC, which controls the expression of citZ and citB (401) and for LevR of L. casei, which regulates the expression of the levABCDX operon (534). Similarly, the acoABCL operon of B. subtilis is controlled by the transcription activator AcoR, the gene of which is preceded by a cre recognized by CcpA (12). While a cre site also precedes citZ and the L. casei levABCDX operon, a cre seems to be absent from acoABCL, which is therefore only indirectly regulated by CcpA. A cre site has also been identified within the sigL gene of B. subtilis, which codes for σ⁵⁴, which is necessary for the transcription from several “−12, −24” promoters (128).

The regM gene of S. mutans encodes a homolog of CcpA. Surprisingly, the disruption of regM increased the glucose repression of α-galactosidase, mannitol-1-P dehydrogenase, and P-β-galactosidase, suggesting that RegM functions as a positive regulator for the expression of the corresponding genes. In addition, regM inactivation affected neither diauxic growth in media containing glucose and a less efficiently metabolizable sugar (816) nor expression of the fructanase-encoding fruA gene, although deletion or mutation of the cre preceding fruA led to a relief from CCR (946). Moreover, FBP, which stimulates HPr kinase activity in most bacteria, slightly inhibited HPr kinase from S. mutans (869). It therefore seems that S. mutans follows a CCR mechanism different from that operative in most other gram-positive organisms.

Although most catabolite-repressed genes and operons of B. subtilis are partly or completely relieved from CCR in both ccpA and ptsH1 mutants, the degree of relief can be quite different for the two mutants. For example, CCR of the amyE gene and the hut, cta, xyn, and iol operons is only slightly affected by the ptsH1 mutation (less than 20%), whereas these five transcription units are completely relieved from CCR in ccpA mutants (184, 250, 488, 924, 985). Similarly, CCA of the phosphotransacetylase-encoding pta gene is not affected in a ptsH1 mutant but is completely lost in a ccpA strain (685). Less pronounced differences in CCR or CCA between ptsH1 and ccpA mutants were observed for the glucitol dehydrogenase-encoding gutB (184), the acetyl-CoA synthetase-encoding acs (984), and the acetate kinase-encoding ackA (886) genes as well as for the lev operon (250, 526). These results suggested that in B. subtilis, the binding of CcpA to certain cre's is regulated not only by P-Ser-HPr but that a second corepressor might exist.

Evidence for a potential second corepressor came from the B. subtilis genome sequencing project, which led to the discovery of a gene encoding an HPr-like protein (219). Although this protein exhibits 45% sequence identity to HPr, there is a striking difference: His-15, the site of PEP-dependent phosphorylation in HPr, is replaced with a Gln in the HPr paralog. As a consequence, the HPr-like protein is not phosphorylated by PEP and EI (250). Nevertheless, the sequence around Ser-46 resembles that in HPr of gram-positive bacteria, and HprK/P-mediated, ATP-dependent, FBP-stimulated phosphorylation at Ser-46 in Crh could be demonstrated (250). These results suggested that the phosphorylated HPr paralog might play a role in CCR similar to that of P-Ser-HPr, and it was therefore called catabolite repression HPr (Crh).

However, when the crh gene is disrupted, no effect on CCR is observed. The participation of P-Ser-Crh in CCR can be detected only in a ptsH1 background. The residual CCR of the lev, iol, xyn (250), and hut (985) operons as well as the remaining CCA of the pta (685) and ackA (886) genes observed in ptsH1 mutants disappeared almost completely when crh was inactivated. Complete relief of the acsA (984), xyn (249), and lev (522) operons from CCR was also observed in a ptsH1 crh1 double mutant in which Ser-46 of both HPr and Crh was replaced with Ala.

In vitro, Crh forms a mixture of monomers and dimers (644), which, similar to EI, are in slow equilibrium. The structure of the monomer and the interface of the dimer were determined by NMR spectroscopy (222). The NMR data suggested that domain swapping of the β1 strand is implicated in B. subtilis Crh dimer formation, and this was confirmed by the crystal structure (389). The Crh monomer strongly resembles HPr, except for a few specific differences. Their structural similarity explains why Gln15His mutant Crh can be phosphorylated by PEP and EI (524). When the Gln15His mutant Crh is overproduced in a B. subtilis ptsH deletion strain, it partly restores PTS transport activity and growth on PTS substrates or glycerol and allows the full activation of several transcription regulators possessing a PTS regulation domain (PRD) (156, 524). GlpK and PRD-containing transcription regulators (see “REGULATORY FUNCTIONS MEDIATED BY P∼His-HPr AND P∼EIIBs”) are stimulated by PEP-dependent phosphorylation via EI and HPr (158, 844).

To determine the binding characteristics of the P-Ser-Crh:CcpA complex, footprint experiments were carried out with cre's from transcription units, which are not completely relieved from CCR in ptsH1 mutants. The xyn cre site was indeed specifically protected by the P-Ser-Crh:CcpA complex, but the effect was slightly weaker than that with P-Ser-HPr:CcpA (249). The use of various mixtures of P-Ser-HPr and P-Ser-Crh did not increase the protective effect compared to the protection obtained with only a single corepressor. For a few catabolic genes, the binding of P-Ser-Crh:CcpA to the cre's was stimulated by FBP, and in some cre's, certain positions became hypersensitive towards DNase I digestion after binding the repressor:corepressor complex. P-Ser-HPr:CcpA and P-Ser-Crh:CcpA protected identical DNA regions, and FBP always had comparable effects on the DNA binding affinity of the two protein complexes. For example, FBP increased the protective effect of the P-Ser-HPr:CcpA and P-Ser-Crh:CcpA complexes against DNase I digestion of the xyn cre (249) and the pta cre (685). In contrast, FBP did not enhance the protective effect of both CcpA:corepressor complexes in footprint experiments with the lev cre (523). Both protein complexes rendered several nucleotides located outside the lev cre hypersensitive towards DNase I digestion. Expression of the lev operon is regulated by the transcription activator LevR, which binds to an upstream activating sequence (UAS) centered at about 130 bp upstream from the lev promoter. LevR was therefore assumed to exert its stimulating effect on lev operon expression by DNA bending (523). The lev cre is located between the UAS and the σ⁵⁴-dependent “−12, −24” promoter. Binding of the P-Ser-HPr:CcpA or P-Ser-Crh:CcpA complexes to the lev cre supposedly prevents LevR-induced DNA bending by altering the DNA structure, which might be responsible for the hypersensitivity towards cleavage during footprint experiments. Although in the case of the pta cre, only weak protection by P-Ser-HPr:CcpA or P-Ser-Crh:CcpA was observed, several nucleotides located outside the cre site became hypersensitive to DNase I digestion. When FBP was included in the reaction mixtures, the hypersensitive bands disappeared, and the protective effect on the pta cre was enhanced (685). The physiological significance of these in vitro observations remains to be determined.

In addition to the fact that certain cre's are not recognized by the P-Ser-Crh:CcpA complex (when the ptsH1 mutation leads to a complete relief from CCR or CCA), P-Ser-Crh was reported to have a 10-fold-lower affinity for CcpA than P-Ser-HPr. These differences are surprising, as, with the exception of two amino acids (His-15 and Thr-20), all P-Ser-HPr residues involved in the interaction with CcpA are conserved in Crh, and the crystal structures of the two proteins revealed a very similar fold. A possible explanation was that the dimerization of Crh (389) might have an influence on the formation of the P-Ser-Crh:CcpA complex and its interaction with cre sites. However, the crystal structure of the triple complex P-Ser-Crh(B. subtilis):CcpA(B. megaterium):cre (same cre as that used for the P-Ser-HPr:CcpA structure) (787) revealed that the two P-Ser-Crh molecules bound to the CcpA dimer are present exclusively in monomeric form (789). The binding of P-Ser-Crh:CcpA to the cre induces a kink (DNA bend angle of 31°) in the central conserved C-G that is almost identical to that observed in the triple complex with P-Ser-HPr (DNA bend angle of 35°). Nevertheless, specific differences between P-Ser-HPr:CcpA and P-Ser-Crh:CcpA occur in the contact region including His-15/Gln-15 and Ala-20/Thr-20 of HPr/Crh, respectively. In the complex with P-Ser-HPr, His-15 hydrogen bonds with the side-chain carboxyl of Asp-296 of CcpA. In contrast, Gln-15 of Crh makes only a weak contact to Asp-296 (distance of 4 Å), but instead, hydrogen bonds with the side chain of Arg-324, which, compared to the P-Ser-HPr:CcpA structure, needs to be rotated to come sufficiently close to the interface to allow the contact with Gln-15. Interestingly, an equivalent of Arg-324 seems to be present in CcpAs of only those organisms possessing Crh (789). If this assumption holds, Bacillus pseudofirmus, Bacillus sphaericus, and Exiguobacterium sibiricum 255-15 should contain a Crh ortholog, as their CcpAs contain an equivalent of Arg-324, whereas G. stearothermophilus, in contrast to Geobacillus kaustophilus, should be devoid of Crh, as its CcpA is missing an equivalent of Arg-324 (replaced with a Glu). The presence of Crh in the above-mentioned four organisms is uncertain, as no or only incomplete genome sequences are presently available.

In conclusion, the structures of the complexes of P-Ser-HPr and P-Ser-Crh with CcpA and a cre exhibit extensive similarity, and the few differences that were observed do not explain why the B. subtilis P-Ser-HPr:CcpA complex binds to several cre sites, which, according to genetic data, are not or only barely targeted by the P-Ser-Crh:CcpA complex. The corresponding transcription units are partly or completely relieved from CCR in ptsH1 and hprK mutants (gnt, mtl, etc.) (184). To better understand the different affinities of the two corepressor:CcpA complexes for certain cre's, it might be useful to determine the structure of a triple complex of P-Ser-HPr, CcpA, and a cre that does not interact with the P-Ser-Crh:CcpA complex.

In addition to B. subtilis (250) and B. megaterium (794), Crh has been detected in most other bacilli (Bacillus anthracis, B. clausii, B. cereus, B. halodurans, B. licheniformis, B. thuringiensis, and B. weihenstephanensis) as well as in G. kaustophilus and O. iheyensis, which are both closely related to bacilli. The various Crhs exhibit a high degree of sequence identity (between 61 and 83%), whereas they show only 41 to 47% sequence identity to the HPrs of their respective organisms. Interestingly, B. licheniformis possesses two HPr-like proteins containing a Gln in place of His-15. One of them has a Ser-46 and therefore corresponds to Crh (83% sequence identity with Crh of B. subtilis). The other protein exhibits only 44% sequence identity to B. subtilis Crh, and its Ser-46 is replaced with an Asp, which probably mimics permanent phosphorylation at position 46 (Deutscher, unpublished). The function of the latter protein is unknown. crh is the last gene (or penultimate gene in B. subtilis) of the yvc operon. The last gene of the B. subtilis yvc operon (250), yvcN, is absent from the other organisms, and O. iheyensis also misses the first gene, yvcI.

The genome sequences of other gram-positive bacteria revealed that staphylococci, clostridia, lactobacilli, lactococci, enterococci, etc., also have a yvc operon, but none of them contains a crh gene. Although the in vivo and in vitro results unequivocally established that P-Ser-Crh can contribute to CCR in B. subtilis, the fact that crh is present only in bacilli and a few close relatives and that crh inactivation in wild-type B. subtilis has no effect on CCR raises serious doubts about whether CCR is the primary function of Crh. P-Ser-HPr seems to be sufficient to mediate CCR in gram-positive organisms, independently of whether they possess Crh or not. In addition, numerous B. subtilis operons are completely relieved from CCR in a ptsH1 mutant, suggesting that their cre is not recognized by the P-Ser-Crh:CcpA complex. So far, only one P-Ser-Crh-specific function could be established. Expression of the Mg²⁺-citrate transporter-encoding B. subtilis citM gene is strongly repressed during growth on glucose but is also slightly repressed when grown on glutamate/succinate as the sole carbon source. Whereas repression of citM by glucose is mediated via CcpA in complex with either P-Ser-HPr or P-Ser-Crh, repression by glutamate/succinate completely disappeared in a crh-disrupted strain (934). Interestingly, crh expression, which is much weaker than ptsH expression, is highest in cells grown on one of the two TCA cycle intermediates, citrate or succinate (286).

It is possible that additional functions of Crh are related to the genes cotranscribed with crh. It is noteworthy that the yvcJ gene organized in an operon with crh resembles yhbJ from the E. coli rpoN operon. YhbJ possesses Walker motifs A and B, but its function is not known. Because in many hprK-containing gram-negative bacteria, yhbJ and other genes of the E. coli rpoN region are organized in an operon with hprK, it is tempting to assume that Crh might somehow play a role in the regulation of SigL (σ⁵⁴), the RpoN ortholog of B. subtilis.

An HPr paralog containing a conserved Ser-46 but missing His-15 (His-15 is replaced with a Phe) is also present in the mollicute Ureaplasma urealyticum (also called Ureaplasma parvum serovar 1) (308). The sequence around Phe-15 (amino acids 12 to 22) completely differs from all known HPrs, while the other parts of the protein resemble HPr from gram-positive and gram-negative bacteria. In contrast to other mollicutes, U. urealyticum lacks EI as well as an HPr with His-15. Mollicutes generally lack CcpA but contain an hprK gene, as is also the case for U. urealyticum (308). The role of P-Ser-HPr formation in these organisms therefore remains unknown (307).

Surprisingly, in certain pathogens, P-Ser-HPr formation affects the activity of transcription regulators of virulence genes. The best-studied example is the Crp/Fnr-type transcription activator PrfA of L. monocytogenes (207). It regulates the expression of several virulence genes, including the hemolysin-encoding hly gene and the phospholipase-encoding plcA gene, which together with the prfA gene are clustered within a 10-kb region on the chromosome (129). Growth of L. monocytogenes on rapidly metabolizable carbon sources such as glucose and fructose leads not only to CCR of catabolic genes (50) but also to repression of the PrfA-controlled genes (558). While repression of catabolic genes requires CcpA, repression of the virulence genes is CcpA independent, since it persists in ccpA mutants (50). Nevertheless, P-Ser-HPr, the corepressor of CcpA, seems to play a role in the regulation of PrfA activity. A B. subtilis strain containing L. monocytogenes prfA under the control of the spac promoter and the lacZ reporter gene fused to the PrfA-activated hly promoter exhibits lacZ expression only in the presence of IPTG (804). Expression of lacZ and thus PrfA activity are inhibited by glucose and fructose, indicating that CCR of PrfA-controlled genes is also operative in B. subtilis (179, 326). Interestingly, introduction of the hprK(Val267Phe) mutation in the B. subtilis strain, which leads to the conversion of more than 95% of the HPr into P-Ser-HPr (571), strongly inhibits PrfA activity even in the absence of a repressing sugar (326). Replacement of Ser-46 in HPr with an alanine (ptsH1 mutation) prevented the inhibitory effect of the hprK(Val267Phe) mutation, suggesting that P-Ser-HPr is directly or indirectly involved in PrfA regulation. In contrast, inactivation of ccpA or introduction of the crh1 mutation in the hprK(Val267Phe) strain did not stimulate PrfA activity (326). The effect of P-Ser-HPr on PrfA activity seems to be indirect and based primarily on its poor phosphorylation by enzyme I, which probably leads to barely phosphorylated EIIAs and EIIBs. In agreement with this concept, deletion of ptsH or the enzyme I-encoding ptsI also led to strong inhibition of PrfA activity (326). PrfA therefore seems to require a fully functional PTS and might be either stimulated by P∼His-HPr, P∼EIIAs, or P∼EIIBs or inhibited by dephosphorylated EIIAs or EIIBs. A connection between the PTS components and PrfA was also suggested by the observation that the overproduction of PrfA inhibits glucose utilization via the PTS (517). In contrast, the utilization of glucose-6-P via the hexose phosphate transport protein (UhpT) was not affected by PrfA overproduction. In addition, in contrast to glucose, fructose, cellobiose, etc., glucose-1-P did not inhibit PrfA activity, although it is probably converted to glucose-6-P and metabolized via glycolysis (728). The failure of glucose-1-P, which is not transported via the PTS, to inhibit PrfA activity might therefore be related to its different mode of transport. In conclusion, these results suggest a direct or indirect effect of PTS components necessary for the uptake of glucose, fructose, cellobiose, etc., on PrfA activity.

In contrast to PrfA, the S. pyogenes virulence gene regulator Mga seems to be controlled by the classical CcpA-dependent CCR mechanism. Mga functions as a transcription activator for the expression of numerous genes encoding mainly surface-associated proteins involved in pathogenesis. Expression of the Mga regulon is growth phase dependent, and the presence of rapidly metabolizable carbon sources such as glucose stimulates the synthesis of members of the M-protein family (emm, mrp, arp, and enn) (424, 535). In addition, Mga controls genes encoding fibronectin-binding proteins (sof and fbx) and a collagen-like protein (sclA) (14, 15). Mga also controls its own expression by interacting with two 59-bp binding motifs located within the mga promoter region (536). In addition, the −35 region of the first of the two mga promoters is preceded by a DNA sequence that almost perfectly matches the cre consensus sequence. Purified CcpA was found to bind to the presumed cre site (K. S. McIver, personal communication). Its location upstream from the −35 promoter region suggests that mga is submitted to CCA. This is in agreement with the previous observation that the presence of glucose stimulates the synthesis of the M protein.

The Streptococcus pneumoniae CcpA ortholog RegM represses α-glucosidase and β-galactosidase activities, but the inactivation of ccpA (regM) also attenuates the virulence of this organism. The mutation has no impact on the expression of several established pneumococcal virulence genes but, rather, affects the expression of genes from the capsular polysaccharide synthesis (cps) locus (273). Capsular polysaccharides are known to be important factors for host cell adhesion. Indeed, the ccpA mutant is severely attenuated for nasopharyngeal colonization and lung infection in mice (366). This effect might partly be related to the differences in surface protein composition observed between the wild type and the ccpA mutant.

Genes encoding proteins with significant similarity to HprK/P are present in numerous gram-negative bacteria (α-, β-, γ-, and δ-proteobacteria), spirochetes, green sulfur bacteria, Fibrobacteres, and fusobacteria (41, 64). The existence of HprK/P homologs in gram-negative bacteria came as a surprise. Due to the absence of Ser-46 phosphorylation in the HPrs of E. coli and other Enterobacteriaceae, HprK/P was originally thought to exist only in gram-positive organisms. Interestingly, HprK/P-containing gram-negative bacteria possess an HPr in which the sequence around Ser-46 differs from the corresponding sequence in HPrs of Enterobacteriaceae and instead resembles those in HPrs from gram-positive organisms (64) (Fig. 4). For the gram-negative bacteria Neisseria gonorrhoeae and N. meningitidis (64), the spirochete T. denticola (278), and the α-proteobacteria Agrobacterium tumefaciens (S. Poncet, A. Khemiri, I. Mijakovic, A. Bourand, and J. Deutscher, unpublished results) and B. melitensis (S. Poncet, M. Dozot, A. Bourand, J. Deutscher, J. J. Letesson, and X. de Bolle, unpublished results), ATP-dependent phosphorylation of HPr by their HprK/Ps has been demonstrated in vitro. In several of the above-mentioned bacteria, ptsH and sometimes also ptsI are located in the hprK operon. Nevertheless, most of these bacteria seem to contain neither CcpA nor an EIIB or EIIC, excluding the possibility that P-Ser-HPr plays a role in the regulation of PTS transport activity or CcpA-mediated CCR in these bacteria.

In addition to ptsH and ptsI, the hprK operons of gram-negative bacteria frequently contain a gene encoding an EIIA of either the fructose/mannitol or the mannose class PTS (64, 345). The organization of the hprK operon in certain gram-negative pathogens and symbionts suggests a role of HprK/P in cell adhesion and virulence (64). In these proteobacteria, the hprK operon usually contains between 1 and 10 genes of the E. coli rpoN region or genes encoding a two-component system. Genes coding for an EIIA-like protein (PtsN) and an HPr-like protein (PtsO or NPr) are also present in the rpoN operon of E. coli and other enterobacteria (553) (for the regulatory functions of RpoN, PtsN, and PtsO, see “SOME UNUSUAL PTS PATHWAYS AND PROTEINS”). In hprK-containing β-, γ-, and δ-proteobacteria, hprK is usually located between rpoN and yhbJ (64). In a few α-proteobacteria, the hprK and yhbJ genes are also located next to each other. Interestingly, the crh gene of B. subtilis is also organized in an operon with a yhbJ homolog (64), while in Ruminococcus albus, hprK is followed by murB and a yhbJ-like gene (Deutscher, unpublished). These observations strongly suggest a functional connection between the PTS and the nucleotide binding protein YhbJ.

For β- and γ-proteobacteria containing a combined hprK/rpoN operon, HprK/P was proposed to control the phosphorylation state of the EIIA^Fru-like PtsN (64, 724), the gene of which often precedes hprK. In these organisms, signals causing enhanced HPr kinase and diminished P-Ser-HPr phosphorylase activity were assumed to lead to elevated amounts of P-Ser-HPr, which would slow the phosphorylation of the EIIA (180, 723). In N. meningitidis, such a phosphorelay system seems to somehow control the interaction with host cells, as the inactivation of hprK strongly reduces cell adhesion. The inactivation of hprK possibly leads to the enhanced formation of P∼PtsN. Dephospho-PtsN was therefore proposed to participate in a signal transduction pathway controlling cell adhesion (64).

Similarly, in several α-proteobacteria, the colocalization of hprK with genes encoding a two-component system suggests a role of HprK/P in the regulation of cell adhesion (64, 345). The A. tumefaciens EnvZ/OmpR-like two-component system ChvG/ChvI, which is encoded by the genes preceding hprK, controls the expression of the virulence genes virB and virE (472), which encode components of a type IV secretion system delivering oncogenic DNA to susceptible plant cells (100). Inactivation of chvG or chvI prevents tumor formation on the host plants (108). Similarly, inactivation of the chvG/chvI-like bvrR and bvrS genes of the pathogenic animal endosymbiont Brucella abortus affects cell invasion and virulence of this organism (821). Inactivation of the ChvG-like sensor kinase ExoS in the plant symbiont Sinorhizobium meliloti prevents the production of succinoglycan, the main exopolysaccharide necessary for the successful invasion of nodules on the host plant alfalfa (126). It has been proposed that HprK/P and the PTS proteins might somehow control the activity of the two-component systems (64). In α-proteobacteria, the hprK operon usually contains a gene encoding an EIIA of the mannose class PTS and sometimes also the genes for EI and HPr, whereas the gene for an EIIA of the fructose class PTS is located next to the rpoN gene (S. Poncet, unpublished observation). HprK/P was thought to control the phosphorylation state of the EIIAs. An EIIA or P∼EIIA was in turn suggested to interact with the two-component system (64), or alternatively, a P∼EIIA might phosphorylate one of the proteins of the two-component system. Interestingly, during a search for genes affecting the virulence of Brucella melitensis, inactivation of the EIIA^Fru-encoding gene was found to lower the pathogenicity of this organism (169).

α-Proteobacteria as well as Coxiella burnetii contain a truncated HprK/P missing the N-terminal domain, which resembles the N-terminal part of MurE (13), the enzyme catalyzing the formation of cytoplasmic precursors for cell wall synthesis (284). Similar to the artificially truncated L. casei HprK/P (229), the natural absence of the first 130 amino acids of A. tumefaciens HprK/P does not affect the known HprK/P functions. Nevertheless, the A. tumefaciens enzyme is not able to efficiently use PP_i as a phosphoryl donor and to dephosphorylate P-Ser-HPr in the presence of P_i (I. Mijakovic, A. Khemiri, A. Bourand, S. Poncet, and J. Deutscher, unpublished results). However, the absence of these functions from A. tumefaciens HprK/P is likely due to sequence differences in the central loop compared to HprK/P from gram-positive organisms. Mutations affecting this loop in L. casei or B. subtilis HprK/P strongly diminish the phosphorylase activity (571).

Interestingly, all known bacteria with a truncated HprK/P possess an EI with an N-terminal extension resembling the N-terminal DNA binding domain of NifA, a transcription activator for certain σ⁵⁴-dependent promoters. EI^Ntr (PtsP) of E. coli, which, together with the HPr paralog PtsO, is assumed to catalyze the phosphorylation of PtsN, also contains this NifA domain (see “SOME UNUSUAL PTS PATHWAYS AND PROTEINS”).

Inducer expulsion is a regulatory phenomenon that has been known for at least 40 years. Halpern and Lupo observed that the presence of certain carbon sources accelerates the exit of α-MG from E. coli cells that had accumulated this nonmetabolizable PTS substrate (309). Glucose had the strongest effect and accelerated α-MG expulsion about sixfold compared to cells to which no carbon source was added. When discussing the expulsion of nonmetabolizable carbohydrates, at least two different forms need to be distinguished, as they follow different mechanisms: the expulsion of non-PTS sugars and PTS sugars. The latter are accumulated as phospho derivatives inside bacterial cells and are first dephosphorylated before they are expelled. We will discuss the expulsion of PTS sugars, because ATP-dependent HPr phosphorylation was discovered in connection with TMG expulsion from S. pyogenes cells (185, 710).

Streptococci and lactococci transport nonmetabolizable sugar derivatives such as TMG or 2DG via the PTS and accumulate these carbohydrates as P derivatives. Interestingly, these nonmetabolizable P-sugars are rapidly expelled when the cells are exposed to a well-metabolizable carbon source such as glucose, sucrose, or mannose (712, 872). Expulsion turned out to be a two-step process: the accumulated P-sugars are first intracellularly dephosphorylated before the dephosphorylated sugars are expelled (710). The expulsion could be prevented when the ATP synthesis was inhibited by poisoning the cells with arsenate, fluoride (712), or iodoacetate (872). It has subsequently been established that in poisoned cells, the dephosphorylation of the accumulated P-sugars is inhibited (710). Arsenate- or fluoride-poisoned S. pyogenes cells synthesize little ATP, but they can metabolize arginine via the deiminase pathway and thereby generate ATP. The presence of arginine indeed restored sugar-P dephosphorylation and inducer expulsion in arsenate- or fluoride-poisoned S. pyogenes cells (710). It was therefore concluded that the first step of inducer expulsion depends on the presence of intracellular ATP and that the sugar-P phosphatase catalyzing this step might be regulated directly or indirectly via ATP-dependent protein phosphorylation. The search for this presumed P-protein led to the discovery of P-Ser-HPr: under conditions provoking inducer expulsion, a low-molecular-weight protein became phosphorylated by ATP (185, 710). This protein was identified as P-Ser-HPr (185), and it was therefore assumed that P-Ser-HPr might play a role in inducer expulsion, probably by controlling the activity of the sugar-P phosphatase (708, 710).

Experiments aimed at studying in vitro TMG-6-P expulsion by L. lactis vesicles supported this concept. When we discussed the effect of P-Ser-HPr formation on PTS-catalyzed carbohydrate uptake, we mentioned that vesicles prepared from L. lactis cells contain EI, HPr kinase, and a sugar-P phosphatase but had lost most of the HPr (970). As a result, L. lactis vesicles exhibit only about half the TMG uptake rate measured with intact cells (970). The full TMG uptake rate was restored when 100 μM B. subtilis HPr was electroporated into the L. lactis vesicles. Coelectroporation of wild-type or mutant HPrs with various metabolites was used to test whether phosphorylation of HPr at Ser-46 affects expulsion of TMG from the vesicles. L. lactis vesicles were able to accumulate TMG-6-P, which, similar to in intact cells, was expelled when glucose was added. Interestingly, vesicles into which HPr had been electroporated exhibited glucose-induced TMG expulsion that was about three times faster than that of vesicles that had been electroporated in the absence of HPr or in the presence of Ser46Ala mutant HPr (970). In addition, a cytoplasmic sugar-P phosphatase was activated about sevenfold when glucose was added to toluenized vesicles into which HPr had been electroporated prior to toluenization. Coelectroporation of FBP and HPr similarly enhanced the sugar-P phosphatase activity, and stimulation was even stronger when ATP was included. Because glucose and FBP stimulate the ATP-dependent phosphorylation of HPr at Ser-46 in L. lactis vesicles, it was assumed that the sugar-P phosphatase catalyzes the first step in inducer expulsion and that it is allosterically regulated by P-Ser-HPr. In agreement with this concept, electroporation of the P-Ser-HPr-resembling Ser46Asp HPr stimulated the intravesicular sugar-P phosphatase even in the absence of FBP (970), whereas only a weak increase of the phosphatase activity was observed with FBP and Ser46Ala mutant HPr. In addition to this cytoplasmic phosphatase, a membrane-associated L. lactis sugar-P phosphatase was also reported to be activated by P-Ser-HPr (973). Similarly, S. bovis contains a cytoplasmic and a membrane-associated sugar-P phosphatase (138). However, the activity of only the latter enzyme was stimulated by Ser46Asp mutant HPr. Membrane-associated, Ser46Asp HPr-activated sugar-P phosphatases were also detected in S. pyogenes and E. faecalis, two other organisms exhibiting inducer expulsion (712, 967). Inducer expulsion in all these bacteria was therefore assumed to be triggered by P-Ser-HPr-mediated activation of the membrane-associated sugar-P phosphatase. However, as heterologous systems were used for the in vitro electroporation experiments (HPr from B. subtilis and vesicles from streptococci, lactococci, or enterococci), these results needed to be confirmed by in vivo experiments.

Similar to L. lactis ML₃, L. casei strains 64H and BL23 transport TMG via a lactose-specific PTS, accumulate it as TMG-6-P, and expel it from the cells as TMG when glucose or another rapidly metabolizable carbon source is added (111, 198). An L. casei mutant synthesizing Ser46Ala HPr (ptsH1) was constructed from strain BL23. Although P-Ser-HPr cannot be formed in this strain, it expelled accumulated TMG-6-P similarly to the wild-type strain when it was exposed to glucose or mannose. Identical results were obtained with ptsH mutants in which Ser-46 or Ile-47 had been replaced with a threonine (198). These results established that the expulsion of TMG in L. casei does not depend on P-Ser-HPr. To exclude the possibility that an HPr-like protein capable of substituting for P-Ser-HPr in inducer expulsion might exist in L. casei, TMG expulsion was also studied in an L. casei hprK mutant (198). Compared to inducer expulsion by a wild-type strain, no effect of the hprK mutation on glucose- or mannose-triggered expulsion of accumulated TMG-6-P was observed.

Because most experiments suggesting a role for P-Ser-HPr in inducer expulsion were carried out with L. lactis vesicles, a ptsH1 mutant of this organism was constructed and tested for the expulsion of preaccumulated TMG-6-P and 2DG-6-P. Expulsion of both nonmetabolizable sugar derivatives was observed in the ptsH1 mutant. However, whereas 2DG expulsion was nearly identical in the wild type and the mutant, TMG expulsion was slowed by about 2.5-fold in the ptsH1 strain (569). Because L. lactis possesses only HPr and no HPr-like proteins (68), these results unequivocally established that inducer expulsion in this organism does not depend on the presence of P-Ser-HPr. Nevertheless, HPr seems to play an indirect role in TMG expulsion in L. lactis, which might partly explain the vesicle results, which led to the conclusion that P-Ser-HPr participates in inducer expulsion. Differences in the phosphorylation state of the EIIBC^Lac, which was proposed to catalyze not only TMG uptake but also TMG expulsion (719), in the wild-type and the ptsH1 strains were suggested to be responsible for the indirect effect of the ptsH1 mutation on TMG expulsion (569).

As explained above (see REGULATION OF CARBON METABOLISM IN GRAM-NEGATIVE ENTERIC BACTERIA), the inducer exclusion mechanism in enteric bacteria is mediated by dephospho-EIIA^Glc, which binds to several target proteins (permeases and catabolic enzymes), thereby inhibiting their activity and preventing the entry or the intracellular formation of the corresponding inducer. Several lines of evidence indicated that P-Ser-HPr might play a similar role in inducer exclusion in gram-positive bacteria (Fig. 6). In L. brevis, the galactose/H⁺ symporter GalP also catalyzes the uptake of TMG (193, 739). The addition of glucose arrests TMG uptake and causes the rapid efflux of preaccumulated TMG from the cells. Both effects of glucose were ascribed to P-Ser-HPr, as its structural homolog, Ser46Asp HPr (from B. subtilis) (952), labeled with ¹²⁵I was able to bind to the galactose/H⁺ symporter of L. brevis (972). Galactose as well as galactose derivatives stimulated the binding of ¹²⁵I-labeled Ser46Asp HPr to the galactose/H⁺ symporter about 10-fold. Coelectroporated metabolites known to stimulate the kinase activity of HprK/P (FBP, gluconate-6-P, and glycerate-2-P) as well as extravesicular glucose inhibited TMG uptake by L. brevis vesicles preloaded with B. subtilis HPr by electroporation about 10-fold (969). No inhibitory effect of glucose or its metabolites on TMG uptake was observed when Ser46Ala mutant HPr was used in place of wild-type HPr. Moreover, Ser46Asp mutant HPr slowed TMG uptake even in the absence of extravesicular glucose or intravesicular metabolites. Studies of 2DG uptake by L. brevis vesicles electroporated with wild-type or mutant HPrs and with glycolytic intermediates provided results that were almost identical to those obtained for TMG uptake (968). In addition, by expressing the L. brevis galP gene (encodes the galactose/H⁺ symporter) in a B. subtilis wild-type strain and an hprK mutant, both B. subtilis strains gained the capacity to efficiently take up TMG. When the B. subtilis strains also synthesized L. brevis wild-type or Ser46Ala or Ser46Asp mutant HPr, a strong inhibition of TMG uptake was observed with the strain producing Ser46Asp HPr (193), suggesting that the P-Ser-HPr-resembling Ser46Asp HPr causes inducer exclusion. However, when the strain producing L. brevis wild-type HPr was grown in the presence of glucose, which stimulates the formation of P-Ser-HPr, no inducer exclusion occurred. In addition, since the experiments with L. brevis vesicles were carried out in vitro with B. subtilis wild-type or mutant HPrs instead of the physiologically relevant L. brevis P-Ser-HPr, the proposed implication of P-Ser-HPr in inducer exclusion needed to be confirmed in vivo.

The importance of ATP-dependent HPr phosphorylation for inducer exclusion could be established by studying this regulatory process in L. casei and L. lactis mutants that are unable to form P-Ser-HPr. To detect a potential influence of the ptsH1 and hprK mutations on inducer exclusion in L. casei, transport studies with maltose and ribose were carried out (198, 919). In L. casei, maltose and ribose are taken up by ABC transport systems (570). When wild-type cells are grown in a medium containing glucose and either ribose or maltose, diauxic growth with a long-lasting lag phase is observed. In addition, maltose uptake is immediately arrested when glucose is added to maltose-transporting L. casei cells (198, 919), suggesting that glucose inhibits maltose uptake via an inducer exclusion mechanism. Interestingly, in ptsH1 (919) and hprK (198) mutants, glucose no longer exerts its inhibitory effect on maltose transport. These results imply that the phosphorylation of HPr at Ser-46 plays a role in inducer exclusion in L. casei. Inactivation of the ccpA gene had no effect on inducer exclusion (919), confirming that the P-Ser-HPr-dependent inhibitory effect of glucose on maltose uptake is due exclusively to an inhibition of the transport step and not to CCR of the maltose operon mediated via the P-Ser-HPr:CcpA complex. Additional evidence for P-Ser-HPr-mediated maltose exclusion from L. casei cells came from direct measurements of sugar consumption in the growth medium. In wild-type cells, maltose consumption is instantaneously arrested when glucose is added to the incubation mixture and does not restart before glucose is exhausted. In contrast, in the hprK and ptsH1 mutants, the addition of glucose causes only a short transient stop of maltose consumption, and the two sugars are subsequently simultaneously utilized (198, 919).

Interestingly, ribose transport was stimulated by the presence of glucose. Glucose stimulation of ribose uptake has also been reported for Lactobacillus sakei. Dephosphorylation of the PTS proteins in the presence of glucose was suggested to be responsible for the stimulating effect, as the inactivation of ptsI also led to enhanced ribose uptake (834, 835).

The repressive effect of glucose on maltose uptake and maltose consumption had also disappeared in an L. casei strain producing Ser46Thr mutant HPr, which is a poor substrate for HprK/P. Unexpectedly, inducer exclusion in L. casei was also prevented by the ptsH(Ile47Thr) mutation, although the amount of P-Ser-HPr detected in the S. salivarius ptsH(Ile47Thr) mutant was similar to that in the parental strain (263). The hydrophobic patch on the surface of HPr, which is formed by Ile-47, Met-48, and Met-51 in gram-positive bacteria (375), is involved in the interaction of P-Ser-HPr with CcpA (381, 787) and in the interaction of HPr with EI (260) and various EIIAs (139, 691, 949). The finding that the L. casei ptsH(Ile47Thr) mutant does not exhibit inducer exclusion suggests that this hydrophobic patch might also be important for the interaction of P-Ser-HPr with non-PTS permeases regulated by inducer exclusion.

Inducer exclusion studies were also carried out with an L. lactis ptsH1 mutant. Similar to L. casei, this organism takes up maltose and ribose via specific ABC transport systems (68). The inhibitory effect of glucose on the transport of the two non-PTS sugars in the wild-type strain had completely disappeared in the ptsH1 mutant (569).

An approximately twofold decrease in the P-Ser-HPr level was reported to lead to an almost complete loss of inducer exclusion in S. salivarius cells (870). The lower amount of P-Ser-HPr was due to specific point mutations in the ptsH gene or the ptsH leader sequence, which did not affect the uptake rate of PTS substrates, although the metabolism of PTS sugars was slowed. Wild-type S. salivarius cells immediately stop the uptake of the non-PTS sugar lactose or galactose when glucose or fructose is added to the growth medium and resume the metabolism of the non-PTS sugars only when glucose or fructose is exhausted (660). In contrast, the uptake of lactose or galactose by the ptsH mutants containing twofold-lower amounts of P-Ser-HPr was not affected by the presence of the PTS sugar glucose or fructose (870). To exert inducer exclusion, gram-positive bacteria therefore seem to require high levels of P-Ser-HPr.

In summary, the above-described results support the concept that inducer exclusion in gram-positive bacteria is regulated via the metabolite-activated formation of P-Ser-HPr. Considering the similarities between the maltose uptake systems from E. coli and those from the lactic acid bacteria L. casei and L. lactis, it is tempting to assume that the exclusion of maltose follows a mechanism analogous to that in gram-negative organisms, except that P-Ser-HPr instead of dephospho-EIIA^Glc binds to MalK (Fig. 6) (Table 1). This assumption is supported by a comparison of the sequences of the nucleotide binding and regulatory domains of MalKs from E. coli (117) and L. casei BL23 (570). The two proteins are composed of very similar N-terminal nucleotide binding domains (amino acids 1 to 241) (60% identity), while the C-terminal regulatory domains (amino acids 242 to 371) exhibit only 19% sequence identity (Deutscher, unpublished). The sequence difference in the regulatory domain might reflect the binding of different effectors: EIIA^Glc in gram-negative bacteria and P-Ser-HPr in gram-positive bacteria.

In addition to the most common mechanisms regulating gene expression, i.e., transcription activation or repression, bacteria also developed a mechanism based on transcription termination/antitermination, i.e., transcription attenuation, which was first described for the E. coli trp operon (60). Two principal mechanisms of transcription termination have been reported for prokaryotes: factor-dependent termination and intrinsic termination (965). In the first case, the RNA polymerase is halted at a terminator located at the end of a transcription unit. Interaction with a ρ factor is necessary to dissociate the DNA:RNA polymerase complex and to terminate transcription. In contrast, transcription units submitted to intrinsic termination contain a ρ-independent terminator. When the ρ-independent terminator is located between the transcription start site and the start codon (Fig. 8), the expression of the corresponding gene or operon usually occurs from a constitutive promoter. However, under noninducing conditions, transcription stops when the terminator is formed on the nascent mRNA. Formation of the terminator leads to the disruption of the elongation complex and to the release of short, incomplete transcripts. Under inducing conditions, an antiterminator binds to the mRNA in front of the terminator or to a region overlapping the terminator, thereby preventing the formation of the disruptive stem-loop structure and allowing the RNA polymerase to complete transcription of the corresponding gene or operon (Fig. 8). Antiterminators can be either proteins or oligonucleotides. When tRNAs function as antiterminators, binding to their target site on the mRNA is usually prevented when they are charged with their corresponding amino acid (136). Polypeptide antiterminators frequently respond to changes in the concentration of intracellular metabolites, which serve as cofactors and allow the corresponding antiterminator to bind to its target on the mRNA. For example, glycerol-3-P binds to and activates the B. subtilis antiterminator GlpP (749). Another mode of regulation was reported for antiterminators controlling the expression of genes and operons encoding either sugar-specific PTS components or enzymes catalyzing the extracellular degradation of polysaccharides to mono- or oligosaccharides, which are subsequently taken up via the PTS. This group of polypeptide antiterminators is regulated by reversible PTS-catalyzed phosphorylation. They possess an N-terminal RNA binding domain (first 55 amino acids) usually followed by two regulatory domains, each containing two conserved histidyl residues (Fig. 7), which are phosphorylated or dephosphorylated depending on whether the corresponding carbohydrate is present in the growth medium.

The B. subtilis levansucrase-encoding sacB gene (31, 806) and the E. coli bgl operon, which encodes a β-glucoside-specific EIIBCA (bglF), a 6-P-β-glucosidase (bglB) (504) (Table 4), and a carbohydrate-specific outer membrane porin (20), were the first two PTS-related transcription units for which experimental evidence for regulation by antitermination has been obtained. The B. subtilis sacB leader region contains an imperfect 28-base-long inverted repeat called sacR (833). When part of sacR was deleted or when a point mutation was introduced, sacB was expressed constitutively, and sacR was therefore proposed to function as a transcription terminator (31, 806). A sucrose-inducible sacB′-′lacZ fusion also became constitutive when sacR was mutated, which further supported the concept of transcription termination (806). Finally, S1 nuclease mapping revealed that in the absence of sucrose, mainly small transcripts terminating within sacR were formed. The presence of the inducer sucrose did not change the total number of transcripts but increased the number of long transcripts extending beyond the terminator sequence sacR (806).

An inverted repeat between the transcription initiation site and the start codon of the bgl operon of E. coli was also detected (505). In the noninduced state, more than 90% of the transcripts, which were initiated at the bgl promoter, were already terminated at the inverted repeat. Expression of a bgl-lacZ fusion containing the inverted repeat required an intact bglG (formerly bglC) gene, which is the first gene of the bgl operon, while bgl-lacZ fusions devoid of the inverted repeat exhibited constitutive, bglG-independent expression (505). Based on these results, it was proposed that the inverted repeat located between the transcription initiation site and the start codon of the E. coli bgl operon serves as a ρ-independent transcription terminator and that the protein encoded by bglG exerts a positive effect on bgl expression by functioning as an antiterminator. In agreement with this hypothesis, bglG disruption caused drastically lowered expression from the bgl promoter (504). However, a few bglG point mutations, including the one present in the bglC9 strain described previously (682), can lead to the constitutive expression of the bgl operon.

Terminators similar to those preceding B. subtilis sacB and the E. coli bgl operon have been found in front of many other transcription units encoding sugar-specific PTS components of the glucose/sucrose class. Mutation or partial deletion of the terminator sequences preceding the B. subtilis sacPA (24), bglPH (430), or ptsGHI (448) operon also led to the constitutive expression of the corresponding transcription unit and thus confirmed their negative role in mRNA synthesis.

Sequencing of the bgl operon allowed the detection of a second terminator located in the intercistronic region between bglG and bglF. Interestingly, the sequences preceding the two bgl terminators are very similar. They were assumed to form weak stem-loops partly overlapping the downstream terminator (785). Insertion of 6 bp in the region upstream from the terminator preceding bglG indeed diminished the induction of a bgl-lacZ fusion (505), suggesting that the region upstream from this terminator is important for antitermination and might represent the binding site for BglG. The conserved sequences preceding the terminators were called ribonucleotidic antiterminator targets (RATs) (32). In in vitro experiments, BglG was indeed able to specifically bind to an RNA probe containing the entire E. coli bgl RAT sequence but only the 5′ half of the terminator (344). In contrast, if an RNA probe containing the entire RAT sequence and the complete terminator was used, very weak binding of BglG was observed. The formation of the terminator was therefore assumed to prevent the formation of the BglG binding site, i.e., the RAT stem-loop. In agreement with this concept, BglG binding to the longer RNA probe was improved when a DNA fragment, which was complementary to the 3′ part of the inverted repeat and which therefore prevented formation of the terminator, was included in the assay mixtures. These results suggested that there is competition between the formation of the terminator and the RAT stem-loop (344). Under noninducing conditions, BglG cannot bind to its mRNA, and the terminator will therefore be formed. Calculations revealed that the formation of the terminator releases much more free energy than the formation of the RAT stem-loop. Under inducing conditions, the binding of the “activated” BglG is assumed to favor the formation of the RAT stem-loop. BglG thereby prevents the formation of the terminator partly overlapping the RAT sequence and allows the synthesis of long transcripts (344).

In B. subtilis, sequences very similar to those of the E. coli bgl RATs precede the above-mentioned terminator sacR (833) and the terminators located in front of the B. subtilis 1,3-1,4-β-d-glucan endoglucanase-encoding bglS (formerly licS) gene (585, 785), the sacPA operon (164), which encodes a sucrose-specific EIIBC (sacP) and an endocellular sucrase (sacA) (32), and ptsG (847), which encodes the glucose-specific EIIBCA (Table 4). RAT sequences are also present in front of PRD-containing antiterminator-controlled genes from other organisms (964). An extensive genetic analysis of the RAT sequence preceding the sacB gene confirmed its importance for sacB induction. Expression of sacB′-lacZ fusions containing mutations at positions assumed to be important for the formation of the RAT stem-loop was barely inducible with sucrose (32).

The antiterminators regulating the expression of the four above-mentioned B. subtilis transcription units have been identified. SacY regulates sacB transcription (832), SacT controls the expression of sacPA (164), LicT is necessary for bglS but also for bglPH expression (430), and GlcT controls ptsGHI transcription (847) (Table 4). They all exhibit significant sequence similarity to BglG of E. coli. The new family of antiterminators was called the BglG/SacY family, as the B. subtilis sacB gene and the E. coli bgl operon, which are regulated by SacY and BglG, respectively, were the first transcription units for which this termination/antitermination mechanism was established.

The RAT sequences preceding sacB and sacPA differ in only three positions (Table 4), and consequently, cross talk has been observed: SacY also controls sacPA expression, and SacT regulates the transcription of sacB (32, 146). In gel shift experiments, not only SacT but also SacY could bind to RNA fragments containing the RAT sequence of sacPA (23). In vitro interactions of the RNA binding domains of LicT, SacY, and GlcT with their respective RAT sequences on bglPA, sacB (168), and ptsGHI (448) mRNA have been demonstrated by carrying out surface plasmon resonance experiments. The dissociation constants determined in these experiments varied from 3 μM (SacY) to 10 nM (LicT). The S. carnosus GlcT homolog regulates the expression of glcA and glcB, which encode a glucose- and a β-glucoside-specific EIICBA, respectively (131). The RAT sequence in the leader region of glcAB of S. carnosus strongly resembles the RAT sequence of B. subtilis ptsGHI. As a consequence, S. carnosus GlcT interacts with the RAT sequence of B. subtilis ptsGHI, as was shown by mutant complementation assays and surface plasmon resonance experiments (413). The high affinity between LicT and its bglPA RAT allowed the determination of the solution structure of the LicT catalytic domain:mRNA complex (964). The antiterminator interacts mainly with the minor groove of the double-stranded RNA formed by two internal loops and the stem in between. In summary, the above-described results make it clear that RAT sequences are the binding sites for BglG/SacY-type antiterminators.

In addition to regulating sacB expression, SacY controls its own synthesis by regulating the expression of the sacS locus containing the two genes sacX and sacY (993). The sacX gene encodes an EIIBC with strong similarity to sucrose-specific EIIs. It is preceded by a RAT sequence and a terminator, which are separated by about 100 bp, whereas in all other transcription units regulated by BglG/SacY-type antiterminators, the RAT sequence and the terminator are overlapping or located next to each other. Nevertheless, both regulatory elements are functional, as mutations within the RAT sequence prevent the induction of sacXY expression, while a partial or complete deletion of the terminator sequence causes constitutive expression (885). It was therefore proposed that the formation of the RAT stem-loop preceding sacXY is stabilized by binding SacY, which in turn prevents the formation of the distant terminator via not-further-specified long-range effects (885).

After RAT sequences were identified as target sites for proteins of the BglG/SacY family, the signal controlling the affinity of the antiterminators for the RAT elements remained to be determined. The early studies on SacY-regulated sacB and BglG-controlled bgl operon expression had already suggested that these antiterminators might be controlled by the PTS. In the late 1970s, it was discovered that the inactivation of B. subtilis EI causes “constitutive” expression of sacB (i.e., synthesis of full-length mRNA) (264, 635). In fact, it was this property of the PTS that was used to successfully clone the B. subtilis ptsHI operon (283). A promoterless kanamycin resistance gene put under the control of the sacB promoter conferred sucrose-inducible kanamycin resistance after integration into the B. subtilis genome. After transposon mutagenesis was carried out, a strain expressing the kanamycin resistance gene in the absence of sucrose was isolated. The mutant had the transposon inserted into the ptsHI operon (283), thus confirming that ptsHI inactivation leads to “constitutive” expression from the sacB promoter.

Another study from the same year showed that the inactivation of E. coli bglF, the gene located downstream from bglG, caused “constitutive” expression of the bgl operon (i.e., synthesis of full-length mRNA) (504). Because bglF encodes a β-glucoside-specific EIIBCA (79, 785), it was concluded that BglG/SacY-type antiterminators might be controlled by a phosphorylation cascade formed by the general PTS proteins EI and HPr and those sugar-specific EIIAs and EIIBs, the synthesis of which is regulated by the antiterminator. In vitro phosphorylation of BglG by P∼EIIBCA^Bgl was indeed reported (18). However, the regulation of antiterminators by PTS-catalyzed phosphorylation turned out to be very complex. First, EI and HPr were later found to be able to phosphorylate most antiterminators in the absence of EIIA and EIIB. Second, the activity of some antiterminators depends on functional EI and HPr, and the inactivation of ptsI therefore does not lead to the “constitutive” expression of the genes controlled by these antiterminators but, on the contrary, lowered or prevented their expression. Third, BglG/SacY-type antiterminators are usually phosphorylated at more than one of the four conserved histidyl residues in their PRDs. Extensive genetic, biochemical, and structural studies carried out mainly with the four B. subtilis antiterminators were necessary to understand in detail how BglG/SacY-type antiterminators are regulated.

In principle, two types of phosphorylation with antagonistic effects on the antiterminator activity have to be distinguished. P∼EIIB-requiring phosphorylation in PRD1 inactivates the antiterminator (site of negative regulation). The absence of phosphorylation in PRD1 leads to full-length expression of the corresponding transcription units. By contrast, phosphorylation by EI and HPr in PRD2 stimulates the activity of antiterminators (site of positive regulation). The absence of phosphorylation in PRD2 leads to CCR of the corresponding genes and operons via a CcpA-independent mechanism (Fig. 8).

“Constitutive” expression from the promoter of an antiterminator-controlled operon following inactivation of the related EII has been observed not only for E. coli bglF (504) but also for several B. subtilis transcription units as well as for the lac operon of L. casei (289). In B. subtilis, disruption of sacX (147), bglP (430, 454), or ptsG (847) leads to inducer-independent expression of sacB, bglPH, and ptsGHI, respectively. As mentioned previously, B. subtilis ptsI mutants exhibit “constitutive” sacB transcription (264, 283, 635), and the deletion of ptsH was reported to lead to the synthesis of full-length ptsGHI mRNA (847). It therefore appeared that a functional phosphorylation cascade composed of the general PTS proteins EI and HPr and sugar-specific EIIAs and EIIBs leads to the inactivation of the corresponding antiterminator.

Interestingly, SacX, the inactivation of which also results in “constitutive” sacB expression (147), contains EIIB and EIIC but no EIIA domain, and it was therefore concluded that EIIB, the last link in the PTS phosphorylation chain (Fig. 1), regulates the induction of antiterminator-controlled transcription units. Experiments with the L. casei lac operon confirmed the importance of P∼EIIB for antiterminator-mediated induction, as the inactivation of lacE as well as lacF (encoding EIICB^Lac and EIIA^Lac, respectively) led to the synthesis of full-length lacTEGF mRNA (289). A different mechanism had originally been proposed for the regulation of E. coli BglG, which was based on the erroneous assumption that His-306 of E. coli EIIBCA^Bgl would be the phosphorylation site in the EIIB domain (784). The replacement of His-306 with nonphosphorylatable amino acids prevents β-glucoside transport but, in contrast to bglF disruption, does not lead to the synthesis of full-length bgl mRNA. It was therefore proposed that EIIA and not EIIB of EIIBCA^Bgl would regulate BglG activity, and EIIA^Glc was also suggested to phosphorylate and inhibit BglG (783). This concept proved to be wrong when the phosphorylation site in the EIIB domain of E. coli EIICB^Glc was identified as being Cys-421 (547), which is equivalent to Cys-24 in EIIBCA^Bgl. A strain containing a bglF(Cys24Ser) mutation not only was unable to transport β-glucosides but also expressed a bgl-lacZ fusion in the absence of an inducer (β-glucoside) (120), confirming that E. coli BglG is also regulated via the EIIB domain of the corresponding PTS permease.

To determine which of the usually four conserved histidyl residues in the PRDs would be the target for P∼EIIB-mediated regulation, extensive genetic analyses were carried out by replacing the histidyl residues in the B. subtilis antiterminators SacY and LicT with various amino acids and by measuring the effect of the mutations on reporter gene fusions in different genetic backgrounds. Replacement of His-99 in PRD1 of SacY with a Tyr (883) or Ala (884) causes constitutive expression of the sacB gene. In contrast, replacing the three other conserved histidyl residues in SacY with Tyr has little effect on the antitermination activity (883). Similarly, replacement of His-100 in B. subtilis LicT (884) or His-101 in L. casei LacT (288), the equivalents of His-99 of SacY, with alanine leads to the synthesis of full-length bglPH or lacTEGF mRNA, respectively. These results strongly suggest that phosphorylation of the first conserved histidyl residue in PRD1 by the corresponding P∼EIIB prevents induction by PRD-containing antiterminators. Attempts to mimic phosphorylated antiterminators by replacing the first phosphorylatable histidine in PRD1 with negatively charged Asp or Glu were only partly successful. Putting a negative charge at position 99 by replacing His-99 with Glu or Asp indeed inactivates SacY and abolishes sacB induction (884). Similarly, the His101Asp replacement in L. casei LacT prevents lac operon expression (288). However, although His100Asp LicT exhibited significantly lower activity than His100Ala LicT, it was not completely inactive (884), indicating that in LicT, the His100Asp replacement is not completely phosphomimetic. Moreover, replacement of His-104 with an Asp in B. subtilis GlcT even led to strongly “constitutive” expression of a ptsG′-′lacZ fusion (35), suggesting that the mutant protein does not at all resemble GlcT phosphorylated at the first conserved histidine in its PRD1.

The original concept for the regulation of BglG/SacY-type antiterminators implied that P∼EIIBs directly phosphorylate and thereby inactivate their corresponding antiterminator (18). The finding that the first conserved histidine in PRD1 of several antiterminators is phosphorylated in vitro by P∼His-HPr therefore came as a surprise (413, 781, 883, 884). For instance, His-99, the site of negative regulation in B. subtilis SacY, is efficiently phosphorylated in vitro by EI and HPr in the absence of SacX (883). Because P∼His-HPr is also necessary for the phosphorylation of EIIBs, it was not clear whether P∼EIIBs directly phosphorylate their corresponding antiterminators or merely stimulate the P∼His-HPr-mediated phosphorylation. It proved difficult to distinguish between the two possibilities, as it was not easy to obtain P∼EIIB preparations that were free of HPr (EIIBs are often fused to their membrane-integrated EIICs, and HPr usually sticks to membranes). Because the inactivation of EI as well as SacX leads to constitutive expression of the SacY-controlled sacB gene, it was proposed that in the cells, efficient P∼His-HPr-mediated phosphorylation of SacY at His-99 would require the presence of P∼SacX. GlcT of S. carnosus also becomes phosphorylated in vitro by EI and HPr in the absence of an EIIB, and mass spectrometry with proteolytic fragments obtained from phosphorylated and dephospho-GlcT revealed that phosphorylation occurs primarily at His-105 (413), the equivalent of His-99 in SacY. In the case of S. carnosus GlcT, phosphorylation in PRD1 was assumed to stimulate the antitermination activity. In contrast, in experiments with GlcT from B. subtilis, P∼His-HPr barely phosphorylated PRD1 but mainly transferred its phosphoryl group to His-210 in PRD2 (781) (for PRD2 phosphorylation, see the next section). In fact, the first conserved histidine in PRD1 of B. subtilis GlcT has been shown to become phosphorylated by P∼EIIB^Glc. When the soluble B. subtilis EIIBA^Glc domains were synthesized without the membrane-integrated EIIC^Glc (to which they are normally fused), HPr-free P∼EIIBA^Glc could be prepared, which could phosphorylate wild-type GlcT but not GlcT(His104Ala). The antiterminator became phosphorylated in PRD2 only when HPr was added to the phosphorylation mixture containing P∼EIIBA^Glc and GlcT(His104Ala), thus confirming the nearly complete absence of HPr from the P∼EIIBA^Glc preparation (781). It therefore seems that phosphorylation of some antiterminators (B. subtilis SacY and S. carnosus GlcT) at PRD1 is catalyzed by P∼His-HPr, and this reaction is stimulated by the corresponding P∼EIIB, while phosphorylation in PRD1 of other antiterminators (GlcT of B. subtilis) occurs via the corresponding P∼EIIB.

Based on genetic studies with licT, it was suggested that P∼EIIB domains might also sequester their corresponding antiterminator (884). A similar mechanism is operative for the E. coli transcription regulator Mlc, which is sequestered by the unphosphorylated EIIB domain of EIICB^Glc (458, 588, 856) (see “REGULATION OF CARBON METABOLISM IN GRAM-NEGATIVE ENTERIC BACTERIA”). Preliminary evidence for the formation of a complex between an antiterminator and an EII was obtained for S. carnosus GlcT, which exists as an active dimer and an inactive monomer. Dimerization is favored by P∼His-HPr-mediated phosphorylation at His-105. Membrane fragments containing EIICBA^Glc (and possibly P∼EIICBA^Glc) specifically interacted only with the dimers in a GlcT monomer/dimer mixture (413). Calorimetric studies had established that the binding of sugar molecules to their respective EIIs, which leads to the dephosphorylation of the EIIB domain, causes extensive structural changes (546). In the case of S. carnosus EIICBA^Glc, the structural changes accompanying the binding of glucose were assumed to lead to a release of active P∼GlcT dimers, thus allowing the expression of the glcAB operon in response to the presence of a substrate.

Unlike antiterminators from B. subtilis and L. casei, the negative regulation site of BglG from E. coli was originally thought to be located in PRD2, and His-208 was reported to be phosphorylated by P∼EIIBCA^Bgl (121). However, the phosphorylation mixtures for BglG phosphorylation contained not only P∼EIIBCA^Bgl but also EI and HPr. Like most antiterminators, BglG is phosphorylated by EI and HPr in PRD2 (287) (see the next section), and Chen et al. (121) probably misinterpreted the P∼His-HPr-dependent modification as being P∼EIIBCA^Bgl-dependent phosphorylation. In fact, the negative regulation site of E. coli BglG has been identified as His-101, and phosphorylation at this amino acid requires the presence of P∼EIIBCA^Bgl (285).

In summary, it can be concluded that induction via most BglG/SacY-type antiterminators is regulated by P∼EIIB-controlled or -mediated phosphorylation of the first conserved histidyl residue in PRD1, which inhibits the activity of the antiterminator. In the presence of the corresponding inducer, i.e., the carbohydrate transported by (or, in the case of B. subtilis, SacX probably bound to) the EII regulating the antiterminator, the antiterminator will barely be phosphorylated at its first conserved histidine in PRD1, because phosphorylation of the inducer is probably faster than phosphorylation of the antiterminator (482) or rephosphorylation of the EIIB by its corresponding P∼EIIA (Fig. 8). For B. subtilis SacY, the presence of sucrose in the growth medium indeed significantly reduced the extent of its in vivo phosphorylation (359). Antiterminator molecules that are not phosphorylated at the first conserved histidine in PRD1 are able to bind to their RAT sequence, thus preventing the formation of the terminator and allowing the synthesis of full-length mRNA of their target gene or operon. If the inducer is absent, P∼EIIB will be formed, which leads to phosphorylation of the first histidine in PRD1 and/or sequestration of the corresponding antiterminator. Both effects prevent the binding of the antiterminator to its RAT sequence and therefore the induction of the corresponding gene or operon. According to genetic analyses with licT of B. subtilis (884) and bglG of E. coli (285), phosphorylation at the second conserved histidine in PRD1 plays a detectable but minor role in the induction process. By contrast, replacing either His-101 or His-159 in L. casei LacT with alanine leads to “constitutive” expression of the lac operon, suggesting that in this antiterminator, the two conserved histidines in PRD1 contribute equally to the induction process (288).

Based on the effect of ptsI or ptsH mutations on the activity of BglG/SacY-type antiterminators, two classes of transcription attenuators can be distinguished. The first class includes B. subtilis SacY and GlcT. Mutations in ptsI or ptsH result in the “constitutive” expression of their target genes sacB (147, 264, 635) and ptsGHI (847), respectively. In these mutants, neither P∼His-HPr nor P∼EIIB is formed, which prevents the inactivating phosphorylation at PRD1 of the antiterminator in the absence of an inducer. The second class of antiterminators includes B. subtilis SacT and LicT and E. coli BglG. The synthesis of the full-length mRNA of their target genes sacPA (24) and bglPH in B. subtilis (429, 430, 454, 482) and bglGFB in E. coli (287) is almost completely prevented when ptsH or ptsI is inactivated. These results were surprising, as we had learned in the previous section that the inactivation of, for example, the EIIBCA^Bgl-encoding bglP leads to the synthesis of full-length mRNA from the LicT-controlled bglPH promoter (430, 454). Because EI and HPr are essential for the phosphorylation of EIIBCA^Bgl, which in turn inactivates LicT by phosphorylating it at His-100 in PRD1, the disruption of ptsH or ptsI should also lead to the synthesis of full-length bglPH mRNA. The observed dependence of LicT, SacT, and BglG activity on functional EI and HPr therefore suggested that the second class of antiterminators is subject to dual control by PTS proteins: positive regulation by P∼His-HPr and negative control by their corresponding P∼EIIBs. Because the specific replacement of His-15, the site of EI-catalyzed phosphorylation in HPr, with an alanine also prevented LicT activity (429), it was likely that EI and HPr exert their positive effect by phosphorylating this class of antiterminators. Indeed, SacT (23), LicT (482), and BglG (287) are phosphorylated by PEP, EI, and HPr. In LicT, all four conserved histidines are phosphorylated by the general PTS proteins, with the main site of phosphorylation being His-207 in PRD2 (482, 884). Because the P∼EIIB-dependent negative effect is mediated via the first conserved His in PRD1 (see the previous section), it was likely that the positive effect of phosphorylation by EI and HPr is mediated via PRD2. Experiments with SacY/LicT hybrid proteins confirmed this assumption. When PRD2 of the EI/HPr-independent SacY was replaced with PRD2 of LicT, the resulting hybrid protein was EI and HPr dependent (884). This concept was further supported by an extensive genetic analysis, during which the conserved histidines in PRD2 of B. subtilis LicT were replaced with various amino acids and the effect of the mutations on bglP′-lacZ expression was measured in different pts genetic backgrounds. Replacement of His-207 or His-269 or both conserved histidines in PRD2 with Ala led to a complete loss of LicT antitermination activity, i.e., prevented the synthesis of full-length mRNA from the bglPH promoter even in the presence of its inducer salicin or in a bglP background. In contrast, in a bglP mutant, if either one of these two histidines or both were replaced with an Asp, which probably mimics a phosphorylated histidine, expression from the bglPH promoter remained “constitutive” irrespective of whether ptsHI were functional or deleted (884). These results demonstrate that PEP-, EI-, and HPr-mediated phosphorylation of PRD2 in antiterminators of the second class stimulates their activity. This is also true for E. coli BglG, where His-208, the only conserved phosphorylatable histidine in PRD2, was first claimed to be the site of negative regulation by P∼EIIBCA^Bgl (121) but was later identified as the site of positive regulation by P∼His-HPr (285, 287). Interestingly, E. coli FruB, a fusion protein composed of HPr- and EIIA^Fru-like domains (267), can replace HPr in BglG phosphorylation and activation (287).

Surprisingly, although the activity of B. subtilis SacY and GlcT does not depend on EI and HPr, they are phosphorylated by PEP, EI, and HPr in their PRD2 (781, 883). It seems that during the course of evolution, SacY and GlcT developed a PTS-independent antitermination activity but retained the ability to become phosphorylated in PRD2. On the contrary, GlcT of S. carnosus, which is probably also EI/HPr independent, is not phosphorylated in its PRD2, as only one major P∼His-HPr-dependent phosphorylation site, His-105 in PRD1, could be detected (413). Interestingly, EI/HPr-independent antiterminators seem to regulate the expression of transcription units that are insensitive to CCR, whereas the EI/HPr-dependent LicT and SacT control operons that are preceded by cre's recognized by the P-Ser-HPr:CcpA complex (429). It was therefore assumed that phosphorylation of LicT and SacT by PEP, EI, and HPr might serve as an additional, CcpA-independent CCR mechanism.

The first evidence for antiterminator-mediated CCR came from studies with a B. subtilis ccpA mutant. In this mutant, expression of a bglP′-lacZ fusion was not completely relieved from CCR. The residual CCR (about 20%) disappeared when the terminator preceding the bglPH operon was deleted, suggesting that bglPH is regulated by two CCR mechanisms: one that is CcpA dependent and one that is terminator dependent (429). Interestingly, the residual CCR also disappeared when a ptsH1 mutation (which prevents the formation of P-Ser-HPr) was introduced into the ccpA mutant. It was therefore assumed that the CcpA-independent CCR mechanism operative for the bglPH operon was based on diminished phosphorylation in PRD2 of LicT due to the formation of P-Ser-HPr (a poor substrate for PEP-dependent phosphorylation by EI) during growth on a rapidly metabolizable carbon source. This assumption was supported by studies with B. subtilis mutants producing LicTs in which either one of the two conserved histidines in PRD2 was replaced with Asp. Expression of a bglP′-lacZ fusion remained inducible in both mutants, but β-galactosidase activity was less severely repressed by glucose than in a strain producing wild-type LicT (909). Unequivocal proof for the antiterminator-mediated CCR mechanism came from studies with strains producing EI/HPr-independent mutant LicTs, which were called LicT(Pia) (Pia stands for PTS-independent antitermination). Starting from a strain carrying a bglPH′-aphA3 (aphA3 confers kanamycin resistance to bacteria) and a bglP′-lacZ fusion as well as the LicT-inactivating ptsGHI deletion, several spontaneous mutants that had gained the ability to express the LicT-dependent bgl fusions despite the absence of functional EI and HPr were isolated (483). The isolated mutants were all affected in licT, and the various mutations apparently render LicT activity independent of functional EI and HPr. The licT(Pia) mutants can be divided into two classes: mutations affecting PRD1 (they lead to constitutive expression from the bglPH promoter) and mutations affecting PRD2 (they remain inducible). The latter type of mutant LicTs resembles the naturally EI/HPr-independent SacY and GlcT. Interestingly, the residual CCR of the bglPH operon detectable in ccpA mutants disappeared when wild-type licT was replaced with one of the licT(Pia) alleles (483), thus confirming that it is the poor phosphorylation of PRD2 in LicT during the uptake of rapidly metabolizable carbon sources that triggers CcpA-independent CCR.

Based on the results obtained with the various LicT(Pia) proteins, distinct amino acids were proposed to be important for the transfer of the activating signal from PRD2 via PRD1 to the RNA binding domain (483), which was called CAT (coantiterminator) (964). When the crystal structures of the two PRDs of the constitutively active His207Asp, His269Asp mutant LicT (909) and of inactive wild-type LicT (292) were solved, a more detailed understanding of the structural changes involved in LicT activation was possible. In fact, the structures of the PRD2 of the two crystallized LicT forms are completely different, while the amino acid backbones of their PRD1s are almost identical. In inactive wild-type LicT, the two PRD2s of the LicT dimer make no contact to each other but are in contact with the two PRD1s. The four phosphorylatable histidines in the PRD2s are exposed to the surface and easily accessible for phosphorylation. In the activated His207Asp, His269Asp mutant LicT, a 180° swing movement of the C-terminal domain leads to extensive structural changes, placing the aspartyl residues (which replace the four histidines) in the center of PRD2, thus rendering them inaccessible from the outside. Two major rotations are responsible for the rearrangement of the PRD2s. Both take place in the interdomain region between PRD1 and PRD2, with one occurring at residues Leu-165 and Asn-166 and the other occurring at Met-169. As a consequence, the two PRD2s of His207Asp, His269Asp mutant LicT are in close contact, whereas most interactions with PRD1 are suspended. These alterations are probably responsible for the changes in the interdomain region connecting PRD1 and CAT, which in turn are thought to cause rearrangements of the CAT domain, ultimately leading to the activation of LicT (292). The monomer/dimer transition does not seem to be important for LicT activation, as the RNA binding domain CAT and active His207Asp, His269Asp mutant LicT as well as inactive dephospho-LicT form dimers (964). Nevertheless, the monomer/dimer transition might be important for the regulation of other antiterminators (56).

The structure of the RNA binding domain of the two antiterminators SacY and LicT has been intensively studied by NMR (510) and X-ray crystallography (910). The solution structure of the RNA binding domain of LicT (first 55 amino acids) attached to its mRNA has also been solved (964). However, the structure of an entire BglG/SacY-type antiterminator has so far not been determined.

Based on the above-described results, the following model for CCR mediated by EI/HPr-dependent antiterminators can be proposed. In the presence of a carbon source rapidly transported via the PTS, the antiterminators will have to compete with the corresponding sugar-specific EIIA for the common phosphoryl donor P∼His-HPr. P∼His-HPr probably transfers its phosphoryl group primarily to EIIAs, thus leaving the antiterminators in the unphosphorylated, less active state. In in vitro experiments, P∼His-HPr indeed phosphorylates EIIAs much faster than antiterminators (482). In addition, the ATP-dependent phosphorylation of HPr at Ser-46 in low-G+C gram-positive bacteria, which is triggered by the presence of rapidly metabolizable carbon sources, drastically slows the PEP-dependent phosphorylation of HPr at His-15 and therefore further diminishes the activation of LicT by P∼His-HPr-dependent phosphorylation. The proposed complex mechanism of induction and CCR of operons controlled by EI/HPr-dependent antiterminators is outlined in Fig. 8. The model requires that during PTS sugar uptake, the corresponding EIIB domain is present mainly in the dephospho form and therefore does not phosphorylate the first conserved His in PRD1 of LicT. On the other hand, P∼His-HPr must be present in sufficient amounts to allow the phosphorylation in PRD2, which implies that PTS operons, the expression of which is regulated by an EI- and HPr-dependent antiterminator, encode low-capacity transport systems, as strong autorepression would otherwise occur. Probably in order to prevent autorepression, the operons for the B. subtilis high-capacity glucose and sucrose PTS are regulated by EI- and HPr-independent antiterminators.

In a several-years-old review on PRD-containing transcription regulators, the BglG/SacY antiterminator family was reported to contain 13 members (844). In addition to the above-mentioned antiterminators, it included LicT of Bacillus amyloliquefaciens, AbgG of Clostridium longisporum (87), ArbG of Pectobacterium chrysanthemi (formerly Erwinia chrysanthemi) (208), BglR of L. lactis (42), CasR of Klebsiella oxytoca (446), and SurT of G. stearothermophilus (473). Owing to the many genome sequences completed in the meantime, a simple BLAST search yields numerous new potential family members. A few of these novel PRD-containing antiterminators have been studied in more detail. BvrA of L. monocytogenes controls the expression of the bvrABC operon encoding the antiterminator, a β-glucoside-specific EIIBCA, and a presumed ADP-ribosylglycohydrolase (81), respectively. In L. monocytogenes, repression of several PrfA-controlled virulence genes by the β-glucosides cellobiose and salicin requires an intact bvr locus, while repression by arbutin, glucose, or fructose persists in a bvrAB mutant. BglG of L. plantarum was reported to control the expression of the bglGPT operon encoding the antiterminator BglG, a β-glucoside-specific EII, and a 6-P-β-glucosidase (513), while LicT of S. mutans regulates an esculin-specific PTS (141). C. acetobutylicum ScrT was proposed to control the expression of the scrAKB operon encoding a sucrose-specific EII, a fructokinase, and a sucrose-6-P hydrolase, respectively (860). In the actinobacterium Corynebacterium diphtheriae, a BglG/SacY-type antiterminator-encoding gene is located downstream from the EIIBCA^Glc-encoding ptsG gene (631). BglT of Pectobacterium carotovorum probably controls the expression of an operon that encodes a β-glucoside-specific PTS and strongly resembles the arb operon of P. chrysanthemi (19). BglG/SacY-type antiterminators are especially abundant in clostridia. With nine BglG/SacY homologs, Clostridium difficile seems to be the organism that contains the highest number of PRD-containing antiterminators, followed by L. plantarum, which contains five such proteins. In fact, BglG/SacY-type antiterminators are present in most firmicutes including bacilli, clostridia, enterococci, lactobacilli, lactococci, listeriae, leuconostocs, staphylococci, streptococci, etc. BglG/SacY-type antiterminators occur less frequently in gram-negative bacteria. So far, in addition to E. coli, PRD-containing antiterminators have been detected in proteobacteria such as P. carotovorum (BglT) (19), P. chrysanthemi (ArbG) (208), Klebsiella oxytoca (CasR) (446), Klebsiella aerogenes (BglG) (690), Shigella flexneri, Shigella sonnei, Shigella boydii, Photorhabdus luminescens, and Yersinia enterocolitica. Interestingly, a gene coding for a BglG/SacY-type antiterminator is also present in the pathogenicity island of a uropathogenic E. coli strain (393). It is preceded by a gene encoding an EIICB resembling the glucose-specific EII (PtsG) of E. coli. However, whether this system is related to the virulence of this organism is not known. BglG/SacY homologs are also present in the actinobacteria Bifidobacterium longum, Corynebacterium diphtheriae, Propionibacterium acnes, and Symbiobacterium thermophilum. The antiterminator-controlled expression of a fructose PTS in Bifidobacterium breve has been studied in more detail (534a).

Transcription activators are DNA binding proteins usually containing a helix-turn-helix motif. They bind upstream from their target promoter, interact with a specific RNA polymerase holoenzyme, and thereby stimulate transcription. Based on the σ factor associated with the RNA polymerase, different classes of transcription activators can be distinguished. The first class comprises NifA/NtrC-type transcription activators. They usually interact with RNA polymerase containing σ⁵⁴ (also called RpoN in E. coli and SigL in B. subtilis). They are composed of an N-terminal regulatory domain, which is usually a receiver domain of a two-component system, a central domain interacting with σ⁵⁴, and a C-terminal helix-turn-helix motif. A nucleotide binding site (Walker motifs A and B) is present in the central domain. In the case of NtrC, phosphorylation of its receiver domain by the sensor histidine kinase NtrB (396, 608) strongly stimulates its ATPase activity, which allows NtrC to enhance the formation of open complexes by the RNA polymerase/σ⁵⁴ holoenzyme (943). NifA/NtrC-type transcription activators bind to the DNA at about 100 bp upstream from their target promoters, and DNA bending is necessary to allow an interaction with σ⁵⁴ (705). In a few cases, the binding site for the transcription activator is located even more than 1 kb away from the target promoter (698, 809). Owing to their mode of action, these transcription activators are also called enhancer DNA binding proteins (EBP).

A second class of transcription activators interacts with the major σ⁷⁰-containing RNA polymerase holoenzyme. This class is very large and heterogenous and includes regulator proteins, which, depending on the location of their binding site, function as transcription activators (binding upstream from the promoter) and/or repressors (the binding site overlaps the promoter or is located downstream from it). This class includes the DeoR family of transcription regulators containing repressors such as E. coli GlpR (790) and SrlR (formerly GutR) (961), B. subtilis IolR (981), and L. lactis LacR (986) but also transcription activators such as E. coli FucR (125), B. anthracis AcpA (414), and S. pyogenes VirR (537). Often, DeoR-like transcription regulators do not contain a clearly identifiable nucleotide binding site. Recognition of their target sequence is controlled by low-molecular-weight effectors such as sugar phosphates, which interact with the C-terminal ligand-binding domain.

Several transcription activators belonging to one of the above-described classes are regulated neither by ligand-binding (DeoR type) nor via phosphorylation by a sensor kinase (NifA/NtrC type) but instead possess PRDs, suggesting that their activity is controlled by PEP-dependent, PTS-catalyzed phosphorylation. PRDs thus seem to be used not only to regulate the RNA binding activity of antiterminators but also to control the DNA binding function of some members of the above-described classes of transcription activators (Fig. 7). Interestingly, all PRD-containing transcription activators also possess an EIIA and an EIIB domain. The EIIB domain in transcription activators always exhibits similarity to EIIBs of the galactitol class PTS (295). In contrast, the EIIA domain belongs to the mannose class PTS in regulators with an NifA/NtrC-like σ⁵⁴ binding domain (178) and to the mannitol/fructose class PTS in regulators with a DeoR-like DNA binding domain. The type of EIIA domain present in transcription regulators seems to be related to the specificity of the PTS that they control. Three cases can be distinguished: (i) transcription activators containing NifA/NtrC- and EIIA^Man-like domains control the expression of operons coding for PTSs belonging to the mannose or lactose class; (ii) transcription activators with DeoR- and EIIA^Mtl-like domains regulate the expression of operons coding for PTSs of the mannitol, lactose, or glucitol class; and (iii) PRD-containing antiterminators (see the previous section) are devoid of an EIIA domain and primarily regulate the expression of operons coding for PTSs belonging to the glucose/sucrose class but also several PTSs of the lactose class.

The best-studied PTS-controlled transcription activator is B. subtilis LevR, which regulates the expression of the lev operon (165). The levR gene is located just upstream from the lev operon, which encodes the EIIs for a low-capacity fructose/mannose-specific PTS (LevD, LevE, LevF, and LevG) and an extracellular levanase (SacC) capable of degrading fructose polymers such as levan (110, 519, 522). Expression of the lev operon occurs from a σ⁵⁴ (σ^L)-dependent −12, −24 promoter, is induced by low concentrations of fructose, and is repressed by rapidly metabolizable carbon sources. Although the Lev PTS transports fructose, high concentrations of this sugar repress lev operon expression, because B. subtilis also possesses a high-capacity fructose PTS encoded by the fruA gene (437).

LevR stimulates the activity of the RNA polymerase/σ⁵⁴ holoenzyme after binding to a UAS, which was identified by footprint experiments (521) and is centered at about 110 bp upstream from the lev promoter. LevR contains an N-terminal helix-turn-helix motif (positions 34 to 54) (521) and thus differs from NifA/NtrC-type transcription activators with a receiver domain of two-component systems, which possess the DNA binding motif at the C terminus. The second domain of LevR resembles the central domain in NifA/NtrC-type transcription activators (165). This domain probably interacts with RNA polymerase associated with the alternate sigma factor σ⁵⁴ and contains Walker motifs A (148-GPTGSGKS-155) and B (221-GILFMDEI-228) for ATP binding and hydrolysis. In LevR, the NifA/NtrC-like central domain is followed by a complete PRD. A second, truncated PRD containing only one conserved histidyl residue is located at the C terminus of LevR (see Fig. 7 for the domain structure of LevR).

The region separating the complete and the truncated PRD of LevR (from positions 575 to 820) is much longer than that in BglG/SacY-type antiterminators. A BLAST search with the sequence separating the two PRDs in LevR revealed that the region from amino acids 578 to 707 exhibits significant similarity to EIIAs of the mannose class PTS (178) (Fig. 7). A histidine equivalent to the phosphorylatable histidine of E. coli EIIA^Man (110, 840) is present in position 585. The strongest similarity was detected with EIIA^Man of Vibrio furnissii. In addition, the sequence between the EIIA^Man-like domain and the truncated PRD from positions 708 to 807 was found to exhibit significant similarity to EIIBs from the galactitol class PTS (GatB, SgaB, and SgcB of E. coli) (295). These EIIBs contain a conserved cysteyl residue close to the N terminus, and an equivalent cysteyl residue is present in the EIIB-like domain of LevR at position 718. It therefore seems that LevR evolved by numerous domain fusions and insertions. It is tempting to assume that the two PRDs were added to the central domain of a NifA/NtrC-type regulator before the EIIA and EIIB domains were inserted between the PRDs. LevR contains a total of five potential PTS phosphorylation sites, two in PRD1, one in the EIIA and EIIB domains, and one in PRD2 (Fig. 7).

Interestingly, the L. casei lev operon (levABCDX) is not preceded by a −12, −24 promoter but is preceded by a −10, −35 promoter. It was therefore assumed that L. casei LevR, which controls the expression of the lev operon of this organism, does not interact with σ⁵⁴. Indeed, while the disruption of levR strongly diminished lev operon expression, inactivation of the σ⁵⁴-encoding rpoN gene had no effect (534). In addition, the amino acid sequence motif -GAFTGA-, known to be essential for the interaction of NifA/NtrC-type EBPs with σ⁵⁴, is absent from the central domain of L. casei LevR. Nevertheless, the domain organization of L. casei LevR is identical to that in B. subtilis LevR, and the two proteins exhibit 41% sequence similarity. So far, L. casei LevR is the only known protein of this family that does not interact with σ⁵⁴ but that interacts with another σ factor (presumably σ⁷⁰).

The binding of B. subtilis LevR to its UAS and/or its interaction with σ⁵⁴ was thought to be regulated by PTS-catalyzed phosphorylation (165). This assumption was based on the observation that mutations affecting LevD or LevE, the fructose-specific EIIA and EIIB of the Lev PTS, caused strong constitutive expression from the lev promoter (520, 522). Mutants carrying deletions of the EI- and HPr-encoding ptsI and ptsH genes or synthesizing His15Ala or Ser46Asp mutant HPr (the latter is a poor substrate for EI) also exhibit elevated LevR activity, which, however, is considerably lower than that in levD (EIIA^Lev) and levE (EIIB^Lev) mutants (526, 846). Deletion of ptsHI in a levD or levE strain diminished the constitutive LevR activity, indicating that EI and HPr exert a positive effect on LevR function (846). These results suggest that similar to the second, PTS-dependent class of BglG/SacY-type antiterminators, LevR might also be regulated by multiple PTS-catalyzed phosphorylations in its PRDs and possibly also in the EIIA and EIIB domains (Table 1).

Indeed, PEP-dependent, EI- and HPr-mediated phosphorylation of B. subtilis LevR occurs at His-585, which was originally thought to be part of PRD1 (520) but was later identified as the phosphorylatable His in the EIIA^Man domain (178). His-488 of L. casei LevR is equivalent to His-585 of B. subtilis LevR and was accordingly identified as a phosphorylation site by P∼His-HPr (534). Replacement of this phosphorylatable His in LevR of B. subtilis or L. casei with an alanine diminishes lev operon expression, thus supporting the concept that EI/HPr-catalyzed phosphorylation exerts a positive effect on the transcription activation function of LevR (520, 534).

A second PEP-dependent phosphorylation requiring EIIA^Lev and EIIB^Lev (in addition to EI and HPr) occurs at the single conserved histidyl residue present in the C-terminally truncated PRD: His-869 in B. subtilis LevR (520) and His-776 in L. casei LevR (534). Experiments with B. subtilis [³²P]∼EIIB^Lev, which had been separated from EI, HPr, and EIIA^Lev after its phosphorylation by these proteins, strongly suggested that P∼LevE directly phosphorylates LevR at His-869 (520). The fact that His869Ala mutant LevR was not phosphorylated by the [³²P]∼EIIB^Lev preparation proved that it was almost free of [³²P]∼His-HPr. Inactivation of levD (EIIA^Lev) and levE (EIIB^Lev) as well as the replacement of His-869 of B. subtilis LevR (or His-776 of L. casei LevR) with an alanine cause constitutive expression from the lev promoter of both organisms. These results confirm that phosphorylation in PRD2 mediated by PEP, EI, HPr, EIIA^Lev, and EIIB^Lev exerts an inhibitory effect on the transcription activation function of LevR.

Mutation of the first conserved histidine (His-506) in PRD1 also slightly affects B. subtilis LevR activity. However, B. subtilis and L. casei mutant LevRs in which both identified phosphorylatable histidines had been replaced with alanine were not phosphorylated in vitro either by EI and HPr or by EI, HPr, EIIA^Lev, and EIIB^Lev (520, 534). These results suggest that in wild-type LevR, neither the two conserved histidyl residues in PRD1 nor the conserved cysteyl residue in the EIIB^Gat domain is phosphorylated.

The results described above establish that the transcription activator LevR in B. subtilis and L. casei is regulated by at least two PTS-catalyzed phosphorylation reactions exerting antagonistic effects on LevR activity. Control of LevR therefore appears to be a complex kinetic phenomenon similar to LicT regulation. Although the kinetics of the phosphoryl group transfer reactions are barely understood, a model can be proposed on the basis of the above-mentioned biochemical and genetic results. In in vitro experiments, B. subtilis P∼EIIB^Lev phosphorylates not only fructose (110) but also His-869 in PRD2 of LevR (520). According to genetic data, the latter reaction inhibits LevR activity, and it is therefore likely that the induction of the lev operon requires the dephosphorylation of His-869. In fact, the uptake of fructose via the Lev PTS will lead to the dephosphorylation of not only P∼EIIB^Lev but, as a consequence of the reversibility of the phosphoryl transfer, probably also P∼His(869)-LevR. Because the phosphorylation of HPr by EI is very fast and because the Lev-PTS is a low-capacity fructose transport system, one can assume that cells transporting fructose via the Lev-PTS probably contain relatively high concentrations of P∼His-HPr, which are probably sufficient for the phosphorylation of LevR at His-585 in the EIIA^Man-like domain. As a consequence, during Lev-PTS-catalyzed fructose uptake, a major fraction of LevR will probably be present in the fully active form, i.e., phosphorylated at His-585 and dephosphorylated at His-869. Dephosphorylation of His-585 will occur when a rapidly metabolizable carbon source, such as glucose, is present in the growth medium. The uptake of glucose will lower the P∼His-HPr level and, consequently, the activity of LevR, even when it is dephosphorylated at His-869. Indeed, LevR activity in levD ΔptsH or levE ΔptsH double mutants was reported to be about 15-fold lower than that in levD or levE single mutants (846). Poor phosphorylation of His-585 during growth on a rapidly metabolizable carbon source serves as a CcpA-independent CCR mechanism and accounts for the residual CCR, which was observed when a ccpA disruption was introduced into an inducer-independent mutant (levE or levR8). The residual CCR almost completely disappeared in a ccpA ptsH1 double mutant (526), which, due to the inability to form P-Ser-HPr, probably contains a larger amount of P∼His-HPr.

Because LevR contains a NifA/NtrC-like central domain, it is likely that LevR activates transcription by a mechanism similar to that operating in non-PRD-containing NifA/NtrC-type transcription activators. In the latter proteins, phosphorylation of the receiver domain by a sensor kinase stimulates the ATPase function located in the central domain and, consequently, the formation of open complexes by RNA polymerase (943). As mentioned above, the central domain of LevR also contains a nucleotide binding site. It is therefore tempting to assume that phosphorylation of the EIIA^Man-like domain by EI and HPr leads to increased LevR ATPase activity, while phosphorylation in PRD2 inhibits it. Nevertheless, how the inactivating/activating signals are transmitted from PRD2 and the EIIA^Man-like domain, respectively, to the NifA/NtrC-like central domain remains an open question. The structure of LevR or one of its homologs has so far not been solved.

For a long time, the only known homolog of LevR was CelR from G. stearothermophilus (445). CelR has the same regulatory domain organization as B. subtilis LevR (Fig. 7), to which it exhibits 47% sequence similarity. Histidyl residues equivalent to the two phosphorylation sites in B. subtilis LevR (His-585 and His-869) are also present in CelR. The celR gene precedes an operon encoding a cellobiose-specific PTS (lactose class) and a presumed 6-P-β-glucosidase (446). CelR was therefore assumed to control the expression of this operon by a mechanism similar to that used by LevR for the regulation of the B. subtilis lev operon.

The L. monocytogenes lmo1721 gene encodes a LevR-like regulator (called CsrA, but it is unrelated to E. coli CsrA), the inactivation of which prevents the repressive effect of cellobiose on the activity of PrfA, a transcription activator of numerous virulence genes in this organism, while the repressive effect of glucose on PrfA persisted in a csrA mutant (558a). The csrA gene precedes and probably regulates an operon containing a potential −12, −24 promoter and three genes, which code for an EIIA and an EIIB of a lactose/cellobiose class PTS and an outer surface protein. However, this operon is missing an EIIC, and it is therefore not clear whether the EIIA and EIIB are involved in cellobiose transport. Interestingly, L. monocytogenes contains an operon encoding a complete potential cellobiose-specific PTS (EIIA, EIIB, and EIIC, lmo2683 to lmo2685), but no regulator gene precedes or follows this operon. Nevertheless, this operon is preceded by a potential −12, −24 promoter identical to the one located in front of the csr operon [TGGCAC(N)5TTGCAT] (Deutscher, unpublished). It is possible that this operon is also regulated by CsrA and that csrA inactivation therefore prevents the expression of both potential cellobiose operons and, consequently, the synthesis of the cellobiose-specific PTS components and cellobiose transport. However, more specific effects of CsrA and/or the EIIA and EIIB encoded by the csr operon can presently not be excluded. It is noteworthy that cellobiose-specific inhibition of PrfA also disappeared in mutants affected in bvrAB, which code for a PRD-containing antiterminator and a β-glucoside-specific EIIBCA (81) (see the preceding section).

Homologs of B. subtilis LevR are present in many other low-G+C gram-positive organisms such as bacilli, clostridia, enterococci, lactobacilli, listeriae, Pediococcus pentosaceus, and Thermoanaerobacter tengcongensis, but they are also present in the actinobacterium S. thermophilum and in gram-negative bacteria such as S. enterica serovar Typhimurium, Salmonella enterica, K. pneumoniae, and Salmonella enteritidis, although they are much less frequent in the latter organisms. The highest number of LevR-like proteins (seven) was detected in C. difficile. The large number of LevR homologs in several bacteria suggests that these proteins are not used exclusively for the regulation of the expression of operons coding for fructose-, mannose-, or cellobiose-specific PTS. Indeed, the L. casei esu operon is regulated by a LevR-like protein and encodes a PTS that probably transports a sugar ester (976). Most of the approximately 100 LevR-like proteins detected have a size similar to that of B. subtilis LevR and exhibit between 40 and 60% sequence similarity to each other. They contain conserved equivalents of the two phosphorylatable histidyl residues, confirming the importance of these two amino acids in the regulation of the activity of LevR-like transcription activators. In contrast, an equivalent of His-506 of B. subtilis LevR, the first conserved histidine in PRD1, is absent from several LevR homologs, implying that this amino acid does not play an essential role in the regulation of LevR-like proteins. One E. faecalis and one C. difficile LevR homolog contain a complete second PRD, thus possessing an additional potentially phosphorylatable histidyl residue.

A protein containing two PRDs and an N-terminal domain resembling the DNA binding domain in transcription activators of the E. coli DeoR family was first detected in G. stearothermophilus (324). The corresponding gene is located in the mtlARFD operon, in which mtlA codes for a mannitol-specific EIICB, mtlR codes for the PRD-containing regulator of the operon, mtlF codes for an EIIA^Mtl, and mtlD codes for a mannitol-1-P dehydrogenase. Soon afterwards, another protein with a DeoR-like N-terminal DNA binding domain followed by two PRDs was found in B. subtilis (878). This protein, LicR, controls the expression of the licRBCAH operon, which encodes the transcription activator LicR; the EIIB, EIIC, and EIIA components of a β-glucoside-specific PTS (belonging to the lactose class); and a presumed 6-P-β-glucosidase, respectively. The genome sequence revealed that B. subtilis contains two other transcription activators possessing a DeoR-like N-terminal domain and PRDs: MtlR (YdaA) and ManR (YjdC). They were proposed to regulate the expression of a mannitol-specific and a mannose-specific PTS, respectively (844). Similar proteins in other bacteria have been described. They include MtlR (342) and SrlR (78) of S. mutans and MtlR of C. acetobutylicum (51). Extensive genetic and biochemical studies have been carried out only with MtlR of G. stearothermophilus and LicR from B. subtilis.

The transcription activators MtlR of G. stearothermophilus and LicR of B. subtilis possess an N-terminal DNA binding domain (Fig. 7), which exhibits significant similarity (around 50%) to the first approximately 180 amino acids of transcription activators of the DeoR family such as E. coli FucR (125), B. anthracis AcpA (414), and S. pyogenes VirR (537). In most DeoR-type transcription activators, a traditional helix-turn-helix motif is located at the beginning of the N-terminal domain. However, in a few PTS-controlled regulators, the helix-turn-helix motif is degenerated and difficult to detect (51). The suggested presence of a second helix-turn-helix motif in LicR (879) has so far not been confirmed. The DNA binding domain is followed by two PRDs, with PRD1 beginning at about amino acid 185 (Fig. 7). Each PRD usually contains two conserved histidyl residues. The PRDs are followed by an EIIB^Gat domain containing a conserved and potentially phosphorylatable cysteyl residue (Cys-423 in G. stearothermophilus MtlR and Cys-413 in B. subtilis LicR) (295). The last domain of DeoR-type PRD-containing transcription activators is an EIIA domain belonging to the mannitol/fructose class PTS. In EIIA^Mtl of S. carnosus, His-62 has been identified as a PEP-dependent phosphorylation site (703). An equivalent histidyl residue is present in the EIIA^Mtl/Fru-like domains of LicR of B. subtilis (His-559) (879) and MtlR of G. stearothermophilus (His-598) (323). In total, these regulators contain six potential PTS phosphorylation sites (Fig. 7).

Phosphorylation of G. stearothermophilus MtlR was observed in the presence of PEP, EI, and HPr (323). The P∼His-HPr-mediated phosphorylation was completely prevented when either one of the two conserved histidyl residues in PRD2 was replaced with an alanine. However, His348Ala, His405Ala, and His348Ala/His405Ala mutant MtlRs were phosphorylated when, in addition to EI and HPr, EIIA^Mtl and EIICB^Mtl were also present in the phosphorylation assay (323). The phosphorylation of MtlR in the presence of EIIA^Mtl and EIICB^Mtl was strongly diminished when the conserved His-598 in the C-terminal EIIA^Mtl/Fru-like domain was replaced with an alanine. The residual weak phosphorylation of His598Ala mutant MtlR was probably due to phosphorylation in PRD2 by the low amounts of P∼His-HPr present in the reaction mixture. In conclusion, MtlR becomes phosphorylated by EI and HPr at one or both conserved histidyl residues in PRD2 and in an EI-, HPr-, EIIA^Mtl-, and EIICB^Mtl-requiring reaction at His-598 in the EIIA^Mtl/Fru-like domain. Phosphorylation of MtlR therefore clearly differs from B. subtilis LevR phosphorylation, where EI and HPr phosphorylate the EIIA^Man-like domain and phosphorylation in the presence of EIIA^Lev and EIIB^Lev occurs in PRD2 (520).

Similar to LevR, MtlR binds to a DNA region located upstream from the promoter, as was demonstrated in footprint experiments with G. stearothermophilus MtlR and a DNA fragment extending about 125 bp upstream and downstream from the transcription start site of the mtl operon. Apparently, MtlR does not interact with a continuous DNA stretch, as within a sequence of about 50 bp, five small regions were protected against DNase I digestion in the footprint experiments (325). The phosphorylation state of MtlR had a strong effect on its DNA binding activity. MtlR phosphorylated by EI and HPr in PRD2, but dephosphorylated in the EIIA^Mtl/Fru-like domain, exhibited an approximately 100-fold-higher affinity for its target DNA than did unphosphorylated MtlR (323). The DNA binding affinity of neither the His348Ala nor the His405Ala mutant MtlRs was enhanced by the presence of P∼His-HPr (as mentioned above, mutation of either His prevents phosphorylation in PRD2). In contrast, phosphorylation in the presence of EI, HPr, EIIA^Mtl, and EIICB^Mtl at His-598 in the EIIA^Mtl/Fru-like domain and in PRD2 lowered the affinity of MtlR for its DNA target about 10-fold compared to the unphosphorylated regulator. Surprisingly, not only phosphorylation of His-598 but also its replacement with an alanine had a negative effect on DNA binding, which could not be overcome by P∼His-HPr-mediated phosphorylation (323). It is possible that the mutation causes structural changes leading to the inactivation of the transcription regulator. In conclusion, MtlR is controlled by at least two PTS-mediated phosphorylation events that exert antagonistic effects. Similar to LicT and LevR, the absence of EI- and HPr-mediated phosphorylation in PRD2 of MtlR serves as an alternate CCR mechanism. In contrast, phosphorylation in the EIIA^Mtl/Fru-like domain, which requires the additional presence of EIIA^Mtl and EIICB^Mtl, exerts a negative effect on MtlR activity. This phosphorylation is probably prevented when mannitol is present in the growth medium, and it is therefore likely that dephosphorylation at His-598 serves as an induction mechanism for the mtl operon. The role of PRD1 in MtlR regulation is not clear. Although the replacement of either one of the conserved histidyl residues in PRD1 diminished the stimulatory effect of P∼His-HPr on MtlR activity, phosphorylation of these histidines could not be detected. Similar to what was proposed for LevR, PRD1 in MtlR might be important only for the signal transfer from PRD2 to the DNA binding domain.

Expression of the mtl operon in S. carnosus (232) and B. subtilis seems to be regulated by a mechanism resembling that in G. stearothermophilus, as the mtl operon of both organisms contains a 50-bp-long upstream activating sequence similar to that recognized by MtlR of G. stearothermophilus (325). In addition, the mtl operon of S. aureus exhibits a gene order (mtlARFD) identical to those in G. stearothermophilus and S. mutans (341, 342). In contrast, ydaA, which encodes the B. subtilis MtlR homolog, is a monocistronic transcription unit located about 15 kb downstream from the mtl operon (437). YdaA was renamed MtlR (844), as it exhibits significant sequence identity (39%) to MtlR from G. stearothermophilus. MtlR of S. mutans exhibits only 28% sequence identity and 48% similarity to MtlR of G. stearothermophilus. Surprisingly, only a polar insertion into mtlR was reported to prevent mannitol utilization by S. mutans, whereas a nonpolar insertion had no effect (342). Therefore, whether MtlR of S. mutans carries out functions similar to those of MtlR of G. stearothermophilus is questionable. Finally, an MtlR homolog of C. acetobutylicum was reported to control the expression of the mtlARFD operon of this organism (51).

The DNA binding site for LicR, the second PRD-containing DeoR-type transcription activator in B. subtilis, was determined by carrying out a deletion analysis with the promoter region of the lic operon. A sequence with a dyad symmetry precedes the lic promoter, which appears to be the target site for LicR (879). Similar to MtlR, the activity of LicR is negatively controlled by its corresponding sugar-specific PTS proteins, EIIA^Cel (LicA) and EIIB^Cel (LicB). Inactivation of EIIA^Cel causes constitutive expression of a licB′-lacZ fusion. In contrast, deletion of the ptsHI genes prevents expression of the licB′-lacZ fusion, suggesting a positive role for the general PTS proteins in LicR regulation (878). The four conserved histidyl residues in the two PRDs of LicR were proposed to be the targets of positive regulation by P∼His-HPr, as the replacement of either one of them with a nonphosphorylatable amino acid results in inactive LicR (879). In contrast, replacement of His-559 in the EIIA-like domain of LicR with a glycine causes constitutive expression of the licB′-lacZ fusion. His-559 therefore appears to be the site of negative control of LicR activity. Based on the genetic results, it was proposed that P∼EIIB^Cel transfers its phosphoryl group to His-559 in the EIIA^Mtl/Fru-like domain of LicR (879). Although EIIB^Cel and EIIA^Mtl/Fru belong to the same superfamily of PTS proteins (glucose-fructose-lactose), their phosphorylation sites do not exhibit significant sequence similarity (231, 703). In addition, the structures of EIIAs from these two PTS classes are completely different (730), and “cross”-phosphorylation between the EIIAs and EIIBs from the different PTS classes has not been reported. It therefore remains to be experimentally confirmed that the EIIA^Mtl/Fru-like domain in LicR can indeed be recognized and phosphorylated by P∼EIIB^Cel. Alternatively, His-559 in LicR could be phosphorylated by P∼His-HPr but only in the presence of P∼EIIB^Cel. This mechanism would resemble the regulation of the SacY antiterminator by P∼His-HPr-mediated phosphorylation at His-99 in the presence of P∼SacX (883).

A third gene, yjdC, encoding a protein containing a DeoR-like DNA binding domain followed by two PRDs and EIIB^Gat- and EIIA^Mtl/Fru-like domains is present in B. subtilis (437). The sequence identities between YjdC and the other two related transcription activators in B. subtilis were 26% to LicR and 21% to MtlR. The yjdC gene is monocistronic and is located in front of the yjdDEF operon encoding an EIIBCA of a fructose/mannitol class PTS (yjdD), an enzyme exhibiting similarity to mannose-6-P isomerase (yjdE) and a protein of unknown function (yjdF). YjdC was called ManR (844), although the sugar specificity of the encoded PTS remains to be determined. Another DeoR-type transcription regulator, SrlR, containing PRDs and EIIB^Gat- and EIIA^Mtl/Fru-like domains has been described for S. mutans. SrlR controls the expression of the srlDRMAEB operon (78). Besides the putative transcription activator, the operon encodes a glucitol (sorbitol)-specific PTS (srlAEB) and a glucitol-6-P dehydrogenase (srlD). Surprisingly, the operon also contains a homolog of E. coli SrlM, a small protein that has been reported to function as an activator for the srlR-less E. coli srl operon (961).

Homologs of G. stearothermophilus MtlR and B. subtilis LicR are found in the genomes of numerous gram-positive bacteria, including other bacilli, clostridia, enterococci, lactobacilli, listeriae, streptococci, staphylococci, Exiguobacterium spp., Geobacillus kaustophilus, Leuconostoc mesenteroides, L. lactis, Moorella thermoacetica, O. iheyensis, T. tengcongensis, Thermoanaerobacter ethanolicus, etc. In total, more than 100 proteins were found to exhibit significant sequence similarity to G. stearothermophilus MtlR or B. subtilis LicR. In many of these proteins, including S. mutans SrlR, one or both conserved histidines in PRD1 are missing, whereas the phosphorylatable histidines in PRD2 and the EIIA^Mtl/Fru-like domain were always present, confirming the importance of the EIIA^Mtl/Fru domain and PRD2 for the regulation of MtlR/LicR-type transcription activators. The organisms with the most DeoR-type PRD-containing transcription activators were C. difficile (13) and L. innocua (10). A few LicR/MtlR homologs are also present in gram-negative organisms such as E. coli, P. carotovorum, S. enterica serovar Typhimurium, S. enterica, S. enteritidis, S. paratyphi, Vibrio fischeri, and Yersinia enterocolitica.

The results obtained with the three best-characterized PRD-containing transcription activators LevR, MtlR, and LicR make it clear that the molecular mechanisms regulating their activity can vary largely. In LicR of B. subtilis, all four conserved histidyl residues in the two PRDs seem to be phosphorylated by P∼His-HPr, and their phosphorylation stimulates LicR activity. In MtlR of G. stearothermophilus, only the two histidyl residues present in PRD2 seem to have an equivalent function. Finally, B. subtilis LevR is activated by P∼His-HPr-mediated phosphorylation of a histidyl residue present in the EIIA^Man-like domain. For each transcription activator, the absence of the stimulatory phosphorylation(s) during the uptake of a rapidly metabolizable PTS substrate initiates a CcpA-independent CCR mechanism. The three PRD-containing transcription activators are inhibited by P∼EIIB-requiring phosphorylation in their C-terminal domain. The presence of the appropriate inducer (PTS substrate) prevents this phosphorylation and leads to the induction of the corresponding operon. Inhibition of LicR of B. subtilis is mediated by phosphorylation at His-559 in the EIIA^Mtl/Fru-like domain. Although the detailed mechanism of this phosphorylation reaction is not known, it is probably mediated by P∼His-HPr in the presence of P∼EIIB^Cel. Similarly, MtlR of G. stearothermophilus is inactivated by P∼His-HPr- or P∼EIIB^Mtl-mediated phosphorylation at His-598 in the EIIA^Mtl/Fru-like domain. In contrast, inactivation of LevR apparently occurs by P∼EIIB^Lev-mediated phosphorylation of a conserved histidyl residue in the second, truncated PRD.

It is tempting to assume that the four regulatory domains present in PRD-containing transcription activators are organized in a row, similar to the PRDs in antiterminators (909). The inactivating signal elicited by phosphorylation in the C-terminal domain therefore has to be transmitted through the other three domains to affect DNA binding. Structure determination of a transcription activator will hopefully allow a better understanding of the signal transmission.

Lactose transport in S. thermophilus is catalyzed by LacS, an integral homodimeric membrane protein that exhibits similarities to the melibiose transporters of E. coli and other bacteria (673). Although LacS is an H⁺/lactose symporter (235), it contains a C-terminal EIIA^Glc-like domain, which becomes phosphorylated by PEP, EI, and HPr (672). In E. coli and S. enterica serovar Typhimurium, the melibiose transporter is inhibited by an allosteric interaction with dephospho-EIIA^Glc, whereas P∼EIIA^Glc exhibits no affinity for the melibiose permease (439, 621) (also see the section on EIIA^Glc-mediated inducer exclusion above). Regulation of LacS-mediated lactose transport in S. thermophilus was hypothesized to follow a similar mechanism, except that the transporter is already fused to its regulatory EIIA^Glc domain. EI/HPr-catalyzed phosphorylation of the EIIA^Glc domain (302) was thought to prevent the inhibition of LacS. However, deletion of the EIIA^Glc domain of LacS or mutation of its phosphorylatable histidyl residue (His-552) did not affect the H⁺/lactose symport activity of LacS. In fact, when the lactose transport rate of LacS reconstituted into liposomes was measured, the presence of PEP, EI, and HPr even slightly inhibited proton motive force-driven lactose uptake.

More detailed studies revealed that monomeric LacS of S. thermophilus (244) catalyzes lactose uptake via a lactose/galactose exchange reaction (915). Galactose produced by the β-galactosidase-catalyzed hydrolysis of intracellular lactose cannot be metabolized by S. thermophilus and therefore needs to be expelled. Galactose export is also catalyzed by LacS and is coupled to the import of lactose (counterflow). The lactose/galactose exchange reaction has a higher V_max than LacS-catalyzed lactose/H⁺ symport and should therefore represent the major entry route for lactose. The V_max of the counterflow reaction was two- to threefold higher when LacS was phosphorylated by PEP, EI, and HPr, and the K_m was twofold lower (301). Thus, PEP-dependent phosphorylation of LacS specifically stimulates lactose uptake via the lactose/galactose exchange reaction (Fig. 6).

A lactose transporter exhibiting 95% sequence identity to LacS from S. thermophilus is present in S. salivarius. In vitro studies revealed that LacS of S. thermophilus and S. salivarius becomes efficiently phosphorylated at His-552 not only by P∼His-HPr but also by doubly phosphorylated (P∼His,P-Ser)-HPr (134, 468), which is present in large amounts in oral streptococci grown on glucose or lactose (660, 893) and in lactose-grown S. thermophilus cells (134). However, in contrast to S. thermophilus, S. salivarius did not expel galactose during growth on lactose. In addition, when grown in medium containing lactose or lactose plus galactose, an S. salivarius ptsI mutant showed the same growth characteristics as the wild-type strain. The role of LacS phosphorylation in S. salivarius is presently not known.

Transporters of the galactoside-pentose-hexuronide family, to which LacS belongs, are abundant in bacteria. It appears that only a few of them (those that transport lactose or raffinose in gram-positive organisms) gained a regulatory domain by fusing EIIA^Glc to their C-terminal end, similar to the PTS-regulated NifA/NtrC- and DeoR-type transcription activators discussed in the previous section. In addition to S. thermophilus and S. salivarius, transporters exhibiting similarity to the E. coli melibiose permease and containing an EIIA^Glc domain fused to the C terminus have been described for a few other gram-positive organisms. They include the lactose transport protein LacS of L. delbrueckii subsp. bulgaricus (467) and Leuconostoc lactis (914). L. plantarum contains not only a LacS homolog but also a raffinose transporter (RafP) with an EIIA^Glc-like domain (815). Likewise, Pediococcus pentosaceus and Lactobacillus johnsonii possess both LacS and RafP homologs. S. thermophilus LacS-like proteins are also present in L. mesenteroides, Lactobacillus acidophilus, Lactobacillus reuteri, and Lactobacillus helveticus as well as in the proteobacterium Mannheimia succiniciproducens (MelB). The presence of an EIIA^Glc-like domain in these proteins indicates that P∼His-HPr-mediated phosphorylation might represent a general mechanism for the stimulation of their lactose/galactose or raffinose/galactose exchange activities.

Glycerol enters bacterial cells by facilitated diffusion, an energy-independent transport mechanism (321). The protein that catalyzes the equilibration of glycerol gradients across the bacterial cytoplasmic membrane is called the glycerol facilitator (GlpF) (481). GlpF, which has been studied extensively in E. coli (584, 849), is present in many other bacteria, archaea, and eukaryotes. It is a member of the major intrinsic protein family of transmembrane channels (39, 628). In addition to GlpF, the major intrinsic protein family includes other membrane-spanning bacterial proteins such as the water channel protein AqpZ from E. coli and several other bacteria (99) and PduF, a presumed propanediol diffusion facilitator of S. enterica serovar Typhimurium (119). A channel protein accepting glycerol and water as substrates was found in L. lactis (246). When the E. coli glpF gene was functionally expressed in Xenopus laevis oocytes, glycerol uptake by the oocytes exhibited the characteristics of diffusion by a pore-type mechanism (532). Because the glycerol facilitator catalyzes the reversible diffusion of glycerol across the cytoplasmic membrane, it is the subsequent metabolism of glycerol that allows a net (steady-state) flux of glycerol into the cell. The first metabolic step is catalyzed either by glycerol dehydrogenase, which converts glycerol into dihydroxyacetone, or by GlpK, which uses ATP to transform intracellular glycerol into glycerol-3-P (656). Glycerol-3-P is not a substrate for the glycerol facilitator and therefore remains entrapped in the cell. It functions as an inducer of the glp regulon by inactivating a repressor (GlpR) in E. coli (790) or by activating an antiterminator (GlpP) in B. subtilis (749). GlpP binds to the leader region in the mRNA of glpD but probably also of glpFK and glpTQ (274). The glpD gene encodes glycerol-3-P dehydrogenase (339), glpT encodes a glycerol-3-P transporter, and glpQ encodes a glycerophosphoryl diester phosphodiesterase.

Although glycerol is not transported by the PTS, mutants of gram-positive and gram-negative bacteria defective in one of the general PTS proteins EI or HPr had lost the ability to grow on glycerol as the sole carbon source (52, 265, 282, 600, 711, 741, 766). The inhibition of S. enterica serovar Typhimurium and E. coli glycerol kinase by unphosphorylated EIIA^Glc and the resulting dependence of glycerol metabolism on a functional PTS are described above (see “REGULATION OF CARBON METABOLISM IN GRAM-NEGATIVE ENTERIC BACTERIA”) (166, 676, 899). A mechanism completely different from that operating in E. coli and S. enterica serovar Typhimurium seems to control glycerol metabolism in most low-G+C gram-positive bacteria.

In contrast to the Enterobacteriaceae E. coli and S. enterica serovar Typhimurium, most gram-positive organisms do not contain free soluble EIIA^Glc (130, 350). In these bacteria, either EIIA^Glc is fused to EIICB^Glc, the membrane component of the glucose-specific PTS, or, when mannose and glucose are taken up by the same mannose class PTS, they possess no EIIA^Glc at all. Even in Mycoplasma capricolum, which is one of the few gram-positive organisms possessing free EIIA^Glc, the PTS protein did not inhibit GlpK activity (992). Moreover, while the inactivation of the EIIA^Glc-encoding E. coli or S. enterica serovar Typhimurium crr gene enabled ptsHI mutants to grow on glycerol as the sole carbon source, deletion or disruption of the 3′ part of the B. subtilis ptsG gene encoding the EIIA^Glc domain of EIICBA^Glc had no such effect (282). Nevertheless, suppressor mutants that restored growth of a B. subtilis ptsHI mutant on glycerol have been obtained (52). They were not affected in ptsG but, rather, were affected in glpK (937), and glycerol uptake in gram-positive bacteria was therefore assumed to be regulated by an EIIA^Glc-independent mechanism.

The observation that GlpK from E. faecalis is phosphorylated by PEP, EI, and HPr (174, 186) also suggested that glycerol metabolism in gram-positive bacteria is regulated by a mechanism different from that operating in gram-negative bacteria. PEP-dependent phosphorylation occurred at the N-3 position of a histidyl residue and stimulated GlpK activity about 10-fold (186). A similar PTS phosphorylation-mediated stimulation was observed for GlpK from Enterococcus casseliflavus (109) and B. subtilis (158). No specific protein phosphatase dephosphorylating P∼GlpK could be detected. Instead, dephosphorylation of purified P∼GlpK was observed when it was incubated together with an excess of HPr, indicating that P∼GlpK can transfer its phosphoryl group back to HPr (186). The site of PEP-dependent, EI- and HPr-catalyzed phosphorylation was determined to be His-232 in GlpK of E. casseliflavus (109) and His-230 in GlpK of B. subtilis (158). Interestingly, in one of the B. subtilis suppressor mutants that were able to grow on glycerol despite the absence of active EI and HPr, His-230 of GlpK was replaced with an arginine, and in another mutant, the neighboring Phe-232 was replaced with a serine (937). These mutations were thought to cause structural changes similar to those triggered by phosphorylation, and the ability of these glpK mutants to grow on glycerol was assumed to be due to PTS-independent elevated activity of the mutant GlpKs. Indeed, GlpK activity in the four B. subtilis ptsHI suppressor mutants was about 10-fold higher than the activity measured in a ptsHI strain producing wild-type GlpK (158). In addition, replacing the phosphorylatable His-232 in E. casseliflavus GlpK or His-230 in B. subtilis GlpK with an arginine led to an increase in activity similar to that caused by phosphorylation (109, 158). In contrast, E. casseliflavus (or B. subtilis) mutant GlpKs, in which His-232 (or His-230) had been replaced with an alanine or glutamine, exhibited low catalytic activity, which could not be stimulated by phosphorylation with PEP, EI, and HPr (109, 158). Compared to EIIAs, the usual phosphoryl acceptors of P∼His-HPr within the PTS phosphorylation cascade, phosphorylation of enterococcal GlpKs was about 100 times slower. Based on these in vitro results, it was concluded that PEP-dependent, EI- and HPr-catalyzed phosphorylation of GlpK serves to regulate glycerol metabolism in response to the presence or absence of a PTS substrate. In the absence of a PTS sugar, GlpK was assumed to be present primarily in the active, phosphorylated form, allowing the rapid uptake and metabolism of glycerol. However, when bacteria grow in a medium containing a PTS substrate, the phosphoryl transfer flux through the PTS will be high, and the concentration of the common phosphoryl donor P∼His-HPr will be low. As a result, GlpK will be present mainly in the less active dephospho form (Fig. 6).

E. faecalis GlpK and P∼GlpK migrate to slightly different positions on sodium dodecyl sulfate-polyacrylamide gels (175). By carrying out Western blot analyses with crude extracts from E. faecalis cells grown in the presence of various carbon sources, the influence of the growth conditions on the GlpK/P∼GlpK ratio could be determined. Growth in glycerol-containing medium induced the synthesis of GlpK, which was present primarily in the phosphorylated form under these conditions. If the growth medium contained glycerol plus glucose or mannitol, the synthesis of GlpK was strongly repressed, with the remaining GlpK being present mainly in the dephospho form. When cells were grown in glycerol-containing medium and glucose was added to one half of the culture 20 min before the cells were harvested, the amount of GlpK was similar to that in cells grown in the presence of glycerol only. However, while bacteria that continued to grow only on glycerol contained primarily P∼GlpK, the presence of glucose in the other half of the culture caused nearly complete dephosphorylation of P∼GlpK. These results support the above-described concept that the dephosphorylation of GlpK in response to the presence of a rapidly metabolizable PTS sugar inhibits glycerol uptake and metabolism. This regulatory mechanism can be considered a second form of “PTS-mediated inducer exclusion,” as the presence of a PTS substrate prevents the GlpK-catalyzed formation of the inducer glycerol-3-P.

The P∼GlpK dephosphorylation-based inducer exclusion mechanism represents a CcpA-independent CCR mechanism and has been studied in detail in B. subtilis, where expression of the glpFK operon is also submitted to P-Ser-HPr:CcpA-mediated CCR (158). In fact, in ptsH1, hprK (both cannot form P-Ser-HPr), and ccpA mutants, expression of glpK or a glpF′-lacZ fusion was still strongly repressed by glucose (158, 184), confirming that a second, P-Ser-HPr:CcpA-independent CCR mechanism is operative for the glp operon. Interestingly, the repressive effect of glucose on glpF′-lacZ expression was absent in a ccpA mutant producing His230Arg mutant GlpK (158), which is highly active even without being phosphorylated by PEP, EI, and HPr. These results provide additional evidence that the CcpA-independent CCR mechanism for the B. subtilis glpFK operon is based on poor phosphorylation of His-230 in GlpK. As discussed above, the prevalence of unphosphorylated, barely active GlpK during the rapid uptake of a PTS sugar is expected to lead to low concentrations of the inducer glycerol-3-P (Fig. 6). Growth of a ccpA mutant in the presence of glycerol-3-P, which is taken up by GlpT, should therefore also prevent CCR of glpF′-lacZ expression, and this has indeed been observed (158). In B. subtilis, the inducer glycerol-3-P activates the antiterminator GlpP, which prevents the formation of the ρ-independent terminator t_glpFK located in the glpFK leader region and consequently leads to the induction of the glp operon. Deletion of t_glpFK from the glpF′-lacZ fusion in a wild-type background therefore caused constitutive lacZ expression, which, however, was still strongly repressed by glucose (CcpA-mediated CCR). By contrast, when t_glpFK was deleted in a ccpA or hprK mutant, the repressive effect of glucose on glpF′-lacZ expression completely disappeared (158).

The crystal structure of GlpK from E. casseliflavus (977) revealed that it forms dimers in which the phosphorylatable His-232 of each monomer is located about 25 Å from the active center on a loop that is part of the dimer interface. As a consequence, the two phosphorylatable histidyl residues of a GlpK dimer are located close to each other (about 5 Å apart). Because more than 80% of GlpK is present as P∼GlpK in cells grown in glycerol-containing medium (175), doubly phosphorylated GlpK dimers probably represent the activated form. Electrostatic repulsion due to the two neighboring negatively charged phosphoryl groups is likely to drive the structural changes leading to elevated activity of P∼GlpK. Owing to the instability of the P∼His bond, the structure of P∼GlpK could not be determined. Nevertheless, the E. casseliflavus His232Arg and Phe234Ser mutant GlpKs exhibit elevated activity even when they are not phosphorylated (109) and are therefore expected to possess a structure similar to that of P∼GlpK. It will therefore be interesting to determine the structure of one of the mutant proteins, work that is presently going on in the laboratories of J. Yeh, P. Briozzo, and J. Deutscher.

The overall structure of GlpK from E. casseliflavus is very similar to that of GlpK from E. coli. The loop containing the phosphorylatable histidine in E. casseliflavus GlpK protrudes less in E. coli GlpK. In addition, the corresponding loop in E. coli GlpK does not contain a histidine. While E. coli and E. casseliflavus GlpKs exhibit 60% overall sequence identity, the sequences corresponding to this loop are completely different in GlpKs from gram-positive and gram-negative bacteria. Interestingly, FBP, an allosteric inhibitor of GlpK from gram-negative bacteria, binds to this loop in E. coli GlpK (622), thereby connecting two GlpK dimers and leading to the formation of inactive GlpK tetramers. It therefore seems that GlpKs from gram-positive and gram-negative bacteria use the same regulatory loop but that the mechanisms controlling GlpK activity via this loop are completely different.

GlpKs from most other gram-positive bacteria with low G+C content, such as E. faecalis, S. pyogenes (109), L. lactis (67), E. sibiricum, L. monocytogenes, L. sakei, O. iheyensis, S. pneumoniae, S. aureus, etc., also contain a histidyl residue equivalent to His-232 of GlpK from E. casseliflavus, and the surrounding sequence, including the three aromatic amino acids, is well conserved. Interestingly, L. plantarum contains two GlpKs that exhibit 76% sequence identity. However, only GlpK1 possesses a regulatory loop with a phosphorylatable His (in boldface type) (T-K-D-Y-H-F-F-G-S), which is replaced with a Leu (in boldface type) in GlpK2 (T-K-D-Y-L-L-Y-G-S), implying that only GlpK1 is regulated by the PTS (Deutscher, unpublished). The gram-positive bacteria M. pneumoniae, Mycoplasma genitalium, and Carboxydothermus hydrogenoformans possess a GlpK that exhibits about 45% sequence identity to GlpK from E. casseliflavus and contains a histidine equivalent to His-232, but the surrounding amino acids are poorly conserved. Therefore, whether these three enzymes are phosphorylated and regulated by P∼His-HPr is questionable. Other gram-positive bacteria with low G+C content, such as clostridia, Desulfitobacterium hafniense, and T. tengcongensis, contain a GlpK exhibiting between 50% and 65% sequence identity to GlpK from E. casseliflavus, but they lack an equivalent of His-232, and the sequence corresponding to the regulatory loop is completely different. Surprisingly, the regulatory loop sequence of C. acetobutylicum and Clostridium perfringens GlpK is almost identical to that of E. coli GlpK, suggesting that these enzymes are regulated by FBP-stimulated tetramer formation and not by P∼His-HPr-mediated phosphorylation. In contrast, the site of interaction with EIIA^Glc (Pro-472 to Tyr-481 in E. coli GlpK) (355) is not conserved in any known GlpK of gram-positive bacteria. These local sequence differences reflect the different modes of regulation of GlpK activity in gram-positive and gram-negative bacteria. Interestingly, amino acids Pro-472 to Tyr-481 are well conserved in GlpKs of only a few gram-negative bacteria including, in addition to E. coli and salmonellae, P. carotovorum, Photorhabdus luminescens, shigellae, and yersiniae, implying that the EIIA^Glc-mediated regulation of GlpK is not very widespread.

At first glance, the mechanisms regulating GlpK activity in gram-positive bacteria (stimulation by P∼His-HPr-dependent phosphorylation of GlpK) and certain gram-negative bacteria (inhibition by EIIA^Glc, which is prevented by P∼His-HPr-dependent phosphorylation at His-90 of EIIA^Glc) seem to be completely different. Nevertheless, the two distinct mechanisms of GlpK regulation, which apparently developed in gram-positive and gram-negative organisms after their separation during evolution, serve identical physiological functions; i.e., they slow glycerol uptake and metabolism when a rapidly metabolizable carbohydrate is transported via the PTS.

The thermostable GlpKs from Thermus flavus (347, 348) and Thermus aquaticus (346), which belong to the Thermus/Deinococcus group, exhibit more than 80% sequence identity to GlpK from B. subtilis. Other functionally related proteins from T. flavus or T. aquaticus and B. subtilis show a much lower degree of sequence identity (30 to 55%), implying that the GlpK-encoding gene present in T. flavus and T. aquaticus was gained by horizontal gene transfer from a gram-positive organism. Their GlpKs contain an equivalent of the phosphorylatable His-232, and the surrounding sequence is well conserved. It was therefore not surprising that EI and HPr from B. subtilis could phosphorylate T. flavus GlpK (157). In addition, T. flavus crude extracts were found to contain EI and HPr activity and were capable of phosphorylating T. flavus GlpK (157). However, phosphorylation of T. flavus GlpK did not increase its activity. A similar observation was made with G. stearothermophilus GlpK (714), and it was therefore hypothesized that the modification of GlpK from these two thermotolerant organisms affects their thermostability (157).

Glycerol kinase is a member of a large family of carbohydrate kinases including xylulose kinase, gluconate kinase, l-fuculose kinase, and many others (954). A data bank search revealed that none of the presently known eubacterial members of this family contains a histidyl residue equivalent to His-232 of GlpK from E. casseliflavus. Nevertheless, some bacteria such as L. rhamnosus (17) and L. casei BL23 (570) possess a glpK-like gene in addition to glpK in the glpFK operon (up to 80% identity at the protein level), which was called gykA. GykA of both organisms contains a P∼His-HPr phosphorylation site in position 231 (His231, in boldface type; -Y-H-F-F-), which is identical to that in several other GlpKs. In L. casei, gykA is monocistronic, and its function is unknown (570). Because the L. rhamnosus gykA gene was not induced during growth on glycerol, it is possible that GykA phosphorylates a substrate other than glycerol. Nevertheless, the PTS-mediated activation mechanism based on P∼His-HPr-dependent phosphorylation at a regulatory loop in the target enzyme seems to have almost exclusively evolved for GlpK of low-G+C gram-positive bacteria and certain members of the Thermus/Deinococcus group.

In E. coli, the regulation of nitrogen metabolism appears to be coupled to specific “classical” PTS components as well as to the PTS^Ntr. The key enzyme and key regulators for the utilization of nitrogen by E. coli are located in the glnALG operon, which encodes glutamine synthethase and the two-component system NtrB and NtrC, respectively (607, 704). Under conditions of nitrogen deprivation, P∼NtrC blocks transcription from glnAp₁ and induces transcription from the σ⁵⁴-dependent glnAp₂ promoter (607, 704). Although the weak σ⁷⁰-dependent glnAp₁ promoter requires activation by Crp/cAMP, glnA expression was found to be much higher during growth on glucose or glucose-6-phosphate than during growth on fructose or glycerol. It was assumed that Crp/cAMP stimulates glnALG expression from glnAp₁ but represses expression from glnAp₂ and that the stimulatory effect is relatively stronger than the effect of repression (875). In line with this assumption, it was found that the addition of cAMP to glucose-grown cells lowered GlnA activity by about sevenfold (506). However, further analysis showed that the effect of glucose is probably not mediated via Crp/cAMP but that it is mediated indirectly via the uridylylation state of GlnB. Furthermore, under nitrogen-limiting conditions, growth of a ptsN mutant on glucose and glutamine had a small but significant negative effect on expression from glnAp₂ (680). The addition of cAMP under these conditions had no effect. E. coli ptsN mutants grow poorly on alanine, and this effect was accentuated by the presence of several sugars or TCA cycle intermediates (680). Repression is relieved by supplying the cells with ammonium salts or by expressing a ptsN gene in trans. Disruption of the gene preceding ptsN (probably also leading to the inactivation of ptsN) causes a slight reduction in transcription from glnAp₂ and pnifL when K. pneumoniae cells are grown on glucose (plus formate) under nitrogen limitation (serine) (553). In contrast, when serine is replaced with ammonium as the nitrogen source, expression from the glnAp₂, pnifL, and pnifH promoters is significantly higher than that in the wild type (553). A similarly elevated expression from glnAp₂ can be observed in a genuine ptsN mutant, whereas the inactivation of npr leads to a decrease in transcription from glnAp₂, pnifL, and pnifH (554). Overexpression of K. pneumoniae ptsN in an E. coli rpoN⁺ strain inhibits growth on glucose minimal medium with either glutamine or ammonium as the nitrogen source but had no effect on an rpoN mutant (49). In R. etli, the inactivation of ptsN slightly reduces growth on alanine or serine in the presence of mannitol or glucose, whereas growth is inhibited when the TCA cycle intermediates succinate and malate are present (555). In addition, a reduction in melanin production was observed. The activity of the pnifH promoter decreases, while the level of NifA, the activator controlling the expression of nifH, is not affected by the mutation. Mutation of ptsA, the gene that precedes the frw operon and codes for EI^Frw in E. coli (345, 720), has effects similar to those of the inactivation of ptsN. A functional link between EI^Frw and EIIA^Ntr was therefore suggested (555).

A connection between EIIA^Ntr and the regulation of branched-chain amino acid metabolism in E. coli was reported (456). The genes encoding the three heterodimeric E. coli acetohydroxy acid synthases, which catalyze the first step in leucine, isoleucine, and valine biosynthesis, are located in three different operons (891). One of the gene pairs (ilvGM) is normally not expressed due to a frameshift mutation. Among the other two gene pairs (ilvBN and ilvIH), the former is responsible for the majority of the enzyme activity (276, 277). The proteins encoded by ilvBN and ilvIH are sensitive to feedback inhibition by valine. As a consequence, wild-type cells exposed to valine as the sole amino acid or nitrogen source cannot make isoleucine and therefore cannot grow (452). Exposure to leucine-containing dipeptides also leads to an impairment of growth and a considerable increase in the intracellular leucine concentration (862). Besides abolishing the growth on leucine-containing dipeptides altogether, inactivation of ptsN completely prevents E. coli cells from growing on leucine and serine and reduces growth on isoleucine (456). Introduction of a plasmid encoding EIIA^Ntr partly restores growth on these compounds, whereas transformants with a plasmid encoding His73Ala mutant EIIA^Ntr (cannot be phosphorylated) grow even better than wild-type cells. Further experiments showed that unphosphorylated EIIA^Ntr mediates derepression of the ilvBN operon and thereby enables the cells to synthesize the other branched-chain amino acids when grown on leucine, isoleucine, or serine.

In P. putida, EIIA^Ntr affects the regulation of gene expression from the TOL plasmid. The TOL plasmid harbors the genes necessary for the metabolism of toluene and xylenes and contains four transcription units: a so-called upper pathway and a meta pathway operon flanked by the promoters Pu and P_M, respectively, and the regulator-encoding genes xylS and xylR connected to the promoters P_S and P_R, respectively (28, 388). Transcription from Pu and P_S1 is σ⁵⁴ dependent and is subject to CCR: it is low when cells grow exponentially on rich medium and is also down-regulated by glucose or gluconate but not by fructose (22, 101, 103). Repression of Pu (101, 103, 104) and P_S1 (199, 281) transcription by rich medium or glucose is partially relieved (recovery of 50% of the activity) by the inactivation of ptsN or the catabolite repression control protein-encoding gene, crc (22). Relief of repression is achieved only in the presence of the inducer o-xylene (22), which increases the affinity of the activator XylR for Pu and P_S1 (57, 514). Moreover, expression of xylR is repressed by rich medium, and this repression is absent in a ptsN mutant, whereas a crc mutation has no effect (22, 514). Thus, it seems that EIIA^Ntr interferes with the activity of the transcription activator XylR, whereas repression via Crc should follow another mechanism (22). The involvement of Crc in the regulation of several sugar catabolic pathways (747) suggests a connection between the various metabolism-controlling regulatory networks (22). This complies with an observed down-regulation of Pu expression mediated via products of the Entner-Doudoroff pathway, where 6-phosphogluconate and/or 2-dehydro-3-deoxyphosphogluconate was identified as the intermediate that exerts repression (916).

Studies on active-site mutants suggest that the phosphorylated form of EIIA^Ntr mediates repression of Pu and P_S1. It was found that a mutant producing His68Asp EIIA^Ntr is permanently repressed, the aspartyl residue is thought to mimic a phosphorylated histidine, whereas a mutant producing His68Ala EIIA^Ntr is not (103). Surprisingly, inactivation of npr does not relieve inhibition by glucose but leads to repression in the absence of glucose, a phenotype resembling that of the His68Asp ptsN mutant (104). A ptsN npr double mutant has a ptsN-like phenotype, and a mutant in which the conserved histidine of NPr has been replaced with an alanine exhibits the same phenotype as the npr-disrupted mutant. Although a P. putida ptsP gene has been identified, its inactivation had no effect. Taking all of these findings into account, it appears that EIIA^Ntr of P. putida is partly phosphorylated during growth on glucose, contrary to what is seen for EIIA^Glc in E. coli. The presumed partial phosphorylation of the PTS components in the presence of glucose might be explained by the fact that glucose is not a PTS sugar in P. putida (103). Because the uptake of glucose by P. putida does not use but yields PEP, it is possible that the PEP-to-pyruvate ratio is raised by the glucose catabolism, which in turn might increase the phosphorylation state of PTS proteins, including EIIA^Ntr. At the same time, the lack of repression by fructose can be explained by the fact that in P. putida, fructose uptake occurs via a PTS and thus lowers the concentration of P∼EIIA^Ntr.

Inactivation of ptsN (and thereby also npr) in Pseudomonas aeruginosa impairs growth on minimal medium A (no carbon source mentioned) (377). Growth can be restored by the addition of glutamine, but not arginine, histidine, proline, or glutamate, and by trans-complementation (with only npr). There was no effect of ptsN overexpression or inactivation on the expression of the σ⁵⁴-regulated pilin or flagellin genes. Likewise, in Caulobacter crescentus, expression from several σ⁵⁴-dependent flagellar genes was unaffected by a ptsN mutation, although expression from the fljK promoter was slightly lower (371). Several bacteria, including numerous proteobacteria and the spirochetes Treponema pallidum, T. denticola, and Leptospira interrogans, the green photosynthetic bacterium Chlorobium tepidum, and the δ-proteobacterium Geobacter metallireducens, encode an EI, an HPr, an HPrK/P, and either one or two EIIA^Ntr homologs within their genome but lack any known EIIB and EIIC (64, 345, 841). In T. pallidum, the EI-encoding gene seems to be an inactive pseudogene. Furthermore, it was shown in vitro that the T. denticola PTS^Ntr proteins are functional in phosphotransfer and that HPr is phosphorylated by HPrK/P requiring a much lower concentration of fructose-1,6-bisphosphate than the enzyme from B. subtilis for full activity (278).

EI^Ntr, which can easily be detected in numerous proteobacteria due to its typical N-terminal extension, exerts diverse functions. Azotobacter vinelandii does not transport carbohydrates via the PTS (44, 740), and a ptsP mutation hardly affects growth on carbon sources like fructose, mannitol, sucrose, acetate, pyruvate, or succinate (793). However, growth of a ptsP mutant on glucose or glycerol is drastically slowed, although glucose uptake and phosphorylation are not affected. The impeded growth was traced to the inactivation of the enzyme nitrogenase, and the suggestion was made that respiratory protection of nitrogenase under carbon-limiting conditions is absent in the ptsP mutant (793). At the same time, it was observed that the inactivation of the ptsP gene in A. vinelandii reduces the capacity of the organism to accumulate poly-β-hydroxybutyrate (793), a carbohydrate storage molecule made from acetyl-CoA building blocks (397). The signaling pathway might also include NPr and EIIA^Ntr, as genome sequencing has revealed the presence of both ptsN and npr in the A. vinelandii rpoN operon (551). Similarly, “leaky” Ralstonia eutropha phbI and phbH mutants show diminished accumulation of poly-β-hydroxybutyrate (687). A close relationship between phbI and phbH and the ptsP and npr genes of E. coli was established (680), although the PhbI protein of R. eutropha lacks the NifA-like extension typical of other EI^Ntrs.

Mutants of P. aeruginosa (854) or Legionella pneumophila (333) affected in ptsP do not show an obvious phenotype when grown in rich medium. However, the ptsP mutants have almost completely lost their virulence due to strongly reduced growth after infection of nematodes, plants, mice, guinea pigs, or human epithelial cells. trans-Complementation with ptsP restores pathogenicity, which proves that the loss of virulence is due to the loss of functional EI^Ntr. L. pneumophila stores poly-β-hydroxybutyrate in its cytoplasm as a nutrition source for long-term starvation survival (370). ptsP mutants showed normal accumulation of the carbon storage molecule when grown in rich medium, but unfortunately, poly-β-hydroxybutyrate levels were not monitored after infection of host cells (333).

In B. japonicum, ptsP is located next to lysC (encoding an aspartokinase), but the two genes do not form an operon (412). Nevertheless, inactivation of lysC or ptsP prevented the utilization of Pro-Gly-Gly and other oligopeptides. In vitro “pull-down” experiments revealed that EI^Ntr interacts with the aspartokinase. Purified EI^Ntr is phosphorylated by ATP in the presence of cell extracts, and the phosphorylation is prevented when aspartokinase is added (412).

The discovery of paralogous PTS proteins associated with the transcriptional regulation of genes involved in nitrogen assimilation has provided candidates for a regulatory connection between nitrogen and carbon metabolism. In fact, a potential PTS^Ntr phosphorylation cascade is present in many bacteria, but a coherent concept for a signaling pathway is lacking. Growth of ptsN null mutants on poor nitrogen sources is impaired, especially when a TCA cycle intermediate is used as the sole carbon source, whereas growth on rich nitrogen media (glutamine and NH₄⁺) is not affected. The PTS^Ntr therefore seems to enable bacteria to grow (faster) on poor nitrogen sources and TCA cycle intermediates. A similar role in adaptation to changes in growth conditions can be attributed to EI^Ntr of P. aeruginosa and L. pneumophila, considering that ptsP mutants of these pathogens grow normally in rich medium but grow very slowly after infection of a host cell. The effect of ptsN, npr, and ptsP mutations on gene expression is less clear. A connection to σ⁵⁴-directed transcription is conceivable, because EIIA^Ntr and NPr are encoded within the rpoN operon and EI^Ntr has a putative regulatory domain present in other σ⁵⁴-interacting proteins, but no data showing exactly how σ⁵⁴-regulated genes are controlled by the PTS^Ntr are available. The amount of σ⁵⁴ present in the cell does not seem to be controlled by PTS^Ntr, as it is always low and is affected neither by growth on glucose nor by ptsN or npr mutations (388). Many effects of the ptsN, npr, or ptsP mutations, such as those on melanin production, poly-β-hydroxybutyrate accumulation, Pu promoter expression, or era activity, seem to be unrelated to nitrogen metabolism. Two-dimensional gel electrophoresis with protein extracts prepared from P. putida wild-type or rpoN and ptsN mutant strains indicated that the role of EIIA^Ntr in σ⁵⁴-mediated transcription can be only minor. On the other hand, in a P. putida ptsN mutant, the level of about 9% of the proteins is altered (102).

Also, whether the PTS^Ntr indeed forms a functional phosphorylation cascade remains uncertain. Some organisms contain only part of the system. In R. etli, the rpoN operon contains ptsN but seems to lack npr, while ptsA (EI^Frw) was identified based on sequence similarity (555). Inactivation of ptsA has effects similar to those of the inactivation of ptsN, suggesting a connection between EI^Frw and EIIA^Ntr. Moreover, genome analysis revealed that several proteobacteria, chlamydiae, and spirochetes lack any known EIIC permease but do contain EI, HPr, or both in combination with one or more components of the putative PTS^Ntr (41, 64). Although the PTS^Ntr is thought to require the phosphoryl group transfer P∼EI^Ntr→P∼NPr→P∼EIIA^Ntr in order to carry out its regulatory functions, the mixed appearance of the components in several bacteria suggests functional cross talk between the PTS^Ntr and “regular” PTS components. In fact, EIIA^Ntr of the spirochete T. denticola (278) and EIIA^Ntr and NPr of E. coli (680) can be phosphorylated in vitro by the regular PTS proteins. Interestingly, Powell et al. (680) found that in vitro NPr almost completely dephosphorylates P∼EIIA^Ntr, implying that phosphoryl group transfer occurs in the opposite direction. In that case, EIIA^Ntr, which is phosphorylated via EI and HPr, could communicate the phosphorylation state of the PTS proteins to NPr and EI^Ntr. Nevertheless, as NPr was present in about an eightfold excess of both EI and EIIA^Ntr in the phosphorylation assays, one might expect the dephosphorylation of P∼EI and P∼EIIA^Ntr independent of which phosphoryl transfer direction is favored. Moreover, the finding that unphosphorylated EIIA^Ntr mediates derepression of the ilvBN operon in E. coli and that the inactivation of ptsP or npr renders the derepression constitutive suggests a “normal” direction of the group transfer (456).

A comparison of the crystal structures of E. coli EIIA^Ntr and the EIIA^Mtl domain shows that the fold of both proteins as well as the structure around the phosphorylation site, His-73 in EIIA^Ntr and His-65 in EIIA^Mtl (703), are nearly identical (73, 905). However, the second conserved histidine, His-120 in EIIA^Ntr and His-111 in EIIA^Mtl, respectively, is differently oriented. His-111 of EIIA^Mtl is part of the active site, whereas His-120 of EIIA^Ntr is located in the vicinity but points away from the active site. Similar observations were made with the crystal structure of N. meningitidis EIIA^Ntr (724). Interestingly, EIIA^Ntr of H. influenzae lacks equivalents of the two histidines of the phosphorylation site altogether (717). These findings are consistent with a regulatory role of EIIA^Ntr through binding rather than through phosphoryl group transfer. Perhaps phosphorylation of the PTS^Ntr proteins serves only to alter binding affinities. On the other hand, the solution structure of the NPr:EIIA^Ntr complex suggests that the protein interface resembles that in the EIIA^Glc:HPr complex (927).