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The PTS consists of the general components, enzyme I (EI) and histidine protein (HPr), and the sugar-specific components (EIIA and EIIB) that transfer the phosphoryl group of phosphoenolpyruvate (PEP) ultimately onto the incoming sugar, which enters the cytoplasma through the transmembrane transport channel EIIC (chapter 75). In this scheme, the common proteins EI and HPr are the soluble cytoplasmic enzymes carrying the phosphoryl group at their conserved His residues and transferring them onto EIIA, again at a conserved His residue. Several transporters share their soluble EIIA, most often the glucose EIIA (EIIAGlc), others have their own. In some cases, the respective EIIAs are covalently linked to their cognate membrane-bound transporters. The transporters minimally consist of the EIIC domain, the transmembrane component, and the usually covalently linked EIIB domain exposed to the cytoplasm. The phosphoryl group of the EIIB domain, positioned last on a conserved Cys residue (in rare instances, a His residue), is finally transferred onto the incoming sugar.
The PTS is not only a means of transporting sugar under simultaneous phosphorylation, it also connects the transport of PTS sugars, notably glucose, to the regulation of gene expression, an effect called catabolite repression. In this scheme, the state of phosphorylation of EIIAGlc determines the activity of adenylate cyclase to produce cAMP, which, in turn, binds to the catabolite activator protein (CAP or CRP) (382, 463, 464, 536). Binding of the cAMP/CAP complex to the promoter of catabolite-sensitive genes (most of the genes encoding sugar transporters and sugar-degradative enzymes) is the prerequisite for their transcription. The nonphosphorylated EIIAGlc (predominantly present under conditions of glucose transport) also controls (i.e., inhibits) the activity of several non-PTS sugar transport systems, an effect called inducer exclusion (219, 394, 461, 462, 464).
Of the eight existing aldo-hexoses (allose, altrose, glucose, mannose, gulose, idose, galactose, and talose) and the four keto-hexoses (psicose, fructose, sorbose, and tagatose) only six are known to be carbon sources of E. coli and Salmonella. The ubiquitous hexoses D-glucose, D-fructose, and D-mannose and the rare sugar L-sorbose are transported via PTS and are described in this chapter. D-Allose is transported by an ABC system (see below) and L-galactose is a fortuitous substrate of the fucose PMF system (see below). D-Galactose can be transported either by an ABC system or by a PMF system (see below).
Glucose uptake and phosphorylation by PtsG initiate the "glucose effect," the curbing of the utilization of sugars, which are additionally present in the growth medium. Yet, under conditions where glucose becomes limiting the glucose effect disappears even at glucose concentrations that are considerably above the Km of PtsG for glucose. Under these conditions glucose will enter the cell without phosphorylation, preferentially via the binding protein-dependent methylgalactoside (Mgl) ABC transporter (107). This also dramatically affects the expression of malto(glyco)porin (108) as well as OmpC/F porins (307) (chapter 5). However, recently, Ferenci and coworkers have found that ptsG expression was highest with low (i.e., micromolar) external glucose levels in glucose-limited chemostats. They concluded that PtsG is primed to contribute to glucose scavenging under starvation-response conditions (498).
Of the six existing hexitols (allitol, altritol, glucitol, mannitol, galactitol, and iditol), only three, mannitol, glucitol, and galactitol, occur naturally. All these hexitols are transported by the PTS producing their 6-phospho derivatives.
Galactitol.
GlcN also can be transported and phosphorylated by the latter PTS to yield glucosamine 6-phosphate (256). Normally, GlcN is not a substrate for the general glucose PTS (EIIBCGlc and EIIAGlc, encoded by ptsG and crr, respectively; see above). However, a mutation in the EIIC membrane domain allows a strain defective in the hexose/mannose PTS to grow on GlcN (356, 386). The mutation (G176D) not only affects sugar specificity but also enhances expression of the ptsG, ptsHIcrr, and manXYZ operons (386). Since the mannose and the glucose PTS both act on glucose, it is not surprising that this sugar powerfully inhibits the uptake of GlcN (256, 295, 599, 600). The GlcNAc and GlcN pathways merge when GlcNAc 6-phosphate is converted by a deacetylase, encoded by nagA, to GlcN 6-phosphate (391, 598, 601), which, in turn, is converted by the GlcNAc-6-phosphate deaminase/isomerase, encoded by nagB, to fructose 6-phosphate (89, 90, 598, 601, 609, 610, 611). NagB acts on sugar phosphate and ammonium in a reversible way, following a rapid-equilibrium random mechanism (6, 70, 363). In vitro, it was found that NagB was stimulated by GlcNAc 6-phosphate, the allosteric activator (70, 89, 90, 330). Like most allosteric enzymes, GlcN-6-phosphate deaminase has two structural states, one displaying high substrate affinity (R state), and the other with low affinity for GlcN 6-phosphate, the T state. The crystallographic structures of the R and T forms of NagB have been solved (233, 363) and allowed a description of the reaction mechanism and allosteric transition of the enzyme (e.g., see references 290, 334, 335, and 456). The transition of the T to R state, which activates the enzyme, can be driven by substrate binding producing positive cooperativity (homotropic activation) and also through GlcNAc 6-phosphate binding to the allosteric site (heterotropic activation). The T state, which is characteristic of the ligand-free enzyme, does not have functional allosteric (GlcNAc 6-phosphate binding) sites, which are only formed in the intersubunits of the NagB hexamer as a consequence of the transition to the R state. There has also been a considerable amount of protein and enzymology work done on site-directed mutants of NagB to dissect the reaction mechanism (e.g., see references 68 and 86 and references therein).
Trehalose.
However, trehalose is also effectively utilized as carbon source at high-medium osmolarity. This is due to the presence of TreA, a periplasmic trehalase encoded by treA, which is not part of the treBC operon but strongly induced by high osmolarity (44) and partially depends on RpoS (214). Trehalose entering the periplasm will thus be cleaved by TreA and the released glucose will be taken up by PtsG, the glucose-specific PTS transporter. Mutants lacking TreA are unable to grow on trehalose at high osmolarity but are still able to grow on it at low osmolarity (45). The opposite is not true. Thus, mutants lacking the treBC system but expressing treA are still able to grow on trehalose. Trehalose produced at high osmolarity can be secreted, most likely by a PMF-dependent exit pump. Therefore, an alternate view for the function of the periplasmic trehalase is to scavenge the precious carbon source via the hydrolysis to glucose and its subsequent PtsG-mediated uptake (530).
Sucrose.
The Mng System.
The MalX System.
Aromatic β-Glucosides.
Cellobiose.
Maltose/Maltodextrins.
The maltose system contains four enzymes. The two most important ones are amylomaltase (603), a dextrinyl glucanotransferase encoded by malQ (402), and a maltodextrin phosphorylase encoded by malP (360, 495, 588). malP malQ form an operon oriented divergently to malT (411) (Fig. 8). It is significant that the malPQ operon is not directly under cAMP/CAP control, indicating a different dependence on catabolite repression than the transport genes (188).
Two other enzymes are known to act on maltose or maltodextrins but are not part of the maltose regulon. The first was discovered through the in vivo activity of a maltose transacetylase (55) and is encoded by maa. The purified enzyme was shown not only to acetylate maltose, but also to acetylate glucose even better. The structure of the protein has been elucidated by X-ray crystallography and additional substrates have been identified (308). The enzyme is similar to thiogalactoside transacetylase, the product of the lacA gene, the last gene of the lac operon (587). Considering the function of maltose transacetylase in the maltose system, it may act as a safety valve when the cytoplasmic concentrations of either maltose or glucose exceed healthy levels. When acetylated they will passively diffuse through the membrane.
Regulation of the maltose system is complicated. The transcriptional activator MalT is essential for the expression of all mal genes and malT null mutants cannot grow on maltose. In wild-type cells, the presence of maltose or any linear maltodextrin that can be taken up induces the system about 15- to 20-fold as compared with the expression level when grown on glycerol. Yet, in vitro, MalT can only be activated by maltotriose (only linear maltodextrins were tested) (412). Thus, it appears that the action of amylomaltase and maltodextrin phosphorylase is responsible for the production of sufficient maltotriose from any maltodextrin to induce the system (494). The complication in the understanding of mal gene regulation will be discussed on three different levels.
The first level concerns endogenous inducer synthesis. In the absence of external maltodextrins, the system can be induced by other carbon sources, for instance, trehalose (282). For this to happen, trehalose-6-phosphate hydrolase, the treC gene product that produces glucose 6-phosphate and glucose from trehalose 6-phosphate, is necessary (cf. Fig. 5). The same occurs in a pgm mutant. Thus, the two sugars glucose 6-phosphate and glucose must be used to produce maltotriose (or another unknown inducer) without the involvement of activated glucose (ADP- or UDP-glucose). Curiously, derepression of treC alone (without the presence of trehalose 6-phosphate in the cytoplasm or trehalose in the medium) will also induce the system even though less strongly. This indicates that trehalose-6-phosphate hydrolase with an unknown side reaction might be involved in the synthesis of inducer, possibly not maltotriose (109). Endogenous inducer, most likely maltotriose, is also produced during growth on limiting amounts of glucose, conditions under which glucose enters the cell without phosphorylation (107, 108). In fact, when glucose is taken up in a ptsG mutant by the mgl-encoded system, it induces the maltose system (110). The role of free internal glucose in the synthesis of inducer is underlined by the observation that overproduction of glucokinase interferes with glucose-mediated induction (329). Similarly, growth on galactose or lactose, particularly in pgm mutants (378, 494), induces the maltose system. This will be discussed further when dissecting galactose metabolism (see below). The curious phenomenon that sugars unrelated to the maltose system are able to induce it finds its explanation in the necessity of induction of the λ receptor for the effective diffusion of these sugars through the outer membrane. There is another source of internal inducer, that is, glycogen. Mutants lacking amylomaltase accumulate several maltodextrins including the inducer maltotriose (129), provided that they are able to synthesize glycogen. Thus, malQ mutants will exhibit nearly constitutive mal gene expression (110). Apparently, the role of amylomaltase is to reshuffle these glycogen-derived maltodextrins (including maltotriose), routing them again into degradation by the mal-encoded enzymes and, consequently, keeping endogenous induction low. The overexpression of glucokinase in malQ mutants has no effect on the semiconstitutive mal gene expression, demonstrating that the glycogen-derived inducer synthesis is different from the one derived from glucose/glucose 6-phosphate (329). The pathway of glycogen degradation (chapter 67) leading to the formation of maltodextrins is not entirely clear. The cluster of genes involved in glycogen synthesis (400), glgA for glycogen synthase (627), glgC for ADP-glucose pyrophosphorylase (24), and glgB for the branching enzyme (21), also harbors genes encoding enzymes that are thought to be involved in the degradation of glycogen. Thus, the product of glgX will act as a debranching enzyme removing α-1,6-linked chains from the "main strand" that are maximally 4 glucosyl residues long ("phosphorylase limit dextrin") (624), whereas glycogen phosphorylase, the product of glgP (85), will shorten the extended chains sequentially from their nonreducing end maximally to the length seen in the "phosphorylase limit dextrin." Thus, most likely, the glgX-encoded debranching enzyme releases maltotetraose from glycogen and plays an important role in the production of endogenous inducer. Since glycogen degradation affects the endogenous induction of the maltose system, the regulation of glycogen synthesis may be connected to the regulation of the maltose system as well. The major players in the expression of the glg genes are the cAMP/CAP complex (450), the stress sigma factor RpoS (213), and the CsrA/B system (449). Other regulatory effects can be introduced by modulation of enzyme activities involved in glycogen synthesis. Thus, glycogen phosphorylase is stimulated by fructose 1,6-diphosphate (24) and by dephosphorylated HPr of the PTS (500). Also, there is the possibility to control the availability of ADP-glucose, the substrate of glycogen synthase. The product of aspP is an ADP-glucose pyrophosphatase that could control glycogen synthesis (338).
The uptake of galactose can be mediated by two different systems. The first system is a binding protein-dependent ABC transporter encoded by the mgl genes at 48.2 min on the E. coli chromosome in the following order: mglB (galactose-binding protein), mglA (ATP-hydrolyzing subunit, a hybrid dimer), and mglC (the transmembrane component) (203, 224) (Fig. 11). The name for the transporter is derived from its alternate substrate β-methyl-D-galactopyranoside (454). The substrate specificity of the transport system reflecting the binding specificity of the binding protein includes also glucose and (2R)-glyceryl-β-D-galactopyranoside (42, 509). The mgl genes are regulated by the repressor GalS encoded by gene galS, which is located in front of and tandemly oriented to the mgl genes (163, 594). D-Galactose acts as an inducer inactivating GalS. Considering that D-glucose is a high-affinity substrate of the system being taken up preferentially by the Mgl system under limiting glucose concentrations (107), one wonders whether GalS may recognize glucose as well. The Mgl system does not entail genes encoding galactose-utilizing enzymes or enzymes hydrolyzing β-galactosides, rather, it serves primarily as a scavenger (614). Consistent with this role is the involvement of the galactose-binding protein in galactose chemotaxis (208, 263). As in the maltose/maltodextrin system the mgl system is present in Salmonella in identical form (29, 146, ). The metabolism of galactose will be dealt with below when discussing GalP-mediated uptake of galactose.
Ribose 5-phosphate isomerases (EC 5.3.1.6) interconvert ribose 5-phosphate and ribulose 5-phosphate. This reaction allows the synthesis of ribose from the pentose phosphate pathway and represents a means for the salvage of carbohydrates after nucleotide breakdown. However, such a dual role for a metabolic pathway is unusual and presents problems for effective metabolic regulation (511). Two unrelated types of enzymes can catalyze the reaction (102). RpiA is highly conserved and present in almost all organisms. In E. coli and Salmonella, the enzyme is constitutively expressed (238). The second type of ribose isomerase, RpiB, is sometimes referred to as AlsB because it can also take part in the metabolism of the rare sugar allose (see above). E. coli strains defective in rpiA are ribose auxotrophs, despite the presence of wild-type rpiB. Ribose prototrophs of an rpiA genetic background arise spontaneously and map in rpiR or result from the cloning of rpiB on a multicopy plasmid (515). RpiA exhibits an α/β/(α/β)/β/α fold, some portions of which are similar to proteins of the alcohol dehydrogenase family (416, 632). The two subunits of the dimer in the asymmetric unit have different conformations, representing the opening/closing of a cleft. The enzyme presumably acts by an acid-base catalysis mechanism.
Secondary transporters use the energy derived from ion gradients across the membrane to facilitate transport of solutes (chapters 7 and 19). Cation-sugar cotransport mechanisms in prokaryotes are generally energized by the electrochemical gradient of protons, the proton motive force (PMF), whereas those of eukaryotes are energized by the gradient of sodium ions.
Lactose.
The discovery of the lactose transport system also goes back to the early days of transport studies (440). The proof that the lacY gene product was in fact a protein was for quite some time one of the most cited publications in the transport literature (149), and the use of the lactose permease was instrumental in the establishment of the chemiosmotic theory of Peter Mitchell (597). The culmination of the many studies on the lactose transporter (done predominantly in the laboratory of Ron Kaback) was the recent resolution of its crystal structure (2).
As β-galactosidase is, the Lac permease is rather permissive in substrate recognition. Many different β-galactosides can be transported and subsequently hydrolyzed by β-galactosidase, a convenient method getting different aglycons into the cell; for example, it is possible to produce N-acetylglucosamine (GlcNAc) within the cell by transport (through LacY) of N-acetyl lactosamine (LacNAc, Gal β-1,4-GlcNAc), which is further hydrolyzed by β-galactosidase (J. Sattler and C. Mayer, unpublished results). However, these β-galactosides usually are not able to induce the lac system.
The utilization of lactose is strongly reduced by glucose in the medium because of catabolite repression and inducer exclusion, which are known to act on many carbohydrate-metabolizing systems. The molecular explanation is based on the control of adenylate cyclase by EIIAGlc-phosphate (stimulation) and the inhibition of the respective transport systems by dephosphorylated EIIAGlc of the phosphotransferase system (394). In the lactose system, the major portion of the "glucose effect" appears to be due to exclusion (280).
Considering nomenclature it is unlikely that lactose is the natural substrate of the E. coli lactose system. E. coli in the gut of mammals will encounter lactose only during the time of milk intake. However, mammals will digest plant leaves during their entire life span. Leaves of green plants contain large amounts of galactolipids, the major lipids of chloroplasts (471). After deacylation of the fatty acids, (2R)-glyceryl-β-D-galactopyranoside is formed in great quantity. This compound is an excellent substrate of the lactose permease, the LacI repressor, β-galactosidase, and even the lacA gene product (509) and may in fact be regarded as the physiological substrate of the lactose system (Fig. 11).
Besides the ABC transporter MglABC (see above), a second transporter for galactose is encoded by galP at 66.5 min on the E. coli chromosome, a typical PMF-dependent transporter (13, 242, 313, 322, 327). galP is part of the galactose regulon that is controlled by a repressor, the product of galR (61, 175, 593). Derepression mediated by galactose binding to GalR may not involve the dissociation from its cognate operator (74, 586). GalR is also the repressor for the galETKM operon encoding the enzymes involved in galactose metabolism (Fig. 11). Galactose, once inside the cell, is first phosphorylated at the 1-position, the reducing end, by the ATP-dependent galactokinase encoded by galK (501). The substrate of the kinase is α-D-galactose, which is phosphorylated under retention of the anomeric carbon to α-D-galactose 1-phosphate. Alternatively, β-galactose released from internal β-galactosides by β-galactosidase is too slow in its spontaneous mutarotation for the demand of galactokinase. This deficit in the utilization of β-galactose is overcome by the product of galM, a mutarotase (51). Since galM is part of the gal operon it may as well be involved in the utilization of galactose transported by either of the transporters, GalP or Mgl. Both α- and β-galactose may be transported equally fast, at least by the ABC transporter (581), and about 60% of galactose is in the β-form under equilibrium conditions. Galactose-1-phosphate uridylyltransferase (288, 457, 590) encoded by galT then catalyzes the reversible transfer of the uridine 5'-monophosphoryl moiety of UDP-glucose to galactose l-phosphate to form UDP-galactose and α-glucose 1-phosphate. UDP-glucose is re-formed from UDP-galactose by the action of UDP-galactose-4-epimerase (3, 549, 550) encoded by galE, the first gene in the gal operon. Thus, one would predict that the transformation of α-glucose 1-phosphate into glucose 6-phosphate by phosphoglucomutase (encoded by pgm) would be essential for the utilization of galactose as a carbon source since this seems to be the only path for α-glucose 1-phosphate to enter glycolysis. However, this is not the case. In the reverse reaction, maltodextrin phosphorylase, the product of malP, will form long maltodextrins (staining blue with iodine) from α-glucose 1-phosphate (cf. Fig. 8). The degradation of these maltodextrins by the enzymes of the maltose system releases glucose for the phosphorylation to glucose 6-phosphate by glucokinase; glucose 6-phosphate then enters glycolysis (494). In particular, maltodextrin glucosidase, encoded by malZ, reveals its essential nature in forming cytoplasmic glucose. Similarly, pgm mutants can still grow on maltose but only if they contain a functional MalZ enzyme (378). pgm mutants should also be unable to produce α-glucose 1-phosphate gluconeogenically from glucose 6-phosphate. Since UDP-glucose, the activated sugar needed for the utilization of galactose, is made from glucose 1-phosphate it follows that pgm mutants should not grow on galactose simply for this reason. Since this is not the case, there must be some source, even if minute, of internal α-glucose 1-phosphate in the absence of Pgm. On the other hand, UTP-α-glucose-1-phosphate uridylyltransferase, the enzyme producing UDP-glucose from UTP and α-glucose 1-phosphate (595), is an essential enzyme for galactose utilization. This is reflected in the name of the encoding gene, galU, even though this gene is not part of the galactose regulon. galU mutants cannot grow on galactose but are sensitive to it when grown on another carbon source. This has been explained by the accumulation of galactose 1-phosphate when the cells are exposed to galactose. Growth inhibition is a common phenomenon when phosphorylated sugars accumulate to high internal concentrations, but the reason for it remains elusive (40, 262, 448, 467). Similar to galU mutants, galT and galE mutants are sensitive to the presence of galactose in the growth medium. The former will also accumulate α-glucose 1-phosphate, whereas the latter will accumulate UDP-galactose. galE mutants are well suited to label galactose-containing polysaccharides by the addition of trace amounts of radioactively labeled galactose to the medium (417). UDP-14C-galactose will be formed internally and not further metabolized. Thus, any transferase will use the radioactive precursor for its reaction.
A projection structure at 8-Å resolution of the melibiose permease displayed an asymmetric protein unit consisting of two domains lining a central and curve-shaped cleft (192, 239). A secondary structure model of melibiose permease was obtained by studying MelB-PhoA fusions, and the previously predicted topology of 12 transmembrane domains was supported (49, 397, 398). These 12 transmembrane helices of MelB are connected by periplasmic and cytoplasmic loops, with both the C and N termini located on the cytoplasmic side of the membrane. Thirty-six amino acids could be removed from the hydrophilic carboxyl domain without loss of sugar specificity, facilitated diffusion, uphill transport, or H+-coupling or Na+-coupling characteristics (50). These results are consistent with the hypothesis that the sugar/cation-binding site is formed by the interaction of the transmembrane helices 3, 4, 6, 9, and 10 and does not involve the carboxyl-terminal portion of the protein. The cytoplasmic loop connecting helices 10 and 11 (loop 10–11), the largest cytoplasmic loop in the membrane-bound melibiose carrier of E. coli, is a functional reentrant loop (113). Photolabeling and limited proteolysis studies suggested that one of the four basic residues of the N-terminal loop 4–5 has a potential role in its symport function (7). Cys replacement analysis indicated that two of the four basic residues, Arg-141 and Arg-149, but not Lys-138 and Arg-139, are essential for MelB transport activity. Replacement of Arg-141 by neutral residues (Cys or Gln) inactivated transport and energy-independent carrier-mediated flows of substrates (counterflow, efflux), whereas it had a limited effect on cosubstrate binding (1).
After entering the cytoplasm melibiose is hydrolyzed by MelA, an α-galactosidase forming glucose and galactose. MelA is a member of family 4 of glycosyl hydrolases that requires NAD+ and Mn2+ for its hydrolytic activity and proceeds by a so-far-unknown mechanism (215, 551, 552). Among the α-galactosidases, melibiose seems exceptional in that it might also induce and be transported by the lactose permease LacY (301). However, the compound cannot be hydrolyzed by the β-galactosidase LacZ (301).
The activity of MelA is controlled by Aes, an esterase of unknown physiological function. Complexes between Aes and α-galactosidase are reduced in α-galactosidase activity but increased in esterase activity (318). Aes is also involved in controlling MalT, the central activator of the maltose/maltodextrin system (255, 379). In both cases, the interaction with Aes seems to prevent the self-multimerization of either MalT or α-galactosidase that is required for their activity (490).
Contrary to general concepts of bacterial saccharide metabolism, melibiose and fructose accumulated as extracellular intermediates during the catabolism of raffinose by ethanologenic recombinants of E. coli B, K. oxytoca, and Erwinia chrysanthemi. Both hydrolysis products (melibiose and fructose) were subsequently transported and further metabolized by all three organisms (333). The mechanism of sugar escape remains unknown but may involve downhill leakage via permeases that transport precursor saccharides or novel sugar export proteins. If sugar escape occurs in nature with wild organisms, it could facilitate the development of complex bacterial communities that are based on the sequence of saccharide catabolism and the hierarchy of sugar utilization.
Sucrose.
Fructoselysine.
Sialic acid (SA) is a general term for a family of unique 9-carbon monosaccharides, with N-acetylneuraminic acid (NeuAc, 2-keto-3-deoxy 5-acetamido-D- glycero-D- galacto-nonulosonic acid) being the most common natural form of SA, primarily found in animal tissue glycoconjugates and in the antigens of some bacterial strains (476, 574). The K-1 capsule polysaccharide colominic acid is an α-2,8-linked polymer of NeuAc (36, 447, 573) and confers invasiveness to E. coli, causing meningitis during the neonatal period (41, 276, 277, 323). Moreover, LPS-containing SAs have been reported in several Enterobacteriaceae, including E. coli and Salmonella strains (25, 158, 159). In contrast to the limited phylogenetic distribution of sialate biosynthesis, many commensal and pathogenic bacteria use sialic acids as sources of carbon, nitrogen, and energy or as sources of amino sugars for cell wall and membrane components (264, 390, 572). Microbial sialic acid metabolism has now been firmly established as a virulence determinant in a range of infectious diseases (574). In general, sialidases (or neuraminidases; EC 3.2.1.18) remove sialic acids from sialoglycoconjugates and have been found in viruses, bacteria, trypanosomes, and mammalian cells (541). There are two families of bacterial enzymes, distinguished by their requirement for a divalent metal ion for maximal activity (548). The sialidase of serovar Typhimurium (NanH) does not require metal ions and has a molecular mass of 42,000 Da (240, 241, 543). Recently, the three-dimensional structure has been determined at 1.6-Å resolution (94, 95). Serovar Typhimurium LT2 is able to use sialyl-α-2,3-lactose as the sole carbon and energy source, while a nanH strain cannot (Steenbergen and Vimr, unpublished data), consistent with a primarily nutritional function of bacterial sialidases (240).
NeuAc pyruvate lyase also catalyzes the reverse reaction, the direct condensation of ManNAc and pyruvate in vitro (144, 443). However, the synthesis of NeuAc for colominic acid biosynthesis (by CMP-NeuAc synthetase, EC 2.7.7.43) in vivo depends on NeuB, the sialic acid synthase (442, 568). In eukaryotic cells, NeuAc synthesis implies the formation of ManNAc 6-phosphate and NeuAc 9-phosphate (476). The value of NeuAc lyase as a reagent for the synthesis of therapeutically useful sialic acid derivates has been proven (574) and stimulated the biotechnical work on NanA (306, 583). The three-dimensional structure of the native NeuAc aldolase has been determined and demonstrated its organization as a homotetramer of 33-kDa polypeptides (243). For further degradation, ManNAc is phosphorylated at the C-6 position by a ManNAc kinase encoded by nanK. The product of the reaction, ManNAc 6-phosphate, is then isomerized to GlcNAc 6-phosphate by a putative isomerase encoded by nanK (390). In the consequence, the NeuAc, ManNAc, and GlcNAc metabolisms merge at the level of ManNAc 6-phosphate or GlcNAc 6-phosphate (390) (Fig. 14).
Xylitol.
The carbohydrate moieties of ubiquitous macromolecules, for example, DNA, RNA, phospholipid, glycogen, and oligosaccharides of the cell surface, have all been shown to serve E. coli and Salmonella as sole carbon and energy sources; however, they are not covered within this chapter (see chapters 5, 6, 9, and 21). Also not included in this survey are the dissimilation pathways of sugar acids (e.g., hexuronides, hexuronates, and hexonates), phosphorylated sugars, carboxylates, and glycerol, which are described in chapter 20 and Chapter Catabolism of Hexuronides, Hexuronates, Aldonates, and Aldarates.
Despite the vast quantity of information available regarding the sugar and sugar alcohol dissimilation of E. coli and Salmonella, as pointed out in this chapter, it is known from genomic sequence data that there are still some gene clusters that have been assigned to sugar metabolism but to which no substrate has yet been attributed. Without a claim for completeness, some of the unkown gene clusters assigned to sugar and sugar alcohol metabolism will be introduced here.
It is expected that with the genomic sequences in hand, approaches that use "reverse genetics" will be a promising means to discover the function of yet unknown sugar metabolic pathways.
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Module3.4.1
