2 Rational Drug Design Program and Department of Bacteriology and Immunology, Biomedicum and Haartman Institute, PO Box 63, FIN-00014 University of Helsinki, Helsinki, Finland; 3 HUCH Laboratory Diagnostics, Helsinki University Central Hospital, PO Box 401, FIN-00029 HUCH, Helsinki, Finland
Received on September 12, 2003; revised on October 18, 2003; accepted on November 25, 2003
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Abstract |
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Key words: 6-deoxy-talose / fucose / rhamnose / nucleotide sugars
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Introduction |
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For decades tremendous efforts have been made to determine the exact chemical structure of various cell surface glycans in bacteria. Recently, much effort has also been put toward elucidating the enzymology involved in the biosynthesis of these glycans. Glycans are synthesized from nucleotide sugars by specific glycosyltransferases that catalyze the transfer of monosaccharides to an acceptor molecule, for example, the growing polysaccharide structure. In general, there is at least one specific glycosyltransferase for each monosaccharide and specific linkage formed between a monosaccharide and an acceptor molecule. To date a large number of prokaryotic glycosyltransferases with different donor, acceptor, and linkage specificities have been cloned and characterized (reviewed in Thorson et al., 2001; Unligil and Rini, 2000
).
Nucleotide sugars are activated sugar metabolites that act as donors in glycosylation reactions. Drugs inhibiting the enzymes involved in the nucleotide sugar pathways specific for bacteria might not interfere with human metabolic pathways and thus might not be harmful for humans. Hence the biosynthetic pathways of bacterial nucleotide sugars could be considered potential targets for interventions of antibacterial therapeutics.
In this review, the biosynthetic pathways of the guanosine diphosphate (GDP)- and deoxythymidine diphosphate (dTDP)-activated fucose, rhamnose and 6-deoxy-talose (Figure 1) and the corresponding glycosyltransferases in bacteria are discussed. Fucose, rhamnose, and 6-deoxy-talose are 6-deoxyhexoses. In general, deoxysugars are an important class of carbohydrates that are formed from common monosaccharides by replacement of one or more hydroxyl groups with hydrogen, for example, in the formation of 6-deoxyhexoses the C6 carbon is deoxygenated. 6-Deoxyhexoses are formed from nucleoside diphosphate-activated hexoses via a 4-keto-6-deoxy intermediate (Figure 1). In bacteria, deoxysugars can be found in lipopolysaccharides (LPSs), extracellular polysaccharides (EPSs), glycoproteins, and in different classes of glycosylated secondary metabolites, for instance, antibiotics (reviewed in He and Liu, 2002; Kren and Martinkova, 2001
; Power and Jennings, 2003
; Thorson et al., 2001
; Trefzer et al., 1999
).
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Fucosylated glycans |
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L-fucose is an important component of the complex glycoconjugates of species ranging from bacteria to mammals. In humans, L-fucose is an essential component of N-linked and O-linked glycoproteins and glycolipids that mediate intercellular adhesion and recognition processes and thereby play a role in embryogenesis, immunity, inflammation, and metastasis of malignant cells. Specifically, L-fucose is present in the human ABO blood group antigens and Lewis (Le) glycans (reviewed in Becker and Lowe, 2003; Tonetti et al., 1998b
).
L-fucose is also a constituent of the cell wall and capsule structures of Gram-negative and Gram-positive bacteria. For instance, most Escherichia coli strains as well as the other species of the Enterobacteriaceae family are known to produce L-fucose containing EPS colanic acid (reviewed in Whitfield and Roberts, 1999). In addition, L-fucose has been shown to be a substantial cell wall component of several human pathogens, for example, H. pylori (reviewed in Appelmelk et al., 2000
), E. coli O:157 (Barua et al., 2002
; Wang and Reeves, 1998
), Campylobacter fetus (Moran et al., 1994
), and Yersinia enterocolitica O:8 and Yersinia pseudotuberculosis O:3 (Skurnik and Zhang, 1996
). L-fucose is also a structural component of the Nod factors that are important signals in the nodulation of the genera of plant microbes Azorhizobium and Rhizobium (reviewed in Carlson et al., 1994
; Trefzer et al., 1999
).
L-fucose in H. pylori LPS
Helicobacter pylori, a microaerophilic bacterium, is an important human gastric pathogen, infecting about half the world's population. Helicobacter pylori is a major cause of chronic gastritis and plays a role in the other gastric diseases, such as duodenal ulcers, gastric ulcers, gastric cancer, and gastric lymphoma (reviewed in Marshall, 2002). The O-antigen of LPS in most H. pylori strains contains fucosylated Le glycans (Table I), predominantly the type II antigens Le x and Le y (Simoons-Smit et al., 1996
; Wirth et al., 1996
). Le antigens expressed by H. pylori are structurally similar to human Le blood group antigens. This restricted diversity in O-antigen expression of H. pylori is unusual and suggests a role for specific epitopes in pathogenesis. Additionally, H. pylori Le antigens can undergo phase variation, which is the random, reversible high-frequency switching of phenotype within a single strain (reviewed in Wang et al., 2000
). A recent study demonstrated that acidic environment can trigger phase variation from Le x to Le y expression (Moran et al., 2002
). The antigenic phase variation is most probably beneficial to H. pylori in adaptation to different environments and environmental changes, but its role in colonization and pathogenesis is unclear. However, phase variation has been demonstrated to contribute to the virulence of bacteria, such as Neisseria sp. (van Putten, 1993
) and Haemophilus influenzae (Humphries and High, 2002
; Weiser and Pan, 1998
).
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GDP-L-fucose metabolism
GDP-L-fucose is synthesized in vivo via two different metabolic pathways. In the salvage pathway, which is not found in bacteria, GDP-L-fucose is synthesized from L-fucose derived from exogenous sources by the action of an L-fucose kinase (EC 2.7.1.52) and a GDP-fucose pyrophosphorylase (EC 2.7.7.31) (reviewed in Becker and Lowe, 2003).
The de novo pathway of GDP-L-fucose (Figure 2), which is evolutionarily conserved, was first identified in bacteria in the 1960s (Ginsburg, 1960) and then described in plants (Liao and Barber, 1972
), mammals (Overton and Serif, 1981
), and invertebrates (Bulet et al., 1984
). This pathway starts from GDP-D-mannose, which is converted into GDP-L-fucose in three steps by two enzymes. The first reaction step is a dehydration of GDP-D-mannose, which leads to formation of an intermediate product, GDP-4-keto-6-deoxy-D-mannose and is catalyzed by a GDP-D-mannose 4,6-dehydratase (GMD, EC 4.2.1.47) (reviewed in Allard et al., 2002
). GMD catalyzes oxidation of the C4 group of the mannose ring to a keto form and the subsequent reduction of the C6 group to a methyl group. During the concerted action of oxidation and reduction, an intramolecular hydride transfer occurs from the C4 position to the C6 position on the mannose ring by a tightly bound nicotinamide adenine dinucleotide phosphate (NADP) cofactor, which is essential for the activity of GMD (Oths et al., 1990
).
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GMD
GMDs have been obtained at various degrees of purity from different sources, both from bacteria, such as Klebsiella pneumonie (Yamamoto et al., 1993) and mammals, such as porcine thyroid (Broschat et al., 1985
). Cloning and characterization of GMD have been performed from bacteria, such as E. coli (Sturla et al., 1997
) and H. pylori (Järvinen et al., 2001
; Wu et al., 2001
), Arabidopsis thaliana (known as MUR1) (Bonin et al., 1997
), and human (Ohyama et al., 1998
; Sturla et al., 1997
; Sullivan et al., 1998
). Multimerization of the purified and cloned GMDs vary as well as the molecular masses of the monomers (
4055 kDa). In general, the reported mature GMDs are homodimers consisting two
42-kDa subunits. Amino acid sequence GILFNHES is commonly found in the characterized GMDs, but the possible function of this sequence is unknown.
The resolved quaternary structure of E. coli GMD was shown to be a homodimeric protein with two 42-kDa monomers composed of two domains (Somoza et al., 2000
). The larger N-terminal domain contains the NADP(H) cofactor binding site in a classical Rossman fold topology and the smaller C-terminal domain harbors the binding site for the nucleotide sugar substrate. These two domains form a pocket where the cofactor and the substrate bind and catalysis of the reaction occurs. Both subunits are suggested to be catalytically competent. The dimeric structures have also been reported for the K. pneumonie (Yamamoto et al., 1993
) and human GMDs (Bisso et al., 1999
). However, the recently resolved 3D structure of A. thaliana GMD (named MUR1) is a tetramer (Mulichak et al., 2002
), and there are reports suggesting that H. pylori GMD is also a tetramer (Wu et al., 2001
) and that porcine thyroid GMD might be a hexamer (Broschat et al., 1985
).
The E. coli GMD model shows that GMD is a member of a short chain dehydrogenase/reductase (SDR) protein family. The SDR protein family is a very large family of enzymes, most of which are known to be NADH/NADPH-dependent oxidoreductases (Jörnvall et al., 1995). The 3D structure of E. coli GMD is closely related to the resolved homodimeric structures of E. coli GMER (Rosano et al., 2000
) and E. coli uridine diphosphate (UDP)-galactose 4-epimerase (GalE) (Thoden et al., 1996
), both of which are also members of the SDR protein family. GalE catalyzes the interconversion of UDP-galactose and UDP-glucose. In addition, the GMD model and site-directed mutagenesis studies show that the Ser-Tyr-Lys catalytic triad common to enzymes of the SDR protein family is found in E. coli GMD, with the exception that Thr replaces Ser. Most GMDs also have Thr in the usual place of Ser in the catalytic triad. This replacement of Ser to Thr is an unusual feature among the SDR protein family. However, the consequences of the exchange are not known. On the basis of the GMD model and mutational analysis, the Glu135 is proposed to play a role in deprotonation/reprotonation of the mannose ring, thus acting as an active-site base during the catalysis (Somoza et al., 2000
).
GDP-L-ß-fucose, which is the end product of the pathway, has been demonstrated to inhibit the activity of both human and bacterial GMD via a specific feedback inhibitory mechanism (Bisso et al., 1999; Sturla et al., 1997
). In E. coli GMD, GDP-L-ß-fucose also shows competitive inhibition in respect to GDP-D-mannose, indicating that GDP-L-ß-fucose probably binds to the same site on the enzyme as GDP-D-mannose. The competitive inhibition also indicates that GDP-L-ß-fucose regulates its own biosynthesis through the activity of GMD (Somoza et al., 2000
). In contrast, GDP-L-ß-fucose shows no competitive inhibition in human GMD (Bisso et al., 1999
). GDP-L-
-fucose, an anomer of GDP-L-ß-fucose, does not inhibit the activity of human and bacterial GMD (Bisso et al., 1999
; Sturla et al., 1997
).
A defective synthesis of GDP-L-fucose and especially defects in GMD activity has been linked to stem shoot development in plants (Bonin et al., 1997). Humans deficient in fucosylation suffer from the rare immune disorder leukocytes adhesion deficiency type II that is characterized by defective selectin ligand formation, recurrent infections, and severe mental and growth retardation (reviewed in Becker and Lowe, 2003
). Recently the molecular mechanism for this disorder has been located to the GDP-L-fucose transporter gene (Etzioni et al., 2002
; Lubke et al., 2001
; Luhn et al., 2001
). In contrast, increased fucosylation of glycoconjugates has been observed in metastasis of cancer patients (Martin-Satue et al., 1998
; Ura et al., 1997
) or inflammatory diseases (Kirveskari et al., 2000
; Toppila et al., 1999
, 2000
).
GMER
GMER was first purified from porcine thyroid (Chang et al., 1988) followed by cloning the corresponding gene from humans (named FX) (Tonetti et al., 1996
), E. coli (also named Fcl/WcaG) (Tonetti et al., 1998
), A. thaliana (named GER1) (Bonin and Reiter, 2000
), and H. pylori (Järvinen et al., 2001
; Wu et al., 2001
). Mature human and E. coli GMERs are homodimers of two
34-kDa subunits. The monomers of A. thaliana and H. pylori GMERs are also
35-kDa, but their possible multimerization is unsolved.
The 3D structure of native and three mutated E. coli GMERs have been elucidated (Rizzi et al., 1998; Rosano et al., 2000
; Somers et al., 1998
). GMER is a homodimeric protein with each monomer composed of two domains. The N-terminal domain contains NADP(H) cofactor binding site in a modified Rossman fold topology, and the C-terminal domain harbors the binding site for the nucleotide sugar substrate. GMER is a member of the SDR protein family and is closely related to the resolved homodimeric structures of E. coli GMD (Somoza et al., 2000
) and E. coli GalE (Thoden et al., 1996
). Additionally, the GMER model and site-directed mutagenesis studies show that the catalytic mechanism of GMER is based on the concerted action of Ser-Tyr-Lys residues forming the conserved catalytic triad of the SDR protein family (Rizzi et al., 1998
; Rosano et al., 2000
). The Cys109 and His179 residues are suggested to play a primary role in the epimerization reaction (Rosano et al., 2000
).
Among the SDR protein family, the ability of GMER to catalyze two different reactions (epimerization and reduction) at the same active site appears unique. Furthermore it is not common in the NADH/NADPH-dependent SDR protein family that the epimerization catalyzed by GMER occurs in the absence of its cofactor, NADPH (Menon et al., 1999). The bifunctional nature of GMER also separates the GDP-L-fucose biosynthesis from that of other deoxy or dideoxy sugars in which the epimerization and reduction reactions are catalyzed by separate enzymes.
Defects in GMER activity has been linked to nodulation (Lamrabet et al., 1999; Mergaert et al., 1997
) and survival of bacteria (McGowan et al., 1998
). In H. pylori, the gmer gene (also known as wbcJ) is up-regulated in response to an acidic environment. The acid-induced up-regulation suggests that an acidic pH may stimulate LPS synthesis, which may be substantial in decreasing the host gastric acid secretion and in masking the antigenic surface epitopes of H. pylori and thereby facilitating early colonization (McGowan et al., 1998
). Moreover, the H. pylori gmer knockout mutant is demonstrated to be more sensitive to acid stress than the wild-type strain, suggesting that O-antigen expression might contribute to acid survival of H. pylori (McGowan et al., 1998
).
FucTs
GDP-L-fucose and dTDP-D-fucose are used as substrates for specific fucosyltransferases that are responsible for transferring L-/D-fucose at different positions of an acceptor molecule. To date no D-fucosyltransferases has been identified, whereas 10 human L-fucosyltransferases (FucTs) responsible for 1,2-,
1,3-,
1,4-, or
1,6-linkages on glycans (FutTs IIX) (reviewed in de Vries et al., 2001
) or O-linkage to Ser or Thr have been identified (O-FucT-1) (Wang et al., 2001
). In addition, Roos and colleagues (2002)
have proposed candidates representing two novel human FucTs named FutX and XI. The candidates are proposed on the basis of in silico analysis, and they have not been experimentally verified. FucTs responsible for
1,2-,
1,3-,
1,4-, or
1,6-linkages on glycans have been identified only in a limited number of bacteria, for example, H. pylori and some Rhizobium species (reviewed in Oriol et al., 1999
; Wang et al., 2000
).
Three futT genestwo 1,3-fucTs (or
1,3/4-fucTs) and one
1,2-fucTare present in each H. pylori isolate investigated to date (Alm et al., 1999
; Rasko et al., 2002a
, 2000b
; Tomb et al., 1997
; Wang et al., 1999). The encoded FucTs from different H. pylori strains have varying levels of activity and different Type I or Type II acceptor specificities.
As in mammalian cells, the synthesis of Le antigens in H. pylori is directed by a series of glycosyltransferases that act sequentially on a precursor molecule. FucTs are responsible for the final steps in this process. It has been demonstrated that the Le x synthesis occurs similarly in H. pylori as observed in mammalian cells; L-fucose is added to the Type II precursor LacNAc by 1,3-FucTs (Figure 3) (Chan et al., 1995
). H. pylori NCTC 11637 strain has two
1,3-FucT genes, futA and futB, encoding two
1,3-FucTs with different specificities. Appelmelk and colleagues (1999)
have suggested that in H. pylori NCTC 11637 strain the futA gene product fucosylates internal LacNAc and the futB gene product fucosylates the terminal LacNAc of O-antigen.
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H. pylori and eukaryotic FucTs are highly divergent in overall amino acid sequences and in domain structures. All the mammalian FucTs have a domain structure containing a short N-terminal cytoplasmic tail, a transmembrane domain, and a stem region followed by a large globular C-terminal catalytic domain. The N-terminal cytoplasmic tail and transmembrane domain have a role in the Golgi localization and in the retention of the enzyme. On the contrary, the amino acid sequences of H. pylori FucTs are devoid of a N-terminal transmembrane domain. The H. pylori 1,3 FucT has been shown to be membrane associated, whereas the
1,2 FucT has been found to be a soluble protein (reviewed in Wang et al., 2000
). In addition, a recent study by Ma and colleagues (2003)
demonstrated that the C-terminal domain of H. pylori FucTs is responsible for the type I and II acceptor recognition, whereas in human FucTs the acceptor specificity is mainly determined by the N-terminal stem domain. Breton and co-workers (1998)
have suggested that because all fucosyltransferases utilize GDP-L-fucose as an L-fucose donor, their specificity will probably reside in the recognition of the acceptor and in the type of linkage formed. Detailed studies on H. pylori (already discussed) and human FucTs (Niemelä et al., 1998
; Toivonen et al., 2002
) support this hypothesis.
1,3/4-fucTs genes (also known as futA/B) of H. pylori contain characteristic polyadeninepolycytosine tracts at the 5' end that have been identified as the cause of DNA slippage leading to the on/off switch of the target gene at the translational level (Appelmelk et al., 1999
). The
1,3/4-fucTs genes also contain a 21-mer repeat region at the 3' end that may play a role in the dimerisation and translational control of the FucT proteins (Ge et al., 1997
). The
1,2-fucT gene (also named futC) contains similar polycytosine tract like the
1,3/4-fucTs genes as well as an adenine-rich sequence at midregion of the gene that may be responsible for slipped strand mispairing leading to the on/off status of
1,2-fucT (Wang et al., 1999a
, 1999b
). These cis-elements of H. pylori fucT genes lead to the phase variation of Le antigens. As mentioned, the antigenic phase variation is most probably beneficial to H. pylori in adaptation to different environments and environmental changes, but its role in colonization and pathogenesis is unclear.
Genetic organization of genes related to L-fucose metabolism
In bacteria, the genes required for the synthesis of nucleotide sugars are generally scattered within the same gene cluster for a particular bacterial polysaccharide. GDP-L-fucose pathway genes have been reported to belong to the wb (formely named rfb) gene cluster that encodes O-antigen synthesis (Zhang et al., 1996) or the CA gene cluster that encodes colanic acid synthesis (Stevenson et al., 2000
). It has been suggested that similarity in the 3D structures of E. coli GMD and GMER with the observation that the corresponding genes are adjacent to each other on the bacterial genome might be due to a common ancestor (Somoza et al., 2000
).
In H. pylori, O-antigen synthesizing genes are not clustered in one locus, as is the case in other bacteria. Instead, only manC, the D-mannose-6-phosphate isomerase/GDP-D-mannose pyrophosphorylase gene (also known as rfbM), gmd (also known as rfbD), and gmer (also known as wbcJ) genes are clustered, and the other genes required for the O-antigen synthesis are distributed throughout the genome (Alm et al., 1999; Tomb et al., 1997
). Berg and co-workers (1997)
have suggested that this unusual genetic organization in H. pylori may be due to the special feature of H. pylorihost interaction that may favor the exchange of LPS-synthesizing genes en bloc. Furthermore, the genetic organization is also in accordance with a proposed assembly mechanism of H. pylori O-antigen that differs from the mechanism used by other Gram-negative bacteria. Instead of transferring subunits consisting of several monosaccharides onto the growing O-antigen, H. pylori O-antigen may be assembled by sequential addition of a single monosaccharide (Berg et al., 1997
; Rasko et al., 2000).
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Rhamnosylated glycans |
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The more common L-rhamnose is found in a wide variety of both Gram-negative and Gram-positive bacteria, in which it is a common component of cell wall and capsule structures (reviewed in Giraud and Naismith, 2000). The widespread prevalence of L-rhamnose and its relevance for several clinically significant pathogens has made the biosynthetic pathway of L-rhamnose an appealing target for novel therapeutic interventions. It has been demonstrated that the disruption of the dTDP-L-rhamnose pathway has attenuated the virulence or influenced the viability of pathogenic bacteria, such as P. aeruginosa, Mycobacterium tuberculosis, Vibrio cholerae, Enterococcus feacalis, and Streptococcus mutans (Chiang and Mekalanos, 1999
; Yamashita et al., 1999
; Rahin et al., 2000
; Xu et al., 2000
). These observations elucidate the importance in bacterial pathogenicity of enzymes involved in the biosynthesis of nucleotide sugars. Humans neither synthesize nor utilize L-rhamnose.
dTDP-L-rhamnose metabolism
The activated nucleotide sugar form of L-rhamnose is dTDP-L-rhamnose, which is synthesized from glucose-1-phosphate and deoxythymidine triphosphate (dTTP) via a biosynthetic pathway requiring four enzymes RmlAD (Figure 1). The quaternary structures of RmlAD enzymes have been solved, and the structural data on substrate complexes have recently been discussed in detail elsewhere (Dong et al., 2003).
The first enzyme in the dTDP-rhamnose pathway is glucose-1-phosphate thymidylyltransferase (RmlA, EC 2.7.7.24), which catalyzes the transfer of a thymidylmonophosphate nucleotide to glucose-1-phosphate. The catalytic activity of RmlA is allosterically regulated by the end product of the pathway, dTDP-L-rhamnose (Blankenfeldt et al., 2000a, 2000b
). The similar specific feedback inhibitory mechanism has also been detected in the biosynthetic pathways of GDP-L-fucose (Sturla et al., 1997
) and GDP-D-mannose (Wu et al., 2002
). RmlA is a homotetramer with the monomer consisting three functional domains: one core domain that shares the sequence similarity with nucleotidyltransferases and two other domains that contain the recognition and binding sites for the nucleotide and sugar-phosphate. The active center of the RmlA is located in a pocket formed by the core and sugar binding domains (Blankenfeldt et al., 2000).
The second enzyme, dTDP-D-glucose 4,6-dehydratase (RmlB, EC 4.2.1.46), catalyzes an oxidation of the C4 hydroxyl group of the D-glucose residue that is followed by dehydration, which leads to the formation of dTDP-4-keto-6-deoxy-D-glucose. Mechanistically, the catalytic action of RmlB is closely related to that of GMD. RmlB uses NAD(H) as a cofactor, in contrast to GMD utilizing NADP(H) as a cofactor (Allard et al., 2001; Somoza et al., 2000). RmlB is a homodimer, and each monomer exhibits an
/ß structure and is composed of two domains. The N-terminal domain contains dinucleotide-binding motif, the Rossmann-type fold, and the C-terminal domain is responsible for the binding of dTDP-D-glucose. The highly conserved catalytic residues Tyr and Lys identify RmlB as a member of the SDR protein family (Allard et al., 2000
, 2001).
The third enzyme, dTDP-4-keto-6-deoxy-D-glucose 3, 5-epimerase (RmlC, EC 5.1.3.13), catalyzes a double epimerization reaction at the C3 and C5 positions of the 4-keto-6-deoxy-D-glucose ring (Graninger et al., 1999). GMER also catalyzes a double epimerization at positions C3 and C5 of the 4-keto-6-deoxy-D-mannose ring, but RmlC shows no structural similarity to GMER (Christendat et al., 2000
; Giraud et al., 2000
; Rosano et al., 2000
). RmlC is a homodimer, and the monomer is predominantly formed of ß-sheets with a jelly rolllike topology. The active center of the RmlC is located at the entrance of ß-barrel formed by both monomers (Giraud et al., 2000
). The cofactor (NADPH) and the nucleotide sugar binding sites of GMER have no counterpart in RmlC, and thereby Giraud and colleagues (2000)
suggested that the mechanisms of epimerization are likely to be different in these enzymes.
Finally, in the synthesis of dTDP-L-rhamnose the fourth enzyme is dTDP-4-keto-6-deoxy-L-mannose reductase (RmlD, EC 1.1.1.133), which reduces the C4 keto group of the 4-keto-6-deoxy-L-mannose moiety and leads to the formation of dTDP-L-rhamnose (Graninger et al., 1999). The functional form of RmlD is a dimer. Each monomer consists two domains: the N-terminal domain that is dominated by the Rossmann-like fold is responsible for the cofactor NAD(P)H binding, and the C-terminal domain is involved in the substrate binding. Moreover RmlD requires Mg2+ to stabilize the dimer interface and for the full enzyme activity (Blankenfeldt et al., 2002
). RmlD contains the characteristics of the SDR protein family, that is, the catalytic triad of Thr-Tyr-Lys and Wierenga motif (Jörnvall et al., 1995
), and it shares sequence homology with GalE, GMD, and GMER in those amino acids known to have catalytic function (Blankenfeldt et al., 2002
; Rosano et al., 2000
; Somoza et al., 2000
; Thoden et al., 1996
). RmlD also shares amino acid identity with its direct analog, a GDP-4-keto-6-deoxy-D-mannose reductase (RMD) involved in the synthesis of GDP-D-rhamnose (Kneidinger et al., 2001
).
The RmlAD enzymes are highly conserved among microorganisms. The genes encoding the RmlAD enzymes are located in the wb (formely known as rfb) gene cluster together with genes required for the assembly of particular bacterial polysaccharide. However, the four rml genes may not necessarily be clustered next to each other (reviewed in Giraud and Naismith, 2000).
Rhamnose in P. aeruginosa LPS
P. aeruginosa is a natural soil inhabitant with a very versatile metabolic potential, permitting it to survive in a number of environments. It is also a human opportunistic pathogen, infecting mainly immunocompromised patients and causing a range of severe infections. A new clinical problem is the rapidly increasing resistance of Pseudomonas aeruginosa against commonly used antibiotics (Kiska and Gilligan, 1999). Hence the virulence factors and the pathogenesis of P. aeruginosa have recently gained much interest.
LPS play an important role in the virulence of this ubiquitous bacterium. Pseudomonas aeruginosa synthesizes concomitantly two chemically and antigenically distinct forms of O-antigen, known as a common A-band and a serotype-specific B-band (reviewed in Rocchetta et al., 1999). The constitutively expressed A-band O-antigen is a homopolymer consisting of D-rhamnose sugar residues arranged as repeating trisaccharide units (
3D-Rha
1
2D-Rha
1
3D-Rha
1
)n (Arsenault et al., 1991
). In contrast, the B-band O-antigen is a heteropolymer composed of repeating di- to pentasaccharide units of many different monosaccharides (Knirel and Kochetkov, 1994
). The B-band O-antigen has been shown to impede phagocytosis and is responsible for resistance to host serum (Engels et al., 1985
). D-rhamnose is found only in the A-band O-antigen, whereas L-rhamnose is found in the core oligosaccharide (Rahim et al., 2000
), B-band O-antigen, and rhamnolipids of P. aeruginosa (reviewed in Maier and Soberon-Chavez, 2000
; Rocchetta et al., 1999
). The gene clusters responsible for the synthesis of A- and B-band LPS have been identified and their genetics and regulatory mechanisms as well as their relevance to the pathogenesis of P. aeruginosa are under extensive study. The availability of the whole genome sequence of P. aeruginosa has contributed to these studies (Stover et al., 2000
).
Pseudomonas aeruginosa is commonly isolated from the specimens obtained from the lungs of patients suffering the congenital monogenic disease cystic fibrosis. The respiratory P. aeruginosa isolates from the chronic cystic fibrosis infection are devoid of O-antigen or mainly express the D-rhamnosylated A-band O-antigen (reviewed in Lyczak et al., 2002; Rocchetta et al., 1999
). Hitherto no specific condition has been identified to influence the synthesis or regulation of A-band LPS. Interestingly, exactly the same D-rhamnan polysaccharide structure as observed in A-band LPS of P. aeruginosa has also been found from the other opportunistic pathogens Burkholderia cepacia and Stenotrophomonas maltophilia, also associated with cystic fibrosis with severe pulmonary manifestation (Cerantola and Montrozier, 1997
; LiPuma, 2000
; Winn and Wilkinson, 1998
). In addition, B. cepacia and S. maltophilia have also emerged as important multidrug-resistant pathogens and a cause of nosocomial infections (reviewed in Mahenthiralingam et al., 2002
; Zhang et al., 2000
).
GDP-D-rhamnose metabolism
Markovitz (1964) proposed a biosynthetic pathway for GDP-D-rhamnose in the 1960s. Recently, Kneidinger and colleagues (2001)
provided evidence for the proposal by cloning the enzymes required for the synthesis of GDP-D-rhamnose. The GDP-D-rhamnose pathway starts from GDP-D-mannose (Figure 4), which is converted into GDP-4-keto-6-deoxy-D-mannose by GMD, as is the case in the synthesis of GDP-L-fucose (Figure 2). The following step in the GDP-D-rhamnose pathway is the specific reduction of the 4-keto group of GDP-4-keto-6-deoxy-D-mannose, which leads to formation of GDP-D-rhamnose and is catalyzed by GDP-4-keto-6-deoxy-D-mannose reductase (RMD) (EC 1.1.1.187). During the action of reduction RMD can use either NADH or NADPH as hydride donors. The reduction activity of RMD is analogous to the activity of the RmlD enzyme, which is responsible for the reduction of the C4 keto group of dTDP-4-keto-6-deoxy-L-mannose in the synthesis of dTDP-L-rhamnose (Giraud et al., 1999
). Kneidinger and colleagues (2001)
identified two enzymes, GMD and RMD, responsible for the GDP-4-keto-6-deoxy-D-mannose reductase activity from nonpathogenic, Gram-positive A. thermoaerophilus L42091T bacterium. D-rhamnose is a constituent of the surface layer glycoprotein of this bacterium. The specific function of D-rhamnosylated S-layer of A. thermoaerophilus L42091T is unknown. The RMD enzyme is also functionally characterized from P. aeruginosa (Mäki et al., 2002
).
|
Genes encoding enzymes involved in the synthesis of nucleotide sugars are usually found in the gene cluster for a particular bacterial polysaccharide. In the genome of A. thermoaerophilus or P. aeruginosa, the rmd is located directly adjacent to the gmd gene. The characterized A. thermoaerophilus and P. aeruginosa RMDs act only as GDP-4-keto-6-deoxy-D-mannose reductases in the synthesis of GDP-D-rhamnose. The molecular masses of the monomers of RMDs are 35 kDa, whereas the possible multimerization of the enzymes are unknown. The amino acid sequences of RMDs contain a N-terminal dinucleotide-binding motif ThrGlyXXGlyXXGly, and an active-site Ser-Tyr-Lys triad common to the SDR protein family (Kneidinger et al., 2001
; Mäki et al., 2002
). Other residues involved in GMD catalysis (Somoza et al., 2000
), with the exception of Glu135, are not found in RMDs (Kneidinger et al., 2001
; Mäki et al., 2002
). In GMD catalysis, Glu135 is proposed to play a role in the deprotonation/protonation of the hydroxyl group of the C5 position of GDP-4-keto-6-deoxy-D-mannose (Somoza et al., 2000
), but this C5 atom is not involved in the reduction reaction catalyzed by RMD (Kneidinger et al., 2001
).
Rhamnosyltransferases
GDP-D-rhamnose and dTDP-L-rhamnose are used as substrates for specific rhamnosyltransferases that are responsible for transferring L-/D-rhamnose to an acceptor molecule via a specific linkage. Currently very little is known about the bacterial rhamnosyltransferases. L-rhamnosyltransferases, which use dTDP-L-rhamnose as a donor, have been reported in several bacteria (reviewed in Giraud and Naismith, 2000), whereas putative D-rhamnosyltransferases, which use GDP-D-rhamnose as a donor, have only been identified from P. aeruginosa (Rocchetta et al., 1998
). On the basis of mutagenesis studies, it has been shown that three putative P. aeruginosa D-rhamnosyltransferases (WbpZ, WbpY, and WbpX) participate in the synthesis of the linear A-band O-antigen. Specific functions or detailed acceptor specificities of WbpZ, WbpY, and WbpX have not been experimentally verified. However, Rocchetta and co-workers (1998)
proposed that in the synthesis of repeating D-rhamnan structure (
3D-Rha
1
2D-Rha
1
3D-Rha
1
)n. WbpZ may be responsible for transferring the first D-rhamnose moiety to the acceptor molecule via an
1,3 linkage. The initial A-band O-antigen acceptor molecule is probably L-rhamnose, which is also the initial B-band O-antigen acceptor molecule (Rahim et al., 2000
). WbpY may be responsible for transferring the following two
1,3-linked D-rhamnose moieties to the first D-rhamnose. WbpX may subsequently add one D-rhamnose moiety to the A-band polymer via an
1,2-linkage. Both WbpY and WbpX would then continue to synthesize the A-band O-antigen. This hypothesis is consistent with the genetic organization of the wbpX, wbpY, and wbpZ genes, which are located in the opposite order to which the encoded enzymes act. A similar genetic arrangement coupled to the sequential order of the O-antigen assembly has been seen for other glycosyltransferases (Kido et al., 1995
).
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Talosylated glycans |
---|
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---|
6-Deoxy-D-talose is an enantiomer of 6-deoxy-L-talose. GDP-6-deoxy-D-talose is an activated nucleotide sugar form of 6-deoxy-D-talose that has been found only in EPS or LPS structures of a limited number of Gram-negative bacteria, such as A. actinomycetemcomitans serotype a and Burkholderia (Pseudomonas) plantarii strain DSM 6535 (Shibuya et al., 1991; Weckesser et al., 1973
; Zahringer et al., 1997
).
6-Deoxy-D-talose in A. actinomycetemcomitans LPS
An O-acetylated, linear homopolysaccharide of 6-deoxy-D-talose has been found in the serotype aspecific polysaccharide antigen (SPA) of A. actinomycetemcomitans (Perry et al., 1996; Shibuya et al., 1991
) and the EPS of the B. plantarii strain DSM 6535 (Zahringer et al., 1997
). The 6-deoxy-D-talan polymer of A. actinomycetemcomitans serotype a is composed of repeating disaccharide units (
3 6dTal
1
2 6dTal
1
)n, whereas 6-deoxy-D-talan of B. plantarii strain DSM 6535 is composed of repeating trisaccharide units (
3 6dTal
1
2 6dTal
1
2 6dTal
1
)n. Both 6-deoxy-D-talan polymers are acetylated at the O2 position of
1
3-linked 6-deoxy-D-talose. The specific roles of 6-deoxy-D-talan O-polysaccharide (O-PS) of human opportunistic pathogen A. actinomycetemcomitans serotype a and 6-deoxy-D-talan EPS of plant pathogen B. plantarii strain DSM 6535 are unknown.
In addition to 6-deoxy-D-talose and 6-deoxy-L-talose, other deoxyhexoses such as L-rhamnose and D-fucose are found in different SPAs of A. actinomycetemcomitans (Amano et al., 1989; Perry et al., 1996
). A. actinomycetemcomitans is a coccobacillus that colonizes the human oral cavity and has been implicated in the etiology of localized juvenile periodontitis, adult periodontitis, and severe nonoral infections (reviewed in Henderson et al., 2002
). The serologic specificity of A. actinomycetemcomitans strains is defined by six structurally and antigenically distinct O-PS components of their respective LPS (Gmur et al., 1993
; Kaplan et al., 2001
; Saarela et al., 1992
). These serotype-specific polysaccharides (af) constitute the outermost surface of the bacterium and have thus been suggested to play a role in the virulence of A. actinomycetemcomitans (reviewed in Fives-Taylor et al., 2000
). 6-Deoxy-D-talan-containing A. actinomycetemcomitans serotype a strain has been isolated from healthy subjects as well as patients with localized juvenile or adult perodontitis (Zambon et al., 1983
).
GDP-6-deoxy-D-talose metabolism
Markovitz proposed the biosynthetic pathway of GDP-6-deoxy-D-talose in 1960s, although the corresponding enzymes were not specifically identified (Markovitz, 1964). Like the other two GDP-6-deoxyhexose pathways introduced, the GDP-6-deoxy-D-talose pathway starts from GDP-D-mannose, which is converted into GDP-4-keto-6-deoxy-D-mannose by GMD (Figure 4). The following step in the GDP-6-deoxy-D-talose pathway is the specific reduction of the 4-keto group of GDP-4-keto-6-deoxy-D-mannose, which leads to the formation of GDP-6-deoxy-D-talose. 6-Deoxy-D-talose is an epimer of D-rhamnose and differs from it only in the orientation of the hydroxyl group at the C4 position (Figure 4). The stereoselectivity of GDP-4-keto-6-deoxy-D-mannose reductase determines which one of these two epimers is synthesized. Recently, the GDP-4-keto-6-deoxy-D-mannose reductase responsible for the synthesis of GDP-6-deoxy-D-talose has been identified and characterized from A. actinomycetemcomitans serotype a (Mäki et al., 2003
; Suzuki et al., 2002
). Other enzymes related to talosylation, for example, deoxytalosyltransferases, as well as the regulatory mechanisms of talosylation remain to be elucidated.
GDP-6-deoxy-D-talose (GTS)
The gene cluster associated with the biosynthesis of 6-deoxy-D-talan SPA of A. actinomycetemcomitans SUNYaB 75 contains 14 open reading frames, 9 of which were not found in SPA-associated gene clusters of the other serotypes (Kaplan et al., 2001; Suzuki et al., 2000
). The gts gene (also known as tld) has been identified from the serotype aSPA-associated gene cluster. The characterized A. actinomycetemcomitans GTS is acting only as a GDP-4-keto-6-deoxy-D-mannose reductase in the synthesis of GDP-6-deoxy-D-talose. The molecular mass of the monomer of GTS is
34 kDa, and the possible multimerization of the enzyme is unknown. The amino acid sequence of GTS contains the most typical motifs of the SDR protein family (Jörnvall et al., 1995
): a N-terminal coenzyme-binding pattern ThrGlyXXGlyXXGly and the conserved triad of Ser-Tyr-Lys (Mäki et al., 2003
; Suzuki et al., 2000
).
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Conclusions |
---|
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---|
L-fucose is the only representative of these deoxyhexoses also found in eukaryotes. Nevertheless, the biosynthetic de novo pathway of GDP-L-fucose could also be considered as a potential therapeutic target, because humans as well as other eukaryotes have an alternative pathway for GDP-L-fucose biosynthesis. This pathway is a result of the salvage metabolism, which uses fucose derived from lysosomal degradation or nutrition (reviewed in Becker and Lowe, 2003). Smith and colleagues (2002)
have shown that the de novo pathway knockout mice grew normally when fed with fucose-supplemented food.
The prevalence of some of these nucleotide sugars is not high, but narrow-spectrum drugs may be of interest in certain disease states, such as chronic infections that require long-term treatment. Furthermore, narrow-spectrum agents would also minimize the spread of drug resistance to other pathogens. It is also worth mentioning that in addition to the GDP-activated L-fucose, D-rhamnose, and 6-deoxy-D-talose pathways, the inhibition of GMD activity would also influence the biosynthesis of GDP-D-perosamine and GDP-colitose. D-perosamine is a constituent of glycoconjugates in several Gram-positive and Gram-negative bacteria, for example, E. coli O:157 and V. cholerae O:1 (Bilge et al., 1996; Villeneuve et al., 2000
). Colitose is a constituent of LPS in Gram-negative bacteria, for example, Salmonella enterica O:35 and E. coli O:111 (Wang and Reeves, 2000
). Humans lack metabolism for both of these nucleotide sugars. Furthermore, in addition to the dTDP-activated D-fucose, L-rhamnose, and 6-deoxy-L-talose pathways, the inhibition of RmlB activity would also influence the biosynthetic pathways of numerous dTDP-activated deoxyhexoses used in the biosynthesis of the secondary metabolites, mainly polyketide antibiotics, in bacteria (reviewed in Amann et al., 2001
; Thorson et al., 2001
).
The preparative synthesis of nucleotide sugars might aid research groups studying the relevance of glycoforms of various bacteria. These groups might benefit from the availability of building blocks required for the synthesis of glycosylated molecules. Before these molecules can be synthesized in vitro, the activated nucleotide sugars and the corresponding glycosyltransferases catalyzing the formation of specific glycosidic linkages are needed. The lack of effective tools for the synthesis and analysis of glycans has thus far been a bottleneck for the progress in glycobiology research. However there are several enzymatic synthesis strategies for the economic and effective preparative synthesis of nucleotide sugars using some bacteria and yeast strains as an expression host for the nucleotide sugar or glycosyltransferase overexpression (reviewed in Albermann et al., 2000; Amann et al., 2001
; Mäki et al., 2002
; Stein et al., 1998
). Furthermore an automated enzyme-assisted chemical transformation in conjunction with combinatorial organic synthesis has now become a tool that can yield sufficient amounts of glycans for the studies of their biological roles in various events, such as cellcell interactions, or the other applications of glycobiology, such as carbohydrate microarray (reviewed in Feizi et al., 2003
; Koeller and Wong, 2001
; Marcaurelle and Seeberger, 2002
; Plante et al., 2001
; Sears and Wong, 2001
). Preparative synthesis of nucleotide sugars might also contribute to the detailed characterization of known or novel glycosyltransferases or the synthesis of glycosylated antibacterial therapeutics or other pharmaceutically important glycodrugs, such as antitumor antibiotics.
Another troublesome point along these multistep glycosylation reactions is the consumption of very expensive nucleotide sugars, such as GDP-fucose, GDP-rhamnose, and GDP-6-deoxy-talose. Nahalka and co-workers (2003) have proposed an interesting alternative to overcome this problem. They immobilize the relevant enzymes on the superbeads to (re)generate nucleotide sugars and to synthesize oligosaccharides. These kinds of innovations could pave the way toward lower-cost synthesis of glycans in the future.
Understanding the mechanism of biosynthesis of bacterial nucleotide sugars should aid in the rational design of drugs with potential in antibacterial chemotherapy or engineering the biosynthetic pathways for the prepative synthesis of nucleotide sugars. Hence the detailed studies of the discrete catalytic mechanisms of the reactions and the specific regulation sites of nucleotide sugars synthesizing enzymes are needed. Analysis of these enzymes would also contribute to the understanding of catalytic mechanism of other dehydratases, epimerases, or reductases belonging to the SDR protein family.
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Footnotes |
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Abbreviations |
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References |
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