Dissection of the Bifunctional Escherichia coli N-Acetylglucosamine-1-phosphate Uridyltransferase Enzyme into Autonomously Functional Domains and Evidence That Trimerization Is Absolutely Required for Glucosamine-1-phosphate Acetyltransferase Activity and Cell Growth*

Frédérique PompeoDagger , Yves Bourne§, Jean van HeijenoortDagger , Florence Fassy, and Dominique Mengin-LecreulxDagger ||

From the Dagger  Laboratoire des Enveloppes Bactériennes et Antibiotiques, UMR 8619, CNRS, Université Paris-Sud, Bâtiment 430, 91405 Orsay Cedex, § Architecture et Fonction des Macromolécules Biologiques (AFMB)-CNRS, 13 chemin Joseph Aiguier, 13402 Marseille Cedex 20, and  Aventis Pharma-Hoechst Marion Roussel, 102 route de Noisy 93230, Romainville, France

Received for publication, June 2, 2000, and in revised form, October 26, 2000



    ABSTRACT
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The bifunctional N-acetylglucosamine-1-phosphate uridyltransferase (GlmU) enzyme catalyzes both the acetylation of glucosamine 1-phosphate and the uridylation of N-acetylglucosamine 1-phosphate, two subsequent steps in the pathway for UDP-N-acetylglucosamine synthesis in bacteria. In our previous work describing its initial characterization in Escherichia coli, we proposed that the 456-amino acid (50.1 kDa) protein might possess separate uridyltransferase (N-terminal) and acetyltransferase (C-terminal) domains. In the present study, we confirm this hypothesis by expression of the two independently folding and functional domains. A fragment containing the N-terminal 331 amino acids (Tr331, 37.1 kDa) has uridyltransferase activity only, with steady-state kinetic parameters similar to the full-length protein. Further deletion of 80 amino acid residues at the C terminus results in a 250-amino acid fragment (28.6 kDa) still exhibiting significant uridyltransferase activity. Conversely, a fragment containing the 233 C-terminal amino acids (24.7 kDa) exhibits acetyltransferase activity exclusively. None of these individual domains could complement a chromosomal glmU mutation, indicating that each of the two activities is essential for cell viability. Analysis of truncated GlmU proteins by gel filtration further localizes regions of the protein involved in its trimeric organization. Interestingly, overproduction of the truncated Tr331 protein in a wild-type strain results in a rapid depletion of endogenous acetyltransferase activity, an arrest of peptidoglycan synthesis and cell lysis. It is shown that the acetyltransferase activity of the full-length protein is abolished once trapped within heterotrimers formed in presence of the truncated protein, suggesting that this enzyme activity absolutely requires a trimeric organization and that the catalytic site involves regions of contact between adjacent monomers. Data are discussed in connection with the recently obtained crystal structure of the truncated Tr331 protein.



    INTRODUCTION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

UDP-N-acetylglucosamine (UDP-GlcNAc), the nucleotide-activated form of N-acetylglucosamine, plays a very important role in the biochemistry of all living organisms. In bacteria, it is required for the biosynthesis of essential cell-envelope components, namely peptidoglycan (1), lipopolysaccharides (2, 3), and teichoic acids (4), and for the formation of the enterobacterial common antigen (5).

Conditional-lethal mutants of Escherichia coli altered in the biosynthesis of this essential precursor are characterized by a cell-lysis phenotype (6-10). The four-step formation of UDP-GlcNAc from fructose-6-P has been now completely elucidated in this bacterial species (6, 7, 9, 11-13). It involves the successive actions of GlcN-6-P synthase, phosphoglucosamine mutase, GlcN-1-P acetyltransferase, and GlcNAc-1-P uridyltransferase (GlmU1; also named UDP-GlcNAc pyrophosphorylase). We showed earlier that the two latter activities were carried by a single 456-amino acid protein, the product of a gene we named glmU located just upstream from the GlcN-6-P synthase glmS gene at 84 min on the E. coli chromosome (7, 14). The glmU gene has been identified in some other bacterial species, in particular Neisseria gonorrhoeae (15) and Bacillus subtilis (16).

The bifunctional E. coli GlmU enzyme has been purified to homogeneity, and its kinetic parameters were determined (7, 13, 17). A complete loss of acetyltransferase activity was observed following incubation of the enzyme in the absence of reducing agent or treatment with thiol-specific reagents. Site-directed mutagenesis experiments further demonstrated the important role of two of the four cysteines of GlmU, namely residues Cys307 and Cys324, for acetyltransferase activity (17). The GlmU protein has been shown to exhibit a number of characteristics, which suggested that the acetyltransferase and uridyltransferase activities may reside in separate catalytic domains: (i) the substrates, products, and effectors of the acetyltransferase reaction did not inhibit the uridyltransferase activity and vice versa (13, 18); (ii) the intermediate GlcNAc-1-P was clearly released from the acetyltransferase domain prior to transformation by the uridyltransferase domain (18); (iii) portions of the GlmU amino acid sequence showing similarities with that of other previously characterized XDP-sugar pyrophosphorylase and acetyltransferase activities were located in the N-terminal portion and the second third of the protein, respectively (13, 15, 19); (iv) the acetyltransferase but not the uridyltransferase was shown to be inactivated by thiol-specific reagents, and the two cysteine residues whose alteration resulted in dramatic decreases of acetyltransferase activity were identified in the second moiety of the protein sequence (17); (v) mutagenesis of residues that are important for uridyltransferase activity did not affect acetyltransferase at all (Ref. 20 and data not shown); (vi) the fusion of a His6 tag at the C terminus of the protein resulted in a 20-fold decrease of acetyltransferase activity, without change in its uridyltransferase activity (17).

The crystal structure of a truncated form of the GlmU enzyme, GlmU-Tr331, was recently resolved at 2.25- and 2.3-Å resolution in the absence or presence of UDP-GlcNAc, respectively (20). The crystal structure is composed of two distinct domains connected by a long alpha -helical arm: a N-terminal domain resembling the dinucleotide-binding Rossmann fold and a C-terminal domain adopting a left-handed parallel beta -helix structure (Lbeta H) that is also found in homologous bacterial acyl- and acetyltransferases (21-24).

We here report the construction of truncated forms of this enzyme, which confirms that the bifunctional enzyme is composed of two autonomously folding and functional domains of roughly equivalent sizes, the N-terminal one exhibiting uridyltransferase activity and the C-terminal one acetyltransferase activity. It is also shown that trimer organization is essential for expression of the acetyltransferase activity and that the catalytic site of the latter should be formed by complementary regions from adjacent monomers.


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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
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Bacterial Strains and Growth Conditions-- E. coli strains JM83 (ara Delta [lac-proAB] rpsL thi Phi 80 dlacZ Delta M15) (25) and DH5alpha (supE44 Delta lacU169 hsdR17 recA1 endA1 gyrA96 thi-1 relA1 Phi 80 dlacZ Delta M15) (Life Technologies, Inc.) were used as hosts for plasmids and for the overproduction of wild-type and mutant GlmU enzymes. Strain UGS83 (JM83 glmU::kan [pGMU]), which carries an inactivated copy of the glmU gene on the chromosome and a wild-type copy of glmU on a plasmid whose replication is thermosensitive, was used in complementation experiments (7). The plasmid vector pTrcHis30 for expression of proteins under a N-terminal histidine-tagged form has been described previously (17). 2YT (26) was used as culture medium, and growth was monitored at 600 nm with a Shimadzu UV-1601 spectrophotometer. For strains carrying drug resistance genes, antibiotics were used at the concentrations of 100 (ampicillin), 35 (kanamycin), and 25 (chloramphenicol) µg·ml-1.

Construction of Expression Plasmids-- Standard procedures for molecular cloning (27) and E. coli cell transformation (28) were used. The pFP1 and pFP3 plasmids, allowing expression of the wild-type and N-terminal His6-tagged forms of GlmU, respectively (under control of the strong trc promoter), have been described previously (17). DNA fragments encoding C-terminally truncated enzymes (i.e. GlmU-Tr227, -Tr250, and -Tr331) were generated by PCR using the following oligonucleotides as primers: 5'-GGACGGGATCCTTGAATAATGCTATGAGCGTAGTGA-3' as a sense primer, 5'-GCAGCTGCAGTCACACGCCTTCTACTTCGCT-3' for Tr227, 5'-GCATCTGCAGTCATAACAGCAGTTTTTCAGCCTG-3' for Tr250, and 5'-CTCAGCTGCAGGACGCAATCAGGCAAACGG-3' for Tr331 as antisense primers, respectively. The resulting PCR products were purified, digested with BamHI and PstI (in bold), and inserted into the pTrcHis30 vector. DNA fragments encoding N-terminally truncated enzymes (i.e. GlmU-del78, -del130, -del227, -del233, and -del250) were similarly generated by PCR using the following primers: 5'-AGCCCTGCAGAATCACTTTTTCTTTACCGG-3' as an antisense primer, and 5'-GTGCTTGGATCCGAGCAGCTGGGTACGGGT-3' for del78, 5'-ATTGGTGGATCCACGGTGAAACTGGATGATCCG-3' for del130, 5'-GTAGAAGGATCCAATAACCGCCTGCAACTC-3' for del227, 5'-CGCCTGGGATCCTCCCGTCTGGAGCGTGTT-3' for del233, and 5'-AAACTGGGATCCGCAGGCGTTATGCTGCGC-3' for del250 as sense primers, respectively. PCR products were purified, digested with BamHI and PstI (in bold), and inserted into the pTrcHis30 vector. The pFP1kan plasmid was constructed by inserting the 1.28-kilobase pair HincII kanamycin resistance cartridge from pUC4K (Amersham Pharmacia Biotech) at the ScaI site lying within the ampicillin resistance gene from pFP1.

Preparation of Crude Extracts and Enzyme Purification-- E. coli cells carrying plasmids described in this work were grown at 37 °C in 2YT-ampicillin medium (0.5-liter cultures). When the optical density (OD) of the culture reached 0.4, IPTG was added at a final concentration of 1 mM and growth was continued for 3 h. Cells were harvested and washed with 40 ml of cold 20 mM potassium phosphate buffer, pH 7.4, containing 0.5 mM MgCl2 and 0.1% beta -mercaptoethanol. The cell pellet was suspended in 5 ml of the same buffer supplemented with a mixture of protease inhibitors: 1 mM leupeptin, 1 mM benzamidine, 1 mM phenylmethylsulfonyl fluoride, and 20 µg·ml-1 trypsin inhibitor. Cells were disrupted by sonication in the cold and the resulting suspension was centrifuged at 4 °C for 30 min at 200,000 × g. The supernatant was dialyzed against 100 volumes of the same buffer. SDS-PAGE analysis of proteins were performed as described previously (29). The different His6-tagged enzymes were purified as reported recently (17), basically following the steps in the manufacturer's (Qiagen) recommendations: binding of His6-GlmU on Ni2+-nitrilotriacetate-agarose (Ni2+-NTA) and extensive washing with 20 mM potassium phosphate buffer, pH 7.4, containing 0.1% beta -mercaptoethanol, 0.5 mM MgCl2, 300 mM NaCl, and 20-100 mM imidazole to remove impurities; elution of His6-GlmU with imidazole (100-300 mM) added to washing buffer; dialysis of His6-GlmU eluate against 100 volumes of the same phosphate buffer supplemented with 10% glycerol. The His6-tagged GlmU enzymes prepared in this manner were all at least 90% pure, as estimated by SDS-PAGE. Protein concentrations were determined by the method of Bradford, with bovine serum albumin as a standard (30).

Extraction and Quantitation of Peptidoglycan Precursors-- Cells of JM83(pFP3-Tr331) were grown at 37 °C in 2YT medium (1-liter cultures). At OD = 0.1 (~6 × 107 cells·ml-1), IPTG was added to one culture at a final concentration of 1 mM. As soon as the first effects on cell growth were observed in induced cells (~2 h later, final OD = 0.7), cultures were stopped by rapid chilling to 4 °C, and cells were harvested in the cold. Cultures of JM83 cells carrying the pTrcHis30 vector were made in parallel as a control. The extraction of peptidoglycan nucleotide precursors as well as the analytical procedure used for their quantitation were as described previously (31). UDP-GlcN was purified from cell extracts by first using the same two-step chromatographic procedure that is commonly used to purify the peptidoglycan nucleotide precursors: a gel filtration on Sephadex G-25 followed by HPLC on a column of µBondapak C18 (7.8 × 300 mm), where it is eluted in mixture with UDP-GlcNAc (31). The separation of UDP-GlcN and UDP-GlcNAc was then achieved by a second step of HPLC on the same column, using this time an elution with 50 mM triethylammonium formate, pH 4.75, at a flow rate of 3 ml·min-1 (their retention times were 9 and 21 min, respectively).

Isolation of Sacculi and Quantitation of Peptidoglycan-- Cells of JM83(pFP3-Tr331) and JM83(pTrcHis30) were grown and induced with IPTG as described above (0.5-liter cultures). Harvested cells were washed with a cold 0.85% NaCl solution and centrifuged again. Bacteria were then rapidly suspended under vigorous stirring in a hot (95-100 °C) aqueous 4% sodium dodecyl sulfate (SDS) solution (20 ml) for 30 min. After standing overnight at room temperature, the suspensions were centrifuged for 30 min at 200,000 × g in a Beckman TL100 centrifuge and the pellets were washed several times with water. Final suspensions were made in 2 ml of water, and aliquots (100 µl) were hydrolyzed and analyzed with a Biotronik model LC2000 amino acid analyzer. The peptidoglycan content of the sacculi was expressed in terms of its muramic acid content (31, 32).

Enzymatic Assays-- Assays for both activities of GlmU were performed as described previously (17), after appropriate dilutions of the enzyme in 20 mM potassium phosphate buffer, pH 7.4, containing 1 mg·ml-1 BSA, 0.5 mM MgCl2, and 0.1% beta -mercaptoethanol. One unit of enzyme activity was defined as the amount that catalyzed the formation of 1 µmol of product/min. UDP-GlcN, which is used as an alternative substrate of GlmU in some acetyltransferase assays, was synthesized enzymatically by using UDP-glucose pyrophosphorylase, as described by Gehring et al. (18). This compound was purified by HPLC as described above, and its authenticity was confirmed by determination of its hexosamine content after acid hydrolysis.

Estimation of Molecular Weight-- Pure samples of the different truncated GlmU proteins (50 µl of 1 mg·ml-1 solutions) were applied onto a column of Superdex 200 HR 10/30 connected to a fast pressure liquid chromatography apparatus (Amersham Pharmacia Biotech). The column was equilibrated with 50 mM potassium phosphate buffer, pH 7.4, containing 150 mM NaCl and 0.1% beta -mercaptoethanol, and samples were eluted at 0.5 ml·min-1. Calibration was carried out with cytochrome c, myoglobin, alpha -chymotrypsinogen A, ovalbumin, and BSA, and the void and total volumes were determined with blue dextran 2000 and tyrosine, respectively. Elution was followed by measurement of the absorbance at 280 nm.


    RESULTS
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
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Construction of Various Truncated Forms of GlmU and Their Activity-- As attempts to crystallize the full-length GlmU protein remained unsuccessful, crystallization of individual domains was therefore envisaged. This prompted us to more precisely define the size of the two putative autonomous domains. Plasmids for high-level overexpression of GlmU proteins truncated either in the N- (del constructs) or in the C-terminal (Tr constructs) region were constructed (Fig. 1). All of these proteins were expressed in a His6-tagged form (N-terminal Met-His6-Gly-Ser extension), allowing their convenient one-step purification. Most of them were successfully overproduced in a soluble form and were purified to near homogeneity (Fig. 2). Others (del26, del182, del250, and Tr227) either appeared very poorly produced, due probably to structural instability and rapid intracellular degradation, or formed insoluble inclusion bodies.



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Fig. 1.   Schematic structure of various truncations of the GlmU protein generated in the present study. Truncated proteins are represented by cross-hatched regions, and numbers indicate amino acid residue numbers within the wild-type protein sequence. All truncated proteins were expressed with a N-terminal His6 tag extension, as shown for the full-length protein.



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Fig. 2.   SDS-polyacrylamide gel electrophoresis of purified truncated forms of the E. coli GlmU protein. Full-length and truncated GlmU proteins were overproduced in E. coli cells in the His6-tagged form (N-terminal Met-His6-Gly-Ser extension). Their one-step purification on Ni2+-NTA was performed as described in the text, and aliquots (2 µg) were analyzed by SDS-PAGE. Lane A, full-length GlmU; lane B, del78; lane C, del130; lane D, Tr331; lane E, del227; lane F, del233; lane G, Tr250. The position and molecular mass (kilodaltons) of marker proteins are indicated on the left.

However, none of these different plasmids could complement the thermosensitive glmU mutant strain UGS83, indicating that the engineered truncations had resulted in the loss of at least one of the two activities of GlmU. This was confirmed by assays of the pure proteins for acetyl- and uridyltransferase activities (Table I). The uridyltransferase activity of proteins truncated in the N-terminal region (del78, del130, del227, and del233) was decreased by a factor of at least 1000, but surprisingly a residual and almost invariant activity (kcat = 0.1-0.3 s-1) was retained by all of them. As discussed below, this residual activity was due to contaminating wild-type GlmU enzyme (originating from chromosomal gene expression) that could form heterotrimers with these His6-tagged truncated proteins. The deletion of the first 130 N-terminal amino acid residues of GlmU resulted in only a 50% decrease of its acetyltransferase activity. Truncation of 100 more residues (del233) was accompanied by a more important decrease (98%) of activity, but the residual acetyltransferase activity remained relatively high with a kcat of 25 s-1. Gehring et al. (18) also previously constructed a truncated form of GlmU (glutathione S-transferase fusion), deleted in that case of the first 179 residues. Its acetyltransferase activity was 150-fold reduced as compared with full-length GlmU with a kcat of 0.5 s-1. As shown in Table I, an inverse pattern of activities was observed for proteins carrying deletions in the C-terminal region; Tr331 and Tr250 proteins lacking the last 125 and last 206 amino acid residues, respectively, had undetectable acetyltransferase activity but retained significant uridyltransferase activity (42% and 2.5% of wild-type enzyme activity, respectively).


                              
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Table I
Enzymatic activities and oligomerization state of truncated GlmU proteins
Full-length and truncated GlmU proteins were overproduced in E. coli cells in the His6-tagged form (N-terminal Met-His6-Gly-Ser extension). They were purified on Ni2+-NTA and assayed for both enzyme activities of GlmU. The values represent the means of three determinations. Their oligomerization state was determined by gel filtration as described under "Experimental Procedures." Yields of these different truncated proteins were similar, from 2.5 to 10 mg of protein per 0.5 liter of culture.

Trimer Organization-- As reported previously, chromatography of full-length GlmU protein on gel filtration was consistent with a homotrimer arrangement (13). The oligomerization state of the different truncated GlmU fragments generated here was then investigated (Table I). The ability of the proteins to trimerize was clearly correlated to the presence of at least the initial part of the C-terminal domain. The fact that the two del227 and Tr331 proteins were trimers suggested that residues involved in the oligomerization process might be located between these two sites of truncation. With the exception of the Tr250 protein, which turned out to be a monomer, all other truncated proteins generated in the present work consisted of trimers.

As shown above, a very low but detectable uridyltransferase activity of 0.1-0.3 s-1 was consistently detected with all preparations of proteins carrying either partial or complete deletions of the N-terminal domain (del78 to del233). It should be noted that these proteins, which carry only one of the two activities of GlmU, could not complement the glmU mutant strain UGS83 and have consequently been expressed in a wild-type E. coli strain. The simplest interpretation was the presence in these preparations of contaminating full-length (non-His-tagged) GlmU enzyme originating from chromosomal expression. That the wild-type enzyme could not be eliminated by extensive washings of the His6-tagged truncated proteins adsorbed on Ni2+-NTA with buffers containing 300 mM NaCl and 20-100 mM imidazole suggested a very tight association (formation of heterotrimers) between the full-length and the truncated proteins. It was effectively controlled that the pure wild-type (non-His-tagged) GlmU protein had by itself no particular affinity for the Ni2+-NTA matrix, i.e. was detected essentially in the pass-through fraction and had been completely eliminated from the column by washing with 20 mM imidazole containing buffers, as confirmed by SDS-PAGE analysis and enzymatic assays (data not shown).

Toxic Effects of the Overproduction of GlmU-Tr331 on E. coli Cell Growth-- JM83 cells carrying the plasmid pFP3-Tr331 were grown in rich medium and expression of the truncated gene was induced with 1 mM IPTG. It resulted in a significant overproduction of the truncated protein, as observed by SDS-PAGE. Interestingly, after 2 h of induction, a rapid slow down of cell growth occurred which was followed by cell lysis about 1 h later (Fig. 3). Observation of cells by optical microscopy showed morphological changes characteristic of an arrest of peptidoglycan synthesis: progressive change of cell shape from rods to greatly enlarged ovoids and cell lysis, as indicated by the presence of many ghosts in the culture. It was reminiscent of the effects previously observed in a thermosensitive glmU mutant strain when grown under restrictive conditions. Cell lysis was clearly due to an arrest of peptidoglycan synthesis as the peptidoglycan content of induced JM83(pFP3-Tr331) cells appeared about 40% lower than that of noninduced cells (at the time where the first effects on cell growth were observed) (Table II). An analysis of the pool levels of peptidoglycan precursors revealed the significant reduction of the pools of UDP-GlcNAc and UDP-MurNAc-pentapeptide in induced cells (Table II), consistent with the inhibition of an early step leading to the formation of UDP-GlcNAc (6, 7). Interestingly, the significant accumulation of a compound which was subsequently identified as UDP-glucosamine was observed in these cells (see below). The morphology, peptidoglycan content, and levels of precursors in noninduced JM83(pFP3-Tr331) cells were similar to those of control JM83(pTrcHis30) cells (Table II).



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Fig. 3.   Effects of the overproduction of the truncated GlmU-Tr331 protein on E. coli cell growth. Cells of JM83(pFP3-Tr331) were grown at 37 °C in 2YT-ampicillin medium. At the time indicated by an arrow (optical density = 0.1), the overproduction of the Tr331 protein was induced with IPTG (1 mM) and growth of induced (open symbols) and not induced (filled symbols) cells was monitored at 600 nm. Growth of cells carrying as a control the plasmid vector pTrcHis30 was unaffected by IPTG and paralleled that of noninduced JM83(pFP3-Tr331) cells (data not shown).


                              
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Table II
Effects of the overproduction of the truncated GlmU-Tr331 protein on peptidoglycan synthesis and activities of the GlmU enzyme in wild-type E. coli cells
Cells were grown in 2YT-ampicillin medium at 37 °C. At OD = 0.1, IPTG (1 mM) was eventually added and incubation was continued until the first effects on cell growth were observed, about 2 h later (Fig. 3). At this time, peptidoglycan and its precursors were extracted and quantitated as described in the text. Crude protein extracts were also prepared and were assayed for GlcN-1-P acetyltransferase and GlcNAc-1-P uridyltransferase activities. The values are means of determinations obtained in two independent experiments.

The levels of the acetyltransferase and uridyltransferase activities of GlmU were determined in crude extracts from cells of JM83(pFP3-Tr331) grown in the absence or presence of IPTG. In the absence of IPTG, JM83(pFP3-Tr331) cells contained about 8-fold more uridyltransferase activity but 30-fold less acetyltransferase activity than control JM83(pTrcHis30) cells. The overproduction of uridyltransferase was due to basal expression from the plasmid promoter of the truncated form of GlmU (which has a normal uridyltransferase activity). The concomitant dramatic decrease of acetyltransferase activity clearly indicated that the activity of newly synthesized wild-type GlmU molecules originating from the chromosomal gene could only be partially detected, due probably to the formation of inactive heterotrimers. When IPTG was added, this effect was exacerbated as a 200-fold increase of uridyltransferase activity but a null acetyltransferase activity were measured in induced JM83(pFP3-Tr331) cells. The loss of acetyltransferase activity and the arrest of UDP-GlcNAc and consequently peptidoglycan biosynthesis were clearly correlated features. It should be noted that the effects on cell morphology and peptidoglycan metabolism were observed only in the presence of IPTG. The fact that the 30-fold decrease of acetyltransferase activity detected in noninduced cells had no apparent effect on cell growth was consistent with the previous demonstration that this activity of GlmU was in great excess in E. coli cells as compared with its specific requirements (13).

Formation of a mixture of four different heterotrimers Wt(3), Wt(2)-Tr(1), Wt(1)-Tr(2), and Tr(3) is expected to occur in vivo, whose proportions theoretically reflect relative abundance of wild-type (Wt) and truncated (Tr) monomers. In conditions where the truncated monomer is largely predominant (at least a 200: 1 ratio could be estimated in IPTG-induced cells from the increase of uridyltransferase activity), Wt(3) and Wt(2)-Tr(1) species are most probably very rarely generated, and wild-type monomers should be exclusively in the Wt(1)-Tr(2) form (model shown in Fig. 4). The fact that the acetyltransferase activity of the wild-type enzyme was completely undetectable in these conditions therefore suggested that more than one full-length monomer per trimer was required for exhibition of the latter activity. However, the question of the acetyltransferase activity of the two Wt(2)-Tr(1) and Wt(1)-Tr(2) species remained. To generate significant amounts of these two heterotrimers, JM83 cells were transformed by two plasmids, pFP1kan and pFP3-Tr331, for concomitant expression of the wild-type (Wt) and the His6-tagged truncated (Tr) proteins, respectively. Analysis of crude extracts prepared from these cells confirmed the large accumulation of the two proteins (data not shown). Purification on Ni2+-NTA was then performed as described above, using extensive washing steps for complete elimination of the non-His-tagged Wt(3) form. The resulting purified material appeared composed of both Wt and Tr proteins in roughly equivalent amounts, as judged by SDS-PAGE (data not shown). Unfortunately, attempts to separate the different His-tagged heterotrimers present in this mixture by gel filtration techniques failed, due to their very close molecular masses, 111, 123, and 135 kDa for Tr(3), Wt(1)-Tr(2), and Wt(2)-Tr(1), respectively. However, this mixture, which theoretically contains the two Wt(1)-Tr(2) and Wt(2)-Tr(1) heterotrimers, exhibited a high uridyltransferase activity (120 s-1) but had no detectable acetyltransferase activity, as observed for the pure Tr(3) homotrimer. This finding was thus consistent with the hypothesis (discussed below) that three full-length monomers should be present in the trimer for expression of the latter activity.



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Fig. 4.   Ribbon diagram of the GlmU-Tr331 trimer viewed perpendicular to the Lbeta H axis. The additional coils have been modeled (dark gray) only in one subunit and contain residues Pro328 to Gly424.

Overexpression of other truncated proteins described in the present work had no effects on cell growth. It should be noted that most of them exhibited significant acetyltransferase activity. In fact, the only other engineered protein with undetectable acetyltransferase activity, Tr250, appeared unable to oligomerize and consequently could not trap the full-length GlmU enzyme into inactive heterotrimers.

Interestingly, IPTG-induced JM83(pFP3-Tr331) cells were shown to accumulate large amounts of a compound, which was identified as UDP-GlcN (Table II). This unexpected finding was clearly correlated with the depletion of the acetyltransferase activity of GlmU in these cells. It suggested that in the absence of the latter enzyme activity GlcN-1-P molecules had been transformed into UDP-GlcN by the still present (and highly overproduced) uridyltransferase activity of GlmU. GlmU-catalyzed uridylyltransfer to glucosamine-1-P was previously reported to be undetectable (<0.0001 s-1) (18). In our hands, however, the pure GlmU enzyme could catalyze the synthesis of UDP-GlcN from GlcN-1-P and UTP with a very good efficiency (kcat = 23 s-1, as compared with 330 s-1 when GlcNAc-1-P is used as the substrate), a finding consistent with the in vivo accumulation of this compound.


    DISCUSSION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Data obtained in the present study confirm that the bifunctional GlmU protein is organized in two autonomously folding and functional domains. As judged by the specific activities exhibited by the various truncated forms of GlmU described here, the size of the two individual domains might be roughly equivalent, each one representing about half of the protein. In the light of the recently established crystal structure of a truncated form of GlmU (Tr331), the connection between the two domains is achieved by a long alpha -helical arm located between residues Asn228 and Ala250 (20). It suggests that this bacterial protein evolved by fusion of two uridyltransferase and acetyltransferase fragments. In eukaryotes, the biosynthesis of UDP-GlcNAc occurs by a slightly different route (via GlcNAc-6-P) in which GlcN-6-P acetyltransferase and GlcNAc-1-P uridyltransferase activities are carried by two distinct monofunctional enzymes (33-37). The selective advantage (if any) conferred to bacteria by this unique bifunctional enzyme remains an enigma. In particular, there is no apparent requirement for a common regulation of the component activities at the level of transcription or translation. It was previously shown that both activities were in great excess in cells as compared with specific requirements in UDP-GlcNAc molecules of the peptidoglycan and lipopolysaccharide pathways (7, 13). As demonstrated by Plumbridge et al. (38), the glmU gene is cotranscribed with the downstream GlcN-6-P synthase glmS gene in E. coli and seems to be expressed at a high constitutive level whatever the growth conditions used. The construction of various mutated forms of GlmU enzyme affected in either of the two activities also showed that the ratio of acetyltransferase and uridyltransferase activities (which is about 5 for wild-type enzyme) could be greatly modified in vivo without detectable effect on the functioning of this pathway. Additionally, there is no apparent advantage for GlmU to be a bifunctional protein in terms of reaction mechanism. By using radiolabeled substrates, Gehring et al. (18) demonstrated that GlcNAc-1-P was released by the enzyme before being used as substrate by the second enzyme activity. In addition, the thermosensitive glmU mutant strain UGS83 was shown to accumulate large amounts of GlcNAc-1-P when grown at the restrictive temperature (7). These results were a priori not consistent with the hypothesis of a concerted action of the two enzyme activities.

Gehring et al. (18) reported previously that the GlmU enzyme was unable to catalyze uridylyltransfer from UTP to GlcN-1-P but could catalyze acetyltransfer from acetyl-CoA to UDP-GlcN, although at a 12-fold reduced rate. It was one of the arguments why these authors conclude that acetyltransfer precedes uridylyltransfer in the two-step formation of UDP-GlcNAc by GlmU. We effectively confirmed here that GlmU could catalyze acetyltransfer to UDP-GlcN and the kcat value we determined was 170 s-1 (9-fold lower as compared with 1500 s-1 for GlcN-1-P). However, we here observed that GlmU could also efficiently catalyze uridylyltransfer to GlcN-1-P, at a 15-fold reduced rate (kcat = 23 s-1) as compared with that with GlcNAc-1-P (350 s-1). In our hands, the GlmU enzyme therefore appears theoretically capable to catalyze a two-step synthesis of UDP-GlcNAc in which uridylyltransfer precedes acetyltransfer but the greatly reduced kinetic parameters confirmed the previous assumption that these two reactions occur in the inverse order under normal physiological conditions (13, 18). As shown in the present report, a significant accumulation of UDP-GlcN was observed in cells in which the acetyltransferase activity of GlmU has been inhibited. This finding confirmed that GlmU could effectively catalyze the uridylyltransfer from UTP to GlcN-1-P in vivo, but it is clear that this only occurred because of very particular physiological conditions in which the availability of the preferred substrate GlcNAc-1-P was impaired.

GlmU and its truncated derivative Tr331 are trimeric proteins, as judged by their behavior on gel filtration. The recently obtained crystal structure of GlmU-Tr331 also showed a trimeric arrangement around the long dimension of the Lbeta H prism (20). The trimeric association of the particular Lbeta H domain is highly conserved between GlmU-Tr331 and other previously characterized bacterial acetyl- or acyltransferases, namely LpxA, PaXAT, DapD, and Cam (20-24). However, it is not known whether a trimeric organization of these proteins is absolutely required for expression of their acetyl- or acyltransferase activities. This also raised the question of the activity of the various heterotrimers that could be generated from a mixture of wild-type and truncated monomers. To answer this question, we followed the growth of a wild-type E. coli strain during the overexpression of GlmU-Tr331, a truncated protein with null acetyltransferase activity but still able to associate in trimers. Its overproduction by a factor of about 200-fold as compared with the wild-type protein level resulted in the complete disappearance of acetyltransferase activity in cells, which was followed by deleterious effects on peptidoglycan biosynthesis and cell growth. This finding strongly suggested that the wild-type enzyme could not exhibit acetyltransferase activity under a monomer form in vivo. It also indicated that the activity of this wild-type protein originating from chromosomal expression could no more be detected once trapped within heterotrimers formed in presence of the predominantly expressed truncated protein (model shown in Fig. 4). An organization in trimer is therefore required for exhibition of the acetyltransferase activity but the data here obtained could be interpreted in several ways. (i) Each monomer carries a complete catalytic site, the active conformation of which is formed only during trimer assembly; (ii) each catalytic site is made of specific complementary regions belonging to more than one monomer, suggesting that one to three catalytic site(s) could exist per trimer unit. To date, only the crystal structure of the truncated form of GlmU has been determined (20) and the exact position and number of binding-sites of the substrate acetyl-CoA are not known. It should be noted that, in the homologous structures of acetyltransferases DapD and PaXAT (21, 22), three binding sites for the substrate acetyl-CoA were detected, each one being located between two subunits on the exterior face of the trimeric Lbeta H domains. This positioning could also be adopted for GlmU. The finding that a mixture of Tr(3), Wt(1)-Tr(2), and Wt(2)-Tr(1) heterotrimers has undetectable acetyltransferase activity further suggests that the catalytic site(s) of the active trimer Wt(3) should be formed by adjacent and complementary regions from three full-length monomers. The confirmation of this hypothesis now requires the elucidation of the three-dimensional structure of the entire GlmU protein. Trimerization is clearly not essential for expression of the uridyltransferase activity of GlmU. The latter activity was retained by the Tr250 protein, which is unable to associate in trimers. However, the 40-fold reduced uridyltransferase activity of this protein suggests that trimerization or at least interactions between regions of the two domains may participate in the folding and stability of the N-terminal domain. In the crystal structure, the only observed contacts between the two domains were van der Waals interactions between the surface loop Ala31-Gly32 in the N-terminal domain and the Arg263 side chain in the C-terminal domain (20). Residues within the long alpha -helical arm (Asn228-Ala250) also established numerous interactions with residues in the two domains (20). The abolishment of at least the van der Waals interactions in the Tr250 protein could partially account for the decreased activity of this protein. However, the positioning of the long alpha -helical arm seems to play an important role for the uridyltransferase activity with the helix dipole aligned with the position of the key positively charged Arg18 residue. Whether an incorrect positioning of the alpha -helical linker in the Tr250 protein could explain for its decreased activity remains to be elucidated.

As mentioned above, the pathway for UDP-GlcNAc biosynthesis appears significantly different in eukaryotes. In the latter, acetyltransfer occurs on GlcN-6-P and not on GlcN-1-P, and, most importantly, acetyltransferase and uridyltransferase activities are carried by two distinct monofunctional enzymes that show little sequence homology with GlmU (35-37). The GlmU enzyme, which is essential and specific of the bacterial world, should therefore be considered as an interesting target for the search of new antibiotics. Biochemical and crystallographic investigations are now developed to gain more information on active sites of this bifunctional enzyme. The present demonstration that a trimer organization is absolutely required for acetyltransferase activity of GlmU could also open the way for a search of inhibitors based on the inhibition of the oligomerization process.


    FOOTNOTES

* This work was supported by CNRS Grant EP1088, Grant 97.C.0177 "Biotechnologies" from the Ministère de l'Education Nationale de la Recherche et de la Technologie, and by a grant-in-aid from Hoechst Marion Roussel (to F. P.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

|| To whom correspondence should be addressed. Tel.: 33-1-69-15-61-34; Fax: 33-1-69-85-37-15; E-mail: dominique.mengin-lecreulx@ebp.u-psud.fr.

Published, JBC Papers in Press, November 17, 2000, DOI 10.1074/jbc.M004788200


    ABBREVIATIONS

The abbreviations used are: GlmU, N-acetylglucosamine-1-phosphate uridyltransferase; IPTG, isopropyl-1-thio-beta -D-galactopyranoside; del, deletion; Tr, truncated; Wt, wild-type; OD, optical density; HPLC, high performance liquid chromatography; NTA, nitrilotriacetic acid; PAGE, polyacrylamide gel electrophoresis.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES


1. van Heijenoort, J. (1996) in Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (Neidhardt, F. C. , Curtis, R., III , Ingraham, J. L. , Lin, E. C. C. , Low, K. B. , Magasanik, B. , Reznikoff, W. S. , Riley, M. , Schaechter, M. , and Umbarger, H. E., eds) , pp. 1025-1034, American Society for Microbiology, Washington, D. C.
2. Raetz, C. R. H. (1996) in Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (Neidhardt, F. C. , Curtis, R., III , Ingraham, J. L. , Lin, E. C. C. , Low, K. B. , Magasanik, B. , Reznikoff, W. S. , Riley, M. , Schaechter, M. , and Umbarger, H. E., eds) , pp. 1035-1063, American Society for Microbiology, Washington, D. C.
3. Stevenson, G., Neal, B., Liu, D., Hobbs, M., Packer, N. H., Batley, M., Redmond, J. W., Lindquist, L., and Reeves, P. (1994) J. Bacteriol. 176, 4144-4156[Abstract]
4. Fisher, W. (1990) in Glycolipids, Phosphoglycolipids, and Sulfoglycolipids (Kates, M., ed) , pp. 123-234, Plenum Press, New York
5. Kuhn, H.-M., Meier-Dieter, U., and Mayer, H. (1988) FEMS Microbiol. Rev. 54, 195-222
6. Mengin-Lecreulx, D., and van Heijenoort, J. (1996) J. Biol. Chem. 271, 32-39[Abstract/Free Full Text]
7. Mengin-Lecreulx, D., and van Heijenoort, J. (1993) J. Bacteriol. 175, 6150-6157[Abstract]
8. Sarvas, M. (1971) J. Bacteriol. 105, 467-471[Medline] [Order article via Infotrieve]
9. White, R. J. (1968) Biochem. J. 106, 847-858[Medline] [Order article via Infotrieve]
10. Wu, H. C., and Wu, T. C. (1971) J. Bacteriol. 105, 455-466[Medline] [Order article via Infotrieve]
11. Dobrogosz, W. J. (1968) J. Bacteriol. 95, 578-584[Medline] [Order article via Infotrieve]
12. Dutka-Malen, S., Mazodier, P., and Badet, B. (1988) Biochimie 70, 287-290[CrossRef][Medline] [Order article via Infotrieve]
13. Mengin-Lecreulx, D., and van Heijenoort, J. (1994) J. Bacteriol. 176, 5788-5795[Abstract]
14. Walker, J. E., Gay, N. J., Saraste, M., and Eberle, A. N. (1984) Biochem. J. 224, 799-815[Medline] [Order article via Infotrieve]
15. Ullrich, J., and van Putten, J. P. M. (1995) J. Bacteriol. 177, 6902-6909[Abstract]
16. Hove-Jensen, B. (1992) J. Bacteriol. 174, 6852-6856[Abstract]
17. Pompeo, F., van Heijenoort, J., and Mengin-Lecreulx, D. (1998) J. Bacteriol. 180, 4799-4803[Abstract/Free Full Text]
18. Gehring, A. M., Lees, W. J., Mindiola, D. J., Walsh, C. T., and Brown, E. D. (1996) Biochemistry 35, 579-585[CrossRef][Medline] [Order article via Infotrieve]
19. Vaara, M. (1992) FEMS Microbiol. Lett. 97, 249-254[CrossRef]
20. Brown, K., Pompeo, F., Dixon, S., Mengin-Lecreulx, D., Cambillau, C., and Bourne, Y. (1999) EMBO J. 18, 4096-4107[Abstract/Free Full Text]
21. Beaman, T. W., Binder, D. A., Blanchard, J. S., and Roderick, S. L. (1997) Biochemistry 36, 489-494[CrossRef][Medline] [Order article via Infotrieve]
22. Beaman, T. W., Sugantino, M., and Roderick, S. L. (1998) Biochemistry 37, 6689-6696[CrossRef][Medline] [Order article via Infotrieve]
23. Kisker, C., Schindelin, H., Alber, B. E., Ferry, J. G., and Rees, D. C. (1996) EMBO J. 15, 2323-2330[Abstract]
24. Raetz, C. R. H., and Roderick, S. L. (1995) Science 270, 997-1000[Abstract]
25. Yanisch-Perron, C., Vieira, J., and Messing, J. (1985) Gene (Amst.) 33, 103-119[CrossRef][Medline] [Order article via Infotrieve]
26. Miller, J. H. (1972) Experiments in Molecular Genetics , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
27. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual , 2nd Ed. , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
28. Dagert, M., and Ehrlich, S. D. (1979) Gene (Amst.) 6, 23-28[CrossRef][Medline] [Order article via Infotrieve]
29. Laemmli, U. K., and Favre, M. (1973) J. Mol. Biol. 80, 575-599[Medline] [Order article via Infotrieve]
30. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
31. Mengin-Lecreulx, D., Flouret, B., and van Heijenoort, J. (1982) J. Bacteriol. 151, 1109-1117[Medline] [Order article via Infotrieve]
32. Mengin-Lecreulx, D., and van Heijenoort, J. (1985) J. Bacteriol. 163, 208-212[Medline] [Order article via Infotrieve]
33. Cabib, E., Roberts, R., and Bowers, B. (1982) Annu. Rev. Biochem. 51, 763-793[CrossRef][Medline] [Order article via Infotrieve]
34. Hofman, M., Boles, E., and Zimmermann, F. K. (1994) Eur. J. Biochem. 221, 741-747[Abstract]
35. Mio, T., Yabe, T., Arisawa, M., and Yamada-Okabe, H. (1998) J. Biol. Chem. 273, 14392-14397[Abstract/Free Full Text]
36. Mio, T., Yamada-Okabe, T., Arisawa, M., and Yamada-Okabe, H. (1999) J. Biol. Chem. 274, 424-429[Abstract/Free Full Text]
37. Wang-Gillam, A., Pastuszak, I., and Elbein, A. D. (1998) J. Biol. Chem. 273, 27055-27057[Abstract/Free Full Text]
38. Plumbridge, J. A., Cochet, O., Souza, J. M., Altamirano, M. M., Calcagno, M. L., and Badet, B. (1993) J. Bacteriol. 175, 4951-4956[Abstract]


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