Search tips
Search criteria 


Logo of jbcThe Journal of Biological Chemistry
J Biol Chem. 2011 August 19; 286(33): 28688–28696.
Published online 2011 June 24. doi:  10.1074/jbc.M111.239673
PMCID: PMC3190676

The LptD Chaperone LptE Is Not Directly Involved in Lipopolysaccharide Transport in Neisseria meningitidis*An external file that holds a picture, illustration, etc.
Object name is sbox.jpg


The biosynthesis of lipopolysaccharide (LPS) in Gram-negative bacteria is well understood, in contrast to the transport to its destination, the outer leaflet of the outer membrane. In Escherichia coli, synthesis and transport of LPS are essential processes. Neisseria meningitidis, conversely, can survive without LPS and tolerates inactivation of genes involved in LPS synthesis and transport. Here, we analyzed whether the LptA, LptB, LptC, LptE, LptF, and LptG proteins, recently implicated in LPS transport in E. coli, function similarly in N. meningitidis. None of the analyzed proteins was essential in N. meningitidis, consistent with their expected roles in LPS transport and additionally demonstrating that they are not required for an essential process such as phospholipid transport. As expected, the absence of most of the Lpt proteins resulted in a severe defect in LPS transport. However, the absence of LptE did not disturb transport of LPS to the cell surface. LptE was found to be associated with LptD, and its absence affected total levels of LptD, suggesting a chaperone-like role for LptE in LptD biogenesis. The absence of a direct role of LptE in LPS transport was substantiated by bioinformatic analyses showing a low conservation of LptE in LPS-producing bacteria. Apparently, the role of LptE in N. meningitidis deviates from that in E. coli, suggesting that the Lpt system does not function in a completely conserved manner in all Gram-negative bacteria.

Keywords: Bacteria, Lipopolysaccharide (LPS), Lipoprotein, Membrane, Membrane Biogenesis, Membrane Trafficking, Outer Membrane Biogenesis


Lipopolysaccharide (LPS) is a glycolipid, uniquely present at the cell surface of Gram-negative bacteria. It consists of a hydrophobic membrane anchor, called lipid A, which is embedded in the outer membrane (OM),2 and an extracellular oligosaccharide core extended in some bacteria with a polysaccharide moiety, the O-antigen. The lipid A part can evoke strong, often toxic, innate immune responses; hence, it is also known as endotoxin. LPS is an essential component of the OM of most Gram-negative bacteria. Therefore, molecules involved in its biogenesis may represent attractive antimicrobial targets, as demonstrated by the successful development of antibiotics targeting LpxC, an enzyme involved LPS biogenesis (1), and LptD, an OM protein involved in LPS transport (2). However, much of the LPS biogenesis is not understood yet. The complete lipid A biosynthetic pathway, which takes place in the cytoplasm and at the inner leaflet of the inner membrane, has been elucidated in Escherichia coli and Salmonella. The ubiquitous presence of the lipid A biosynthetic enzymes in most other Gram-negative organisms suggests that this process is well conserved (3). The transport process of LPS from its site of synthesis to the cell surface is much less well understood (47).

The ATP-binding cassette transporter MsbA is thought to transport the lipid A-core moiety of newly synthesized LPS over the inner membrane (8, 9). At the OM, two proteins have been shown to be involved in the transport of LPS to the cell surface: the integral OM protein designated LptD (formerly Imp/OstA) (10) and the lipoprotein LptE (formerly RlpB), which is associated with LptD (11). A soluble periplasmic protein, designated LptA, was also postulated to be involved in LPS transport, because an E. coli strain depleted for this protein demonstrated a similar phenotype as a strain depleted for LptD with LPS accumulating in aberrant membrane-like structures in the periplasm (1214). As LptA was shown to bind lipid A (15), it might act as a chaperone, assisting the amphipathic LPS molecules to pass through the aqueous periplasm. Furthermore, the inner membrane proteins LptB, LptC, LptF, and LptG, which form a complex (16), were also implicated in LPS transport because of their essential nature and because their depletion resulted in a phenotype similar to that of strains depleted for LptD or LptE (13, 17). Furthermore, LptC was shown to bind LPS (18). All seven Lpt proteins were found to interact (19), suggesting that they form a transenvelope complex, which is consistent with the observation that transport of LPS to the OM still proceeds in spheroplasted E. coli cells (20). If, indeed, they form such a complex, this complex might not only mediate the transport of LPS to the cell surface, but also the transport of phospholipids to the inner leaflet of the OM.

In E. coli, LPS is an essential component of the OM, because it is impossible to knock out the lpxA gene, which encodes the first enzyme involved in LPS biosynthesis (21). Also, all of the LPS transport-related proteins are essential in E. coli (11, 17, 22, 23). Surprisingly however, an lpxA mutant of Neisseria meningitidis turned out to be viable while completely devoid of LPS (24), and also the LptD and MsbA proteins are not essential in this organism (10, 25). In meningococcal lptD and msbA mutants, LPS levels are severely decreased likely by the action of a feedback mechanism that inhibits LPS synthesis when transport is hampered. Because clean knock-out mutants affected in LPS synthesis and transport can be generated, N. meningitidis is an ideal organism to study LPS biogenesis. We proved this notion previously by identifying, for the first time, the function of LptD in LPS transport (10) and by demonstrating that MsbA is not involved in phospholipid transport in N. meningitidis (25).

The discovery of novel putative LPS transport components in E. coli prompted us to test whether these components would be present and functioning in the same process also in N. meningitidis. The nomenclature for proteins involved in LPS transport in E. coli has recently been changed to Lpt (LPS transport) proteins. Unfortunately, the designation LptA was already given to the enzyme lipid A phosphoethanolamine transferase in N. meningitidis (26). Therefore, throughout this paper, we will use the designation LptH for the neisserial homolog of E. coli LptA.


Bacterial Strains and Growth Conditions

The bacterial strains used in this study are listed in Table 1. E. coli strains were grown on LB agar plates at 37 °C or in liquid LB medium. When necessary, an appropriate antibiotic (25 μg/ml chloramphenicol, 100 μg/ml ampicillin, or 50 μg/ml kanamycin) was added. N. meningitidis strains were grown at 37 °C in candle jars on GC agar plates (Oxoid), supplemented with Vitox (Oxoid) and, when necessary, with an antibiotic (10 μg/ml chloramphenicol or 80 μg/ml kanamycin). Liquid cultures were grown in tryptic soy broth (BD Biosciences). To achieve depletion of proteins encoded by genes cloned behind an isopropyl-β-d-1-thiogalactopyranoside (IPTG)-inducible promoter, N. meningitidis cells, grown overnight on plates containing 1 μm IPTG, were resuspended in tryptic soy broth without IPTG to an absorbance at 550 nm (A550) of 0.1 and grown for 6 h. To induce expression of IPTG-regulated genes in N. meningitidis, 0.5 mm IPTG was added to the medium. To obtain sialylated cells, 80 μm CMP-N-acetylneuraminic acid (Sigma) was added for 2 h to the medium of bacteria growing in the mid-log phase.

Strains and plasmids used in this study

Plasmid and Mutant Constructions

Plasmids and primers used in this study are summarized in Table 1 and supplemental Table 1, respectively. Primers were designed based on the genome sequence of serogroup B strain MC58, which belongs to the same clonal complex as the strain used in this study, H44/76. Deletion constructs of genes or operons were obtained by amplifying DNA fragments upstream and downstream of these loci by PCR using genomic DNA of an unencapsulated derivative of H44/76, i.e. strain HB-1, as template and primer pairs indicated with Up-For/Up-Rev and Down-For/Down-Rev in supplemental Table 1. The fragments were cloned into pCRII-TOPO and joined together in one plasmid using the AccI sites that were introduced via the primers and the XbaI and/or SpeI sites in the vector. A kanamycin-resistance gene (kanR) cassette including the neisserial DNA uptake sequence, obtained from pMB25, was inserted in each plasmid after AccI restriction. All inactivation constructs contained the kanR in the same transcriptional direction as the gene or operon it replaced. N. meningitidis was transformed as described (10) with PCR fragments obtained from the gene replacement constructs using primer pair M13Rev and M13For. The transformants were checked for the presence of the mutant alleles by PCR using the corresponding Up-For and Down-Rev primers and for the absence of the wild-type alleles by PCR using primers annealing within the removed coding sequence (indicated with -int in supplemental Table 1) and the corresponding Down-Rev primer. Intact genes were amplified by PCR using genomic DNA of HB-1 as template and primers indicated with NdeI and AatII (supplemental Table 1). The resulting PCR products were cloned into pCRII-TOPO and subcloned into the neisserial replicative plasmid pEN11-Imp via NdeI/AatII restriction and ligation. In this vector, expression of the inserted gene is driven by tandem lac/tac promoter/operator sequences. To engineer a His tag at the C terminus of LptE, the lptE gene was amplified from HB-1 DNA using primers LptE-NdeI and LptE-Rev-XhoI. The resultant PCR product was cloned into pCRII-TOPO followed by subcloning into pET26b via NdeI and XhoI restriction, yielding pET26b-LptE. This plasmid was then used as template in a PCR with primers LptE-NdeI and pET26-Rev-AatII. After cloning the PCR product into pCRII-TOPO, an NdeI-AatII fragment from this construct was used to replace the lptD gene in pEN11-imp generating pEN11-LptE-His, which encodes LptE with a C-terminal His6 tag.

LPS Localization

Assessment of cell surface localization of LPS using modification by PagL or accessibility to extracellular neuraminidase was performed as described (10).

Cell Fractionations

To collect proteins from extracellular media, bacterial cultures were centrifuged for 5 min at 10,000 × g in an Eppendorf centrifuge. One ml of supernatant was treated with 7% trichloroacetic acid for 1 h at 4 °C and centrifuged for 30 min at 16,000 × g. The resulting pellet was washed twice with ice-cold acetone and boiled in SDS-PAGE sample buffer. Cell envelopes were prepared as described (27).

Pulldown Experiment Using LptE-His

Derivatives of strain HB-1 containing either pEN11-LptE or pEN11-LptE-His were grown in 100 ml of tryptic soy broth plus IPTG to an A550 of 2.5. Cells were harvested and processed to cell envelopes. These were incubated with 1.25% Elugent (Calbiochem) in 20 mm Tris-HCl, 150 mm NaCl, pH 7.5 (TBS) for 2 h at room temperature and centrifuged for 1 h at 100,000 × g. The supernatant was incubated with Ni2+-NTA agarose beads (Qiagen) in TBS containing 0.1% Elugent and 20 mm imidazole (wash buffer) for 16 h at 4 °C. The beads were washed five times with wash buffer. Bound material was eluted with wash buffer containing 300 mm imidazole. Samples were analyzed by SDS-PAGE and immunoblotting.

SDS-PAGE Analysis and Immunoblotting

Protein samples were analyzed by regular SDS-PAGE and immunoblotting as described (10). Semi-native SDS-PAGE was performed as described (27). For LPS analysis, samples denatured in SDS-PAGE sample buffer were treated with 1 mg/ml proteinase K at 55 °C for 1 h, boiled for 10 min, and subjected to regular SDS-PAGE or Tricine-SDS-PAGE (10) and stained with silver (28).

Vancomycin Sensitivity

Meningococci grown overnight on GC agar plates were resuspended in 100 μl of tryptic soy broth to an A550 of 0.2 and plated on GC agar plates. Paper discs containing 30 μg of vancomycin (BD Biosciences) were placed on top of the agar. The plates were incubated at 37 °C for 24 h, after which growth inhibition zones around the discs were measured in millimeters from the rim of the disc. All tests were repeated at least three times.

Bioinformatic Analyses

The Comprehensive Microbial Resource data base was used to find putative LptE homologs. The genome data base at NCBI was used to search for the presence of COG family members in individual genomes. The NCBI website was used to assess the presence of COG families in different bacterial phylae. Secondary structure predictions were performed using the PSIPRED protein structure prediction server.


Putative LPS Transport-associated Loci in N. meningitides

In E. coli, the newly identified genes involved in LPS transport are present in three different loci on the chromosome: one comprising lptA, lptB, and lptC, another comprising lptF and lptG, and the third one containing lptE. By performing Blast searches, we found homologs of all of these genes in the N. meningitidis genome sequences in a generally similar organization, although some of the neighboring genes are different (Fig. 1). Homologs of the lptC, lptA, and lptB genes correspond to locus tags NMB0354, NMB0355, and NMB0356, respectively, in the genome sequence of strain MC58. As explained above, the neisserial homolog of E. coli LptA will be referred to as LptH. The genes encoding LptF and LptG were identified as NMB1570 and NMB1571, respectively. They overlap by four nucleotides, as in E. coli (Fig. 1B). The lptE homolog was identified as NMB0707, which is located upstream of the holA gene, separated by only one nucleotide, whereas the two genes overlap by one nucleotide in E. coli. Upstream of lptE in E. coli, the leuS gene encoding leucyl-tRNA synthetase is found, which is present at a different location in the MC58 genome (encoded by NMB1897) (Fig. 1C). Upstream of the lptE gene, the locus tag NMB0706 is annotated as a homolog of yfiH, encoding a conserved protein annotated as a laccase.

Organization of genetic loci implicated in LPS transport in E. coli and N. meningitidis. Annotations were taken from strains MG1655 (E. coli) and MC58 (N. meningitidis) from the JCVI Comprehensive Microbial Resource. ORF lengths are not to scale. A, ...

Construction of lpt Mutants

We created individual gene inactivation constructs for lptB, lptC, lptE, lptF, lptG, and lptH by replacing the main part of each coding sequence by a kanR cassette with preservation of the overlapping regions between kdsC/lptC and lptC/lptH. In addition, a construct was created where both lptF and lptG were simultaneously replaced by one kanR cassette. Subsequently, strain HB-1 was transformed with PCR products comprising the various gene inactivation constructs. Transformants were tested by PCR for the presence of the inactivated allele and the absence of the wild-type allele. Correct mutants were obtained with the lptB, lptC, lptH, lptF, lptG, and lptFG inactivation constructs but not with the lptE inactivation construct. The failure to obtain a ΔlptE mutant suggested that this gene could be essential or that a polar effect of the deletion on the downstream gene, i.e. holA, encoding subunit delta of DNA polymerase III, prevented the isolation of a ΔlptE mutant. To distinguish between these options, we transformed strain HB-1 with a series of plasmids containing lptE and/or holA under IPTG control and tested whether we could inactivate the chromosomal copy of lptE in the resulting strains in the presence of IPTG. Plasmids containing the lptE-holA locus or only holA allowed for the recombination of lptE::kan on the chromosome, but not a plasmid containing only lptE. This demonstrates that LptE is not an essential protein, but that replacement of part of the lptE gene by a kanR cassette has a polar effect on holA. Possibly, the promoter in the kanR cassette is of a different strength than the normal promoter driving holA expression, or the holA promoter is located in the deleted portion of lptE. Thus, none of the tested Lpt proteins is essential in N. meningitidis, as expected if they only function in LPS transport.

Phenotypes of lpt Mutants

To establish a role for the Lpt-related proteins in LPS transport, we first tested the mutants obtained for the most striking feature observed previously in meningococcal mutants defective in LPS transport, i.e. a severely decreased production of LPS (10, 25). We analyzed LPS levels by SDS-PAGE of proteinase K-treated cell lysates followed by silver staining. This experiment showed dramatically decreased cellular levels of LPS in all lpt mutants, except in the ΔlptE strain (Fig. 2A). Note that we had to load much more of the mutant cell lysates to detect LPS. Then, we tried to complement the phenotype by transforming the mutants with plasmids containing the relevant gene under IPTG control. Only after several attempts, we obtained a few transformants for the ΔlptH and ΔlptFG mutants, which were designated HB-1ΔlptH(pLptH) and HB-1ΔlptFG(pLptFG). LPS levels were restored to wild-type levels after growth of these strains in the presence of IPTG (Fig. 2B). Despite many tries, we were unable to transform the other lpt mutants with the complementing plasmids in a similar manner. Apparently, the lpt mutations render the cells less competent. This is likely related to their low LPS content, because a meningococcal lpxA mutant, which does not produce any LPS, was also reported to be severely impaired in competence (29). Therefore, we created conditional strains by first introducing a plasmid containing the relevant gene under IPTG control in strain HB-1, followed by inactivation of the chromosomal gene copy in the presence of IPTG. These conditional strains, HB-1ΔlptB(pLptB), HB-1ΔlptC(pLptC), and as a control strain HB-1ΔlptH(pLptH)-1 created in a similar manner, were subsequently grown in the presence or absence of IPTG after which LPS levels were assessed by SDS-PAGE and silver staining. Growth in the presence of IPTG resulted in LPS levels comparable with those in strain HB-1 (data not shown). As shown in Fig. 2C, depletion of LptB, LptC, or LptH after growth in the absence of IPTG resulted in a drastic decrease in cellular LPS levels. These levels appeared not to be as low as those found in the null mutants, likely because the tandem lac-tac promoter controlling lpt expression in these conditional strains is never completely shut off (30). Together, these data show that the decreased LPS levels in the ΔlptB, ΔlptC, ΔlptH, and ΔlptFG mutants are directly related to the absence of the Lpt protein and not a consequence of secondary mutations.

Phenotypes of lpt mutants. A–C, proteinase K-treated cell lysates were analyzed by SDS-PAGE and silver staining to assess cellular LPS levels. The relative loadings, based on absorbance, are indicated below the lanes. The presence or absence of ...

N. meningitidis mutants, producing no or low levels of LPS, such as ΔlpxA, ΔlptD, and ΔmsbA mutants, show a characteristic enhanced, stable colony opacity on plates (10, 25, 45). All lpt mutant strains also showed this phenotype, except for the ΔlptE mutant (data not shown). Furthermore, all lpt mutants, except the ΔlptE strain, grew considerably slower than the parent strain (data not shown) and released high levels of proteins in the medium (Fig. 2D), similar to what was observed before for the ΔlptD mutant (10). For comparison, we included in Fig. 2D the extracellular protein profile of a ΔrmpM strain, a mutant known to excessively shed OM blebs (31), to demonstrate that the increased amounts of extracellular proteins in the lpt mutants do not result from extensive blebbing. The apparent leakiness of the cells was also indicated from their enhanced sensitivity toward the antibiotic vancomycin, with the ΔlptE mutant showing an intermediate sensitivity (Fig. 2E). These leaky phenotypes were restored in the strains with the complementing plasmids after growth in the presence of IPTG (data not shown). Overall, the phenotypes of the mutants lacking LptB, LptC, LptH, and LptF, and LptG are consistent with the proposed role of these proteins in LPS transport, but, interestingly, we found no evidence for a role for LptE in this process.

Assessment of Cell Surface Localization of LPS

To show that the mutants were indeed affected in LPS transport and not only in LPS biosynthesis, we created lpt mutations in two different backgrounds that allow for assessment of the presence of LPS at the cell surface. First, we used a strain containing an IPTG-inducible copy of Bordetella bronchiseptica pagL. PagL is an OM enzyme that can remove an acyl chain from LPS when LPS is present at its proper location, i.e. the outer leaflet of the OM. We used this modification previously to demonstrate lack of LPS cell surface exposure in an N. meningitidis lptD mutant (10). When PagL is expressed in the parent strain HB-1, its LPS migrates faster in Tricine-SDS-PAGE, consistent with the removal of an acyl chain (Fig. 3A). In contrast, the expression of PagL in the lptH, lptC, lptF, and lptG mutants did not affect LPS mobility (Fig. 3A), demonstrating that the residual LPS present in these strains does not reach the cell surface. The lptB mutation resulted in such low levels of LPS that we could not reliably assess differences in its LPS migration. We could not create an lptE knock-out mutation in the strain expressing PagL because both pagL and holA are present on similar plasmids, and there are no known compatible plasmid systems for N. meningitidis. Therefore, and to confirm the conclusions from the PagL assay for the other mutants, we also created lpt mutations in a wild-type N. meningitidis strain expressing the L3 immunotype LPS, which is characterized by the presence of a lacto-N-neotetraose unit in the outer core. This moiety can be capped with sialic acid when the bacteria are grown in the presence of CMP-N-acetylneuraminic acid. This sialic acid can be removed from LPS by extracellularly added neuraminidase provided that the LPS is properly exposed at the cell surface (10). Fig. 3B shows the appearance of L3 LPS in the lpt mutants after growth in the presence of CMP-N-acetylneuraminic acid and subsequent treatment, or not, of intact cells with neuraminidase. Only the LPS of the wild type and of the ΔlptE mutant was surface-accessible as shown by the higher electrophoretic mobility of the LPS after neuraminidase treatment. The LPS of other lpt mutants was not affected by neuraminidase treatment, showing that it is not exposed at the cell surface, consistent with the results of the PagL modification assay. Thus, LptE has no apparent role in LPS transport in N. meningitidis, a finding that is at odds with that reported for E. coli.

Localization of LPS. Proteinase K-treated cell lysates were analyzed by Tricine-SDS-PAGE and silver staining. Relative loadings, based on absorbance, for wild-type and ΔlptE strains were one-fifth of those of the other strains. A, the indicated ...

Function of LptE in N. meningitidis

To test whether also other features of E. coli LptE might be different from those of N. meningitidis LptE, we examined the interaction between LptE and LptD. In E. coli, LptD was shown to interact with a C-terminally His-tagged LptE in pulldown assays (11). To assess this interaction in N. meningitidis, we constructed a strain, designated HB-1(pLptE-His), that produces LptE with a His tag on its C terminus from plasmid. Affinity purification using Ni2+-NTA beads on membrane extracts resulted in co-purification of LptD with LptE (Fig. 4A). This interaction was specific because no Omp85/BamA was co-purified and no LptD was purified when membranes containing overexpression levels of LptE without a His tag were used (Fig. 4A). Thus, in this respect, neisserial LptE behaves the same as E. coli LptE.

Interaction between LptE and LptD. A, association of LptE and LptD. Pulldown experiments using cells expressing His6-tagged LptE (LptE-His) or LptE without a tag as control (LptE). The two left panels represent the cell envelopes used as starting material, ...

To study the interaction between LptD and LptE further, we assessed the levels of LptD in the ΔlptE mutant and, as controls, also in the other lpt mutants. Cellular LptD levels were similar in all mutants except in the ΔlptE strain where they were severely decreased (Fig. 4B). This defect was completely restored in a strain complemented with a copy of lptE under IPTG control on plasmid (Fig. 4B, compare lanes 5 and 6). Together, these data show that LptE is associated with LptD and that it functions in the biogenesis and/or the stability of LptD, i.e. like a chaperone, but that it has no direct role in the transport of LPS to the cell surface. The amounts of LptD in the OM of the ΔlptE mutant are apparently not sufficiently decreased to cause serious defects in LPS transport to the cell surface.

Conformation of LptD in Wild-type and ΔlptE Strains

To investigate further whether LptE is essential for correct folding of LptD, we assessed the electrophoretic mobility of LptD in wild-type and ΔlptE cells by semi-native SDS-PAGE. When neisserial cell envelopes are subjected to this procedure, LptD is detected as two high molecular weight (HMW) bands, indicating that the protein is present in complexes (32) (Fig. 5A). These HMW forms of LptD were present in the same positions in the gel when cell envelopes of the ΔlptE mutant were analyzed, which shows that LptE is not part of these complexes. Apparently, LptE dissociated from LptD, presumably by the presence of SDS in the sample buffer. Consistent with the lower cellular LptD levels, both HMW bands were less intense in the ΔlptE mutant than in the parent or in the complemented strain (Fig. 5A).

Conformations of LptD. Cell envelopes derived from the indicated strains were subjected to semi-native SDS-PAGE followed by immunoblotting with anti-LptD antiserum. The positions of molecular mass markers (in kDa) are shown at the left. A and C, samples ...

LptD is an unusual OM protein because it contains multiple cysteine residues. In E. coli, it is the only OM protein with more than two, i.e. four, cysteine residues (33). Interestingly, LptD of N. meningitidis contains an uneven number of cysteine residues, namely five. To test whether disulfide bridges are involved in the formation of the HMW LptD complexes, we compared the migration of LptD in reducing versus nonreducing conditions. As shown in Fig. 5B, addition of β-mercaptoethanol to unboiled samples resulted in a shift of almost all LptD to a position in the gel lower than that of the fully denatured protein. This form likely represents a partially folded LptD monomer, where the faster migration is imparted by the heat-modifiability of the β-barrel domain, a common characteristic of OM proteins (34). This observation demonstrates that disulfide bridges are required for the appearance of LptD in HMW complexes. To assess whether these disulfide bonds are intra- or intermolecular, cell envelopes were boiled in the absence of a reducing agent before loading on the gel. Under those conditions LptD migrated in approximately the same position as the fully denatured protein, indicating that LptD is not present in the HMW forms as disulfide-linked monomers (Fig. 5B). In fact, the similar positions in the gel of reduced and oxidized heat-denatured LptD suggest that LptD is not bound to any protein via disulfide bridges.

To test whether any of the other Lpt proteins is present in the HMW complexes, we performed semi-native SDS-PAGE analysis of cell envelopes derived from the lpt mutants. As shown in Fig. 5C, the migration of the LptD complexes was unaffected by the absence of any of the Lpt proteins, demonstrating that they are not part of this complex. Also, the complete absence of any LPS, as assessed by using an lpxA mutant, did not affect the appearance of the HMW LptD complex (Fig. 5C, lane 7). Altogether, these data show that intramolecular disulfide bridges are required for formation of the HMW LptD complexes and that LptD does not form intermolecular disulfide bridges. The absence of LptE did not grossly affect any of the electrophoretic properties of LptD, although some unfolded LptD was detected in unboiled preparations of the ΔlptE mutant cell envelopes, consistent with the proposed role of LptE in biogenesis and/or stability of LptD.

Lack of Conservation of LptE in LPS-producing Proteobacteria

Recent bioinformatic studies demonstrated that not all LPS-producing bacteria contain homologs of all E. coli Lpt proteins (35, 36). Specifically, homologs of LptC and LptE appear to be poorly conserved. The absence of LptE in LPS-producing bacteria would substantiate our finding that this protein is not essential for LPS transport in all bacteria. Possibly, if LptE is mainly involved in chaperoning LptD, there might be less constraint on its sequence conservation. To test this idea, we used identification criteria other than significant Blast hits to find potential LptE-like proteins, i.e. by exploiting its genetic context. A comparison of the lptE gene among the γ-proteobacteria showed a striking conservation in its genetic location: it was invariably found in between leuS and holA. Sequence alignments of LptE homologs from the γ-proteobacteria showed little sequence conservation, except for the first three amino acids of the predicted mature part of the protein: the invariant cysteine residue is almost always followed by a glycine and an aromatic residue (supplemental Fig. 1). Furthermore, the γ-proteobacterial LptE proteins belong to the COG2980 cluster of orthologous groups of proteins, as identified by the Conserved Domain Data base (CDD) (37). Using these criteria, we analyzed representative genomes from all classes of the proteobacteria for the presence of putative LptE homologs (Table 2). When we used LptE as query in Blast searches, we found no hits in all α- and ϵ-proteobacteria and some δ-proteobacteria. However, in all cases analyzed, we found a putative lipoprotein-encoding gene downstream of leuS and/or upstream of holA. These putative lipoproteins from the α-, δ-, and ϵ-proteobacteria did not always belong to the COG2980 family. The Conserved Domain Data base detected either no or a different COG, for example COG5468 in the α-proteobacteria. However, despite the lack of sequence conservation, the predicted secondary structures of the identified lipoproteins were all consistent with the determined three-dimensional structures of three LptE orthologs (Protein Data Base ID codes 2R76, 2JXP, and 3BF2), with mostly β-strands and a distinct long α-helix at the C terminus of the protein (data not shown). Thus, LPS-producing bacteria can be found whose genomes do not appear to encode LptE orthologs. However, the highly conserved presence of a putative lipoprotein encoded by a gene adjacent to the leuS and/or holA genes in these bacteria, with LptE-like predicted secondary structure, is striking and might indicate that this lipoprotein represents a functional LptE homolog apparently undergoing extensive evolution.

Features of putative LptE-like proteins in representative proteobacterial genomes


N. meningitidis is a highly informative model organism for the study of LPS biogenesis, because it is able to survive without LPS. Consequently, clean knock-out mutants can be generated for genes involved in LPS biosynthesis. Also, mutants lacking genes involved in LPS transport are viable because LPS does not accumulate in aberrant locations, as is the case in E. coli, apparently due to an unknown feedback mechanism that lowers LPS synthesis in such N. meningitidis mutants (10, 25). Such clean knock-out mutants generally allow for a clearer interpretation of mutant phenotypes as opposed to those of depletion strains, where it can be more difficult to discriminate between direct and indirect effects. In the current study, we addressed the role in N. meningitidis of several newly identified proteins, implicated in LPS transport in E. coli. The phenotypes of N. meningitidis strains lacking LptB, LptC, LptH (homolog of E. coli LptA), LptF, and LptG were identical to those lacking LptD or MsbA, i.e. the knock-out mutants were viable but leaky and produced only very little LPS, which was not present at the cell surface. These findings substantiate a general role of these proteins in LPS transport. Moreover, the fact that none of the Lpt proteins is essential in N. meningitidis also indicates that they are not involved in the transport of phospholipids to the OM. This obviously essential process is still poorly understood. In E. coli, the MsbA protein is implicated in the translocation of both LPS and phospholipids across the inner membrane as inferred from the phenotype of an msbA depletion mutant (9). However, we showed that this is not the case in N. meningitidis, because a meningococcal msbA mutant is viable and forms an OM (25). It is unknown whether the proteins more downstream in the LPS transport pathway are also involved in phospholipid transport in E. coli, but our current data clearly indicate that this is not the case in N. meningitidis.

Interestingly, the absence of LptE did not affect LPS levels or its presence at the cell surface, demonstrating that LptE is not directly involved in LPS transport in N. meningitidis, in contrast to its reported role in E. coli. However, similar as in E. coli, LptE appears to be associated with LptD in N. meningitidis. As we observed drastically decreased amounts of LptD in the absence of LptE, we conclude that LptE may serve to stabilize LptD or assist in its biogenesis, which is also suggested from the low amounts of unfolded LptD present in membrane preparations of the lptE mutant after semi-native SDS-PAGE (Fig. 5B). This suggests that LptE functions as an LptD chaperone. Apparently, the reduced amounts of correctly assembled LptD in the lptE mutant are still sufficient to maintain normal LPS transport. In E. coli, LptE was also found to be required for the assembly of LptD (38) and for its proper oxidation (39). Interestingly, in E. coli, LptE was shown to reside within the LptD β-barrel (40) and to bind LPS in vitro (41). However, our finding that LPS is properly transported to the cell surface in the absence of LptE indicates that LptE cannot be the final docking site for LPS at the OM, at least not in N. meningitidis. LPS can be transferred from LptC to its structural homolog LptA, but not vice versa, consistent with the expected directionality in the LPS transport pathway (18). LptA was shown to bind to LptC (42, 43) but not to LptE (42), which would be consistent with the idea that LptE is not the LPS recipient at the OM. A more likely recipient for LPS could be the N terminus of LptD because it shows sequence similarity to LptA (6).

Neisserial LptD is present in HMW complexes, detectable in semi-native SDS-PAGE. As we showed in this study, none of the other Lpt proteins is part of these complexes, so they likely represent LptD homo-oligomers. The observation that the lipoprotein LptE did not remain associated with the HMW LptD complexes during semi-native SDS-PAGE is reminiscent of previous findings on the Bam complex in N. meningitidis. The Bam complex, which inserts β-barrel proteins into the OM, consists of the central component BamA/Omp85 (a β-barrel protein) with associated lipoproteins. When this complex from N. meningitidis was subjected to similar semi-native SDS-PAGE conditions as used here, the lipoprotein components also detached from the BamA protein (27). The putative LptD multimers observed in semi-native SDS-PAGE require folded and oxidized forms of LptD because they fall apart upon heating or reduction of the samples (Fig. 5B). Judging from the migration of the folded monomeric form of LptD, the complexes might represent tetramers and heptamers. These findings constitute another difference with E. coli LptD, which was not found in HMW complexes, but where LptD and LptE form a very tight complex that withstands SDS-PAGE conditions (41). Possibly, the LPS-receiving complex in the OM can be formed by a tight LptD-LptE complex as in E. coli, or by an LptD multimer that does not need LptE for function as in N. meningitidis.

Several bioinformatics studies report that LptE and LptC are the least conserved proteins of the Lpt system, as in many LPS-producing bacteria no homologs of E. coli LptC and LptE were found (35, 36, 44). However, our searches, using criteria other than significant sequence homology, actually revealed the presence of putative LptE-like proteins in these bacteria. It will be very interesting to determine whether these putative LptE-like proteins indeed function as components of the Lpt machinery.

In summary, we show that LptB, LptC, LptF, LptG, and LptH(LptA) are essential components of the LPS transport system in the important model organism for OM biogenesis, N. meningitidis, as they are in E. coli. However, we show that LptE is not directly involved in LPS transport, but only assists in the assembly of LptD. Furthermore, we report on the presence in LPS-producing bacteria of putative LptE proteins that are divergent in sequence from their E. coli counterparts. These findings underscore the relevance of studying conserved mechanisms in different model systems.

Supplementary Material

Supplemental Data:


We thank Ingrid Schilders for technical assistance.

*This work was supported by the Netherlands Research Council for Earth and Life Sciences of the Netherlands Organization for Scientific Research.

An external file that holds a picture, illustration, etc.
Object name is sbox.jpgThe on-line version of this article (available at contains supplemental Fig. 1 and Table 1.

2The abbreviations used are:

outer membrane
high molecular weight
kanamycin resistance gene
nickel-nitrilotriacetic acid


1. McClerren A. L., Endsley S., Bowman J. L., Andersen N. H., Guan Z., Rudolph J., Raetz C. R. H. (2005) Biochemistry 44, 16574–16583 [PMC free article] [PubMed]
2. Srinivas N., Jetter P., Ueberbacher B. J., Werneburg M., Zerbe K., Steinmann J., Van der Meijden B., Bernardini F., Lederer A., Dias R. L., Misson P. E., Henze H., Zumbrunn J., Gombert F. O., Obrecht D., Hunziker P., Schauer S., Ziegler U., Käch A., Eberl L., Riedel K., DeMarco S. J., Robinson J. A. (2010) Science 327, 1010–1013 [PubMed]
3. Raetz C. R., Whitfield C. (2002) Annu. Rev. Biochem. 71, 635–700 [PMC free article] [PubMed]
4. Doerrler W. T. (2006) Mol. Microbiol. 60, 542–552 [PubMed]
5. Sperandeo P., Dehò G., Polissi A. (2009) Biochim. Biophys. Acta 1791, 594–602 [PubMed]
6. Bos M. P., Robert V., Tommassen J. (2007) Annu. Rev. Microbiol. 61, 191–214 [PubMed]
7. Ruiz N., Kahne D., Silhavy T. J. (2009) Nat. Rev. Microbiol. 7, 677–683 [PMC free article] [PubMed]
8. Zhou Z., White K. A., Polissi A., Georgopoulos C., Raetz C. R. (1998) J. Biol. Chem. 273, 12466–12475 [PubMed]
9. Doerrler W. T., Gibbons H. S., Raetz C. R. (2004) J. Biol. Chem. 279, 45102–45109 [PubMed]
10. Bos M. P., Tefsen B., Geurtsen J., Tommassen J. (2004) Proc. Natl. Acad. Sci. U.S.A. 101, 9417–9422 [PubMed]
11. Wu T., McCandlish A. C., Gronenberg L. S., Chng S. S., Silhavy T. J., Kahne D. (2006) Proc. Natl. Acad. Sci. U.S.A. 103, 11754–11759 [PubMed]
12. Sperandeo P., Cescutti R., Villa R., Di Benedetto C., Candia D., Dehò G., Polissi A. (2007) J. Bacteriol. 189, 244–253 [PMC free article] [PubMed]
13. Sperandeo P., Lau F. K., Carpentieri A., De Castro C., Molinaro A., Dehò G., Silhavy T. J., Polissi A. (2008) J. Bacteriol. 190, 4460–4469 [PMC free article] [PubMed]
14. Ma B., Reynolds C. M., Raetz C. R. H. (2008) Proc. Natl. Acad. Sci. U.S.A. 105, 13823–13828 [PubMed]
15. Tran A. X., Trent M. S., Whitfield C. (2008) J. Biol. Chem. 283, 20342–20349 [PMC free article] [PubMed]
16. Narita S., Tokuda H. (2009) FEBS Lett. 583, 2160–2164 [PubMed]
17. Ruiz N., Gronenberg L. S., Kahne D., Silhavy T. J. (2008) Proc. Natl. Acad. Sci. U.S.A. 105, 5537–5542 [PubMed]
18. Tran A. X., Dong C., Whitfield C. (2010) J. Biol. Chem. 285, 33529–33539 [PMC free article] [PubMed]
19. Chng S. S., Gronenberg L. S., Kahne D. (2010) Biochemistry 49, 4565–4567 [PMC free article] [PubMed]
20. Tefsen B., Geurtsen J., Beckers F., Tommassen J., de Cock H. (2005) J. Biol. Chem. 280, 4504–4509 [PubMed]
21. Galloway S. M., Raetz C. R. H. (1990) J. Biol. Chem. 265, 6394–6402 [PubMed]
22. Braun M., Silhavy T. J. (2002) Mol. Microbiol. 45, 1289–1302 [PubMed]
23. Sperandeo P., Pozzi C., Dehò G., Polissi A. (2006) Res. Microbiol. 157, 547–558 [PubMed]
24. Steeghs L., den Hartog R., den Boer A., Zomer B., Roholl P., van der Ley P. (1998) Nature 392, 449–450 [PubMed]
25. Tefsen B., Bos M. P., Beckers F., Tommassen J., de Cock H. (2005) J. Biol. Chem. 280, 35961–35966 [PubMed]
26. Cox A. D., Wright J. C., Li J., Hood D. W., Moxon E. R., Richards J. C. (2003) J. Bacteriol. 185, 3270–3277 [PMC free article] [PubMed]
27. Volokhina E. B., Beckers F., Tommassen J., Bos M. P. (2009) J. Bacteriol. 191, 7074–7085 [PMC free article] [PubMed]
28. Hitchcock P. J., Brown T. M. (1983) J. Bacteriol. 154, 269–277 [PMC free article] [PubMed]
29. Albiger B., Johansson L., Jonsson A. B. (2003) Infect. Immun. 71, 155–162 [PMC free article] [PubMed]
30. Long C. D., Tobiason D. M., Lazio M. P., Kline K. A., Seifert H. S. (2003) Infect. Immun. 71, 6279–6291 [PMC free article] [PubMed]
31. Steeghs L., Berns M., ten Hove J., de Jong A., Roholl P., van Alphen L., Tommassen J., van der Ley P. (2002) Cell. Microbiol. 4, 599–611 [PubMed]
32. Volokhina E. B., Grijpstra J., Stork M., Schilders I., Tommassen J., Bos M. P. (2011) J. Bacteriol. 193, 1612–1621 [PMC free article] [PubMed]
33. Denoncin K., Vertommen D., Paek E., Collet J. F. (2010) J. Biol. Chem. 285, 29425–29433 [PMC free article] [PubMed]
34. Dekker N., Merck K., Tommassen J., Verheij H. M. (1995) Eur. J. Biochem. 232, 214–219 [PubMed]
35. Sutcliffe I. C. (2010) Trends Microbiol. 18, 464–470 [PubMed]
36. Haarmann R., Ibrahim M., Stevanovic M., Bredemeier R., Schleiff E. (2010) J. Phys. Condens. Matter 22, 454124. [PubMed]
37. Marchler-Bauer A., Anderson J. B., Cherukuri P. F., DeWeese-Scott C., Geer L. Y., Gwadz M., He S., Hurwitz D. I., Jackson J. D., Ke Z., Lanczycki C. J., Liebert C. A., Liu C., Lu F., Marchler G. H., Mullokandov M., Shoemaker B. A., Simonyan V., Song J. S., Thiessen P. A., Yamashita R. A., Yin J. J., Zhang D., Bryant S. H. (2005) Nucleic Acids Res. 33, D192–196 [PMC free article] [PubMed]
38. Chimalakonda G., Ruiz N., Chng S. S., Garner R. A., Kahne D., Silhavy T. J. (2011) Proc. Natl. Acad. Sci. U.S.A. 108, 2492–2497 [PubMed]
39. Ruiz N., Chng S. S., Hiniker A., Kahne D., Silhavy T. J. (2010) Proc. Natl. Acad. Sci. U.S.A. 107, 12245–12250 [PubMed]
40. Freinkman E., Chng S. S., Kahne D. (2011) Proc. Natl. Acad. Sci. U.S.A. 108, 2486–2491 [PubMed]
41. Chng S. S., Ruiz N., Chimalakonda G., Silhavy T. J., Kahne D. (2010) Proc. Natl. Acad. Sci. U.S.A. 107, 5363–5368 [PubMed]
42. Bowyer A., Baardsnes J., Ajamian E., Zhang L., Cygler M. (2011) Biochem. Biophys. Res. Commun. 404, 1093–1098 [PubMed]
43. Sperandeo P., Villa R., Martorana A. M., Samalikova M., Grandori R., Dehò G., Polissi A. (2011) J. Bacteriol. 193, 1042–1053 [PMC free article] [PubMed]
44. Silander O. K., Ackermann M. (2009) BMC Res. Notes 2, 2. [PMC free article] [PubMed]
45. Bos M. P., Tommassen J. (2005) Infect. Immun. 73, 6194–6197 [PMC free article] [PubMed]
46. Tzeng Y. L., Datta A., Strole C., Kolli V. S., Birck M. R., Taylor W. P., Carlson R. W., Woodard R. W., Stephens D. S. (2002) J. Biol. Chem. 277, 24103–24113 [PubMed]

Articles from The Journal of Biological Chemistry are provided here courtesy of American Society for Biochemistry and Molecular Biology