Molecular and Cellular Biology Program, Division of Infectious Diseases1 and Comparative Medicine Program, Department of Medicine,2 University of Maryland, Baltimore, School of Medicine, Baltimore, MD 21201, USA
Institut für Mikrobiologie und Genetik, Georg-August Universität, Grisebachstrasse 8, 37077 Göttingen, Germany3
Author for correspondence: Michael S. Donnenberg. Tel: +1 410 706 7560. Fax: +1 410 706 8700. e-mail: mdonnenb{at}umaryland.edu
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ABSTRACT |
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Keywords: type IV fimbriae, periplasm, bundle-forming pili, bundlin
Abbreviations: BFP, bundle-forming pilus/pili; EPEC, enteropathogenic Escherichia coli
a These authors contributed equally to this paper.
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INTRODUCTION |
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Despite the identification of many genetic loci that are required for type IV pilus expression, little insight has been gained into the process of type IV pilus morphogenesis. The functions of only three of the proteins involved in this process are well understood. The pilin protein itself has been studied in detail. This protein makes up most, if not all, of the subunits of the pilus, where it is arranged in a helical array (Paranchych & Frost, 1988 ). The pilin proteins of several bacterial species are known to be modified by glycosylation and/or phosphorylation (Virji et al., 1993
; Stimson et al., 1995
; Castric, 1995
; Forest et al., 1999
). The crystal structure of the complete pilin from Neisseria gonorrhoeae has been solved and has been shown to resemble a ladle, with a long, amino-terminal handle composed of an uninterrupted hydrophobic
-helix and a head composed predominantly of anti-parallel ß-sheets (Parge et al., 1995
). It is postulated that the handles form the core of the pilus, while certain antigenic regions of the rest of the protein protrude from the pilus surface. According to a theoretical model of bundlin (Protein Database accession no. 1QT2; http://www.rcsb.org/pdb/), the type IV pilin of enteropathogenic Escherichia coli (EPEC), highly variable amino acids of bundlin from different EPEC strains are also predominantly located on the surface (Blank et al., 2000
). The pre-pilin peptidase is another type IV pilus biogenesis protein that is well understood. In P. aeruginosa this protein is a bi-functional cytoplasmic transmembrane protein which cleaves off the short, hydrophilic pre-pilin signal sequence and N-methylates the new amino terminus of the pilin (Strom et al., 1993
). Pre-pilin peptidases compose a unique class of bi-lobed aspartate proteases (LaPointe & Taylor, 2000
). A third protein, the secretin, has been analysed in some detail in the P. aeruginosa type IV pilus system and also in EPEC. This outer-membrane protein forms multimeric ring-shaped structures that are postulated to allow the passage of intact pili through the outer membrane (Bitter et al., 1998
; Schmidt et al., 2001
; Wolfgang et al., 2000
).
Mutants defective in the production of type IV pili with disruptions in numerous other genes have been described, but analyses of these mutants have shed little light on the process of type IV pilus biogenesis. A principal reason for this lack of illumination is that these mutants all have similar phenotypes. In most cases, the pre-pilin protein is produced and processed, no pili are made and fractionation studies reveal that the pilin co-purifies with membrane components (Nunn et al., 1990 ; Alm & Mattick, 1995
; Alm et al., 1996a
, b
; Anantha et al., 2000
). In striking contrast to chaperone-usher pilus biogenesis systems, best exemplified by P fimbriae, no periplasmic phase of export has been described for type IV pilus biogenesis. Thus, no mutations of type IV pilus biogenesis genes have resulted in the accumulation of free pilin in the periplasmic space and no periplasmic chaperones have been described. This has led some investigators to propose models for type IV pilus biogenesis that include no periplasmic phase of export, but instead envisage a structure that spans the inner and outer membranes (Fussenegger et al., 1997
; Iredell & Manning, 1994
; Hobbs & Mattick, 1993
). Recently, the use of conditional double-knockout mutants has allowed the dissection of type IV pilus formation into three different phases pilin processing, pilus formation and pilus extrusion (Wolfgang et al., 1998
, 2000
). To reveal these phases, it was essential to inactivate PilT, a putative ATP-binding protein involved in pilus retraction (Merz et al., 2000
). This protein seems to act in quality control by preventing fibre formation and, in turn, minimizing growth defects that may occur in the absence of essential biogenesis components.
Another striking feature of type IV pilus biogenesis systems is their similarity to type II secretion systems (Pugsley, 1993 ; Hobbs & Mattick, 1993
; Russel, 1998
; Nouwen et al., 1999
, 2000
). Type II secretion systems are responsible for the export of a variety of extracellular toxins and enzymes through the outer membrane. These exported proteins are first secreted into the periplasmic space by the Sec translocation system. Among the components shared between type IV pilus and type II secretion systems are a polytopic cytoplasmic transmembrane protein, a cytoplasmic nucleotide-binding protein, an outer-membrane secretin, a pre-pilin peptidase and several pre-pilin peptidase substrates. This extensive conservation of components implies that the two systems share common mechanisms.
We, and others, have previously described a cluster of 14 bfp genes from EPEC that is associated with the production of the type IV bundle-forming pilus (BFP) of this organism (Stone et al., 1996 ; Sohel et al., 1996
). When cloned into a laboratory strain of E. coli and placed under the control of an artificial promoter, these 14 genes are sufficient to direct the synthesis of the BFP. Thus, the BFP system is an excellent model for the study of type IV pilus biogenesis, since all of the required components are known. Four of the 14 genes of the bfp cluster have no known homologues in other type IV pilus systems. One of the most intriguing of these four genes is bfpU, which is predicted to encode a 17 kDa protein that has a typical type I signal peptidase cleavage site (Stone et al., 1996
). As the rest of the protein is predicted to be hydrophilic, it seems likely that BfpU resides in the periplasmic space. Thus, we decided to test the hypothesis that bfpU encodes a periplasmic protein required for type IV pilus biogenesis.
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METHODS |
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Construction of a bfpU mutant.
A fragment of the bfp gene cluster (GenBank accession no. Z68186; Donnenberg et al., 1992 ; Stone et al., 1996
) extending from nucleotide 3586 to 4350 and containing an XmaI site at its 3' end was amplified from plasmid pKDS5.1 using primers Donne-198 and the T7 primer from the pBluescript vector. Similarly, a fragment extending from nucleotide 4560 to 5702 of the bfp gene cluster and containing an XmaI site at its 5' end was amplified using primer Donne-199 and the universal reverse primer. These fragments were cloned separately into pCR-Script and then joined by their XmaI sites, to produce pKDS5.3. The aphA-3 kanamycin-resistance cassette was cloned from pUC18K2 into the XmaI site of pKDS5.3, to produce pKDS5.4. The 3·3 kb SalI fragment from pKDS5.4 was cloned into positive-selection suicide vector pCVD442 to create pKDS5.5. Allelic exchange was performed as described previously (Donnenberg et al., 1993
). PCR was performed using primer pairs Donne-28/Donne-29, Donne-110/Donne-111 and Donne-121/Donne-122, to verify the size of the bfpA, bfpI and bfpU genes, respectively. Details for all of the Donne- primers used in this study can be found in Table 2
.
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Antibody production.
The purified N-terminal hexahistidine-tagged BfpU was used for the production of mAbs in BALB/c mice using standard procedures. Ascites was prepared commercially (BioWorld, Dublin, OH) and purified using an E-Z Sep kit (Amersham Pharmacia Biotech), according to the manufacturers instructions. Mouse polyclonal anti-BfpU serum was obtained at the same time as spleen cells were harvested for production of the mAbs and absorbed against an acetone powder prepared from the bfpU mutant. All procedures were part of a protocol approved by the IACUC of the University of Maryland, School of Medicine. Animals were housed in a facility accredited by the Association for Accreditation and Assessment of Laboratory Animal Care (AAALAC), International, and directed by a board-certified Laboratory Animal Veterinarian.
Localized adherence and auto-aggregation assays.
Assays for localized adherence to HEp-2 cells and for auto-aggregation were performed as described previously (Donnenberg & Nataro, 1995 ; Anantha et al., 1998
).
Immunoblotting.
Proteins were separated by SDS-PAGE and transferred to PVDF membranes by the method of Towbin et al. (1979) . Primary antibodies were used at the following dilutions: anti-BfpU mAb 8C8, 1:60000; polyclonal anti-bundlin (Zhang & Donnenberg, 1996
), 1:5000; monoclonal anti-bundlin ICA4 (Girón et al., 1995
), 1:30000; anti-maltose-binding protein (New England Biolabs), 1:10000; anti-ß-galactosidase (5' to 3'; Boulder, CO), 1:1000; anti-penicillin-binding protein 1B (a gift of Nanne Nanninga, University of Amsterdam), 1:2000; anti-intimin (Jerse & Kaper, 1991
), 1:1000. Secondary goat-anti-mouse IgG or goat-anti-rabbit IgG antiserum conjugated to horseradish peroxidase (Amersham Pharmacia Biotech) was used at a dilution of 1:30000. Blots were developed by enhanced chemiluminescence (Amersham Pharmacia Biotech). When necessary, blots were stripped according to the manufacturers instructions.
For quantitative immunoblotting, protein concentrations of a whole-cell lysate of wild-type EPEC strain E2348/69 grown under BFP-expressing conditions and of purified His-tagged proteins were determined using a bicinchoninic acid assay (Pierce). Prior to cell lysis, the c.f.u. count for the wild-type culture was determined by plating out serial dilutions of the culture and enumerating the number of colonies formed. Samples with a range of protein concentrations were separated by 15% SDS-PAGE and analysed by immunoblotting, as described above. The blots were scanned on a Hewlett Packard ScanJet 4c machine and analysed using Scion Image software (Scion). Standard curves of relative intensity versus the logarithm of the number of molecules of His-tagged proteins present were generated using Microsoft Excel. The correlation coefficients of these curves were >0·95. From these curves, the number of molecules of bundlin and BfpU in the whole-cell preparations was calculated and the relative amounts of these proteins were determined.
Electron microscopy.
For negative-staining of BFP, bacteria were grown and prepared as described previously (Anantha et al., 2000 ). Cells used in the immunolocalization experiments were grown in DMEM until visible aggregation occurred. For plasmolysis, the cells were harvested by centrifugation, resuspended in 1/10 vol. of an ice-cold sucrose solution (0·3 M sucrose, 0·15 M Tris) and incubated on ice for 30 min. After adding 10 vols of an EDTA solution (0·1 M, pH 8), the cells were collected by centrifugation and washed twice. They were then chemically fixed (at 4 °C for at least 1 h) in PBS (10 mM sodium phosphate, 137 mM sodium chloride, pH 7·0) with 0·2% (w/v) formaldehyde and 0·3% (w/v) glutaraldehyde and washed twice in the same buffer containing 10 mM glycine. Lowicryl K4M resin was used for embedding the cells (Gerberding & Mayer, 1988
; Roth et al., 1981
). Sections were cut with an Ultracut E (Reichert-Jung) ultramicrotome, mounted on nickel grids and kept, face down, on drops of PBS until they were immunolabelled. Gold-conjugates (10 nm) (goat anti-mouse; BBI) at a dilution of 1:100 were used to detect bound primary antibody (polyclonal mouse BfpU antiserum, 1:10). A 4% (w/v) aqueous uranyl acetate solution (pH 4·8) was used for post-staining of the sections.
Cell fractionation.
Bacteria were grown under BFP-inducing conditions and centrifuged (4000 g, 15 min at 4 °C). IPTG (1 mM) was added to the cultures for the last hour of incubation to induce ß-galactosidase expression. The supernatants were passed through a 0·2 µm filter and concentrated by precipitation with 10% (v/v) trichloroacetic acid. The precipitate was resuspended in 1/200 vol. of SDS loading buffer containing 10% saturated Tris base. The pellets of the culture were resuspended in 1/200 vol. of TEX buffer [50 mM Tris/HCl (pH 8·0), 3 mM EDTA, 0·1% (v/v) Triton X-100] and held on ice for 30 min. After centrifugation (10000 g, 10 min at 4 °C), the supernatant was carefully removed and another volume of TEX buffer was added without resuspending the pellet. The supernatant of a second centrifugation was combined with the first to yield the periplasmic fraction, and the pellet was resuspended in SDS-PAGE loading buffer to yield a volume of the cellular fraction that was equivalent to that of the periplasmic fraction.
Sucrose-floatation density-gradient centrifugation.
Sucrose-floatation density-gradient centrifugation was performed as described previously (Anantha et al., 2000 ).
N-terminal sequencing.
Affinity-purified C-terminal hexahistidine-tagged BfpU was electroblotted onto a PVDF membrane, visualized with Sypro Ruby Red Blot Stain (Bio-Rad) and subjected to Edman degradation (Biomolecular Research Facility, University of Virginia).
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RESULTS |
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Complementation of the bfpU mutant
To verify that the phenotypes of the bfpU mutant were due to mutation of the bfpU gene, we attempted to complement the mutant with a variety of plasmids containing bfpU. We were unable to demonstrate complementation with pMSD232 containing bfpU alone cloned in the low-copy-number vector pWKS30 (data not shown). To better control BfpU expression we attempted to complement the bfpU mutant with pMSD201. This plasmid is a pBR322 derivative containing a BamHI fragment that extends from 273 bp upstream of bfpA, which includes the promoter of the bfp operon (Tobe et al., 1996 ), to include the 5' end of the sixth gene (bfpD), the locus immediately downstream of bfpU (Fig. 1
). Plasmid pMSD201 also failed to complement the mutant (data not shown). However, the same DNA fragment cloned in a lower-copy-number vector (pWKS30) to produce pMSD233 complemented the bfpU mutant to restore BfpU expression (Fig. 2
), auto-aggregation (51±41%), localized adherence (Fig. 3E
) and morphologically indistinguishable BFP production (Fig. 3F
). Note that pMSD233 does not include the entire bfpD locus, and previous studies have demonstrated that it is insufficient to complement a bfpD mutant (Anantha et al., 2000
). As a control, we introduced a similar plasmid into the bfpU mutant, except that instead of extending beyond bfpU into the next cistron, this plasmid (pRPA103) contains a BamHIXbaI fragment that ends within bfpU (Fig. 1
). In contrast to pMSD233, pRPA103 failed to restore BfpU expression (Fig. 2
) and to complement the mutant for auto-aggregation (-1±1%), localized adherence (Fig. 3G
) or BFP production (Fig. 3H
). These results verify that the failure of the bfpU mutant to exhibit localized adherence and to produce BFP is due solely to the mutation in bfpU and not to polar effects on downstream genes. We attribute the absence of complementation by the other plasmids to stoichiometric imbalances, a problem we have encountered in other mutations within the bfp operon (Anantha et al., 2000
).
BfpU does not alter the expression, processing or localization of bundlin in the membrane
To determine whether the bfpU mutant is capable of expressing pre-bundlin (the BFP pre-pilin encoded by bfpA) and processing the protein to mature bundlin, we performed Western blotting on whole-cell lysates. We found that the bfpU mutant remained capable of expressing and processing pre-bundlin in a manner indistinguishable from that of the wild-type strain (Fig. 4).
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To determine the localization of BfpU, we analysed the distribution pattern of BfpU in sucrose-floatation gradients (Fig. 5a) and found that BfpU remained in the high-density fractions that contained soluble and insoluble proteins. To distinguish between these two alternatives, we grew wild-type EPEC strain E2348/69 and mutant strain UMD922 under BFP-inducing conditions and separated the cell lysates into soluble and insoluble fractions by ultracentrifugation. The quality of the separation was confirmed by probing immunoblots against the soluble protein ß-galactosidase and the outer-membrane protein intimin. As shown in Fig. 6
, BfpU was released entirely into the soluble fraction, indicating its location in the cytoplasm, the periplasm or both of these compartments.
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DISCUSSION |
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While our experiments do not demonstrate the precise function of BfpU in the biogenesis of BFP, they do shed some light on the process. BfpU appears to have a function that is different from those of other known periplasmic proteins involved in pilus biogenesis. For example, DsbA is a periplasmic protein required for BFP and P fimbrial biogenesis. In the absence of DsbA, bundlin and the PapD chaperone are unstable and are rapidly degraded (Zhang & Donnenberg, 1996 ; Jacob-Dubuisson et al., 1994
). However, the absence of BfpU does not lead to bundlin instability nor does it alter the membrane localization of bundlin. Thus, despite its localization in the periplasm, BfpU does not appear to be a specific pilus chaperone, and thus is not analogous to PapD in the P fimbriae system (Lindberg et al., 1989
). Instead, our results suggest that BfpU is part of the machinery necessary either to form pili from pilin monomers or to extrude the pilus structure through the outer membrane. Whether BfpU interacts with the outer-membrane protein components BfpB and BfpG (Schmidt et al., 2001
) of the BFP assembly apparatus or forms a bridge between inner-membrane proteins such as BfpE (Blank & Donnenberg, 2001
) and outer-membrane proteins is also amenable to experimental investigation. Nevertheless, these interactions are either very weak or exist only temporarily during the biogenesis process, as no BfpU was detected in the inner- or outer-membrane fractions in sucrose-floatation gradients and no other Bfp proteins co-purify with BfpU. One indication that such interactions may occur is our observation that the cellular abundance of BfpU is reduced by approximately 75% in a bfpB mutant strain, suggesting that BfpB may stabilize BfpU (data not shown).
We calculated that there are on average approximately 4x105 molecules of bundlin and 9x103 molecules of BfpU per EPEC cell when the bacteria are grown under the optimal conditions for BFP expression. This number of bundlin molecules would account for approximately 6·5% of the total cellular protein. This estimate appears to be accurate, since we calculated that a crude pilus preparation containing 2550% bundlin is enriched fivefold for bundlin (data not shown). Thus, there are approximately 40 molecules of bundlin for each molecule of BfpU. If one assumes that there is one biogenesis apparatus per pilus (with the pilus protruding through the secretin), and more than 40 bundlin monomers per pilus, this estimate indicates that there is more than one molecule of BfpU per pilus-assembly machine.
It is not yet clear whether the results we report herein extend to other type IV pilus systems. No proteins with primary amino-acid sequences similar to that of BfpU have been described in other pilus biogenesis or related systems. Although the Vibrio cholerae TcpH, TcpQ and TcpS proteins are all similar in size to BfpU, and like BfpU they contain signal peptidase I leader sequences and have been proposed to be periplasmic (Iredell & Manning, 1994 ), these proteins share no significant sequence similarities with BfpU. Similarly, the PilM protein from the R64 thin (type IV) pilus (Kim & Komano, 1997
) and CofG from CFA/III of enterotoxigenic E. coli (Taniguchi et al., 2001
) are predicted to be periplasmic proteins of roughly the same size as BfpU, but without sequence similarities. It has been established that PilM is necessary for R64 thin pilus biogenesis (Yoshida et al., 1999
), but it remains to be determined whether any of these proteins is a functional homologue of BfpU. Interestingly, the R64 thin pilus, CFA/III and TCP are the three type IV pilus systems most closely related to the BFP system (Spangenberg et al., 1997
; Kim & Komano, 1997
). In each of these so-called class B type IV pilus systems the genes are arranged in a tandem cluster, the pilin proteins contain long leader peptides and the amino-terminal residue of the mature pilin is not phenylalanine. Despite these similarities, we found that the BFP apparatus is not capable of assembling TCP (McNamara & Donnenberg, 2000
). It is tempting to speculate that the differences between BfpU and the putative periplasmic TCP proteins could be related to specificity in pilus assembly. The fact that similar periplasmic proteins have not been identified in the class A type IV pilus systems could reflect differences in the mechanisms of pilus biogenesis of these classes. Alternatively, our ignorance of such proteins in class A type IV systems could simply reflect the fact that the genes responsible for type IV pilus biogenesis in these systems are not clustered and have yet to be fully catalogued.
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ACKNOWLEDGEMENTS |
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Received 4 March 2002;
revised 4 April 2002;
accepted 19 April 2002.