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Dipartimento di Scienze e Tecnologie Biomediche, Sez. Microbiologia Medica, Università di Cagliari, Via Porcell, 4, 09100 Cagliari, Italy,1 Dipartimento di Biologia, Università "Tor Vergata," Via della Ricerca Scientifica, I-00133 Roma, Italy,2 Istituto di Biologia e Patologia Molecolare CNR, Roma, Italy,3 Dipartimento di Citomorfologia, Università di Cagliari, Monserrato, Cagliari, Italy,4 Dipartimento di Bioinformatica, SharDna Life Sciences, 09100 Pula, Cagliari, Italy5
Received 30 July 2006/ Accepted 2 November 2006
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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These proteins include the cell division initiator proteins FtsZ and FtsA, which are required at the early stages of the process (25, 29, 32), and some of the later proteins, DivIB/FtsQ, DivIC/FtsB, FtsL, FtsW, PBP 2X, and PBP 1A (29, 32, 33, 38), which are the septal markers for S. pneumoniae cells. Recent studies have confirmed that, overall, the major events in septation are conserved in S. pneumoniae. However, other aspects related to the division process, such as the associated morphological changes, the correct choice of the division site, and proper chromosome segregation, and the factors that regulate these aspects remain largely unknown.
We have described characterization of a chromosome region in S. pneumoniae, downstream of the ftsZ gene, that is well conserved among gram-positive bacteria and is physically and transcriptionally related to the division and cell wall (dcw) cluster. We showed that functional inactivation of each of the five genes in the region resulted in defects in cell morphology, chromosome segregation, and/or cell division (13), and the importance of these genes in other species has been confirmed (18, 23, 30). In S. pneumoniae, the most dramatic effect on division was observed after inactivation of a gene designated divIVA, after its closest homolog in B. subtilis (13, 29).
Clear orthologs of DivIVA are found in a wide range of gram-positive bacteria and in other phylogenetically distinct species, including Deinococcus radiodurans and cyanobacteria. Multiple-sequence alignment of the DivIVA proteins has shown that despite great variation in length, the N-terminal part of DivIVA is significantly conserved, while the C-terminal part is much more varied. However, the part C-terminal contains predicted repetitive coiled-coil regions, supporting the notion that DivIVA is a coiled-coil protein (11).
Despite the significant degree of sequence similarity of the DivIVA proteins, studies of the physiological role of DivIVA in cell division and related processes have not revealed a unified function. Indeed, in B. subtilis DivIVA (DivIVABS) was proposed to be the equivalent of the missing MinE determinant and was shown to be involved in division site selection, attracting the MinCD cell division inhibitors away from midcell (3, 10, 28). Other studies have shown that DivIVABS also has a second, quite distinct function during sporulation, in which it is involved in chromosome segregation, attracting another complex that consists of the chromosome origin and the DNA-binding protein RacA together with Spo0J and Soj (1, 48, 51). In Synechococcus elongatus, which has both MinE and DivIVA, disruption of minE resulted in filamentation, while disruption of divIVA resulted in cells that were two or three times longer than the wild-type cells and exhibited reduced frequency and misplacement of the Z ring (30).
Gene inactivation or depletion in other species that lack MinC, MinD, and MinE homologs has revealed a variety of other phenotypes. In Streptomyces coelicolor and Brevibacterium glutamicum, DivIVA has been shown to be essential for growth, and both protein depletion and overexpression studies have shown that its function is related to polar growth and morphogenesis (14, 44). In S. pneumoniae, disruption of divIVA resulted in the formation of chains of unseparated, morphologically altered cells with incomplete septa, often devoid of nucleoids. This complex phenotype suggested that DivIVA has a role in cell shape, septum assembly, and completion, as well as chromosome segregation through an unknown mechanism (13). A similar phenotype was observed for Enterococcus faecalis when DivIVA was depleted (43). Finally, in Staphylococcus aureus, a mutant in which the divIVA gene was inactivated did not have any distinctive phenotype related to morphology, growth, chromosome partitioning, and division, suggesting that in this species DivIVA is dispensable (42).
In agreement with the proposed function(s) in the different host backgrounds, localization of DivIVA by immunofluorescence and/or by green fluorescent protein (GFP)-DivIVA fusion revealed that the protein localizes at the cell division site and is retained or localizes directly at the cell poles (10, 11, 14, 44), except in S. aureus, where it localizes only at the cell center (42).
An additional characteristic of DivIVABS is its ability to form oligomers in vitro (34, 46), and E. faecalis DivIVA (DivIVAEF) can self-interact in vivo (43). Nothing is known about the interaction(s) of DivIVA with proteins other than itself.
In this paper, we report the cytological and interaction properties of DivIVASPN. We found that in addition to its unique pattern of localization simultaneously at the cell center and the cell poles, DivIVASPN interacts with a number of different proteins and that some of these interactions are crucial for its function.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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E. coli strain DH5
was used as a host for cloning, and E. coli strain BL21(DE3)AI (Invitrogen) was used as a host to overexpress the DivIVASPN fusion protein. Luria-Bertani (LB) broth and agar supplemented with ampicillin (150 µg/ml), when required, were used for routine growth of E. coli at 37°C.
E. coli strain R721 was used as a host for the two-hybrid assay. LB broth and agar were used for bacterial cultures and plating, and suspension medium was used for dilution as described previously (7, 8). Ampicillin (50 µg/ml), chloramphenicol (34 µg/ml), and kanamycin (30 µg/ml) were added when necessary.
Plasmid construction.
For cloning and expression purposes, the divIVASPN gene was amplified by PCR from S. pneumoniae Rx1 chromosomal DNA (13), using the spnF_divIVA and spnR_divIVA primers (Table 1); the only difference was the incorporated restriction sites used for cloning, BamHI (forward primer) and EcoRI (reverse primer). The amplified 848-bp DNA fragments were purified, double digested with BamHI and EcoRI (Roche Molecular Biochemicals, Mannheim, Germany), repurified, and ligated to the BamHI-EcoRI-digested pRSETA vector (Invitrogen). The ligation mixture was transformed into E. coli DH5 competent cells (Invitrogen), and transformants were selected after overnight growth on LB medium plates with ampicillin. Clones containing recombinant plasmid pRSETA_divIVASPN were verified by restriction analysis and DNA sequencing of the inserts and were used as sources of plasmid DNA to transform E. coli strain BL21(DE3)AI (Invitrogen) to produce the corresponding protein fused to a His tag at the N-terminal end (see below).
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Finally, the recombinant plasmids used in the E. coli two-hybrid assays were constructed by cloning the genes of interest into the pcIP22 and pcI434 vectors (7). Each gene was amplified by PCR using the specific oligonucleotides shown in Table 1 carrying at the ends compatible restriction sites for SalI (or XhoI) (forward primer) and BamHI (reverse primer). The resulting recombinant plasmids were designated pcI434(x) and pcIP22(y), where x is the gene to be tested and y is divIVA or vice versa.
Purification of DivIVASPN-His6 and production of anti-DivIVASPN antibodies. E. coli BL21(DE3)AI (Invitrogen) cells were used to overexpress the protein from plasmid pRSETA_divIVA. The pellet obtained from a 1.5-liter culture induced with 0.2% (wt/vol) arabinose for 3.5 h was disrupted as described above. Purification was performed with a Ni2+-charged column, using a His-Bind kit (Novagen) according to the manufacturer's instructions. The integrity and purity of the His-tagged DivIVA fusion were verified by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and the fusion was quantified using the Bradford method (Bio-Rad).
Polyclonal antibodies against the purified DivIVASPN protein were obtained from Custom Hybridoma.com (PickCell Laboratories, Leiden, The Netherlands). No cross-reactions in any of the preimmune sera were observed with any S. pneumoniae or E. coli protein. The DivIVA protein was detected in streptococcal cells by Western blotting as previously described (25). S. pneumoniae cells were cultured in TSB, and the growth was monitored turbidimetrically until the optical density at 650 nm (OD650) was 0.45. Cultures (100 ml) were rapidly chilled in an ice bath, centrifuged with a Sorvall centrifuge (20,000 x g, 15 min, 4°C), washed once in 10 mM phosphate (pH 7.0), and resuspended in 1 ml of the same buffer. Cells were disrupted mechanically using glass beads and a Hybaid Ribolyser (Hybaid Ltd., Ashford, United Kingdom) according to the manufacturer's instructions. After cells were broken, they were mixed 1:1 with Laemmli buffer (Bio-Rad) and boiled for 5 min. Crude extracts were stored at –20°C until they were used. Proteins were separated on 12.5% Criterion precast gels (Bio-Rad) and transferred to a nitrocellulose membrane (Protran BA83; Schleicher & Schuell) at 100 V for 30 min with a Criterion Blotter apparatus (Bio-Rad). After transfer, the membranes were stained with Ponceau S (Sigma-Aldrich) and then incubated in 5% nonfat dry milk in PBST (phosphate-buffered saline [PBS] containing 0.05% Tween) overnight at 4°C. After blocking, membranes were incubated with an anti-DivIVA antibody solution (1:25,000) in PBST for 1 h at room temperature and, after washing, with horseradish peroxidase-conjugated antibodies (Bio-Rad). FtsA and FtsZ antibodies were used as previously described (25). Chemiluminescent bands were detected using an Immun-Star horseradish peroxidase chemiluminescence kit (Bio-Rad) and Kodak Biomax light film.
Cellular immunolocalization of the DivIVA protein. For immunofluorescence, exponentially growing cells were fixed and treated as described by Lara et al. (25). Cells were then transferred onto poly-L-lysine-coated slides (Sigma). The slides were washed twice with PBS, air dried, dipped in methanol at –20°C for 10 min, and allowed to dry. After rehydration with PBS, the slides were blocked for 1 h at room temperature with 2% (wt/vol) bovine serum albumin (BSA) and 0.2% Triton X-100 (vol/vol) in PBS (BSA-PBST) and for 1 h with appropriate dilutions of anti-DivIVA or other antibodies in BSA-PBST. The slides were then washed five times with PBST and incubated for 30 min with a 1:500 dilution of anti-rabbit immunoglobulin (IgG) Alexa Fluor 488 (Invitrogen, Molecular Probes) or with a 1:500 dilution of anti-mouse IgG Alexa Fluor 488 or Alexa Fluor 594 in BSA-PBST. To check the specificity of the secondary antibodies, incubation with primary antibodies was omitted in negative controls. Preparations were finally stained with a fluorescence antifade solution containing propidium iodide (0.5 µg/ml) or 4',6-diamidino-2-phenylindole (DAPI) (0.2 µg/ml) and 2% (wt/vol) 1,4-diazabicyclo[2.2.2] octane (DABCO), all obtained from Sigma. Slides were observed using a Zeiss Axioplan 2 equipped with a 100x Achroplan and fluorescence objective and standard filter sets (Zeiss no. 01, no. 09, no. 15, and no. 24). Photographs were taken with a Canon Powershot G6 digital camera, acquired with a Canon Zoom Browser, and processed with Adobe Photoshop 6.0.
A double-immunofluorescence analysis was performed as described by Morlot et al. (32), using polyclonal rabbit anti-DivIVA antibodies in combination with polyclonal mouse anti-FtsZ antibodies, kindly provided by T. Vernet.
For immunoelectron microscopy, S. pneumoniae Rx1 cells were cultured in TSB and growth was monitored turbidimetrically until the OD650 was 0.6. Cultures (4 ml) were rapidly chilled in an ice bath, centrifuged (10,000 x g, 15 min, 4°C), washed in 10 mM phosphate (pH 7.0), and fixed in 1 ml of a solution containing 2% paraformaldehyde and 0.05% glutaraldehyde in 1x PBS for 4 h at 4°C. The fixed cells were washed with PBS and stored at 4°C until they were used. Small pellets of fixed cells were cryoprotected with glycerol, applied to small pieces of filter paper, and quickly frozen in liquid ethane. Frozen specimens were transferred to a Reichert-Jung AFS freeze-substitution unit (Leica, Vienna, Austria) for 48 h at –90°C in a mixture of methanol and 0.5% (wt/vol) uranyl acetate for complete substitution of the water in the sample. After freeze-substitution, samples were infiltrated using Lowicryl K4M (EML Laboratories, Berkshire, United Kingdom) at 30°C and polymerized with UV light. Ultrathin sections of the samples were collected on 200-mesh gold grids covered with Formvar and carbon and processed for immunogold labeling as follows. After 40 min of blocking with Tris buffer-gelatin (TBG) (30 mM Tris-HCl [pH 8.0], 150 mM NaCl, 0.1% bovine serum albumin, 1% gelatin), sections were incubated for 1 h in TBG containing anti-DivIVA (1:750) and then washed with PBS. Next, grids were floated on three drops of TBG and incubated for 10 min on the last drop before a 45-min incubation with 10 nM gold-labeled goat anti-rabbit IgG (1:40) in TBG. The grids were then washed in PBS and distilled water before they were stained with saturated uranyl acetate for 30 min, followed by lead citrate for 1 min. Images were collected with a JEOL 1200-EXII electron microscope (Zeiss) operating at 100 kV. Electron micrographs were scanned using an Epson Stylus PHOTO 890 scanner and the Picture Publisher 8 software.
Bacterial two-hybrid assay. All the different combinations of recombinant plasmids coding for the chimeric repressors, obtained as described above, were cotransformed into the recipient E. coli strain R721, and a ß-galactosidase assay was performed as described previously (8). Bacterial cultures were grown at 34°C in Luria-Bertani medium supplemented with 0.1 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) to an OD600 of 0.5, and the residual ß-galactosidase activity was evaluated for each strain. E. coli R721 without plasmids and E. coli R721 with plasmids pcI434434 and pcIP22434 were used as negative and positive controls, respectively. Plasmids pcIP22(divIVA)/pcI434434 and pcI434(divIVA)/pcIP22434 were used as additional negative controls to exclude self-interactions. Residual ß-galactosidase activity that was less than 50% indicated that there was repression and hence a protein-protein interaction, whereas when the activity was greater than 50%, the interaction was uncertain or null. The use of these values has been discussed previously (8).
Coimmunoprecipitation.
Cultures of the E. coli DH5 strain containing the pairs of recombinant plasmids to be tested [pTTQ18(gfp) and p434(cI) derivatives] were grown overnight in LB medium supplemented with the appropriate antibiotic and then diluted 100-fold in fresh prewarmed medium. To induce expression, 0.1 mM (final concentration) IPTG was added to a culture, and the culture was incubated for 4 h. After this, the cells were centrifuged, washed and resuspended 1/300 (vol/vol) in lysis buffer (1 mM EDTA [pH 8], 25 mM HEPES [pH 7.6], 0.1 mg/ml lysozyme), and cooled on ice for 30 min. Membrane fractions were then prepared as described by Buddelmeijer and Beckwith (2). Soluble fractions were prepared by centrifuging a sample at 50,000 x g for 45 min following lysis and recovering the supernatant. Proteins concentrations were determined by the Bradford method.
The immunoprecipitation experiments were similar to those described by Duong and Wickner (9). The assays were performed using commercial antibodies, whose specificity was determined. The beads (protein A-Sepharose) were divided into 50- to 100-µl aliquots in microcentrifuge tubes, and 10 µl of anti-GFP mouse monoclonal antibodies (1:100; Santa Cruz Biotechnology, Inc.) was added to each tube and incubated for 15 to 60 min at room temperature with gentle mixing on a shaker. After centrifugation, samples were washed with 1 ml washing buffer (20 mM HEPES buffer [pH 7.5], 150 mM NaCl, 0.1% Triton X-100, 10% glycerol). Then 0.1 to 1 ml of cell lysate (corresponding to about 1 mg of protein) was added to each tube. Samples were incubated for 90 min to overnight at 4°C with gentle mixing on a shaker. Immunoprecipitated complexes were collected by centrifugation at 3,000 x g for 2 min at 4°C. Electrophoresis and Western blotting were performed as described above. The nitrocellulose membranes were probed with anti-cI rabbit polyclonal antibodies (1:6,000) and detected with an ECF Western blotting kit (Amersham Biosciences).
Allelic replacement mutagenesis.
S. pneumoniae genomic DNA isolated from strain Rx1 as previously described (13) was used as a template for PCR experiments. Constructs for allelic replacement were obtained by a two-step PCR method similar to the method described previously for gene insertion/deletion mutagenesis (13, 25). However, in these experiments four sets of primers (PF1-PR2, PF3-PR4, PF5-PR6, and PF7-PR8 [sequences available on request]) were used for each construct. The desired T
C mutation at nucleotide position 232, which resulted in an A78T substitution in the recombinant protein, was introduced into the PF3 primer. Two additional substitutions (A C and GA at positions 229 and 230, respectively), resulting in silent changes, were also introduced in order to saturate and inhibit the mismatch repair system (4). PCR-derived fragments were purified using a QIAquick PCR purification kit (QIAGEN), mixed at a ratio of 1:1:1:1, and reamplified in a second PCR, using the external primers. Constructs for gene mutagenesis, obtained as described above, were used to transform S. pneumoniae Rx1 competent cells (13). Transformants were selected after 24 h of growth on tryptone soya blood agar plates containing chloramphenicol. The presence of the mutation was confirmed by PCR using specific primers, by a restriction digestion profile analysis of the PCR products using the PvuII restriction enzyme, and finally by sequence analysis of the entire constructs. As a control, a wild-type transformant containing the cat cassette inserted at same position was constructed by using the same procedure.
Growth studies, viability, and microscopy. For physiological studies, bacterial cultures were inoculated from glycerol stocks into prewarmed TSB and incubated at 37°C in an atmosphere containing 5% CO2. Growth was monitored turbidimetrically at 650 nm every 30 min for 9 h with an Ultrospec 3100 (Amersham Pharmacia Biotech). Viable counts were determined every 90 min by serial dilution of the cultures and spot plating 20 µl on tryptone soya agar plates supplemented with 5% defibrinated sheep blood. At selected times during the exponential phase of growth, samples (100 µl) were transferred into Eppendorf tubes and fixed with 1% formaldehyde for 15 min at room temperature. Aliquots (10 µl) were transferred to poly-L-lysine-coated slides to which 2 µl of DAPI (0.5 µg/ml) was added. Samples were incubated for 5 to 10 min at room temperature in the dark before microscopic examination using a Zeiss Axioskop HBO 50 equipped with 100x phase-contrast and fluorescence objectives. To determine the presence of dead cells in the various strains, exponentially growing unfixed cells were washed and resuspended in 10 mM phosphate (pH 7.0) and then treated with propidium iodide (3 µg/ml) and DAPI (0.5 µg/ml). Cells were then incubated at room temperature for 15 min in the dark, transferred to poly-L-lysine-coated slides, and examined with the microscope as described above. Photographs were taken with a Zeiss MC100 Spot camera.
For scanning electron microscopy S. pneumoniae cells were cultured in TSB and growth was monitored turbidimetrically until the OD650 was 0.45. Cultures were rapidly chilled in an ice bath, centrifuged (10,000 x g, 15 min, 4°C), washed in 10 mM phosphate (pH 7.0), and fixed in 1% paraformaldehyde and 1.25% glutaraldehyde in 0.15 M sodium cacodylate buffer for 4 h at room temperature. Fixed cells were washed with PBS, transferred onto a circular cover glass, and dehydrated using increasing concentrations of acetone; this was followed by critical-point drying using CO2. Samples were then coated with platinum using an Emitech 575 turbo sputtering apparatus and examined with an FE Hitachi S4000 scanning electron microscope operating at 15 to 20 kV.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The subcellular localization of DivIVA in S. pneumoniae Rx1 cells was then examined by both immunofluorescence microscopy and immunoelectron microscopy using exponentially growing cells that were fixed and processed as previously described (25). Using both techniques, DivIVA was observed in more than 80% of the cells at the cell division septum and the cell poles simultaneously (Fig. 1).
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Immunoelectron microscopy confirmed the pattern of localization observed by immunofluorescence microscopy. A representative dividing cell is shown in Fig. 1B. The distribution of the gold beads in the 360 cells examined showed that around 50% of the beads were at midcell and 24% were at the poles, primarily localized near the membrane. About 10% of the beads were outside the cells, and the remaining beads (around 16%) were observed in the cytoplasm, occasionally in a central position.
Differential localization and timing of DivIVA with respect to FtsZ during septation. In dividing S. pneumoniae cells, FtsZ is targeted to the septum at early stages of cell division and remains there throughout the process (25, 32). Some cell division proteins, such as FtsA, exhibit the same pattern of localization (25; unpublished results), while others, such as PBP 2X, PBP 1A, FtsW, and DivIC, are targeted to the division site after FtsZ or, as is the case for DivIB/FtsQ and FtsL, are present transiently only during septation (32, 33, 38).
To determine the relative timing of assembly of DivIVASPN with respect to FtsZ during the cell cycle, we used double immunofluorescence with rabbit anti-DivIVA and mouse anti-FtsZ polyclonal antibodies, combined with DAPI nucleoid staining. Representative cells immunostained for FtsZ, DivIVA, FtsZ/DivIVA, and nucleoids are shown in Fig. 2. Six distinguishable patterns, in agreement with the patterns obtained previously for single localization, were observed. Immunocolocalization experiments showed that DivIVA is recruited to the septum at a later stage than FtsZ. Using the reconstruction of Morlot et al. (32) and assuming that all the cells were growing and dividing, the percentage in each stage was related to the duration of the phase. For cells growing with a generation time of 32 min, a delay of about 5 min was observed between FtsZ localization and DivIVA localization. Consistently, when FtsZ was already localized to the septum or relocalized to the equator of the daughter cells, DivIVA was still visible at the center and retained at the poles, and it was subsequently recruited to the future division sites only after the Z ring was assembled (Fig. 2).
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Unexpectedly, we observed no structurally intact anucleate cells in the immunofluorescence images of the divIVA null mutant, which we anticipated from a previous characterization of this mutant (13), suggesting that such cells could have been lost during the immunofluorescence procedure. This hypothesis was tested by performing a live/dead assay with unfixed, exponentially growing cells, which revealed that the anucleate cells observed in the divIVA null mutant (corresponding to 15% to 20% of the population) were indeed dead cells in the process of lysing (see below).
S. pneumoniae DivIVA protein interacts with itself and with other proteins of the septation machinery. The septal localization of DivIVASPN suggested that the protein could interact with other components of the streptococcal division machinery; however, polar localization did not exclude the possibility that there were interactions with other proteins involved in chromosome segregation. To determine the interaction partners of DivIVASPN, we used a bacterial two-hybrid assay that is a powerful system for analyzing the protein-protein network of the E. coli components of the divisome (8).
Among the most prominent candidates, we tested the cell division proteins that are known or are thought to participate in divisome formation in S. pneumoniae, like FtsZ, FtsA, FtsK, FtsL, FtsQ/DivIVB, FtsB/DivIC, FtsW, PBP 2X, and PBP 1A (25, 32, 33, 38), as well as homologs of proteins that have been shown to have a role in division in other bacteria, like ZapA (17) and EzrA (26). In addition, we tested interactions with two other proteins, PcsB and LytB; PcsB is an essential putative murein hydrolase, whose precise function remains unclear (35, 36, 37), while LytB is a endo-ß-N-acetylglucosaminidase involved in the separation of daughter cells (6). Finally, we examined the possible interactions with the homolog of Spo0J (15) and PfrA (SP1020/Spr0924), whose orthologs are involved in chromosome segregation in B. subtilis (22, 39).
The divIVASPN gene was fused in frame with the segment encoding the N-terminal portion of the phage 434 cI repressor (cI434-divIVA), while the various genes tested were fused in frame with the segment encoding the N-terminal portion of the phage P22 cI repressor (cIP22-y), as described in Materials and Methods. Pairs of the resulting recombinant plasmids, expressing the chimeric repressors, were then cotransformed into the reporter strain E. coli R721 carrying the lacZ gene under control of the hybrid 434-P22 promoter/operator, and ß-galactosidase activity was measured as described previously (8).
As shown in Table 2, DivIVASPN was able to interact with itself. Moreover, high levels of interaction were observed with the cell division initiator proteins FtsZ, FtsA, and ZapA and also with FtsK and FtsL, as well as with the chromosome segregation protein Spo0J and the putative morphogenic determinant PcsB. Lower, but still significant, levels of interaction were detected with EzrA, DivIC/FtsB, DivIB/FtsQ, FtsW, and LytB. Finally, no interactions between DivIVA and PBP 2X, PBP 1A, or PrfA were detected.
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repressor (cI) or GFP, cloned and expressed in E. coli, and using commercial antibodies. Figure 4 shows the results of the coimmunoprecipitation experiments, which confirmed that there were interactions between FtsZ and DivIVA and between Spo0J and DivIVA (Fig. 4A) and that there were not interactions between DivIVA and PfrA (Fig. 4B), as detected by the two-hybrid assay. No cI-tagged proteins were detected in the pellets of control strains lacking GFP-tagged proteins (data not shown).
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DISCUSSION |
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Double-immunofluorescence analysis using anti-FtsZSPN antibodies and anti-DivIVASPN antibodies showed that DivIVA is recruited to the septum about 5 min (i.e., between one-sixth and one-seventh of the generation time) after FtsZ is recruited (Fig. 2). All the other cell division proteins that we tested were localized in the divIVA null mutant, indicating that they do not require DivIVA for localization and that, on the contrary, DivIVA may be dependent on them. However, proper placement of FtsZ and DivIVA in other cell division null mutants of S. pneumoniae, like Rx1 ftsK::cat and ftsQ::cat, which are viable (O. Massidda, unpublished results), suggests either that the formation of the Z ring alone is sufficient to target DivIVA to the septum and then to the new forming poles or that DivIVA is recruited to these sites independently. The presence of fluorescence at the division septa and, in particular, at the poles of E. coli cells expressing DivIVASPN-GFP (O. Massidda and P. Ghelardini, unpublished observations), as seen previously for DivIVBS-GFP (11), may be consistent with divisome-independent polar targeting, although the possibility of heterologous interactions with E. coli septal components cannot be excluded at this time.
While the localization profile observed for DivIVASPN was consistent with what was observed for the protein in B. subtilis, it was even more puzzling given the absence of both the MinC/MinD system and sporulation. On the one hand, septal localization strongly suggested that the protein could be necessary at the septum, raising the possibility that it interacts with other components of the streptococcal division machinery. On the other hand, polar localization did not allow us to exclude other functions, like chromosome segregation.
The ability of DivIVABS to oligomerize in vitro, forming elongated particles, the so-called "doggy-bone" structures, has been described previously (34, 46). Moreover, DivIVAEF was recently found to self-interact in vivo (43). In addition, it has been proposed that DivIVA could also interact with one or more components of the cell division machinery, as well as with other proteins (1, 11, 13, 34, 43, 51). Very recently, some interactive partners for DivIVABS have been determined (41). Therefore, as a step to further characterize DivIVASPN, we tested a number of different proteins using a two-hybrid assay.
As anticipated, DivIVASPN interacted with itself, indicating that it should at least dimerize efficiently. Moreover, in accordance with its localization profile and coiled-coil nature, DivIVASPN interacted directly or indirectly with a number of different proteins (Table 2). Although some interactions could be expected, we were surprised by the large number of them.
DivIVASPN interactions were detected with most components required at the early or late stage of septosome assembly but not with the septal peptidoglycan biosynthetic enzymes, PBP 2X and PBP 1A. However, DivIVASPN interacted with PcsB and LytB (6, 35, 36, 37), which were included because of their likely role in streptococcal cell division and because of the divIVA null phenotype. Inactivation of LytB resulted in chains of morphologically normal cells, consistent with the polar localization of this protein and its proposed function in daughter cell separation (6). In S. pneumoniae pcsB is essential, but reduced expression also results in chains of cells with abnormal morphology (36) that resemble the divIVA null cells remarkably. PcsB is a putative peptidoglycan hydrolase, conserved among streptococci, that is believed to be involved in cell growth and morphology, although an enzymatic activity on peptidoglycan has not been demonstrated yet (35, 36). Interestingly, LytB and PcsB are thought to cooperate in maintaining membrane and cell wall integrity and are positively regulated by the essential YycFG/VicRK two-component system (31, 35, 36, 37). We also found that DivIVASPN interacts with Spo0J, the only ortholog of the Spo0J/Soj/RacA complex of B. subtilis that could be identified in S. pneumoniae. Interactions between DivIVA and Spo0J were somewhat expected, given the chromosome segregation defects of both S. pneumoniae and E. faecalis that lacked divIVA or in which divIVA was depleted. Additionally, this interaction was postulated to occur in B. subtilis, certainly for the two proteins during sporulation and possibly also during vegetative growth (40, 51). Interestingly, Perry and Edwards have demonstrated that in B. subtilis DivIVABS coimmunoprecipitates with FtsZ and MinD and that a mutant allele, DivIVABSR18C, also coimmunoprecipitates with Spo0J (41).
To understand the biological meaning of the DivIVASPN interactions, we generated a missense isogenic S. pneumoniae DivIVA A78T mutant and compared it with the isogenic DivIVA wild-type strain in terms of growth, morphology, cellular localization, and interaction profile. The DivIVA A78T mutant was of particular interest since it has been shown that in B. subtilis this amino acid substitution correlates with cell division defects, possibly because it is located in the hydrophobic core of the first predicted coiled-coil domain (3, 10, 34). The DivIVA A78T mutant had an intermediate phenotype compared with the phenotypes of the DivIVA wild-type strain and the null mutant (Fig. 6), suggesting that DivIVA A78T is at least partially functional. Immunofluorescence revealed an altered localization profile, and although the protein was still visible at the septum and the poles, the majority was located diffusely around the cell. The two-hybrid assay showed that DivIVASPN A78T was able to self-interact, consistent with the finding of Muchova et al. (34) that DivIVABS A78T oligomerizes, and this suggests that oligomerization per se is not sufficient for the function of the protein.
Some of the interactions were also observed with the A78T protein, while others were significantly reduced or absent (Table 2), suggesting that they are crucial for proper function. The interaction profile of DivIVASPN A78T correlates well with the S. pneumoniae DivIVA A78T mutant phenotype. The DivIVA-PcsB and DivIVA-Spo0J interactions, both of which occurred in the A78T mutant, appear to be sufficient to reduce and eliminate, respectively, the cell morphology and nucleoid segregation defects evident in the null mutant. On the other hand, the DivIVA-LytB interactions, which did not occur with the mutated protein, could explain the chainy phenotype. Finally, while the interactions with the early cell division proteins may correlate with some localization of DivIVA A78T seen at the expected sites, the complete absence of interactions with the other division components that follows may be responsible for the abnormal localization of this mutant, although we cannot exclude other possible explanations.
In agreement with the original phenotype of the S. pneumoniae divIVA null mutant (13), these results indicate that DivIVA has a multifaceted role in controlling cell morphology and completion of cell division and separation, as well as chromosome segregation, through a complex interaction web. We speculate that DivIVA is a cytoplasmic cytoskeleton-like element that acts like a scaffold to direct cross wall formation and ultimately the formation of mature poles. Additionally, since this protein is stably present at the poles, it is not surprising that it may also cooperate with a putative segregation system that includes Spo0J to ensure proper chromosome partitioning in each daughter cell. Interestingly, a similar scaffolding role was postulated for DivIVA in S. coelicolor (14).
One possible model is that DivIVASPN arrives at the septum at some point during septation and assembles with the rest of the cell division proteins. Once there, DivIVA then ensures correct division through positioning the peptidoglycan hydrolytic enzymes necessary for septum splitting and possibly the late cell division proteins, determining the formation and the maturation of the distinctive pointed streptococcal poles. In the absence of DivIVA, this organization is lost, resulting in cells whose shape and diameter are altered, in which septum assembly and closure are delayed and (in some cells) chromosome segregation is impaired. In support of this is the fact that streptococci have rigid metabolically inert poles that are necessary to maintain cell shape (5) and the fact that the divIVA null cells form clearly defective poles (Fig. 7) (13). This view agrees well with the model for streptococcal growth and division of Higgins and Shockman (20) that predicts that in dividing cells, after initial inward growth of the cross wall, centripetal penetration remains relatively constant until the two newly synthesized internal hemispheres have reached the size of the external hemispheres. Moreover, in a subsequent study, the area of a streptococcal pole was found to be larger than the area expected for a pole formed from a flat, although complete, septum, suggesting that stretching alone is not sufficient to form a complete pole (21). This led to Koch's proposal that to form a prolate pole, like that in streptococci, new cell wall material should be added, in addition to stretching, in accordance with a "split and splay" model rather than the "split and stretch" model proposed for bacilli (24).
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ACKNOWLEDGMENTS |
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We thank Loredana Onidi for small-scale purification of the DivIVASPN protein and Stefania Collu for assistance with immunofluorescence. We thank A. Riva, Dipartimento di Citomorfologia, Università di Cagliari, for his help with the electron microscopy and Miguel Vicente for helpful comments. We thank Luciano Paolozzi for his constant support and advice. Finally, we thank Thierry Vernet for the generous gift of anti-FtsZ and other polyclonal antibodies and for lively discussions.
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FOOTNOTES |
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Published ahead of print on 10 November 2006.
Supplemental material for this article may be found at http://jb.asm.org/.
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