Influence of core divisome proteins on cell division in Streptomyces venezuelae ATCC 10712

Abstract

The sporulating, filamentous soil bacterium Streptomyces venezuelae ATCC 10712 differentiates under submerged and surface growth conditions. In order to lay a solid foundation for the study of development-associated division for this organism, a congenic set of mutants was isolated, individually deleted for a gene encoding either a cytoplasmic (i.e. ftsZ) or core inner membrane (i.e. divIC, ftsL, ftsI, ftsQ, ftsW) component of the divisome. While ftsZ mutants are completely blocked for division, single mutants in the other core divisome genes resulted in partial, yet similar, blocks in sporulation septum formation. Double and triple mutants for core divisome membrane components displayed phenotypes that were similar to those of the single mutants, demonstrating that the phenotypes were not synergistic. Division in this organism is still partially functional without multiple core divisome proteins, suggesting that perhaps other unknown lineage-specific proteins perform redundant functions. In addition, by isolating an ftsZ2p mutant with an altered −10 region, the conserved developmentally controlled promoter was also shown to be required for sporulation-associated division. Finally, microscopic observation of FtsZ-YFP dynamics in the different mutant backgrounds led to the conclusion that the initial assembly of regular Z rings does not per se require the tested divisome membrane proteins, but the stability of Z rings is dependent on the divisome membrane components tested. The observation is consistent with the interpretation that Z ring instability likely results from and further contributes to the observed defects in sporulation septation in mutants lacking core divisome proteins.

Introduction

Since the 1960s, Streptomyces coelicolor A3(2) has been used as the model system to study the filamentous sporulating soil bacteria in the genus Streptomyces [1]. The streptomycetes are true mycelial organisms of ecological importance, and are responsible for the synthesis of many important biologically active compounds, including a wide range of antibiotics [2]. To facilitate dispersal in the environment, and to enable long-term survival, streptomycetes produce semi-dormant spores. Spores are formed as part of a complex developmental life cycle, in which the streptomycete initially grows vegetatively as branching hyphae while nutrients are available, forming a vegetative mycelium, also referred to as substrate mycelium. Eventually, specialized spore-forming aerial hyphae emerge on the colony surface. The apical parts of such aerial hyphae are partitioned into chains of prespores via synchronous formation of several tens of cell division septa in each hypha, and thereafter prespores mature to form pigmented and thick-walled spores with condensed nucleoids and a hydrophobic outer surface layer, before being released into the environment (reviewed in e.g. [3–5]). When conditions are favourable, spores germinate and grow to form a new mycelium. The formation of aerial mycelia and spores is governed by a regulatory cascade of transcription factors that has mainly been studied in S. coelicolor , but more recently much progress has been made in mapping the developmental regulatory networks in the new model organism Streptomyces venezuelae [3–5].

Cell division and its regulation are of central importance in the Streptomyces life cycle, which involves two distinct types of division. Both types are formed by a typical bacterial cell division machinery, organized by the tubulin homologue FtsZ [6, 7]. In vegetatively growing hyphae, cell division is infrequent and gives rise to widely spaced hyphal cross-walls. Intriguingly, vegetative hyphae can even grow in the absence of such cross-walls and cell division is not essential for proliferation of these organisms; the key cell division gene ftsZ is dispensable for growth and viability in S. coelicolor and S. venezuelae [8, 9]. The ftsZ null mutants do not make cross-walls at all, but can still grow as branching hyphae. On the other hand, cell division is absolutely essential for spore formation. The septa that divide aerial hyphae into prespore compartments are formed by the same ftsZ-dependent core cell division machinery as the vegetative cross-walls. However, sporulation septa differ from hyphal cross-walls. Structurally, sporulation septa are thicker, eventually leading to full constriction and separation of daughter cells, and sporulation septatation is subject to spatial and temporal regulation [10].

Cell division genes are developmentally regulated and directly controlled by several of the key transcriptional regulators that govern morphological differentiation and sporulation. For example, ftsZ has a developmentally regulated promoter, ftsZ2p, that is critical for spore formation [11]. This promoter is repressed by the master regulator BldD in complex with cyclic di-GMP; c-di-GMP-bound BldD negatively controls many important genes related to aerial mycelium formation and sporulation [12, 13]. In S. venezuelae , both the ftsZ2p promoter and promoters for cell division genes ftsW, ftsK and sepF2 are activated by the proteins WhiA and WhiB, which presumably act as a complex that controls a large regulon of genes involved in spore formation [14, 15]. In addition, ssgB and, indirectly, ssgA are controlled by the developmental regulator BldM [16]. SsgA and SsgB belong to an actinobacteria-specific protein family and affect the selection of septation sites in S. coelicolor [17]. Finally, two dynamin-like proteins are involved in septation, specifically in sporulating aerial hyphae, and developmental upregulation of the corresponding genes, dynA and dynB, depends on the sporulation-specific GntR-family regulator WhiH in S. venezuelae [18].

The bacterial cell division process is primarily controlled at the level of the assembly of FtsZ into cytokinetic polymers and their formation of a ring-shaped pattern, the Z ring, which serves to recruit and organize most other proteins involved in cell constriction and septum formation [19–21]. Interestingly, streptomycetes, as well as Actinobacteria in general, lack obvious homologues of most of the proteins that are known to regulate Z ring formation, including MinC, MinD, SulA and Noc, or to stabilize FtsZ polymers or tether them to the cytoplasmic membrane, like FtsA and ZipA [6, 7]. The exception is SepF, which links FtsZ to membranes and facilitates Z ring formation in Gram-positive bacteria such as Bacillus subtilis [22], and which is present as three homologues in S. venezuelae [18]. The regulation of Z ring assembly in Streptomyces remains poorly understood. During sporulation, SsgA and SsgB proteins have been reported to mark the sites of Z ring formation in S. coelicolor [17], and DynA and DynB are recruited to septation sites and help stabilize Z rings in S. venezuelae [18]. DynA and DynB interact with each other, and DynB also interacts with both SsgB and one of the three SepF proteins in S. venezuelae [18]. Analyses with two-hybrid systems have further suggested that the three SepF proteins interact with each other, SepF interacts with FtsZ and SepF2 interacts with both DynB and SsgB [18], indicating a sophisticated protein interaction network affecting Z ring assembly. Additional proteins have been suggested to affect this critical step in cell division, including SepG and CrgA, but details remain unclear [23, 24].

Once formed, the Z ring recruits further division proteins that collectively are referred to as the divisome. Present in streptomycetes are orthologues of conserved divisome proteins FtsQ, FtsL and DivIC(FtsB), which are known to form a complex with a structural and/or regulatory role in Escherichia coli and B. subtilis [19, 25, 26], and FtsW and FtsI, which encode a peptidoglycan transglycosylase of the shape, elongation, division and sporulation (SEDS) family, and a cognate penicillin-binding protein with transpeptidase activity, respectively [27]. In addition, the DNA translocase FtsK and ABC transporter proteins FtsEX are encoded by streptomycete genomes [7]. Genetic studies in S. coelicolor show that none of these proteins are essential for growth or viability, which is consistent with the finding that cell division is dispensable in streptomycetes. However, although mutants for ftsQ, ftsL, divIC, ftsW and ftsI are largely defective in spore formation, none of these genes are absolutely needed for cell division, with all mutants being able to form hyphal cross-walls and some sporulation septa [28–31]. For the last four genes, the mutant phenotypes were found to be conditional, leading to suppression of the septation defect on minimal medium or low osmolarity medium. Mutants lacking ftsK or ftsEX show apparently regular sporulation septation, but ftsK mutants have a defect in chromosome stability, presumably related to the role of FtsK in clearing trapped chromosomes from the closing septa [7, 32–34].

The fact that Streptomyces cell division is non-essential, developmentally regulated, disconnected from vegetative growth and involves previously unknown mechanisms for control of septum formation, makes streptomycetes attractive model systems to study the division process and its regulation [6, 7]. This distinction is further accentuated by the recent development of live cell imaging systems in the new model organism S. venezuelae that allow time-lapse visualization of the cell division in great detail through the entire life cycle [3, 35]. In order to further establish S. venezuelae as a cell division model system, we report the isolation and characterization of null mutants for key cell division genes ftsZ, ftsQ, ftsL, divIC, ftsW and ftsI for this organism, and we clarify the effect of late divisome components on assembly of Z rings and cell division in S. venezuelae .

Methods

Bacterial strains and growth conditions

The S. venezuelae strains used in this study were derived from S. venezuelae strain ATCC 10712, acquired from Dr Colin Stuttard (Dalhousie University, Halifax, Canada) (Table S1, available in the online version of this article). S. venezuelae strains were cultivated at 30 °C on maltose yeast extract medium (MYM) agar plates or in MYM liquid medium [36], as described by Bush et al. [14]. S. venezuelae transconjugants were selected on either MYM or R2S agar after interspecies conjugation, as described previously [37]. Culture conditions and antibiotics followed previously described procedures for streptomycetes [38]. E. coli strain TG1 was used for cloning, construction and propagation of vectors [39]. E. coli strain BW25113/pIJ790 [40, 41] was used to create cosmid derivatives containing insertion–deletion mutations. E. coli strain ET12567/pUZ8002 was used for mobilization of oriT-containing cosmids and plasmids into S. venezuelae [37, 38]. E. coli strain BT340 was used to express yeast Flp recombinase in E. coli to excise antibiotic resistance markers flanked by FRT sites [42]. Culture conditions, antibiotic concentrations and genetic manipulations generally followed those previously described for E. coli [39].

Plasmids and general DNA techniques

The plasmids and cosmids used in this study are listed in Table S2. DNA restriction and modifying enzymes were used according to the manufacturer’s recommendations (New England BioLabs). Phusion DNA polymerase (Thermo Fisher Scientific) was used according to the manufacturer’s instructions. S. venezuelae total DNA preparations were obtained using the Wizard genomic DNA purification kit (Promega). λRED-mediated recombineering, modified for Streptomyces , was used in E. coli to replace S. venezuelae genes on cosmids with mutagenic linear DNA cassettes. The apramycin-resistance gene cassette [oriT acc(3)IV] was amplified by PCR from plasmid pIJ773. The oligonucleotide primers used in this study are listed in Table S3. When necessary, the bla gene of the cosmid backbone was replaced by recombineering with a bla homology-flanked oriT acc(3)IV-cassette from pIJ799. Plasmid pMS82 was used to create genetic complementation plasmids for site-specific integration in the chromosome at the ΦBT1 attachment site [43]. DNA sequences of unmarked in-frame deletions were verified using BigDye cycle sequencing analysed on an ABI 3130 Prism Genetic Analyzer (Applied Biosystems).

Isolation of strains mutant for cell division genes

Using PCR product-directed recombineering of cosmid inserts, insertion–deletion mutations were created in which all or crucial portions of the coding regions of S. venezuelae division genes were replaced with an apramycin-resistance cassette [oriT acc(3)IV] from pIJ773 (Table S2). Care was taken when designing the 3′ endpoints of in-frame deletions to minimize the potential polar effects on expression of the downstream co-transcribed gene(s). Mutagenized cosmids were confirmed by restriction enzyme digestion and PCR amplification with primers flanking the introduced mutations. These mutagenized cosmids were introduced into S. venezuelae by interspecies conjugation and marked null mutants, generated by double homologous recombination events, were identified among primary transformants by their apramycin-resistant, kanamycin-sensitive phenotypes. Genomic DNA from mutant candidates was analysed by PCR amplification using primers flanking the mutations.

Most mutations were designed to introduce unique XbaI and SpeI sites flanking the oriT acc(3)IV cassette that was inserted into cosmids to generate the marked insertion–deletions. These restriction sites facilitated the isolation of unmarked deletion mutant strains. The mutagenized cosmids were digested with XbaI and SpeI and religated, removing the oriT acc(3)IV cassette, leaving a 6 bp in-frame scar with the sequence of ACTAGA. Alternatively, the antibiotic-resistance cassette was removed by site-specific recombination, resulting in an 81 bp frt scar for unmarked ftsZ and ftsI mutations. Subsequently, a linear oriT acc(3)IV cassette was used to replace the bla gene on the cosmid backbone, allowing conjugation into S. venezuelae and selection of exconjugants by apramycin resistance marker in the vector backbone. Primary exconjugants generated by a single homologous recombination incident were screened by PCR for gene conversion events in which resident wild-type alleles were replaced with the introduced unmarked mutagenized ones. Generally, 5–10 % of exconjugants had undergone gene conversion events. Exconjugants that were homozygous for the mutant allele were restreaked without selection to allow loss of integrated cosmids and progeny colonies were screened for apramycin sensitivity, indicating intramolecular homologous recombination events and the loss of the cosmid. All of the mutants chosen for further characterization were checked by PCR amplification from genomic DNA with primers flanking the introduced mutation to confirm the presence of only the unmarked deletion allele. Double mutants were constructed by introducing marked insertion–deletion mutations into unmarked single mutants, as described above for isolating single mutants. A triple mutant for ftsL, ftsQ and divIC was isolated in a similar fashion from an unmarked double mutant strain. A double mutant strain for the adjacent ftsL and ftsI genes was obtained by combining recombineering primers used for single mutation isolation (i.e. using the 5′ ftsL primer and the 3′ ftsI primer). The resulting ΔftsIL::apra mutation was introduced into the chromosome in the same way as for single mutant isolation.

For generation of a non-sporulating strain by manipulating the developmentally controlled ftsZ2p, the TAGTGT residues of the −10 motif on cosmid Sv-4-G01 were replaced with an oriT acc(3)IV cassette flanked by introduced SpeI and XbaI sites. Restriction digestion of the mutagenized cosmid with SpeI and XbaI and religation left ACTAGA in place of the native −10 sequence. Exconjugants were selected as described above for unmarked mutations and identified by a PCR analysis using oligonucleotides specific for each of the two promoter sequences.

Construction of genetic complementation plasmids

For genetic complementation, a series of DNA fragment inserts were generated from cosmids by restriction digestion or amplification by PCR (Figs 1a and S8) and cloned into site-specific integration vectors pMS82. The resulting plasmids were introduced into S. venezuelae mutant strains by conjugation and integrated in trans into the chromosome at the ΦBT1 attachment site.

Fig. 1.

Construction and complementation of S. venezuelae strains mutant for core division genes. (a) A physical map of the dcw cluster in S. venezuelae and the genetic locus of divIC. Maps of two regions of the S. venezuelae chromosome are shown that contain genes encoding core proteins of the divisome. In each of the two loci, all genes are in the same orientation as the divisome genes. Regions replaced with an apramycin-resistance cassette or an unmarked in-frame deletion mutation are shown above the maps. DNA fragments used for constructing genetic complementation plasmids are shown below the map. (b) Phase-contrast microscopy of wild-type and mutant phenotypes and mutant phenotypes following genetic complementation. All images are phase-contrast micrographs of cover slip impressions from cultures grown for 2 days at 30 °C on MYM agar. The top row contains wild-type S. venezuelae strain containing the empty complementation vector on the left (WT). Immediately adjacent are shown seven division mutants containing the empty complementation vector pMS82. In the bottom row are shown the seven division mutants containing a complementing fragment cloned into pMS82, which restores sporulation to wild-type levels. Scale bar, 5 µm. (c) Transmission electron micrographs reveal septation and cell wall defects in the ftsZ, ftsI and ftsL mutants. Cells were grown for 2 days at 30 °C on MYM agar and thin sections were viewed by transmission electron microscopy. Mainly spores were observed for the wild-type strain (WT). No examples of vegetative cross-walls and sporulation septa were observed for the ftsZ null mutant. White arrow heads indicate formed unresolved sporulation septa in aerial hyphae for the ftsL and ftsI mutants. Scale bar, 500 nm.

Construction of strains expressing fluorescent FtsZ–YPet fusion proteins

Plasmid pKF351, carrying an ftsZ–ypet fusion in a vector that integrates at the ΦC31 attachment site [44], was introduced into relevant mutants by interspecies conjugation, as described above.

Microscopy

For phase-contrast microscopy, bacteria were grown as confluent patches on MYM agar. Cover slips were touched to the surface of sporulated patches and the material lifted was mounted on pads of 1 % agarose in phosphate-buffered saline (PBS). Samples were visualized using a Nikon Eclipse E400 with a Nikon 100×/1.25 oil immersion objective and a MicroPublisher 5.0 RTV high-resolution CCD camera (Qimaging).

For staining of cell walls and nucleoids, cultures were grown on MYM agar or in MYM liquid medium, and samples were fixed in ice-cold methanol for 5 min, washed twice in PBS and mounted in 100 μg ml⁻¹ propidium iodide (Molecular Probes) and 10 μg ml⁻¹ WGA–FITC (Molecular Probes) in 50 % glycerol. Fixed and stained samples were then spotted onto pads of 2 % agarose in PBS and sealed with petroleum jelly. Fluorescence imaging was performed with a Leica SP2 TCS confocal microscope using a Leica 63×/1.4 glycerin immersion objective.

In order to visualize fluorescent FtsZ–YPet fusion protein, cells were grown in liquid MYM, harvested and fixed with 2.28 % formaldehyde and 0.018 % glutaraldehyde, washed in PBS and mounted on 1 % agarose in PBS. To follow FtsZ dynamics, microfluidics-based time-lapse microscopy was performed using the CellASIC ONIX system and B04A-03 microfluidic plates (Merck Millipore), as described previously [35, 45]. The live-cell time-lapse experiments were repeated twice for each strain. Imaging was performed on a Zeiss AxioObserver.Z1 microscope with a Zeiss Plan-Apochromat 100×/1.4 Oil Ph3 objective, ZEN software (Zeiss) and an ORCA Flash 4.0 LT camera (Hamamatsu). Images and movies were processed using ImageJ/Fiji [46].

For transmission electron microscopy (TEM), cells were grown as lawns on MYM agar and fixed in 2.5 % glutaraldehyde in 0.05 M cacodylate buffer (pH 7.2) and incubated for 1 h in 2 % osmium tetroxide. Cells were dehydrated by successive transfer in five steps from 50–100 % ethanol. Cells were washed in propylene oxide and then in a 1 : 1 solution of propylene oxide and Spurr’s before being incubated in Spurr’s resin overnight at 60°C. Thin sections were stained with 2 % uranyl acetate and 1 % lead citrate. Samples were visualized using a JEOL JEM-1210 equipped with a Hamamatsu Orca-HR CCD camera.

Results and discussion

Generation of ftsZ null mutants

The earliest acting divisome protein FtsZ is required for cell division and viability in most bacteria. Previously, it had been demonstrated that ftsZ null strains could be isolated for S. coelicolor [8]. However, it was not clear whether this would be a general property of streptomycetes. Therefore, the procedure for isolating a deletion–insertion mutation for S. coelicolor ftsZ was replicated in S. venezuelae by replacing 844 bp, starting 16 bp upstream of the ftsZ start codon, with an apramycin-resistance cassette (Fig. 1a). The mutants described in this paper were isolated in S. venezuelae ATCC 10712 acquired from Dr Colin Stuttard (Dalhousie University, Halifax, Canada). In three independent experiments, we were able to isolate S. venezuelae null mutants for ftsZ. However, in contrast to the majority of the other cell division mutants isolated in S. venezuelae (described below), obtaining the ftsZ null mutants was not as straightforward. The initial ftsZ null mutant colonies grew very slowly after selection on conjugation plates, therefore, waiting up to 8 days before picking colonies was necessary to identify these mutants as ‘specks’ or ‘flecks’ on the agar surface. These primary mutant colonies on conjugation plates did not increase in size upon prolonged incubation, and nor did they sector. Subsequently, mutants were restreaked several times on selective media. ftsZ null mutants were unhealthy and grew slowly on the nalidixic acid-containing medium used for counter-selection of the conjugation donor E. coli, perhaps contributing to the difficulty when isolating them initially and, similarly, when introducing an empty vector as control for genetic complementation studies (see below). Once nalidixic acid counter-selection was no longer needed, the single colonies were uniform in appearance and the phenotype was stable. Colonies of the purified ftsZ null strains appeared on plates at similar incubation times relative to the wild-type, although plating efficiency was much lower (Figs S1 and S2); mature colonies were smaller and took much longer to develop an aerial mycelium, as judged by the surface of the colonies becoming white. Isolated independent strains had similar microscopic phenotypes and the insertion–deletion mutation of ftsZ was confirmed by PCR from genomic DNA. One representative strain was picked for further analysis (DU500). As expected, Western blot analysis verified that FtsZ was not detected in a whole-cell extract from DU500 (Fig. S3). As anticipated, phase-contrast and TEM microscopic analyses of the ftsZ null mutant grown on agar showed that aerial hyphae did not differentiate to produce spores and the vegetative hyphae were devoid of the normal cross-walls (Fig. 1b, c). The ftsZ null mutant was also completely blocked in both cell division and spore production when grown in liquid cultures and little cell material accumulated for the mutant under these growth conditions (data not shown). To confirm that the observed division phenotype was the result of the introduced mutation, genetic complementation studies were carried out. A restriction fragment containing ftsQ, ftsZ and the native ftsZ promoters in the intergenic region between the genes was integrated at the chromosomal attΦBT1 site (pJS8; Fig. 1a). The complementation vector rescued the division phenotype of the ftsZ null mutants, as judged by phase-contrast microscopy (Fig. 1b), and also restored growth and colony size on agar medium (Figs S2 and S3). We conclude that the deletion–insertion mutation is not polar on downstream gene expression, and FtsZ-dependent cell division is dispensable for growth and viability of S. venezuelae . Nonetheless, an unmarked ftsZ null strain was also generated and it had an identical microscopic phenotype as the insertion–deletion mutant, but was not used further in this study (DU665, data not shown). The fact that an unmarked null mutant can be isolated by gene conversion (see the Methods section) argues that the ftsZ mutants are difficult to distinguish from background on primary conjugation plates, not that they can only be isolated by very strong selection for marker replacement by double homologous recombination.

Using the same procedure that is described above, we have also isolated an ftsZ null mutant (DU669) in the S. venezuelae strain NRRL B-65442 obtained from Dr Mark Buttner (John Innes Centre, Norwich, UK) [47]. The mode of growth of this ftsZ null mutant DU699 (NRRL B-65442 background) in the absence of cell division and hyphal cross-walls has been described elsewhere [9]. As observed for other cell division mutants (data not shown), the macroscopic and microscopic phenotypes of ftsZ null mutants in the two independent wild-type backgrounds were essentially indistinguishable and could be genetically complemented, revealing no overt differences at the phenotypic level between the S. venezuelae parent strains obtained from different sources. All experiments described in the rest of this paper were carried out in the ATCC 10712 strain background. (Nonetheless, about a dozen core divisome mutant strains were also isolated in the NRRL B-65442 background. Their strain designations and genotypes are listed in Table S1.)

The developmentally regulated promoter of ftsZ is required for sporulation-associated cell division in S. venezuelae

For S. coelicolor , three promoters for ftsZ have been mapped to the 288 bp intergenic region between ftsZ and ftsQ, and one of them, ftsZ2p, is developmentally regulated [11]. It has been shown that BldD, a transcriptional regulator that plays a key role in Streptomyces development, binds to the developmentally regulated ftsZ2p promoter and acts to repress expression of ftsZ during vegetative growth [12]. In S. venezuelae it has been shown that expression of ftsZ is dependent on WhiA and WhiB, which are transcriptional regulators required for the transition from growth of aerial hyphae to sporulation [14, 15]. ChIP experiments indicated that a WhiA binding target lies around 158 bp from the predicted start codon of ftsZ [14], and in this region is a sequence that is identical to the −10 region TAGTGT of the S. coelicolor ftsZ2p [11] and Streptomyces griseus P_spo [48]. The intergenic region upstream of ftsZ is highly conserved between S. coelicolor and S. venezuelae with sequence conservation at the three mapped promoter regions, including the ftsZ2p promoter (Fig. S4a). To test whether the importance of this developmental promoter for sporulation was also retained in S. venezuelae , a strain was generated that was mutant for this presumed ftsZ2p promoter region. In the unmarked mutant, the TAGTGT residues at the −10 region of this presumed promoter were changed to ACTAGA (Fig. S4b). The resulting strain (DU523) had reduced plating efficiency compared to the wild-type, but grew robustly and formed an abundant aerial mycelium (Fig. S1). However, the ftsZ2p mutant was unable to efficiently convert the aerial hyphae to spore chains during growth on solid medium, and mature spores were observed much less frequently compared to the wild-type. Instead, longer spore-like compartments of irregular length were produced from aerial hyphae (Fig. 1b). Thus, this promoter mutant can form functional division septa that result in complete division events with cell separation that lead to formation of spore-like aerial hyphal fragments. However as shown by the absence of regularly septated spore chains, the mutant has a greatly reduced frequency of cell division compared to wild-type. The phenotype of the S. venezuelae ftsZ2p mutant is consistent with a failure to up-regulate the expression of ftsZ, and this up-regulation is required for developmentally associated cell division, similarly to what has been observed in S. coelicolor and S. griseus [11, 48].

Null mutants for core divisome genes

In order to clarify the roles of some of the conserved core divisome proteins for S. venezuelae , unmarked in-frame null mutants for ftsQ (strain DU629), divIC (strain DU613), ftsL (strain DU520), ftsW (strain DU521) and ftsI (strain DU679) were isolated (Fig. 1a). These genes are broadly conserved among bacteria and their products are membrane proteins required for coordinating the cytoplasmic Z ring with the peptidoglycan synthesis machinery [19, 25]. In other bacteria, FtsQ, FtsL and DivIC form a subcomplex that is recruited to the divisome. A 1 MDa complex containing those proteins, along with FtsZ, has been identified for E. coli [49]. Likewise, FtsI and FtsW form a subcomplex involved in septal peptidoglycan synthesis as a transpeptidase and transglycosylase, respectively [27]. While it has not been demonstrated directly for Streptomyces , it is reasonable to expect that the protein subcomplexes are conserved.

Since most of the core divisome genes are part of the complex dcw gene cluster (Fig. 1a) and are likely co-transcribed with other cell wall biosynthetic genes, unmarked in-frame deletions were generated to avoid polar effects on downstream genes. The mutants were readily isolated and showed consistent macroscopic and microscopic phenotypes that were strikingly similar to one another on MYM agar, but were distinct from the ftsZ null and ftsZ2p mutants. In contrast to the ftsZ null mutant, plating efficiency was unaffected in these mutants (Fig. S1), suggesting that vegetative cross-wall formation was not severely impaired. Likewise, aerial mycelium development was essentially unaffected, but aerial hyphae were not efficiently converted into chains of spores. By 48 h of incubation on solid medium, the majority of aerial mycelia had been converted into spores for the wild-type. For the ftsQ, ftsL, divIC, ftsW and ftsI null mutants, a mixture of spores, hyphal fragments and frequent lysed compartments were observed (Figs 1b and S5). Aerial hyphae often contained frequent and regularly spaced constrictions, reminiscent of sporulation septa (Fig. 1b). Some separated spores were produced and fragments of varying lengths were also observed for each of these mutants, showing that the products of these genes are not absolutely required for cell division for S. venezuelae (Figs 1b and S5). Each mutation was genetically complemented using site-specific integration plasmids with inserts shown in Fig. 1a. The complemented strains sporulate similarly to the wild-type parent, indicating that the phenotypes are associated with the introduced null mutations (Figs 1b and S1). Combining ftsQ, divIC and ftsL mutations as either double mutants or as a triple mutant did not have a synthetic effect on the observed phenotypes, as judged by phase-contrast microscopy (Fig. 2), suggesting that the loss of all the parts of the putative subcomplex formed by their gene products is no more deleterious than the loss of any one component. This result is consistent with the interpretation that missing any one component must inactivate the remainder of the tripartite complex. In addition, a double mutant lacking both ftsW and ftsI was also constructed and the phenotype was indistinguishable from the individual ftsW and ftsI mutants (Fig. 2), suggesting that the loss of both parts of the putative subcomplex formed by their gene products is no more deleterious than the loss of either one of the components. Finally, deleting both adjacent ftsL and ftsI genes together resulted in a mutant with a similar phenotype to an ftsI single mutant (Fig. 2), indicating that removal of parts of both putative divisome subcomplexes is no more deleterious than the loss of one part. The similarity of the core mutant phenotypes and lack of synergism when combining divisome mutations seem to support a model where there is no apparent hierarchy of assembly of the core divisome components in S. venezuelae . Further experimentation will be needed to clarify the situation and define each contribution.

Fig. 2.

Double and triple divisome mutants do not have additive or synergistic division phenotypes. The strains were grown for 2 days on MYM agar medium at 30 °C. Shown are phase-contrast images from impression coverslips of aerial hyphae for double and triple mutant strains. Aerial hyphae of double and triple mutants frequently contain evenly spaced constrictions, as do the single mutants. The double and triple mutant phenotypes are strikingly similar to those of the single mutants (Fig. 1b) and do not result in synthetic division phenotypes. Scale bar, 5 µm.

Spores could be isolated from the aerial mycelium of surface-grown cultures despite the fact that development-associated division was impaired for core divisome single mutants, with a reduction in the number of spores produced relative to the wild-type. In order to quantify the severity of reduction in sporulation-associated division in these core divisome mutants, measurements were made of mature spores and hyphal fragments harvested in a typical fashion from agar plates after 4 days’ incubation. For the wild-type, the material harvested consisted almost entirely of spores with an average length of 1.00 (±0.23) μm (Fig. S6). In contrast, the average lengths of the spores and hyphal fragments for the ftsQ, divIC and ftsL mutants were similar at 2.13 (±1.76), 2.75 (±2.67) and 2.49 (±3.92) μm, respectively, suggesting that when development-associated division resulting in cell separation occurred, every other to every third septum was formed with cell separation in some aerial hyphae. In contrast, the average spore-type compartment lengths were greater for the ftsW and ftsI mutants, 7.18 (±8.32) and 5.73 (±6.00) μm, respectively (Fig. S6), suggesting that in some hyphae with development-associated division leading to cell separation, every sixth to seventh septum may have been completed all the way to detachment of cells.

Phase-contrast microscopy showed that the divisome mutants were capable of division leading to cell separation, but did not provide detail on the septum morphology when division failed. Of the five isolated single mutants, ftsL and ftsI single mutants were selected for observation by electron microscopy as representative examples of mutants affecting the putative FtsQ–DivIC–FtsL complex and the FtsW–FtsI complex. TEM analysis for an ftsL null mutant indicated that the evenly spaced constrictions in aerial hyphae observed by light microscopy represented complete invaginations with very thick peptidoglycan (Fig. 1c), while ftsI null mutants produced more normal looking septa (Fig. 1c), but at a lower frequency.

While extremely rare in the wild-type strain, branching within nascent spore chains was commonly observed for the ftsQ, divIC, ftsL, ftsW and ftsI mutants. Both the cell division defect and the observed branching phenotype in aerial hyphae were rescued in genetic complementation studies, confirming that the phenotypes are associated with the deletion of these genes and not the result of unlinked mutations (Fig. 1b).

Core divisome proteins are not absolutely required for genome segregation in S. venezuelae

To further characterize the cell division and sporulation defects in ftsZ, ftsZ2p, ftsQ, ftsL, divIC, ftsW and ftsI mutants of S. venezuelae , the cell wall was stained using WGA–FITC and nucleoids were stained by propidium iodide (Figs 3 and S7). Cell wall staining confirmed that hyphae of the ftsZ null mutant had no signs of invagination, vegetative cross-walls or sporulation septa. In addition, there was no evidence of DNA condensation either (Fig. 3). In contrast, partial nucleoid condensation and segregation was observed for the ftsZ2p developmental promoter mutant (Fig. 3).

Fig. 3.

DNA segregation and cell wall phenotypes of S. venezuelae division mutants. Cells were grown for 2 days at 30 °C on MYM agar and cover slips were pressed onto confluent lawns. Samples of aerial hyphae were stained for cell wall (green) and DNA (red) and viewed by epifluorescence microscopy. The top row contains corresponding DIC transmitted light images. Wild-type samples mainly contained spores and spore chains. Examples of aerial hyphae of mutant strains ΔftsZ, ftsZΔ2p, ΔftsL and ΔftsI are shown. Scale bar, 5 µm.

Consistent with observations from phase-contrast and TEM microscopy, ftsQ, ftsL, divIC, ftsW and ftsI mutants showed very similar patterns of cell wall and DNA staining (Figs 3 and S7). In aerial hyphae with constrictions visible by light microscopy, ladders of nascent septal wall material could be seen. However, these ladders were often not as regular as the evenly spaced ones seen for the wild-type. DNA segregation was not grossly affected by the loss of any of these division genes, but often was less uniform than for the wild-type. Overall, under the laboratory conditions that we tested, we conclude that S. venezuelae is able to lay down cell division septa and segregate their genomes even in the absence of the core divisome genes ftsQ, ftsL, divIC, ftsW or ftsI. Future avenues of research will be necessary to understand why these genes are conserved, yet their products are not essential for septum formation for this filamentous bacterium.

Assembly of FtsZ into ladder-like arrays of Z rings in sporogenic hyphae does not require the core divisome genes ftsQ, ftsL, divIC and ftsW

Next, using a subset of the mutants, we investigated the extent to which the core divisome mutations affected the localization and dynamics of FtsZ rings in S. venezuelae . In order to do this, ftsZ–ypet (pKF351) was introduced into the ΦBT1 att site of ftsQ, ftsL, divIC and ftsW mutants, as well as the wild-type strain, leading to the production of FtsZ fused to the yellow fluorescent protein YPet in addition to the native FtsZ. In vegetative hyphae sampled from standard liquid medium cultures at different stages along the growth curve, we observed an apparently normal distribution of Z rings in vegetative hyphae (not shown, but also see the microfluidics data below). Further, sporulating hyphae with multiple, closely and regularly spaced Z rings were observed in both the wild-type strain and the ftsQ, ftsL, divIC and ftsW mutants (Fig. 4), albeit examples of sporulating hyphae were observed at a lower frequency in the mutants than in the wild-type.

Fig. 4.

Z ring assembly in sporogenic hyphae of S. venezuelae divisome mutants. Batch cultures were grown in a standard fashion in liquid MYM at 30 °C and samples were fixed by formaldehyde treatment before cells were mounted for microscopy. Representative micrographs of sporulating hyphae with FtsZ ladders are shown, visualized using YPet-tagged FtsZ. Shown are the wild-type control strain and the indicated divisome mutants into which plasmid pKF351(P_ftsZ-ftsZ–ypet) had been introduced. Scale bars, 2 µm.

In order to observe more clearly how mutations in core divisome genes affect FtsZ dynamics and Z ring formation, we used microfluidics-based fluorescence live cell imaging, as described previously [35, 45]. Representative micrographs of FtsZ ladders formed in the wild-type and ftsQ and ftsW mutants under these conditions are shown in Fig. 5. Time-lapse images were also acquired to visualize the FtsZ dynamics. In the wild-type, during the vegetative stage, typical Z rings are observed that are highly dynamic, as shown by their movement along the hyphae before they stabilize at fixed positions and then increase in fluorescence intensity (Movie S1). Presumably, these observed intense Z rings mark sites of vegetative cross-wall formation. A very similar pattern was seen for the formation of Z rings at early growth timepoints for vegetative hyphae of the ftsQ and ftsW mutants (Movies S2 and S3, respectively). FtsZ dynamics were also visualized during the sporulation stage for the wild-type parent, where examples of the assembly of evenly spaced Z rings in ladder-like patterns could be seen in sporogenic hyphae (Movie S1, >10 h of growth in this sample). Intriguingly, similar development-associated FtsZ dynamics were observed in the mutants for ftsQ (Movie S2) and ftsW (Movie S3) and the assembly of evenly spaced FtsZ ladders occurred. Closer inspection of the stability of FtsZ ladder persistence was accomplished by constructing a montage from timepoint images for the wild-type parent and the ftsQ and ftsW mutants (Fig. 6). The FtsZ ladders persist for approximately 2 h for the wild-type, but the ladders do not show the same dynamics for the mutants. Certain FtsZ rings are lost over time in the mutants, with the rungs of FtsZ ladders in the ftsW mutant being the least stable. These relative FtsZ ladder stabilities correlate with the average lengths of mature spores that can be harvested from surface grown cultures (Fig. S6), with the spores for ftsQ and ftsW mutants being approximately 2× and 7× the length of those produced by the wild-type.

Fig. 5.

Live-cell imaging of Z ring assembly in sporogenic hyphae of S. venezuelae wild-type and ΔftsQ and ΔftsW mutants. Cultures were grown in MYM at 30 °C using a microfluidic system. Representative micrographs of unfixed sporulating hyphae with FtsZ ladders are shown, visualized using YPet-tagged FtsZ. Shown are the wild-type control strain and the indicated divisome mutants into which plasmid pKF351(P_ftsZ−ftsZ–ypet) had been introduced. Scale bars, 2 µm.

Fig. 6.

FtsQ and FtsW stabilize Z rings during sporulation-specific cell division. Shown is a montage of representative time series documenting FtsZ dynamics during spore formation. Strains were grown in liquid MYM at 30 °C using a microfluidic system. Fluorescence images of FtsZ–YPet signal were obtained from time-lapse microscopy (top) and the corresponding phase-contrast images are also shown (bottom). Shown are montages of the wild-type control strain and the indicated divisome mutants into which plasmid pKF351(P_ftsZ-ftsZ–ypet) had been introduced. Time intervals between images were kept at 20 min. In addition, 0 min was considered as the time wherein the shown hyphae had undergone arrest of tip extension before sporulation septation began. Scale bars, 2 µm.

Physiological relevance of cumulative results

Overall, the results clearly show that the products of the divisome genes ftsQ, ftsL, divIC and ftsW are not required for Z ring assembly, for the single Z rings that are formed in vegetative hyphae (normally leading to hyphal cross-walls in the wild-type), and in sporogenic hyphae, where ladders of regularly spaced Z rings are typically formed as part of sporulation septation. In some instances, Z ring formation appeared essentially normal in the ftsQ, ftsL, divIC and ftsW mutants. The results are consistent with the previously observed ability of corresponding mutants in S. coelicolor to form cross-walls and septa, at least under certain conditions [28–31]. It has been speculated previously that ftsW may be required for Z ring assembly, and may provide a membrane attachment for FtsZ in both S. coelicolor and Mycobacterium smegmatis [31, 50]. Our results presented here for S. venezuelae show that ftsW is not required for Z ring formation.

The fact that core cell division proteins FtsQ, FtsL, DivIC, FtsW and FtsI are not strictly required for cell division in Streptomyces spp. gives rise to interesting questions to be investigated in future studies. For example, how is it possible to carry out cell division in the absence of the FtsQ–FtsL-DivIC complex? Either the divisome in Streptomyces spp. can be stable and functional without these proteins, or there are other proteins that can replace or reinforce these core divisome proteins. In the latter case, such proteins would be pertinent to identify. Interestingly, co-immunoprecipitation experiments identified 63 FtsQ-interacting proteins for Mycobacterium tuberculosis and may point to homologues for further investigation [51].

Similarly, how can cell division occur in the absence of FtsW and its cognate transpeptidase FtsI? The transpeptidase activity of FtsI has been shown to be nonessential for some other Gram-positive bacteria, although the protein is still physically required [52–54]. In the absence of FtsI, perhaps FtsW functions with one or more of the many PBPs encoded for S. venezuelae . FtsI transpeptidase activity can be supplied by other PBPs in B. subtilis [54]. FtsW co-purifies with two different PBPs in a potential trimeric complex in E. coli [55]. FtsW has recently been identified as a peptidoglycan transglycosylase (essentially a peptidoglycan polymerase) [27], in similarity to related RodA SEDS proteins [56–58]. These are essential activities for the formation of a cell division septum, and the results presented here suggest that another peptidoglycan polymerase is likely recruited to Z rings at division sites in order for the S. venezuelae ftsW mutant to form septa. As one possibility, perhaps transglycosylase activity can be provided by an autonomous bifunctional class A PBP and not by a SEDS protein. In support of that notion, evidence for intimate participation of bifunctional PBPs in septum peptidoglycan synthesis has been accumulating [59]. Recent evidence suggests that pneumococcal peptidoglycan is synthesized, in part, by bifunctional PBPs [60]. As another possibility, one of the other three SEDS proteins encoded by streptomycete genomes [29, 31] may be active either at the same time as FtsW and/or be induced in its absence. It will be interesting to see which protein(s) functions in a ftsW mutant and how it would be recruited to the divisome. Of final note, we also have constructed strains individually expressing EFGP fusions to each protein of the FtsQ–FtsL–DivIC or FtsI–FtsW complexes and the fluorescent localization signals are not strong enough to publish (data not shown). Potentially, the weak fluorescence signal is indicative of a low intracellular concentration. Again, future work will have to be performed to learn if small amounts of the proteins are needed for normal function, if another protein can substitute, or if multiple SEDS–PBP pairs work simultaneously during sporulation septum formation.

Summary and conclusions

In this study, we have established the contributions of known central cell division proteins in the coordinated process of sporulation septation in S. venezuelae . Knowing the null phenotypes for mutants lacking known players in cell division will be essential for future studies as we continue to peel back the novel lineage-specific layers of controls evolved to govern the concerted development-associated control of essential cell biological processes in streptomycetes.

In this study, we have taken advantage of the benefits of S. venezuelae to visualize the synchronous events being orchestrated within sporogenic hyphae by live-cell time-lapse microscopy because this species undergoes differentiation under submerged growth conditions. The data show that ladder-like assemblages of evenly spaced FtsZ rings typically form in all of the characterized core divisome mutants. Thus, the tested divisome components are not required for that early coordinated event. However, once formed, the Z rings appear to be unstable and a number of rings prematurely disband. The loss of coordination results in irregular spacing between completed septa and irregular spore size, as seen in the divisome mutants.

Evidence has accumulated for subcomplex formation of FtsQ–FtsL–DivIC and FtsW–FtsI before participation in the divisome. For S. venezuelae , combining mutations of genes encoding these components does not result in synthetic phenotypes. This result is consistent with the interpretation that the loss of any one component disrupts the function of the subcomplex. While the subcomplexes are not absolutely required, they do contribute to the stability of the synchronous tandem arrays of divisome complexes, as visualized by FtsZ–YPet. Recently, analysis of bacterial dynamins DynA and DynB for S. venezuelae showed that they interact with the division machinery [18] and contribute to Z ring stability, and mutants encoding those proteins have somewhat similar phenotypes to the divisome mutants reported here. Future work will be needed to understand how these components interact and are regulated to synchronously coordinate sporulation septum formation.