The spread of antimicrobial resistance and vaccine escape in the human pathogen Streptococcus pneumoniae can be largely attributed to competence-induced transformation. Here, we studied this process at the single-cell level. We show that within isogenic populations, all cells become naturally competent and bind exogenous DNA. We find that transformation is highly efficient and that the chromosomal location of the integration site or whether the transformed gene is encoded on the leading or lagging strand has limited influence on recombination efficiency. Indeed, we have observed multiple recombination events in single recipients in real-time. However, because of saturation and because a single-stranded donor DNA replaces the original allele, transformation efficiency has an upper threshold of approximately 50% of the population. The fixed mechanism of transformation results in a fail-safe strategy for the population as half of the population generally keeps an intact copy of the original genome.
The opportunistic human pathogen Streptococcus pneumoniae (the pneumococcus) kills over a million individuals each year, despite the introduction of several vaccines targeting its capsule (Croucher et al., 2018; O'Brien et al., 2009; Prina et al., 2015). Because of its ability to take-up DNA from its environment by competence activation, genes associated with capsule biosynthesis are rapidly transferred from one strain to the other thereby contributing to vaccine escape (Salvadori et al., 2019). In addition, antibiotic resistance remains a cause of concern and competence-dependent recombination plays an important role in the spread of drug resistance (Sw et al., 2019). For example, one of the main genetic sources for penicillin resistance in S. pneumoniae is DNA acquired from non-pathogenic Streptococci from the viridans group such as S. mitis that also lives in the human nasal and oral cavities (Bryskier, 2002; Janoir et al., 1999). Consistently, antibiotic- resistant pneumococci and vaccine-escape variants remain an important cause of invasive infections in spite of the introduction of the conjugate vaccines (Fenoll et al., 2018; Levy et al., 2019; Ouldali et al., 2018).
Although pneumococcal competence is one of the best studied bacterial regulatory systems (Gómez-Mejia et al., 2018; Johnston et al., 2014; Lin and Kussell, 2017; Salvadori et al., 2019; Shanker and Federle, 2017; Straume et al., 2015; Veening and Blokesch, 2017), and pneumococcal transformation was already discovered in the early twentieth century (Avery et al., 1944; Griffith, 1928), we have a poor understanding on how competence-dependent transformation drives pneumococcal population dynamics, serotype displacement and the spread of antibiotic resistance. Importantly, horizontal gene transfer (HGT) via natural transformation is not only conserved in Streptococci but is present in many human pathogens where it promotes the spread of virulence determinants and antibiotic resistance (Brockhurst et al., 2019; Dubnau and Blokesch, 2019; Johnston et al., 2014). For this reason, it is crucial to understand what the main bottlenecks are during the take-up and recombination of exogenous DNA that leads to transformed new genotypes.
In contrast to many other competent pathogens such as Acinetobacter spp. and Neisseria meningitidis in which competence is constitutively expressed, competence development in S. pneumoniae is only activated under specific conditions (Blokesch, 2016; Claverys et al., 2006). Pneumococcal competence is under control of a two-component quorum sensing system (Figure 1). ComC is cleaved and exported by the peptidase-containing ATP-binding cassette transporter ComAB (Chandler and Morrison, 1988; Håvarstein et al., 1995; Hui et al., 1995). Cleaved ComC autoinducer is commonly referred to as competence stimulating peptide (CSP) (Alloing et al., 1996; Håvarstein et al., 1996; Håvarstein et al., 1995). CSP is recognized by the membrane-bound histidine kinase ComD (Håvarstein et al., 1996). Once a certain threshold level of CSP has been reached, as the culture reaches higher densities, or when other environmental factors increase local CSP concentrations (Domenech et al., 2018; Moreno-Gámez et al., 2017), ComD will autophosphorylate and transfer the phosphoryl group to the response regulator ComE (Martin et al., 2013). Phosphorylated ComE then dimerizes (Boudes et al., 2014; Sanchez et al., 2015) and binds to a specific DNA sequence (Martin et al., 2013; Pestova et al., 1996; Slager et al., 2019; Ween et al., 1999). The comCDE and comAB operons are under direct control of ComE, setting up a positive feedback loop. The genes under control of ComE are called the early com genes (Figure 1). Importantly, phosphorylated ComE also activates expression of the gene encoding the alternative sigma factor ComX. ComX activates transcription of the so-called late com genes, which includes the genes required for DNA uptake and integration (Campbell et al., 1998; Dagkessamanskaia et al., 2004; Luo et al., 2003; Pestova and Morrison, 1998; Slager et al., 2019; Figure 1). While regulation of competence is highly diverse between naturally transformable bacteria, the actual DNA uptake and integration machinery is largely conserved (Chen and Dubnau, 2004; Johnston et al., 2014).
During pneumococcal competence, exogenous double stranded DNA (dsDNA) is bound by a type IV-like pilus (Laurenceau et al., 2013) and subsequently sequestered to the DNA uptake machinery (Figure 1). Note that in contrast to some other competent bacteria, pneumococcus binds and takes up DNA of any sequence, including non-kin DNA (Mell and Redfield, 2014). Next, the dsDNA is processed into single-stranded DNA (ssDNA) by the EndA nuclease and internalized through a membrane pore consisting of ComEC. Once inside, the ssDNA is bound by a competence-specific ssDNA-binding protein, SsbB, and stabilized by DprA and RecA (Attaiech et al., 2011; Bergé et al., 2003; Figure 1). This complex undergoes homology scanning and forms a temporal hetero-duplex during strand invasion which can lead to homologous recombination (Mortier-Barrière et al., 2007). The exact details on the kinetics of this process, as well as how the hetero-duplex is resolved in most cells remains elusive. The competent transformation state in S. pneumoniae is transient as DprA interacts with phosphorylated ComE to inhibit its activity (Mirouze et al., 2013). In addition, several key Com proteins are rapidly turned over after their synthesis, leading to a window of DNA uptake of approximately 15 min (Liu et al., 2019; Tomasz, 1966; Weng et al., 2013).
As most work on pneumococcal competence and transformation has been performed using bulk assays, it is unclear what the actual bottlenecks are during competence development and why one cell will be transformed whereas another one will not. Here, we have set up single-cell transformation assays that allow us to quantify successful recombination events in real-time. This study provides direct evidence for several decade-old models underpinning bacterial transformation, and offers new insights that help explain why competence-induced transformation is so effective in changing global pneumococcal population structures.
To quantify pneumococcal transformation efficiency and determine at which step potential bottlenecks arise, we systematically analyzed every stage during the process: (1) competence development, (2) production of the DNA uptake machinery, (3) binding of exogenous DNA, and (4) recombination and expression of the newly acquired genetic information (Figure 1). While competence development in B. subtilis is limited to approximately 10% of the population (Maamar and Dubnau, 2005; Smits et al., 2005), up to 100% of cells within pneumococcal populations have been reported to become competent when induced with exogenously added synthetic CSP or when grown on semi-solid surfaces (Bergé et al., 2017; Domenech et al., 2018; Litt et al., 1958; Martin et al., 2010; Moreno-Gámez et al., 2017; Slager et al., 2014).
To quantify competence development in clonal pneumococcal populations in a systematic fashion, we constructed a set of reporters. First, we assessed the timing of both naturally induced and artificially induced competence (by the addition of synthetic CSP) at the population level utilizing a firefly luciferase reporter under the control of the late competence ssbB promoter (strain DLA3). Cells were grown in C+Y medium at 37°C (see Materials and methods) and growth and luciferase activity were measured every 10 min. As expected, under these experimental conditions, the population rapidly activates ssbB in the presence of added CSP, while in the absence of externally added CSP, the ssbB promoter peaks after approximately 100 min (Figure 2A). To determine which fraction of the cells switch on the competence pathway, we fused the ssbB promoter to a fast folding yellow fluorescent protein (msfYFP) and integrated this construct at the native ssbB locus (strain VL2219). As shown in Figure 2B–C,~97% of the population was positive for ssbB expression 20 min after addition of synthetic CSP as determined by fluorescence microscopy followed by automated image analysis (see Materials and methods for details). Importantly, spontaneous competence without the addition of synthetic CSP was reached in 92% of the population showing that almost all pneumococci, regardless of their cell length and cell-cycle status become naturally competent (Figure 2—figure supplement 1).
To test whether competent cells actually produce the machinery required for DNA uptake, we constructed translational msfYFP fusions to three essential components of the transformation machinery: ComGA (ATPase driving the DNA uptake pilus), ComEA (DNA receptor), and ComFA (ATPase driving DNA import) as the only copy integrated at their native locus. After 20 min of incubation with synthetic CSP, cells were collected for fluorescence microscopy. In line with the fraction of cells that become competent, msfYFP-ComEA, ComFA-msfYFP, and ComGA-msfYFP were also expressed in the majority of the cells (~92,~99, and~99%, respectively) (Figure 2D and Figure 2—figure supplement 2). A double-labeled strain (strain OVL2536: PssbB-mScarlet-I, ComGA-msfYFP) demonstrated that all competent cells indeed produce the DNA uptake machinery (Figure 2E).
Finally, to assess whether the proteins required for recombination and chromosomal integration of exogenous DNA also were expressed in the majority of the population, we constructed translational fusions to RecA and the recombination mediator protein DprA. Similar to the DNA-uptake proteins, RecA and DprA were induced in most competent cells (Figure 2D and Figure 2—figure supplement 2).
During pneumococcal competence, the capture of extracellular DNA by the ComGC pilus is an essential step for transformation (Bergé et al., 2002). To examine which proportion of cells is capable of binding DNA during competence, we labeled extracellular DNA (285 bp S. pneumoniae DNA fragment, see Materials and methods) fluorescently with the Cy3 dye. After induction of competence with synthetic CSP of cells mutated for EndA (to prevent degradation of the exogenous DNA),~90% of the population bound extracellular DNA as visualized by fluorescence microscopy (Figure 2F). Even without additional CSP, spontaneous competence also led to most cells (89.6%) binding exogenous DNA (Figure 2G). As observed before in an unencapsulated R6 strain (Bergé et al., 2013), we note that also in the encapsulated serotype 2 D39V strain, DNA mainly bound to the mid-cell positions of the cell, corresponding to the localization of the DNA uptake machinery particularly the ComEA receptor (Figure 2F and Figure 2—figure supplement 1; Bergé et al., 2013). Collectively, these data validate by direct single-cell observations that pneumococcal competence development, the subsequent production of the DNA uptake and integration machinery, as well as DNA binding is highly efficient and occurs in nearly every cell of the population regardless of their cell-cycle state.
Having established that there are no significant bottlenecks during the process of both induced and natural competence development and DNA uptake, we next set out to develop a system that allows for the direct visualization of successful recombination. Traditionally, transformation efficiencies are evaluated using antibiotic selection methods. However, these selection methods have limitations because they depend on the counting of colony forming units, which can lead to the overestimation of transformation efficiencies, due to inefficient separation of transformed from non-transformed daughter cells and nongenetic inheritance of antibiotic resistance (Dalia and Dalia, 2019; Domenech et al., 2018; Ephrussi-Taylor, 1962; Ephrussi-Taylor, 1958; Figure 3—figure supplement 1). In order to overcome these concerns and analyze successful recombination events during transformation at the single-cell level, we developed a fluorescence-based reporter system inspired by a system previously used to observe natural transformation in S. pneumoniae (Bergé et al., 2013) and other bacterial species (Boonstra et al., 2018; Corbinais et al., 2016; Godeux et al., 2018). To do so, we utilized a fluorescent donor strain in which the gene encoding the abundant histone-like protein HlpA (aka HU) was fused in frame with the gene encoding the red fluorescent protein mScarlet-I integrated at the native hlpA locus at 169° on the circular chromosome (Keller et al., 2019) (strain VL1780) (Figure 3A). A recipient, non-fluorescent strain was constructed (strain VL1784) in which hlpA was separated from mScarlet-I by a stop codon mutation (G > T) (Figure 3A–C and Figure 3—figure supplement 2A, hlpA-stop-mScarlet-I). Upon uptake, integration and expression of exogenous transforming DNA (tDNA) containing the donor construct (intact hlpA-mScarlet-I), successfully transformed recipient cells will produce functional HlpA-mScarlet-I that can be quantified by fluorescence microscopy or flow cytometry (Figure 3B and C and Figure 3—figure supplement 2B). As this is a recombination event between highly similar but not identical DNA (except for the SNP causing a stop codon), this is called a homeologous recombination event (Humbert et al., 1995; Petit et al., 1991). Note that this reporter system does not affect growth regardless of the presence of the stop codon (Figure 3—figure supplement 3) and that flow-cytometry analysis slightly overestimates the real transformation efficiencies due to cell chaining (Figure 4 and Figure 3—figure supplement 4, see below).
As mScarlet-I is a fast folding red fluorescent protein (Bindels et al., 2017), this reporter system should allow for the real-time detection of homeologous recombination during transformation. To test this, we provided competent recipient cells that besides the hlpA-stop-mScarlet-I allele also constitutively expressed sfGFP (strain VL1832) with intact hlpA-mScarlet-I as donor tDNA in the presence of CSP and then performed time-lapse microscopy (see Materials and methods for details). As shown in Figure 3D and Videos 1 and 2, recipient cells do not display any red fluorescence in the beginning and then gradually start to express red fluorescence. When quantifying the fluorescence signals and superimposing this on a cell lineage tree constructed using a set of new scripts written in BactMAP (van Raaphorst et al., 2020) (see Materials and methods), it becomes apparent that the initial recipient cell already expresses HlpA-mScarlet-I right after the addition of tDNA before the first cell division as red fluorescent signals above background levels can be detected (Figure 3E). Notably, only half of the recipients’ descendants appear to strongly express HlpA-mScarlet-I (Figure 3E, right lineage). Contrary, after three more divisions the non-transformed lineage no longer expresses red fluorescence (Figure 3E, left lineage). These results are in line with a recent study in Vibrio cholerae that showed a period of nongenetic inheritance in daughter cells during transformation (Dalia and Dalia, 2019). Similar observations were made when using a different transformation reporter system (Figure 5C and Video 3, see below). In line with current models of transformation (Davidoff-Abelson and Dubnau, 1971; Ephrussi-Taylor and Gray, 1966; Fox and Allen, 1964; Gabor and Hotchkiss, 1966; Lacks, 1962; Méjean and Claverys, 1984; Piechowska and Fox, 1971), these observations are consistent with a model in which recombination occurs by direct integration of the ssDNA donor and forms a hetero-duplex. Therefore, at least one round of DNA replication and division is required to generate two different homo-duplex chromosomes in progeny cells (Figure 3F). The fact that we initially also observe fluorescence in the untransformed lineage suggests that phenotypic expression derived from the acquired allele might occur prior to forming a homo-duplex. In this case, the transformed ssDNA likely replaced the anti-sense, noncoding strand so functional hlpA-mScarlet-I could be immediately transcribed after integration via RecA-directed homeologous recombination (mismatched pairing between exchanged DNA strands that are tolerated during the process of homologous recombination). Alternatively, phenotypic expression in these cells can occur if the transformed locus gets replicated, resulting in two homo-duplexes (transformed and original allele), and then transcribed before division of the cell (Dalia and Dalia, 2019).
The constructed system now allows us to quantify successful homeologous recombination events at the single-cell level, without the bias introduced by traditional plating assays. Previously, it was shown that the concentration of donor DNA as well as the length of the homology regions strongly influences transformation efficiency (Keller et al., 2019; Lee et al., 1998). To examine recombination bottlenecks in our single-cell setup, we treated our reporter recipient strain VL1784 with CSP and used intact hlpA-mScarlet-I donor tDNA with various lengths of homology surrounding the stop codon (fragments of 2.7, 5, or 7 kb) at a range of different concentrations (0.0032, 0.032, 0.32, or 3.2 nM). Then, after 4 hr incubation in liquid medium to allow for complete homo-duplex allele formation and dilution of nongenetically inherited HlpA-mScarlet-I, cells were separated from chains by vigorously shaking on a bead beater devise (see Figure 3—figure supplement 4). Finally, transformation efficiencies were quantified by flow-cytometry (Figure 5A). In line with studies using classical plating methods to assess transformation efficiencies (Keller et al., 2019; Lee et al., 1998), higher transformation frequencies were observed at higher donor DNA concentrations and with longer homology regions (Figure 5B). Interestingly, the frequency of transformation plateaued at ~50% regardless of the concentration of donor DNA and sequence homology length (Figure 5B). This is in contrast to reported transformation frequencies using traditional plating assays where transformation frequencies of higher than 75% (Ephrussi-Taylor, 1958) and up to 100% (Marie et al., 2017) have been reported. This discrepancy can be explained by the lack of separation of transformed from non-transformed cells within the counted colony (Figure 3—figure supplement 1). To exclude the possibility that the observed limitation in transformation frequency is due to an unique feature of the hlpA-stop-mScarlet-I reporter, we constructed an alternative reporter cassette in which we translationally fused the superfolder green fluorescent protein (sfGFP) and SPV_1159, a nonessential small membrane protein under control of the strong constitutive P3 promoter (Keller et al., 2019; Sorg et al., 2015) cloned into the transcriptionally silent CEP locus at 295° on the circular chromosome (Figure 5C, strain VL1786). Based on this construct, a recipient strain was constructed containing a stop codon mutation in the linker between spv_1159 and sfGFP (strain VL1788). Indeed, this spv_1159-sfGFP-based transformation reporter demonstrated similar transformation characteristics as the hlpA-mScarlet-I reporter in time-lapse microscopy and flow-cytometry analysis (Figure 5—figure supplement 1 and Video 3). The transformation frequency of the spv_1159-sfGFP reporter was also dependent on donor DNA concentration and never exceeded ~50% (Figure 5B).
These data show that there is a limit on the maximum efficiency of transformation, despite the fact that most cells become competent and bind extracellular DNA (Figure 2) and support a model in which in general only one of the recipient allele strands is replaced by the donor DNA (Ephrussi-Taylor, 1966; Figure 3F). Importantly, these experiments indicate that during competence-dependent transformation, given the donor DNA is of sufficient (homology) length and concentration (see Discussion), in principle all targeted loci can be replaced at least on one strand.
Previous work showed that pneumococcal genetic transformation involves the DNA mismatch repair (MMR) system, which is mediated by HexA (Claverys and Lacks, 1986), and it was suggested that certain alleles upon transformation might be particularly prone to repair (Ephrussi-Taylor, 1966). To test whether hexA plays a role in our reporter system, we quantified transformation efficiencies in a hexA mutant background. This showed no significant recombination differences compared to the wild-type background (Figure 4—figure supplement 2). To test if our transformation reporter system depends on the competence-induced homologous recombinase, RecA, we depleted RecA expression level using CRISPR interference (Liu et al., 2017; Figure 5D). In control strain VL3485 (Plac_dcas9, without sgRNA), induction of dCas9 by IPTG did not affect the transformation efficiency with hlpA-mScarlet-I tDNA. However, when RecA expression was depleted by induction with IPTG in strain VL3486 (Plac_dcas9+sgRNA-recA), the transformation frequency was decreased in an IPTG-dose dependent manner. Note that although RecA is known to be critical for optimal growth in S. pneumoniae (Mortier-Barrière et al., 1998), the CRISPRi depletion levels during competence did not affect bacterial growth (Figure 5—figure supplement 3). Collectively, these data show that our fluorescence-based transformation assay faithfully reflects RecA-dependent homeologous recombination events.
It was previously suggested that the genomic location and the cell-cycle state might influence transformation efficiency as each hetero-duplex needs to be resolved to a homo-duplex by DNA replication and loci located close to oriC will have multiple copies (Bergé et al., 2013; Dalia and Dalia, 2019; Ephrussi-Taylor and Gray, 1966; Porter and Guild, 1969). To test whether the genomic location of the recombination site and the read orientation of the newly acquired functional allele influences transformation efficiency, we introduced the spv_1159-stop-sfGFP reporter at four different positions on the chromosome: on the right arm of the chromosome at 101° of the circular chromosome (bgaA locus), near the terminus at 169° (hlpA locus), on the left arm of the chromosome at 295° (cep locus) and near oriC at 359° (comCDE locus) (Figure 4A and B). In addition, spv_1159-stop-sfGFP was introduced on both the positive and negative strand on the left arm of the chromosome (cep locus at 295°) and on the right arm of the chromosome (bgaA locus at 101°) (Figure 4C). As shown in Figure 4, and Figure 4—figure supplement 1, transformation efficiencies were of a similar order across all tested loci and genetic orientations, with a maximal recombination efficiency of approximately 50%. We do note that certain loci consistently demonstrate higher transformation efficiencies than others (cf. CEP locus vs bgaA locus, Figure 4—figure supplement 1), but no significant differences were observed regarding the orientation of the construct (see Discussion).
By performing time-lapse microscopy and tracking cell fates across several generations, we can, in principle, tell whether there was a preference for integration at either the leading or lagging strand (Figure 4—figure supplement 2A). By placing the direction of transcription of the reporter on the leading strand, RNAP will thus use the noncoding strand as template. In this situation, only if the noncoding strand is replaced by the donor DNA, fluorescence will be apparent during the first cell cycle upon transformation. If the donor DNA is integrated at the coding strand, it will take one more replication cycle before the hetero-duplex is resolved and the noncoding strand contains the intact reporter and fluorescence will be observed later than in the first case (Figure 4—figure supplement 2B). Indeed, we can observe all different scenarios with transformants rapidly expressing HlpA-mScarlet-I (possible noncoding strand or double stranded recombinants) and cells that only express HlpA-mScarlet-I after the first cell division (possible coding strand recombinants) (Figure 4—figure supplement 2C, Video 4). As we did not simultaneously track DNA replication in these cells, we cannot exclude the possibility that after transformation, a round of replication occurs before phenotypic expression. Nevertheless, together with the ‘bulk’ (FACS) single-cell transformation experiments described above, the time-lapse data strongly suggest that there is no preference for integration at either the leading or lagging strand and that this is an unbiased event. These findings correspond with work done in the 1960s and 1970s that showed that either strand of the incoming dsDNA is degraded randomly by EndA and either strand has a similar chance of being integrated (Puyet et al., 1990). Recent work in V. cholerae demonstrated that 7% of transformation events occurred at both strands of the integration site, and it was speculated that this was because of integration of multiple donor ssDNA’s replacing both the leading and lagging strand of the recipient. By recording 76 single-cell transformation events using time-lapse microscopy, we found six cases in which both daughter cells (7.8%) expressed fluorescence, suggestive of double transformation events on both strands. These findings also indirectly indicate that hetero-duplex DNA can be transcribed by RNAP and do not necessarily require a round of DNA replication to form homo-duplex DNA (see below and [Uptain and Chamberlin, 1997]). Together, these data show that hetero-duplexes with exogenous DNA are made across all available loci regardless of reading strand or distance to oriC.
The previous experiments demonstrated that, under ideal conditions with long flanking homology regions and high DNA concentrations, all available recombination sites are transformed on at least one of the strands. Previous studies demonstrated that pneumococcal natural transformation is capable to deal with multiple donor DNAs for genetic recombination (Dalia et al., 2014; Lam et al., 2020). Also, it has been reported that the DNA uptake and recombination process in S. pneumoniae is complete within 15 min (Bergé et al., 2003), which is a shorter time window than the doubling time (Ephrussi-Taylor, 1966). In order to investigate the possibility of visualizing multiple recombination events, we constructed a dual reporter strain (strain VL1803), which harbors both hlpA-stop-mScarlet-I and spv_1159-stop-sfGFP at distinct chromosomal locations (Figure 6A). Transformation efficiencies of this reporter strain with each single donor DNA at the saturated concentration typically reached 50% for both hlpA-mScarlet-I and spv_1159-sfGFP as quantified by microscopy (Figure 6C–D). When both donor DNAs were provided, double transformants were observed (15.6 ± 4.4%) as well as single hlpA-mScarlet-I transformants (20.2 ± 8.9%) and single spv_1159-sfGFP transformants (15.2 ± 4.9%). Time-lapse imaging of competent recipient VL1803 cells with both donor DNAs clearly demonstrated that single recipients could successfully recombine both fragments (Figure 6B and C, Videos 5,6). We note that, on average, the fraction of non-transformed cells is close to 50% (48.9 ± 9.5%), implying that each recombination event is not independent from the next or that there is an upper limit to the number of successful recombinations, otherwise we would expect the fraction of non-transformed cells to decrease with multiple donor DNAs (Figure 6—figure supplement 1). An alternative model could be that each recombination event is independent from the next but due to recombination events outside the stop codon SNP, which cannot be quantified in our setup, a reduced transformation efficiency is recorded (see Discussion).
To further explore whether transformation efficiency indeed has a plateau, we constructed a triple reporter strain (VL3127) that harbors ftsZ-stop-mTurquoise2, spv_1159-stop-msfYFP, and hlpA-stop-mScarlet-I at three different genomic locations (Figure 7A). Beside the fact that the fluorescent proteins used are spectrally distinct, every fluorescent reporter also has a specific cellular localization, facilitating automated image analyses of successful recombination. The triple reporter strain was transformed with donor tDNA fragments ftsZ-mTurquoise2, spv_1159-msfYFP, and hlpA-mScarlet-I. After 4 hr of incubation for fluorescent protein maturation and chromosomal segregation, cells were assessed by fluorescence microscopy. As shown in Figure 7B and Video 7, multiple transformed cells with double or triple acquired fluorescence signals were readily observed. Next, we performed single-cell transformation assays with strain VL3127 providing one tDNA or all three tDNAs and automatically quantified recombination efficiencies using Oufti and BactMAP-based image analysis (Paintdakhi et al., 2016; van Raaphorst et al., 2020; Figure 7C). In line with our previous observations, each single transformation with a saturated concentration of donor tDNA resulted in a recombination efficiency not higher than 50% (Figure 7C). Interestingly, every possible recombination event happened within the population: cells were observed in which just a single recombination event took place (the most occurring type of transformation), two recombination events (2.2 ± 0.9%, 4.1 ± 2.7% or 2.1 ± 1.8% for each possible combination) or even three recombination events (1.5 ± 1.1% of all cells). Nevertheless, more than half of the population (58.7 ± 13.4%) did not show any fluorescence when simultaneously transformed with three tDNAs. These observations support a model in which each transformation is in principle independent from the next (Figure 6—figure supplement 1).
The data so far show that in principle every locus in S. pneumoniae can be efficiently transformed to a maximum of 50% of the cells when providing tDNA at high DNA concentrations and with long homology arms. However, when providing multiple tDNAs, the untransformed fraction even increases and becomes greater than 50%. Recently, it was shown using whole genome sequencing of transformation events occurring during contact-dependent DNA uptake, that a single recipient could have at least 29 different recombination events (Cowley et al., 2018). Together, this suggests that many recombination events are likely going unnoticed in our single-cell transformation assay, and that these recombination events become limiting, as we can only detect successful recombination when the stop codon in our fluorescent reporter is replaced for a functional allele. If this is true, adding non-specific DNA would compete with donor tDNA resulting in reduced transformation efficiencies. To test this, we utilized homology-unrelated E. coli-derived DNA fragments of 5 kb with a similar GC content to S. pneumoniae as competing donor DNA. Indeed, as shown in Figure 7E, co-transformation of strain VL1803 (hlpA-stop-mScarlet-I, spv_1159-stop-sfGFP) with E. coli DNA significantly reduced the transformation efficiency. When 0.32 μM of hlpA-mScarlet-I tDNA alone was used as 7 kb donor DNA, approximately 43% of cells were transformed. However, when 0.32 μM of hlpA-mScarlet-I tDNA was given in the presence of saturating amounts of E. coli DNA (3.2 μM), only 3% of transformants were observed. Together, these data suggest that the fact that we never reach transformation efficiencies higher than 50% of the population even in the presence of multiple tDNAs, is because of saturation of the DNA uptake and integration machinery. The saturation can be caused by non-successful recombination events with donor DNA or successful recombination events with the donor DNA but outside of the stop SNP that cannot be detected in the single-cell assay (Figure 6—figure supplement 1).
So far, we demonstrated that transformation is highly efficient under ideal and saturated experimental settings in which pure PCR products were used as donor DNA. Previous studies showed that natural environments also promote efficient HGT (Cowley et al., 2018; Domenech et al., 2020). Indeed, in pneumococcal biofilms, which are a model for nasopharyngeal colonization, competence and fratricide are strongly induced, resulting in high relative concentrations of DNA. To assess transformation potential under more realistic conditions, we tested transformation efficiency in a co-culture system in which two pneumococcal strains are grown together on a semi-solid surface without adding synthetic CSP (Figure 8A; see Materials and methods section for detail procedure). HGT in such systems is based on genomic DNA released by dead cells followed by DNA uptake and transformation of competent recipient cells (Domenech et al., 2020). Here, we used strain R895 (cmR) as recipient and strain R4692 (∆comCDE, smR, novR) as donor. Both are genetically identical unencapsulated R800 derivatives (Lefevre et al., 1979) except for a single SNP conferring streptomycin (SNP in rpsL) or novobiocin (SNP in gyrB) resistance and a chloramphenicol resistance cassette present in the recipient R895 strain (Figure 8A). Strain R4692 is also unable to activate competence due to a comCDE deletion so transformation can only occur in one direction from donor (R4692) to recipient (R895). R895 and R4692 were pre-cultured separately until early exponential phase and then mixed in an approximate ratio of 3:7 (see Materials and methods). The mixture was immediately spotted on agar plates followed by incubation at 37°C for 4 hr to allow spontaneous competence development and transformation between strains. Cells were collected by scraping them from the plates and separated by sonication. Serial dilutions of the resulting cell suspension were plated with 4.5 µg/mL of chloramphenicol (for the recovery of the total number of viable recipient cells) and with combinations of chloramphenicol plus streptomycin (10 µg/mL) and/or novobiocin (4 µg/mL) (for the recovery of the single or double transformed recipient cells). As shown in Figure 8B, also in this more realistic model, very high transformation efficiencies are obtained with a single transformation efficiency with smR or novR of 5.70 × 10−2% (SD, 5.70 × 10−2%) or 1.75 × 10−2% (SD, 1.68 × 10−2%), respectively. Double transformation efficiency with both smR and novR was 8.01 × 10−5% (SD, 9.92 × 10−5%), which is close to the product of the single transformation efficiencies (5.70 × 10−2% × 1.75×10−2% = 9.9×10−4%). As a control, we also performed experiments using strain R4574 as donor (same genotype as R4692, but not harboring smR or novR allele), which never generated streptomycin nor novobiocin-resistant R895, demonstrating that de novo mutations conferring resistance do not occur in this experimental setup. Together, these experiments support our single- cell observations that multiple transformation events occur efficiently and independently even in more realistic settings within sessile co-cultures.
The species of S. pneumoniae is vastly diverse with a core genome of approximately 500–1100 orthologues genes and a pan-genome of 5000–7000 orthologs (Hiller and Sá-Leão, 2018). In addition, many genes are mosaic such as several genes encoding for penicillin-binding proteins in penicillin-resistant clinical strains (Hakenbeck et al., 2012). One of the main reasons for the high level of genome plasticity and rapidly changing population dynamics is because of the highly conserved competence-based transformation system present in nearly all pneumococcal genomes (Croucher et al., 2016). Indeed, rapid spread of antibiotic resistance alleles and capsule loci have been observed among human populations under selective pressure (Chewapreecha et al., 2014). Here, we investigated the molecular basis for competence-dependent transformation at the single-cell level and show that the uptake, integration, and expression of tDNA are highly efficient and are largely independent from the recipient’s cell-cycle stage or of the chromosomal position of the target locus. This was made possible by the setup of a sensitive real-time detection system to quantify successful homeologous recombination events. A major benefit of the here established single-cell approach over traditional plate-based assays is that it allows for the detection of more subtle effects and offers better resolution to study the kinetics of the processes involved. Indeed, using the system developed here, we could visualize and quantify the recombination of three different tDNAs in single recipient cells demonstrating the efficiency of the pneumococcal transformation process.
Genome sequencing has indicated that up to 29 recombination events may have taken place in a single round of transformation in the same cell when selecting for the transfer of an antibiotic resistance allele in S. pneumoniae (Cowley et al., 2018), while 40 recombination events have been reported in B. subtilis (Carrasco et al., 2016). Our work now provides direct evidence that this is not an anomaly and that multiple recombination events are possible during a single transformation episode, even in the absence of selection. Besides shedding light on the efficiency by which transformation can happen in S. pneumoniae, by imaging transformation at the single-cell level, we provide direct evidence that typically only one recipient strand is replaced during competence-dependent transformation, and that there is no bias toward replacement of the leading or lagging strand. As observed in V. cholerae, in approximately 7% of transformants, both strands can be replaced, which is likely caused by DNA repair leading to removal of the recipient strand on the hetero-duplex or by integration of multiple tDNAs (Dalia and Dalia, 2019). This is in line with predictions made using unlinked antibiotic resistance alleles (Porter and Guild, 1969). In addition, our single-cell observations suggest that the replaced noncoding strand by recombination within the hetero-duplex is immediately transcribed by RNAP and can lead to lineages of cells with nongenetic inherited phenotypes, or that the transformed allele is replicated and transcribed well before cell division occurs (Figure 3).
We show that any site regardless of its chromosomal position or orientation with regard to DNA replication can be efficiently transformed, although not with the exact same efficiencies (Figure 4—figure supplement 1). Possible explanations for local difference in recombination efficiency could be the levels of DNA compaction or transcription activity. As RecA-mediated DNA strand exchange is a reversible reaction in vitro (Dutreix et al., 1991; Konforti and Davis, 1990), under steady state conditions DNA strand exchange rarely reaches 50% efficiency. However, in vivo, when providing a single tDNA to competent cells, we readily reach 50% DNA strand exchange, again highlighting that this process is highly efficient under our experimental conditions.
Interestingly, we find that the percentage of untransformed cells is lower when three tDNAs are provided instead of two tDNAs (~58% vs ~49% of untransformed cells, respectively: Figures 6 and 7). Together with the observation that the presence of non-homologous DNA reduced our observed transformation efficiency (Figure 7E), suggests that, in principle, every recombination event is independent of the next, but that many unsuccessful recombination events and successful recombination events outside the stop codon of our reporter are taking place and that this limits the efficiency of site-specific recombination (Figure 6—figure supplement 1 and Figure 7F).
The overall biological implication of the limitation on competence-dependent transformation is that this mechanism ensures that in most cases one copy of the original recipient DNA remains unaltered. This might represent a fail-safe scenario so that in case a deleterious tDNA is incorporated, at least one daughter cell will survive. While this might be considered as a 'spandrel’ effect: a characteristic that flows inevitably from a selected phenotype but has not been selected for directly (Gould and Lewontin, 1979), being able to safely sample from a large pan-genome might contribute to the vast genome plasticity and genome diversity as observed in natural pneumococcal populations. Interestingly, we also find highly efficient HGT and independent transfer of genetic markers between a donor and recipient pneumococcal strain growing together on agar plates (Figure 8), indicating that our single-cell observations under laboratory conditions also reflect settings that depend on lysis of the donor cell and uptake of chromosomal DNA. It will be interesting to see how efficient competence-dependent transformation and HGT is under more realistic conditions such as within polymicrobial community within a host. Future single-cell work will allow the investigation of the localization of the enzymes involved in transformation, how strand exchange during transformation occurs and what the dynamics of the molecular machines are during DNA uptake, integration, and expression of tDNA.
All pneumococcal strains used in this study are derivatives of serotype 2 S. pneumoniae D39V (Avery et al., 1944; Slager et al., 2018) unless specified otherwise. See Supplementary file 1 for a list of the strains used. S. pneumoniae was grown in C+Y (pH 6.8) medium at 37°C. C+Y was adapted from (Adams and Roe, 1945) and contained the following compounds: adenosine (68.2 mM), uridine (74.6 mM), L-asparagine (302 mM), L-cysteine (84.6 mM), L-glutamine (137 mM), L-tryptophan (26.8 mM), casein hydrolysate (4.56 g L−1), BSA (729 mg L−1), biotin (2.24 mM), nicotinic acid (4.44 mM), pyridoxine (3.10 mM), calcium pantothenate (4.59 mM), thiamin (1.73 mM), riboflavin (0.678 mM), choline (43.7 mM), CaCl2 (103 mM), K2HPO4 (44.5 mM), MgCl2 (2.24 mM), FeSO4 (1.64 mM), CuSO4 (1.82 mM), ZnSO4 (1.58 mM), MnCl2 (1.29 mM), glucose (10.1 mM), sodium pyruvate (2.48 mM), saccharose (861 mM), sodium acetate (22.2 mM), and yeast extract (2.28 g L−1).
To construct YFP reporter for ssbB transcription, monomeric yfp (myfp) was introduced immediately downstream of ssbB at the native ssbB locus together with an RBS. myfp gene was amplified with OVL1414 and OVL1417 from genomic DNA of MK308 (parB::parB-yfp) (van Raaphorst et al., 2017). Upstream and downstream fragments were amplified with primer pairs of OVL166/OVL1196 and OVL1199/OVL167 using genomic DNA of VL599 (ssbB::ssbB_luc,kanR) (Slager et al., 2014) as template, respectively. The three resulting fragments were digested with BsmBI, ligated and transformed into S. pneumoniae D39V to obtain strain VL2219.
To construct translational fusion of comGA and msfYFP, linker-mYFP was amplified by PCR with OVL351/OVL690 from genomic DNA of MK308 (parB::parB-yfp) (van Raaphorst et al., 2017). ‘Upper’ and ‘downer’ fragments containing comGA were amplified with primer pairs OVL391/OVL392 and OVL691/OVL394 using genomic DNA of D39V as template, respectively. The three resulting fragments were fused by overlap PCR and transformed into S. pneumoniae D39V. Transformed clones were screened by PCR and sequenced. Resulting strains were additionally transformed by ssbB::ssbB_mScarlet-I, kanR fragments, obtaining strain VL2536.
To construct transcriptional fusion of ssbB and mScarlet-I, mScarlet-I was amplified by PCR with OVL1415/OVL1418 from genomic DNA of VL1787 (cep:spv_1159-mScarlet-I, spcR) (Keller et al., 2019). Upper and downer fragments were amplified with OVL166/OVL1168 and OVL1199/OVL167 using genomic DNA of VL599 (ssbB::ssbB_luc, kanR) as template, respectively. The three resulting fragments were fused by Golden Gate assembly using BsmBI and transformed into S. pneumoniae strain, comGA::comGA-msfYFP. Transformed clones were selected with kanamycin and sequenced, obtaining strain VL2536.
To construct translational fusion of comEA and msfYFP, mYFP-linker was amplified with OVL2029/OVL2028 from genomic DNA of VL1818 (comEC::msfYFP-comEC) (Veening lab collection). Upper and downer fragments containing comEA were amplified with OVL354/OVL1664 and OVL2026/OVL357 using genomic DNA of VL870 (comEA::mNeonGreen-comEA) (Veening lab collection) as template, respectively. The three resulting fragments were fused by Golden Gate assembly using BsmBI and transformed into S. pneumoniae D39V. Transformed clones were screened by PCR and sequenced, obtaining strain VL2537.
To construct a translational fusion of comGA and msfYFP, linker-mYFP gene was amplified with OVL351/OVL690 from genomic DNA of MK308 (parB::parB-yfp) (van Raaphorst et al., 2017). Upper and downer fragments containing comFA were amplified with OVL358/OVL521 and OVL1129/OVL361 using genomic DNA of D39V as template, respectively. The three resulting fragments were fused by overlap PCR and transformed into S. pneumoniae D39V. Transformed clones were screened by colony PCR and sequenced, obtaining VL2538 strain.
To construct a translational fusion of dprA and msfYFP, linker-mYFP gene was amplified with OVL3481/OVL3482 from genomic DNA of cep::spcR, P3_spv_1159-msfYFP (codon-optimized) strain (Rueff AS and Veening JW, unpublished). Upper and downer fragments containing dprA were amplified with OVL3487/OVL3488 and OVL3489/OVL3490 using genomic DNA of D39V as template, respectively. Erythromycin resistance marker (eryR) was amplified OVL2549/OVL2771 using genomic DNA of hexA::eryR strain (Veening lab collection). The four fragments were fused by Golden Gate assembly using BsmBI and transformed into S. pneumoniae D39V. Transformed clones were selected by erythromycin and sequenced, obtaining strain VL3355.
To construct hlpA-mScarlet-I, the hlpA-mScarlet-I gene was introduced downstream of the original hlpA gene at its own locus as a second copy of hlpA. Upper and downer fragments were amplified by PCR with OVL43/OVL44 and OVL45/OVL46 using genomic DNA of MK119 (hlpA::hlpA_hlpA-mKate2, cmR) (Beilharz et al., 2015), respectively. mScarlet-I gene was amplified by PCR with OVL55 and OVL56 using codon-optimized synthetic mScarlet-I gene as template (Keller et al., 2019). The three resulting fragments were fused by overlap PCR and transformed into S. pneumoniae. Transformed clone was selected by chloramphenicol, obtaining VL1780.
To disrupt translation between hlpA and mScarlet-I, on hlpA::hlpA_hlpA-mScarlet-I, cmR construct, single nucleotide mutation was introduced in domain breaking linker between hlpA and mScarlet-I. Upper or downer fragments were amplified by PCR with OVL43/OVL724 or OVL873/OVL46 using genomic DNA of VL1780 as template. The resulting fragments were fused by overlap PCR and transformed into S. pneumoniae D39V. Transformed clone was selected by chloramphenicol, obtaining VL1784.
To obtain VL1832 (hlpA::hlpA_hlpA-mScarlet-I, cmR; CEP::sfGFP, spcR), CEP::P3_sfGFP, spcR fragment was amplified by PCR with OVL37/OVL40 using genomic DNA of D-PEP33 (CEP::spcR, P3_sfGFP) (Sorg et al., 2015). The fragment was transformed into VL1784, and transformed clone was selected by spectinomycin, obtaining VL1832.
To construct membrane localizing sfGFP, hypothetical protein with transmembrane domain, spv_1159, was translationally fused to sfGFP under the control of synthetic constitutive promoter P3 at the CEP locus of the S. pneumoniae chromosome. Upper and downer fragments were amplified by PCR with OVL37/OVL631 and OVL634/OVL40 using genomic DNA of VL430 (CEP::spcR, P3_spv_1159-sfGFP) (Keller et al., 2019), respectively. spv_1159 was amplified by PCR with OVL632 and OVL633 using genomic DNA of D39V. The three resulting fragments were fused by Golden Gate assembly with BsmBI and transformed into S. pneumoniae D39V. Transformants were selected on Colombia agar plates containing spectinomycin, obtaining strain VL1785.
To obtain a dual-labeled strain, the CEP::spcR, P3_spv_1159-sfGFP fragment was amplified by PCR with OVL37/OVL40 using genomic DNA of VL1785, and transformed into VL1780 to obtain VL1800 (hlpA::hlpA_hlpA-mScarlet-I,cmR; CEP::spcR, P3_spv_1159-sfGFP).
To disrupt translation between spv_1159 and sfGFP, on the CEP::spcR, P3_spv_1159-sfGFP construct, a single nucleotide mutation was introduced in domain breaking linker between spv_1159 and sfGFP. Upper and downer fragments were amplified by PCR with OVL37/OVL724 or OVL723/OVL40 using genomic DNA of VL1785 as template. The resulting fragments were fused by overlap PCR and transformed into S. pneumoniae D39V. Transformants were selected by chloramphenicol, obtaining strain VL1786.
To obtain dual reporter strain VL1800, the CEP::spcR, P3_spv_1159-stop-sfGFP fragment was amplified by PCR with OVL37/40 using genomic DNA of VL1786, and transformed into VL1784 to obtain VL1800 (hlpA::hlpA_hlpA-stop-mScarlet-I, cmR; CEP::spcR, P3_spv_1159-stop-sfGFP).
To insert the spcR, P3_spv_1159-sfGFP construct downstream of the hlpA locus, spcR, P3_spv_1159-sfGFP was amplified by PCR with OVL2855/OVL2856 using genomic DNA of VL1785 (CEP::spcR, P3_spv_1159-sfGFP) as template. Upper and downer fragments were amplified by PCR with OVL2868/OVL2859 and OVL2860/OVL2869 using genomic DNA of D39V, respectively. The three resulting fragments were fused by Golden Gate assembly with BsmBI and transformed into S. pneumoniae D39V. Transformants were selected by spectinomycin, obtaining strain VL3096.
To disrupt translation between spv_1159 and sfGFP, on hlpA::spcR, P3_spv_1159-sfGFP construct, single nucleotide mutation was introduced in domain breaking linker between spv_1159 and sfGFP. Upper or downer fragments were amplified by PCR with OVL2868/OVL724 or OVL723/OVL2869 using genomic DNA of VL3096 as a template. The resulting fragments were fused by overlap PCR and transformed into S. pneumoniae D39V. Transformants were selected on Colombia agar plates containing chloramphenicol, obtaining VL3097.
To insert the spcR, P3_spv_1159-sfGFP construct right downstream of the comCDE locus, spcR, P3_spv_1159-sfGFP was amplified by PCR with OVL2855/OVL2856 using genomic DNA of VL1785 (CEP::spcR,P3_spv_1159-sfGFP) as template. Upper and downer fragments were amplified by PCR with OVL371/OVL2861 and OVL2862/OVL2870 using genomic DNA of D39V, respectively. The three resulting fragments were fused by Golden Gate assembly with BsmBI and transformed into S. pneumoniae D39V. Transformed clone was selected by spectinomycin, obtaining VL3098.
To disrupt translation between spv_1159 and sfGFP, on comCDE::spcR, P3_spv_1159-sfGFP construct, single nucleotide mutation was introduced in domain breaking linker between spv_1159 and sfGFP. Upper or downer fragments were amplified by PCR with primers OVL371/OVL724 or OVL723/OVL2870 using genomic DNA of VL3096 as template. The resulting fragments were fused by overlap PCR and transformed into S. pneumoniae D39V. Transformants were selected on Colombia agar plates containing, obtaining VL3099.
To insert the spcR, P3_spv_1159-sfGFP construct right at the bgaA locus, spcR, P3_spv_1159-sfGFP was amplified by PCR with OVL2855/OVL2856 using genomic DNA of VL1785 (CEP::spcR, P3_spv_1159-sfGFP) as template. Upper and downer fragments were amplified by PCR with OVL1312/OVL2863 and OVL2864/OVL2871 using genomic DNA of D39V, respectively. The three resulting fragments were fused by Golden Gate assembly with BsmBI and transformed into S. pneumoniae D39V. Transformed clone was selected by spectinomycin, obtaining VL3100.
To obtain strain VL3348, the bgaA::spcR, P3_spv_1159-sfGFP fragment was amplified by PCR with OVL1312/2871 using genomic DNA of VL3100, and transformed into VL1780 to obtain VL3348 (hlpA::hlpA_hlpA--mScarlet-I, cmR; bgaA::spcR, P3_spv_1159-sfGFP).
To disrupt translation between spv_1159 and sfGFP, on bgaA::spcR, P3_spv_1159-sfGFP construct, single nucleotide mutation was introduced in domain breaking linker between spv_1159 and sfGFP. Upper or downer fragments were amplified by PCR with OVL1312/OVL724 or OVL723/OVL2871 using genomic DNA of VL3096 as template. The resulting fragments were fused each other by overlap PCR and transformed into S. pneumoniae D39V. Transformed clone was selected by chloramphenicol, obtaining VL3101.
To obtain dual reporter strain, the bgaA::spcR, P3_spv_1159-stop-sfGFP fragment was amplified by PCR with OVL1312/2871 using genomic DNA of VL1786, and transformed into VL1784 to obtain VL3349 (hlpA::hlpA_hlpA-stop-mScarlet-I, cmR; bgaA::spcR, P3_spv_1159-stop-sfGFP).
To re-introduce spcR, P3_spv_1159-sfGFP in inverted direction at CEP locus, spcR, P3_spv_1159-sfGFP was amplified by PCR with OVL3358/OVL3359 using genomic DNA of VL1785 (CEP::spcR, P3_spv_1159-sfGFP) as template. Upper and downer fragments were amplified by PCR with OVL37/OVL3390 and OVL3391/OVL40 using genomic DNA of D39V, respectively. The three resulting fragments were fused by Golden Gate assembly with BsmBI and transformed into S. pneumoniae VL1784. Transformed clone was selected by spectinomycin, obtaining VL3346 (hlpA::hlpA_hlpA-mScarlet-I, cmR; CEP::spcR, P3_spv_1159-sfGFP(inverted)).
To disrupt translation between spv_1159 and sfGFP, on CEP::spcR, P3_spv_1159-sfGFP(inverted) construct, single nucleotide mutation was introduced in domain breaking linker between spv_1159 and sfGFP. Upper or downer fragments were amplified by PCR with OVL37/OVL723 or OVL724/OVL40 using genomic DNA of VL3346 as template. The resulting fragments were fused each other by overlap PCR and transformed into S. pneumoniae VL1784. Transformed clone was selected by chloramphenicol, obtaining VL3347 (hlpA::hlpA_hlpA-stop-mScarlet-I, cmR; CEP::spcR, P3_spv_1159-stop-sfGFP(inverted)).
To re-introduce spcR, P3_spv_1159-sfGFP in inverted direction at CEP locus, spcR, P3_spv_1159-sfGFP was amplified by PCR with OVL3358/OVL3359 using genomic DNA of VL1785 (CEP::spcR, P3_spv_1159-sfGFP) as template. Upper and downer fragments were amplified by PCR with OVL1312/OVL2863 and OVL2864/OVL2871 using genomic DNA of D39V, respectively. The three resulting fragments were fused by Golden Gate assembly with BsmBI and transformed into S. pneumoniae VL1784. Transformed clone was selected by spectinomycin, obtaining VL3346 (hlpA::hlpA_hlpA-mScarlet-I ,cmR; CEP::spcR, P3_spv_1159-sfGFP(inverted)).
To disrupt translation between spv_1159 and sfGFP, on bgaA::spcR, P3_spv_1159-sfGFP(inverted) construct, single nucleotide mutation was introduced in domain breaking linker between spv_1159 and sfGFP. Upper or downer fragments were amplified by PCR with OVL1312/OVL723 or OVL724/OVL2871 using genomic DNA of VL3346 as template. The resulting fragments were fused each other by overlap PCR and transformed into S. pneumoniae VL1784. Transformed clone was selected by chloramphenicol, obtaining VL3347 (hlpA::hlpA_hlpA-stop-mScarlet-I, cmR; CEP::spcR, P3_spv_1159-stop-sfGFP(inverted)).
To construct a triple labeled strain, upper or downer fragments were amplified with primer pair OVL452/OVL1921 or OVL1922/OVL1441 using genomic DNA of ftsZ::ftsZ-mTurquoise2, spcR strain (Gallay C and Veening JW, unpublished) as template. These fragments were fused by overlap PCR to remove the spcR gene. The fused fragment was used for transformation in D39V and spectinomycin-susceptible clone was selected.
To construct triple labeled strain, hlpA::hlpA_hlpA-mScarlet-I, cmR was amplified with OVL43/OVL46 using genomic DNA of VL1780 and cep::spcR, P3_spv_1159-msfYFP was amplified with OVL37/OVL40 using genomic DNA of cep::spcR, P3_spv_1159-msfYFP (codon-optimized) strain (Rueff AS and Veening JW, unpublished). These two DNA fragments were transformed into the strain (ftsZ::ftsZ-mTurquoise2) and transformed clone was selected by chloramphenicol and spectinomycin, obtaining triple labeled strain VL3126 (ftsZ::ftsZ-mTurquoise2; hlpA::hlpA_hlpA-mScarlet-I, cmR; cep::spcR, P3_spv_1159-msfYFP).
To construct triple reporter strain, upper or downer fragments were amplified with primer pair of OVL452/OVL724 or OVL723/OVL1441 using genomic DNA of VL3126 as template. These fragments were fused each other by overlap PCR to introduce stop codon between ftsZ and mTurquoise2. The fused fragment ftsZ::ftsZ-stop-mTurquoise2 was transformed in ftsZ::ftsZ-mTurquoise2 strain and clone that lost mTurquoise2 fluorescence was screened by fluorescence microscopy.
To disrupt translation between spv_1159 and msfYFP, on CEP::spcR, P3_spv_1159-msfYFP construct, single nucleotide mutation was introduced in domain breaking linker between spv_1159 and msfYFP. Upper or downer fragments were amplified by PCR with OVL37/OVL724 or OVL723/OVL40 using genomic DNA of VL3126 as template. While, hlpA-::hlpA-stop-mScarlet-I, cmR was amplified by PCR with OVL43/OVL46 using VL1784. The resulting two fragments were transformed into ftsZ::ftsZ-stop-mTurquoise2 and transformed clone was selected by chloramphenicol and spectinomycin, obtaining VL3128.
mCherry-eryR was amplified with RR93/RR94 using VL371 (RR27) (van Raaphorst et al., 2017). Upper or downer fragments were amplified by PCR with RR91/RR92 or RR93/RR94 using genomic DNA of D39V as template. The three fragments were assembled using Gibson one-step ISO assembly (Gibson, 2011) and transformed into D39V. Transformed clone was selected by erythromycin.
To monitor competence development, strains containing a transcriptional fusion of the firefly luc gene with the late competence gene ssbB were used. Cells were pre-cultured in C+Y (pH 6.8) at 37°C to an OD595 nm of 0.2. Right before inoculation, cells were collected by centrifugation (6000 x g for 3 min) and resuspended in fresh C+Y at pH 7.9, which is permissive for natural competence. Luciferase assays were performed in 96-wells plates with a Tecan Infinite 200 PRO illuminometer (TECAN) at 37°C as described before (Slager et al., 2014). Luciferin was added at a concentration of 0.45 mg/mL to monitor competence by means of luciferase activity. Optical density (OD595nm) and luminescence (relative luminescence units [RLU]) were measured every 10 min.
Microscopy acquisition was performed using a Leica DMi8 microscope with a sCMOS DFC9000 (Leica) camera and a SOLA light engine (Lumencor) and a 100x/1.40 oil-immersion objective. Images were primarily processed using LAS X (Leica). For snap shot imaging, cells were concentrated 10x by centrifugation (6000x g, 3 min) and 0.5 µL of cells were spotted on 1% agarose/PBS. For time-lapse microscopy, a semi-solid growth surface was prepared with C+Y (pH 7.9) containing 1% agarose in Gene Frame (Thermo Fischer) (de Jong et al., 2011). As C+Y medium has some background fluorescence, the C+Y agar pad was pre-exposed on a UV illuminator for 1 min to bleach the background fluorescence.
Phase contrast images were acquired using transmission light with 100 ms exposure for snap shot and 50 ms exposure for time-lapse. Fluorescence was usually acquired with 700 ms exposure for snap shot, and 200–500 ms exposure (17–30% of power from SOLA light engine) for time-lapse using filter settings described below. Time-lapses images were recorded by taking images every 5 or 10 min.
Leica DMi8 filters set used are as followed: mTurquoise2 (Ex: 430/24 nm Chroma, BS: LP 455 Leica 11536022, Em: 470/24 nm Chroma ET470/24 nm or Ex: 430/29 nm Chroma, BS: 455 (450–490) Chroma 69008, Em: 470/26), sfGFP (Ex: 470/40 nm Chroma ET470/40x, BS: LP 498 Leica 11536022, Em: 520/40 nm Chroma ET520/40 m), msfYFP (Ex: 500/20 nm Chroma ET500/20x, BS: LP 520 Leica 11536022, Em: 535/30 nm Chroma ET535/30 m or Ex:495/25 nm Chroma ET495/25x, BS520 (510–560) Chroma 69008, Em: 533/30 nm) and mScarlet-I (Chroma 49017, Ex: 560/40 nm, BS: LP 590 nm, Em: LP 590 nm or Ex: 575/35 nm, BS: 595 (590–670) nm Chroma 69008, Em: 635/70 nm). Microscopy images are available at the BioImages Archive (accession S-BIAD26).
For quantitative image analysis of single cells, obtained microscopic images were processed by FIJI software (Schindelin et al., 2012). Single-cell segmentation and fluorescence signal intensity measurement were performed by Oufti (Paintdakhi et al., 2016). The generated celllist files were analyzed in R (https://www.r-project.org/), using BactMAP (van Raaphorst et al., 2020) for statistical analysis and visualization. After celllist file were imported into R, cells were filtered between 0.7–1.2 µm in width length to exclude false events derived from noise or miss-segmentation. Threshold of fluorescence of signal intensity was defined based on negative or positive control for each experiment setting. >500 cells were analyzed at least for each replicate. To exclude the possibility of overlap in detection of fluorescence (particularly mTurquoise2/msfYFP and msfYFP/mScarlet-I) in multi-fragments transformation, we ensured that single transformation experiments did not show any signal in the other channels and this was confirmed by looking at the protein localization patterns.
For generating cell lineage trees from time-lapse imaging, the stacked time-lapse images were processed by FIJI and stabilization between time frames was performed by Huygens (Scientific volume imaging). Single-cell segmentation and fluorescence intensity acquisition were performed by SuperSegger (Stylianidou et al., 2016). The resultant data set was analyzed using BactMAP (van Raaphorst et al., 2020).
Analysis of DNA binding was performed in an endA mutant background (strain D39V ssbB::luc (cam) endA::kan), to favor accumulation of transforming DNA at the surface of competent cells. In wild- type, endA+ cells, surface-bound DNA is immediately internalized into the cytosol or degraded otherwise, which makes surface-bound DNA accumulation hard to be visualized as previously shown (Bergé et al., 2013). After gently thawing stock cultures, aliquots were inoculated at an OD550 of 0.006 in C+Y medium, supplemented with 20 mM HCl to prevent spontaneous competence development, and grown at 37°C to an OD550 of 0.3. These precultures were inoculated (1/50) in C+Y medium (pH 7.8) and incubated at 37°C. In these conditions, competence developed spontaneously and reached its maximal level in the population after 55–60 min. At 35 min, 1 ml samples were collected and induced, or not, with synthetic CSP (50 ng/ml). At 50 min, these samples were incubated for 5 min with 10 ng of a 285 bp DNA fragment labeled with a Cy3 fluorophore at its 5’ extremities (Bergé et al., 2013). Cells were pelleted (3000 x g, 3 min), washed twice in 500 µL C+Y, and resuspended in 20 to 50 µL C+Y medium before microscopy. Two μL of this suspension was spotted on a microscope slide containing a slab of 1.2% C+Y agarose as described previously (de Jong et al., 2011).
Phase contrast and fluorescence microscopy were performed with an automated inverted epifluorescence microscope Nikon Ti-E/B, a phase contrast objective (CFI Plan Apo Lambda DM 100X, NA1.45), a Semrock filter set for Cy3 (Ex: 531BP40; DM: 562; Em: 593BP40), a LED light source (Spectra X Light Engine, Lumencor), and a sCMOS camera (Neo sCMOS, Andor). Images were captured and processed using the Nis-Elements AR software (Nikon). Cy3 fluorescence images were false colored red and overlaid on phase contrast images. Overlaid images were further analyzed to quantify the number of cells bound with Cy3-labeled DNA. Single cells were first detected using the threshold command from Nis-Elements and cells bound or not to DNA were manually classified using the taxonomy tool.
To quantify the efficiency of transformation with tDNA fragments, reporter cells were pre-cultured in C+Y (pH 6.8) at 37°C to an OD595 nm of 0.2. Right before inoculation, cells were collected by centrifugation (6000 x g for 3 min) and resuspended in fresh C+Y at pH 7.9, adjusted to OD = 0.1. Competence was induced by incubation in the presence of CSP (100 ng/µL) at 37°C for 10 min and then donor tDNA was provided the indicated concentration. After an additional 4 hr of incubation at 37°C for complete cell division to form homo-duplex and maturation of fluorescence proteins, cells were placed on ice to stop cell growth and were directly analyzed by fluorescence microscopy or flow-cytometry.
Donor tDNA was designed in such a way that the single nucleotide mutation is positioned in the middle of the entire fragment so that the left and right homology arms are of equal length. Preparation of the tDNA was performed by PCR using primer pairs indicated in Supplementary file 2, using the corresponding parent strain as template. For competition experiments using unrelated tDNA as shown in Figure 7, a DNA fragment that has no homology to the pneumococcal genome but is of equal size (5 kb) and GC content (~40% of GC) to the hlpA-mScarlet-I/spv_1159-sfGFP fragments, was amplified by PCR from genomic DNA of E. coli DH5alpha (Hanahan et al., 1991), using primers OVL3527 and OVL3528.
Cells were collected by centrifugation (6000 x g for 3 min) and resuspended in filtered (0.22 µm) PBS adjusted to a cell density of approximately 1.0 × 105 to 1.0 × 106 cells/mL. As encapsulated S. pneumoniae D39V cells tend to form chains particularly during competence, we separated cells by bead beating (BioSpec) without any glass beads. At least >1.0×104 events were analyzed on a Novocyte flow cytometer (ACEA bioscience) harboring 488 nm and 561 nm lasers. Fluorescence filters used were: FITC (Ex: 488 nm, Em: 530/45 nm) for sfGFP and PE.Texas.Red (Ex: 561 nm, Em: 615/20 nm) for mScarlet-I. Obtained raw data were imported and analyzed in R. Non-bacterial particles were excluded by gating the FSC and SSC values. A threshold was determined so that positive events counted in a negative control strain were <1% and validated with both negative (no DNA control) and positive (parent strain without the point mutation) control for each experimental setting.
We used strain R895 as a recipient strain and R4692 and R4574 as donor strains. R895 is a naturally competent derivative of unencapsulated strain R6 and carries a chloramphenicol resistance marker (Chastanet et al., 2001). R4692 is unable to develop competence and carries point mutations conferring resistance to streptomycin, novobiocin and rifampicin. R4574 was used as a negative control donor strain. All strains were pre-cultured in C+Y (pH 6.8) at 37°C to an OD550 nm of 0.1. Cells were then collected by centrifugation (6000 x g for 3 min) and concentrated 3-fold in fresh C+Y at pH 7.9. Mixed inoculates containing 30 µL of donor strain R895 and 70 µL of recipient strain (R4574 or R4692) were subsequently spotted onto Petri dishes containing 3% horse blood CAT-agar (1% agar) supplemented with catalase (300 U/mL). Different ratios of donor and recipient were tested (1:1, 1:10 and 3:7) and a 3:7 ratio showed the least HGT variation across experiments. After 4 hr incubation at 37°C, cells were scraped off the plates and separated by sonication in an ultrasonic cleaner (90 s, 50 Hz). Serial dilutions were plated with 4.5 µg/mL of chloramphenicol and with combinations of chloramphenicol plus streptomycin (10 µg/mL) and/or novobiocin (4 µg/mL). Transformation efficiency was calculated by dividing the number of transformants by the total number of viable recipient count. Four independent biological replicates were performed.
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Petra Anne LevinReviewing Editor; Washington University in St. Louis, United States
Gisela StorzSenior Editor; National Institute of Child Health and Human Development, United States
In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.
This elegant and insightful paper develops a powerful new set of quantitative assays for tracking recombination in single cells to answer fundamental questions about natural transformation in the pathogenic bacterium Streptococcus pneumoniae; and thereby, resets paradigms in this research area. The paper shows that any site or orientation with regard to DNA replication can be transformed in pneumococcal cells, including multiple chromosomal insertions; however, there is an intrinsic limitation to the efficiency of recombination, possibly related to the level of off-marker recombination. This limitation likely has implications to pneumococcal genome evolution, affecting exchange of genes that determine virulence and antibiotic resistance.
Decision letter after peer review:
Thank you for submitting your article "Unbiased homeologous recombination during pneumococcal transformation allows for multiple chromosomal integration events" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by Petra Levin as Reviewing Editor and Gisela Storz as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Jason Rosch (Reviewer #1).
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, we are asking editors to accept without delay manuscripts, like yours, that they judge can stand as eLife papers without additional data, even if they feel that they would make the manuscript stronger. Thus the revisions requested below only address clarity and presentation.
All three reviewers felt that the manuscript was experimentally sound and praised the authors for their use of single cell analysis to tackle the question of why some cells are transformed and others are not in a population of genetically competent Pneumococci. The thoughtful presentation of complicated and extensive data was appreciated by all. Two reviewers were enthusiastic about the study's conclusions regarding bet hedging and the potential for an intrinsic limit on recombination efficiency, reducing the potential for off-marker recombination which, as reviewer #3 notes, might have implications to pneumococcal evolution. At the same time, reviewer #2 had some reservations about the significance of the data in light of previous studies of Pneumococcus and other naturally competent organisms. Most importantly, this reviewer questions whether the finding that only a portion of bacteria incorporate exogenous DNA is a particularly novel and, regardless, whether the saturating DNA concentrations used in the study are representative of a "natural" environment.
Given the discrepancy between the three reviewers with regard to the papers potential impact, we would like to request that you and your co-authors revise the paper, paying special attention to ensuring that the major conclusions are contextual both in terms of previous work and with regard to physiological relevance. I am including reviewer #2's comments in their entirety at the end of this letter to ensure you are aware of all concerns in this regard.
In this work Kurushima et al. use recently developed fluorescent labelling techniques to study natural transformation in the human pathogen Streptococcus pneumoniae. Previously, genetic marker analyses have been used to study the different aspect of this process, but with these new techniques the process can now be studied at the single cell level. The authors used the single cell analysis to identify new transformation bottlenecks and tried to determine why some cells are genetically transformed and others are not. Related experiments have been performed in the past using classic genetics and Kurushima et al. were able to confirm these studies. In that sense, in my opinion, the novelty is limited and no important new molecular insights are provided. They found that the number of cells that are ultimately transformed is plateauing at approximately 50%, despite the fact that most cells bind DNA. This is partially the result of the heteroduplex formed after recombination followed by separation by strand replication, combined with the fact that the DNA binding sites on cells are limited so that there is a competition between DNA markers at saturating DNA concentrations. The authors argue that this mechanism entails a "fail-safe strategy for the population as half of the population generally keeps an intact copy of the original genome". I find this conclusion far-fetched for two reasons. Firstly, the DNA recombination event followed by DNA replication will automatically assure that only half the population will inherit the mutation, and to speak of a strategy implies that the organism has specifically evolved this system, but we are dealing here with a well-known and general recombination system found in many organisms that will generally result in a 50/50 distribution. Maybe more importantly, under natural conditions it is highly unlikely that cells encounter saturating levels of tDNA. In their experiments the authors use 3.2 nM DNA for transformation. If my calculation is correct, this would amount to 19xE11 DNA molecules per ml, which seems a bit high when assuming tDNA comes from lysed bacteria. In nature, this number will be much (much) smaller therefore there is no need for the bacterium to come up with a dedicated strategy to assure that not all cells in a population are being transformed. Finally, the results are very well presented and the paper makes easy reading.https://doi.org/10.7554/eLife.58771.sa1
In this work Kurushima et al. use recently developed fluorescent labelling techniques to study natural transformation in the human pathogen Streptococcus pneumoniae. Previously, genetic marker analyses have been used to study the different aspect of this process, but with these new techniques the process can now be studied at the single cell level. The authors used the single cell analysis to identify new transformation bottlenecks and tried to determine why some cells are genetically transformed and others are not. Related experiments have been performed in the past using classic genetics and Kurushima et al. were able to confirm these studies. In that sense, in my opinion, the novelty is limited and no important new molecular insights are provided. They found that the number of cells that are ultimately transformed is plateauing at approximately 50%, despite the fact that most cells bind DNA. This is partially the result of the heteroduplex formed after recombination followed by separation by strand replication, combined with the fact that the DNA binding sites on cells are limited so that there is a competition between DNA markers at saturating DNA concentrations. The authors argue that this mechanism entails a "fail-safe strategy for the population as half of the population generally keeps an intact copy of the original genome". I find this conclusion far-fetched for two reasons. Firstly, the DNA recombination event followed by DNA replication will automatically assure that only half the population will inherit the mutation, and to speak of a strategy implies that the organism has specifically evolved this system, but we are dealing here with a well-known and general recombination system found in many organisms that will generally result in a 50/50 distribution. Maybe more importantly, under natural conditions it is highly unlikely that cells encounter saturating levels of tDNA. In their experiments the authors use 3.2 nM DNA for transformation. If my calculation is correct, this would amount to 19xE11 DNA molecules per ml, which seems a bit high when assuming tDNA comes from lysed bacteria. In nature, this number will be much (much) smaller therefore there is no need for the bacterium to come up with a dedicated strategy to assure that not all cells in a population are being transformed. Finally, the results are very well presented and the paper makes easy reading.
We thank the reviewer for hinting to us to look at more environmentally relevant scenarios. To this end, we performed a set of new experiments (new Figure 8) in which we grew two pneumococcal strains together on agar plates in which one of the strains cannot become competent and carries SNPs conferring streptomycin and novobiocin resistance, respectively. In this scenario, horizontal gene transfer occurs via lysis of the recipient and by transformation of the then liberated environmental chromosomal DNA (new Figure 8A). One could argue this experiment more closely mimics the natural polymicrobial biofilm-like niche pneumococci encounter in the human nose, compared to our in vitro experiments using PCR products as donor DNA. As shown in new Figure 8B, also in this experimental setup, transformation is very efficient, up to 5.70 x 10-2 (~0.05% of recipient cells are transformed) and markers transform independent of each other as demonstrated by our single cell analysis. We think this new experiment underscores that competence-based transformation is a main driver for pneumococcal genome evolution. We like to note that we do not think that the fail-safe has evolved as a dedicated strategy, but is rather a consequence of the recombination mechanism. This was mentioned in the Discussion as being a ‘spandrel effect’ and we have now further clarified this in the revised manuscript and changed fail-safe mechanism to fail-safe scenario.https://doi.org/10.7554/eLife.58771.sa2
- Jan-Willem Veening
- Jan-Willem Veening
- Patrice Polard
- Jun Kurushima
- Jan-Willem Veening
- Patrice Polard
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank Jelle Slager and Arnau Domenech for critically reading this manuscript, two anonymous referees, Juan Carlos Alonso and all members of the Veening lab for stimulating discussions. Work in the Veening lab is supported by the Swiss National Science Foundation (SNSF) (project grant 31003A_172861), a JPIAMR grant (40AR40_185533) from SNSF and ERC consolidator grant 771534-PneumoCaTChER. Work in the Polard lab is supported by the Centre National de la Recherche Scientifique, Université Paul Sabatier, and the Agence Nationale de la Recherche (grants ANR-10-BLAN-1331 and ANR-17-CE13-0031). Jun Kurushima was supported by The Naito Foundation.
- Gisela Storz, National Institute of Child Health and Human Development, United States
- Petra Anne Levin, Washington University in St. Louis, United States
- Jason Rosch
© 2020, Kurushima et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.
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