replication, partition/segregation, and cell wall growth/cell division (Helmstetter et al., 1968). The initiation of chromosome replication immediately precedes the initiation of chromosome partitioning and chromosome movement into separate cell spaces that will eventually become the daughter cells at cell division (Toro and Shapiro, 2010). This close temporal link suggests that it would be especially advantageous to co-regulate replication and partitioning. In a previous review article from our lab, we argued that eubacteria use one origin of replication (*ori*) per chromosome not because they are simpler organisms, but because a single *ori* allows for a more rapid and efficient control of replication (Marczynski et al., 2015). Bacterial chromosomes with one *ori* can more easily respond to many inputs both from outside and from inside the cell. We will argue that input/signals from inside the cell and crosstalk/ signals with chromosome partitioning (*par*) systems are especially important. While most studies illustrate *par* components signaling replication, we will also highlight recent studies of crosstalk in the reverse direction. However, before presenting concrete examples of crosstalk, we will first outline the basic features of both *ori* and *par* systems and emphasize their potential for regulation.

# ORIGINS OF REPLICATION RECEIVE SIGNALS AND DYNAMIC PROTEIN ASSEMBLIES

Like transcription promoters, bacterial *oris* are platforms for assembling replication proteins and their regulators (Kornberg and Baker, 1992). The *Escherichia coli oriC* and DnaA model for initiating chromosome replication has revealed the most detailed molecular mechanisms that operate inside *oris* (Kaguni, 2011; Skarstad and Katayama, 2013; Kaur et al., 2014). In broad outline, a bacterial *ori* is a specific place where the DnaA protein binds an array of DnaA boxes to self-assemble and then to promote the assembly of the downstream replication proteins (Wolanski et al., 2014a,b).

In *E. coli*, chromosome replication starts from one "*oriC*" when a threshold level of activated DnaA (ATP bound ATP-DnaA) is reached (Katayama et al., 2010; Skarstad and Katayama, 2013). Both forms of DnaA, ATP-DnaA and ADP-DnaA, bind to the strong/high affinity DnaA boxes in *oriC*, but only the activated ATP-DnaA proteins will bind to weak DnaA box motifs and oligomerize on *oriC* through neighboring AAA+ domains (McGarry et al., 2004; Kawakami et al., 2005; Erzberger and Berger, 2006; Grimwade et al., 2018). Such DnaA self-assembly starts from strategically placed "anchor" DnaA boxes (Rozgaja et al., 2011), and the resulting protein-DNA structure (and possibly a helix) causes DNA unwinding and a further altered structure with new protein surfaces that recruit downstream replication proteins. More specifically, *oriC* DNA unwinding allows DnaA to recruit DnaB (the replicative DNA helicase) bound to DnaC, the helicase escort/loader, on to the single-stranded DNA of the AT-rich region (Mott and Berger, 2007). It is likely that two types of DnaA protein-DNA structures form on *oriC*; one that unwinds and keeps the AT-rich region open and single stranded and another DnaA-DNA structure that recruits and loads two DnaB hexamers around the single-stranded DNA. Once loaded, the two DnaB hexamers move apart, expanding the single-stranded DNA region, thereby permitting the recruitment of primase DnaG. Next, the DNA polymerase III holoenzyme composed of the Pol III and the beta-clamp (DnaN) is recruited, and together with the clamp-loading proteins, these form the "replisome" that synthesizes the complementary DNA strands (Kaguni, 2011; Skarstad and Katayama, 2013; Katayama, 2017).

Since most eubacteria use the DnaA protein to initiate chromosome replication (Wolanski et al., 2014a,b), DnaA and the assembly reactions at *oriC* are major targets for the regulators of chromosome replication (Wolanski et al., 2014a,b). Recent reviews have described many proposed and established regulators of replication, and an especially good review with fine graphic summaries was provided by Katayama et al. (2010).

Most importantly for our topic, DnaA assembly at *oriC* is dynamic, and *in vivo* there is probably both back and forth assembly and dis-assembly of DnaA until the critical amount of DnaA oligomerization and active structure formation is reached (Leonard and Grimwade, 2011; Kaur et al., 2014). This dynamic feature of *E. coli* replication initiation implies that there are many ways to shift the assembly versus dis-assembly of DnaA and DnaB. This process has the potential to integrate many signals that can be constantly added or subtracted in real time before the final commitment to replication is made. We will describe below how this view of dynamic *oriC*/DnaA assemblies helps us to understand the regulatory crosstalk with chromosome partitioning.

# DNA PARTITIONING SYSTEMS

Many bacteria use systems often called "*parABS"* for mitoticlike chromosome separation and partitioning into cell compartments, and their proximity to origins of replication (*oris*) suggests functional linkages (Livny et al., 2007). These partitioning systems were originally studied on large low-copy plasmids, and they account for faithful and consistent plasmid distribution to both progeny cells (Austin and Abeles, 1983; Ogura and Hiraga, 1983; Gerdes et al., 1985). Despite much effort, exactly how the *parABS* systems work to move and to position plasmid and chromosome DNAs remains incompletely understood and in parts controversial (Gerdes et al., 2010). Here we want to present the basic information and sketch what appear to us the most relevant models for our topic. Knowledge of the detailed mechanisms is required not just to understand how *parABS* systems work to partition DNA but also to understand and speculate how evolution has harnessed these systems for other functions and particularly for crosstalk with chromosome replication. With respect to deep evolutionary potentials, *parABS* systems have also been harnessed for protein positioning and localization, as, for example, organizing chemotaxis proteins and other large protein assemblies (Vecchiarelli et al., 2012).

In bare outline, the three-component *parABS* system works as follows: The *parS* DNA acts as a "centromere-like" locus with specific DNA sequences that bind and hold ParB proteins. ParA protein binds and hydrolyses ATP, and it somehow imparts motion to the ParB-*parS* complex through interactions with ParB. These basic functions need to be controlled and organized by regulators and structures that change during the cell cycle. As a main topic, we will address some key regulators and structures below, including, for example, the cell-pole proteins that anchor the chromosome ParB-*parS* complexes.

There are several significant variations to the above bare outline of *parABS* system. For example, some bacteria apparently use several *parS* loci, while others appear to use just one. *Bacillus subtilis* probably uses 10 *parS* loci and 8/10 loci cluster toward the *oriC* side of the chromosome (Breier and Grossman, 2007). *Myxococcus xanthus* may use as many as 22 *parS* loci, likewise near its *oriC*, for partitioning its exceptionally large 9.1 Mb chromosome (Iniesta, 2014). In contrast, the *Caulobacter crescentus* (Mohl and Gober, 1997) and the *Vibrio cholerae* (Espinosa et al., 2017) chromosomes appear to use just one *parS* per chromosome, and these single *parS* loci are likewise closely linked to their corresponding *oriCs*. Why does one bacterium need one *parS* and another several? There is no good answer yet, but this distinction may be too simplistic. For example, a recent study showed that *C. crescentus* has several yet substantially weaker ParB-binding sites (Tran et al., 2018), and it may be more correct to speak of a "*parS* region" surrounding the origin of replication as described further below.

There are also significant variations in how ParB binds DNA to create a "centromere-like" locus. ParB binds specifically to *parS* DNA and less specifically to other parts of the chromosome. First, ParB binds specifically to an inverted DNA repeat that is typical of many standard dimeric helix-turn-helix DNA-binding proteins, and these sites are easily found and used to identify *parS* sites in most bacterial genomes (Livny et al., 2007; Iniesta, 2014). However, ParB is reported to have additional modes of DNA binding. *In vivo* cross-linking and transcription reporter experiments imply that ParB binds to *parS* sites and then spreads to adjacent DNA as if forming a filament across the DNA to distant sites. It is not likely that "spreading" is an experimental artifact because spreading is required for partitioning. ParB mutants that do not spread do not partition DNA (Rodionov et al., 1999; Graham et al., 2014).

The exact DNA/protein structure(s) of these "spreading" ParB molecules is not known, but interactions can be inferred from crystal structures (Chen et al., 2015). ParB can bind other ParB molecules through lateral contacts that reach adjacent DNA and through bridging contacts that bring distant DNAs together with loops. This capacity for non-specific DNA binding suggests that ParB can be classified as one among many nucleoid-associated proteins (NAPs) that compact and organize bacterial chromosomes. Recently, a ParB "caging model" has been proposed whereby *parS* organizes a large chromosome subdomain through dynamic ParB-ParB and ParB-DNA interactions (Funnell, 2016). This model is further supported by *in vitro* experiments with magnetic tweezers, which suggest that the overall ParB-DNA complex is not well ordered and vaguely resembles a phase separation from the rest of the nucleoid (Taylor et al., 2015). In summary, considering the proximity of *parS* to *oriC*, ParB protein certainly has the potential to influence chromosome replication, and we will describe specific examples below.

#### PARTITION PROTEIN PARA CAN BE A MOTOR AND A REGULATOR

The preceding observations argue that ParA imparts motion not just to a small ParB-*parS* locus but also to a large ParB-DNA subdomain of the chromosome. Exactly how ParA drives ParB-DNA motion also remains controversial. However, ParA has several established and speculative properties that enable it to serve both as a propeller and as a regulator. We will focus below on two properties required for regulation: First, we explain that ParA can act like a "molecular switch" and second, we explain that ParA (like ParB) can bind and influence large domains of DNA.

ParA "switches" within a biochemical cycle: ParA monomers bind ATP, the ParA-ATP dimerizes, and this form binds DNA non-specifically. ATP hydrolysis creates ParA-ADP molecules, which disassociate from the DNA as monomers. When ParA binds ParB, specific protein-protein contacts stimulate ATP hydrolysis, thereby resetting the ParA-ATP/DNA binding versus ParA-ADP/DNA release cycle (Vecchiarelli et al., 2010). A protein contact switch seems ideal for regulation, and as an interesting example, we will describe below how *Bacillus subtilis* has harnessed ParA to also regulate chromosome replication through direct contacts with DnaA.

Exactly how this ParA cycle drives ParB-DNA motion remains controversial. It is also not clear if propulsion and switching/ regulation are separable functions. Here we can only superficially comment on this literature, and we will focus on how ParA binds to the nucleoid. For example, it has been proposed that ParA binds ParB and then retracts to pull the ParB-DNA along its path. This could be an active process where ParA imparts the force of motion or it could be a more passive mechanism, for example, a "catch and release" mechanism whereby ParA guides and biases a random "DNA flapping" motion. ParA may be organized as "microtubule-like" or as "cloud-like" structures that move forward and recede by assembly and dis-assembly. The literature is not consistent. However, there are credible reports that during partition, ParA forms dynamic cloud-like patterns on the surface of the nucleoid, and this pattern is interpreted as a gradient that recedes and seems to draw the ParB bound to *parS* (Hatano and Niki, 2010; Ah-Seng et al., 2013). Nucleoid patterning by ParA proteins resembles membrane patterning by the *E. coli* MinCDE system (Vecchiarelli et al., 2012), which imparts positional information for cell division, so that the septum forms at mid-cell (Lutkenhaus, 2007). Furthermore, the ParA and Min proteins belong to the same class of ATPases, and their mechanisms for molecular positioning may be fundamentally similar (Vecchiarelli et al., 2012).

Ietswaart et al. have presented an important synthesis between what seemed at first to be distinct and contradictory par mechanisms (Ietswaart et al., 2014). They demonstrate that the *par* system stimulates plasmid DNA motion above the random Brownian motion kinetics, thereby demonstrating that the *par* system can impart an active force and does not simply bias a random motion. Also, very importantly, Ietswaart et al. have argued that nucleoid structure plays an essential role in ordering the bound ParA-ATP structures. For example, helical nucleoid folds might provide grooves for channeling ParA-ATP aggregates into filaments or elongated clouds. Their model requires linear arrays of DNA-bound ParA-ATP and not necessarily that they be microtubule-like filaments. In other words, ParA-ATP linearity imparts the directionality to DNA motion and short disjoint filaments or elongated clouds (where individual ParA-ATP dimers bound to the nucleoid need not touch) will equally satisfy their model. In summary, the *par* literature argues that both ParA and ParB shape and respond to the structure of the nucleoid. Consequently, NAPs should significantly impact both chromosome replication and its partitioning. We will therefore discuss NAPs as regulators further below.

# ESTABLISHED EXAMPLES OF CROSSTALK: THE *BACILLUS SUBTILIS* SYSTEM

*Bacillus subtilis* provides clear examples of crosstalk and a series of papers provide the best and earliest evidence. For example, early studies showed that *B. subtilis*, Spo0J(ParB), is required for the normal positioning of the *oriC* region and for restricting its replication. Wild type cells prior to replication place their *oriC* regions at the lateral mid-cell position and when they duplicate their *oriC* regions, they position them around the cell quarter-length positions. However, in *spo0J(parB)-*null strains, the duplicated *oriC* regions are positioned significantly closer together and toward the mid-cell. Interestingly, these *spo0J(parB)* null strains had more *oriC* DNA per cell, as determined by flow cytometry. Apparently, *spo0J(parB)-*null cells had increased chromosome content from an excessive and/or an asynchronous initiation of DNA replication from *oriC* (Lee et al., 2003).

One general question is whether asynchronous firing of *B. subtilis oriC* was caused indirectly by *oriC* mislocalization or whether the ParAB system directly interacts with the replication system. Later studies showed that the *B. subtilis* ParAB proteins directly target DnaA (Murray and Errington, 2008). Using fluorescence-tagged ParA and ParB proteins, Murray and Errington showed that these proteins dynamically localize as specific foci (spots) near *B. subtilis* cell poles and nucleoids and that ParA can both inhibit and activate DnaA to alter chromosome replication. The inferred cytogenetic interactions between ParA and DnaA were supported by direct *in vivo* crosslinking and two-hybrid assays. In addition to this direct mechanistic link, this article also made several other interesting observations: For example, *parA*-null mutants behave like wildtype cells arguing for redundant or multiple regulatory inputs. Revealing the hidden cell-cycle interactions required assaying mutant protein forms. For example, revealing DnaA-dependent ParA foci at *oriC* required expressing a fluorescent ParA protein that bound ATP but did not bind DNA. Presumably, the weaker binding of ParA to DnaA protein at *oriC* would be otherwise obscured by its stronger binding to the larger/bulkier chromosome DNA. Similarly, revealing ParB-dependent ParA foci required fluorescent ParA that was deficient for ATPase and therefore apparently remained bound for longer times to the DNA.

Furthermore, the cell-cycle roles of ParAB were originally hidden because *parAB* mutants were first classified as sporulation genes. ParB was called Spo0J because null alleles were blocked in the earliest 0-stage of sporulation. ParA was called Soj, "suppressor of spo gene J," because its null alleles allowed sporulation of spo0J null strains (Ireton et al., 1994; Quisel and Grossman, 2000). We now know that sporulation is inhibited by ParA (Soj), which requires ParA-ATP dimerization and that ParB (Spo0J) counteracts ParA (Soj) by stimulating ParA-ATP hydrolysis. Murray and Errington also showed that ParA (Soj) acts through the Sda-dependent DNA replication checkpoint (Murray and Errington, 2008). Sporulation is not just a simple response to starvation. Sporulation also requires passing several checkpoints and conditions that perturb chromosome replication block sporulation by expressing a sporulation inhibitor, Sda (Ruvolo et al., 2006). Most interestingly, the transcription promoter of *sda* has many DnaA boxes, and like *oriC*, it essentially acts as a sensor for DnaA activity. In other words, one had to look through one layer of regulation (Sda check point regulation) to see the other layer of *oriC*/DnaA regulation. Note also that both sporulation and chromosome replication are long processes that require a "full commitment" following a "deliberation process" with multiple inputs, and that evolution has recruited DnaA in both cases as an integrating component.

Subsequent studies showed how ParA changes DnaA oligomerization at the *B. subtilis oriC*. For example, Scholefield et al. showed that the initiation of chromosome replication is inhibited by monomeric ParA-ADP (Soj) and conversely activated by dimeric ParA-ATP (Scholefield et al., 2011). This study also identified specific amino-acid contacts on coregulator ParB (Spo0J) that touch ParA and "flip the switch" to its inactive form. Next, in their following paper, Scholefield et al. demonstrated specific amino-acid contacts between ParA and DnaA with both molecular-genetic and biochemical (e.g. SPR sensorgram and crosslinking) experiments. Most impressively, this study showed that monomeric ParA represses *oriC* replication by depolymerizing DnaA (Scholefield et al., 2012). These experiments used a functional double-cysteine version of DnaA (DnaA-CC) that allowed stable crosslinking of the DnaA-CC oligomers during *in vitro* and *in vivo* experiments. These oligomers presumably reflect the assembly of the DnaA*oriC* DNA complexes, and their summary model implies that monomer ParA acts as a negative input during the dynamic assembly and dis-assembly process that tips *oriC* either toward or away from replication.

Recent microscopic studies have more directly confirmed this rapid assembly and dis-assembly model of DnaA at *B. subtilis oriC* and the proposed regulatory roles of ParA (Soj) in this dynamic process (Schenk et al., 2017). More specifically, "FRAP" fluorescence recovery and photobleaching analysis of a functional fluorescent YFP-DnaA protein showed that DnaA is bound to *oriC* with a short half-time of only 2.5 s. As predicted, a genetic deletion of *parA* (*soj*) increased the DnaA residence time at *oriC* and this in turn caused over-replication of the chromosome, presumably by shifting the equilibrium more frequently toward DnaA-*oriC* DNA complex formation. Furthermore, single-molecule YFP-DnaA microscopy showed that DnaA oscillates between polar-oriented *oriC* foci with a very short ~2 s periodicity. This last observation unexpectedly shows that DnaA can behave more like the *par* and *min* (cell division) system proteins than previously suspected (Schenk et al., 2017).

The overall view that emerges from these studies is that ParA (Soj) is an important *oriC*/DnaA regulator or more accurately, a key regulatory input. This regulation is not essential but instead seems to fine tune the cell cycle in growing cells and their timely exit into sporulation. ParA (Soj) can either delay or advance the start of *oriC* replication depending on its monomer versus dimer states and its contacts with ParB (Spo0J). However, exactly how these factors link *oriC*/DnaA regulation to chromosome movements and perhaps to other cell-cycle processes remains vague and speculative.

# ESTABLISHED EXAMPLES OF CROSSTALK: THE *VIBRIO CHOLERAE* SYSTEM

*Vibrio cholerae* presents another interesting, evolutionary very divergent and well-studied system for addressing chromosome replication and partitioning. This topic has recently been well reviewed (Espinosa et al., 2017). *V. cholerae* is closely related to *E. coli*, and while these bacteria have expected similarities, they also have some very surprising differences. For example, the *V. cholerae oriC* and the *E. coli oriC* seem to function and use DnaA very similarly. However, unlike *E. coli*, *V. cholerae* has two chromosomes, one replicated by an *E. coli*-like *oriC* (Chrom I) and the other by a distinct plasmid-like origin of replication (Chrom II). The *V. cholerae* Chrom I and *E. coli oriCs* have identical DnaA box distributions, and as expected, DnaA is the primary initiator (Egan and Waldor, 2003). In contrast, the *V. cholerae* Chrom II *ori* has only one DnaA box, and it instead uses an "iteron" organization, i.e., a long array of binding sites for the initiator protein RctB (Gerding et al., 2015). Yet, despite such major differences both Chrom I and II are well integrated into the *V. cholerae* cell cycle, and their replication is strictly timed (Espinosa et al., 2017).

*V. cholerae* Chrom I and II have evolved separate and specific replication and partitioning crosstalk systems. For example, the control of chromosome replication through ParA and ParB, seen above in *B. subtilis*, also seems to apply to the large chromosome (Chrom I) of *V. cholerae* (Kadoya et al., 2011). Interestingly, each Chrom I and II has its own chromosomespecific *parABS* system. Accordingly, Chrom I has corresponding *parA1*, *parB1,* and *parS1* linked to its *E. coli*-like *oriC*. Prior to the start of replication, this *V. cholerae par/oriC* DNA region is positioned at the cell pole. Deletion of either *parA1* or *parS1* caused delocalization away from the cell pole. Deletion of *parB1* caused a similar delocalization as expected, plus an increased *oriC* copy number indicating that lack of ParB1 causes overreplication. Therefore, as in *B. subtilis*, ParB1 limits ParA1 activity, which then presumably targets *oriC* through DnaA. This view is supported, by, for example, double *parB1* and *parA1* deletions, which reduce and restore approximately normal levels of *oriC* replication presumably by eliminating the stimulus of ParA1-ATP dimers. Unfortunately, direct evidence for ParA1 and *V. cholerae* DnaA interactions is lacking. It is tempting to speculate that like *B. subtilis* ParA (Soj), the *V. cholerae* ParA1 also directly contacts the AAA+ domain of DnaA and more specifically that it too both stabilizes and destabilizes the DnaA structure on *oriC*. However, there are many ways to regulate DnaA activity, and considering the evolutionary distance between Gram (+) and Gram (−) bacteria, other mechanisms are likely, and the details of this broad outline need to be investigated.

The *V. cholerae* (*Vc*) Chrom II system is significantly different from Chrom I: Its *ori* is flanked by two genetic loci *rctA* and *rctB* (Egan and Waldor, 2003). While *rctB* simply encodes the DNA-binding initiator protein, the *rctA* locus seems to be a complex regulatory system with the *Vc parS2* "centromere" embedded among its regulatory elements (Gerding et al., 2015). Also, Chrom II seems to have an interesting parallel regulation with that of the *Caulobacter crescentus* (*Ccr*) chromosome, which will be described further below: As with most *parABS* systems, the *Vc parS* centromere locus in *rctA* binds *Vc* ParB2 and the *Ccr parS* binds *Cr* ParB. However, very interestingly, both centromere loci also bind their main replication initiator proteins, *Vc* RctB (Gerding et al., 2015) and *Ccr* DnaA, respectively (Mera et al., 2014). This is probably an example of convergent functional evolution, since *Vc* RctB and *Ccr* DnaA are otherwise unrelated.

Despite these two clear examples of crosstalk, the details of their mechanisms, as far as they are known, appear to be very different. The details of *Ccr parS* and *Ccr* DnaA interactions will be described further below in the context of cell-cycle control. Here we will note some mechanistic similarities and differences. For example, the *rctA/parS2* locus of *Vc* Chrom II binds RctB protein and seems to repress replication by titrating RctB away from the nearby origin of replication (Yamaichi et al., 2011). This is clearly different than *Ccr* DnaA protein that binds *parS* to apparently trigger DNA movement. Also, RctB has at least two separate DNA-binding domains (Yamaichi et al., 2011), one to bind *rctA* DNA and the other to bind the iteron motifs inside the adjacent Chrom II *ori*. In contrast, DnaA uses its single domain IV to bind DnaA boxes in both *parS* and *ori* DNA (Mera et al., 2014). Moreover, *rctA/parS2* seems to be a more complex locus. Its small ORF does not seem to encode a functional protein, and instead, it seems to function by providing an RNA molecule, or as a platform for transcription activity (perhaps to alter DNA topology), and as a platform for binding proteins, including ParB2 (at the main *parS2* sequences) and RctB. Both ParB2 and RctB can bind and simultaneously occupy *rctA* DNA in what appears to be adjacent binding zones (Yamaichi et al., 2011). ParB2 binding to *rctA* DNA counteracts *rctA* repression of replication, yet ParB2 protein does not seem to displace the bound RctB protein. This last observation argues that simple RctB protein titration away from the *ori* does not obviously explain how the *rctA* locus acts through RctB protein to repress replication or how ParB2 binding counteracts this effect. A fuller explanation is needed, and it may need to invoke altered protein and DNA structures.

Separate studies confirm the preceding antagonistic relationships among *rctA/parS2*, ParB2, and RctB, but the inferred mechanism does not involve RctB titration (Venkova-Canova et al., 2013). Instead, it was argued that RctB binds short 12-mer DNA sequences to activate replication and to longer 39-mer DNA sequences to repress replication. Apparently, ParB2 has two ways to relieve this repression. In the first way, ParB2 binds at *rctA/parS2* and spreads laterally across the DNA into a nearby 39-mer, thereby displacing RctB and relieving its repression. In the second way, ParB2 has a secondary intrinsic affinity for the 39-mer DNA, and so ParB2 competes for RctB repressor binding at a distant 39-mer without the spreading mechanism from *parS2*.

Furthermore, RctB and ParB2 provide a second level of crosstalk since they control transcription of the downstream *parAB2* operon. As observed in similar *par* systems, ParB2 binds *parS2/rctA* and auto-represses the *parAB* operon. However, RctB binding stimulates transcription, thereby increasing ParB2. Therefore, RctB and ParB2 have mutually antagonistic effects on both *parAB2* operon transaction and on Chrom II replication (Yamaichi et al., 2011; Gerding et al., 2015). Overall, these observations suggest a dynamic back and forth switch between *par* and *ori* control that is yet to be fully understood.

In summary, the *V. cholerae* two chromosome system provides interesting examples of *ori* and *par* crosstalk. At Chrom I, evolution has apparently conserved the ParB1, ParA1, and DnaA signaling pathway between *parS1* and the origin of replication. However, at Chrom II, evolution has modified the paralogous ParB2 protein to interact more directly with a very different type of origin of replication through direct contact or through competition with its iteron-binding protein RctB.

## THE *CAULOBACTER CRESCENTUS* CELL-CYCLE MODEL FOR CROSSTALK

*Caulobacter crescentus* provides further evidence of *ori* and *par* crosstalk. As a chief advantage, this bacterium allows crosstalk studies in the context of a synchronized and well-studied cell cycle (**Figure 1A**). This is a "di-morphic" cell cycle where the transition from the "swarmer cell" to the "stalked cell" also marks the key steps of replication and chromosome partitioning. Conceptually, the cell cycle starts with the motile and non-replicating swarmer cell. Its chromosome replication is blocked by the CtrA regulator with five-binding sites inside the *C. crescentus* origin of replication (*Cori*) (Siam and Marczynski, 2000). The *C. crescentus parS* is only about 8 kb from *Cori* (Mohl and Gober, 1997), and this whole region of the chromosome is polarized and held near the flagellated cell pole by *parS-*binding ParB, which in turn is bound to a polar matrix protein called "PopZ" (Bowman et al., 2008). We will describe PopZ further below and argue that it can serve as a "hub" for many regulatory interactions, but the most conspicuous role for PopZ is to serve as the substrate that binds ParB, which thereby anchors the *parS* and *Cori* region in the swarmer cell (Bowman et al., 2008).

The cell-cycle transition from swarmer cell to stalked cell coincides with many molecular events that suggest *Cori* and *parS* crosstalk (Marczynski et al., 2015). While the swarmer cell ejects its flagellum and starts to grow its stalk (a tubular cell wall outgrowth), the CtrA protein is inactivated (dephosphorylated) and degraded, the *parS*-*Cori* region detaches from the cell pole, and the chromosome initiates replication from *Cori* (Toro and Shapiro, 2010). The initiation of chromosome partitioning is practically simultaneous with the initiation of chromosome replication, and both processes continue through most of the cell division cycle. Note especially that the dividing cell poles are different, one pole has a stalk, while the other is building a new flagellum, so this is an "asymmetric" cell division cycle (**Figure 1**). Eventually, one whole chromosome is placed in the nascent swarmer cell compartment, while the other chromosome is placed in the stalked cell compartment. In other words, with respect to chromosome replication, one chromosome will be placed into an inactive swarmer cell, and the other chromosome will be placed into an active stalked cell.

Such asymmetric cell division implies that the initiation of chromosome replication and partitioning coincide with the critical chromosome symmetry-splitting step of the cell cycle (**Figure 1A**). Time-lapse fluorescence microscopy showed that *parS-Cori* region DNA partitioning (visualized with fluorescent ParB) is a complex process involving at least the following steps: First *parS-Cori* separation, then *parS-Cori* discrimination, such that one *parS-Cori* region seems to be chosen for reattachment to the stalked pole. Then, the other (apparently unattached) *parS-Cori* region moves slowly away from the stalked pole to approximately the quarter-cell length position before moving more rapidly to the new swarmer pole (Shebelut et al., 2010). Further analysis showed that only the last fastmovement phase requires ParA ATPase activity and that the early slow movement of *parS-Cori* to the quarter-cell length position occurs faithfully when a dominant-negative ParA allele is expressed (Shebelut et al., 2010). Since fluorescently labeled ParB is bound to *parS* during this early slow-movement phase, then how does *parS*-ParB move without ParA?

More importantly, how do these early partitioning steps faithfully split chromosome symmetry to channel them toward two different cell fates (**Figure 1A**)? What are the regulators and the motors during the early partitioning steps? How do they communicate with chromosome replication? These questions are starting to be addressed in the following paragraphs.

(*Cori*). The swarmer cell next differentiates into a non-motile and replicating stalked cell (St). Coincident with this cell differentiation, the chromosome replication and partitioning phases initiate apparently simultaneously, and they continue together for much of the cell cycle. The partitioning movement of *parS-Cori* has an initial slow phase that uses GapR protein and a later fast phase that requires the partitioning protein ParA (see text for further details). This slow partitioning phase overlaps the chromosome "symmetry breaking" step (\*) of the cell cycle, which symmetrically channels the duplicated *parS-Cori* regions and eventually the entire chromosomes into distinct replicating (stalked cell) and non-replicating (swarmer cell) compartments. The blue cytoplasmic shading represents the activity (presence and phosphorylation) of the master cell-cycle regulator CtrA, which among many functions bind *Cori* to repress replication in swarmer cells. Asymmetric cell division (Div) proceeds with the return of CtrA activity and the building of a new polar flagellum. Eventually the two distinct cell types are formed. (B) A closer look at the cell poles during the above cell cycle. On the left, an early stalked cell pole where the *parS-Cori* region has been released from the PopZ matrix protein (not shown) and where rising DnaA activity first acts at *parS* DnaA boxes before acting at the *Cori* DnaA boxes. Although GapR binds broad regions of the chromosome, its strongest peaks are around the *parS-Cori* DNA. Next, a stalked cell pole immediately following the initiation of chromosome replication. One duplicated *parS-Cori* region reattaches to the old PopZ matrix at the stalked pole (symbolized by the broad arrow, the ParB bridge is not shown). The other duplicated *parS-Cori* region moves slowly away with the aid of GapR before its fast movement driven by ParA toward the other cell pole. On the right, both poles of a dividing cell. At the swarmer pole, the translocated *parS-Cori* region is attached to the new PopZ matrix that formed coincidentally with its arrival. At the opposite stalked pole, the *parS-Cori* region is released from PopZ roughly coincident with stalked cell reentry into another round of chromosome replication.

# *C. CRESCENTUS* DNAA ALSO SIGNALS CHROMOSOME PARTITIONING

A study by Mera et al. implicated *C. crescentus* DnaA in chromosome partitioning (Mera et al., 2014). A conditional DnaA expression strain failed to initiate chromosome replication when DnaA was shut-off, as expected (Gorbatyuk and Marczynski, 2001), and as expected kept a single fluorescent ParB-*parS* centromere complex at the old stalked cell pole while the cell attempted to grow and divide. However, and very surprisingly, DnaA expression at low levels that could not initiate chromosome replication could still initiate and complete *parS-Cori* partitioning. Under these low DnaA conditions, many cells that had only a single, i.e. an un-replicated ParB-*parS* centromere complex could still move it completely from the old stalked pole to the new swarmer cell pole. Mera et al. clearly showed that DnaA binds the *parS* and that DnaA-ATP is required for this partitioning since a DnaA allele that does not bind ATP does not support partitioning. The view suggested by these results is that as DnaA activity rises (as both protein abundance and DnaA-ATP) during the swarmer cell to stalked cell transition, DnaA first acts at *parS* to perhaps commit the chromosome to partitioning before acting at *Cori* to commit it to chromosome replication (**Figure 1B**). This view is attractive considering that DnaA often acts as a global regulator of cell-cycle gene expression (Hottes et al., 2005) and chromosome replication (Gorbatyuk and Marczynski, 2001) and now apparently chromosome partitioning as well.

## CONTROL BY NUCLEOID-ASSOCIATED PROTEINS

Nucleoid organization can theoretically impact both chromosome replication and partitioning (Badrinarayanan et al., 2015). Unlike eukaryotic cells, bacteria do not possess histones, and instead, several small proteins called nucleoidassociated proteins (NAPs) compact and organize their genomes (Luijsterburg et al., 2006; Stavans and Oppenheim, 2006). Bacterial NAPs are not always conserved, but they share many features such as a small size, a high expression level, and a tight DNA binding (Krogh et al., 2018). NAPs impact DNA topology, which must be regulated for efficient transcription and replication (Donczew et al., 2014; Dorman and Dorman, 2016). For example, negatively supercoiled genes are more efficiently transcribed than positively supercoiled genes suggesting transcriptional control by NAPs (Sobetzko et al., 2012). The most investigated and one of the most conserved NAPs is the HU protein of *E. coli* (Ali Azam et al., 1999). HU exists as a homo- or hetero-dimer of *α* and β chains depending on the growth phase. DNA-binding affinity is different for each dimer, leading to differential nucleoid compaction and differential transcription between the growth phases. HU also stabilizes the pre-replication complex essential for the initiation of *E. coli oriC* replication (Chodavarapu et al., 2008). Other NAPs also influence the initiation of DNA replication. In *E. coli*, NAPs "FIS" and "IHF" repress and stimulate the initiation of DNA replication (Ryan et al., 2004; Wolanski et al., 2014a,b). In *B. subtilis*, the NAP "ROK" recruits and interacts with the bacterial replication initiator DnaA. ROK thereby directs DnaA to repress transcription and to help shape the nucleoid (Seid et al., 2017).

# *C. CRESCENTUS* GAPR IS A NOVEL NAP THAT AIDS CHROMOSOME REPLICATION AND PARTITIONING

In *C. crescentus*, the recently identified and now best characterized NAP "GapR" is implicated in cell-cycle control including chromosome replication and partitioning. GapR is an essential 89 amino-acid protein exclusively found in the alphaproteobacteria, which are also known for their asymmetric cell division (Brilli et al., 2010). It is therefore tempting to speculate that GapR contributes to the chromosome asymmetry of *C. crescentus* (**Figure 1A**). Recent papers report that GapR has several relevant properties. For example, GapR binds DNA in AT-rich regulatory regions and next to highly expressed genes. Interestingly, the bulk distribution of GapR on the chromosome forms a gradient that decreases from the *parS-Cori* region to the terminus region (Arias-Cartin et al., 2017). Recently, another NAP "HupB" in *M. smegmatis* (Holowka et al., 2017) was shown to have a similar chromosome-wide gradient distribution. In the absence of GapR, both DNA replication and cell division are impaired (Arias-Cartin et al., 2017; Taylor et al., 2017). However, depletion of GapR only slightly affects global gene expression and most of the genes that are overexpressed belong to the DNA damage stress response and could be induced by indirect DNA damage. These observations argue that GapR is not primarily a transcription regulator. ChIP-seq analysis and fluorescence microscopy have shown that binding of GapR on the chromosome is dynamic and changes throughout the cell cycle. The strongest GapR peaks accumulate near *Cori* and downstream of *parB* near *parS* during the initiation of DNA replication (Taylor et al., 2017). Through this binding GapR somehow enhances the early slow phase of chromosome partitioning (**Figure 1B**), because without GapR, the *parS-Cori* region duplicates and then collapses into one focus before repeating the separation/partitioning process. This is the critical time when separation of the two chromosomes directs them to their alternative fates (Taylor et al., 2017). Subsequently, as the chromosome replicates and partitions, GapR localization correlates with the moving replisome and the replication fork seems to displace the protein from the DNA (Arias-Cartin et al., 2017). Consistent with these observations, X-ray protein crystallography has shown that two GapR dimers assemble to encircle DNA that must be overly twisted to fit inside the hole (Guo et al., 2018). Such overly twisted DNA is either found in front of the replication forks or downstream highly transcribed genes. Although the molecular details have still to be explored, it was proposed that once bound to the overly twisted DNA, GapR enhances or recruits the gyrase activity to dissipate (+) supercoiled DNA produced by replication forks and by RNA polymerase (Guo et al., 2018). Therefore, unlike most NAPs that primarily compact the nucleoid, GapR seems to primary facilitate nucleoid replication and partitioning perhaps at least in part by strategically directing DNA gyrase and perhaps other "molecular machines" including RNA and DNA polymerases.

# *C. CRESCENTUS* PROTEIN POPZ IS A POLAR ORGANIZING "HUB"

Multiple cell-cycle regulators act through the cell poles, and PopZ is their polar "hub protein" acting at the heart of chromosome replication and partitioning (Bowman et al., 2010). PopZ is an intrinsically disordered network protein that fills and forms special apical zones in the cytoplasm. Molecular recognition features "MoRFs" (Holmes et al., 2016) allow PopZ to engage and to localize many cell-cycle proteins. PopZ is initially found at the cell poles, where it binds ParB to anchor *parS* (Bowman et al., 2008; Ebersbach et al., 2008). In addition to this key function, PopZ serves as a platform for other cell-cycle regulators. For example, CtrA and its kinases CckA regulate chromosome replication. CtrA binds *Cori* and both CtrA and CckA are recruited to the stalked cell pole in a PopZ-dependent manner (Bowman et al., 2010; Holmes et al., 2016). Moreover, PopZ sequesters and restrains the CtrA-targeting protease ClpXP (Joshi et al., 2018). In the absence of PopZ, ClpXP exhibits unprecedently high CtrA degradation rates. Under normal conditions, the PopZ-recruited adaptor protein CdpR modulates ClpXP activity also by CckA-mediated phosphorylation. When PopZ is lost, CckA localization is hindered, and CdpR remains in its "active" dephosphorylated state. Consequently, overly active CdpR recruits more ClpXP to accelerate the proteolysis of CtrA. Interestingly, over-expression of PopZ also stimulates the proteolysis of CtrA but by a different mechanism. Under these abnormal conditions, CtrA and ClpXP are thought to concentrate at the cell pole and directly interact without using the CdpR adaptor (Joshi et al., 2018).

While CtrA inactivation is required for the initiation of chromosome replication in stalked cells (**Figure 1A**), its re-accumulation and phosphorylation in late S-phase are also required for cell-cycle transcription control and to prevent premature replication in the new swarmer cell compartment (Sanselicio et al., 2015). Accordingly, MopJ (motility PAS domain associated with DivJ) emerged as an important enhancing factor for CtrA accumulation (Sanselicio et al., 2015). At the cell poles, MopJ attenuates the DivJ-DivK-DivL kinase pathway that is also involved in the downregulation of CtrA through ClpXP. Once again, PopZ lies at the heart of this molecular interaction because the PopZ polar matrix localizes DivJ to the stalked pole, which in turn drives the polarization of DivK, DivL, and MopJ (Ebersbach et al., 2008).

During chromosome replication, the role of PopZ in partitioning switches from passive anchoring to an active participation in the movement of *parS-Cori*. Co-Immunoprecipitation experiments revealed that PopZ interacts directly with ParB, and a PopZ-ParB-*parS* complex presumably accounts for the initial polar anchoring/tethering at the early stalked pole (Bowman et al., 2008). Somehow the *parS-Cori* region is released from PopZ, and upon replication initiation, the duplicated DNA regions are separated such that one region seems to reattach, while the other moves slowly toward the quarter cell-length position. This corresponds to the slow phase of chromosome partitioning (**Figure 1**) that, as we described above, requires GapR but not ParA (Taylor et al., 2017). This is also the symmetry splitting point in the cell cycle that determines the subsequent fates of the chromosomes. Once this step is reached, the subsequent fast phase of partitioning uses ParA-ATPase activity. As the ParB*parS* chromosome complex contacts DNA-bound ParA-ATP, the stimulated ATP hydrolysis causes subsequent ParA release. Such repeated interactions of binding and unbinding presumably cause the movement toward the new pole (Laloux and Jacobs-Wagner, 2013; Ptacin et al., 2014). Interestingly, the PopZ matrix directly sequesters the DNA-released ParA subunits at the new pole and then revives their ATP-bound state and their affinity for nucleoid DNA (Ptacin et al., 2014). This "recycling" or "rejuvenating" function of PopZ presumably enhances partitioning, since by concentrating and reactivating ParA-ATP dimers, PopZ will create a sharper ParA gradient that leads to the new cell pole. Interestingly, another cell pole "landmark" protein "TipN" shares functional redundancy with PopZ as it also recruits ParA to prevent reversal of the segregating ParB-*parS* complex (Ptacin et al., 2014). Accordingly, the ΔtipNΔpopZ double mutation is synthetically lethal (Schofield et al., 2010), and TipN polar localization is disrupted in the absence of PopZ (Ebersbach et al., 2008).

Further studies suggest an added layer of communication between ParA and PopZ. The redistribution of PopZ to the new swarmer pole (**Figure 1**) is coordinated with the arrival of the second ParB-*parS* focus at the new pole (Bowman et al., 2008; Ebersbach et al., 2008; Laloux and Jacobs-Wagner, 2013). Therefore, the ParA-dependent partitioning process somehow also drives the bi-polar organization of PopZ. In support of this notion, delayed partitioning caused by TipN depletion postponed PopZ accumulation at the new pole (Laloux and Jacobs-Wagner, 2013). ParA participates in the formation of the new PopZ matrix, as its loss disrupts PopZ bi-polarity. While other means of PopZ localization have been suggested such as self-organization by nucleoid occlusion (Ebersbach et al., 2008; Saberi and Emberly, 2010), these are clearly not enough, and a ParA-mediated PopZ-localization mechanism is required. If basal levels of ParA initiate PopZ recruitment, this may trigger a positive-feedback loop where ParA and PopZ will accumulate together through mutual support (Laloux and Jacobs-Wagner, 2013). As mentioned above, TipN also recruits ParA, and therefore, this polar landmark protein may also start or contribute to the growth of the PopZ matrix.

PopZ interactions are certainly complex yet robust, and however, this happens in wild-type *C. crescentus* cells, a new PopZ matrix always forms in time to meet and anchor the ParB-*parS* complex arriving at the new swarmer pole (**Figure 1B**). Interestingly, this cell-cycle pattern is very similar to that of the *V. cholerae* Chrom I, which is anchored through *parS1*-ParB1 to a polar PopZ-like protein called "HubP" (Yamaichi et al., 2012). Yet despite such a striking functional correspondence, HubP and PopZ are otherwise evolutionarily unrelated proteins.

The cell-cycle regulated zinc-finger protein ZitP offers yet another mechanism to control PopZ, independent of the *parABS* system (Berge et al., 2016). When ZitP is removed in a strain expressing a variant of PopZ that cannot bind ParB, bi-polar ParB fluorescent foci are rarely seen. However, the resupply of ZitP restores ParB foci at both cell poles, which implies the restoration of localized PopZ anchors (Berge et al., 2016). In this situation, the chromosome anchoring function may rely solely on ZitP since the PopZ variant is unable to bind ParB, but in wild-type cells, the role of ZitP in anchoring would be considered supportive. Normally, PopZ-bound ZitP indirectly binds to *parS*-flanking sites, where it functions to enhance ParB nucleation on the *parS* DNA. This assembly of ZitP-PopZ-ParB on the chromosome effectively restrains segregation (Berge et al., 2016).

It seems that the common theme for this multifaceted PopZ protein is its capacity for two-way interactions with many regulating and cell organizing proteins. For example, ZitP also relies on PopZ to recruit and position pilus biogenesis and swarming motility systems (Mignolet et al., 2016). In summary, PopZ is certainly a "hub" for cell-cycle communication that is yet to be fully explored as a mediator of crosstalk. Future studies promise new insights and new mechanisms of crosstalk between chromosome replication, partitioning, and probably the other landmarks of the cell cycle.

#### AUTHOR CONTRIBUTIONS

GM proposed, organized, and wrote the bulk of this review. KP wrote the section on GapR. PP wrote the section on PopZ.

#### REFERENCES

#### FUNDING

This work was funded by the Canadian Institutes for Health Research (CIHR) operating grant MOP-12599.

centromere segregation. *Proc. Natl. Acad. Sci. USA* 111, E2046–E2055. doi: 10.1073/pnas.1405188111

implications for chromosome segregation. *Nucleic Acids Res.* 43, 719–731. doi: 10.1093/nar/gku1295

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2019 Marczynski, Petit and Patel. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Where and When Bacterial Chromosome Replication Starts: A Single Cell Perspective

Damian Trojanowski† , Joanna Hołówka† and Jolanta Zakrzewska-Czerwinska ´ \* †

Department of Molecular Microbiology, Faculty of Biotechnology, University of Wrocław, Wrocław, Poland

Bacterial chromosomes have a single, unique replication origin (named oriC), from which DNA synthesis starts. This study describes methods of visualizing oriC regions and the chromosome replication in single living bacterial cells in real-time. This review also discusses the impact of live cell imaging techniques on understanding of chromosome replication dynamics, particularly at the initiation step, in different species of bacteria.

Keywords: replication initiation, oriC, replisome, single-cell, bacterial chromosome

#### Edited by:

Feng Gao, Tianjin University, China

#### Reviewed by:

Julia Grimwade, Florida Institute of Technology, United States Christian J. Rudolph, Brunel University London, United Kingdom

#### \*Correspondence:

Jolanta Zakrzewska-Czerwinska ´<EMAIL_ADDRESS>
†These authors have contributed equally to this work

#### Specialty section:

This article was submitted to Evolutionary and Genomic Microbiology, a section of the journal Frontiers in Microbiology

Received: 30 September 2018 Accepted: 02 November 2018 Published: 26 November 2018

#### Citation:

Trojanowski D, Hołówka J and Zakrzewska-Czerwinska J (2018) ´ Where and When Bacterial Chromosome Replication Starts: A Single Cell Perspective. Front. Microbiol. 9:2819. doi: 10.3389/fmicb.2018.02819

#### INTRODUCTION

DNA replication is an enormously intricate process, in which a few dozen enzymes catalyze a series of reactions, including DNA unwinding and the synthesis of sister DNA strands. This process must be highly precise and accurately timed to prevent any unnecessary loss of energy and to ensure that DNA is faithfully and completely replicated only once per cell-division cycle (Leonard and Grimwade, 2015). In all three domains of life, chromosomal replication is mainly regulated at the initiation step (Nielsen and Løbner-Olesen, 2008; Aves, 2009; Skarstad and Katayama, 2013), an important cell cycle checkpoint guaranteeing that DNA replication begins at the right place and time.

Most bacterial genomes consist of one covalently closed chromosome (**Figure 1**). In a few bacteria, however, the genetic information is distributed on two [e.g., Vibrio cholerae (Trucksis et al., 1998)] or even more [e.g., Paracoccus denitrificans (Winterstein and Ludwig, 1998)] chromosomes. Interestingly, some bacteria possess linear chromosomes [e.g., Streptomyces (Lin et al., 1993)].

In contrast to eukaryotes, bacterial chromosomes have a single, unique origin of replication (oriC) (Bird et al., 1972; Kaguni and Kornberg, 1984; Gao and Zhang, 2008; Masai et al., 2010; Méchali, 2010; Katayama, 2017). DNA synthesis is initiated at this unique oriC, generating a single replication eye per chromosome (**Figure 1**). Cooperative binding of the initiator protein, DnaA, to multiple DnaA-recognition sites (DnaA boxes) within the oriC region triggers separation of the DNA strands at the DNA unwinding element (DUE), providing an entry site for the machinery of replication (replisome, **Figures 1**, **2A**; Skarstad et al., 1986, 1990; Bach et al., 2008; Leonard and Grimwade, 2011; Wolanski et al., 2014 ´ ; Richardson et al., 2016).

Enormous progress has been made in recent years toward understanding the mechanisms of replication initiation, particularly the organization and function of oriC regions in different bacteria (Donczew et al., 2012; Makowski et al., 2016; Jaworski et al., 2018; Midgley-Smith et al., 2018; Samadpour and Merrikh, 2018). Less is known, however, about the subcellular localization of replication processes during the cell cycle in various bacterial species. The development of sophisticated cell biology techniques has allowed examination of when and where the replication machinery is assembled within the bacterial cells, and how the initiation of replication is coordinated with the cell cycle (Donczew et al., 2012; Harms et al., 2013; Santi and McKinney, 2015;

Trojanowski et al., 2015; Böhm et al., 2017). This process is particularly interesting in bacteria with two chromosomes (V. cholerae) (Demarre et al., 2014; Ramachandran et al., 2018) and in those that undergo complex cell differentiation (Caulobacter crescentus) (Jensen et al., 2001; Toro et al., 2008) and/or exhibit complicated life cycles, e.g., Myxococcus xanthus (Harms et al., 2013; Lin et al., 2017) and Streptomyces species (Kois-Ostrowska et al., 2016). In these bacteria, the regulatory networks that control replication initiation are likely to be intricate and require specific mechanisms that can synchronize the initiation of chromosomal replication with developmental processes.

The main goal of this review is to highlight imaging techniques that allow the determination of the subcellular location of oriC regions and the initiation of chromosome replication (i.e., assembly of the replication machinery) in single living bacterial cells in real time. This review also discusses the impact of real-time single-cell imaging on understanding of chromosome replication dynamics, particularly at the initiation step, in different bacteria.

### VISUALIZATION OF REPLICATION INITIATION AND REPLISOME DYNAMICS IN LIVE CELLS

The development of live cell imaging techniques has allowed the visualization of replisomes (**Figure 2A**; Jensen et al., 2001; Reyes-Lamothe et al., 2008; Wang and Sherratt, 2010; Harms et al., 2013; Santi and McKinney, 2015; Trojanowski et al., 2015; Mangiameli et al., 2017) in live cells and the study of DNA replication dynamics, including the timing and localization of replication initiation, in real time at the single-cell level. Microscopic analysis of live cells has several advantages over analysis of fixed samples. Fixing the cells, a process that involves dehydration and/or intracellular cross-linking, may influence the localization of proteins or subcellular structures of interest. Moreover, some fusions with fluorescent proteins (FP) are sensitive to the harsh conditions used during fixation. For example, different sample preparation of Mycobacterium smegmatis cells results in ParA-EGFP localizing either apically or as a cloud arising from the new cell pole (Ginda et al., 2013, 2017). Furthermore, permeabilization of the bacterial cell wall during immunostaining may contribute to a loss of cytoplasmic content or, due to cellular crowding, may generate high background noise or alter the localization of large immunocomplexes, particularly when using secondary antibodies for signal amplification. Although several high quality studies of fixed samples have provided invaluable data, the conditions found in cells fixed on a coverslip only approximate the conditions found in live cells.

Replication is visualized primarily by the fusion of different replisome (DNA polymerase III) subunits (**Figure 2A**) to a variety of FP. The choice of subunit to create the fusion protein should be guided by the specific application and the specific type of bacterium. Escherichia coli is the best characterized bacterial model for tracking live replication (Kongsuwan et al., 2002; Bates and Kleckner, 2005; Fossum et al., 2007; Reyes-Lamothe et al., 2008, 2010; Su'etsugu and Errington, 2011; Wang et al., 2011; Moolman et al., 2014; Beattie et al., 2017).  Findings of these studies may help in the construction of fluorescent fusions of replisome components in other bacteria. It is also important to consider alternative N- and C-terminal fusion, as one, or sometimes both, ends of target proteins may be implicated in inter- or intra-molecular interactions. The sliding clamp (**Figure 2A**) is the protein of choice in most studies and both N- and C-terminal fusions proved to be functional in a range of species (Kongsuwan et al., 2002; Reyes-Lamothe et al., 2010; Su'etsugu and Errington, 2011; Moolman et al., 2014; Santi and McKinney, 2015; Trojanowski et al., 2015; Arias-Cartin et al., 2017; Böhm et al., 2017; Mangiameli et al., 2017; Hołówka et al., 2018). However, the sliding clamp also participates in processes other than DNA replication, including recombination and DNA repair, possibly altering the distribution of DnaN-FP (or FP-DnaN) foci in these cells. This is not usually a concern in wild-type-like fluorescent reporter strains, under both optimal and minimal conditions, but may be of concern in knock-out/overproducing mutant strains, involving, for example, genes engaged in DNA repair, or when studying replication dynamics under stress-inducing conditions such as in the presence of antibiotics, mutagenic compounds like

mitomycin, and replication inhibitors. In these experiments, choosing another replisome component may be advisable. Beside the siding clamp, DnaX (Lemon and Grossman, 2000; Bates and Kleckner, 2005; Berkmen and Grossman, 2006; Vallet-Gely and Boccard, 2013; Raaphorst et al., 2017) (particularly its C-terminal fusion) is frequently used as a replisome localization marker. The dnaX gene encodes two alternative proteins, τ – the full-length protein encoded by the dnaX gene, and γ, which originates from ribosome switching during translation, resulting in premature termination of translation and generating a truncated protein.  Monitoring replisome dynamics in strains expressing fusion proteins encoded on an episomal plasmid is not recommended, as plasmid replication is triggered mainly by the same protein components that trigger chromosomal replication. Fusion with catalytic core subunits (Lemon and Grossman, 1998; Migocki et al., 2004; Trojanowski et al., 2017) is also possible, although additional cargo attached to core Pol-DNA III may affect nucleotide incorporation rates and influence the kinetic parameters of the entire replication complex. This was shown for M. smegmatis, where the C-terminal fusion of a catalytic alpha subunit to EYFP prolonged the C-period (Trojanowski et al., 2017). Thus proteins other than the catalytic core complex may be a better choice for studies of replisome dynamics. Other fusions successfully used for replisome tracking include DnaB (DNA helicase) (Jensen et al., 2001; Beattie et al., 2017), DnaQ (Reyes-Lamothe et al., 2008, 2010; Wallden et al., 2016; Mangiameli et al., 2017), and χ and δ 0 subunits (Jensen et al., 2001; Reyes-Lamothe et al., 2008). When designing a fluorescent fusion for replisome visualization, additional features should be taken into account, especially oligomerization status, fluorescence yield and

spectral properties. FP (especially GFP derivatives) are likely to form low-affinity oligomers (Costantini et al., 2012), which may influence the dynamics of the studied protein complex, especially when the fusion protein is produced at a high level. Thus, choosing a fluorescent variant with a lower tendency to undergo oligomerization (e.g., mCherry, mCherry2, mCitrine, and mScarlett) is recommended. Spectral characteristics and brightness are essential, especially when replisomes are localized together with other cellular components (e.g., chromosome and membrane) (Shaner et al., 2005). Importantly, FP are sensitive to pH and cannot be utilized to analyze anaerobic bacteria, as maturation of the chromophore requires oxygen molecules (Shaner et al., 2005; Landete et al., 2015). Fluorescent fusion proteins are suitable for both qualitative long-term live cell imaging and quantitative analysis. For example, Y-Pet fusion with a variety of replisome subunits was used to quantify the numbers of copies of particular proteins within a replication eye in vivo (Reyes-Lamothe et al., 2010). However, most of these variants lacked the properties required for super-resolution imaging. In the latter case, proteins of interest should be fused with photoactivated or photoconvertible proteins. Recently published studies may provide hints regarding single-molecule resolution microscopy of replication complexes (Georgescu et al., 2012; Stracy et al., 2014; Liao et al., 2016; Lewis et al., 2017). The fusion of replisome subunits with HaloTag may be an alternative to FP. The size of HaloTag is similar to that of FP, but the ligands that bind to HaloTag have better fluorescence yield, resulting in a higher signal compared with standard FPs (HaloTag <sup>R</sup> Protein Purification System, 2018). The advantage of using direct fluorescent ligands (e.g., dTMR and dR110) is that they do not need to be washed out before acquisition. Halo ligands are also suitable for high-resolution microscopy.

Replication tracking (particularly initiation of replication) is often accompanied by localization of nascent oriCs (**Figure 2B**).

The fluorescence repressor operator system (FROS) or ParB/parS is frequently used for live cell tracking (Lau et al., 2003). The FROS system (**Figure 2B**) consists of two components: operator sequences (usually lacO or tetO arrays repeated up to several hundred times in tandem and interspersed by oligonucleotide spacers) and an FP-tagged repressor protein (LacI-FP or TetR-FP), which binds to the operator sequences. FROS was efficiently used to localize chromosomal loci, including oriC, terminus and other specific loci on both replichores in a variety of species (Viollier et al., 2004; Fogel and Waldor, 2005; Frunzke et al., 2008; Liu et al., 2010; Vallet-Gely and Boccard, 2013; Wang et al., 2014; Santi and McKinney, 2015). However, it is often difficult to insert the large operator arrays into the chromosome, particularly in highly transcribed regions such as oriC (Le and Laub, 2014). Moreover, overexpression of repressor may result in replication/transcription hold-up or alteration in segregation of replicated regions (Possoz et al., 2006; Mettrick and Grainge, 2016). Thus, low levels of repressor should be produced, usually by using inducible promoters. Additionally, tracking oriCs together with replisomes requires delivery of the repressor-FP fusion protein from the chromosomal locus, either as a part of an operator array construct or inserted into an attachment site. Although FROS may provide invaluable data, its instability is a major drawback.

The ParB-FP/parS system (which originated from naturally existing chromosome and/or plasmid partitioning strategies) (**Figure 2B**) represents an easier alternative to FROS. This system uses an intrinsic feature of ParB, its binding to centromerelike parS sequences (Wang et al., 2011; Reyes-Lamothe et al., 2012; Badrinarayanan et al., 2015). Most bacterial species possess the ParABS chromosome segregation system, except for several well-studied Gammaproteobacteria, including E. coli. Because most chromosomal parS sites are localized proximal to the oriC-proximal regions (Livny et al., 2007), introduction of fluorescent ParB, which oligomerizes within parS sequences, addresses all of the system requirements for successful oriC labeling. This approach has been shown effective in a number of bacteria, including Mycobacterium, M. xanthus (Harms et al., 2013), Streptomyces (Donczew et al., 2016; Kois-Ostrowska et al., 2016), C. crescentus (Laloux and Jacobs-Wagner, 2013), and C. glutamicum (Donovan et al., 2010; Böhm et al., 2017). In bacteria lacking a chromosomal ParABS system (e.g., E. coli), plasmid-derived partitioning components (phage P1 or Yersinia pestis MT1ParB/parS systems) are frequently used (Youngren et al., 2000; Li et al., 2002; Nielsen et al., 2006, 2007). The use of plasmid-derived parS/ParB is also beneficial, as it does not interfere with the endogenous chromosomal ParABS system or another plasmid-derived parS/ParB system (P1/MT1), allowing the simultaneous localization of multiple chromosomal loci. Its major advantage compared with FROS is that insertion of only a few copies of parS is sufficient for strong fluorescent signals after ParB-FP binding.

Determination of the specific point (and subcellular localization) at which replication is initiated requires long-term imaging of living cells (from several minutes to hours, depending on the bacterial growth rate and the conditions being tested, e.g., rich versus minimal medium). The simplest way to analyze replication at the single-cell level is to spread the cells of the reporter strain on the agar pad (a thin agar layer between the microscope slide and the cover glass) or on the bottom of solidified medium inside culture dishes (Joyce et al., 2011;Dhar andManina, 2015). Although simple and low-cost, this approach is not always applicable (e.g., labeling and medium changing). Microfluidic flow chambers are used for the latter purposes, as well as for rapidly changing culture conditions (e.g., applying stress). Various microfluidic chips and plates are commercially available from an increasing number of companies, whereas custom made (usually PDMS) chips are a cost-reducing alternative and also allow for more personalized applications (Wang et al., 2010; Cattoni et al., 2013; Dhar and Manina, 2015; Trojanowski et al., 2015; Wallden et al., 2016). The architecture of microfluidic chips and plates varies among studies and choosing the right one should be dictated by the specific study purpose and the availability of additional equipment, e.g., peristaltic/syringe/pressure pumps, flow controllers, or automation.

## SPATIOTEMPORAL LOCALIZATION OF THE REPLISOME DURING REPLICATION INITIATION

Localization of the replication machinery at the beginning of DNA synthesis is dependent on oriC position, and is therefore connected with the spatial arrangement of the chromosome. In bacteria having oriC and ter regions positioned at the mid-cell, the intervening chromosomal regions (i.e., the left and right chromosomal arms) are stretched out toward opposite cell poles, creating a left-ori-right pattern, whereas cells having oriC and ter regions localized to opposite poles show an ori-ter chromosomal arrangement (Wang and Rudner, 2014). Replisomes in the cells exhibiting a left-ori-right configuration are assembled in the midcell region of the chromosome. This pattern has been observed in E. coli cells (Postow et al., 2004; Valens et al., 2004; Boccard et al., 2005) and during the vegetative growth of B. subtilis (the chromosome in B. subtilis is oscillating between left-ori-right and ori-ter configuration) (Wang et al., 2014; **Figure 3A**). During sporulation, however, the B. subtilis chromosome adopts an oriter orientation to segregate an entire copy of the chromosome within each spore. Positioning of the oriC at the mid-cell of B. subtilis and E. coli is maintained by the condensins SMC and MukB (a structural homolog of SMC), respectively (Niki et al., 1992; Danilova et al., 2007; Sullivan et al., 2009). SMC can compact large chromosomal regions, and, by interacting with ParB protein, organizes the oriC-proximal regions in B. subtilis, with ParB binding to parS sequences located near oriC (Gruber and Errington, 2009). The interaction of MukB with the nucleoid associated protein HU ensures proper oriC positioning in E. coli cells (Lioy et al., 2018). After initiation, E. coli replisomes oscillate near the cell center, while newly replicated oriCs are segregated toward the cell poles (Reyes-Lamothe et al., 2008). In comparison, B. subtilis replisomes colocalize throughout replication (Migocki et al., 2004), and are therefore visible as a single fluorescent focus. Replisome positioning in the cell center can be also found in oval-shaped S. pneumoniae (Kjos and Veening, 2014; van

Raaphorst et al., 2017), which, similar to many other bacteria including B. subtilis, encodes an SMC homolog.

Some bacteria, such as M. smegmatis (Santi and McKinney, 2015; Trojanowski et al., 2015) and M. xanthus (Harms et al., 2013), exhibit off-center replisome localization during the initiation of replication (see **Figure 3B**). In M. smegmatis, segregation of the newly replicated oriCs starts immediately after initiation of replication, with one oriC remaining near the old cell pole and the other traveling toward the opposite pole (Ginda et al., 2017; Hołówka et al., 2018). Replisomes oscillate in the

old-pole-proximal cell half during most of the replication process, but localize closer to the new cell pole prior to termination (Trojanowski et al., 2015). A slight asymmetry in mycobacterial replisome positioning is associated with the apical growth mode of these bacteria. Positioning of oriC region(s) in Mycobacterium depends on the interaction of ParB with ParA protein, which in turn interacts with the polar growth determinant, DivIVA protein (Ginda et al., 2013).

As a result of the asymmetric location of oriC, M. xanthus replisomes are positioned at the subpolar regions (**Figure 3B**; Harms et al., 2013). Although M. xanthus contains a DivIVA homolog, suggesting analogous interactions at the pole as described for Mycobacterium, deletion of this homolog does not affect cell division or chromosome segregation. Rather, localization of the ParA and ParB-parS complexes (and thus the oriC region) in M. xanthus is controlled by the bactofilins BacNOP, through the direct interactions of ParA and ParB with the scaffold created by BacNOP (Lin et al., 2017).

Bacteria exhibiting complex life cycles often show an ori-ter chromosome orientation (**Figure 3C**). In C. crescentus stalked cells, chromosome replication starts at the old cell pole (Jensen et al., 2001). The anchorage of the chromosome at the old cell pole is maintained by the protein PopZ (Bowman et al., 2008). Similarly, in V. cholerae, the origin (oriI) of one of the two chromosomes, chrI, is attached to the old pole by HubP protein (Yamaichi et al., 2012), thereby setting the subcellular position for assembly of the replication machinery. In contrast, the origin (oriII) of the second, smaller chromosome (chrII) is located at mid-cell. Replication of V. cholerae chrII starts later than that of chrl to synchronize the termination of replication of both chromosomes (Demarre et al., 2014; Ramachandran et al., 2018). As a result of the subpolar localization of C. crescentus and V. cholerae (chrI) replisomes near the old cell pole, one of the newly replicated oriC regions travels across the chromosome to the opposite cell pole with the assistance of the ParABS system (Toro et al., 2008; Ramachandran et al., 2014). Interestingly, in P. aeruginosa exhibiting ori-ter orientation, the chromosome is apparently not anchored to the cell pole, as shown by the cytoplasmic gap between oriC and the cell pole (Vallet-Gely and Boccard, 2013).

The multiploid and apically growing bacterial species S. coelicolor, exhibits another mode of spatiotemporal replisome localization, in which replication is initiated during vegetative growth (**Figure 3D**; Kois-Ostrowska et al., 2016). Replication of multiple copies of the S. coelicolor chromosome starts asynchronously, and newly replicated sister chromosomes follow the extending hyphal tip. Similar to Mycobacterium, positioning of the tip-proximal oriC (and hence the replisomes) is maintained through ParA interactions with the polarisome complex, which includes the proteins ParB, DivIVA, and Scy (Flärdh et al., 2012; Ditkowski et al., 2013). In the closely related and diploid species C. glutamicum, replisomes are assembled on each chromosome asymmetrically, in proximity to the cell poles (**Figure 3D**; Böhm et al., 2017). Fluorescently tagged ParB attaches to the cell poles, suggesting an ori-ter-ter-ori spatial orientation of C. glutamicum chromosomes.

Described differences among bacteria in the positioning of oriC regions during the replication initiation reflect the different modes of chromosome segregation. Mid-cell replisomes location results in symmetric segregation of oriCs toward the opposite cell poles, while polar and offcenter replisome positioning imply asymmetric segregation of the newly replicated oriC regions. Furthermore, polar localization requires the complex system to either anchor oriC directly at the pole (e.g., PopZ and HubP proteins) or to maintain the subpolar position by protein complexes (e.g., the interaction of ParABS system with the DivIVA or the BacNOP). Such variety in the composition of multiprotein complexes involved in oriC(s) positioning provides an opportunity for the discovery of novel genus/species-specific drug targets.

#### CONCLUSION

Single-cell fluorescence imaging and fluorescence tagging techniques allow researchers to precisely visualize proteins and their complexes inside living bacterial cells in real time. These techniques revealed that many proteins are targeted to distinct subcellular positions, where they participate in various cellular processes including chromosome replication. Recent studies using advanced live-cell imaging demonstrated that chromosome replication is coordinated with other key steps of the cell cycle, such as chromosome segregation and cell division. Proteins (or protein complexes) involved in condensation (i.e., SMC/MukB), chromosome segregation (i.e., ParAB in Gram-negative and Gram-positive bacteria) and/or cell division (DivIVA in Gram-positive bacteria) take part directly or indirectly in oriC positioning, thus indicating the site of replisome assembly. Additionally, other proteins guiding the oriC region have been recently identified. Interestingly, they vary significantly among different bacteria, e.g., PopZ (C. crescents), HubP (V. cholerae, chromosome I), and bactofilins (M. xanthus). The diversity and complexity of the systems involved in oriC (and thus replisome) subcellular positioning suggest the possibility of developing new antimicrobial therapies and/or altering existing treatments (Kaguni, 2018).

#### AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct, and intellectual contribution to the work, and approved it for publication.

# FUNDING

This study was supported by the National Science Center, Poland (MAESTRO Grant 2012/04/A/NZ1/00057 and OPUS Grant 2017/25/B/NZ1/00657). The cost of publication was supported by the Wrocław Centre of Biotechnology under the Leading National Research Centre (KNOW) program, 2014–2018.

# ACKNOWLEDGMENTS

We apologize that numerous original papers could not be cited due to space limitations.

#### REFERENCES

fmicb-09-02819 November 23, 2018 Time: 17:10 # 7

chromosome replication in gram-negative bacteria. Nucleic Acids Res. 40, 9647–9660. doi: 10.1093/nar/gks742

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Trojanowski, Hołówka and Zakrzewska-Czerwinska. This is an ´ open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Functionality of Two Origins of Replication in Vibrio cholerae Strains With a Single Chromosome

Matthias Bruhn<sup>1</sup> , Daniel Schindler<sup>2</sup> , Franziska S. Kemter<sup>1</sup> , Michael R. Wiley<sup>3</sup> , Kitty Chase<sup>3</sup> , Galina I. Koroleva<sup>3</sup> , Gustavo Palacios<sup>3</sup> , Shanmuga Sozhamannan4,5 \* and Torsten Waldminghaus<sup>1</sup> \*

<sup>1</sup> LOEWE Centre for Synthetic Microbiology-SYNMIKRO, Philipps-Universität Marburg, Marburg, Germany, <sup>2</sup> Manchester Institute of Biotechnology, The University of Manchester, Manchester, United Kingdom, <sup>3</sup> United States Army Medical Research Institute of Infectious Diseases, Frederick, MD, United States, <sup>4</sup> Defense Biological Product Assurance Office, Frederick, MD, United States, <sup>5</sup> The Tauri Group, LLC, Alexandria, VA, United States

#### Edited by:

Alan Leonard, Florida Institute of Technology, United States

#### Reviewed by:

Dhruba Chattoraj, National Institutes of Health (NIH), United States Ole Skovgaard, Roskilde University, Denmark Gregory Marczynski, McGill University, Canada

#### \*Correspondence:

Shanmuga Sozhamannan<EMAIL_ADDRESS>Torsten Waldminghaus Torsten.Waldminghaus@ SYNMIKRO.Uni-Marburg.de

#### Specialty section:

This article was submitted to Evolutionary and Genomic Microbiology, a section of the journal Frontiers in Microbiology

Received: 28 September 2018 Accepted: 14 November 2018 Published: 30 November 2018

#### Citation:

Bruhn M, Schindler D, Kemter FS, Wiley MR, Chase K, Koroleva GI, Palacios G, Sozhamannan S and Waldminghaus T (2018) Functionality of Two Origins of Replication in Vibrio cholerae Strains With a Single Chromosome. Front. Microbiol. 9:2932. doi: 10.3389/fmicb.2018.02932 Chromosomal inheritance in bacteria usually entails bidirectional replication of a single chromosome from a single origin into two copies and subsequent partitioning of one copy each into daughter cells upon cell division. However, the human pathogen Vibrio cholerae and other Vibrionaceae harbor two chromosomes, a large Chr1 and a small Chr2. Chr1 and Chr2 have different origins, an oriC-type origin and a P1 plasmid-type origin, respectively, driving the replication of respective chromosomes. Recently, we described naturally occurring exceptions to the two-chromosome rule of Vibrionaceae: i.e., Chr1 and Chr2 fused single chromosome V. cholerae strains, NSCV1 and NSCV2, in which both origins of replication are present. Using NSCV1 and NSCV2, here we tested whether two types of origins of replication can function simultaneously on the same chromosome or one or the other origin is silenced. We found that in NSCV1, both origins are active whereas in NSCV2 ori2 is silenced despite the fact that it is functional in an isolated context. The ori2 activity appears to be primarily determined by the copy number of the triggering site, crtS, which in turn is determined by its location with respect to ori1 and ori2 on the fused chromosome.

Keywords: DNA replication, secondary chromosome, plasmid, multipartite genome, replication initiation, pathogens, cholera

# INTRODUCTION

The generally accepted paradigm of chromosome replication in bacteria is elucidated in Escherichia coli. Replication is initiated at a unique singular sequence, the origin of replication (oriC) by DnaA, proceeds bidirectionally along the chromosome and ends at the terminus diametrically opposite to oriC on the circular chromosome. In E. coli and related bacteria, immediate re-initiation of chromosome replication is hindered due to the hemi-methylated status of the sister chromosomes and sequestration of oriC by SeqA which has a high binding affinity to hemimethylated ori sequences (Lu et al., 1994; Slater et al., 1995; Waldminghaus and Skarstad, 2009). Most bacteria have single chromosomes and follow this general replication paradigm. However, about 10% of bacterial species have more than one chromosome and exhibit some deviation from this norm (Fournes et al., 2018). Among these, Vibrio cholerae with chromosome 1 (Chr1, ∼3 Mbps) and

chromosome 2(Chr2, ∼1 Mbps) has served as a model system for studies pertaining to multi-chromosome replication mechanisms, and in recent years, an extensive body of information has been accumulated on various aspects of Chr1 and Chr2 replication (Egan et al., 2005; Jha et al., 2012; Val et al., 2014b; Espinosa et al., 2017; Ramachandran et al., 2017).

Chr1 in V. cholerae is similar to the E. coli chromosome in that the replication follows the same pattern: replication origin, ori1, contains multiple DnaA boxes, which are bound by DnaA that unwinds the DNA and initiate replication (Duigou et al., 2006). The similarity is so striking that V. cholerae ori1 can functionally substitute the E. coli replication origin oriC (Egan and Waldor, 2003; Demarre and Chattoraj, 2010; Koch et al., 2010; Kamp et al., 2013).

In contrast, the V. cholerae Chr2 appears to have an origin that resembles those of low copy number plasmids such as P1 and F (Fournes et al., 2018). The ori2 contains an array of repeats (iterons) where the Chr2 specific initiator protein, RctB, binds and unwinds the DNA for ori2 firing (Egan and Waldor, 2003; Duigou et al., 2008) but also exerts a form of negative regulation, termed 'handcuffing,' originally discovered in plasmids (Venkova-Canova and Chattoraj, 2011). Although ori2 has plasmid-like features, Chr2 resembles typical chromosomes in some respects: (1) Participation of SeqA and Dam in regulation of ori2 (Saint-Dic et al., 2008; Demarre and Chattoraj, 2010; Koch et al., 2010; Stokke et al., 2011). (2) Indispensability of Chr2, unlike plasmids, for cell survival because it harbors essential genes (Heidelberg et al., 2000; Kamp et al., 2013). (3) High level of coordination of replication between Chr1 and Chr2 in order to prevent over replication of Chr2 and ensure a guaranteed inheritance of a single copy of both chromosomes (Baek and Chattoraj, 2014; Val et al., 2016; Ramachandran et al., 2018). This raises the question on how coordination between Chr1 and Chr2 with respect to their timing of replication initiation is achieved given the disparity in their sizes and mechanisms of replication.

Chr1 replication is initiated at the onset of the replication period while initiation of Chr2 is delayed and occurs only when 2/3rd of Chr1 replication has been completed. Since Chr2 is 1/3rd the size of Chr1, both chromosomes consequently terminate their replication roughly at the same time (Rasmussen et al., 2007; Stokke et al., 2011). This termination synchrony appears not to be accidental but is selected for during evolution and is conserved within Vibrionaceae despite differing ratios of chromosome sizes (Kemter et al., 2018). This synchrony occurs through the crtS (Chr2 replication triggering Site) present on Chr1 that positively regulates ori2 initiation (Val et al., 2016). Translocation of the crtS locus on Chr1, either closer to ori1 or farther away, resulted in a corresponding shift in Chr2 initiation time as revealed by marker frequency analysis (Val et al., 2016), indicating that the native position of crtS sets the timing of Chr2 replication initiation such that its replication terminates synchronously with Chr1 (Kemter et al., 2018). The exact mechanism of crtS action remains to be elucidated but may include physical contacts between crtS and ori2 as well as sequestration of Chr2 replication initiator protein, RctB (Baek and Chattoraj, 2014; Val et al., 2016). Recently, the global transcription factor Lrp was shown to bind to the crtS site and to facilitate RctB binding (Ciaccia et al., 2018). Heterologous E. coli systems have been established based on ori2 mini-chromosomes demonstrating that crtS provided in trans increases mini chromosome copy number indicating a positive role played by crtS in ori2 firing (Baek and Chattoraj, 2014; Schallopp et al., 2017; de Lemos Martins et al., 2018). Recently, it was demonstrated that the copy number of crtS rather than the act of replicating the crtS is critical in the triggering of ori2 firing (de Lemos Martins et al., 2018; Ramachandran et al., 2018).

In order to assess the differential genetic requirements of Chr1 and Chr2 replication, an artificial single chromosome V. cholerae strain has been created by genetic engineering in which ori1 drives the replication of the fused chromosome (Val et al., 2012). In this strain, designated MCH1, the sequences to the left and right of ori2 were fused to the terminus of Chr1. In this arrangement the direction in which chromosome arms are replicated is conserved to minimize conflicts between DNA replication and transcription. The MCH1 strain was instrumental in establishing the essentiality of Dam methyltransferase in V. cholerae because of its role in ori2 function which was first shown by Demarre and Chattoraj (2010). This conclusion was further supported by the finding that depletion of Dam leads to spontaneous chromosomal fusion (Val et al., 2014a). In this case, the entire genome is replicated from ori1 which can tolerate the absence of Dam (Val et al., 2014a).

Recently, we described two naturally occurring V. cholerae strains (NSCV1 and NSCV2) containing both ori1 and ori2 on the same chromosome (Chapman et al., 2015; Xie et al., 2017). In these strains, Chr1 and Chr2 are fused at two different locations (**Figure 1**). The locations of relevant features such as ori1, ori2, and crtS sites are indicated in **Figure 1A**. In NSCV1, crtS is located about 670 kbs away from ori1 (**Figure 1**). It is similar to the standard two-chromosome reference strain N16961 with respect to distance, where crtS is located 695 kbs away from ori1. In NSCV2, the distance between ori1 and crtS is 1,566 kbs due to a large inversion that has occurred around the terminus region of the chromosome (**Figure 1C**).

This genomic architecture raised interesting questions about whether both origins are functional in the same cell or one or the other ori is silenced since in principle a single origin should suffice to replicate the fused chromosome. The strains also allowed us to ask if the chromosomal fusions are maintained without resorting to genome splitting which is the predominant genome configuration in Vibrionaceae. We found that in NSCV1 both origins are active in the same cell whereas in NSCV2 ori2 appears to be silent. Further, these chromosomes appear to be in a locked configuration since even after prolonged continuous growth they remain fused without splitting into two.

# RESULTS

#### Activity of ori1 and ori2 in V. cholerae NSCV Strains

In general, the replication in bacteria relies on one origin of replication for one replicon. Fusion of two replicons, such as seen in NSCV1 and NSCV2, would initially give rise to a chromosome with two functional replication origins where one could be

superfluous. This raises the question of whether both, ori1 and ori2 are active in the fused chromosomes of NSCV1 and NSCV2, or only one of the origins is active and the other is silent. Inspection of the sequence of the origins and the replication initiator genes revealed no obvious mutational changes that could indicate non-functionality of one or the other of the origins in the two NSCV strains (**Figure 2**) (Xie et al., 2017). Compared to strain N16961, both strains possess complete ori1 sequences,

with seven SNPs (NSCV1) and four SNPs (NSCV2) spanning the 474 bps long gidA-mioC intergenic region and none of the DnaA boxes were affected by mutations. The 5,656 bps long ori2 regions (including genes parB, parA, and rctB) are also intact, with 54 SNPs in NSCV1 and 57 SNPs in NSCV2. Notably, the RctB-binding iteron sequences are not affected by mutations.

To test the activity of replication origins experimentally, we carried out marker frequency analyses (MFA). It is known that actively growing cells have a higher copy number of ori proximal sequences compared to ori distal/ter proximal sequences. We employed next generation sequencing technology to obtain whole genome sequences of DNAs isolated from logarithmic and stationary phase cultures of NSCV1 and NSCV2 and analyzed the sequence data for marker frequency. Read data from stationary phase DNAs were used for normalization and read mapping plots were created with ori1 repositioned at the center of the plot to represent bidirectional replication as well as for easy visualization of ori activity (**Figure 3**). Both NSCV1 and NSCV2 exhibited a maximum copy number of reads close to ori1 and a decreasing gradient on either side of ori1 moving toward the terminus creating a tent-shape (Skovgaard et al., 2011), indicating an active ori1 and bidirectional replication in both strains. In strain NSCV1, a local higher marker frequency was observed at the ori2 position, indicating that ori2 also is active in this strain. In contrast, no such local peak in marker frequency was found at the ori2 position in NSCV2 consistent with a silent ori2. In addition, an almost 3X lower ori1/ter ratio was observed in NSCV2 compared to NSCV1 indicating less or no overlap of replication cycles. The rationale for this interpretation is as follows: On a replicon with overlapping replication, the ori/ter ratio would be four since replication is initiated twice before termination can occur. A replicon that initiates only once would consequently have an ori/ter ratio of two. Considering that the culture represents a mixed population of cells before and after termination an ori/ter ratio of higher than two, as in the case of NSCV1, indicates overlapping replication cycles while values less than two indicate no overlap of replication cycles.

# NSCV1 and NSCV2 Carry a Functional ori2

The apparent inactivity of ori2 in strain NSCV2 raises the question of whether this origin of replication is functional but silenced or non-functional. To answer this question, we cloned the origins into a mini replicon and assessed independent replication in a plasmid backbone. Plasmid pMA135 carries oriR6K and can replicate conditionally in E. coli strains that provide in trans, the replication initiator protein, Pir, from a lambda prophage. In addition pMA135 can be transferred by conjugation from a donor to a recipient (Messerschmidt et al., 2015). In the absence of λpir, no exconjugants are obtained unless replication is driven by another fully functional origin of replication. The ori2 fragments of NSCV1 and NSCV2, including the core ori2 plus the genes rctB and parAB2, were cloned into pMA135 independently and the number of exconjugants was enumerated in an E. coli strain that does not contain λpir. The ori2 minichromosome constructs from all three strains (N16961, NSCV1, and NSCV2) yielded exconjugants at a frequency of about 1% (10−<sup>2</sup> ) of the recipients, as did the positive control F plasmid ori (**Table 1**). Minichromosomes based on oriII of strain N16961 have been shown not to integrate into the E. coli chromosome (Messerschmidt et al.,

†All replicons possess an oriR6K and conjugated from E. coli strain WM3064 to MG1655. ‡Efficiency of one representative experiment is given as ratio of conjugant CFU over total recipient CFU.

2016). To test if the replicons based on NSCV oriII are also replicating autonomously, we performed a plasmid isolation procedure for five individual clones for each of the two tested NSCV replicons. In all cases, we were able to isolate the corresponding minichromosomes as evidenced by agarose gel electrophoresis (data not shown), verifying autonomous replication without integration into the primary chromosome. The oriR6K replicon by itself did not yield any exconjugant. We conclude that both NSCV1 and NSCV2 carry a functional ori2.

#### Genetic Stability of Chromosome Fusions in NSCV1 and NSCV2

An active ori2 in strain NSCV1 could potentially allow the chromosome fusion to be reversed. Similarly, the silent ori2 in NSCV2 might be inactive only in the context of a fused chromosome; in either case, splitting of Chr1 and Chr2 is conceivable. It was observed that V. cholerae with an artificially fused chromosome grew slower compared to the two-chromosome parental strain indicating a negative fitness burden on the bacterium (Val et al., 2012). Similarly, we observed an increased doubling time of strains NSCV1 (20 ± 0.5 min) and NSCV2 (29 ± 1.3 min) compared to the two-chromosome strain N16961 (16 ± 0.2 min) (**Supplementary Table S4**). In addition, microscopic examination showed that strain NSCV2 cells exhibited a distinct phenotype. While the two-chromosome reference strain N16961 and NSCV1 cells are comma-shaped as typical for V. cholerae, NSCV2 cells are much more curled and occasionally S-shaped (**Figure 4**). It is unknown whether this phenotype is related to the chromosome fusion.

It is conceivable that the slower growth of the fused chromosome strains may have negative fitness value leading to instability, thus promoting genome splitting. On the other hand, if the fused chromosome were under a positive selection pressure to remain fused even at a greater cost in terms of slower growth rate or if they are locked in the fusion configuration due to genetic defects that prevent splitting, then the fusion would be stable even after long term continuous culturing. This led us to test whether the two NSCV strains potentially could revert back to a two-chromosome arrangement upon prolonged continuous culturing. If splitting of the fused chromosomes occurs and if this splitting leads to a fitness advantage due to increased growth rates one would expect twochromosome clones to appear in a population of NSCV1 and NSCV2 and these clones could potentially replace the fused chromosome cells after prolonged growth by clonal expansion. To test this hypothesis, we cultured the two NSCV strains for 16 days in 100 ml of liquid medium with replenishment of fresh medium every 24 h resulting in approximately 160 generations of growth in total. To examine if the NSCV strains reverted back to a two-chromosome arrangement we isolated DNA from long-term grown cells and performed long-read DNA sequencing using the PacBio technology to be able to detect long range chromosomal rearrangements such as the chromosomal fusion junctions/or junctions (∼24–51 kbs) of the split chromosomes. A de novo genome assembly led to one single contig for both NSCV1 and NSCV2 reflecting the original one-chromosome configuration. In conclusion, the chromosome fusions in NSCV1 and NSCV2 appear to be stable and chromosome splitting is not a frequent event or the fused state is probably under positive selection pressure.

#### Replication Origins Are Active in an Engineered System Resembling the NSCV Arrangement

Why is ori2 of strain NSCV2 silenced while it is active in NSCV1? One obvious difference between the two strains is the differential positioning of the two replication origins ori1 and ori2 to one another. While the distance between ori1 and ori2 is 1.828 Mbps in NSCV1 it is only 1.118 Mbps in the genome of NSCV2.

To test experimentally, if the ori positioning of NSCV1 and NSCV2 influences the initiation outcome, we re-constructed the ori1 to ori2 arrangement in a genetically accessible system since NSCV1 and NSCV2 are recalcitrant for genetic manipulations. To accomplish this, we transferred a functional hapR to V. cholerae strain MCH1 to render it naturally competent (Lo Scrudato and Blokesch, 2012, 2013). Strain MCH1 was derived from the prototype V. cholerae strain N16961 by fusion of the two chromosomes with a deletion of ori2 (Val et al., 2012). We inserted a copy of ori2 including the flanking genes

parAB and rctB into MCH1 at positions analogous to NSCV1 and NSCV2 with respect to the distance from ori1 giving rise to strains VC61 and VC62, respectively, and performed marker frequency analysis of exponentially grown cultures (**Figure 5**).

The MFA plot of strain VC61 resembled the pattern seen in NSCV1 (**Figure 5A**, compared to **Figure 3A**). In contrast, a clear local peak of marker frequency was seen at the ori2 position of strain VC62 (**Figure 5B**). Thus, even though the origin arrangement is similar in NSCV2 and VC62, the ori2 copy appears to be active only in the engineered strain VC62 and not in NSCV2. To test if the ori2 positioning at the NSCV2 position has more negative effect on growth compared to the ori2 insertion at the NSCV1 position we measured the doubling times of strains VC61 (26 ± 0.1 min) and VC62 (27 ± 0.5 min) (**Supplementary Table S4**). The difference in doubling time was only marginally affected suggesting that the differences in DNA replication between the two strains do not result in severe impairment in growth. We conclude that an ori2 insertion at positions analogous to those in the NSCV strains relative to ori1 can be active as determined by MFA.

## Dam Methyltransferase Is Functional in NSCV Strains

We have shown above that ori2 in strain NSCV2 is functional but appears to be not active. One potential cause of ori2 silencing could be an inactive Dam methylation system. Methylation of the adenine within the Dam recognition site 'GATC' present at ori2 locus is a prerequisite for ori2 activity (Demarre and Chattoraj, 2010). Sequence analyses showed intact dam genes in both strains (Xie et al., 2017). The methylation status of GATC sites can be analyzed by the differential sensitivities of the genomic DNA to DpnI (cleaves only methylated/hemimethylated GATC sites), DpnII (cleaves only unmethylated GATC sites) and Sau3A1 (cleaves both unmethylated and methylated GATC sites) restriction enzymes.

Illumina sequencing. Gray dots represent log numbers of normalized reads as mean values for 1 kbp windows relative to the stationary phase sample. Vertical dotted black lines mark the locations of replication origins of replication and the crtS sites. The solid black lines represent the fitting of regression lines and the green line corresponds to the Loess regression (F = 0.05). Maxima are highlighted by red and minima as blue dots. Plots of biological replicates are shown in Supplementary Figure S1.

Genomic DNAs of NSCV1 and NSCV2 were sensitive to DpnI and Sau3A1 and resistant to DpnII, indicative of full methylation at GATC sites (**Figure 6**). This result was further confirmed by the PacBio sequence data. In PacBio SMRT (Single Molecule Real Time) sequencing, presence of modified base (A in GATC) in the DNA template results in a delayed incorporation of the corresponding T nucleotide, i.e., longer inter-pulse duration (IPD) compared to template lacking the modification (Flusberg et al., 2010). These kinetic measurements create specific signatures for different types of base modifications. Analyses of the PacBio sequence data for modified bases indicated that both NSCV1 (38571/38572 sites) and NSCV2 (37573/37590) have fully methylated (>99.99%) GATC sites. We conclude that Dam is functional in both NSCV strains and a lack of methylation cannot explain the inactivity of ori2 in NSCV2.

#### crtS Sites of NSCV Strains Are Functional

Another possibility for ori2 silencing in NSCV2 is through alterations of crtS activity. This short DNA sequence is found on Chr1 in all available whole genome sequences of two-chromosome strains of Vibrionaceae and its replication triggers ori2 firing on Chr2 (Baek and Chattoraj, 2014; Val et al., 2016; Kemter et al., 2018). Consequently, a nonfunctional crtS could lead to an inactive ori2. To decipher the functionality of crtS sites in NSCV1 and NSCV2

cleaves GATCs independent of the methylation state. DNA cleavage is evident from the disappearance of high molecular weight band and appearance of low in silico, we extracted the respective sequences and aligned them to the consensus sequence we established recently (**Figure 7A**) (Kemter et al., 2018). Notably, all highly conserved parts of the crtS sequence are also conserved in the crtS sequences of NSCV1 and NSCV2 (**Figure 7A**). To test the functionality of crtS in the context of a fused chromosome experimentally, we deleted the crtS in strain VC62 (resulting in strain VC71) and performed marker frequency analysis

(**Figure 7B**). As expected, no local peak of copy number increase was seen at the ori2 position of the 1crtS strain VC71 in contrast to the MFA of the parental strain VC62 (compare **Figures 5B**,**7B**) confirming the necessity of a functional crtS site for ori2 firing. Incidentally, the doubling time of this strain is not much different (25 ± 0.6 min) from that of VC61 and VC62 (**Supplementary Table S4**). To analyze the functionality of the crtS sites from NSCV1 and NSCV2, we inserted these sequences into the genome of E. coli and transformed the corresponding strains with the ori2-based minichromosome synVicII. The rationale behind this approach is the observation of a copy number increase of an ori2-based minichromosome in E. coli strains carrying a functional crtS site (Baek and Chattoraj, 2014; Kemter et al., 2018). The copy number of ori2 minichromosomes was measured in E. coli strains either carrying the crtS sites of strain N16961, NSCV1, NSCV2 or no crtS (**Figure 7C**). In this case, we tested the resistance level of the respective strains to ampicillin as an indirect measure of the minichromosome copy number as described previously (Schallopp et al., 2017). The crtS sites of both NSCV strains increased the copy number of the ori2 minichromosome compared to the strain lacking any crtS site and to a similar extent as the crtS site from the two-chromosome V. cholerae strain N16961 used as positive control (**Figure 7C**). We conclude that crtS sites in NSCV1 and NSCV2 are functional in a heterologous system and a defective crtS might not explain the silent ori2 in strain NSCV2.

## Relative ori1, ori2, and crtS Locations Determine ori2 Activity in NSCV Strains

In a canonical two-chromosome V. cholerae, replication of the crtS on Chr1 triggers the initiation at ori2 on Chr2. This scenario can in principle be analogous in a fused chromosome such as NSCV1 in which ori1, ori2 and crtS are present on the same molecule. Here the crtS is located about 677 kbps away from ori1 and is replicated first with replication forks originating at ori1 which subsequently triggers ori2 firing, which is further downstream of crtS. The organization is different in strain NSCV2 where the crtS lies about 416 kbps downstream of ori2. Thus, in NSCV2, ori2 precedes crtS. This arrangement is the consequence of a large inversion of the Chr1 part of the genome (**Figure 1**). The replicative outcome of this arrangement is difficult to predict considering the known triggering role of crtS; i.e., ori2 being replicated first by replication forks originating at ori1, subsequently upon duplication of the crtS site. In other words, crtS can also retrospectively trigger replication of already replicated ori2 that crtS replication is supposed to trigger in the

molecular weight streak of DNA.

first place. To test this scenario, we moved the crtS site in strain VC62 to the position analogous to strain NSCV2 (resulting in strain VC73). VC73 did not exhibit any significant difference in doubling time compared to other strains (26 ± 0.2 min) (**Supplementary Table S4**). Respective MFA analysis revealed an active ori2, suggesting that an ori2 copy on a fused chromosome can be triggered by a crtS site located downstream relative to ori2, contrary to what was observed in NSCV2 (compare **Figures 3B– 8**). The peak at the ori2 position is not very strong but it is important to note that the fitting of regression lines is automated based on the maxima found within the mapped reads (Kemter et al., 2018). There is no pre-selection of ori positions and the fact that the peak is detected at the ori2 position by the computational fitting model implies that it is an active origin as also seen in the biological replicate (**Supplementary Figure S1F**).

#### DISCUSSION

Vibrio cholerae has partitioned its genome between a true bacterial chromosome and a "domesticated" plasmid replicon (Heidelberg et al., 2000; Venkova-Canova and Chattoraj, 2011; Val et al., 2014b). Unlike most bacteria, this two-replicon arrangement is conserved within the family of Vibrionaceae (Okada et al., 2005; Val et al., 2014b; Ramachandran et al., 2017). It was postulated that the bipartite genome in Vibrio species enables varying the copy numbers of both chromosomes in a niche-specific manner under certain environmental conditions as an adaptation strategy (Heidelberg et al., 2000; Schoolnik and Yildiz, 2000; Srivastava and Chattoraj, 2007). Alternatively, the two-chromosome setup in Vibrio species can be considered as an adaptive feature that enables rapid genome duplication if multiple chromosomes are replicated simultaneously. The latter assumption fits with the observation that Vibrio species is one of the fastest growing bacteria known, a feature that has led to a recent proposal for using the non-pathogenic V. natriegens as the new workhorse for bioengineering (Weinstock et al., 2016; Dalia et al., 2017; Hoffart et al., 2017). However, it brings up the question of how the two replicons are coordinating their replication and regulate over or under replication in order to ensure inheritance of the just one copy of the full complement of the genome into daughter cells upon cell division. Furthermore, the evolutionary driving

force (or forces) that led to the two-chromosome setup in Vibrionaceae remain speculative. One way to better understand the evolutionary significance of this unique feature of Vibrio species is to study naturally occurring exceptions to the two chromosome rule: strains which have evolved into a single chromosome by chromosomal fusion. We have previously identified two Natural Single Chromosome Vibrio (NSCV) cholerae strains (Chapman et al., 2015; Johnson et al., 2015; Xie et al., 2017) and in this study we addressed the functionality and activity of the two origins of replication on the same chromosome.

Fortuitously, the two NSCV strains have different fusion junctions and other genetic rearrangements that provided an opportunity to compare and contrast their ori functionality. We used marker frequency analysis as an indirect measure of ori activity and found ori1 to be active in both strains. In strain NSCV1, we detected an additional, replication activity peak at the ori2 locus indicating that this second origin is active as well. Since the MFA experiments are population based, we cannot exclude that some cells use ori1 only and other cells ori2 only to replicate the fused chromosome. However, we consider it more likely that both origins are firing within the same cell. Interestingly, the copy number ratio of ori2/ter was lower than for ori1/ter consistent with a time delayed initiation of ori2 on the fused chromosome. Such an origin initiation differential has also been observed in conventional two-chromosome Vibrio strains resulting in synchronous termination of replication of the two chromosomes despite their different sizes (Rasmussen et al., 2007; Stokke et al., 2011; Kemter et al., 2018). Conservation of the orchestrated replication timing of the two origins in NSCV1 indicates that the single-chromosome setup does not lead to any origin interference. In two-chromosome Vibrio species, replication of the crtS site located on Chr1 has been shown to trigger ori2 firing (Baek and Chattoraj, 2014; Val et al., 2016; Ramachandran et al., 2018). In V. cholerae N16961, the time between crtS replication and ori2 initiation corresponds to the time the replication fork needs to replicate about 200 kbps of DNA (Val et al., 2016). Considering a similar delay in strain NSCV1, ori2 would have fired long before replication fork originating at ori1 reaches it since the distance between crtS to ori2 is more than 1 Mbps. As a consequence, replication forks originating at ori1 and ori2 will meet at some position of the fused chromosome, the timing of which will dependent on ori1 location and its regulation just as it occurs in two-chromosome V. cholerae.

The scenario is much different in NSCV2. Here, ori2 is replicated before crtS by replication forks originating at ori1 (**Figure 1A**). Intuitively, this would lead to a chaotic replication perturbance because ori2 firing precedes crtS duplication. In the two-chromosome context, crtS duplication occurs first which then triggers ori2 firing. Hence, if replication of the crtS triggers ori2 initiation also in this genomic arrangement it would happen on two copies of the already replicated ori2. In addition, the replication forks coming from this newly initiated ori2 copies have the potential to replicate the crtS in just a few minutes because of their close proximity and could potentially lead to additional rounds of ori2 initiation and thus an uncontrolled ori2 firing. If this were to happen, not only replication control at ori2 would be severely perturbed but also ori1 functioning can be interfered because ori1 might be replicated passively from replication forks, coming from ori2. Interestingly, what we observed in strain NSCV2 is not a replication out of control but instead a simple silencing of ori2 activity. Our data clearly demonstrate that ori2 of NSCV2 is functional in an isolated context as shown by its ability to drive replication of a mini-chromosome in E. coli. In addition, critical factors involved in regulation of ori2 appear to be fully functional in NSCV2, namely the Dam methylation system and the crtS site. Paradoxically, ori2 is not used to initiate DNA replication in the fused chromosome strain, NSCV2.

A recent study offers a simple explanation for our observation on ori2 silencing in NSCV2 (Ramachandran et al., 2018). These authors showed that it is the doubling of the crtS dosage rather than the process of replicating the crtS that triggers ori2 initiation which results in an even number of crtS and ori2 copies. This tendency of the regulatory system to produce similar copy numbers of crtS and ori2 have also been found in engineered systems with multiple crtS sites (de Lemos Martins et al., 2018). If the crtS site is duplicated before ori2 as it is the case in twochromosome V. cholerae strains, the ori2 copy number will be lower compared to crtS and consequently the initiation at ori2 will be triggered to restore crtS/ori2 copy number balance. If the crtS site is replicated after ori2 has been copied by replication forks coming from ori1 as in the case of NSCV2, there is no need to initiate at ori2 because the copy number of crtS and ori2 are in balance already. The lack of initiation at ori2 in NSCV2 is therefore fully consistant with and confirmatory to the findings of the aforementioned studies (de Lemos Martins et al., 2018; Ramachandran et al., 2018). However, in contrast to the expectation of the crtS-to-ori2 copy control model, we observed that ori2 is active in the engineered strain VC73 in which the crtS lies downstream of ori2 analogous to the arrangement in NSCV2 (**Figure 8**). However, it is important to note that although the positioning of ori1, ori2 and crtS are similar in strains NSCV2 and VC73, the genomic contexts of ori2 and crtS are entirely artificial compared to their native positions and furthermore, VC73 does not share the large chromosomal inversion found in NSCV2. More precisely, in VC73, ori2 lies in a region of the original Chr1 and crtS within a region of the original Chr2 (**Figure 8**, left panel). The discrepancy in ori2 firing between the naturally occurring NSCV2 and the engineered strain VC73 may therefore be explained by their different genomic context.

One interesting question that arises is if the two V. cholerae strains with fused chromosomes are evolutionarily stable. It could also be that the fusion occurred only transiently as an artifact of lab cultivation or during the original strain isolation from a patient sample. In fact, chromosome fusions in V. cholerae have been observed previously to occur frequently within a population but the two chromosome configuration seems to provide a selective advantage leading to rapid elimination of the fused chromosomes cells from the population (Val et al., 2014a). Based on this observation, we grew the NSCV strains for about 160 generations and expected a splitting of the

fused chromosome that may provide a selective advantage thereby eliminating the cells with single chromosomes from the population. Contrary to the expectation, we found that cells retained the fused chromosomes indicating that they are in fact, locked in this configuration and not easily revertible. An alternative explanation would be that the fused state is under positive selection pressure. It remains to be seen what the significance of the fused chromosome status in NSCV1 and NSCV2 is, in contrast to vast majority of other V. cholerae strains where two chromosome status is the norm. In any case, the single-chromosome V. cholerae appears to be more frequent than expected as yet another NSCV strain was discovered recently (Yamamoto et al., 2018). We expect future studies of NSCV strains which are a deviation from the norm, would to lead to a better understanding of why most V. cholerae carry their genome split into two as the norm.

# MATERIALS AND METHODS

#### Strains, Plasmids, Oligonucleotides, and Growth Conditions

All strains, plasmids and oligonucleotides used in this study are listed in the **Supplementary Tables S1–S3**. Unless indicated otherwise, the bacterial cells were grown in LB medium at a temperature of 37◦C. Antibiotic selection was performed at the following concentrations, if not indicated differently: Ampicillin 100 µg/ml, Kanamycin 35 µg/ml for E. coli and 70 µg/ml for V. cholerae, Spectinomycin 100 µg/ml, Gentamicin 20 µg/ml, Chloramphenicol 35 µg/ml. Where needed, diaminopimelic acid (DAP) was added to the medium at a concentration of 300 µM. To determine doubling times, cells were grown in LB medium in a 96-well plate at 37◦C in a microplate reader (Infinite M200 pro multimode microplate reader, Tecan). OD<sup>600</sup> was measured every 5 min for 18 h. Doubling times were calculated in exponential phase for OD<sup>600</sup> values between 0.01 and 0.1. For continues cultivation of NSCV strains to investigate potential chromosome splitting strains were inoculated in the morning and grown in 100 ml liquid medium for 24 h. The next morning, new flasks were inoculated 1:1,000 from the previous cultures. After 4 working days (on day 5), 1 ml samples of the cultures were frozen at −80◦C as glycerin stocks. Two days later, new flasks were inoculated from these glycerin stocks and the procedure was continued for a total of 16 days. Before sequencing, these were re-streaked on TCBS medium to verify that it is V. cholerae and to obtain single colonies. For each strain one single colony was used to inoculate a culture for DNA isolation and sequencing.

### Construction of Replicons and Strains

NSCV minichromosomes were constructed by PCR amplifying the respective origins of replication using the primers 1227/1228 on V. cholerae NSCV1 genomic DNA for pMA739 or V. cholerae NSCV2 genomic DNA for pMA755. The vector pMA135 was digested with AscI and the PCR products were integrated in the linearized vector using Gibson Assembly (Gibson et al., 2009) used to transform E. coli WM3064. For construction of chromosomal integrations and deletions, integration cassettes were assembled by using the MoClo system as described previously (Weber et al., 2011; Milbredt et al., 2016; Schindler et al., 2016). The ori2 insertion cassettes on pMA735 and pMA736, as well as the crtS deletion and insertion cassettes on pMA748 and pMA749 were assembled by MoClo reactions using the plasmids indicated in **Supplementary Table S2**, which themselves were assembled by MoClo reactions of the respective PCR products into the respective backbones (primers, templates, and backbones indicated in **Supplementary Table S2**). The linear cassettes were released by restriction enzyme digestions with BsaI. Triparental mating was performed to deliver the plasmids pUXBF13 and pGP704-mTn7-hapR\_ATN from E. coli S17-1 λpir to V. cholerae MCH1 (Bao et al., 1991; Meibom et al., 2005; Val et al., 2012). The created strain V. cholerae VC49 was naturally

competent and by transforming it with the ori2 insertion cassettes released from pMA735 or pMA736, respectively, followed by transformation with pBR-flp and flippase reaction strains VC61 and VC62 were created (De Souza Silva and Blokesch, 2010). V. cholerae strain VC71 was constructed by deletion of the crtS sequence by transforming V. cholerae VC49 with the crtS deletion cassette from pMA748 and a flippase reaction to excise the resistance marker. Subsequently, strain VC71 was transformed with the crtS insertion cassette from pMA749 and the flippase reaction was performed to create strain VC73. To construct plasmid pMA449, crtS was amplified with primers 1439/1440 from gDNA of V. cholerae NSCV1 and for pMA450 with primers 1439/1440 from gDNA of V. cholerae NSCV2. PCR products were assembled in pMA349 by MoClo assembly as described (Schindler et al., 2016) and used to transform E. coli TOP10 cells. Details of further MoClo assemblies are provided in **Supplementary Table S2**. Assemblies were used to transform E. coli DH5α λpir. The derived integration cassettes were cut out with BsaI, integrated into the chromosome of E. coli AB330 and transferred to E. coli MG1655 per P1-transduction. FRT recombination was used to remove the resistance marker.

#### Sequence Comparison

Sequences from V. cholerae N16961, NSCV1 and NSCV2 were compared by multiple sequence alignment using Clustal Omega (Goujon et al., 2010; Sievers et al., 2011) to find single nucleotide polymorphisms. To analyze the whole genomes of the V. cholerae strains NSCV1 and NSCV2 for sequence homology, the alignment free chromosome comparison tool SMASH (Pratas et al., 2015) was used with a minimum block size setting of 5,000 bp.

## Preparation of Genomic DNA From Bacteria

The desired amount of culture (between 0.1 ml and 5 ml) was mixed 1:1 with ice-cold killing buffer and centrifuged at maximum speed and 4◦C for 3 min. The pellet was resuspended in 300 µl TE, 40 µl SDS and 3 µl 0.5 M EDTA. After 5 min incubation at 65◦C, 750 µl Isopropanol were added and the sample was centrifuged 5 min at maximum speed. The pellet was resuspended in 500 µl TE and 2 µl RNAse were added. After 30 min at 37◦C, 2 µl of Proteinase K were added and incubated for another 15 min. The sample was purified via twofold Phenol-Chloroform extraction and precipitated over night at −20◦C by mixing it with 1 ml pure ethanol and 40 µl 3 M sodium acetate.

The next day, DNA was spinned down at maximum speed for 10 min, the pellet was washed with 70% ethanol and centrifuged another 10 min. The pellet was dried and resuspended in 50 µl pure water.

#### Short Read Sequencing (Illumina MiSeq)

V. cholerae genomic DNA was isolated as described (Kemter et al., 2018) from various strains grown under different conditions (log phase vs. stationary) were quantitated using the Qubit fluorimeter (Thermo Fisher) and adjusted to 0.2 ng/µL with nuclease-free water. Sequencing libraries were prepared using the Illumina Nextera XT kit, processed and pooled according to manufacturer's instructions (Illumina). The final, pooled sample was paired-end sequenced (2×300 bp) using the Illumina MiSeq with a v3 chemistry, 600 cycle kit. Post sequencing processing was performed the systems software packages and the final demultiplexed fastq reads produced by the instrument were used for MFA against the reference genome. Raw sequencing data are available on request.

# Long Read Sequencing (PacBio)

Whole genome sequencing was performed on a Pacific Biosciences RSII platform. The sequencing library was prepared using the SMRTbellTM Template Prep Kit (Pacific Biosciences, Menlo Park, CA, United States) following manufacturer's protocol. 5 µg of DNA was fragmented using gTUBE (Covaris Inc., Woburn, MA, United States) to ∼20 kb. After DNA damage repair and ends repair, blunt hairpin adapters were ligated to the template. Non-ligated products were digested by ExoIII and ExoVII exonucleases. Resulting SMRTbell template were purified with AMPure PB beads and size selected on BluePippin system (Sage Science, Beverly, MA, United States), using 0.75% dye-free agarose cassette, with 4–10 kb Hi-Pass protocol and lower cut set on 4 kb. Size selected purified libraries were quantified by Qubit dsDNA High Sensitivity assay. After primer annealing, and P6 polymerase binding, templates were bound to MagBeads for loading. Each sample was sequenced on two SMRT cells, using C4 sequencing kit and 360-min movies per SMRT cell. Presence of unidentified contaminant in two libraries (NSV2 16 days) inhibited sequencing reactions, which manifested in extremely low P1 (2%) and super short reads. Both libraries were subjected to the cleanup procedure that involves binding annealed SMRTbell libraries to magnetic beads, washing the bound annealed DNA SMRTbell templates to remove potential contaminants, and eluting the purified, annealed DNA SMRTbell templates from the magnetic beads. The purified SMRTbell templates were then re-quantified by Qubit and prepared for sequencing on the PacBio RSII according to the Binding Calculator. After cleanup procedure, PacBio RS II instrument sequencing yields were comparable to the other samples. Raw sequencing data are available on request.

# Marker Frequency Analysis

Sequencing reads from a NGS were mapped to the respective genome using the program Geneious (Biomatters Ltd.; Kearse et al., 2012). Read densities were extracted and plotted using custom R scripts as described previously (Kemter et al., 2018).

# Semiquantitative Conjugation

All used replicons possess an oriR6K and were conjugated from E. coli strain WM3064 to MG1655. For overnight cultures of donor and recipient strains the OD600 was determined and the amount of cells corresponding to 1 ml of OD600 = 1 was centrifuged 1 min at 13,000 × g. The cells were washed twice in TBS and resuspended in 100 µl TBS. From each donor strain, 50 µl were mixed with 50 µl of the recipient strain and dropped on LB agar including DAP. After 6 h cells were scraped off the plate, washed twice in TBS. The total CFU of recipient cells was

determined by plating dilutions on LB, while the CFU of plasmid bearing recipients was determined by plating the same dilutions on selective media. The selective CFU was then normalized to total CFU.

#### Microscopy

Differential interference contrast (DIC) microscopy was performed on 1% (w/v) agarose pads in PBS buffer using the Nikon Ti fluorescence microscope (100× objective, NA 1.45).

#### Quantification of Replicon Copy Number via Antibiotic Sensitivity

The copy-up effect of crtS was measured as described (Messerschmidt et al., 2016). Cells were grown in LB medium with either 100 or 500 µg/ml ampicillin at 37◦C in 96-well plates in a microplate reader (Infinite M200 pro multimode microplate reader, Tecan). The main culture (150 µL) was inoculated 1:1,000 and growth curves recorded for 15 h. For better visualization, 1 divided by the time needed to reach an OD<sup>600</sup> of 0.1 was defined as measure of the copy number.

## AUTHOR CONTRIBUTIONS

MB, DS, SS, and TW contributed to the conceptualization of the study. MB, DS, FK, MW, KC, GK, and GP contributed to the methodology. MB, DS, KC, and GK contributed to the software. MB, DS, FK, MW, KC, GK, GP, SS, and TW contributed to the formal analysis of the study. MB, DS, FK, MW, KC, GK, and GP executed the investigation for the study. MB, SS, and TW contributed to the writing of the original draft. SS and TW

#### REFERENCES

reviewed and edited the manuscript. DS, SS, and TW provided the supervision. SS, GP, and TW contributed to the funding acquisition.

# FUNDING

This work was supported within the LOEWE program of the State of Hesse and a grant of the Deutsche Forschungsgemeinschaft (Grant No. WA2713/4-1). Funding for the sequencing work was provided by National Strategic Research Institute (NSRI, University of Nebraska) under the contract number FA4600-12- D-9000 – TOPR 0042 – TO0059).

#### ACKNOWLEDGMENTS

We thank all current and former members of the Waldminghaus lab for help and fruitful discussions. Nadine Schallopp is acknowledged for excellent technical assistance and Mehryad Mataei for experimental support. We are grateful to Melanie Blokesch for advice in V. cholerae genetics and for providing strains and plasmids and to Didier Mazel for providing V. cholerae strain MCH1.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb. 2018.02932/full#supplementary-material

recombination. Plasmid 64, 186–195. doi: 10.1016/j.plasmid.2010. 08.001

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Bruhn, Schindler, Kemter, Wiley, Chase, Koroleva, Palacios, Sozhamannan and Waldminghaus. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Commentary: Functionality of Two Origins of Replication in *Vibrio cholerae* Strains With a Single Chromosome

Bhabatosh Das <sup>1</sup> and Dhruba K. Chattoraj <sup>2</sup> \*

*<sup>1</sup> Translational Health Science and Technology Institute, Faridabad, India, <sup>2</sup> Center of Cancer Research (CCR), National Cancer Institute (NCI) and National Institute Health (NIH), Bethesda, MD, United States*