Identification of MSMEG_0004 and Rv0004

In previous work we probed the transcriptional responses of M. smegmatis, a non-pathogenic model organism for Mtb, to double-stranded DNA (dsDNA) breaks [13]. We identified MSMEG_0004 as being upregulated in response to dsDNA breaks that occurred during logarithmic (log) but not stationary phase. By measuring the expression of MSMEG_0004 during log versus stationary growth phase in the absence of induced DNA damage, we found that MSMEG_0004 is also more highly expressed in log phase in the absence of stress (Fig 1A). We observed a similar result for the Mtb homolog Rv0004 (Fig 1B). Together, these data suggest that expression of 0004 genes is important while the bacteria are actively growing and dividing.

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Fig 1. 0004 (dciA) is regulated by growth phase and is conserved throughout Actinobacteria.

(A,B) Fold change in MSMEG_0004 (dciA_Msm) transcript in M. smegmatis (A) and Rv0004 (dciA_Mtb) transcript in Mtb (B) normalized to 16S rRNA in stationary phase versus logarithmic (log) phase, log phase levels were set to 1. Mean ODλ₆₀₀ ± SD is indicated. Bars represent means ± SEM (n = 3). **** p <0.0001, ** p <0.01, statistical significance was determined by Student’s unpaired t-test. (C) Multiple sequence alignment of 0004 (DciA) protein sequences from different Actinobacteria. Red residues are identical across all sequences, while yellow residues have global similarity scores of at least 0.7 calculated by MultAlin. Within yellow residues, bold letters signify the most common residue. The black line indicates the region of the protein that constitutes DUF721. (D) Genomic context of the dciA (0004) gene in M. smegmatis and Mtb.

https://doi.org/10.1371/journal.pgen.1007115.g001

MSMEG_0004 homologs, which are not present in eukaryotes, E. coli, or B. subtilis, encode hypothetical proteins that contain a domain of unknown function (DUF721; PFAM05258, Fig 1C) and are predicted to be nucleic acid-binding proteins (COG5512 family members) [14]. Brezellec et al. recently identified DUF721 as being widely conserved in proteins they termed DciA [12]. Based on the presence of DUF721 in Rv0004 and MSMEG_0004, we will refer to Rv0004 as dciA_Mtb and MSMEG_0004 as dciA_Msm.

dciA_Msm is located in an operon next to oriC that also contains dnaN and recF, which encode the DNA Pol III beta clamp and a DNA repair protein, respectively (Fig 1D) [15,16]. This operon is located between dnaA and the gyrB-gyrA operon, which encode the replication initiator protein and bacterial gyrase, respectively. This genome structure is conserved between M. smegmatis and Mtb except for MSMEG_0002, a gene that is predicted to encode a 6-phosphogluconate dehydrogenase and is encoded at a separate genomic location in Mtb (Fig 1D) [17]. The orientation of the DNA replication and repair genes dnaA, dnaN, recF, gyrB, gyrA near oriC is a conserved feature in many bacteria [18]. Like dciA_Msm, dnaA, dnaN, and recF were all more highly expressed in log phase versus stationary phase, consistent with roles in DNA replication (S1 Fig). The genomic location and the increased expression of dciA_Mtb and dciA_Msm during log phase support a link between the mycobacterial DciA proteins and DNA replication or repair.

dciA is essential for M. smegmatis viability in culture

In order to study the roles for dciA_Msm and dciA_Mtb in mycobacteria, we took a reverse genetic approach. Attempts to delete dciA_Msm from M. smegmatis and dciA_Mtb from Mtb were unsuccessful, suggesting these genes are essential for viability. To study the dciA genes, we constructed a merodiploid strain in which the dciA_Mtb gene was integrated at the M. smegmatis attB site under the control of a promoter that contains tet operator sites and is linked to a kanamycin resistance cassette (Fig 2A, S1 and S2 Tables). We then deleted dciA_Msm from its endogenous locus (Fig 2B), resulting in the strain ΔdciA_Msm attB::tetdciA_Mtb (Fig 2A). We used the ΔdciA_Msm attB::tetdciA_Mtb strain to study dciA_Mtb in the context of M. smegmatis. Unfortunately, we were unable to engineer similar strains in Mtb, likely due to the difficulty of manipulating the genome near oriC. To confirm that dciA is essential in M. smegmatis, we used a gene-switching technique [19] to replace the kanamycin resistance cassette-linked dciA_Mtb allele at the attB site in the ΔdciA_Msm attB::tetdciA_Mtb strain with a zeocin-resistant plasmid that was either empty or expressed dciA_Mtb. We were able to switch in the zeocin-resistant plasmid expressing dciA_Mtb but we were unable to recover a dciA gene null mutant (Fig 2C), further supporting that dciA_Msm is essential for viability.

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Fig 2. dciA is essential for growth in M. smegmatis.

(A) Construction of ΔdciA_Msm attB::tetdciA_Mtb. (B) Southern blot of wild-type M. smegmatis (wt Msmeg) and ΔdciA_Msm attB::tetdciA_Mtb genomic DNA. After digestion with PvuII, wt Msmeg yields a 693 bp band and ΔdciA_Msm yields a 3033bp band. (C) Agar plates 3 days after transformation of ΔdciA_Msm attB::tetdciA_Mtb with a gene-switching, attB-integrating plasmid encoding either dciA_Mtb (left) or not (right). After incubation of the empty vector plate for over 10 days, a dciA null strain was never recovered. This experiment was repeated at least five times. (D,E) Fold change in dciA_Mtb transcript normalized to sigA during Tet-DciA growth in the absence of ATc where the level at 3 hours (h) for each culture is set to 1. (D) Time course from Tet-DciA cultures grown -ATc, with each point representing n = 4 except the 16 h and 27 h points, which each represent n = 3. (E) dciA_Mtb transcript levels at 16 h growth -ATc compared to 3 h. Center values and error bars represent mean ± SEM. The fold change of 0.040 ± 0.009 is equivalent to a 96.0% ± 0.9% reduction in transcript at 16 h. (F) Representative standard growth curve of Tet-DciA grown +ATc (n = 3) or -ATc (n = 3). Standard growth curves were performed four times. (G) Doubling times from 24 h to 36 h of standard growth curves calculated using the exponential growth equation. Bars indicate means ± SEM (Tet-DciA +ATc n = 6; Tet-DciA -ATc n = 9). (H) Representative continual log growth curve of Tet-DciA grown +ATc (n = 3) or -ATc (n = 3). Continual growth curves were repeated eight times. (I) Doubling times from 24 h to 36 h of continual log growth curve calculated using the exponential growth equation. Bars indicate means ± SEM (Tet-DciA +ATc n = 18; Tet-DciA -ATc n = 21). **** p <0.0001, ** p <0.01, statistical significance determined by Student’s unpaired t-test.

https://doi.org/10.1371/journal.pgen.1007115.g002

Depletion of dciA_Mtb results in inhibition of growth

To study the loss of dciA expression in M. smegmatis, the ΔdciA_Msm attB::tetdciA_Mtb strain was transformed with an episomal plasmid expressing a Tet-ON repressor (TetR) [20] (S1 and S2 Tables). In this M. smegmatis ΔdciA_Msm attB::tetdciA_Mtb +pTetR strain, referred to as Tet-DciA, dciA_Mtb transcript is only expressed in the presence of anhydrotetracycline (ATc). We monitored dciA_Mtb depletion by diluting cultures of Tet-DciA grown in the presence of ATc (+ATc) into liquid media lacking ATc (-ATc) and collecting RNA at 3, 8, 16, 24, 28, and 32 hours after growth -ATc. After 16 hours in depleting conditions, dciA_Mtb transcript levels were 96% lower than at 3 hours (Fig 2D and 2E). To characterize the requirement of dciA_Mtb expression for growth, we monitored the growth of the Tet-DciA strain during dciA_Mtb transcript depletion. Over the first 24 hours, there was no significant difference in the growth of Tet-DciA in depleting (-ATc) or replete (+ATc) conditions (Fig 2F). However, after diluting the stationary phase culture back to early log phase at 24 hours, depleted cells grew more slowly than controls (Fig 2F). We calculated the doubling times of Tet-DciA cultured ± ATc and found that after 24 hours, M. smegmatis depleted of dciA_Mtb grew significantly more slowly than the replete controls (5.8 hour versus 3.7 hour doubling time) (Fig 2G).

To determine whether the growth defect of dciA_Mtb-depleted cells was due to an inability to recover from stationary phase versus DciA_Mtb playing a critical role in growth during log phase, we performed a continual log liquid growth curve where cultures were diluted to early log phase every 8 hours to ensure that they did not enter stationary phase. Similar to the standard growth curve experiments, M. smegmatis depleted of dciA_Mtb did not show a significant growth defect until after 24 hours of growth–ATc (Fig 2H). We calculated the doubling times of Tet-DciA cultured ± ATc and found that after 24 hours of continual log growth, M. smegmatis depleted of dciA_Mtb grew significantly more slowly than the replete controls (4.7 hour versus 3.4 hour doubling time) (Fig 2I). After 40 hours of growth in depleting conditions, suppressors of the Tet-ON system are selected for and the levels of dciA_Mtb transcript are no longer controlled by ATc. Together, these data demonstrate that DciA is important for M. smegmatis growth.

Depletion of dciA_Mtb results in a block in cell cycle progression

We have shown that dciA is essential for growth in culture and is involved in responding to DNA damage in M. smegmatis. However, a role in DNA damage responses alone cannot explain the essentiality of dciA, since genes key for mycobacterial DNA repair pathways are not essential in vitro [21–24]. The location of mycobacterial dciA genes in an operon with and adjacent to genes involved in DNA replication raises the question of whether DciA plays a role in this essential process in mycobacteria. In addition, P. aeruginosa DciA (DciA_Pa) has been implicated in DNA replication [12]. To investigate a role for mycobacterial DciA in DNA replication and the bacterial cell cycle, we monitored the cellular morphology of Tet-DciA cultured ± ATc in the continual log growth curve (Fig 2H). By 24 hours of depletion, Tet-DciA cultured without ATc were on average 12.9% (p <0.01) longer than Tet-DciA cultured in dciA_Mtb-replete conditions (S2 Fig). After 36 hours, Tet-DciA cultured without ATc averaged over 60% longer (p <0.0001) than Tet-DciA grown in dciA_Mtb-replete conditions (Fig 3A, 3B and 3D). The elongated cellular morphology observed during dciA_Mtb depletion indicates a block in cell cycle progression.

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Fig 3. Depletion of DciA leads to a block in cell cycle progression.

(A) Fluorescence microscopy of Tet-DciA cells grown for 36 hours (h) +ATc (top row) or -ATc (bottom four rows) pictured in brightfield, stained with DAPI for DNA, or stained with FM1-43FX for membranes. Blue arrows indicate areas free of DNA staining and white chevrons indicate anucleate cells. Scale bars are 10 μm. (B) Cell lengths of Tet-DciA grown ± ATc at 36 h. (C) % DNA occupation, which is the nucleoid length divided by the cell length times 100, of Tet-DciA grown ± ATc at 36 h. (D) Means ± SD from data in (B) and (C). (E) TEM of Tet-DciA grown ± ATc at 36 h. Arrows indicate septa. Scale bars are 500 nm.

https://doi.org/10.1371/journal.pgen.1007115.g003

To characterize where in the cell cycle dciA_Mtb-depleted M. smegmatis is blocked, we analyzed nucleoid morphology and septum formation. To observe nucleoid morphology, Tet-DciA was grown ± ATc, DNA was stained with DAPI, and cells were visualized by fluorescent microscopy (Fig 3A). The nucleoid in dciA_Mtb-replete Tet-DciA (+ATc) appears as several distinct puncta throughout the length of the cell (Fig 3A, top row, second panel), as has been reported previously for M. smegmatis [25]. Following dciA_Mtb depletion, the DAPI-stained nucleoid still appeared as several distinct puncta, but was not distributed throughout the length of the cell. Instead, there were areas free of DNA staining found at the poles (Fig 3A). We quantified the abnormal nucleoid morphology using “% DNA occupation,” which represents the nucleoid length as a percentage of the total cell length. Tet-DciA depleted for dciA_Mtb exhibit significantly lower % DNA occupation starting at 16 hours of depletion (Fig 3C and 3D; S2 Fig). The observation that depletion of dciA_Mtb leads to significantly lower % DNA occupation at 16 hours even though cell lengths are not significantly longer at this time point (S2 Fig) indicates that abnormal nucleoid morphology is the earlier phenotype, and that a DNA-related function causes the cell cycle block and slowed growth (Fig 2I).

We also observed the presence of 9% anucleate cells by DAPI staining upon dciA_Mtb depletion (Fig 3A). Anucleate cells result when DNA replication and cellular division are uncoupled. The presence of anucleate cells also indicates that cell division is still able to occur. To confirm that septum formation was intact, we used transmission electron microscopy (TEM) and found that cells depleted for dciA_Mtb were still able to form normal septa (Fig 3E).

To visualize cell division of dciA_Mtb-depleted M. smegmatis in real time, we performed live-cell imaging [26] of Tet-DciA grown ± ATc in a constant-flow microfluidic device (S1 and S2 Movies). Using FM4-64 membrane stain, we observed that dciA_Mtb-depleted cells can form septa and undergo cell division (S2 Movie). We also detected increased cell death in dciA_Mtb-depleted cells compared to controls, where dead cells stop elongating and take up more FM4-64 dye, thus exhibiting a rapid increase in fluorescence (S2 Movie). 9.4% of dciA_Mtb-depleted cells displayed these cell death characteristics. In support of our fixed fluorescent microscopy, Tet-DciA cells grown -ATc were on average 56% longer at birth and division than Tet-DciA grown +ATc (S3 Fig, S1 and S2 Movies, p <0.0001). However, despite the increased average length of dciA_Mtb-depleted cells, we also observed division of unusually small cells (S3E Fig), many of which died shortly after division. In general, we observed greater variability in birth length in dciA_Mtb depleted versus replete cells, with coefficients of variation of 41.7% and 18.2%, respectively (S3C Fig). The increased heterogeneity among cell birth lengths during dciA_Mtb depletion indicates a disruption in the coordination of septum formation with cell growth, leading to the dysregulation of cell size. Thus, time-lapse imaging demonstrates that in addition to dciA_Mtb-depleted cells being elongated on average, they are also characterized by increased variation in cell size and frequency of death.

Together, our data show that cell division is intact but chromosome replication or segregation is blocked during dciA_Mtb depletion. In support of this conclusion, the abnormal nucleoid morphologies observed in M. smegmatis depleted of dciA_Mtb phenocopy those of M. smegmatis depleted of DnaA, the chromosomal replication initiator protein [25] (Fig 4A–4D, S4 Fig), but not M. smegmatis depleted of FtsZ, the protein that comprises the Z-ring precursor of the septum [27] (Fig 4E–4H, S5 Fig).

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Fig 4. dciA_Mtb depletion phenocopies dnaA depletion and not ftsZ depletion in M. smegmatis.

(A) Fluorescence microscopy of rgm36, an acetamide-inducible dnaA depletion M. smegmatis strain [25], grown in the presence (top row) or absence (bottom two rows) of the acetamide (acet.) inducer pictured in brightfield, stained with DAPI for DNA, and stained with FM1-43FX for membrane. Blue arrows indicate areas free of DNA staining, while white chevrons indicate anucleate cells. Scale bars are 10 μm. (B) Cell length and (C) DNA occupation histograms of rgm36 cells after 24 hours of growth in depleting (-acet.) or replete (+acet.) conditions. (D) Table displays averages ± standard deviations from data depicted in histograms (B,C), along with p-values determined by Student’s t-test. (E) Fluorescence microscopy of csm362, a TetOn FtsZ depletion M. smegmatis strain, in the presence (top row) or absence (bottom two rows) of ATc pictured in brightfield, stained with DAPI for DNA, and stained with FM1-43FX for membrane. Scale bars are 10 μm. (F) Cell length and (G) % DNA occupation histograms of csm362 cells after 24 hours of growth in depleting (-ATc) or replete (+ATc) conditions. (H) Table displays averages ± standard deviations from data depicted in histograms (F,G), along with p-values determined by Student’s t-test.

https://doi.org/10.1371/journal.pgen.1007115.g004

dciA depletion results in decreased DNA synthesis

The data so far support a model that DciA homologs are required for either DNA replication or chromosome segregation. To determine if dciA_Mtb-depleted M. smegmatis is defective in DNA replication, we directly measured rates of DNA synthesis using a nucleotide incorporation assay [25,28]. Specifically, we determined the rates of [5,6-³H]-thymidine incorporation into DNA by Tet-DciA cells grown ± ATc in continual log phase. We found that the rate of DNA synthesis in M. smegmatis was significantly lower at 24 hours (Fig 5A and 5C) and 36 hours (Fig 5B and 5C) in Tet-DciA grown -ATc relative to Tet-DciA grown +ATc, proving that the rate of DNA replication itself decreases upon dciA_Mtb depletion. To further confirm this defect in DNA replication, we stained Tet-DciA grown ± ATc with DAPI and measured DNA content using flow cytometry. dciA_Mtb-depleted cells had lower DNA content per cell based on the mean fluorescence intensity (MFI) of DAPI staining relative to controls (Fig 5D and S6A Fig). Together, these data demonstrate that DciA_Mtb is involved in DNA replication, but do not differentiate between which step(s) of DNA replication DciA_Mtb acts.

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Fig 5. Depletion of DciA leads to decreased DNA synthesis.

(A,B) ³H incorporation in Tet-DciA as measured by ³H counts per minute (cpm) per colony forming unit (cfu) at 24 (A) and 36 (B) hours (h) growth ± ATc. Symbols represent mean ± SEM (n = 3). (C) Rates of ³H incorporation calculated from slopes of the lines of the linear regression from (A) and (B), symbols represent each biological replicate. (D) Geometric mean fluorescence intensity (MFI) of DAPI stained Tet-DciA grown ± ATc for 36 h relative to an unstained sample determined by flow cytometry. Representative flow cytometry histograms are found in S6A Fig. Symbols represent each biological replicate, center values and error bars represent means ± SEM. ** p <0.01, * p <0.05, statistical significance determined by Student’s unpaired t-test.

https://doi.org/10.1371/journal.pgen.1007115.g005

DciA_Mtb binds DNA

To determine how DciA proteins function in DNA replication, we investigated macromolecular interaction partners of DciA_Mtb. DciA_Mtb has a calculated isoelectric point (pI) around 12, indicating that the protein is positively-charged at neutral pH. Other proteins in mycobacteria with high isoelectric points include histone-like proteins (H-NS, HupB) and integration host factor (IHF), which are all known to bind DNA [29–32]. DciA_Mtb, as a member of COG5512, is also predicted to be a nucleic-acid binding protein [14]. Binding to nucleic acid could be relevant to the role for DciA in DNA replication given the numerous protein-nucleic acid complexes that form during this process. We tested the DNA binding activity of purified DciA_Mtb protein (S7A Fig) in electromobility shift assays (EMSAs) with ³²P-radiolabeled DNA. Due to its role in DNA replication (Figs 3 and 5), we first tested whether DciA_Mtb was able to bind oriC DNA in vitro. DciA_Mtb was able to bind and shift a 553 basepair (bp) dsDNA fragment containing Mtb oriC DNA [33] (Fig 6A). Given its high isoelectric point and the negative charge of DNA, we hypothesized that DciA_Mtb would be able to bind any DNA sequence and not just oriC. Indeed, we found that DciA_Mtb was also able to shift a 333 bp dsDNA sequence from a site in the genome distantly located from oriC (S7B Fig). DciA_Mtb’s DNA binding activity is not limited to dsDNA sequences as DciA_Mtb was also able to bind and shift a 72 nucleotide single-stranded DNA (ssDNA) oligo (S7C Fig). These data indicate that DciA_Mtb can bind diverse double and single stranded DNA molecules in vitro, including oriC. This sequence independent DNA-binding activity could relate to the role of DciA_Mtb in DNA replication.

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Fig 6. DciA_Mtb binds DNA, interacts directly with DnaB, and affects the DnaB-DnaA interaction.

(A) Autoradiograph of EMSA with DciA_Mtb protein and 4.8ng oriC_Mtb dsDNA separated by native PAGE. All lanes contain ³²P-labeled oriC_Mtb DNA. Amount of DciA_Mtb in each lane is indicated. (B) Proteins involved in DNA replication or repair that were identified by MS analysis as co-immunoprecipitating with HA-DciA_Mtb. (C) GelCode Blue-stained SDS-PAGE of pull-down experiments using HA-DciA_Mtb, DnaB, and DnaA. Lanes 1–3 after ladder contain purified proteins used in pull-downs. Lanes 4–7 contain eluates from pull downs with immobilized HA-DciA_Mtb incubated with DnaA (lanes 4–5) or DnaB (lanes 6–7). Proteins used were purified in the absence (lanes 4,6) or presence (lanes 5,7) of Benzonase (Benz.). Top arrow indicates the size of DnaA and DnaB. Bottom arrow indicates the size of HA-DciA_Mtb. (D) Western blot analysis of pull-downs with DnaB-FLAG as bait and DciA_Mtb as prey. (E) Representative curves from BLI showing the association and dissociation of DciA_Mtb at the indicated concentrations with immobilized biotinylated-DnaB. (F,G) Representative western blot of pull-downs with (F) DnaA-HA as bait and DnaB-FLAG as prey or (G) DnaB-FLAG as bait and DnaA-HA as prey and either no (lanes 1,6), 0.5x (lanes 2,7), 1x (lanes 3,8), 2x (lanes 4,9), or 4x (lanes 5,10) molar DciA_Mtb relative to the bait. (H) Quantification of the ratio of prey-to-bait for triplicate western blots like those shown in (F) and (G) where the ratio for lanes with no DciA_Mtb is set to 1 and the ratio for all other samples is relative to the lane with no DciA_Mtb. Symbols represent each replicate, center values and error bars represent mean ± SEM. (I) Fold enrichment in levels of DNA fragments containing oriC (oriC1-3, see S8A Fig), the promoter of the rplN operon (rplN), or the sigA coding sequence (sigAIN) in samples immunoprecipitated with an anti-HA antibody (αHA-IP) or input samples (input) from strains expressing untagged DciA_Mtb (No tag, black bars) and HA-DciA_Mtb (grey bars) strains. Data is represented as fold enrichment relative to No tag levels. Bars represent mean ± SEM (n = 5 except No tag sigAIN, which is n = 4). **** p <0.0001, * p <0.05, ns is not significant, statistical significance was determined by one-way ANOVA and Tukey’s multiple comparison test.

https://doi.org/10.1371/journal.pgen.1007115.g006

DciA_Mtb associates with proteins involved in DNA replication and repair

We next identified the mycobacterial proteins that associate with DciA_Mtb by performing co-immunoprecipitation mass spectrometry (co-IP/MS). We engineered a strain of M. smegmatis that encodes an HA-tagged version of dciA_Mtb (HA-DciA_Mtb) as its only dciA allele (S1 Table). We generated cell lysate from this strain and immunoprecipitated HA-DciA_Mtb along with associated protein complexes using an anti-HA antibody conjugated agarose (Sigma). After eluting with HA peptide, eluates were separated by SDS-PAGE, silver-stained, and bands specific to the HA-DciA_Mtb lane (S7D Fig) were isolated and analyzed by MS. We observed a similar banding pattern when we performed these experiments with DNAse-treated cell lysates (S7E Fig). The most abundant band on the silver-stained gel contained ClpX (S7D Fig), a component of the essential ClpXP protease. The association of DciA_Mtb with Clp protease may explain our inability to detect native DciA proteins by western blot. In addition to components of the Clp protease, we also found that DciA_Mtb associates with a number of proteins involved in DNA replication and repair (Fig 6B, S4 Table).

DciA_Mtb interacts directly with DnaB

Since we have shown that DciA_Mtb is involved in DNA replication (Fig 5), we sought to confirm the association of DciA_Mtb with the co-immunoprecipitated DNA replication proteins. These proteins included gyrase, DnaX (τ clamp-loader subunit of DNA Pol III), DnaB (replicative helicase), and DnaA (replication initiator protein). To prioritize our studies, we identified DNA replication proteins that are conserved in E. coli and B. subtilis but do not yet have known homologs in mycobacteria, namely DnaA regulators and DnaB helicase loaders (Table 1). Since DnaA and DnaB were both found to associate with DciA_Mtb through co-IP/MS, we first tested whether DciA_Mtb directly interacts with these proteins.

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Table 1. Review of E. coli and B. subtilis DNA replication proteins that are conserved in Mtb.

https://doi.org/10.1371/journal.pgen.1007115.t001

We performed pull-down experiments similar to the co-IP approach described earlier, but using purified recombinant HA-DciA_Mtb, DnaB, and DnaA. We immobilized HA-DciA_Mtb onto anti-HA agarose and added DnaA or DnaB. Analysis of the protein complexes eluted with HA peptide showed that HA-DciA_Mtb pulls down both DnaA (Fig 6C lane 4) and DnaB (Fig 6C lane 6), but not a negative control protein, Rel_Mtb^1-394 (S7F Fig). Since HA-DciA_Mtb, DnaA, and DnaB can all bind DNA, we tested whether the interactions between these proteins were dependent on nucleic acid by performing the same pull-down experiments using recombinant proteins purified from Benzonase-treated lysates to degrade nucleic acids. When proteins purified from Benzonase-treated lysates were used, HA-DciA_Mtb failed to pull down DnaA (Fig 6C lane 5) but retained its interaction with DnaB (Fig 6C lane 7). Therefore, DciA_Mtb interacts directly with DnaB but depends on nucleic acid to associate with DnaA.

We confirmed that DciA_Mtb and DnaB directly interact by performing the reciprocal pull-down with purified FLAG-tagged DnaB (DnaB-FLAG) immobilized as bait and DciA_Mtb as prey (Fig 6D). We performed Bio-Layer Interferometry (BLI) to quantify the affinity of the interaction between DnaB and DciA_Mtb. The association and dissociation of varying concentrations of DciA_Mtb to biotinylated DnaB loaded onto streptavidin-coated biosensor pins (ForteBio) were measured (Fig 6E) and an affinity constant (K_D) of 210.9 ± 8.19 nM was calculated (fit R² = .9018) (S7G Fig). These data demonstrate that DnaB and DciA_Mtb interact in a dose-dependent manner. The calculated affinity constant in the nanomolar range suggests that the interaction could occur under physiological conditions, although cellular concentrations of DnaB and DciA_Mtb need to be quantified to confirm this.

DciA_Mtb can affect the interaction between DnaB and DnaA

In E. coli, the interaction between DnaA and DnaB is required for efficient loading of DnaB [51,52]. The result that DciA_Mtb directly binds DnaB and can indirectly associate with DnaA led us to probe whether DciA_Mtb affects DnaB-DnaA complex formation. Mtb DnaA has been shown to interact with the N-terminus of DnaB (residues 1–206) [36], and we confirmed that the full length Mtb DnaB-FLAG and DnaA-HA proteins directly interact in our system (Fig 6F lane 1 and Fig 6G lane 6). To determine the effect of DciA_Mtb on the DnaA-DnaB interaction, we performed pull-downs with DnaA-HA as bait and DnaB-FLAG as prey in the presence of varying amounts of DciA_Mtb. All pull-downs were performed using proteins purified from Benzonase-treated lysates to exclude contributions from nucleic acid interactions. As increasing amounts of DciA_Mtb were added, DnaA-HA pulled down more DnaB-FLAG (Fig 6F and 6H). These results indicate that DciA_Mtb facilitates DnaA-DnaB complex formation. In contrast, when DnaB-FLAG was immobilized as bait and DnaA-HA was added as prey, increasing the amount of DciA_Mtb present did not change the amount of DnaB-FLAG that associated with DnaA-HA (Fig 6G and 6H). Therefore, DciA_Mtb can affect DnaB-DnaA complex formation, but this is dependent on which protein is immobilized. One possible explanation for this observation is that DnaB, which functions as a hexamer [34], is unable to properly hexamerize while immobilized as bait, affecting DciA_Mtb activity. The ability of DciA_Mtb to positively affect the DnaB-DnaA interaction suggests that DciA_Mtb promotes rather than inhibits DNA replication, which is consistent with the observations that dciA_Mtb depletion leads to decreased DNA synthesis and DNA content (Fig 5).

DciA_Mtb is enriched at oriC in M. smegmatis

The ability of DciA_Mtb to affect the association of DnaB with DnaA suggests that DciA functions during DNA replication initiation. To test whether DciA is enriched at oriC where initiation occurs, we performed chromatin immunoprecipitation quantitative PCR (ChIP-qPCR) experiments. These experiments were performed with log-phase cultures of M. smegmatis strains that express HA-tagged DciA (HA-DciA), untagged DciA (no tag), and HA-tagged CarD (HA-CarD), a mycobacterial transcription factor that associates with RNA Polymerase and is known to localize to every promoter throughout the M. smegmatis genome [53]. Protein-nucleic acid complexes were immunoprecipitated from each culture using anti-HA resin and co-immunoprecipitated DNA was probed for sequences specific for oriC (S8A Fig), the rplN promoter, and internal to the sigA gene using qPCR. Enrichment of sequences within a given sample was determined relative to the no tag control.

As expected, only DNA fragments containing promoters (oriC and rplN promoter) were specifically and significantly enriched for following immunoprecipitation with HA-CarD (S8B Fig). The only DNA fragments that were significantly enriched for following immunoprecipitation of HA-DciA_Mtb were those containing the oriC (Fig 6I). As a control, no sequences were enriched in input samples before immunoprecipitation (Fig 6I and S8C Fig). The specific enrichment of DciA_Mtb at oriC and not at other areas of the chromosome indicates that DciA_Mtb is involved in DNA replication initiation, which is consistent with its role in affecting the interaction between DnaB and the DnaA initiator protein (Fig 6F–6H).

The C-terminus of DciA_Mtb is similar to the DnaA N-terminal domain

To investigate how DciA_Mtb facilitates the association between DnaA and DnaB, we used structural prediction tools to gain further insight into the protein architecture. Both Phyre2 [54] and I-TASSER [55] predicted that a region in the C-terminus of DciA_Mtb within DUF721 (~92–142 aa) is structurally similar to the N-terminal domain (NTD) of DnaA in B. subtilis (DnaA_Bs, PDB: 4TPSD) and E. coli (DnaA_Ec, PDB: 2E0GA) (Fig 7A and 7B). We named this region of DciA_Mtb the DnaA NTD-Like (DANL) domain. The DnaA NTD is responsible for many protein-protein interactions important for DNA replication, including the interaction of DnaA_Ec with DnaB, DiaA, and other DnaA_Ec monomers, the interaction of DnaA_Bs with SirA, and the interaction between H. pylori DnaA and HobA [51,52,56–58]. Phenylalanine 46 (F46) in DnaA_Ec, which is equivalent to F49 in DnaA_Bs, is specifically important for DnaA_Ec to load DnaB [57]. Though shifted by one residue, our structural alignment shows that a tryptophan at position 113 in DciA_Mtb (W113) is the closest aromatic amino acid to F49 in DnaA_Bs (Fig 7A, 7C and 7D). Consurf alignment [59] reveals that the region around W113 is one of two highly conserved regions of the DciA_Mtb protein that are both located in DciA_Mtb’s C-terminus (Fig 7E). The position of W113 in the predicted structural model and the conservation of this region across DciA homologs suggest that W113 may be important for DciA_Mtb activity.

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Fig 7. The C-terminus of DciA contains a structurally predicted DnaA NTD-Like (DANL) domain.

(A) Alignment generated by Phyre2 of the DciA_Mtb DANL domain with B. subtilis DnaA NTD from PDB:4TPSD. Red boxes indicate identical residues, while structural predictions above the alignment refer to the predicted structure of DciA_Mtb (arrows for beta sheets, loops for alpha helices, and TT for the turn between the two beta sheets). (B) Structural model of DciA_Mtb DANL domain generated by Phyre2. DciA_Mtb DANL (residues 92–142, magenta) and the B. subtilis DnaA NTD (residues 29–80, gold, PDB:4TPSD) were aligned using PyMol. (C) The structural alignment is shown in the context of the SirA-DnaA complex (PDB:4TPS) with SirA in blue and the inter-facial locations of F49 of DnaA and W113 of DciA_Mtb highlighted. (D) Zoomed in image of (C). (E) Consurf analysis based on 150 non-redundant homologs of DciA_Mtb protein. Highly conserved residues are in magenta and variable residues are teal. The W113 residue is underlined. Due to a probable misannotation of the start codon, the beginning of actinobacterial DciA proteins share poor homology (Fig 1C). Therefore, Consurf analysis of DciA begins at the glycine that marks the beginning of homology across mycobacterial DciA proteins (at residue 21 in Fig 1C).

https://doi.org/10.1371/journal.pgen.1007115.g007

A single point mutation in the DciA_Mtb DANL domain results in similar phenotypes as dciA depletion

To test if W113 in the DciA_Mtb DANL domain is important for DciA_Mtb function, we engineered M. smegmatis to expresses a version of DciA_Mtb where the W113 is mutated to an alanine (W113A) as its only allele of dciA. The W113A mutation leads to a growth defect (Fig 8A and 8B), elongated cellular and abnormal nucleoid morphologies (Fig 8C–8F), and decreased DNA content (Fig 8G; S6B Fig). The observation that the mutation of a single residue can cause similar phenotypes to those observed during dciA_Mtb depletion confirms that the dciA_Mtb-depletion phenotypes were not due solely to the depletion of an essential protein. These experiments also show that the W113 residue, which is located within the region of DciA_Mtb that is predicted to be structurally similar to the protein-protein interaction domain of DnaA, is important for DciA_Mtb activity.

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Fig 8. Mutating a conserved tryptophan in the DciA DANL domain causes similar phenotypes as dciA_Mtb depletion.

(A) Representative growth curve of the M. smegmatis W113A strain (n = 3) and the wild-type control (wt ctrl, n = 4). Symbols represent means ± SEM. (B) Doubling times calculated from three independent experiments, bars indicate means ± SEM (wt ctrl n = 9; W113A n = 9, p <0.0001). (C) Fluorescence microscopy of wt ctrl (top row) or W113A (bottom two rows) stained with DAPI for DNA. Blue arrows indicate areas free of DNA staining and the white chevron indicates an anucleate cell. Scale bars are 10 μm. Cell lengths (D) and % DNA occupation (E) of wt ctrl or W113A cells. (F) Means ± SD from data in (D) and (E). (G) Geometric mean fluorescence intensity (MFI) of DAPI-stained wt ctrl or W113A relative to an unstained sample determined by flow cytometry. Representative flow cytometry histograms are found in S6B Fig. Symbols represent each replicate, center values and error bars represent mean ± SEM, * p <0.05, **** p <0.0001, statistical significance determined by Student’s unpaired t-test. (H,I) Western blot analyses of eluates from pull-downs with DnaB-FLAG as prey and either HA-DciA_Mtb^Δ89–167 (N-terminus) or HA-DciA_Mtb^Δ1–88 (C-terminus) (H) or HA-DciA_Mtb^Δ1–88 or HA-DciA_Mtb^{W113AΔ1–88} (I) as bait. Each pull down was repeated at least three times with different preparations of purified proteins.

https://doi.org/10.1371/journal.pgen.1007115.g008

The W113A mutation affects the binding of the DciA_Mtb C-terminus to DnaB

In order to determine how W113 is contributing to DciA_Mtb function, we purified DciA_Mtb^W113A protein and tested its ability to perform the functions we have assigned to DciA_Mtb. DciA_Mtb^W113A was able to bind and shift DNA (S9A Fig) at similar concentrations to DciA_Mtb wild-type protein (Fig 6A). DciA_Mtb^W113A was also able to bind DnaB in the presence (S9B Fig) and absence (S9C Fig) of nucleic acid, as well as affect DnaA-DnaB complex formation similarly to wild-type DciA_Mtb (S9D–S9F Fig). The DNA and DnaB binding activities of DciA_Mtb^W113A indicate that the DciA_Mtb^W113A protein is functional, structurally intact, and not grossly misfolded.

Given the location of W113 in the predicted DANL protein-protein interaction domain, we were surprised that the W113A mutation did not affect the interaction between DciA_Mtb and DnaB. To test whether the DANL domain itself was involved in the interaction with DnaB, we purified the HA-tagged DciA_Mtb N-terminus (HA-DciA_Mtb^Δ89–167) and HA-tagged DciA_Mtb C-terminus (HA-DciA_Mtb^Δ1–88, containing the DANL domain) protein truncations, and tested their ability to bind DnaB. We found that both the N-terminus (HA-DciA_Mtb^Δ89–167) and the C-terminus (HA-DciA_Mtb^Δ1–88) of DciA_Mtb can individually bind DnaB-FLAG (Fig 8H). While we predicted the DANL-domain within the C-terminus of DciA_Mtb would bind DnaB based on its similarity to the DnaA NTD, we did not anticipate that the DciA_Mtb N-terminus would also interact with DnaB. This is particularly interesting since there are no known or predicted structures for the N-terminus of DciA_Mtb. We were unable to isolate viable M. smegmatis dciA_Mtb^Δ89–167 or dciA_Mtb^Δ1–88 mutants using the gene-switching approach described earlier, indicating that both halves of the protein contribute to DciA_Mtb’s essential cellular function. Therefore, the ability of both the N- and C-terminus of DciA to associate with DnaB is unlikely due to redundant roles for these two regions of the protein.

We tested whether the DnaB-binding activity provided by the N-terminus of DciA_Mtb was precluding our ability to assess the contribution of W113 to the interaction with DnaB. To investigate this, we purified the HA-tagged DciA_Mtb^W113A C-terminus (HA-DciA_Mtb^{W113AΔ1–88}) and tested its ability to bind DnaB-FLAG. We found that HA-DciA_Mtb^{W113AΔ1–88} associated with more DnaB-FLAG than the wild-type HA-DciA_Mtb^Δ1–88 (Fig 8I). Therefore, the W113A mutation in DciA_Mtb affects the association of the C-terminal DANL domain with DnaB and it is possible that the W113A M. smegmatis mutant strain could have defects due to altered DnaB binding by DciA_Mtb^W113A.

Bacterial strains and growth conditions

Mtb Erdman strain was grown at 37°C in Middlebrook 7H9 supplemented with 0.5% glycerol, 0.05% Tween 80, and 10% oleic acid/albumin/dextrose/catalase. M. smegmatis mc²155 and its derivatives were grown on LB agar plates supplemented with 0.5% dextrose and 0.5% glycerol and in LB broth supplemented with 0.5% dextrose, 0.5% glycerol, and 0.05% Tween 80 except in the [5,6-³H] uracil experiment, in which cells were grown in Middlebrook 7H9 supplemented with 0.5% dextrose, 0.5% glycerol, and 0.05% Tween 80 and for live-cell imaging, in which M. smegmatis was grown in Middlebrook 7H9 supplemented with 0.05% Tween 80, 10% ADC (albumin, dextrose, catalase), and 0.2% glycerol. All bacterial strains, plasmids, and primers used in this study are described in detail in S1–S3 Tables.

Quantitative real-time PCR (qRT-PCR)

RNA was extracted from mycobacteria using Trizol (Invitrogen) followed by high salt and isopropanol precipitation. Contaminating genomic DNA was removed using the TURBO DNA-free kit (ThermoFisher Scientific), cDNA was synthesized using Superscript III (Invitrogen), and iTaq Universal SYBR Green Supermix (Bio-Rad) was used in qRT-PCR reactions. Primers used to amplify 16S rRNA from Mtb and M. smegmatis, dciaA_Mtb (Rv0004), dciA_Msm (MSMEG_0004), and M. smegmatis dnaA, dnaN, MSMEG_0002, recF, gyrB and sigA are found in S3 Table. Levels of dciA_Msm, dciaA_Mtb, dnaA, dnaN, MSMEG_0002, recF, and gyrB, were normalized to either 16S rrnA or sigA transcript levels as previously described [68].

Fluorescence microscopy

M. smegmatis was collected, washed once with equal volume Phosphate Buffered Saline (PBS), resuspended in equal volume 1 μg/ml FM1-43FX (Thermo Fisher) diluted in PBS and incubated for 20 minutes at 37°C shaking. Cells were then fixed with 3% paraformaldehyde in PBS for 30 minutes shaking at 37°C. Fixed cells were applied to 0.1% poly-L-lysine (Sigma) treated multitest slides (MP Biomedicals) and then washed once with PBS. Cells were permeabilized by treatment with 2 mg/ml lysozyme (Sigma) at 37°C for 30 minutes followed by 0.1% Triton-X 100 (Sigma) for exactly five minutes at room temperature [69]. Cells were rinsed with PBS and stained with DAPI (Thermo Fisher) diluted to 5μg/ml with Slow Fade Antifade Equilibration Buffer (ThermoFisher) and mounted using the Slow Fade Antifade Kit according to the manufacturer’s instructions. Slides were visualized with a Zeiss Axioskop 2 Mot Plus equipped with an Axiocam MRm monochrome camera and a 100X, 1.4 numerical aperature Zeiss Plan Apochromat oil objective and images were acquired using Axiovision 4.6 software, or using an upright Zeiss Axio Imager M2 fluorescence microscope and the Zen Blue image acquisition software.

Live-cell imaging

M. smegmatis cells were grown to log phase overnight with shaking at 37°C. M. smegmatis cells were filtered through a 10 μm filter to remove clumps before being loaded into a custom polydimethylsiloxane (PDMS) microfluidic device, as before [26]. The viewing device incorporates a main microfluidic channel for continuous flow for growth media, with a height of approximately 10–17 μm, and viewing chambers with a diameter of 60 μm and a height of 0.8−0.9 μm. 2% DMSO and 0.0625 mg/ml FM4-64 are present in the flowed medium, to stain septal membranes.

The microfluidics device was placed on an automated microscope stage housed within an environmental chamber maintained at 37°C. M. smegmatis cells were imaged for up to 40 hours using a widefield DeltaVision PersonalDV (Applied Precision, Inc) with a hardware-based autofocus. Cells were illuminated with an InsightSSI Solid State Illumination system every 15 minutes: FM4-64 was visualized with 475nm excitation and 679 nm emission wavelengths, and cells were imaged using transmitted light brightfield imaging. Each set of images was illuminated with identical imaging conditions that were optimized to decrease phototoxicity inherent in long-term fluorescent imaging of live cells.

Tet-DciA was grown in the presence of hygromycin and ATc overnight and for the duration of the DciA_Mtb replete control movies (S1 Movie). Tet-DciA was grown in the presence of hygromycin and ATc overnight followed by growth in the presence of hygromycin alone for 10.5 hours prior to imaging and for the duration of imaging (S2 Movie). For analysis, all Tet-DciA cells grown in depleting conditions were born at least 24 hours after ATc was removed from the media.

Transmission electron microscopy with thin-sectioning

M. smegmatis was fixed in 2% paraformaldehyde/2.5% glutaraldehyde (Polysciences Inc.) in 100 mM sodium cocadylate buffer, pH 7.2 for 1 hour at room temperature. Samples were washed in sodium cacodylate buffer and postfixed in 1% osmium tetroxide (Polysciences Inc.) for 1 hr. Samples were then rinsed extensively in dH₂0 prior to en bloc staining with 1% aqueous uranyl acetate (Ted Pella Inc.) for 1 hr. Following several rinses in dH₂0, samples were dehydrated in a graded series of ethanol and embedded in Eponate 12 resin (Ted Pella Inc.). Sections of 95 nm were cut with a Leica Ultracut UCT ultramicrotome (Leica Microsystems Inc.), stained with uranyl acetate and lead citrate, and viewed on a JEOL 1200 EX transmission electron microscope (JEOL USA Inc.) equipped with an AMT 8 megapixel digital camera and AMT Image Capture Engine V602 software (Advanced Microscopy Techniques).

[5,6-³H]-thymidine incorporation assays

Assays were carried out similarly to published reports [25,28], with a few modifications. M. smegmatis cells were collected at ODλ₆₀₀ = 0.3, washed once with 7H9 media, and inoculated into fresh 7H9 media containing 1μCi/ml [5,6-³H]-uracil (Perkin Elmer). [5,6-³H]-uracil is converted to [5,6-³H]-thymidine and incorporated into DNA in mycobacteria [25,28].

After 20 and 60 minutes of labeling, 3 ml aliquots of cells were taken for processing to measure counts per minute (cpm) of ³H incorporated into DNA or for enumeration of bacterial counts by colony forming units (cfu). A 7H9 media plus [5,6-³H]-uracil sample was also collected. Aliquots for ³H cpm were treated with 0.3M KOH to hydrolyze RNA and incubated at 37°C for 24 hours. Macromolecules were then precipitated with ice cold 10% trichloroacetic acid (Sigma) and filtered onto glass 25 mm GF/C filters (GE Healthcare). Filters were dried under a heat lamp and submerged in 15 ml of Ultima Gold liquid scintillation fluid (Perkin Elmer). ³H counts per minute (cpm) were measured using a Beckman LS6000IC scintillation counter that was programmed to read each sample for 5 minutes. All presented data indicates the cpm of that sample minus the cpm from the processed media + [5,6-³H]-uracil control to subtract background, relative to the cfu of that sample. The efficiency of RNA hydrolysis by KOH treatment was confirmed by including a KOH-untreated control.

Measuring DNA content by flow cytometry

Cells were fixed and permeabilized as described for fluorescent microscopy [69] with an additional RNaseI_f treatment (NEB), and stained with 100 μM DAPI diluted in water for 15 minutes at room temperature. Cells were then resuspended in PBS, sonicated, and passed through a 30μm filter to remove clumps before flow cytometry analysis. Samples were analyzed using a FACSAria (Becton Dickinson) and data were processed with FlowJo (Treestar). All cells (events) were included in the analysis of DAPI intensity.

Protein purification

Mtb dciA_Mtb, HA-dciA_Mtb, dciA_Mtb^W113A, HA-dciA_Mtb^W113A, truncations of HA-dciA_Mtb and HA-dciA_Mtb^W113A, dnaA, and HA-dnaA were cloned into pGEX-6P (GE Healthcare Life Sciences, S2 and S3 Tables). The plasmids were transformed into BL21(DE3) (Novagen). Mtb DnaB and DnaB-FLAG were amplified from Mtb genomic DNA following the published cloning scheme[35] to exclude the native intein, cloned into pET SUMO (Invitrogen) and transformed into BL21(DE3) cells (Novagen). Rel_Mtb^1-394 was cloned into pET SUMO [70]. Transformed cells were grown to mid-logarithmic phase (ODλ₆₀₀ 0.6–0.8) and induced with 1mM isopropyl β-D-1-thiogalactopyranoside (IPTG) (GoldBio) for 3 hours at 37°C. Cells were harvested by centrifugation and resuspended in lysis buffer supplemented with 1 mg/ml lysozyme and flash frozen. The lysis buffer for DnaB constructs was 50mM Tris-HCl pH8, 300mM NaCl, 5mM imidazole pH8 and 1mM β-mercaptoethanol (BME). The lysis buffer for DnaA was PBS, and for the DciA_Mtb constructs the lysis buffer was 0.25M Urea, 750mM NaCl, 2.7mM KCl, 10mM sodium phosphate dibasic heptahydrate, 2mM potassium phosphate supplemented with .01% Nonidet P-40 (Sigma) and .1% TritonX 100. After thaw, cells were sonicated, treated with 9 units per ml Benzonase (Sigma) and 6mM MgCl₂ when indicated, and spun at 10,000g to clarify the lysate.

For the purification of DnaB and Rel_Mtb^1-394 lysate was incubated with Ni-NTA agarose (Qiagen), washed with 50mM Tris-HCl pH8, 300mM NaCl, 20mM imidazole pH8, and 1mM BME, and proteins were eluted with 50mM Tris-HCl, pH8, 300mM NaCl, 250mM imidazole pH8, and 1mM BME. The His-SUMO tags were cleaved with His-tagged Ulp1 enzyme. A second incubation with Ni-NTA agarose was used to bind His-Ulp1 and the cleaved His-SUMO tag, and un-tagged recombinant protein was collected as flow-through.

For the purification of DnaA and DciA_Mtb constructs, lysate was incubated with Protino Glutathione Agarose 4B (Macherey-Nagel), washed with wash buffer (PBS or DciA_Mtb lysis buffer plus .01% Nonidet P-40), and cleaved on beads with GST-tagged PreScission Protease (GE Healthcare) in 50mM Tris-HCl pH7, 150mM NaCl, 1mM EDTA, and 1mM DTT, with un-tagged recombinant protein collected as flow-through.

Electromobility shift assays

DNA fragments (see S3 Table) containing oriC_Mtb [33] and rrnAPL [19] were amplified by PCR and products were gel-purified using QIAquick column (Qiagen). 250 ng of gel-purified dsDNA or 3x FLAG ssDNA oligo (see S3 Table) were labeled with T4 polynucleotide kinase (NEB) and [γ-³²P]-ATP, and unincorporated [γ-³²P]-ATP was removed using Illustra ProbeQuant G-50 microcolumns (GE Healthcare). 20,000 cpm of labeled probe, 10 μg BSA, and the indicated amounts of DciA_Mtb or DciA_Mtb^W113A were mixed with buffer (50mM Tris-HCl pH7, 150mM NaCl, 1mM EDTA, 1mM DTT) in a total volume of 12μl and incubated for 20 minutes at room temperature. Samples underwent native electrophoresis in 4–20% nondenaturing TBE polyacrylamide gels (Invitrogen). The gels were dried and exposed to film for detection by autoradiography.

Immunoprecipitation and pull-down experiments

For immunoprecipitation from M. smegmatis cell lysates, 1 liter cultures of mid-logarithmic phase M. smegmatis were washed with PBS and frozen at -80°C until processing. Cell pellets were resuspended in 10mls NP-40 Buffer (10 mM sodium phosphate, pH8, 150 mM NaCl, 1% Nonidet P-40, and Roche EDTA-free Complete protease inhibitor cocktail) and lysed using a Constant Systems Cell Disruptor (4 passes at 40k psi), and spun at 50,000g for 30 minutes to generate lysate. When indicated, some lysates were treated with DNase I (NEB) according to manufacturers instructions. 1 ml of lysate was added to 50 μL monoclonal anti-HA agarose (Sigma) and rotated at 4°C overnight. The anti-HA agarose was then washed 3 times with NP-40 buffer and immune complexes were eluted with 500 μg/ml HA peptide (Roche) in IP elution buffer (50 mM Tris-HCl, pH 7.5, 50 mM NaCl with Roche EDTA-free Complete protease inhibitor cocktail).

For pull-down experiments with purified protein, 0.3114 nmol of purified bait protein in 100 μL NP-40 buffer was bound to either 50 μL anti-HA agarose (Sigma) or 40 μL anti-FLAG M2 affinity gel (Sigma) and rotated for 5–6 hours 4°C. The bait-bound matrix was then washed 3 times with NP-40 buffer and twice molar excess (0.6228 nmol) of prey protein was added (unless otherwise indicated) in 275 μL NP-40 buffer and rotated at 4°C overnight. The matrix was then washed three times and immune complexes were eluted with either 500 μg/ml HA peptide or 150 μg/ml FLAG peptide in IP elution buffer as described above.

Antibodies and western blot

For western blot analyses, FLAG-tagged proteins were detected with mouse monoclonal anti-FLAG clone M2 antibody (Sigma) and HA-tagged proteins were detected with mouse monoclonal anti-HA clone HA-7 (Sigma). To detect DciA_Mtb, we used a rabbit polyclonal anti- DciA_Mtb antibody that was generated by Cocalico Biologicals, Inc. by raising rabbit anti-sera against purified DciA_Mtb protein. Western blots were visualized on a ChemiDoc Touch Imaging System (Biorad) as well as film and quantified using ImageLab software version 5.2.1 software (Biorad).

Chromatin immunoprecipitation quantitative PCR (ChIP-qPCR)

50 mL cultures of M. smegmatis expressing either HA-tagged DciA_Mtb (HA-DciA_Mtb), HA-tagged CarD (HA-CarD), or untagged DciA_Mtb (No tag) were grown to an ODλ₆₀₀ of 0.6. Protein-nucleic acid complexes were crosslinked with 2% formaldehyde for 30 min at room temperature, crosslinking was quenched with 0.125M glycine, and cells were lysed in ChIP lysis buffer (50mM HEPES-KOH pH7.5, 140mM NaCl, 1mM EDTA, and 1% TritonX 100, 1X protease inhibitors (Roche), and 2mg/ml lysozyme). After sonication, 0.5 ml of lysate was kept as input fractions and the remaining lysate was rotated with anti-HA agarose (Sigma) overnight at 4°C. Anti-HA agarose was washed 2 times each with ChIP lysis buffer, ChIP lysis buffer plus 360mM additional NaCl, ChIP wash buffer (10mM Tris-HCl pH8, 250mM LiCl, 0.5% NP-40, 0.5% sodium deoxycholate, and 1mM EDTA), and TE buffer (10mM Tris-HCl pH8 and 1mM EDTA). All buffers were supplemented with 1X protease inhibitors. Protein-nucleic acid complexes were eluted from the anti-HA agarose twice with ChIP elution buffer (50mMTris-HCl pH8, 10mM EDTA, 1% sodium dodecyl sulfate, 1X protease inhibitors) for 10 min at 65°C with agitation. Inputs and eluates were incubated at 65°C overnight to reverse crosslinking. Two phenol-chloroform extractions were performed consecutively on inputs and eluates and DNA was precipitated with ethanol and resuspended in TE buffer. Inputs and eluates were diluted to 0.5896 ng/μl and 0.426 ng/μl, respectively. Quantitative PCR (qPCR) was performed with 1 μl of the diluted DNA using primers to amplify three DNA fragments in oriC (oriC1, oriC2, oriC3), a fragment in the promoter of rplN, and a fragment within the sigA coding region (sigAIN) (See S3 Table and S8A Fig).

Biolayer interferometry

Recombinant DnaB and DciA_Mtb were purified from E. coli as described above. An additional dialysis step to remove Tris from the buffer was performed into 150mM NaCl, 10mM NaPO₄ pH8, 1mM BME. DnaB was non-specifically biotinylated in vitro using EZ-Link NHS-PEG₄-Biotin (ThermoFisher). Five molar excess of biotin relative to DnaB was used and incubated for 30 minutes at room temperature. Excess biotin was removed using Zeba Spin Desalting columns (ThermoFisher).

The Octet RED96 System was used to attain biolayer interferometry (BLI) progress curves. Assay buffer for all steps consisted of 150mM NaCl, 0.02% Tween-20, 0.1% bovine serum albumin, and 10mM NaPO₄ pH8. Briefly, streptavidin (SA) biosensor pins (ForteBio) were first equilibrated by being dipped into assay buffer for a 180 second baseline step, and then captured 200nM biotinylated DnaB during a 200 second loading step, followed by a 180 second baseline step in assay buffer. After performing an additional 60 second baseline step in assay buffer, pins were dipped into DciA_Mtb protein samples for a 300 second association step, followed by a 300 second dissociation step in assay buffer. This series of 60 second baseline, 300 second association, and 300 second dissociation steps was performed for each concentration of DciA_Mtb. Curves were corrected by subtracting double reference of both biotin-coated pins dipped into the DciA_Mtb wells and DnaB-coated pins dipped into buffer only. Data was analyzed on ForteBio Data Analysis 6.4. Processed data were fit globally for all concentrations of DciA_Mtb in a 1:1 kinetic binding model.

Mass spectrometry

Bands generated by SDS-PAGE followed by staining with ProteoSilver Plus Silver Stain Kit (Sigma), were cut out and destained according to manufacturer’s instructions, and submitted to the Proteomics & Mass Spectrometry Facility at the Danforth Plant Science Center for trypsin digestion followed by LC-MS/MS analysis.

Statistics

Prism6 (Graphpad Software, Inc.) was used to determine statistical significance of differences. Unpaired two-tailed Student’s t-test was used to compare two groups with similar variances. Unpaired two-tailed Student’s t-test with Welch’s correction was used to compare two groups with different variances. One-way analysis of variance (ANOVA) and Tukey’s multiple comparison test were used to determine significance when more than two groups were compared. When utilized, center values and error bars represent mean ± SEM. * p <0.05, ** p <0.01, *** p<0.001, **** p <0.0001.