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Published in final edited form as: Curr Biol. 2025 Mar 31;35(8):1816–1827.e3. doi: 10.1016/j.cub.2025.03.009

Live tracking of replisomes reveals nutrient-dependent regulation of replication elongation rates in Caulobacter crescentus

Inchara S Adhikashreni 1,#, Asha Mary Joseph 1,3,*,#, Sneha Phadke 1, Anjana Badrinarayanan 1,4,5,*
PMCID: PMC7617702  EMSID: EMS205641  PMID: 40168985

Summary

In bacteria, commitment to genome replication (initiation) is intricately linked to nutrient availability. Whether growth conditions affect other stages of replication beyond initiation remains to be systematically studied. To address this, we assess the replication dynamics of Caulobacter crescentus, a bacterium that undergoes only a single round of replication per cell cycle, by tracking the replisome across various growth phases and nutrient conditions. We find that the replication elongation rates slow down as cells transition from exponential (high-nutrient) to stationary (low-nutrient) phase, and this contributes significantly to the overall cell-cycle delay. Although elongation rates are correlated with growth rates, both properties are differentially influenced by nutrient status. This slowdown in replication progression is reversed via supplementation with dNTPs and is not associated with increased mutagenesis or upregulation of the DNA damage responses. We conclude that growth conditions not only dictate the commitment to replication but also the rates of genome duplication. Such regulation appears to be distinct from stress-induced replication slowdown and likely serves as an adaptive mechanism to cope with fluctuations in nutrient availability in the environment.

Graphical abstract.

Graphical abstract

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Introduction

Chromosome replication is essential for the propagation of life and is carried out by a multi-subunit complex called the replisome.1,2 In bacteria, replication initiates at the origin of replication (“ori”). Two replisomes carry out DNA synthesis bidirectionally along the chromosome arms and replication terminates when the replisomes meet at the terminus (“ter”).3–9 These steps of replication can be used to define the bacterial cell-cycle stages: (1) initiation (B period) entails the time from cell birth until the start of DNA replication. (2) Elongation (C period) is the period of DNA synthesis and (3) division (D period) is the time between completion of replication and cell division.10 DNA replication must be integrated with cell growth and environmental fluctuations to ensure the transmission of a fully replicated copy of the genome at the time of cell division. Although this appears to be a challenging task, given large variations in growth rates and/or nutrient availability, replication is remarkably robust and accurate.11 How bacteria maintain control and coordination of DNA replication and cell-cycle progression across diverse growth conditions is still not completely understood.

Although many bacteria lack dedicated cell-cycle-checkpoint mechanisms (as observed in eukaryotes), the commitment to replication (initiation) is highly regulated via multiple mechanisms.12–14 For example, in fast-growing bacteria such as E. coli, which divide faster than their C period in nutrient rich conditions, replication initiation occurs in preceding rounds of the cell cycle.15 Control of initiation is centered on the replication initiation protein DnaA, and this has been extensively studied, both during steady-state growth in different nutrient conditions as well as in contexts of stress, such as starvation and stationary phases.16–23 Apart from initiation, it is unclear whether other stages of replication are also regulated based on nutrient availability.

Our understanding of replication control was originally derived from population-level experiments from which a systems-level understanding of the intricate relationship between cell size, growth rate, and cell-cycle regulation also emerged.15,24,25 However, precise measurement of elongation rates across diverse growth phases/conditions has been challenging. Earlier studies estimating C period at the population-level in E. coli proposed invariance across varied nutrient conditions. However, later studies showed that this applied to only fast growth conditions, with both C and D periods also increasing as growth rates decreased.10,15,26–32 Paradoxically, most studies that have probed replication timing are limited to bacteria like E. coli and Bacillus subtilis that pose an additional challenge as these systems, depending on growth conditions, initiate single or multiple rounds of replication within one cell cycle.

Such ensemble measurements also mask cell-to-cell variations within the population, which have become evident with more recent single-cell analysis. Apart from recapitulating growth and size laws at the level of single cells, these studies have provided valuable mechanistic insights into regulatory principles and have identified deviations from idealized population averages.11,31,33–35 Incidentally the focus of these experiments has also been restricted to initiation control or to broadly infer an overall cell-cycle timing. It is possible that direct visualization of individual replication cycles and comparison of C periods across nutrient conditions becomes difficult in bacteria that can go through multiple initiation events within the same cell cycle. Indeed, it would be surprising if growth conditions do not impact other stages of replication, as this would imply that elongation would need to progress (at the same rates) to completion regardless of the growth context.

Here, we used Caulobacter crescentus, a bacterium that undergoes only a single round of replication in every cell cycle,36 to precisely and directly measure replication timings. Using single-cell fluorescence time-lapse microscopy, we track the replisome across changing growth phases (from exponential to stationary) and nutrient conditions (rich vs. minimal media). We find that the time between consecutive cell divisions increases as cells transition from exponential (high-nutrient) to stationary-phase (low-nutrient) growth. Apart from shutdown of replication initiation, replication elongation times also increase, significantly contributing to the overall cell-cycle delay. We observe a strong correlation between replication elongation rates and cellular growth rates, with both rates changing differentially in response to nutrient availability. Consistently, shifting cells from low-nutrient to high-nutrient conditions results in faster progression of replication elongation and increased cellular growth rates. We find that the nutrient-dependent slowdown of replication as cells enter stationary phase is not dependent on starvation-inducible nucleotide (p)ppGpp. Instead, we show that dNTPs (deoxynucleoside triphosphate) can revert this slowdown. Such alterations in replication rates do not result in increased mutagenesis or upregulation of DNA damage responses, which are typically seen much later in stationary phase. Together, our study provides the first direct measure of replication progression as cells transition from exponential to stationary phase. Using this paradigm, we uncover a relationship between replication speeds and nutrient availability, highlighting that the growth environment not only governs the commitment to replication but also the rates of genome duplication.

Results

Measurement of Caulobacter cell-cycle stages via single-cell tracking of replisome dynamics

To track replication dynamics in real time, we imaged the β-clamp subunit of the replisome (DnaN) fused to YFP (yellow fluorescent protein), expressed from its endogenous locus. This strategy has been used previously to track replisome mobility inside single cells.8,9,34,37–39 As reported earlier, we observed no localizations (fluorescent focus) of DnaN (diffuse cytoplasmic signal) in new-born Caulobacter cells.6,9 Following this, a single fluorescent focus of DnaN was observed at a cell pole (where the ori is located), indicative of replication initiation. As replication progressed, the DnaN fluorescent focus moved toward the opposite pole and, on completion of replication, dissociated around mid-cell where the ter is located. Such dissociation of DnaN fluorescent focus is indicative of replication completion and is followed by successful cell division (Figures 1A and S1A–S1C). Though replication is mediated simultaneously by two replisomes on both chromosome arms, due to chromosome inter-arm alignment, only a single fluorescent focus was visualized during most part of the replication cycle.9

Figure 1. Measurement of Caulobacter cell-cycle stages via single-cell tracking of replisome dynamics.

Figure 1

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(A) (Top) Representative montage showing progression of replication during Caulobacter cell cycle. The montage captures localization of DnaN-YFP (appearing as a fluorescent focus) at different points of the cell cycle for a cell growing in exponential phase. Time (mins) from the start of the cell cycle is indicated on top left corner of each frame in the montage. Initiation and completion of replication are denoted with arrow and asterisk, respectively (scale bar, 2 μm). The cell whose cell cycle is tracked is outlined with yellow dashed line. (Bottom) Schematic showing progression of replication during Caulobacter cell cycle. Red dots indicate the representative spatial positioning of replisomes along the duration of cell cycle. Duration of each stage of replication (B, C, and D periods) within one cell cycle are also depicted. B period is defined as the time to form a fluorescent DnaN focus after cell birth. C period is defined as the time period for which DnaN remains localized and is mobile in the cell. D period is defined as the time between dissociation of the DnaN fluorescent focus and cell division. “Interdivision” time or “cell-cycle” time is the total time taken between two cell divisions (B+C+D periods).

(B) Growth curve of Caulobacter crescentus batch cultures growing in PYE at 30 C. The graph shows mean with SD for optical density measurements (OD600) from three independent experiments. The colors denote the growth phases I–mid-exponential, OD600 – 0.2–0.4; II—transition from exponential to stationary phase, OD600 0.8–1.4; and III—early stationary phase, OD600 - 1.4 for a period of 4 h that were characterized for replication dynamics in this study.

(C) Interdivision time of bacteria from the three growth phases indicated in 1B. Individual dots in the scatterplot show data from single cells, and the boxplots show median with interquartile ranges for each distribution here and in similar graphs elsewhere. The bold line on some of the boxplots denotes median overlapping with one of the interquartile ranges here and in similar instances elsewhere. n ≥ 292 cells pooled from three independent experiments.

See also Figure S1 and Tables S1 and S2.

Because only a single round of replication is initiated in each cell cycle, by tracking the formation, progression, and dissociation of the replisome focus, we can accurately measure each stage of replication—the time to form a fluorescent DnaN focus after cell birth (B period), the time period for which DnaN remains localized and is mobile in the cell (C period), and the time between dissociation of the DnaN fluorescent focus and cell division (D period). We defined “interdivision” time or “cell cycle” time as the total time taken between two cell divisions (B+C+D periods) (Figures 1A and S1A–S1C). For example, in exponentially growing cells (OD600 0.2–0.4), a B period of 4.6 ± 4 min, C period of 69.8 ± 7.4 min, and D period of 5.8 ± 4.6 min correspond to an interdivision time of 80.2 ± 8.8 min.

We employed this strategy to track cell-cycle timing across three distinct phases across the growth curve, with active replication (as deduced by the presence of DnaN fluorescent foci) and representing progressively reducing nutrient availability: phase I—mid-exponential (OD600 – 0.2–0.4), phase II—transition from exponential to stationary phase (OD600 0.8–1.4), and phase III—early stationary phase (OD600 – 1.4 for a period of 4 h) (Figures 1B, S1A–S1C, and S2A–S2B). For this, cells from appropriate growth phases (in batch culture) were imaged via time-lapse microscopy (see STAR Methods section for details about the imaging setup). We restricted our analysis to single cells where a complete cell cycle was observed (from cell birth until division) to ensure that the B, C, and D periods could be estimated. Furthermore, we only analyzed the cell cycles of stalked cells (longer cells during asymmetric Caulobacter cell divisions) for accurate estimation of the B period from a single cell type. Measurement of interdivision times for these phases indicated that cells slowed down their cell cycle significantly upon entry to stationary phase (interdivision time of 132.4 ± 16.2 vs. 80.2 ± 8.8 min in early stationary [III] and exponential [I] phases, respectively) (Figures 1C and S1A–S1C).

Entry into stationary phase is associated with slowdown in replication elongation rates

Slowdown in overall cell-cycle timing could be due to a delay in B, C, or D periods. For example, previous studies have shown that Caulobacter cells shut down replication initiation in stationary phase via regulating DnaA levels.21 However, these experiments were conducted much later in stationary phase (approx. 24 h after reaching OD600 – 1.4), and it is unclear whether initiation shutdown alone can explain an overall slowdown in the cell cycle (as cells still commit to replication at OD600 – 1.4). Two observations from our data suggest an additional layer of replication regulation likely acting on the C period as well: (1) after entry into stationary phase (OD600 – 1.4), for a period of 6 h (∼4 generation times of exponential-phase growth), ∼40% of the population harbored replisome foci (DnaN-mCherry), and (2) complete replication shutdown (when <5% of the population harbored replisome foci) occurred only much later in stationary phase (24 h after reaching OD600 – 1.4) (Figure S2A). To corroborate these observations, we assessed the localization of another replisome marker, single-stranded DNA-binding protein (SSB). We found that SSB and DnaN were co-localized across all three growth phases (Figure S2C).

We next measured the timing of each replication stage across the three growth phases to assess the contribution of the same toward the cell-cycle time. We found that the B, C, and D periods were comparable for phase I and phase II cells—B period was found to be 4.6 ± 4 and 6.7 ± 4.1 min, C period 69.8 ± 7.4 and 76.1 ± 7.1 min, and D period 5.8 ± 4.6 and 5.5 ± 4.3 min, respectively, for phases I and II (Figures 2A–2C). This is consistent with the observation that interdivision times also did not vary much between these two phases of growth (Figure 1C). In contrast, upon entry into stationary phase (phase III), cells significantly slowed down all three replication stages, with a B, C, and D period of 17.7 ± 9, 103.7 ± 12.9, and 11 ± 6 min, respectively (Figures 2A–2C).

Figure 2. Entry into stationary phase is associated with slowdown in replication elongation rates.

Figure 2

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(A–C) B, C, and D periods of bacteria from the three growth phases indicated in Figure 1B.

(D–F) B, C, and D periods of individual bacteria plotted as a function of interdivision time across growth phases I, II, and III. Pearson’s correlation coefficients (r) with CI for B (0.40 [0.30 0.49] for phase I, 0.49 [0.40 0.57] for phase II, and 0.51 [0.41 0.60] for phase III), C (0.73 [0.67 0.78] for phase I, 0.75 [0.69 0.79] for phase II, and 0.79 [0.73 0.83] for phase III), and D (0.39 [0.29 0.48] for phase I, 0.37 [0.27 0.46] for phase II, and 0.26 [0.13 0.37] for phase III) periods against interdivision time in each growth phase are denoted in respective colors. n ≥ 292 cells pooled from three independent experiments for graphs in (A)–(F).

See also Figure S2 and Tables S1 and S2.

Across the three growth phases, we observed a weak correlation between interdivision time and B/D periods. For B period, the Pearson’s correlation coefficient (r) was 0.40 with confidence interval CI [0.30 0.49] in phase I and this changed to r 0.51 CI [0.41 0.6] in phase III. Similarly, for D period, the Pearson’s correlation coefficient (r) was 0.39 with CI [0.29 0.48] in phase I and this changed to (r) 0.26 with CI [0.13 0.37] in phase III. On the other hand, across all growth phases, the strongest relationship was observed between interdivision time and C period (r 0.73 with CI [0.67 0.78], r 0.75 [0.69 0.79], and r 0.79 [0.73 0.83], respectively, in growth phases I, II, and III) (Figures 2D–2F).

Replication slowdown is associated with slower growth rates

We next asked whether growth rates also slowed down as replication elongation times increased. This was based on the observations that, despite variations in interdivision/cell-cycle timings, the average length of cells at division was comparable across the three growth phases. We saw only a modest increase in cell length in growth phases II and III in comparison with growth phase I (mean cell lengths at division being 4.04 ± 0.41, 4.31 ± 0.39, and 4.3 ± 0.37 μm for growth phases I, II, and III, respectively) (Figure 5A). Thus, we estimated single-cell growth rates (per cell division) for all three growth phases (Figure 3A). We found that growth rates did reduce from 0.023 ± 0.004 μm/min in exponential-phase cells to 0.015 ± 0.002 μm/min in stationary-phase cells (Figure 3B).

Figure 5. Replication slowdown is not associated with induction of the DNA damage response or increased mutagenesis.

Figure 5

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(A) Pre-divisional cell lengths of bacteria from growth phases I–III and post-media shift (associated with Figures S4 and 4). As a positive control, cell lengths of cells treated with 0.5 μg/mL DNA damaging agent mitomycin C (MMC) for 2 h are provided, which show filamentation under damage stress. n ≥ 292 cells pooled from three independent experiments.

(B) Expression of yfp from an SOS-inducible promoter (PsidA). Total fluorescence intensity normalized to cell area is plotted for bacteria from growth phases I–III. n ≥ 300 cells pooled from two independent experiments. As a positive control, PsidA-yfp intensity from cells treated with DNA damaging agent (MMC) are provided to show that sidA promoter is induced upon DNA damage stress.

(C) Intensity of SSB-YFP foci normalized to total fluorescence intensity within the cell for bacteria in growth phases I – III. n ≥ 216 cells from two independent experiments.

(D) Frequency of rifampicin-resistant colonies in wild-type (WT) and ΔmutL cultures from growth phases I – III. Each dot represents frequency calculated from an independent experiment and the black lines indicate medians.

See also Tables S1 and S2.

Figure 3. Replication slowdown is associated with slower growth rates.

Figure 3

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(A) Schematic showing growth rate calculation from single cells.

(B) Growth rate of single cells from different growth phases.

(C) C periods of single cells from growth phases I, II, and III plotted as a function of growth rate. Pearson’s correlation coefficient (r) with CI for C period against growth rate (−0.76 [−0.77 −0.71]) is indicated in black.

(D) Replication elongation rates of single cells from growth phases I, II, and III plotted as a function of growth rate. Pearson’s correlation coefficient (r) with CI for replication elongation rate against growth rate (0.74 [0.70 0.77]) is indicated in black. n ≥ 292 cells pooled from three independent experiments for graphs in (A)–(D).

See also Tables S1 and S2.

We next compared the cell growth rates with their respective C periods across the three growth phases. We observed a striking relationship between growth rates and replication elongation rates across these populations (r 0.74 [0.70 0.77]), with faster-growing cells having a faster replication elongation rate (Figures 3C–3D). We did note that B period additionally showed a weak correlation with growth rate (r −0.60 [−0.64 −0.56] and r −0.35 [−0.41 −0.30] for B and D periods, respectively, Table S1). Cells with a slower growth rate also tended to take longer to initiate new rounds of replication and, in some cases, divided slower after the completion of replication. However, we did not observe any of the individual periods co-varying with each other, suggesting that they are likely regulated independently and/or by distinct mechanisms (Table S1).

Impact of nutrient status on replication progression

We wondered what factors contributed to the slowdown in replication elongation. Transition from exponential to stationary phase is characterized by alterations in cellular physiology due to shifts in nutrient availability and metabolism, which can potentially impact growth dynamics of bacteria.40–42 We thus asked whether variation in nutrient status in the growth medium would reflect in variation in replication elongation rates. For this, we compared replication progression in exponentially growing 1820 Current Biology 35, 1816–1827, April 21, 2025 steady-state cells across three standard laboratory media used for Caulobacter growth: PYE (peptone yeast extract; rich, complex media) and minimal salts media supplemented with two carbon sources (M2G—with 0.2% glucose and M2X—with 0.2% xylose) (Figure S3A). We observed that cells grown in M2G or M2X had significantly longer C periods when compared with PYE, with M2X cells showing the slowest replication elongation rates (976 ± 101, 751 ± 75, and 722 ± 73 bp/s for PYE, M2G, and M2X, respectively) (Figures S3B and S3C). Consistently, measurement of interdivision time revealed longer cell-cycle duration in minimal media conditions (Figure S3D) and the correlation between C period and growth rates was also recapitulated (r −0.78 [−0.81 −0.76]) (Figure S3B). As in the case of different growth phases, interdivision time displayed the strongest relationship with the C period compared with the B or D periods (Figures S3E and S3F; r 0.53 [0.45 0.61], r 0.81 [0.77 0.85], and r 0.29 [0.18 0.39] for B, C, and D periods, respectively, in M2G and r 0.58 [0.50 0.65], r 0.80 [0.76 0.84], and r 0.15 [0.03 0.25] for B, C, and D periods, respectively, in M2X).

To directly assess the impact of nutrient availability on replication elongation rates, we designed an experiment to supplement stationary-phase cells with additional nutrients in the growth media. For this, we shifted cells growing in stationary phase (OD600 – 1.4) into growth media extracted from exponentially growing cells (OD600 − 0.2) and assessed the timing of the very first cell cycle of these “media-shifted” cells in their new growth environment (Figure S4A).

We observed an immediate reduction in the C period of these media-shifted cells (103.7 ± 12.9 and 85.3 ± 12 min in stationary and media-shifted cells, respectively) (Figures 4A and S4B). This was also reflected in the interdivision time of these media-shifted cells from 132.4 ± 16.2 min in stationary phase to 102.6 ± 16.2 min in media-shifted conditions (Figures 4B and S4B). We did also observe an increase in growth rates—but not to the same extent as seen for the change in interdivision time or C period (growth rate of 0.015 ± 0.002 and 0.017 ± 0.004 μm/min in stationary-phase and media-shifted cells, respectively) (Figures 4C and 4D). This decoupling was also reflected in the cell lengths, with media-shifted cells having a modestly smaller mean cell length when compared with stationary-phase cells (4.08 ± 0.37 and 4.3 ± 0.37 μm, respectively) (Figure 5A). These observations suggest that although there is concurrence in replication rates and growth rates in steady-state conditions, their regulation may occur independently. This is also consistent with observations in cells transitioning from exponential phase toward stationary phase. In this case we found that the proportion of cells initiating replication already started to decline from mid-exponential phase even though growth rates remained unchanged until the transition from exponential to stationary phase (Figures 3B and S2A).

Figure 4. Impact of nutrient status and dNTP on replication progression.

Figure 4

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(A) C periods of bacteria from growth phases I and III compared with those shifted from growth phase III to media from growth phase I (MS—media shift). Data for growth phases I and III have been reproduced from Figure 2B for comparison.

(B) Interdivision time of bacteria from growth phases I and III compared with those shifted from growth phase III to media from growth phase I (MS). Data for growth phases I and III have been reproduced from Figure 1C for comparison.

(C) Growth rate of single cells from growth phases I and III compared with those shifted from growth phase III to media from growth phase I (MS). Data for growth phases I and III have been reproduced from Figure 3B for comparison.

(D) C periods of bacteria shifted from growth phase III to media from growth phase I (MS), plotted as a function of growth rate. Data for growth phases I–III have been reproduced from Figure 3C for comparison. Pearson’s correlation coefficient (r) with CI for C period against growth rate (−0.68 [−0.71 −0.65]) is indicated in black. n ≥ 300 cells pooled from three independent experiments for the media shift experiment.

(E) C periods of bacteria from growth phase I,with and without treatment with 3 mM HU.

(F) Growth rate of bacteria from growth phase I, with and without treatment with 3 mM HU. n ≥ 100 cells pooled from three independent experiments for (E) and (F).

(G) C periods of bacteria from growth phases I and III, with and without treatment with 100 mM dNTPs.

(H) Interdivision time of bacteria from growth phase III, with and without treatment with 100 mM dNTPs. n ≥ 150 cells pooled from three independent experiments for (G) and (H).

See also Figures S3 and S4 and Tables S1 and S2.

To test whether this is indeed the case, we treated cells with a sub-lethal dose of hydroxyurea (HU), an inhibitor of ribonucleotide reductase (RNR) activity. We hypothesized that HU treatment should result in an immediate slowdown of replication progression, without a significant impact on growth rates. For this, we grew cells on agarose pads containing 3 mM HU and analyzed cells where a complete cell cycle (from cell birth until cell division) was trackable. We found that the C period significantly increased in these cells (67.8 ± 6.3 min in wild-type exponential phase to 84.1± 11.6 min in HU-treated cells) (Figure 4E). In addition to the prolonged C period, we also found that the interdivision times increased significantly in HU-treated cells (Figure S4C). However, the growth rates remained similar between HU-treated and untreated conditions (Figure 4F). The prolonged interdivision time in combination with the unchanged growth rates was reflected in cell length distributions, with HU-treated cells having a longer average cell length in comparison with wild-type cells (Figure S4D). These observations suggest that replication elongation rates (and interdivision times) can vary independent of changes in growth rates.

Replication elongation rates change in the presence of dNTPs

Together, our observations are consistent with replication speed variation in a nutrient-dependent manner. An important factor in mediating replication regulation is the alarmone (p)ppGpp. (p) ppGpp levels have been shown to influence replication initiation commitment as well as replication elongation rates in B. subtilis and E. coli.43–49 We hence assessed the role of (p)ppGpp in the observed replication elongation slowdown in our conditions via tracking replication in cells deleted for the bifunctional enzyme SpoT/Rel that is essential for ppGpp synthesis as well as degra-dation. Similar to observations in wild-type cells, we found that the replication elongation rates still decreased significantly as DΔpoT cells entered stationary phase (OD600 – 1.4) (Figure S4E), suggesting that (p)ppGpp does not regulate the slowdown in replication elongation rates.

Another source of variation could be via a change in the levels of precursors required for replication elongation. We thus asked whether directly supplementing stationary-phase cells with dNTPs in the stationary-phase growth media would rescue the observed delay in the C period. For this, we first confirmed that supplementation of the growth media with 100 mM dNTP could mitigate a HU-associated growth phenotype. Our observations were consistent with a previous report showing that dNTPs are taken up in Caulobacter cells directly from the growth media and that this could rescue a HU-induced growth defect50 (Figure S4F). Based on these findings, we next evaluated the impact of dNTP supplementation on replication times for cells growing in exponential (OD600 − 0.2) and stationary (OD600 − 1.4) phases. We found that dNTP supplementation significantly affected the C period of cells in stationary phase, with the replication elongation time reducing from 131.5 ± 24.5 min without dNTP supplementation to 98.1 ± 17.2 min upon dNTP addition (Figure 4G). In contrast, replication elongation times in exponential phase did not change upon dNTP supplementation, consistent with this being a nutrient-replete growth environment. Interestingly along with a faster C period, we also noted that the interdivision time of stationary-phase cells supplemented with dNTPs also reduced, without a change in the B or D periods of these cells (Figures 4H, S4G, and S4H). These data indicate that replication elongation rates may be directly influenced by dNTP availability, likely determined by the nutrient status of the growth environment.

Replication slowdown is not associated with induction of the DNA damage response or increased mutagenesis

Could this slowdown reflect a perturbed replication state, resulting in DNA damage or increased mutagenesis? Interestingly, both of these features have been observed during late stationary-phase growth,51,52 at time points where we do not observe replisome foci in Caulobacter cells (Figure S2A). Three pieces of evidence suggested this may not be the case as cells transition into stationary-phase growth. (1) A hallmark of DNA damage stress is cell filamentation due to cell division inhibition.53–55 In contrast, we did not observe cell filamentation, with cell lengths remaining similar across the three growth phases (Figure 5A). (2) In support of this, SOS response induction was not observed, via assessment of yfp expression from the DNA-damage-responsive promoter PsidA (Figure 5B). Under DNA damage, the extent of PsidA induction as well as cell filamentation was much more pronounced (Figures 5A and 5B). (3) Replication-blocking lesions would reveal long stretches of single-stranded DNA (ssDNA) due to continued unwinding of double-stranded DNA by the helicase despite replisome stalling.56 Our measurement of single-stranded DNA-binding protein (SSB) fluorescent focus intensities did not show a significant increase in cells where replication elongation rate decrease was observed (Figure 5C). On the contrary, we observed a modest decrease in SSB fluorescent focus intensities in growth phase III, suggesting a concomitant slowing down of the entire replisome (including the helicase).

A second key feature of perturbed replication is an increase in stress-induced mutagenesis. Indeed, such mutagenesis mediated via error-prone polymerases is reported to occur in the context of late stationary-phase cells (grown beyond 48 h).57–59 To assess whether the replication slowdown observed in the conditions tested here is associated with an increase in mutagenesis, we quantified frequency of rifampicin-resistant mutants from different phases of growth. We observed similar mutant frequencies across all growth phases tested for wild-type cells. Furthermore, an ∼100-fold increase in mutation frequencies was observed when mismatch repair was lacking (ΔmutL cells), and this, too, remained invariant across all growth phases (Figure 5D).

Together, these data indicate that replication slowdown observed as cells enter stationary phase is not associated with the SOS response and does not result in increased mutagenesis. Hence, replication elongation rates are influenced by the growth environment in a manner that is distinct from replication pertur-bation under conditions of stress.

Discussion

Response of individual bacteria to fluctuations in the growth environment (such as nutrient availability) can collectively shape the fitness of bacterial populations and diversity within microbial communities.60,61 The most frequent responses to many environmental fluctuations include alterations in bacterial metabolism, growth, and cell cycle; whereas resource abundance results in fast growth and division, resource depletion leads to slowing down or inhibition of both.40,61,62

The transition between fast and slow growth modes as observed within natural niches can be recapitulated to some extent in batch cultures of bacteria growing under optimal laboratory conditions. As resource depletion and other stresses set in, the exponentially multiplying bacterial population enters stationary phase, where bacterial physiology changes and the viable bacterial count in the population starts to stagnate.41,42,63 In fact, in nature, bacteria are thought to go through similar cycles of feast and famine, with a large proportion of bacteria being in physiological states comparable with stationary phase.41,63,64 The progressive deterioration in growth conditions during exponential to stationary phase transition provides an excellent window to assess temporal regulation of cellular processes during transitions between growth modes.

Indeed, during transitions between growth modes, one of the most critical processes to be regulated is DNA replication. To ensure a complete duplicated copy of the genome prior to every division, it is imperative that replication is coordinated with growth and cell division. In our current work, we systematically characterized replication dynamics in single Caulobacter cells across different growth contexts, encompassing bacterial growth phases as well as media conditions. Our findings directly demonstrate, at the level of single cells, the regulation of replication elongation rates on transition from the exponential to the stationary phase, which is also recapitulated under media conditions of varied richness. This work has revealed a clear link between nutrient status, bacterial growth, and rate of genome duplication. We consider a few possible mechanisms driving this regulation of replication elongation rates below.

Nutrient-dependent regulation of replication

Although the speed of DNA synthesis is inherent to the polymerase, multiple factors can affect the processivity of the holoenzyme. One consideration is the role of the alarmone (p)ppGpp, which mediates stringent response in bacteria. It has been observed that induction of the stringent response affects not only replication initiation44,46–48,65 but also elongation rates.43,45,49 In Bacillus subtilis and E. coli, (p)ppGpp inhibits the activity of primase DnaG, either by binding to it or lowering levels of GTP. However, (p)ppGpp production and the associated stringent response are generally triggered under severe carbon and amino acid deprivation.43,45,48,49 Our observations in ΔspoT cells suggest that (p)ppGpp may not contribute to the observed slowdown in replication elongation times in early stationary phase. We did note that ΔspoT cells tended to shut down the commitment to replication in stationary phase slower than wild-type cells, with 63 ± 3% and 77 ± 4% cells having replication foci in stationary phase for wild-type and ΔspoT cells, respectively. Such a misregulation of replication initiation is consistent with previous reports in Caulobacter implicating a role for (p)ppGpp in regulating replication initiation.46,48

Our data instead suggest that availability of precursors for DNA synthesis can directly influence elongation rates without impacting initiation (B period). Consistently, replication elongation rates in cells treated with HU also show this to be the case, without concomitant alterations in growth rates. Indeed, earlier studies in E. coli have demonstrated that titrating the expression of RNR enzymes can lead to alterations in dNTP pools and a substantial reduction in the C period duration.66,67 At the level of transcriptional control of RNR genes involved in dNTP biosynthesis, it is interesting to note that DnaA has been implicated as a regulator.68 However, we did not observe significant differences in transcript levels of RNR genes involved in dNTP biosynthesis (nrdE, nrdB, nrdJ, or CCNA_01420) between Caulobacter in the exponential phase (phase I) and during the onset of the stationary phase (phase III) (RNA sequencing [RNA-seq] experiment, data not shown). Alternatively, reduction in dNTP levels could simply be a consequence of resource depletion as cells enter stationary phase. It would be important now to understand whether there are active mechanisms by which dNTP levels are regulated in stationary phase and how the variation in dNTP levels in turn affects replication elongation rates.

Going fast and slow

Although commitment to replication (initiation) is an important and highly regulated event in the bacterial cell cycle, our observations suggest that initiation regulation may not be the only determinant of overall cell-cycle timing in Caulobacter (as B period is not always strongly correlated with interdivision time). Instead, it appears that replication dynamics are tuned at the stage of replication elongation as well. This modulation is independent of cell growth rates and appears to contribute to the overall cell-cycle timing. Such replication speed variation in single Caulobacter cells is also consistent with observations made in E. coli and B. subtilis, underscoring the conserved nature of this plasticity.

Variations in replication elongation rates can have important consequences. On the one hand, given the trade-off between speed and accuracy, it can be anticipated that slow replisomes may have lower mutation rates.69–71 On the other hand, slow replisomes can result in generation of longer stretches of ssDNA between the stalled replisome and the helicase unwinding the double-stranded DNA ahead of it. This could trigger induction of the SOS response, which results in the expression of error-prone polymerases, contributing to an increase in mutagenesis. For example, treatment of cells with HU, which depletes cellular dNTP pools, can lead to generation of ssDNA tracts that elicit an SOS response.56 Conversely, SOS response itself can also be a source of replication slowdown. For instance, studies have shown that cells encounter oxidative stress in late stationary phase. This can result in DNA damage and expression of error-prone polymerases in turn impacting replication progression.72 Our results show that the replication slowdown we observe is neither a source of DNA damage and associated mutagenesis nor is this phenomenon a consequence of damage.

Instead, our observations suggest that there is a “Goldilocks” zone of replication rates with a large range (660 ± 80 to 976 ± 101 bp/s in present study) in which variation is tolerated without significant impact on mutagenesis. Furthermore, these rates are also readily reversible (dependent on nutrient availability), indicating the plasticity of this process. It is possible that there are tolerable levels of environmental fluctuations within which such plasticity is observed and, beyond a threshold, fluctuations could be associated with more permanent changes. Thus, although replication slowdown mediated by genotoxic stress is often associated with activation of stress responses and stress-induced mutagenesis,56,73 the replisome speed variation that occurs in response to nutrient availability (such as observed here) is likely a robust adaptation to fluctuating environmental conditions.

We speculate that such changes in replication elongation rates might be essential to coordinate replication with cell growth as well as other processes that rely on metabolic states. This could also contribute to a prolonged stationary phase (observed here) where replication is still active, prior to a complete shutdown. Such a slowdown could enable cells to rapidly return to fast growth in case nutrients become readily available in the environment. In human cell line studies, it was observed that the inability of replication forks to sense cellular metabolic states and slow down replication resulted in a loss of genome integrity, high-lighting the importance of modulating replication elongation rates.74 Similarly in bacteria, nutrient-dependent regulation of replication rates could be a strategy to cope with transient fluctuations in natural niches without requiring the induction of stress responses or mutagenesis. The mechanisms that regulate replication rates and/or the threshold of nutrient fluctuations beyond which stress responses and mutagenesis are induced are exciting and important open questions.

Resource Availability

Lead contact

Requests for further information and resources should be directed to and will be fulfilled by the lead contact, Anjana Badrinarayanan (anjana@ncbs.res.in).

Materials availability

All reagents generated in this study are available from the lead contact without restriction.

Star ⋆ Methods

Key Resources Table

REAGENT or RESOURCE SOURCE IDENTIFIER
Bacterial and virus strains
NABC12 - dnaN-mCherry::kanR This study N/A
NABC124 – dnaN-YFP::specR This study N/A
NABC247 - Pxyl-ssb-GFP::specR; dnaN-mCherry::kanR Joseph et al.37 N/A
NABC508 – Pxyl-ssb-YFP::kanR This study N/A
NABC581 - PsidA-YFP::kanR Joseph et al.75 N/A
NABC580 - ΔmutL Joseph et al.75 N/A
NABC967 - ΔspoT, dnaN-YFP::specR This study N/A
Chemicals, peptides, and recombinant proteins
Hydroxyurea (HU) Sigma Product number H8627
Deoxynucleotide (dNTP) solution mix New England Biolabs Catalog # N0447L
Mitomycin C (MMC) AG Scientific Catalog #M-1108
GTG Agarose NuSieve Product code 182178
Rifampicin Sigma Catalog # R3501
Kanamycin Sigma Catalog #60615
Spectinomycin Sigma Catalog # PHR1441
Oligonucleotides
dnaN forward oligo - TTATCATATGAAGCTTACGATCGAACGGGCGGCG This study N/A
dnaN reverse oligo - TTATGAATTCCGGACCCGCAGCGGCATCAGCAC This study N/A
Recombinant DNA
pNABC198 Joseph et al.37 N/A
pNABC419 Joseph et al.37 N/A
pNABC1001 This study N/A
pCHYC2 Thanbichler et al.76 N/A
Software and algorithms
MATLAB R2020a MathWorks RRID:SCR_002285
Oufti Paintdakhi et al.77 RRID:SCR_016244
Fiji (ImageJ) Schindelin et al.78 RRID:SCR_002285
MicrobeJ (ImageJ plugin) Ducret et al.79 RRID:SCR_023914
RStudio (R 4.3.3) Rtools 4.3 RRID:SCR_000432
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Experimental Model and Study Participant Details

Bacterial strains

All experiments were performed on bacterial strains derived from Caulobacter crescentus NA1000. All bacterial strains and plasmids used in this study are available in key resources table. Strains and plasmids generated in this study are detailed below.

For construction of pNABC1001 plasmid, full length dnaN was amplified using dnaN forward and reverse oligos, digested with Nde1/EcoR1 and ligated to Nde1/EcoR1 digested pCHYC2. dnaN and mCherry in this construct was separated by 36 bp resulting in a 12 amino acid linker in the fusion protein.

For construction of NABC12 (dnaN-mCherry::kanR), CB15N was transformed with pNABC1001 to integrate dnaN-mCherry linked to kanR at the endogenous locus.

For construction of NABC124 (dnaN-YFP::specR) strain, CB15N was transformed with pNABC198 plasmid to integrate dnaN-YFP linked to specR at the endogenous locus.

For construction of NABC508 (Pxyl-ssb-YFP::kanR), CB15N was transformed with pNABC419 plasmid to integrate ssb-YFP linked to kanR under the Pxyl promoter at xyl locus.

For construction of NABC967, CB15N ΔspoT strain21 was transformed with pNABC198 plasmid to integrate dnaN-YFP linked to specR at the endogenous locus.

Media and growth conditions

Caulobacter crescentus strains were routinely grown in PYE (0.2% peptone, 0.1% yeast extract and 0.06% MgSO4) supplemented with antibiotics as required. While growing strain expressing fluorescently-tagged SSB under Pxyl, 0.3% xylose was added to the culture 3 hrs before the experiment commenced. For experiments in minimal media bacteria were grown in M2 minimal media (1X M2 salts - 0.087% Na2HPO4, 0.53% KH2PO4, and 0.05% NH4Cl, 0.01 mM FeSO4 and 0.01 mM CaCl2) supplemented with either 0.2% glucose (M2G) or 0.2% xylose (M2X). For imaging experiments with HU or dNTPs, cultures at appropriate OD600 values were pre-treated with 3 mM HU or 100 mM dNTPs for 30 min prior to spotting on pads containing the same concentrations of either of them. All experiments were performed in PYE unless otherwise indicated.

Method Details

Growth curve measurements

For growth curve measurements, overnight cultures were back diluted to OD600 – 0.025 (PYE) or 0.05 (M2G and M2X). OD600 measurements were recorded manually every half an hour using Hitachi UH5300 Spectrophotometer. The measurements were carried out in at least three independent cultures under each growth/media regime. For growth curve, mean with SD from three independent experiments were plotted.

Fluorescence microscopy—sample preparation and imaging

For time course microscopy, aliquots were taken from cultures growing at specified conditions at different time points, pelleted down and resuspended in appropriate volumes of growth medium. 2 μl of the cell suspension was spotted on 1% agarose pads (prepared in water) with approximate dimensions of 0.5 X 0.5 cm.

For time lapse microscopy, two independent cultures were grown to the same optical density – one for imaging and the other for extracting spent media. The culture used for extraction of spent media was grown to required OD600 approximately 3 hrs ahead of the culture used for imaging. This culture was then filtered with a 0.22 μm membrane filter and the filtrate (spent media) was used to prepare 1.5% GTG agarose pads. 3 mM HU or 100 mM dNTPs was added to the pads for experiments where HU or dNTP treatments were done (in Figures 4 and S4). At specified OD600 an aliquot taken from the culture to be imaged was pelleted down, concentrated in their respective spent media. 2 μl of the cell suspension was spotted on the pads made from their respective spent media (of similar dimensions as mentioned above), grown inside an OkoLab incubation chamber maintained at 30°C and imaged at 5 min intervals for a duration of 6 hrs.

Imaging was done on Nikon Eclipse Ti2 wide-field microscope equipped with a motorized XY stage, using a 60X plan apochromat objective (NA 1.4) and illumination from pE4000 light source (CoolLED). Image capture was performed with Hamamatsu Orca Flash 4 camera using Nikon’s NIS Elements software. Infrared-based Perfect Focusing System ensured maintenance of focus during time lapse imaging. Fluorescence imaging of GFP, YFP and mCherry were done with excitation at 470 nm, 490 nm and 550 nm respectively, for exposure times of 400 ms.

Mutagenesis assay

Overnight cultures of WT or ΔmutL strains were back diluted to OD600 of 0.1. Aliquots were drawn from these cultures at OD600 – 0.4, 0.8 and 1.4, and plated on 100 μg/ml rifampicin plate. At each of these time points a separate aliquot was drawn to calculate viable counts in the volume of culture plated on rifampicin plate. Colonies were counted after 48 hrs of incubation at 30°C. Rif mutant frequencies were calculated by normalizing Rif resistant colonies to viable counts in each culture.

Survival assay

Cultures were grown in PYE to OD600 - 0.3. Serial dilutions were made in 10 fold increments and 5 μl of each dilution (10−1 to 10−8) were spotted on PYE agar (1.5%) with or without 4 mM HU, supplemented with 0 or 100 mM dNTPs. These plates were incubated at 30°C for 48 hours and images were captured. Growth was assessed by comparing the dilution factor of the last visible spot in each treatment regime with no treatment control.

Quantification and Statistical Analysis

Image analysis

Percentage of cells with DnaN foci from time course microscopy was manually analysed using CellCounter plugin on ImageJ. For calculating interdivision time, B, C and D periods from time lapse imaging, independent crops of cells in which complete cell cycle could be tracked were made. These crops were manually analysed for each period and interdivision time using ImageJ. For cell length analysis, the same cell crops were segmented using Oufti, and cell lengths were extracted using a MATLAB (version – 2020a) script. The same data was used to analyse growth rate using the formula indicated in Figure 3A. Replication rates were calculated by dividing whole genome length by replication time of individual cells. For fluorescence intensity analysis of YFP expression from PsidA promoter, cells were segmented with MicrobeJ plugin on ImageJ and total fluorescence intensity normalized to cell area was extracted. For calculating SSB foci intensity, each focus was segmented using MicrobeJ plugin on ImageJ with maxima (foci) function, and total intensity of each focus was normalized to total fluorescence intensity within that cell. For calculating distance between DnaN-mCherry and nearest SSB-GFP focus, each focus was separately segmented using MicrobeJ plugin on ImageJ with maxima (foci) function. Difference in position was calculated and plotted as a cumulative frequency distribution. Difference between DnaN and random positions in the cell were generated using an R script.

Statistical analysis

All graphs were plotted on RStudio (R version – 4.3.3). For calculating statistical significance (Figures 1C, 2A–2C, 3B, 4A–4C, 4E–4H, 5A–5C, S3D, S4C–S4E, S4G, and S4H) data from three independent biological replicates were pooled, and outliers were first removed using the z-score method. Brown-Forsythe test was used to ensure similarity in variance and one-way ANOVA with Bonferroni correction was performed (assuming these distributions to be normal). Pearson’s correlation coefficients (r) were obtained to assess correlations in Figures 2D–2F, 3C, 3D, 4D, S3B, S3C, S3E, S3F, and S4B. Results of all statistical tests are consolidated in Tables S1 and S2.

Supplementary Material

Supplemental information can be found online at https://doi.org/10.1016/j.cub.2025.03.009.

Supplementary Material

In brief.

Adhikashreni et al. use quantitative live-cell imaging to track cell-cycle dynamics in hundreds of individual bacteria. They observe nutrient-dependent regulation of replication elongation rates that is linked to dNTP availability and suggest this to be a mechanism for robust adaptation to fluctuations in the growth environment.

Highlights.

  • DNA replication slows down during transition to stationary phase

  • Replication rates are linked to nutrient status and availability of dNTPs

  • Despite strong correlation, replication and growth-rate regulation are decoupled

  • Such replication slowdown is not associated with mutagenesis or DNA-damage stress

Acknowledgments

The authors are grateful to the reviewers for the constructive feedback and experimental suggestions. The authors thank Dr. Kristina Jonas for sharing the ΔspoT strain and Dr. Rodrigo Reyes-Lamothe, Dr. Petra Levin, and Dr. Sunish Radhakrishnan for feedback on the work. This work was supported by funding to I.S.A. and S.P. (TIFR graduate student fellowship) and to A.B. via the India Alliance Intermediate Grant (grant no. IA/I/21/1/505630) and intramural funding via NCBS-TIFR (Department of Atomic Energy, Government of India, project identification no. RTI 4006).

Footnotes

Author contributions

A.M.J. and A.B. conceived the project. I.S.A. led the project, carried out most of the experiments pertaining to growth phases, conducted data analysis, and performed all statistical analysis. S.P. carried out experiments across different growth media, in ΔspoT strain, with HU and dNTP, and conducted a part of the image analysis. A.M.J. conducted mutagenesis experiments and image analysis for ΔspoT and dNTP experiments. A.M.J., I.S.A., and S.P. contributed to tool generation. A.M.J. and A.B. supervised the project. A.B. procured funding. A.M.J. and A.B. wrote the manuscript with input from all authors.

Declaration of interests

The authors declare no competing interests.

Data and code availability

  • All data are provided in the manuscript.

  • No unique codes were specifically generated for the study.

  • Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Material
EMS205641-supplement-Supplementary_Material.zip (23.3MB, zip)

Data Availability Statement

  • All data are provided in the manuscript.

  • No unique codes were specifically generated for the study.

  • Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

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