Evidence for a DNA-relay mechanism in ParABS-mediated chromosome segregation

Abstract

The widely conserved ParABS system plays a major role in bacterial chromosome segregation. How the components of this system work together to generate translocation force and directional motion remains uncertain. Here, we combine biochemical approaches, quantitative imaging and mathematical modeling to examine the mechanism by which ParA drives the translocation of the ParB/parS partition complex in Caulobacter crescentus. Our experiments, together with simulations grounded on experimentally-determined biochemical and cellular parameters, suggest a novel 'DNA-relay' mechanism in which the chromosome plays a mechanical function. In this model, DNA-bound ParA-ATP dimers serve as transient tethers that harness the elastic dynamics of the chromosome to relay the partition complex from one DNA region to another across a ParA-ATP dimer gradient. Since ParA-like proteins are implicated in the partitioning of various cytoplasmic cargos, the conservation of their DNA-binding activity suggests that the DNA-relay mechanism may be a general form of intracellular transport in bacteria.DOI: http://dx.doi.org/10.7554/eLife.02758.001.

Keywords: Caulobacter crescentus; DNA partitioning; Par system; chromosome dynamics; intracellular transport.

Figure 1.. Biochemical analysis of ParA ATPase cycle.

(A) Gel filtration analysis of purified wild-type ParA (left) and DNA-binding deficient mutant ParA_R195E (right) in the presence and absence of ATP. (B) Specific activity of ParA (concentration fixed at 6 µM) measured as a function of DNA concentration (0–2.0 mg/ml). Michaelis–Menten equation fit to the data (black) gives K_DNA = 0.15 mg/ml. (C) Effects of DNA and ParB on ATPase activities of ParA (cyan) and ParA_R195E (yellow). (D) ADP production rate as a function of ParA concentration measured with fixed ATP (2.5 mM) and DNA (1.5 mg/ml) concentrations. Linear fit to the data (black line) gives k_cat = 5.8 hr⁻¹. (E) Dependence of ATPase activity of ParA upon ATP concentration measured with fixed ParA (10 µM) and DNA (1.5 mg/ml) concentrations. Fitting the Michaelis–Menten equation (black line) gives k_cat = 5.9 hr⁻¹ and K_M = 150 µM. (F) Dependence of ParA (3 µM) ATPase activity on ParB concentrations. Michaelis–Menten equation fit to the data (black line) gives k_cat = 120 hr⁻¹ and K_M = 80 µM. (G) Model of ParA ATPase cycle. ATP-binding promotes ParA dimerization. Upon binding DNA, ParA-ATP dimers presumably adopt an ATP-hydrolysis competent state. A high concentration of ParB is required to stimulate ATP hydrolysis of DNA-associated ParA-ATP dimers. The cycle reinitiates through nucleotide exchange. SDS-PAGE images showing the purity of the purified protein preparations and the results from experiments comparing the stimulatory effects of wild-type (WT) ParB and mutant ParB_L12A on ParA ATPase activity are presented in Figure 1—figure supplements 1 and 2. Experiments performed to correlate the biochemical activities measured in vitro with in vivo conditions are presented in Figure 1—figure supplements 3–8. DOI: http://dx.doi.org/10.7554/eLife.02758.003

Figure 1—figure supplement 1.. SDS-PAGE analysis of purified protein preparations used in this study.

(A) ParA, (B) ParA_R195E, (C) ParB-His6, (D) ParB_L12A-His6, and (E) ParA_G16V-YFP. Approximately 5–15 µg of proteins were loaded in each lane. DOI: http://dx.doi.org/10.7554/eLife.02758.004

Figure 1—figure supplement 2.. L12A mutation, which prevents ParA–ParB interaction, severely compromises ParB ability to activate ParA ATPase activity.

ATPase rates were measured from reactions containing 3 µM ParA and 1.5 mg/ml salmon sperm DNA in the absence or presence of 90 µM of either wild-type ParB or ParB_L12A. The measured rates were compared to the rate of the reaction lacking ParB. Error bars represent the standard deviations (SD) of results measured in duplicates. DOI: http://dx.doi.org/10.7554/eLife.02758.005

Figure 1—figure supplement 3.. Measurement of GFP-ParB distribution at the subcellular level.

(A) Representative fluorescence image showing autofluorescence in wild-type (CB15N) cells and subcellular distribution of GFP-ParB fluorescence in MT174 cells, which produce GFP-ParB from the native parB promoter on the chromosome as the only ParB copy. (B) Single-cell total fluorescence and cell areas were quantified using MicrobeTracker and a histogram of normalized fluorescence (total fluorescence divided by cell area) is shown. The two-peak distribution was fit by a bimodal Gaussian (black), representing fluorescence detected in two populations of cells: wild-type (magenta) and GFP-ParB-producing cells (cyan). The mean of the magenta curve was designated as the mean normalized autofluorescence (AF_mean), which was used for background subtraction. (C) Histogram showing the fraction of GFP-ParB associated with parS in single cells. The experimental distribution (grey) was fitted with a Gaussian function (dashed black curve), giving an average of 80% ParB molecules associated with the partition complex. (D) Western blot of a lysate made from an MT174 culture using an anti-GFP monoclonal antibody (JL-8) or an anti-ParB polyclonal antibody (data not shown). GFP-ParB and a degradation fragment were detected. (E) The fraction of GFP-ParB fluorescence associated with parS stays constant as a function of cell length. Single-cell raw data (grey circles) and average fraction GFP-ParB fluorescence associated with parS (black circles) ± SD across 0.25 μm intervals are shown. (F) Subcellular volume occupied by a ParB/parS complex as determined by PALM microscopy. mEos3.2-ParB synthesis in CJW4978 cells was induced with 0.03% xylose for 1 hr in M2G before imaging using PALM. Fluorescent emitters within six polar partition complexes were analyzed. The histogram was fitted with a Gaussian function with σ = 39 nm (solid line). Radial distribution of mEos3.2-ParB from the foci were then combined and plotted as a histogram (n = 440 localizations). This means that 95% of the molecules were localized within 78 nm (2σ) from the center of the spot. We used 78 nm as the radius of the spherical volume occupied by ParB dimers within a partitioning complex. Such a sphere occupied a volume of 2 × 10⁻³ fL. DOI: http://dx.doi.org/10.7554/eLife.02758.006

Figure 1—figure supplement 4.. Quantification of ParB and ParA abundance in cell lysates by quantitative Western blotting.

(A) Top, a Western blot of a wild-type swarmer cell lysate (CB15N producing endogenous ParB) and indicated amounts of purified ParB-His6 using a polyclonal antibody against ParB. Bottom, the optical densities (OD) of the bands corresponding to ParB-His6 standards were quantified (black circles) and were fit by a linear function (black line). The amount of ParB in the cell lysate (red circle) was calculated from the calibration. This measurement was repeated twice. The number of ParB per cell was calculated using this formula: $\frac{ParB\hspace{0.17em}amount\hspace{0.17em}in\hspace{0.17em}lysate\times Avogadro\hspace{0.17em}number}{ParB\hspace{0.17em}moleculr\hspace{0.17em}weight\times \#cells\hspace{0.17em}in\hspace{0.17em}lysate}=720\pm 80$ ParB molecules per cell. (B) Top, a Western blot of a swarmer cell lysate of CJW3010 strain (in which parA-yfp replaces parA at the native location in the chromosome) and indicated amounts of purified ParA_G16V-YFP-His6 using a monoclonal antibody against GFP (JL-8). Bottom, the ODs of the bands corresponding to ParA_G16V-YFP-His6 standards were quantified (black circles) and fit with a linear function to create a calibration curve (black line). Red circle represents the calculated amount of ParA-YFP in the cell lysate. This measurement was repeated three times. The number of ParA-YFP per cell was calculated using the formula $\frac{ParA-YFP\hspace{0.17em}amount\hspace{0.17em}in\hspace{0.17em}lysate\times Avogadro\hspace{0.17em}number}{ParA-YFP\hspace{0.17em}moleculr\hspace{0.17em}weight\times \#cells\hspace{0.17em}in\hspace{0.17em}lysate}=170\pm 20$ ParA-YFP molecules per cell. DOI: http://dx.doi.org/10.7554/eLife.02758.007

Figure 1—figure supplement 5.. His₆-ParB forms a dimer in solution.

Purified His6-ParB was subjected to HPLC size exclusion chromatography (SEC) coupled with UV, on-line laser light scattering (LS) and refractive index (RI) detectors (SEC-UV/LS/RI) to measure its absolute MW. The dashed lines indicate the elution profile of His6-ParB from an RI detector while the dots are MW calculated from LS measurements at 1-s intervals. The MW measurement showed an average MW of 69 kDa, which is consistent with His6-ParB (monomeric MW = 34 kDa) forming a dimer in solution under this experimental condition. DOI: http://dx.doi.org/10.7554/eLife.02758.008

Figure 1—figure supplement 6.. Fluorescence-based quantification of ParA-YFP abundance in C. crescentus cells.

Purified ParA_G16V-YFP-His6 molecules were immobilized on a poly-Lysine-coated cover slip and imaged by stream acquisition using 2-s integration time. ParA_G16V-YFP-His6 molecules were observed as immobile diffraction-limited spots. Shown in (A) are time lapse sequences representing the three most observed spot behaviors: (i) one-step photobleaching, (ii) two-step photobleaching and (iii) blinking. (B) Intensity profiles of single diffraction-limited spots shown in (A) showing 1-step photobleaching, 2-step photobleaching and blinking of YFP molecules. ΔI was calculated as the difference between the averaged fluorescence intensities before and after each stepwise photobleaching. (C) Histogram showing the magnitude of fluorescence loss following 1-step photobleaching (n = 219). The mean of a Gaussian fit to the histogram gives the fluorescence value of a single YFP molecule. (D) Number of YFP molecules per cell as a function of cell length. Swarmer cells from CJW3010 strain, which expresses parA-yfp from the native promoter of parA, were imaged on the same pad as wild-type swarmer cells. The number of YFP molecule per cell was calculated by dividing single-cell YFP fluorescence values by single YFP fluorescence. The cells were color-coded based on the YFP count using 50 as a cut-off (blue: parA-yfp-expressing cells, n = 251; red: wild-type, n = 248). The black lines represent linear fits to the red (y1 = 1.6x; autofluorescence in wild type cells on average equals to 1.6 YFP molecules per 1 µm cell length) and blue (y2 = 56x; fluorescence in parA-yfp cells on average equals to 56 YFP molecules per 1 µm cell length) data points, respectively. Typically, C. crescentus is ∼2.3 µm in length when parS translocation commences. Thus, there are 2.3 × (56–1.6) = 120 ParA-YFP molecules at the beginning of parS translocation. DOI: http://dx.doi.org/10.7554/eLife.02758.009

Figure 1—figure supplement 7.. Estimation of the relative abundance of ParA and ParB in vivo.

(A) Cultures (1.5 ml) of CJW3010 (parA-yfp-expressing) and MT174 (gfp-parB-expressing) cells at OD₆₆₀ = 0.25 were harvested and lysed by boiling in equal volumes of SDS buffer followed by sonication. Duplicates of 10 µl ParA-YFP lysate were resolved on the same gel as GFP-ParB lysates (in varied volumes as indicated) and subjected to Western blot analysis using the JL-08 monoclonal α-GFP antibody (Clontech). The first lane separated by the dashed line was moved from a different region of the same blot for easier comparison. (B) All bands were quantified using ImageJ and the intensities of the GFP-ParB bands were plotted as a function of volume of lysate loaded. The solid line represents the best linear fit, which we used to calculate that 10 µl of ParA-YFP lysate (the red dot represents the mean intensity of two lanes on the same blot) contained approximately the same amount of GFP fusion molecules as those in ∼3 µl of GFP-ParB lysate. In other words, assuming that YFP/GFP fusions do not affect protein levels, the ParA:ParB ratio in swarmer/early stalked cell stage is approximately 1:3. This method estimated that there are 720 ÷ 3 = 240 ParA molecules per cell. DOI: http://dx.doi.org/10.7554/eLife.02758.010

Figure 1—figure supplement 8.. Maximum lengths of different possible ParA polymer configurations.

(A) A surface representation of the Thermus thermophilus Soj dimer crystal structure showing the dimensions of the dimer along the x- and y- axes (Leonard et al., 2005). The red areas represent the positions of arginine 182 which corresponds to arginine 195 in C. crescentus ParA and which has been shown to be critical for DNA-binding (Ptacin et al., 2010; Schofield et al., 2010). (B) Bottom view of the T. thermophilus Soj dimer showing the dimensions of the dimer along the x- and z- axes. (C) Three possible polymer configurations considering polymerization along the x-, y- and z-axes of a ParA dimer. The maximal achievable length of each polymer (L) was calculated as the product of the physiological abundance of ParA (90 dimers) and the unit length of a ParA dimer along each axis. DOI: http://dx.doi.org/10.7554/eLife.02758.011

Figure 2.. Two-dimensional dynamics of the partition complex.

(A) Representative 2-D trajectory (Δt = 20 s) of duplicated GFP-ParB/parS complexes in a CJW4762 cell. Positions of each partition complex detected in consecutive frames were joined with solid lines while dotted lines were used to connect positions interrupted by frame(s) with failed localization. (B) Trajectory of GFP-ParB/parS complexes in (A) along the long cell axis (black, old pole-proximal ParB/parS complex; for the distal partition complex: blue, ‘slow’ phase, red, ‘fast’ phase, and green, the anchored phase). x₀ is the position of the distal partition complex at the moment the second GFP-ParB/parS spot appeared in a cell. x_finish is the position of the distal partition complex at the moment it became anchored at the new pole. x_start is the position when the partition complex transitions from the ‘slow’ phase into the ‘fast’ phase. Run length is the distance between x_finish and x_start along the long cell axis. Run period is the time taken to travel from x_start to x_finish along the long axis. Also shown is a table summarizing the mean ± SD values for x₀, x_start and x_finish. There were more counts for x_start and x_finish (n = 141) than for x₀ (n = 96) because in some cells the GFP-ParB/parS complex had already duplicated prior image acquisition. These values were used to define the start position for the ParB/parS complex in our computer simulations and the fast phase in the analysis. (C) Histogram showing the distribution of run lengths (n = 141). (D) Histogram showing the distribution of run periods (n = 141). (E) Histogram showing the distribution of translocation speeds of partitioning ParB/parS complexes during the fast phase (n = 141). (F) Histogram showing the distribution of short-axis displacements of segregating partition complexes during the slow (blue), fast (red) and pole-anchored (green) phases. DOI: http://dx.doi.org/10.7554/eLife.02758.013

Figure 3.. Subdiffraction visualization of ParA-Dendra2 in live cells.

(A) Representative super-resolution PALM images of cells producing ParA-Dendra2 at native levels in place of ParA (strain CJW4915), ParA-Dendra2 at xylose-inducible levels in addition to ParA-Dendra2 expressed from the native promoter (CJW5154), crescentin-Dendra2 at native levels (CJW4902) and ribosome-associated L1-Dendra2 at native levels (CJW5156). Synthesis of ParA-Dendra2 in CJW5154 cells was induced with 0.3% xylose for 1 hr before swarmer cells were isolated for imaging. Scale bar = 1 µm. (B) Positional distributions of single emitters along the short-cell axis for cells expressing ParA-Dendra2 (CJW5154, green), crescentin-Dendra2 (CJW4902, blue) and L1-Dendra2 (CJW5156, red). For each cell, a 0.5–0.8 μm segment containing at least 600 emitters was selected. Emitter positions along the short cell axis were calculated relative to the mean position. Additional super-resolution microscopy images and analysis are presented in Figure 3—figure supplement 1. DOI: http://dx.doi.org/10.7554/eLife.02758.014

Figure 3—figure supplement 1.. Comparison of ParA and crescentin localization by super-resolution imaging.

(A) Left, super-resolution image of CJW5154 cells producing ParA-Dendra2 from both the native and xylose promoters. Cells were imaged at 100 frames/s for 30 s using a custom-built microscope setup equipped with a sCMOS camera (Huang et al., 2013). ParA-Dendra2 synthesis was induced with 0.3% xylose for 1 hr before swarmer cells were isolated for imaging. The data set was processed using a sCMOS-specific algorithm (Huang et al., 2013). Right, line profiles of emitter positions in the white boxes show the spread of ParA-Dendra2 across the cell width near the new pole (i and ii) and the distribution of ParA-Dendra2 localized at the cell pole (iii). (B) Same as A, except for the CJW4902 strain, which produces crescentin-Dendra2 from the native creS promoter, were imaged. Scale bars = 500 nm. For visualization purpose, the upper bounds in the color map have been adjusted for A and B separately. AU, arbitrary units. DOI: http://dx.doi.org/10.7554/eLife.02758.015

Figure 4.. The DNA-relay model results in robust translocation of the ParB/parS complex.

(A) Left, graphical representation of the ‘diffusion-binding’ model: the diffusing ParB/parS complex (green disk) interacts with DNA-immobilized ParA-ATP dimers (filled red disks), stimulates ATPase activity, resulting in the dissociation of ParA from the DNA (open red disks). Right, graphical representation of the ‘DNA-relay model’: same as in (A) except that DNA-bound ParA-ATP dimers fluctuate randomly according to the movement of the associated DNA. When the ParB/parS complex is associated with one or more DNA-bound ParA-ATP dimers, it experiences the elastic force governing the dynamics of the DNA loci associated with ParA dimers. (B) Averaged positions of the simulated ParB/parS complex as a function of time. (C) Fraction of trajectories that completed translocation as a function of time. A summary of all parameters used in the simulations is presented in Table 2. Additional experiments performed to obtain these parameter values are presented in Figure 4—figure supplements 1 and 2. DOI: http://dx.doi.org/10.7554/eLife.02758.016

Figure 4—figure supplement 1.. Average ParA-YFP fluorescence profiles during ParB/parS segregation.

Cells expressing ParA-YFP and CFP-ParB (CJW3367 strain) were imaged, analyzed with MicrobeTracker and SpotFinderY, and sorted based on the distance of the translocating partition complex from the new pole at 100-nm intervals. Shown in the figure are the average fluorescence profiles of ParA-YFP between the translocating CFP-ParB/parS partition complex and the new pole (red curves) of cells sorted into the 2.0 ± 0.05 and 1.6 ± 0.05 µm groups. The grey bars mark the average distance of the partition complex from the new pole. DOI: http://dx.doi.org/10.7554/eLife.02758.017

Figure 4—figure supplement 2.. Estimation of the diffusion coefficient of the ParB/parS complex, D_PC.

(A) Three segments were identified in the trajectories of GFP-ParB/parS when the partition complexes were not engaged in the fast, ParA-dependent movement (see ‘Materials and methods’ for details). (B) The displacement along the long cell axis within each segment was collected from all cells and the distributions were plotted. Closed circles indicate data while solid lines indicate Gaussian fits to displacement distributions for each of the segments. (C) Mean square displacement (MSD) was plotted for each of the segments (closed circles). Solid lines represent linear fits (MSD = D_PC t + C, with a variable offset C, as described previously [Bakshi et al., 2011; English et al., 2011]) to the data points. DOI: http://dx.doi.org/10.7554/eLife.02758.018

Figure 5.. Chromosomal loci exhibit elastic dynamics.

(A) Representative trajectory of a LacI-CFP-labeled groESL locus tracked in live CJW2966 cells showing the dynamics of a chromosomal locus. The DNA locus was imaged every 2 s for 180 s. The displacements were colored as a function of time and overlaid with the cell outline (green). The zoomed trajectory in the inset shows the magnitude of displacements. To induce LacI-CFP expression, cells were incubated for 1 hr with 0.03% xylose prior to imaging. (B) Left, two-dimensional distribution of the groESL locus positions relative to the mean position of each trajectory. The scatter plots were generated from locus positions of 641 trajectories while the mesh surface is the best 2-D asymmetrical Gaussian fit (with σ_long = 0.06 µm and σ_short = 0.04 µm). The lower plane is a heatmap of the experimental data. Right, the same data set is represented as 1-D distributions of the groESL locus positions relative to the mean position of each trajectory along the long and short cell axes in single cells. Experimental data (filled circles, long axis; open circles, short axis) and Gaussian fits (solid line, long axis; dashed line, short axis) are shown. (C) 1-D distributions of the 139_lac and 165_lac locus positions (Viollier et al., 2004) relative to the mean position of each trajectory along the long and short cell axes in single cells. Left, asymmetrical Gaussian fit gives σ_long = 0.06 µm and σ_short = 0.03 µm for the 139_lac locus in which the lac operators are inserted at position 1599540 on the C. crescentus chromosome (CJW5466 strain). Right, asymmetrical Gaussian fit gives σ_long = 0.05 µm and σ_short = 0.03 µm for the 165_lac locus in which the lac operators are inserted at position 2481399 on the C. crescentus chromosome (CJW5468 strain). DOI: http://dx.doi.org/10.7554/eLife.02758.022

Figure 6.. The ParB/parS complex transiently adopts an extended conformation during the fast segregation phase.

(A) A fraction of GFP-ParB/parS complexes (red arrows) adopts extended conformation that appears as non-diffraction limited spots in fluorescent images of live CJW4762 cells. (B) Same as (A) except that cells were fixed with 4% formaldehyde prior imaging. (C) The aspect ratio (AR) of CFP-ParB/parS complexes in live CJW3367 cells was calculated as the ratio of the longest dimension, l, to the shortest dimension, d, for each partition complex signal. A representative cell is shown with the fluorescent CFP-ParB/parS signals outlined in yellow. (D) The propensity of a CFP-ParB/parS partition complex (PC) to display a decompacted conformation is shown with respect to cellular position. Individual CFP-ParB/parS complexes were binned according to their relative positions in the cell with old pole = 0 and new pole = 1 (see ‘Materials and methods’ for pole discrimination). The fraction of CFP-ParB/parS complexes with AR >1.5 in each bin are shown. (E) Top, representative time-lapse sequence (4-s intervals) showing de-compaction and recoiling of a segregating partition complex (marked by GFP-ParB) in a CJW4762 cell. Bottom, ARs of the anchored (red) and segregating (blue) ParB/parS complexes as a function of time is shown. For all the experiments indicated above, the expression of ParB fusion proteins was induced with 0.03% xylose in M2G for 60–75 min prior to synchronization and imaging. All scale bars = 1 µm. DOI: http://dx.doi.org/10.7554/eLife.02758.024

Figure 7.. Appropriate ParB-stimulated ParA ATPase rates are important for the robustness of the DNA-relay model.

(A) Averaged trajectories of the partition complex along the long cell axis during the fast ParA-dependent phase were simulated using varied ParB-stimulated ParA ATPase rates (k_cat) and a fixed diffusion coefficient for the ParB/parS complex (D_PC) of 0.0001 μm²/s. (B) Same data set as (A), except that for each k_cat, the fraction of trajectories that completed translocation are shown as a function of time. DOI: http://dx.doi.org/10.7554/eLife.02758.025