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Cell Reports

Report The Bacterial SMC Complex Displays Two Distinct Modes of Interaction with the Chromosome Luise A.K. Kleine Borgmann,1 Jonas Ries,2 Helge Ewers,2 Maximilian H. Ulbrich,3,4,* and Peter L. Graumann1,4,5,* 1Microbiology,

Faculty for Biology, University of Freiburg, Scha¨nzlestraße 1, 79104 Freiburg, Germany Zu¨rich, Institute of Biochemistry, Wolfgang-Pauli-Str. 10, HCI, F 209 8093 Zu¨rich, Switzerland 3Institute of Physiology II, University of Freiburg, Scha ¨ nzlestraße 18, 79104 Freiburg, Germany 4BIOSS Centre for Biological Signalling Studies, Albert-Ludwigs-Universita ¨ t Freiburg, Scha¨nzlestraße 18, 79104 Freiburg, Germany 5SYNMIKRO, LOEWE-Zentrum fu ¨ r Synthetische Mikrobiologie, Hans-Meerwein-Straße, Mehrzweckgeba¨ude, C6, 35043 Marburg, Germany *Correspondence: [email protected] (M.H.U.), [email protected] (P.L.G.) http://dx.doi.org/10.1016/j.celrep.2013.04.005 2ETH

SUMMARY

The bacterial SMC (structural maintenance of chromosomes) complex binds nonspecifically to DNA in vitro and forms two discrete subcellular centers in vivo, one in each cell half. How this distribution is maintained is unclear. We show by time-lapse imaging of single molecules that the localization is achieved through limited, yet rapid movement of the SMC subunits through the nucleoid. Accessory ScpAB subunits mediate the arrest of 20% of SMC molecules at the center of a cell half and do not move together with the 80% mobile SMC molecules. Only free SMC, but not the preformed SMC/ScpAB complex, was able to bind to DNA in vitro, revealing distinct functions of SMC fractions. Thus, whereas SMC alone dynamically interacts with many sites on the chromosome, it forms static assemblies together with ScpAB complex partners. Our findings reveal two distinct modes of interaction of SMC with the chromosome and indicate that limited diffusion within a confined space and transient arrest may be a general mechanism for positioning proteins within a chromosome and within a noncompartmentalized cell. INTRODUCTION The bacterial chromosome is highly compacted and has a defined spatial arrangement in those bacterial cells that have been investigated so far (Toro and Shapiro, 2010). Compaction, replication, and transcription occur in parallel throughout the chromosome and are achieved by a multitude of DNA-binding proteins and proteins indirectly associated with the nucleoid. The condensation of the bacterial chromosome and the proper segregation of duplicated chromosome regions depend on several DNA-binding proteins, such as topoisomerases and small histone-like proteins, like HU, H-NS, FIS, and IHF that bind to DNA with little sequence specificity (Browning et al., 2010; Dame, 2005; Dillon and Dorman, 2010). Recently, the

global transcription silencer H-NS has been shown to form two discrete centers within each nucleoid by superresolution microscopy (Wang et al., 2011). Center formation is based on oligomerization of H-NS, which leads to the sequestration of regulated operons to the H-NS clusters. A further important complex that is essential for chromosome compaction and segregation in prokaryotic and eukaryotic cells is the SMC (structural maintenance of chromosomes) complex consisting of an SMC homo- or heterodimer and several non-SMC subunits (Hirano, 2006; Nasmyth and Haering, 2005). In eukaryotes, condensin is essential for compaction during prophase, whereas cohesin tethers sister chromosomes together from S phase into Metaphase. SMC proteins consist of an ATP-binding cassette domain called head domain, an extended stretch of coiled coils and a hinge domain, which mediates homo- or heterodimerization (Lo¨we et al., 2001). The bacterial SMC complex (MukBEF complex in E. coli) consists of the SMC protein, ScpA, and ScpB (Graumann and Knust, 2009). Although SMC forms a dimer, ScpA and ScpB form a subcomplex that binds to the SMC head domains (Volkov et al., 2003) and appears to regulate DNA binding of the SMC dimer (Hirano and Hirano, 2004). In analogy, non-SMC subunits of condensin bind to the SMC 2/4 head domains (Yoshimura et al., 2002). DNA binding involves closing of the SMC dimer into a ring-like structure (Cuylen et al., 2011; Gruber et al., 2003; Petrushenko et al., 2010; Volkov et al., 2003) and depends on proper supercoiling of the chromosome and ongoing replication (Mascarenhas et al., 2005). Ring closure is most likely mediated through dimerization of SMC head domains based on the sandwiching of two ATP molecules (Lammens et al., 2004), and ATPase activity may lead to an opening of the ring and dissociation from DNA. On the other hand, MukE and MukF form an elongated dimeric complex that binds MukB heads to also constitute a closed ring-like structure (Woo et al., 2009). ATP binding was found to disrupt the MukEF/MukB interaction in vitro, indicating an intricate interplay of SMC proteins with their complex partners and ATP. The bacterial SMC complex forms two discrete subcellular centers termed ‘‘condensation centers’’ (as seen by standard fluorescence microscopy), one within each cell half, which are essential for the proper arrangement as well as segregation of the chromosome (den Blaauwen et al., 2001; Lindow et al., Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors 1483

2002; Mascarenhas et al., 2002). Biochemical and cell biological experiments on condensin, bacterial SMC, and MukBEF complexes have shown that ScpA and ScpB/MukE and MukF are both essential for complex formation and for the formation of the subcellular condensation centers and that condensin/SMC/ MukB form high-order clusters that bind nonspecifically to DNA and can bridge distant DNA segments (Mascarenhas et al., 2005; Petrushenko et al., 2010; She et al., 2007; Strick et al., 2004). Additionally, strong ties exist between SMC/MukB and chromosome supercoiling via topoisomerases (Hayama and Marians, 2010; Kimura and Hirano, 1997; Li et al., 2010; Sawitzke and Austin, 2000; Tadesse et al., 2005); however, it is still unclear how the SMC/MukB complex can compact an entire chromosome. Many protein complexes are located at defined regions within cells, but the molecular basis for their specific subcellular localization is unclear. We wished to gain insight into the establishment and dynamic organization of SMC centers in the model bacterium Bacillus subtilis and, therefore, performed singlemolecule fluorescence microscopy. With this technique, fluorescent protein (FP) tags genetically fused to the target protein are excited through a focused laser beam (and rapidly bleach), while images are captured in stream acquisition in a wide-field fluorescence microscope. The amount of molecules present in the cell can be accurately determined from the bleaching decay (Leake et al., 2006), and when only few nonbleached molecules are left, their movement can be tracked. The intensity of a single FP molecule is determined from singular blinking events, i.e., from a single stationary FP whose fluorescence is lost in a single bleaching step within a cell. We show that cells contain two distinct fractions of SMC, 20% static and 80% dynamic molecules at any given time, whereas ScpAB complex partners were largely statically positioned in the centers of the cell halves. Contrarily, four regions on the chromosome (origin, terminus, 90 , and 270 ) localized statically, each within a different region on the nucleoid. Thus, SMC dimers dynamically interact with many chromosomal sites or, alternatively, with ScpAB complex partners within one cell half, which changes the view of how SMC interacts with the chromosome. RESULTS AND DISCUSSION Two Distinct Fractions of SMC Molecules We imaged an SMC-YFP fusion that is expressed from the original gene locus as a sole version of SMC under the control of the original smc promoter, and can functionally replace the wild-type protein, in live B. subtilis cells using 29 ms acquisitions and single-molecule microscopy (see Supplemental Information). Only full-length SMC-YFP was produced in the cells; no degradation product or free YFP was detectable (Figure S1). In the first frames of the stream acquisitions (see real-time Movies S1 and S2), fluorescence of all SMC-YFP molecules was apparent, followed by stochastic bleaching of molecules, after which only few or one molecule(s) remained, and the position of the fluorescent spots in one cell could be identified. In the final frames, a single spot remained that eventually bleached in a single step or occasionally blinked a few times before its final loss of fluorescence (Figure S2). Such single events of bleaching can only be 1484 Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors

explained through the presence of a single YFP molecule, and thus, the fluorescence intensity of a single FP could be determined. Because FPs bleached randomly, the chance of an SMC dimer containing two active FPs in the last frames was extremely low. When FP fusions moved out of the focal plane during the experiments, spots broadened, and fluorescence appeared weaker (Movies S1 and S2), but this does not influence the brightness estimates because the total integrated fluorescence stayed constant. From the visual inspection of the recorded movies, we found that SMC-YFP molecules consist of two apparent fractions, of rather stationary molecules and highly mobile ones, and the fraction of mobile SMC molecules appeared to be considerably larger than that of static molecules (Movies S1 and S2). To obtain more information on the observed behavior and the different dynamics of SMC molecules, we performed automated singlemolecule tracking and computational analysis of trajectories (Ewers et al., 2005). Fluorescent spots were recognized by a peak-finding algorithm, and closest center positions of consecutive frames were connected to tracks. Tracks shorter than seven frames were discarded. Resulting trajectories followed the fluorescent spots well (Figures 1A and 1B; Movie S1). For later analysis, we manually connected some tracks that were interrupted for a few frames but by visual inspection belonged to the same single SMC-YFP molecule (Figures 1C and 1D). In the missing frames, the presence of fluorescence could often be seen by eye, but the peak-finding algorithm failed to detect the molecule, probably due to the movement out of the focal plane. Tracks were also extracted from strains expressing ScpA-YFP, SMC-YFP with ScpA and ScpB deleted (DscpAB), and SMC-YFP after addition of mitomycin C (MMC), which will be described later (Figure 1E). SMC-YFP was never observed close to the cell poles but only to the central part of cells that is occupied by the nucleoid, in agreement with the findings that SMC is a nonspecifically DNA-binding protein (Figure 1E). As a measure of movement of single YFP-labeled molecules, we determined for each track the diameter of a minimum-bounding circle (Figure 2A). Spots that stayed within a circle of a three pixel diameter (300 nm) with a maximum of two outliers were considered as the immobile fraction (Figure 2B), and this definition agreed well with the visual impression of immobility. We found that 80% of SMC-YFP molecules were mobile and 20% immobile. To determine the speed of movement of mobile SMC molecules and to test for possible transitions between the mobile and immobile state, we analyzed the longest trajectories (>30 frames, i.e., longer than 900 ms). These included the tracks that we had manually connected from separate tracks, as described above (Figure 1C). Even for the longest tracks of more than 50 frames, we never observed the transition of a static molecule to a mobile mode or of a mobile molecule to the static mode. These data indicate that static and mobile SMC fractions exchange in a time frame of at least minutes. The diffusion coefficient of mobile SMC deduced from ten long trajectories was 0.31 ± 0.16 (SD) mm2/s, two orders of magnitude below that of freely diffusing GFP molecules (Kumar et al., 2010) (Table S1). This difference cannot be explained by the difference in hydrodynamic radius of the SMC-YFP molecule alone. Thus, SMC

Figure 1. Single-Molecule Microscopy and Tracking in Exponentially Growing B. subtilis Cells (A) Example track (green line) for a mobile SMC-YFP spot that moves for 0.58 s and then photobleaches. The outline of the cell is indicated by the white marking. (B) Two cells with three tracked SMC-YFP molecules. The beginning of the yellow and red tracks in earlier frames is not shown. The molecule with the purple track was classified as immobile; the other two as mobile. First frame shows counterstaining with SYTO60 that is the basis for the cell outlines in the second frame. (C) Example of a long track of SMC-YFP (in a cell whose outlines are indicated by the white line) that was connected from three shorter tracks (indicated by different shades of red), between which fluorescence had disappeared for only one or two frames. (D) Examples of tracks that last at least 20 frames: upper-three panels show mobile SMC-YFP molecules, lower-two panels static molecules, and outlines of cells are indicated by white lines. (E) Examples of overlays of single-molecule YFP trajectories obtained in cells (outlined by white lines); only trajectories longer than ten frames were analyzed. i, SMC-YFP; ii, ScpA-YFP; iii, SMC-YFPDscpAB; iv, SMC-YFP + 50 ng of MMC added for 30 min before imaging. Numbers in (A) and (B) state acquisition time in seconds. Scale bars, 1 mm. See also Figures S1, S2, and Movie S1.

molecules do not undergo free Brownian motion in the cell but are constrained in their motion. Similarly, the diffusion of a nonspecifically bound lactose repressor dimer has been reported to be also 0.4 mm2/s, which was attributed to 1D diffusion along DNA and 3D movement to other DNA loops based on nonspecific DNA binding (Elf et al., 2007). To support our finding that SMC-YFP displayed two populations with different mobilities, we used a second approach that is based on fitting the cumulative density function (CDF) of the distribution of single-molecule step sizes after a defined time,

thus defining the fraction of molecules whose displacement remains below a certain length within a given time (see Experimental Procedures section). We first fitted the CDF for a single homogeneous population, but the fit deviated strongly from the observed distribution (data not shown). The fit with two distributions having distinct diffusion coefficients matched the experiment very well (Figure 2C), with a fraction of 17% ± 1.4% (SD) in the immobile state and diffusion coefficients D1 = 0.012 ± 0.003 mm2/s (immobile) and D2 = 0.30 ± 0.01 mm2/s (mobile) (Figure 2D), in good agreement with the diffusion coefficient for the Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors 1485

Figure 2. Mobility of SMC and ScpA in B. subtilis (A) Beeswarm plot of the frequency of diameter of movement (ScpA-YFP, n = 106; SMC-YFP, n = 156; SMC-YFPDscpAB, n = 129; SMC-YFP MMC, n = 241). The x axis shows the density/ number of the data points such that the longer branches represent domains with higher density of data points, and the shorter branches are at lower densities. The diameter of movement was calculated from trajectories with a minimum length of 11 frames. (B) Bar chart of the percentage of tracks with a diameter less than three pixels (300 nm), which we defined as immobile. Error bars represent 95% confidence intervals for prevalence (binomial exact, Clopper-Pearson). (C) Cumulative percentage of step length of two steps (0.06 s). Green curves indicate two-distribution fitting of data (blue dots); red curve shows one-distribution fitting. (D) Diagram (log scale) of diffusion coefficient D in mm2/s, assuming a two-distribution fitting for SMC-YFP, ScpA-YFP, and SMC-YFP plus MMC, and a one-step fitting for SMC-YFPDscpAB, which is more appropriate because of the missing static fraction (see A). The area corresponds to the fraction of molecules with low- or high-diffusion coefficient. See also Figures S1, S2, and Table S1.

mobile fraction determined from the long tracks. The diffusion of the immobile fraction is so low that it cannot be distinguished from nonmovement (diffusion of one pixel per second is within the experimental uncertainty) and may thus be due to limited localization precision. Thus, our analysis shows that about 80% of all SMC molecules are highly dynamic, and 20% stay immobile. The latter fraction comprises the condensation centers seen by conventional epifluorescence microscopy, whereas the surprisingly large fraction of dynamic SMC has not been anticipated before and must be considered for any model of activity of SMC. The previously observed formation of stationary subcellular condensation centers using standard epifluorescence microscopy (Danilova et al., 2007; Lindow et al., 2002; Mascarenhas et al., 2002) had suggested that most SMC molecules would be present in these bipolar structures, but a majority of molecules (i.e., the mobile ones) was missed. The Dynamic Fraction of SMC Molecules Moves throughout the Chromosome Next, we investigated whether movement of SMC is due to rapid movement of chromosome regions. We monitored the mobility of four different regions on the circular chromosome (origin, 359 , 90 , 180 , and 270 ) in real time using LacI-GFP that binds to an operator cassette of about 250 lacO repeats that was integrated at the specific chromosome locus (Figure 3A). In Movies S3 and S4 (origin and 270 ), mostly stationary bright foci are apparent, as well as few rapidly moving spots. Although the former consist of many lacO-bound LacI tetramers, the latter are diffusing LacI molecules that move much faster than SMC1486 Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors

YFP molecules. All four chromosome regions showed very little displacement, revealing that chromosome regions are stably positioned at defined subcellular locations (Figures 3B–3E). Although this has previously been inferred from slow-speed time-lapse experiments (Viollier et al., 2004; Webb et al., 1998), our data show that positioning is static also on a much shorter timescale of 29 ms. Taking into consideration the rather static behavior of the origin regions (Figure 3B) and similarly of the centromere-binding Spo0J protein (Figure S3) that binds to at least ten specific sites surrounding the origin of replication, oriC (Lin and Grossman, 1998), we conclude from the observed movement of SMC molecules that the dynamic fraction (80%) rapidly moves between all regions of the chromosome. We next determined the distribution of single-molecule tracks in relation to their position within a cell (Figure 4A). The distribution of the tracks’ centroids along the cell’s major axis (Figure 4B) confirms previous epifluorescence microscopy observations of SMC predominance close to defined foci that lie around the middle of a cell half (Lindow et al., 2002; Mascarenhas et al., 2002; Volkov et al., 2003). In B. subtilis, DNA is duplicated in the cell center, where both replication forks are located for a long time during the cell cycle (Lemon and Grossman, 1998). Origin regions move away from the cell center soon after their duplication, sequentially followed by all other regions after their duplication (Teleman et al., 1998; Webb et al., 1998). Interestingly, CHIP on chip experiments, in which the position of proteins on the chromosome after crosslinking with the DNA is analyzed, have shown an accumulation of SMC at sites surrounding origin regions (Gruber and Errington, 2009; Sullivan et al., 2009), where the specific Spo0J-binding sites are located (Lin and Grossman,

Figure 3. Loci on the Chromosome Are Stationary (A) Schematic drawing of the B. subtilis chromosome. The regions are defined as corresponding degrees on a circle. A plasmid carrying a tandem repeat of lacO sequences is integrated at any position in the chromosome and can be visualized as LacI-GFP binds to this region. (B–E) Overlays of single-molecule trajectories of different chromosome loci obtained in exponentially growing B. subtilis cells, showing that all four chromosome regions show very little displacement. Only trajectories longer than ten frames were analyzed. Scale bars, 2 mm. See also Figure S3.

1998). It has been shown that Spo0J facilitates the binding of SMC to DNA in vitro, thus possibly acting as an SMC ‘‘loading’’ factor in vivo. Our findings that 80% of SMC molecules move throughout the entire nucleoid whereas oriC regions remain stationary reveal that the association of SMC with origin regions is not static but rather suggest that SMC moves away from oriC after initial binding (Figure 4G), to transiently interact with many chromosomal sites. This is in agreement with earlier findings that the SMC complex centers (i.e., the static SMC molecules) do not colocalize with oriC throughout the cell cycle (Volkov et al., 2003). The Subunits of the SMC Complex Show Different Patterns of Dynamics To better understand the regulation of SMC activity, we next analyzed the mobility of ScpA and ScpB molecules, which bind to SMC head domains and are thought to downregulate ATPase activity of SMC (Hirano and Hirano, 2004). Again, we expressed the YFP-tagged molecules at the original locus under the control of the original promoter. Strikingly, we found that the movement of ScpA-YFP was much more stationary compared to what we observed for SMC, and only some ScpA-YFP molecules showed rapid movement (Movies S5 and S6). This is apparent from the analysis of the diameter of the bounding circle (Figure 2A): 86.7% of all ScpA-YFP molecules are static (Figure 2B). The step-length distribution of ScpA-YFP molecules, when fitted with the CDF for two populations, displayed equal fractions (50% ± 8%) of very slow (D1 = 0.096 ± 0.001 mm2/s) and slightly faster (D2 = 0.047 ± 0.008 mm2/s) molecules (Figures 2C and 2D). However, the mobile fraction was a factor of six slower than the mobile fraction of SMC-YFP, implying strongly constrained diffusion because ScpA-YFP (56 kDa) is much smaller than SMC-YFP (162 kDa). In fact, apparent movement can in part be due to impaired tracking accuracy, generating artificial mobility for static molecules, as for the slow SMC-YFP fraction. Few ScpA-YFP molecules showed highly rapid movement, similar to nonconstrained diffusion, which is apparent as cloud-like fluorescence or extremely rapidly moving spots in Movies S5 and S6. The exact number of molecules is difficult to determine; we estimate that 2%–5% of the ScpA-YFP molecules show free diffusion. ScpB-YFP was more difficult to track due to lower fluorescence intensity, but in general, movement of ScpB was very

similar to that of ScpA and dissimilar to SMC (data not shown). These experiments show that a majority of SMC molecules move without ScpA and ScpB because these remain rather stationary, whereas 80% of the SMC molecules are highly mobile. Therefore, the different components of the SMC complex have a different localization pattern when tracked with high temporal resolution. A Fraction of SMC Is Arrested in Its Movement through Its Complex Partners Neither ScpA nor ScpB nor the ScpAB complex binds to DNA, whereas SMC binds nonspecifically to DNA in vitro (Volkov et al., 2003). The previously demonstrated colocalization and biochemical interaction of SMC with ScpAB (Hirano and Hirano, 2004; Mascarenhas et al., 2005) suggest that SMC changes from diffusion through the chromosome in the absence of ScpAB to stationary SMC bound to ScpAB at the center of the nucleoid. If this was true, we would expect the static SMC fraction to be lost due to a reduction of the stationary mode in the absence of ScpAB. Indeed, in an scpAB deletion strain, the overall diffusion rate rose significantly because the fraction of stationary SMC molecules was basically absent (Figures 2A–2D; Movie S7). The CDF of the step-length distribution could be fitted well with a single population with a diffusion coefficient of D = 0.42 ± 0.03 mm2/s (SD), similar to the fast fraction of molecules in the SMC-YFP strain with intact ScpAB. Also, movement of SMC was seen close to the cell poles, i.e., molecules moved throughout the whole cells because nucleoids are decondensed in scpAB mutant cells (Mascarenhas et al., 2002; Soppa et al., 2002) (Figure 1E). To test whether the increase in mobility of SMC could be caused by chromosome decondensation, we treated the cells with MMC, which induces DNA damage and chromosome decondensation (Kidane et al., 2004). Interestingly, diffusion rates of SMC-YFP were considerably lower for MMCtreated compared with nontreated cells (Figure 2C), possibly due to an increase in roadblocks caused by repair complexes. Slower diffusion was accompanied by a marked increase in the static fraction (Figure 2A; Movie S8), which forms the ‘‘SMC condensation centers’’ (see below). The position close to the cell quarter sites of these centers did not change after MMC treatment (data not shown). The slower diffusion due to MMC treatment suggests that a change in chromosome structure induced by DNA damage can also affect the movement Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors 1487

Figure 4. Movement of SMC around Condensation Centers (A) Schematic drawing of a bacterial cell. The red dots represent the outer edges of the cell. Connecting these points creates the major axis (thin dashed line) from where the middle line (thick dashed line), which divides the cell halves, is calculated. The colored lines represent sample tracks. (B) Histogram of distribution of SMC-YFP tracks’ centroids along the cells’ major axis (n = 234). (C) Overlay of fluorescence centers obtained from the first ten frames (red circle), with tracks of static molecules (diameter less than three pixels) that could be tracked later in the experiment (note that many tracks are too short for analysis and, thus, are not shown). Example of tracks that colocalize with a center and example of tracks that do not colocalize with a center, which constitutes a more rarely observed second center within a cell half. (D) Histogram of the distance of centroids of static SMC-YFP tracks (diameter less than three pixels) to the next condensation center. (E) Histogram of the distance of centroids of mobile SMC-YFP tracks (diameter greater than three pixels) to the next condensation center. (F) Cartoon of the architecture of the SMC complex. (G) Model for the dynamics of SMC and the SMC complex; the cell wall is depicted in brown. Replication of the chromosome (orange) by the replication machinery (red), which is located at the middle of the cell, starts at the origin of replication (purple). Ensuing, newly synthesized DNA moves toward opposite cell poles. Binding of SMC (gray arrows) to the DNA is facilitated by Spo0J (gray) around the origin of replication, and SMC dimers (blue) are moving along the DNA. Together with ScpA (dark green) and ScpB (light green), a subfraction of SMC forms a static condensation center in the middle of each cell half. See also Figures S1 and S4.

of SMC molecules but has an effect opposite to the loss of ScpA and ScpB, further supporting the finding that the two interactors directly mediate pausing of SMC. If the stationary fraction of SMC molecules arises through pausing in their movement via binding to ScpAB molecules, which are present at the condensation centers, the static SMC molecules should also be found at the condensation centers. To verify this, we plotted the centroids of the tracks of static or mobile SMC molecules relative to the position of the centers, which were determined from an overlay of the first ten frames smoothened by a Gaussian filter and peak finding of fluorescence (Figure 4C). This method gave results equal to manual selection of peaks after adjustment of the contrast to visualize fluorescence centers (Figure S4A). Figure 4D shows that the 1488 Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors

static molecules (diameter of movement less than three pixels) were mostly close to the centers, whereas the mobile molecules (movement greater than three pixels) were further away (Figure 4E). Figure 4C shows an example of a cell containing a static track colocalizing with a condensation center and a mobile track moving at a distance to a center. In some cells, static molecules were also found at positions away from the main centers (Figure 4D); this agrees with the findings from earlier studies that one (rarely) to three SMC-YFP centers can be observed in a cell half (Volkov et al., 2003). As a second test, we plotted the displacement of SMC-YFP molecules between 29 ms acquisition intervals, dependent on proximity (three pixels and less) or distance (five pixels or more) from the centers. Figure S4B shows that SMC-YFP molecules close to the centers tended to move

Figure 5. SPR Analysis of DNA Binding of SMC Response units, as a measure of the mass bound to the surface of a sensor chip, are plotted versus time (s). A 500 bp double-stranded DNA is immobilized in the chip. (A) Incubation of 250 nM SMC with increasing molar amounts of copurified ScpA and ScpB, as indicated by ‘‘ratio’’ (SMC:ScpAB). (B) Two concentrations of purified SMC or purified SMC complex.

less than molecules away from these subcellular sites (bearing in mind that mobile SMC-YFP molecules can also stop in their movement for a few frames or move past a condensation center). Thus, stationary molecules tend to be present in the condensation centers, whereas mobile molecules mostly move at positions away from the centers. The SMC Complex Fails to Bind to DNA De Novo In Vitro Any complex is in an equilibrium between free and bound subunits; the ratio of bound versus unbound depends on affinity and on abundance of subunits. We have previously shown that in cell extract, SMC is present in a high molecular weight complex together with ScpA and ScpB but also as a free dimer (Mascarenhas et al., 2005). Our tracking data now reveal that both fractions are not due to cell rupturing but indeed exist in vivo and that both fractions are stable for a time frame of minutes or more (i.e., do not interchange rapidly). Likewise, ScpB has been shown to be present as dimer, in complex with ScpA, as well as in complex with SMC and ScpA in cell extract (Mascarenhas et al., 2005). Purified ScpA and ScpB form different high molecular weight complexes (Mascarenhas et al., 2005), indicating that ScpAB multimers free of SMC may be present in the stationary centers at the nucleoid center. Freely diffusing ScpA-YFP molecules (and also ScpB-YFP; data not shown) are visible in Movies S5 and S6 as described above. Thus, the biochemical characterization of the SMC complex is in good agreement with the localization data. Hirano and Hirano have shown that purified ScpA and ScpB reduce the DNA-binding activity of SMC in vitro (Hirano and Hirano, 2004). To gain more insight into the nature of this negative effect, we employed surface plasmon resonance (SPR) experiments. SMC-his and SMC-strep were purified separately via nickel-NTA or via streptavidin affinity chromatography (SMC-his yielded more protein and was used for most experiments, but qualitatively, SMC-his and SMC-strep behave similarly) and ScpA and ScpB-strep by coexpression and purification via streptavidin affinity. The entire complex was purified by coexpression of SMC with ScpA and ScpB-strep, and analogous strep purification. As reported previously, SMC-his had low intrinsic ATPase activity, which was reduced but still present in the SMC/ScpA/ScpB complex (data not shown). Addition of ScpAB to SMC reduced DNA binding, albeit very inefficiently. At 4-fold molar excess, binding was reduced by 25%, and at 8-fold excess, to 50%. Even 32-fold

excess of the interactors could not fully suppress DNA binding by SMC (Figure 5A). Interestingly, when the entire SMC complex was purified under identical conditions and the fraction containing SMC, ScpA, and ScpB (750 kDa on gel filtration) was used, the complex failed to bind to DNA (Figure 5B), whereas purified SMC bound very efficiently. Neither purified ScpB-strep nor ScpA-his alone could reduce DNA binding of SMC (data not shown). These results suggest that the addition of ScpAB to SMC leads to inefficient formation of the complex (this was shown in Mascarenhas et al., 2005; Volkov et al., 2003), and the reduction in DNA binding is due to two fractions: free SMC that binds to DNA, and ScpAB-bound SMC that does not bind to DNA at all. These data suggest that de novo DNA binding in vivo can be achieved by the free SMC fraction and that SMC within the condensation centers may be bound to DNA (when it previously was free to interact with the chromosome) but that no new DNA binding will occur when SMC is in complex with ScpAB. Conclusions Our results show that a large fraction (80%) of SMC molecules is mobile and moves throughout the nucleoid, whereas the other fraction is halted in its movement through its SMC-complex partners. This provides an intriguing concept for the function of SMC because SMC is not constantly present at the origin regions but most likely loaded (i.e., its binding to the chromosome is facilitated) at this region to then diffuse away and move throughout the nucleoid, interacting with many sites on the chromosome. The basis of the bipolar condensation centers is a transient interaction with ScpA and ScpB partners of the SMC complex. SMC dimers change between mobile and static phases in a time frame of at least minutes, and the larger fraction of SMC molecules, those that are ScpAB-free, must also be accounted for active chromosome condensation because it can reach all sites on the chromosome. Similarly, SMC-like MukB has been shown to be composed of a large mobile and smaller static fraction in E. coli cells (Badrinarayanan et al., 2012), revealing striking similarities between these related condensation proteins. We show that SMC binds to DNA only in the absence of its interaction partners in vitro, but not when in complex with ScpAB; addition of ScpAB confers an inefficient negative effect on DNA binding of SMC, most likely because the addition of purified ScpAB to SMC leads to only a partial complex formation, and also reduces Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors 1489

ATPase activity (Hirano and Hirano, 2004). Thus, the SMC/ ScpAB complex in the condensation centers is unable to bind to DNA de novo. These data show that the dynamic, ScpABfree fraction of SMC molecules is contributing to DNA condensation via de novo DNA binding, whereas the static SMC/ScpAB complex fraction is at a central position, where SMC (possibly bound to DNA) is immobilized (Figure 4G) but where no new DNA binding occurs. Thus, our data showing a dual mode of movement of SMC and interaction with ScpAB strongly change the view on how SMC acts in live cells, revealing that DNA binding occurs throughout the nucleoid in a seconds timescale, whereas condensation centers are rather stable SMC complex places (Figure 4G). Exchange between the two fractions may be based on the ATPase cycle of SMC, which is essential for its function (Hirano et al., 2001; Schwartz and Shapiro, 2011). Possibly, the conserved eukaryotic condensin complex (Hirano, 2006; Nasmyth and Haering, 2005) also consists of highly dynamic molecules that are centered via interaction with their corresponding ScpA-like kleisin and other complex partners. Alternating movement and transient pauses based on dynamic interaction with complex partners may be the basis of the localization of many protein complexes apparently positioned at specific places within the chromosome, the cytosol, or the cell membrane in noncompartmentalized cells and within cell compartments.

To obtain the distribution of molecules along the cell, cell ends were determined visually from the SYTO 60 counterstaining and used to define the middle line dividing the cell into two halves. To investigate the mobility in dependence on the distance of a molecule to the nearest focus, the center of the region containing the focus was determined. For every frame, the distance of a localized molecule to the nearest focus was calculated. In addition, we determined the displacement of the molecules to the next frame as a measure for their mobility. Molecules closer than three pixels to the focus were categorized as ‘‘close to focus,’’ where as molecules further away than five pixels were categorized as ‘‘far away from focus.’’ For both cases, histograms of the displacement between adjacent frames were calculated and normalized.

EXPERIMENTAL PROCEDURES

Protein Expression and Sample Collection For purification of a 63His-tagged version of SMC, M15 cells harboring the expression plasmid pQE60 were inoculated to OD600 0.05 into LB with the required antibiotics and incubated on a shaking platform at 37 C till OD600 0.9 is reached. Expression was induced with 0.5 mM IPTG. Cells were harvested by centrifugation after 1 hr and lysed by French pressing in 50 mM HEPES (pH 7.5) and 300 mM NaCl. SMC was bound to a Ni-NTA column, eluted with a step gradient, and the peak fractions were rebuffered by gel filtration (Superose 6 30/100 GL) in BIAcore buffer (50 mM HEPES [pH 7.5], 100 mM NaCl, 1 mM MgCl2, and 0.005% Tween 20). The ScpAB complex or the full SMC complex was expressed from pETDuet (ScpA/B-Strep) and pCDFDuet(SMC) in Rosetta 2 (DE3) pLysS cells and was purified by strep affinity purification. The cells were incubated on a shaking platform at 37 C till OD600 0.9 is reached and induced with 0.5 mM IPTG. After 1 hr, cells were harvested by centrifugation and lysed by French pressing. ScpB-Strep was bound to a Strep-Tactin column pulling down the untagged ScpA or ScpA and SMC, respectively. The complex was eluted, concentrated with a MonoQ column, and rebuffered to BIAcore buffer by gel filtration (Superdex 200 30/100 GL).

Single-Molecule Tracking and Track Analysis Images were recorded using Andor Solis software and were subsequently analyzed by using ImageJ software Version 1.43 m (W. Rasband, National Institutes of Health, Bethesda, MD, USA; http://rsb.info.nih.gov/ij/). Single YFP molecules were identified, and the intensity of a bleaching step of a single fluorophore was measured. Trajectories of single molecules were tracked using the ImageJ plugin Particle Tracker (Sbalzarini and Koumoutsakos, 2005). Trajectories consisting of less than seven frames were excluded from further analysis. Trajectory analysis was done with custom software written in MATLAB and Mathematica. As done by others, the single-molecule-based mean square displacement (MSD) of a trajectory was calculated under the assumption that averaging multiple subtrajectories of a single particle will yield a good approximation for an ensemble average (Meier et al., 2001). Therefore, we used the following equation:

SPR A 500 bp fragment of DNA containing a native parS site was amplified from chromosomal B. subtilis DNA with 50 -biotin-labeled primers. The fragment was passed across one flow cell of a streptavidin-coated BIAcore chip surface (sensor chip SA; GE Healthcare) at 20 ml/min in 0.5 M NaCl. The surface was washed by injection of 10 ml wash buffer (500 mM NaCl, 100 mM NaOH) to remove nonspecifically adsorbed DNA. The analytes were passed across the surface of two flow cells at 20 ml/min in BIAcore buffer (50 mM HEPES [pH 7.5], 100 mM NaCl, 1 mM MgCl2, and 0.005% Tween 20). The signal of the flow cell without DNA (‘‘control’’) was subtracted from the flow cell with the DNA-coated surface (‘‘sample’’). Surfaces were regenerated by the injection of 5 ml wash buffer. For additional methods and strains, please consult Table S2 and the Extended Experimental Procedures.

MSDðDt = n dtÞ = ðN  nÞ1

Nn  X

 ðxi + n  xi Þ2 + ðyi + n  yi Þ2 ;

SUPPLEMENTAL INFORMATION

i=1

where xi and yi are the particle position of a trajectory with N frames at frame i with a frame time interval of dt. To estimate the diffusion coefficients (D) from individual single-molecule tracks, the first 40% of the MSD versus time (Dt) curve was fitted linearly. To extract the diameter of the minimumbounding circle, the smallest circle around all localizations within a single track was fitted while neglecting the two localizations with the largest distance from the center of mass because most tracks contained one or two outliers. Cumulative probability density functions of all observed step lengths were fitted to extract diffusion coefficients from all observed single molecules. The expected CDF is cdf1(x) = 1Exp(x2/(4Dt)) when there is a single population with diffusion coefficient D, t = 0.06 s being the time between three frames, and x the distance between center positions in frames n and n+2. With two populations having diffusion coefficients D1 and D2 and relative fractions a and 1a of the total, the CDF becomes cdf2(x) = 1  a 3 Exp(x2/(4D1t))  (1a) 3 Exp(x2/(4D2t)). For all parameter fit values, the SD was calculated as a measure for the deviation of the fitted curve from the experimental data points.

1490 Cell Reports 3, 1483–1492, May 30, 2013 ª2013 The Authors

Supplemental Information includes Extended Experimental Procedures, four figures, two tables, and eight movies and can be found with this article online at http://dx.doi.org/10.1016/j.celrep.2013.04.005. LICENSING INFORMATION This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial-No Derivative Works License, which permits non-commercial use, distribution, and reproduction in any medium, provided the original author and source are credited. ACKNOWLEDGMENTS We would like to thank Professor Dr. Dieter Hauschke (University Medical Centre of Freiburg) and Frederick Person (University of Uppsala) for help with the statistical analysis or determination of diffusion coefficients. This work has been supported by the Deutsche Forschungsgemeinschaft (IRTG

1478), the Excellence Initiative of the German Federal and State Governments (EXC 294), and the Swiss NCCR Neural Plasticity and Repair. J.R. is a Marie Curie Fellow. Received: February 28, 2012 Revised: October 26, 2012 Accepted: April 4, 2013 Published: May 9, 2013 REFERENCES Badrinarayanan, A., Reyes-Lamothe, R., Uphoff, S., Leake, M.C., and Sherratt, D.J. (2012). In vivo architecture and action of bacterial structural maintenance of chromosome proteins. Science 338, 528–531. Browning, D.F., Grainger, D.C., and Busby, S.J. (2010). Effects of nucleoidassociated proteins on bacterial chromosome structure and gene expression. Curr. Opin. Microbiol. 13, 773–780. Cuylen, S., Metz, J., and Haering, C.H. (2011). Condensin structures chromosomal DNA through topological links. Nat. Struct. Mol. Biol. 18, 894–901.

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