Introduction

Rapid and specific interference with protein activity and dynamics in living cells is essential for studying and understanding biological mechanisms. However, current approaches—primarily based on small-molecule inhibitors—remain laborious, time-consuming, and costly. The advent of conformation-specific synthetic nanobody (sybody) selection (Zimmermann et al., 2018), and more recently, computational design of protein binders (Pacesa et al., 2025), offer unique opportunities for protein-based inhibition and modulation of cellular targets. Here, we generate sybodies against the SMC complex in B. subtilis to efficiently inhibit its function within cells.

Structural Maintenance of Chromosomes (SMC) complexes are multi-subunit, ring-shaped ATP-hydrolyzing DNA motors that structure chromosomal DNA by loop extrusion. They are essential for chromosome organization and segregation, gene expression, DNA repair, and defense against non-self DNA across all domains of life loop (Yatskevich, Rhodes and Nasmyth, 2019). Each SMC protein (Smc in B. subtilis, bsuSmc) is a long polypeptide with its N- and C-termini folding together to form a globular ATP binding cassette (ABC) “head” domain. The head is connected to a “hinge” dimerization domain via a ∼50 nm antiparallel coiled coil “arm”, creating an elongated dimer (Figure 1Aii and 1Aiii). ATP binding promotes head engagement, bringing the heads together to form a functional ATPase. Non-SMC subunits, a kleisin (ScpA in B. subtilis) and a dimer of KITE (ScpB in B. subtilis) or two HAWK proteins, bridge the heads, together forming a ring around DNA (Figure 1Aiii). In B. subtilis, Smc-ScpAB is recruited to the origin of replication (oriC) region on the chromosome by ParB, a DNA-binding protein that recognizes centromere-like parS sequences (Figure 2A). Once loaded, Smc-ScpAB translocates at a rate of ∼1 kb/s onto flanking DNA, aligning the two chromosome arms and individualizing nascent sister chromosomes (Gruber et al., 2014; Bürmann and Gruber, 2015; Wang et al., 2017), likely bypassing obstacles on the chromosome through the SMC hinge gate (Liu et al., 2025). Null mutants of smc, scpA, or scpB fail to segregate chromosomes properly and lose viability under conditions promoting rapid growth (Sullivan, Marquis and Rudner, 2009; Gruber et al., 2014; Wang et al., 2014).

Figure 1:

Figure 2:

Several models have been proposed for the mechanism of DNA loop extrusion by SMC complexes, including the “segment capture” model, in which DNA segments are transiently trapped between the SMC arms in partially open states and fused into larger loops through iterative ATP-driven cycles (Figure 1Aiii and S1B) (Minnen et al., 2016; Diebold-Durand et al., 2017; Marko et al., 2019). However, detailed structural understanding and experimental testing in vivo are lacking.

Antibodies can block biochemical reactions by stabilizing reaction intermediates. Synthetic single-domain antibodies called sybodies are small, robust antigen-binding proteins that were engineered based on three camelid nanobody structures (Zimmermann et al., 2018). They are selected from synthetic libraries that encode a wide diversity of binding surfaces and epitope shapes based on the length and geometry of their CDR3 loop, allowing strong binding of diverse antigen surfaces (Figure 1Ai). Their small size (∼15 kDa), stability, and ability to bind transient epitopes make them ideal tools for targeting specific states of biomolecules (Rasmussen et al., 2011). Selection is carried out using purified and immobilized Smc-ScpAB by ribosome display and phage display, typically followed by an ELISA-based screening (Zimmermann et al., 2018).

Here, we demonstrate that sybodies can be used to interfere with the function of B. subtilis Smc-ScpAB in vivo. We first isolate binders that specifically target purified Smc-ScpAB in vitro and then select those that eliminate Smc function when expressed in B. subtilis. Fourteen sybodies were found to disrupt chromosome segregation, mimicking smc deletion phenotypes. Biochemical assays confirmed that selected sybodies alter the ATP hydrolysis rate of Smc-ScpAB and its stimulation by DNA addition, likely stabilizing otherwise rare intermediates of the conformational cycle. Mapping experiments using chimeric Smc constructs revealed that most binders, unexpectedly, target the coiled coil, specifically in a defined region near the Smc 4N arm-to-arm contact, highlighting the potential of sybodies and designed binders, as genetic tools to identify essential functional domains and underscoring the importance of coiled coil dynamics for Smc function.

Results

Generation of bsuSmc-ScpAB-specific sybodies

To isolate sybodies that impede the function of B. subtilis Smc-ScpAB complex, we first performed in vitro selections with purified Smc-ScpAB under conditions favoring an ATP-bound state, using 40 bp duplex DNA, and an ATPase-deficient mutant (E1118Q) of bsuSmc biotinylated at the hinge at residue R643C for immobilization (Figure 1A and S1A) (Hirano and Hirano, 2004; Bürmann et al., 2017; Zimmermann et al., 2018; Zimmermann et al., 2020). Starting from three synthetic sybody libraries encoding distinct epitope-binding geometries (denoted as concave, loop, convex, respectively, Figure 1A), we performed one round of ribosome display followed by two rounds of phage display, thereby enriching sybodies that bind to ATP- and DNA-bound Smc-ScpAB (Table 1). From each library, 95 sybody-expressing E. coli clones were randomly chosen for in vivo characterization.

Identification of bsuSmc-ScpAB-blocking sybodies through phenotypic screening in B. subtilis

Despite containing a conserved disulfide bond, some sybodies can be expressed inside cells and bind their targets in the reducing environment of the cytoplasm, likely owing to their robust folding (Deneka et al., 2021). To assess whether the selected sybodies interfere with Smc function in vivo, each of the 285 sybody sequences was cloned under a xylose-inducible promoter (P_xyl) into an E. coli– B. subtilis shuttle vector and integrated at the amyE locus of the wild-type 1A700 strain (Figure 1B) (Diebold-Durand, Bürmann and Gruber, 2019). Fourteen sybodies from the loop library failed to yield transformants in the presence (but not absence) of xylose, indicating cell lethality due to sybody inhibition of Smc-ScpAB (Figure 1C, S2A). These Smc-disruptive sybodies (denoted as Sb002, 003, 006, 010, 015, 020, 021, 031, 156, 160, 162, 169, 188, and 194, respectively) harbor fourteen distinct sequences, showing the absence of duplicates (Table S1). Intriguingly, no such disruptive sybodies were isolated from the concave and convex libraries (Figure S2B), suggesting that the CDR3 geometry as present in the loop library is particularly effective at targeting bsuSmc-ScpAB. Alternatively, the other libraries did not produce good binders or these sybodies were not stably expressed in B. subtilis. Notably, tendencies of preferential isolation of binders from one of the three libraries have previously been observed, although at milder levels (Zimmermann et al., 2018; Hutter et al., 2019; Deneka et al., 2021).

Selected sybodies target bsuSmc-ScpAB function in vivo

To determine whether the fourteen sybodies indeed impair chromosome segregation, we used fluorescence microscopy to monitor oriC positioning in a B. subtilis strain expressing a ParB-GFP fusion protein together with a P_xyl-inducible sybody gene. In these strains, ParB-GFP marks the replication origin region by binding to parS sites near oriC (Figure 2Ai). As expected, the Δsmc strain displayed fewer ParB-GFP foci and elongated cells, consistent with delayed oriC separation (Figure 2Ai) (Table S2) (Gruber et al., 2014; Wang et al., 2014). A time course experiment using sybody Sb006 revealed chromosome organization defects as early as 30 minutes post induction, with a significant reduction in ParB-GFP foci per µm cell length at 40 minutes (p = 0.0145) (Figure 2Aii). Subsequent imaging was performed ∼35 minutes after induction of sybody expression.

All disruptive sybodies reduced ParB-GFP foci density compared to the control (1.03 foci/µm), with values ranging from 0.56 foci/µm (Sb020) to 0.22 foci/µm (Sb003). These defects were all robustly detected, while being less severe than the Δsmc mutant (∼0.05 foci/µm), conceivably due to the short induction time or incomplete inhibition. Milder defects were also observed without induction, likely attributed to leaky expression from the P_xyl promoter (Figure S3A). Sybody expression also caused cell elongation and growth delays, both hallmarks of impaired Smc activity, consistent with chromosome segregation defects that delay cell division (Figure S3B–C). Altogether, these results show that all selected sybodies induce Δsmc-like phenotypes likely by interfering with bsuSmc-ScpAB activity in vivo.

Selected sybodies specifically target B. subtilis Smc-ScpAB

To test whether sybodies specifically target B. subtilis Smc-ScpAB, we assessed their effects in a strain expressing Streptococcus pneumoniae (Spn) Smc-ScpAB and ParB proteins in place of the endogenous B. subtilis proteins. The Spn sequences can functionally replace the corresponding Bsu sequences despite significant sequence divergence (∼38% sequence identity) (Bock et al., 2022). Strikingly, none of the sybodies impaired growth in this background, even though a cognate ParAB pair is absent in this strain (S. pneumoniae lacks parA), sensitizing cells to chromosome segregation defects (Figure 2C). These results confirm that the sybodies are specific to bsuSmc-ScpAB and that off-target toxicity is not noticeable.

bsuSmc-ScpAB-disrupting sybodies target two distinct coiled coil regions adjacent to the bsuSmc 4N arm-to-arm contact

To map the sybody-binding site on the bsuSmc-ScpAB complex, we utilized five chimeric Smc constructs (Smc Chimera 1-5), in which the hinge and progressively longer segments of the adjacent coiled coils were replaced with the corresponding sequences from the S. pneumoniae Smc protein (Figure 3A). Chimeric junctions were designed to preserve coiled coil integrity based on available crystal structures, coiled coil predictions by DeepCoil, and AlphaFold2 structural models (Zimmermann et al., 2018; Gabler et al., 2020; Jumper et al., 2021). All chimeric Smc strains retained the native B. subtilis scpA, scpB, and parB genes and were viable under conditions promoting fast growth, demonstrating proper protein folding and functioning of chimeric Smc-ScpAB complexes (Figure S4A).

Figure 3:

Figure 4:

Chimeras 1 and 2, which carry the central hinge and coiled coil sequences of Spn origin (residues 363-812 and 339-836, respectively), showed the same sybody-induced growth defects as the wild-type strain, indicating that the sybodies bind further away from the hinge, within the remaining B. subtilis portion comprising of the Smc head and the first ∼230 coiled coil residues, and ScpAB (Figure 3A). By contrast, Chimera 5 (residues 248–927 of Spn origin), supported robust growth even in the presence of the bsuSmc-disrupting sybodies, indicating that the region comprising the heads and first ∼50 residues of the coiled coil (including the joint) does not comprise the binding site. The binding region for the sybodies is thus located on the coiled coil between residues 248 and 339 (or 836 and 927), demonstrating that all fourteen sybodies bind to a central region of the coiled coil.

Chimera 3 (spnSmc residues 318-857) and chimera 4 (spnSmc residues 276-899) revealed two distinct sybody responses. Sb003, 031, 156, 160, 162, and 169 impaired growth of both strains, mapping their binding sites to B. subtilis residues 248-276 (and 899-927), present in chimeras 1–4 but absent in chimera 5. In contrast, Sb002, 006, 010, 015, 020, 021, 188, and 194 had no impact on the viability of chimera 3 or 4, suggesting they target residues 318-339/836-857 (Figure 3A, 3D).

In sum, these experiments mapped the binding sites of all fourteen disruptive sybodies to one of two ∼twenty amino acid segments in the central region of the bsuSmc coiled coil (Figure 3D). These regions harbor relatively poorly conserved sequences but flank the 4N arm-arm contact (∼at residue 295), previously proposed to serve as an essential structural switch between closed and open conformations (Vazquez Nunez et al., 2021). These results reveal that all isolated sybodies target the same functional element of the Smc arms, and that interfering with this element disrupts Smc-dependent chromosome organization in vivo.

Smc-disrupting sybodies affect the ATPase activity in one of two ways

To investigate how sybodies may influence bsuSmc-ScpAB, we next measured ATP hydrolysis rates using purified proteins in the presence and absence of DNA. SMC ATPase activity serves as an indirect readout for state transitions, including arm opening and head engagement. In absence of DNA, bsuSmc-ScpAB hydrolyzes ATP at ∼4 ATP/min/Smc (Vazquez Nunez et al., 2021). DNA binding stimulates ATP hydrolysis to ∼9 ATP/min/Smc, presumably by DNA binding at the hinge promoting arm opening and head engagement, creating a more open conformation.

All disruptive sybodies measurably affected the ATP hydrolysis rate of bsuSmc-ScpAB. Ten maintained a near-basal ATPase rate even in presence of DNA (Sb002, Sb003, Sb010, Sb031, Sb156, Sb160, Sb162, Sb169, Sb188, and Sb194) (Figure 3B-C and S4B). This suggests that they stabilize the complex in a closed or partially closed conformation, thus hindering DNA-dependent ATPase stimulation. Notably, in this group, Sb002 and Sb194 showed slightly elevated ATPase rates with DNA, hinting at a mildly more flexible conformation.

In contrast, the other sybodies (Sb006, Sb015, Sb020, and Sb021) stimulated ATP hydrolysis in the absence of DNA, with Sb006 reaching levels comparable to DNA-induced activation (∼10 ATP/min). This implies that these sybodies stabilize Smc-ScpAB in a more open state promoting head engagement. Curiously, addition of DNA in the presence of Sb006 and Sb021 reduced—rather than stimulated—ATPase activity (Figure 3B-C), suggesting that concurrent DNA and sybody binding may trap the complex in a non-productive conformation, perhaps by favoring the open state excessively. These findings are consistent with the idea that 4N contact region plays a critical role in arm opening and ATPase stimulation.

Interestingly, the observed ATPase profiles correlate with distinct binding regions along the Smc arms. All ATPase-stimulating sybodies (Sb006, Sb015, Sb020, Sb021) bind above the 4N contact (“Opening^UP”), while sybodies that hinder DNA stimulation fall into two subgroups: those binding above the 4N contact (Sb002, Sb010, Sb188, Sb194—”Closing^UP”) and those binding below it (Sb003, Sb031, Sb156, Sb160, Sb162, Sb169—”Closing^DOWN”) (Figure 3D). These results indicate that the sybodies perturb bsuSmc-ScpAB in two principal ways: either by favoring the opening of the arms or by constraining the arms in a more closed state. The results highlight arm opening and closure as central regulatory features of bacterial Smc-ScpAB (Soh et al., 2015; Minnen et al., 2016; Diebold-Durand et al., 2017; Vazquez Nunez, Ruiz Avila and Gruber, 2019) and confirm the key role of the 4N arm contact in balancing the opening and closing of SMC arms (Vazquez Nunez et al., 2021).

Discussion

Over the past three decades, sybodies and their synthetic counterparts have transformed biomedical research and structural biology by enabling the targeting and stabilization of transient protein conformations. Despite these advances, their application to probing protein function in living cells has remained limited (Seeger et al., 2012; Deneka et al., 2021). Here, we expand the utility of synthetic sybody selection by employing it as a tool to screen for genetic probes targeting the function of selected molecular machines in vivo. This strategy combines the advantages of controlled biochemical reconstitution with the ability to study the function of proteins in their native context. Importantly, sybody selection requires no prior knowledge of vulnerable regions, providing an unbiased means to interrogate protein function that is difficult to achieve through rational design. Our study based on the B. subtilis Smc-ScpAB DNA motor as a model uncovers a central regulatory region in the coiled coil arms of Smc and highlight how distinct arm conformations shape SMC function in loop extrusion and chromosome segregation.

Revealing Smc arm dynamics through synthetic binders

All fourteen Smc-disrupting sybodies share the loop library design. Moreover, their Smc binding sites map to the two short coiled coil segments flanking the conserved 4N arm-to-arm contact, a region previously implicated in key conformational transitions during arm opening (Vazquez Nunez et al., 2021). While the fourteen sybodies harbour different epitope binding sequences, we suspect that they may all bind to the Smc dimer in an analogous manner. With the long CDR3 loops known to bind narrow structural cavities (Rasmussen et al., 2011; Kruse et al., 2013), we speculate that these sybodies may intercalate between the two coiled coil arms (Figure 3D). Structural studies are needed to test this hypothesis directly, however, our attempts at X-ray crystallography and electron microscopy of Smc-sybody complexes have failed so far.

Notably, while the coiled coil dimer appears similarly structured from the elbow to the hinge (Diebold-Durand 2017) (Figure 3D), no Smc-disrupting sybodies mapped to regions further away from the 4N contact (Figure 3D). This indicates that opening and closure of the Smc arms might be dispensable at these places, although formal testing would require isolating binders efficiently targeting these regions, potentially facilitated by binder design, and expressing them in B. subtilis. This parallels with findings with the bacterial SMC defense system Wadjet, where hinge-proximal arm opening is dispensable for loop extrusion, but essential for obstacle bypass (Liu et al., 2025). Alternatively, these arm-to-arm contacts might be more stable and thus less likely to be interfered with by sybody binding. Similarly, sybodies targeting below the 4N contact all reduce the ATPase rate (or rather its DNA stimulation), indicating that artificial opening at this position may not disrupt Smc function (Figure S5). Again, these hypotheses need to be tested more directly. We envision that the design of protein binders may allow us to confirm these observations and further dissect the underlying mechanisms.

This work establishes sybodies as precision tools for blocking ATP-driven machines inside bacterial cells. By targeting allosteric control points in Smc, they enable mechanistic dissection of loop extrusion and open new avenues for studying dynamic complexes in vivo. More broadly, the study demonstrates how synthetic binders can trap, stabilize, or block transient conformations of active chromatin-associated machines, providing a powerful means to probe their mechanisms in living cells. Looking ahead, the rational design of protein binders with tailored geometry and allosteric potential could allow researchers to manipulate and visualize specific conformational states with even greater control and refinement, across both prokaryotic and eukaryotic organisms, while the generation and in vivo screening of sybodies remains an attractive approach to gain unexpected biological insights in an unbiased manner.

Material and methods

Bacillus subtilis strains construction and growth

B. subtilis strains used in this study were derived from the parental strain 1A700. Allelic replacement was achieved via double-crossover recombination at the endogenous smc locus using the natural competence method described by (Bürmann et al., 2013). Chromosome-integration of sybody genes were performed by transforming a vector containing an amyE homologous regions on each side of the recombinant gene of interest, as described in (Diebold-Durand, Bürmann and Gruber, 2019). Transformants were selected on ONA solid medium containing the appropriate antibiotics. Following transformation, strains were purified by single colony isolation and verified through a combination of marker testing, phenotype assessment, PCR, and Sanger sequencing of the smc locus, when appropriate. For spot assays, cells were cultured to stationary phase in liquid LB medium, and 9^-2 and 9^-5 dilutions were then spotted onto solid ONA medium with chloramphenicol (5 ug/mL) and xylose (0.5% w/v) when induction of the P_xyl promoter was required. The strains, primers and plasmids used in this study are listed in Table S3, Table S4 and Table S5, respectively.

For viability assessment, 200 LB medium was inoculated with a 1:1000 dilution of a dense B. subtilis culture and grown to stationary phase. The culture was then serially diluted in LB medium. From each dilution (9⁻¹ to 9⁻⁸), 5 µL was spotted onto LB agar plates containing the appropriate antibiotic selection. Plates were incubated, and viability was assessed by imaging the spots at 16- and 24-hours post-incubation.

Sybody selection

Sybody selection was performed following the protocol by (Zimmermann et al., 2020), with modifications to enrich binders specific for the ATP/DNA-stabilized “open” form of the bsuSmc-ScpAB complex.

Ribosome Display

The standard WTB buffer (50 mM Tris/acetate pH7.4, 150 mM NaCl, 50 mM MgAc2) and its derivatives were adjusted to 50 mM NaCl to facilitate bsuSmc-ScpAB binding to dsDNA. The target complex was assembled by mixing 500 nM bsuSmc(C119S, C437S, C826S, E1118Q, R643C)-ScpAB, 2 mM ATP, and 5 µM 40 bp dsDNA in WTB-D-BSA buffer (WTB including 0.5% (w/v) BSA and 0.1% Tween-20), incubated for 15 min at room temperature (RT). The ribosome display panning solution was supplemented with 2 mM ATP and 5 µM dsDNA to maintain binding conditions. Solution panning and biotinylated target capture were conducted at RT to optimize selection for the “open” complex. Washing steps incorporated 2 mM ATP and 5 µm 40 bp dsDNA, while elution, reverse transcription, and cDNA amplification followed the original protocol. Reverse transcribed RNA molecules were quantified by qPCR.

First round of phage display

Phagemid libraries were cloned and electroporated into E. coli SS320 as previously described (Zimmermann et al., 2020). Phage production was performed using M13KO7 helper phage. The phage display buffer (TBSM) consisted of 20 mM Tris-HCl (pH 7.4), 50 mM NaCl, and 2 mM MgCl2. Target preparation involved incubating 500 nM biotinylated bsuSmc(C119S, C437S, C826S, E1118Q, R643C)ScpAB with 2 mM ATP and 5 µM dsDNA in TBSM-BSA-D (TBSM including 0.5% (w/v) BSA and 0.05% Tween-20). Phages (10¹²/ml) were incubated with the target at 50 nM for 20 min at RT before immobilization on neutravidin-coated plates. Washing (with added 2 mM ATP and 5 µm 40 bp dsDNA) and elution otherwise followed the original protocol, with enrichment assessed by qPCR. Amplified phages were used for a second selection round.

Second round of phage display

Purified phages from the first round were used at 5 × 10¹³ phages/ml, and the target was prepared as in the first round. Phage-target binding occurred at 50 nM in the presence of ATP and dsDNA, followed by capture on magnetic beads. A competition step using non-biotinylated bsuSmc-ScpAB (1 µM) was included before washes with TBSM-D. Phages were eluted and enrichment was determined by qPCR. Phagemids were purified, sub-cloned into the pNb_init vector, and transformed into E. coli MC1061.

Growth curve analyses

Growth curves were generated from B. subtilis strains grown overnight to exponential phase in LB medium supplemented with 0.5% glucose, incubated at 30°C with shaking. The following day, fresh 5 mL LB cultures were inoculated with the overnight culture to an OD of 0.005 and incubated at 37°C with shaking until reaching an optical density (OD) of 0.05. Subsequently, each culture was subjected to a twofold dilution in a 96-well plate (Costar #3596), achieving a final volume of 200 µL per well. Where necessary, cultures were induced with 0.5% xylose. Plates were incubated at 37°C with continuous shaking in a Thermo Scientific Multiskan FC plate reader. Growth curves were determined by light scattering at 620 nm. The BactEXTRACT app was used to perform Analysis and visualization of the data (Dénéréaz and Veening, 2024).

Protein purification

Purification of bsuSmc and biotinylated bsuSmc(C119S, C437S, C826S, E1118Q, R643C)

bsuSmc proteins were purified according to (Bürmann et al., 2017). pET-22 or pET-28 plasmids encoding the Smc recombinant sequences were transformed into E. coli BL21-Gold (DE3) cells. Protein expression was carried out in ZYM-5052 autoinduction medium for 23 hours at 24°C. The cells were harvested and resuspended in lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, 10% (w/v) sucrose) supplemented with protease inhibitor cocktail. Cell lysis was achieved by sonication, and the lysate was clarified by centrifugation. The supernatant was filtered through a 0.45 µm membrane and loaded onto two HiTrap Blue HP 5 mL columns connected in series. Elution was performed using lysis buffer with 1 M NaCl. The main peak fractions were pooled and diluted with salt-free buffer (50 mM Tris-HCl, pH 7.5; 1 mM EDTA; 1 mM DTT) to a final conductivity equivalent to 50 mM NaCl (∼8 mS/cm). This diluted sample, supplemented with protease inhibitor cocktail, was applied to a HiTrap Heparin HP 5 mL column, and the proteins were eluted using a linear gradient up to 2 M NaCl. The main peak fractions (∼ 5 mL) were collected and subjected to further purification by gel filtration using an XK 16/70 Superose 6 PG column equilibrated with 50 mM Tris-HCl, pH 7.5, 200 mM NaCl, 1 mM EDTA, and 1 mM TCEP. The peak fractions were collected, concentrated using a Vivaspin 15 10K MWCO filter, flash frozen in liquid nitrogen, and stored at -80°C. Protein concentration was determined by absorbance, utilizing theoretical molar absorption and molecular weight values.

For the biotinylated bsuSmc(C119S, C437S, C826S, E1118Q, R643C) protein used in the sybody selection, the protein was first purified as described above, except that the reducing agent (TCEP) was omitted from the final gel filtration buffer. Labelling was performed by incubating 600 µL of bsuSmc(C119S, C437S, C826S, E1118Q, R643C) (83 µM in 50 mM Tris-HCl pH 7.5, 200 mM NaCl) with 1 mM PEG2-biotin maleimide for 10 minutes at 4 °C. The reaction was quenched by addition of 0.5 mM 2-mercaptoethanol. Excess label was removed using Zeba spin desalting columns (Thermo Fisher) in multiple parallel runs due to volume limitations. The final yield was 600 µL at 72.35 µM (subunit concentration). Labelled bsuSmc(C119S, C437S, C826S, E1118Q, R643C) was loaded onto a Superose 6 10/300 Increase column pre-equilibrated in 50 mM Tris-HCl pH 7.5, 200 mM NaCl. Peak fractions were pooled and mixed with ScpA and ScpB at a 1:1:2 molar ratio relative to the bsuSmc dimer. The final concentration of the full complex was adjusted to 5.7 µM. Complexes were aliquoted in 100 µL portions, flash-frozen in liquid nitrogen, and stored at −80 °C.

Purification of bsuScpA

ScpA was purified using the method described by (Vazquez Nunez, Ruiz Avila and Gruber, 2019). E. coli BL21-Gold (DE3) cells, transformed with a pET-28 derived plasmid encoding the ScpA protein, were used for expression. Cultivation was performed in ZYM-5052 autoinduction medium at 16°C for 28 hours. Cells were then harvested and resuspended in lysis buffer (50 mM Tris-HCl, pH 7.5; 200 mM NaCl; 5% glycerol) supplemented with protease inhibitor cocktail. The cells were lysed by sonication, and the lysate was clarified by centrifugation. The supernatant was applied to a 5 mL HiTrap Q ion exchange column and eluted with a gradient up to 2 M NaCl. Peak fractions were pooled and adjusted to a final concentration of 3 M NaCl by mixing with 4 M NaCl buffer. This mixture was loaded onto a HiTrap Butyl HP column and eluted with a reverse gradient to 50 mM NaCl. Eluted peak fractions were concentrated to 5 mL using Vivaspin 15 10K MWCO filters and further purified by size exclusion chromatography (SEC) on a HiLoad 16/600 Superdex 75 pg column equilibrated with 20 mM Tris-HCl, pH 7.5, and 200 mM NaCl. The purified protein was concentrated, flash frozen, and stored at -80°C.

Purification of bsuScpB

ScpB was purified following the protocol outlined by (Vazquez Nunez, Ruiz Avila and Gruber, 2019). The coding sequence of ScpB, cloned into a pET-22 derived plasmid, was transformed into chemically competent BL21-Gold (DE3) E. coli cells. These cells were cultivated in ZYM-5052 autoinduction medium at 24°C for 23 hours. Subsequently, the cells were harvested and resuspended in lysis buffer (50 mM Tris-HCl, pH 7.5; 150 mM NaCl; 1 mM EDTA; 1 mM DTT) supplemented with protease inhibitor cocktail. Cell lysis was performed by sonication, followed by centrifugation to remove cell debris. The resulting supernatant was diluted to a final NaCl concentration of 50 mM and loaded onto a 5 mL HiTrap Q HP column. Elution was achieved using a gradient up to 2 M NaCl. The eluate was then diluted with lysis buffer containing 4 M NaCl to achieve a final concentration of 3 M NaCl. This sample was applied to two 5 mL HiTrap Butyl columns connected in series, and the protein was eluted with a reverse gradient to 50 mM NaCl. The peak fractions from this column were concentrated and subjected to size exclusion chromatography (SEC) using a HiLoad 16/600 Superdex 200 pg column equilibrated with 50 mM Tris-HCl, pH 7.5, 100 mM NaCl, and 1 mM DTT. Fractions containing ScpB were concentrated, flash frozen, and stored at -80°C.

Medium scale sybody purification

Individual sybody plasmids were transformed into chemically competent E. coli MC1061 cells via heat shock at 42 °C for 45 s, followed by recovery in LB medium at 37 °C. Transformed cells were cultured overnight in TB medium supplemented with 25 µg/ml chloramphenicol. Precultures were used to inoculate 50 mL TB cultures, which were grown at 37 °C before shifting to 22 °C. Expression was induced with 0.02% (wt/vol) L-(+)-arabinose and continued overnight at 22 °C with shaking. Cells were harvested by centrifugation and resuspended in periplasmic extraction buffer. Following incubation at 4 °C, cells were pelleted, and the supernatant was supplemented with imidazole to 15 mM. The extract was incubated with His MultiTrap HP resin, followed by centrifugation and washing with TBS containing 30 mM imidazole. Elution was performed using TBS with 300 mM imidazole. Purified sybodies were analyzed on a Sepax SRT-10C SEC100 column at 1 ml/min flow rate. Monomeric sybodies eluted between 11–12.5 ml, while retention volumes <11 ml indicated oligomerization, and >14 ml suggested strong column interaction. Non-expressed, oligomeric, or highly interacting sybodies were discarded. Typical yields ranged from 200 µg to 1 mg (Zimmermann et al., 2020).

Fluorescence imaging of ParB-GFP strains Image acquisition

B. subtilis cells were first cultured in LB supplemented with 5 µg/mL chloramphenicol and 0.5% glucose at 30°C, from which a day culture was inoculated to 0.005 in LB with 5 µg/mL chloramphenicol. When needed, cultures were induced at OD₆₀₀ ∼ 0.02 with 0.5% xylose and grown an extra 30 minutes until imaging at an OD₆₀₀ of 0.04. For microscopy analysis, 0.5 μL of the cell suspension was spotted onto agarose-coated microscopy slides. Images were acquired using a Leica DMi8 microscope equipped with an sCMOS DFC9000 (Leica) camera, a SOLA light engine (Lumencor), and a 100×/1.40 oil-immersion objective. Exposure time for image acquisition was set to 600 ms. Acquired images were processed using LAS X Office software (v.1.4.7.28982, Leica Microsystems).

Image analysis

Microscopy images were analyzed using a fully automated image processing pipeline. Cell segmentation was performed with the pretrained Omnipose model (Cutler et al., 2022) implemented in Cellpose (Stringer et al., 2021), which is optimized for bacterial morphologies. Prior to segmentation, a normalization step was applied to standardize image contrast. Following segmentation, intracellular foci were detected using Spotiflow (Mantes et al., 2024), a dedicated model trained for spot identification. Each detected spot was assigned to an individual cell based on spatial location, enabling quantification of the number of foci per cell. Cell morphology analysis was also performed to estimate cell length.

Quantitative results were compiled into a structured dataset, including the number of cells per image, percentage of cells lacking foci, number of foci per cell, cell length, and number of foci per micron. Data visualization and statistical analysis was performed using GraphPad Prism 10.4.1 (627).

ATPase measurements

ATPase activity was assessed using the coupled pyruvate kinase/lactate dehydrogenase reaction as described by (Bürmann et al., 2017). ADP production was monitored over a period of 1 hour by measuring the absorbance changes of NADH at 340 nm. Data collection was performed using a Synergy Neo Hybrid Multi-Mode Microplate Reader. The reaction mixture comprised 1 mM NADH, 3 mM phosphoenolpyruvic acid, 100 U pyruvate kinase, 20 U lactate dehydrogenase, and varying concentrations of ATP. For assays requiring double-stranded oligonucleotides, a 40 bp oligonucleotide (5’-TTAGTTGTTC GTAGTGCTCG TCTGGCTCTG GATTACCCGC-3’) was added to a final concentration of 3 mM. The final protein concentration in the assay was 0.15 μM bsuSmc dimers in ATPase assay buffer (50 mM HEPES-KOH, pH 7.5; 50 mM NaCl; 2 mM MgCl2). All measurements were conducted at 25°C.

Data availability

All raw data will be made available at Mendeley Data DOI: 10.17632/k6k62p7z2s.1.

Supplementary figures

Figure S1:

Figure S2:

Figure S3:

Figure S4:

Figure S5:

Acknowledgements

We are grateful to Roberto Vazquez Nunez for the biotinylated bsuSmc(C119S, C437S, C826S, E1118Q, R643C) protein preparation, Arianna Ravera from the DCSR computing facility for help in image analysis and data extraction of the fluorescence microscopy images. Members of the Gruber Lab for critical feedback of the manuscript and for stimulating discussions.

Additional information

Funding

This study was supported by internal funding and by the European Research Council (Horizon 2020 ERC CoG 724482).

Author contributions

O.G. performed all in vivo experiments from sybody selection, fluorescence microscopy imaging, chimeric phenotypic screen. M.T. and L.H-H. performed the in vitro sybody screening (ribosome display, phage display, and first sybody purification). O.G. performed protein purifications, cross-linking and ATPase measurements, as well as data analysis, figures and writing. O.G. wrote the first manuscript draft. All authors reviewed and edited the manuscript. Funding acquisition and supervision, S.G.

Funding

EC | Horizon Europe | Excellent Science | HORIZON EUROPE European Research Council (ERC)

https://doi.org/10.3030/724482

• Stephan Gruber

Additional files

Table 1. Sybody enrichments at different steps of the selection procedure. Ribosome display output was quantified by qPCR, while phage display results include final phage titers and enrichment values from rounds one and two, also measured by qPCR.

Table S1. Sybody sequences.

Table S2. Percentage of cells lacking ParB-GFP foci, indicating the absence of chromosome due to segregation defect. In presence and absence of xylose for sybody induction.

Table S3, S4 and S5. Table S3: Strains. Table S4: Plasmids. Table S5: Oligonucleotides