Antiparallel stacking of Csu pili drives Acinetobacter baumannii 3D biofilm assembly

Domain: neuroscience
Published: 2026-02-07
Editorial outcome: minor_revision

Abstract
Many Gram-negative nosocomial pathogens rely on adhesive filaments, known as archaic chaperone-usher pili, to establish stress- and drug-resistant, multi-layered biofilms. Here, we uncover the mechanism by which these pili build three-dimensional (3D) biofilm architectures. In situ analyses of Acinetobacter baumannii biofilms using electron microscopy (EM) reveal an extensive network of ultrathin, flat stacks of archaic Csu pili interconnecting bacterial cells in 3D space. Cryo-EM structures of a single native pilus, pilus pairs, and two types of multi-pilus stacks show that the pili pack into antiparallel sheets, with their rods connected laterally by junctions at their zigzag corners. This antiparallel arrangement ensures that contacts form primarily between pili from interacting cells rather than pili from the same cell. With a remarkably short helical repeat, archaic chaperone-usher pili spontaneously establish a high density of junctions that determines the biofilm’s 3D architecture. Our findings may help develop new therapies against multidrug-resistant bacterial infections by targeting pilus-pilus interactions.

Introduction
The formation of multi-layered biofilms is an effective survival strategy for many bacterial pathogens1–3. Dense 3D biofilms shelter bacteria4 from anti-bacterial drug treatments and harsh environments5, while promoting horizontal gene transfer6 and development of resistance mutations through exposure to lowered drug doses7, often leading to poor bacterial eradication and chronic infections8,9. A variety of extracellular polymeric substances (EPS), including filamentous proteins, polysaccharides, and extracellular DNA (eDNA), have been found to play a role in 3D biofilm formation in different bacteria10. However, the biofilm matrix composed of these EPS is not easily amenable to high-resolution structural analysis, and therefore little is known about the mechanisms of bacterial cell-cell association and 3D biofilm architecture10.

Adhesive pili, or fimbriae, are hair-like surface appendages that mediate bacterial spread and colonisation of host tissues. In Gram-negative bacteria, most adhesive pili assemble from protein subunits via the chaperone-usher pathway (CUP)11–13. CUPs are subdivided into three groups comprising six major phylogenetic clades: alternative (α-fimbriae), classical (β-, γ-, κ-, and π-fimbriae), and archaic (σ-fimbriae)12. While classical and alternative CUPs are primarily found in β- and γ-proteobacteria, archaic CUPs are more prevalent and present across a wider range of phyla12.

Classical and alternative CUPs primarily facilitate initial, highly specific adhesion. In contrast, archaic CUP systems are associated with the establishment of mature 3D biofilms14–17. Among the most troublesome Gram-negative bacteria, Acinetobacter baumannii and Pseudomonas aeruginosa18,19 rely on archaic Csu and CupE pili, respectively, to develop stress and drug-resistant mushroom or mountain-like 3D biofilms on a diverse array of biotic and abiotic surfaces14–17,20. Although the central role of Csu and CupE pili in 3D biofilm formation has been firmly established through deleterious mutagenesis of structural or assembly genes from both archaic systems14–16, the underlying mechanism of this process remained elusive.

The initial surface attachment of A. baumannii is mediated by hydrophobic finger-like loops at the tip of the two-domain subunit CsuE21, which is displayed by adaptor subunits CsuA and CsuB at the tip of a long Csu pilus rod made of subunit CsuA/B22. Both Csu and CupE pili are assembled through the archaic CUP20,21,23,24 by donor strand complementation (DSC), in which the incoming pilin subunit inserts its donor strand into the hydrophobic groove of the preceding subunit, thereby completing its immunoglobulin-like fold25–29. Unlike the classical and alternative CUP pili that have either tube-like helical30–32 or flexible rods33, the Csu and CupE pili share a characteristic zigzag architecture with short 7-subunit and 5-subunit repeats, respectively, allowing them to act as super-elastic springs under tensile forces22,34.

Here, we elucidate the mechanism of biofilm maturation into multi-layered structures using a combination of scanning, negative stain and cryogenic electron microscopy of A. baumannii biofilms and isolated networks of Csu pili. We reveal that Csu pilus rods organise into flat antiparallel sheets linking neighbouring cells in three-dimensional space. Meanwhile, poly-N-acetylglucosamine (PNAG) fills the gaps within the Csu stack networks, collectively providing protection to the bacteria.

Results
Flat stacks of Csu pili interconnect A. baumannii cells in biofilms
Earlier studies of A. baumannii biofilms using scanning electron microscopy (SEM) have shown that bacterial cells are interconnected by Csu pili20,35. However, our analysis of SEM images indicates that the thickness of these ‘pili’ is often much greater than the width of a single Csu pilus rod filament determined by cryo-EM22. In the examples shown in Fig. 1, the apparent width is ~30 nm, which is ten-fold greater than that of a single pilus. This prompted us to examine the Csu pili on bacteria grown in liquid and on solid media at higher resolution using negative staining transmission electron microscopy (TEM) (Fig. 2).

A. baumannii 19096T translucent phenotype cells grown in a liquid medium display Csu pili that are mostly separate and pointed radially in every direction (Fig. 2a). In contrast, in colonies of these cells, most Csu pili stack upon each other, merging into flat sheets (Fig. 2c–e, Supplementary Fig. 1a‒c). This generates a network of stacks in the tight gaps of a bacterial colony, with relatively thin, local stacks that connect bacterial neighbours (Fig. 2g, i) merging into long and wide “superstacks” (Fig. 2f, h, Supplementary Fig. 1b, c) that interconnect multiple bacteria (Fig. 2b, Supplementary Fig. 1d) and essentially wrap around each bacterium, akin to a sheath (Fig. 2c, Supplementary Fig. 1d‒h). Stacks may contain more than 15 pili, with pili often forming lateral interactions extending over half of their length (Supplementary Fig. 1a‒c). Superstacks, continually incorporating new pili from the surfaces of multiple bacteria, can exceed 12 μm in length (Supplementary Fig. 1d‒h), clumping bacteria in 3D structures (Supplementary Fig. 2). These structures formed equally efficiently under both ambient and physiological temperatures (22‒37°C) and across a broad range of pH (6.0‒8.5) and salt concentration (42.8‒400 mM) (Supplementary Fig. 2).

To study the Csu pilus stacks in their native state, we analysed the suspended A. baumannii colonies using cryo-electron tomography (cryo-ET) (Fig. 3). 3D segmentation of reconstructed tomograms revealed individual pili stacking into flat, single-layer sheets (Fig. 3d–i, Supplementary Videos 1, 2). These sheets widen as they extend from the bacterial surface and merge into larger sheets, interconnecting multiple bacteria. Thus, altogether, our results suggest that Csu pili connect bacterial cells in A. baumannii biofilms through a complex 3D network of flat stacks.

Csu pilus–mediated bacterial interlinking and adhesion to surfaces depend on different structural elements of the pilus
While both piliated A. baumannii and csu-expressing Escherichia coli typically exist as single cells in liquid cultures, they occasionally form small clumps, resembling the initial stages of the formation of a surface pellicle36. To investigate the nature of these clumps, we analysed them using negative staining TEM (Supplementary Fig. 3). The cells within these clumps had interconnecting Csu pilus stacks with a similar architecture to the stacks seen in 3D biofilms. This suggests that prior adhesion to surfaces is not strictly necessary for Csu pili to link the bacteria together.

This observation was further supported by the detection of similar Csu pilus stack-tethered bacterial clumps (or rafts) in liquid cultures of E. coli strain expressing recombinant Csu pili rendered unable to facilitate bacterial adhesion to hydrophobic surfaces due to a mutant tip-subunit CsuE with a compromised hydrophobic tip-finger site21 (Supplementary Fig. 4a‒d). Similarly, extensive Csu pilus stacks formed in the induced colonies of the recombinant E. coli strain, regardless of whether it possessed the wild type or the mutant CsuE (Supplementary Fig. 4e, f). Collectively, these results indicate that Csu pilus–mediated surface adhesion and bacterial interlinking are distinct processes that depend on different structural elements of the pilus. Nevertheless, both processes are essential for 3D biofilm formation.

Csu pili spontaneously form a network of pilus stacks in vitro
To study the mechanism of Csu pilus stack formation, we isolated Csu pili from A. baumannii and recombinant E. coli cells and examined these samples at different time points using negative staining TEM. While freshly isolated samples mostly contained individual pili, with a small fraction of pilus pairs, samples incubated for over a week contained a significant portion of pilus pairs as well as a few multi-pilus superstacks (Fig. 4a, b). After one month, the samples formed a gel-like structure (Supplementary Fig. 5a), comprising a dense network of flat Csu pilus stacks (Fig. 4c, d). The structure of these stacks and the architecture of the network formed in vitro closely resembled those observed between bacteria in 3D biofilms (Figs. 1–3, Supplementary Figs. 1, 2) with the Csu pili arranged into flat stacks of variable width that gradually merge into central structures comprising tens of pili.

The striking similarity between the in vitro and in vivo networks, along with the progressing nature of their assembly observed in vitro, suggests that 3D biofilm formation is a spontaneous process inherently programmed within the structure of the pili.

Structure of Csu pilus stacks
Csu pili from A. baumannii may have post-translational modifications in regions that can potentially affect both Csu pilus architecture and stack formation. To explore this possibility, we determined the structure of isolated A. baumannii pili at 3.28 Å resolution using single particle (SPA) cryo-EM (Fig. 5a, Supplementary Fig. 6a–d, h and Supplementary Table 1). The native pilus rod has the same symmetry and clinch structure as the recombinantly produced pili (Supplementary Fig. 6d), supporting our previous findings22. The resolution and quality of the map were sufficient to rule out any repeated post-translational modifications, suggesting that stack formation depends on the inherent structure of the Csu pilus itself.

Pilus pairs, representing the minimal pilus stack, are common within the 3D biofilms. Their relative abundance in the examined in vitro pilus samples allowed us to select enough images for SPA cryo-EM structure determination. Since the pilus pair lacks helical symmetry, we generated an initial model based on the structure of the single pilus rod by optimising the overlap between its projections and a 2D class average (Supplementary Figs. 14–16). A filament-based SPA reconstruction yielded an 8.91 Å resolution density map, into which we fitted the model using rigid body real-space refinement (Fig. 5b, Supplementary Fig. 6e, i and Supplementary Table 1). The two Csu pilus rods form an antiparallel pair connected laterally by binding junctions at their zigzag corners (Fig. 6, Supplementary Video 3).

To determine the structure of multi-pilus stacks, we used a sample of isolated pili incubated for extended periods (Supplementary Fig. 5). We produced the cryo-EM maps of two alternative stacking architectures with the 7.58 Å map of type 1 stack showing three 18-subunit rod fragments, and the 4.70 Å map of type 2 stack showing four 17-subunit rod fragments (Fig. 5c, d, Supplementary Fig. 6f, g, j, k). Similar to the Csu pilus pair, the adjacent pilus rods in both types of stacks run antiparallel to each other, ensuring that contacts form primarily between pili from interacting cells rather than among the pili extending from the same cell (Fig. 6, Supplementary Videos 4, 5).

Although pilus pairs exhibit more flexibility with smooth bending, the structure of the rods and their contacts appear to be identical between the pair and the two types of stacks (Fig. 5b–d). Furthermore, these structures show only marginal differences from the structure of a single pilus rod (Fig. 5a), and to model the stacks, the helical twist of the native Csu pilus rod structure had to be adjusted less than 0.3°, representing less than a 0.2% change. This suggests that interacting pili can form contacts without significant induced fit in their overall structure, which may explain their high propensity to form stacks.

As with the single pilus, the Csu pilus rods in the stacks have a 188.66 Å long repeat with 3-turns-per-7-subunits symmetry, giving the pilus a heptagonal end projection with seven potential binding sites at 51.43° angles between them (Fig. 6, green and magenta arrows around the single pilus and pair, see also Supplementary Fig. 7a, b). The binding junctions in an interacting pilus pair form on the plane of their helical axes, which are 52.35 Å apart. Their bulk prevents a third pilus from binding at a 51.43° angle from the first junction (magenta arrows), but it can still bind at 154.29° and 102.86° angles (green arrows), corresponding to type 1 and type 2 stacks, respectively. The 154.29° angles make the type 1 stack slightly corrugated with 66.33 Å thickness and junctions at 12.86° angles to the underlying surface, whereas 102.86° angles generate the more corrugated type 2 stack, which is 84.58 Å thick with 38.57° angles to the surface (Fig. 6).

Both types of stacks consist of identical pilus rod pairs. However, adjacent pilus pairs are shifted relatively to each other by one subunit in type 1 stacks and three subunits in type 2 stacks, respectively (Fig. 6, side views). Consequently, in type 1 stacks, the binding junctions cluster into lanes perpendicular to the rod axes and separated by four uninvolved subunits. In contrast, junctions in type 2 stacks are more evenly distributed along the pilus axes, with each junction separated from the next by two uninvolved subunits on one side and one subunit on the other.

The binding junctions consist of two subunit pairs, one from each adjacent, antiparallel rod (Figs. 5–7). The 4.70 Å resolution density map for the type 2 stack (Fig. 7a) and the 3.28 Å resolution structure of a single native Csu pilus (Fig. 5a) enable accurate modelling of the main chain and most side chains involved in the contact (Fig. 7b). The junction exhibits two-fold symmetry due to the complementarity of the two identical sides. The binding junction consists of a central region with two Thr14 and two Gln142 residues forming hydrogen bonds, and of two distal regions with Thr66 forming hydrogen bonds with Asn87 and Gln142 as well as hydrophobic interactions with Thr88, Ala89, and Gly143. Additionally, Ser120 may also interact with Ala89, though, being part of the flexible D′′–E1-loop, its orientation is less restricted. The distal regions constitute a larger portion than the central region of the total interactive surface of ∼415.95 Å2 per junction. The charged residues around the binding site are too distant to form strong electrostatic interactions (Fig. 7c).

We next examined the contribution of individual residues to stack formation. Most mutations designed to have an effect at the pilus stacking interfaces also severely disrupted pilus assembly (Supplementary Table 3). However, the double mutant T66D–A89D, designed to introduce repulsive negative charges, and its variant T66D–ΔP68–A89D, featuring a shortened B″–C1 loop to position Asp66 closer to Asp89 and to prevent Asp66 from forming hydrogen bonds with the main-chain of the opposing Csu pilus (Supplementary Fig. 8a, b), were well tolerated. Although these mutations did not abolish antiparallel interactions in vivo, the more sensitive in vitro assays using purified pili showed that the mutants had smaller, less organised stacks with shorter continuous stacked regions compared to the wild type (Supplementary Fig. 8c, d), consistent with our structural data.

Sequence analysis of the interactive regions in CsuA/B subunits from 29 Acinetobacter species revealed two distinct groups (Supplementary Fig. 9a, Supplementary Fig. 10). The first group, comprising 18 sequences, differs from the A. baumannii CsuA/B sequence only by point mutations. The second group, consisting of nine sequences, all have a characteristic single residue deletion in the B′′–C1-loop and a two-residue deletion in the C2–D-loop, both central to the binding junction. Modelling the junction for a member of the first group, with several point mutations to its interactive residues, resulted in similar interactions (Supplementary Fig. 9b, c). In contrast, the conserved deletions and point mutations in the second group created a uniform 2 Å gap throughout the antiparallel junction in the models (Supplementary Fig. 9d). However, manually moving the rods closer led to a tight contact due to surface complementarity. Thus, despite relatively low sequence conservation of the interactive residues, the contacts between rods appear to be conserved at the structural level, consistent with the considerable resilience of the junction to mutagenesis.

For more distantly related archaic CUP pili, the modelling of antiparallel binding becomes more challenging due to the uncertainty in the angle between subunits forming the zigzag architecture. However, this was possible for CupE pili from P. aeruginosa, as the structure of a single CupE pilus rod has been determined (PDB: 8CIO)34. Like Csu pili16, CupE pili are necessary for P. aeruginosa to form mushroom-like 3D biofilms15. Furthermore, they also appear to form mesh-like arrays in vitro34. Interestingly, in the published cryo-EM density map (EMDB: EMD-16683) of a single CupE pilus rod, the C2–D-loop is linked to an unexplained density that disappears when the loop is mutated34. Its position indicates it could be part of an antiparallel CupE pilus included in the helical reconstruction, which allowed us to model the potential antiparallel junction (Supplementary Fig. 11b). Only a small adjustment to the helical twist (−1.33°, <1%) was necessary to generate the perfect 2-turns-per-5-subunits repeat capable of assembling into regular stacks (Supplementary Fig. 11c, d).

CupE has even shorter repeats (165.57 Å) than Csu pili (188.66 Å), as it consists of only five subunits, two of which our model predicts to be engaged in a binding junction (Supplementary Fig. 11b). Such unusually short repeats are possible only because of the zigzag architecture of the pilus, which generates a high density of binding junctions. For example, in a typical stack observed in vivo, a traced Csu pilus ran ~792 nm antiparallel to adjacent pili forming ~42 binding junctions with a predicted total interactive surface of around 17470 Å² (Supplementary Fig. 1b). Therefore, it is plausible that the archaic CU pilus zigzag architecture, which enables the formation of flat stacks, evolved to shape bacterial communities.

Poly-N-acetylglucosamine and eDNA embed pilus stacks in mature biofilms
To study the kinetics of pilus stack assembly in vivo, we performed SEM analysis of A. baumannii biofilm formation on glass slides at different time points. Stack formation was unexpectedly fast, progressing simultaneously with 3D biofilm growth, and the stack network was essentially complete within 24–72 h (Fig. 1). However, over the course of days, we observed the stack network gradually becoming embedded in a less structurally defined material (Fig. 8a–d). This observation is consistent with previous findings that A. baumannii biofilms tend to accumulate significant amounts of PNAG and eDNA14,16. Higher resolution analysis using negative stain TEM revealed a close association of the pilus stacks with this extracellular substance, which essentially enwraps the individual stacks (Fig. 8e). Gold-labelled wheat germ agglutinin (WGA), extensively used to detect bacterial PNAG capsules, specifically binds to this substance, suggesting that it indeed contains a significant amount of PNAG (Fig. 8f, g).

While most bacteria seen in our negative stain TEM samples formed dense clumps with a plenty of Csu pilus stacks on the surfaces and PNAG or eDNA filling the tight gaps (Fig. 9a–c, Supplementary Fig. 2), others formed loose, yet cohesive pilus-linked rafts with otherwise empty, wide gaps (Fig. 9d–f). The distinctive lack of PNAG and eDNA in these rafts indicates that these EPS contribute little to clump cohesion. To test this further, we grew A. baumannii AB5075 mutant pgaC::T26, which is unable to produce PNAG37, and treated suspended colonies with DNase I and the DNA-intercalating agent chloroquine to degrade eDNA38. Subsequent dilution and washing steps revealed several highly organised rafts, with tight, clear gaps and extensive Csu pilus stacks traversing them (Fig. 9g–i), further supporting the conclusion that Csu pilus stacks link the bacteria in early 3D biofilms independently of PNAG and eDNA.

Mechanism of Csu pilus-mediated 3D biofilm formation
Formation of pilus stacks in 3D biofilms is much faster than the assembly process observed in samples of purified pili, which may take months to complete. This suggests that something facilitates this process in vivo. We observed that in liquid cultures, the Csu pilus stacks are common at cell division zones as well as at the ends of bacterial rods, possibly as remnants of cell divisions (Supplementary Fig. 3c–e). During cell division, the cell membrane grooves inward around the septum, positioning Csu pili around the septum antiparallel to each other, which seems to promote pilus stack formation (Fig. 10). Connecting daughter cells by pilus stacks is likely a major step in the formation of a 3D biofilm, allowing the stacks to form simultaneously with bacterial growth (Supplementary Video 6).

Although this process can occur both in liquid and solid media, the formation of stacks linking bacteria together is dramatically more efficient in 3D biofilms than in liquid cultures. A. baumannii in biofilms produce more pili than liquid culture cells39, but since clumps are also rare in E. coli liquid cultures overexpressing Csu pili, the higher number of pili is unlikely to be the main factor here.

Potentially, bacterial attachment to surfaces can promote subsequent formation of 3D biofilms, even though these two processes can occur independently, mediated by different parts of the pili (Fig. 10, Supplementary Fig. 4e, f). First, the stacks formed between daughter cells attached to a surface would experience lower levels of turbulence, therefore preserving the contacts. In contrast, A. baumannii cells growing in a liquid medium are less likely to clump, probably because turbulence, moving both Csu pili and the cells around, inhibit pilus stack formation and breaks most pairs that form (Supplementary Figs. 3, 4 and 12). Second, the narrow gaps between neighbouring cells in a 3D biofilm help position Csu pili antiparallel to each other, guiding the formation of extensive Csu pilus stacks (Figs. 9, 10). Such extensive stacks between daughter cells and neighbouring cells could help maintain their spatial orientation, allowing the biofilm to grow taller without breaking apart. In liquid cultures, such neighbouring cells are most often absent (Supplementary Fig. 12), unless the liquid culture is stagnant, making its conditions similar to solid media and allowing the formation of a surface pellicle with CsuA/B as its most abundant protein component36. Finally, our results show that 3D biofilms on solid media accumulate PNAG, eDNA, or other secreted materials that fill the narrow gaps between the cells, further fixing cell positions and stabilising the pilus stack network (Figs. 8, 10).

Methods
Production and purification of Csu pili from liquid cultures
Csu pili were extracted from either E. coli BL21-AI (F− ompT hsdSB(r − m −) gal dcm araB::T7RNAP-tetA; Invitrogen) transformed with the pBAD-Csu plasmid21 or the Acinetobacter baumannii CCUG 19096T strain53, which we selected for the translucent phenotype54, as it is generally associated with a higher number of pili35. The BL21-AI preculture was grown overnight in LB medium containing 100 μg/ml ampicillin at 37°C with shaking at 225 rpm. Subsequently, 7 ml of this preculture was inoculated into 300 ml of LB medium with 100 μg/ml ampicillin and grown with shaking at 225 rpm, until an optical density (OD) at 595 nm of around 1.0 was reached. The shaking speed was then reduced to 150 rpm, and Csu pilus production was induced by adding 0.2% (w/v) L-arabinose for an additional 2.5 h. For the A. baumannii cells, cultures were grown without antibiotics in 300 ml of LB medium at 22°C with shaking at 150 rpm for 26‒28 h. The same procedures were applied when growing liquid cultures for negative stain transmission electron microscopy (TEM), except that liquid cultures used to observe bacterial clump formation were left stagnant for around 1 h at 22°C before taking samples for TEM.

The extraction of Csu pili followed the same protocol for both sources. Cells were first centrifuged at 15,971 × g for 15 min. The resulting pellet was resuspended in 3.6 ml of 20 mM HEPES, 75 mM NaCl (pH 7.4), and the Csu pili were extracted by heating the suspension to 65°C for 60 min. Following this, the mixture was centrifuged at 17,700 × g for 15 min, and the supernatant was collected and dialysed against 20 mM HEPES, pH 7.4. The sample was then filtered using either a 0.22 μm or 0.45 μm filter and applied to a column packed with 20 ml of Source 30Q (Cytiva), using either the Äkta Purifier UPC10 or Äkta Pure (Cytiva). Csu pili were eluted at a flow rate of 0.9 ml/min using a 50% gradient of 1.0 M NaCl in 20 mM HEPES, pH 7.4, over 50 min. Fractions with the highest concentrations of CsuA/B and least impurities were selected based on SDS-PAGE analysis and used without further purification.

Growing A. baumannii and recombinant E. coli colonies for Csu pilus stack analysis
We initially cultivated A. baumannii CCUG 19096T strain colonies using a ten-fold dilution series to achieve approximately 100–200 colonies per 0.5x LB agar plate (composed of 2.5 g NaCl, 5 g tryptone, 2.5 g yeast extract, and 8 g agar, pH 7.4, in 1 L of distilled water) to ensure the manifestation of the translucent phenotype. We later observed that even seemingly opaque colonies had enough phenotype-switched cells to produce large pilus-linked clumps of bacteria. For the early negative stain transmission electron microscopy (TEM) experiments, the bacteria were grown for 16–20 h at 37°C until the colonies reached a diameter of approximately 1.0–1.5 mm, and then stored at 8°C to keep them from growing further. Under this condition, colonies could be stored for 2–3 weeks, and remained usable even after three months, with the only apparent change being the slow accumulation of amorphous materials, presumably PNAG or eDNA. For the later TEM experiments using A. baumannii AB5075 strain colonies and for cryo-ET using A. baumannii CCUG 19096T strain colonies, we grew them at 22°C for three days before using them immediately for the experiments without any storage. Finally, for the TEM experiments focusing on stack formation in variable growth conditions, one variable was growing them at 30°C or 37°C for less than 18 h before preparing the grids. The other growth condition experiments grew for three days at 22°C, but had pH adjusted to 8.5 or 6.0 instead of 7.4, or NaCl concentration of 200 mM or 400 mM instead of 42.8 mM. The assembly of Csu pili in recombinant E. coli colonies was induced by growing them for 15–17 h at 37°C in a low concentration of L-arabinose (0.001% w/v) to limit the number of Csu pili produced.

Single-particle cryo-EM
To determine the structure of the native Csu pilus rod, 3.5 μl of sample purified from A. baumannii CCUG 19096T strain was applied to glow-discharged 300-mesh gold holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified using a Vitrobot Mark IV (Thermo Fisher Scientific) at 4°C and 100% humidity. The data were collected on a Titan Krios transmission electron microscope (Thermo Fisher) operating at 300 kV equipped with a K3 direct electron detector (Gatan) in counting mode with a BioQuantum GIF energy filter (slit width of 20 eV) at a magnification of ×81,000 corresponding to a pixel size of 1.08 Å at the specimen level. 60-frame videos with a dose rate of ~15 electrons per pixel per second and a total accumulated dose of 54.8 e-/Å2 were collected using the Latitude-S (Gatan) single-particle data acquisition programme. The nominal defocus values were set from −0.8 to −2.5 μm.

To determine the structure of the Csu pilus pair and the initial structure of a Csu pilus stack, 4 μl of sample purified from E. coli BL21-AI with pBAD-Csu plasmid was applied to glow-discharged 300-mesh copper holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified in liquid ethane using a Vitrobot Mark IV (Thermo Fisher) at 4°C and 100% humidity. The data were collected on a 300 kV Titan Krios electron microscope (Thermo Scientific) equipped with a Gatan K2 Summit direct electron detector operated in super-resolution mode with a pixel size of 0.851 Å and a defocus range of −0.8 to −2.5 μm. A total dose of 50 e-/Å² was applied and equally divided among 40 frames to allow for dose weighting.

To determine the final Csu pilus stack structures, we used highly concentrated fractions of Csu pili, which we purified from E. coli BL21-AI with the pBAD-Csu plasmid and then stored at 4°C for over a year. The sample comprised an upper liquid phase and a lower gel-like phase containing Csu pilus stacks (Supplementary Fig. 5a). 3 μl of the gel phase was applied to glow-discharged 300-mesh copper grids with a carbon film (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified using a Vitrobot Mark IV (Thermo Fisher Scientific) at 4°C and 100% humidity. The data were collected using a 300 kV Titan Krios electron microscope (Thermo Fisher Scientific) equipped with a Gatan K3 direct electron detector in super-resolution mode, using a pixel size of 0.825 Å and a defocus range of −0.4 to −1.6 μm. After manually picking the grid holes (Supplementary Fig. 5b–d), we used automated cryo-EM data collection utilising EPU v3.6.0 collection software (Thermo Fisher Scientific). The total dose was 59.848 e-/Å², split evenly across 45 frames, giving each frame a dose of 1.330 e-/Å².

Cryo-EM image processing and reconstruction
The movie frames of the native Csu pilus rods were subjected to beam-induced motion correction using Patch Motion Correction in CryoSPARC v4.5.355 followed by determination of Contrast Transfer Function (CTF) parameters in Patch CTF. Micrographs with CTF fit better than 3.5 Å were used for further analysis. For the native Csu pilus rod, the particles picked with Filament Tracer (189,846 particles) were subjected to 2D classification followed by Ab Initio model generation with two classes. The best class from Ab Initio (50,502 particles) was used for non-uniform refinement, local motion correction and reference-based motion correction, generating a map with a global resolution of 3.28 Å (Supplementary Fig. 17a).

The movies of the Csu pilus pair were subjected to beam-induced motion correction using MotionCor256 followed by determination of CTF in CTFFind-4.157 in RELION 3.058. Micrographs with CTF fit better than 3.5 Å were used for filament picking. The filament pairs were manually picked in RELION (35,938 particles), extracted with a box size of 300 pixels and imported to CryoSPARC. The particles were then subjected to 2D classification, followed by Ab Initio model generation and homogeneous refinement. The final particle stack (12,883 particles) generated a map with a global resolution of 8.91 Å (Supplementary Fig. 17a).

The movie frames from the initial test set of Csu pilus stacks were processed using Patch Motion Correction and Patch CTF in CryoSPARC v4.5.355 for beam-induced motion correction and CTF estimation. Micrographs with a CTF fit better than 3.5 Å were selected for particle picking using Filament Tracer, yielding 136,628 particles. Only about a quarter of the filaments represented pilus stacks, with the remainder corresponding to single pilus rods. The particles were then subjected to 2D classification followed by Ab Initio model generation and homogeneous refinement, generating an 8.5 Å map (Supplementary Fig. 17a) that served as the initial model for processing a larger Csu pilus stack dataset (Supplementary Fig. 17b‒h).

The movies from the larger dataset of Csu pilus stacks were subjected to beam-induced motion correction using Patch Motion Correction in CryoSPARC v4.5.355, followed by determination of Contrast Transfer Function (CTF) parameters in Patch CTF. In Manually Curate Exposures, we rejected 736 and left 7205 exposures. Filament tracer gave 32,866 particles after removing bad picks with Inspect Picks, Extract Micrographs and two rounds of 2D Classification and Select 2D Classes. The best 2D classes were used in template-based picking, which resulted in 367,173 particles after removing bad picks.

Homogeneous Refinement using our initial 8.5 Å map of the type 1 stack revealed that there are two types of stacks. We produced models of the two stacks to create 16 Å simulated maps in UCSF Chimera59, and used ChimeraX60 and Volume Tools in CryoSPARC to produce masks for two Homogeneous Refinements. Using the resulting maps to sort the particles in Heterogeneous Refinement revealed a need for more particles. A second round of template-based picking gave two mixed particle sets, which we sorted with three pairs of Heterogeneous Refinement using the previous two Homogeneous Refinement maps, and every time, combining the particles sorted for one type of stack in one group. This resulted in 710,036 particles for type 1 and 924,343 for type 2 stacks after the Remove Duplicate Particles jobs.

Both types of stacks had issues with pseudosymmetry due to pilus helical symmetry, and stacks always resembling a row of pillars, even if flipped 180° in any direction. This caused the binding junctions to disappear as the increasing number of flipped particles biased the search maps. Remnant particles representing the wrong type of stack further exacerbated the issue. We reduced the issue for the type 1 stack by splitting the particles into seven sets, refining each with Homogeneous Refinement, and excluding two sets with the weakest binding junction densities. The main solution to the pseudosymmetry issue was rotating a 16 Å simulated map 180° in every direction to generate seven search models to sort the particles in Heterogeneous Refinement based on their orientations. This required having few contaminant particles and deliberately lowering the O-EM learning rate, so that occasional flipped particles would not bias the maps. Re-orienting the search models with their sorted particles back to the initial orientation and running Local Refinement with the combined dataset, while limiting particle rotation, yielded the final maps. The Local Refinement for type 1 stack used 504,940 particles. We used a type 1 stack simulated map also as a decoy class, when sorting the particle orientations for type 2 stack, reducing its final number of particles used for Local Refinement to 687,088 (Supplementary Fig. 17b‒h, Supplementary Table 1).

Model building and refinement
To construct the model for the native A. baumannii Csu pilus rod, we fit subunits from our previous recombinant Csu pilus rod structure (PDB: 7ZL4) into the density map using the Fit in Map tool in UCSF Chimera. First, as previously22, we adjusted the angle between two subunit models to match the average angle between two subsequent subunits in multiple positions along the pilus rod density map. Due to the map voxel size, this still left a small gap between the subunits, so in the second step, we shifted the two subunits closer in Coot 0.8.9.261 to link the lower subunit to its donor strand, and used the shifted pair to adjust the map voxel size in Chimera. We iterated these two steps until subunit positions stabilised. After further manual adjustments in Coot, the structure was refined using the rigid body and per-residue B factor real space refinement with PHENIX v1.20.1-448762.

The native pilus structure served as the basis for building models of the Csu pilus pair and both types of stacks, with minor adjustments to the helical twist. The angle between rod pairs in all structures was determined based on our overlap analysis with a 2D class (Supplementary Figs. 14–16, Supplementary Note 1). The binding junction was modelled using the 4.70 Å map of the type 2 stack, and this structure was replicated for the Csu pilus pair and the type 1 stack, as the experimental maps for these structures had lower resolution. The structures were refined as described above for the native Csu pilus structure. Refinement statistics are shown in Supplementary Table 1.

We determined the interactive surface of a binding junction using the CCP4i implementation of AREAIMOL63 by first calculating the sum of individual surfaces from two antiparallel pilus fragments and then reducing from that the surface of the fragments combined in a single model.

Cryo-ET data collection and processing
For cryo-ET sample preparation, A. baumannii colonies were suspended in a solution containing 10 nm colloidal gold beads (Sigma-Aldrich, Australia) pre-coated with 1% BSA. The mixture was applied to glow-discharged R2/2 Quantifoil holey carbon grids (Quantifoil Micro Tools GmbH, Jena, Germany). Freezing was performed using Vitrobot (FEI Thermo Fisher Scientific), maintained at 100% humidity. Excess liquid was removed with Whatman filter paper, and the grids were plunge-frozen in liquid ethane. Imaging was conducted on the Titan Krios G4 cryo-EM system equipped with a Gatan energy filter and a K3 Summit direct detector at an acceleration voltage of 300 kV. Tilt-series were acquired with Tomography 5 software version 5.14 (Thermo Fisher Scientific) over a tilt range of −45° to 45° (3° increments), with a cumulative electron dose of 120 e−/Å2, a defocus of −8 μm, and a pixel size of 3.39 Å.

Three-dimensional reconstructions of tilt-series were performed using the IMOD software package64. After binning the tilt series to 1k, alignment was completed using IMOD, followed by SIRT-based reconstruction using Tomo3D65. To enhance interpretability, the tomograms were deconvolved using IsoNet version 0.266.

Tomogram segmentation
For the better visualisation of the tomograms, Dragonfly software, version 2022.2 (https://www.theobjects.com/dragonfly/index.html) was used for 3D segmentation. Tomograms were initially filtered using built-in filters to improve their clarity. Manual segmentation was conducted on 15 to 20 slices from individual tomograms, which were then used to train a neural network employing the U-Net architecture with 2.5D input dimensions (5 slices)67. The trained model was subsequently applied to segment the full tomograms, with manual corrections implemented as required. 2D images and 3D movies were generated using the built-in functions of the Dragonfly software.

Negative stain transmission electron microscopy
All samples (4 μl) were prepared on Formvar/Carbon 200 mesh copper grids (Ted Pella, Inc.), glow-discharged with either a Tergeo Plasma Cleaner (Pie Scientific LLC) using water vapour or GLOQUBE plus (Quorum) using air.

Samples of the extracted Csu pili taken from the upper liquid phase (see above) were, in some cases, diluted 1:10 with Milli-Q water, depending on the pilus concentration. Stacks at the bottom of the tube were partially resuspended with a pipette tip before pipetting onto the grid undiluted. The grids were incubated for 1 min for diluted samples and 2 min for undiluted samples. Grids with undiluted samples were washed with 4 μl of Milli-Q water for 30 s to remove salts. For bacterial liquid cultures, a 1:5 dilution with Milli-Q water was prepared immediately before applying the samples to the grids, followed by a 2-min incubation. For the initial colony samples, a piece of a colony measuring 1.0 to 1.5 mm in width was picked using a 1 μl loop and suspended in 20 μl of Milli-Q water by spinning the loop for 1‒2 min. For later samples, including all the A. baumannii AB507568 samples, the spinning time was only 15‒20 s. When assessing for the presence of PNAG in A. baumannii colonies, undiluted Triticum vulgare lectin (WGA), colloidal gold conjugate (15 nm, CD BioGlyco) was used for suspension instead of Milli-Q water. The samples were then added to the grids and incubated for 2 min. When testing Csu pilus stack stability in the absence of PNAG and eDNA, colonies of A. baumannii AB5075 pgaC:T26 mutant lacking PNAG were incubated for 1 h in 20 μl of 25 mM Tris-HCl, 6.25 mM CaCl2, 1 mM MgSO4, pH 7.5 with added 1 mg/ml DNase I grade II from bovine pancreas (04716728001, Roche) and 5 μM chloroquine (C6628-25G, Sigma-Aldrich). Chloroquine had the purpose of changing eDNA in Z-DNA conformation into B-DNA that DNase I can break. To reduce DNase I concentration after the reaction, each reaction mixture was diluted with 20 μl of Milli-Q water, then added to the grids in 4 μl volume and incubated for 2 min, before washing the grid for 30 s with 4 μl of Milli-Q water.

All samples were stained with 4 μl of 1% (w/v) uranyl acetate for 1 min. Images were recorded on a 120 kV JEM−1400 Plus transmission electron microscope (JEOL Ltd.) equipped with an OSIS Quemesa 11 Mpix bottom-mounted digital camera.

Scanning electron microscopy analysis of A. baumannii 3D biofilms
Acinetobacter baumannii clinical isolates expressing Csu pili involved in biofilm formation (A. baumannii A10035, A. baumannii AB507568, A. baumannii ATCC 19606T)53 were cultured on LB agar at 30°C with duration as mentioned in the respective figure legends. Pieces of agar containing bacterial colonies were sampled and fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate overnight at 4°C. The specimens were subsequently dehydrated in a graded series of ethanol treatment and coated with 5 nm gold/palladium as described earlier35. Images of the bacterial cells were obtained with a field-emission scanning electron microscope (Carl Zeiss Merlin FESEM) using secondary electron detectors at an accelerating voltage of 4 kV and probe current of 50–100 pA.

Phylogenetic analysis of CsuA/B subunits from different Acinetobacter species
For the analysis, we used 29 sequences of the CsuA/B subunit from various Acinetobacter species, along with nine homologous sequences from different genera as an outgroup. Secretory signal peptides predicted using the SignalP 5.0 server69 were removed. The sequences were aligned using the MUSCLE algorithm in MEGA 1170, with parameters set to a −3.50 gap open penalty, −0.30 gap extend penalty, 1.50 hydrophobicity multiplier, and 18 maximum iterations. These settings effectively aligned the invariant disulphide pair and the other highly conserved residues and hydrophobic sites.

We constructed a maximum likelihood phylogenetic tree using MEGA 11, applying the Whelan and Goldman + Freq. model71. The initial tree was built by employing the Neighbour-Joining and BioNJ algorithms on a matrix of pairwise distances derived from the JTT model71, selecting the topology with the highest log-likelihood value. To model evolutionary rate differences, we used five discrete Gamma categories, with some sites treated as evolutionarily invariable. The resulting tree features branches supported by over 50% of the 3000 replicates. For further details, see Supplementary Figs. 9a, 10.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Malmi et al. report a novel mechanism contributing to biofilm formation in Acinetobacter baumannii, a common pathogen in developing countries associated with various human infections, including pneumonia as well as skin and urinary tract infections. A. baumannii relies on its abundant Csu pili for adherence to epithelial cells and subsequent biofilm formation, making them a major virulence factor in disease progression. While the structure of Csu pili has been recently determined— revealing a highly elastic, zig-zagged assembly with regularly spaced clinches of unknown biological significance—the precise mechanism by which they promote biofilm formation has remained unclear.\nIn this study, Malmi et al. use a combination of scanning and transmission electron microscopy to propose that antiparallel stacking between adjacent Csu pili is the primary driver of biofilm formation in A. baumannii. Although the study lacks in vivo data to support this hypothesis, the authors employ state-of-the-art cryo-electron tomography and microscopy to provide structural evidence of stacking in in vitro purified Csu pili samples following prolonged incubation at low temperatures. Their findings suggest that the clinches identified in the atomic structure of Csu pili play a critical role in the reported stacking.\nOverall, while the structural data strongly suggest the proposed stacking mechanism, without appropriate in vivo validation, this conclusion cannot be fully substantiated. Therefore, in its current form, the manuscript is not yet suitable for publication.\nIf in vivo evidence supporting the structural findings were provided, the study would represent a significant and novel contribution to understanding biofilm formation and pathogenesis in A. baumannii. Under those conditions, I would recommend the manuscript for publication given its potentially high impact and relevance.\nMajor Comments:\nFirstly, the absence of mutational validation is particularly striking given that Figure 6 identifies specific residues involved in stacking. This validation is essential to confirm the role of Csu pili stacking in A. baumannii biofilm formation and to disentangle its contribution from other polar effects, such as PNAG or eDNA accumulation. Mutants should be analysed via electron microscopy alongside biofilm formation assays to determine the precise impact of stacking. Furthermore, since the authors propose a similar mechanism for CupE pili, a comparable experimental approach would strengthen the claim that stacking is a widespread feature of archaic chaperone-usher pili, thereby enhancing the study’s significance. Moreover, due to the high impact and novelty of the findings, I encourage the authors to consider fluorescence imaging of the pili to show the establishment and progression of the Csu stacking.\nSecondly, the authors’ claim that stacking is the ‘key’ mechanism of biofilm formation is not fully supported, as the extent of pili-pili interactions remains unclear. While the authors demonstrate that stacking occurs significantly in vitro under prolonged low-temperature incubation, the SEM images (Fig. 1) suggest that stacking contacts are limited to short stretches rather than spanning the entire pilus length. Consequently, it is uncertain how much these interactions contribute to in vivo biofilm formation. As the authors suggest, the presence of PNAG and eDNA likely reinforces these interactions, the primary role of Csu pili in biofilm formation could be solely to sterically trap large amounts of PNAG and eDNA rather than stacking with itself. To clarify this, the authors should generate a non-stackable Csu mutant (as mentioned previously) to determine whether stacking is the ‘key’ or rather a ‘supporting’ mechanism. Another approach would be to assess biofilm formation in the presence and absence of PNAG/eDNA using enzymatic digestion. This would allow for the isolation of pili-pili interactions, helping to determine whether stacking is indeed the key mechanism or a secondary effect.\nSpecific comments\nL71-73: The authors state that the measured thickness of Csu pili is ~30 nm (Fig. 1). However, it is evident from the image that this thickness likely results from the bundling of filaments rather than individual pili. Could the authors clarify how these measurements were obtained? How many bundles were considered? Was there a range of values? Presenting this data as mean ± SD would enhance transparency, particularly given that this observation prompted the study.\nL76: The authors base their in vivo observations on a translucent phenotype, which is generally associated with increased pilus production. To ensure that the observed stacking is not an artefact of higher pili numbers, could the authors provide similar images to those in Figure 2 for a wild-type A. baumannii strain?\nL77-81: Authors show that stacks are not formed effectively in liquid cultures and are present in colonies. Would that not indicate that it is rather the accumulation of ECM (possible only under colony growth and not in liquids) that is primarily responsible for pili bundling and cell aggregation rather than the inherent pili-pili interaction which should be capable of forming even in liquid environments?\nL82-87: In the cryo-ET results (Fig. 3), there appears to be no continuity between pili from neighbouring cells to confirm that stacks are intercellular rather than intracellular, suggesting that the shown stacks originate from the same cell. Given the claims of exclusively anti-parallel stacking, how do the authors reconcile this observation?\nL99-101: The authors state that a CsuE mutant with a compromised tip finger site impairs adhesion but does not affect Csu pili stacking, and consequently, biofilm formation remains unaffected. This statement requires quantification—biofilm formation levels should be explicitly compared to support the claim of unaltered levels.\nL112-115: The progressing nature of assembly has been only shown at low temperatures in vitro and therefore should not be mentioned confidently unless authors could show it through in vivo imaging. Moreover, the assertion that the biofilm-like structures observed in vitro indicate that biofilm formation is a “spontaneous process inherently programmed within the structure of the pili” is strong but lacks consideration of alternative explanations. Could environmental factors, such as salinity or pH, also contribute to stack formation? Testing whether stacks still form under high ionic strength conditions would help exclude non-specific interactions driven by low ionic strength. Additionally, is stacking reversible under certain environmental conditions?\nL135-141: During grid preparation and freezing, did the authors observe any dependence of stacking on ice thickness? Could the thin ice required for SPA induce or perhaps also limit the formation of Csu pili stacks?\nL189-202: I strongly encourage the authors to provide mutational validation for their observations regarding CupE pili. This would significantly strengthen the novelty of their claims, allowing for extrapolation of their findings to all archaic pili based on experimental data, increasing impact.\nL204: The claim that PNAG embeds pilus stacks in mature biofilms is based on WGA labelling. To confirm this, could the authors treat samples with PNAG-hydrolyzing enzymes and analyze the effect via TEM imaging? Additionally, how do biofilm levels compare in the presence and absence of this enzyme? Could authors discuss the possibility that trapping PNAG and eDNA is the main involvement of Csu pili in biofilm formation rather than a side effect of stacking.\nL237-240: Are there any documented differences in the production of Csu pili in liquid vs biofilm growth that could support this?\nL251-252: Has it been tested—either in this study or elsewhere—whether non-zigzagged pili are unable to form gel-like structures after prolonged incubation at low temperatures? If not, this statement should be softened.\nL269-270: Structurally, if one site on the pilus rod is occupied by another to form a stack, why would the remaining binding sites become inaccessible for another junction? Can the authors confidently exclude the possibility of three-dimensional stacking instead of exclusively flat stacking? Is there any in vivo evidence supporting the formation of flat connections rather than thick, circular bundles? If flat stacks were the predominant arrangement, wouldn’t they be visible in SEM images? While the cryo-ET data suggests flat stacks, without subtomogram averaging, this conclusion remains uncertain.\nL233-236: The authors suggest that increased turbulence explains why liquid cultures do not exhibit the same levels of Csu-mediated biofilm formation. Could they provide evidence demonstrating the impact of growth conditions on Csu stack formation in vitro? Additionally, could they visualize the effect of turbulence on pre-assembled Csu stacks?", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "I. Overall review:\nThe archaic chaperone usher Csu pili formed by Acinetobacter baumannii are known to play essential roles in the formation of biofilms. Csu pili have been previously shown to biofilms on abiotic surfaces as well as biofilm pellicles and play crucial roles in adhesion to mammalian tissues. This study by Malmi et. al uses a variety of electron microscopy techniques (SEM, Negative stain TEM, Cryo-ET, Cryo-EM, and gold conjugation labeling) to investigate the role that Csu pili play in biofilm formation at multiple length scales from the micro/nanoscale of SEM and cryo-ET to sub-nanometer and atomistic details provided by single particle cryo-EM. Using these length scales the authors are able to strongly suggest how the Csu pili are involved in biofilm formation. The cryo-EM structural details of the pilus pairs and stacks, while low-resolution, are able to show novel anti-parallel packing interactions between pili. When attached to cells these anti-parallel interactions would presumably mostly occur between different cells which is plausible. Upon examination of the atomic models and cryo-EM density maps, I am confident in the authors’ interpretation of most of their cryo-EM structures. Any concerns are noted in the next section of my review. With all of this data the authors propose a plausible model for Csu pili-mediated biofilm formation.\nOverall, this study is rigorous, well-written, well-presented, and should have appeal to a broad scientific readership. Upon sufficient response to my concerns from the authors, I think this paper would be an excellent fit for publication at Nature Communications. I have several points which I would strongly like to see the authors address as well as some minor suggestions that could help improve their story/results.\nII. Major points to be addressed:\n1. In the Methods on page 16 lines 30-31 it is stated: “Homogeneous Refinement using our earlier 8.5 Å map (unpublished) of type 1 stack revealed that there are two types of stacks”. This seems very unlikely that two types of stacks were revealed this way. Do the authors mean “Heterogenous Refinement”?\n2. For the Csu pili pairs, the results section on page 5 lines 30-32 state that: “Since the pilus pair lacks helical symmetry, we generated an initial model based on the structure of the single pilus rod by optimizing the overlap between its projections and a 2D class average (Supplementary Fig. 10–12)”. This model was built prior to reconstruction of the pilus pairs map and then docked into the map. Do the authors find any significant differences if they build the model by docking individual subunits into the low-resolution map?\n3. Map:Map FSC curves for each of the structures from cryoSPARC should be presented as supplementary figures with resolution and the either the 0.143 or 0.5 FSC line being indicated.\n4. Workflows for each cryo-EM structure should be presented so readers can easily follow how the maps were generated. These should start with particle picking and show each step in the reconstruction process (heterogenous refinement, homogenous refinement, post processing, .etc).\n5. The model in Figure 7 proposed for Csu pili-mediated biofilm formation focuses on cell division as the means from which more cells are introduced into this biofilm. Acinetobacter baumannii have been shown to have T4P mediated twitching motility. Could this also significantly contribute to biofilm formation? If so, such considerations should be introduced into the paper. If not, I would suggest that the authors also address this briefly in the manuscript discussion.\n6. Building off of this, type IV pili appear to not be mentioned at all in the paper despite their presence in this strain of Acinetobacter baumannii (I believe). Were any T4P detected during cryo-EM image processing (2D classification steps, .etc)? Given the robust results, I have little doubt that the stacks/networks are formed by the Csu pili, I am just curious whether any T4P were detected at all.\n7. In figures 2-3 the pili stacks/bundles are quite hard to see in the negative stain TEMs (Fig. 2d-f mostly) as well as in the tomogram slices (Fig. 3a-c). To an untrained eye, which would include much of the broad readership of Nature communications, some of these panels might difficult to interpret even with the helpful annotations. Is there any way that higher contrast images can be used for some of these panels? For figure 2, I have absolutely no idea how to interpret panel 2f. I can detect no difference between stacks and networks in that panel despite the presence of the diamonds. For figure 3 a-c it might be possible to average together several adjacent Z-slices to generate better contrast. Would this be feasible?\nIII. Additional suggestions which may improve the story/results\n1. Supplementary Fig. 7 might be better as a main figure as its results are key to the model proposed in current main Figure 7.\n2. The number of particles used to obtain the type 1 and type 2 pili stacks was quite a lot given the relatively low resolutions achieved. Perhaps heterogeneity is the limiting factor in terms of resolution. I wonder if more classes of pili stacks would be revealed with additional heterogeneity analyses such as cryoSPARC’s 3D variability analysis (cluster output might be most useful) or cryoDRGN?"}, "author_response": "Detailed responses to the reviewers’ comments\nResponses to reviewer 1, pages 1‒19\nResponses to reviewers 2 and 3, pages 20‒32\nReferences, pages 32‒34\nReviewer 1\nWe were pleased to read that the reviewer found that our work is has “potentially high impact and relevance” and “would represent a significant and novel contribution to understanding biofilm formation and pathogenesis in A. baumannii”. We thank the reviewer for a thorough assessment of our manuscript and helpful suggestions.\nGeneral points:\nOverall, while the structural data strongly suggest the proposed stacking mechanism, without appropriate in vivo validation, this conclusion cannot be fully substantiated. Therefore, in its current form, the manuscript is not yet suitable for publication. If in vivo evidence supporting the structural findings were provided, the study would represent a significant and novel contribution to understanding biofilm formation and pathogenesis in A. baumannii. Under those conditions, I would recommend the manuscript for publication given its potentially high impact and relevance.\nWe initially chose not to include additional in vivo validation because the critical role of Csu pili in A. baumannii, as well as CupE pili in P. aeruginosa, in 3D biofilm formation has already been demonstrated in three independent studies, as summarized in Box 1. We apologize for not stating this more clearly in the Introduction.\nAlthough the mechanism of archaic CUP pilus-mediated 3D biofilm formation was previously not known, these studies firmly established the importance of this phenomenon and explored its relationship to PNAG and eDNA—addressing several points raised by the reviewer.\nBox 1. Summary of in vivo evidence from previously published studies.\n(Romero et al, 2022) demonstrated that deletion of the Csu pilus assembly platform (CsuD usher) in A. baumannii strain ATCC 17978 abolishes the formation of mushroom-shaped 3D biofilms (see Fig. 2 in their paper). Given that this study observed the natural accumulation of A. baumannii cells from a liquid culture onto a glass plate and the subsequent biofilm development over four days, it offers strong in vivo evidence.\n(Ahmad et al, 2023) covered this topic even more thoroughly through various deletion mutants. As can be seen in their Fig. 2A‒B, the deletion of Csu pilus neck-subunit CsuA in the strain ATCC 17978 turns the normally thick patches of cells on glass plates into flat monolayers, which is also apparent in the relative OD595 and measured biomass. This also directly correlates with the presence of Csu pili on the bacterial surface, as they show in their Supplementary Fig. 2A. Note that for the Fig. 2A, they also tested a deletion of the chaperone CsuC, which also collapsed biofilm mass, and restoration of CsuA through a plasmid, which brought biofilm mass back to wild type levels. Furthermore, as shown by their Fig. 2A‒B and Supplementary Fig. 2A‒C, their deletion of Type IV pilus main subunit PilA causes an increase in CsuA/B expression, more Csu pili on the surface, as well as thicker biofilms with higher biomass. Finally, they also repeated these results in the hyper-virulent strain AB5075, whose wild type form produces an abundance of Csu pili, as shown by their AFM image in Fig. 3C. Their Fig. 3F and Supplementary Fig. 2D‒E show that transposon insertions into the genes for Csu pilus main subunit CsuA/B and the PilA in strain AB5075 matched the effects of the equivalent deletions in the strain ATCC 17978. The authors also observed accumulation of PNAG in mountain-like biofilm structures (Fig 1D in the paper); however, their mutagenesis data clearly demonstrate the dominant role of Csu pili in 3D biofilm formation.\nSimilarly, by deleting genes in the cluster encoding CupE pili in P. aeruginosa, (Giraud et al, 2011) demonstrated the key role of CupE pili in cohesive biofilm structuring: the absence of CupE pili abolished the formation of three-dimensional, mushroom-shaped bacterial assemblies (see Fig. 6 in their paper). Like Csu pili in A. baumannii, CupE pili in P. aeruginosa are assembled via the archaic CUP, suggesting a broader, possibly conserved role for archaic CUP systems in 3D biofilm formation.\n“Classical and alternative CUPs primarily facilitate initial, highly specific adhesion. In contrast, archaic CUP systems are associated with the establishment of mature 3D biofilms. (Ahmad et al., 2023; Ching et al, 2024; Giraud et al., 2011; Romero et al., 2022) Among the most troublesome Gram-negative bacteria, Acinetobacter baumannii and Pseudomonas aeruginosa (Cerqueira & Peleg, 2011; Tuon et al, 2022) rely on archaic Csu and CupE pili, respectively, to develop stress and drug-resistant mushroom or mountain-like 3D biofilms on a diverse array of biotic and abiotic surfaces (Ahmad et al., 2023; Ching et al., 2024; Giraud et al., 2011; Romero et al., 2022; Tomaras et al, 2003). Although the central role of Csu and CupE pili in 3D biofilm formation has been firmly established through deleterious mutagenesis of structural or assembly genes from both archaic systems (Ahmad et al., 2023; Giraud et al., 2011; Romero et al., 2022), the underlying mechanism of this process remained elusive.”\nMajor Comments:\n(1.1) Firstly, the absence of mutational validation is particularly striking given that Figure 6 identifies specific residues involved in stacking. This validation is essential to confirm the role of Csu pili stacking in A. baumannii biofilm formation and to disentangle its contribution from other polar effects, such as PNAG or eDNA accumulation. Mutants should be analysed via electron microscopy alongside biofilm formation assays to determine the precise impact of stacking.\nThe importance of Csu pili in establishing 3D biofilms has already been extensively validated in vivo by two independent studies (see our detailed response to General points above and Box 1). These studies firmly establish that Csu pili are critical to 3D biofilm formation and explore their relationship with PNAG and eDNA. However, we agree with the reviewer that mutagenesis of the stacking interfaces is valuable, as it would also provide important mechanistic insight on how the binding junction forms. In fact, we sought to identify the binding junction residues through mutagenesis before we achieved sufficient resolution in the cryo-EM maps to model the binding contacts. This approach, however, faced two major challenges.\nFirst, designing mutants that completely disrupt binding is challenging because the junction residues are surrounded by highly conserved regions that are critical for donor strand exchange and pilus assembly (see “Supplementary Notes,” section “Antiparallel Binding Site Adjustment”). This limits the use of larger mutations, such as deletions or insertions. Conversely, introducing point mutations at the binding residues presents a different challenge: the binding junction itself appears to be tolerant, accommodating multiple residue combinations. This is supported by our phylogenetic analysis of CsuA/B from different species (current Supplementary Fig. 9a and 10), which predicts considerable sequence variability at these positions. Our modelling (Supplementary Fig. 9b‒d) also indicates that these variants should work similarly well requiring no major changes to subunit orientations or structure.\nSecond, the high heterogeneity of stack networks limits the techniques that could be utilized to assess the effects of mutations. Each stack can include Csu pili of varying lengths, each forming a variable number of binding junctions with neighbouring pili. Additionally, the number of Csu pili per stack can vary significantly, making biophysical experiments impractical, as they cannot be reliably reproduced. The pili are also too thin to be individually visualized using fluorescence microscopy and our anti-CsuA/B polyclonal antibodies with two binding sites each would likely get in the way of stack formation, while also connecting the pili randomly. This leaves negative stain TEM and cryo-EM as the only viable methods for detecting the presence of stacks. However, these techniques cannot assess the strength of the binding interactions. Therefore, unless the mutant prevents the binding entirely, we need to look for various cues, such as the accumulation rates of pairs and stacks 2 compared to the number of individual pili, which is quite open to interpretation. This challenge is further compounded by variability in the rate of stack formation: while stacks form within hours-days in biofilms (in vivo), they may take over a month to develop in purified samples (in vitro).\nBecause of these limitations, we adopted two alternative approaches to inhibit pilus stacking. In the first approach, we introduced aspartate and glutamate residues opposite to each other in the binding junction to create electrostatic repulsion between the two surfaces. The second approach leveraged the predicted differences between the binding junctions of A. baumannii ATCC 19606 and A. defluvii WCHA30 (Supplementary Fig. 9). Specifically, A. defluvii WCHA30 has a two-residue deletion in the C2–D loop and a single-residue deletion in the B′′–C1 loop, and modelling these into the A. baumannii binding junction results in an even, approximately 2 Å wide gap between the pili, which is predicted to close by the pili moving closer. We aimed to replicate this effect by introducing only one of these deletions into the binding junction, thereby inhibiting stack formation by preventing the gap from closing.\nThe results of this study are presented in the new Supplementary Table 3. Although most of the mutations either prevented or severely impaired pilus assembly/secretion, rendering them unsuitable for further analysis, the T66D-A89D double mutation and the same mutation with an additional deletion ΔP68 were well tolerated. Both mutations negatively affected the formation of larger stacks in vitro, with the effect being somewhat clearer in the triple mutant (new Supplementary Fig. 8). These results provide independent support for the proposed mechanism of pilus stacking and highlight the adaptability of binding contacts, as predicted by our phylogenetic analysis. We have added the following sentences to the main text (page 7):\n“We next examined the contribution of individual residues to stack formation. Most mutations at the pilus stacking interfaces also severely disrupted pilus assembly (Supplementary Table 3). However, the double mutant T66D–A89D, designed to introduce repulsive negative charges, and its variant T66D–ΔP68–A89D, featuring a shortened B″–C1 loop to position Asp66 closer to Asp89 and to prevent Asp66 forming hydrogen bonds with the main-chain of the opposing Csu pilus (Supplementary Fig. 8a, b), were well tolerated. Although these mutations did not abolish antiparallel interactions in vivo, the more sensitive in vitro assays using purified pili showed that the mutants had smaller, less organized stacks with shorter continuous stacked regions compared to the wild type (Supplementary Fig. 8c, d), consistent with our structural data.”\n(1.2) Furthermore, since the authors propose a similar mechanism for CupE pili, a comparable experimental approach would strengthen the claim that stacking is a widespread feature of archaic chaperone-usher pili, thereby enhancing the study’s significance.\nWe fully agree with the reviewer. Unfortunately, the P. aeruginosa CupE system is challenging to work with. While A. baumannii Csu pili are expressed in large quantities across many A. baumannii strains, the expression of CupE pili is tightly regulated and occurs only under specific conditions that strongly favour the sessile growth of this normally flagellated bacterium. Additionally, P. aeruginosa expresses several other chaperone-usher pathway (CUP) systems, necessitating the use of deletion mutants to prevent confusion during analysis.\nOur prediction is based on the excellent work by (Bohning et al, 2023), who observed a “crisscross mesh-like array” of CupE pili in vitro (Supplementary Figure 1D in their paper), which, in our opinion, closely resembles the Csu pilus stacks. We also clearly observe pilus pairs in their images (Figure 1A; Supplementary Figure 1B). To increase CupE expression and enable structural analysis, Böhning et al. engineered a triple mutant strain (ΔpilA ΔfliC ΔmvaT). Given that Böhning et al. are conducting similar studies and possess greater expertise with this organism, we plan to collaborate with their group to solve the structure of CupE pilus stacks, which we intend to publish in a separate manuscript. 3\nTo address the reviewer’s concern, we have tempered our claim regarding the similar mechanism for CupE pili. Specifically, we added “potential” to the following sentence on page 8:\n“Its position indicates it could be part of an antiparallel CupE pilus included in the helical reconstruction, which allowed us to model the potential antiparallel junction (Supplementary Fig. 11b).”\nIn the following paragraph, we also changed “it is quite likely” to “is plausible”:\n“Therefore, it is plausible that the archaic CU pilus zigzag architecture, which enables the formation of flat stacks, evolved to shape bacterial communities.”\n(1.3) Moreover, due to the high impact and novelty of the findings, I encourage the authors to consider fluorescence imaging of the pili to show the establishment and progression of the Csu stacking.\nWe greatly appreciate this valuable suggestion. However, Csu pili are too thin to individually visualize under confocal microscopy. Implementing such experiments would require the development of an entirely new experimental setup and this is therefore beyond the scope of the current manuscript. However, we are actively pursuing this direction in our ongoing work. Recently, we discovered that Csu pili (and likely most archaic CUP systems) form biofilms on oil surfaces due to the unique hydrophobic finger-like structure at the tip subunit, CsuE. Analysing biofilms on oils using confocal microscopy offers an excellent system to study Csu-mediated adhesion to surfaces and 3D biofilm formation (see Fig. R1 below). However, a new method is still needed to visualize pilus stacks in this context. Once the complete method is established, we plan to carry out these experiments and publish a separate paper on Csu-mediated biofilm formation on oils, as it will include a substantial amount of new data.\nFig. R1 | Csu pilus mediated A. baumannii biofilm formation on a drop of mineral oil. The first (0 min) and the last (2 h 40 min) frames of a video recorded on a confocal microscope.\nWe have added the following sentences to the main text in Discussion on page 11:\n“Csu pilus stack formation is a surprisingly rapid process, occurring simultaneously with 3D biofilm growth. Our results suggest that the fast kinetics may be explained by the initiation of pilus stacking around the septa of dividing cells. Pilus–pilus contacts may form quickly through a zipper-like mechanism, in which the establishment of each binding junction facilitates the formation of the next. The high cooperativity of this process may depend on the relatively flexible tips of the B′′–C1 and 4 D′′–E1 loops (Supplementary Fig. 7c), which, according to our bioinformatic analysis and mutagenesis data, can accommodate a range of binding geometries. Further studies will be needed to clarify this mechanism.”\n(2.1) Secondly, the authors’ claim that stacking is the ‘key’ mechanism of biofilm formation is not fully supported, as the extent of pili-pili interactions remains unclear. While the authors demonstrate that stacking occurs significantly in vitro under prolonged low-temperature incubation, the SEM images (Fig. 1) suggest that stacking contacts are limited to short stretches rather than spanning the entire pilus length. Consequently, it is uncertain how much these interactions contribute to in vivo biofilm formation.\nResponse on temperature:\nBefore addressing to the reviewer’s comment, we would like to clarify a major misunderstanding regarding the temperature that likely arose from our failure to specify clearly the temperatures at which Csu pilus stacks form in vivo. Unlike in our in vitro experiments, Csu pilus stacking networks form very rapidly in vivo across a broad range of ambient and physiological (body) temperatures, progressing in parallel with 3D biofilm formation. The sole purpose of storing the plates at 8°C was to prevent overgrowth of the colonies.\nTo address this point, we analysed stack formation in A. baumannii colonies immediately after growth at 22°C, 30°C, and 37°C using negative stain TEM (new Supplementary Fig. 2a‒c). Extensive networks of pilus stacks were observed at all three temperatures in the hypervirulent strain AB5075 of A. baumannii.\nFurthermore, our cryo-electron tomography experiment (Fig. 3) was conducted on an A. baumannii 19096T colony grown for three days at 22°C, without any low-temperature storage. Likewise, A. baumannii liquid cultures grown at 22°C (Supplementary Fig. 3; please, note that figure numbers have changes) and recombinant E. coli liquid cultures grown at 37°C were not stored prior to analysis. The same applies to all samples in our new negative stain TEM experiments (new Fig. 8 and Supplementary Fig. 1‒2, 4).\nWe have added the following sentence to the main text on page 4:\n“These structures formed equally efficiently under both ambient and physiological temperatures (22‒ 37°C) and across a broad range of pH (6.0‒8.5) and salt concentration (42.8‒400 mM) (Supplementary Fig. 2).”\nWe have also clarified the conditions of cell growth in the Methods section and relevant figure legends.\nResponse on “the SEM images (Fig. 1) suggest that stacking contacts are limited to short stretches rather than spanning the entire pilus length”\nWe thank the reviewer for this observation. Unfortunately, SEM offers low resolution and is prone to dehydration artefacts. Moreover, most of the longer stacks in colonies are not straight but instead follow the curvature of the bacteria, akin to a sheath, which makes them particularly difficult to capture with SEM. Therefore, to address this point, we focused on the analysis of negative-stain TEM and cryo-electron tomography.\nBoth our negative stain TEM and cryo-electron tomography in vivo experiments reveal highly extensive networks of Csu pili, with stacks forming extremely long and continuous contacts. In our 5 new Supplementary Figure 1b, we highlight a traced 7920 Å section of a Csu pilus within a stack, showing that ~6980 Å of this section is engaged in interactions—roughly half the pilus length. This correlates to around 42 binding junctions with a combined interactive surface of 17,470 Å². Moreover, stacks formed in vivo can be rather wide, with the one we observed in Supplementary Fig. 1c consisting of 15 adjacent Csu pili. Finally, the local Csu pilus stacks may also connect with one another to form superstacks, linear regions of stacked pili far exceeding the length of individual Csu pili (>12 μm vs. 1.7 μm) by continuously receiving new pili from the bacterial surfaces at regular intervals, wrapping around from multiple bacteria (see Fig. 2f; Supplementary Figure 1d‒h).\n Please note that the resolution of the TEM figures in the automatically generated pdf file is not optimal. The images are better seen in the docx source files at a 125 % zoom.\nWe have inserted the following sentences into the main text page 4:\n“Stacks may contain more than 15 pili, with pili often forming lateral interactions extending over half of their length (Supplementary Fig. 1a‒c). Superstacks, continually incorporating new pili from the surfaces of multiple bacteria, can exceed 12 μm in length (Supplementary Fig. 1d‒h), clumping bacteria in 3D structures (Supplementary Fig. 2). These structures formed equally efficiently under both ambient and physiological temperatures (22‒37°C) and across a broad range of pH (6.0‒8.5) and salt concentration (42.8‒400 mM) (Supplementary Fig. 2).”\nand page 8:\n“Such unusually short repeats are possible only because of the zigzag architecture of the pilus, which generates a high density of binding junctions. For example, in a typical stack observed in vivo, a traced Csu pilus ran ~792 nm antiparallel to adjacent pili forming ~42 binding junctions with a predicted total interactive surface of around 17470 Å² (Supplementary Figure 1b). Therefore, it is plausible that archaic CU pilus zigzag architecture, enabling the formation of flat stacks, evolved to shape bacterial communities.”\n(2.2) As the authors suggest, the presence of PNAG and eDNA likely reinforces these interactions, the primary role of Csu pili in biofilm formation could be solely to sterically trap large amounts of PNAG and eDNA rather than stacking with itself. To clarify this, the authors should generate a non-stackable Csu mutant (as mentioned previously) to determine whether stacking is the ‘key’ or rather a ‘supporting’ mechanism. Another approach would be to assess biofilm formation in the presence and absence of PNAG/eDNA using enzymatic digestion. This would allow for the isolation of pili-pili interactions, helping to determine whether stacking is indeed the key mechanism or a secondary effect.\nWe thank the reviewer for this insightful suggestion. While previous in vivo studies have established that Csu pili are essential for linking bacterial cells in 3D biofilms (see response above and Box 1), they did not conclusively demonstrate that this occurs independently of PNAG and eDNA. As the reviewer pointed out, pili could instead facilitate the accumulation of PNAG and eDNA, which would then glue the bacteria together. Resolving this issue requires direct proof that Csu pili and their stacks alone can hold bacteria together in the absence of PNAG and eDNA. We agree that non-stackable mutants would provide particularly direct evidence. However, due to the technical challenges of generating mutants that completely abolish stacking without impairing pilus assembly or secretion (see response to Main Comment 1), we were unable to pursue this approach.\nInstead, we implemented the reviewer’s alternative suggestion by assessing biofilm formation in the absence of PNAG and eDNA. Using an A. baumannii transposon mutant (pgaC::T26) lacking PNAG and enzymatic treatment with DNase I (with chloroquine to enhance eDNA degradation), we observed that bacterial clumps and rafts still formed extensive networks of stacked Csu pili even in 6 the complete absence of PNAG and eDNA (new Fig. 8). These findings demonstrate that Csu pili stacking alone provides strong cohesion and structural integrity, while PNAG and eDNA serve only as additional, non-essential reinforcements.\nTogether, these results support the conclusion that stacking is not a secondary effect, but rather the primary mechanism by which Csu pili mediate bacterial linkage and 3D biofilm formation.\nWe have inserted the following sentences into the main text on page 9:\n“While most bacteria seen in our negative stain TEM samples formed dense clumps with aplenty of Csu pilus stacks on the surfaces and PNAG or eDNA filling the tight gaps (Fig. 8a–c, Supplementary Fig. 2), others formed loose, yet cohesive pilus-linked rafts with otherwise empty, wide gaps (Fig. 8d–f). The distinctive lack of PNAG and eDNA in these rafts indicates that these EPS contribute little to clump cohesion. To test this further, we grew A. baumannii AB5075 mutant pgaC::T26, which is unable to produce PNAG (Gallagher et al, 2015), and treated suspended colonies with DNase I and the DNA-intercalating agent chloroquine to degrade eDNA (Buzzo et al, 2021). Subsequent dilution and washing steps revealed several highly organized rafts, with tight, clear gaps and extensive Csu pilus stacks traversing them (Fig. 8g–i), further supporting the conclusion that Csu pilus stacks link the bacteria in early 3D biofilms independently of PNAG and eDNA.”\nAt the same time, we cannot exclude a possible role of other EPS at different stages of A. baumannii 3D biofilm formation. Therefore, we added the following text in Discussion (page 11):\n“Our results support the dominant role of archaic Csu pili in clumping bacteria into 3D structures. However, a role for other EPS, such as PNAG and eDNA, at different stages of this process cannot be excluded. Similarly, several protein surface receptors, including OmpA (Gaddy et al, 2009), Ata (Bentancor et al, 2012) as well as Biofilm-Associated Protein (Bap) and its closely related Blp1 and Blp2 (Brossard & Campagnari, 2012; De Gregorio et al, 2015) facilitate biofilm formation. As these receptors have smaller structures, they likely mediate more localized interactions compared to the Csu pili. In addition, A. baumannii produces Type IV pili, and its different variants promote motility in AB5075 strain and biofilm formation in ACICU strain (Ronish et al, 2019). Interestingly, the latter strain has a transposase gene inserted between csuA/B and csuA within its csu operon. In our colonies of A. baumannii 19096T and AB5075, we observed little to no filaments wide enough (72 Å) (Wilharm et al, 2013) to be interpreted as Type IV pili. Only colonies grown at 30 °C and 37 °C displayed a few bacteria with such filaments, while most cells expressed abundant Csu pili. This agrees with the observation that cyclic AMP acts as a master regulator, suppressing the expression of Csu pilus subunits, PNAG-synthesizing enzymes and quorum sensing, and thus biofilm formation in general, while strongly upregulating Type IV pilus subunits and other motility-related genes (Harkova et al, 2024). Therefore, while Type IV pili may contribute to biofilm formation by mediating adhesion to certain surfaces, they likely lose relevance once A. baumannii switches to producing Csu pili. However, the expression regulation could be different in other species. Studies of other archaic CUP pili, such as the P. aeruginosa CupE system, may help clarify this.”\nSpecific comments\n(3) L71-73: The authors state that the measured thickness of Csu pili is ~30 nm (Fig. 1). However, it is evident from the image that this thickness likely results from the bundling of filaments rather than individual pili. Could the authors clarify how these measurements were obtained? How many bundles were considered? Was there a range of values? Presenting this data as mean ± SD would enhance transparency, particularly given that this observation prompted the study.\n“Earlier studies of A. baumannii biofilms using scanning electron microscopy (SEM) have shown that bacterial cells are interconnected by Csu pili (Ahmad et al, 2019; Tomaras et al., 2003). However, our analysis of SEM images indicates that the thickness of these ‘pili’ is often much greater than the width of a single Csu pilus rod filament determined by cryo-EM (Pakharukova et al, 2022). In the example shown in Figure 1, the apparent width is ~30 nm, which is 10-fold greater than that of a single pilus. This prompted us to examine the Csu pili on bacteria grown in liquid and on solid media at higher resolution using negative staining transmission electron microscopy (TEM) (Fig. 2).”\nFor quantitative analysis of in vivo stacks, we rely on negative stain TEM and cryo-ET, which preserve the native state and allow us to assess stack organization in detail. These reveal that stack width varies considerably with different numbers of Csu pili being included in it at different positions in the stacking network, making a mean ± SD for a group of stacks misleading. Representative measurements in vivo revealed a stack of 15 pili with the width of 72.7±1.6 nm (n=5) (new Supplementary Fig. 1c), which most closely correlates to a type 1 stack. Thus, the SEM observation is consistent with our higher-resolution analyses and the measured ~30 nm thickness reflects the presence of stacked pili.\nWe have inserted the following sentences into the main text page 4:\n“Stacks may contain more than 15 pili, with pili often forming lateral interactions extending over half of their length (Supplementary Fig. 1a‒c). Superstacks, continually incorporating new pili from the surfaces of multiple bacteria, can exceed 12 μm in length (Supplementary Fig. 1d‒h), clumping bacteria in 3D structures (Supplementary Fig. 2). These structures formed equally efficiently under both ambient and physiological temperatures (22‒37°C) and across a broad range of pH (6.0‒8.5) and salt concentration (42.8‒400 mM) (Supplementary Fig. 2).”\nand we also mentioned the width of the 15-pilus stack (72.7±1.6 nm, n=5) in the legend to Supplementary Fig. 1c.\n(4) L76: The authors base their in vivo observations on a translucent phenotype, which is generally associated with increased pilus production. To ensure that the observed stacking is not an artefact of higher pili numbers, could the authors provide similar images to those in Figure 2 for a wild-type A. baumannii strain?\nThe opaque and translucent phenotypes of A. baumannii are both considered wild type, as there is no underlying genetic difference between them—only variations in expression levels regulated by several transcription factors (Tipton et al, 2015). For our purposes, the key distinction is that the translucent phenotype produces an abundance of Csu pili (Ahmad et al., 2019), promoting biofilm formation (the native sessile form), whereas the opaque phenotype predominantly expresses Type IV pili, which facilitate motility of individual cells (the native motile form). A recent article by (Harkova et al., 2024) revealed that cyclic AMP (cAMP) most likely controls this switch in gene expression. While this phenotypic difference may seem ideal for our experiments, the high rate of switching between the two states poses a challenge. Moreover, even opaque colonies retain some expression 8 of Csu pili (Ahmad et al., 2019), possibly due to the constant phenotype switching of individual bacteria (Tipton et al., 2015), which poses a challenge for a reproducible analysis if not considered while picking bacteria from colonies for the experiments.\nWe initially selected for the translucent phenotype following the protocol of (Anderson & Rather, 2019) due to two reasons: (1) to ensure efficient yield of Csu pili from A. baumannii 19096 grown in a liquid culture to facilitate structural determination of a single Csu pilus from A. baumannii (Fig. 5) and (2) to avoid inconsistency between experiments that may occur due to colony phase variation switch. The selection process is summarized in Box 2.\nBox 2. Selection of translucent phenotypes of A. baumannii\n1. Pick a colony representing a chosen phenotype (thus most cells belong to that phenotype), suspend it in medium, and grow new colonies so small that it is unlikely that the bacteria in them have switched their phenotype yet. The phenotype is not yet apparent in these barely visible colonies.\n2. Grow several 5 ml liquid cultures, each using one of the tiny colonies, but stop their growth before OD595 reaches 0.1, so the bacteria are less likely to have switched their phenotype yet.\n3. Make glycerol stocks of each liquid culture. Later, check by plating on Petri-dishes that most colonies formed from a glycerol stock match the desired phenotype. Keep only the good stocks.\nThe instructions provided very specific guidance to ensure that the bacteria would not have an opportunity to switch phenotypes. However, our examination of fully-grown colonies from both phenotypes revealed that many bacteria had already undergone phenotypic switching. Negative stain TEM of opaque colonies showed bacteria forming large, tight clumps as well as loose rafts with Csu pilus stacks and pairs running through the intercellular gaps (see Fig. R2 below), similar to what we show in our new Fig. 8.\nWe made the following change to “Growing A. baumannii and recombinant E. coli colonies for Csu pilus stack analysis” in the Methods section, page 17:\n“We initially cultivated A. baumannii CCUG 19096T strain colonies using a 10-fold dilution series to achieve approximately 100–200 colonies per 0.5x LB agar plate (composed of 2.5 g NaCl, 5 g tryptone, 2.5 g yeast extract, and 8 g agar, pH 7.4, in 1 L of distilled water) to ensure the manifestation 10 of the translucent phenotype. We later observed that even seemingly opaque colonies had enough phenotype-switched cells to produce large pilus-linked clumps of bacteria.”\n(5) L77-81: Authors show that stacks are not formed effectively in liquid cultures and are present in colonies. Would that not indicate that it is rather the accumulation of ECM (possible only under colony growth and not in liquids) that is primarily responsible for pili bundling and cell aggregation rather than the inherent pili-pili interaction which should be capable of forming even in liquid environments?\nWe thank the reviewer for this point. Csu pili do form stacks in liquid cultures leading to small clumps of bacteria with little to no ECM between them (Supplementary Figures 3 and 4a–d). However, these networks are far less extensive than in colonies, where surface adhesion, higher cell density, and possibly ECM facilitate stacking. To address this point as well as major comment 2.2 (see above), we performed experiments with PNAG-deficient (pgaC::T26) mutants and DNase I treatment (Fig. 8d–i) that show that bacterial clumps and rafts remain cohesive and interconnected by extensive Csu pilus stacks even in the complete absence of PNAG and eDNA. Together with prior deletion studies showing that biofilms fail to form without Csu pili despite intact ECM (see Box 1 and our response to general comments), these findings indicate that pili–pili interactions are the primary driver of bacterial cohesion, while ECM plays only a supportive role.\nWe modified the text and added a new figure (Fig. 8). Please, see our response to major comment 2.2.\n(6) L82-87: In the cryo-ET results (Fig. 3), there appears to be no continuity between pili from neighbouring cells to confirm that stacks are intercellular rather than intracellular, suggesting that the shown stacks originate from the same cell. Given the claims of exclusively anti-parallel stacking, how do the authors reconcile this observation?\nIn the cryo-ET experiment thickness of vitrified ice is critical for the quality of data (the thinner the better). Since regions with large and tight bacterial clumps had much higher thickness of vitrified ice, we focused on collecting data in a region between less tightly clumped bacteria. The main purpose of this cryo-ET experiment was not to depict a connection between two bacteria, but to reveal the morphology (flatness) of the stacks. The sample and grid preparation protocol for the cryo-ET experiment followed the exact same steps as when we prepared colony samples for negative stain TEM (with the only exception being the use of 1 % (w/v) Uranyl Acetate staining vs. cryo-plunging using Vitrobot IV). This means that the connections seen in the negative stain TEM images are the same as in the cryo-ET experiments. We have now added plenty of negative stain TEM images (see Fig. 2 and 8; Supplementary Fig. 1‒4), showing both tight and more loose bacterial clumps connected by Csu pilus stacks as well as superstacks connecting multiple bacteria, and with these, we believe that this phenomenon should now be well documented.\nWe have now also highlighted that we used mostly the same sample preparation protocols for our negative stain TEM and cryo-ET samples by writing about them together in the following text from “Growing A. baumannii and recombinant E. coli colonies for Csu pilus stack analysis” in the Methods section, page 17:\n“For the early negative stain transmission electron microscopy (TEM) experiments, the bacteria were grown for 16–20 hours at 37°C until the colonies reached a diameter of approximately 1.0–1.5 mm, and then stored at 8°C to keep them from growing further. Under this condition, colonies could be stored for 2–3 weeks, and remained usable even after three months with the only apparent change being the slow accumulation of amorphous materials, presumably PNAG or eDNA. For the later TEM experiments using A. baumannii AB5075 strain colonies and for cryo-ET using A. baumannii CCUG 19096T strain colonies, we grew them at 22°C for three days before using them immediately 11 for the experiments without any storage.”\nWe similarly now highlight that the cryo-ET preparation method used suspended A. baumannii colonies in “Cryo-ET data collection and processing” in the Methods section, page 21:\n“For cryo-ET sample preparation, A. baumannii colonies were suspended in a solution containing 10 nm colloidal gold beads (Sigma-Aldrich, Australia) pre-coated with 1% BSA.”\n(7) L99-101: The authors state that a CsuE mutant with a compromised tip finger site impairs adhesion but does not affect Csu pili stacking, and consequently, biofilm formation remains unaffected. This statement requires quantification—biofilm formation levels should be explicitly compared to support the claim of unaltered levels.\nWe believe the reviewer’s question stems from our title for this section “Csu pilus-mediated 3D biofilm assembly is independent of bacterial adhesion to surfaces”. We admit that the title is confusing and invites a misunderstanding of us stating that CsuE has no role in the formation of 3D biofilms. We apologise for our poor wording. We have now changed the title to be more specific:\n“Csu pilus–mediated bacterial interlinking and adhesion to surfaces depend on different structural elements of the pilus”.\nFor A. baumannii 3D biofilms, both bacterial adhesion to surfaces and bacterial cross-linking are essential. In our previous study (Pakharukova et al., 2018), we demonstrated that mutations disrupting the hydrophobic tip finger site in the CsuE subunit impair adhesion and thereby prevent biofilm formation from the start. Within this framework of A. baumannii biofilm definition, the statement “consequently, biofilm formation remains unaffected” is not entirely accurate. In all classical biofilm assays that assess bacterial attachment to solid surfaces, the hydrophobic tip finger mutant yields negative results.\nWe believe the reviewer is, in fact, interested to know whether the CsuE hydrophobic tip finger site controls not only the bacterial adhesion to surfaces but also bacterial interlinking. Therefore, it is important to show that stack formation mediated by pilus rods can form and maintain a link between bacteria even without the CsuE contributing.\nTo address this, we investigated whether surface adhesion-deficient mutant Csu pili can cross-link recombinant E. coli into clumps. Negative stain TEM of liquid cultures from the mutant strain revealed Csu pilus–cross-linked clumps similar to those observed in native A. baumannii and wild-type recombinant E. coli strains, demonstrating that hydrophobic tip fingers are not required for bacterial cross-linking. This experiment further supports the conclusion that adhesion (monolayer formation) and 3D biofilm maturation (multilayer formation) are mechanistically distinct processes, mediated by two separate structural features of the Csu pilus—the hydrophobic tip fingers and the pilus rod, respectively.\nIn the revised version, we have added a new Supplementary Fig. 4 to demonstrate bacterial clump formation by the mutant in liquid cultures and in colonies. We also revised the text to describe this result explicitly and refined the terminology in the main text on page 5 to avoid potential confusion:\n“This observation was further supported by the detection of similar Csu pilus stack-tethered bacterial clumps (or rafts) in liquid cultures of E. coli strain expressing recombinant Csu pili rendered unable to facilitate bacterial adhesion to hydrophobic surfaces due to a mutant tip-subunit CsuE with a compromised hydrophobic tip-finger site (Pakharukova et al, 2018) (Supplementary Fig. 4a‒d). Similarly, extensive Csu pilus stacks formed in the induced colonies of the recombinant E. coli strain, 12 regardless of whether it possessed the wild type or the mutant CsuE (Supplementary Fig. 4e, f). Collectively, these results indicate that Csu pilus–mediated surface adhesion and bacterial interlinking are distinct processes that depend on different structural elements of the pilus. Nevertheless, both processes are essential for 3D biofilm formation.”\nAdditionally, wherever we refer to “biofilm” formed by Csu pilus rod stacking, we now specify that it is a “3D” biofilm, whenever it is necessary to ensure the clarity of the text.\n(8) L112-115: The progressing nature of assembly has been only shown at low temperatures in vitro and therefore should not be mentioned confidently unless authors could show it through in vivo imaging. Moreover, the assertion that the biofilm-like structures observed in vitro indicate that biofilm formation is a “spontaneous process inherently programmed within the structure of the pili” is strong but lacks consideration of alternative explanations. Could environmental factors, such as salinity or pH, also contribute to stack formation? Testing whether stacks still form under high ionic strength conditions would help exclude non-specific interactions driven by low ionic strength. Additionally, is stacking reversible under certain environmental conditions?\nWe thank the reviewer for this excellent suggestion. To address this point, we thoroughly investigated Csu pilus stack formation in vivo at different temperatures, salinities, and pH levels. The effect of temperature has already been discussed above. We analysed stack formation in A. baumannii colonies immediately after growth at 22°C, 30°C, and 37°C using negative stain TEM (new Supplementary Fig. 2b‒c). At all three temperatures, we observed identical and extensive networks of pilus stacks using the hypervirulent strain AB5075 of A. baumannii. These results demonstrate that in vivo, Csu pilus stacking networks form very rapidly and in parallel with colony growth, and do so robustly across a wide range of both ambient and physiological (body) temperatures.\nWe also tested the effect of pH (ranging from 6.0 to 8.5) and salinity (from 42.8 mM to 200 mM NaCl), and observed no noticeable effect on Csu pilus stack formation or bacterial clumping in vivo (new Supplementary Fig. 2d‒g). However, growth at a high salt concentration of 400 mM significantly affected A. baumannii. Under this condition, the colony diameters were only a fourth of the ones grown at 42.8 mM NaCl, the bacteria expressed fewer Csu pili, and the resulting bacterial clumps were smaller and less dense. Nonetheless, Csu pilus stacks still formed wherever there were sufficient numbers of Csu pili near one another, highlighting the strong intrinsic tendency of these pili to assemble into stacks even under suboptimal conditions.\nWe have inserted the following sentence into the main text, page 4:\n“Stacks may contain more than 15 pili, with pili often forming lateral interactions extending over half of their length (Supplementary Fig. 1a‒c). Superstacks, continually incorporating new pili from the surfaces of multiple bacteria, can exceed 12 μm in length (Supplementary Fig. 1d‒h), clumping bacteria in 3D structures (Supplementary Fig. 2). These structures formed equally efficiently under both ambient and physiological temperatures (22‒37°C) and across a broad range of pH (6.0‒8.5) and salt concentration (42.8‒400 mM) (Supplementary Fig. 2).”\nSince Csu pilus stacks form spontaneously in vivo in such short amounts of time (less than 3.5 hours after induction; see Supplementary Fig. 4) that observing their actual progress would be difficult, we also adjusted the sentence on page 5:\n“The striking similarity between the in vitro and in vivo networks, along with the progressing nature of their assembly observed in vitro, suggests that 3D biofilm formation is a spontaneous process 13 inherently programmed within the structure of the pili.”\n(9) L135-141: During grid preparation and freezing, did the authors observe any dependence of stacking on ice thickness? Could the thin ice required for SPA induce or perhaps also limit the formation of Csu pili stacks?\nNo, we did not observe any dependence of stacking on ice thickness. Stack formation appears to be a highly robust process, as it was unaffected by any of the conditions we tested (see our answer above). Moreover, the stacks used for single-particle cryo-EM had already formed well before being added to the cryo-EM grid, as they were visible as a gel at the bottom of the sample tube (Extended Data Fig. 4a). We also confirmed that this gel consisted of extensive Csu pilus stacks using negative stain TEM (Fig. 4d). Obtaining the same result with two different methods rules out any possible influence of vitrified ice thickness on stack formation.\n(10) L189-202: I strongly encourage the authors to provide mutational validation for their observations regarding CupE pili. This would significantly strengthen the novelty of their claims, allowing for extrapolation of their findings to all archaic pili based on experimental data, increasing impact.\nAs discussed in point 1.2, the technical challenges associated with expressing CupE pili place experimental work on CupE pili beyond the scope of the present manuscript; this will be addressed in future studies.\nWe have added the following text in Discussion (pages 11‒12):\n“Our results support the dominant role of archaic Csu pili in clumping bacteria into 3D structures. However, a role for other EPS, such as PNAG and eDNA, at different stages of this process cannot be excluded. Similarly, several protein surface receptors, including OmpA (Gaddy et al., 2009), Ata (Bentancor et al., 2012) as well as Biofilm-Associated Protein (Bap) and its closely related Blp1 and Blp2 (Brossard & Campagnari, 2012; De Gregorio et al., 2015) facilitate biofilm formation. As these receptors have smaller structures, they likely mediate more localized interactions compared to the Csu pili. In addition, A. baumannii produces Type IV pili, and its different variants promote motility in AB5075 strain and biofilm formation in ACICU strain (Ronish et al., 2019). Interestingly, the latter strain has a transposase gene inserted between csuA/B and csuA within its csu operon. In our colonies of A. baumannii 19096T and AB5075, we observed little to no filaments wide enough (72 Å) (Wilharm et al., 2013) to be interpreted as Type IV pili. Only colonies grown at 30 °C and 37 °C displayed a few bacteria with such filaments, while most cells expressed abundant Csu pili. This agrees with the observation that cyclic AMP acts as a master regulator, suppressing the expression of Csu pilus subunits, PNAG-synthesizing enzymes and quorum sensing, and thus biofilm formation in general, while strongly upregulating Type IV pilus subunits and other motility-related genes (Harkova et al., 2024). Therefore, while Type IV pili may contribute to biofilm formation by mediating adhesion to certain surfaces, they likely lose relevance once A. baumannii switches to producing Csu pili. However, the expression regulation could be different in other species. Studies of other archaic CUP pili, such as the P. aeruginosa CupE system, may help clarify this.”\n(11) L204: The claim that PNAG embeds pilus stacks in mature biofilms is based on WGA labelling. To confirm this, could the authors treat samples with PNAG-hydrolyzing enzymes and analyze the effect via TEM imaging? Additionally, how do biofilm levels compare in the presence and absence of this enzyme? Could authors discuss the possibility that trapping PNAG and eDNA is the main involvement of Csu pili in biofilm formation rather than a side effect of stacking.\nThis is an excellent suggestion. Please see our extended answer to a related point 2.2. We 14 attempted to purchase the PNAG-hydrolysing enzyme Dispersin B from Kane Biotech Inc., but they refused to sell it to us due to regulatory restrictions. Fortunately, we were able to implement the second approach suggested by the reviewer earlier, mutagenesis. Specifically, we obtained an A. baumannii pgaC::T26 mutant, which is unable to produce PNAG because the glycosyltransferase gene pgaC is disrupted by a T26 transposon. We then grew this mutant and suspended three-day-old colonies in DNase I solution to degrade eDNA. After dilution and washing, the remaining bacterial clumps, or “tight rafts” (Fig. 8g), were highly organized and resembled wild type clumps, but displayed clear gaps devoid of any material resembling PNAG or eDNA (Fig. 8h), while extensive Csu pilus stacks were observed traversing these gaps (Fig. 8i). The fact that the clumps retained their macroscopic structure in the absence of PNAG and eDNA demonstrates that these components are not essential for maintaining bacterial cohesion.\nWe have inserted the following sentences into the main text on page 9:\n“While most bacteria seen in our negative stain TEM samples formed dense clumps with aplenty of Csu pilus stacks on the surfaces and PNAG or eDNA filling the tight gaps (Fig. 8a–c, Supplementary Fig. 2), others formed loose, yet cohesive pilus-linked rafts with otherwise empty, wide gaps (Fig. 8d–f). The distinctive lack of PNAG and eDNA in these rafts indicates that these EPS contribute little to clump cohesion. To test this further, we grew A. baumannii AB5075 mutant pgaC::T26, which is unable to produce PNAG (Gallagher et al., 2015), and treated suspended colonies with DNase I and the DNA-intercalating agent chloroquine to degrade eDNA (Buzzo et al., 2021). Subsequent dilution and washing steps revealed several highly organized rafts, with tight, clear gaps and extensive Csu pilus stacks traversing them (Fig. 8g–i), further supporting the conclusion that Csu pilus stacks link the bacteria in early 3D biofilms independently of PNAG and eDNA.”\n(12) L237-240: Are there any documented differences in the production of Csu pili in liquid vs biofilm growth that could support this?\nWe assume the reviewer is referring to the following statement: “Second, the small gaps between neighbouring cells in a biofilm help position Csu pili antiparallel to each other, promoting stack formation (Fig. 7). Csu pilus stacks between daughter cells and neighbouring cells could help maintain their spatial orientation, allowing the biofilm to grow taller without breaking apart. In liquid cultures, such neighbouring cells are most often absent (Supplementary Fig. 8)”.\nWe refer to such study (ref. 39, Rumbo-Feal et al., 2013) and discuss this issue a few sentences above:\n“Although this process can occur both in liquid and solid media, the formation of stacks linking bacteria together is dramatically more efficient in 3D biofilms than in liquid cultures. A. baumannii in biofilms produce more pili than liquid culture cells (Rumbo-Feal et al, 2013), but since clumps are also rare in E. coli liquid cultures overexpressing Csu pili, the higher number of pili is unlikely the main factor here.”\nOur conclusion is largely based on observations from negative stain TEM analysis of suspended colonies, where it is widespread that extensive Csu pilus stacking networks follow the curvature of bacteria some distance away from the cell surface (Fig. 2 and Supplementary Fig. 1d‒h, see also Fig. R2e, h above). These superstacks are consistently present in samples from both A. baumannii and E. coli colonies and are most visible on the exposed surfaces of the bacteria. In contrast, in our liquid culture samples, both from A. baumannii and E. coli, the Csu pili tend to point in every direction with only a few stacking interactions. However, in the bacterial clumps found in liquid cultures, while the Csu pili on the outer surfaces still point directly outwards, the Csu pili between the bacteria show 15 stacking interactions (Supplementary Fig. 3 and 4) with many of them running parallel to the cell surfaces, especially when the gaps are narrow (Supplementary Fig. 3).\n(13) L251-252: Has it been tested—either in this study or elsewhere—whether non-zigzagged pili are unable to form gel-like structures after prolonged incubation at low temperatures? If not, this statement should be softened.\nThe reviewer is referring to the following statement: “We show that such flat assemblies require helical filaments with very short repeats, suggesting that the zigzag architecture of archaic CUP pili has likely evolved to mediate 3D biofilm formation.”\nCsu pilus stacks do not require “prolonged incubation at low temperatures” to form in vivo. On the contrary, stack formation is a surprisingly rapid process that occurs simultaneously with 3D biofilm growth across a wide range of temperatures (see our answers above). We propose that the zigzag architecture provides multiple features beneficial for 3D biofilm formation, including the unusually short helical repeat that enables a high density of binding junctions between pili. Classical and alternative CUP pili, by contrast, have different architectures and are typically not involved in 3D biofilm formation; instead, they generally mediate highly specific adhesion to biological targets. Nevertheless, one example of classical CUP pili contributing to 3D biofilm formation has been thoroughly documented (E. coli common pili), and it is quite possible that other classical or alternative CUP systems have also been adapted for bacterial cross-linking in 3D biofilms. Documented cases in which both CUP and non-CUP pili participate in 3D biofilm formation are discussed in the Discussion section on page 12:\n“Archaic zigzag pili represent the first example of using highly structured, regular flat sheets for 3D biofilm assembly. Several other EPS scaffolds reported in bacteria and archaea are also DSC-linked fibres, but they mostly form twisted (Garnett et al, 2012),(Gohl et al, 2006) or cable-like bundles (Bohning et al, 2022; Gaines et al, 2022; Sonani et al, 2025; Wang et al, 2022). E. coli common pili, assembled through classical CUP mechanisms (Garnett et al., 2012), and archaeal bundling pili (ABP) (Wang et al., 2022) from Pyrobaculum calidifontis involve antiparallel interactions, while archaeal Tafi pili from Natrinema sp. J7-2 appear to involve both parallel and antiparallel interactions (Sonani et al., 2025). The directionality remains unknown for Acinetobacter sp. strain BD413 classical Acu pili (Gohl et al., 2006), Sulfolobus acidocaldarius archaeal thread filaments (Gaines et al., 2022) and TasA pili of the Gram-positive bacterium Bacillus subtilis (Bohning et al., 2022). Further studies are necessary to understand how these bundles spontaneously assemble to support 3D biofilms.”\n(14) L269-270: Structurally, if one site on the pilus rod is occupied by another to form a stack, why would the remaining binding sites become inaccessible for another junction? Can the authors confidently exclude the possibility of three-dimensional stacking instead of exclusively flat stacking? Is there any in vivo evidence supporting the formation of flat connections rather than thick, circular bundles? If flat stacks were the predominant arrangement, wouldn’t they be visible in SEM images? While the cryo-ET data suggests flat stacks, without subtomogram averaging, this conclusion remains uncertain.\nWe split the comment in two parts and address each part separately below.\n(14.1) Structurally, if one site on the pilus rod is occupied by another to form a stack, why would the remaining binding sites become inaccessible for another junction? Can the authors confidently exclude the possibility of three-dimensional stacking instead of exclusively flat stacking?\nTheoretically, despite the geometric constraints outlined in the new Supplementary Fig. 7a, a wide variety of structures are possible (Supplementary Fig. 7b), including three-dimensional assemblies (Supplementary Fig. 7e, f). However, despite extensive analyses using negative stain TEM, single-particle cryo-EM, and cryo-ET, and examining Csu pilus stacks in both in vivo (colonies, liquid cultures) and in vitro (extracted pili) contexts, we have never observed any structures that could be confidently interpreted as 3D stacks. This stands in sharp contrast to the abundant evidence for pairs and flat stacks under both conditions. Thus, while 3D stacks remain a theoretical possibility, they are likely to be extremely rare.\nA plausible explanation for their rarity is that such structures would compromise the remarkable stretching capacity of individual Csu pili (Pakharukova et al., 2022). As shown in the current Supplementary Fig. 13a–d, the ability of a single Csu pilus to nearly double its length by opening its clinch-structures is largely preserved in pilus pairs and in both types of flat stacks, with type 2 stacks potentially also allowing lateral stretching. In contrast, none of the predicted 3D stacks would accommodate this property. For example, the helical tube variant of the type 1 stack lacks subunits that can stretch without disrupting multiple binding junctions (Supplementary Fig. 7e; end view), making it not only extremely rigid but also inhospitable to slightly pre-stretched pili attempting to join the structure. Similarly, the second type of 3D stack contains only two stretchable subunits per seven-subunit repeat (Supplementary Fig. 7f; orange and green subunits). While this configuration would allow limited extension, it would still be far more rigid than flat stacks or pairs.\nIt is also important to consider the abundant “random bundles” frequently observed in our samples. These are twisted, disorganized versions of flat stacks and pairs, sometimes stabilized by additional pili wrapping around them and forming junctions that help maintain their orientation (Fig. 4b). Occasionally, a pilus may transiently contact three antiparallel pili simultaneously—similar to the geometry of predicted 3D stacks (Supplementary Fig. 7f)—but these interactions never give rise to regular 3D architectures. Instead, we consistently find only pairs and flat stacks embedded within such bundled regions. Competition with these disorganized random bundles is therefore another likely reason why highly ordered 3D stacks are so rare.\nWe have added the following sentences in Discussion (page 11):\n“This extensive stretching capability may explain why Csu pili tend to form only flat stacks, even though the available binding sites theoretically allow for much greater variation (Supplementary Fig. 7a, b). It appears that, apart from the wavy type 1 stack variant that may have a somewhat reasonable ability to stretch (Supplementary Fig. 7c), the spacing of binding junctions in others would exclude most of the subunits from stretching (Supplementary Fig. 7d‒f). Importantly, the high elasticity of stacks would make established 3D bacterial clumps more resistant to turbulent environments by absorbing stress. Interestingly, the stacks embedded into extracellular material made of PNAG and eDNA are reminiscent of reinforced concrete, as in this combination, the stack may act like steel bars or mesh, providing resistance to tensile and shear stresses, while PNAG or eDNA would provide resistance to compression. Further experimental biophysical studies will be required to investigate the predicted properties.”\n(14.2) Is there any in vivo evidence supporting the formation of flat connections rather than thick, circular bundles? If flat stacks were the predominant arrangement, wouldn’t they be visible in SEM images? While the cryo-ET data suggests flat stacks, without subtomogram averaging, this conclusion remains uncertain.\nBy contrast, the limited SEM images we included are not representative of stack structures. Their purpose was primarily to provide an overview and to visualize PNAG accumulation. For more comprehensive SEM data on Csu pili in A. baumannii biofilms, earlier studies by Tomaras et al. (2003) and Ahmad et al. (2019) are more informative, showing that stacks can be far more extensive than in our selected image. Moreover, SEM is a low-resolution technique requiring extensive sample preparation, which is well known to introduce artefacts. Since Csu pilus stacks form in a hydrated environment, drying and coating samples with heavy metals is highly likely to distort their native architecture (e.g., flat stacks could roll into tube-like structures or collapse under the stain). Additionally, SEM provides only 2D projections and therefore cannot reliably resolve 3D structures. In contrast, cryo-ET produces complete, rotatable 3D segmentations of structures preserved in vitrified liquid, capturing them in their native state. Our cryo-ET data were obtained from freshly grown colonies, suspended and applied to grids within minutes of removal from Petri dishes, leaving little to no opportunity for artefacts to arise. For these reasons, cryo-ET provides far more reliable evidence of stack morphology than SEM.\nRegarding the reviewer’s suggestion that subtomogram averaging is required to validate our cryo-ET data, we would like to point out that, for subtomogram averaging to be meaningful, it is necessary to align and average large numbers of near-identical particles (or helical assemblies) to obtain higher resolution (often nanometre or sub-nanometre) structures. Csu pilus stacks are too sparse to provide a sufficient number of near-identical subvolumes for subtomogram averaging, rendering this method unsuitable. Nevertheless, in all our cryo-ET reconstructions, the stacks are consistently several-fold wider than they are thick, as demonstrated in Supplementary Video 1. Moreover, we observed these flat stacks at various orientations relative to the imaging plane, with individual pili visible in the same tomograms, making it implausible that the observed stacks are imaging artefacts. To further support this conclusion, we have added a series of tilt images of our segmentation (Fig. 3i), showing a single stack rotated 90° in four steps, directly illustrating its flatness.\n(15) L233-236: The authors suggest that increased turbulence explains why liquid cultures do not exhibit the same levels of Csu-mediated biofilm formation. Could they provide evidence demonstrating the impact of growth conditions on Csu stack formation in vitro? Additionally, could they visualize the effect of turbulence on pre-assembled Csu stacks?\nAgain, an interesting point and excellent direction for future studies. We have partially addressed this point with our new in vivo experiments (1) and some in vitro observations (2):\n(1) The effect of environmental conditions on Csu pilus assembly is much easier to test in vivo as forming them in vitro takes significantly more time, possibly due to turbulence or lack of narrow bacterial gaps guiding the pilus orientations. As we mentioned in our answer to the reviewer’s comment 8, we conducted an experiment of growing A. baumannii colonies on standard (0.5x LB, 42.8 mM NaCl and pH 7.4, 3 days at 22°C) and adjusted conditions, changing NaCl concentrations (200 mM and 400 mM) and pH values (pH 6.0 and 8.5) (Supplementary Fig. 2a, d‒g). We also grew colonies at 30°C and 37°C overnight to confirm the effect of temperature and time on Csu pilus stack formation (Supplementary Fig. 2a‒c). Our results indicated that Csu pilus stacks form rapidly in all tested conditions with the only limiting factor being high NaCl concentrations that reduce bacterial growth and the number of Csu pili per bacteria. Even then, stacks formed when enough of pili were sufficiently close to one another, so the excess NaCl did not affect the stack formation itself. 18\n(2) When it comes to the effects of turbulence on pre-assembled Csu pilus stacks in vitro, the process of pipetting stacks that accumulated at the bottom of a fraction tube (Supplementary Fig. 5a) and adding them onto a TEM grid is typically enough to temporarily resuspend most of the visible accumulated substance into the solution. However, it seems insufficient to break the stacks themselves, as the negative stain TEM grids made from such samples tend to have aplenty of regular stacks (Fig. 4c, d). The resuspended stacks will also settle back to the bottom of the tube, allowing us to use the same stack sample as long as enough stacks remain, which was the case, when we checked for the presence of stacks using negative stain TEM (Fig. 4d) before using the sample to make our cryo-EM grids for structure determination (Supplementary Fig. 5). Therefore, the Csu pilus stacks are quite stable, once they have formed.\nWe believe that analysing the role of turbulence in stack formation requires a dedicated study using appropriate methods. Such work could potentially suggest new strategies to prevent or disrupt the formation of Csu pilus–mediated 3D biofilms and may have considerable medical relevance.\nWe are very pleased that the reviewers found our study to be “rigorous, well-written, well-presented, and of interest to a broad scientific readership,” and that they supported our interpretation of most cryo-EM structures. We are grateful for the reviewer’s thorough assessment and for the many excellent suggestions and corrections, which have helped us to improve the manuscript. In response, we have improved the description of our structure refinement protocol, added the missing FSC curves, examined the fit of individual subunits into the pilus pair map, performed 3D variability analysis, and prepared a detailed workflow figure. To further enhance clarity and interpretation of our negative stain TEM and cryo-ET data, we improved several figures, added four new figures illustrating different aspects of Csu pilus stack networks, revised the figure legends, and included additional references. Finally, we re-examined both old and new samples to assess the presence of Type IV pili, and our data indicate that Csu pili and Type IV pili are not co-expressed, as discussed in the revised manuscript.\nReviewer #2 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.\nReviewer #3 (Remarks to the Author):\nI. Overall review:\nThe archaic chaperone usher Csu pili formed by Acinetobacter baumannii are known to play essential roles in the formation of biofilms. Csu pili have been previously shown to biofilms on abiotic surfaces as well as biofilm pellicles and play crucial roles in adhesion to mammalian tissues. This study by Malmi et. al uses a variety of electron microscopy techniques (SEM, Negative stain TEM, Cryo-ET, Cryo-EM, and gold conjugation labeling) to investigate the role that Csu pili play in biofilm formation at multiple length scales from the micro/nanoscale of SEM and cryo-ET to sub-nanometer and atomistic details provided by single particle cryo-EM. Using these length scales the authors are able to strongly suggest how the Csu pili are involved in biofilm formation. The cryo-EM structural details of the pilus pairs and stacks, while low-resolution, are able to show novel anti-parallel packing interactions between pili. When attached to cells these anti-parallel interactions would presumably mostly occur between different cells which is plausible. Upon examination of the atomic models and cryo-EM density maps, I am confident in the authors’ interpretation of most of their cryo-EM structures. Any concerns are noted in the next section of my review. With all of this data the authors propose a plausible model for Csu pili-mediated biofilm formation.\nOverall, this study is rigorous, well-written, well-presented, and should have appeal to a broad scientific readership. Upon sufficient response to my concerns from the authors, I think this paper would be an excellent fit for publication at Nature Communications. I have several points which I would strongly like to see the authors address as well as some minor suggestions that could help improve their story/results.\nII. Major points to be addressed:\n1. In the Methods on page 16 lines 30-31 it is stated: “Homogeneous Refinement using our earlier 8.5 Å map (unpublished) of type 1 stack revealed that there are two types of stacks”. This seems very unlikely that two types of stacks were revealed this way. Do the authors mean “Heterogenous Refinement”?\nWe thank the reviewer for this observation. We first observed indications of two distinct stack types during Homogeneous Refinement (see Fig. R3 below), which were subsequently confirmed through 20 multiple rounds of Heterogeneous Refinement (now outlined in the new workflow figure, Supplementary Fig. 17).\nThe Csu pilus pairs are nearly identical in both stack types, differing only in how they bind to additional pili (Fig. 5b, Supplementary Fig. 7a, b). As a result, refinement programs often overlapped one pair from each type, producing a combined density map where the two stacks intersected at 51.43° (360°/7) angle. In this overlap, type 1 stacks could be recognized by 154.43° (360°/7×3) angles between three adjacent pili, whereas type 2 stacks had 102.86° (360°/7×2) angles. The sharper geometry of type 2 stacks was evident from the end views, and type 1 stack models did not fit well into type 2 stack densities. Importantly, the combined maps showed unequal density distribution, consistent with two distinct flat stacks being superimposed rather than the pili forming a single, unified 3D stack (Supplementary Fig. 17e and 7f; Fig. R3b). By fitting modified type 1 stack overlap models into these combined maps, we confirmed that type 2 stacks use alternative subunits for pilus–pilus contacts rather than using different types of binding junctions altogether (Fig. R3c, d). This density pattern was not unique to this initial refinement run but recurred whenever we used unsorted particle sets, and the final map orientations (Supplementary Fig. 17h) still reflect this characteristic ~51.43° overlap.\nThus, although the job in question was a Homogeneous Refinement, the presence of two nearly identical pilus pairs caused the algorithm to merge features from both stack types, revealing their coexistence in the dataset. This overlap, together with pseudosymmetry from 180° flipped orientations, represent the two main sources of heterogeneity in our stack density maps (see response to point 4).\n2. For the Csu pili pairs, the results section on page 5 lines 30-32 state that: “Since the pilus pair lacks helical symmetry, we generated an initial model based on the structure of the single pilus rod by optimizing the overlap between its projections and a 2D class average (Supplementary Fig. 10–12)”. This model was built prior to reconstruction of the pilus pairs map and then docked into the map. Do the authors find any significant differences if they build the model by docking individual subunits into the low-resolution map?\nWe thank the reviewer for the suggestion. We docked individual donor-strand complemented CsuA/B subunits into the Csu pilus pair density map. The overall result was very similar to docking the full pilus pair model (Fig. R4a, b).\n3. Map:Map FSC curves for each of the structures from cryoSPARC should be presented as supplementary figures with resolution and the either the 0.143 or 0.5 FSC line being indicated.\n4. Workflows for each cryo-EM structure should be presented so readers can easily follow how the maps were generated. These should start with particle picking and show each step in the reconstruction process (heterogenous refinement, homogenous refinement, post processing, .etc).\nWe thank the reviewers for the suggestion. We have made a new figure (Supplementary Fig. 17) showing the map refinement workflows for the Csu pilus rod segment, the Csu pilus rod pair, the initial map of Csu pilus type 1 stack as well as the combined map refinement workflow for Csu pilus type 1 and 2 stacks. The first three workflows (Supplementary Fig. 17a) are quite standard, but the workflow for pilus stacks 1 and 2 is more complex (Supplementary Fig. 17b‒h), as explained below:\nThe reconstruction of pilus stacks type 1 and type 2 faced two main issues. First, Csu pilus pairs were nearly identical between the two stack types, and since each pair constituted a large portion of the map, refinement programs frequently misassigned particles. This produced composite maps in which density patterns from the two stacks crossed at 51.43°, while non-shared junctions showed weaker density. Second, particle sets contained mixtures of 180° flipped orientations, as the pillar-like stacks and zigzag features of individual pilus rods appeared similar when inverted. This pseudosymmetry caused overlaps during refinement, evident in 2D classes showing adjacent sharp and diffuse regions (Supplementary Fig. 17d).\nTo separate particles from the two stack types, we ran iterative Heterogeneous Refinement job pairs, pooling particles assigned to the same type for subsequent rounds. Residual misassignments were addressed by subdividing type 1 stack particles into seven subsets and excluding the two poorest based on Homogeneous Refinement results. For type 2 stacks, we instead used a type 1 model as a decoy class to clean the set, as type 1 stack maps appeared less prone to snatch type 2 stack particles than vice versa.\nPseudosymmetry was further resolved by using multiple 180°-flipped versions of the stack maps as search models in Heterogeneous Refinement, while deliberately lowering the O-EM learning rate so occasional flipped particles would not bias the maps (Supplementary Fig. 17g). After refinement, we rotated results to match the original model and combined them in Local Refinement to avoid orientation randomization (Supplementary Fig. 17h).\n5. The model in Figure 7 proposed for Csu pili-mediated biofilm formation focuses on cell division as the means from which more cells are introduced into this biofilm. Acinetobacter baumannii have been shown to have T4P mediated twitching motility. Could this also significantly contribute to biofilm formation? If so, such considerations should be introduced into the paper. If not, I would suggest that the authors also address this briefly in the manuscript discussion.\nWe thank the reviewers for raising this point. To address it, we re-examined all our old and new negative stain TEM micrographs of A. baumannii colonies from two strains (CCUG 19096 and AB5075) grown under different conditions (temperature, pH, salt concentration, and incubation time). We tested colonies of the translucent phenotype (most samples), mixed phenotype (AB5075 during revision), and opaque phenotype (see Fig. R2 in our responses to Reviewer 1). Type IV pili in A. baumannii (~72 Å in diameter; (Wilharm et al., 2013)) are notably wider than Csu pili, which vary from ~55 Å at their widest point to less than half of that (Fig. 4b), making their average width around half the width of Type IV pili. In most TEM samples, we observed no filaments wide enough to be identified as Type IV pili. Only in colonies grown at 30°C or 37°C did we detect a few filaments too wide to be Csu pili (Fig. R5a), which could possibly represent Type IV pili. By contrast, all colonies displayed abundant Csu pili forming extensive stacks. These findings suggest that in colonies, Csu pili overwhelmingly dominate, and the rare appearance of Type IV pili likely reflects a few cells switching to (or retaining) the opaque phenotype.\nThis result is consistent with prior studies showing that Csu and Type IV pili are antagonistically regulated. Recently, (Harkova et al., 2024) demonstrated that cyclic AMP (cAMP) acts as a master regulator controlling the switch between two phenotypes, affecting ~234 differentially expressed genes (see their Fig. 2–3). Specifically, cAMP suppresses expression of Csu pilus subunits, PNAG biosynthetic enzymes and quorum-sensing, and thus biofilm formation overall, while strongly upregulating Type IV pilus subunits and other motility genes. Thus, Csu pili and Type IV pili are not expected to be co-expressed: Csu pili promote sessile biofilm growth, while Type IV pili mediate twitching motility—processes that are physiologically incompatible. This conclusion is also supported by (Giles et al, 2015), who showed that cAMP degradation by phosphodiesterase CpdA increased CsuA/B expression and promoted surface pellicle formation, and by (Nait Chabane et al, 2014), who identified CsuA/B as the most abundant protein in the pellicles. The cAMP-dependent switch likely underlies the opaque–translucent phase variation, with the translucent phenotype producing more Csu pili and displaying reduced motility (Ahmad et al., 2019; Tipton et al., 2015).\nAlthough Type IV pili appear dispensable in colonies, they may contribute more significantly to pellicle formation in liquid cultures, which may better reflect conditions for motile, opaque-phase bacteria. However, all our liquid culture experiments used translucent-phase stocks to ensure efficient Csu pilus expression, and we currently lack opaque-phase stocks. Furthermore, it would be difficult to determine whether Type IV pili in pellicles derive from pre-existing opaque cells or from phenotype switching during growth. Likewise, small clumps of cells with Type IV pili in liquid culture could reflect phenotype switching rather than true accumulation. It is also possible that motile cells lose their Type IV pili upon joining a pellicle. While we recognize this as an interesting direction for future work, it is beyond the scope of the present study.\nAs an informative comparison, our TEM samples of Pseudomonas aeruginosa PA14 colonies grown under similar conditions at 22°C showed bacteria with multiple flagella and abundant wide filaments consistent with Type IV pili (Fig. R5b), with few other pili present. This suggests that under our standard growth conditions, the equilibrium in P. aeruginosa strongly favours the motile phenotype. 26\nFig. R5 | Type IV pili in Acinetobacter baumannii and Pseudomonas aeruginosa colonies and their absence among pili purified from E. coli pBAD-Csu overexpression system. a, Possible Type IV pili observed on A. baumannii AB5075 colony grown at 30°C overnight. There were no filaments wide enough to interpret as Type IV pili in any of our liquid cultures or colonies grown at 22°C. While colonies grown at 30°C and 37°C did have a few bacteria that seemed to express Type IV pili, these were rare, and the vast majority of bacteria produced Csu pili assembling into extensive stacks (Supplementary Fig. 2b, c). b, P. aeruginosa PA14 grown on identical 0.5x LB agar plates as A. baumannii formed rapidly spreading, flat colonies with the bacteria covered in flagella as well as numerous filaments that we believe are Type IV pili due to their width, while producing very few other types of filaments. This suggests that under these growth conditions, P. aeruginosa strongly exhibits a motile phenotype. Since CupE are necessary for the formation of stationary, mushroom-like 3D biofilms in P. aeruginosa (Giraud et al., 2011), their expression is likely suppressed in these motile phenotype cells. c, Csu pilus fractions purified for cryo-EM from E. coli pBAD overexpression system using a Source30Q anion exchange column and then checked with SDS-PAGE for purity. Since the Csu pili were produced in E. coli and then purified with an ion exchange column, there were very few other types of proteins present in the fractions used for cryo-EM. Naturally, none of the cryo-EM processing steps showed any Type IV pili.\nWe have added the following text to the Discussion section on pages 11‒12:\n“Our results support the dominant role of archaic Csu pili in clumping bacteria into 3D structures. However, a role for other EPS, such as PNAG and eDNA, at different stages of this process cannot be excluded. Similarly, several protein surface receptors, including OmpA (Gaddy et al., 2009), Ata (Bentancor et al., 2012) as well as Biofilm-Associated Protein (Bap) and its closely related Blp1 and Blp2 (Brossard & Campagnari, 2012; De Gregorio et al., 2015) facilitate biofilm formation. As these receptors have smaller structures, they likely mediate more localized interactions compared to the Csu pili. In addition, A. baumannii produces Type IV pili, and its different variants promote motility 27 in AB5075 strain and biofilm formation in ACICU strain (Ronish et al., 2019). Interestingly, the latter strain has a transposase gene inserted between csuA/B and csuA within its csu operon. In our colonies of A. baumannii 19096T and AB5075, we observed little to no filaments wide enough (72 Å) (Wilharm et al., 2013) to be interpreted as Type IV pili. Only colonies grown at 30 °C and 37 °C displayed a few bacteria with such filaments, while most cells expressed abundant Csu pili. This agrees with the observation that cyclic AMP acts as a master regulator, suppressing the expression of Csu pilus subunits, PNAG-synthesizing enzymes and quorum sensing, and thus biofilm formation in general, while strongly upregulating Type IV pilus subunits and other motility-related genes (Harkova et al., 2024). Therefore, while Type IV pili may contribute to biofilm formation by mediating adhesion to certain surfaces, they likely lose relevance once A. baumannii switches to producing Csu pili. However, the expression regulation could be different in other species. Studies of other archaic CUP pili, such as the P. aeruginosa CupE system, may help clarify this.”\n6. Building off of this, type IV pili appear to not be mentioned at all in the paper despite their presence in this strain of Acinetobacter baumannii (I believe). Were any T4P detected during cryo-EM image processing (2D classification steps, .etc)? Given the robust results, I have little doubt that the stacks/networks are formed by the Csu pili, I am just curious whether any T4P were detected at all.\nTo obtain most of our SPA cryo-EM structures, including the Csu pilus stacks, we expressed Csu pili using our E. coli BL21 pBAD-Csu overexpression system. The only exception was the high-resolution structure of the native single Csu pilus, which we purified directly from the A. baumannii CCUG 19096T strain through multiple rounds of purification, yielding a highly pure Csu pilus preparation. As a result, our cryo-EM micrographs did not contain Type IV pili, and none were detected during 2D classification or other image processing steps.\n7. In figures 2-3 the pili stacks/bundles are quite hard to see in the negative stain TEMs (Fig. 2d-f mostly) as well as in the tomogram slices (Fig. 3a-c). To an untrained eye, which would include much of the broad readership of Nature communications, some of these panels might difficult to interpret even with the helpful annotations. Is there any way that higher contrast images can be used for some of these panels? For figure 2, I have absolutely no idea how to interpret panel 2f. I can detect no difference between stacks and networks in that panel despite the presence of the diamonds. For figure 3 a-c it might be possible to average together several adjacent Z-slices to generate better contrast. Would this be feasible?\nWe thank the reviewers for this suggestion. We have now revised Fig. 2 and 3 to improve clarity and logical presentation. Please note that the resolution of the TEM figures in the automatically generated pdf file is not optimal. The images are better seen in the docx source files at a 125 % zoom.\nWe replaced panel 2e with another view that depicts a local network of stacks at a reasonably high resolution. In addition, we added three close-up views (2g‒i) of the bundle from panel 2f, which we now call a “superstack”.\nWe have also revised the figure legend for clarity:\n“Fig. 2 | Csu pili form a network of stacks interlinking bacteria in 3D biofilms. a, A. baumannii 19096T cells from liquid cultures are mostly covered in separate Csu pili pointing radially in every direction. b–i, A negative stain TEM image of a suspended A. baumannii colony reveals a network of Csu pilus stacks (b), including local stacks wrapping around the bacteria (c‒e) and a long, fully exposed superstack (f‒i) linking two bacteria on the left to a clump of bacteria on the right, which they likely detached from during sample preparation. Typically, the pili growing in the narrow gaps between bacteria wrap closely around each bacterium (c) by forming pairs and flat stacks (d,e) that 28 further merge into extensive networks connecting multiple bacteria (b,f). The ends of the exposed superstack connecting to the two groups of bacteria consist of Csu pilus pairs and stacks emerging from multiple directions (g,i) that then merge into a unified stack closer to the centre (h). Blue dots highlight examples of separate pili, while red dots border pilus pairs or stacking networks, and blue lines in d indicate individual pili in a multi-pilus stack. Diamonds in e‒i highlight the local networks of pilus stacks merging into the superstack.”\nTo help confirm that the terms we use above are now clear, we have added a more detailed description of them below. The terms “a network of Csu pilus stacks” and “stacking network” both refer to all Csu pili in a region that are connected to one another through stacking interactions. Due to their length, each individual pilus may take part in multiple separate pairs and stacks, and since the other pili in those pairs and stacks may form even further stacks, this creates an extensive network branching in every direction inside the biofilm. To describe better the large stack connecting the two groups of bacteria, we have now introduced the term “superstack”, which we used to replace the confusing term “bundle” in all of our descriptions of large, mostly organized Csu pilus stacks. The term refers to particularly dense sections of the stacking network that appear in overviews as long, mostly linear filaments. These are quite common, though their low contrast in TEM images and other obscuring factors often prevent us showing them in full. We used the term “superstack” also in our new Supplementary Fig. 1 that depicts stack dimensions observed in our in vivo samples.\nFinally, to give further examples of Csu pilus stacks in TEM images as well as to answer questions from Reviewer 1, we added a total of four new figures featuring Csu pilus stacks seen in TEM images. First figure depicts the dimensions of the stacks that we have observed in vivo, as we mentioned above (Supplementary Fig. 1). Second figure depicts stacks formed in various environmental conditions (pH, excess salt, and different temperatures) (Supplementary Fig. 2). Third figure depicts Csu pilus stacks tethering bacteria in recombinant E. coli liquid culture clumps, even when the tip-subunit CsuE has a mutation hindering its ability to bind into hydrophobic surfaces (Supplementary Fig. 4). The fourth one depicts how the Csu pilus stacks link the bacteria together independently of PNAG and eDNA (Fig. 8). These additional figures should make it easier to interpret the contents of Fig. 2.\nWhen it comes to the tomogram Z-slices in Fig. 3, the stacks seen at the tomogram are all at an angle to the plane of the Z-slices, which causes an issue with detecting them, as the reviewers have well pointed out. To make them easier to understand, we averaged 20‒25 Z-slices as the reviewers suggested, showing the pili in the stacks slightly better, though unfortunately, the improvement is not much. We also updated the figure legend to indicate, what the tomogram Z-slices now show:\n“a–c, A representative tomographic slice depicting cell-cell interactions in a fresh, suspended A. baumannii colony grown at 22oC (a) and close-up views (b,c) (red rectangles) with 20‒25 Z-slices averaged to show cross-sections of the Csu pilus stacks, as indicated (yellow arrowheads).”\nSince averaging several Z-slices had less of an effect than we had hoped, we added an extra video (new Supplementary Video 2) to depict, what the Csu pili and their stacks look like in the tomogram Z-slices. The video tilts back and forth around the view seen in Fig. 3c, while stopping to point out with yellow arrows the Csu pilus stacks that remain in view longer than the individual Csu pili.\nFinally, as an additional change to Fig. 3, we added panel 3i, which depicts a stack from the 3D segmentation tilted from 0° frontal view to a 90° side view in four 30° increments. This is similar to what we already had in the current Supplementary Video 1, but we decided it would be good to have it shown in Fig. 3, as well. This should highlight the flatness of the stacks, as from a certain angle they look as narrow as a single pilus.\nIII. Additional suggestions which may improve the story/results\n29\n1. Supplementary Fig. 7 might be better as a main figure as its results are key to the model proposed in current main Figure 7.\nWe have transferred the former Supplementary Fig. 7 to the Main figures as new Fig. 7, as the reviewers had suggested. Following that, we have added a new Fig. 8 depicting how Csu pilus stacking networks are able to maintain cohesion in bacterial rafts even in the complete absence of PNAG and eDNA.\n2. The number of particles used to obtain the type 1 and type 2 pili stacks was quite a lot given the relatively low resolutions achieved. Perhaps heterogeneity is the limiting factor in terms of resolution. I wonder if more classes of pili stacks would be revealed with additional heterogeneity analyses such as cryoSPARC’s 3D variability analysis (cluster output might be most useful) or cryoDRGN?\nWe agree with the reviewers that heterogeneity is a major limiting factor for the resolution of our maps. However, according to our observations, this heterogeneity is mostly from three sources:\n1. CryoSPARC misidentifying the near-identical pilus pairs from the type 1 and 2 stacks and combining them into one map by overlapping pilus pairs from one type of stack at variable pair positions in the other stack map (Supplementary Fig. 17e, see also Fig. R3 above). This results in weakened binding junctions in places where the binding junction positions from the other type of stack do not match the correct stack, as well as weak pilus densities poking from the main map. While we sorted out most of the incorrect particles, some of them remain even in the final datasets (see Fig. R6 below).\n2. CryoSPARC misidentifying stack orientations and merging 180° flipped particles with the correctly oriented particles (Supplementary Fig. 17d). We recognized this as a similar mixture of flipped views had occurred earlier, when we tried to use SPRING image processing package in our initial attempts to solve the structure of a single Csu pilus rod (Pakharukova et al., 2022). We reduced this error by sorting the stack orientations between 180° flipped search models in Heterogeneous Refinement, while using low O-EM learning rate to prevent the still flipped particles from biasing the maps, before rotating the sorted particles and merging them in Local Refinement (Supplementary Fig. 17b, g, see also our answer to Point 4). However, the issue remains to an extent, resulting in averaged density maps with smeared resolution and weakened binding junctions.\n3. Some of the originally picked particles included pairs and individual pili crossing underneath or above the observed stacks (Supplementary Fig. 5). Filament Tracer can avoid these, but it ended up picking far too few particles (leaving out repeating stack regions perpendicular to pilus axes etc.), so we had to use Template Picking instead. Some of these crossing pili remain as background noise.\nAs the reviewers suggested, we ran cryoSPARC 3D Variability analysis with cluster output for both type 1 and type 2 stack maps. The program revealed no separate clusters that could be identified as novel stack classes (heterogeneity) or structural flexibility, as indicated by the Clustered Scatterplot and the corresponding maps (see Fig. R6 below). The minor variability detected appears to result from the remaining misidentification of the two types of stacks and stack flipping. In addition, 3D Variability analysis seems to interpret the degrading type 2 stack pilus resolution near the edge of the picked region as variation.\nWe do have additional evidence to suggest that there should not be other major stack types apart from the type 1 and 2 stacks. As we stated in our answer to the reviewers’ Point 1, our first Homogeneous Refinement of the combined stack data (Supplementary Fig. 17b) generated a map, in which the two stack density patterns are clearly visible at the same time, while overlapping at a 51.43° angle by sharing one Csu pilus pair (Supplementary Fig. 17e, see also Fig. R1b‒d above). The type 1 stack end view showed distinct 154.29° angles between three adjacent pili, while the type 2 stack had 102.86° angles, which made it easy to notice that they were different (Supplementary Fig. 17e). Refinement jobs utilizing mixed particle sets repeated this error consistently (Supplementary Fig. 7a, b, see also Fig. R3b above), so this tendency to overlap pairs from different stacks should be very strong, and therefore any unknown stack classes using the same binding junctions should produce similar unexplained density patterns around the central pair. At no stage of the refinement did we observe such unexplained densities. Furthermore, the final stack maps from cryoSPARC Local Refinement (before cropping the maps around the models) connect to additional pilus densities through binding junctions extending in the directions predicted by our stack models, while all other density blobs lack such connections to the stack densities (Supplementary Fig. 17h). This suggests that there should be no additional types of stacks, at least not in significant quantities.\nReferences used in the responses:\nAhmad I, Karah N, Nadeem A, Wai SN, Uhlin BE (2019) Analysis of colony phase variation switch in Acinetobacter baumannii clinical isolates. PLoS One 14: e0210082\nAhmad I, Nadeem A, Mushtaq F, Zlatkov N, Shahzad M, Zavialov AV, Uhlin BE (2023) Csu pili dependent biofilm formation and virulence of Acinetobacter baumannii. NPJ Biofilms Microbiomes 9: 101\nAnderson SE, Rather PN (2019) Distinguishing Colony Opacity Variants and Measuring Opacity Variation in Acinetobacter baumannii. Methods in molecular biology 1946: 151-157\nBentancor LV, Camacho-Peiro A, Bozkurt-Guzel C, Pier GB, Maira-Litran T (2012) Identification of Ata, a multifunctional trimeric autotransporter of Acinetobacter baumannii. J Bacteriol 194: 3950-3960\nBohning J, Dobbelstein AW, Sulkowski N, Eilers K, von Kugelgen A, Tarafder AK, Peak-Chew SY, Skehel M, Alva V, Filloux A et al (2023) Architecture of the biofilm-associated archaic Chaperone-Usher pilus CupE from Pseudomonas aeruginosa. PLoS Pathog 19: e1011177\nBohning J, Ghrayeb M, Pedebos C, Abbas DK, Khalid S, Chai L, Bharat TAM (2022) Donor-strand exchange drives assembly of the TasA scaffold in Bacillus subtilis biofilms. Nat Commun 13: 7082\nBrossard KA, Campagnari AA (2012) The Acinetobacter baumannii biofilm-associated protein plays a role in adherence to human epithelial cells. Infect Immun 80: 228-233\nBuzzo JR, Devaraj A, Gloag ES, Jurcisek JA, Robledo-Avila F, Kesler T, Wilbanks K, Mashburn-Warren L, Balu S, Wickham J et al (2021) Z-form extracellular DNA is a structural component of the bacterial biofilm matrix. Cell 184: 5740-5758 e5717\nCerqueira GM, Peleg AY (2011) Insights into Acinetobacter baumannii pathogenicity. 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J Bacteriol 197: 2027-2035\nGarnett JA, Martínez-Santos VI, Saldaña Z, Pape T, Hawthorne W, Chan J, Simpson PJ, Cota E, Puente JL, Girón JA et al (2012) Structural insights into the biogenesis and biofilm formation by the Escherichia coli common pilus. Proceedings of the National Academy of Sciences 109: 3950-3955\nGiles SK, Stroeher UH, Eijkelkamp BA, Brown MH (2015) Identification of genes essential for pellicle formation in Acinetobacter baumannii. BMC Microbiol 15: 116\nGiraud C, Bernard CS, Calderon V, Yang L, Filloux A, Molin S, Fichant G, Bordi C, de Bentzmann S (2011) The PprA-PprB two-component system activates CupE, the first non-archetypal Pseudomonas aeruginosa chaperone-usher pathway system assembling fimbriae. Environmental microbiology 13: 666-683\nGohl O, Friedrich A, Hoppert M, Averhoff B (2006) The thin pili of Acinetobacter sp. strain BD413 mediate adhesion to biotic and abiotic surfaces. 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J Bacteriol 195: 4146-4153\nSecond round\nReviewer 1\nThe authors have successfully implemented all necessary changes and improvements, resulting in a significantly better and more substantial paper. I recommend its publication in its current form.\nWe are pleased that the reviewer appreciated the revised version of the paper. We sincerely thank the reviewer for the thorough evaluation and for the many valuable suggestions and corrections, which have significantly improved the manuscript.\nReviewers 2 and 3\nThe authors have more than addressed all of my concerns. I view this work as impactful and within the scope of Nature Communications and I recommend this manuscript for publication. From a cryo-EM perspective the reconstruction of the different classes of pili stacks was technically challenging and impressive, despite the fact that the authors only achieved low to modest resolution of the stacks. Their high-resolution structure of the pilus rod as well as the fact that they use isolated stacks from a Csu E. coli expression system leave little ambiguities to the sequence of the pilin subunit in these structures. I have several minor suggestions that I think would improve the manuscript.\nWe are very pleased to read the reviewer’s comment that “the authors have more than addressed all of my concerns.” We are grateful for the reviewer’s thorough assessment and for the many excellent suggestions and corrections. We have addressed the new minor suggestions as follows:\n1. In Figure 5A the authors should include text displaying the resolution of each structure shown.\nThe resolution is now indicated in parenthesis next to the name of each structure shown in Figure 5. To comply with the editor’s request that each figure occupy one page, Figure 5 was divided into Figure 5 (formerly Figure 5a) and Figure 6 (formerly Figure 5b).\n2. The authors stated in their rebuttal \"To obtain most of our SPA cryo-EM structures, including the Csu pilus stacks, we expressed Csu pili using our E. coli BL21 pBAD-Csu overexpression system\". It should be made very clear in the methods which expressed systems were used for which structures.\nThis information has been added to the following sentences in the “Single-particle cryo-EM” section:\n“To determine the structure of the native Csu pilus rod, 3.5 µl of sample purified from A. baumannii CCUG 19096T strain was applied to glow-discharged 300-mesh gold holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified using a Vitrobot Mark IV (Thermo Fisher Scientific) at 4 °C and 100% humidity.”\n“To determine the structure of the Csu pilus pair and the initial structure of a Csu pilus stack, 4 µl of sample purified from E. coli BL21-AI with pBAD-Csu plasmid was applied to glow-discharged 300-mesh copper holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified in liquid ethane using a Vitrobot Mark IV (Thermo Fisher) at 4 °C and 100% humidity.”\n“To determine the final Csu pilus stack structures, we used highly concentrated fractions of Csu pili, which we purified from E. coli BL21-AI with pBAD-Csu plasmid and then stored at 4°C for over a year.”\n1"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have successfully implemented all necessary changes and improvements, resulting in a significantly better and more substantial paper. I recommend its publication in its current form.", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "The authors have more than addressed all of my concerns. I view this work as impactful and within the scope of Nature Communications and I recommend this manuscript for publication. From a cryo-EM perspective the reconstruction of the different classes of pili stacks was technically challenging and impressive, despite the fact that the authors only achieved low to modest resolution of the stacks. Their high-resolution structure of the pilus rod as well as the fact that they use isolated stacks from a Csu E. coli expression system leave little ambiguities to the sequence of the pilin subunit in these structures. I have several minor suggestions that I think would improve the manuscript.\n1. In Figure 5A the authors should include text displaying the resolution of each structure shown.\n2. The authors stated in their rebuttal \"To obtain most of our SPA cryo-EM structures, including the Csu pilus stacks, we expressed Csu pili using our E. coli BL21 pBAD-Csu overexpression system\". It should be made very clear in the methods which expressed systems were used for which structures."}, "author_response": "We are pleased that the reviewer appreciated the revised version of the paper. We sincerely thank the reviewer for the thorough evaluation and for the many valuable suggestions and corrections, which have significantly improved the manuscript.\nWe are very pleased to read the reviewer’s comment that “the authors have more than addressed all of my concerns.” We are grateful for the reviewer’s thorough assessment and for the many excellent suggestions and corrections. We have addressed the new minor suggestions as follows:\n1. In Figure 5A the authors should include text displaying the resolution of each structure shown.\nThe resolution is now indicated in parenthesis next to the name of each structure shown in Figure 5. To comply with the editor’s request that each figure occupy one page, Figure 5 was divided into Figure 5 (formerly Figure 5a) and Figure 6 (formerly Figure 5b).\n2. The authors stated in their rebuttal \"To obtain most of our SPA cryo-EM structures, including the Csu pilus stacks, we expressed Csu pili using our E. coli BL21 pBAD-Csu overexpression system\". It should be made very clear in the methods which expressed systems were used for which structures.\nThis information has been added to the following sentences in the “Single-particle cryo-EM” section:\n“To determine the structure of the native Csu pilus rod, 3.5 µl of sample purified from A. baumannii CCUG 19096T strain was applied to glow-discharged 300-mesh gold holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified using a Vitrobot Mark IV (Thermo Fisher Scientific) at 4 °C and 100% humidity.”\n“To determine the structure of the Csu pilus pair and the initial structure of a Csu pilus stack, 4 µl of sample purified from E. coli BL21-AI with pBAD-Csu plasmid was applied to glow-discharged 300-mesh copper holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) and vitrified in liquid ethane using a Vitrobot Mark IV (Thermo Fisher) at 4 °C and 100% humidity.”\n“To determine the final Csu pilus stack structures, we used highly concentrated fractions of Csu pili, which we purified from E. coli BL21-AI with pBAD-Csu plasmid and then stored at 4°C for over a year.”"}]

Decision letter