Subcellular reorganization upon phage infection reveals stepwise assembly of viral particles from membrane-associated precursors

Domain: biology
Published: 2026-04-02
Editorial outcome: minor_revision

Abstract
Viruses are obligate intracellular parasites and viral infections lead to massive host cell rearrangement to support the rapid generation of progeny. Host take-over and remodelling include formation of viral-induced compartments for viral genome replication and/or assembly. While viruses infecting bacteria, bacteriophages or phages, have been extensively characterized in vitro, the molecular mechanisms underlying the viral cycle inside the crowded cytoplasm remain unclear. Here, we investigate the spatial reorganization of SPP1-infected bacteria under near-native conditions by electron cryo tomography. The most prominent feature is the formation of a large viral DNA (vDNA) compartment from which ribosomes are excluded. In SPP1 infection, there is no membrane nor proteinaceous shell surrounding these compartments. Also, we identify novel key intermediates in virus assembly: open precursors of procapsid lattice are found at the cytoplasmic membrane in a process that requires expression of the portal protein. Next, DNA-free procapsids relocate inside the vDNA compartment where vDNA is packed in a stepwise manner. Finally, DNA-filled capsids segregate to the periphery of the compartment for assembly completion and storage. Collectively, we provide comprehensive mechanistic insights into the complete viral assembly pathway of SPP1 directly in cellula and show how specific steps are coordinated inside the reorganized bacterial cell.

Introduction
Viruses are obligatory parasites that can infect bacteria, archaea or eukaryotes, and show co-evolution with their hosts. Several viral lineages are deeply rooted with common ancestry such as tailed bacterial viruses (bacteriophages or phages) together with herpesviruses1. During lytic infection, viruses massively hijack the resources of the host towards a rapid generation of progeny. This take-over promotes reorganization of the infected cell, frequently leading to formation of compartments dedicated to viral genome replication, progeny virion assembly and/or evasion from host defence systems2. Several strategies for viral-induced compartmentation are known: membrane-bound, confined by a protein cage, formed by liquid-liquid phase separation or result of more complex phase separations as polymer compaction (e.g., DNA molecules) upon macromolecular crowding3–5.

In the case of bacteriophages, the main steps of viral multiplication were studied extensively in vitro. However, it remains largely unclear how well this represents the molecular processes actually taking place in the complex cytoplasmic environment inside infected cells. Recently, jumbo phages were shown to form a proteinaceous shell making a nucleus-like compartment, which physically separates replication and transcription of the viral genome from translation, viral particle assembly, and metabolic reactions inside the host bacterium6. This feature of giant jumbo phages (genome sizes above 200 kbp) distinguishes them from the vast majority of tailed viruses that have smaller genomes, most frequently in the 30-60 kbp size range7. Notably, infection by siphoviruses lambda8,9 and SPP110–12 also lead to the formation of a viral DNA (vDNA) compartment that is localized asymmetrically within the bacterial host cell9,11. In the case of SPP1, which infects the Gram-positive soil bacterium Bacillus subtilis13, hybrid replisomes are found at several independent locations within the vDNA compartment. Those replisomes composed of phage (gp40) and bacterial (PolC, DnaE, DnaX, DnaN, DnaG, SsbA and DnaB) replication proteins are hypothesized to represent foci of active vDNA replication events taking place in parallel10. At late stages of SPP1 infection, procapsids co-localize with vDNA for packaging10. Interestingly, the vDNA compartment does not protect the viral genome against bacterial defence mechanisms12. After vDNA packaging termination, DNA-filled viral particles are stored in so-called warehouses found outside the vDNA compartment10.

Assembly of SPP1 particles is initiated by formation of the DNA-free procapsid I, which requires three essential components: the major capsid protein gp13, the scaffolding protein gp11 that assists with gp13 polymerization, and the portal protein gp6 located at one single vertex of the icosahedral procapsid14. Release of gp11 from the procapsid I interior is accompanied by a major conformational change leading to expansion of the capsid lattice to the procapsid II state15. The portal protein gp6 forms a dodecamer with a central channel that serves as the doorway for vDNA packaging into procapsids II. Gp1, the terminase small subunit (TerS), binds specifically to the SPP1 DNA sequence pac that is subsequently cleaved by gp2, the terminase large subunit (TerL)16,17. Docking of the terminase-DNA complex to the procapsid portal vertex leads to assembly of the DNA packaging motor that translocates DNA to the procapsid interior16,17. DNA packaging is terminated by cleavage of the SPP1 DNA concatemer and departure of the terminase. Sequential binding of gp15 and gp16 to the capsid portal vertex leads to formation of the connector18 that provides the interface for attachment of the SPP1 long flexible tail, built in an independent assembly pathway19. Assembly of SPP1 infectious viral particles follows a pathway conserved among siphoviruses. However, there is a critical lack of knowledge on intermediate assembly states and how these processes are regulated in the bacterial cell. Here, we investigate the spatial reorganization of the host bacterial cell induced upon infection by the bacteriophage SPP1 and characterize key individual steps of the viral particle assembly pathway in cellula under near-native conditions. Using cellular electron cryo tomography (cryoET), we could identify assembly intermediates previously undetected in in vitro studies: formation of procapsid precursors at the cellular membrane as well as subsequent packaging of vDNA into the procapsid structure. Importantly, we also find that these viral molecular processes correlate with a specific spatial partitioning within the rearranged infected cell either at the plasma membrane, inside the vDNA compartment or confined to its periphery. Altogether, we provide comprehensive mechanistic insights into the complete assembly pathway from membrane-associated precursors of procapsids to DNA-filled head-and-tail capsids directly in the bacterial host cell.

Results
Electron cryo tomography applied to study SPP1-infected bacteria
We developed an optimized sample preparation workflow to follow SPP1 infection in cellula (Supplementary Fig. 1) and measure its impact on the remodelled infected bacteria under near-native conditions20. The well-characterized siphovirus SPP1 infects the Gram-positive soil bacterium Bacillus subtilis13. Under optimal infection conditions, SPP1 DNA is found inside bacterial cells within 3 min post infection (p.i.)21. Transcription and replication of the vDNA genome are initiated rapidly afterwards22, while expression of late genes coding for morphogenetic proteins starts at 10 to 12 min p.i.21. From 30 min p.i. onwards, infectious viral particles are formed and then released by cell lysis21. In order to visualize all states of virus assembly, we generated specimens of SPP1-infected B. subtilis cells at 15 and 25 min p.i. by plunge-freezing (see Methods). Then, thin lamellae (i.e., sections of ~100 nm in thickness) were micromachined under cryogenic conditions using a scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM) (Supplementary Fig. 1)23. Each lamella (~12 µm in width) contained cross-sections of several bacteria (Supplementary Fig. 1f) infected either with wild type (wt) or mutant SPP1 phages that were investigated by cryoET (Supplementary Table 1).

SPP1 infection leads to extensive remodelling of B. subtilis cells and formation of a prominent vDNA compartment
B. subtilis cells were imaged before and after infection with SPP1 using cryoET (Fig. 1; Supplementary Fig. 2). Tomograms of uninfected control cells display a homogeneous distribution of ribosomes in the cytoplasm (Fig. 1a; Supplementary Fig. 2a; Supplementary Movie 1), which starkly contrasts with infected cells where ribosomes are mostly excluded from a large area of the cytoplasm (Fig. 1b; Supplementary Fig. 2b; Supplementary Movie 2). There, a prominent compartment is formed that contains the vDNA (Supplementary Fig. 2b-h). Indeed, fluorescence microscopy of ribosomal proteins L1 and S2 showed that reorganization of the cytoplasmic space correlated with the exclusion of ribosomes from the phage vDNA compartment, and late in infection with the formation of adjacent warehouses where mature infectious virions are stored (Supplementary Fig. 3). Notably, these two types of viral-induced compartments are neither delineated by a lipid bilayer nor by a proteinaceous shell (Fig. 1; Supplementary Fig. 2).

Spatial coordination of the different stages of viral particle assembly inside the host bacterial cell
Direct visualization of individual phage particles in cellula using cryoET overcomes a long-standing bottleneck in the field posed by the inability to image those structures by conventional electron microscopy (EM) approaches after sample preparation on chemically fixed bacteria following infection10. In wt infections, we observed that the majority of viral particles can be assigned to several intermediates of the capsid assembly process, namely procapsid I, procapsid II, DNA-filled capsid or mature virus (Fig. 1c-f). Structures of three of those assembly states have previously been determined from purified particles by cryoEM single particle analysis15. Accordingly, procapsids I are spherically-shaped icosahedra, 55 nm in diameter, with the scaffolding proteins (SP, highlighted in blue in Fig. 1c) in the lumen. Procapsids II exhibit an angular icosahedral structure with a diameter of 65 nm and were here frequently filled with various amounts of DNA (Fig. 1d). Finally, DNA-filled capsids with similar dimensions to procapsids II (Fig. 1e) can be found at the periphery of the vDNA compartment, together with mature virions attached to a long flexible tail (Fig. 1f; Supplementary Table 1).

Precursors of SPP1 procapsids assemble at the bacterial cell membrane
Notably, partial SPP1 procapsid I-like structures were consistently associated to the inner side of the bacterial cell membrane (Fig. 2; Supplementary Fig. 4). We consider these incomplete procapsids as early precursors of SPP1 assembly. They account for more than 10% of viral particles observed in our tomograms at 15 min p.i. and about 3% at 25 min p.i. (Supplementary Table 1). Based on the statistical distribution of precursors with different degrees of completeness, it appears that initial capsomere assembly is rapid but subsequent procapsid completion and membrane dissociation might be the limiting steps (Fig. 2i). The stages of precursor assembly observed vary from about one third of an opened sphere (Fig. 2a,e; Supplementary Fig. 4a), up to closed procapsid I-like shells still bound to the membrane (Fig. 2d,h; Supplementary Fig. 4d). The diameter of those precursors (52 nm on average, SD=0.7 nm, n=20) as well as the curvature of the thick outer capsid lattice are similar to the unexpanded lattice of procapsids I observed in cellula (this work) and in the high-resolution structure previously determined in vitro15 (Supplementary Fig. 5). The lumen of precursors consists of one or two layers of, presumably, internal scaffolding proteins that serve for capsomere assembly (SP, highlighted in blue in Fig. 2e-h). A pattern of radial striations is indeed observed within the outer shell, giving it the appearance of individual concentric rods (Fig. 2a-h,j). Sub-volume averaging was performed on procapsids I found in our tomograms in order to determine the symmetry of the scaffolding layers with alignment focusing either on the icosahedral shell or on the protein scaffold (see Methods). In both cases, no structural detail of the scaffolding proteins was resolved, regardless of whether 12-fold icosahedral symmetry was introduced or not, implying that the scaffold does not follow a defined regular icosahedral organization in these procapsids but a different and possibly asymmetrical shell packing (Supplementary Fig. 5a,b).

Role of the portal protein in membrane-associated procapsid assembly
All capsid precursors observed were in close proximity to the bacterial cell membrane and connections are sometimes visible between the inner side of the cell membrane and the capsid lattice being assembled (Fig. 2a-h; Supplementary Fig. 4). These suggest that the host membrane serves as an anchor point for nucleation of procapsid assembly initiation. We hence hypothesized that the portal protein gp6 could nucleate SPP1 procapsid assembly at the bacterial membrane. Consistent with this hypothesis, it was already reported that gp6 is soluble in solution24 but can also form nanopores in membranes under certain conditions in vitro25. In order to probe the localization of isolated SPP1 portal proteins in cellula, we engineered a B. subtilis strain producing gp6 whose carboxyl terminus was fused to the fluorescent protein mCitrine (see Methods). Gp6-mCitrine is functional for SPP1 assembly as assessed in a trans-complementation assay (Supplementary Fig. 6). These cells were infected by a gp6 deletion mutant strain (SPP1gp6-), ensuring that the sole source of portals for incorporation in procapsids is the bacterial-encoded gp6-mCitrine. Epifluorescence microscopy observations showed a weak punctate fluorescence distribution in non-infected bacteria (Fig. 3a). We then used total internal reflection fluorescence (TIRF) microscopy to assess the dynamics of gp6-mCitrine at different stages of infection26. Early in infection and in non-infected bacteria, puncta of gp6-mCitrine are highly dynamic, but become immobilised in larger clusters at 23 min p.i., which coincides with viral particle assembly (Fig. 3a,b; Supplementary Movies 3 and 4). Interestingly, these fluorescent foci localised to the periphery of the vDNA compartment (white arrows in Fig. 3b). We conclude that, at this stage of infection, gp6-mCitrine proteins are mostly incorporated in viral procapsids that mature into virions and are subsequently stored in immobile warehouse compartments that were previously characterized10.
We next investigated whether procapsid assembly occurs at the membrane in absence of the portal protein. In B. subtilis cells infected with SPP1gp6-, meaning conditions under which the portal is not produced (Fig. 3c; Supplementary Fig. 2c; Supplementary Movie 5), we found that only procapsid I-like particles (Fig. 3e), smaller procapsids (Fig. 3f) and some aberrant particles (Fig. 3g) were present (Supplementary Table 1). These observations are consistent with former studies showing that SPP1 procapsid-like structures can assemble in absence of the portal protein but that the portal is necessary for faithful size determination of procapsid size14. Strikingly, no procapsid assembly intermediates were observed to be associated with the cell membrane in the 58 tomograms analysed (Supplementary Table 1). We hence hypothesize that the incomplete, open particles rarely spotted in the cytoplasm and in the vDNA compartment (n=5) represent intermediates of an alternative, gp6-independent assembly pathway under these specific conditions (e.g., Fig. 3h,i). This previously unsuspected requirement of gp6 for localization of procapsid precursors at the inner side of the bacterial cell membrane strongly suggests that it plays a key role in regulating and driving the early stages of SPP1 procapsid assembly. Such finely tuned mechanism would favour in particular the incorporation of one portal at the initiation of assembly, prior to the icosahedral lattice polymerization step.

Concomitant DNA packaging and capsid expansion occur after relocalization of procapsids to the vDNA compartment
To determine the spatial distribution of procapsids in absence of any vDNA packaging events, we imaged bacteria infected by SPP1 conditional mutants defective for expression of the viral terminase components TerS or TerL (gp1 and gp2 in SPP1, respectively; see Methods). In SPP1gp1- and SPP1gp2- infections, capsid precursors are found associated to the membrane and procapsids I in the vDNA compartment, in a manner similar to wt virus infection (Fig. 4a; Supplementary Fig. 2d,e; Supplementary Movie 6). Hence, detachment of fully assembled procapsids I from the membrane and their relocation to the vDNA compartment occur independently of subsequent vDNA encapsidation events (Supplementary Table 1). Notably, procapsid-like structures without portal assembled during SPP1gp6- infection are also found in the vDNA compartment (blue in Fig. 3c; Supplementary Fig. 2c) demonstrating that their specific targeting to the compartment is independent of the portal, and likely relying on the procapsid icosahedral lattice properties.
The transition from procapsid I to procapsid II is marked by capsid expansion and release of the scaffolding proteins15. The topography of the particle surface changes to a thinner and smoother lattice in procapsid II, while maintaining the overall electrostatics potential distribution (Supplementary Fig. 7). Procapsids II were present neither in SPP1gp1- nor in SPP1gp2- infections (Fig. 4a,b; Supplementary Fig. 2d,e; Supplementary Table 1) showing that vDNA packaging is essential to trigger expansion of the procapsid lattice in cellula (Fig. 4c-g). Nevertheless, in a previous study, expanded procapsids II were found in extracts of lysed bacteria from a SPP1gp1- infection15. Those procapsids II did not contain scaffolding proteins associated to capsid hexamers but the ones bound to the vertex pentamers were still present. Given our new findings, it seems that the disruption of the cellular environment by lysis was the trigger for procapsid I to procapsid II transition in absence of vDNA packaging. This discrepancy to earlier findings highlights the limitations of relying uniquely on in vitro structural biology to investigate such complex molecular processes.
In SPP1wt infection, procapsids II, compared to procapsids I, display a larger, more angular shape similar to the structures determined in vitro15. In snapshots of vDNA packaging found in tomograms, the genome appears as thin threads inside procapsids II, which all have the same dimensions (63 nm in diameter ±1.1 nm; n=37). Altogether, our findings suggest that expansion is essentially a single transition event triggered by vDNA packaging initiation. Different steps of genome encapsidation were visible from nearly empty to almost full icosahedral capsids (Fig. 4d-g). The viral genome appears to occupy the overall internal procapsid II space adopting progressively a more compact organization and no common or regular DNA encapsidation pattern was observed inside those packaging intermediates (Supplementary Fig. 8).

Segregation of DNA-filled capsids to the periphery of the vDNA compartment and binding to pre-assembled tails
We noticed that procapsids I and II are confined at 97% (n=1478) and 94% (n=225), respectively, to the vDNA compartment consistent with the localization of vDNA encapsidation events mediating the transition between the two procapsid stages (Supplementary Table 1). In contrast, 40% of DNA-filled capsids (n=604) distinguishable by their electron dense inner content (Figs. 1e, 4g) and the vast majority of tailed virions, 84% (n=113), were located outside of this compartment, mostly at its periphery (Fig. 5a,b; Supplementary Table 1). We conclude that capsids depart from the vDNA compartment after packaging of the phage genome and that tail attachment to DNA-filled capsids takes place outside of the vDNA compartment but adjacent to it (Supplementary Fig. 9a,b). These late events of viral assembly lead to the appearance of foci detected with a fluorescently labelled marker of DNA-filled capsids10 such as gp12-mCitrine27. By fluorescence microscopy, the foci are located in the vicinity of the vDNA compartment (Supplementary Fig. 3). In our tomograms, those regions appear as locally enriched in individual phage particles next to some pre-assembled tails in SPP1wt infections (Fig. 5a; Supplementary Fig. 2f; Supplementary Movie 7). In contrast, during infection with a SPP1gp12- mutant strain, defective in production of the capsid surface protein gp12, viral capsids are packed as large and ordered three-dimensional arrays termed warehouses observed in 15 out of 25 tomograms (Fig. 5b; Supplementary Fig. 2g; Supplementary Movie 8). We confirm that formation of phage warehouses is a result of capsid-capsid interactions depending on their surface properties and on the presence of the auxiliary protein gp1227. Indeed, particles lacking gp12 arrange as warehouses or clusters, which are significantly more compact than those containing gp12 (compare Fig. 5a and b).
Free tails are rare in infected cells when the whole assembly process of viral particles takes place such as in wt infections but can sometimes be observed at the periphery of the vDNA compartment (Fig. 5a). In contrast, clusters of tails are visible in the infected cell cytoplasm - again next to the vDNA compartment - when capsid assembly is arrested prior to tail-capsid association (Fig. 5c; Supplementary Fig. 2h; Supplementary Movie 9), as reported previously10. We conclude that phage tails are pre-assembled outside the vDNA compartment and bind to DNA-filled capsids after they depart from the vDNA compartment, leading to the formation of infectious particles in a spatially coordinated manner (Fig. 6).

Methods
Bacterial strains and growth conditions
Bacterial strains used in this work are listed in Supplementary Table 2.
B. subtilis GSY10027 was constructed by transformation of B. subtilis GSY10004 with plasmid pAL27 linearized with ScaI. Double cross-over led to insertion of the gene coding for gp6-mCitrine under control of the inducible promoter P at the amyE locus.
xyl
B. subtilis GSY10082 and GSY10087 were constructed in three steps. First, GSY10074 and GSY10085 were constructed by transformation of B. subtilis YB886 with plasmids pLG51 and pLG50, respectively, linearized with ScaI. Double cross-over led to insertion of genes coding for RpsB-mCFP and RplA-mCFP, respectively, under control of the inducible promoter P at the amyE locus. Then, GSY10075 and GSY10086 were constructed by transduction of GSY10074 and GSY10085, respectively, with a lysate of SPP1 wild type (wt) amplified in strain GSY10004. The resulting strains encode LacIΔ11-mCherry under control of the constitutive promoter P at the pen thrC locus. Finally, GSY10082 and GSY10087 were constructed by transduction of GSY10075 and GSY10086, respectively, with a lysate of SPP1wt amplified in strain GSY10024. The resulting strains encode gp12-mCitrine under control of the inducible promoter P at the sacA locus. Expression from promoters P and P was induced with 1% (w/v) xylose and 1 mM IPTG, respectively.
spac xyl spac
Transformation of competent B. subtilis cells was performed by a two-step starvation protocol using Spizizen medium for growth44. Linearized plasmid (1 µg) was added to 500 µL competent cells for 30 minutes at 37°C, plated on LB plates supplemented with 5 µg/mL kanamycin for GSY10027 or 100 µg/mL spectinomycin for GSY10074 and GSY10085 and incubated overnight at 37°C.
Transduction of B. subtilis cells was performed using a previously published protocol11. Briefly, to produce SPP1 lysates, donor strains GSY10004 and GSY10024 were grown overnight at 30°C in LB medium. Cultures were diluted 1:100 in 10 mL of LB medium and grown at 37°C. At an OD of 0.8, cultures were supplemented with 10 mM CaCl , infected with SPP1wt at an input multiplicity of 5 pfu/cfu (i.e., plaque forming unit over colony forming unit) and incubated for 2 h at 30°C. Cell debris were sedimented by centrifugation at 4°C for 15 min at 8,000 g and the supernatant was stored at 4°C. Then, the SPP1 transducing lysates were used to infect 300 µL of a late-logarithmic-phase culture (OD of ~1.6) of the receptor strain supplemented with 10 mM CaCl at an input multiplicity of 1. After 10 minutes at 37°C, the infected culture was mixed with 1.5 mL of prewarmed LB medium and further incubated with shaking for 10 minutes. Bacteria were sedimented and resuspended in 300 µL of prewarmed LB medium containing 40 μL of anti-SPP1 serum to inactivate free phages. Bacteria were incubated for 30 minutes, sedimented, and resuspended in 300 mL of prewarmed LB medium. Serial dilutions of the culture were plated in solid medium supplemented with appropriate antibiotics and incubated at 37°C overnight.
Antibiotic selection was 12.5 µg/mL lincomycin + 0.5 µg/mL erythromycin (mls resistance) and 5 µg/mL kanamycin (kan resistance).

Plasmid construction
Plasmids and oligonucleotides used in the present study are listed in Supplementary Table 2 with references.
SPP1 gene 6, coding for the portal protein, was amplified by PCR from SPP1lacO64 DNA with primers 55 and 56. The PCR product digested with ClaI-XhoI was ligated to pAL21 cut with the same restriction enzymes to generate plasmid pAL27.
Genes rplA, coding for ribosomal protein L1, and rpsB, coding for ribosomal protein S2, were amplified by PCR from B. subtilis strain YB886 DNA with primers 20 and 28, 22 and 29, respectively. PCR products digested with KpnI-EcoRI for rplA or with KpnI-XhoI for rpsB were ligated to pAL29 cut with the same enzymes, generating pLG50 and pLG51, respectively. These constructs encode ribosomal proteins L1 and S2, respectively, fused to CFP at their carboxyl terminus. The cfp synthetic gene was codon-optimized for B subtilis. Fusion genes were expressed under the control of a xylose-inducible promoter.
Plasmid constructions were transformed into Escherichia coli DH5α and selected on LB plates supplemented with 100 µg/mL ampicillin. Clones were checked by DNA sequencing.

Construction of phage strains
Phage strains used in this work are listed in Supplementary Table 2.
Phage SPP1delX110lacO64sus115 (abbreviated SPP1lacO64gp6-) was generated by co-infecting the permissive B. subtilis strain HA101B with phages SPP1delX110lacO64 (abbreviated SPP1lacO64) and SPP1sus115 (abbreviated SPP1gp6-) at an input multiplicity of 10 pfu/cfu for each phage strain. Co-infection was carried out in LB medium supplemented with 10 mM CaCl at 37°C for 2 hours. The cell lysate was centrifuged at 12,000 g at 4°C for 20 minutes and the supernatant was titrated using bacterial strain HA101B. Ninety-six isolated phage plaques were collected and resuspended separately in 200 µL of TBT buffer (100 mM Tris-Cl pH 7.5, 100 mM NaCl, 10 mM MgCl ) in a 96-well plate. Phage clones were spotted in the non-permissive strain YB886, in the permissive strain HA101B and in the complementing strain YB886 (pSPW7) (Supplementary Table 2). Phages carrying mutation sus115 were identified by not multiplying in strain YB886. These clones were then screened by PCR to identify those carrying insertion lacO64 (primers 3185 and 3682; Supplementary Table 2) and deletion delX combined with insertion lacO64 (primers 2677 and 3383; Supplementary Table 2)11 Selected clones were re-titrated to obtain single plaques of pure clones, their genotype confirmed as above and amplified in bacterial strain HA101B.

Fluorescence microscopy
Fluorescence microscopy of GSY10082 and GSY10087 strains was performed as previously described10. Briefly, cultures of B. subtilis strains grown overnight at 30°C were diluted 1:100 in fresh LB medium with 1 mM IPTG and 0.5% xylose and grown at 37°C. At an OD of 0.8, cultures were supplemented with 10 mM CaCl and infected with SPP1lacO64gp6- at an input multiplicity of 1. Infected cultures were grown at 37°C with orbital shaking for 22 min. Infected cells were mounted on agarose pads and microscopy was carried out subsequently at room temperature. Image acquisitions were performed at early (25-35 min p.i.) and late (50-60 min p.i.) time-points after phage infection on a Zeiss Axio Observer Z1 microscope with 63X oil objective.
For Total Internal Reflection Fluorescence Microscopy (TIRFM) experiments, samples of bacterial cells were cultivated and infected with an input multiplicity of 2 pfu/cfu. After ~23 minutes of incubation at 37°C with orbital shaking, cells were spotted on a 1% agarose pad (thickness of 2 mm) diluted in CH medium45. Initial single exposure images were obtained with conventional epifluorescence images using 100 ms-long exposures at 488 nm and 561 nm. Then, time-lapse images were collected with TIRF using 100 ms-long exposures at 488 nm and 561 nm with an image taken every 2 seconds for 1 or 2 minutes. Final control images illuminated by epifluorescence were collected from a single exposure of 100 ms at 488 nm. TIRFM imaging was performed using a Nikon Ti-Eclipse microscope with a 100X oil objective (NA: 1.3; WD: 0.2 mm; CFI Plan Fluor - Phase - Nikon) and an EMCCD camera (Andor). Snapshots were extracted from time-lapse movies.

Electron cryo tomography
Sample preparation: B. subtilis strain GSY10024 was cultivated in LB medium supplemented with 1 mM IPTG at 37°C. At OD =0.8 (~108 cfu/mL), the culture was supplemented with 10 mM CaCl and cells were infected by SPP1 phages at 37°C for ~15 minutes (early time point) or ~25 minutes (late time point) at an input multiplicity of 5 pfu/cfu. Cells were then centrifuged at 16,200 g for 1 minute at room temperature and pellets were resuspended in MIII medium46 in 1/10 of the culture initial volume. Non-infected cells were treated the same way as a control. For electron cryo tomography (cryoET), 4 µL of infected or non-infected bacteria were added to electron microscopy (EM) grids (Quantifoil Micro Tools GmbH; Cu R2/2) before blotting for 20 s on one side and plunge freezing using a Leica EM GP plunger (Leica Biosystems Nussloch GmbH) (Supplementary Table 3). Grids were clipped using C-clips and autoGrids in the AutoGrid assembly workstation (ThermoFisher Scientific). Lamellae were prepared in an Aquilos 2 (ThermoFisher Scientific) equipped with a gallium focused ion beam (FIB in short), used to make thin sections of cells called lamellae by ablating cellular material around the area of interest. For this, sample preparation, milling and polishing steps were done automatically using Maps and AutoTEM software (ThermoFisher Scientific) with a milling angle target of 8 degrees (2 degrees of tolerance) and a final lamella thickness set to 120 nm. The grids were retrieved after polishing of the lamellae and stored for a minimal amount of time in liquid nitrogen prior to data collection.
Data acquisition: Grids with milled lamellae were transferred for tilt-series acquisition to a Titan Krios G3 operated at 300 kV (ThermoFisher Scientific), equipped with a K3 direct electron detector and a post-column BioQuantum energy filter (Gatan). Grids were screened and data acquired using SerialEM47. Grid scans were acquired to locate the lamellae. Tilt-series were collected using a dose-symmetric tilt scheme48 from 60 to -60 degrees specimen tilt range with a 3 degree increment step starting with 7-10 degree of pre-tilt (according to the specific milling angle of each lamella). Frames were acquired at 15 e-/pixel/s in electron counting mode with a pixel size of 3.36, 2.15 or 1.37 Å and aligned using SerialEM implementation (see Supplementary Table 3). A 70 μm objective aperture and a 20 eV energy slit were inserted during data acquisition. A nominal defocus range between 4 and 5 µm was used.
Tomogram reconstruction: Reconstruction of tomograms from tilt-series was done using IMOD version 4.12.4049 or Aretomo50 when no fiducial-like feature were visible on the projection images. For visualization purposes, we typically used a binning of 4 and deconvolution using the standard parameters of ISONET without missing wedge compensation51.
Sub-volume averaging of procapsid I: 572 procapsids I were manually picked using IMOD49 from tomograms binned by 4 (n = 30; pixel size = 8.6 Å) and separated in two halves for independent processing done in PEET 1.14.152,53. For each dataset, the initial reference was the in vitro determined map of Procapsid I (EMD-4717)15 lowpass filtered to 55 Å. This reference was used for averaging with particles having randomized orientations using custom scripts based on TEMPy54 with a box size of 80*80*80 voxels. The major capsid protein lattice of this average was segmented using the TSegger tool55 in Chimera56, to obtain a mask for subsequent alignment. The whole capsid average volume using this mask converged to (stabilised on) an icosahedral structure. Alignment was then focused on the 12 icosahedral vertices using custom scripts based on TEMPy, resulting in 6864 particles extracted with a box size of 80*80*80 voxels. Further iterative alignment steps were done until convergence. The final average volume reached a resolution estimated to 26.5 Å by Fourier Shell Correlation using PEET calcUnbiasedFSC.
Volume visualisation: Surface rendering of volumes was performed using ChimeraX. Segmentation of bacterial cell membrane was done using drawing tools in 3dmod. Average volumes of procapsids I, procapids II, mature virions, and ribosomes were placed back within each tomogram using coordinates and orientations obtained by sub-volume averaging with PEET 1.14.152,53 using custom scripts based on TEMPy54 as described above. Here, sub-volume averaging was performed using the following references: procapsid I (EMD-4717), procapsid II (EMD-10002), mature virions (EMD-4716), and ribosomes (EMD-5787). Template matching for ribosomes was done by converting heavily filtered and thresholded volumes to model point coordinates and aligning these to a filtered reference. After cleaning by cross-correlation and removal of duplicate particles, the resulting sub-volumes were manually curated from false positive and plotted back in the original tomogram coordinate system.

Statistical analysis
Statistical analysis was performed using RStudio software. All data is represented as a boxplot: the full line represents the value of the median and each point corresponds to one tomogram. Shapiro and Bartlett’s tests were used to evaluate normality and variance homogeneity. As the data distribution was not normal (p-value<0.05), non-parametric tests were used for statistical analysis. Statistical significance for the percentage of the number of events observed for the different assembly intermediates inside the vDNA compartment was evaluated using a Kruskal-Wallis test (non-parametric one-way ANOVA test) followed by a pairwise, two-sided, Wilcoxon post-hoc test with Benjamini-Hochberg (BH) correction. Tomograms presenting aggregates of viral particles (i.e. warehouses) were removed from this Kruskal-Wallis analysis for accurate quantification of viral particles. In addition, statistical significance of the ratio of precursors compared, this time, to the total number of capsids was evaluated using an unpaired, two-sided Wilcoxon test (non-parametric t-Student test). Statistical significance is assessed by a p-value < 0.05.

Data visualization
The capsid surface electrostatic potential was calculated and displayed using PyMOL57. Figures were prepared with Adobe Illustrator, BioRender, Pymol and Chimera56.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Corroyer-Dulmont et al. report cryo-EM results of seven different SPP1-infected Bacillus samples (gp1⁻, gp2⁻, g12⁻, gp6⁻_15min, gp6⁻_25min, wt_15min, and wt_25min), derived from more than 200 cryo-electron tomograms. Their perseverance in collecting such a large dataset is admirable—particularly since all samples were prepared as FIB-milled lamellae to achieve sufficient contrast for downstream analysis.\nPrevious studies of siphovirus capsid maturation have relied primarily on in vitro structural analyses or in vivo low-resolution fluorescence microscopy. In contrast, electron cryo-tomography (ECT) provides in situ, high-resolution visualization. Their ECT observations are consistent with earlier in vitro studies (procapsid to DNA-filled capsid transition) and in vivo microscopy (viral DNA compartment formation). However, ECT uniquely allows direct in situ visualization and quantitative analysis.\nKey findings from their ECT study include:\n1. The membrane-less viral DNA compartment does not require a proteinaceous shell.\n2. Procapsids are located within the vDNA compartment, while mature capsids are positioned at the periphery.\nThe most novel discovery (in my opinion) is the suggestion that gp6 initially localizes to the cell membrane, where it initiates procapsid assembly. This study represents a landmark demonstration of how ECT, when combined with optimized sample preparation, can reveal subcellular reorganization during phage assembly within the host.\nWhile the manuscript presents valuable findings, several issues should be addressed before publication.\nMajor\n1. Background Information – The manuscript lacks sufficient background prior to presenting results. Since phage protein nomenclature (gene products) is phage-specific, the authors should describe the overall capsid and portal architecture, including the relevant gp proteins. Prior “in vitro” work on the procapsid-to-mature capsid transition should also be summarized. A bit more in-depth summarization of previous in vitro structural and biochemical studies would help readers appreciate the novelty of the present data.\n2. Virus-Induced Compartmentation – In the first two paragraphs, the authors focus on viral-induced compartmentation, emphasizing proteinaceous shells (giant jumbo phage), but provide minimal context on SPP1. Only two sentences touch upon previous SPP1 studies, without explaining what was proposed or observed. Since SPP1 vDNA compartmentation has been reported previously, more detail is required before presenting the “near-native” high-resolution cryoET results. This would make the breakthrough clearer to the field.\n3. SPP1 vDNA compartment– While cryoET results demonstrate that SPP1 vDNA compartments are neither membrane- nor shell-bound, the authors should discuss potential mechanisms separating vDNA from host cytosol. Some form of validation or supporting data would strengthen the interpretation.\n4. Procapsid Assembly – The gp6⁻ mutant results clearly suggest gp6 localizes to the cytosolic membrane to initiate procapsid assembly. However, portal assembly involves additional proteins, including motor proteins. What evidence supports the claim that only gp6 is required for initiation?\n5. Oversimplified Figure 6 – Figure 6 illustrates capsid maturation, but the model may oversimplify the process. CryoET lacks sufficient resolution to identify all components involved, and such conclusions often require knockout mutants combined with cryoET. The discussion should acknowledge these limitations and somehow illustrate them in figure 6 to avoid over-interpretation.\n6. Unexplained Observations – Several findings require further explanation or discussion:\no Procapsid positioning: Why are procapsids inside the vDNA compartment while mature capsids remain outside? How does DNA packaging drive this spatial transition? What surface properties change, and the evidence/reference for that?\no gp6 dynamics: If gp6 initially binds the membrane to initiate procapsid assembly, how is it released so procapsids can enter the vDNA compartment? Are additional proteins involved? References should be cited.\no Other assembly factors: Proteins such as terminase and motor proteins are required for DNA packaging. Where are they localized during infection? Again, those known components for capsid maturation should be included in Figure 6 and discussed in the discussion part.\nMinor\n1. Extended Data Fig. 6 – The scale bar (20 nm) in panel a appears inconsistent in length compared with b and c.\n2. Line 306 – The phrase “Altogether, we new complementary findings provide a more comprehensive understanding of virus-host interactions” should be revised. The only virus–host interaction (in my opinion) described is gp6 binding to the host membrane.\n3. References – Line 3042 mentions “phage have been studied over the past 6 decades,” yet only 31 references are cited. Additional relevant literature should be included.\n4. Tomogram Data –Providing Z-stack series with segmentation would strengthen the manuscript.\n5. Number of Bacteria Observed – Authors note that each tomogram contains several bacteria. It would be better to summarize the number of bacteria observed in the extended data figure 2 (this should be table, not figure)\n6. Warehouse Observation– The “tightly packed warehouse” shown in gp12- strain (Fig. 5, Extended Data Fig. 3g). Was this observed only once, or repeatedly? The frequency of observation should be reported.", "reviewer_2": "The authors used cryo-electron tomography of Bacillus subtilis infected with bacteriophage SPP1 to investigate the assembly pathway of the virus in cellula. They show that the portal protein gp6 associates with the inner bacterial membrane, where it initiates nucleation of the procapsid. The assembled procapsid then relocates to the viral DNA (vDNA) compartment, where genome packaging takes place. Capsid auxiliary protein Gp12 prevents the aggregation of DNA-filled capsid. DNA-filled capsids subsequently attach to tails that are assembled separately in the cytoplasm. The observations of membrane-associated procapsid formation and genome packaging within a distinct vDNA compartment are novel and provide important insights into phage assembly in cellula. While the authors have done an excellent job visualizing the phage assembly pathway using cryo-ET, several aspects that could not be directly addressed by this technique remain unclear and would benefit from further discussion.\nHow does the vDNA compartment differ conceptually and functionally from the ‘replication centers’ described in other systems, given that it lacks any defined physical boundary?\nA discussion of possible mechanisms leading to boundary-less compartmentalization of vDNA may help. Can it be DNA condensation or liquid-liquid phase separation-like processes, as proposed for other phage systems?\nAdditional discussion would be helpful to clarify whether the vDNA compartment functions to concentrate viral or host enzymes involved in DNA replication and packaging, to locally enrich nucleotides or ATP required for DNA translocation, or to protect intermediates from host defense mechanisms. It is possible that resolving this is far beyond the scope of the present paper but this should be stated.\nProcapsid formation at the inner membrane: It would strengthen the discussion to include potential functional advantages of initiating procapsid assembly at the inner membrane. For example, the membrane could provide a scaffold that spatially organizes portal and capsid protein assembly, ensuring the incorporation of a single portal vertex per procapsid and preventing the formation of incomplete or aberrant structures.\nWhile gp6 appears to play a key role in membrane-associated initiation of procapsid assembly, the molecular nature of its interaction with the membrane remains speculative. Have authors analyzed the gp6 12-mer portal structures for the protein surface that can potentially interact with the membrane?\nFurthermore, the mechanism by which the portal protein detaches from the membrane after procapsid completion is not discussed. Could changes in membrane tension, capsid curvature, or scaffold release serve as potential triggers?\nMinor Comments\n• In Figure 1 (panels b1–b3), the vDNA compartment is highlighted in red. The same highlighting could be added to the back-plotted tomogram shown in Figure 1b for clarity. In fact, highlighting the vDNA compartment in all subsequent back-plotted tomogram figures would improve consistency and readability throughout the paper.\n• The authors report using C12 symmetry for subvolume averaging of the procapsid I (lines 134–140). It should be clarified why only C12 symmetry was applied, and whether other symmetry options (e.g., C5, C6, or asymmetric reconstructions) were tested.", "reviewer_3": "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_4": "In the manuscript “Subcellular reorganization upon phage infection reveals stepwise assembly of viral particles from membrane-associated precursors,” Corroyer-Dulmont et al. describe the in situ assembly of the SPP1 phage particle using cryo-electron tomography (cryoET). The excellent cryoET work is supported by well-executed fluorescence microscopy experiments and functional assays. The authors demonstrate that the SPP1 assembly pathway is consistent with those of other long-tailed phages, which were previously inferred from in vitro studies and thus carried a certain degree of ambiguity. Notably, in this manuscript, many steps of the assembly pathway have been visualized under native conditions for the first time.\nThe authors find that different components of the virion are assembled at spatially separated areas of the cell, which they refer to as “compartments.” However, these “compartments” are not separated from the rest of the cytoplasm by either proteinaceous or lipid “walls,” unlike those of jumbophages. In principle, there would be no need to bring up jumbophages if the SPP1 “compartments” were referred to by terms that do not imply a physical barrier - such as “area,” “region,” or “subvolume.”\nThis is my only, and very minor, comment on an otherwise excellent manuscript. I am happy to congratulate the authors on this outstanding study!"}, "author_response": "We thank the referee for the detailed revision of our work and the summary highlighting its novelty. We address below the different queries raised that helped to improve the manuscript.\nMajor\nQ1.R1 (query 1, referee 1): Background Information – The manuscript lacks sufficient background prior to presenting results. Since phage protein nomenclature (gene products) is phage-specific, the authors should describe the overall capsid and portal architecture, including the relevant gp proteins. Prior “in vitro” work on the procapsid-to-mature capsid transition should also be summarized. A bit more in-depth summarization of previous in vitro structural and biochemical studies would help readers appreciate the novelty of the present data.\nA1.R1 (answer 1, referee 1): We thank the referee for her/his useful suggestion. We agree that the background we initially provided on capsid assembly and its effectors was limited. We have now included a paragraph in the Introduction containing such information for SPP1 in more details in the revised version of the manuscript. It provides current knowledge on capsid assembly of tailed phages together with references that are relevant to the work presented in our manuscript (lines 72-88 of the revised manuscript with visible modifications).\nQ2.R1: 2. Virus-Induced Compartmentation – In the first two paragraphs, the authors focus on viral-induced compartmentation, emphasizing proteinaceous shells (giant jumbo phage), but provide minimal context on SPP1. Only two sentences touch upon previous SPP1 studies, without explaining what was proposed or observed. Since SPP1 vDNA compartmentation has been reported previously, more detail is required before presenting the “near-native” high-resolution cryoET results. This would make the breakthrough clearer to the field.\nA2.R1: We thank the referee for this suggestion to highlight previous research that revealed SPP1-induced compartmentalization of the Bacillus subtilis cytoplasm. The initial findings that made the foundation for our work are now described in more detail in lines 61-71 of the Introduction part in the revised version of the manuscript.\nQ3.R1: 3. SPP1 vDNA compartment– While cryoET results demonstrate that SPP1 vDNA compartments are neither membrane- nor shell-bound, the authors should discuss potential mechanisms separating vDNA from host cytosol. Some form of validation or supporting data would strengthen the interpretation.\nA3.R1: This is indeed an important question in our opinion too. Our current hypothesis is that the large vDNA compartment is generated by folding of DNA to a phase condensed state leading to separation from the rest of the components of the cytosol. However, local regions within this compartment might have different biophysical properties. We address these issues in the Discussion of the revised text (lines 289-307): “Nevertheless, the SPP1 vDNA compartment still excludes most of the cytoplasmic content such as ribosomes (Fig. 1b; Extended DataSupplementary Fig. 34) and consequently proteinwith their associated translation reactionsmachinery, as well as certain stages of viral particle assembly (Fig. 6). The vDNA compartment is a large, and somehow irregular in shapefeature that persists throughout infection (Supplementary Fig. 2b-h). Its asymmetric position within mono-infected bacteria, there is a single vDNA compartment that persists mainly immobile throughout infection. Its asymmetric positioning in the cell is probably defined by the positionmay be linked to the site of viral DNA ejection during entry10,11. Overall, This global behaviorthis virus-induced cellular compartmentalization can be explained by could arise from the folding of the large replicated SPP1 DNA concatemers tointo a phase-separated condensed state within the confined space of the crowded cellular environmentcytoplasm410, as predictedin line with predictions byfrom polymer dynamics simulations5,104,5. SuchAn analogous mechanism was also proposed to drive bacterial nucleoid condensation (Surovetsev and Jacobs-Wagner 2018). Notably, tThe previous finding that replisome proteins form discrete foci associated withto vDNA (Labarde et al 2021) is compatible with the hypothesis that the compartment might havecontains regionssub regions with differentdistinct biophysical properties, including potentially more dynamic zones arising fromof liquid-liquid phase separation5,28. Further investigation will be necessary to assess such global and local properties leading tothat drive compartmentalization of vDNA and its associated transactions during infection.”\nTesting of this model will require extensive investigation of the dynamics of DNA molecules and different protein components located in the vDNA compartment using a combination of approaches like FRAP, FCS and SPT that represent future research avenues of high interest.\nQ4.R1: 4. Procapsid Assembly – The gp6⁻ mutant results clearly suggest gp6 localizes to the cytosolic membrane to initiate procapsid assembly. However, portal assembly involves additional proteins, including motor proteins. What evidence supports the claim that only gp6 is required for initiation?\nA4.R1: In our opinion, there are three lines of evidence, which taken together, support that the SPP1 portal protein gp6 is the initiator of procapsid assembly. First, co-production of gp6, the scaffolding protein gp11 and the major capsid protein gp13 in the heterologous bacterium Escherichia coli is necessary and sufficient to assemble procapsids that are competent to package phage DNA leading to the formation of infectious phages (Ref. 14: Dröge et al 2000). This identifies the minimal set of phage proteins necessary for SPP1 procapsid assembly. Second, the SPP1 portal is necessary to regulate the size of the viral procapsid icosahedron (Ref. 14: Dröge et al 2000; Fig. 3), a feature that is determined at an early stage of procapsid lattice formation. Third, in the present work, it is shown that procapsid initiation at the membrane occurs in infection with mutants SPP1gp1- and SPP1gp2- that are recombinant strains targeting the SPP1 DNA packaging motor proteins (Fig. 4a; Supplementary Table 1). Finally, extensive in vitro work on bacteriophages phi29 and P22 made by the teams of Peter Prevelige and Carolyn Teschke also showed that the portal initiates procapsid assembly in these phages (lines 330-335 of the revised manuscript Discussion).\nQ5.R1: 5. Oversimplified Figure 6 – Figure 6 illustrates capsid maturation, but the model may oversimplify the process. CryoET lacks sufficient resolution to identify all components involved, and such conclusions often require knockout mutants combined with cryoET. The discussion should acknowledge these limitations and somehow illustrate them in figure 6 to avoid over-interpretation.\nA5.R1: We acknowledge the view of the reviewer. We would like to emphasize that figure 6 is only meant as a schematic representation of our proposed model, which is indeed based on cryoET analysis combined with characterization of recombinant mutant viruses. To highlight the limitations pointed out by the referee, Figure 6 was slightly improved and we added more information to the figure legend. This includes a sentence on the limited resolution to image some of the viral components including the terminase for example: “This schematic representation is based on our observations by cryo electron tomography of B. subtilis cells infected by SPP1 wild type combined with the characterization of virus recombinant strains SPP1gp6-, SPP1lacO64gp2-, SPP1gp1-, SPP1lacO64gp12-. Note that macromolecular complexes such as the terminase powering DNA translocation through the portal of procapsid II is certainly associated to those structures as represented in the panel (gp1 and gp2) but our cryoET imaging is limited in resolution and does not allow their direct visualization.”.\n6. Unexplained Observations – Several findings require further explanation or discussion:\nQ6.R1: o Procapsid positioning: Why are procapsids inside the vDNA compartment while mature capsids remain outside? How does DNA packaging drive this spatial transition? What surface properties change, and the evidence/reference for that?\nA6.R1: Our data show that procapsids I are strongly enriched in the vDNA compartment and that such localization is independent of the presence of the portal. This localization depends hence exclusively on procapsid I properties and conceivably of its external lattice that is in contact with the surrounding DNA molecules. Analysis of the electrostatic potential of procapsid I, procapsid II and the DNA-filled capsid, did not reveal any striking difference that could explain the association/dissociation to DNA of those structures. The properties of the diffusion of such structures inside or outside the vDNA compartment remains to be determined. These problematics are now also addressed on lines 356-367 of the revised Discussion.\nQ7.R1: o gp6 dynamics: If gp6 initially binds the membrane to initiate procapsid assembly, how is it released so procapsids can enter the vDNA compartment? Are additional proteins involved? References should be cited.\nA7.R1: This is a great question. Presently we can only speculate. Our hypothesis is that closure of the procapsid lattice to fully embed the portal will promote release of the complete structure from the membrane: “Lattice cClosure of the procapsid lattice around the portal is proposed to promote release of procapsid I from the membrane.” (lines 330-331 of revised Discussion).\nQ8.R1: o Other assembly factors: Proteins such as terminase and motor proteins are required for DNA packaging. Where are they localized during infection? Again, those known components for capsid maturation should be included in Figure 6 and discussed in the discussion part.\nA8.R1: We anticipate that the viral DNA packaging proteins gp1 (TerS; specific SPP1 DNA binder) and gp2 (TerL; endonuclease and DNA packaging motor) are associated to procapsids in SPP1 DNA packaging intermediates, a hypothesis that is schematized in the revised Fig. 6. However, the resolution of our cryoET data does not allow their direct localization, a limitation that is now stressed in the figure legend (see also A5.R1).\nMinor\nQ9.R1: 1. Extended Data Fig. 6 – The scale bar (20 nm) in panel a appears inconsistent in length compared with b and c.\nA9.R1: We thank the referee to spot this mistake, which has now been corrected.\nQ10.R1: 2. Line 306 – The phrase “Altogether, we new complementary findings provide a more comprehensive understanding of virus-host interactions” should be revised. The only virus–host interaction (in my opinion) described is gp6 binding to the host membrane.\nA10.R1: We agree with the referee. The sentence was revised accordingly to “…comprehensive understanding of virus multiplication within the bacterial cell context…” (lines 381-382 of revised Discussion)\nQ11.R1: 3. References – Line 3042 mentions “phage have been studied over the past 6 decades,” yet only 31 references are cited. Additional relevant literature should be included.\nA11.R1: We focused our citations on major scientific and technological advances that contributed to the study of phage assembly and structure (lines 374-377 of revised version). Among the vast literature on the topic, we selected three additional references even though we are conscious that they cover only very incompletely the amazing discoveries made in this field of phage biology in the past 60 years.\nQ12.R1: 4. Tomogram Data –Providing Z-stack series with segmentation would strengthen the manuscript.\nA12.R1: We prepared segmentation of tomograms manually for the figure panels, which is a powerful and meaningful representation but remains somehow subjective. Therefore we think that representing the tomogram data as 3D rendering of the segmentation should be complemented with Z-stack series of the raw data from reconstructed tomograms (and filtered to a certain extent for display purposes), which we provide as supplementary figure panels or videos.\nQ13.R1: 5. Number of Bacteria Observed – Authors note that each tomogram contains several bacteria. It would be better to summarize the number of bacteria observed in the extended data figure 2 (this should be table, not figure)\nA13.R1: We agree with the reviewer and extended data figure 2 is now a table and referenced as one throughout the text. We are sorry if it was not clear from our description of the workflow that each tomogram is taken at high magnification on a small region of a bacteria, however, it is actually the sections, the so-called FIB-milled lamellae, which contain several bacteria as written in the text: “Each lamella (~12 µm in width) contained cross-sections of several bacteria” (line 112). We hope this helps to clarify this aspect.\nQ14.R1: 6. Warehouse Observation– The “tightly packed warehouse” shown in gp12- strain (Fig. 5, Extended Data Fig. 3g). Was this observed only once, or repeatedly? The frequency of observation should be reported.\nA14.R1: Warehouses are found in 15 out of 25 tomograms taken on SPP1gp12--infected bacteria. The fact that they are tightly packed correlate with fluorescence microscopy observations showing phage particles foci much brighter in SPP1gp12- infected cells than in SPP1 wild type infections (Ref. 10: Labarde et al 2021). We have further clarified this point in the revised manuscript (line 267).\nReviewer #2 (Remarks to the Author):\nThe authors used cryo-electron tomography of Bacillus subtilis infected with bacteriophage SPP1 to investigate the assembly pathway of the virus in cellula. They show that the portal protein gp6 associates with the inner bacterial membrane, where it initiates nucleation of the procapsid. The assembled procapsid then relocates to the viral DNA (vDNA) compartment, where genome packaging takes place. Capsid auxiliary protein Gp12 prevents the aggregation of DNA-filled capsid. DNA-filled capsids subsequently attach to tails that are assembled separately in the cytoplasm. The observations of membrane-associated procapsid formation and genome packaging within a distinct vDNA compartment are novel and provide important insights into phage assembly in cellula. While the authors have done an excellent job visualizing the phage assembly pathway using cryo-ET, several aspects that could not be directly addressed by this technique remain unclear and would benefit from further discussion.\nWe thank the reviewer for the revision and interest, including the clear summary and suggestions to improve our manuscript.\nQ1.R2: How does the vDNA compartment differ conceptually and functionally from the ‘replication centers’ described in other systems, given that it lacks any defined physical boundary?\nA1.R2: In eukaryotes, the terminology “replication centers” has been used for intracellular compartments established inside the host infected cell that contain the viral DNA genome and the factors necessary for its replication (Schmid et al, 2014; doi: 10.1128/JVI.02046-13). In case of bacteriophage SPP1, viral DNA replication takes place inside independent foci within the vDNA compartment (Ref. 10: Labarde et al 2021; lines 61-71 of the revised manuscript). We hypothesize that these foci are conceptually and functionally the genome “replication centers”. Here, both viral DNA replication and the stepwise assembly of DNA packaging events also occur in this compartment. Therefore, we decided to name the compartment according to the fact that it contains the viral DNA genome in high copy number rather than to assign it to a specific function.\nQ2.R2: A discussion of possible mechanisms leading to boundary-less compartmentalization of vDNA may help. Can it be DNA condensation or liquid-liquid phase separation-like processes, as proposed for other phage systems?\nA2.R2: This is an important and timely question that was also raised by referee 1. As described above in A3.R1., our current hypothesis is that the large vDNA compartment is generated by folding of DNA to a phase condensed state leading to separation from the cytosol but that local regions within the compartment might have different biophysical properties (lines 289-307 of revised Discussion). Of note, phage lambda has most likely a similar strategy to compartmentalize its vDNA genome and possibly coordinate DNA encapsidation events (Ref. 8: Trinh et al, 2020). In contrast, DNA replication of phage Phi29 relies on interactions with the host’s MreB cytoskeleton to position phage DNA and the replication machinery (Munoz-Espin et al, 2009; https://doi.org/10.1073/pnas.0906465106). Its DNA replication process is thus constrained in the cell space to the MreB perimembrane localization. Other than that, we did not find in the literature studies for other phage systems and we are not aware of liquid-liquid phase separation-like processes during infection. We would be most thankful to the referee if he/she advices on relevant references that we missed.\nQ3.R2: Additional discussion would be helpful to clarify whether the vDNA compartment functions to concentrate viral or host enzymes involved in DNA replication and packaging, to locally enrich nucleotides or ATP required for DNA translocation, or to protect intermediates from host defense mechanisms. It is possible that resolving this is far beyond the scope of the present paper but this should be stated.\nA3.R2: This is a good suggestion. We showed previously that the SPP1 helicase gp40 together with host replication proteins PolC, DnaE, DnaX, DnaN, DnaG, SsbA and DnaB are massively recruited to discrete foci within the vDNA compartment (Ref. 10: Labarde et al, 2021). We added this information to the revised manuscript (Introduction; lines 61-71). Imaging of DNA packaging motor proteins is more problematic due to poor signal-to-noise ratio in cellula but we would like to tackle these aspects and address their role in future studies.\nQ4.R2: Procapsid formation at the inner membrane: It would strengthen the discussion to include potential functional advantages of initiating procapsid assembly at the inner membrane. For example, the membrane could provide a scaffold that spatially organizes portal and capsid protein assembly, ensuring the incorporation of a single portal vertex per procapsid and preventing the formation of incomplete or aberrant structures.\nA4.R2: We thank the reviewer for this insightful comment. Based on this suggestion, we develop further, in the revised Discussion, the mechanistic implications of assembly at the membrane associated-portal. First, that it provides a platform to initiate and direct polymerization of the scaffolding and major capsid proteins in a cooperative manner (lines 330-335). Second, what we consider most important: to ensure incorporation of one, and only one, portal in the procapsid that is a strict requirement to assemble a functional structure, meaning infectious viral particles ultimately (lines 329-337 of the revised manuscript).\nQ5.R2: While gp6 appears to play a key role in membrane-associated initiation of procapsid assembly, the molecular nature of its interaction with the membrane remains speculative. Have authors analyzed the gp6 12-mer portal structures for the protein surface that can potentially interact with the membrane?\nA5.R2: This is an interesting point raised by the reviewer. Indeed, quoting the referee: the nature of the interaction of gp6 with the membrane remains unknown. We have analyzed high-resolution structures available for the SPP1 portal to provide some hints on the molecular basis. The size of the portal itself make it difficult to apply molecular dynamics simulations to predict potential gp6-membrane interactions more reliably using conventional software like PMIPred (https://pmipred.fkt.physik.tu-dortmund.de/).\nOn the other end, the electrostatics potential displayed using Pymol for the structures of gp6 purified (left, 13-mer) or determined directly in the DNA-filled capsid (right, 12-mer), highlight a region on the gp6 stem that alternates polar and hydrophobic residues (figure below and new figure extended 6). This observation could explain why under certain conditions gp6 can insert into membranes (Zhou et al 2016; DOI: 10.1016/j.biomaterials.2016.08.002 ). Interestingly, a similar “oily” interface was reported for the stem of phage phi29 portal (Simpson et al 2000, doi: 10.1038/35047129). Although this could suggest that the gp6 stem enters the membrane, we feel that experimental data are necessary to probe and validate such hypothesis.\nQ6.R2: Furthermore, the mechanism by which the portal protein detaches from the membrane after procapsid completion is not discussed. Could changes in membrane tension, capsid curvature, or scaffold release serve as potential triggers?\nA6.R2: Our hypothesis is quite simple and relies on closure of the procapsid lattice around the portal that could promote release of procapsid I from the membrane (lines 330-331 of revised Discussion). This would explain that only fully assembled procapsids I are released from the membrane, which appears to happen quite quickly based on the fact that such procapsids are never observed anchored at the membrane in our extensive datasets. Of course, local membrane curvature or tension could also be a factor, which can not be assessed in our tomograms. Capsid curvature however appears less likely to play a role given that positioning of hexamers and pentamers leading to formation of a uniformly sized T=7 capsid, normally requires maintaining a specific curvature throughout capsid lattice growth. Scaffolding proteins release is also unlikely as it occurs during the transition from procapsid I to procapsid II, triggered during DNA packaging. Although, this model is generally accepted in the field, it is true that we cannot fully exclude the possibility that some scaffolding proteins are locally associated to the portal and would be specifically released when the procapsid lattice closes around the portal.\nMinor Comments\nQ7.R2: • In Figure 1 (panels b1–b3), the vDNA compartment is highlighted in red. The same highlighting could be added to the back-plotted tomogram shown in Figure 1b for clarity. In fact, highlighting the vDNA compartment in all subsequent back-plotted tomogram figures would improve consistency and readability throughout the paper.\nA7.R2: We actually tried what the reviewer is proposing and were not convinced. We prepared segmentation of tomograms manually for the figure panels, which makes it very arbitrary to determine the contour of the vDNA compartment. This is due to the fact that it is not delimited by a membrane nor a shell here and instead, we think that ribosome exclusion helps delineate the boundaries of the compartment in an indirect but also much more accurate manner using template matching for automatic detection and plot back in the coordinate system of the original tomogram.\nQ8.R2: • The authors report using C12 symmetry for subvolume averaging of the procapsid I (lines 134–140). It should be clarified why only C12 symmetry was applied, and whether other symmetry options (e.g., C5, C6, or asymmetric reconstructions) were tested.\nA8.R2: Thank you for noticing this, this is a typographical error. Icosahedral, not C12 symmetry, was introduced in an approach further outlined in image methods. This should also clarify the reasoning behind introducing this symmetry. Briefly, unsymmetrized alignment was first used to assess the structure with as little bias as possible. 12-fold icosahedral symmetry operators were introduced primarily as a means of increasing the amount particles for alignment, It is also plausible that a 1:1 stoichiometry of scaffold:capsomeres exists and it is the combination of low particle numbers and minor lattice flexibility prevented this structural relationship from emerging in the unsymmetrised case. Applying any symmetry operator comes with the real possibility of generating symmetrical features out of noise, however, this is where our use of PEET alignment is particularly useful: symmetry was not forced, but rather particles are added at symmetry-related orientations and are allowed to move independently. That there was no apparent correspondence between scaffold and capsid subunits then serves as a stronger evidence of their mismatch.\n“In both cases, no structural detail of scaffolding proteins was resolved, regardless of whether 12-fold icosahedral symmetry was introduced or not” (lines 165-169).\nReviewer #3 (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.\nWe thank the reviewer for her/his input and careful co-revision of the manuscript.\nReviewer #4 (Remarks to the Author):\nIn the manuscript “Subcellular reorganization upon phage infection reveals stepwise assembly of viral particles from membrane-associated precursors,” Corroyer-Dulmont et al. describe the in situ assembly of the SPP1 phage particle using cryo-electron tomography (cryoET). The excellent cryoET work is supported by well-executed fluorescence microscopy experiments and functional assays. The authors demonstrate that the SPP1 assembly pathway is consistent with those of other long-tailed phages, which were previously inferred from in vitro studies and thus carried a certain degree of ambiguity. Notably, in this manuscript, many steps of the assembly pathway have been visualized under native conditions for the first time.\nQ1.R4: The authors find that different components of the virion are assembled at spatially separated areas of the cell, which they refer to as “compartments.” However, these “compartments” are not separated from the rest of the cytoplasm by either proteinaceous or lipid “walls,” unlike those of jumbophages. In principle, there would be no need to bring up jumbophages if the SPP1 “compartments” were referred to by terms that do not imply a physical barrier - such as “area,” “region,” or “subvolume.”\nA1.R4: This is an important point raised by the reviewer. We define compartment as a region in the (bacterial) cell that is enriched in some viral or cellular proteins and/or nucleic acids, here, while excluding other components. This broad definition is now relatively established in cell biology to include membrane-less regions that compartmentalize macromolecules or factors and biochemical reactions in the cell. Such compartments not delimited by a physical barrier result most frequently of phase separation that can have liquid, gel or quasi-solid biophysical properties (Azaldegui et al 2021; doi: 10.1016/j.bpj.2020.09.023; King JT, Shakya A 2021; doi: 10.1016/j.bpj.2021.01.033). We discuss in the revised version the potential biophysical mechanisms leading to formation of the vDNA focus and its inner DNA replication centers in the case of SPP1-infected Bacillus subtilis cells in the discussion in particular (lines 289-307).\nThis is my only, and very minor, comment on an otherwise excellent manuscript. I am happy to congratulate the authors on this outstanding study!\nWe are delighted by the referee’s enthusiasm for our work"}, {"round": 2, "reviewer_comments": {"reviewer_1": "Corroyer-Dulmont et al. report cryo-ET results of seven different SPP1-infected Bacillus samples (gp1=, gp2-, gp12- , gp6- 15min, gp6- 25min, WT 15min and WT 25min), based on 195 cryo-electron tomograms. Their perseverance in collecting such an extensive dataset is admirable—particularly given that all samples were prepared as FIB-milled lamellae to achieve sufficient contrast for downstream analyses.\nPrevious studies of siphovirus capsid maturation have relied primarily on in vitro structural analyses or in vivo low-resolution fluorescence microscopy. In contrast, electron cryo-tomography (ECT) enables in situ, high-resolution visualization. The authors’ ECT observations are consistent with earlier in vitro findings (the transition from procapsid to DNA-filled capsid) and in vivo microscopy studies (viral DNA compartment formation). Importantly, ECT uniquely provides direct in situ visualization and enables quantitative analysis.\nKey findings from this study include:\n1. The membrane-less viral DNA compartment does not require a proteinaceous shell.\n2. Procapsids are located within the vDNA compartment, whereas mature capsids are positioned at the periphery.\nThe most novel discovery, in my opinion, is the suggestion that gp6 initially localizes to the cell membrane, where it initiates procapsid assembly. Overall, this study represents a landmark demonstration of how ECT, when combined with optimized sample preparation, can be used to reveal subcellular reorganization during phage assembly within the host.\nThis revised manuscript adequately addresses all of my previous concerns. If I may offer one remaining comment:\nSupplementary Table 1 suggests that procapsid I can still form in the absence of gp6, whereas Figure 6 implies that gp6 is essential for procapsid I formation. This apparent discrepancy may be confusing to readers. It would be helpful if the authors could clarify this point when discussing their major findings (e.g., around lines 299 or 306).", "reviewer_2": "The authors have done an excellent job of responding to all of the reviewers and revising the paper accordingly.", "reviewer_3": "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_4": "In the opinion of this scientist, the authors of the MS have adequately addressed all concerns raised by the review panel."}, "author_response": "Point-by-point response to the reviewers’ queries:\nReviewer #1 (Remarks to the Author):\nCorroyer-Dulmont et al. report cryo-ET results of seven different SPP1-infected Bacillus samples (gp1=, gp2-, gp12- , gp6- 15min, gp6- 25min, WT 15min and WT 25min), based on 195 cryo-electron tomograms. Their perseverance in collecting such an extensive dataset is admirable—particularly given that all samples were prepared as FIB-milled lamellae to achieve sufficient contrast for downstream analyses.\nPrevious studies of siphovirus capsid maturation have relied primarily on in vitro structural analyses or in vivo low-resolution fluorescence microscopy. In contrast, electron cryo-tomography (ECT) enables in situ, high-resolution visualization. The authors’ ECT observations are consistent with earlier in vitro findings (the transition from procapsid to DNA-filled capsid) and in vivo microscopy studies (viral DNA compartment formation). Importantly, ECT uniquely provides direct in situ visualization and enables quantitative analysis.\nKey findings from this study include:\n1. The membrane-less viral DNA compartment does not require a proteinaceous shell.\n2. Procapsids are located within the vDNA compartment, whereas mature capsids are positioned at the periphery.\nThe most novel discovery, in my opinion, is the suggestion that gp6 initially localizes to the cell membrane, where it initiates procapsid assembly. Overall, this study represents a landmark demonstration of how ECT, when combined with optimized sample preparation, can be used to reveal subcellular reorganization during phage assembly within the host.\nWe like to thank the reviewer for the detailed revision of our work and once more, for the nice summary highlighting its novelty. We address below her/his final query.\nQ1.R1 (query 1, referee 1): This revised manuscript adequately addresses all of my previous concerns. If I may offer one remaining comment: Supplementary Table 1 suggests that procapsid I can still form in the absence of gp6, whereas Figure 6 implies that gp6 is essential for procapsid I formation. This apparent discrepancy may be confusing to readers. It would be helpful if the authors could clarify this point when discussing their major findings (e.g., around lines 299 or 306).\nA1.R1 (answer 1, referee 1): This is indeed an important issue that justifies additional clarification in the manuscript text. Accordingly, we have now added some information in Results (lines 188-191) and Discussion (lines 306-308) of the revised manuscript quoting that “SPP1 procapsid-like structures with two different sizes and aberrant particles can assemble in absence of the portal (Fig. 3c, e-i). However, no intermediates of their off-assembly pathway were found associated to the bacterial membrane (Fig. 3c, e-j; Supplementary Table 1). “\nReviewer #2 (Remarks to the Author):\nThe authors have done an excellent job of responding to all of the reviewers and revising the paper accordingly.\nWe thank the reviewer for her/his interest in our work and the useful suggestions and comments that improved our manuscript very much throughout the revision process.\nReviewer #3 (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.\nWe thank the reviewer for her/his input in co-reviewing the manuscript.\nReviewer #4 (Remarks to the Author):\nIn the opinion of this scientist, the authors of the MS have adequately addressed all concerns raised by the review panel.\nWe thank the reviewer for his/her great enthusiasm for our work along with the useful remarks during the review process that helped us improve the manuscript."}]

Decision letter