Spatial organization of biological processes allows for variability in molecular outcomes and coordinated development. Here, we investigate how organization underpins phage lambda development and decision-making by characterizing viral components and processes in subcellular space. We use live-cell and in situ fluorescence imaging at the single-molecule level to examine lambda DNA replication, transcription, virion assembly, and resource recruitment in single-cell infections, uniting key processes of the infection cycle into a coherent model of phage development encompassing space and time. We find that different viral DNAs establish separate subcellular compartments within cells, which sustains heterogeneous viral development in single cells. These individual phage compartments are physically separated by the E. coli nucleoid. Our results provide mechanistic details describing how separate viruses develop heterogeneously to resemble single-cell phenotypes.
Organization is a fundamental part of life. For complex organisms, the spatial development of body parts is controlled for proper function1. In the cells comprising these organisms, separate organelles are organized by membranes2. Bacterial cells utilize proteins to localize processes in lieu of intracellular membranes3. Furthermore, the physical inhomogeneity of cytoplasmic DNAs, RNAs, and proteins favor the segregation of components for different processes4,5,6. Viruses are even simpler than cellular life, but have also been reported to organize their development within cytoplasmic inclusions or proteinaceous compartments7,8.
Bacteriophages, bacterial viruses, are among the simplest biological systems and serve as models for advanced cellular processes. High-resolution fluorescence microscopy and mathematical modeling have been used to examine the phage lambda lysis–lysogeny paradigm for cellular decision-making to uncover surprising phenomena9,10,11. By labeling single virus particles, we characterized that separate votes for decisions by co-infecting lambda phages determined cell fates12. We next provided distinct voices for separate phages to expound upon our voting model by incorporating different lytic and lysogenic reporters into different phages, finding that phage DNAs compete and cooperate as subcellular individuals13. It is curious how identical viral DNA molecules can commit to divergent trajectories while inhabiting a single cellular environment. While stochasticity is understood to definitely influence development, where noisy elements of gene expression might partially explain differential subcellular behaviors14, it is also evident that properly targeted, high-resolution studies can elicit molecular mechanisms beyond the intrinsic stochasticity of cellular biochemistry15,16,17. We hypothesize that subcellular organization allows different phages to develop as individuals in a single cell and expect to detect the underlying subcellular heterogeneity by specifically investigating the spatial distribution of viral/host biomolecules. Here, we show that cascading events during viral transcription, DNA replication, and gene expression combine to establish an organized subcellular unit of phage development. This organization permits multiple individual viruses in the same cell to develop via different pathways in separate areas of that cell.
Live-cell fluorescent reporters of phage development
To work toward a unified model of individual lambda development in cellular space, we used live-cell time-lapse microscopy targeting the initial infecting phage DNAs, host cell’s replication resources, replicated phage DNAs, and phage decisions (Fig. 1a). To visualize the initial infecting phage DNA, we modified cells to be dam− and carry a seqA-mKO2 (mKO2 signal defined as yellow) translational fusion18. This system has been validated previously to correctly label single DNA molecules from infecting phages11,13,18. Upon ejection of methylated phage DNA into the host, SeqA-mKO2 binds exclusively to the ejected phage DNA and any DNA copies retaining the methylated parent strands with single-DNA labeling sensitivity, but not subsequent replicated DNA copies (SeqA system, Fig. 1b)11,18. Since the SeqA system cannot target all replicated phage DNAs, we recombineered an array of tetO sequences into the phage genome. With the host cell harboring a TetR-mCherry plasmid (mCherry signal defined as red), all phage genomes are bound and labeled at tetO sites by TetR (Tet system, Fig. 1d) (Supplementary Discussion). The Tet-labeling scheme lacks single-DNA sensitivity under our experimental conditions, so we used both Tet and SeqA systems to target phage DNA. As lambda depends on host factors for viral DNA replication, we translationally fused the Escherichia coli helicase19, DnaB, with mTurquoise2 (mTurquoise2 signal defined as blue) by replacing the native dnaB gene with dnaB-mTurquoise2 on the E. coli chromosome (Fig. 1c; Supplementary Fig. 2f). DnaB is essential for phage/E. coli DNA replication and directly interacts with lambda P (analog of E. coli DnaC)20. The DnaB construct does not appear to impose major detriment on E. coli or phage growth (Supplementary Fig. 2g–j). Finally, we reported lambda lysis–lysogeny decision-making using previously developed systems13. Briefly, we modified phages with a D-mNeongreen (mNeongreen signal defined as green) translational fusion, reporting the lytic pathway because progeny phages are assembled with green gpD, and a cI-mKO2 transcriptional fusion, reporting the lysogenic pathway because the cI operon(s) are expressed during lysogeny (Fig. 1a). Accordingly, we developed a data analysis framework for these reporters to detail the spatial organization of subcellular events during infection (Supplementary Fig. 1). Notably, all presented images of individual cells unambiguously represent single cells, because early expression of the Kil protein by lambda inhibits cell division during infection21.
Organization of resources and replication by individual phages in single cells
We expect that compartmentalization of biomolecules reflects the establishment and perpetuation of individualistic development by lambda. Uninfected cells (LZ1557) displayed diffuse blue, yellow, and red fluorescence, indicating that DnaB, SeqA, and TetR do not compartmentalize without phage infection (Supplementary Fig. 2a; Supplementary Movie 1). Functional DnaB is required for growing cells19; thus, DnaB localization under cell growth conditions represents a basal DnaB state. Importantly, active DnaB-mTurquoise2 does not localize as a focus under phage-free, normal growth conditions. When infecting these cells with our phages (λLZ1576), the fluorescence patterns change significantly from the phage-free state. We primarily focus on analyzing lytic cells because extensive DNA replication is required for successful lytic propagation and note that lysogenic development markedly differs from lytic behaviors (Supplementary Fig. 3b, c; Supplementary Movie 2; Supplementary Discussion). SeqA foci appear within the cells, typically early during infection (Supplementary Fig. 3a), indicating that phage DNA entered the cell (Fig. 1e). As phage DNA replication demands host resources, we tracked DnaB. DnaB foci typically formed after SeqA foci and were commonly colocalized with SeqA foci over time (Fig. 1e, f, i, j; Supplementary Fig. 4a, b). This indicates that phage DNA directly alters the natural behavior of DnaB by collecting essential resources to its own location. DnaB foci represent multiple DnaB complexes, specifically aggregated to single phage DNAs for extensive phage DNA replication. In lytic cells, blue foci remained over time (median: 90 min), suggesting that lambda persistently recruits resources for itself (Supplementary Fig. 3b).
Resource recruitment precedes phage DNA replication. Red fluorescence is initially spread throughout the cell, representing free TetR as background signal, not phage DNA signal (Supplementary Fig. 2c, e, at 40 min). TetR signal later rearranged into small clusters over time, signifying the production of additional phage DNAs nearby the specific locations of the single DNAs and resources (Fig. 1e–h; Supplementary Movie 3). These subcellular local maxima, or clusters, of TetR signal represent phage DNA. DnaB was pre-localized near the eventual red clusters and remained within the clusters as they expanded within the cell, suggesting that replicating phage DNAs predictably arise at the location where previous DNAs gathered resources, and then maintain this sequestration (Fig. 1k, l; Supplementary Figs. 3d and 4c, d). In lytic cells, gpD (green) signal increased throughout the cell over time (Fig. 1e–h). This signal is initially diffused throughout the cell, but later, green foci, corresponding to phage capsids, preferentially formed in the red clusters (Supplementary Fig. 5a). This suggests that the locations of phage DNA determine where progeny phages are assembled, consistent with the characterized mechanisms of phage DNA packaging22. Altogether, the data indicate that a single phage DNA organizes its own subcellular phage factory, persistently maintaining its clones and resources proximal to itself. We designated this phage-derived, subcellular compartment as a “phactory” for future reference.
As the single phage DNA had the capacity to assemble its own compartment, we predicted that multiple, individual phage DNAs could form separate phactories. We identified cells with single phage DNAs in different locations (Fig. 1g, h; Supplementary Movie 4), where each DNA collected its own stockpile of DnaB. Separate red clusters appeared and grew at each DnaB location, and finally, green foci grew into clusters nearby the separated DNA clusters. Intriguingly, the phage-related biomolecules remain separated in space as different viral microenvironments, where the levels and identities of the phage DNA, RNA, and protein in the microenvironments may change and be exchanged while still maintaining spatial separation. These data suggest that each subcellular phage DNA can be its own entity and can organize its own phactory within single cells. In conditions that specifically stimulate spatially segregated phage DNAs to develop, lysogenic induction (Fig. 2a; Supplementary Discussion), multiple phactories separately form and progress in single cells (Fig. 2b–d; Supplementary Figs. 5b and 6). Phactories have unequal DNA and lytic reporter levels over time, suggesting that heterogeneous development is sustained within individual phactories (Fig. 2e; Supplementary Figs. 7 and 8).
Host nucleoids maintain separation of individually developing phages
Our results indicate that phage DNAs in different phactories remain separated as the basis of their individuality, but do not indicate obvious barriers that segregate phage DNAs, obscuring the detailed mechanisms of their heterogeneous development. We observed that the growth of phage DNA clusters decreased as the cell filled with phage DNA, as if their size might be physically limited by something in the cell (Supplementary Fig. 9a, c; Supplementary Discussion). We hypothesized that replicating phage DNAs are physically separated from E. coli DNA, a presumed barrier shaping the phactories. We tested this hypothesis by labeling E. coli attB with a lacO array and LacI-EYFP construct, adapted from previous work11. We infected these cells (LZ1643) with phages (λLZ1629) (Fig. 3a). Phage DNAs and capsids behave similarly to the above infections with λLZ1576 and LZ1557 (Supplementary Figs. 5c, 9b, d and 10). The terminus of lytic cells is cell lysis, so we tracked the location of attB in the final time points to determine how preceding lytic development affects E. coli DNA, finding that attB locations were biased near the poles (Fig. 3h). Conversely, the terminus of non-lytic development is cell division, and attB was localized between the mid-cell and quarter-cell in non-lytic cells, suggesting differential development (Fig. 3h; Supplementary Fig. 11a; Supplementary Movie 5). We subcategorized the lytic cells based on the interaction of phage DNA with attB (push, spread, and squeeze, Fig. 3b-g; Supplementary Fig. 11b-d; Supplementary Movies 6–8). In the largest class (push, 60%, 108 out of 179 lytic cells), we found that attB was pushed towards one side of the cell, away from a single expanding phage DNA cluster (Fig. 3b, c; Supplementary Movie 6). Within this population of cells, as phage DNA cluster sizes increased over time, attB moved closer to cell poles (Fig. 3i, j). Furthermore, after phage DNA clusters expand near attB, they do not move past it, suggesting that phage and bacterial DNA do not mix together (Supplementary Fig. 11e, f; Supplementary Discussion). These data agree with our previous observations regarding the polar movement of attB in lytic cells11, provide a clear phage-active mechanism, where the spatial expansion of phage DNA due to replication explains the results, and corroborate our hypothesis that E. coli DNA helps determine phactory localization and segregation. Because the two phactories do not merge around the bacterial marker (Fig. 3f), it suggests that bacterial DNA is a physical barrier which allows subcellular viruses to maintain different identities.
We next used alternate techniques to examine phage infection, utilizing bacteria/phages without genetically engineered reporters to minimize the likelihood that the genetic modifications influenced biological behaviors. We performed single-molecule fluorescence in situ hybridization (FISH) to characterize the spatial aspect of phage transcription and to examine phage DNA replication in fixed cells, to better characterize how different subcellular viruses develop23,24,25.
To compare FISH with our live-cell techniques, we labeled phage λLZ613 DNA during infection using lambda DNA-specific probes. At early time points, phage DNA existed as small foci/clusters in cells (Fig. 4a). The localization of FISH foci resembles that of SeqA foci in live cells (Fig. 4p, q). At later time points, we found that DNA signal increased with time, and appeared as larger clusters, similar to our Tet reporter system (Fig. 4b). The amount of DNA per cell and within clusters varied (Fig. 4c, d; Supplementary Fig. 12a). To study bacterial DNA, we treated the cells with DAPI in FISH experiments. While DAPI stains DNA non-specifically, the size of E. coli DNA (4.6 Mbp) far outstrips that of lambda DNA (48.5 kbp); thus, DAPI will primarily stain bacterial DNA at early time points before extensive phage DNA replication, allowing phage DNA FISH signal and DAPI to be mutually exclusive (Supplementary Discussion). We observed that the spatial distribution of the DAPI signal in E. coli leaves distinct nucleoid-free zones (Fig. 4h, p; Supplementary Fig. 13b), and phage DNAs preferred these areas in early time points (Fig. 4g, h, j, p; Supplementary Fig. 13a, b), supporting earlier observations of negative bacterial and phage DNA spatial correlation (Fig. 3). Phage DNA and DAPI signals overlap in later time points, after presumed phage DNA replication (Fig. 4j; Supplementary Figs. 12b–e and 13a, b, g). Furthermore, we generated probes against E. coli attB, and found that attB generally localizes with DAPI (Supplementary Fig. 13d–f), but avoids phage DNA locations (Fig. 4g, i, k; Supplementary Fig. 13a, c, h), congruent with our live-cell data (Supplementary Discussion). These results suggest that the organization of heterogeneous, separated phage DNA units exists and is reported faithfully by both live-cell and fixed-cell methods.
Phages maintain localized transcription
Given the organization of segregated phage DNAs, our next step was to investigate the organization of phage mRNAs because gene expression is a key component of phage development and decision-making. We studied phage transcription with RNA FISH24, where we first targeted pR, an early transcriptional unit comprising genes for decision-making and phage DNA replication26. At early time points, pR transcripts existed as small clusters (Fig. 4e). We found that pR localizes to areas with lower DAPI signal (Fig. 4l–o; Supplementary Fig. 14a–d), near poles and/or mid-cells, regions well-characterized to typically be nucleoid-free27,28, suggesting that phage mRNAs, like phage DNAs, reside away from nucleoids. Remarkably, even at later time points, pR retains subcellular localization as opposed to diffusing evenly throughout the cell (Supplementary Fig. 14a). Notably, DAPI signals increase in pR locations at later time points following presumed phage DNA replication. From the convergence of our experimental data, we concluded that the locations of pR transcripts in FISH also represent the locations of phage DNA. These results suggest that intracellular phactories possess their own gene expression profiles in single cells. This surprising degree of mRNA spatial organization for lambda may be promoted by the physical attachment of mRNAs to phage DNAs during transcript elongation29, and whole-cell diffusion may be discouraged by the relatively short lifetimes of mRNAs in E. coli30, combined with similarly localized ribosomes translating the transcripts27,31. Notably, phage mRNA localization remains even after transcription is finished and further transcription is blocked with rifampicin treatment32, demonstrating that phage RNAs are continuously localized with phage DNAs in the phactory (Supplementary Fig. 15).
Separate decisions made in different locations of the same cell
Downstream of phage DNA replication and early gene expression is decision-making, and we predicted that the organization of upstream processes would impact the execution of specific cell-fate expression programs26. The pRʹ transcript encodes the lysis and morphogenesis proteins during lytic development; the pRE/pRM (referred to as pRE) transcripts encode CI to establish and maintain lysogeny. Therefore, we targeted the pRʹ and pRE transcripts with FISH to characterize subcellular decision-making in subcellular space (Fig. 5a, b).
We performed a set of FISH experiments at 15 min after infection because decision-making is expected to occur around this timeframe24. In support of our model, pR levels vary at different locations of the same cell, more so than pR varies between different cells (Supplementary Fig. 14g). These data reiterate that pR expression is localized and suggest that the variation of localized pR mRNA levels may lead to different localized decisions. The lysogenic decision will eventually repress transcription from pR due to the action of CI26. Accordingly, we found that cellular pRE levels negatively correlate with cellular pR levels (Fig. 5c). Furthermore, focusing on the spatial aspect of transcription, we determined that the cellular coordinates of the brightest pRE signals are offset from the brightest pR signals (Supplementary Fig. 14e, h). These data corroborate earlier hypotheses that pRE/pR head-on transcription might commit the lambda genetic circuit towards lysogeny when pRE dominates33. The data suggest that lysogenic decisions may be locally initiated by individual DNAs, where pRE activation halts pR transcription in a single phage genome, and that these localized actions precede cell-wide repression by CI. As for the pRʹ transcripts, our FISH data show that the brightest pRʹ and pR signals coincide within the cell. Altogether, the data suggest that decisions are enacted by separate phage DNAs in specific subcellular areas (Supplementary Fig. 14f, i).
Our model for lambda decision-making involves voting for decisions by co-infecting phages, signifying that individual lambda DNAs are capable of heterogeneous development in single cells12,13. Given our conclusions up until now, we would expect that these voting behaviors occur in separate subcellular areas. Accordingly, we found that in our FISH experiments, pRʹ transcripts can exist in a subset of intracellular pR clusters (Fig. 5d, e). We also found that pRE and pRʹ transcripts can coexist in different locations in single cells, suggesting that one intracellular phactory might vote differently from a neighboring phactory (Fig. 5f, g). The organization underpinning these behaviors persists even after new transcription initiation is blocked (Supplementary Fig. 15f–k, m). These data indicate that multiple intracellular lambda DNAs can execute divergent developmental pathways, where the spatial organization of phage/bacterial biomolecules and processes supports their individuality (Fig. 5h).
We described how multiple aspects of lambda development coalesce via spatial organization, allowing multiple viruses to develop separately in a cell. We characterized how the single phage DNA collects DnaB, essential for DNA replication, to its own location, begetting its clones and their corresponding development to its own coordinates. We showed that phage DNAs organize multiple transcripts locally at early time points, suggesting that different gene expression patterns are enacted by separate sets of viral DNA in separate cellular spaces, altogether resulting in individual, subcellular compartments of phage development, or phactories. The individual phactory is formed both due to the phage’s own actions, including DnaB recruitment and phage mRNA organization from active and finished transcripts, and also due to the physical separation of the viral biomolecules by the bacterial chromosome. It is unclear to what degree phage proteins are separated during the infection cycle, which is another important element contributing to divergent development in single cells (Fig. 5h).
It is intriguing that even simpler viruses, such as lambda, have evolved their own means of organization, which allows relatively complex behaviors to be achieved. When compared to particularly complex phages, such as Pseudomonas phage 201phi2-1, with a genome over 300 kbp encoding over 400 putative genes, lambda is simplistic34. 201phi2-1 houses its own cytoskeletal genes that position and build a protein-enclosed compartment encapsulating its DNA replication and transcription processes separately from its translation, evoking comparison to a nucleus8. Simpler phages, such as phage phi29, may have increased reliance or exploitation of host-specific proteins and processes. Efficient phi29 DNA replication relies on interactions with the host’s MreB cytoskeleton to position phage DNA and replication machinery35. Phi29 DNA replication also depends on the organization of a replication resource, phage DNA polymerase, via the phage-driven actions of its terminal protein36. Phage lambda has a similar strategy to organize replication, as it both utilizes its own proteins to actively reorganize its essential resource, DnaB, and also depends on its host’s architecture, the location of the bacterial nucleoid. The relative simplicity of lambda can also be compared to some high-copy number plasmids lacking partitioning systems, which tend to localize to non-nucleoid locations, perhaps passively37. There are major differences between lambda and plasmid replication. Our data demonstrate that lambda retains its DNA into a single phactory, which pushes the nucleoid as it grows in space. This would obviously be detrimental for plasmids, because a single plasmid cluster in one cellular position would become easily lost during cell division, and if plasmids interfered with bacterial DNA localization, it would disrupt cell viability. Additionally, lambda actively commandeers DnaB, which clearly diverges from plasmid behavior, since this would kill the cell.
The application of high-resolution methods to phage lambda has the potential to reveal distinct and novel mechanisms buried within the long-standing lambda paradigm and could be equally applicable in other phage systems for comparison38. Further investigations into the biophysical interplay of viral and bacterial biomolecules, especially those which leverage continually advancing high-resolution techniques capable of fine spatial discrimination, promise new insights into the varied mechanisms behind viral development.
Strains, plasmids, and primers
Bacterial strains, phages, and plasmids used in this study are listed in Supplementary Table 1.
Primers and homology regions used in this study are listed in Supplementary Table 3.
Single-cell infection imaging assay
Host cells, LZ1557, were grown from a −80 °C frozen permanent stock in 1 ml of M9 + 0.4% maltose (M9M), supplemented with antibiotics, 100 μg/ml ampicillin (Amp100) + 50 μg/ml kanamycin (Kan50) + 10 μg/ml chloramphenicol (Cm10) in a shaker at 37 °C, 265 r.p.m., overnight for ~24 h. This overnight culture was then diluted 1:1000 into 5 ml of M9M + antibiotics as above and grown under the same conditions for ~16–18 h, to OD600 ~0.3–0.4. Then, 1 ml of this culture was pelleted in a tabletop centrifuge, 3000 × g for 4 min at room temperature. During this time, 20 μl of purified reporter phage (λLZ1576) at ~3–4 × 1010 pfu/ml is pipetted into a microcentrifuge at room temperature. After the cells were centrifuged, the supernatant was pipetted away, and the pellet was resuspended in 200 μl of room temperature M9M. Twenty microliters of this suspension was then mixed with the 20 μl of phage solution via gentle pipetting, resulting in an average phage input (API, the phage:bacterium ratio) of ~4, and then 80 μl of room temperature M9M was added to the mixture and mixed via gentle pipetting. This new mixture was then moved to a pre-warmed 35 °C water bath for 4 min to allow for phage adsorption and DNA ejection. During this time, a small (1–1.5 cm2) section from a room temperature M9M agarose pad (all pads in all experiments were 1.5% agarose), freshly made, was set onto a small No. 1 coverslip (18 × 18 mm). Following the incubation of the mixture, 1 μl of the mixture was deposited onto the M9M pad. After the mixture visibly dried, ~1 min, a larger No. 1 coverslip (24 × 50 mm) was overlaid onto the M9M pad, sandwiching it, and the sample was moved to the microscope for time-lapse imaging at 30 °C.
For the experiment using bacterial strain LZ1643 and phage λLZ1629, the M9M is supplemented with Amp100 + Kan50 for the bacterial growth. A colony from a plate was grown overnight for ~16–18 h. This overnight culture was then diluted 1:100 into fresh media and grown until OD600 ~0.3–0.4, about 3–4 h. The cells were then pelleted and resuspended as described above. In these experiments, after the phage and cell suspensions were mixed, the mixture was set on ice for 30 min to pre-adsorb the phages before moving the mixture to the 35 °C water bath for 5 min. The following steps up to imaging are identical to the above description.
Induction imaging assay
A lysogen colony, LZ1596, from a plate was grown overnight in 1 ml of LB + 10 mM MgSO4 (LBM) supplemented with Amp100 + Cm10, ~16–18 h in a 30 °C shaker, 225 r.p.m. The overnight culture was then diluted 1:100 into 5 ml of fresh LBM with the above antibiotics and grown under the same conditions until OD600 ~0.3–0.4. For the 0 min time point, 1 μl of the culture was taken from the flask and deposited onto a phosphate-buffered saline (PBS) agarose pad, similarly as described above with the M9M pad, and then imaged. The lysogen culture was then induced by moving the flask to a 42 °C (our phages bear the cI857 temperature-sensitive allele39), 225 r.p.m. shaking water bath for 15 min. The 5-min time point occurs after 5 min at 42 °C, and the sample was processed and imaged at that time, as the 0 min sample was. The 15-min time point occurs at the end of the 42 °C incubation. While the 15-min time point sample was processed, the induction culture was moved to a 37 °C, 225 r.p.m. shaking water bath, and imaging of the 15-min time point occurred during this 37 °C incubation. The remaining time points were taken as the culture was shaking at 37 °C.
Bacterial growth and lysogen induction assays
To generate bacterial growth curves, bacterial strains were plated on a standard LB agar plate supplemented with appropriate antibiotics. A single colony was used to inoculate a 1 ml overnight culture in LB or M9 + 0.4% maltose (M9M) in a 37 °C, 265 r.p.m. shaker. The overnight culture was then diluted 1:100 into 30 ml of LB or M9 + 0.4% maltose (M9M) supplemented with appropriate antibiotics in a flask and grown in a 37 °C, 265 r.p.m. shaking water bath. OD600 was measured using a spectrophotometer.
For lysogen induction assays, a lysogen colony from an LB plate was grown overnight in 1 ml of LB + 10 mM MgSO4 (LBM) supplemented with appropriate antibiotics in a 30 °C, 180 r.p.m. shaking water bath. An overnight culture was then diluted 1:100 into 25 ml of fresh LBM with appropriate antibiotics in a flask and grown under the same conditions until an OD600 of ~0.3, which serves as the 0-min time point. The lysogen culture was then transferred to a 42 °C, 180 r.p.m. shaking water bath for 15 min for thermal induction. The OD600 of the culture was measured at the end of the 42 °C incubation, serving as the 15-min time point. The induction culture was moved to a 37 °C, 180 r.p.m. shaking water bath. The OD600 of the culture was measured every 5 min until lysis with an OD600 of ~0.05 using a spectrophotometer. Subsequently, 5 ml of the induction culture was taken from the flask and mixed with chloroform to a final concentration of 2% (vol/vol). The culture was agitated using a nutator for 15 min at room temperature and then centrifuged at 3000 × g for 10 min to obtain the phage lysate. The concentration of the phage lysate was determined via a standard phage titration assay.
For DNA FISH, probes for lambda DNA were produced by PCR amplifying ~3 kbp of the lambda genome (f-lambda-dnafish and r-lambda-dnafish primer pair), using a phage lysate as the template, and treating the purified PCR product with a PromoFluor500-dUTP nick translation kit (PromoCell) to generate DNA-PromoFluor500 fragments ranging from 100 to 500 bp. To generate probes for the E. coli attB region, an ~3 kbp region of the E. coli genome, including the attB region, was amplified with PCR (f-attb-dnafish and r-attb-dnafish primer pair) and treated with a PromoFluor640-dUTP nick translation kit (PromoCell). Equal amounts of these probes were mixed together to form a probe mixture.
To perform DNA FISH on infection samples, cells (MG1655) were first grown from a colony overnight, ~16–18 h, in LBMM (LB + 0.2% maltose + 10 mM MgSO4). The overnight was then diluted 1:1000 into 50 ml of fresh LBMM and grown at 37 °C, 265 r.p.m., until OD600 ~0.3–0.4, about 3–3.5 h. The culture was then pelleted via tabletop centrifuge (2000 × g, 4 °C, 15 min), the supernatant was discarded, and the pellet was resuspended in LBM at 1/10th the original volume to concentrate the cells. Four milliliters of the cells were placed on ice, and ~40 μl of λLZ613 phage, at ~1 × 1011 pfu/ml, was added to the cells and gently mixed. 2 × 500 μl aliquots of cells were also separated as a control without phages. After leaving the infection mixture for 30 min on ice, the tube was moved to a 35 °C water bath for 5 min for phage DNA ejection. At this point, 500 μl of the infection mixture was aliquoted into a culture tube with 4.5 ml of LB + 0.2% glucose + 10 mM MgSO4 (LBGM) for each time point, all tubes were then moved to a 30 °C shaker at 265 r.p.m. At each given time, a tube was taken and fixed by pouring the mixture into a 15 ml centrifuge tube with 550 μl of 37% formaldehyde. This tube was left to shake on a nutator for 30 min, and then centrifuged at 4000 × g for 3 min to pellet the cells. The control sample was fixed after the 35 °C incubation.
Details of fixation, permeabilization, and hybridization are detailed in other studies25. Briefly, the fixed cells were washed with 1 ml of ice-cold 1× PBS three times and resuspended in 1 ml of GTE solution (50 mM glucose, 20 mM Tris-HCl [pH 7.5], 10 mM EDTA). For the control sample, three separate 500 μl aliquots of the cell suspension were then mixed with 10 μl of 0.01 μg/μl lysozyme solution and incubated at room temperature for 2, 4, and 6 min followed by three washes with GTE, pelleting the cells via centrifugation at 10,000 × g for 30 s. The cells were then resuspended in ~150 μl of GTE. For each control sample, 1 μl of the cells was deposited onto a PBS agarose pad and imaged. The lysozyme treatment time yielding ~90–95% intact cells (~1–5% lysed cells) represents the optimal treatment time for the samples. The actual time point samples, from the initial GTE wash, were then processed as the control was, using the optimal lysozyme time. For each time point, 10 μl of cells were deposited onto poly-l-lysine-coated large coverslips (24 × 50 mm), then covered with a smaller, normal, coverslip (22 × 22 mm). The coverslips were then immersed in 1× PBS and the smaller coverslip was removed, leaving only the sample coverslip. The cells were then dehydrated by immersing the coverslip in increasing concentrations of ethanol (70, 90, then 100%). Samples were then ready for hybridization.
For each sample, approximately 160 μg of the probe mixture was combined with 10 μl of hybridization solution (50% formamide, 10% dextran sulfate, 50 mM NaPO4/pH 7, 2× SSC). The dsDNA probes were denatured at 75 °C in a thermocycler, then placed on ice. Ten microliters of the denatured probe mixture were then deposited onto the center of the sample on the coverslip and overlaid with a small coverslip (22 × 22 mm). The small coverslip was then sealed with nail polish, forming a sample chamber. The chambers were incubated at 80 °C for 5 min to denature the cellular DNA, and then placed on Kimwipes over ice for 5 min. The chambers were then incubated in a 37 °C incubator overnight to complete hybridization.
The next day, the chambers were immersed in 2× SSC until the smaller coverslip dislodged. The remaining coverslips were soaked in wash solution (2× SSC, 50% formamide) for 20 min at 37 °C twice. The coverslips were then washed with a series of increasing SSC concentration washes (1, 2, then 4×), each for 5 min at room temperature. A DAPI solution was then made by mixing 1 μl of 10 mg/ml DAPI to 1 ml of 4× SSC. For each sample, 500 μl of the DAPI solution was added over the sample, covering it, and incubated for 5 min at room temperature. After drying the coverslip, 10 μl of 2× SSC was added over the sample and overlaid with a small coverslip (22 × 22 mm). The samples were then imaged.
Different probes were synthesized to target different phage transcripts (Biosearch Technologies). Probes targeting pR and pRE were designed following previous studies23,24, labeled with Cy5 and TAMRA, respectively. Probes targeting pR′ followed the same design principles as pR and pRE, and were labeled with AlexaFluor488 (pR′ probes listed in Supplementary Table 2).
To perform RNA FISH, we follow the same infection protocols as described above for DNA FISH. At given time points, the cells were fixed in formaldehyde and pelleted. In one set of experiments, samples were taken between 6 and 40 min, after infection by phages at ~2 × 1010 pfu/ml. In another set of experiments, samples were taken at 15 min, after infection by phages at ~1, 2, 3, and 4 × 1011 pfu/ml. The processing of the samples is detailed in our previous study24. Briefly, after fixation, the cells were washed three times with 1× PBS. Subsequently, the cells were permeabilized by resuspension in 70% ethanol for 1 h at room temperature and centrifuged to collect the cells. The pellet was then resuspended in wash solution (40% formamide, 2× SSC) and incubated for 5 min at room temperature, and pelleted again, ready for hybridization.
The cells were then resuspended in 25 μl hybridization solution (40% formamide, 2× SSC, 1 mg/ml E. coli tRNA, 2 mM ribonucleoside-vanadyl complex, and 0.2 mg/ml BSA) with each set of probes reaching a final concentration of 1 μM. The samples were then incubated in a 30 °C water bath overnight. The next day, the cells were washed three times using wash solution by incubating the cell pellet for 30 min in a 30 °C water bath. After the final wash, the cells were resuspended in wash solution + 10 μg/ml DAPI and incubated for 10 min at room temperature. This suspension was then pelleted and resuspended in 2× SSC. The sample was then ready for imaging.
For the infections with rifampicin, rifampicin was added at a final concentration of 50 μg/ml to a 50 ml infection mixture in a flask at 15 min after the 35 °C step. Instead of pre-aliquoting separate infection tubes, 5 ml of the infection mixture was withdrawn at each given time point after addition of rifampicin for fixation and further processing as described.
E. coli nucleoid imaging after rifampicin treatment
Bacterial cells (MG1655) were inoculated from a colony into 1 ml of LB and grown at 37 °C, 265 r.p.m. for overnight. The overnight culture was then diluted 1:1000 into 5 ml of LB and grown under the same conditions until an OD600 of ~0.3. To study the effect of rifampicin on E. coli nucleoid morphology, rifampicin was added to the culture aliquots at final concentrations of 50, 100, and 300 μg/ml, and treated the cells for 15 min and 30 min at 37 °C, 265 r.p.m. One milliliter of the cell cultures was then fixed in 3.7% formaldehyde for 30 min at room temperature, followed by a washing step with 1 ml of PBS. The cell pellet was resuspended in 100 μl of PBS to reach an optimal cell density for microscopy imaging. For nucleoid imaging, 10 μl of cells were mixed with 10 μl of 20 μg/ml DAPI (to reach the final concentration of 10 μg/ml DAPI) for 10 min at room temperature. One microliter of the DAPI-stained cells were spotted onto a PBS agarose pad and imaged.
SeqA cell lysogenization culture
An overnight culture of cells bearing the SeqA reporter (LZ1557) were diluted 1:1000 and grown in 10 ml of LBMM + Kan50 + Amp100 + Cm10 to OD600 ~0.4. The culture was centrifuged, and the pellet was resuspended in 1 ml of LBM to concentrate the cells by 10-fold. Two hundred and fifty microliters of cells were mixed with phage (λLZ1576) to reach an API of ~4. The infection mixture was placed on ice for 30 min, and then moved to a 35 °C water bath for 5 min. The mixture was then diluted into 5 ml of fresh LBM and incubated in a 30 °C shaking water bath at 265 r.p.m. Samples were withdrawn at given time points for imaging.
Each set of experiments (live-cell with λLZ1576/LZ1557, live-cell with λLZ1629/LZ1643, lysogen induction, DNA FISH, and RNA FISH) had its own set of imaging parameters according to the specific strains and fluorophores employed. All imaging was performed on a Nikon Eclipse Ti inverted epifluorescence microscope using a 100× objective (Plan Fluo, NA 1.40, oil immersion) with a 2.5× TV relay lens, using a mercury lamp as the light source (X-Cite 200DC, Excelitas Technologies), within a cage incubator (InVivo Scientific) at 30 °C, and acquired using a cooled EMCCD (electron multiplying charge-coupled device) camera (iXon3 897; Andor, Belfast, United Kingdom). The software images each stage through each filter sequentially for each time point before moving to the next stage. For the induction and fixed-cell experiments, stages with abundant cells were chosen for imaging. The stages were imaged under phase-contrast and specific filter cubes. The fluorescent filters used in the study were as follows (X, Y [excitation bandwidth] excitation filter/dichroic beamsplitter wavelength/X, Y [emission bandwidth] emission filter/company, product #): DAPI (350 nm, 50ex/400 nm/460 nm, 50em/Nikon, 96310), blue (436 nm, 20ex/455 nm/480 nm, 40em/Nikon, 96361), custom green (490 nm, 20ex/505 nm/525 nm, 30em/Chroma, custom 49308), yellow (500 nm, 20ex/515 nm/535 nm, 30em/Nikon, 96363), orange (539 nm, 21ex/556 nm/576 nm, 31em/Chroma, 49309), Cy3 (545 nm, 30ex/570 nm/610 nm, 75em/Nikon, 96323), red (560 nm, 40 nm/585 nm/630 nm, 75 nm/Nikon, 96365), far red (592 nm, 21ex/610 nm/630 nm, 30em/Chroma, 49310), and Cy5 (615 nm, 70ex/660 nm/700 nm, 75em/Nikon, 96366).
For imaging each experiment, samples were exposed to the named filter cube in this order (with this exposure time, against this target).
Infection movies using λLZ1576/LZ1557: phase-contrast (100 ms), blue (1 s, DnaB), orange (100 ms, single phage DNA), far red (200 ms, replicated phage DNA), and green (40 ms, capsid). This cycle of imaging occurred automatically once every 10 min for at least 3 h in each movie.
Infection movies using λLZ1629/LZ1643: phase-contrast (100 ms), blue (100 ms, capsid), yellow (100 ms, attB), and red (200 ms, replicated phage DNA). This cycle of imaging occurred automatically once every 5 min for at least 2 h in each movie.
Lysogen induction: phase-contrast (100 ms), blue (1 s, DnaB), far red (200 ms, replicated phage DNA), and green (40 ms, capsid). This was not a movie, so the imaging cycle was repeated for each stage/time point in the dataset.
DNA FISH: phase-contrast (100 ms), yellow (200 ms, phage DNA), Cy5 (200 ms, attB), and DAPI (30 ms, DAPI). This imaging cycle was repeated for each stage in the dataset.
RNA FISH: phase-contrast (100 ms), Cy5 (200 ms, pR), Cy3 (200 ms, pRE), yellow (200 ms, pRʹ), and DAPI (30 ms, DAPI). This imaging cycle was repeated for each stage in the dataset.
E. coli nucleoid imaging after rifampicin treatment: phase-contrast (100 ms) and DAPI (100 ms).
Microscopy images were analyzed using the cell recognition program Schnitzcells (gift of Michael Elowitz, California Institute of Technology), the spot recognition program MicrobeTracker (gift of Christine Jacob-Wagner, Yale University), and homemade scripts in Matlab (Supplementary Discussion).
To construct the phages with tetO arrays, phages bearing fluorescent reporters (λLZ1269, λLZ1369, λLZ1527, details for constructing these types phages were reported in previous studies) served as the parents13. Then a tetO-recombination plasmid was constructed to replace the bor::KanR region of the fluorescent phages with a bor::CmR 24×tetO array construct, using homology regions (upstreambor and downstreambor) The tetO array was derived from another study40, and inserted adjacent to a CmR cassette. Phages were titered onto host cells bearing the tetO-recombination plasmid (pBR322 24×tetO bor::CmR) and a pLate*D plasmid, then lysogenized the resulting plate lysate into MG1655, selecting for Cm resistance and Kan sensitivity in single-integration lysogens. The genomic construct was then verified by PCR.
To construct the triple reporter strain, bearing SeqA (single phage DNA), TetR (replicated phage DNA), and DnaB reporters, strain LZ1383 (MG1655 seqA-mKO2 CmR-FRT) served as the parent. Plasmid PCP20 was transformed into this strain to recombine out the CmR cassette flanked by FRT sites41, imparting Cm-sensitivity to the cell (LZ1535). Then, MG1655 dnaB-mTurquoise2-CmR-FRT (LZ1510) was generated by first constructing the plasmid, pdnaB-mTurquoise2-CmR-H (H is the downstream homologous region of dnaB). Next, PCR was performed to create a linear dsDNA of the dnaB-mTurquoise2-CmR-H region, which was used for red-recombination42, to generate LZ1510. Then P1 transduction43 was performed to move the dnaB-mTurquiose2 reporter from LZ1510 (donor) to LZ1535 (recipient), making a strain with both the SeqA and DnaB reporters (LZ1552). Next, the Δdam-KanR marker was transduced from LZ1386 to LZ1552 to complete the SeqA reporter (LZ1555). This new strain was transformed with pACYC177 pFtsKi tetR-mCherry to complete the triple reporter strain, LZ1557.
To construct the attB reporter strain, MG1655 served as the parent, and a 96× lacO array was inserted via red-recombination. This was done by first inserting upstream (E. coli genome region amplified with the f-up-attb and f-up-attb primer pair) and downstream (E. coli genome region amplified with the f-down-attb and f-down-attb primer pair) homology regions flanking the 96× lacO KanR region of a plasmid40, and then digesting this plasmid (pattB 96× lacO KanR) within the homology regions to produce linear dsDNA for red-recombination. This strain was then transformed with pACYC177 pFtsKi tetR-mCherry lacI-eyfp.
To construct the lysogen with DnaB, TetR, and gpD reporters, LZ1510 (dnaB-mTurquoise2 CmR-FRT) served as the parent. The CmR cassette was removed using PCP20 to generate LZ1511 (dnaB-mTurquoise2) and then transformed pACYC177 pFtsKi tetR-mCherry into the strain. This host was then lysogenized with λLZ1575 (cI857 D-mNeongreen bor::CmR 24×tetO), producing LZ1596.
The purification protocol was adapted from other sources12,44,45. Briefly, a single colony of desired lysogens was grown with appropriate antibiotics at 30 °C overnight. The overnight culture was then diluted into 500 ml and induced. The phages were then precipitated using 10% PEG8000 + 1 M NaCl. The resulting phage pellet was soaked in a total of 8 ml of cold SM buffer and incubated at 4 °C overnight, ~16 h. An organic extraction was performed by mixing the SM suspension gently with an equal volume of chloroform and centrifuging at 3000 × g for 15 min at 4 °C. The supernatant was removed to exclude the PEG pellet, and the extraction step was done two more times, to finally yield a clear supernatant containing the phage. A step gradient was made for each desired phage using 1.5 ml each of 1.3, 1.5, and 1.7 g/ml CsCl + SM buffer solutions, and the phage (~8 ml) was layered on top in a 13.2 ml ultraclear tube (Beckman Coulter), then ultracentrifuged in a Beckman SW41Ti rotor at 24,000 r.p.m. for 6–8 h at 4 °C. The phages migrated to a band and were then extracted from the side wall of the tube. This phage extraction was then loaded into a 5 ml ultraclear tube (Beckman Coulter) and then filled with a 1.5 g/ml CsCl + SM buffer and ultracentrifuged in a Beckman SW50 rotor at 35,000 r.p.m. for 24 h at 4 °C, and then was extracted in the same manner. This new phage extraction was then dialyzed 1:1000 against SM buffer over ~24 h. This phage solution was then extracted and stored away from light and at 4 °C to be used in the experiments.
Further information on research design is available in the Nature Research Reporting Summary linked to this Article.
All strains and raw experimental data are available from the corresponding author upon request. Source data are provided with this paper.
Data analysis scripts are available from public repositories at https://doi.org/10.5281/zenodo.3906930.
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We are grateful to Craig Kaplan and Ryland Young for commenting on the earlier versions of the manuscript. We thank David J. Sherratt for gifting strains. This research was supported by National Institutes of Health (grant R01GM107597), National Science Foundation (grant 2013762), and Texas A&M University X-Grant (grant 290386) to the Zeng Laboratory.
The authors declare no competing interests.
Peer review information Nature Communications thanks Mathew Stracy, and other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.
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Trinh, J.T., Shao, Q., Guan, J. et al. Emerging heterogeneous compartments by viruses in single bacterial cells. Nat Commun 11, 3813 (2020). https://doi.org/10.1038/s41467-020-17515-8
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