Two unrelated Pseudomonas aeruginosa phages require the exopolysaccharide Psl for infection

Two unrelated Pseudomonas aeruginosa phages require the exopolysaccharide Psl for infection | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article Two unrelated Pseudomonas aeruginosa phages require the exopolysaccharide Psl for infection Kristen Amyx-Sherer, Leila C. Awasthi, Amanda Zheng, Anna Johannesman, and 2 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-6623495/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 18 Nov, 2025 Read the published version in npj Biofilms and Microbiomes → Version 1 posted 9 You are reading this latest preprint version Abstract Bacteria commonly protect themselves from a variety of threats by forming biofilms, which are communities of bacteria that are tightly packed together within an extracellular matrix. Biofilm formation has generally been thought to protect bacteria from phage infection. The opportunistic pathogen Pseudomonas aeruginosa produces biofilm matrices that can contain three distinct exopolysaccharides that contribute to the difficulty in treating infected patients. Here, we demonstrate that two diverse P. aeruginosa phages have evolved to exploit this biofilm matrix to access the bacterial cells by both binding to and degrading a major biofilm exopolysaccharide, Psl. We examined the effect of these phages on biofilms in different in vitro biofilm models and found that both phages prevent bacterial surface attachment, but only one of the two phages can disrupt a mature biofilm. The phages also rapidly lead to the emergence of bacterial strains that produce reduced amounts of Psl and are unable to adhere to surfaces. These phages may be useful therapeutically by driving bacteria away from producing biofilms and shifting P. aeruginosa cells into the more treatable planktonic growth state. Biological sciences/Microbiology/Bacteria Biological sciences/Microbiology/Biofilms Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Introduction A major obstacle to the treatment of bacterial infections is the growth of bacteria in protective aggregates called biofilms, which occur in roughly 80% of human bacterial infections 1 . Within biofilms, bacteria are encased in a complex extracellular matrix that may be composed of a range of macromolecules such as exopolysaccharides, extracellular DNA, and proteins. This matrix serves as a barrier that can exclude a wide range of threats, ranging from antibiotics to immune molecules 2 . As a result, when pathogenic bacteria form biofilms in humans, these infections often become chronic and untreatable. The opportunistic pathogen Pseudomonas aeruginosa , a model for the study of biofilms, frequently causes biofilm-involved infections in a range of settings, including the lungs of people with cystic fibrosis, catheter or ventilator-associated infections, and chronic wounds. The P. aeruginosa biofilm matrix can vary in composition, with many clinical isolates of P. aeruginosa from the lungs of people with cystic fibrosis dominated either by alginate or by a mixture of the exopolysaccharides Pel and Psl 3 , 4 , as is the case for rugose small colony variants (RSCV). RSCVs are a hyper-biofilm-forming variant, which commonly arise in chronic infections and are especially recalcitrant to treatment 5 , 6 . The biofilm matrix is generally thought to exclude bacteriophages (phages), the viruses that infect bacteria, thus limiting the utility of phage therapy for biofilm-involved infections 7 , 8 . However, few studies have examined these interactions at the cellular or molecular level. Phage propagation and characterization are typically performed using a double overlay assay in which bacteria are suspended in a gel-like agar matrix that is normally associated with bacterial motility 9 , rather than biofilm formation, and is thus not well suited to biofilm-phage interaction studies. Only a small handful of studies have investigated phages using in vitro biofilm models 10 – 13 . Commonly used static biofilm assays only represent one aspect of biofilm development and may miss some of the different effects that phages can exert on biofilms. For example, in a static biofilm assay as opposed to biofilms grown under flow (e.g., flow cell or flow tube biofilms), the phages do not have to contend with flow, which may wash away less adherent phages, but they are faced with relatively high concentrations of extracellular bacterial waste products 14 . Further, bacterial physiology, which matters for the efficacy of phage infection, varies depending on the biofilm model that is used in the experiment. For these reasons, it has been suggested that phages be tested against multiple biofilm models in order to gain a better understanding of phage-biofilm interactions 15 . Interestingly, many phages are known to produce depolymerase enzymes as part of their tail fibers. These depolymerases have been shown to interact with specific cell-associated extracellular sugars such as lipopolysaccharides (LPS) or the capsule-associated exopolysaccharides (CPS) 16 . However, although a few phages have been demonstrated to specifically interact with biofilm matrix exopolysaccharides 17 ,1819 , phages that degrade biofilm matrix components, including exopolysaccharides, have not been reported. Indeed, given the diversity of both bacterial species and their phages, we have very little knowledge about how most phages interact with bacterial biofilms or whether biofilms do indeed represent a first line of defense against phage infection. A set of studies, albeit with only a single bacterial and phage species, highlight the complexity of biofilm-phage interactions and provide motivation for further studies to probe these interactions. Specifically, in some instances, the spatially segregated, heterogeneous bacterial population present in biofilms supports bacterial co-existence with phages 12 . Studies using E. coli biofilms that contain the functional bacterial amyloid called curli observed that the model T7 phage is trapped by curli, but depending on the location of the phages and the biofilm age, this interaction can both prevent T7 particles from accessing bacteria at the biofilm aggregate interior and also serve as a phage reservoir for newly arriving bacterial cells 10 , 11 . Here, we describe the discovery of two unrelated P. aeruginosa phages – one podo- and one siphophage – that both require the exopolysaccharide Psl for infection. We find that both of these Psl-dependent phages require Psl for adsorption to bacteria and degrade Psl to access the bacterial surface. Further, using an in vitro biofilm model, we find that both phages can prevent bacterial adherence to a surface. However, only one of the phages can disrupt a mature biofilm. Selective pressure of these phages leads to the rapid emergence of P. aeruginosa escape mutants with severely reduced Psl production and an inability to adhere. Results Isolation of two Psl-dependent P. aeruginosa phages To identify phages that may have exopolysaccharide-degrading properties, we set out to test whether any phages in our collection formed haloes on an exopolysaccharide overproducing strain, ∆ wspF wherein the deletion of the negative regulator wspF leads to the constitutive upregulation of the exopolysaccharides, Pel and Psl 20 . To our surprise, we found that two of our phages (PaStL1M and PaStL2M) displayed an increase in titer on the ∆ wspF strain (Fig. 1 A), with distinct plaquing morphology. Based on the plaquing patterns, we hypothesized that each of our phage stocks actually contained two distinct phages, and that these two phages had different replication capacities on PAO1 compared with ∆ wspF . To determine whether the phage with a preference for ∆ wspF was dependent on a particular exopolysaccharide, we also tested the phage mixtures on strains lacking genes encoding for key early enzymes in the Psl and Pel biosynthetic pathways ( Table S1 ), and found that plaques on ∆ wspF ∆psl resembled those on PAO1, suggesting that the second phage in the mixture was Psl-dependent and that only one of the two phages in each mixture was replicating on ∆ wspF ∆psl ( Fig S1 ). We also tested plaquing on a clinical isolate, CF127, previously shown to overproduce Psl 4 , and its derivative CF127 ∆ psl , and noted reduced plaques on CF127, and no plaques on CF127 ∆ psl , suggesting that only the Psl-dependent phage in each case could replicate on CF127. We thus purified the original PaStL1M and PaStL2M phage stocks using both ∆ wspF ∆psl and CF127. This resulted in four isolated phages: two Psl-dependent phages (PaStL1 and PaStL2) that can replicate on ∆ wspF and CF127, but not PAO1, and two Psl-independent phages (PaStL3 and PaStL4), that can replicate on all strains except CF127 and CF127 ∆ psl (Fig. 1 B). We were unaware of any previously reported Psl-dependent phages at the time, and thus decided to further characterize PaStL1 and PaStL2. Since both phages were unable to form plaques on PAO1, we wondered how they were present at high titers when titered on an exopolysaccharide overproducing strain such as ∆ wspF or CF127. These observations suggested that they must be able to replicate in planktonic culture on the wild-type PAO1 strain that we had used for all enrichment and isolation steps. Indeed, when we added these phages at a low multiplicity of infection (MOI) of 0.01 to PAO1 and monitored bacterial growth under standard planktonic growth conditions, we observed a typical infection curve with a loss of bacterial density beginning around 2.5 h after phage addition (Fig. 1 C). Planktonically growing PAO1 thus produces sufficient levels of Psl under standard laboratory growth conditions in rich media to sustain infection by these Psl-dependent phages, while Psl is likely down-regulated in double overlay assays, which mimic conditions typically used for bacterial swarming assays (0.6% agar) where production of biofilm exopolysaccharides is reduced 21 . Sequencing these phages revealed that PaStL1 and PaStL2 are both new species of distinct genera: PaStL1 is a Bruynogevirus, while PaStL2 is in the Iggyvirus genus of the Queuovirus subfamily. Transmission electron microscopy of these phages further confirmed a podo- and siphophage morphology, consistent with the sequencing data (Fig. 1 D). We note that these phages are similar to reports of Psl-dependent phages that were made in the course of our investigations 22 – 24 . Sequencing of PaStL3 and PaStL4 confirmed that the co-isolated, non-Psl-dependent phages are unrelated: PaStL3 is a new species of Yuavirus, while PaStL4 is a strain of Pbunavirus Ab28. PaStL1 and PaStL2 bind and degrade Psl We used an adsorption assay to assess whether phage attachment to bacterial cells was dependent on Psl. Specifically, phages were incubated with bacteria that did not express Psl (i.e., PAO1 ∆ wspF ∆ pel ∆ psl ) or expressed high levels of Psl via an arabinose-inducible promoter (i.e., PAO1 ∆ wspF ∆ pel pBAD psl ). The phages PaStL1 and PaStL2, which both required bacterial production of Psl for infectivity in a plaque assay, displayed high levels Psl-dependent adsorption (Fig. 2 A). In contrast, the presence of Psl did not impact adsorption of either PaStL3, which did not adsorb well to either strain, or PaStL4, in which 90% of the phage adsorbed to both strains. Together, these results support that PaStL2 and PaStL3 uniquely use Psl as a receptor for binding to the bacterial cells, while the two non-Psl-dependent, co-isolated phages do not require Psl for adsorption. Next, we determined if the phages were able to depolymerize Psl, as has been observed for other phages that bind and degrade the surface glycan, o-antigen 16 . To do so, we incubated purified phage with cell-free supernatant preparations from the Psl- and non-Psl-producing strains, PAO1 ∆ wspF and PAO1 ∆ wspF ∆ psl ∆ pel ∆ algD (referred to as PAO1 ∆ wspF ∆EPS), respectively. Following 6h of incubation at 37°C, we detected Psl levels using an anti-Psl immunoblot (Fig. 2 B). Incubation with phages PaStL1 and PaStL2 decreased Psl levels. In contrast, incubation with either PaStL3 or PaStL4 did not discernibly change Psl levels relative to the no-phage control. As is expected for depolymerization of Psl by an enzyme, heat inactivation of the phages prior to incubation with the cell-free supernatant abolished the ability of the phages to depolymerize Psl. We surveyed the phage genomes for depolymerases using DePolymerase Predictor (DePP) 25 and identified putative depolymerases encoded in the genomes of both PaStL1 and PaStL2. These putative depolymerases had different sequences as well as strikingly different structures, as predicted by AlphaFold ( Fig S2 ). Notably, there were no similar depolymerases predicted that are shared between PaStL1 and PaStL2, suggesting that these two phages have independently evolved mechanisms for binding to and degrading Psl. These results support that both PaStL1 and PaStL2 bind and depolymerize Psl, but they likely do so via different depolymerases. PaStL1 and PaStL2 bind and degrade Psl Given the Psl-dependence of the phages, we predicted that they might be effective against biofilms. To test this, we grew biofilms statically in 96-well microtiter dishes for 24 h, removed any unattached biomass, and treated the remaining adherent biomass with the phages. We tested each of the phages individually as well as the two original phage mixtures, PaStL1M (i.e., PaStL1 and PaStL3) and PaStL2M (i.e., PaStL2 and PaStL4). As evidenced by the decrease in adherent biomass relative to the no phage control, both Psl-dependent phages, PaStL1 and PaStL2, were more effective at clearing biofilms formed by both PAO1 and PAO1 ∆ wspF than their co-isolated counterparts (Fig. 2 C). The original phage mixtures showed differing results, with PaStL1M being similarly effective as PaStL1 alone and PaStL2M being less effective than PaStL2 alone. Similar trends were observed for phage treatment of static biofilms formed by PAO1 ∆ wspF ∆pel . However, the Psl-dependent phages had a reduced ability to clear biofilms formed by PAO1 ∆ wspF ∆psl . Interestingly, some of the phage-treated PAO1 ∆ wspF ∆psl biofilms had increased adherent biomass relative to the respective no-phage control. This could be due to the release of extracellular DNA if the phages lysed a subset of the bacteria, or due to the phage-induced bacterial stress triggering an increase in biofilm formation, as has been previously described 26 . Overall, this static biofilm assay supports that, in addition to both PaStL1 and PaStL2 requiring Psl for infection, both phages clear biofilms in a Psl-dependent manner. Further, this result shows that when PAO1 is cultured as a biofilm and is making high levels of Psl, we observe similar Psl-dependence of these two phages as for PAO1 ∆ wsp F biofilms. Only PaStL2 clears mature biofilms cultured in flow cells Given the role of Psl in P. aeruginosa surface attachment, we investigated whether the Psl-dependent phages, PaStL1 and PaStL2, could block bacterial surface attachment. To do so, we performed a bacterial attachment assay in microfluidic devices called flow cells. In this experiment, PAO1 ∆ wspF bacteria that constitutively expressed the fluorescent protein GFP (PAO1 ∆ wspF GFP + ) were allowed to attach to the surface of a flow cell for 10 min while the flow was stopped prior to the addition of phage. Purified phages were added and incubated with the bacteria for an additional 30 min, and then flow was resumed. The flow cells were imaged by confocal microscopy, and the numbers of attached bacteria were quantified using BiofilmQ 27 . We observed that incubation with both Psl-dependent phages, PaStL1 and PaStL2, decreased bacterial attachment to the flow cell surface (Fig. 3 A). To test the impact of the phages on established biofilms, we cultured PAO1 ∆ wspF GFP + in flow cells for 72 h, and then phages were added and statically incubated with the biofilms for 30 min before resuming flow. In this experiment, in addition to monitoring the bacteria, we also monitored Psl levels by staining with the fluorophore-conjugated Psl-specific lectin, Hippeastrum hybrid lectin (HHL)-TRITC. Biofilms were stained and imaged 2, 10, 16, and 26 h following phage addition (Fig. 3 B-D). Separately cultured biofilms were stained and imaged for each of the time points to ensure that staining with the Psl-specific lectin did not impact the results. Without phage incubation, typical biofilms formed with bacterial aggregates that extend into the flow cell channel approximately 60 mm and have peripherally localized Psl (Fig. 3 B-C). At 2 and 10 h after the addition of phage, the flow cell biofilms appeared similar to one another, regardless of which phage had been added (Fig. 3 B, S3A). By 16 h after the addition of phage, biofilms that had been incubated with PaStL2 began to clear, and by 26 h, biofilms that had been incubated with PaStL2 were entirely cleared (Fig. 3 C, S3B). Biofilms incubated with PaStL1 did not clear, even by 26 h after the addition of phage. We quantified the overall biomass (GFP fluorescence) and Psl levels (HHL-TRITC fluorescence) using BiofilmQ, which allowed us to make two additional observations (Fig. 3 D-E). First, for biofilms incubated with PaStL2, the Psl levels appeared to decrease more rapidly than the bacteria, suggesting that degradation of Psl precedes overall biofilm clearing. The second observation was that biofilms that were incubated with PaStL1 showed slightly increased levels of bacteria relative to biofilms that were not incubated with phage. Possibly, this slight increase in biofilm bacteria could be due to the lysis of the bacteria by the phage, resulting in increased extracellular DNA levels and, thus, bacteria within the biofilm aggregates. As controls, we performed two additional flow cell experiments. In the first experiment, we tested whether the ∆ wspF mutation impacted the ability of PaStL2 to clear the biofilms. To do so, we cultured flow cell biofilms of PAO1 for 72 h, added PaStL2, and then observed the flow cell biofilms 26 h later. We found that PaStL2 also cleared PAO1 flow cell biofilms, indicating that the ∆ wspF mutation was not necessary for clearing by PaStL2 ( Fig S4 ). This control experiment was important since the ∆ wspF mutation has impacts on the bacteria beyond simply increasing Psl production. In the second control experiment, we tested whether Psl was required for clearing of flow cell biofilms by PaStL2. To do so, we cultured flow cell biofilms of PAO1 ∆ wspF ∆ psl for 72 h, added PaStL2, and checked for clearing 26 h later. We observed that PaStL2 was unable to clear PAO1 ∆ wspF ∆ psl biofilms ( Fig S5 ). As previously reported, PAO1 ∆ wspF ∆ psl flow cell biofilms appear different than PAO1 ∆ wspF flow cell biofilms, even without phage infection, due to the reliance on Pel rather than Psl 28 . Together, these findings show that PaStL2 is able to clear both PAO1 and PAO1 ∆ wspF flow cell biofilms, and that bacterial production of Psl is required for it to clear. PaStL2 propagates in biofilms grown under flow Given the unexpected difference in the abilities of PaStL1 and PaStL2 to clear flow cell biofilms, we wondered if PaStL1 had a reduced ability, compared to PaStL2, to propagate in bacteria cultured under flow. To test this hypothesis, we cultured PAO1 ∆ wspF GFP + biofilms for 72 h in silicone tubing, which was cut to have a similar internal volume to the flow cells used in the prior experiment, and then statically incubated phage with the biofilms for 30 min before resuming flow. The biofilms were harvested 26 h later, and both colony-forming units (CFUs) and plaque-forming units (PFUs) were determined. CFUs were determined using the entire harvested biofilm, and PFUs were determined using the cell-free supernatant of the biofilm. To avoid confounding effects due to possible phage killing during the CFU assay, we determined CFUs indirectly by measuring the GFP fluorescence of the biofilms, which we correlated to CFUs via a standard curve of CFU versus GFP fluorescence that was obtained for PAO1 ∆ wspF GFP + bacteria that were cultured in the absence of phage ( Fig S6 ). Incubation of PaStL2, but not PaStL1, resulted in a decrease of the biofilm bacteria ( p < 0.05), although the decrease in this growth system was not as dramatic as was observed in the flow cell experiment (Fig. 4 A). Specifically, the CFUs for biofilms with PaStL1 added were ~ 5 x 10 9 CFU/mL and with PaStl2 added, only around 1 x 10 9 CFU/mL. A much more dramatic difference was observed for PFUs. For the biofilms treated with PaStL1, we recovered only ~ 5x10 4 PFUs/mL, and for those treated with PaStL2, we recovered 2x10 10 PFUs/mL (Fig. 4 B). To determine the dynamics of PaStL2 phage infection, we also harvested separately cultured biofilms at 2, 10, and 16 h post-phage addition, and similarly determined both CFUs and PFUs (Fig. 4 C). We observed that PaStL2 titers increased at 16 h, which preceded the decrease in biofilm bacteria CFUs. Given that the titer of the phage stock added to the biofilm was ~ 10 8 PFUs/mL, with a similar volume as the recovered cell-free supernatant of the biofilm (300 µL vs ~ 250 µL, respectively) these data support that PaStL2 is able to propagate within a biofilm cultured under flow, and PaStL1 is not. P. aeruginosa evades PaStL1 and PaStL2 via loss of Psl We noted that both in planktonic growth curves and on top agar, resistant bacteria typically emerged following infection with PaStL1 and PaStL2. We isolated individual colonies that emerged in planktonic growth of PaStL1 as well as within plaques of PaStL2 (Fig. 1 B) and then retested these for sensitivity to both phages. A subset of these clones was sensitive to phage infection, suggesting a regulatory response to phage infection that reverted upon reculturing. However, many of these clones were persistently resistant, and we found that in all cases, resistant to both PaStL1 and PaStL2. We hypothesized that these cells had likely lost the ability to produce Psl. To test this idea, we performed dot blots and found that all of the escape mutants produced significantly less Psl than either the PAO1 or ∆ wspF strain (Fig. 5 A). We also found that these escape mutants all had reduced ability to adhere to a surface as measured in a static biofilm assay (Fig. 5 B). We sequenced a subset of these escape mutants and found that ∆ wspF-PaStL1EA , that arose via resistance to PaStL1 had a mutation in pslA that led to a truncation of the protein, while surprisingly ∆ wspF-PaStL1B had only a substitution in a PA1327, a predicted protease (Table 1 ). The escape mutants isolated from double overlays had all acquired a 7-base duplication in the wspR gene that encodes the regulator controlling the Wsp pathway, and in some cases additional mutations as well (Table 1 , S2). Taken together, these data demonstrate that both PaStL1 and PaStL2 provide a strong selective pressure that leads to the loss of Psl production in P. aeruginosa . Table 1 Mutations present in bacterial escape mutants to PaStL1 and PaStL2 genes mutated designation phage locus Mutation(position) locus Mutation(position) a PaStL1 pslA frameshift b PaStL1 PA1327 R245P c PaStL2 wspR frameshift d PaStL2 wspR frameshift pilY1 truncation (553) e PaStL2 wspR frameshift f PaStL2 wspR frameshift Discussion Despite that numerous studies have pursued phage or phage cocktails as therapeutics for recalcitrant bacterial infections, including biofilm infections, there has been a general lack of mechanistic insight into interactions of phage with biofilms. In this study, we discovered two phages that require the P. aeruginosa biofilm matrix exopolysaccharide Psl for attachment to their bacterial hosts. PaStL1 is related to recently reported Psl-dependent phages Clew-1 22 and LUZ24 24 , while PaStL2 is related to the siphophage Knedl 23 , 29 . We find that both phages, using different depolymerase proteins, can both bind to and degrade Psl, and that as a result, they are able to prevent bacteria from establishing biofilms by eliminating the ability of the cells to attach to the surface. We further find that only PaStL2 can disrupt a fully formed biofilm that is cultured under flow, a finding not reported in other related studies that did not assess phage impact on this biofilm model. We further demonstrate that both phages strongly select for non-biofilm-forming bacteria that do not produce Psl and are unable to adhere to surfaces to establish biofilms, even in the absence of phage. These results demonstrate the utility of these phages in therapeutic applications where they could both disrupt biofilms, as well as drive bacteria into a planktonic growth state where they would be more amenable to treatment. This study highlights the importance of considering phage isolation methodology when performing environmental phage isolations: phages will only replicate and form plaques on strains that are producing necessary receptors. Both of the Psl-dependent phages that we isolated were originally co-isolated with another non-Psl-dependent phage. This co-isolation was essentially what enabled us to discover these phages, as without the second phage present, our initial plaque assays on PAO1 would have been blank and falsely suggested that there was no phage present in our sample. The proteins and exopolysaccharides present on the cell surface are highly dynamic, as this first line of defense against many threats is critical to bacterial survival. Using conditions in which bacteria produce proteins and exopolysaccharides present during infections is critical for finding phages that are likely to be useful in therapeutic applications. In addition to requiring Psl for absorption to the bacterial cell, we found that both PaStL1 and PaStL2, but not PaStL3 or PaStL4, result in the degradation of Psl following their incubation with Psl-containing culture supernatants. We predict that both phages contain depolymerases, as predicted by bioinformatic analyses and analogous to the depolymerases that phages employ to degrade LPS and CPS. Unlike LPS and CPS in many bacterial species, P. aeruginosa Psl is generally believed to be chemically identical or similar across P. aeruginosa isolates due to the extremely high conservation of the Psl operon 30 . As such, Psl may serve as a better target for phage-based therapies as opposed to LPS or CPS, since bacteria cannot simply modify Psl to escape phage infection. Rather, as we observed in our experiments, bacteria that evolve to escape infection by PaStL1 or PaStL2 lose the ability to produce Psl altogether, which makes them unable to form biofilms, likely rendering them susceptible to more typical antibacterial therapeutics. This study also highlights the need to evaluate phage-biofilm interactions in multiple biofilm models since phages can have dramatically different effects on different types of biofilms 15 . For example, we observed that both Psl-dependent phages, PaStL1 and PaStL2, cleared biofilms that were cultured statically in microtiter dishes. However, only PaStL2, and not PaStL1, could clear mature biofilms that were grown under flow growth conditions (i.e., flow cell biofilms, tube biofilms). While these are still very much simplified in vitro models, biofilms cultured under flow likely better mimic turbulent or dynamic conditions encountered in some infections (e.g., infections of catheters, heart valve infections, etc), and thus, PaStL2 might be a better therapeutic candidate than PaStL1. The reason(s) for the different abilities of PaStL1 and PaStL2 to clear biofilms cultured under flow could include that PaStL1 does not bind as well as PaStL2 or perhaps the bacterial physiology in these biofilms is less amenable to PaStL1 propagation. Our results point toward both of these options as possibilities, as more PaStL2 absorbed to Psl-producing bacteria compared to PaStL1, and PaStL2 was able to propagate in tube biofilms while PaStL1 did not, although additional experiments are required to say definitely. Overall, in this study, we discovered two different phages that require Psl for infection and can degrade Psl and clear P. aeruginosa biofilms. In addition to providing candidates for phage therapy of chronic P. aeruginosa infections, the predicted Psl depolymerases could serve as anti-biofilm candidates on their own, similar to other hydrolases that degrade matrix exopolysaccharides and help to break apart biofilms, potentially broadening their potential impact beyond the narrow strain profile of these phages. This is an interesting avenue for future research, as currently, nearly all candidate hydrolases for anti-biofilm therapeutics come from either bacteria or fungi, and phage may be an untapped source of novel enzymes to break apart biofilms. Lastly, our study serves as a roadmap for future discovery of additional biofilm-targeting phages. Methods Bacterial strain growth. Planktonic cultures of P. aeruginosa were routinely grown on Lysogeny broth (LB) or TSB medium at 37°C with constant shaking (225 rpm) unless indicated otherwise. Strains are listed in Table S1 . Phage isolation . Wastewater from a local treatment facility was centrifuged to remove particulates, then filtered with a 0.22 µm filter to remove bacterial cells. The resulting supernatant was mixed with concentrated LB broth for a final 1x concentration of LB and mixed with an overnight culture of PAO1 at 1:100. The suspension was incubated overnight at 37°C with aeration. The following day, the culture was centrifuged to pellet bacteria and filtered, and the supernatant was spotted on double overlay plates prepared with 4 mL of 0.6% LB agar with 200 µL of an overnight PAO1 culture and poured onto a 1.2% LB agar plate. Plates were examined for clearing after 6–16 h, and for lysates that displayed clearing, the culture was serially diluted to determine an estimate of titer as described below with the spot titer assay, then plated for single plaque isolations using the full plate assay. Individual plaques were picked with a P10 pipet tip and added to 100 µL of a 1:10 dilution of an overnight PAO1 culture in LB and incubated overnight at 37°C in a microcentrifuge tube. The resulting lysate was centrifuged and plaque purifications were performed 3x. Phage titer assays. Phage concentration was determined by measuring plaque-forming units / mL (PFU / mL) using either a spot titer or full plate assay. 200 µL of an overnight bacterial culture was mixed with 4 mL of melted 0.6% LB agar (55°C) and poured on top of a 1.2% LB agar plate. Once solidified, 2–40 µL of a 10x serially diluted phage was spotted and allowed to dry before incubation of plates for 5–16 h at 37°C. Alternatively, 100 µL of phage was added directly to the top agar. PFUs / mL were calculated by counting the number of plaques in the lowest serial dilution and then multiplying by the dilution factor. Phage propagation and purification. P. aeruginosa phages were produced by back-diluting overnight cultures of PAO1 to an OD 600 of 0.1 in LB, adding an aliquot of phage stock to a final multiplicity of infection (MOI) of 0.1, and then incubating the cultures for 5 h at 37°C with constant shaking (225 rpm). After incubation, bacterial cells were separated from the supernatant by centrifugation (6,000 x g for 20 min) at 4°C. The supernatant was passed through a 0.22 µm filter and used directly for tittering assays and adsorption experiments. For all biofilm experiments, the filtered lysate was further purified via ultracentrifugation through a 35% (w /v) sucrose cushion at 23,5000 x g for 50 min. The supernatant was removed via decanting, and the pellet was suspended in 1 mL of 1x PBS. The sample was centrifuged at 6,000 x g for 20 min and then dialyzed (8–10 kDa MWCO Float-A-Lyzer G2 Dialysis Device) (MilliporeSigma) against 1x PBS overnight at 4°C. Afterwards, the sample was centrifuged at 17,000 x g for 10 min. The phage was then either used in assays directly or further subjected to size-exclusion chromatography (1x PBS; HiPrep 16/60 sephacryl S-500 HR column) (Cytiva). Phage-containing fractions were concentrated 10-fold using a centrifugal filter (50 kDa MWCO) (MilliporeSigma). Transmission Electron Microscopy. For analyses of phages at the ultrastructural level, samples were allowed to absorb onto freshly glow discharged formvar/carbon-coated copper grids (200 mesh, Ted Pella Inc., Redding, CA)) for 10 min. Grids were then washed two times in dH2O and negative stained with 1% aqueous uranyl acetate (Ted Pella Inc.) for 1 min. Excess liquid was gently wicked off and grids were allowed to air dry. Samples were viewed on a JEOL 1200EX transmission electron microscope (JEOL USA, Peabody, MA) equipped with an AMT 8 megapixel digital camera (Advanced Microscopy Techniques, Woburn, MA). Static biofilm assay. Static biofilm formation was assessed by performing a crystal violet assay 31 . Biofilms were cultured in Nunc Bacti 96-well microtiter plates (Fisher Scientific) using TSB. 100 mL of mid-log culture was used to inoculate each well of the microtiter plate. The plates were statically incubated for 24 h at 37°C. Non-adherent cells were removed, and wells were washed with 150 mL 1x PBS. Then 100 mL of phage (5x10 7 PFUs/mL, 1x PBS buffer) or buffer alone was added to each well and statically incubated for 4 h at 37°C. Non-adherent cells were removed by pipetting, and the wells were washed three times with 150 mL of ddH 2 O. The 96-well plate was inverted and allowed to dry for 30 min before the addition of 150 mL of 0.1% (w/v) crystal violet (Fisher Scientific). The plate was statically incubated with crystal violet for 15 min. The crystal violet solution was removed, and the wells were washed three times with 150 mL of ddH 2 O. The plate was inverted and allowed to dry overnight. Finally, the crystal violet was solubilized with 200 mL of 95% ethanol, and 100 mL was transferred to a fresh plate and the absorbance at 595 nm was measured (Varioskan LUX multimode microplate reader, Thermo Fisher Scientific). Phage DNA sequencing. Phage DNA was extracted using a Norgen Phage DNA Extraction kit and short read Illumina sequencing was either performed by SeqCoast Genomics (Portsmouth, NH, USA) or libraries were prepared using Illumina Tagmentation reagents with a low input protocol 32 , and sequenced at the DNA Sequencing and Innovation Lab at the Edison Family CGS&SB Sequencing Center. Reads were de novo assembled in Geneious Prime (version 2025.1.1) and the TaxMyPhage 33 tool was used to determine taxonomy. Phage genomic sequencing data is available at PRJNA1260455. Adsorption assays. Overnight bacterial cultures were back-diluted into LB media supplemented with 1% (w/v) L-arabinose, and then grown overnight with shaking (225 rpm) at 37°C. A 1-mL aliquot of the culture was centrifuged at 5000 x g for 10 min, the supernatant discarded, and the pellet suspended in 1 mL of 1x PBS. Phage was added to achieve a multiplicity of infection (MOI) of 0.01 and incubated for 15 min at room temperature with occasional inversion. 20 µL was removed for serial dilution; the remaining sample was centrifuged at 8000 x g for 3 min and a 20 µl aliquot of the supernatant was collected for serial dilutions. 20–40 µL of these dilutions were spotted on double overlay assays to determine relative titers. Psl degradation assay. Overnight cultures of PAO1 ∆ wspF and PAO1 ∆ wspF ∆ EPS in TSB were back diluted 100-fold and grown for 16 h at 37°C with shaking (225 rpm). Cultures were normalized to OD 600 of 1.0 and the centrifuged for 2 min at 15060 rpm. The supernatant was then incubated with bacteriophage (10:1 supernatant to phage, 1x10 8 PFUs/mL) at 37°C for 6 h. Immediately after incubation, samples were boiled at 98°C for 5 min and cooled to 12°C. Then samples were treated with proteinase-K (2 mg/mL) (Qiagen) at 60°C for 1 h, 80°C for 30 min, and then cooled down to 12°C. The presence of Psl was assessed by anti-Psl immunoblot. Anti-Psl Immunoblot. To assess for the presence of Psl, samples were analyzed by anti-Psl immunoblot as previously described 35 . First, samples were treated with proteinase-K (2 mg/mL) (Qiagen) at 60°C for 1 h, 80°C for 30 min, and then cooled down to 12°C. After proteinase K treatment, 5 µL of sample was loaded onto a nitrocellulose membrane (Bio-Rad) (0.2 mm) and allowed to dry for 10 min. The membrane was blocked for 1 h with 5% (w/v) milk, 10 mM Tris pH 7.5, 150 mM NaCl, 0.1% Tween 20 (TBST). Then the membranes were probed with anti-Psl primary antibodies (0.635 mg/mL WapR-001, 0.371 mg/mL WapR-016, 0.867 mg/mL Cam-003) (MedImmune) for 1 h 36 , 37 . Membranes were washed three times with TBST. The membranes were probed with horseradish peroxidase (HRP) conjugated goat anti-human antibody (Bio-Rad) (2 mL in 25 mL TBST) for 1 h. Membranes were washed three times with TBST. Detection was performed with Supersignal West Pico PLUS chemiluminescent substrate (Thermo Fisher Scientific) and blots were imaged with AlphaImager HP (Alpha Innotech). Attachment assay. Flow cells were inoculated with PAO1 ∆ wspF Tn7 Gm::P(A1/04/03)::GFP bacteria that were grown to mid-log phase in TSB and then diluted to OD 600 of 0.1 in 1% TSB. Bacterial cells were allowed to attach under static conditions in an inverted flow cell for 10 min and then incubated with 300 µL of phage (1x10 8 PFUs/mL) for 30 min. Then, non-attached cells were washed away by initiating media flow (40 mL/h) through the flow cell for 20 min. Flow was reduced to 3 mL/h prior to imaging. Attached cells were visualized on Nikon Eclipse Ni-E confocal laser scanning microscope (20x objective), and six fields of view were captured (Ex 488 nm, Em 499–551 nm). Images were analyzed with BiofilmQ 27 . To extract 3D objects from the imported microscopy data, the biofilm images were segmented. To remove background fluorescence and fluorescence in between cells, the top-hat filter was set to 15 vox. Afterward, the Otsu thresholding method with a sensitivity of 0.25 was set. Cubes were used for the dissection method and were approximately 1 vox in length. The images for the GFP fluorescent channel were segmented with these parameters and then global biofilm properties were calculated to collect the bacterial volume. Flow cells imaging of biofilms. Biofilms were cultured in flow cells as previously described 38 . The flow cells were made of polysulfone, and the microfluidic chamber was 1.125 inches long, 0.185 inches wide, and 0.162 inches deep. A microscope coverglass (24x60 mm, 0.175 mm thick) (Fisher Scientific) was fixed to the top of the chamber using clear adhesive sealant (Permatex silicone RTV 80050). After allowing the sealant to dry, the flow cells were autoclaved. Sterile flow cells were inoculated with bacterial cultures of PAO1 ∆ wspF Tn7 Gm::P(A1/04/03)::GFP or as specified, that were grown to mid-log phase in TSB and then diluted to OD 600 of 0.01 in 1% TSB. Bacterial cells were allowed to attach under static conditions in an inverted flow cell for 1 h before the start of media flow. Biofilms were grown in 1% TSB for 72 h at 25°C under a constant flow rate (10 mL/h). After 72 h, flow was stopped, and the biofilm was statically incubated with 300 µL of phage in 1x PBS (1x10 8 PFUs/mL) for 30 min prior to resuming media flow. Biofilms were monitored 2, 10, 16, and 26 h after phage incubation. Biofilms were stained with 0.1 mg/mL HHL-TRITC (GlycoMatrix) to visualize Psl. After staining, flow cells were washed with medium at 10 mL/h for 5 min and visualized on Nikon Eclipse Ni-E confocal laser scanning microscope using 20X or 40X objective (GFP: Ex 488 nm, Em 499–551 nm; TRITC: Ex 561nm, Em 571–625 nm). Images were analyzed with BiofilmQ 27 . To extract 3D objects from the imported microscopy data, the biofilm images were segmented. The images were scaled down 0.25x for pre-processing, and to remove background fluorescence and fluorescence in between cells, the top-hat filter was set to 15 vox. Afterwards, Otsu thresholding method with a sensitivity of 0.25 was set. Cubes were used for the dissection method and were approximately 1 vox in length. The images for each fluorescent channel were segmented with these parameters, and then global biofilm properties were calculated to collect the bacterial and Psl volume. Flow tubes. Biofilms were cultured as tube biofilms as previously described with some modifications 39 . PAO1 ∆ wspF Tn7 Gm::P(A1/04/03)::GFP bacteria were grown to mid-log phase in TSB and then diluted to OD 600 of 0.01 in 1% TSB. This back-diluted culture then was used to inoculate a sterile 4.5-cm long Nalgene 50 platinum-cured silicon tubing (1/8 in interior diameter and ¼ in exterior diameter) (Thermo Fisher Scientific). Note that the actual tube length in which the biofilm grew after accounting for the lengths of the connecting luers was 3 cm. Bacterial cells were allowed to attach to the tubes under static conditions for 1 h before the start of media flow. Biofilms were grown in 1% TSB for 72 h at room temperature (approximately 25°C) under a constant flow rate (10 mL/h). After 72 h, flow was stopped, and the biofilm was statically incubated with 300 µL of bacteriophage (1x10 8 PFUs/mL) for 30 min. Then flow was resumed. Biofilms were collected at 2, 10, 16, and 26 h after phage incubation. The biomass inside of the tube was collected by using the handle of a sterile, disposable L-spreader (Fisher Scientific) to push out the biomass into a collection tube. The sample was vortex mixed for 15 s, and then fluorescence (Ex 488 nm, Em 520 nm) was recorded (Varioskan LUX multimode microplate reader) (Thermo Fisher Scientific). The sample was centrifuged for 2 min at 15060 rpm. The supernatant was removed and used for plaque assays. GFP standard curve for CFU calculation. To determine the colony forming units (CFUs) in biomass collected from flow tubes incubated with phage, the correlation between GFP fluorescence (due to constitutive expression of GFP by the bacteria) and CFUs was analyzed to create a standard curve. To do so, overnight cultures of PAO1 ∆ wspF Tn7 Gm::P(A1/04/03)::GFP that were grown in TSB were serially diluted, and then the fluorescence (Ex 488 nm, Em 520) was measured in a 96-well black plate (Thermo Fisher Scientific) with Varioskan LUX multimode microplate reader (Thermo Fisher Scientific). In parallel, CFUs of the samples were determined. The CFU values were plotted against GFP fluorescence to obtain a best line fit equation (y = 5.2545x10 7 (x) – 1.3505x10 7 ), where y is CFUs, and x is fluorescence. Bacterial escape mutant sequencing. Resistant mutants were used in titer assays to confirm resistance to PaStL1 and PaStL2. Resistant strains were further cultured and bacterial genomic DNA was extracted using a Qiagen Puregene Kit. Genomic DNA was sent to SeqCoast for short-read (Illumina) sequencing. Reads were mapped to PAO1 using Geneious Prime (version 2025.1.1) and analyzed for mutations using the Geneious Variant Finder tool, with the variant frequency cutoff set to 60% and coverage cutoff of 5. All mutations were compared with sequencing of the ∆ wspF background strain, and only mutations unique to the escape mutants were further analyzed. All sequencing data were deposited to PRJNA1260455. Declarations Author Contribution M.L. and C.R. designed the research; K.A.S., L.C.A., A.Z., and A.J. performed research; K.A.S., M.L., and C.R. analyzed data; and M.L. and C.R. wrote the manuscript. All authors reviewed the manuscript. Acknowledgement This work was supported by an Office of the Vice Chancellor for Research Interdisciplinary Seed Grant to M.L. and C.R., and unrestricted funds from Washington University School of Medicine to M.L. The Psl-specific antibodies were obtained by C.R. from AstraZeneca (Dr. DiGiandomenico). We thank Wandy Beatty for assistance and training on TEM imaging. We thank Forrest Walker for feedback and discussion. Data Availability All sequencing data were deposited to PRJNA1260455. Additional data is provided within the manuscript or supplementary information files. References Flemming, H.-C. & Wuertz, S. Bacteria and archaea on Earth and their abundance in biofilms. Nat. Rev. Microbiol. 17, 247–260 (2019). Karygianni, L., Ren, Z., Koo, H. & Thurnheer, T. Biofilm matrixome: extracellular components in structured microbial communities. Trends Microbiol. 28, 668–681 (2020). Colvin, K. M. et al. The Pel polysaccharide can serve a structural and protective role in the biofilm matrix of Pseudomonas aeruginosa . PLOS Pathog. 7, e1001264 (2011). Colvin, K. M. et al. The Pel and Psl polysaccharides provide Pseudomonas aeruginosa structural redundancy within the biofilm matrix. Environ. Microbiol. 14, 1913–1928 (2012). Häußler, S., Tümmler, B., Weißbrodt, H., Rohde, M. & Steinmetz, I. Small-colony variants of Pseudomonas aeruginosa in cystic fibrosis. Clin. Infect. Dis. 29, 621–625 (1999). Starkey, M. et al. Pseudomonas aeruginosa rugose small-colony variants have adaptations that likely promote persistence in the cystic fibrosis lung. J. Bacteriol. 191, 3492–3503 (2009). Ferriol-González, C. & Domingo-Calap, P. Phages for biofilm removal. Antibiotics 9, 268–16 (2020). Danis-Wlodarczyk, K. M., Wozniak, D. J. & Abedon, S. T. Treating Bacterial infections with bacteriophage-based enzybiotics: In vitro, in vivo and clinical application. Antibiot. Basel Switz. 10, 1497 (2021). Ha, D.-G., Kuchma, S. L. & O’Toole, G. A. Plate-based assay for swarming motility in Pseudomonas aeruginosa . Methods Mol. Biol. Clifton NJ 1149, 67–72 (2014). Vidakovic, L., Singh, P. K., Hartmann, R., Nadell, C. D. & Drescher, K. Dynamic biofilm architecture confers individual and collective mechanisms of viral protection. Nat. Microbiol. 3, 26–31 (2018). Bond, M. C., Vidakovic, L., Singh, P. K., Drescher, K. & Nadell, C. D. Matrix-trapped viruses can prevent invasion of bacterial biofilms by colonizing cells. eLife 10, e65355 (2021). Simmons, E. L. et al. Biofilm structure promotes coexistence of phage-resistant and phage-susceptible bacteria. mSystems 5, 385–17 (2020). Winans, J. B., Wucher, B. R. & Nadell, C. D. Multispecies biofilm architecture determines bacterial exposure to phages. PLOS Biol. 20, e3001913 (2022). Rumbaugh, K. P. & Whiteley, M. Towards improved biofilm models. Nat. Rev. Microbiol. 23, 57–66 (2025). Abedon, S. T., Danis-Wlodarczyk, K. M., Wozniak, D. J. & Sullivan, M. B. Improving phage-biofilm in vitro experimentation. Viruses 13, (2021). Knecht, L. E., Veljkovic, M. & Fieseler, L. Diversity and function of phage encoded depolymerases. Front. Microbiol. Volume 10-2019, (2020). Gutiérrez, D. et al. Role of the pre-neck appendage protein (Dpo7) from phage vB_SepiS-phiIPLA7 as an anti-biofilm agent in Staphylococcal species. Front. Microbiol. 6, (2015). Wu, Y. et al. A novel polysaccharide depolymerase encoded by the phage SH-KP152226 confers specific activity against multidrug-resistant Klebsiella pneumoniae via biofilm degradation. Front. Microbiol. 10, (2019). Cornelissen, A. et al. Identification of EPS-degrading activity within the tail spikes of the novel Pseudomonas putida phage AF. Virology 434, 251–256 (2012). Hickman, J. W., Tifrea, D. F. & Harwood, C. S. A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. Proc. Natl. Acad. Sci. U. S. A. 102, 14422–14427 (2005). Yan, J., Monaco, H. & Xavier, J. B. The ultimate guide to bacterial swarming: An experimental model to study the evolution of cooperative behavior. Annu. Rev. Microbiol. 73, 293–312 (2019). Walton, B. et al. A biofilm-tropic Pseudomonas aeruginosa bacteriophage uses the exopolysaccharide Psl as receptor. 2024.08.12.607380 Preprint at https://doi.org/10.1101/2024.08.12.607380 (2024). Maffei, E., Manner, C., Jenal, U. & Harms, A. Complete genome sequence of Pseudomonas aeruginosa phage Knedl. Microbiol. Resour. Announc. 13, e0117423 (2024). Billaud, M. et al. Complementary killing activities of Pbunavirus LS1 and Bruynoghevirus LUZ24 phages on planktonic and sessile Pseudomonas aeruginosa PAO1 derivatives. 2025.04.09.647956 Preprint at https://doi.org/10.1101/2025.04.09.647956 (2025). Magill, D. J. & Skvortsov, T. A. DePolymerase Predictor (DePP): a machine learning tool for the targeted identification of phage depolymerases. BMC Bioinformatics 24, 208 (2023). LeRoux, M. et al. Kin cell lysis is a danger signal that activates antibacterial pathways of Pseudomonas aeruginosa. eLife 4, (2015). Hartmann, R. et al. Quantitative image analysis of microbial communities with BiofilmQ. Nat. Microbiol. 6, 151–156 (2021). Jennings, L. K. et al. Pel is a cationic exopolysaccharide that cross-links extracellular DNA in the Pseudomonas aeruginosa biofilm matrix. Proc. Natl. Acad. Sci. 112, 11353 (2015). Manner, C. et al. A genetic switch controls Pseudomonas aeruginosa surface colonization. Nat. Microbiol. 8, 1520–1533 (2023). Tabor, D. E. et al. Pseudomonas aeruginosa PcrV and Psl, the molecular targets of bispecific antibody MEDI3902, are conserved among diverse global clinical isolates. J. Infect. Dis. 218, 1983–1994 (2018). O’Toole, G. A. Microtiter dish biofilm formation assay. J. Vis. Exp. 2437 (2011) doi: 10.3791/2437 . Baym, M. et al. Inexpensive multiplexed library preparation for megabase-sized genomes. PloS One 10, e0128036 (2015). Millard, A. et al. taxmyPHAGE: Automated taxonomy of dsDNA phage genomes at the genus and species level. 2024.08.09.606593 Preprint at https://doi.org/10.1101/2024.08.09.606593 (2024). Bouras, G. et al. Pharokka: a fast scalable bacteriophage annotation tool. Bioinformatics 39, btac776 (2023). Byrd, M. S. et al. Genetic and biochemical analyses of the Pseudomonas aeruginosa Psl exopolysaccharide reveal overlapping roles for polysaccharide synthesis enzymes in Psl and LPS production. Mol. Microbiol. 73, 622–638 (2009). DiGiandomenico, A. et al. Identification of broadly protective human antibodies to Pseudomonas aeruginosa exopolysaccharide Psl by phenotypic screening. J. Exp. Med. 209, 1273–1287 (2012). Li, H. et al. Epitope mapping of monoclonal antibodies using synthetic oligosaccharidesuncovers novel aspects of immune recognition of the Psl exopolysaccharide of Pseudomonas aeruginosa. Chem. – Eur. J. 19, 17425–17431 (2013). Passos Da Silva, D. et al. The Pseudomonas aeruginosa lectin LecB binds to the exopolysaccharide Psl and stabilizes the biofilm matrix. Nat. Commun. 10, 2183 (2019). Peterson, S. B. et al. Different methods for culturing biofilms in vitro. in Biofilm infections (eds. Bjarnsholt, T., Jensen, P. Ø., Moser, C. & Høiby, N.) 251–266 (Springer, New York, NY, 2011). Additional Declarations No competing interests reported. Supplementary Files PhageBiofilmSupplementalv2.pdf Cite Share Download PDF Status: Published Journal Publication published 18 Nov, 2025 Read the published version in npj Biofilms and Microbiomes → Version 1 posted Editorial decision: Revision requested 22 Jun, 2025 Reviews received at journal 19 Jun, 2025 Reviews received at journal 16 Jun, 2025 Reviewers agreed at journal 02 Jun, 2025 Reviewers agreed at journal 28 May, 2025 Reviewers invited by journal 28 May, 2025 Editor assigned by journal 22 May, 2025 Submission checks completed at journal 13 May, 2025 First submitted to journal 08 May, 2025 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-6623495","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Article","associatedPublications":[],"authors":[{"id":463009843,"identity":"f845ce1c-9146-4d6a-a5da-a3c5bb564bad","order_by":0,"name":"Kristen Amyx-Sherer","email":"","orcid":"","institution":"Washington University in St. Louis","correspondingAuthor":false,"prefix":"","firstName":"Kristen","middleName":"","lastName":"Amyx-Sherer","suffix":""},{"id":463009844,"identity":"886b6b87-6a75-4b2d-945d-9673535d890f","order_by":1,"name":"Leila C. Awasthi","email":"","orcid":"","institution":"Washington University in St. Louis School of Medicine","correspondingAuthor":false,"prefix":"","firstName":"Leila","middleName":"C.","lastName":"Awasthi","suffix":""},{"id":463009845,"identity":"8268dd80-f4df-4989-b34e-9ad5c95176d8","order_by":2,"name":"Amanda Zheng","email":"","orcid":"","institution":"Washington University in St. Louis","correspondingAuthor":false,"prefix":"","firstName":"Amanda","middleName":"","lastName":"Zheng","suffix":""},{"id":463009846,"identity":"fd91e730-89c8-4014-954f-71cbb5e05b26","order_by":3,"name":"Anna Johannesman","email":"","orcid":"","institution":"Washington University in St. Louis School of Medicine","correspondingAuthor":false,"prefix":"","firstName":"Anna","middleName":"","lastName":"Johannesman","suffix":""},{"id":463009847,"identity":"43a531dd-68cc-460f-8734-c1ab9b449af9","order_by":4,"name":"Michele LeRoux","email":"","orcid":"","institution":"Washington University in St. Louis School of Medicine","correspondingAuthor":false,"prefix":"","firstName":"Michele","middleName":"","lastName":"LeRoux","suffix":""},{"id":463009848,"identity":"04730e4f-db50-4c5b-8521-86028bf25aa9","order_by":5,"name":"Courtney Reichhardt","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAA5ElEQVRIie3NsQrCMBCA4ZTAuUSzRhx8hYqgiIivohTcioPgIhRBOBfdC/ogji0Bp+Is1EEpODnoIggOJuAosW4O+SGEg/s4Qmy2Py1yppH6es5RTyX1RE7iUVdvQx5C3gRELsJXMo4em0O1OR9uJ0UMAiA0TpmBiN2gFy+Tc22dZJAWUaor4LVNxGXMlQ5KJxQepD5GirBGxUz4VZOuJiMfA0X4/QthRJO+JtRHqq+AkYgE3HiJ0gtZVq88d7KMFOqttYHwBc2uD5SdsNA/3cJxwHlhdtpfDORD9Ld1m81ms33oBX1pSAhjgSVyAAAAAElFTkSuQmCC","orcid":"","institution":"Washington University in St. Louis","correspondingAuthor":true,"prefix":"","firstName":"Courtney","middleName":"","lastName":"Reichhardt","suffix":""}],"badges":[],"createdAt":"2025-05-08 21:08:16","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-6623495/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-6623495/v1","draftVersion":[],"editorialEvents":[{"content":"https://doi.org/10.1038/s41522-025-00841-4","type":"published","date":"2025-11-18T15:57:32+00:00"}],"editorialNote":"","failedWorkflow":false,"files":[{"id":83752198,"identity":"59870dff-71bb-4a5b-baa3-86359d0ca526","added_by":"auto","created_at":"2025-06-02 07:12:41","extension":"jpg","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":76950,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eTwo Psl-dependent phages were co-isolated with a non-Psl-dependent phage. (A)\u003c/strong\u003e Phage plaques of two phage stocks on PAO1 and PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e as determined via serial dilutions of the indicated phage stock that were spotted on plates made with the indicated bacterial strain. \u003cstrong\u003e(B)\u003c/strong\u003e Serial dilutions of each purified phage were spotted on a double overlay plate prepared with the indicated bacterial strain.\u003cstrong\u003e (C) \u003c/strong\u003eBacterial growth was monitored via absorbance at OD\u003csub\u003e600\u003c/sub\u003e\u003cstrong\u003e \u003c/strong\u003eafter addition of the indicated phage at an MOI of 0.01. \u003cstrong\u003e(D) \u003c/strong\u003eTEM images of the indicated phages.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003c/p\u003e","description":"","filename":"Picture1.jpg","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/3bd4bc3abd8c89280c9eac13.jpg"},{"id":83752204,"identity":"037c4322-f8f6-4e7f-921f-031a282b2d48","added_by":"auto","created_at":"2025-06-02 07:12:41","extension":"jpg","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":1283576,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003ePaStL1 and PaStL2 bind to and degrade Psl. (A) \u003c/strong\u003eBacterial adsorption was measured following ten minutes of incubation at room temperature with the indicated strain for each phage. The percentage of bound phage was calculated using 100% * (total - unbound) / total. \u003cstrong\u003e(B) \u003c/strong\u003ePsl degradation by each phage was determined by anti-Psl immunoblot. \u003cstrong\u003e(C) \u003c/strong\u003eAdherent biomass following incubation with phages was determined using a crystal violet assay. Data represent the means of results from 3 biological replicates, and error bars represent standard deviation.\u003c/p\u003e","description":"","filename":"Picture2.jpg","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/c21b4ee07ecff64d20d1adc9.jpg"},{"id":83752201,"identity":"5ac6cf22-f866-4a79-8c5a-01d1d3a5e918","added_by":"auto","created_at":"2025-06-02 07:12:41","extension":"jpg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":607872,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003ePaStL2 clears mature flow cell biofilms. (A) \u003c/strong\u003eThe impact of phages on the initial attachment of PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+\u003c/sup\u003e bacteria to flow cells was evaluated by confocal microscopy\u003cstrong\u003e. \u003c/strong\u003ePhages were added to mature flow cell biofilms of PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+\u003c/sup\u003e, and \u003cstrong\u003e(B)\u003c/strong\u003e 2 h or \u003cstrong\u003e(C) \u003c/strong\u003e26 h post-phage addition, confocal microscopy images of the biofilms were collected to determine whether the phages cleared the biofilms. Fluorescence due to GFP (bacteria) is shown in green, and HHL-TRITC (Psl) is shown in magenta. Typical images for each time point are shown.\u003cstrong\u003e (D) \u003c/strong\u003eThe volume of bacteria in the flow cells for each time point and phage condition was determined using BiofilmQ, normalized to the volume of bacteria in the no-phage control at 2 h post-phage addition.\u003cstrong\u003e (E) \u003c/strong\u003eThe volume of Psl in the flow cells was similarly determined and normalized to the volume of Psl in the no-phage control at 2 h post-phage addition. Data represent the means of results from 3 biological replicates (3 flow cells with 6 images collected per flow cell), and error bars represent standard deviation.\u003c/p\u003e","description":"","filename":"Picture3.jpg","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/cc709b60e69a4be79d7fdefd.jpg"},{"id":83752831,"identity":"da0e7736-3cdf-4518-b4d0-756e7b7815bc","added_by":"auto","created_at":"2025-06-02 07:28:41","extension":"jpg","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":356739,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003ePaStL2 replicates in a flow tube biofilm. (A)\u003c/strong\u003e Colony forming units (CFUs/mL) were determined for mature flow tube biofilms formed by PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+ \u003c/sup\u003e26 h after the addition of phage. \u003cstrong\u003e(B) \u003c/strong\u003ePlaque forming units (PFUs/mL) within the supernatant of the tube biofilm were determined 26 h after the addition of phage.\u003cstrong\u003e \u003c/strong\u003en.d. = none detected.\u003cstrong\u003e (C) \u003c/strong\u003eBoth CFUs and PFUs were determined at 2, 10, 16, and 26 h after the addition of PaStL2, and the values were compared to those of the no-phage control. \u003cstrong\u003e(D) \u003c/strong\u003eData represent the means of results from 3 biological replicates, and error bars represent standard deviation.\u003c/p\u003e","description":"","filename":"Picture4.jpg","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/2bf9a5d807c78526b8a8bb67.jpg"},{"id":83752359,"identity":"f634e6ca-eec6-40cb-ae64-e5a765e48aa3","added_by":"auto","created_at":"2025-06-02 07:20:41","extension":"jpg","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":94036,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eBacteria resist PaStL1 and PaStL2 by halting Psl production. (A) \u003c/strong\u003eBacterial adherence as measured in a static biofilm assay for indicated control strains, in addition to the bacterial escape mutants. \u003cstrong\u003e(B) \u003c/strong\u003ePsl production was measured with an anti-Psl dot blot for each of the indicated strains. Bars represent the mean of 3 independent replicates with error bars representing standard deviation.\u003c/p\u003e","description":"","filename":"Picture5.jpg","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/471cb3572dab35e91c5fb9be.jpg"},{"id":96650096,"identity":"d6748731-3672-4766-bfb9-b3008fd7fcc1","added_by":"auto","created_at":"2025-11-24 16:07:16","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":3409715,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/9c65db25-42ac-4ae4-a4aa-9f2964c6b7a8.pdf"},{"id":83752203,"identity":"c83902aa-7090-4f4d-9978-49250e4c50d4","added_by":"auto","created_at":"2025-06-02 07:12:41","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":1944794,"visible":true,"origin":"","legend":"","description":"","filename":"PhageBiofilmSupplementalv2.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6623495/v1/291cfce8117cc0b3a543de25.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"Two unrelated Pseudomonas aeruginosa phages require the exopolysaccharide Psl for infection","fulltext":[{"header":"Introduction","content":"\u003cp\u003eA major obstacle to the treatment of bacterial infections is the growth of bacteria in protective aggregates called biofilms, which occur in roughly 80% of human bacterial infections\u003csup\u003e\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e\u003c/sup\u003e. Within biofilms, bacteria are encased in a complex extracellular matrix that may be composed of a range of macromolecules such as exopolysaccharides, extracellular DNA, and proteins. This matrix serves as a barrier that can exclude a wide range of threats, ranging from antibiotics to immune molecules\u003csup\u003e\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e\u003c/sup\u003e. As a result, when pathogenic bacteria form biofilms in humans, these infections often become chronic and untreatable. The opportunistic pathogen \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e, a model for the study of biofilms, frequently causes biofilm-involved infections in a range of settings, including the lungs of people with cystic fibrosis, catheter or ventilator-associated infections, and chronic wounds. The \u003cem\u003eP. aeruginosa\u003c/em\u003e biofilm matrix can vary in composition, with many clinical isolates of \u003cem\u003eP. aeruginosa\u003c/em\u003e from the lungs of people with cystic fibrosis dominated either by alginate or by a mixture of the exopolysaccharides Pel and Psl\u003csup\u003e\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e,\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e\u003c/sup\u003e, as is the case for rugose small colony variants (RSCV). RSCVs are a hyper-biofilm-forming variant, which commonly arise in chronic infections and are especially recalcitrant to treatment\u003csup\u003e\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e,\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eThe biofilm matrix is generally thought to exclude bacteriophages (phages), the viruses that infect bacteria, thus limiting the utility of phage therapy for biofilm-involved infections\u003csup\u003e\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e,\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e\u003c/sup\u003e. However, few studies have examined these interactions at the cellular or molecular level. Phage propagation and characterization are typically performed using a double overlay assay in which bacteria are suspended in a gel-like agar matrix that is normally associated with bacterial motility\u003csup\u003e\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e\u003c/sup\u003e, rather than biofilm formation, and is thus not well suited to biofilm-phage interaction studies. Only a small handful of studies have investigated phages using in vitro biofilm models\u003csup\u003e\u003cspan additionalcitationids=\"CR11 CR12\" citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e\u003c/sup\u003e. Commonly used static biofilm assays only represent one aspect of biofilm development and may miss some of the different effects that phages can exert on biofilms. For example, in a static biofilm assay as opposed to biofilms grown under flow (e.g., flow cell or flow tube biofilms), the phages do not have to contend with flow, which may wash away less adherent phages, but they are faced with relatively high concentrations of extracellular bacterial waste products\u003csup\u003e\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e\u003c/sup\u003e. Further, bacterial physiology, which matters for the efficacy of phage infection, varies depending on the biofilm model that is used in the experiment. For these reasons, it has been suggested that phages be tested against multiple biofilm models in order to gain a better understanding of phage-biofilm interactions\u003csup\u003e\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e\u003c/sup\u003e. Interestingly, many phages are known to produce depolymerase enzymes as part of their tail fibers. These depolymerases have been shown to interact with specific cell-associated extracellular sugars such as lipopolysaccharides (LPS) or the capsule-associated exopolysaccharides (CPS)\u003csup\u003e\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e\u003c/sup\u003e. However, although a few phages have been demonstrated to specifically interact with biofilm matrix exopolysaccharides\u003csup\u003e\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e,1819\u003c/sup\u003e, phages that degrade biofilm matrix components, including exopolysaccharides, have not been reported.\u003c/p\u003e \u003cp\u003eIndeed, given the diversity of both bacterial species and their phages, we have very little knowledge about how most phages interact with bacterial biofilms or whether biofilms do indeed represent a first line of defense against phage infection. A set of studies, albeit with only a single bacterial and phage species, highlight the complexity of biofilm-phage interactions and provide motivation for further studies to probe these interactions. Specifically, in some instances, the spatially segregated, heterogeneous bacterial population present in biofilms supports bacterial co-existence with phages\u003csup\u003e\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e\u003c/sup\u003e. Studies using \u003cem\u003eE. coli\u003c/em\u003e biofilms that contain the functional bacterial amyloid called curli observed that the model T7 phage is trapped by curli, but depending on the location of the phages and the biofilm age, this interaction can both prevent T7 particles from accessing bacteria at the biofilm aggregate interior and also serve as a phage reservoir for newly arriving bacterial cells\u003csup\u003e\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e,\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eHere, we describe the discovery of two unrelated \u003cem\u003eP. aeruginosa\u003c/em\u003e phages \u0026ndash; one podo- and one siphophage \u0026ndash; that both require the exopolysaccharide Psl for infection. We find that both of these Psl-dependent phages require Psl for adsorption to bacteria and degrade Psl to access the bacterial surface. Further, using an in vitro biofilm model, we find that both phages can prevent bacterial adherence to a surface. However, only one of the phages can disrupt a mature biofilm. Selective pressure of these phages leads to the rapid emergence of \u003cem\u003eP. aeruginosa\u003c/em\u003e escape mutants with severely reduced Psl production and an inability to adhere.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003e \u003cb\u003eIsolation of two Psl-dependent\u003c/b\u003e \u003cb\u003eP. aeruginosa\u003c/b\u003e \u003cb\u003ephages\u003c/b\u003e\u003c/p\u003e \u003cp\u003eTo identify phages that may have exopolysaccharide-degrading properties, we set out to test whether any phages in our collection formed haloes on an exopolysaccharide overproducing strain, ∆\u003cem\u003ewspF\u003c/em\u003e wherein the deletion of the negative regulator \u003cem\u003ewspF\u003c/em\u003e leads to the constitutive upregulation of the exopolysaccharides, Pel and Psl\u003csup\u003e\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u003c/sup\u003e. To our surprise, we found that two of our phages (PaStL1M and PaStL2M) displayed an increase in titer on the ∆\u003cem\u003ewspF\u003c/em\u003e strain (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA), with distinct plaquing morphology. Based on the plaquing patterns, we hypothesized that each of our phage stocks actually contained two distinct phages, and that these two phages had different replication capacities on PAO1 compared with ∆\u003cem\u003ewspF\u003c/em\u003e. To determine whether the phage with a preference for ∆\u003cem\u003ewspF\u003c/em\u003e was dependent on a particular exopolysaccharide, we also tested the phage mixtures on strains lacking genes encoding for key early enzymes in the Psl and Pel biosynthetic pathways (\u003cb\u003eTable \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e\u003c/b\u003e), and found that plaques on ∆\u003cem\u003ewspF ∆psl\u003c/em\u003e resembled those on PAO1, suggesting that the second phage in the mixture was Psl-dependent and that only one of the two phages in each mixture was replicating on ∆\u003cem\u003ewspF ∆psl\u003c/em\u003e (\u003cb\u003eFig \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e\u003c/b\u003e). We also tested plaquing on a clinical isolate, CF127, previously shown to overproduce Psl\u003csup\u003e\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e\u003c/sup\u003e, and its derivative CF127 ∆\u003cem\u003epsl\u003c/em\u003e, and noted reduced plaques on CF127, and no plaques on CF127 ∆\u003cem\u003epsl\u003c/em\u003e, suggesting that only the Psl-dependent phage in each case could replicate on CF127. We thus purified the original PaStL1M and PaStL2M phage stocks using both ∆\u003cem\u003ewspF ∆psl\u003c/em\u003e and CF127. This resulted in four isolated phages: two Psl-dependent phages (PaStL1 and PaStL2) that can replicate on ∆\u003cem\u003ewspF\u003c/em\u003e and CF127, but not PAO1, and two Psl-independent phages (PaStL3 and PaStL4), that can replicate on all strains except CF127 and CF127 ∆\u003cem\u003epsl\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eB).\u003c/p\u003e \u003cp\u003eWe were unaware of any previously reported Psl-dependent phages at the time, and thus decided to further characterize PaStL1 and PaStL2. Since both phages were unable to form plaques on PAO1, we wondered how they were present at high titers when titered on an exopolysaccharide overproducing strain such as ∆\u003cem\u003ewspF\u003c/em\u003e or CF127. These observations suggested that they must be able to replicate in planktonic culture on the wild-type PAO1 strain that we had used for all enrichment and isolation steps. Indeed, when we added these phages at a low multiplicity of infection (MOI) of 0.01 to PAO1 and monitored bacterial growth under standard planktonic growth conditions, we observed a typical infection curve with a loss of bacterial density beginning around 2.5 h after phage addition (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC). Planktonically growing PAO1 thus produces sufficient levels of Psl under standard laboratory growth conditions in rich media to sustain infection by these Psl-dependent phages, while Psl is likely down-regulated in double overlay assays, which mimic conditions typically used for bacterial swarming assays (0.6% agar) where production of biofilm exopolysaccharides is reduced\u003csup\u003e\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e\u003c/sup\u003e. Sequencing these phages revealed that PaStL1 and PaStL2 are both new species of distinct genera: PaStL1 is a Bruynogevirus, while PaStL2 is in the Iggyvirus genus of the Queuovirus subfamily. Transmission electron microscopy of these phages further confirmed a podo- and siphophage morphology, consistent with the sequencing data (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eD). We note that these phages are similar to reports of Psl-dependent phages that were made in the course of our investigations\u003csup\u003e\u003cspan additionalcitationids=\"CR23\" citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e\u003c/sup\u003e. Sequencing of PaStL3 and PaStL4 confirmed that the co-isolated, non-Psl-dependent phages are unrelated: PaStL3 is a new species of Yuavirus, while PaStL4 is a strain of Pbunavirus Ab28.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003ePaStL1 and PaStL2 bind and degrade Psl\u003c/h2\u003e \u003cp\u003eWe used an adsorption assay to assess whether phage attachment to bacterial cells was dependent on Psl. Specifically, phages were incubated with bacteria that did not express Psl (i.e., PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003epel\u003c/em\u003e ∆\u003cem\u003epsl\u003c/em\u003e) or expressed high levels of Psl via an arabinose-inducible promoter (i.e., PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003epel\u003c/em\u003e pBAD\u003cem\u003epsl\u003c/em\u003e). The phages PaStL1 and PaStL2, which both required bacterial production of Psl for infectivity in a plaque assay, displayed high levels Psl-dependent adsorption (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). In contrast, the presence of Psl did not impact adsorption of either PaStL3, which did not adsorb well to either strain, or PaStL4, in which 90% of the phage adsorbed to both strains. Together, these results support that PaStL2 and PaStL3 uniquely use Psl as a receptor for binding to the bacterial cells, while the two non-Psl-dependent, co-isolated phages do not require Psl for adsorption.\u003c/p\u003e \u003cp\u003eNext, we determined if the phages were able to depolymerize Psl, as has been observed for other phages that bind and degrade the surface glycan, o-antigen\u003csup\u003e\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e\u003c/sup\u003e. To do so, we incubated purified phage with cell-free supernatant preparations from the Psl- and non-Psl-producing strains, PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e and PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003epsl\u003c/em\u003e ∆\u003cem\u003epel\u003c/em\u003e ∆\u003cem\u003ealgD\u003c/em\u003e (referred to as PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆EPS), respectively. Following 6h of incubation at 37\u0026deg;C, we detected Psl levels using an anti-Psl immunoblot (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB). Incubation with phages PaStL1 and PaStL2 decreased Psl levels. In contrast, incubation with either PaStL3 or PaStL4 did not discernibly change Psl levels relative to the no-phage control. As is expected for depolymerization of Psl by an enzyme, heat inactivation of the phages prior to incubation with the cell-free supernatant abolished the ability of the phages to depolymerize Psl. We surveyed the phage genomes for depolymerases using DePolymerase Predictor (DePP)\u003csup\u003e\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e\u003c/sup\u003e and identified putative depolymerases encoded in the genomes of both PaStL1 and PaStL2. These putative depolymerases had different sequences as well as strikingly different structures, as predicted by AlphaFold (\u003cb\u003eFig S2\u003c/b\u003e). Notably, there were no similar depolymerases predicted that are shared between PaStL1 and PaStL2, suggesting that these two phages have independently evolved mechanisms for binding to and degrading Psl. These results support that both PaStL1 and PaStL2 bind and depolymerize Psl, but they likely do so via different depolymerases.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003ePaStL1 and PaStL2 bind and degrade Psl\u003c/h3\u003e\n\u003cp\u003eGiven the Psl-dependence of the phages, we predicted that they might be effective against biofilms. To test this, we grew biofilms statically in 96-well microtiter dishes for 24 h, removed any unattached biomass, and treated the remaining adherent biomass with the phages. We tested each of the phages individually as well as the two original phage mixtures, PaStL1M (i.e., PaStL1 and PaStL3) and PaStL2M (i.e., PaStL2 and PaStL4). As evidenced by the decrease in adherent biomass relative to the no phage control, both Psl-dependent phages, PaStL1 and PaStL2, were more effective at clearing biofilms formed by both PAO1 and PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e than their co-isolated counterparts (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC). The original phage mixtures showed differing results, with PaStL1M being similarly effective as PaStL1 alone and PaStL2M being less effective than PaStL2 alone. Similar trends were observed for phage treatment of static biofilms formed by PAO1 ∆\u003cem\u003ewspF ∆pel\u003c/em\u003e. However, the Psl-dependent phages had a reduced ability to clear biofilms formed by PAO1 ∆\u003cem\u003ewspF ∆psl\u003c/em\u003e. Interestingly, some of the phage-treated PAO1 ∆\u003cem\u003ewspF ∆psl\u003c/em\u003e biofilms had increased adherent biomass relative to the respective no-phage control. This could be due to the release of extracellular DNA if the phages lysed a subset of the bacteria, or due to the phage-induced bacterial stress triggering an increase in biofilm formation, as has been previously described\u003csup\u003e\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e\u003c/sup\u003e. Overall, this static biofilm assay supports that, in addition to both PaStL1 and PaStL2 requiring Psl for infection, both phages clear biofilms in a Psl-dependent manner. Further, this result shows that when PAO1 is cultured as a biofilm and is making high levels of Psl, we observe similar Psl-dependence of these two phages as for PAO1 ∆\u003cem\u003ewsp\u003c/em\u003eF biofilms.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e\n\u003ch3\u003eOnly PaStL2 clears mature biofilms cultured in flow cells\u003c/h3\u003e\n\u003cp\u003eGiven the role of Psl in \u003cem\u003eP. aeruginosa\u003c/em\u003e surface attachment, we investigated whether the Psl-dependent phages, PaStL1 and PaStL2, could block bacterial surface attachment. To do so, we performed a bacterial attachment assay in microfluidic devices called flow cells. In this experiment, PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e bacteria that constitutively expressed the fluorescent protein GFP (PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+\u003c/sup\u003e) were allowed to attach to the surface of a flow cell for 10 min while the flow was stopped prior to the addition of phage. Purified phages were added and incubated with the bacteria for an additional 30 min, and then flow was resumed. The flow cells were imaged by confocal microscopy, and the numbers of attached bacteria were quantified using BiofilmQ\u003csup\u003e\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e\u003c/sup\u003e. We observed that incubation with both Psl-dependent phages, PaStL1 and PaStL2, decreased bacterial attachment to the flow cell surface (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA).\u003c/p\u003e \u003cp\u003eTo test the impact of the phages on established biofilms, we cultured PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+\u003c/sup\u003e in flow cells for 72 h, and then phages were added and statically incubated with the biofilms for 30 min before resuming flow. In this experiment, in addition to monitoring the bacteria, we also monitored Psl levels by staining with the fluorophore-conjugated Psl-specific lectin, Hippeastrum hybrid lectin (HHL)-TRITC. Biofilms were stained and imaged 2, 10, 16, and 26 h following phage addition (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB-D). Separately cultured biofilms were stained and imaged for each of the time points to ensure that staining with the Psl-specific lectin did not impact the results. Without phage incubation, typical biofilms formed with bacterial aggregates that extend into the flow cell channel approximately 60 mm and have peripherally localized Psl (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB-C). At 2 and 10 h after the addition of phage, the flow cell biofilms appeared similar to one another, regardless of which phage had been added (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB, S3A). By 16 h after the addition of phage, biofilms that had been incubated with PaStL2 began to clear, and by 26 h, biofilms that had been incubated with PaStL2 were entirely cleared (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC, S3B). Biofilms incubated with PaStL1 did not clear, even by 26 h after the addition of phage. We quantified the overall biomass (GFP fluorescence) and Psl levels (HHL-TRITC fluorescence) using BiofilmQ, which allowed us to make two additional observations (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eD-E). First, for biofilms incubated with PaStL2, the Psl levels appeared to decrease more rapidly than the bacteria, suggesting that degradation of Psl precedes overall biofilm clearing. The second observation was that biofilms that were incubated with PaStL1 showed slightly increased levels of bacteria relative to biofilms that were not incubated with phage. Possibly, this slight increase in biofilm bacteria could be due to the lysis of the bacteria by the phage, resulting in increased extracellular DNA levels and, thus, bacteria within the biofilm aggregates.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eAs controls, we performed two additional flow cell experiments. In the first experiment, we tested whether the ∆\u003cem\u003ewspF\u003c/em\u003e mutation impacted the ability of PaStL2 to clear the biofilms. To do so, we cultured flow cell biofilms of PAO1 for 72 h, added PaStL2, and then observed the flow cell biofilms 26 h later. We found that PaStL2 also cleared PAO1 flow cell biofilms, indicating that the ∆\u003cem\u003ewspF\u003c/em\u003e mutation was not necessary for clearing by PaStL2 (\u003cb\u003eFig S4\u003c/b\u003e). This control experiment was important since the ∆\u003cem\u003ewspF\u003c/em\u003e mutation has impacts on the bacteria beyond simply increasing Psl production. In the second control experiment, we tested whether Psl was required for clearing of flow cell biofilms by PaStL2. To do so, we cultured flow cell biofilms of PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003epsl\u003c/em\u003e for 72 h, added PaStL2, and checked for clearing 26 h later. We observed that PaStL2 was unable to clear PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003epsl\u003c/em\u003e biofilms (\u003cb\u003eFig S5\u003c/b\u003e). As previously reported, PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003epsl\u003c/em\u003e flow cell biofilms appear different than PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e flow cell biofilms, even without phage infection, due to the reliance on Pel rather than Psl\u003csup\u003e\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e\u003c/sup\u003e. Together, these findings show that PaStL2 is able to clear both PAO1 and PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e flow cell biofilms, and that bacterial production of Psl is required for it to clear.\u003c/p\u003e\n\u003ch3\u003ePaStL2 propagates in biofilms grown under flow\u003c/h3\u003e\n\u003cp\u003eGiven the unexpected difference in the abilities of PaStL1 and PaStL2 to clear flow cell biofilms, we wondered if PaStL1 had a reduced ability, compared to PaStL2, to propagate in bacteria cultured under flow. To test this hypothesis, we cultured PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+\u003c/sup\u003e biofilms for 72 h in silicone tubing, which was cut to have a similar internal volume to the flow cells used in the prior experiment, and then statically incubated phage with the biofilms for 30 min before resuming flow. The biofilms were harvested 26 h later, and both colony-forming units (CFUs) and plaque-forming units (PFUs) were determined. CFUs were determined using the entire harvested biofilm, and PFUs were determined using the cell-free supernatant of the biofilm. To avoid confounding effects due to possible phage killing during the CFU assay, we determined CFUs indirectly by measuring the GFP fluorescence of the biofilms, which we correlated to CFUs via a standard curve of CFU versus GFP fluorescence that was obtained for PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e GFP\u003csup\u003e+\u003c/sup\u003e bacteria that were cultured in the absence of phage (\u003cb\u003eFig S6\u003c/b\u003e). Incubation of PaStL2, but not PaStL1, resulted in a decrease of the biofilm bacteria (\u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026lt;\u0026thinsp;0.05), although the decrease in this growth system was not as dramatic as was observed in the flow cell experiment (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA). Specifically, the CFUs for biofilms with PaStL1 added were ~\u0026thinsp;5 x 10\u003csup\u003e9\u003c/sup\u003e CFU/mL and with PaStl2 added, only around 1 x 10\u003csup\u003e9\u003c/sup\u003e CFU/mL. A much more dramatic difference was observed for PFUs. For the biofilms treated with PaStL1, we recovered only\u0026thinsp;~\u0026thinsp;5x10\u003csup\u003e4\u003c/sup\u003e PFUs/mL, and for those treated with PaStL2, we recovered 2x10\u003csup\u003e10\u003c/sup\u003e PFUs/mL (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB). To determine the dynamics of PaStL2 phage infection, we also harvested separately cultured biofilms at 2, 10, and 16 h post-phage addition, and similarly determined both CFUs and PFUs (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eC). We observed that PaStL2 titers increased at 16 h, which preceded the decrease in biofilm bacteria CFUs. Given that the titer of the phage stock added to the biofilm was ~\u0026thinsp;10\u003csup\u003e8\u003c/sup\u003e PFUs/mL, with a similar volume as the recovered cell-free supernatant of the biofilm (300 \u0026micro;L vs\u0026thinsp;~\u0026thinsp;250 \u0026micro;L, respectively) these data support that PaStL2 is able to propagate within a biofilm cultured under flow, and PaStL1 is not.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cb\u003eP. aeruginosa\u003c/b\u003e \u003cb\u003eevades PaStL1 and PaStL2 via loss of Psl\u003c/b\u003e\u003c/p\u003e \u003cp\u003eWe noted that both in planktonic growth curves and on top agar, resistant bacteria typically emerged following infection with PaStL1 and PaStL2. We isolated individual colonies that emerged in planktonic growth of PaStL1 as well as within plaques of PaStL2 (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eB) and then retested these for sensitivity to both phages. A subset of these clones was sensitive to phage infection, suggesting a regulatory response to phage infection that reverted upon reculturing. However, many of these clones were persistently resistant, and we found that in all cases, resistant to both PaStL1 and PaStL2. We hypothesized that these cells had likely lost the ability to produce Psl. To test this idea, we performed dot blots and found that all of the escape mutants produced significantly less Psl than either the PAO1 or ∆\u003cem\u003ewspF\u003c/em\u003e strain (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA). We also found that these escape mutants all had reduced ability to adhere to a surface as measured in a static biofilm assay (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eB). We sequenced a subset of these escape mutants and found that ∆\u003cem\u003ewspF-PaStL1EA\u003c/em\u003e, that arose via resistance to PaStL1 had a mutation in \u003cem\u003epslA\u003c/em\u003e that led to a truncation of the protein, while surprisingly ∆\u003cem\u003ewspF-PaStL1B\u003c/em\u003e had only a substitution in a PA1327, a predicted protease (Table\u0026nbsp;\u003cspan refid=\"Tab1\" class=\"InternalRef\"\u003e1\u003c/span\u003e). The escape mutants isolated from double overlays had all acquired a 7-base duplication in the \u003cem\u003ewspR\u003c/em\u003e gene that encodes the regulator controlling the Wsp pathway, and in some cases additional mutations as well (Table\u0026nbsp;\u003cspan refid=\"Tab1\" class=\"InternalRef\"\u003e1\u003c/span\u003e, S2). Taken together, these data demonstrate that both PaStL1 and PaStL2 provide a strong selective pressure that leads to the loss of Psl production in \u003cem\u003eP. aeruginosa\u003c/em\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cdiv class=\"gridtable\"\u003e\u003ctable float=\"Yes\" id=\"Tab1\" border=\"1\"\u003e \u003ccaption language=\"En\"\u003e \u003cdiv class=\"CaptionNumber\"\u003eTable 1\u003c/div\u003e \u003cdiv class=\"CaptionContent\"\u003e \u003cp\u003eMutations present in bacterial escape mutants to PaStL1 and PaStL2\u003c/p\u003e \u003c/div\u003e \u003c/caption\u003e \u003ccolgroup cols=\"6\"\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c1\" colnum=\"1\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c2\" colnum=\"2\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c3\" colnum=\"3\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c4\" colnum=\"4\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c5\" colnum=\"5\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c6\" colnum=\"6\"\u003e\u003c/div\u003e \u003cthead\u003e \u003ctr\u003e \u003cth align=\"left\" colname=\"c1\"\u003e\u0026nbsp;\u003c/th\u003e \u003cth align=\"left\" colname=\"c2\"\u003e\u0026nbsp;\u003c/th\u003e \u003cth align=\"left\" colspan=\"4\" nameend=\"c6\" namest=\"c3\"\u003e \u003cp\u003egenes mutated\u003c/p\u003e \u003c/th\u003e \u003c/tr\u003e \u003c/thead\u003e \u003ctbody\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003edesignation\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ephage\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003elocus\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eMutation(position)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e \u003cp\u003elocus\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e \u003cp\u003eMutation(position)\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003ea\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePaStL1\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003e\u003cem\u003epslA\u003c/em\u003e\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eframeshift\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e\u0026nbsp;\u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e\u0026nbsp;\u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eb\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePaStL1\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003ePA1327\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eR245P\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e\u0026nbsp;\u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e\u0026nbsp;\u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003ec\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePaStL2\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003ewspR\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eframeshift\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e\u0026nbsp;\u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e\u0026nbsp;\u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003ed\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePaStL2\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003ewspR\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eframeshift\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e \u003cp\u003epilY1\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e \u003cp\u003etruncation (553)\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003ee\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePaStL2\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003ewspR\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eframeshift\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e\u0026nbsp;\u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e\u0026nbsp;\u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003ef\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePaStL2\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003ewspR\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eframeshift\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c5\"\u003e\u0026nbsp;\u003c/td\u003e \u003ctd align=\"left\" colname=\"c6\"\u003e\u0026nbsp;\u003c/td\u003e \u003c/tr\u003e \u003c/tbody\u003e \u003c/colgroup\u003e \u003c/table\u003e\u003c/div\u003e \u003c/p\u003e"},{"header":"Discussion","content":"\u003cp\u003eDespite that numerous studies have pursued phage or phage cocktails as therapeutics for recalcitrant bacterial infections, including biofilm infections, there has been a general lack of mechanistic insight into interactions of phage with biofilms. In this study, we discovered two phages that require the \u003cem\u003eP. aeruginosa\u003c/em\u003e biofilm matrix exopolysaccharide Psl for attachment to their bacterial hosts. PaStL1 is related to recently reported Psl-dependent phages Clew-1\u003csup\u003e22\u003c/sup\u003e and LUZ24\u003csup\u003e24\u003c/sup\u003e, while PaStL2 is related to the siphophage Knedl\u003csup\u003e\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e,\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e\u003c/sup\u003e. We find that both phages, using different depolymerase proteins, can both bind to and degrade Psl, and that as a result, they are able to prevent bacteria from establishing biofilms by eliminating the ability of the cells to attach to the surface. We further find that only PaStL2 can disrupt a fully formed biofilm that is cultured under flow, a finding not reported in other related studies that did not assess phage impact on this biofilm model. We further demonstrate that both phages strongly select for non-biofilm-forming bacteria that do not produce Psl and are unable to adhere to surfaces to establish biofilms, even in the absence of phage. These results demonstrate the utility of these phages in therapeutic applications where they could both disrupt biofilms, as well as drive bacteria into a planktonic growth state where they would be more amenable to treatment.\u003c/p\u003e \u003cp\u003eThis study highlights the importance of considering phage isolation methodology when performing environmental phage isolations: phages will only replicate and form plaques on strains that are producing necessary receptors. Both of the Psl-dependent phages that we isolated were originally co-isolated with another non-Psl-dependent phage. This co-isolation was essentially what enabled us to discover these phages, as without the second phage present, our initial plaque assays on PAO1 would have been blank and falsely suggested that there was no phage present in our sample. The proteins and exopolysaccharides present on the cell surface are highly dynamic, as this first line of defense against many threats is critical to bacterial survival. Using conditions in which bacteria produce proteins and exopolysaccharides present during infections is critical for finding phages that are likely to be useful in therapeutic applications.\u003c/p\u003e \u003cp\u003eIn addition to requiring Psl for absorption to the bacterial cell, we found that both PaStL1 and PaStL2, but not PaStL3 or PaStL4, result in the degradation of Psl following their incubation with Psl-containing culture supernatants. We predict that both phages contain depolymerases, as predicted by bioinformatic analyses and analogous to the depolymerases that phages employ to degrade LPS and CPS. Unlike LPS and CPS in many bacterial species, \u003cem\u003eP. aeruginosa\u003c/em\u003e Psl is generally believed to be chemically identical or similar across \u003cem\u003eP. aeruginosa\u003c/em\u003e isolates due to the extremely high conservation of the Psl operon\u003csup\u003e\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e\u003c/sup\u003e. As such, Psl may serve as a better target for phage-based therapies as opposed to LPS or CPS, since bacteria cannot simply modify Psl to escape phage infection. Rather, as we observed in our experiments, bacteria that evolve to escape infection by PaStL1 or PaStL2 lose the ability to produce Psl altogether, which makes them unable to form biofilms, likely rendering them susceptible to more typical antibacterial therapeutics.\u003c/p\u003e \u003cp\u003eThis study also highlights the need to evaluate phage-biofilm interactions in multiple biofilm models since phages can have dramatically different effects on different types of biofilms\u003csup\u003e\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e\u003c/sup\u003e. For example, we observed that both Psl-dependent phages, PaStL1 and PaStL2, cleared biofilms that were cultured statically in microtiter dishes. However, only PaStL2, and not PaStL1, could clear mature biofilms that were grown under flow growth conditions (i.e., flow cell biofilms, tube biofilms). While these are still very much simplified in vitro models, biofilms cultured under flow likely better mimic turbulent or dynamic conditions encountered in some infections (e.g., infections of catheters, heart valve infections, etc), and thus, PaStL2 might be a better therapeutic candidate than PaStL1. The reason(s) for the different abilities of PaStL1 and PaStL2 to clear biofilms cultured under flow could include that PaStL1 does not bind as well as PaStL2 or perhaps the bacterial physiology in these biofilms is less amenable to PaStL1 propagation. Our results point toward both of these options as possibilities, as more PaStL2 absorbed to Psl-producing bacteria compared to PaStL1, and PaStL2 was able to propagate in tube biofilms while PaStL1 did not, although additional experiments are required to say definitely.\u003c/p\u003e \u003cp\u003eOverall, in this study, we discovered two different phages that require Psl for infection and can degrade Psl and clear \u003cem\u003eP. aeruginosa\u003c/em\u003e biofilms. In addition to providing candidates for phage therapy of chronic \u003cem\u003eP. aeruginosa\u003c/em\u003e infections, the predicted Psl depolymerases could serve as anti-biofilm candidates on their own, similar to other hydrolases that degrade matrix exopolysaccharides and help to break apart biofilms, potentially broadening their potential impact beyond the narrow strain profile of these phages. This is an interesting avenue for future research, as currently, nearly all candidate hydrolases for anti-biofilm therapeutics come from either bacteria or fungi, and phage may be an untapped source of novel enzymes to break apart biofilms. Lastly, our study serves as a roadmap for future discovery of additional biofilm-targeting phages.\u003c/p\u003e"},{"header":"Methods","content":" \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003cp\u003e \u003cb\u003eBacterial strain growth.\u003c/b\u003e Planktonic cultures of \u003cem\u003eP. aeruginosa\u003c/em\u003e were routinely grown on Lysogeny broth (LB) or TSB medium at 37\u0026deg;C with constant shaking (225 rpm) unless indicated otherwise. Strains are listed in \u003cb\u003eTable \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e\u003c/b\u003e.\u003c/p\u003e \u003cp\u003e \u003cb\u003ePhage isolation\u003c/b\u003e. Wastewater from a local treatment facility was centrifuged to remove particulates, then filtered with a 0.22 \u0026micro;m filter to remove bacterial cells. The resulting supernatant was mixed with concentrated LB broth for a final 1x concentration of LB and mixed with an overnight culture of PAO1 at 1:100. The suspension was incubated overnight at 37\u0026deg;C with aeration. The following day, the culture was centrifuged to pellet bacteria and filtered, and the supernatant was spotted on double overlay plates prepared with 4 mL of 0.6% LB agar with 200 \u0026micro;L of an overnight PAO1 culture and poured onto a 1.2% LB agar plate. Plates were examined for clearing after 6\u0026ndash;16 h, and for lysates that displayed clearing, the culture was serially diluted to determine an estimate of titer as described below with the spot titer assay, then plated for single plaque isolations using the full plate assay. Individual plaques were picked with a P10 pipet tip and added to 100 \u0026micro;L of a 1:10 dilution of an overnight PAO1 culture in LB and incubated overnight at 37\u0026deg;C in a microcentrifuge tube. The resulting lysate was centrifuged and plaque purifications were performed 3x.\u003c/p\u003e \u003cp\u003e \u003cb\u003ePhage titer assays.\u003c/b\u003e Phage concentration was determined by measuring plaque-forming units / mL (PFU / mL) using either a spot titer or full plate assay. 200 \u0026micro;L of an overnight bacterial culture was mixed with 4 mL of melted 0.6% LB agar (55\u0026deg;C) and poured on top of a 1.2% LB agar plate. Once solidified, 2\u0026ndash;40 \u0026micro;L of a 10x serially diluted phage was spotted and allowed to dry before incubation of plates for 5\u0026ndash;16 h at 37\u0026deg;C. Alternatively, 100 \u0026micro;L of phage was added directly to the top agar. PFUs / mL were calculated by counting the number of plaques in the lowest serial dilution and then multiplying by the dilution factor.\u003c/p\u003e \u003cp\u003e \u003cb\u003ePhage propagation and purification.\u003c/b\u003e \u003cem\u003eP. aeruginosa\u003c/em\u003e phages were produced by back-diluting overnight cultures of PAO1 to an OD\u003csub\u003e600\u003c/sub\u003e of 0.1 in LB, adding an aliquot of phage stock to a final multiplicity of infection (MOI) of 0.1, and then incubating the cultures for 5 h at 37\u0026deg;C with constant shaking (225 rpm). After incubation, bacterial cells were separated from the supernatant by centrifugation (6,000 x g for 20 min) at 4\u0026deg;C. The supernatant was passed through a 0.22 \u0026micro;m filter and used directly for tittering assays and adsorption experiments. For all biofilm experiments, the filtered lysate was further purified via ultracentrifugation through a 35% (w /v) sucrose cushion at 23,5000 x g for 50 min. The supernatant was removed via decanting, and the pellet was suspended in 1 mL of 1x PBS. The sample was centrifuged at 6,000 x g for 20 min and then dialyzed (8\u0026ndash;10 kDa MWCO Float-A-Lyzer G2 Dialysis Device) (MilliporeSigma) against 1x PBS overnight at 4\u0026deg;C. Afterwards, the sample was centrifuged at 17,000 x g for 10 min. The phage was then either used in assays directly or further subjected to size-exclusion chromatography (1x PBS; HiPrep 16/60 sephacryl S-500 HR column) (Cytiva). Phage-containing fractions were concentrated 10-fold using a centrifugal filter (50 kDa MWCO) (MilliporeSigma).\u003c/p\u003e \u003cp\u003e \u003cb\u003eTransmission Electron Microscopy.\u003c/b\u003e For analyses of phages at the ultrastructural level, samples were allowed to absorb onto freshly glow discharged formvar/carbon-coated copper grids (200 mesh, Ted Pella Inc., Redding, CA)) for 10 min. Grids were then washed two times in dH2O and negative stained with 1% aqueous uranyl acetate (Ted Pella Inc.) for 1 min. Excess liquid was gently wicked off and grids were allowed to air dry. Samples were viewed on a JEOL 1200EX transmission electron microscope (JEOL USA, Peabody, MA) equipped with an AMT 8 megapixel digital camera (Advanced Microscopy Techniques, Woburn, MA).\u003c/p\u003e \u003cp\u003e \u003cb\u003eStatic biofilm assay.\u003c/b\u003e Static biofilm formation was assessed by performing a crystal violet assay \u003csup\u003e\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e\u003c/sup\u003e. Biofilms were cultured in Nunc Bacti 96-well microtiter plates (Fisher Scientific) using TSB. 100 mL of mid-log culture was used to inoculate each well of the microtiter plate. The plates were statically incubated for 24 h at 37\u0026deg;C. Non-adherent cells were removed, and wells were washed with 150 mL 1x PBS. Then 100 mL of phage (5x10\u003csup\u003e7\u003c/sup\u003e PFUs/mL, 1x PBS buffer) or buffer alone was added to each well and statically incubated for 4 h at 37\u0026deg;C. Non-adherent cells were removed by pipetting, and the wells were washed three times with 150 mL of ddH\u003csub\u003e2\u003c/sub\u003eO. The 96-well plate was inverted and allowed to dry for 30 min before the addition of 150 mL of 0.1% (w/v) crystal violet (Fisher Scientific). The plate was statically incubated with crystal violet for 15 min. The crystal violet solution was removed, and the wells were washed three times with 150 mL of ddH\u003csub\u003e2\u003c/sub\u003eO. The plate was inverted and allowed to dry overnight. Finally, the crystal violet was solubilized with 200 mL of 95% ethanol, and 100 mL was transferred to a fresh plate and the absorbance at 595 nm was measured (Varioskan LUX multimode microplate reader, Thermo Fisher Scientific).\u003c/p\u003e \u003cp\u003e \u003cb\u003ePhage DNA sequencing.\u003c/b\u003e Phage DNA was extracted using a Norgen Phage DNA Extraction kit and short read Illumina sequencing was either performed by SeqCoast Genomics (Portsmouth, NH, USA) or libraries were prepared using Illumina Tagmentation reagents with a low input protocol\u003csup\u003e\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e\u003c/sup\u003e, and sequenced at the DNA Sequencing and Innovation Lab at the Edison Family CGS\u0026amp;SB Sequencing Center. Reads were de novo assembled in Geneious Prime (version 2025.1.1) and the TaxMyPhage\u003csup\u003e\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e\u003c/sup\u003e tool was used to determine taxonomy. Phage genomic sequencing data is available at PRJNA1260455.\u003c/p\u003e \u003cp\u003e \u003cb\u003eAdsorption assays.\u003c/b\u003e Overnight bacterial cultures were back-diluted into LB media supplemented with 1% (w/v) L-arabinose, and then grown overnight with shaking (225 rpm) at 37\u0026deg;C. A 1-mL aliquot of the culture was centrifuged at 5000 x g for 10 min, the supernatant discarded, and the pellet suspended in 1 mL of 1x PBS. Phage was added to achieve a multiplicity of infection (MOI) of 0.01 and incubated for 15 min at room temperature with occasional inversion. 20 \u0026micro;L was removed for serial dilution; the remaining sample was centrifuged at 8000 x g for 3 min and a 20 \u0026micro;l aliquot of the supernatant was collected for serial dilutions. 20\u0026ndash;40 \u0026micro;L of these dilutions were spotted on double overlay assays to determine relative titers.\u003c/p\u003e \u003cp\u003e \u003cb\u003ePsl degradation assay.\u003c/b\u003e Overnight cultures of PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e and PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e ∆\u003cem\u003eEPS\u003c/em\u003e in TSB were back diluted 100-fold and grown for 16 h at 37\u0026deg;C with shaking (225 rpm). Cultures were normalized to OD\u003csub\u003e600\u003c/sub\u003e of 1.0 and the centrifuged for 2 min at 15060 rpm. The supernatant was then incubated with bacteriophage (10:1 supernatant to phage, 1x10\u003csup\u003e8\u003c/sup\u003e PFUs/mL) at 37\u0026deg;C for 6 h. Immediately after incubation, samples were boiled at 98\u0026deg;C for 5 min and cooled to 12\u0026deg;C. Then samples were treated with proteinase-K (2 mg/mL) (Qiagen) at 60\u0026deg;C for 1 h, 80\u0026deg;C for 30 min, and then cooled down to 12\u0026deg;C. The presence of Psl was assessed by anti-Psl immunoblot.\u003c/p\u003e \u003cp\u003e \u003cb\u003eAnti-Psl Immunoblot.\u003c/b\u003e To assess for the presence of Psl, samples were analyzed by anti-Psl immunoblot as previously described\u003csup\u003e\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e\u003c/sup\u003e. First, samples were treated with proteinase-K (2 mg/mL) (Qiagen) at 60\u0026deg;C for 1 h, 80\u0026deg;C for 30 min, and then cooled down to 12\u0026deg;C. After proteinase K treatment, 5 \u0026micro;L of sample was loaded onto a nitrocellulose membrane (Bio-Rad) (0.2 mm) and allowed to dry for 10 min. The membrane was blocked for 1 h with 5% (w/v) milk, 10 mM Tris pH 7.5, 150 mM NaCl, 0.1% Tween 20 (TBST). Then the membranes were probed with anti-Psl primary antibodies (0.635 mg/mL WapR-001, 0.371 mg/mL WapR-016, 0.867 mg/mL Cam-003) (MedImmune) for 1 h\u003csup\u003e\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e,\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e\u003c/sup\u003e. Membranes were washed three times with TBST. The membranes were probed with horseradish peroxidase (HRP) conjugated goat anti-human antibody (Bio-Rad) (2 mL in 25 mL TBST) for 1 h. Membranes were washed three times with TBST. Detection was performed with Supersignal West Pico PLUS chemiluminescent substrate (Thermo Fisher Scientific) and blots were imaged with AlphaImager HP (Alpha Innotech).\u003c/p\u003e \u003cp\u003e \u003cb\u003eAttachment assay.\u003c/b\u003e Flow cells were inoculated with PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e Tn7 Gm::P(A1/04/03)::GFP bacteria that were grown to mid-log phase in TSB and then diluted to OD\u003csub\u003e600\u003c/sub\u003e of 0.1 in 1% TSB. Bacterial cells were allowed to attach under static conditions in an inverted flow cell for 10 min and then incubated with 300 \u0026micro;L of phage (1x10\u003csup\u003e8\u003c/sup\u003e PFUs/mL) for 30 min. Then, non-attached cells were washed away by initiating media flow (40 mL/h) through the flow cell for 20 min. Flow was reduced to 3 mL/h prior to imaging. Attached cells were visualized on Nikon Eclipse Ni-E confocal laser scanning microscope (20x objective), and six fields of view were captured (Ex 488 nm, Em 499\u0026ndash;551 nm). Images were analyzed with BiofilmQ\u003csup\u003e\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e\u003c/sup\u003e. To extract 3D objects from the imported microscopy data, the biofilm images were segmented. To remove background fluorescence and fluorescence in between cells, the top-hat filter was set to 15 vox. Afterward, the Otsu thresholding method with a sensitivity of 0.25 was set. Cubes were used for the dissection method and were approximately 1 vox in length. The images for the GFP fluorescent channel were segmented with these parameters and then global biofilm properties were calculated to collect the bacterial volume.\u003c/p\u003e \u003cp\u003e \u003cb\u003eFlow cells imaging of biofilms.\u003c/b\u003e Biofilms were cultured in flow cells as previously described\u003csup\u003e\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e\u003c/sup\u003e. The flow cells were made of polysulfone, and the microfluidic chamber was 1.125 inches long, 0.185 inches wide, and 0.162 inches deep. A microscope coverglass (24x60 mm, 0.175 mm thick) (Fisher Scientific) was fixed to the top of the chamber using clear adhesive sealant (Permatex silicone RTV 80050). After allowing the sealant to dry, the flow cells were autoclaved. Sterile flow cells were inoculated with bacterial cultures of PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e Tn7 Gm::P(A1/04/03)::GFP or as specified, that were grown to mid-log phase in TSB and then diluted to OD\u003csub\u003e600\u003c/sub\u003e of 0.01 in 1% TSB. Bacterial cells were allowed to attach under static conditions in an inverted flow cell for 1 h before the start of media flow. Biofilms were grown in 1% TSB for 72 h at 25\u0026deg;C under a constant flow rate (10 mL/h). After 72 h, flow was stopped, and the biofilm was statically incubated with 300 \u0026micro;L of phage in 1x PBS (1x10\u003csup\u003e8\u003c/sup\u003e PFUs/mL) for 30 min prior to resuming media flow. Biofilms were monitored 2, 10, 16, and 26 h after phage incubation. Biofilms were stained with 0.1 mg/mL HHL-TRITC (GlycoMatrix) to visualize Psl. After staining, flow cells were washed with medium at 10 mL/h for 5 min and visualized on Nikon Eclipse Ni-E confocal laser scanning microscope using 20X or 40X objective (GFP: Ex 488 nm, Em 499\u0026ndash;551 nm; TRITC: Ex 561nm, Em 571\u0026ndash;625 nm). Images were analyzed with BiofilmQ \u003csup\u003e\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e\u003c/sup\u003e. To extract 3D objects from the imported microscopy data, the biofilm images were segmented. The images were scaled down 0.25x for pre-processing, and to remove background fluorescence and fluorescence in between cells, the top-hat filter was set to 15 vox. Afterwards, Otsu thresholding method with a sensitivity of 0.25 was set. Cubes were used for the dissection method and were approximately 1 vox in length. The images for each fluorescent channel were segmented with these parameters, and then global biofilm properties were calculated to collect the bacterial and Psl volume.\u003c/p\u003e \u003cp\u003e \u003cb\u003eFlow tubes.\u003c/b\u003e Biofilms were cultured as tube biofilms as previously described with some modifications \u003csup\u003e\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e\u003c/sup\u003e. PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e Tn7 Gm::P(A1/04/03)::GFP bacteria were grown to mid-log phase in TSB and then diluted to OD\u003csub\u003e600\u003c/sub\u003e of 0.01 in 1% TSB. This back-diluted culture then was used to inoculate a sterile 4.5-cm long Nalgene 50 platinum-cured silicon tubing (1/8 in interior diameter and \u0026frac14; in exterior diameter) (Thermo Fisher Scientific). Note that the actual tube length in which the biofilm grew after accounting for the lengths of the connecting luers was 3 cm. Bacterial cells were allowed to attach to the tubes under static conditions for 1 h before the start of media flow. Biofilms were grown in 1% TSB for 72 h at room temperature (approximately 25\u0026deg;C) under a constant flow rate (10 mL/h). After 72 h, flow was stopped, and the biofilm was statically incubated with 300 \u0026micro;L of bacteriophage (1x10\u003csup\u003e8\u003c/sup\u003e PFUs/mL) for 30 min. Then flow was resumed. Biofilms were collected at 2, 10, 16, and 26 h after phage incubation. The biomass inside of the tube was collected by using the handle of a sterile, disposable L-spreader (Fisher Scientific) to push out the biomass into a collection tube. The sample was vortex mixed for 15 s, and then fluorescence (Ex 488 nm, Em 520 nm) was recorded (Varioskan LUX multimode microplate reader) (Thermo Fisher Scientific). The sample was centrifuged for 2 min at 15060 rpm. The supernatant was removed and used for plaque assays.\u003c/p\u003e \u003cp\u003e \u003cb\u003eGFP standard curve for CFU calculation.\u003c/b\u003e To determine the colony forming units (CFUs) in biomass collected from flow tubes incubated with phage, the correlation between GFP fluorescence (due to constitutive expression of GFP by the bacteria) and CFUs was analyzed to create a standard curve. To do so, overnight cultures of PAO1 ∆\u003cem\u003ewspF\u003c/em\u003e Tn7 Gm::P(A1/04/03)::GFP that were grown in TSB were serially diluted, and then the fluorescence (Ex 488 nm, Em 520) was measured in a 96-well black plate (Thermo Fisher Scientific) with Varioskan LUX multimode microplate reader (Thermo Fisher Scientific). In parallel, CFUs of the samples were determined. The CFU values were plotted against GFP fluorescence to obtain a best line fit equation (y\u0026thinsp;=\u0026thinsp;5.2545x10\u003csup\u003e7\u003c/sup\u003e(x) \u0026ndash; 1.3505x10\u003csup\u003e7\u003c/sup\u003e), where y is CFUs, and x is fluorescence.\u003c/p\u003e \u003cp\u003e \u003cb\u003eBacterial escape mutant sequencing.\u003c/b\u003e Resistant mutants were used in titer assays to confirm resistance to PaStL1 and PaStL2. Resistant strains were further cultured and bacterial genomic DNA was extracted using a Qiagen Puregene Kit. Genomic DNA was sent to SeqCoast for short-read (Illumina) sequencing. Reads were mapped to PAO1 using Geneious Prime (version 2025.1.1) and analyzed for mutations using the Geneious Variant Finder tool, with the variant frequency cutoff set to 60% and coverage cutoff of 5. All mutations were compared with sequencing of the ∆\u003cem\u003ewspF\u003c/em\u003e background strain, and only mutations unique to the escape mutants were further analyzed. All sequencing data were deposited to PRJNA1260455.\u003c/p\u003e \u003c/div\u003e"},{"header":"Declarations","content":"\u003ch2\u003eAuthor Contribution\u003c/h2\u003e\u003cp\u003eM.L. and C.R. designed the research; K.A.S., L.C.A., A.Z., and A.J. performed research; K.A.S., M.L., and C.R. analyzed data; and M.L. and C.R. wrote the manuscript. All authors reviewed the manuscript.\u003c/p\u003e\u003ch2\u003eAcknowledgement\u003c/h2\u003e\u003cp\u003eThis work was supported by an Office of the Vice Chancellor for Research Interdisciplinary Seed Grant to M.L. and C.R., and unrestricted funds from Washington University School of Medicine to M.L. The Psl-specific antibodies were obtained by C.R. from AstraZeneca (Dr. DiGiandomenico). We thank Wandy Beatty for assistance and training on TEM imaging. We thank Forrest Walker for feedback and discussion.\u003c/p\u003e\u003ch2\u003eData Availability\u003c/h2\u003e\u003cp\u003eAll sequencing data were deposited to PRJNA1260455. Additional data is provided within the manuscript or supplementary information files.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\u003cli\u003e\u003cspan\u003eFlemming, H.-C. \u0026amp; Wuertz, S. Bacteria and archaea on Earth and their abundance in biofilms. Nat. Rev. Microbiol. 17, 247\u0026ndash;260 (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKarygianni, L., Ren, Z., Koo, H. \u0026amp; Thurnheer, T. Biofilm matrixome: extracellular components in structured microbial communities. Trends Microbiol. 28, 668\u0026ndash;681 (2020).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eColvin, K. M. \u003cem\u003eet al.\u003c/em\u003e The Pel polysaccharide can serve a structural and protective role in the biofilm matrix of \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e. PLOS Pathog. 7, e1001264 (2011).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eColvin, K. M. \u003cem\u003eet al.\u003c/em\u003e The Pel and Psl polysaccharides provide \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e structural redundancy within the biofilm matrix. Environ. Microbiol. 14, 1913\u0026ndash;1928 (2012).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eH\u0026auml;u\u0026szlig;ler, S., T\u0026uuml;mmler, B., Wei\u0026szlig;brodt, H., Rohde, M. \u0026amp; Steinmetz, I. Small-colony variants of \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e in cystic fibrosis. Clin. Infect. Dis. 29, 621\u0026ndash;625 (1999).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eStarkey, M. \u003cem\u003eet al. Pseudomonas aeruginosa\u003c/em\u003e rugose small-colony variants have adaptations that likely promote persistence in the cystic fibrosis lung. J. Bacteriol. 191, 3492\u0026ndash;3503 (2009).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFerriol-Gonz\u0026aacute;lez, C. \u0026amp; Domingo-Calap, P. Phages for biofilm removal. Antibiotics 9, 268\u0026ndash;16 (2020).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDanis-Wlodarczyk, K. M., Wozniak, D. J. \u0026amp; Abedon, S. T. Treating Bacterial infections with bacteriophage-based enzybiotics: In vitro, in vivo and clinical application. Antibiot. Basel Switz. 10, 1497 (2021).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHa, D.-G., Kuchma, S. L. \u0026amp; O\u0026rsquo;Toole, G. A. Plate-based assay for swarming motility in \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e. Methods Mol. Biol. Clifton NJ 1149, 67\u0026ndash;72 (2014).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eVidakovic, L., Singh, P. K., Hartmann, R., Nadell, C. D. \u0026amp; Drescher, K. Dynamic biofilm architecture confers individual and collective mechanisms of viral protection. Nat. Microbiol. 3, 26\u0026ndash;31 (2018).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBond, M. C., Vidakovic, L., Singh, P. K., Drescher, K. \u0026amp; Nadell, C. D. Matrix-trapped viruses can prevent invasion of bacterial biofilms by colonizing cells. eLife 10, e65355 (2021).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSimmons, E. L. \u003cem\u003eet al.\u003c/em\u003e Biofilm structure promotes coexistence of phage-resistant and phage-susceptible bacteria. mSystems 5, 385\u0026ndash;17 (2020).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWinans, J. B., Wucher, B. R. \u0026amp; Nadell, C. D. Multispecies biofilm architecture determines bacterial exposure to phages. PLOS Biol. 20, e3001913 (2022).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRumbaugh, K. P. \u0026amp; Whiteley, M. Towards improved biofilm models. Nat. Rev. Microbiol. 23, 57\u0026ndash;66 (2025).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eAbedon, S. T., Danis-Wlodarczyk, K. M., Wozniak, D. J. \u0026amp; Sullivan, M. B. Improving phage-biofilm in vitro experimentation. Viruses 13, (2021).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKnecht, L. E., Veljkovic, M. \u0026amp; Fieseler, L. Diversity and function of phage encoded depolymerases. Front. Microbiol. Volume 10-2019, (2020).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGuti\u0026eacute;rrez, D. \u003cem\u003eet al.\u003c/em\u003e Role of the pre-neck appendage protein (Dpo7) from phage vB_SepiS-phiIPLA7 as an anti-biofilm agent in \u003cem\u003eStaphylococcal\u003c/em\u003e species. Front. Microbiol. 6, (2015).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWu, Y. \u003cem\u003eet al.\u003c/em\u003e A novel polysaccharide depolymerase encoded by the phage SH-KP152226 confers specific activity against multidrug-resistant \u003cem\u003eKlebsiella pneumoniae\u003c/em\u003e via biofilm degradation. Front. Microbiol. 10, (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCornelissen, A. \u003cem\u003eet al.\u003c/em\u003e Identification of EPS-degrading activity within the tail spikes of the novel \u003cem\u003ePseudomonas putida\u003c/em\u003e phage AF. Virology 434, 251\u0026ndash;256 (2012).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHickman, J. W., Tifrea, D. F. \u0026amp; Harwood, C. S. A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. \u003cem\u003eProc. Natl. Acad. Sci. U. S. A.\u003c/em\u003e 102, 14422\u0026ndash;14427 (2005).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eYan, J., Monaco, H. \u0026amp; Xavier, J. B. The ultimate guide to bacterial swarming: An experimental model to study the evolution of cooperative behavior. Annu. Rev. Microbiol. 73, 293\u0026ndash;312 (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWalton, B. \u003cem\u003eet al.\u003c/em\u003e A biofilm-tropic \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e bacteriophage uses the exopolysaccharide Psl as receptor. 2024.08.12.607380 Preprint at \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://doi.org/10.1101/2024.08.12.607380\u003c/span\u003e\u003cspan address=\"10.1101/2024.08.12.607380\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e (2024).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMaffei, E., Manner, C., Jenal, U. \u0026amp; Harms, A. Complete genome sequence of \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e phage Knedl. Microbiol. Resour. Announc. 13, e0117423 (2024).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBillaud, M. \u003cem\u003eet al.\u003c/em\u003e Complementary killing activities of Pbunavirus LS1 and Bruynoghevirus LUZ24 phages on planktonic and sessile \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e PAO1 derivatives. 2025.04.09.647956 Preprint at \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://doi.org/10.1101/2025.04.09.647956\u003c/span\u003e\u003cspan address=\"10.1101/2025.04.09.647956\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e (2025).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMagill, D. J. \u0026amp; Skvortsov, T. A. DePolymerase Predictor (DePP): a machine learning tool for the targeted identification of phage depolymerases. BMC Bioinformatics 24, 208 (2023).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eLeRoux, M. \u003cem\u003eet al.\u003c/em\u003e Kin cell lysis is a danger signal that activates antibacterial pathways of Pseudomonas aeruginosa. \u003cem\u003eeLife\u003c/em\u003e 4, (2015).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHartmann, R. \u003cem\u003eet al.\u003c/em\u003e Quantitative image analysis of microbial communities with BiofilmQ. Nat. Microbiol. 6, 151\u0026ndash;156 (2021).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eJennings, L. K. \u003cem\u003eet al.\u003c/em\u003e Pel is a cationic exopolysaccharide that cross-links extracellular DNA in the \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e biofilm matrix. \u003cem\u003eProc. Natl. Acad. Sci.\u003c/em\u003e 112, 11353 (2015).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eManner, C. \u003cem\u003eet al.\u003c/em\u003e A genetic switch controls \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e surface colonization. Nat. Microbiol. 8, 1520\u0026ndash;1533 (2023).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTabor, D. E. \u003cem\u003eet al. Pseudomonas aeruginosa\u003c/em\u003e PcrV and Psl, the molecular targets of bispecific antibody MEDI3902, are conserved among diverse global clinical isolates. J. Infect. Dis. 218, 1983\u0026ndash;1994 (2018).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eO\u0026rsquo;Toole, G. A. Microtiter dish biofilm formation assay. J. Vis. Exp. 2437 (2011) doi:\u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003e10.3791/2437\u003c/span\u003e\u003cspan address=\"10.3791/2437\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBaym, M. \u003cem\u003eet al.\u003c/em\u003e Inexpensive multiplexed library preparation for megabase-sized genomes. PloS One 10, e0128036 (2015).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMillard, A. \u003cem\u003eet al.\u003c/em\u003e taxmyPHAGE: Automated taxonomy of dsDNA phage genomes at the genus and species level. 2024.08.09.606593 Preprint at \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://doi.org/10.1101/2024.08.09.606593\u003c/span\u003e\u003cspan address=\"10.1101/2024.08.09.606593\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e (2024).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBouras, G. \u003cem\u003eet al.\u003c/em\u003e Pharokka: a fast scalable bacteriophage annotation tool. Bioinformatics 39, btac776 (2023).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eByrd, M. S. \u003cem\u003eet al.\u003c/em\u003e Genetic and biochemical analyses of the \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e Psl exopolysaccharide reveal overlapping roles for polysaccharide synthesis enzymes in Psl and LPS production. Mol. Microbiol. 73, 622\u0026ndash;638 (2009).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDiGiandomenico, A. \u003cem\u003eet al.\u003c/em\u003e Identification of broadly protective human antibodies to Pseudomonas aeruginosa exopolysaccharide Psl by phenotypic screening. J. Exp. Med. 209, 1273\u0026ndash;1287 (2012).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eLi, H. \u003cem\u003eet al.\u003c/em\u003e Epitope mapping of monoclonal antibodies using synthetic oligosaccharidesuncovers novel aspects of immune recognition of the Psl exopolysaccharide of Pseudomonas aeruginosa. Chem. \u0026ndash; Eur. J. 19, 17425\u0026ndash;17431 (2013).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePassos Da Silva, D. \u003cem\u003eet al.\u003c/em\u003e The \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e lectin LecB binds to the exopolysaccharide Psl and stabilizes the biofilm matrix. Nat. Commun. 10, 2183 (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePeterson, S. B. \u003cem\u003eet al.\u003c/em\u003e Different methods for culturing biofilms in vitro. in \u003cem\u003eBiofilm infections\u003c/em\u003e (eds. Bjarnsholt, T., Jensen, P. \u0026Oslash;., Moser, C. \u0026amp; H\u0026oslash;iby, N.) 251\u0026ndash;266 (Springer, New York, NY, 2011).\u003c/span\u003e\u003c/li\u003e\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"npj-biofilms-and-microbiomes","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"npjbiofilms","sideBox":"Learn more about [npj Biofilms and Microbiomes](http://www.nature.com/npjbiofilms/)","snPcode":"41522","submissionUrl":"https://submission.springernature.com/new-submission/41522/3","title":"npj Biofilms and Microbiomes","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"NPJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"","lastPublishedDoi":"10.21203/rs.3.rs-6623495/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-6623495/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eBacteria commonly protect themselves from a variety of threats by forming biofilms, which are communities of bacteria that are tightly packed together within an extracellular matrix. Biofilm formation has generally been thought to protect bacteria from phage infection. The opportunistic pathogen \u003cem\u003ePseudomonas aeruginosa\u003c/em\u003e produces biofilm matrices that can contain three distinct exopolysaccharides that contribute to the difficulty in treating infected patients. Here, we demonstrate that two diverse \u003cem\u003eP. aeruginosa\u003c/em\u003e phages have evolved to exploit this biofilm matrix to access the bacterial cells by both binding to and degrading a major biofilm exopolysaccharide, Psl. We examined the effect of these phages on biofilms in different \u003cem\u003ein vitro\u003c/em\u003e biofilm models and found that both phages prevent bacterial surface attachment, but only one of the two phages can disrupt a mature biofilm. The phages also rapidly lead to the emergence of bacterial strains that produce reduced amounts of Psl and are unable to adhere to surfaces. These phages may be useful therapeutically by driving bacteria away from producing biofilms and shifting \u003cem\u003eP. aeruginosa\u003c/em\u003e cells into the more treatable planktonic growth state.\u003c/p\u003e","manuscriptTitle":"Two unrelated Pseudomonas aeruginosa phages require the exopolysaccharide Psl for infection","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-06-02 07:12:36","doi":"10.21203/rs.3.rs-6623495/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2025-06-22T05:14:56+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-06-19T15:23:18+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-06-17T00:25:34+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"20076385731388020372789620604899159795","date":"2025-06-02T13:48:38+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"253307639120069634642373035779665315102","date":"2025-05-28T10:06:06+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-05-28T06:00:15+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-05-22T05:30:09+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-05-13T07:10:04+00:00","index":"","fulltext":""},{"type":"submitted","content":"npj Biofilms and Microbiomes","date":"2025-05-08T21:05:36+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"npj-biofilms-and-microbiomes","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"npjbiofilms","sideBox":"Learn more about [npj Biofilms and Microbiomes](http://www.nature.com/npjbiofilms/)","snPcode":"41522","submissionUrl":"https://submission.springernature.com/new-submission/41522/3","title":"npj Biofilms and Microbiomes","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"NPJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"50adbe51-c251-4ca4-bc2b-f2b73e68e625","owner":[],"postedDate":"June 2nd, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[{"id":49163467,"name":"Biological sciences/Microbiology/Bacteria"},{"id":49163468,"name":"Biological sciences/Microbiology/Biofilms"}],"tags":[],"updatedAt":"2025-11-24T16:00:54+00:00","versionOfRecord":{"articleIdentity":"rs-6623495","link":"https://doi.org/10.1038/s41522-025-00841-4","journal":{"identity":"npj-biofilms-and-microbiomes","isVorOnly":false,"title":"npj Biofilms and Microbiomes"},"publishedOn":"2025-11-18 15:57:32","publishedOnDateReadable":"November 18th, 2025"},"versionCreatedAt":"2025-06-02 07:12:36","video":"","vorDoi":"10.1038/s41522-025-00841-4","vorDoiUrl":"https://doi.org/10.1038/s41522-025-00841-4","workflowStages":[]},"version":"v1","identity":"rs-6623495","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-6623495","identity":"rs-6623495","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}