{"paper_id":"27109d3e-7715-41a7-ae41-e0bf2b18f6f5","body_text":"Viral isolation reveals novel and diverse phages infecting natural stream biofilms 1 \n 2 \nWai Hoe Chin1, Martin Boutroux1, Akira Harding1, Davide Demurtas2, Florian Baier1, & Hannes 3 \nPeter1 4 \n1 River Ecosystems Laboratory, Alpine & Polar Environmental Research Centre  (ALPOLE), Swiss 5 \nFederal Technology Institute of Lausanne (EPFL) 6 \n2 Interdisciplinary Centre for Electron Microscopy  (CIME), Swiss Federal Technology Institute of 7 \nLausanne (EPFL) 8 \n 9 \nSummary 10 \nBacteriophages of environmental bacteria remain underrepresented, lending paucity to 11 \nphage-biofilm research beyond clinical and model species domains. Here, we present the 12 \nAlpine Lotic Phage (ALP) collection, curated through an isolation campaign from biofilm-13 \nforming bacteria of alpine streams. We obtained 57 phage isolates, which were dereplicated 14 \nto 28 unique genomes following sequencing. The collection consists of tailed phages infecting 15 \n14 bacterial host species with genomes spanning 37 to 363 kb while exhibiting diverse plaque 16 \nmorphologies, depolymerase activity, and distinct impacts on host biofilm architecture.  17 \nComparative analyses against public viral genomes and a curated planetary -scale contig  18 \ndatabase revealed limited sequence similarity, underscoring the novelty of ALP phages. 19 \nFunctional annotation resolved 9 – 54% of predicted genes  which encoded viral structural 20 \ncomponents, nucleotide metabolism functions, anti-defence mechanisms, and auxiliary genes 21 \nthat facilitate viral  infection and replication. Together, the ALP collection represents a 22 \nfoundational resource for investigating phage evolution and ecology in natural bacterial 23 \ncommunities.  24 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nIntroduction 25 \nBiofilms are ubiquitous microbial assemblages 1, consisting of complex, spatially structured 26 \nmicrobial communities that lend emergent properties such as resource retention and resilience 27 \nagainst environmental stress 2. These sessile communities  have been under a negative 28 \nspotlight in medical and industry settings but they play important roles in maintaining 29 \nfundamental functions across a wide range of ecosystems . For example, benthic biofilms in 30 \nstreams and rivers orchestrate carbon and nutrient cycling at the base of lotic food webs 3,4 31 \nwhile often acting as sentinels for monitoring climate-induced changes to the ecosystem5,6.  32 \n 33 \nAs the dominant microbial lifestyle for millienia7, biofilms also possess a long eco-evolutionary 34 \nhistory with bacteriophages (phages for short), which are viruses that infect bacteria. Virulent 35 \nphages adopt the lytic cycle exclusively where infected cells are lysed to release assembled 36 \nvirions, while temperate phages alternate between active lytic replication and dormant 37 \nlysogeny as the phage genome integrates with the bacterial chromosome 8. Through these 38 \nlifecycles, phages regulate bacterial abundance and foster microbial diversity9, co-evolution, 39 \nand horizontal gene transfer 10,11. In doing so, phage s are also the main source of microbial 40 \nmortality in natural systems, affecting broader biogeochemical processes by liberating organic 41 \nmatter via cell lysis 12,13 while also shifting metabolic demands of infected populations by 42 \nreprogramming their hosts14,15. Phage predation is therefore expected to shape the ecology 43 \nand evolution of biofilm communities 15. However, the complex interactions between phages 44 \nand biofilm-dwelling bacteria in nature remain poorly understood.  45 \n 46 \nMajor strides in unravelling phage-biofilm interactions were largely made within the motives of 47 \nphage therapy research16. Key insights were derived from  single-species biofilms of model 48 \nand clinically-relevant bacterial species such as Escherichia coli, Staphylococcus epidermidis, 49 \nand Vibrio cholerae. These studies have uncovered the protective effects of the biofilm matrix 50 \nagainst phages17, the impact of cell packing density on viral spread across the biofilm 18, and 51 \ncommunal signalling amongst biofilm-dwellers under phage invasion19,20. Few have extended 52 \nthese insights  to mixed-species biofilm 21,22, yet these systems remain confined to model  53 \nbacteria, thereby representing a narrow subset of phage-biofilm interactions. In contrast, field-54 \nbased metagenomics of native biofilms have revealed robust coupling  and host-specificity 55 \nbetween phages and bacteria that extend to global scales23,24, often reporting microbial taxa 56 \nthat are underrepresented in contemporary biofilm research. However, a mechanistic basis 57 \nunderlying these field observations often remain unresolved due to a paucity of matched, 58 \nculturable phage -bacteria pairs necessary for empirical investigation and validation. For 59 \ninstance, leveraging synthetic assemblages to mimic environmental biofilms and investigate 60 \nviral-driven mechanisms in spatially structured heterogeneous communities. Many extensive 61 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nand well -coordinated initiatives   have been undertaken to develop and maintain phage 62 \nrepositories globally25 such as PhagesDB26, BASEL27, and CPL28. However, these collections 63 \nremain largely focused on model and clinically relevant bacterial species with environmental 64 \nstrains remain comparatively underrepresented.  65 \n 66 \nTo advance the field beyond these confines , we present  the Alpine Lotic Phage (ALP) 67 \ncollection: a  resource compris ing of 28 unique phage isolates infecting 1 4 environmental 68 \nbiofilm-forming bacterial species, spanning three taxonomic classes (α- and γ-proteobacteria, 69 \nand Flavobacteriia). By integrating morphological, phenotypic, and genomic characterization, 70 \nthe ALP collection represents a highly novel and diverse culturable fraction of viruses 71 \npreviously hidden within the viral “dark matter” of natural stream biofilms. We envisage this 72 \nresource to provide a foundation for empirical and computational studies aimed at 73 \nunderstanding phage–biofilm dynamics, adaptation, and ecosystem function  within 74 \nenvironmentally relevant context s. This collection also offers a valuable reference for 75 \nadvancing phage gene annotation and exploring the biotechnological potential of alpine 76 \nphages.  77 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nResults 78 \nPhage isolation workflow and overall composition of the ALP collection  79 \nTo facilitate the isolation of phages infecting biofilm-forming bacteria in nature, we selected a 80 \npanel of 3 7 bacterial isolates from a recently established in-house collection derived from 81 \nstream biofilms. This panel spans 24 genera across eight taxonomic classes , representing 82 \nbacterial taxa commonly reported in freshwater biofilm communities  (Table S1). Notably, 83 \ngenera such as Rhodoferax, Flavobacterium, Massilia, and Sphingomonas are well -84 \ndocumented inhabitants of streambed biofilms, with Rhodoferax and Flavobacterium reported 85 \nas abundant and widespread across glacier-fed streams globally29,30. These taxa contribute to 86 \nkey ecosystem processes, including carbon and nutrient cycling 24, and are increasingly 87 \nsusceptible to climate-driven environmental change6,31.  88 \n 89 \nAll bacterial isolates (and their corresponding phages) grew on standard R2A media and 90 \nencompassed a spectrum of phenotypes relevant to biofilms including a range of growth rates, 91 \nsurface motility, aggregative behaviour (i.e. floc -forming), and pigment production. Stream 92 \nwater was collected from the confluence of a groundwater-fed stream (La Vièze) and a glacier-93 \nfed tributary (La Saufla) (Fig.S1), which was subsequently concentrated using tangential flow 94 \nfiltration, followed by 0.45 μm dead-end filtration to remove most prokaryotic and eukaryotic 95 \ncells while retaining the concentrated viral fraction (Fig.1A). Phages were isolated using soft-96 \nagar overlays  of the concentrated water samples where observed plaques were picked , 97 \ndouble plaque -purified, and amplified for downstream characterisation by whole-genome 98 \nsequencing and transmission electron microscopy (TEM) (Fig.1A).  99 \n 100 \nIn total, 57 phage isolates were recovered across 14 bacterial host genera (Fig.2A). Rahnella 101 \ninusitata yielded the highest number of isolates (n = 22), followed by Pseudomonas cyclaminis 102 \n(n = 7) and Pseudomonas haemolytica (n = 6). All isolates were culturable except a single 103 \nSphingomonas phage, which could not be amplified sufficiently for DNA extraction and 104 \ntransmission electron microscopy (TEM). High-quality genomic DNA was obtained from 43 of 105 \nthe 56 culturable isolates, whereas the remaining isolates  - particularly those infecting P. 106 \ncyclaminis - were recalcitrant to DNA extraction. Dereplication based on genome identity 107 \nresulted in a final  collection of 28 unique phage s (Fig. 2A;  Table S2). All 28 assembled 108 \ngenomes were independently classified as viral by both VIBRANT and geNomad. CheckV 109 \nassessment indicated that 26 genomes were high quality and 2 (Rahnella phages B311P5 110 \nand B311P9) were medium quality (Table S3). All genomes were assigned by geNomad to the 111 \nclass Caudoviricetes, with eight further resolved at the family level: six Autographiviridae, one 112 \nCasjensviridae, and one Schitoviridae (Table S3). Across host genera, between 1 and 9 unique 113 \nphages were recovered per bacterial genus. Notably, the 22 initial Rahnella-infecting isolates 114 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\ndereplicated into nine distinct phages, indicating frequent re-isolation of closely related viruses 115 \ntargeting this host (Fig. 2A). 116 \n 117 \n118 \nFig.1 Phage isolation workflow and bioinformatic assembly pipeline of viral genomes. A) ~120 L 119 \nof stream water was collected from an alpine stream and was filtered to remove prokaryotes with a 0.45 120 \nµm filter. The viral fraction retained was concentrated by ~100-fold via tangential flow. The concentrated 121 \nstream water was screened for phages using soft-agar overlay mixed with bacterial host broth culture. 122 \nVisible plaques were picked, double-purified, and amplified in host broth culture to yield pure high-titre 123 \nphage lysates. These lysates were then leveraged to characterise phages via transmission electron 124 \nmicroscopy (TEM) and whole-genome sequencing. B) Summary of DNA sequencing and bioinformatic 125 \npipeline to yield unique, high -quality, and complete phage genomes which constitutes the ALP 126 \ncollection. Hybrid assemblies with long -read Oxford Nanopore and short -read MiSeq sequencing 127 \nplatforms were adopted for selected isolates with marginal DNA quantity due to inherent challenges 128 \nwith DNA extraction. Tools adopted in each step of the pipeline are indicated by bolded grey text, where 129 \ntool versions are referenced in online methods.   130 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nAlpine stream phages exhibit diverse viral and plaque morphologies  131 \nUnique viral isolates were imaged with TEM  to provide a first glance at the morpho types 132 \npertaining to phages infecting stream biofilms . Of the 28 phage isolates, we were able to 133 \nacquire electron micrographs for 18  (Table S2). TEM images revealed a collection of tailed 134 \nviruses comprising 7 siphoviruses, 6 podoviruses, and 5 myoviruses , highlighting the  135 \nheterogeneous viral population infecting sessile bacterial communities in streams (Fig.2B). 136 \nTwo myoviruses, Rahnella phage B311P2 and Comamonas phage B146P1, exhibited large 137 \nvirion dimensions with capsid diameters exceeding 100 nm, consistent with structural 138 \ndimensions expected of jumbophages32.  139 \n 140 \nIn addition to virion structure, plaque morphologies were assessed for the 18 imaged isolates 141 \nto capture basic phage phenotypes under semi-solid growth conditions. Here, plaque sizes 142 \nvaried markedly across the collection, ranging from small punctate plaques to large clearings 143 \nup to 280 μm in diameter (Fig.2B). Plaques exceeding 200 µm in diameter were predominantly 144 \nproduced by podoviruses, in line with their smaller virion dimensions which enhances  145 \ndiffusivity in structured matrices relative to larger myo- and siphoviruses. Recent biophysical 146 \ninsights further suggest that larger plaque sizes may also be associated with shorter viral latent 147 \nperiods which lends higher phage virulence33. Plaques also differed in qualitative features, 148 \nincluding central turbidity and translucent halos extending beyond the plaque margin. Most 149 \nisolates produced clear plaques  reflecting their effective host -killing capacity. H owever, 150 \nMassilia phage B343P1 and Pseudomonas phage B427P1 formed turbid plaques, indicative 151 \nof reduced lytic capacity relative to other ALP isolates.  152 \n 153 \nIn addition, 8 of the 18 imaged phages produced translucent halos surrounding the plaque 154 \nborder, with radii ranging from 30 to 250 µ m (Fig.2B). These halos are consistent with 155 \ndepolymerase activity, whereby phage -encoded enzymes degrade bacterial surface 156 \npolysaccharides including biofilm exopolysaccharides to facilitate access to host receptors34. 157 \nIntriguingly, Janthinobacterium phages B503P1 and B503P2 were dereplicated as near-158 \nidentical genomes yet, the isolates exhibited distinct plaque morphologies  (Fig.2B). Isolate 159 \nB503P2 produced larger plaques with more extensive depolymerase halos compared to 160 \nB503P1, indicating phenotypic divergence despite 99.9% genome identity. Closer inspection 161 \nrevealed a single nucleotide substitution in the tail fibre gene, resulting in a serine -to-162 \nasparagine substitution at position 590 (S590N; Fig. S2). Although the structural 163 \nconsequences of this substitution is not known, it occurs adjacent to a region partially 164 \nhomologous to a pectate lyase domain , which was associated with depolymerase activity in 165 \nAcinetobacter phages35. Collectively, these observations reveal the substantial morphological 166 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nand phenotypic diversity of lotic phages where the latter underscores the various viral infection 167 \ntraits leveraged against structured sessile microbial communities.  168 \n 169 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\n170 \nFig.2 Plaque and virus morphologies of alpine stream phages. A) A total of 28 unique phage 171 \nisolates were derived from 14 alpine bacterial species in our viral screening effort. Coloured legend 172 \nrepresents the bacterial host taxonomic class that were targeted by phages while phage icons represent 173 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nthe number of genomes after dereplication (e.g. 2 unique phages were isolated for Flavobacterium sp.) 174 \nthat were isolated per bacterial species. Question mark designates the unknown number of unique 175 \nphages as dereplication was not possible without successful phage genome extraction (except 176 \nSphingomonas sp. phage which only had one isolate but DNA extraction was unsuccessful). B) Phage 177 \nplaques were assayed using 0.4% soft agar overlay with brightfield images demonstrating a variety of 178 \nplaque sizes and morphologies. Haloed plaques demonstrating potential depolymerase activity were 179 \nmarked with asterisks. Transmission electron microscopy images of 18 phage isolates revealed a 180 \ncollection of tailed phages consisting of myoviruses, podoviruses, and siphoviruses . Only the 181 \nSphingomonas phage isolate was neither successfully sequenced nor imaged due to poor culturability 182 \nin vitro.   183 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nThe ALP collection represents an emerging repository of novel viruses infecting stream 184 \nbiofilm-forming bacteria 185 \nGiven the understudied nature of viruses associated with natural stream biofilms, we assessed 186 \nthe genomic novelty represented by the ALP collection. Briefly, phage genomes were BLAST-187 \nqueried against the NCBI nucleotide database where we retained the top five hits based on 188 \nsequence coverage and nucleotide identity. The majority of isolates exhibited limited similarity 189 \nto previously described viruses, with <20% genome coverage and a mean nucleotide identity 190 \nof ~ 80% (Table S4). Only 12 of the 28 phages showed >20% coverage against publicly 191 \navailable viral genomes, indicating that most ALP phages lack close relatives in existing 192 \nreference databases and public collections  (Table 1). Among these, Pseudomonas phage 193 \nB508P1 demonstrated the strongest match (94% genome coverage and nucleotide identity ) 194 \nwith a cold-active podovirus VSW-3 infecting Pseudomonas fluorescens in wetlands36. This is 195 \nfollowed by Rahnella phage B311P4 with 96% coverage and 87% identity against phage  196 \nKLB24 infecting plant-associated Klebsiella sp. which was isolated from farmland rain 197 \npuddles37. Further adding to the relatedness of ALP isolates with other aquatic phages , 198 \nPseudomonas phage B072P2 displayed 81–86% genome coverage and ~85% nucleotide 199 \nidentity to podoviruses also infecting P. fluorescens, notably the  psychrophilic freshwater 200 \nphage phiGM22-338. The remaining phages infecting Rahnella, Massilia, Brevundimonas, and 201 \nRhodoferax exhibited varying similarity (23–50% coverage, >74% identity) to known phages, 202 \nexcept Rahnella phage B311P3 which was partially found  amongst Serratia bacterial 203 \ngenomes, implying a possible distant prophage relative within this genus. We also note that 204 \nRahnella jumbophage B 311P2 appears distantly related to two other  jumbophages 205 \nCronobacter phage vB_CsaM_GAP32 and Escherichia phage PBECO 4 , suggesting that 206 \nthese large viruses could share core genomic segments. 207 \n 208 \nTable 1: List of top two BLAST with >20% query coverage of ALP phages against NCBI database 209 \nPhage (φ) isolate Closest hits \nQuery \ncoverage \n% \nidentity Accession # \nBrevundimonas φ B307P1  \nCaudoviricetes sp. (MAG)a 38% 80.94% OR222513.1 \nCaulobacter phage Seuss 14% 77.03% NC_047757.1 \nBrevundimonas φ B307P2 \nCaudoviricetes sp. (MAG)a 36% 81.23% OR222513.1 \nCaulobacter phage Seuss 14% 77.02% NC_047757.1 \nMassilia φ B343P2 \nCaudoviricetes sp. (partial MAG)a 68% 85.50% BK020520.1 \nCaudoviricetes sp. (MAG)a 1% 75.35% OR222459.1 \nPseudomonas φ B072P2 \nPseudomonas phage phiGM22-3 86% 84.73% MW627366.1 \nPseudomonas phage phi2 81% 85.57% NC_013638.1 \nPseudomonas φ B508P1 \nPseudomonas phage VSW-3 94% 94.39% NC_041885.1 \nProvidencia phage PSTNGR1 2% 74.38% MW145136.1 \nPseudomonas φ B529P1 Pseudomonas phage phiPsa300 50% 82.01% NC_073687.1 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nPseudomonas phage phiPsa347 49% 81.99% NC_073685.1 \nRahnella φ B311P2 \nCronobacter phage \nvB_CsaM_GAP32 33% 72.00% JN882285 \nEscherichia phage PBECO 4 15% 68.00% KC295538 \nRahnella φ B311P3 \nSerratia fonticolab 23% 79.02% CP013913.1 \nSerratia sp. JSRIV004b 22% 79.07% CP074140.1 \nRahnella φ B311P4 \nBacteriophage sp. (partial MAG)a 96% 87.22% OP072654.1 \nKlebsiella phage vB_KM5a1-KLB24 96% 87.13% PP554338.1 \nRahnella φ B311P5 \nEnterobacter phage EC151 27% 74.50% MW464860.1 \nKlebsiella phage vB_Ko_K4PH164 8% 73.92% OY979482.1 \nRahnella φ B311P7 \nEnterobacter phage EC151 36% 76.76% MW464860.1 \nKlebsiella phage vB_Ko_K4PH164 22% 75.07% OY979482.1 \nRhodoferax φ B534P2 \nVariovorax phage Gard 38% 80.25% PV920657.1 \nVariovorax phage VAC_51 32% 79.67% OX359471.1 \n 210 \na MAG denotes metagenome-assembled genomes that are either complete or partial. 211 \nb Closest hits with bacterial origins i.e. non-viral hits.   212 \n 213 \nTo further contextualise novelty at a global scale, the ALP genomes were queried against 214 \nLOGAN, which is a planetary -scale contig database curated from public sequence read 215 \narchives39. Using a permissive k -mer coverage threshold of 0.25, 16 out of 28 ALP phages 216 \nreturned positive matches (Fig.3A). These matches were geographically widespread, 217 \npredominantly originating from the European continent but also spanning Asia  and North 218 \nAmerica (Fig.3A, Table S5). The matches were also associated with diverse environment al 219 \nsources ranging from  freshwater ecosystems including rivers, glaciers, groundwater and  220 \nalpine streams to other environments such as soil, plants, and terrestrial animals, followed by 221 \nanthropogenic food and wastewater sources  (Figs.3A and B) . Overall, 7 phage isolates 222 \nmatched with contigs from a single source while the remaining 9 isolates were associated with 223 \ncontigs from multiple environments of up to 6 sources (Fig.3B). Despite this broad distribution, 224 \nindividual matches generally exhibited low similarity, with a median k-mer coverage score of 225 \n0.36 (range: 0.25 –0.85) against LOGAN contigs  (Fig.3A). Collectively, the sparse 226 \nrepresentation of phages in both reference and global metagenomic data bases underscores 227 \nthe substantial novelty behind viruses associated with stream biofilms.  228 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\n 229 \nFig.3 Matches of ALP genome sequences against planetary -wide contig database. A) Total 230 \nnumber of phage isolates with matches to contigs within the LOGAN database and a histogram 231 \ndepicting the spread of k-mer coverage scores of all positive hits to the database. The map represents 232 \nthe geographical distribution of these hits while the colours represent the source of the contigs identified 233 \nwithin the database. Asterisk denote matches to “aquatic (unknown)” sources do not possess any 234 \ngeographical metadata and are thus, not located on the map. B) Cumulative k -mer coverage of 235 \nsequence matches between a 1kb sequence of the ALP isolates to the LOGAN contigs, where colours 236 \nalso represent the contig source stipulated within the database.   237 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nAuxiliary coding sequences in phages facilitate host takeover for infection and 238 \nenvironmental resilience 239 \nAcross the 28 phage isolates, genome sizes spanned over an order of magnitude from 37 to 240 \n363 kb  (Fig.4A). Two isolates, Comamonas phage B146P1 (206 kb) and Rahnella phage 241 \nB311P2 (363 kb), exceeded 200 kb and were hence, classified as jumbophages consistent 242 \nwith their large virion dimensions (Fig.2B). Differences in phage genome size were matched 243 \nby expansions in capsid size,  with phages below 83 kb exhibiting relatively similar capsid  244 \ndiameters (69 nm on average) . Meanwhile, capsids enlarged progressively as genome size 245 \nexceed 111 kb, reaching 133 nm in the Rahnella jumbophage B311P2 with a 363 kb genome. 246 \nAllometric analysis also revealed proportional scaling between genome size and estimated 247 \ncapsid volume (allometric coefficient α = 0. 91), indicating that DNA packing density was 248 \nrelatively conserved across the ALP collection  following structural principles of icosahedral 249 \ndsDNA viral capsids40 (Fig.4B, Table S6). Furthermore, coverage patterns from read mapping 250 \nidentified DNA packaging mechanisms41 for 14 phage isolates: 1 utilized a 3’ cos terminus, 4 251 \nharboured pac sites, and 9 employed direct terminal repeats (DTR) ( Table S7), where t he 252 \nRahnella jumbophage B311P2 possessed a remarkably long DTR of  19,824 bp. Together, 253 \nthese features illustrate the genomic architectural diversity, packaging constraints, and 254 \nmechanisms among isolates of the ALP collection.  255 \n 256 \nGenome annotation resolved putative functions for 9 – 54% of predicted coding sequences 257 \n(CDS), with gene functions  predominantly associated  with virion structure , nucleotide 258 \nmetabolism, and host lysis (Fig.4A, Table S6). Only one isolate – Rahnella phage B311P3 – 259 \nwas identified as a temperate phage, encoding both integrase and excisionase while also 260 \npossessing genes consistent with lysogenic capacity such as CII- and CIII-like transcriptional 261 \nregulators and Ren-like superinfection exclusion factor. In addition, Rahnella phage B311P3 262 \nwas also detected as an integrated prophage within the Rahnella host genome. Meanwhile, 263 \nlarger phages generally encoded substantial tRNAs relative to the entire collection, comprising 264 \n5 – 11% of total CDS (Fig.4A). This pattern aligns with previous observations that phages with 265 \nlarger genomes  leverage self-proprietary tRNAs to sustain viral r eplication as host 266 \ntranslational capacity becomes limiting during late-stage infection42. 267 \n 268 \nBeyond core functions, morons and auxiliary metabolic genes (AMGs) were predicted in 20 of 269 \n28 phage genomes, ranging from 1 – 8 genes per genome  except for the Rahnella 270 \njumbophage, which encoded 33 auxiliary genes (Figs.4A and C, Table S8). The majority of 271 \nthese genes however, corresponded to membrane proteins and biocatalytic enzymes of broad 272 \nunclassified functions. Among fully annotated genes, functions associated with host takeover 273 \nand anti-defence were prevalent, such as the host mRNA-modulating HicA-like toxin-antitoxin 274 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nsystem43, NAD synthesis and scavenging factors, followed by anti-restriction and anti-CRISPR 275 \nsystems. Several phages also encoded enzymes implicated in phage DNA modification, such 276 \nas 2 -oxoglutarate/Fe(II)-dependent oxygenases 44 and phosphoheptose isomerases 45, 277 \nconsistent with strategies to evade intracellular host restriction and CRISPR -Cas immunity 278 \n(Fig.4C). 279 \n  280 \nIn contrast, relatively few AMGs were predicted to directly enhance host environmental 281 \nresilience. These included tellurite resistance genes encoded by the Rahnella jumbophage 282 \nand Pseudomonas phage B529P1, as well as genes  involved in antioxidative protection and 283 \ncellular energetics  in other isolates  (e.g. gluthathionylspermidine synthase and porphyrin 284 \nbiosynthesis; Fig.4C). These AMGs are consistent with promoting survival and maintaining 285 \ncellular metabolism in biofilm-associated microenvironments characterised by accumulation 286 \nof trace metal ions46,47, steep redox gradients 48, and chronic oxidative stress49. Notably, 4 287 \nRahnella phage isolates encoded enzymes within queuosine biosynthesis pathway alongside 288 \na queuine tRNA -ribosyltransferase, suggesting roles in phage DNA modification for host 289 \nimmune evasion 50 and maintaining translational efficiency  to sustain bacterial cells  in 290 \nbiofilms51. Despite expectations that temperate phages preferentially retain host -beneficial 291 \ngenes52, only a single AMG was detected in the temperate Rahnella phage B311P3, encoding 292 \na sporulation factor of unclear relevance to the non-sporulating Rahnella genus. Collectively, 293 \nthese findings reveal that stream-biofilm phages encode diverse auxiliary gene functions 294 \nprimarily associated with host takeover and antiviral defence evasion, with a smaller subset 295 \nlinked to host cellular and biofilm stress tolerance. The large unannotated portion of ALP 296 \ngenomes also represents a resource for exploring  additional phage-encoded adaptations or 297 \neven, gene products of potential biotechnological relevance.  298 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\n 299 \nFig.4 Genomic features of ALP isolates and profile of auxiliary coding sequences. A) Genome 300 \nanalysis of sequenced phage isolates where the red and green line plots correspond to genome size 301 \n(kb) and capsid diameter (nm) respectively, which are plotted on the top x-axis, while the coloured bar 302 \nchart indicates the proportion of genes predicted , corresponding to the bottom x-axis. B) Log -303 \ntransformed plot of capsid volume (approximated as spherical volume:  [p ÷ 6] ´ D3, where D is the 304 \ncapsid diameter) against genome length. The regression line is shown in blue with the accompanying 305 \nallometric exponent (a i.e. gradient of the log-transformed regression), R-squared value, and p-value. 306 \nC) Frequency table of morons and AMGs predicted in 20 phage isolates using nucleotide-based 307 \n(pharokka) and protein structure-based (phold) algorithms. Gene functions are broadly categorised into 308 \n6 groups. Each cell within the table is separated to four quadrants which are colour -coded based on 309 \nphold annotation confidence i.e. (blue = high, green = medium, and low = orange), while the bottom -310 \nright red quadrant indicates genes predicted by the nucleotide-based algorithm, pharokka. The number 311 \nwithin each quadrant shows the frequency of gene functions predicted in accordance with the type of 312 \nalgorithm employed and confidence level for phold annotations.   313 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nPhage infections in microfluidics reveal host -specific architectural responses of 314 \nenvironmental isolate biofilms.  315 \nMicrofluidic platforms have emerged as powerful systems to resolve phage-biofilm interactions 316 \nwith high spatial and temporal precision , lending crucial insights into mechanistic responses 317 \nof biofilms under phage infection 17,20,21,53 and viral propagation strategies in structured 318 \ncommunities18,54. Yet, their application to environmentally derived phage -host systems 319 \nremains limited. To demonstrate the utility of the ALP collection, we performed time -course 320 \ninfections of environmental isolate biofilms in microfluidic devices. We selected three 321 \nrepresentative bacterial isolates exhibiting high, moderate, and low biofilm-forming capacities 322 \n(Massilia sp., R. inusitata, and P. fluorescens, respectively), based on crystal violet staining of 323 \nstatic overnight biofilms (Fig.5A). These isolates were seeded into microfluidic channels and 324 \nallowed to establish for 24 h ours under continuous flow prior to infection with their 325 \ncorresponding phages: Massilia phage B343P1, Rahnella jumbophage B311P2, and 326 \nPseudomonas phage B508P1 (Fig.5B).  327 \n 328 \nOver the course of 96 hours following infection, biofilms were monitored using brightfield 329 \nmicroscopy. These environmental isolates are non -fluorescent and display heterogeneous 330 \ngrowth patterns under fluid flow (Fig.5B), highlighting the exploratory potential  of extending 331 \nmicrofluidic phage-biofilm assays beyond fluorescently tagged model systems. Here, time -332 \ncourse imaging revealed distinct architectural characteristics across the three biofilm species. 333 \nMassilia sp., a floc -forming isolate, developed heterogeneous biofilms with microcolonies 334 \nreaching up to ~200 µm wide. Meanwhile, environmental P. fluorescens, despite limited biofilm 335 \nformation in static assays, formed dense communities with patterned fronts that aligned with 336 \nthe direction of flow. In contrast, R. inusitata formed comparatively homogeneous and low-337 \ndensity lawn-like biofilms spanning the channel surface (Fig.5B). Notably, Massilia sp. and P. 338 \nfluorescens biofilms exhibited quasi -cyclical development even in the absence of phage, 339 \ncharacterized by initial disruption of heterogeneous biofilms under flow between 48 – 72 hours 340 \nfollowed by structured re-growth. 341 \n 342 \nPhage predation reduced biofilm surface area across all systems by 48 h post -infection and 343 \naltered the structure of Massilia sp. and P. fluorescens communities. Despite continuous viral 344 \npressure, Massilia sp. and P. fluorescens biofilms recovered but adopted architectures distinct 345 \nfrom both their early developmental states and uninfected controls. Massilia sp. biofilms 346 \nformed sparser  microcolonies, whereas the structured fronts of P. fluorescens appeared 347 \ndistorted despite unchanged flow conditions (Fig.5B, 96 hpi). These changes imply potential 348 \ntrade-offs between biofilm formation and phage resistance in these two isolates. In contrast, 349 \nR. inusitata biofilms failed to recover under sustained jumbophage pressure, and surviving 350 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\ncells frequently exhibited filamentation  similar to infections caused by E. coli  jumbophage 351 \nSharanji55. Together, these findings demonstrate that phages can actively reshape biofilm 352 \narchitecture and induce host-specific responses that dictate biofilm recovery under sustained 353 \nviral pressure. By extending microfluidic infection assays beyond model systems, these 354 \nexperiments illustrate  the tractability of environmentally derived phage -host pairs within 355 \ncontemporary biofilm platforms and provides a framework for investigating virus -biofilm 356 \ndynamics in structured natural communities. 357 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\n 358 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nFig.5 Biofilm forming capacity of environmental isolates and time -course phage infection in 359 \nmicrofluidics. A) Crystal violet absorbance A590 values for static biofilms grown in 96-well plates for 24 360 \nhours with three replicates (n = 3) per bacterial species. Species highlighted in bold were adopted for 361 \nmicrofluidic experiments as representative for their varying capacities to form  biofilm (i.e. high, 362 \nmoderate, and low). Error bars represent the SEM of triplicate A590 values. B) Brightfield images of 24 363 \nh-old biofilms established in microfluidic devices which were subsequently infected with 106 phages/mL 364 \nunder continuous flow at 0.1 µL/min and monitored up to 96 hours post-infection. Massilia sp. biofilms 365 \nwere infected with phage B343P1, R. inusitata with jumbophage B311P2. and P. fluorescens with phage 366 \nB508P1. Arrowheads in Massilia sp. biofilm highlights examples of microcolonies observed over the 367 \nexperimental duration while arrowheads in R. inusitata community indicate cell filamentation under 368 \nsustained jumbophage pressure.   369 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nDiscussion 370 \nBiofilms represent the dominant microbial lifestyle on Earth, forming highly structured and 371 \nubiquitous communities that underpin essential ecosystem processes1. As obligate bacterial 372 \nparasites, phages have coexisted with biofilms over evolutionary timescales , and 373 \nmetagenomic surveys consistently reveal strong coupling between viral and bacterial 374 \ncommunity composition across diverse environments8,23,24,56. Despite the central role of 375 \nphages in driving microbial diversity, carbon cycling, and horizontal gene transfer57, our 376 \nunderstanding of phage-biofilm interactions remains disproportionately informed by model and 377 \napplied phage-host systems. Consequently, the  diversity, biology, and ecological roles of 378 \nphages in natura l multispecies biofilms remain poorly understood. The ALP collection  379 \naddresses these gaps by  providing a culturable, full -genome resource of novel , 380 \nenvironmentally derived phages that extend our current understanding of viral diversity and  381 \ntraits beyond model and clinical biofilms.  382 \n 383 \nMountain stream  ecosystems are structured and sustained by benthic biofilms that drive 384 \nprimary production, respiration, and nutrient cycling3. Through these processes, they support 385 \nthe functioning o f river networks that supply freshwater to meet the demands of millions 386 \nworldwide58. Moreover, stream biofilms are also sensitive indicators of environmental change 387 \nand are increasingly affected by climate warming6,29,31. Therefore, there is growing urgency to 388 \nunderstand how biofilms and their associated viral communities respond with accelerating 389 \nclimatic pressures. By targeting native stream bacteria from benthic biofilms , the ALP 390 \ncollection captures a fraction of viral diversity that is largely absent from existing phage and 391 \ngenome repositories. Although derived from alpine stream systems, its ecological relevance 392 \nextends broadly as benthic biofilms are pervasive across aquatic ecosystems. 393 \n 394 \nThe limited representation of freshwater biofilm -associated phages in reference databases 395 \nand sequence archives underscores the novelty observed across the ALP collection. Only 5 396 \nof 28 isolates exhibited >50% genome coverage with previously described freshwater phages 397 \n(Table 1), while queries against the curated LOGAN database revealed sparse and low -398 \ncoverage matches across geographically diverse environments (Fig.3). The latter also 399 \nindicates that potential relatives – or perhaps the ALP isolates themselves – are rarely 400 \nrecovered as near -complete genomes in metagenomic datasets , reflecting  persistent 401 \ntechnical challenges in viral  metagenomic assemblies 59,60. In contrast , DNA from purified 402 \nisolates are more amenable to yield complete genomes thereby, providing greater confidence 403 \nin identifying features such as terminal repeats that may inform phage DNA packaging 404 \nstrategies41. Moreover, the ALP isolates were sequenced and characterised with minimal 405 \nlaboratory passage, providing a complementary perspective of genomic content that reflect 406 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\ntheir recent life history in nature and minimising genetic drift caused by extended propagation 407 \nunder laboratory conditions 61. In these contexts, the ALP collection not only expands 408 \ntaxonomic representation but also offers high-resolution genomes for facilitating bioinformatic 409 \nanalyses and ecological interpretation of biofilm-associated viruses in freshwaters.  410 \n 411 \nConsistent with the global predominance of tailed dsDNA viruses 24,62, the ALP collection 412 \ncomprises myo-, sipho-, and podovirus morphotypes supported by TEM imaging and genome 413 \nannotation (Figs.2B and 4A).  Capsid-genome scaling of the ALP isolates also followed 414 \nallometric relationships consistent with icosahedral dsDNA phage s, where scaling models 415 \nreported proportional expansion in capsid volume  with increasing viral genome sizes 40,63 416 \n(Fig.4B). This ensures that DNA packing density is maintained to preserve the internal capsid 417 \npressure required for genome ejection 40,63, thereby reflecting the conserved infection 418 \nmechanobiology amongst ALP isolates. Consequently, the ALP collection also expands the 419 \nempirical basis of these capsid -genome scaling models 63 by contributing physical 420 \nmeasurements of virions derived from these novel isolates. Importantly, the conserved capsid-421 \ngenome allometry also underscores the constraints of phages operating within environmental 422 \nbiofilms. For instance, larger capsids are expected to reduce phage diffusivity in biofilm 423 \nmatrices64, yet simultaneously accommodate larger genomes that may encode beneficial 424 \nauxiliary genes, such as those enhancing host metabolism to support viral replication. While 425 \nnot directly tested here, this hints that biofilm-associated phages likely occupy a constrained 426 \nmultidimensional trait space, maintaining internal capsid pressure  while balancing  gene 427 \ncontent, and diffusion through biofilms. Whether matrix-degrading depolymerases can partially 428 \nalleviate diffusional constraints also remains unclear. This framework  can be extended to 429 \nstream environments where hydraulic forces (e.g. turbulent flow and shear) and oligotrophy 430 \nmay further shape viral structural stability and auxiliary gene content. 431 \n 432 \nThe compact and densely packed nature of phage genomes impose s strong constraints on 433 \ngene content, which limits retention of genes beyond core replicatory functions65. Accordingly, 434 \nmorons and AMGs were sparse among ALP isolates and were largely associated with host 435 \ntakeover, metabolic reprogramming, and suppression of antiviral defences (Figs.4A and 4C). 436 \nThis pattern aligns with the predominance of virulent phages in the collection, prioritising 437 \ngenes that optimise the lytic cycle  which typically occur at shorter timescales  than lysogeny 438 \n(Fig.4C). In contrast, only a minor subset of AMGs was  putatively linked to longer-term host 439 \nenvironmental resilience, notably metalloid resistance and oxidative stress mitigation. These 440 \nfunctions are congruent with overcoming stream biofilm-associated stressors, where EPS 441 \nmatrices are known to  concentrate heavy metals and resident cells experienc ing chronic 442 \noxidative stress46,48. Within the context of virulent phages,  these AMGs may enhance phage 443 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nproduction indirectly by temporarily sustaining host physiology under environmental stress 444 \nrather than promoting long-term host persistence. Nonetheless, the true functional diversity of 445 \nthe ALP collection is likely underestimated given the overwhelming proportion of unknown 446 \ngene functions (Fig.4A), despite recent major strides with protein structure-based annotations 447 \nin revealing the functions of >50% of genes within an average phage genome66.  448 \n 449 \nBeyond genomic features, the presence of multiple infection phenotypes especially amongst 450 \nphages infecting the same species (Fig.2B; Rhodoferax and Massilia phages), coupled with 451 \nhost-specific structural responses to phage infection (Fig.5B) suggests that viral invasion of 452 \nbiofilms is unlikely dictated by a dominant infection strategy. Instead, competing viral traits and 453 \nunique host-specific adaptations in biofilms may stabilise microbial and viral diversity 56 by 454 \npromoting localised negative frequency -dependent “kill-the-winner” dynamics and limiting 455 \nselective sweeps across the structured community. These effects are also likely amplified 456 \nwithin natural multispecies biofilms as several taxa are competing within a spatially 457 \nheterogenous niche, generating a mosaic of localised virus -microbe interactions where viral 458 \nsuppression is tempered and bacterial adaptation is dependent on the microenvironment 67 459 \nand local phage pressure. In this context, environmental phages function not only as agents 460 \nof mortality but as active architects of biofilm structure (Fig.5B) and community composition, 461 \npromoting turnover and preserving functional resilience of environmental biofilm by 462 \nmaintaining microbial community diversity.  463 \n 464 \nIn summary, the ALP collection captures substantial genotypic and phenotypic diversity among 465 \nbiofilm-associated phages in streams. This curated set of 28 unique phage isolates expands 466 \nupon the viral “dark matter” that remains poorly represented in current culture collections and 467 \ndatabases, particularly among underexplored natural environments where biofilms are 468 \npervasive. Crucially, the collection provides a foundation for necessary empirical 469 \ninvestigations into virus-microbe interactions across scales. For example, integrating the ALP 470 \ncollection with synthetic communit ies68,69 and microfluidic platform s18,21 offers a tractable 471 \nframework to finely dissect viral -driven mechanisms in heterogeneous biofilms. These 472 \nexperiments are also extendable to mesoscales such as flume models 70 or interconnected 473 \nfluidic systems of biofilm networks 71 to explore and validate emergent ecological properties 474 \nand evolutionary dynamics as phages propagate across broader biofilm landscapes. Beyond 475 \nexperimental applications, the viral genomes offer valuable references for advancing virome 476 \nresearch and computational tools, with the prospect of uncovering environmental gene 477 \nfunctions that may provide the biotechnological innovation required to navigate  our rapidly 478 \nchanging ecosystems in a warming planet.  479 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nMaterials and methods 480 \nBacterial host and phage culture. All bacterial host isolates used in this study were 481 \ncultivated at ambient room temperature (~22°C) in 1´ Reasoner’s 2a (R2A) medium (Neogen, 482 \nUSA) which was prepared following manufacturer’s protocol. Agar (Merck, Germany) was 483 \nadded to a final concentration of 0.4% or 1.5% for soft-agar overlay assay and standard agar 484 \nmedia plates, respectively. Phages were propagated via broth or soft-agar medium depending 485 \non the viral isolate. Broth amplification was started with a 1:50 dilution of an overnight host 486 \nculture in 10 mL of fresh media broth followed by spiking with 20 µL of phage lysate (minimum 487 \n~106 PFU/mL). The mixture is then incubated at room temperature with agitation overnight or 488 \nuntil complete lysis i.e. broth clearance. The amplification reaction is centrifuged at 5000 ´g 489 \nfor 10 minutes to pellet bacterial debris and the viral supernatant is sterilised with 0.45 µm 490 \nsyringe filters (Sartorius, Germany). Amplification via soft-agar medium begins with mixing 500 491 \nµL of overnight host culture with 50 µL of serially-diluted phage lysate and 2 mL of 0.4% R2A 492 \nagar. The 0.4% mixture is casted on 1.5% agar media to form a soft-agar overlay and the plate 493 \nis incubated at room temperature overnight. Plates of phage serial dilutions exhibiting near 494 \ncomplete lysis are then scraped with 5 – 10 mL of sterile 1´ PBS (Invitrogen, Lithuania) and 495 \ntransferred to a 50 mL Falcon tube. The scrapings were vortexed vigorously followed by 496 \ncentrifugation at 5000 ´g for 10 minutes to pellet agar debris and the viral supernatant was 497 \nsterilised with 0.45 µm syringe filters. The phage lysates were stored at 4 °C for short -term 498 \npurposes and at -80°C in 20% glycerol final concentration for long-term storage.  499 \n 500 \nPhage isolation from stream water. The water column in lotic systems functions as a 501 \ndynamic conduit that integrates viral production from upstream biofilms through downstream 502 \ndispersal, where waterborne phages represent a subset of mobile and infectious viruses 503 \ndispersing across biofilm-associated hosts. Hence, concentrating viral particles from the water 504 \ncolumn enables scalable and reproducible recovery of viable phages. 120 L of fresh water 505 \nwas collected from a stream confluence located at Champéry, Switzerland that was fed by 506 \nboth groundwater-fed and glacier-fed streams (Fig.S1: 46.164 N, 6.8610 E; 1051 m above sea 507 \nlevel). Water samples were immediately transported back to our facility and concentrated via 508 \ntangential flow using  a hollow fibre cartridge with 1.15 m 2, 100 000 kDa membrane ( GE 509 \nHealthcare, USA ) to a final volume 2 L which was filtered via 0.45 µm Sterivex (Merck, 510 \nGermany) to exclude eukaryotic and most prokaryotic organisms but not viruses. The filtered 511 \nwater was further concentrated to 500 mL using a tangential flow cassette with 100 000  kDa 512 \n(Sartorius, Germany). The concentrated water is then screened for phages against 40 513 \npreviously isolated bacterial host strains via soft -agar overlay by mixing 500 µL of overnight 514 \nhost culture with 1 mL of concentrated water and 2 mL of 0.4% agar before casting on 1.5% 515 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nagar media plates. Phage plaques identified were cored with sterile wide-bore pipette tips and 516 \nresuspended vigorously in 100 µL of 1 ´ PBS. Phages in PBS were then serially diluted and 517 \nplated using soft agar overlay to perform a subsequent round of coring resulting in double -518 \npurified phage isolates. Purified phages were amplified in broth or soft-agar media to generate 519 \nhigh viral titres (108 – 1010 PFU/mL) for downstream analyses.  520 \n 521 \nTransmission electron microscopy and plaque morphology imaging. TEM was 522 \nperformed with negative stain. Each sample was  adsorbed on a glow -discharged carbon-523 \ncoated copper grid 400 mesh (EMS, Hatfield, PA, USA) washed with  deionized water and 524 \nstained with Uranyl Acetate 1% for 30 seconds. Observations was made using a Talos L120C 525 \nelectron microscope (Thermo Fisher, Hillsboro, USA) operated at 120 kV. Digital images were 526 \ncollected using a CMOS camera Ceta -S (Thermo Fisher, Hillsboro, USA)   4098 ´ 4098 527 \npixels.), using a defocus range between -1.5 µm and -2.5 µm. Images of plaque morphologies 528 \nwere obtained on serially -diluted pure phage cultures on 0.4% soft -agar with a Scan 30 0 529 \nautomatic colony counter (Interscience, France). 530 \n 531 \nPhage DNA extraction and long -read sequencing. 1 mL of phage lysate at 10 8 PFU/mL 532 \nminimum, were initially treated with 10 U of DNase ( Thermo Scientific, USA) with 1´ DNase 533 \nbuffer (diluted from 10´ stock comprising of 0.1 M Tris pH 7.5, 1 mM CaCl2 and 25 mM MgCl2) 534 \nfor 2 h at 37 °C to eliminate bacterial DNA within the lysate. DNase was  then inactivated by 535 \nincubating at 75°C for 30 minutes. Removal of bacterial DNA was verified via 16S rRNA PCR 536 \n(V3/V4 primers 341F: 5’ -TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGCCT 537 \nACGGGNGGCWGCAG-3’ and 785R: 5’- GTC TCGTGGGCTCGGAGATGTGTATAAGAGACA 538 \nGGACTACHVGGGTATCTAATCC-3’) with the following cycling conditions: initial denaturation 539 \nat 95°C for 3 minutes, 30 cycles at 95 °C for 30 seconds, 55 °C for 30 seconds, 72°C for 30 540 \nseconds, 5-minute final extension  using 1 µL of template;  and subsequently, gel 541 \nelectrophoresis with a ~550 bp product if PCR-positive for 16S rRNA. PCR negative lysates 542 \nwere extracted for phage DNA using two methods. For most phage isolates, Norgen Biotek 543 \ncolumn-based phage extraction kit (Norgen Biotek, Canada) was used following 544 \nmanufacturer’s protocol including 20 µL Proteinase K (PanReac AppliChem, Switzerland) 545 \ntreatment and an extended incubation at 55°C for 1 h during viral lysis step. However, isolates 546 \nthat are recalcitrant to column -based extraction were extracted following a modified DNA 547 \nisopropanol precipitation protocol. Here, 10% SDS (Fisher Scientific, UK) was added to the 548 \nphage lysate at a final concentration of 0.5%, mixed well, and allowed to incubate at ambient 549 \nroom temperature for 30 minutes. Sodium acetate ( Sigma-Aldrich, USA) solution was then 550 \nadded to the lysate to a final concentration of 0.3 M followed by 0.6 – 0.7 volume of ice-cold 551 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nisopropanol (Merck,  Germany). The mixture was mixed by inversion and immediately 552 \ncentrifuged at 15 000 ´g at 4°C for 30 minutes. The supernatant was discarded, and the DNA 553 \npellet was washed with 500 µL of freshly prepared 70% ethanol (Fisher  Scientific, UK) and 554 \npelleted at 15 000 ´g at 4°C for 30 minutes, twice. Once residual ethanol from the final wash 555 \nwas dried, the pellets were resuspended in 50 µL TE buffer overnight at 4°C for complete DNA 556 \nresuspension. Extracted DNA concentration, quality, and integrity was assessed using Qubit® 557 \nFluorometer with 1 ´ dsDNA High Sensitivity kit ( Invitrogen, USA ), Nanodrop® ( Scientific, 558 \nUSA), and Genomic TapeStation® (Agilent, USA) following manufacturers’ protocols, 559 \nrespectively. DNA library preparation and MinION long-read sequencing were performed using 560 \nNative Barcoding Kit 24 V14 (Oxford Nanopore Technologies, UK) following manufacturer 561 \nprotocol, using short fragment buffer to maximise read preservation at final wash step prior to 562 \neluting the genomic library for sequencing. When long-read data were insufficient to assemble 563 \ncomplete phage genomes, DNA sequencing was performed on the Illumina MiSeq i100 564 \nplatform using the TruSeq™ DNA PCR-Free Library Prep Kit (Illumina, USA) with a random 565 \nfragmentation (Covaris E220 focused ultrasonicator). We caution that certain phage isolates 566 \nespecially those infecting P. cyclaminis, remain notoriously recalcitrant to DNA extraction while 567 \nthe Sphingomonas phage isolate was limited to low amplification titre of <104 PFU/mL despite 568 \nour best efforts, which not only hampered DNA extraction but also transmission electron 569 \nmicroscopy.  570 \n 571 \nPhage genome assembly, annotation, and bioinformatic analyses . Long-reads were 572 \nbase-called and demultiplexed using Dorado (v0.9.1) with high-accuracy model and remaining 573 \nadapters and barcodes were removed using Porechop (v0.2.4). Any reads mapping to the 574 \nbacterial host genome were also removed. Sequenced reads were assembled into contigs 575 \nwith Flye (v2.9.5) when using long -reads only and with Unicycler (v0.5.1) when performing 576 \nhybrid assembly. Contigs were quality -checked to verify their viral origins using VIBRANT 577 \n(v1.2.1) and geNomad (v1.9.0). Where possible, we also derived taxonomic information of our 578 \ncompleted genomes with geNomad to assess the novelty of our viral collection against publicly 579 \navailable databases. We then complemented CheckV (v1.0.3) with an in-house read-mapping 580 \nvisualiser theBIGbam (v0.1; unpublished) to evaluate phage genome completeness based on 581 \nnucleotide sequence and read mapping features, respectively. We dereplicated our collection 582 \nof complete phage genomes via nucleotide identity using scripts provided by CheckV  with 583 \n85% coverage and 95% nucleotide identity thresholds. Phage contigs assembled as head-to-584 \ntail concatemers were identified as overcomplete genomes, and genome lengths were 585 \nautomatically corrected by retaining a single -repeat unit corresponding to the expected 586 \ncomplete phage genome. Genes were predicted using PHANOTATE (v1.5.0) and annotated 587 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nwith a combination of pharokka (v1.8.2) and phold (v1.1.0) using sequence and structural 588 \nhomology, respectively. DNA packaging mechanisms were also determined with theBIGbam 589 \nusing the logic from PhageTerm33. Nucleotide BLAST to NCBI database was performed with 590 \ndefault settings using the dereplicated genome sequences as the query sequence. 591 \nMeanwhile, queries to planetary -scale LOGAN contig database were initiated by randomly 592 \nsampling 1 kb segments from the genomes of each dereplicated phage isolate. The 1 kb query 593 \nsequence was then interrogated against all available contigs in LOGAN with the threshold 594 \nsetting set to the lowest value at 0.25 k-mer coverage (query date 01-Dec-2025). Cumulative 595 \nk-mer coverage scores were calculated by summing the individual coverage values per 596 \nsource. All data was analysed and plotted with R (v.4.2.0). 597 \n 598 \nCrystal violet static biofilm assay. Bacterial overnight cultures were adjusted to OD600 ~0.02 599 \nwith fresh media and 190 µL was aliquoted into each well of a 96-well flat-bottom polystyrene 600 \nplate (Greiner Bio-One, Switzerland). Biofilms were then grown statically in triplicate for each 601 \nbacterial species for 24 h. Following static incubation, planktonic cells in the supernatant were 602 \ndiscarded by carefully inverting the plate into a waste basin and the biofilms were wash thrice 603 \nwith MilliQ water (Merck Millipore, Germany) by submerging the plate in a separate filled basin 604 \nand discarding the water in waste. The 96 -well plate was then inverted on a paper towel to 605 \nremove excess liquid and is allowed to air-dry. Crystal violet staining was performed by adding 606 \n200 µL of 0.01% Crystal violet solution  (Sigma-Aldrich, USA) in each well and stained at 607 \nambient temperature with low-speed orbital shaking for 30 minutes. Excess Crystal violet was 608 \ndiscarded, and the stained biofilms were washed thrice with MilliQ water and allowed to air -609 \ndry. 200 µL of 70% ethanol (Fisher Scientific, UK) was then added to each well and incubated 610 \nat ambient room temperature for 30 minutes with low orbital shaking to solubilise the Crystal 611 \nviolet. Absorbance values at 590 nm were obtained (A 590) using Biotek Synergy H1 plate 612 \nreader (Agilent, USA) and the datapoints were blanked with A590 values from negative control 613 \nwells containing sterile media.  614 \n 615 \nMicrofluidic biofilm time -course phage infection. Polydimethylsiloxane (PDMS) 616 \nmicrofluidic devices with channel dimension of 40 µm height, 250 µm width, and 1 mm length 617 \nwere purchased (Wunderlichips, Switzerland) and sterilised with 70% ethanol followed by UV 618 \nradiation for 30 minutes. Microfluidic biofilms were established by first seeding the channels 619 \nwith 10 µL of overnight bacterial cultures adjusted to OD 600 ~0.1 in fresh media. Cells were 620 \nallowed to attach under static conditions for 2 h at ambient room temperature. Devices were 621 \nthen connected a media-filled syringe via autoclave-sterile PTFE tubing (ID: 0.56 mm and OD: 622 \n1.07 mm, Fisher Scientific, Switzerland) with the outlet emptying into a waste Eppendorf tube. 623 \nMedia was infused continuously with a syringe pump (New Era Pump Systems, USA) at 0.1 624 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint \n\nµL/min flow rate and biofilms were grown for 2 h at ambient room temperature. Following 625 \novernight growth, sterile media syringes were swapped for 10 6 PFU/mL of phages diluted in 626 \nsterile media and phages were infused continuously at 0.1 µL/min flow rate for a further 96 h. 627 \nControl biofilms not infected with phages were maintained with sterile media infusions. Biofilm 628 \ndevelopment was then monitored over the 96-h duration under default brightfield settings via 629 \nZeiss Axio Zoom.V16 microscope with the Plan-NEOFLUAR Z 1.0 ´/0.25 FWD 56  mm 630 \nobjective and 112 ´ zoom magnification  (Carl Zeiss Microscopy, Germany) . Images were 631 \nacquired and processed with Zen (blue edition) software (Carl Zeiss Microscopy, Germany).  632 \n 633 \nData, code, and phage culture availability. All code used for  long-read preprocessing, 634 \nassembly and genome annotation were integrated in a Snakemake  (v7.32.4) pipeline 635 \navailable at: https://github.com/bhagavadgitadu22/PhageID. Sequencing reads and 636 \nassembled genomes will be deposited in NCBI under BioProject: PRJNA1423127 following 637 \nmanuscript review. Purified phage cultures will also be submitted to CRBIP Biological 638 \nResource Center of Insitut Pasteur following manuscript acceptance. All plots were generated 639 \nwith R and raw data are available via supplementary data tables.  640 \n 641 \nAcknowledgements. We thank Martina Gonzalez and David Touchette for sharing their 642 \nbacterial culture collection from streambed biofilms as bait for our phage isolation efforts. We 643 \nalso thank Florence Jagorel and Marc Monot from the group PhagoMics, Phages.fr, for their 644 \nadvice and support, and the Biomics Platform, C2RT, Institut Pasteur, Paris, France, supported 645 \nby France Génomique (ANR -10-INBS-09) and IBISA.  We finally thank Tom Battin for his 646 \nguidance and provision of laboratory facility in support of this work. This work was supported 647 \nby the Swiss National Science Foundation under grant no.212726 awarded to Hannes Peter. 648 \n 649 \nAuthor contributions . W.H.C and H.P. conceived the study, and W.H.C designed 650 \nexperiments and interpreted data. W.H.C, M.B., and A.H. carried out field expeditions, sample 651 \nprocessing, and phage isolations. W.H.C and A.H. purified phages and maintained the phage 652 \ncollection throughout the study. W.H.C, A.H., and F.B. performed phage genome extractions 653 \nand sequencing. M.B. established bioinformatic pipelines and analysed genomes . W.H.C 654 \nperformed experiments, viral database screens, and data analyses. D.D. performed electron 655 \nmicroscopy imaging. W.H.C prepared figures. W.H.C. and H.P. wrote the manuscript. H.P. 656 \nsupervised the study. All authors reviewed, edited, and commented on the manuscript.  657 \n.CC-BY 4.0 International licenseperpetuity. 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It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted March 26, 2026. ; https://doi.org/10.64898/2026.03.26.713887doi: bioRxiv preprint","source_license":"CC-BY-4.0","license_restricted":false}