Phage therapy has gained rapid traction as a biocontrol strategy in soils and soil–vegetable interfaces, providing a precision, self-amplifying alternative to copper bactericides and antibiotics for pathogens such as Pseudomonas syringae, Xanthomonas spp. and Ralstonia solanacearum (Bisen et al., 2024). High-throughput isolation coupled with long-read genomics was used to certify absence of lysogeny or AMR genes. This has yielded “phage banks,” and three phage active ingredients (marketed as AgriPhage^™) which have cleared U. S. Environmental Protection Agency (USEPA) Microbial Pesticide registration (USEPA, 2011). A separate EPA decision approved phage active-ingredients against Erwinia amylovora (EPA, 2018).

Greenhouse and field assays show that individual phage strains (e.g., ɸsp1) have achieved ~82–88% wilt reduction in tomato seedling assays (Wang et al., 2020). Commercial phage formulations have advanced to orchard-scale applications; however, UV exposure promotes pyrimidine dimerization in DNA, thereby limiting phage persistence. To address this, evening sprays and phage formulations containing UV-shielding adjuvants (such as peptides, aromatic amino acids, polysorbate, kaolin, pregelatinized corn flour, sucrose, skim milk formulations, or pigments like carotenoids and betalaines) have been employed to extend the phage activity sufficiently to overlap with pathogen infection windows (Born et al., 2015; Ke et al., 2024). These findings underline the need for next-generation UV-protective formulations or evening spraying regimes to ensure field trials are successful. Beyond direct pathogen inactivation, metagenomic surveys indicate that soil phages modulate nutrient cycling and contribute to auxiliary metabolic genes that improve host fitness (Wang et al., 2024a). These agronomic and ecological gains position bacteriophages as an emerging component of sustainable agriculture, advancing One-Health objectives by limiting environmental AMR reservoirs while enhancing soil-ecosystem function.

Fuel-ethanol fermentations are commonly run under non-sterile conditions because full steam-sterilization of 100–250 m³ vessels would impose an energy penalty. This compromise invites chronic contamination, wherein next-generation sequencing surveys show that lactic-acid bacteria (LAB) dominate the microbial load. A single bloom of Lactobacillus fermentum can cut ethanol yields by up to 27% and trigger costly shutdowns (Bischoff et al., 2009). Historically, plants have dosed oxidizing chemicals or antibiotics such as virginiamycin; however, Lactobacillus isolates from medicated facilities carry the vat(E) efflux gene (Khatibi et al., 2014), and concerns about antibiotic use are posited due to residues persisting in distillers’ grains. A study showed that phage cocktails can recover ethanol yield in L. fermentum–contaminated fermentations to near control level (Liu et al., 2015); pilot-scale deployments are needed to validate yield recovery at equivalent dosing levels with no adverse effects on yeast viability.

While phage cocktails demonstrate strong standalone performance in recovering ethanol yields, integrating complementary agents such as phage-derived endolysins could further stabilize fermentation processes. Wang et al. (2020) reported recovering ~3.3 g/L of thermostable lysin (TSPphg) from a 20 L E. coli fed-batch, a value which is the current maximum in microbial hosts (Cremelie et al., 2024).

The persistence of hard-to-eradicate microbes including water-borne pathogens, bloom-forming cyanobacteria, filamentous “foaming” bacteria and recalcitrant biofilms, poses an escalating challenge for aquatic systems worldwide. Outbreaks of Vibrio cholerae, Escherichia coli, Legionella spp., Leptospira and Salmonella enterica serovar Typhi are showing geographically shifting incidence patterns tied to environmental change. Since 2010, the Global Task Force on Cholera Control has recorded millions of suspected cholera cases and tens of thousands of deaths, with South Asia bearing the heaviest burden. The WHO priority list 2024 named 15 antibiotic-resistant bacteria for accelerated R&D, ranking water-associated carbapenem-resistant Acinetobacter baumannii and third-generation-cephalosporin/carbapenem-resistant Enterobacterales as “critical” (WHO Bacterial Priority Pathogens List, 2024).

Target-specific phages have been isolated from rivers, wastewater and sewage for many of these pathogens, including E. coli, A. baumannii, Shigella, V. cholerae, Salmonella, Citrobacter freundii, Pseudomonas aeruginosa and Klebsiella, and repeatedly reduce bacterial loads in laboratory and pilot studies (Ji et al., 2021). WHO conducted earliest trials of anticholera phages in vivo, and demonstrated that seasonal cholera epidemics in Bangladesh are modulated by rising environmental vibriophage levels, leading to self-limitation (Marcuk et al., 1971). Anticholera phages lowered in-vivo V. cholerae counts, and long-term environmental monitoring in Bangladesh revealed self-limiting cholera waves as vibriophage titres rose (Faruque et al., 2005). A three-phage cocktail (pSf-1, pSb-1, pSs-1) suppressed Shigella in animal model (Jun et al., 2016). Although no lytic phage for Legionella has yet been recovered, a study identified CRISPR spacer matches to Microviridae-related sequences in Legionella, indicating yet-to-be-isolated phages (Deecker et al., 2021).

Cyanobacterial blooms create multimillion-dollar annual burdens. While strain-specific cyanophages for Microcystis are known (Zhang et al., 2022), field deployment is nascent: a Lake Baroon trial achieved 95% cell lysis but regrowth ensued as resistance emerged (Tucker and Pollard, 2005). Infection of Aphanizomenon flos-aquae by cyanophage vB_AphaS-CL131 demonstrated that phage infection alters ¹⁵N₂ assimilation and thus algae nitrogen metabolism. While these findings underscore the potential of phages for bloom control, they also highlight two critical challenges: the rapid emergence of phage resistance under field conditions, and the need for careful evaluation of ecosystem-level effects following viral-mediated lysis events. Future deployments should integrate adaptive phage strategies and longitudinal environmental monitoring to mitigate unintended consequences with minimal ecosystem disruption (Kuznecova et al., 2020). Lytic phages from Myoviridae family such as SN-phage, GTE7 and SPI1 can suppress foaming filamentous bacteria in activated sludge (Vesga-Baron et al., 2022), but no phage has yet been isolated for the ubiquitous foam forming organism—Microthrix (Batinovic et al., 2019). Bhattacharjee et al. (2015) reported that membrane flux decreased from ~15 to ~47 L/h·m² due to Delftia tsuruhatensis biofilm formation, but the application of phage DTP1 restored membrane flux to 70% of unfouled baseline. Ayyaru et al. (2018) demonstrate that E. coli phage P2 significantly reduce the antibiotic-resistant biofilms on PVDF–graphene membranes. Zhang et al. (2024) show via metagenomics that viral predation in anaerobic digestion predominantly reduces antibiotic resistant bacteria loads, with limited horizontal gene transfer of antimicrobial resistance genes (Zhang et al., 2024). Environmental phage applications are a promising strategy for aquatic pathogen control and bottom-up control of engineered processes.

Host-range precision is both a strength and weakness in phage therapy for environmental biotechnology applications. Surveys show that most lytic phages infect only a few tested strains. In a survey of Klebsiella phages across 138 strains, 42 of 46 phages (≥91%) infected three or fewer strains, averaging under 2% host coverage per phage (Beamud et al., 2023). This implies a key challenge for phage therapy in safeguarding beneficial native microbiota while limiting environmental dispersal within genetically diverse and spatially dynamic bacterial communities (de Jonge et al., 2019). Infection begins when virion adhesins dock onto envelope receptors (O-antigen side-chains, core LPS, wall-teichoic acids, outer-membrane porins, type IV pili or flagella) the molecular “locks” for the phage “key” (Broeker and Barbirz, 2017; Venturini et al., 2022). This is because these structures diversify rapidly, most phages recognize only a narrow phylogenetic subset (Beamud et al., 2023). Such selectivity is valuable in the rhizosphere or activated sludge, where functional components of the microbiome, e.g., nitrifiers and plant-growth promoters should be avoided. For example, in Salmonella phage panels, individual phages often infect only one or a few serovars; a monophage for PT4 fails against PT8 or co-resident Klebsiella strains, illustrating incomplete control (Beamud et al., 2023; Wang et al., 2024b). Studies where broader host range has been engineered via tail-fiber swaps report effective range expansion but phages pay a ‘fitness cost’ due to reduced adsorption rates and burst size (Dunne et al., 2019). Each engineered lineage, however, is a new biological entity with its own safety dossier and regulatory path (Samson et al., 2024). In-silico tools, e.g., PHERI, vHULK predict the probable hosts directly from metagenomic contigs (Amgarten et al., 2022; Baláž et al., 2023). This effectively reduces the wet-lab screening burden and accelerating targeted phage discovery from complex environmental samples.

Rational phage cocktails mitigate these limits. High-throughput screening workflows against broad isolate libraries, plus whole-genome sequencing to exclude lysogeny, toxins and ARGs, yields complementary candidates (Molina et al., 2022). Phages are amplified and produced at ≥10⁷–10⁸ PFU mL⁻¹ per phage, then concentrated/filtered (e.g., ultracentrifugation) and mixed in defined ratios (Jończyk-Matysiak et al., 2021). Most environmental formulations contain single digit numbers of phages, balancing coverage with manufacturing complexity and maintain quality control feasibility (Holtappels et al., 2021). Evolution experiments illustrate the payoff, for example, Martinez-Soto et al. report a five-phage Salmonella cocktail yielding a bacteriophage-insensitive-mutant frequency of 6.22 × 10⁻⁶ PFU mL⁻¹ (much lower than with single phages) over their assays four-phage cocktail targeting O-antigen, porins, flagella and an efflux pump (Martinez-Soto et al., 2024). When resistance finally arises, it often carries fitness costs including truncated LPS or lost motility. Multiple studies show that phage-resistant mutants often bear LPS-structure deletions or motility defects, reducing in vivo fitness and sometimes increasing antibiotic sensitivity (Liu et al., 2015).

Host specificity remains both the strength and the bottleneck of environmental phage therapy; genomics-guided cocktail design, machine-learning host prediction and receptor engineering together offer the most pragmatic route to broaden coverage while preserving ecological precision. Ultimately, balancing host-range precision with breadth of coverage is not merely a formulation challenge, but an important ecological consideration for environmental application of phages therapy.

Industrial-scale phage manufacture now integrates biopharmaceutical–style upstream intensification with vaccine-grade downstream polishing to deliver well-defined, GMP-compliant APIs. Upstream, high-cell-density fed-batch and continuous stirred-tank (CSTR) cultivations routinely achieve high titres up to 10¹¹ PFU/mL, orders of magnitude above traditional batch fermentations (Mancuso, 2020). Many facilities employ single-use, closed bioreactors equipped with in-line optical-density for real-time monitoring of host growth and phage production (Matthew, 2022), and capacitance sensors are used widely in mammalian studies and could be deployed for phages. Emerging intensification strategies, such as perfusion or alternating tangential-flow reactors which are used in vaccine/viral vector contexts could boost productivity while maintaining tight process control, but no published phage-specific data are available.

Downstream, workflows mirror those used for viral vaccines: primary clarification by depth filtration followed by tangential-flow ultrafiltration (100–300 kDa MWCO) to concentrate phages and reduce endotoxin by 1–2 logs. Rebula et al. (2023) showed that an 8 mL CIMmultus OH capture step removed 98% of host proteins and >99% of host DNA with 100% phage recovery. A follow-up polishing step on 8 mL CIMmultus H-Bond or PrimaS columns achieved a 7 log₁₀ reduction in endotoxin while maintaining phage infectivity, as confirmed by HPLC analytics correlated to drop-assay titration. CIMmultus® monoliths also support very high virus binding capacities up to 6 × 10¹³ PFU/mL have been demonstrated, enabling rapid, high-throughput viral purification. Process Analytical Technology tools such as digital PCR for identity/quantification and inline Limulus Amoebocyte Lysate assays for endotoxin are being piloted to accelerate real-time release testing in broader biomanufacturing but are recommended for phage specific production. These tools specifically detect lipopolysaccharide which is the “endotoxin” carried in the outer membrane of Gram-negative bacteria, and is not triggered by viral proteins or nucleic acids.

Validated GMP processes aim for low residuals (<5 EU/mL endotoxin, <10 ng/mL host DNA) based on vaccine monographs, although phage-specific regulatory guidelines have yet to fully evolve. Together, developments in upstream and downstream advances—coupled with genomic safety screening and real-time analytics provide a framework for scalable, cost-effective production of environmental phage products under emerging GMP frameworks.

Regulatory oversight of bacteriophage-based interventions is now the principal rate-limiting step in translating laboratory successes into routine environmental practice. In the therapeutic arena the pathway is well defined: in the United States, any phage intended to treat or prevent disease is classified as a “biological product” and reviewed by the FDA’s Center for Biologics Evaluation and Research under the IND/GMP rubric used for vaccines and monoclonal antibodies (Plaut and Stibitz, 2021). Although no product has yet reached full licensure, several first-in-human and emergency protocols are active; the 2022 IND for Adaptive Phage Therapeutics’ PhageBank which evaluated a dynamic quality-controlled phage library and matching algorithm rather than a static cocktail, represents a critical regulatory milestone. The European Medicines Agency takes the same position: phages are medicinal products, and its 2023 veterinary guideline (EMA, 2023a) formally adopted as agency policy (EMA, 2023b). This requires a traceable master bank, fully annotated genomes free of lysogeny, toxins and AMR genes, plus potency assays demonstrating each phage’s contribution to spectrum and resistance management. Both regulators are piloting “phage-bank” or master-file schemes that would allow a pre-vetted phage to be swapped into a licensed cocktail without reopening the entire regulatory dossier (Lea-Smith et al., 2025).

In contrast to medicinal use of phages, regulation for environmental phage applications remains fragmented and immature (Samson et al., 2024). The U. S. Environmental Protection Agency has approved a handful of agricultural sprays under its microbial-biopesticide pathway (EPA Biopesticide Registration Notices), yet there is still no defined route for water disinfection, corrosion control or biofuel fermenters, and the European Union relies on generic biocide or feed-additive law rather than a phage-specific framework. In practice, applicants everywhere are asked for the same core dossier, i.e., genomic safety, potency and identity over shelf-life for every constituent, and evidence that each phage earns its place, however, timelines and evidentiary depth vary widely. Harmonized guidance for non-clinical sectors, formal adoption of dynamic “phage-bank” licensing, and expanded use of real-time release testing (RTRT) would significantly streamline regulatory approval for environmental phage products. Ongoing EPA biopesticide approvals and EMA draft guidance/concept papers provide a framework, but a dedicated environmental-phage regulatory pathway, ideally harmonized under a One-Health umbrella—will be crucial to move from lab to routine practice (Figure 1).