From Isolation to Application: Utilising Phage‐Antibiotic Synergy in Murine Bacteremia Model to Combat Multidrug‐Resistant Enterococcus faecalis

ABSTRACT

Enterococcus species, natural inhabitants of the human gut, have become major causes of life‐threatening bloodstream infections (BSIs) and the third most frequent cause of hospital‐acquired bacteremia. The rise of high‐level gentamicin resistance (HLGR) in enterococcal isolates complicates treatment and revives bacteriophage therapy. This study isolated and identified forty E. faecalis clinical isolates, with 30% exhibiting HLGR. The HLGR5 isolate, resistant to fosfomycin, vancomycin, and linezolid, was used to isolate the vB_EfaS_SZ1 phage from effluent water. This phage specifically lysed 42% of HLGR isolates. vB_EfaS_SZ1 demonstrated beneficial traits, including thermal stability, acid–base tolerance, a short latent period, and a large burst size. The phage genome comprises a 40,942 bp linear double‐stranded DNA with 65 open reading frames (ORFs). The genome closely resembled Enterococcus phages, classifying it within the Efquatrovirus genus. Phage‐antibiotic synergy was assessed using checkerboard assays and time‐killing analyses, revealing enhanced bacteriolytic activity of ampicillin and fosfomycin, with significant reductions in minimum inhibitory concentration values. In a mouse bacteremia model, phage‐antibiotic combinations significantly reduced E. faecalis liver burden compared to monotherapies. Histopathological analysis confirmed therapeutic synergy, showing reduced inflammation and improved hepatocyte regeneration. These findings underscore the potential of phage vB_EfaS_SZ1 as an adjunct to antibiotic therapy for resistant enterococcal bacteremia.

Isolation, Genome Annotation, Characterisation, and Therapeutic Application of the vB_EfaS_SZ1 Enterococcus Phage.

This study isolated and identified a novel lytic phage that infects HLGR Enterococcus faecalis . vB_EfaS_SZ1 exhibited advantageous traits, including enhanced stability and efficacy. Additionally, phage‐antibiotic synergy significantly improved treatment outcomes for enterococcal bacteremia in a murine model.

1. Introduction

Bloodstream infection (BSI) is a rising public health concern, being one of the most serious nosocomial infections. BSIs cause globally poor patient prognosis, adverse outcomes, exacerbation of healthcare expenditures, and diagnostic ambiguities (Laupland and Leal 2020). BSIs have been classified according to the place of acquisition into community and healthcare‐associated BSIs. Meaningful differences with respect to clinical characteristics, isolated microorganisms, and outcomes were identified between community‐ and healthcare‐associated bacteremia (Friedman et al. 2002). Mortality of BSIs developed in the community is 14%. However, the mortality rate increases drastically to 30% among hospitalised patients with comorbidities. This could be even worse if this infection is caused by antimicrobial‐resistant (AMR) bacteria, “called nightmare bacteria”, since it cannot be successfully tackled by currently available therapies (Santoro et al. 2020). AMR pathogens cause 700,000 global deaths annually, and it is estimated that the global annual number of AMR infections will increase tenfold by 2050 (Tagliabue and Rappuoli 2018). The development of alternative approaches to overcome increased antimicrobial resistance is an urgent global demand and challenge.

Among the AMR pathogens causing healthcare‐associated BSIs is enterococci, one of the ESKAPE pathogens as stated by WHO (Ma et al. 2020). Enterococcal bacteremia mainly affects immunocompromised and elderly patients, leading to high morbidity and mortality, and can be complicated by infective endocarditis (Wisplinghoff et al. 2004). Enterococci are normally found in the gastrointestinal tract of many organisms, including humans (Jett, Huycke, and Gilmore 1994). As a commensal exposed to antimicrobial selective pressure, especially during clinical practice, enterococci have developed a wide array of stress responses and genetic manipulation to survive and subsequently disseminate in the healthcare settings (Miller, Munita, and Arias 2014). Despite rigorous protocols for cleaning and sterilisation in healthcare settings, enterococci bacteria can survive under wide ranges of pH, temperature, and salinity levels. Moreover, enterococcal resistance to detergents such as bile salts and sodium dodecyl sulphate was also stated (Fiore, Tyne, and Gilmore 2019). This robust enterococcal environmental ruggedness plays a major role in their persistence and spread in healthcare settings (Weinstein and Hayden 2000).

Treatment of enterococcal bacteremia is quite challenging owing to enterococcal intrinsic resistance and tremendous genomic plasticity to accumulate resistance genes to different chemotherapeutic agents. Specifically, Enterococcus faecalis ‐causing bacteremia demonstrated the highest number of antibiotic resistance genes (He et al. 2018). Of the intrinsically resistant antibiotics, E. faecalis reduces cellular permeability to aminoglycosides, leading to low‐level intrinsic resistance. Moreover, low penicillin‐binding protein (PBP) affinity to β‐lactams renders β‐lactams bacteriostatic rather than bactericidal (Mundy, Sahm, and Gilmore 2000). The combination of β‐lactams and aminoglycosides restores the bactericidal effect and is considered the gold standard in clinical practice in enterococcal bacteremia therapy (Beganovic et al. 2018). Nevertheless, encoding for aminoglycoside‐modifying enzymes and ribosomal attachment site modification leads to high levels of aminoglycoside resistance (HLAR) by E. faecalis , which eliminates synergism of aminoglycosides with β‐lactams. Notably, more than half of enterococcal infections in healthcare settings were found to be high‐level aminoglycoside resistant (Chow 2000). Enterococci's ultimate resistance profile and their progressive acquisition of antimicrobial resistance necessitate the introduction of phage therapy as a salvage therapy into clinical practice.

Phage therapy, viruses used to treat bacterial infections, has recently been proposed as an alternative treatment approach, especially in the context of infections caused by AMR pathogens. However, the therapeutic applications of phage in monotherapy led to the emergence of phage‐resistant mutants (El Haddad et al. 2019). To overcome the evolutionary bacterial resistance towards either phage or antibiotics, the combination of phage and antibiotics and their potential synergy are being investigated. Interestingly, previous in vitro studies confirmed PAS in combating E. faecalis infection (Bolocan et al. 2019). Moreover, a recent clinical trial revealed that phage‐antibiotic therapy of recurrent enterococcal bacteremia improved patients clinically with reduced intestinal burden of E. faecium (Stellfox et al. 2024). The current study aimed to isolate and provide a comprehensive biological and genomic characterisation of a novel lytic phage, vB_EfaS_SZ1, targeting HLGR Enterococcus faecalis . Furthermore, we evaluated the therapeutic efficacy of phage‐antibiotic co‐therapy against E. faecalis both in vitro and in vivo using a murine bacteremia model.

2. Materials and Methods

2.1. Bacterial Strains

A total of forty E. faecalis isolates were collected from patients admitted to Zagazig University Hospitals from different clinical sources. The isolates were identified using conventional biochemical techniques, in addition to the VITEK system (Biomerieux, Marcy l'etoile, France). The broth microdilution method was employed to assess the potential development of HLGR among clinical E. faecalis isolates in accordance with the guidelines set by the Clinical Laboratory Standards Institute (CLSI). Susceptibility to antimicrobial agents for HLGR isolates was tested using VITEK susceptibility cards. The isolates were preserved in Muller Hinton broth containing 10%–15% glycerol and stored at −80°C.

2.2. Isolation and Enrichment of E. faecalis Phage

Effluent water samples were collected from Zagazig University Hospitals to isolate E. faecalis phage vB_EfaS_SZ1 using the enrichment technique (Melo et al. 2014). In brief, a flask was loaded with 20 mL of the centrifuged water sample, 20 mL of double‐strength Tryptone Soya Broth (TSB), along with 50 μL of exponential phase TSB culture of E. faecalis using HLGR5 as the host bacterium. Then, the flask was incubated overnight at 37°C with shaking. Following incubation, the culture was centrifuged at 10,000 g for 10 min at 4°C, the supernatant was filtered using a 0.22 μm membrane filter (Millipore, USA), and it was screened for the presence of phage using a spot assay.

2.3. Purification and Propagation of E. faecalis Phage

To purify the isolated vB_EfaS_SZ1 phage, a sterile tip was employed to pick an individual plaque, which was resuspended in 1 mL of TSB, subjected to serial dilution, and then plated using the Double Layer Agar (DLA) technique until consistent plaque morphology was achieved. A high‐titre phage stock was obtained, and the confluently lysed plates were soaked overnight in 5 mL of saline magnesium (SM) buffer. The liquid was decanted, and the top agar layer was scraped into a centrifuge tube before being centrifuged, filtered using a 0.22 μm membrane filter, and stored at 4°C (Karumidze et al. 2013).

2.4. Transmission Electron Microscopy (TEM)

In order to visualise vB_EfaS_SZ1 phage particles, TEM was used (Chatterjee et al. 2019). In brief, a high‐titre pure phage was dropped onto a copper grid. Then, the phage particles were negatively stained with 2% phosphotungstic acid before being observed using a JEM‐2100 TEM (JEOL Ltd.).

2.5. Host Specificity Testing

vB_EfaS_SZ1 phage host range was tested against 12 HLGR E. faecalis isolates in addition to different bacterial species taken from the culture collection of the microbiology department, Faculty of Pharmacy, Zagazig University (Table S1). In brief, aliquots of 10 μL of the concentrated phage stock were applied to the surface of the DLA agar pre‐seeded with the tested strains. After incubation, the plates were inspected for the presence of clear, visible plaques (Karumidze et al. 2013).

2.6. Thermal, Chloroform, pH, and Storage at 4°C Stability Testing

To assess vB_EfaS_SZ1 phage stability against different conditions, phage lysate was treated with various temperatures (from 30°C to 90°C), pH values (2–12), and chloroform ratios (from 0% to 80%) for 1 h. The DLA technique was used to determine the phage titre. Additionally, the phage's stability against storage at 4°C was examined, and the titre was calculated at the specified intervals over a 75‐day period (Melo et al. 2014).

2.7. One‐Step Growth Curve

To determine the vB_EfaS_SZ1 phage latent period and burst size, a one‐step growth curve was employed with minor modifications (Melo et al. 2014). Briefly, the bacterial culture in the early log phase was mixed with the phage stock at a multiplicity of infection (MOI) of 0.1 and incubated at 37°C for 5 min. Subsequently, the mixture underwent centrifugation, and the resulting pellet was resuspended in 10 mL of fresh TSB before being re‐incubated. Aliquots of 100 μL were taken every 10 min to assess the phage titre using the DLA technique.

2.8. Checkerboard Assay

vB_EfaS_SZ1 phage synergy with either ampicillin or fosfomycin was tested using a checkerboard assay with some modifications (Kim et al. 2018). In brief, the antibiotics were serially diluted (twofold) starting at 2× minimum inhibitory concentration (MIC) and added to the wells of the microtiter plate. The phage was serially diluted (tenfold) starting at 10⁸ PFU/mL and added in a perpendicular direction. After the bacterial suspension (10⁵ CFU/mL) was added, the microtiter plates were overnight incubated at 37°C and optical density (OD) at 600 nm was measured. The fractional inhibitory concentration indices (FICIs) were calculated.

2.9. Time‐Kill Assay

To confirm phage‐antibiotic synergy, a time‐kill experiment was employed (Kim et al. 2018). Aliquots of 100 μL from each sample; phage alone, antibiotic alone, phage‐antibiotic combination, as well as the control sample were taken at 0, 2, 4, 6, 24, and 48 h of incubation, serially diluted, and plated to determine bacterial counts. A two‐way ANOVA was employed on interaction plots to evaluate possible synergy between phage and antibiotics.

2.10. Genome Sequencing and Annotation

vB_EfaS_SZ1 phage nucleic acid was extracted from phage lysate (2.5 × 10¹² PFU/mL) using a DNeasy Blood and Tissue kit (QIAGEN, Germany) following the manufacturer guidelines with modification (Jakočiūnė and Moodley 2018). Briefly, 450 μL of the filter‐sterilised lysate is incubated with 50 μL of DNase I 10× buffer, 1 μL of DNase I (1 U/μL), and 1 μL of RNase A (10 mg/mL) for 1.5 h at 37°C without shaking to remove any residual bacterial DNA and RNA present in the lysate. Thereafter, 20 μL of 0.5 M EDTA (final concentration 20 mM) is added to inactivate DNase I and RNase A. Proteinase K (20 mg/mL) is then added and incubated for 1.5 h at 56°C without shaking to digest the phage protein capsid. Then, the guidelines of the Qiagen DNeasy Blood and Tissue kit were followed. The concentration and quality of the DNA were assessed using the Thermo Scientific Nanodrop ND‐1000 UV/VIS Spectrometer and Qubit according to the manufacturer's instructions. The DNA elute was further stored at −20°C until use.

The DNA library for sequencing was generated by applying a Nextera XT DNA Library Preparation kit (Illumina, FC‐131‐1024). The DNA was fragmented and then tagged utilising the transposome in the Nextera XT Kit. The whole genome sequencing was performed on a MiSeq Desktop Sequencer (Illumina) using a MiSeq Reagent kit, version 3, for two 75‐bp paired‐end reads (Illumina, MS‐102‐3001). All preparations and the sequencing run were performed according to Illumina manufacturing instructions. The raw sequence within the phage genome was checked for quality with FastQC, reads trimming was performed, and the trimmed reads were de novo assembled using the Basic Variant Detection tool (Qiagen, v.2.2) of CLC Genomic Workbench (Qiagen, v.21.0.3).

For genome annotation and visualisation, Prokka (Seemann 2014) was employed to predict the open reading frames (ORFs), functional annotation of the predicted protein‐coding sequences, and ORFs in the genome, and the existence of tRNA, rRNA, and ncRNA coding genes were analysed by the Bakta (Schwengers et al. 2021), and Phigaro (Starikova et al. 2020) tools of the online Proksee web‐based platform (accessed on 10 May 2024) (Grant et al. 2023) BLASTP at NCBI (http://blast.ncbi.nlm.nih.gov/). Antibiotic resistance genes were checked using the comprehensive antibiotic resistance database (CARD) Resistance Gene Identifier tool, and clustered regularly interspaced short palindromic repeats (CRISPR) arrays were detected using the CRISPR/Cas Finder tool of the Proksee online package (https://proksee.ca). Linear phage genome comparison was performed using Easyfig 2.2.5 (http://mjsull.github.io/Easyfig/ accessed on 28 April 2024) (Sullivan, Petty, and Beatson 2011).

2.11. Phylogenetic Analysis

The phylogenetic analysis was conducted in MEGA11 (Tamura, Stecher, and Kumar 2021) by comparing the whole genome of the isolated vB_EfaS_SZ1 phage with the complete genome of 20 related Enterococcus phages (a cut‐off ≥ 75% query coverage with ≥ 92.43% nucleotide sequence identity). First, we performed multiple alignments of the 21 phage genomes through the MAFFT online service (Katoh, Rozewicki, and Yamada 2019) (accessed on 28 April 2024). The evolutionary history was inferred using the UPGMA method (Schlee 1975); the bootstrap consensus tree inferred from 1000 replicates is taken to represent the evolutionary history of the taxa analysed (Felsenstein 1985). The evolutionary distances were computed using the Maximum Composite Likelihood method (Tamura, Nei, and Kumar 2004).

2.12. Protein Structure Modelling

Proteins representing DNA helicase, DNA polymerase B, and N‐acetylmuramoyl‐L‐alanine amidase (lysin) of vB_EfaS_SZ1 phage were modelled with AlphaFold v2.3.2 (AF2) (Evans et al. 2021) to predict the 3D structure. Parallel proteins WNA13851.1, WNA13866.1, and WNA13869.1, representing the same enzymes from vB_Efa_VP14 (Sequence ID: OR237563.1), were modelled in the same way. The predicted 3D structure of each pair of parallel proteins from the 2 phages was superimposed and visualised using Mol* Viewer (Sehnal et al. 2021).

2.13. Nucleotide Sequence Accession Number

The whole‐genome sequence of vB_EfaS_SZ1 phage was deposited in GenBank (Accession No.: PP848496).

2.14. In Vivo Mouse Infection Model

The effectiveness of vB_EfaS_SZ1 phage, antibiotics, and combination therapy was investigated using a mouse infection model as previously described with some modifications (Oli et al. 2021). A total of 90 female albino mice (20–25 g) were utilised in the current study (permission for in vivo study use ZU‐IACUC/3/F/400/2023). In brief, the E. faecalis HLGR5 isolate was grown in TSB until it reached the log phase of growth. Then, the bacterial cells were pelleted by centrifugation, and the pellet was resuspended in saline. Mice were distributed into nine groups, with each group consisting of ten mice. Mice in group I were infected intravenously with the E. faecalis HLGR5 isolate (10⁹ CFU). Mice in groups II‐VI were also infected and received a single intravenous injection with different treatments: vB_EfaS_SZ1 phage (10¹⁰ PFU/mL), ampicillin (12.5 mg/kg), a combination of vB_EfaS_SZ1 phage (10¹⁰ PFU) and ampicillin (12.5 mg/kg), fosfomycin (75 mg/kg), and a combination of vB_EfaS_SZ1 phage (10¹⁰ PFU) and fosfomycin (75 mg/kg), respectively. Mice in group VII were uninfected and inoculated once with an intravenous injection of vB_EfaS_SZ1 phage (10¹⁰ PFU). Negative controls, mice were either uninfected and received no treatments or were given a single saline injection.

To assess the impact of various treatments on bacterial burden, mice from each group were anaesthetised and sacrificed at 48 and 72 h post‐inoculation. Liver was aseptically retrieved for bacterial CFU and phage PFU counting. In addition, liver tissue fragments were preserved in 10% buffered formalin for histopathological analysis.

2.15. Statistical Analysis

The data were analysed using GraphPad Prism version 8 software on the Microsoft Windows 2019 operating system. All the experiments were performed in triplicate at different time points. Unless otherwise mentioned, the data was analysed using one‐way ANOVA or t‐test and expressed in terms of mean ± standard error of the mean (SEM) for statistical considerations.

3. Results

3.1. HLGR E. faecalis Clinical Isolates Identification

Forty E. faecalis clinical isolates were isolated and identified. All isolates were tested for their high‐level gentamicin resistance using the broth microdilution method. Twelve out of forty (30%) were HLGR isolates showing gentamicin MIC values > 500 μg/mL. HLGR5 E. faecalis isolate was selected as a host for phage isolation. HLGR5 E. faecalis isolate was resistant to fosfomycin, vancomycin, tetracycline, erythromycin, linezolid, and quinupristin/dalfopristin and sensitive to ampicillin, tigecycline, ciprofloxacin, levofloxacin, and nitrofurantoin (Table 1).

TABLE 1.

Antimicrobial susceptibility of the host strain.

Test group	Antimicrobial agent	MIC (μg/mL)	Interpretation
Penicillins	Ampicillin	0.375	S
Glycopeptides	Vancomycin	≥ 32	R
Fosfomycins	Fosfomycin	1024	R
Tetracyclines	Tetracycline	≥ 16	R
	Tigecycline	≤ 0.12	S
Macrolides	Erythromycin	≥ 8	R
Oxazolidinones	Linezolid	≥ 8	R
Quinolones	Ciprofloxacin	1	S
	Levofloxacin	0.5	S
Nitrofurans	Nitrofurantoin	≤ 16	S
Streptogramins	Quinupristin/dalfopristin	≥ 16	R

Abbreviations: R, resistant; S, sensitive.

3.2. Phage vB_EfaS_SZ1 Isolation and Morphology

E. faecalis HLGR5 isolate was used as the host bacterium in isolating bacteriophages from effluent water samples collected from Zagazig University Hospitals, Zagazig, Egypt. During plaque assay (Figure 1A), clear plaques with an average diameter of approximately 3 mm were observed after 18 h (Figure 1B). Further incubation for up to 48 h resulted in the appearance of an obvious translucent halo zone surrounding the clear plaques (Figure 1C). Moreover, purified phage particles were examined under TEM and revealed an icosahedral head of 47 ± 0.78 nm diameter and a non‐contractile tail of 8.16 ± 1.7 nm diameter and 161.5 ± 3 nm length (Figure 1D). Based on the guidelines of the International Committee on Taxonomy of Viruses (ICTV), phage was classified as a double‐stranded DNA virus belonging to the Siphoviridae family (Figure 1E) within the class Caudoviricetes.

FIGURE 1.

Phage morphology. (A) Schematic overview of the plaque assay's methodology. Phage vB_EfaS_SZ1 plaques on DLA after 18 h (B) and 48 h (C). (D) The morphology of vB_EfaS_SZ1 phage particles under TEM. (E) Illustrations of tailed bacteriophages from Podoviridae, Myoviridae, and Siphoviridae families.

3.3. Host Range of vB_EfaS_SZ1 Phage

The lytic spectrum of E. faecalis ‐isolated phage vB_EfaS_SZ1 was tested against twelve HLGR enterococcal clinical isolates. vB_EfaS_SZ1 phage can infect and lyse five HLGR E. faecalis (42%) tested isolates (Table S1). However, vB_EfaS_SZ1 did not have any lytic activity on either Gram‐positive or Gram‐negative species tested. These findings indicate that the phage vB_EfaS_SZ1 has high bacterial specificity, which can evade interference with human normal flora during clinical application.

3.4. Biological Characteristics of vB_EfaS_SZ1 Phage

Phage vB_EfaS_SZ1 thermal stability was assessed over a wide range of temperatures using the DLA technique. Phage titers did not exhibit significant alterations upon exposure to temperatures ranging from 4°C to 40°C. However, at 50°C and 60°C, phage titre declined to 74% and 88%, respectively. Phage vB_EfaS_SZ1 lost its infectivity at temperatures exceeding 60°C (Figure 2A). Furthermore, chloroform treatment more than 20% (v/v) significantly affected vB_EfaS_SZ1 phage titre, though complete inactivation was not observed (Figure 2B). The acid–base tolerance of vB_EfaS_SZ1 phage revealed its stability over a wide range of pH (3 to 11), with maximum stability observed at pH 7. Phage vB_EfaS_SZ1 was totally inactivated at pH lower than 3 and higher than 11 (Figure 2C). Stability of phage lysate during long‐term storage at 4°C was assessed. Storage of vB_EfaS_SZ1 phage over a 75‐day period demonstrated successive declines in phage titers in the initial 40 days of storage. However, from days 45 to 75, vB_EfaS_SZ1 phage remained at 50% of its initial viability (Figure 2D). To identify phage growth cycle and burst size, a one‐step growth curve was conducted. vB_EfaS_SZ1 phage (an MOI of 0.1) demonstrated a relatively short latent period of 20 min and a burst size of approximately 70 PFU per infected cell (Figure 2E). Collectively, these biological characteristics of vB_EfaS_SZ1 phage facilitate its practical application.

FIGURE 2.

Biological characteristics of vB_EfaS_SZ1. (A) vB_EfaS_SZ1 phage was thermally stable between 4°C and 40°C. (B) vB_EfaS_SZ1 phage titre abated following chloroform treatments exceeding 20% (v/v). (C) vB_EfaS_SZ1 phage tolerated a wide range of pH values from 3 to 11. (D) Long‐term storage at 4°C affected vB_EfaS_SZ1 phage infectivity. (E) One step‐growth curve of vB_EfaS_SZ1 phage cocultured with HLGR5 E. faecalis isolate. PFU, plaque‐forming unit.

3.5. vB_EfaS_SZ1 Phage Genome Sequencing and Functional Annotation

Whole‐genome sequencing results revealed that phage vB_EfaS_SZ1 has linear double‐stranded DNA with a length of 40,942 bp and G + C content of 34.5% (Figure 3A). The genome contains 65 ORFs; 45 (69.23%) of the predicted ORFs are located on the positive strand and 20 (30.77%) ORFs on the negative strand (Figure 3A).

FIGURE 3.

Genome characterisation and phylogenetic analysis of Phage vB_EfaS_SZ1 (A) Circular genome map of phage vB_EfaS_SZ1 (GenBank accession no. PP848496) generated by Proksee on line package (https://proksee.ca) indicating the coding sequence (CDS) on the positive and negative strands, GC content, and GC skew (B) Phylogenetic tree of vB_EfaS_SZ1 phage (in blue) and 20 other Enterococcus phages including vB_Efa_VP14 (in red) using MEGA11program based on alignment of the complete genome sequence against related phages.

The whole genome sequence of phage vB_EfaS_SZ1 was searched in the NCBI BLAST nucleotide database (BLASTN suite). As expected, the BLAST nucleotide search identified the queried genome to be closely related to the Enterococcus phages (up to 85% query sequence coverage and 94.55% sequence identity). Based on BLAST search findings and the closely related genomes of the Enterococcus phages, our novel phage belongs to the Efquatrovirus genus of the Caudoviricetes class, matching genotypically the described morphology of the phage.

Compared to other reported Enterococcus phages, whole‐genome‐based phylogenetic analysis clustered our phage vB_EfaS_SZ1 in the same clade with Enterococcus phage vB_Efa_VP14 (Sequence ID: OR237563.1) with a genome size 40,454 bp, query coverage of 85%, and 94.55% identity (Figure 3B).

Genome functional annotation identified that 47.7% (31) of the ORFs encode hypothetical proteins, and the remaining 34 (52.3%) ORFs constitute structural and functional proteins, including structure, lysis, portal, DNA replication, packaging, and assembly (Figure 4A). The genome of vB_EfaS_SZ1 does not harbour any apparent AMR genes, nor does it contain genes coding for tRNAs, rRNAs, ncRNAs, tmRNAs, CRISPR arrays, or Cas genes. Additionally, the phage genome is free from lysogeny‐related genes and integrases, signifying the lytic mode of the isolated phage.

FIGURE 4.

Functional annotation and genome organisation of phage vB_EfaS_SZ1 (A) Circular genome map of phage vB_EfaS_SZ1 generated by Proksee on line package (https://proksee.ca); the inner circle represents genome organisation and degree of identity of vB_EfaS_SZ1 relative to vB_OCPT_CCS4 (Sequence ID: ON113176.1). (B) Pairwise BLAST comparison of vB_EfaS_SZ1 and vB_Efa_VP14 (Sequence ID: OR237563.1) using Easyfig. ORFs of each phage are presented in orange, and the degree of identity of homologous proteins is shown with different grey colours. (C–E) Superimposition of the AF‐predicted structure of DNA helicase, DNA polymerase B, and lysin, respectively of vB_EfaS_SZ1 (in blue) and vB_Efa_VP14 (in red).

Thirteen ORFs encode the structure/assembly and portal proteins, including prohead protease, head‐tail joining protein, head‐tail adaptor protein, major tail protein, major capsid protein, tail assembly protein, tail tape measure chaperone protein, tail tape measure, phage portal proteins, and phage terminase. Eleven ORFs constitute the DNA replication/modification cluster, including DNA primase/helicase, HNH endonucleases, transposase, DNA methyltransferase, nucleotide kinase, and DNA polymerase B‐like protein. Four ORFs are assigned to holins and the peptidoglycan lysin or hydrolase (N‐acetylmuramoyl‐L‐alanine amidase) (Figure 4A). A full description of genome annotation is found in Table S2.

Comparing the genome of vB_EfaS_SZ1 against the complete genomes of both vB_OCPT_CCS4 (top sequence coverage) and vB_Efa_VP14 (top sequence identity), our phage displayed gene rearrangements compared to vB_OCPT_CCS4 (Figure 4A). Although vB_EfaS_SZ1 shared a high sequence identity and similar gene arrangement with vB_Efa_VP14, significant differences were found in gene clusters responsible for DNA replication (e.g., DNA polymerase B) and modification (e.g., DNA helicase). However, both phages exhibited complete identity in most structural genes. When we aligned the peptidoglycan lysin in our phage and vB_Efa_VP14, they displayed 94% identity (Figure S1A). Phylogenetic analysis of vB_EfaS_SZ1 and other Enterococcus phages, based on the protein sequences of tail tape measure, DNA helicase, DNA polymerase B, and lysin. vB_EfaS_SZ1 and vB_Efa_VP14 occupied the same clade in the phylogenetic tree based on the 156.2 kDa tail tape measure protein (Figure S1B–E). In the same context, analysing the hypothetical proteins that make up ~48% of the phage genome, we selected the largest hypothetical protein (ORF no. 50 of 696 amino acids) for phylogenetic analysis. This protein demonstrated significant homology to analogous proteins from other closely related Enterococcus phages. Notably, homologous proteins included those from both vB_OCPT_CCS4 (UQT01251.1, with 96.7% identity) and vB_Efa_VP14 (WNA13873.1, with 96.41% identity). These findings suggest that hypothetical proteins exhibit considerable homology among phages that infect similar hosts, further emphasising their potential functional relevance (Figure S2A,B).

Next, we utilised AlphaFold to model the 3D structures of the inspected proteins and compared them against the modelled paralogous proteins of vB_Efa_VP14. Despite significant variations in their primary structures (Figure S1F), DNA helicase and polymerase B exhibited nearly identical superimposition in both phages (Figure 4C,D). However, the lysin protein, which shared 94% identity at the primary structure level, displayed a noticeable shift when superimposed (Figure 4E). This indicates that changes in the N terminus have a dramatic effect on the lysin's 3D structure.

3.6. In Vitro vB_EfaS_SZ1 Phage‐Antibiotic Synergy Enhances Bacteriolytic Activity

The effectiveness of vB_EfaS_SZ1 phage‐antibiotic interactions was evaluated by checkerboard assay in which the OD was measured and converted to a heat map. Ampicillin and fosfomycin combination with vB_EfaS_SZ1 phage demonstrated synergy with FICI ranging from 0.25 to 0.5 and 0.13 to 0.5, respectively (Figure 5A,B). The MIC value of ampicillin dropped 4 fold in the presence of sub‐inhibitory vB_EfaS_SZ1 phage (at an MOI of 10⁻³). However, fosfomycin combined with vB_EfaS_SZ1 phage (at an MOI of 10⁻¹) showed an 8‐fold reduction in MIC value compared to fosfomycin alone. Moreover, both ampicillin and fosfomycin effectively reduced vB_EfaS_SZ1 phage titre needed for effective bacterial killing.

FIGURE 5.

Checkerboard assay of ampicillin and fosfomycin combined with vB_EfaS_SZ1 phage on HLGR5 E. faecalis isolate. (A) Ampicillin combined treatment with vB_EfaS_SZ1 phage showing synergism. (B) Fosfomycin combined treatment with vB_EfaS_SZ1 phage showing synergism. Wells of combination treatments that demonstrated in vitro phage‐antibiotic synergy, FICI ≤ 0.5, were outlined by red. Combinational treatments, denoted by a star, were further analysed via time‐killing assay. Time‐killing assay demonstrating the bacteriolytic effect of combination vB_EfaS_SZ1 phage (10⁶ PFU/mL) and ampicillin (0.188 μg/mL) (D) or fosfomycin (512 μg/mL) (C). Interaction plot of vB_EfaS_SZ1 phage with ampicillin (E) and fosfomycin (F) confirming the existence of phage‐antibiotic synergy. Reduction (%) = [(OD_HLRG5 − OD_treatment)/OD_HLRG5] × 100. CFU, colony=forming unit; PFU, plaque‐forming unit. Two‐way ANOVA was performed for statistical significance testing for the interaction plot. *p < 0.05.

To address the combinational efficacy of vB_EfaS_SZ1 phage and ampicillin or fosfomycin, the phage‐antibiotic killing dynamic was monitored. The impact of a combination of vB_EfaS_SZ1 phage (10⁶ PFU/mL) with ampicillin (0.188 μg/mL) or fosfomycin (512 μg/mL), as indicated by checkerboard assay, on bacterial viability was assessed. Under combinational treatment, the reduction in bacterial viability was 69% and 62% against 30% and 27% in ampicillin‐ and fosfomycin‐treated bacteria, respectively (Figure 5C,D). Furthermore, the efficiency of vB_EfaS_SZ1 phage in the presence of ampicillin and fosfomycin was increased by 2.2× and 1.8× times compared to its activity alone. Similarly, both ampicillin and fosfomycin activity were augmented by 2.2 times in combinational treatment compared to their activity alone. In this regard, antibiotic combinational treatment with vB_EfaS_SZ1 phage was analysed via interaction plot and yielded statistically significant synergistic interactions (Figure 5E,F).

3.7. In Vivo vB_EfaS_SZ1 Phage‐Antibiotic Synergy Diminished E. faecalis Load Within Liver Tissue

The therapeutic potential of combinational therapy of vB_EfaS_SZ1 phage and ampicillin or fosfomycin was evaluated in a mouse model of bacteremia (Figure 6A). Suboptimal concentrations of each antibiotic were administered to mimic their clinical application. Antibiotic, phage and combination treatments 48 h post‐inoculation showed reduction in E. faecalis burden within the liver (Figure 6B). However, no pronounced synergistic effect was observed between phage and antibiotic treatment, as no statistical difference was detected between monotherapies and combination therapy. On the contrary, 72 h post‐inoculation, phage‐ampicillin synergism successfully reduced bacterial load with a final 3.2, 3 and 2.8 log difference for the growth control, phage‐ and ampicillin‐only groups, respectively (Figure 6C). Similarly, the combination of phage with fosfomycin synergised their bacteriolytic activity by 1.7× and 1.6× times compared to their activity alone. Interestingly, bacterial resurgence was observed in monotherapy groups (bacterial load mean: 6.1, 5.9 and 5.8 log CFU/g for vB_EfaS_SZ1 phage, ampicillin, and fosfomycin vs. 3.1 and 3.6 log CFU/g for vB_EfaS_SZ1 phage‐ampicillin or fosfomycin combinations, respectively). This finding suggests that phage‐ampicillin or fosfomycin synergy achieved a sustained reduction of bacterial load within the liver with low potential for the emergence of bacterial resistant mutants.

FIGURE 6.

In vivo vB_EfaS_SZ1 phage‐antibiotic synergy diminished bacterial load within liver tissue of E. faecalis ‐infected mice. (A) Schematic overview of the experiment methodology where E. faecalis HLGR5 load within liver tissue after 48 h (B) and 72 h (C) post‐inoculation. *p < 0.05, ***p < 0.001.

In parallel with an increase of E. faecalis load within the phage‐treated group, the phage titre was significantly increased. Moreover, effective phage‐antibiotic combinational therapy led to a drop in phage propagation and titre (Figure 7A). The correlation coefficient of 0.427 showed an evident positive relationship between the phage and E. faecalis titers in both phage and combinational treatment (Figure 7B). Notably, phage was undetectable 72 h post‐inoculation in the uninfected phage‐administered group. These results indicate that phage was eliminated successfully from mice, and phage only coexists with its host bacteria. Collectively, our in vivo study demonstrated greater efficacy of combinational treatment of vB_EfaS_SZ1 phage and ampicillin or fosfomycin compared to either antibiotic‐only or phage‐only against E. faecalis bloodstream infection.

FIGURE 7.

In vivo vB_EfaS_SZ1 phage‐antibiotic synergy led to phage titre reduction correlated with bacterial load in the liver. (A) Combinational therapies significantly reduced phage titre compared to phage‐only therapy. (B) A correlation curve between phage titre and bacterial load in liver tissue indicating a positive relationship. Pearson's correlation test indicated a positive correlation (r 0.4272, p 0.0082). *p < 0.05, ***p < 0.001.

3.8. In Vivo vB_EfaS_SZ1 Phage‐Antibiotic Synergy Mitigated Liver Tissue Damage

The therapeutic effects of phage‐antibiotic synergy were also confirmed by histopathological examination of liver tissue. The haematoxylin and eosin (H&E) stained sections showed dramatic pathological changes in the liver tissues of mice infected with E. faecalis HLGR5, including a tremendous inflammatory infiltrate with hepatocyte degeneration, hepatic sinusoidal dilatation, and congestion (Figure 8A).

FIGURE 8.

Phage‐antibiotic synergy resolved pathological changes caused by E. faecalis infection within liver tissue. (A) E. faecalis‐infected mice showing multifocal inflammatory leukocytic infiltrate (arrows), hepatocytic degeneration (star), and bacterial detection (circle). (B) Phage‐treated mice showed a decrease in the inflammatory infiltrate (arrow). (C) Ampicillin‐treated mice showed sparse inflammatory cells (arrows) and mild degeneration of hepatocytes (star). (D) Phage‐ampicillin combination showing normal hepatic structure and disappearance of inflammatory cells. (E) Fosfomycin‐treated mice showing mild inflammation, scattered inflammatory cells (arrow) and apoptotic bodies (triangle). (F) Phage‐fosfomycin combination showing normal appearance of hepatocytes. Normal hepatic architecture, central vein (cv), and normal hepatocytes are observed in uninfected phage‐treated (G), uninfected (H), and uninfected saline‐injected mice (I). All sections were stained with haematoxylin and eosin (H and E) and were photographed at ×20.

However, either antibiotic or vB_EfaS_SZ1 phage treatment partially resolved the liver tissue alterations triggered by the bacterial infection (Figure 8B,C). Notably, mice treated with fosfomycin showed partial improvement in the form of decreased inflammatory infiltrate and multiple apoptotic bodies (Figure 8E). Combinational therapy with ampicillin or fosfomycin and vB_EfaS_SZ1 phage almost completely abolished the inflammation damage and alterations caused by E. faecalis . Moreover, the combinational treatment groups displayed markedly pronounced normal hepatocytes, restoring healthy tissue appearance like uninfected controls and indicating successful phage‐antibiotic synergy (Figure 8D,F–I).

4. Discussion

Bloodstream infections (BSIs) are a serious health concern that can be either community‐ or hospital‐acquired, with nosocomial BSIs having a higher mortality rate (Friedman et al. 2002; Santoro et al. 2020). Among the most often isolated pathogens causing BSIs is enterococci (Ma et al. 2020). The treatment of enterococcal bacteremia is quite complex due to the rising antimicrobial resistance rates, with all enterococci exhibiting inherited resistance to cephalosporins in addition to resistance to glycopeptides, β‐lactams, and aminoglycosides, which is increasingly prevalent. Moreover, acquisition of genes encoding aminoglycoside‐modifying enzymes and ribosomal attachment site modification leads to HLAR E. faecalis . It was also noted that HLGR isolates had a higher rate of multidrug resistance than non‐HLGR isolates (Chow 2000; Dadfarma et al. 2013).

In the present study, among forty clinical E. faecalis isolates, high‐level gentamicin resistance (HLGR) was noted in 30% of the tested isolates. Enterococcal isolates exhibiting HLGR were also previously reported in Egypt, with BSIs being the second most frequent isolation source after urinary tract infections (Diab et al. 2019). In the current study, E. faecalis isolate HLGR5, obtained from blood, exhibited resistance to fosfomycin, vancomycin, and even linezolid, the only approved medications for the treatment of bacteremia caused by vancomycin‐resistant enterococci (VRE) (Narayanan et al. 2019). The emergence of resistant E. faecalis isolates highlights the urgent need for effective therapeutic strategies, such as phage therapy.

In the current study, a newly lytic phage, vB_EfaS_SZ1, which specifically targets HLGR E. faecalis isolates, was isolated from wastewater effluent discharged from Zagazig University Hospitals using E. faecalis isolate HLGR5 as a host. The isolated phage formed clear plaques on the bacterial lawn, surrounded by halo zones. Clear plaques are commonly regarded as a marker of a lytic phage. The phages that produce halos encode polysaccharide‐degrading enzymes that facilitate their penetration of the bacterial exopolysaccharides and/or capsules (Letarov and Letarova 2023). Additionally, TEM revealed that vB_EfaS_SZ1 phage is categorised within the Siphoviridae family, as were previously isolated E. faecalis phages (Azadani et al. 2020). The potential utility of bacteriophages in clinical application was reported to be correlated with their host range (Hyman and Abedon 2010). vB_EfaS_SZ1 phage could lyse 5/12 (42%) of the tested HLGR E. faecalis isolates, including the original host strain. Similarly, a phage targeting HLGR E. faecalis with a limited host range was identified in another investigation (Azadani et al. 2020). The current results suggest that the isolated phage showed high specificity and a considerable host range.

Understanding how phages respond to varying environmental conditions could help ensure their viability during production, storage, and therapeutic use. Phages belonging to the Siphoviridae family are noted for their stability at high temperatures and harsh pH conditions (Lasobras et al. 1997). Temperature is critical for phage attachment, genetic material ejection, and multiplication. vB_EfaS_SZ1 survived temperatures up to 40°C while the titre was undetectable at higher temperatures (< 60°C). Disulphide cross‐links in the phage capsid proteins have been suggested to be essential for phage stability against heat denaturation (Olson, Axler, and Hicks 2004). Several studies revealed E. faecalis phages that endured temperatures up to 60°C (Yang et al. 2020; Lee et al. 2019). Moreover, the acid–base tolerance of phage facilitates their clinical application. Interestingly, vB_EfaS_SZ1 exhibited a wide pH tolerance (3–11). This agreed with that reported in a recent study (El‐Atrees et al. 2022). vB_EfaS_SZ1 lost its stability at pH < 3 and > 11, which could be due to protein coat denaturation and capsid protein dissociation, respectively. In addition, the optimal phage stability was achieved at pH 7, suggesting its possible applications for clinical use, large‐scale industrial manufacturing, and hospital equipment sterilisation (Feng et al. 2003).

Chloroform treatment has been reported to cause a titre drop in nearly 30% of tailed phages. Similarly, vB_EfaS_SZ1 was tolerant to low concentrations of chloroform (< 20%). Storage stability is crucial for the practical application of phages because their low stability during storage will reduce their effectiveness in controlling diseases (Peng et al. 2023). vB_EfaS_SZ1 phage retained half of its viability after 45 days of storage at 4°C during a 75‐day period. Regarding the one‐step growth curve, vB_EfaS_SZ1 phage had a short latency of 20 min and a burst size of approximately 70 PFU/cell. Another study reported E. faecalis phages with different latent periods and burst sizes (Lee et al. 2019). It has been suggested that phages exhibiting a short latent period and a large burst size are good candidates for phage therapy (Amarillas et al. 2017). Hence, vB_EfaS_SZ1 phage's burst size and short latent period strengthen its potential use in clinical applications.

Whole genome sequencing revealed the structural and functional insights of the genome of vB_EfaS_SZ1. Genetic analysis confirmed the observed phenotype regarding the morphology of the Siphoviridae family. Significant changes on the gene clusters regulating DNA replication, modification, and metabolism were detected by comparing the whole genome of vB_EfaS_SZ1 and another 2 phages sharing the same clade (vB_OCPT_CCS4 and vB_Efa_VP14), which may explain the enhanced latency of vB_EfaS_SZ1.

Among the factors that limit the application of phage therapy are the potential contribution of phages to the development of antibiotic resistance and their lysogenising and genome‐editing activities (Enault et al. 2017; Principi, Silvestri, and Esposito 2019). However, a detailed analysis of the vB_EfaS_SZ1 genome did not identify any putative ORFs encoding antibiotic resistance genes, bacterial virulence factors, lysogeny‐related genes, integrases, or CRISPR arrays, thereby eliminating these concerns.

Treatment of systemic enterococcal infections is challenging. Despite the fact that E. faecalis is susceptible to ampicillin, the high treatment failure rate associated with its use as monotherapy in addition to the loss of bactericidal activity necessitates the use of combination therapy involving bactericidal agents. The co‐administration of β‐lactams and aminoglycosides was initially used; however, its clinical efficacy was quickly compromised by the emergence of enterococcal isolates exhibiting HLAR (Beganovic et al. 2018). Currently, ampicillin and ceftriaxone dual therapy are the only available options against HLAR enterococcal bacteremia in clinical settings. However, the continuous use of this combinational therapy led to the evolution of resistance besides pronounced VRE colonisation (Beganovic et al. 2018). Therefore, more research has been performed to investigate other combinational therapies in the treatment of enterococcal bacteremia (Tang et al. 2013; Farina et al. 2012). A further instance of this is the in vitro synergy observed with the combination of fosfomycin and daptomycin against HLGR E. faecalis (9Rice, Eliopoulos, and Moellering Jr 1989). In addition, clinical trials have reported promising activity with fosfomycin and imipenem co‐therapy in the treatment of bacteremia and endocarditis (del Río et al. 2014). Accordingly, the present study investigated the possible synergy between vB_EfaS_SZ1 phage and ampicillin or fosfomycin based on their clinical relevance against enterococcal bacteremia.

In the current study, the combined use of vB_EfaS_SZ1 phage with either ampicillin or fosfomycin displayed synergistic effects that surpassed monotherapy involving vB_EfaS_SZ1 phage or antibiotics alone. One possible explanation is that using sub‐lethal concentrations of ampicillin and fosfomycin could induce cell filamentation and inhibit bacterial peptidoglycan synthesis at an early stage, respectively (Comeau et al. 2007; Silver 2017). This altered morphological state induced by sub‐lethal concentrations of antibiotics led to the accumulation of bacterial precursors, which are beneficially utilised by phage for its multiplication, leading to potential synergy (Comeau et al. 2007). Interestingly, PAS could occur regardless of whether the bacteria are sensitive or resistant to the tested antibiotic (Kamal and Dennis 2015). Based on checkerboard assay results, a time‐kill assay was conducted to measure bacterial killing over several time points.

Meanwhile, the emergence of bacterial resistance was observed with treatments utilising either the phage or antibiotics individually. There are other proposed mechanisms by which PAS may occur, including reduced emergence of phage‐resistant and antibiotic‐resistant mutants; enhanced bacterial susceptibility to antibiotics; decreased MIC of antibiotics; improved diffusion and cell penetration of antibiotics caused by phage enzymes; in addition to the increased burst size, phage proliferation, and plaque size (Łusiak‐Szelachowska et al. 2022). The current findings suggest that vB_EfaS_SZ1 phage could be utilised as a useful adjuvant to ampicillin or fosfomycin, offering a safer alternative to dual β‐lactam therapy for resistant enterococcal BSIs.

Using an in vivo model is a crucial preclinical stage in evaluating the effectiveness of phage‐antibiotic co‐therapy. The current findings revealed that all treated groups initially exhibited reduced liver bacterial load after 48 h. However, by 72 h, the combination therapy was more effective, lowering the bacterial density to a level that enabled immune clearance. The rise in bacterial burden seen with single therapy is possibly attributed to the selection of the bacterial strains that the phage could not adsorb to, leading to the emergence of resistant phenotypes or mutant strains within the host bacterial population (Hyman and Abedon 2010; Amarillas et al. 2017). This suggests that combining distinct selective pressures would work better together in limiting bacterial growth and restricting the emergence of resistance than single treatment. In addition, mice receiving combined therapy had a significantly lower phage titre in liver tissues compared to those treated with vB_EfaS_SZ1 phage alone. This phenomenon observed in combinational therapy could be attributed to the initial drop in bacterial titre, which in turn restricted the phage propagation and sped up their removal from the mice. Furthermore, the uninfected phage‐injected group showed no viable phage 72 h post‐treatment, indicating successful phage clearance and phage persistence alongside their host.

Histopathological analysis of liver tissue further supported the therapeutic benefits of PAS. The liver tissues of the mice group revealed significant pathological alterations suggesting the successful establishment of the infection model. Nevertheless, individual treatments with either antibiotic or vB_EfaS_SZ1 phage partially alleviated the liver tissue changes induced by enterococcal infection. Notably, mice receiving fosfomycin showed a partial recovery, evidenced by a reduced inflammatory infiltration and an abundance of apoptotic bodies. The mice inflammatory status within liver tissues was markedly reduced with the combination therapy, as indicated by the pronounced presence of normal hepatocytes, suggesting successful PAS. Interestingly, a recent study reported that introducing phage therapy to the antibiotic regimens temporarily suppressed recurrent E. faecium bacteremia, reduced hospitalisation, and enhanced patient's quality of life for several months (Stellfox et al. 2024). Consequently, utilising vB_EfaS_SZ1 phage in conjunction with antibiotics would provide a new horizon for combating resistant enterococcal bacteremia in clinical settings, leading to better outcomes. In spite of the insightful findings provided in the current study, there are some noteworthy limitations. Further investigation of phage‐antibiotic interactions with more emphasis on the antibiotic concentration, phage MOI, administration time, and order is required. Additionally, more antibiotics representing different classes need to be explored.

5. Conclusion

In conclusion, our study provided a detailed biological and genomic characterisation of the novel lytic phage vB_EfaS_SZ1, which targets clinical isolates of Enterococcus faecalis exhibiting high‐level gentamicin resistance (HLGR). Synergy assays demonstrated that combining vB_EfaS_SZ1 with ampicillin or fosfomycin significantly enhanced bacteriolytic activity and reduced antibiotic MIC values. In a mouse bacteremia model, these phage‐antibiotic combinations significantly reduced E. faecalis liver burden compared to single therapy of either antibiotic or phage. The phage's specificity, besides its ability to enhance antibiotic efficacy, underscores its potential to reduce bacterial load and prevent resistance development. These findings pave the way for the introduction of phage‐antibiotic co‐therapy into treatment regimens for enterococcal BSIs in clinical settings, thereby improving patient outcomes and addressing the growing challenge of resistance to either phage or antibiotic.

Author Contributions

Fatma Al‐zahraa A. Yehia: conceptualization, investigation, writing – original draft, methodology, validation, writing – review and editing, data curation, formal analysis, resources, visualization, software, project administration. Galal Yahya: conceptualization, investigation, funding acquisition, writing – original draft, writing – review and editing, visualization, validation, methodology, software, formal analysis, supervision, data curation, project administration, resources. Eslam M. Elsayed: investigation, funding acquisition, writing – review and editing, methodology, software, validation, data curation, resources, formal analysis. Javier Serrania: investigation, funding acquisition, methodology, validation, writing – review and editing, software, data curation, resources. Anke Becker: investigation, funding acquisition, methodology, validation, writing – review and editing, software, data curation, resources. Salwa E. Gomaa: conceptualization, investigation, writing – original draft, writing – review and editing, visualization, methodology, validation, software, project administration, formal analysis, data curation, resources, supervision.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1.

Figure S2.

Table S1. Host range determination of vB_EfaS_SZ1 phage.

Table S2. Detailed genome annotation of vB_EfaS_SZ1 phage.

Data Availability Statement

The data supporting this study's findings are available from the corresponding author upon reasonable request.