In recent years, an increase in the prevalence of leek bacterial blight was noted in Flanders, Belgium. The disease is caused by the bacterial pathogen Pseudomonas syringae pv. porri, which was first reported in the United Kingdom by Lelliott (1952). Two decades later, the species was isolated in New Zealand by Hale (1975) who further described the pathogen. Later, the bacteria was classified as a new pathovar, P. syringae pv. porri, based on extensive research by Samson et al. in France in 1998. Currently, the disease has spread globally and is reported in The Netherlands (Janse, 1982; van Overbeek et al., 2010), Italy (Varvaro, 1983), the United States (Koike et al., 1999), Australia (Noble et al., 2006), Greece (Glynos and Alivizatos, 2006), Japan (Goto, 1972), and Korea (Myung et al., 2011, 2012). Typical symptoms include leaf curling and yellowing of the middle vein in young plants and water soaked spots on older leaves and flowering stems (Samson et al., 1998; Noble et al., 2006). Leek (Allium porrum) is the major host, but the pathogen has also been diagnosed on onion (Allium cepa) and shallot (Allium cepa var. aggregatum; Noble et al., 2006). The disease is known to be transmitted by seed (Koike et al., 1999; Ikene et al., 2003), but crop waste also plays a role in contaminating new leek plants in the field (van Overbeek et al., 2010).

The recent rise in leek bacterial blight is consistent with a growing number of Flemish farmers obtaining leek transplants from plant nurseries. High plant densities in those nurseries, combined with plant manipulations such as irrigation and mowing, promote dissemination of the pathogen among the transplants (Koike et al., 1999). In a previous study, the causative agent of recent leek blight epidemics in Flanders was examined, leading to the proposed subdivision of the strains into two groups, with small genomic differences (Rombouts et al., 2015). Knowledge on this intra-pathovar diversity is important to determine the specificity of phages isolated and characterized within this study.

To date, disease management in leek production mainly consists of prevention. Clean seeds, a strict hygiene in plant nurseries and the use of more tolerant varieties are important control measures. However, once the first symptoms of bacterial blight appear, no solution is available because of the ban on streptomycin derivatives to control bacterial plant diseases in the European Union. The use of copper-based agrichemicals is also undesirable in view of ecotoxicity and bacterial resistance development (Cooksey, 1994; Pietrzak and McPhail, 2004; Hwang et al., 2005). Therefore, the use of phage therapy constitutes an attractive alternative. Bacteriophages are viruses that specifically infect bacteria, their replication resulting in the lysis of their bacterial host and the release of newly formed viral particles. Several reviews have previously been published, highlighting the possibilities and limitations of phage therapy in plant disease control (Gill and Abedon, 2003; Jones et al., 2007; Balogh et al., 2010). In agriculture, phage therapy research has been conducted for pathogens in other crops including "Dickeya solani" (Adriaenssens et al., 2012), Erwinia amylovora (Gill et al., 2003; Boulé et al., 2011), Pectobacterium carotovorum (Ravensdale et al., 2007), Ralstonia solanacearum (Fujiwara et al., 2011), Xanthomonas axonopodis pvs. citri and citrumelo (Balogh et al., 2008), and P. syringae pv. actinidiae (Frampton et al., 2014). To date, phage research resulted in a limited number of commercial phage-based products for agricultural use. These products are for control of two tomato pathogens, Xanthomonas campestris pv. vesicatoria and P. syringae pv. tomato (AgriPhage; Flaherty et al., 2000; Balogh et al., 2003) and against potato rot (Biolyse, APS Biocontrol Ltd.).

Phages infecting P. syringae pv. porri have not yet been described. In this manuscript, we focus on the isolation of novel phages which cover the bacterial diversity present in Flemish leek production. Extensive in vitro characterization of the phages was followed by bio-assays and field trials to evaluate the potential of these phages for the biocontrol of bacterial blight in leek.

## MATERIALS AND METHODS

## Bacterial Strains and Growth Conditions

The bacterial strains used in this study are listed in **Table 1**. P. syringae pv. porri strains were isolated from fields in Flanders and covered the existing diversity. Their identification and characterization was previously published (Rombouts et al., 2015). These isolates were supplemented with P55, an isolate from soil from the study of van Overbeek et al. (2010), and reference strains of P. syringae pv. porri from the Collection Française de Bactéries Phytopathogènes (CFBP; Beaucouzé, France), including the type strain and a set of phylogenetic related strains. Bacteria were grown in lysogeny broth with reduced salt concentration (LBrs) (0.5 g/l NaCl) at 26◦C, or on LBrs plates supplemented with 1.5% agar. LBrs with 0.7% agar (soft agar) was used for the plate overlays.

## Bacteriophage Isolation, Amplification, and Purification

Phages were isolated from soil samples taken in 2011 and 2012, from the same fields from which the P. syringae pv. porri strains were isolated. To enrich for phages, a bacterial host (LMG 28495 or LMG 28596) was grown in 25 ml LBrs at 26◦C up to exponential growth phase and 5 g of soil sample was added. After overnight incubation, 250µl chloroform was added and incubation was continued for 1 h to lyse all bacterial cells. Next, the mixture was centrifuged (30 min, 4500 rpm) in a Sorvall Legend RT+ centrifuge with swing-out 4-place rotor, type 75006445 (Thermo Scientific, Waltham, MA, USA). The supernatant was filtered through a Millex R syringe filter with a 0.45µm pore size (Merck Millipore Ltd.) and 10µl was spotted on a soft agar layer that contained the bacterial host. Lysis zones were picked up with sterile toothpicks and suspended in phage buffer (10 mM Tris-HCl; pH 7.5; 10 mM MgSO4; 150 mM NaCl). These suspensions were plated by pooling 250µl overnight bacterial host culture, 100µl phage suspension and 4 ml LBrs overlay agar, according to the overlay agar technique (Adams, 1959). After overnight incubation at 26◦C, single plaques were picked up again. Three successive single plaque isolations were performed to achieve pure phage isolates. Based on their DNA restriction pattern (data not shown) different phages were selected for further analysis.

One bacteriophage (KIL3b) was obtained after a co-cultivation experiment. This experiment consisted of the infection of an

The two symbols (• and ×) indicate two BOX-PCR pattern groups. The arrow gives the position of the discriminative band. Light and dark green: lysis after 24 and 48 h, respectively and yellow: lysis from without. aGBBC: culture collection of plant pathogenic bacteria at ILVO; CFBP, Collection Française de Bactéries Phytopathogènes; INRA Angers; LMG, Belgian Coordinated Collections of Microorganisms at the Laboratory of Microbiology of Ghent University with T as type strains and PT as pathovar reference strains. bGeographical origin. cYear of isolation. d received from van Overbeek et al. (2010).

TABLE 2 | Bacterial host of the new bacteriophages and phage genome characteristics determined by bioinformatic analysis.

exponentially growing liquid culture of P. syringae pv. porristrain CFBP 1687 with phage KIL3 at a multiplicity of infection (MOI) of 0.01. For 4 weeks, 40µl of the liquid culture was transferred to a tube with 4 ml fresh LBrs medium every 2 to 3 days and incubated at room temperature. Supernatant of every two-dayold liquid culture was spot tested against the bacterial strain P55 to select for a broadened lytic activity as none of the other phages were able to lyse this strain.

Phages were amplified by plating 105 plaque forming units (pfu) per plate together with their bacterial host on a soft agar overlay plate. After overnight incubation at 26◦C, the soft agar was scraped off and suspended in phage buffer. Soft agar and cell debris were subsequently removed by centrifugation (20 min, 4000 rpm) in a Sorvall Legend RT+ centrifuge. The supernatant was filtered over a 0.22µm membrane Millex R syringe filter (Merck Millipore Ltd.), after which polyethylene glycol (PEG) 8000 (Sigma-Aldrich) containing 1 M NaCl was added to the suspension to a final concentration of 8%w/v. After minimum 3 h incubation at 4◦C, phages were precipitated by centrifugation (30 min, 4600 rpm) and the pellet was dissolved in 1 ml phage buffer.

Phage purification was performed using CsCl-gradient ultracentrifugation. A phage suspension (15 ml) with a minimum concentration of 1010 pfu/ml was layered on a CsCl step gradient (1.33, 1.45, 1.50, and 1.70 CsCl g/cm3 ) and centrifuged (3 h, 140.000 × g) with a Beckman Coulter Optima L-90K ultracentrifuge (rotor type SW28) (Brea, CA, USA). All phages were collected at the interface between the 1.45 and 1.50 g/cm3 density layer and dialyzed three times for 30 min against 300 volumes of phage buffer.

### Electron Microscopy

To obtain transmission electron microscopy (TEM) images of the bacteriophages, dilutions of the samples were spotted on carbon coated grids (Quantifoil, Großlöbichau, Germany) after glow-discharge, and negatively stained with 2% uranyl acetate. A Philips CM12 microscope was used at 120 kV acceleration voltage. Images were produced using a Gatan Orius 1 k camera.

## Host Range Analysis and General Characterization

Phage host range was tested by spotting a phage suspension on plates with a soft agar layer supplied with a specific bacterial isolate. For all the phages, three different concentrations, ranging from 102 to 106 pfu/ml, were tested to determine infectivity. The bacterial strains used in the host range assay are listed in **Table 1**.

Adsorption experiments were performed according to Adriaenssens et al. (2012). Briefly, bacteria and phage were mixed at an MOI of 0.0001. Next, samples were taken every minute and bacteria were immediately lysed by adding chloroform. The upper phase was titred to determine the number of non-adsorbed phages.

Killing curves at different MOIs were generated by infecting a bacterial culture at an optical density at 600 nm (OD600) of 0.3. The OD600 of infected and uninfected cultures was monitored every 20 min during 6 h. OD600 results were the average of three independent biological repeats.

Phage stability was tested by incubating a phage suspension of 106 pfu/ml in phage buffer at different temperatures (−20, 4, 16, 30, 37, and 50◦C) and at different pH ranging from 1 to 13 (universal buffer consisting of 150 mM KCl, Janssen Chimica, Geel, Belgium; 10 mM KH2PO4, VWR International, Leuven, Belgium; 10 mM sodium citrate, Acros Organics; 10 mM H3BO3, Acros Organics; adjusted to pH 1–13 with NaOH, or HCl). Infectivity of the phages at different temperatures was tested by incubating soft agar plates with spots of different phage concentrations at room temperature (±21◦C), 26 and 30◦C and counting the infective phage titers.

The frequency of bacterial resistance was determined using an adaptation of a previously developed method (Beale, 1948). Using the agar overlay method, phage and bacteria were plated at MOI 1 and MOI 10, in order to lyse all bacteria on the plate. After 72 h of incubation, emerging colonies were counted and about 20 colonies were cultured and re-tested for resistance against the phages in **Table 2**. Phage-resistant isolates were plated on Pseudomonas agar F (PAF, Becton Dickinson) supplemented with glycerol to check for fluorescence under UV-light and analyzed with a multiplex PCR containing specific primers (Ineke Wijkamp, Rijkzwaan, personal communication) to confirm their P. syringae pv. porri identity.

Lysogeny in these resistant bacteria was tested as previously described (Petty et al., 2006). Supernatant from overnight cultures of resistant bacteria were collected and tested for the spontaneous release of phage particles by spotting 5µl drops onto soft agar containing the different bacterial hosts. The supernatants from CFBP 1687 and LMG 28496 were used as negative controls, and pure phage solutions (106 pfu/ml) were used as positive controls for plaque formation.

## Genome and Proteome Analysis

### DNA Isolation and Sequencing

Contaminating bacterial DNA was removed from phage lysates by DNase treatment. Upon inactivation of the DNase by the addition of EDTA, phage particles were disrupted by adding SDS and proteinase K. The released phage DNA was purified by standard phenol-chloroform extraction and subsequent ethanol precipitation according to Sambrook and Russell (2001). DNA concentration and purity was verified with the NanoDrop ND-1000 spectrophotometer (Thermo Fisher Scientific Inc.). DNA integrity and genome size was evaluated on an 0.8% agarose gel. High throughput sequencing was done using the Illumina MiSeq platform. A 2∗ 250 bp paired-end library was prepared for each sample, tagged with a unique adapter sequence. The quality of each library preparation was controlled using the Agilent Bioanalyzer. All library preps were equally pooled and sequenced. After processing, the reads of every library prep were assembled in a single contig with a general coverage over 1000x using the CLC Bio Genomics Workbench de novo assembly algorithm (version 7.5.1) (Aarhus, Denmark).

### "In silico" Analysis

The genomes were scanned for open reading frames (ORFs) with GeneMark.hmm and GeneMarkS software (Lukashin and Borodovsky, 1998; Besemer et al., 2001). Shine-Dalgarno sequences were verified manually upstream of each annotated ORF. Functional annotation was carried out by comparing translated ORFs in a Blastp analysis (Altschul et al., 2005) against the nonredundant GenBank protein database and with the Protein Homology/analogY Recognition Engine v2.0 (PHYRE2) (Kelley and Sternberg, 2009). tRNAs were detected with the programs ARAGORN and tRNAscanSE (Lowe and Eddy, 1997; Laslett and Canback, 2004). Rho-independent terminators were predicted with ARNold, a search program that combines two complementary programs: RNAMotif and Erpin (Gautheret and Lambert, 2001; Macke et al., 2001). Probable promoter sequences were identified by looking for conserved intergenic motifs in the 100 bp upstream of every ORF with MEME and PHIRE (Lavigne et al., 2004; Bailey et al., 2009).

### Protein Clustering and Phylogenetic Analysis

Homologous proteins, deduced from their DNA sequences, were examined in the phages KIL1, KIL2, KIL3, KIL4, KIL5, KIL3b, and a number of fully sequenced bacteriophages using HMMER (version3.0; http://hmmer.janelia.org), together with the ACLAME search (version0.4; http://aclame.ulb.ac.be; as of June 2015). In this manner, we retrieved a total of 4632 predicted protein sequences represented in the genomes of the 26 relevant phages (Table S2). We performed pairwise similarity comparisons for each predicted protein by using the ACLAME database with the database of "viruses" and an E-value < 0.001 (Lima-Mendez et al., 2008). The remaining proteins which could not be assigned into any ACLAME protein family were defined as the unclassified protein families (UPFs) as previously described (Jang et al., 2013). The protein families that were common to all 32 representative phage genomes were identified and their phylogenetic relationships were inferred. After multiple sequence alignment of the conserved proteins using CLUSTALW2, noninformative positions were excluded with the BMGE software package (Criscuolo and Gribaldo, 2010). The two alignments were then concatenated into a FASTA file and a phylogenetic tree was built with MEGA5 (version 5.2.1; http://www.megasoftware. net/; Tamura et al., 2011) using a Jones–Taylor–Thornton (JTT) model. To estimate the robustness of the trees, we used the maximum-likelihood algorithm provided with bootstrap support (n = 1000 replicates).

### Proteome Analysis

Structural proteins of phages KIL3 and KIL5 were identified by SDS-PAGE gel electrophoresis, isolation of gel bands, and subsequent trypsinization and ESI-MS/MS as described previously (Lavigne et al., 2009a). The resulting data were analyzed using Mascot (version 2.3.01) and Sequest (version <IP_ADDRESS>), against a database containing all possible ORFs based on the DNA sequence of the phages.

# Phage Therapy Experiments

### Bio-Assay

In planta activity of the phages was tested by injecting phage and bacterial suspensions into leek leaves. Leek plants of the disease susceptible cultivar Krypton (Nunhems) were grown in separate pots until full development of three leaves. Next, 0.1 ml of bacterial suspension was injected with a syringe into the middle of the leaf. Subsequently, 3 cm above the bacterial injection place 0.1 ml of phage suspension (109 pfu/ml) was injected (Supplementary Figure 1). In the first bio-assay all 41 P. syringae pv. porri strains (107 CFU/ml) were inoculated in leek leaves (three inoculated leaves/strain) and all phage types (109 pfu/ml) were co-inoculated with a P. syringae pv. porri representative (eight inoculations/phage-bacteria combination). This experiment was repeated using the disease susceptible cultivar Striker (Bejo) and a lower bacterial concentration (106 CFU/ml). Phages KIL1, KIL2, KIL3, and KIL3b (109 pfu/ml) were co-inoculated with bacterial strain CFBP 1687 and phages KIL4 and KIL5 with LMG 28496 (10 inoculations/phage-bacteria combination). Plants were covered for 48 h with a plastic bag to maintain humid conditions and kept at a temperature of 25◦C. Lesion lengths were measured after 10 days. Injections with only bacteria, phage or buffer served as controls.

### Field Trials

In a first trial, leek transplants were treated with phages before they were planted in an infested field. First, the field was infested by spraying a solution of CFBP 1687 with a concentration of 106 CFU/ml on the soil at a rate of 1000 l/ha, at 1.5 bar and with 0.01% Tween R 20 as surfactant. The next day, before planting, leek plants were submerged in a solution containing a mixture of the six different phages, each at a concentration of 107 pfu/ml. Control plants were planted without treatment. Four blocks of the phage-treated and non-treated plants were randomly distributed over the field, with each block containing 300 plants. At multiple time points after planting, disease incidence (number of damaged plants) as well as disease severity (% of leaf surface affected) was measured for 20–50 plants per block. This trial was performed simultaneously on three different locations in Flanders: Kruishoutem, PCG (N 50.94337◦ , E 3.52710◦ ), Sint-Katelijne Waver, PSKW (N 51.078120◦ , E 4.528180◦ ), and Beitem, Inagro (N 50.901508◦ , E 3.124464◦ ).

In a second trial, leek transplants were planted in a noninfested field. Two months after planting, bacterial infection was performed with strain CFBP 1687. A suspension with a concentration of 106 CFU/ml was spread over the plants at a rate of 1000 l/ha, at 1.5 bar and with 0.01% Tween R 20 as surfactant. The next day, a mix of the six different phages, each at a concentration of 109 pfu/ml and supplemented with 0.01% Tween R 20, was sprayed over the plants, in a way that each plant was covered with on average 108 pfu of each phage after spraying. Plants were covered with plastic for 48 h and assessed as described. This trial was also performed in parallel on the three different locations in Flanders mentioned above.

### Statistical Analysis of Data

Statistical analyses were performed with IBM SPSS Statistics 20. For the bio-assays, normality of the data was assessed with the Kolmogorov-Smirnov and Shapiro-Wilk test at a significance level of 0.05. Homogeneity of variances was tested using the Levene's test. When the data were distributed normally and homoscedasticity was proven, an one-way ANOVA test was performed to check for differences. If the tests showed significant differences, the data were further analyzed with a post-hoc Tukey HSD test. To test for differences between groups of not normally-distributed data, a Kruskal-Wallis non-parametric test was used. When differences were present, Mann-Whitney U nonparametric test was used to compare the groups, paired two by two. For the field trials, the statistical program ARM (version 2015.3) was used. This program, which is designed specifically for field trials, uses the Bartlett's test to test the homogeneity of variances and the Kurtosis and Skewness tests to check normality. One-way ANOVA with the post-hoc Duncan test was used for normal distributed and homoscedastic data. As a non-parametric test, the Friedman test was applied to check for differences between several variables.

### RESULTS

### Phage Isolation and TEM

Phages were isolated from soil samples taken in 2011 and 2012 from the same fields from which P. syringae pv. porri isolates were obtained (**Table 2**). After enrichment of the phages present in the soil, their capacity to lyse P. syringae pv. porri strain CFBP 1687 was tested. Based on their differential DNA restriction pattern (data not shown) phages were selected for further analysis. By this method, five novel phages were discovered for P. syringae pv. porri and the phages were named KIL1, KIL2, KIL3, KIL4, and KIL5 (referring to the involved research institutes KU Leuven and ILVO) with scientific names vB\_PsyM\_KIL1 etc. as proposed by Kropinski et al. (2009). The host strains used for amplification and characterization of each of these phages are indicated in **Table 2**.

TEM images of a phage representative revealed that they belong to the Myoviridae family with morphological similarity

to the Pseudomonas phage PB1 (genus Pbunalikevirus), as shown for phage KIL1 in **Figure 1**. The other analyzed phages have an identical morphology.

### Host Range Analysis

To investigate the specificity of the phages, a host range analysis was performed (**Table 1**), by using standard serial dilution spotting assays to distinguish infection from lysis from without phenomena, as described previously (Adriaenssens et al., 2012). Some of the bacterial isolates could not be infected by our five phage isolates from soil, therefore the host-range mutant phage KIL3b was developed. The results indicate that the bacterial strains can be divided in three groups. One group consists of five field isolates from the year 2012 and was only infected by phages KIL4 and KIL5. A second group of strains was infected by all the investigated phages and a third group of strains only showed lysis after 48 h and by a limited number of phages. None of the pathovars related to P. syringae pv. porri was infected by the phages demonstrating their specificity for pathovar porri. However, P. syringae pv. garcae (CFBP 1634) showed "lysis from without" at phage concentrations of 106 pfu/ml. In addition, none of the phages infected all P. syringae pv. porri strains tested but the combined host range of the phages covers all 41 isolates tested. A previous BOX-PCR analysis on DNA of all the bacterial strains used in the host range analysis provided information about their genomic diversity and identified two groups within P. syringae pv. porri (Rombouts et al., 2015). These two groups were confirmed by phage profiling, suggesting that differences in the bacterial genome are responsible for the observed host range. Two phages, KIL3 and KIL5 were selected for further characterization.

### General Characteristics

To assess the infection parameters, adsorption experiments were performed for phage KIL3. On average, 84% of phages were irreversibly adsorbed to the host cell after 1 min and 99% after 6 min with an adsorption constant k [k = (2.3/(B∗ t))∗ log(P0/P), with B the bacterial titer at time zero, P the phage titer and t the time] at 1 min of 7.52 × 10−9 ml/min. Compared to other Myoviridae phages, this is slower than reported for phage T4 (2.4 × 10−9 ml/min) but comparable to the adsorption constants of phages LIMEstone1 (9.53 × 10−9 ml/min) and LIMEstone2 (2.05 × 10−8 ml/min) infecting Dickeya (Kasman et al., 2002; Adriaenssens et al., 2012).

Killing curves generated by infecting an exponentially growing cultures of P. syringae pv. porri CFBP 1687 and LMG 28496 with phages KIL3 and KIL5, respectively, at different MOI demonstrated their virulence (Supplementary Figure 2). Optical density of the bacterial cultures at 600 nm decreased after 80 and 100 min for phage KIL3 and KIL5, respectively. KIL5 showed a later but steeper decline reaching OD600 < 0.1 after 180 min instead of 220 min for KIL3. No rise in OD600 appeared within the 5 h of monitoring indicating that resistance did not develop at that point.

The influence of temperature and pH on the viability was tested. As shown for KIL3 and KIL5 (representatives of the two phage clades in the cocktail), the phages were stable between 4 and 37◦C in phage buffer for 24 h, but after 24 h of incubation at 50◦C, a two and one log10 unit decrease was noted for phage KIL3 and KIL5, respectively. After freezing, all viable KIL3 phages were lost, while the titer of KIL5 decreased by three log10 units. Phages were stable from pH 4 to 12 for 24 h.

An analysis of the optimal infection temperature revealed that none of the phages are able to infect their host when grown at temperatures of 30◦C. Phages KIL1, KIL2, KIL3, KIL4, and KIL5 were able to infect at temperatures of 26◦C and room temperature (±21◦C), and KIL3b only infected at room temperature.

For two phage-host interactions, frequency of bacterial resistance development was determined. An exponentially growing culture of strain CFBP 1687 was infected with KIL3 at MOI 1 and after 72 h incubation, an average of 110 phageresistant bacteria appeared on plate. To exclude the possibility of contamination, the identity of 12 colonies was verified with PCR-analysis using specific primers, confirming their identity as P. syringae pv. porri. This resulted in a resistance frequency of 1.83 × 10−6 . A host screen with those resistant bacterial derivatives revealed that five of them were resistant to the six phages, a phenomenon called cross-resistance. The other isolates could still be infected by the host-range mutant phage KIL3b. The same experiment was performed with phage KIL4. After 72 h incubation on average 200 resistant colonies appeared and PCRanalysis was used to confirm the identity of 12 of them, resulting in a calculated resistance frequency of 3.33 × 10−6 . Again, crossresistance was observed for part of the strains (four out of 12), the other resistant strains were still infected by KIL3b.

A spot test with supernatant from an overnight culture of those resistant isolates on strains CFBP 1687 and LMG 28496 suggested there was no induction of lysogens under the conditions tested. In general, as all phages produced clear plaques there was no indication for lysogeny.

### Genome and Proteome Analysis Genome Analysis

The genomes of all six phages were sequenced using the Illumina MiSeq platform and single contigs were obtained. Information about the genome characteristics of the phages is summarized in **Table 2**.

ORFs were identified and similarity at the protein level was verified by Blastp analysis. In addition, a putative function could be assigned to some ORFs without homologs using PHYRE2. All tRNAs were found in the same region spanning 2.7 kb in front of the large terminase subunit, a part of the packaging machinery (**Figure 2**). This is consistent with the genome organization of the related PAK\_P1-like phages (Debarbieux et al., 2010; Henry et al., 2015). When comparing the tRNAs present in the six genomes, phages KIL1, KIL3, and KIL3b encode the same five tRNAs. KIL2, KIL4, and KIL5 also have these five tRNAs in their genomes, supplemented with four additional tRNAs. Depending on the phage, 17 or 18 rho-independent terminators were retained after manual verification and were located across the entire genome. Probable promoter sequences were identified by looking for conserved motifs in the 100 bp upstream region of every ORF. Using MEME, a motif resembling a typical E. coli bacterial promoter sequence (TTGACA-N17−18-TATAAT) was found in all phages. As in the most closely related P. syringae phage PhiPsa374, the 41 bp conserved motif surrounding the bacterial promotor sequences in PAK\_P1-like viruses was not encountered thereby confirming its specificity for P. aeruginosa phages (Henry et al., 2015). No toxin genes, virulence genes or genes related to lysogeny were discovered in the phage genomes, indicating their suitability for phage therapy.

A remarkable feature of all KIL-like phages is the presence of a homopolymeric G-stretch of variable length at one or two locations in the genome (Table S1). Illumina reads at those positions contain 6–13 Gs. Both stretches are found in intergenic regions, yet not clearly associated to known promoter elements. A similar feature was reported previously in PB1-like phages, in a sequence coding for a baseplate protein and in the ORF coding for a DNA adenine methylase in Bordetella phages BPP-1, BIP-1, and BMP-1 (Liu et al., 2004; Ceyssens et al., 2009). It was proposed that the resulting frameshift serves as a control point for expression levels of the protein. Another remarkable region in the KIL-like phage genomes is a long Astretch varying from 14 to 33 nucleic acids in length. In the most related phage phiPsa374 those homopolymeric stretches are not present. A phylogenetic analysis of the six phages based on their genome sequences confirms the subdivision of the phages into two groups, correlated to their host range. The first group constitutes of phage KIL1, KIL2, KIL3, and KIL3b, the second group contains phages KIL4 and KIL5. Between the groups, some differences in the ORFs are predicted, mostly among unknown genes. The difference between the genomes of phage KIL3 and KIL3b, which is a host range mutant of the former, is limited to the number of repeats located in the homopolymeric A- and Gstretches. No clear biological function could be attributed to this phenomenon.

### The Phylogenetic and Proteome-Based Relationships

To further frame these phages in their comparative genomic context, phylogenetic trees were constructed based on the concatenated datasets of two structural genes encoding the major capsid protein and the baseplate protein. These two proteins

are shared among the genomes of four clades: the FelixO1 like viruses, PAK\_P1-like viruses, and KPP10-like viruses within the Felixounalikevirus genus and the rV5-like viruses, as well as phages CR3, ICP1, and our six phage isolates. Each protein was assigned to "Family:virus:314" and "Family:virus:3622" by ACLAME-based clustering (Table S2). Phylogenetic analysis revealed the four clades comprising the KPP10-, PAK\_P1-, FelixO1, and rV5-like phages, respectively (Left, **Figure 3**). Most clades branch deeply and are well supported with bootstrap support values of more than 90%. Within the Felixounalikevirus genus, the Pseudomonas phages KIL1, KIL2, KIL3, KIL4, KIL5, and KIL3b are grouped into a distinct clade, together with a Pseudomonas phage phiPsa374.

To investigate the genome relatedness based on the proportion of conserved homologous proteins between two phages, we performed pairwise comparisons of the 32 phage genomes, which allowed us to assign 5622 protein sequences to 1322 homologous protein families (Table S1). A cut-off value of 40% of homologous proteins was used to assign phage genomes to a putative genus (Lavigne et al., 2008, 2009b). The pattern of proteome-based grouping was highly similar to our phylogenetic results, in which KIL1, KIL2, KIL3, KIL4, KILP5, KIL3b, and phiPsa374 form a conserved group with ≥50% homologous proteins in common to each other's genome (Right, **Figure 3**). These seven phages are also related to three groups of the Felixounalikevirus phages, sharing more than 20% of their homologous proteins, with the PAK\_P1-like phages being the closest relatives (average 34% shared proteins). Notably, of these, phiPsa374 shared the highest overlap percentages of the proteomic content (≥40%) with the PAK\_P1-like phages.

FIGURE 3 | Phylogenetic and proteome-based relationships of phages KIL1, KIL2, KIL3, KIL4, KIL5, and KIL3b. Left, maximum-likelihood tree was constructed upon a concatenation of two structural proteins (major capsid protein and baseplate protein) that are common to the 32 phages. The numbers at the branch represent the bootstrap values. Edges with bootstrap values above 75% are represented. The scale bar indicates the number of substitution per site. Right, in the matrix view, the columns and rows correspond to the phages. Each cell indicates the percentage of shared protein families between two phages. Color tags are used to visualize the genomic relatedness based on the proportion of shared protein families.

### Proteome Analysis

Analysis of the proteome of phages KIL3 and KIL5 (representatives of the two clades) by ESI-MS/MS led to the identification of 51 and 67 proteins, respectively (Table S3). For KIL3 and KIL5, respectively 27 proteins and 26 had a predicted function based on homology to other phage proteins. In both phages, most of the detected proteins are early phage proteins or structural phage proteins. The use of non CsCl-purified phages might explain the presence of the non-structural phage proteins since they can remain in the phage lysate after bacterial lysis. Alternately, these proteins may be considered as candidate proteins which may be co-injected during infection. The most abundant proteins based on the number of unique peptides recovered are the tape measure protein, ORF 46 (no predicted function) and the major capsid protein in phage KIL3. Similarly, the most abundant proteins of phage KIL5 are the capsid protein, ORF 46 (no predicted function) and the tail fiber protein (ORF 76). As was seen by Henry et al. (2015) for the related phage PAK\_P3, the ORF upstream of the capsid protein had the second highest relative abundance (total number of spectra / Molecular Weight) next to the capsid protein itself, and this in both phages. Although the function of this protein could not be predicted, an association with the capsid protein is suspected. The differences in the number of proteins identified for the two phages could be explained by a difference in protein concentration as the number of peptides recovered for proteins of KIL5 were in general higher than those for KIL3. The identification of these peptides by mass spectrometry confirms the in silico ORF predictions. One structural protein, a tail lysozyme (ORF 70), could not be detected in KIL3; however, the corresponding protein in KIL5 (ORF 71) was identified. All other predicted structural proteins could be experimentally confirmed by mass spectrometry.

## Phage Therapy

### Bio-Assay on Leek Leaves

To investigate whether the phages were also capable of lysing their bacterial host in the plant environment, bio-assays were performed on leek leaves.

In a first bio-assay, the bacterial concentration was chosen to be 107 CFU/ml, the amount of bacteria necessary to produce clear symptoms. For the phages, a relatively high concentration of 109 pfu/ml was used to prove efficacy. All 41 P. syringae pv. porri strains were inoculated and showed comparable virulence, presented on the left side of the boxplot (**Figure 4**). Strains CFBP 1687 and CFBP 1770 were chosen as representatives of P. syringae pv. porri and were used as bacterial host in combination with the different phages. Lesion lengths on the phage treated leaves were compared to the lesion lengths induced by all P. syringae pv. porristrains used in this study. In general, each phage reduced the mean lesion length compared to the non-phage treated leaves. The reduction in lesion length was significant for phages KIL1, KIL3, and KIL3b with p-values of 0.003, 0.005, and 0.004, respectively. Phages KIL2, KIL4, and KIL5 reduced the lesion length, but were not able to prevent bacterial infection completely.

In a second bio-assay, the effect of the phages on a lower bacterial concentration of 106 CFU/ml was tested using ten leaves per phage. CFBP 1687 was used in combination with phages KIL1, KIL2, KIL3, and KIL3b, phages KIL4 and KIL5 were co-inoculated with bacterial strain LMG 28496. Again, lesion lengths of the phage-treated leaves were compared to the lengths produced by their respective host without phage. As expected, lesion lengths were smaller in all treatments compared to the previous assay with higher bacterial concentrations. Only phages KIL2 and KIL3b decreased lesion length significantly compared to their bacterial control, with p-values of respectively 0.042 and 0.006. Leaves treated with phage KIL3b showed no symptoms for the most part. The results of this second bio-assay confirm that phages KIL4 and KIL5 are less suitable to reduce lesion length significantly. Only phage KIL3b showed significant reduction in both assays, demonstrating its in planta antibacterial effect. The effect of phages KIL1, KIL2, and KIL3 varied between the assays. Nevertheless, it was considered prudent to maintain all phages in a cocktail for subsequent testing in field trials.

### Field Trials

In a first field trial set-up, the potential of phages to protect leek transplants against bacterial infection at planting was tested. Transplants were submerged in a phage solution containing the six phages before planting them in an infested field. This experiment was performed on three different locations in Flanders in the year 2014 (**Table 3**).

In the trial at PCG, leek plants of cultivar Kenton (Nunhems) were used. The first symptoms developed 25 days after planting in phage-treated and non-treated plants. Two months later, percentages of diseased plants increased to 82% for the phagetreated plants and 92.5% for the control plants. Later on, symptoms disappeared again in both groups and percentages of diseased plants were again lower 4 months after planting. The same trend was apparent when scoring the percentage of leaf surface affected.

The same experiment was performed in PSKW with cultivar Harston (Nunhems). Two and three months after planting, the disease incidence was measured. Percentages of symptomatic plants varied from 29% for the phage-treated plants and 19% for the control plants after 2 months to 18 and 15% after 3 months, respectively. No significant differences were observed.

In the experiment at Inagro, leek plants of cultivar Harston (Nunhems) were used. No disease incidence was recorded 2 months after planting. After 3 months, measurements reported an infection rate of 23.5% of control plants and 18% for the phage treated plants.

In all three experiments, no significant differences in disease occurrence between treated and untreated plants could be noticed. In the trials performed at PSKW and Inagro, disease incidence remained low indicating that infection was not successful. Conclusions on the efficacy of phage treatment could therefore not be made. From the trial results of PCG we can conclude that phage treatment was not successful in protecting the leek plants from infection when planted in an infested field. However, monitoring at an earlier time-point could be better to detect an effect of the phages. It is also possible that more frequent phage applications are necessary to protect the plants against infection.

In a second field trial, phages were sprayed on artificially infected plants, to test for their potential as a crop protection agent in a later growth stage. First, bacteria were sprayed onto the plants 3 months after planting. The day after, a phage cocktail with the six phages present in a concentration of 109 pfu/ml each was sprayed onto the plants. Results are shown in **Table 4**.

In the trial at PCG, again cultivar Kenton (Nunhems) was used and phage treatment was repeated four times every 2 weeks. The first disease assessment was performed 19 days after infection and demonstrated a higher infection rate at the bacteria treated plant in relation to the other plants, meaning that infection was successful. One month after infection, infection rate in the bacteria-treated plants increased and a difference trend could be measured for the phage-treated object in comparison with the non-treated infected object with, respectively, 38.5 and 63% symptomatic plants. In subsequent measurements this trend continued although the differences became less apparent. The same trend was demonstrated by the data displaying the affected leaf surface. At all time-points, infection was slightly higher in the phage-treated control compared to the untreated plants. The trial demonstrated that phages could decrease symptom development but were not able to completely stop infection. A higher phage concentration and more frequent applications can possibly lead to a better disease control.

The same trial was performed at PSKW with plants of the cultivar Harston (Nunhems). Phage treatments were applied every week for 8 weeks with half the dose of phages compared to the trial of Kruishoutem, meaning that a concentration of 0.5 × 109 pfu/ml of each phage was used. Disease incidence was measured one, 2 and 3 months after infection but no differences between the plants could be measured. Even the untreated and infected plants showed the same level of symptoms, indicating

(B) Lesion lengths in the second bio-assay in which KIL1, KIL2, KIL3, and KIL3b (109 pfu/ml) were co-inoculated with bacterial strain CFBP 1687 (106 CFU/ml) and phages KIL4 and KIL5 with LMG 28496 (10 inoculations/phage-bacteria combination).

TABLE 3 | Percentage symptomatic plants after 3 months for the field trial at three different locations where 1200 transplants were submerged in phage solution (107 pfu/ml) before planting in a field infested with CFBP 1687.

the possibility that symptoms were masked by natural infection with another pathogen.

The trial performed at Inagro used plants of the cultivar Harston (Nunhems). Five phage applications were carried out: one a week before infection, one the evening of the day during which bacterial infection was performed, and three more weekly applications after infection. Two months after infection disease incidence was measured showing no differences between the four treatments. Even the uninfected and untreated control and the uninfected phage-treated control displayed an infection of respectively 19.5 and 24% indicating the presence of a natural infection that could not be affected by our phages.

Results of the three trials demonstrate that only at PCG conducive conditions for infection with P. syringae pv. porri were present. From that trial we can conclude that phage application can lead to a reduction in bacterial disease incidence but cannot Rombouts et al. Novel Bacteriophages against Leek Blight

TABLE 4 | Percentage symptomatic plants after 1 (PCG and PSKW) and 2 months (Inagro) for the field trial with *P. syringae* pv. *porri* CFBP 1687 (106 CFU/ml) and the phage cocktail (109 pfu/ml) being sprayed over 1000 leek plants per treatment.

completely prevent it. Other factors such as natural infections with other pathogens make it difficult to predict the effect and reliability of phage therapy in field trials.

### DISCUSSION

The increase of bacterial blight of leek and the related economical losses attracted our interest in the problem a few years ago leading to an extensive investigation of the causative agent, P. syringae pv. porri. Previous research divided the Flemish isolates into two groups based on their BOX-PCR fingerprints (Rombouts et al., 2015). Genomic differences among P. syringae pv. porri strains were also described by Noble et al. (2006), who showed minor differences based on profiles generated with IS50-primer amplification and RFLP of a 16S rDNA fragment. In contrast, these authors as well as Koike et al. (1999) and van Overbeek et al. (2010) did not discriminate different genotypes among their isolates from California, Australia and the Netherland when applying different repetitive-element-PCRs (BOX, REP, and ERIC). This overall bacterial diversity was kept in mind when looking for novel phages capable of lysing P. syringae pv. porri. Host range analysis of the five phages isolated from soil and the H-mutant phage confirmed the BOX-PCR grouping of bacterial strains, suggesting that differences at the bacterial genome are responsible for the observed host range. Previous genome analysis of a bacterial strain of each group demonstrated that variation was mostly situated in prophage content and mobile genetic elements (Rombouts et al., 2015). No specific element could be found that linked bacterial diversity with the phage host range. In general, genomic differences can affect phage infection at different stages. If those genome differences result in an altered phage receptor, different receptor binding proteins (RBP) capable of recognizing the altered receptor of the five bacterial strains in group one could be present among the phages KIL4 and KIL5. Changes in host range due to a spontaneous mutation in a tail fiber protein (replacement of a positively charged lysine by an uncharged asparagine) were previously reported for two related phages, Pseudomonas aeruginosa phages PaP1 and JG004 (Le et al., 2013). However, the exact role of the variation and mutation observed here does not lead to a straightforward hypothesis and requires further investigation. In spite of the overall bacteriophage diversity, all phages we found against P. syringae pv. porri belong to the Myoviridae family. Genome sequence analysis further classified them into a new clade within the Felixounavirinae genus.

The phage characteristics analyzed in this study show that all six phages are suitable for use in phage therapy. First, they are strictly virulent, meaning that infection with the phages results in bacterial lysis minimizing the risk of horizontal transfer of pathogenicity genes as is the risk with lysogenic phages (Penadés et al., 2015). Furthermore, by demonstrating a temperature stability from 4 to 37◦C and pH stability from pH 4 to 12, they are capable of surviving in the plant environment when applied. However, a disadvantage is their limitation to infect only bacteria grown at temperatures of 28◦C or lower. This means that in summer, when temperatures occasionally rise above 28◦C bacteria may not be killed by the phages. Probably, the bacterial receptor necessary for phage infection is not expressed at higher temperatures. Temperature dependent infection was previously reported for phage ϕS1 which is not able to infect Pseudomonas fluorescens cells grown at 37◦C (Sillankorva et al., 2004). It has been shown that expression of bacterial genes can be temperature dependent. For example, the expression of flagellar components and the surfactant syringafactin is reduced in P. syringae strain B728a at a temperature of 30◦C, resulting in reduced swimming and swarming motility (Hockett et al., 2013). Host range analysis demonstrates that none of the phages is capable of lysing all tested P. syringae pv. porri strains, but a combination of different phages in a phage cocktail can cover all strains used. The results indicate that there is some resistance development among P. syringae pv. porri strain CFBP 1687 when challenged with the bacteriophages KIL3 or KIL4. Although the kill curves indicate that in the first 5 h there is no resistance development, after 72 h an average resistance frequency of around 2×10−6 was noted. When phages are applied in a phage cocktail, this resistance development is less likely to occur (Gill and Hyman, 2010; Barbosa et al., 2013). The possibility of resistance through lysogenic conversion was excluded, thereby confirming their suitability for phage therapy for this parameter. In addition, development of the host-range mutant phage KIL3b demonstrates that host-range expanding adaptions are possible to counter bacterial resistance, especially because this phage was still able to infect more than half of the resistant bacteria.

To investigate evolutionary relationships of KIL1, KIL2, KIL3, KIL4, and KIL5 with their close relatives, phylogenetic analysis was combined with a whole genome proteomic approach (Lavigne et al., 2008, 2009b). The overall phylogenetic relationships between the concatenated sequences of the major capsid protein and the baseplate proteins of the Felixounalikevirus phages are consistent with previous taxonomic description (Henry et al., 2015). The phages KIL1, KIL2, KIL3, KIL4, and KIL5 as well as phiPsa374 can clearly be grouped into a new phylogenetic clade. Their location within the Felixounalikevirus genus suggests that the sequences of two structural proteins are highly conserved within their own group and appear to be most closely related to those of the "PAK\_P1-like viruses" and "KPP1-like viruses." In addition, our proteome-based analysis, which is more sensitive and accurate for taxonomic classification for phage genomes (Lavigne et al., 2008), supports the phylogenetic relationships of the Felixounalikevirus phages. Given that shared homologs between the phage genomes can be used as an indication of phage members in the same genus (Lavigne et al., 2008, 2009b), we proposed a possible division of the Felixounalikevirus genus into the four subclades, which include the "KPP10-like viruses" (phages PAO1\_Ab17, PAO1\_Ab03, PAK\_P5, KPP10, CHA\_P1, P3-CHA, and PAK\_P3), "PAK\_P1-like viruses" (PAK\_P4, C2- 10\_Ab1, C2-10\_Ab02, C2-10\_Ab-8, PAK\_P1, PaP1, and JG004), "FelixO1-like viruses" (FelixO1, wV8, phiEa21-4, and phiEa104), and "KIL-like viruses" (KIL1, KIL2, KIL3, KIL4, KILP5, KIL3b, and phiPsa374). Interestingly, although phage phiPsa374 has been suggested as a member of the PAK\_P1-like clade (Frampton et al., 2014; Henry et al., 2015), our results indicate the genome of phiPsa374 aligns with more shared protein families to the KIL-like phages.

No toxin genes, virulence genes or genes related to lysogeny could be discovered in the phage genomes at this time, indicating their suitability for phage therapy. Efficacy of the phages in the plant environment was analyzed with bio-assays in leek leaves. Significant results were only obtained for phages KIL1, KIL2, KIL3, and KIL3b but the broader host range of phages KIL4 and KIL5 led to their incorporation in the phage cocktail for the field trials. When treating transplants with a phage cocktail before planting in an infested field, no significant differences were observed. Spraying of phages on artificially infected plants led to a slightly reduced symptom development in one of the three trials, yet again without statistical significance. Since two trials suffered from natural infections, their impact has most likely masked the effect of the treatments. Although isolation and characterization of phages is reported for several plant pathogens, results of other field trials are scarce. A recent overview has recently been published by Czajkowski (Czajkowski, 2016). A field trial with phages against Dickeya solani on potato also reports of phage therapy resulting in minor differences in disease severity (Adriaenssens et al., 2012). In experiments with phages against X. axonopodis pvs citri and citrumelo, significant control of bacterial spot was reported, yet treatment with copper-mancozeb was more effective (Balogh et al., 2008). As other researchers mentioned, open field (and greenhouse) applications with phages encounter many challenges such as

### REFERENCES

Adams, M. H. (1959). Bacteriophages. New York, NY: Interscience Publishers Inc.

UV-light, desiccation, application method, and the need of a large quantities of phages. Especially the phyllosphere is a harsh environment for phages, making leaf diseases difficult to treat (Gill and Abedon, 2003; Goodridge, 2004; Iriarte et al., 2012). This study indicates that phage therapy is able to reduce bacterial blight symptoms in leek, but to improve the efficacy of the phages in the field, phage persistence in the plant phyllosphere should be improved by the use of protective formulations, addition of non-pathogenic phage-propagating bacterial strains or adapting timing and frequency of application as suggested previously (Jones et al., 2007; Balogh et al., 2010).

### AUTHOR CONTRIBUTIONS

Conceived and designed the experiments: SR, AV, JV, MM, RL. Performed the experiments: SR, AV, SV, BD, DV, CA, CV, HJ, YB, JN, JK. Analyzed the data: SR, DV, YB, MM, RL. Contributed reagents/materials/analysis tools: JN, JK, MM, RL. Wrote the paper: SR. All authors read and approved the final manuscript.

### ACKNOWLEDGMENTS

The authors would like to thank Dr. LS van Overbeek of Wageningen UR (The Netherlands) for providing strain P55. The first author would like to thank Mr. Thijs van den Brande for his help with some of the figures and Mr. Johan Dick and Mrs. Cindy Elisabeth for taking care of the plants. The research was funded by the Agency for Innovation by Science and Technology (IWT, project nr.100881) with co-funding of Boerenbond, INGRO cvba, LAVA cvba, and the seed companies Bejo, Enza, Nunhems, Hazera, Rijk Zwaan, and Syngenta.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2016.00279

the KZ-related group, marine Vibrio phage JM-2012. J. Virol. 87, 12866–12878. doi: 10.1128/JVI.02656-13

a leaf disease of leek in Australia. Eur. J. Plant Pathol. 115, 419–430. doi: 10.1007/s10658-006-9033-4

temperature, host growth phase and media. FEMS Microbiol. Lett. 241, 13–20. doi: 10.1016/j.femsle.2004.06.058

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Rombouts, Volckaert, Venneman, Declercq, Vandenheuvel, Allonsius, Van Malderghem, Jang, Briers, Noben, Klumpp, Van Vaerenbergh, Maes and Lavigne. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Prophylactic Bacteriophage Administration More Effective than Post-infection Administration in Reducing Salmonella enterica serovar Enteritidis Shedding in Quail

Mosab Ahmadi1 , M. Amir Karimi Torshizi1 \*, Shaban Rahimi1 and John J. Dennehy2

1 Department of Poultry Science, Faculty of Agriculture, Tarbiat Modares University, Tehran, Iran, 2 Biology Department, Queens College and The Graduate Center of the City University of New York, New York, NY, USA

### Edited by:

Heather K. Allen, National Animal Disease Center, USA

### Reviewed by:

Yang Shen, ETH Zürich, Switzerland Beata Weber-D ˛abrowska, Immunology and Experimental Therapy, Polish Academy of Sciences, Poland

### \*Correspondence:

M. Amir Karimi Torshizi<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 15 May 2016 Accepted: 28 July 2016 Published: 09 August 2016

### Citation:

Ahmadi M, Karimi Torshizi MA, Rahimi S and Dennehy JJ (2016) Prophylactic Bacteriophage Administration More Effective than Post-infection Administration in Reducing Salmonella enterica serovar Enteritidis Shedding in Quail. Front. Microbiol. 7:1253. doi: 10.3389/fmicb.2016.01253 Infections caused by Salmonella bacteria, often through poultry products, are a serious public health issue. Because of drawbacks associated with antibiotic prophylaxis, alternative treatments are sought. Bacterial viruses (bacteriophages) may provide an effective alternative, but concerns remain with respect to bacteriophage stability and effectiveness. To this end, we assessed the stability of a novel bacteriophage isolated from poultry excreta, siphovirus PSE, and its effectiveness in reducing Salmonella enterica serovar Enteritidis colonization in vitro and in vivo. Moreover, we sought to determine how the timing (prophylactic or therapeutic) and route (oral gavage or vent lip) of PSE administration impacted its effectiveness. Here we report that significant quantities of viable PSE bacteriophages were recovered following exposure to high and low pH, high temperatures, and bile salts, testifying to its ability to survive extreme conditions. In addition, we found that ileal lactic acid bacteria and Streptococcus spp. counts increased, but colibacilli and total aerobe counts decreased, in quail receiving phage PSE through both oral gavage and vent lip routes. In other experiments, we assessed the efficiency of PSE administration, in both prophylactic and therapeutic contexts, via either oral gavage or vent lip administration, on S. Enteritidis colonization of quail cecal tonsils. Our results demonstrate that administration of PSE as a preventive agent could reduce the S. Enteritidis colonization more effectively than post-challenge administration. Furthermore, oral administration of PSE phage is a more effective prophylactic tool for reduction of S. Enteritidis shedding in poultry than is vent lip administration.

Keywords: phage therapy, prophylactic agent, Salmonella Enteritidis, phage administration method, microflora population

## INTRODUCTION

Despite impressive advances in the control of infectious diseases, some bacterial pathogens have acquired antibiotic resistance and are emerging in human populations. Many of these infections are zoonotic and are transmitted from healthy carrier animals to humans through contaminated food (Wegener et al., 2003). For example, Salmonella bacteria, especially the serovars Typhimurium and

Enteritidis, are common contaminants of poultry and eggs, causing food-borne disease and death (Borie et al., 2008). In order to reduce pathogen contamination of the food chain and eliminate food poisoning in human population, the eradication of Salmonella infections before harvest and processing is crucial (Seo et al., 2000; Mølbak and Neimann, 2002). Efforts to control S. Enteritidis in poultry historically have relied on a combination of farm biosecurity and the prophylactic application of antibiotics, but these strategies are increasingly unreliable (Hugas and Tsigarida, 2008; Buncic and Sofos, 2012). First, indiscriminate use of antibiotics has been implicated in a surge in multidrug resistant S. Enteritidis. Second, poultry consumers are increasingly concerned that edible poultry tissues are contaminated with harmful concentrations of drug residues (Donoghue, 2003; Griggs and Jacob, 2005). Third, changes in food production, food rejection, and preventive measures have incurred significant economic losses to poultry producers (Tsonos et al., 2013).

Furthermore, most antibiotics, up to 90% of orally administered doses, are not fully absorbed in the chicken gut, and can be excreted in the feces unchanged (Kumar et al., 2005). As one of the largest food-producing industries in the world, the poultry industry generates enormous amounts of manure and litter, much of which is currently applied to agricultural land (Bolan et al., 2010). The effects of antibiotics contained in manure on soil microbial communities are largely unknown (Ollivier et al., 2010). In addition, poultry litter from contaminated farms may introduce zoonotic pathogens such as S. Enteritidis into the environment (Line and Bailey, 2006). Thus the application of poultry litter as fertilizer can create significant public and environmental health concerns. Because of these issues, there is an urgent need to find novel and effective S. Enteritidis control strategies to minimize the risk of spreading antimicrobial resistance among animal and human populations.

Ideally vaccination would be an optimal control strategy, but S. Enteritidis vaccines do not provide complete protection to infected chickens (Gast et al., 1992). There are more than 2,500 serovars of Salmonella, and vaccines made from any one serovar often do not confer cross-protection against others (Singh, 2009). An old strategy of using naturally occurring bacterial viruses (bacteriophages) to tackle infections, is regaining popularity (Tsonos et al., 2013). The therapeutic and prophylactic application of bacteriophages is generally considered safe (Johnson et al., 2008). Bacteriophages are highly specific to certain bacterial strains, thus presumably have minimal impact on natural human or animal microflora populations (Sulakvelidze et al., 2001). However, bacteriophage therapy has not been consistently effective (Lu and Koeris, 2011). Issues remain in terms of effective delivery routes, host resistance, limited generalizability, and possible interactions with the immune system (Loc-Carrillo and Abedon, 2011). As capsid-based entities, bacteriophages are especially sensitive to environmental conditions, such as that in the upper gastrointestinal tract. Therefore, oral administration of phages may be limited in efficiency unless the selected phages are especially resilient to environmental challenges.

An alternative method of phage administration, the vent lip of the avian cloaca, was previously described by Andreatti Filho et al. (2007). However, the relative efficacy of oral versus vent lip administration has not been comparatively assessed. In this study, a novel lytic bacteriophage against S. Enteritidis, PSE, was isolated and characterized in terms of pH and thermal stability, bile salts tolerance, morphology, host range, and one-step growth. In addition, the therapeutic and prophylactic effects of PSE administration on the bacterial load of quail ilea were compared. Finally, in order to provide additional insight in the feasibility of phage application as an antimicrobial agent in poultry industry, the efficacy of PSE oral versus vent lip administration to reduce S. Enteritidis shedding in experimentally infected Japanese quail was determined.

## MATERIALS AND METHODS

### Bacterial Challenge Strain

The Salmonella Enteritidis (RITCC 1695) was purchased from the Razi Vaccine and Serum Research Institute (Karaj, Iran). For culture preparation, bacteria were grown in nutrient broth (Merck, Germany) at 37◦C overnight. Culture turbidity was adjusted to match McFarland standard 4.

## Isolation, Purification and Enumeration of Bacteriophage

Each poultry excreta sample was collected and suspended 1:10 (w/v) in SM (salt of magnesium) buffer (50 mM Tris-HCl [pH 7.5], 0.10 M NaCl, 8 mM MgSO4·7H2O, 0.01% gelatin [Sigma, Germany]), then suspension was centrifuged at 15,610 × g for 20 min at 4◦C. The resulting supernatant was filtered through a 0.22 µm membrane filter (Jet Biofil, China). Two hundred microliter of each filtered sample was mixed with 50 µl of fresh log-phase S. Enteritidis and 7 ml of top agar (0.7% agar in nutrient broth) and poured over a petri dish containing nutrient agar (1.5% agar; Serva, Germany) and incubated at 37◦C for 24 h. Individual clear plaques were extracted from the agar overlay with a sterile Pasteur pipette, and suspended in 500 µl of SM buffer, which was then re-plated to obtain single plaques. Each isolate was plaque purified 4x to ensure that each isolate represented a single clone.

Phage stocks were serially diluted in SM buffer to achieve a concentration that would produce discrete plaques on a bacterial lawn. Aliquots of 200 µl of any 10 × dilution series, 50 µl of fresh log-phase S. Enteritidis and 7 ml of soft agar (0.7% agar in nutrient broth, 42–45◦C) mixed and poured onto a plate containing nutrient agar (Serva, Germany). Plates were incubated for 24 h at 37◦C and phage plaques were enumerated.

### Phage PSE Lytic Spectrum

To evaluate the lytic spectrum of isolated bacteriophages, the sensitivity of 15 bacterial strains including three Salmonella serovars, four Escherichia coli serotypes, one Campylobacter serotype and seven gram-positive bacteria to isolated bacteriophages was determined by spot plating phage lysates on bacterial lawns (Adams, 1959). The resulting plates were incubated overnight at 37◦C, and subsequently checked for the phage plaque formation on the bacterial lawns.

### One-Step Growth

fmicb-07-01253 August 6, 2016 Time: 16:24 # 3

An overnight culture of S. Enteritidis (1 ml) was inoculated into fresh medium (100 ml) and incubated at 37◦C for 1 h to yield a cell density of 1.5 × 108 cfu ml−1 . To this culture, 1 ml of isolated phage was added, giving an approximate multiplicity of infection of 0.83. Samples were taken at 5 min intervals and immediately chilled until diluting and plating for phage quantitation. Viable bacteria were counted before bacteria and phage were mixed, and subsequently assessed at intervals. Latent period was defined as the time interval between the adsorption and the beginning of the first burst, as indicated by the initial rise in bacteriophage titer. Burst size was estimated from triplicate experiments using the equation described by Jiang et al. (1998).

## Thermal and pH Stability of Bacteriophage

In order to evaluate the stability of isolated bacteriophage at various temperatures, test tubes containing bacteriophage were placed in a water bath at each temperature (30, 40, 50, 60, 70, 80, and 90◦C) for 30 and 60 min (Bao et al., 2011). The surviving phages were diluted and enumerated immediately using the methods described above. In addition, the stability of bacteriophage at different pH was assessed according to the method described by Verma et al. (2009). The pH of nutrient broth was adjusted with either 1 M HCl or 1 M NaOH to obtain solutions with pH ranging from 2 to 11. A total of 1 ml of phage suspension at a titer of 1012 pfu ml−1 was inoculated into 9 ml of pH-adjusted medium to obtain a final concentration of 1011 pfu ml−1 . After incubation for 2 h at 37◦C, the surviving phages were diluted and counted by the methods described above. These experiments were performed three times, and the results are reported as the mean of three observation ± standard deviation.

### Bile Salts Tolerance

To determine bile salts tolerance, 1 ml of bacteriophage (1012 pfu ml−1 ) was placed into 9 ml of nutrient broth supplemented with 0 (control), 0.15, 0.3 and 1.0% W/V of bile salts (Quelab, Canada). After 0.5, 1, 3, and 4 h of incubation at 37◦C, phage survival was assayed by diluting and enumeration of phage using the methods described above. Phage titer in each concentration of bile salts was compared with a bile salt-free control.

### Phage Morphology Examination

A drop of 109 pfu ml−1 bacteriophage was spotted onto a formvar-coated grid and fixed for 2 min with 2.5% glutaraldehyde. Excess sample was removed, and the grid was washed with a drop of double-distilled water. Negative staining was performed by adding 1 drop of 2% uranyl acetate to the grid surface, and excess stain was removed immediately. The grid was allowed to air dry for 60 min and was then observed with a Zeiss-EM 10C transmission electron microscope (TEM) at 100 kV. Phage morphology and dimensions (head diameter, tail length) were determined by the mean of at least 5 measurements.

## Analysis of Bacteriophage Nucleic Acid

Extraction of phage nucleic acid was conducted according to the method of Binetti et al. (2005) with slight modifications. Phage particles were precipitated with 4% polyethylene glycol 6000 in 0.2 M NaCl for 48 h at 4◦C and centrifuged at 15,610 × g for 240 min at 4◦C. The pellet was re-suspended in SM buffer, supplemented with 40 µg/ml RNase A (Sinaclon, Iran) or 1 µg/ml DNase I (Sinaclon, Iran) and incubated at 37◦C for 30 min. Then 80 µl of lysis solutions (0.25 M EDTA [pH 8.1], 0.5 M Tris-HCl [pH 9.6] and 2.5% sodium dodecyl sulfate [SDS]) were added to 400 µl of a concentrated suspension of phage particles, and the mixture was incubated at 65◦C for 30 min. One hundred microliter of 8 M potassium acetate was then added, and the mixture was incubated on ice for 15 min before centrifugation (15,610 × g, 30 min, 4◦C). Phage nucleic acid was precipitated from the supernatant with an equal volume of isopropanol, kept at room temperature for 5 min, then centrifuged again (15,610 × g, 30 min, 4◦C). The pellet was re-suspended in 630 µl of TE (Tris EDTA) buffer (10 mM Tris-HCl; 1 mM EDTA, pH 8.0) in the presence of 0.3 M sodium acetate and precipitated with isopropanol for 5 min, followed by centrifugation (15,610 × g, 30 min, 4 ◦C). The pellet of nucleic acid was washed twice with 70% ice-cold ethanol, dried and re-suspended in 50 µl of TE buffer.

Purified nucleic acid was digested with DNase I or RNase A according to the manufacturer's instructions to determine nucleic acid type. We also attempted digestion with restriction enzymes EcoRI and HindIII according to the manufacturer's instruction (Serva, Germany). Products of digested phage nucleic acid were electrophoresed by 0.8% agarose gel in a TAE (Tris Acetate EDTA) buffer (40 mM Tris-acetate, 1 mM EDTA) and visualized by transillumination with UV light after the gels were stained with ethidium bromide (Serva, Germany).

### Determination of Phage Genome Size by Agarose Gel Electrophoresis

The relatively small genome of PSE made it possible to use agarose gel electrophoresis for determination of its size. The distance of PSE DNA migration on agarose gel electrophoresis was compared to that of un-cut and HindIII lambda DNA fragments as DNA size marker by Photo Capt software version 12.4 (Vilber Lourmat, France) (Brown, 2010).

### Experimental Analysis of Bacteriophage Treatment on S. Enteritidis Infection

This project maintained proper ethical standards and all experiments were approved by the Animal Care Committee of Tarbiat Modares University.

### Experiment 1

To evaluate the effects of isolated phages on bird natural microflora, ninety 33-day-old Salmonella-free quail were

randomly divided into three groups of 30 birds. All birds received an identical diet, and feed and water were supplied ad libitum. A control group (Group 1) did not receive bacteriophage treatment. Group 2 received 109 pfu ml−1 bacteriophage in 100 µl aliquot by oral gavage for 2 days, once every 24 h and group 3 received 10 µl of 1010 pfu ml−1 bacteriophage, via vent lip route for 2 days, once every 24 h.

Bacterial quantitation of the ileal contents of 10 birds per each group was done at 36 days. One gram of the ileal contents of each bird was serially diluted in phosphate-buffered saline from 10−1 to 10−6 . Dilutions were subsequently plated on selective agar media for enumeration of target bacteria. Lactic acid bacteria, colibacilli, streptococci and total aerobes were plated on de Man, Rogosa and Sharpe agar (MRS, Merck, Germany), Mac Conkey agar (Merck, Germany), Kenner Fecal Streptococcus agar (KF Streptococcus, Merck, Germany**)**, and plate count agar (Merck, Germany), respectively. Plates were incubated at 37◦C overnight, and then the resulting colonies were counted.

### Experiment 2

To compare the efficiency of prophylactic versus therapeutic application of phage on S. Enteritidis shedding, one hundred 8 day-old Salmonella-free quail were randomly divided into four groups of 25 birds. Group 1 (prophylactic treatment) received 100 µl of 106 pfu ml−1 bacteriophage via oral gavage for 3 days, once every 24 h, before being challenged orally by 100 µl of 1.2 × 109 cfu ml−1 S. Enteritidis (**Table 1**). Group 2 (therapeutic treatment) received 100 µl of 1.2 × 109 cfu ml−1 S. Enteritidis first, then received bacteriophage as described in group 1 (**Table 1**). Birds of group 3 were challenged like group 2, but did not receive phage treatment (**Table 1**). Neither Salmonella nor bacteriophage was administered to group 4 (**Table 1**).

The presence of S. Enteritidis was assayed at 12, 24 h, and 7 days following S. Enteritidis challenge. Birds were euthanized by CO2 inhalation, and their cecal tonsils were aseptically removed, homogenized and incubated overnight at 37◦C in 10 ml of peptone water buffer (Merck, Germany). Subsequently, 100 µl of cultured peptone water was transferred into Rappaport–Vassiliadis broth (Merck, Germany) at 37◦C for 24 h, and then streaked on xylose-lysine desoxycholate (XLD) agar plates (Merck, Germany). Plates were incubated at 37◦C for 24 h. Any black colonies that formed were identified as Salmonella Enteritidis. Any doubtful colonies were plated on triple sugar iron agar (TSI agar, Merck, Germany) slants. In addition, urease activity was checked by assaying the degradation of urea in urea broth (Stager et al., 1983). The recovery of S. Enteritidis is reported as the number of S. Enteritidis positive samples per number of total samples.

### Experiment 3

The effectiveness of oral gavage versus vent lip administration of phage on S. Enteritidis colonization in cecal tonsils of quail was evaluated in a third experiment. Three hundred, 1-dayold Salmonella-free quail were randomly assigned to five groups of 60 birds. Each group of birds was placed in cages in a controlled environment under strict biosecure conditions. Fecal samples were taken to determine if any pre-existing Salmonella phages were present using the method described above. Feed and water were provided ad libitum, and birds were maintained at an age-appropriate temperature for the duration of the experiment.

Neither Salmonella nor bacteriophages were administered in group 1. Group 2 birds did not receive bacteriophage, but at the age of 4 days were challenged orally by 100 µl of 1.2 × 109 cfu ml−1 S. Enteritidis. Birds in group 3 received 100 µl of 109 pfu ml−1 bacteriophage via oral gavage every 24 h for 6 days, starting 3 days before S. Enteritidis challenge. Group 4 received 5 µl of 2 × 1010 pfu ml−1 bacteriophage, via the vent lip route every 24 h for 6 days, starting 3 days before S. Enteritidis challenge. Birds of group 5 received bacteriophages similar to groups 3 and 4, but at half the concentration, and were not challenged with S. Enteritidis (**Table 2**). Necropsies of birds from each group were performed at 6, 12 h, 1, 2, 3, 7, 14, 28, and 35 days post-challenge. The presence of S. Enteritidis was assayed as described above. In this experiment, recovery of bacteriophage from bird feces was done by using described method for phage isolation.

### Statistical Analysis

The data related to the bacterial enumeration of quail ileal contents were compared between groups using a generalized linear model (GLM) implemented in SAS with subsequent Duncan's multiple range test (SAS Institute, 2003). The significance of differences in the incidence of S. Enteritidis recovery between control and phage-treated experimental groups was determined by the chi-square test (p ≤ 0.05).

### TABLE 1 | Scheme of Experiment 2.

aDay of hatching is given as day 0. bDose of phage given as pfu ml−1 . cDose of S. Enteritidis given as cfu ml−1 .

a Day of hatching is given as day 0. b Dose of phage given as pfu ml−1 . cDose of S. Enteritidis given as cfu ml−1 .

## RESULTS

## Phage PSE Isolation and Lytic Spectrum Assay

One Salmonella-lysing phage, named PSE, was isolated from poultry's excreta using an enrichment strategy (Cross et al., 2015). PSE forms clear 2–3 mm plaques on Salmonella Enteritidis (**Figure 1A**). A lytic spectrum test indicated that PSE was able to lyse three strains of Salmonellae, but was unable to lyse bacteria of other genera (**Table 3**).

## Phage PSE Morphology

Phage PSE has a round head with a diameter of 51.29 ± 3.15 nm and a tail with length of 74.53 ± 0.74 nm. From its morphology, PSE was presumptively identified as a member of the Siphoviridae family with long non-contractile tail in the order of Caudovirales (**Figures 1B,C**).

### Phage PSE One-Step Growth Dynamics

One-step growth curves of PSE were conducted to determine the growth pattern and the number of progeny phages released

FIGURE 1 | Associated plaque (A) and transmission electron micrographs of PSE phage under 100 kV magnification. The bars represent 120 nm (B) and 50 nm (C).

### TABLE 3 | The lytic spectrum of PSE on 15 bacterial strains from 7 genera.

1Razi vaccine and serum research institute, Iran. 2American Type Culture Collection, USA.

by lysing a single bacterial host cell. One-step growth curves for phage PSE showed a latent period and burst size of about 20 min and 66.66 ± 6.67 pfu cell−1 , respectively (**Figure 2**).

## Phage PSE Stability at Different Bile Salts Concentrations, Temperatures and pH

The viability of phage PSE was fully maintained and no reduction in PSE titer was observed after 4 h of exposure to any of the tested bile salts concentrations (data not shown). No obvious effect on PSE activity was observed after 2 h incubation at pH levels ranging from 3 to 11 (**Figure 3**). In addition, the results of thermal stability tests suggested that phage PSE was relatively heat stable up to 60 min at temperatures between 30◦C to 70 ◦C. At 80◦C or higher, the phage titer quickly dropped, no viral particles were detected after 30 min of incubation, and phage activity was completely lost (**Figure 4**).

### Phage PSE Genomic Characterization

The analysis of phage nucleic acid suggested that PSE is a DNA phage as the genome was completely digested by DNase I, but refractory to the activities of RNase A. The estimated genome size of PSE was approximately 35.72 kb (**Figure 5**). Two restriction enzymes used in this study, EcoRI and HindIII, were unable to digest the PSE genome.

### Effect of Phage PSE on Bacterial Frequencies in the Quail Ilea

We assessed the impact of phage PSE administration on quail ileal bacteria. Our results show that lactic acid bacteria and streptococci counts increased relative to controls when PSE was administered via oral gavage and vent lip methods (**Table 4**; P = 0.0001). By contrast, colibacilli and total aerobes counts decreased relative to controls following both treatment methods (**Table 4**; P = 0.0001 and 0.005 respectively). Administration of bacteriophage PSE via oral gavage and vent lip routes increased lactic acid bacteria count by 1.47 and 0.73 log cfu g−1 , respectively (**Table 4**). Also streptococci counts increased by 1.05 and 0.58 log cfu g−1 , when PSE was administered by oral gavage and vent lip routes, respectively (**Table 4**).

FIGURE 3 | Effect of pH on stability of phage PSE.

### Comparison of Preventive versus Therapeutic Phage Treatment

We tested the effectiveness of PSE treatment, either prophylactically prior to S. Enteritidis exposure or therapeutically after S. Enteritidis exposure. One day post-challenge, S. Enteritidis was isolated from 100% of cecal tonsils of positive control and therapeutic groups. By contrast, S. Enteritidis isolation rates in birds of negative control and preventive groups were 0 and 33.3% respectively (**Table 5**). In addition, 7 days post-challenge, S. Enteritidis was isolated from 100% of positive control and therapeutic groups (**Table 5**). By contrast, the S. Enteritidis isolation rate from the preventive group was 20% and birds of the negative control remained free from S. Enteritidis.

### Comparison of Oral versus Vent Lip Administration of Phage PSE

The application of phage PSE, either by oral or vent lip routes, reduced S. Enteritidis recoveries from cecal tonsils of quail in nine consecutive samplings (**Table 6**). Administration of PSE via the oral route was able to eliminate S. Enteritidis at all sampling times except 6 h post-challenge. One week after treatment, S. Enteritidis was recovered from 100% of cecal tonsil samples of

the positive control group, while no S. Enteritidis was found in birds that orally received bacteriophage PSE (**Table 6**). Overall, at all sampling times, S. Enteritidis was isolated from 90.74% of cecal tonsil samples of the positive control group. By contrast, S. Enteritidis isolation rates in birds treated with bacteriophage PSE via oral and vent lip routes, and in the negative control, were 2.77, 17.58, and 0% respectively (**Table 6**). Bacteriophages were recovered from fecal specimens of all bacteriophage-receiving groups, regardless of administration routes (**Table 7**).

### DISCUSSION

A rise in antibiotic-resistant S. Enteritidis bacteria, and concerns about the ecological and environmental effects of unrestricted antibiotic usage, have provided strong motivation to find new and effective prophylactic and therapeutic means of eradicating S. Enteritidis from poultry. Bacteriophages have excellent potential for replacing or supplementing antibiotics, but their effectiveness and stability must be demonstrated experimentally. Here we describe the isolation, characteristics and effectiveness of a novel bacteriophage able to infect S. Enteritidis.

∗SEM and P-values were calculated using a GLM univariate analysis of variance. Post hoc comparisons (Duncan's test) of mean values within each column revealed that all are significantly different (P-value < 0.05) from other groups in the same column except the pair labeled with the superscript ns.

On the assumption that phages isolated from poultry's excreta are more stable in the poultry gastrointestinal tract compared to isolated bacteriophages from other sources, we isolated a bacteriophage, which we called PSE, from poultry excreta. Bacteriophage PSE has a number of characteristics that make it a strong phage therapy candidate. Bacteriophage PSE formed 2– 3 mm round, clear plaques on S. Enteritidis, implying that it is a lytic phage (Yoon et al., 2007). Because they quickly reproduce within and lyse host bacteria, lytic phages are more suitable than lysogenic phages for phage therapy (Abedon, 2008). Moreover, although PSE was isolated using S. Enteritidis, it was also able to infect and lyse S. Typhimurium and S. Pullorum, two other significant bacterial pathogens found in poultry (**Table 3**).

Bacteriophage PSE's latent period was estimated to be about 20 min, and its burst size was approximately 66.66 ± 6.67 pfu cell−1 . There seems to be considerable variation in latent period length and burst size among Salmonella-specific phages. Some Salmonella-specific phages have been reported to possess small burst sizes and long latent periods, for example φ st1 (22 pfu cell−1 , 40 min) (Wong et al., 2014) and phage Felix 01 (14 pfu cell−1 , 60 min) (O'Flynn et al., 2006). On the other hand, some Salmonella-specific phages were isolated with burst sizes range from 100 to 200 pfu cell−1 and shorter latent periods (Ahiwale et al., 2013). Differences in medium, host cell, pH and temperature may affect variations in latent period and burst size of different phage isolates (Müller-Merbach et al., 2007). For the purposes of phage therapy, optimum latent periods and burst sizes have not been established; however, there is some evidence that these parameters can vary considerably depending on host density and physiological status (Abedon, 1989; Abedon et al., 2001; Wang, 2006).

Phages used for therapeutic or prophylactic purposes should be highly stable and remain viable in a wide range of potential environments. Phage PSE meets some of these requirements. It exhibited powerful antibacterial activity at temperatures ranging from 30◦C to 70◦C, and pH levels ranging from 3 to 11. Also PSE was found to be resistant to high bile salt concentrations, remaining viable even after 4 h of exposure to 1% bile solution. These characteristics suggest that phage PSE would remain therapeutically viable inside animal bodies, including the animal gastrointestinal tract, where Salmonella may be prevalent. All of

### TABLE 5 | Frequency of S. Enteritidis positive cecal tonsils per group (%).

TABLE 6 | Efficiency of phage PSE administration methods on S. Enteritidis recovery from cecal tonsils of Japanese quail (S. Enteritidis positive cecal tonsil/ total).

1Hour. 2 Day.

TABLE 7 | Recovery of bacteriophage from feces of birds after 7 days post treatment.

aAverage of 10 birds.

these features point to the utility of phage PSE as a stable agent for control of S. Enteritidis colonization of poultry.

In the intestine, bacterial populations can interact with each other, and phages are expected to have a significant role in driving the biodiversity of this complex ecosystem (Ventura et al., 2011). Our data demonstrate that the PSE phage administration, either by oral or vent lip method, strongly affected ileal bacteria frequencies (**Table 4**). We observed lower frequencies of coliforms in phage-treated birds compared with controls (**Table 4**). The declines in coliforms were matched by increases in lactic acid bacteria and streptococci (**Table 4**). Lactobacillus spp. are often thought to have a positive effect on poultry health by reducing or preventing colonization of the poultry intestinal tract by undesirable bacteria (Karimi Torshizi et al., 2008). However, given the complexities of the gut microbiome, it is difficult to determine the direct mechanism by which PSE treatment may have influenced the relative numbers of coliforms, lactic acid bacteria and streptococci. This finding is nonetheless an intriguing topic for further investigation.

The S. Enteritidis reductions observed in cecal tonsil samples following preventive treatment strongly suggest that phage PSE is a viable prophylactic against S. Enteritidis colonization (**Table 5**). It is commonly believed that bacteriophages administered to treated animals are present for the duration of the infection, but once the bacterial host is eliminated so too is the bacteriophage. In our experiments, this did not happen. Bacteriophage PSE was isolated from feces of all groups that received it, even after S. Enteritidis infection was no longer observed. Interestingly, phage PSE persistence was also observed in the absence of S. Enteritidis challenge (**Table 7**). This finding suggests that phage PSE either remains inactive in gastrointestinal tract for long periods or may have alternate hosts that allow it to proliferate. Similarly, it was reported that, in the absence the primary host, the bacteriophage UZ1 persisted 13 days in a simulated colon (Verthé et al., 2004). This evidence lends credibility to the prophylactic ability of PSE.

This result is in sharp contrast to the observation that the therapeutic PSE treatment of birds infected by S. Enteritidis failed to eradicate S. Enteritidis from bird cecal tonsils relative to controls (**Table 5**). It may be that PSE treatment of extant infections rapidly selected for PSE resistance in S. Enteritidis bacteria or that S. Enteritidis that had previously colonized the treatment animals were resistant to infection (Higgins et al., 2008). This experiment highlights the importance of phage administration as a prophylactic prior to S. Enteritidis infection is a more effective strategy than phage administration as a treatment following S. Enteritidis infection (**Table 5**). Presumably, prophylactic PSE treatment can prevent establishment of S. Enteritidis infection, but is relatively ineffective against established S. Enteritidis infections.

In another experiment, phage PSE was applied to treated birds, either orally or via the vent lip, for 3 days prior to infection with S. Enteritidis. Following S. Enteritidis challenge, all birds were tested for S. Enteritidis infection periodically across 35 days. In birds treated with PSE orally, S. Enteritidis was detected in

the cecal tonsils 6 h following S. Enteritidis challenge, but not subsequently (**Table 6**). Some birds treated with phage PSE via the vent lip periodically tested positive for S. Enteritidis across the study period, but the majority remained free of S. Enteritidis (**Table 6**). Birds in the negative control (no treatments) and birds receiving only bacteriophage treatment remained free from contamination with S. Enteritidis across the entire study period (**Table 6**). These results imply that the bird quarantine was effective and no cross contamination occurred during the entire experimental period. Significantly, while both oral and vent lip treatment routes reduced S. Enteritidis infection compared to the positive control, the oral treatment route was more effective than the vent lip method in treatment of S. Enteritidis infections.

Although some studies suggested that it might be necessary to employ cocktails of bacteriophages to provide protection against S. Enteritidis (Atterbury et al., 2007; Borie et al., 2008), our study demonstrates that treatment with a single phage type can effectively prevent S. Enteritidis colonization. For practical and economic reasons, employing a single phage to achieve therapeutic effect is more desirable than cocktails that contain several types of phage (Sulakvelidze et al., 2001).

### CONCLUSION

Phage PSE shows great promise for the prevention and treatment of S. Enteritidis infection, and it may be a plausible alternative to antibiotics for the reduction of S. Enteritidis shedding in poultry. Phage PSE was most effective when administered prophylactically prior to S. Enteritidis infection than as a treatment for established S. Enteritidis infections. Although we observed a reduction in S. Enteritidis infection in birds prophylactically treated with phage

### REFERENCES

PSE via the vent route, our results indicate that administration of phage PSE via oral route is most effective.

### HIGHLIGHTS

Phage PSE persistence was observed in the absence of S. Enteritidis challenge. Administration of phage PSE as a preventive agent could reduce the S. Enteritidis colonization more effectively than post challenge administration. Administration of phage PSE via oral route is most effective for reducing S. Enteritidis colonization.

### AUTHOR CONTRIBUTIONS

MA and MK designed the study, performed experiments, analyzed the data and wrote the manuscript. SR and JD reviewed the manuscript. All authors read and approved the final manuscript.

### FUNDING

This project was funded by the Research Council of Tarbiat Modares University. The funder had no role in study design, data collection and analysis, or preparation of the manuscript.

### ACKNOWLEDGMENT

We thank the Research Council of Tarbiat Modares University for funding this study.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Ahmadi, Karimi Torshizi, Rahimi and Dennehy. This is an openaccess article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Bacteriophages against Serratia as Fish Spoilage Control Technology

### Igor Hernández\*

AZTI-Tecnalia, Food Research Division, Parque Tecnológico de Bizkaia, Derio, Spain

Bacteria of the genus Serratia, mainly S. proteamaculans and S. fonticola, are important spoilage agents in Atlantic horse mackerel (Trachurus trachurus). In order to evaluate whether bacteriophages against Serratia could delay the spoilage process, 11 viral strains active against this genus were isolated from food and best candidate was applied to fresh mackerel filets. All the phages belong to the Siphoviridae and Podoviridae families and were active at multiplicity of infection (MOI) levels below 1:1 in Long & Hammer broth. The ability of phage AZT6 to control Serratia populations in real food was tested in Atlantic horse mackerel extract and applied to fresh mackerel filets. Treatment with high phage concentration (MOI 350:1, initial Serratia population 3.9 ± 0.3 Log cfu/g) can reduce the Serratia populations up to 90% during fish storage (a maximum of 6 days) at low temperatures (6◦C). Bacterial inhibition was dependent on the bacteriophage dosage, and MOI of 10:1 or lower did not significantly affect the

### Edited by:

Pilar García, Consejo Superior de Investigaciones Científicas (CSIC), Spain

### Reviewed by:

Lorena Rodriguez-Rubio, KU Leuven, Belgium María Ángeles Tormo-Mas, Researcher Ramon y Cajal, Spain

> \*Correspondence: Igor Hernández<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 28 June 2016 Accepted: 03 March 2017 Published: 04 April 2017

### Citation:

Hernández I (2017) Bacteriophages against Serratia as Fish Spoilage Control Technology. Front. Microbiol. 8:449. doi: 10.3389/fmicb.2017.00449 Serratia populations.

 Their ability to reduce bacterial loads has led them to be suggested as therapies for human and animal microbial infections (Brüssow, 2012), but other applications are also possible. In food technology, phages have been proposed for food spoilage control (García et al., 2008; Mahony et al., 2011), and selected phages have been tested in food contaminated with Salmonella (Leverentz et al., 2004), Listeria (Leverentz et al., 2004; Carlton et al., 2005), or Escherichia coli (Carter et al., 2012). However, few references describe phages as a strategy against spoilage bacteria for shelf life extension of food. Bacteriophages against spoilage bacteria have the advantage that natural resistance of bacteria to lysis by phage is not a critical safety concern, while even modest reduction of specific microorganisms can significantly prolong food's shelf life. To the best of our knowledge, the first reference of phages as food "spoilage control agents" was from Ellis et al. (1973), who worked on milk storage. Later, other authors described phages that were active against Pseudomonas in beef (Greer and Dilts, 1990) and in milk (Sillankorva et al., 2008), Listeria brevis in beer (Deasy et al., 2011), Brochothrix thermosphacta in pork (Greer and Dilts, 2002), and Shewanella in fish (Li et al., 2014). In most cases, the target bacteria were inoculated in the food matrix, and the activity against naturally presenting bacteria was not considered; hence, complicates the transfer of the technology to the food industry.

**Abbreviations:** MOI, multiplicity of infection; Pfu/ml, plaque formation units per milliliter.

In previous work, Alfaro and Hernández (2013) determined that Serratia is a dominant bacterial genus (over 25% of the total bacterial load) at sensorial rejection time of Atlantic Horse Mackerel (Trachurus trachurus) filets stored in modified atmospheres (MAPs), and the increase of these bacteria during fish storage has been related to their fresh shelf life (Alfaro et al., 2013). Many bacterial species have been associated with formation of biogenic amines (De Filippis et al., 2013), and they are part of the spoilage microflora of diverse foods (Doulgeraki et al., 2012). The Serratia genus are Gram negative, facultative anaerobic bacteria, which grow in a broad range of temperatures and substrates, including plant surfaces, soil, water, and food products such as fruit juices and fish (Garrity et al., 2005). Many species have been related with food spoilage, and some of them (mainly S. marcescens and S. liquefaciens) have been described as opportunistic human pathogens (Mahlen, 2011).

Bacteriophages for Serratia have been purified before from sewage effluent, mainly using strains of S. marcescens as targets. Those virons have a relatively wide broad range of action, and are even able to infect bacteria of genera other than Serratia (Prinsloo and Coetzee, 1964; Prinsloo, 1966; Evans et al., 2010). Phages against Serratia have been used as molecular biology tools (Petty et al., 2006) and as models for viron abundance estimation in soil (Ashelford et al., 1999, 2003). To the best of our knowledge, no utilization in the food industry has been reported.

This paper describes the isolation, selection, and characterization of bacteriophages against Serratia species, and the evaluation of their ability to control growth of naturally occurring Serratia during spoilage of Mackerel filets. The research reported here provides an example of the potential and limitations of the application of bacteriophages to retard bacterial spoilage in foods.

## MATERIALS AND METHODS

### Bacterial Strains and Culture Media

Serratia fonticola and S. proteamaculans strains were isolated in our facilities from spoiling Atlantic horse mackerel and were characterized by 16S RNA sequencing as previously described (Alfaro and Hernández, 2013). Two strains of S. fonticola (SFO001 and SFO002) and six strains of S. proteamaculans (SPR001, SPR002, SPR004, SPR005, SPR006, and SPR009) were used for phage isolation and characterization. Two S. marcescens strains (CECT 854 and CECT 977) were purchased from the Spanish type culture collection (CECT).

Long & Hammer agar and Long & Hammer broth (Koutsoumanis and Nychas, 1999) were used as general propagation media for Serratia. Strains were cultivated for 24 h at room temperature (22–25◦C) in aerobic conditions. Bacterial populations in fish samples were quantified using: (i) Serratia selective agar (SSA) prepared as described by Starr et al. (1976) for enumeration of Serratia. Incubations were aerobic at 20◦C. (ii) Triptone soya agar (TSA) incubated aerobically at 30◦C for 24 h for total bacterial counts. (iii) Long & Hammer agar incubated aerobically at 12◦C for 72 h for psychrotropic total bacterial counts. Peptone water was used for decimal dilutions.

Spoilage-related bacteria previously isolated and characterized from Atlantic horse mackerel (Alfaro and Hernández, 2013) were used to study the phage specificities. Carnobacterium maltaromaticum, Shewanella putrefaciens. Vibrio ssp., and Yersinia intermedia strains were used for phage specificity determinations. Bacteria in exponential growth phase (in TSB) were treated with phages of interest (MOI 1000:1), and growth was controlled by optical density at 600 nm (OD600 nm) and by colony count in TSA (incubated at 30◦C for 24 h).

## Phage Isolation and Enumeration

The Serratia bacteriophages were isolated from cheese whey obtained from artisanal cheese producers. Briefly, whey samples were mixed with an exponential (OD600 = 0.250–0.300) growth culture of SFO001 (whey:culture: 1:1) and incubated at 20◦C for 7 h. These phage-rich samples were mixed with chloroform (10:1 v/v), centrifuged (3 min, 14.000 × g) and the water fractions were transferred to sterile tubes. Water fractions (or their decimal dilutions) were mixed (1:1) with high-density SFO001 culture (OD = 1.5) and with Long & Hammer soft agar. The mixture was smeared over a plate of Long & Hammer agar and incubated at 20◦C for 24 h until plaques were visible. For further purifications, one plaque from each plate was extracted, dissolved in peptone water and treated as described above for whey samples. Three purification cycles were performed for each phage.

Phage enumeration was done with a spot test and double-layer agar technique (Kropinski et al., 2009; Mazzocco et al., 2009), using exponentially growing SFO001 as sensible strains. Plates (Long & Hammer agar and soft agar) were incubated at 25◦C for 24 h. Samples of 10 µl of phage dilution were used for the spot tests. Phage dilutions were done in sterile saline–magnesium– gelatine buffer (SGM, 0.1 M NaCl, 1 mM MgSO4 ∗ 7H2O, 50 mM Tris–HCl and 0.01% gelatine, pH = 7.5).

SPR009 has a high growth rate at a broad-range of growth temperatures, which facilities the use of this strain for phageactivity characterization in liquid medium. Long & Hammer broth was used.

For stock production of phages, they were cultivated with exponentially growing target strain (SPR009) in fresh Long & Hammer broth at 20◦C for 24 h. Bacteria were hydrolyzed and removed by mixing the culture with chloroform (10:1), vortex and centrifuged (5 min, 25000 × g). Supernatant was spread on a Petri plate under sterile air for 15 min and stored at 4 ◦C or at −80◦C (20% glycerol added) until used. When high volumes of bacteriophage were necessary, samples were purified and concentrated as described by Sambrook and Russell (2001), using modified Long & Hammer broth (gelatin concentration was reduced to 1% and 10 mM CaCl2 was added) as cultivation medium and PEG6000 as precipitation agent.

### Phage Characterization

For molecular characterization, DNA was isolated according to the method described by Sambrook and Russell (2001). For restriction endonuclease analyses, DNA was digested with EcoRI (New England Biolabs) according to the manufacturer's recommendations. Fragment lengths were verified with gel electrophoresis (0.8% agarose in TAE buffer, 70v; Biorad, Madrid,

Spain) and visualized in a BioDoc-it imaging system (UVP, Upland, CA, USA). Generuler 1-kb DNA leader was used as a calibration standard (VWR, Madrid).

Phage morphology was studied by transmission electron microscopy (TEM). For purification, phage dilution (estimated as 1012 pfu/ml) was centrifuged (1500 × g, 60 min, 4◦C), and the pellet was dissolved in sodium acetate solution. Centrifugation was repeated, and the pellet was finally dissolved in sodium acetate. For sample preparation, 5 µl of phage dilution was fixed in an Agar S-160 carbon rack temporally hydrophilized by glow discharge (20 s, 800v DV in vacuum conditions). Phages were stained with uranyl acetate 1% (twice, 30 s each). Phage morphology was examined with TEM (Philips CM120 biofilter) and photographed with an Olympus SIS "Morada" camera. Phage morphology and dimensions (capsid diameter, tail length, and width) were analyzed from electron micrographs with ImageJ software (Rasband, Ver 1.48).

Phage lytic spectra were determined using fresh phage propagated in SFO001 and double agar layer technique as described above. Plaques were counted after 24 h at 25◦C.

## Evaluation of Phage Activity in Fish Broth and Filets

For fish broth preparation, five fresh Atlantic horse mackerel were eviscerated, cut in small pieces and mixed. Then 1 kg of the fish was boiled for 10 min in 1.0 l of distilled water. The broth was filtered through 1 mm pores and autoclaved 15 min at 110◦C. Fresh inoculum of strain SPR009 was used to inoculate the broth (final concentration (3.9 ± 0.2 log cfu/ml) and AZT6 phage was inoculated 10 min later at 6 or 20◦C. After 5 days, a 1 ml sample was removed for bacterial and phage concentration determinations. Bacterial load was estimated on Long & Hammer agar (25◦C, 24–48 h), and phage load was determined by the spottest technique described above. Each experimental condition was studied in triplicate.

For antimicrobial activity on fish filets, T. trachurus filets were used as a model food matrix. These filets were not inoculated with bacteria, and the results reflect reduction of natural microbiota. Two fresh fish were supplied by each of two different local markets, eviscerated, heads-removed under sterile conditions and the bodies divided along the vertebral bones in two similar portions (in total eight "filets"). Four filets were submerged for 60 s in AZT6 phage solution (SMG buffer with different concentrations of phages), dried for 60 s under sterile air and stored in aseptic 140-mm Petri plates with plastic covers. Samples were stored aerobically at 6◦C in a temperature-controlled chamber. After 3 or 6 days at 6◦C, 5–10 g of fish from each filet were aseptically removed, homogenized in APT (fish:broth ratio was 1:10) using a Stomacher R blender (6 × {2000 on and 1000 off}) and used for bacterial counts.

## Statistical Analysis

PSPP V0.6.2 (Free Software Foundation, Inc.) was used for analysis of variance (ANOVA) and least significant difference

(LSD) statistical procedures. A confidence interval of 95% (p ≤ 0.05) was used.

### RESULTS AND DISCUSSION

fmicb-08-00449 March 31, 2017 Time: 18:28 # 4

### Phage Isolation and Characterization

Working with Serratia fonticola SFO001 the as target strain, 11 phages were isolated from cheese whey samples using the doubleagar layer technique. Initial phage purification did not require Ca2+ addition, but addition of 0.5 mM Ca2+ increased the plaque size after the last purification cycle.

Isolated phages consistently produced clear plaques on Long & Hammer agar inoculated with Serratia SFO001 and, therefore, the whey phages were classified as virulent. Analysis showed that DNA of all of the isolated virons was double stranded and produced different DNA digestion profiles with EcoRI (results not shown). We discarded duplicate isolations. Estimated DNA size ranged from 50 to 70 Kbp.

Phages AZT1, AZT2, AZT3, AZT5, AZT6, and AZT9 have icosahedral heads (80–90 nm) and long non-contractile tails (100–120 nm). In contrast, AZT4, AZT7, AZT8, AZT10, and AZT11 presented spherical heads (75–90 nm) and extremely short tails (10–15 nm). All the isolated virons have lytic activity after 10 min of incubation in chloroform, suggesting that a lipid envelope is not a functional requirement. The first group would be classified as type B in Bradley's classification (Bradley, 1967), and they probably belong to the Siphoviridae family. The second group would be C-type phages (Bradley, 1967) classified with the Podoviridae. **Figure 1** presents the typical conformation of each type of phage. Other authors have demonstrated the highly variable morphology of the Serratia phages, even when they are isolated from the same source (Bradley, 1965; Ashelford et al., 2003).

Maximal phage titrations in the medium after a single cultivation round were between 6.9 and 9.3 Log pfu/ml (Supplementary Table 1). All bacteriophages were active after 30 days at −80◦C with losses of viability less than 2.0 Log pfu/ml, except AZT5 and AZT8 that lost viability by more than five orders of magnitude during ultrafreezing. Similar activity losses in pure phage solutions were observed after 30 days at 4◦C (Supplementary Table 2).

Bactericidal effects depend on MOI and, in most of the cases, effective MOI at 20◦C ranged over five orders of magnitude (**Figure 2**, AZT3 as an example). An MOI close to 100:1 produced an initial decrease in bacterial concentration, followed by a normal growth curve. Lower phage doses (when activity is detected) permit initial bacteria growth, followed by decreases in bacterial concentration in later phases (**Figure 2**). In all cases, MOI of 1:1 or higher resulted in significant bacterial reduction (i.e., Supplementary Figure 1 for MOI 1:1) and doses with MOI smaller than 1:1000 were not tested. In most cases, no significant differences were observed in the phage activity while varying calcium concentration (up to 10 mM Ca2+, Supplementary Figure 1) in broth. However, significant increases of inhibitory activity were observed using phages AZT7 and AZT9 when Ca2+ was increased over this concentration (results not shown).

Lytic spectrum (**Table 1**) showed high diversity among isolated phages. AZT6 showed the highest activity against four strains, while AZT5 is specific for SFO001 and AZT1 and AZT8 have a narrow activity range. None of the phages were active against all the considered strains.

Two phages were active against S. marcescens strains. AZT9 reduced populations of CECT977 by 35%, and AZT4 reduced those of CECT854 by 46% when the bacteria were grown in TSB supplemented with 2 mM Mg2+ and 2 mM Ca2+. That lytic activity was low, so those phages have only weak impacts on the tested S. marcescens strains. None of the isolated phages were active against both strains.

Only AZT6 was selected for further studies. Reasons to discard other isolations include quick recovery of maximal growth rate after treatment (phages AZT1, AZT2, and AZT3), no activity at low MOI (AZT4 and AZT9), low stability at −80◦C (AZT5 and AZT8) and maximal titrations below 8.5 Log pfu/ml (AZT7, AZT10, and AZT11).

### AZT6 Characteristics

Four strains of C. maltaromaticum, S. putrefaciens, Y. intermedia and Vibrio spp., isolated in our lab from spoiled fishes (Alfaro and Hernández, 2013), were treated at high MOI to test AZT6 phage specificity. For all strains of these bacteria, our results (not shown) showed that phage-treated and -untreated samples had similar kinetic growth parameters (Lag phase duration, growth rate, maximal yield). As expected, AZT6 was specific for Serratia and did not affect other genera of spoilage bacteria. AZT6 has a broad host range and produced lytic calves with both S. fonticola (SFO001 and SFO002) and three S. proteamaculans (SPR001, SPR006, and SPR009; **Table 1**).

AZT6 reduced the growth rate and delayed the stationary phase of SPR009 when that strain was cultivated between 4 and 20◦C (MOI 1:1, results not shown), demonstrating activity in the temperature range of food storage. At higher temperatures, bactericidal effects disappeared after the initial 24 h, when

FIGURE 2 | Optical density (OD600) evolution of strain SPR009 cultivated in Long & Hammer broth at 20◦C treated with cultures AZT3 at different MOI. (A) No phages, (B) 100:1, (C) 1:1, (D) 1:1000.

### TABLE 1 | Lytic spectra of isolated phages against considered bacterial strains.

aPhage total count was 2 × 106 pfu/ml using SFO001 as sensible strain.

b+++, >105 pfu/ml; ++ 103 to 105 pfu/ml; +, 101 to 103 pfu/ml; −, no visible calves.

bacteria had reached to stationary phase, but 48 h or longer was necessary to verify bacterial growth reduction at low temperatures.

Cultivation broth had an important effect on phage activity. In clear medium (Long & Hammer) MOI of 1:500 or higher produced significant reduction in bacterial counts, but in fish broth, an MOI of 1:3 was required to reduce growth of the Serratia population. Along with other factors, fish extract presents high numbers of particles in suspension that would reduce the activity of the bacteriophages.

Phage titration of AZT6 was stable during storage at cold temperatures. In pure culture, phage concentration was reduced 10-fold in the first 30 days at 6◦C (from 8.9 ± 0.0 to 7.9 ± 0.2 Log pfu/ml), after which constant titration lasted up to 6 months at 4 ◦C (from 7.9 ± 0.2 to 7.7 ± 0.2 Log pfu/ml). Ultra-cold storage (−80◦C, 30 days) in the presence of glycine (20%) resulted in a titration reduction of an order of magnitude.

### Phage Activity in Liquid Medium

In order to select the best strategy for increasing the shelf life of fish products stored at 6◦C, different MOI were tested in Long & Hammer broth. As observed in **Table 2**, MOI of 20:1 or higher have a high impact on bacterial load during initial storage (1–3 days), but then their effect decreased up to day 7, when no reduction was observed. Lower MOI did not initially reduce the bacterial growth or have a significant effect after three or more storage days. These results would agree with the expected mode of action of bacteriophages: at high concentration, bacteriophages cause initially a high bacterial decrees and days after treatment, phage-resistant bacteria growth rapidly. In the other hand, low phage dosage requires some time to propagate the infection and to reduce bacterial count significantly.

Phage activity under real fish-storage conditions (6◦C) was tested in fish extract medium using SPR009 as the sensible strain. High phage:bacteria ratios significant reduced the bacterial concentration, up to 99.9% of inhibition after 5 days (**Table 3**). Use of a MOI lower than 1:1 had no effect on bacterial concentration at the tested storage times. Addition of Ca2+ (up TABLE 2 | Bacterial counts reduction (control – AZT6 treated samples, strain SPR009; Log cfu/ml) in Long & Hammer medium stored at 6◦C using different MOI.

∗Significant differences between treated and control samples (p < 0.05) at different sampling times.

TABLE 3 | Bacterial counts reduction (control – AZT6 treated samples, strain SPR009, Log cfu/ml) in fish extract stored at 6◦C for 5 days using different initial AZT6 and bacterial concentrations.

∗Bacterial reduction using the untreated samples as controls. In all cases, differences are statistically significant.

to 5 mM) to the fish extract did not affect phage activity (results not shown), similar to the results obtained in Long & Hammer broth.

### Phage Activity in Fish Filets

Determine the specific counts of Serratia in a wild environment represented an analytical problem in this research. Medium SSA

Hernández Bacteriophages for Food Spoilage Control

has been described as selective for Serratia (Starr et al., 1976) and does not allow the growth of other representative strains of major bacterial genera in spoiled T. trachurus: Yersinia, Photobacterium, Vibrio, Shewanella, and Carnobacterium. Only Serratia formed visible colonies in that medium (results not shown), which supports that counts in SSA mainly were Serratia. Nevertheless, we cannot fully discount that other bacteria might grow in this medium.

Phage AZT6 was effective when applied to raw fish filets, and it can reduce of the Serratia load after 6 days by more than 90%. When fresh fish filets were treated with high phage–concentration solutions (7.8 Log pfu/g, MOI 350:1) results showed a significant reduction of bacterial counts in SSA after 3 (−0.4 Log cfu/g) and six storage days (−1.1 Log cfu/g) in treated samples compared to control samples (**Table 4**, bach 1).

As expected, reduction in Serratia counts in treated filet samples compared with untreated ones were parallel to significant reductions (p < 0.05) of total bacterial counts after 3 and 6 days in TSA agar (−0.7 and −0.8 Log cfu/g respectively) and after 3 days (0.6 Log cfu/g) in Long & Hammer agar (**Table 4**, bach 1). In this medium, no differences were observed after 6 days, probably because bacterial counts had reached their maxima. Other bacteria genus, like Yersinia, Photobacterium and Carnobacterium, are counted in Long & Hammer agar and have been identified as important spoilage bacteria in T. trachurus (Alfaro and Hernández, 2013). These genera can grow vigorously in refrigeration (Alfaro et al., 2013) and, presumably, compensate the decrease in Serratia counts. A combination of bacteriophages active against different species would be a good strategy to reduce total bacteria and to increase the product's shelf life. To the best of our knowledge, no examples of bacteriophage mixture against different genera in food have been published, but the concept has been proved successfully in pharmacology (Chan and Abedon, 2012) and further research would validate this strategy as food preservation technique.

When fish filets from other batches were treated with two phage dosages (MOI 100:1 or 30:1), the one with the highest phage:bacteria ratio showed significantly lower Serratia concentration after 3 days (−1.5 Log cfu/g), but loads were comparable after six spoilage days (**Table 4**, bach 2). These dosage effect differences were not detected in TSA or Long & Hammer agar (**Table 4**, bach 2).

Nevertheless, no count reductions similar to those in SSA, TSA, or Long & Hammer agar were observed in the filets treated with phages at MOI ≤ 10:1 (results not shown), even in the initial days. Phage AZT6 was active in broth at low MOI, and other researchers have described experience with phages active at low MOI (1:1) in pork adipose tissue (Greer and Dilts, 2002), but AZT6 required an MOI higher than 10:1, as have other phages described in the literature (Guenther and Loessner, 2011). It can be speculated that different strains of Serratia active on different fish samples could modify the bacterial response to phage treatments, that AZT6 is not able to multiply in or remain stable in the mackerel food matrix. As can be observed in **Figure 3**, when MOI ≤ 10:1 were used, AZT6 concentration in fish muscle increased in the first 2 days, and then decreased along fish spoilage; phage titrations went from the initial 5.9 Log pfu/ml to near the detection limit after 6 days. AZT6 titration in buffer solution was stable under the experimental conditions (time, temperature, pH), and no a clear cause was found to explain the decreases on the filets. Stability of other phages tested in food matrices were dependent on the food pH (Leverentz et al., 2001) or, as we expected for AZT6, they were stable in food matrices throughout storage (Guenther et al., 2009). Phages can be stabilized by absorbing them into water–base matrix (Murthy and Engelhardt, 2012) or microencapsulated (Ma et al., 2008;

TABLE 4 | Bacterial concentrations (Log cfu/g) in fish filets treated (MOI 350:1) or not with AZT6 after 3 and 6 days at 6◦C.

∗Significant differences between treated and untreated fish filets (p < 0.05) at each sampling times.

Dini et al., 2012), that would result in a controlled bacteriophage release.

Phage adherence and penetration in fish samples are fast. Just 25 min after immersing fish filets in an AZT6 solution at 7.9 Log pfu/ml, we observed that phage concentration in the muscle increased from an initial value below 3.5–5.9 Log pfu/g (**Figure 3**). Phage AZT6 that settled on the filet surface penetrated by passive diffusion into the fish muscle that seems to be a fast and efficient process.

Our first attempt to test the phage activity in a solid matrix was to inoculate sterile fish cubes (autoclaved at 121◦C for 15 min) with phage-sensitive SFO001 Serratia strain, and then treating them with high phage MOI (1000:1). After 24 and 96 h at 6◦C, there was no bacteriophage effect on bacterial counts (results not shown). It is well-known that phage activity is favored in liquid media, and changes in muscle texture and reduction in food moisture would reduce phage activity. Phages are not motile organisms and their distributions are strongly dependent on conditions in food matrices. That is why most reported uses of bacteriophage involve adding them to liquid foods, like beer (Deasy et al., 2011) or milk (Modi et al., 2001; García et al., 2007). In only a few cases have they been shown to act on food surfaces (Guenther and Loessner, 2011).

Very few papers have been published about using phages for biocontrol of unintentionally contaminated foods. Probably, the report most similar to our study is that by Greer and Dilts (1990), who reported the inability of phages to control the spoilage of beef by wild Pseudomonas. The results presented in this paper not only demonstrate that the reduction of Serratia populations during spoilage is possible, but also demonstrated that many factors have to be adjusted for useful effective applications. Natural resistance is considered a problem for industrial applications. A mixture of phages providing broad spectra toward multiple target strains could to be an effective strategy. Another strategy would be that used by García et al. (2007), who transformed phages against Staphylococcus aureus, obtaining mutant phages that reduced otherwise bacteriophage-insensitive S. aureus variant levels up to 200-fold. A similar strategy would be useful for bacteriophage development against spoilage bacteria. Other factors, like application duration, have been reported as critical for effective phage activity in food (Leverentz et al., 2004).

### REFERENCES

We have described here a preliminary strategy for effective attenuation of Serratia growth during fish spoilage. Phage AZT6 is a promising candidate for application in order to reduce the Serratia population in freshly butchered fish tissues. It is active at low MOI, can be produced at high titration, and is stable for long periods under refrigeration. It shows bactericidal activity for multiple Serratia species, and we demonstrated its activity in real food matrices subject to spoilage. When MOI was 350:1, the bactericidal effect was up to 90% of the Serratia population in fish filets during storage at 6◦C, linked with a bacterial count reduction in TSA and Long & Hammer agar. No Ca2+ additions were necessary for maximum activity, which would simplify their application in the large-scale fish-processing systems.

### AUTHOR CONTRIBUTIONS

IH was responsible of the conception and design of the work, the acquisition, analysis and interpretation of the data presented in this work.

## FUNDING

This work was partially funded by the Basque Government (Department of the Environment, Regional Planning, Agriculture and Fisheries) and LIFE+ program (project LIFE13 ENV/ES/001048 ENVIPHAGE).

## ACKNOWLEDGMENTS

Author thanks Cristina Garcia for her excellent technical support and Begoña Alfaro for her critical review. Transmission microscopy was performed in the Basque Country University (UPV/EHU general service facilities, Leioa, Spain).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2017.00449/full#supplementary-material

application. Regul. Toxicol. Pharmacol. 43, 301–312. doi: 10.1016/j.yrtph.2005. 08.005

Interaction, Vol. 1, eds M. R. J. Clokie and A. Kropinski (New York, NY: Springer).

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Hernández. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Phage Therapy: Combating Infections with Potential for Evolving from Merely a Treatment for Complications to Targeting Diseases

Andrzej Górski1,2,3 \*, Ryszard Mi ˛edzybrodzki1,2,3, Beata Weber-D ˛abrowska1,2 , Wojciech Fortuna1,2, Sławomir Letkiewicz2,4, Paweł Rogóz˙ 1,2, Ewa Jonczyk-Matysiak ´ 1 , Krystyna D ˛abrowska1 , Joanna Majewska1 and Jan Borysowski3

1 Bacteriophage Laboratory, Ludwik Hirszfeld Institute of Immunology and Experimental Therapy, Polish Academy of Sciences, Wroclaw, Poland, 2 Phage Therapy Unit, Ludwik Hirszfeld Institute of Immunology and Experimental Therapy, Polish Academy of Sciences, Wroclaw, Poland, 3 Department of Clinical Immunology, Transplantation Institute, The Medical University of Warsaw, Warsaw, Poland, 4 Katowice School of Economics, Katowice, Poland

### Edited by:

Peter Mullany, University College London (UCL), UK

### Reviewed by:

Victor Krylov, I. I. Mechnikov Research Institute of Vaccines and Sera, Russia Diana R. Alves, Blond McIndoe Research Foundation, UK

> \*Correspondence: Andrzej Górski<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Antimicrobials, Resistance and Chemotherapy, a section of the journal Frontiers in Microbiology

Received: 11 July 2016 Accepted: 09 September 2016 Published: 26 September 2016

### Citation:

Górski A, Mie¸ dzybrodzki R, Weber-Da¸ browska B, Fortuna W, Letkiewicz S, Rogóz P, ˙ Jonczyk-Matysiak E, Da¸ browska K, ´ Majewska J and Borysowski J (2016) Phage Therapy: Combating Infections with Potential for Evolving from Merely a Treatment for Complications to Targeting Diseases. Front. Microbiol. 7:1515. doi: 10.3389/fmicb.2016.01515 Antimicrobial resistance is considered to be one of the greatest challenges of medicine and our civilization. Lack of progress in developing new anti-bacterial agents has greatly revived interest in using phage therapy to combat antibiotic-resistant infections. Although a number of clinical trials are underway and more are planned, the realistic perspective of registration of phage preparations and their entering the health market and significantly contributing to the current antimicrobial crisis is rather remote. Therefore, in addition to planning further clinical trials, our present approach of phage treatment carried out as experimental therapy (compassionate use) should be expanded to address the growing and urgent needs of increasing cohorts of patients for whom no alternative treatment is currently available. During the past 11 years of our phage therapy center's operation, we have obtained relevant clinical and laboratory data which not only confirm the safety of the therapy but also provide important information shedding more light on many aspects of the therapy, contributing to its optimization and allowing for construction of the most appropriate clinical trials. New data on phage biology and interactions with the immune system suggest that in the future phage therapy may evolve from dealing with complications to targeting diseases. However, further studies are necessary to confirm this promising trend.

Keywords: bacteriophage, phage therapy, antibiotic resistance, anti-phage antibodies, inflammation, reactive oxygen species, compassionate use

## IS IT ETHICAL TO CONTINUE NOT PURSUING PHAGE THERAPY? (Henein, 2013)

On May 19, 2016 a review on antimicrobial resistance (AMR) commissioned by the UK Prime Minister was published. AMR – considered to be a challenge to health and our entire civilization – compared to terrorism and global warming – costs some 700,000 deaths annually and, if not controlled, would lead to 10 million deaths by 2050, exceeding the toll of cancer. To prevent medicine being cast back to the dark ages, the review suggests a number of actions promoting