Effectiveness of Bacteriophages Against Biofilm-forming Shiga-toxigenic Escherichia coli In-vitro and on Food-Contact Surfaces

1. Introduction

Shiga-toxigenic Escherichia coli (STEC) are human pathogens responsible for multiple foodborne disease outbreaks. Their infections can range from mild to severe bloody diarrhea and hemorrhagic colitis (HC), leading to life-threatening complications, such as hemolytic-uremic syndrome (HUS) and thrombotic thrombocytopenic purpura [1]. While the STEC serotype, O157:H7, is frequently associated with HC and HUS, other non-O157 serotypes, reported to cause similar infections, have also emerged. These include the top 6 serotypes, O26, O45, O103, O111, O121, and O145, that have been responsible for multiple outbreaks and infections in the last two decades [2,3,4]. According to the Centers for Disease Control and Prevention (CDC) estimates, STEC O157 and non-O157 together cause more than 265,000 illnesses each year in the United States [5]. These infections could be due to the consumption of contaminated fresh produce, undercooked meat, unpasteurized milk, or drinking water [6]. Other factors include, person to person contact, or livestock handling and contact with their environment on the farm, livestock events or through petting zoos [7].

These pathogens can also adapt to adverse environmental conditions by forming biofilms on a wide variety of food-contact surfaces, as well as on fresh produce and meat products [8,9,10]. Biofilms consist of a network of adhesive carbohydrates, exopolysaccharides (EPS), that are difficult to penetrate and can protect the bacteria from stressful conditions and antimicrobials [11,12]. It has been reported that STEC biofilms have an increased tolerance to common sanitizers such as chlorine and quaternary ammonium compounds [10,13,14,15]. This could be due to a combination of bacterial resistance mechanisms such as, diffusional resistance of the EPS matrix, chemical and enzymatic inactivation of sanitizers and disinfectants, physiological changes in the cell, and induction of stress responses in the cell [16]. The ability of these pathogens to form strong biofilms poses a significant threat of cross-contamination to the food-processing industry. It is therefore critical to develop effective strategies to prevent, remove or control biofilms in the food industry for improved food safety.

Bacteriophages are viruses that infect and kill bacteria and have garnered significant attention as antibacterial agents, primarily due to their target-specificity towards the host bacteria [17,18,19]. They are present as commensal microflora in the gastrointestinal tract of animals and have also been isolated from various food and water sources [20,21]. The host-specificity and ubiquitous nature of lytic bacteriophages makes them highly desirable as antibacterial agents [18]. Under in-vitro conditions, virulent bacteriophages have shown the potential for selective elimination of foodborne pathogens [22,23]. Attention has also been given to the use of bacteriophages for removal of biofilms formed by foodborne pathogens on various food-contact surfaces [24,25,26,27]. Viazis et al. [27] tested a phage cocktail against E. coli O157:H7 biofilms on stainless steel (SS), ceramic tile, and high-density polyethylene (HDPE) coupons. The phage cocktail effectively reduced pathogen populations in biofilms on all three surfaces, within an hour of the treatment. In another study, phage treatment on spinach-harvesting blade reduced E. coli O157:H7 in biofilms by 4.5 logs CFU/blade [25]. The majority of these studies have been limited to evaluating bacteriophage potential against E. coli O157:H7 and its biofilms. Very little is known about the effectiveness and applicability of phages against non-O157 STEC and their biofilms. The objective of the present study was to evaluate the efficacy of previously isolated and characterized bacteriophages [28,29] against O157 and non-O157 STEC biofilms in-vitro and on food-contact surfaces.

2. Materials and Methods

2.1. Bacterial cultures

The STEC isolates used in the study included, E. coli O157:H7 (ATCC 43895, wild type (WT): LF4, KF10); O26 (CDC 2003-3014, WT: QF6, BF8); O45 (CDC 2000-3039, WT: EF2, AF1); O103 (CDC 2006-3008, WT: GF6, AF10); O111 (CDC 2010C-3114, ATCC: 2440, 2180), O121 (CDC 2002-3211, ATCC: 2219, 2203); and O145 (CDC 99-3311, ATCC: 2208, 1652). The WT isolates were retrieved from the Jaroni laboratory culture collection, originally isolated from bovine feces or cattle farm environment [30]. Biofilm forming capability of these isolates was previously tested in-vitro [31].

For in-vitro studies, overnight cultures of STEC isolates were prepared in Luria Broth (LB; MP Biomedicals, CA) and incubated while shaking (180 rpm; V.390W, Fisher Scientific, NJ) at 37°C for 16 h. For phage propagation and food-contact surface studies, overnight cultures were prepared in tryptic soy broth (TSB; Bacto™, Difco, BD, Sparks, MD) and incubated statically at 37°C for 18 h. Bacterial cocktail suspensions were prepared by mixing (1:1) isolates of STEC serotypes, at a population of 10⁹ CFU/ml (Table 1,Table 2,Table 3).

2.2. Phage preparation

Phages were propagated by suspending 100 µl overnight culture of host bacterium in molten (0.75%) NZCYM agar (RPI Corp, IL; Fisher Scientific, NJ) and plating via double layer agar method [28]. Phage working-stock solutions were prepared as previously described [28] and stored at 4°C, until further use. Prior to an experiment, phage titers were determined as plaque forming units per ml (PFU/ml), by serially diluting the phage working-stock in phosphate buffered saline (PBS: pH 7.4: NaCl, KCl, NaH2PO4 and KH2PO4, Sigma-Aldrich, MO, USA) and performing a plaque assay [28]. All phage treatments were prepared at a population of 10⁹ PFU/ml.

2.3. STEC biofilm disruption

2.3.1. In vitro biofilm disruption

In-vitro STEC biofilm disruption was determined as previously described [32], in the following three experiments (I, II, III). In Experiment-I, previously isolated and characterized phages (n=52) were used individually to test their efficacy against their host STEC biofilms. Based on the results from Experiment-I, phages were selected to prepare host-specific cocktails (Table 1), which were then tested in Experiment-II. In Experiment-III, a multi-phage cocktail, containing 21-phages (3 phages per STEC serotype), was tested against biofilms of multi-serotype STEC cocktail (Table 2).

Overnight pathogen cultures were diluted (1:100) in M9 medium (MP Biomedicals, CA), supplemented with 0.4% (wt/vol) glucose (Fisher Scientific, NJ) and minerals (1.16 mM MgSO4, 2 µM FeCl3, 8 µM CaCl2, and 16 µM MnCl2; Fisher Scientific, NJ), and incubated with shaking (180 rpm) for 24 h at 37°C. Following incubation, bacterial cultures were further diluted (1:100) in M9 medium (containing glucose and minerals) and allowed to form biofilms in 96-well microtiter plates (Thermo Scientific, Denmark) by aliquoting 150 µl in each well (in triplicates) and incubating the plates at 37°C for 24 h. Wells filled with sterile M9 were used as the negative control. After incubation, the liquid culture was carefully removed using a micropipette and the wells washed three times with PBS (150 µl), without disturbing any biofilm formed at the bottom of the wells. Plates were dried at 37°C for 15 min and 150 µl of the respective bacteriophage treatment (individual or cocktail) or PBS (positive control) was added to the wells. Plates were incubated further at 37°C for 0, 3, and 6 h. After each incubation period, the treatment solution was removed and the wells washed with PBS, as described above. After drying at 37°C for 15 min, biofilms were stained by placing crystal violet (CV; Fisher Scientific, NJ) solution (0.1% in distilled water; wt/vol) in the wells for 2 min, washing thrice with PBS, and drying (37°C for 15 min). The stain was released with 150 µl of ethanol:acetone solution (80:20; vol/vol; Fisher Scientific, NJ) and biofilm disruption quantified by measuring the optical density (OD) at 595 nm (SpectroMax M3; Molecular Devices LLC., CA). Wells filled with ethanol:acetone solution were used as the blank.

2.3.2. STEC biofilm disruption on food-contact surfaces

The efficacy of bacteriophages to disrupt STEC biofilms on food-contact surfaces was studied in two experiments (I and II). In Experiment-I, a cocktail of each STEC serotype (described in 2.1) was used to form biofilms on stainless steel (SS) and high-density polyethylene (HDPE) coupons and treated with their respective phage cocktail for 16 h (Table 3). In Experiment-II, a 21-phage cocktail was evaluated against biofilms formed by a cocktail of 14 STEC isolates (2 isolates per serotype), on SS and HDPE coupons, for 0, 3, 6 and 16 h (Table 2).

2.3.2.1. Preparation of coupons

Stainless steel (304 finish, type 4; Stillwater Steels, OK) and HDPE (1/8”x 24”x 48”; Polymersan, Hialeah, FL) coupons (2 x 5 cm²) were used for the study and prepared as described by Hood and Zottola [33]. Coupons were cleaned by soaking in acetone (30 min), followed by distilled water rinse (5 min), soaking (1 h) in 1 N NaOH (Fisher Scientific, NJ) and sonicating (40KHz; Branson, CT) in distilled water (1 h). Following sonication, coupons were rinsed in distilled water, air-dried and sterilized prior to use.

2.3.2.2. Phage treatment of STEC biofilms

For each STEC serotype, 3 coupons were inoculated, where one coupon was designated as the inoculated, untreated control, one was treated with the respective phage cocktail, and one treated with PBS (control). In each experiment (I and II), coupons were first immersed in 30 ml suspension of bacterial cocktail (10⁹ CFU/ml) in a 50 ml centrifuge tube (Fisher Scientific, GA) and incubated at 25°C for 2.5 h, to facilitate bacterial attachment to the surface. Coupons were gently removed from the suspension using sterile forceps and placed in a sterile 50 ml tube for 5 h in a biosafety hood, allowing further bacterial attachment and biofilm formation. After incubation, coupons were rinsed in 30 ml sterile distilled water to remove unattached bacterial cells from the surface. One coupon of each surface material (SS or HDPE) was sampled to determine the initial pathogen population within the biofilm (inoculated, untreated control). The remaining inoculated coupons were then suspended in 30 ml bacteriophage (10⁹ PFU/ml) treatment (Table 3) or PBS at 37°C for 16 h in Experiment-I, and 0, 3, 6 and 16 h in Experiment-II. After incubation, coupons were sonicated for 5 min at 40 KHz to dislodge bacterial cells from the coupon surface. Immediately following, 3 g of glass beads (Genlantis, CA) were added to the tube and agitated for 1 min, using a vortex (Fisher Scientific, NJ), to remove any remaining attached cells from the coupon [26]. Surviving pathogen populations in the solution were enumerated (CFU/cm²) on TSA or STEC CHROM agar (CHROMagar, Paris, France). Any injured cells were recovered by 24 h enrichment in TSB at 37°C.

2.4. Statistical analysis

All the experiments were repeated 3 times. Surviving STEC populations, recovered after treatments, were converted to log₁₀ CFU/cm² and the mean values of the three replicates obtained. Data were analyzed using General Linear Model (SAS v.9.3 software; SAS Inst., Cary, NC, USA) to determine analysis of variance (ANOVA) for phage treatment effects. Significant differences between the treatment means were separated by the least significant difference (LSD) at P<0.10 for in-vitro studies and P<0.05 for food-contact surface studies.

3. Results and Discussion

3.1. STEC biofilm disruption

Effectiveness of individual and cocktails of phages to disrupt biofilms of 7 STEC serotypes, in-vitro and on food-contact surfaces (SS and HDPE), was determined. Results demonstrated that the phages were more effective in reducing biofilms on food-contact surfaces than in-vitro.

3.1.1. In vitro STEC biofilm disruption

Biofilm disruption by individual, serotype-specific bacteriophages, was determined in-vitro by measuring OD (A₅₉₅) at 0, 3 and 6 h. STEC-biofilm disruption was observed with a reduction in OD from 2.262 nm (0 h) to 0.808 nm (6 h) in phage-treated wells. All phages (n=52) reduced biofilm formation by their respective host bacteria, with 77% showing significant (P<0.10) reductions at 3 and 6 h (Figure 1A-G).

Figure 1. (A). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O157:H7 after treatment for 0, 3, and 6 h.

Figure 1. (A). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O157:H7 after treatment for 0, 3, and 6 h.

Figure 1. (B). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O26 after treatment for 0, 3, and 6 h.

Figure 1. (B). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O26 after treatment for 0, 3, and 6 h.

Figure 1. (C). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O111 after treatment for 0, 3, and 6 h. .

Figure 1. (C). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O111 after treatment for 0, 3, and 6 h. .

Figure 1. (D). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O103 after treatment for 0, 3, and 6 h.

Figure 1. (D). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O103 after treatment for 0, 3, and 6 h.

Figure 1. (E). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O121 after treatment for 0, 3, and 6 h.

Figure 1. (E). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O121 after treatment for 0, 3, and 6 h.

Figure 1. (F). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O45 after treatment for 0, 3, and 6 h. .

Figure 1. (F). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O45 after treatment for 0, 3, and 6 h. .

Figure 1. (G). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O145 after treatment for 0, 3, and 6 h. .

Figure 1. (G). In-vitro biofilm disruption by individual bacteriophages specific to E. coli O145 after treatment for 0, 3, and 6 h. .

In Experiment-I, where individual phages were tested against their host pathogens, varying results were obtained. All the O157-phages (P1-O157 to P7-O157) showed a reduction in pathogen biofilms at 0 h (A₅₉₅ = 0.472 – 0.696), 3 h (A₅₉₅ = 0.284 – 0.441) and 6 h (A₅₉₅ = 0.202 – 0.321), except phages P3, P5 and P7, where an increase in absorbance was observed at 6 h (Figure 1A). At 3 and 6 h, phage P4-O157 showed the highest reduction compared to the rest of the phages. Non-O157 STEC biofilms, treated with phages, also showed significant (P<0.10) reductions at 0 h (A₅₉₅ =1.034 – 3.853), 3 h (A₅₉₅ = 0.506 – 2.631) and 6 h (A₅₉₅ = 0.202 – 3.577) (Figure 1B-G). However, STEC O45-specific phages were more effective after 3 h than after 6 h treatment. A list of the most effective serotype-specific phages is provided in Table 4.

Based on the results from Experiment-I, selected phage cocktails were tested against their respective STEC-serotype biofilms (Experiment-II). A list of the most effective serotype-specific phage cocktails is provided in Table 4. Results revealed that all O157-phage cocktails (CT-1 to CT-4) were able to disrupt E. coli O157:H7 biofilms, reducing OD from 1.990 – 2.329 at 0 h to 0.576 – 0.636 at 6 h (Figure 2A). The CT-3 cocktail showed the highest reduction from 2.250 at 0 h to 0.748 at 3 h and CT-2 showed the highest reduction at 6 h from 2.319 to 0.576. Among the STEC nonO157 phage cocktails, CT5-O26, CT7-O103 and CT12-O145 performed the best by continuing to reduce biofilms at 6 h (Figure 2B,C). Phages from Experiment-II were selected to make a 21-phage cocktail (3 phages per serotype) to treat multi-serotype STEC biofilms (2 bacterial isolates per serotype). Results showed that phage cocktail reduced biofilm after 3 h of treatment, where OD decreased from 2.561 at 0 h to 1.321 at 3 h (Figure 2D). However, a slight increase in absorbance (2.186) was recorded after 6 h of phage-cocktail treatment (Figure 2D).

Figure 2. (A). In-vitro biofilm disruption by bacteriophage cocktails specific to E. coli O157 after treatment for 0, 3, and 6 h.

Figure 2. (A). In-vitro biofilm disruption by bacteriophage cocktails specific to E. coli O157 after treatment for 0, 3, and 6 h.

Figure 2. (B). In-vitro biofilm disruption by bacteriophage cocktails specific to E. coli O121 and O145 after treatment for 0, 3, and 6 h.

Figure 2. (B). In-vitro biofilm disruption by bacteriophage cocktails specific to E. coli O121 and O145 after treatment for 0, 3, and 6 h.

Figure 2. (C). In-vitro biofilm disruption by bacteriophage cocktails specific to E. coli O26, O45, O103 and O111 after treatment for 0, 3, and 6 h.

Figure 2. (C). In-vitro biofilm disruption by bacteriophage cocktails specific to E. coli O26, O45, O103 and O111 after treatment for 0, 3, and 6 h.

Figure 2. (D). In-vitro STEC biofilm disruption by 21-phage cocktail after treatment for 0, 3, and 6 h. Values represent the average of three replications. Different letters a, b etc. represent significant difference (P<0.10) between incubation times (0, 3, 6 h) for the control or phage treatment. The bars with the same letters indicate that there is no significance difference (P<0.10) between incubation times (0, 3, 6 h) for the control or phage treatment..

Figure 2. (D). In-vitro STEC biofilm disruption by 21-phage cocktail after treatment for 0, 3, and 6 h. Values represent the average of three replications. Different letters a, b etc. represent significant difference (P<0.10) between incubation times (0, 3, 6 h) for the control or phage treatment. The bars with the same letters indicate that there is no significance difference (P<0.10) between incubation times (0, 3, 6 h) for the control or phage treatment..

In the current study, a slight increase in OD was observed with some phages and phage cocktails at 6 h treatment. Results of the current study agree with other studies assessing phage treatment effects on bacterial biofilms [34,35]. Chan et al. [34] observed an increase in E. coli biofilms treated with T-4 phage on day 4, after an initial decrease (30%) on day 3. Similarly, Hughes et al. [36] obtained a maximum reduction in biofilm within 2 h or 5 h, depending on the biofilm studied. Factors such as bacterial appendages, used by the bacteria for biofilm attachment, could decrease the efficacy of phages in the biofilm [36,37]. Additionally, changes in bacterial biofilm profile over time, such as inconsistent expression and accumulation of protein through biofilm development stages, could affect phage efficacy [38,39]. Higher protein production at later stages of biofilm formation could impair phage movement within the medium and interfere with phage efficacy against bacterial biofilm [34,40]. Furthermore, static biofilms are known to produce higher polysaccharide and protein content, which could interfere with phage infection efficacies [41]. Vogeleer et al. [32] showed variation in biofilm matrices of individual non-O157 STEC serotypes under static and dynamic conditions. This variation in biofilm matrix could minimize phage effectiveness against biofilm, due to the highly specific nature of its depolymerase enzyme [24,36]. This enzyme is highly specific to EPS produced by host-bacteria, and even a minor change in the EPS composition could prohibit its activity, leading to reduced phage activity against biofilm.

Loss in phage effectiveness after 6 h could also be due to generation of phage resistant mutants [42]. Incubation of biofilms at optimum temperatures (30-37°C) could trigger the rapid growth of bacteria in the biofilm, including phage-resistant bacteria, resulting in decreased efficacy of the phage treatment [37,43]. It has also been shown that STEC serotypes, grown together to form biofilms, could generate an abundance of morphological variants, which could exhibit varying susceptibility to treatments [32,44]. Some newly generated mutants could lose the receptor responsible for bacteriophage susceptibility and become resistant to phages. The use of more diverse phages with the ability to bind to different bacterial receptors could help in controlling the emergence of bacterial mutants by exerting selective pressure on bacterial population in the biofilm and increasing the effectiveness of phage cocktail treatment [45]. At the same time, phage resistance has been shown to be transient in bacterial cells and they could revert to phage susceptible state [46]. Additionally, this decrease in phage effectiveness can be overcome by application of a cocktail of multiple phages [46,47]. Multi-phage cocktails were therefore evaluated in the current study for their effectiveness against STEC biofilms.

3.2. STEC biofilm disruption on food-contact surfaces

Based on the in-vitro experiments, the most effective phage cocktails were selected to conduct application studies on SS and HDPE coupons. In Experiment-I, where STEC serotype-specific cocktails were used to form biofilms on SS and HDPE coupons, phage cocktails significantly (P<0.05) reduced bacterial population in the biofilm when compared to the control (Figure 3A,B). Populations of E. coli O45, O111, O121, and O145 on SS were reduced to undetectable levels after treatment with respective phage cocktails (Figure 3A). A reduction of 3.1 and 3.7 logs was observed in E. coli O26 and O103 populations on SS coupons treated with phages, compared to the control (Figure 4A). On HDPE surface, E. coli O121 population was reduced to undetectable levels after respective phage cocktail treatment (Figure 3B). A reduction between 2.3-5.3 logs was observed in E. coli O26, O45, O103, O111 and O145 populations on HDPE coupons treated with respective phage cocktails, compared to the control (Figure 4B). Phage cocktails for specific STEC serotypes were very effective in reducing pathogen populations on both, SS (1.9-4.1 logs CFU/cm² reduction) and HDPE (2.3-5.6 logs CFU/cm² reduction) surfaces. These cocktails were more effective on SS than on HDPE, with the phage cocktails for E. coli O45, O145, O111, and O121 reducing pathogen populations to undetectable levels. These differences could be due to the surface type or difference in the biofilm composition, where different types of EPS were produced, which could provide increased resistance to bacteria against phage [47,48,49].

In Experiment-II, biofilms of a cocktail of 14 STEC isolates (two isolates per serotype), on SS and HDPE coupons, were treated with a 21-phage cocktail (three phages per serotype). The cocktail showed an immediate (0 h) reduction in STEC populations on HDPE (1.5 logs CFU/cm²) and SS (1.0 log CFU/cm²) (Figure 4A,B). At 3, 6 and 16 h, the 21-phage cocktail showed significant reductions (P<0.05) in STEC populations on both the surfaces when compared to the control (Figure 4A,B). At 3 h, pathogen populations were reduced by 2.8 and 1.7 logs CFU/cm² on SS and HDPE, respectively. At 6 h, pathogen populations were reduced to undetectable levels on SS and by 1.7 logs CFU/cm² on HDPE. At 16 h, STEC populations were reduced to undetectable levels on both surfaces. These results suggest that the tested bacteriophage treatments effectively reduced STEC populations in the biofilms on SS and HDPE surfaces.

Studies have previously shown that bacteriophages can effectively reduce pathogens attached to hard surfaces found in food processing environments [26,50,51]. Reductions between 3.5 to 5.4 logs CFU, in attached Listeria monocytogenes population, were observed on SS after 24 h of phage P100 treatment [52]. In a study by Sharma et al. [26], lytic bacteriophages reduced E. coli O157:H7 populations by 1.2 logs CFU on SS coupon. In the current study, phage treatments reduced E. coli O157:H7 by 1.9 logs CFU/cm² and non-O157 STEC by 3.1-4.1 logs CFU/cm² on SS surface. Studies have shown that phages can diffuse through the biofilm formed by bacteria [53] and that the formation of biofilms does not provide additional protection to bacteria against phage attack [26]. Biofilm break-down mechanism of phage is speculated to be associated with enzymatic means. Studies have shown that phages produce enzymes that can degrade the EPS layer, the major component of a biofilm matrix [24,36]. Hughes et al. [36] showed that bacteriophage specific for Enterobacter agglomerans disrupted biofilm through a combination of lytic activity against bacterial cells and degradation of EPS through phage-associated polysaccharide depolymerase enzyme. The presence of phage-depolymerase in an O45-phage (P-9), used in the current study, has been confirmed (data not shown), suggesting that other phages in the cocktail tested may also produce EPS-degrading enzyme, resulting in reduction of bacteria embedded in the biofilm matrix.

Variations in pathogen attachment and biofilm formation were observed on the food- contact surfaces tested in the present study. Higher STEC attachment was observed on HDPE (up to 5.6 logs CFU/cm²) compared to SS (up to 4.3 logs CFU/cm²) surface. A similar trend has been observed in previous studies. Higher bacterial attachment and biofilm formation has been observed on plastic surfaces (polyethylene, polypropylene, polyvinyl chloride) than on stainless steel or glass surfaces [48]. Variations found in polymer surfaces, such as smoothness, charge, Zeta-potential and active chemical groups, could influence bacterial attachment to the surface [54]. Studies have also shown that a variety of active chemical groups released by HDPE surfaces, such as phenols, quinones, aromatic hydrocarbons, aldehydes, and ketones, could be utilized by bacteria as carbon sources and result into higher bacterial attachment [55,56,57].

5. Conclusions

This study has shown the potential for use of lytic bacteriophages, against 7 STEC serotypes as a treatment to control biofilm formation. Future studies need to be conducted to understand phage-bacteria interactions in biofilm using kinetic modeling. The preliminary results from the current study show promise, due to the fact that phages within the biofilms effectively targeted and lysed STEC and were also able to disperse the extracellular matrix forming the biofilm.