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. 2015 Jun 19;81(14):4841–4849. doi: 10.1128/AEM.00812-15

Liposome-Encapsulated Bacteriophages for Enhanced Oral Phage Therapy against Salmonella spp.

Joan Colom a, Mary Cano-Sarabia b, Jennifer Otero a, Pilar Cortés a, Daniel Maspoch b,c,✉, Montserrat Llagostera a,✉
Editor: M J Pettinari
PMCID: PMC4551199  PMID: 25956778

Abstract

Bacteriophages UAB_Phi20, UAB_Phi78, and UAB_Phi87 were encapsulated in liposomes, and their efficacy in reducing Salmonella in poultry was then studied. The encapsulated phages had a mean diameter of 309 to 326 nm and a positive charge between +31.6 and +35.1 mV (pH 6.1). In simulated gastric fluid (pH 2.8), the titer of nonencapsulated phages decreased by 5.7 to 7.8 log units, whereas encapsulated phages were significantly more stable, with losses of 3.7 to 5.4 log units. The liposome coating also improved the retention of bacteriophages in the chicken intestinal tract. When cocktails of the encapsulated and nonencapsulated phages were administered to broilers, after 72 h the encapsulated phages were detected in 38.1% of the animals, whereas the nonencapsulated phages were present in only 9.5%. The difference was significant. In addition, in an in vitro experiment, the cecal contents of broilers promoted the release of the phages from the liposomes. In broilers experimentally infected with Salmonella, the daily administration of the two cocktails for 6 days postinfection conferred similar levels of protection against Salmonella colonization. However, once treatment was stopped, protection by the nonencapsulated phages disappeared, whereas that provided by the encapsulated phages persisted for at least 1 week, showing the enhanced efficacy of the encapsulated phages in protecting poultry against Salmonella over time. The methodology described here allows the liposome encapsulation of phages of different morphologies. The preparations can be stored for at least 3 months at 4°C and could be added to the drinking water and feed of animals.

INTRODUCTION

Foodborne diseases are a global health problem and include infections with nontyphoidal Salmonella, which is among the most common zoonotic pathogens that affect humans. In fact, this bacterium was responsible for more than 1 million annual cases of food-related illnesses in the United States from 2000 to 2008 (1) and for more than 90,000 cases of salmonellosis (mainly from Salmonella enterica serovars Enteritidis and Typhimurium) diagnosed in the European Union in 2012 (2). The major source of Salmonella infections in humans is poultry products (2), and poultry is the major reservoir of this bacterium. The health risk is exacerbated in the case of broilers, which are asymptomatic carriers that house Salmonella in their gut. In fact, several Salmonella strains persistently colonize chickens but without causing any signs of illness (3). This lack of clinical symptoms facilitates the dissemination of Salmonella within flocks (4), thereby increasing the probability of cross-contamination during the transport, slaughter, and processing of broilers.

Salmonella infection in broilers has been controlled mainly through the use of vaccines (5, 6), probiotics, prebiotics, and synbiotics (7), and antibiotics (8), although the effectiveness of these treatments is limited. Importantly, antibiotics can ultimately increase the severity and frequency of colonization by certain resistant strains of Salmonella (9).

Phages are viruses that infect bacteria, subsequently replicating and then rapidly killing their bacterial host (10). Thus, phage therapy has been proposed as a means to control Salmonella infection in chickens. Indeed, several studies have shown that phages effectively reduce Salmonella enterica colonization in poultry (11–15). The utility of phages in food preservation and safety has also been demonstrated (16–21). In a previous study based on an experimental model of Gallus gallus free of specific pathogens, we showed that a phage cocktail containing the phages UAB_Phi20, UAB_Phi78, and UAB_Phi87 is highly effective in reducing Salmonella colonization; the same results were obtained when diverse foodstuffs were similarly treated (20, 21).

Despite these advances, inherent challenges remain in the practical use of oral phage therapy. For example, phages typically lack stability in the extremely acidic environment of the chicken stomach (22), and their residence time in the intestinal tract is very short (20). These problems translate into a low efficiency of phage infection that can only be compensated for by very frequent treatment, which is costly and time-consuming (20). Thus, the practical use of phages for controlling Salmonella infections in broilers will require phages (or formulations thereof) that are more acid stable and have longer residence times. Two strategies have recently been developed for stabilizing phages in acidic media: (i) the addition of antacids (e.g., CaCO3) to phage suspensions (15) and (ii) the encapsulation of phages into polymeric microcapsules, including those comprising alginate-pectin (23) (size, 1 mm), alginate-chitosan (size, 780 μm) (24), methacrylate (25), and alginate-CaCO3 (size, 900 μm) (26). However, to date, there have been no reports of a method that simultaneously resolved both the acid sensitivity and the limited intestinal residence time of phages to achieve a significant reduction of Salmonella infections in broilers. In considering liposomes, we hypothesized that they would serve two primary functions: (i) as a barrier to protons (27), thus protecting phages against gastric acids, and (ii) as a promoter of mucoadhesiveness, owing to their positively charged surfaces (28, 29), which would prolong the intestinal residence time. We chose cationic liposomes as the encapsulation matrix because they readily allow the encapsulation of negatively charged, nanoscale biological entities (30, 31) such as phages. Furthermore, cationic liposomes favor the dispersion of their biomolecules in aqueous media over prolonged periods of time, facilitate the oral administration of these biomolecules to animals, and are degraded upon contact with intestinal bile salts (32).

In the work reported here, we separately encapsulated the three phages (UAB_Phi20, UAB_Phi78, and UAB_Ph87) in cationic lipid envelopes (liposomes) and then compared the performances of the resulting liposome-encapsulated and nonencapsulated phages as orally administered phage therapy in broilers.

MATERIALS AND METHODS

Bacterial strains and growth conditions.

Salmonella Typhimurium LB5000 (SGSC181; University of Calgary) was used for the propagation of phages UAB_Phi20, UAB_Phi78, and UAB_Phi87, and S. Typhimurium ATCC 14028 Rifr for the colonization of the animals. All bacterial strains were grown in Luria-Bertani (LB) broth or on LB agar plates for 18 h at 37°C. Viable counts of S. Typhimurium ATCC 14028 Rifr were determined by plating on xylose-lysine-deoxycholate (XLD) plates (Laboratorios Conda, Spain) supplemented with rifampin (75 μg/ml), followed by incubation at 37°C for 18 h.

In vitro multiplication of bacteriophages.

UAB_Phi20, UAB_Phi78, and UAB_Phi87 phage lysates were obtained by infecting exponential-phase cultures of S. Typhimurium strain LB5000 grown in LB broth at an input multiplicity of infection (MOI) (ratio of added phages to added bacteria) of 0.01 (33). After a 5-h incubation at 37°C, the cultures were centrifuged at 10,414 × g for 10 min. The supernatants were collected and then filtered first through 0.45-μm and then through 0.22-μm syringe polyestersulfone (PES) filters (Millipore, Carrigtwohill, Ireland). The phage titer was determined by plating serial dilutions (1:10) onto LB plates using the double agar layer method (34).

When high titers of phage lysates were required, 50-ml portions of Salmonella cultures in LB broth at an initial optical density at 550 nm (OD550) of 0.2 were prepared and incubated at 37°C with agitation until an OD550 of 1 was obtained. The cultures were infected with the appropriate phage at an input MOI of 0.01, and the suspensions were then incubated at 37°C for 12 to 14 h (34). Finally, the lysates were recovered by centrifugation as described above. The phage lysates were concentrated by ultracentrifugation (Optima L-80; Beckman, CA, USA) in an 80Ti rotor (Beckman, CA, USA) at 4°C (68,584 × g) for 2 h (35). The pellets were resuspended in aqueous MgSO4 (10 mM, pH 6.1) with overnight shaking. The resulting phage suspensions were filtered through 0.45-μm PES filters and then stored at 4°C. The phage titer was determined as described above.

Encapsulation method.

The phages were encapsulated in lipids using the thin-film hydration method (36). A lipid mixture of 1,2-dilauroyl-rac-glycero-3-phosphocholine (DLPC), cholesteryl polyethylene glycol 600 sebacate (Chol-PEG600), cholesterol (Chol), and cholesteryl 3β-N-(dimethylaminoethyl)carbamate hydrochloride (cholesteryl) was used (1:0.1:0.2:0.7 molar ratio). Briefly, after each lipid was dissolved in chloroform (100 mg/ml), a solution containing 106 μl of DLPC, 17 μl of cholesterol-PEG600, 13 μl of Chol, and 64 μl of cholesteryl was prepared under sterile conditions. The total lipid concentration was 17 mM. The organic solvent was removed under vacuum and nitrogen to afford a dry lipid film, which was then hydrated with 2 ml of the appropriate aqueous suspension of the phage (either UAB_Phi20, UAB_Phi78, or UAB_Phi87) at a concentration of 1011 PFU/ml under stirring for 1 h. Under these conditions, the stacks of liquid crystalline lipid bilayers become fluid and swell, resulting in their detachment during agitation and their self-closure to form large multilamellar vesicles (LMVs). Unilamellar vesicles were obtained by homogenizing the LMV suspension using an extruder (Lipex Biomembranes, Vancouver, Canada) and a polycarbonate membrane (pore size, 400 nm). Particle size distributions of the vesicles were determined using a dynamic light scattering (DLS) analyzer combined with noninvasive backscatter technology (Malvern Zetasizer; Malvern Instruments, United Kingdom). The samples (1 ml) were measured without dilution directly thereafter. The mean of three different measurements was taken as the mean diameter. The stability of the liposome-encapsulated phages was examined by measuring their zeta potential with an electrophoretic mobility and light scattering analyzer (Malvern Zetasizer). The samples were placed into the cuvettes and measured without dilution immediately thereafter. The mean from three different experiments was taken as the mean zeta potential of the dispersed system.

Encapsulation efficiency.

The encapsulation efficiency of the phages was calculated according to the equation EE (%) = 100 − (Cfree/Ctotal) × 100, where EE is encapsulation efficiency, Ctotal is the total phage concentration, and Cfree is the concentration of free phages. To quantify Ctotal, 0.5 ml of liposome-encapsulated phages was treated with 0.5 ml of bile salts (50 mM) (Sigma-Aldrich, MO, USA) to disrupt the liposomes. A preliminary experiment confirmed that this concentration had no significant effect on the infectivity of the phages. The double agar layer method was used to plate appropriate dilutions of the samples onto LB plates with S. Typhimurium strain LB5000, and the Ctotal was then determined. To quantify Cfree, the titers of liposome-encapsulated phages were directly determined by plating serial dilutions (1:10) with strain LB5000. The same method was used to determine the stability of liposome-encapsulated phages that had been stored at 4°C for 3 months.

Microscopy.

Liposome integrity (morphology and lamellarity) was examined by cryogenic transmission electron microscopy (cryo-TEM) using a JEOL-JEM 1400 microscope (JEOL, Japan). The samples were prepared in a controlled environment vitrification system. A 5-μl aliquot of each one was deposited onto carbon-coated film meshes supported by standard copper TEM grids. After 30 s, the grids were gently blotted with a double layer of filter paper to obtain a thin film (thickness, 20 to 400 nm), plunged into liquid ethane at its freezing point (−180°C), and then transferred into liquid nitrogen (−196°C), where they were stored until use. At that time, the vitrified specimens were transferred to the microscope using a cryotransfer and its workstation (626 DH; Gatan).

To confirm that the phages had indeed been encapsulated within liposomes, fluorescently labeled samples were observed by laser confocal microscopy using a Leica TCS SP5 confocal microscope (Leica Microsystems, Germany). Phages were stained with 100× SYBR gold (Molecular Probes, OR, USA) by adding 0.02 ml of SYBR gold to 10 ml of phages suspended in 10 mM MgSO4 to obtain a concentration of 1011 PFU/ml. After an overnight incubation in the dark (37), the samples were ultrafiltered and washed three times at 5,000 × g using Amicon Ultra 50K tubes (Millipore). The fluorescent SYBR gold-labeled phages were encapsulated using the thin-film hydration method described above and using the DLPC–Chol-PEG–Chol–cholesteryl lipid mixture incorporating the Vybrant DiI phospholipid-labeling solution (10 μl of Vybrant DiI per 20 mg lipid). In this case, the liposomes were not extruded through a membrane, as the resulting particles would have been below the resolution limit of the optical microscope. A 30-μl sample was placed on a coated glass slide and observed in resonance scanning mode.

Lyophilization.

For the long-term storage of the liposome-encapsulated phages, the cryoprotectant trehalose was added during liposome synthesis, and the encapsulated phages were subjected to lyophilization as follows. The dried lipid film was hydrated with a suspension containing the phages and trehalose at a lipid/carbohydrate ratio of 1:5 (3.2%, wt/vol). The resulting cryoprotected liposome-encapsulated phages were frozen at −80°C for 2 h and then lyophilized at −40°C for 48 h. For comparison, nonencapsulated phages were also lyophilized with the cryoprotectant trehalose. When needed, the lyophilized samples were resuspended in aqueous MgSO4 (10 mM) and plated as indicated above to determine the pre- and postlyophilization phage titers for use in calculating phage infectivity.

Bacteriophage stability in SGF.

The liposome-encapsulated phages were tested for their stability in simulated gastric fluid (SGF) solution (pH 2.8) comprising pepsin (3 mg/ml) (Sigma-Aldrich, MO, USA) in 0.85% NaCl (38). In all cases, 0.1 ml of encapsulated phages was added to 10 ml of the simulated gastric fluid solution. No change in pH occurred. The samples were then incubated in a water bath at 42°C with agitation to mimic the conditions of the avian stomach. To determinate the total phage concentration, aliquots taken at 0, 30, and 60 min were treated with bile salts (50 mM) and plated as described previously. The same methodology, but without the bile salts treatment, was used to test nonencapsulated phages.

In vivo assays in broilers.

Both the intestinal residence time and the effectiveness of a cocktail of either liposome-encapsulated or nonencapsulated phages against Salmonella were evaluated in vivo in commercial broilers (Gallus gallus, Ross strain 308; Terra-Avant S.A., Girona, Spain). The animals were treated in agreement with the guidelines of the ethics commission (Comissió d'Ètica en l'Experimentació Animal i Humana [CEEAH]) of the Universitat Autònoma de Barcelona (UAB). The study was approved and assigned the authorization number 1953.

Newly hatched chickens were purchased and subsequently transported to a biosafety level 2 animal-testing facility at the UAB's Servei de Granges i Camps Experimentals (Cerdanyola del Vallès, Spain). The broilers were housed in pens with food and water supplied ad libitum. In an attempt to simulate farm conditions and increase the feeding impulse of the animals, they were kept under 23-h/1-h light/dark cycles using green and blue light (39). Prior to each of the experiments, tissue samples were obtained from two euthanized chickens and tested using enrichment protocols to confirm that they were free of Salmonella and phages. For the detection of Salmonella, homogenized tissues were incubated in buffered peptone water (BPW) (Merck) at 37°C for 18 h, after which 0.2 ml of tissue suspension was inoculated in 2 ml of Müller-Kauffman selective broth (Merck, Darmstadt, Germany). Following a 24-h incubation at 37°C, 0.1 ml of this culture was plated onto XLD plates. Phages were detected as previously reported (10). Briefly, 0.2 ml of each homogenate was added to a 2-ml culture of Salmonella Typhimurium in LB at an OD550 of 0.2. After 24 h of incubation at 37°C, the presence of phages was determined by spotting 10 μl of the above-described suspensions onto lawns of S. Typhimurium ATCC 14028 Rifr. The plates were incubated at 37°C for 24 h, and bacterial lysis was then evaluated. By this method, fewer than 200 phages per gram of cecum were detected.

In vivo assays were based on the oral administration to the broilers of 100 μl of a cocktail containing either the liposome-encapsulated phages or the nonencapsulated phages. The cocktail comprised a 1:1:1 mixture of the three phages (UAB_Phi20, UAB_Phi78, and UAB_Phi87) at a concentration of 1011 PFU/ml in MgSO4 buffer without any antacid. All oral inoculations were performed using a curved oral-dosing needle (75 mm, 16 gauge; Veterinary Instrumentation, Sheffield, United Kingdom).

In vivo retention in the intestinal tract.

The intestinal residence time of a cocktail of either liposome-encapsulated or nonencapsulated phages was determined over 72 h as follows. Two groups of 63 1-day-old chickens were housed in two poultry pens. At time zero, one group was treated with 100 μl of the liposome-encapsulated phage cocktail and the other with 100 μl of the nonencapsulated phage cocktail. In both cases, the dose was 1010 PFU/animal. After 2, 48, and 72 h, 21 animals from each group were euthanized and cecum samples were collected. For phage quantification, serial dilutions of homogenized cecum samples were plated onto a lawn of Salmonella Typhimurium ATCC 14028 Rifr using the double agar layer method and then incubated at 37°C for 24 h. When direct detection of phages was not possible, the enrichment procedure was carried out as detailed above.

Assay of the efficacy of bacteriophage therapy.

The efficacy of phage therapy against Salmonella was assessed over 17 days and assayed in chickens orally administered 100 μl of a cocktail containing liposome-encapsulated or nonencapsulated phages. The animals were orally infected on day 0 with 100 μl of a suspension of S. Typhimurium ATCC 14028 Rifr in LB medium at a dose of 107 CFU/animal. Three groups of 84 recently hatched commercial broilers (Gallus gallus, Ross strain 308) were separated in pens. Group 1, which controlled for Salmonella colonization, was untreated. Groups 2 and 3 received the nonencapsulated and encapsulated phages, respectively. In both cases, phages in an oral dose of 1010 PFU/animal were administered once daily for 8 days, from day −1 to day 7 postinfection with Salmonella. The control group was orally inoculated over the same 8 days with an aqueous suspension of MgSO4 (10 mM). In each group, 14 chickens were euthanized on days 1, 3, 6, 8, 10, and 15 postinfection, and cecum samples were obtained for Salmonella quantification. To count the salmonellae, the tissues were weighed, resuspended in 4 ml of BPW, and then mechanically homogenized for 15 min. Serial dilutions of the homogenates were plated on XLD agar plates (Laboratorios Conda) supplemented with rifampin (75 μg/ml). After incubation at 37°C for 24 h, the number of colonies of Salmonella was recorded. For each treatment, the reduction in bacteria was calculated by subtracting their mean cecal concentration (expressed in log units) in groups 2 and 3 from the mean value of the group 1 control.

In vitro studies of bacteriophage release.

Phage release from liposomes in the cecum was studied in 12 1-day-old chickens maintained in pens as described above. After 15 days, the animals were euthanized and the contents of their ceca were harvested. For each phage, 990 μl of cecal content was mixed with 10 μl of liposome-encapsulated phages and incubated at 42°C, the body temperature of the chickens. Phage concentration over time was determined by plating the appropriate dilutions using the double agar layer method.

Statistical analysis.

All results were analyzed using IBM SPSS software. For normally distributed samples, analysis of variance (ANOVA) and Student's t test were applied; in cases of nonnormal distribution, the Kruskal-Wallis and Mann-Whitney tests were used. The polydispersity index (PdI), defined as the square of the ratio of the standard deviation to the mean diameter, was determined as a measure of the width of the particle size distribution.

RESULTS

Characterization of the liposome-encapsulated bacteriophages.

Each lytic phage (UAB_Phi20, UAB_Phi78, or UAB_Ph87) (20, 21) was separately encapsulated into liposomes and characterized. DLS indicated that the mean size of the liposome-encapsulated phages was 308.6 ± 20.9 nm for UAB_Phi20 (PdI = 0.14), 320.6 ± 15.2 nm for UAB_Phi78 (PdI = 0.21), and 325.8 ± 23.1 nm for UAB_Phi87 (PdI = 0.18) (Fig. 1). The surface charges of the liposome-encapsulated phages, determined by measuring their zeta potential at pH 6.1, were +35.1 ± 1.0 mV, +33.0 ± 2.1 mV, and +31.6 ± 1.3 mV, respectively (Table 1).

FIG 1.

FIG 1

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(A) Schematic representation of the liposome-encapsulated bacteriophage system, showing the presence of phages (green) encapsulated inside liposomes or attached to their external surfaces. (B to D) Particle size distribution plots for the liposome-encapsulated phages UAB_Phi20 (B), UAB_Phi78 (C), and UAB_Phi87 (D), as measured by dynamic light scattering.

TABLE 1.

Size and zeta potential of the liposomes and encapsulation yield for the three phages freshly prepared and after their storage at 4°C for 3 monthsa

Phage Fresh
 Stored at 4°C for 3 months
Encapsulation yield (%) Size (nm) ζ potential (mV) Encapsulation yield (%) Size (nm) ζ potential (mV)
UAB_Phi20 49 ± 2.4 308.6 ± 20.9 +35.1 ± 1.0 59 ± 9.2 292.7 ± 15.9 +34.2 ± 1.3
UAB_Phi78 48 ± 6.4 320.6 ± 15.2 +33.0 ± 2.1 55 ± 3.8 338.5 ± 12.1 +35.2 ± 1.1
UAB_Phi87 47 ± 5.1 325.8 ± 23.1 +31.6 ± 1.3 43 ± 2.7 302.6 ± 23.5 +32.5 ± 1.6
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a

Each value represents the average from three independent experiments ± the standard deviation.

The morphology and structure of the resulting liposome-encapsulated phages were imaged by cryo-TEM. As shown in Fig. 2A to C, the phages were encapsulated in unilamellar liposomes. The number of encapsulated phages in a single liposome varied; empty liposomes and nonencapsulated phages were also observed. In the latter, most were attached to the external surface of the liposome. The sizes of the liposomes as determined by cryo-TEM were consistent with those estimated by DLS. The encapsulation efficiency of each phage was determined by preparing six independent encapsulation batches. The encapsulation yields (expressed as a percentage) were 49% ± 2.4% for UAB_Phi20, 48% ± 6.4% for UAB_Phi78, and 47% ± 5.1% for UAB_Phi87 (Table 1). To further characterize the physicochemical structure of the liposome-encapsulated phages, they were labeled with SYBR gold and encapsulated into DiI-labeled liposomes. Figure 2D to F shows the three-dimensional (3D) spatial superimposition of the fluorescence intensities of SYBR-gold-labeled UAB_Phi20, UAB_Phi78, and UAB_Phi87 phages (in green) and those of the DiI-labeled liposomes (in red). The images confirmed that each type of phage had indeed been encapsulated inside the liposomes.

FIG 2.

FIG 2

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(A to C) Cryo-TEM images of liposome-encapsulated phages UAB_Phi20 (A), UAB_Phi78 (B), and UAB_Phi87(C). (D to F) 3D confocal microscopy images of SYBR gold-labeled phages UAB_Phi20 (D), UAB_Phi78 (E), and UAB_Phi87 (F) (green) encapsulated into fluorescent DiI-labeled liposomes (red) 3D images of the liposome surface are shown on the left, and the corresponding cross-sectional images on the right. Scale bars, 5 μm.

We then tested the stability of the encapsulated phages, that is, the extent to which the phages remained inside the liposomes. To this end, both the DLS characteristics and the zeta potentials of the freshly encapsulated phages were determined, together with their concentrations. In addition, samples of each freshly encapsulated phage were stored at 4°C for 3 months and subsequently characterized as described above. No significant differences were found in the values of any of the parameters between the fresh and stored encapsulated phages (Table 1), thereby confirming the stability of the liposome-encapsulated phages under the test conditions (4°C and 3 months).

Lyophilization of the liposome-encapsulated bacteriophages.

With the goal of obtaining a stable product suitable for long-term storage and easily administered in animal water or feed, we evaluated the effect of the lyophilization procedure on liposome-encapsulated phages. Prior to lyophilization, the cryoprotectant trehalose (24) was added to the phage-containing suspension during the synthetic hydration step. Nonencapsulated phages were also lyophilized, for the purpose of comparison. After lyophilization, each resulting powder was reconstituted by resuspension in aqueous 10 mM MgSO4, and the phage titer was determined. Table 2 shows the titers, expressed as a percentage, of both the nonencapsulated and encapsulated phages after lyophilization with respect the titers before lyophilization. The values of the encapsulated phages were consistently much higher than those of the corresponding nonencapsulated phages (P < 0.05): roughly 82% and 84% versus 22% and 47% for encapsulated and nonencapsulated UAB-Phi20 and UAB-Phi87, respectively. In the third phage, UAB_Phi78, which was extremely sensitive to lyophilization, the variation between the values obtained for encapsulated and nonencapsulated forms pre- and postlyophilization was much greater, 15% versus 2%, respectively.

TABLE 2.

Effect of lyophilization on the titers of phages

Phage Encapsulation Titer (%)a
UAB_Phi20 No 22.0 ± 13.4
Yes 82.3 ± 15.4*
UAB_Phi78 No 2.0 ± 1.5
Yes 15.1 ± 9.6*
UAB_Phi87 No 47.5 ± 12.1
Yes 84.4 ± 16.9*
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a

The value is the percentage of the titer after lyophilization with respect to the titer before lyophilization. Each value is the average from six independent experiments ± standard deviation. *, P < 0.05.

Bacteriophage resistance to SGF.

The acid stabilities of liposome-encapsulated and nonencapsulated phages were tested in simulated gastric fluid (SGF) at pH 2.8 for 60 min. For the nonencapsulated phages (Fig. 3), after 30 min of incubation, titer losses of 4.2 log units (UAB_Phi20), 6.2 log units (UAB_Phi78), and 6.1 log units (UAB_Phi87) were recorded. Further decreases occurred after 60 min: 5.7 log units (UAB_Phi20), 8.0 log units (UAB_Phi78), and 7.8 log units (UAB_Phi87). In contrast, all three encapsulated phages were less prone to acidic degradation at both time points (Fig. 3) (P < 0.05). Thus, the titer losses after 30 min of incubation were 3.2 log units (UAB_Phi20), 4.0 log units (UAB_Phi78), and 3.3 log units (UAB_Phi87). After 60 min of incubation, losses of 4.8 log units (UAB_Phi20), 5.4 log units (UAB_Phi78), and 3.7 log units (UAB_Phi87) were recorded.

FIG 3.

FIG 3

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Stability of the nonencapsulated (white bars) and liposome-encapsulated (gray bars) phages in simulated gastric fluid (pH 2.8). (A) UAB_Phi20; (B) UAB_Phi78; (C) UAB_Phi87. Each value is the average from six independent experiments ± standard deviation. *, P < 0.05.

In vivo retention in the intestinal tract.

We studied whether the intestinal residence time of phages UAB_Phi20, UAB_Phi78, and UAB_Phi87 in vivo in broilers was prolonged by their encapsulation in cationic liposomes. Two phage cocktails were prepared: one comprising a 1:1:1 mixture of the three liposome-encapsulated phages and the other a 1:1:1 mixture of the three nonencapsulated phages. In both cases, the final concentration was 1011 PFU/ml. Two groups of 63 broilers each were treated with the first (encapsulated) and second (nonencapsulated) cocktails. For each group, 21 chickens were euthanized 2, 48, and 72 h later, at which time the presence of phages in the cecum of each animal was determined. Figure 4 shows the percentage of chickens from each treatment group that contained phages in the cecum at each of the three time points tested. After 2 h, there was no significant difference between the two groups: 66.7% of the chickens administered the encapsulated phages versus 57.1% of those receiving the nonencapsulated phages. However, after 48 and 72 h the differences were significant: 90.5% versus 38.1% (P < 0.001) and 38.1% versus 9.5% (P < 0.05), respectively.

FIG 4.

FIG 4

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Persistence of nonencapsulated (white bars) and liposome-encapsulated (gray bars) phages in the ceca of broilers. *, P < 0.05; ***, P < 0.001.

Bacteriophage therapy against Salmonella.

The therapeutic efficacies of cocktails of the encapsulated and nonencapsulated forms of the three phages against Salmonella colonization in commercial broilers were tested. As shown in Table 3, cecal Salmonella concentrations were lower in the two treated groups than in the untreated control group, but the two types of cocktails differed in their long-term efficacies. Thus, within the first 6 days postinfection (during which the broilers received their respective cocktails), the decreases in the Salmonella concentration in both treated groups were similar and significant compared to the control. The difference in the decrease in the Salmonella concentration between the nonencapsulated and control groups was significant only on day 8, the first day in which treatment was no longer administered (reduction of 1.5 log10; P < 0.05). In contrast, the differences between the encapsulated and control groups were significant until the end of the experiment (day 15; P < 0.001). In this case, the reductions on days 8, 10, and 15 were 3.8, 3.9, and 1.5 log units, respectively. Moreover, when the efficacies of the two treatments were compared statistically, the reduction in Salmonella achieved using the encapsulated phages was significantly greater from day 8 (P < 0.05) until the end of the experiment (day 15; P < 0.001).

TABLE 3.

Salmonella concentrations in the ceca of broilers treated with encapsulated and nonencapsulated phages

Day postinfection Salmonella concn in cecum (log10 CFU/g)a
Control group Encapsulated group Nonencapsulated group
1 5.8 ± 0.7 3.8 ± 1.2b 2.9 ± 2.3b
3 6.6 ± 0.5 3.3 ± 2.6b 3.3 ± 2.7b
6 6.9 ± 0.8 3.2 ± 2.6b 4.1 ± 2.1b
8 6.7 ± 0.5 2.9 ± 2.8b,c 5.2 ± 2.2b
10 6,4 ± 1.0 2.5 ± 2.8b,c 5.7 ± 1.9
15 5.2 ± 1.3 3.7 ± 1.4b,c 6.3 ± 1.0
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a

Each value is the average from 14 cecum samples ± standard deviation.

b

Statistical significance between the control and each treated group (P < 0.001 at days 1 to 6, P < 0.001 at day 8 for encapsulated group, and P < 0.05 at day 8 for nonencapsulated group).

c

Statistical significance between the two treated groups (P < 0.05 at days 8 and 10 and P < 0.001 at day 15).

In vitro bacteriophage release by cecum content.

The kinetics of the release of liposome-encapsulated phages incubated for up to 60 min with the cecal contents of the chickens were determined. After 30 min, the percentages of released phages were 19.4 ± 2.6, 80 ± 17.2, and 42.7 ± 14.1 for UAB_Phi20, UAB_Phi78, and UAB_Phi87, respectively (Fig. 5). After 60 min, the corresponding percentages were 74.7 ± 5.4, 92.6 ± 12.4, and 56.6 ± 16.7 (Fig. 5).

FIG 5.

FIG 5

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In vitro release of liposome-encapsulated phages UAB_Phi20 (○), UAB_Phi78 (■), and UAB_Phi87 (♢) by the cecal contents of the chickens. Each value is the average from six independent experiments ± standard deviation.

DISCUSSION

Salmonella in poultry production remains a serious problem. The use of phages to fight against these bacteria and its efficient development in the veterinary sector may therefore have a significant economic impact. However, the effectiveness of oral phage applications is compromised by several factors, such as their lack of stability in the acidic environment and their short residence time in the intestinal tracts of animals. With the aim of overcoming the problems inherent in the oral administration of phages to control bacterial pathogens, we tested the efficacy of phages encapsulated in liposomes in broiler chickens. We reasoned that cationic liposomal complexes would promote the release of the encapsulated phages in the target area and therefore their activity against Salmonella.

The three targeted phages (UAB_Phi20, UAB_Phi78, and UAB_Ph87) were encapsulated in cationic lipid envelopes comprising a mixture of lipids with efficiencies of nearly 50%. Our encapsulation procedure does not inactivate the phages. Moreover, it can be used for phages with different morphologies, as phages UAB_Phi20 and UAB_Phi78 have icosahedral heads (60 ± 1.5 nm and 66 ± 1.7 nm, respectively) and noncontractile short tails (13 ± 0.7 nm and 14 ± 0.7 nm, respectively), whereas UAB_Phi87 has an icosahedral head (68 ± 2.7 nm) but a long, contractile tail (114 ± 4.3 nm) (20, 21).

A first obvious advantage of our method is that the liposome-encapsulated phages are stable in aqueous suspensions at 4°C for at least 3 months. In addition, they can be prepared as a dry, stable powder suitable for long-term storage. In previous studies, the lytic activity of lyophilized phages, even those that had been encapsulated, was compromised. This was the case in both alginate-chitosan-encapsulated and poly(lactic-co-glycolic acid)-encapsulated phages (24, 40). However, in a study on the effect of lyophilization of nonencapsulated dysentery phage, in which raw egg yolk was employed as a source of phospholipids, the phage survived lyophilization (35). In our study, the significantly lower susceptibility to the lyophilization process of the encapsulated than the nonencapsulated phages can be attributed to our use of a protective lipid mixture and to the addition of trehalose as a cryoprotectant.

The cationic lipids used in the lipid mixture probably protected the orally administered phages from the acidic environment of the stomach (pH ∼1 to 3) by forming a barrier against protons. In fact, under simulated gastric conditions, the survival of the liposome-encapsulated phages was higher than that of their nonencapsulated counterparts (Fig. 3). In addition, after 48 and 72 h, the intestinal residence time in broilers of an orally administered cocktail of phages was significantly longer for the encapsulated than for the nonencapsulated form (Fig. 4). Consistent with the resistance of the encapsulated phages to gastric pH, at 2 h after their administration, their cecal concentration was significantly higher than that of nonencapsulated phages (data not shown). In addition, in an in vitro experiment (Fig. 5) in which the three phages were tested individually; we were able to show that the cecal contents provoked different release kinetics. Thus, the major part of phage UAB_Phi78 is released during the first 30 min; meanwhile, the release of the other two phages is slower, in particular for phage UAB_Phi87, which has a long and contractile tail. Moreover, different affinities of the viral capsid and tail proteins for the lipids used in the encapsulation can explain these results. Nonetheless, in all three cases the in vivo release of the phages from their lipid capsules was likely due to the activity of bile salts, because their concentrations in the intestinal tracts of broilers are high enough to cause the rupture of the liposomes (38). However, the release kinetics in vivo undoubtedly differ and are more complex than those determined in vitro. Thus, adherence of the liposomes to the intestinal mucosa, the potentially abundant mucus produced by the underlying epithelium, and the concomitant digestion of food could delay the release of phages. All these facts are crucial for the success of the phage therapy because once phages pass through the stomach, they remain retained in the intestinal tract. In this sense, an in vitro study showed that phage can interact with tissue culture cells in the presence of mucus (41). However, it was also described that the concentration of phages in the gut decreased dramatically when the concentration of the bacterial host was reduced (20), such that the phages were retained only briefly by the mucous membrane. We were able to show that one way to address the inherently low retention of phages is to encapsulate them in matrices such as liposomes. Cationic liposomes are positively charged and exhibit mucoadhesive properties, which together result in a much better permeability in the intestinal mucosa and a much longer intestinal retention time than for nonencapsulated formulations (28, 29).

These advantages conferred by the lipid mixture used in this study should translate into an enhanced therapeutic effect of the liposome-encapsulated phages; in fact, their long-term efficacy against Salmonella infection was much higher than that of the nonencapsulated phages (Fig. 4). Specifically, when administered daily for 6 days postinfection, both cocktails similarly protected the chickens against Salmonella colonization. However, once treatment had been stopped, the protective effects of the nonencapsulated phages disappeared after 72 h, whereas those of the encapsulated phages persisted for 1 week, consistent with their higher intestinal residence time and their presumed gradual release from liposomes. However, it must be noted that a total reduction of Salmonella was not reached by this treatment. In this respect, it must be considered that the experimental infection of chickens with Salmonella gives rise to a high initial concentration of bacteria (approximately 6 log10 CFU/g of cecum), and even under these conditions, a total Salmonella reduction was found in the 21% of animals treated with encapsulated phages (data not shown). Therefore, better results would be expected if the initial concentration of Salmonella was lower than that used here, which probably occurs under natural conditions.

It is important to note that our study of the efficacy of phage therapy was done in chickens infected with Salmonella (i.e., the host of the three phages), whereas determination of the in vivo intestinal retention of the phages was performed in noninfected chickens. This difference is crucial in the correct interpretation and comparison of our results, because during the treatment of infected animals with phage, once the phages are released they begin to multiply, thereby generating new progeny (42). This would explain the superior therapeutic effects of the liposome-encapsulated phages.

In summary, we showed that phages with different morphologies can be encapsulated into cationic liposomes and are markedly more stable to acid and to lyophilization in vitro than the corresponding nonencapsulated phages. It is noteworthy that the nanometric size of the liposome-encapsulated phages confers excellent characteristics for their real applications. These include an avoidance of their sedimentation in drinking water and their ready mixture with both the ground feed usually given to younger broilers and the pellets used to feed older chickens, without compromising feed quality. The prolonged intestinal residence time of the encapsulated phages and their efficient release from liposomes in the intestinal tract together resulted in potent and long-lasting therapeutic effects. We are confident that our findings will help researchers to overcome some of the disadvantages of oral phage therapy against other bacterial pathogens.

ACKNOWLEDGMENTS

This work was supported by grants from La Caixa and the Associació Catalana d'Universitats Públiques (2010ACUP00300), AGAUR-ACCIÓ-Generalitat de Catalunya (2010VALOR00114), and AGAUR-Generalitat de Catalunya (2014SGR572). We are grateful to the Servei de Granges i Camps Experimentals and the Servei de Microscòpia of the Universitat Autònoma de Barcelona (UAB) for their support. J.C. and J.O. received predoctoral fellowships from the UAB.

We thank S. Campoy, J. Aranda, and A. Mayola for their help with the animal experiments and S. Escribano for her excellent technical assistance.

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