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Phage preparation FBL1 prevents Bacillus licheniformis biofilm, bacterium responsible for the mortality of the Pacific White Shrimp Litopenaeus vannamei
Aquaculture 484 (2018) 160–167
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Phage preparation FBL1 prevents Bacillus licheniformis biofilm, bacterium responsible for the mortality of the Pacific White Shrimp Litopenaeus vannamei
MARK
Catalina Prada-Peñarandaa, Marcela Salazarb, Linda Güizab, Maria Isabel Pérezc, Chad Leidyc, ⁎ Martha J. Vives-Floreza, a b c
Department of Biological Sciences, Universidad de los Andes, Carrera 1#18A-12 J Building, Bogotá 111711, Colombia Center for Aquaculture Research in Colombia (CENIACUA), Carrera 70F#78A-84, Bogotá 111711, Colombia Biophysics Laboratory, Physics Department, Universidad de los Andes, Cra 1#18A-12, Q Building, Bogotá 111711, Colombia
A R T I C L E I N F O
A B S T R A C T
Keywords: Bacillus licheniformis Biofilm Litopenaeus vannamei Phage therapy Biofilm prevention
Recurrent outbreaks of Bacillus licheniformis strain 52 caused high mortality on juvenile and reproducers of the Pacific White Shrimp Litopenaeus vannamei breeders, in a maturation facility in Colombia. This recurrence led to the suspicion of a permanent contamination source within the tanks. It was hypothesized that biofilms could be that source, but there were no previous reports on the ability of B. licheniformis to form biofilms. Consequently, we tested for the biofilm formation capacity of the bacterium. Using a novel method, a complex biofilm was obtained. Then, phage therapy was assayed as an alternative to control the biofilm. The best outcome was obtained when bacteria and phage preparation FBL1 were inoculated simultaneously, resulting in a 44.77% reduction of the biofilm. Our data indicated that the reduction was probably due to a diminished initial inoculum caused by the phage, instead of the removal of the formed biofilm. This work provides new information that contributes to the understanding of the outbreaks caused by the pathogenic strain 52 in aquaculture systems, and presents promising data supporting the future use of phage therapy.
1. Introduction
industry (Newmark et al., 2009). In the last years, new bacterial pathogens, such as Vibrio parahaemolyticus, responsible for AHPND (Acute Hepatopancreatic Necrosis Disease) have caused billions of dollars in losses. In Colombia, the Colombian Research Center of Aquaculture CENIACUA (Cartagena, Colombia), studied an outbreak of high mortality (up to 70%) of L. vannamei caused by Bacillus licheniformis. Histopathology analysis of moribund shrimps showed a total destruction of the lymphoid organ, and the hepatopancreas in some cases. Microbiological and molecular analyses made from L. vannamei juveniles' haemolymphs identified B. licheniformis as the bacterium responsible for the disease (Gálvez et al., 2016). In Colombia, shrimp farmers use control methods taking advantage of routine management practices, such as frequent water changes to lower the water salinity that could impair bacterial life conditions, decreasing the number of infected shrimps per square meter. Pseudomonads and vibrios are common pathogens in aquaculture productions worldwide but B. licheniformis is not. B. licheniformis is a gram positive, facultative anaerobe bacterium, belonging to the subtilis group, within the Bacillaceae family, closely related to B. subtilis, B.
Shrimp production is an important industry worldwide, valued at around USD 10.6 billion in 2005 (WWF, 1961–2016). According to FAO projections, in 2030 the world shrimp industry could reach 11–18 million tons, doubling its nowadays production (MercoPress, 1997–2016). However, as microbial infections cause mortalities over 90% in shrimp hatcheries (Soto Rodríguez et al., 2006), these represent an important drawback for shrimp farmers. Bacterial infections, the most prevalent during the shrimps' early life stages, are hard to eradicate. Most antibiotics (with the exception of tetracycline and florfenicol) are banned from the cultures with the result that bacteria have become resistant to them (ICA, 1962–2016). Vaccines are not a viable option due to the fact that shrimps do not seem to have a specific immune system; therefore no immunity is generated (Chong and Maha, 2014). Litopenaeus vannamei, the Pacific white shrimp, is the principal cultivated shrimp worldwide and the infectious diseases, caused by viruses and bacteria, are one of the main barriers to the growth of the ⁎
Corresponding author. E-mail addresses: [email protected] (C. Prada-Peñaranda), [email protected] (M. Salazar), [email protected] (L. Güiza), [email protected] (M.I. Pérez), [email protected] (C. Leidy), [email protected] (M.J. Vives-Florez). https://doi.org/10.1016/j.aquaculture.2017.11.007 Received 16 March 2017; Received in revised form 27 October 2017; Accepted 3 November 2017 Available online 07 November 2017 0044-8486/ © 2017 Published by Elsevier B.V.
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standardize a new method for its biofilm quantification.
amyloliquefaciens and B. pumilus. These species produce important enzymes, vitamins and fermented food (Jeyaram et al., 2011; Rey et al., 2004). B. licheniformis produces an assortment of extracellular enzymes, like proteases and amylases, with significant industrial importance (Rey et al., 2004; Sánchez-M and Romero-R, 2010). Several Bacillus species have been reported as capable to decrease pathogen population, being used as probiotics on the shrimp industries (Martínez Cruz et al., 2012); in particular, B. licheniformis is approved and used as probiotic in the aquaculture industries (Martínez Cruz et al., 2012; Sánchez-M and Romero-R, 2010). Probiotic products made from B. licheniformis, that assure inhibition of pathogenic microorganisms, water purification and improvement of health and growth on aquatic animals, poultry, swine, among others, are currently commercialized (Alibaba 1999–2014), as well as mixtures of B. licheniformis and B. subtilis, to control Vibrio alginolyticus (Zhang et al., 2011). Nevertheless, B. licheniformis have been associated to food poisoning and gastroenteritis (Salkinoja-Salonen et al., 1999). Moreover, according to ElSersy and Mohamed (2011) some B. licheniformis strains can produce exotoxins with strong hemolytic activity, causing a reduction in aquaculture products. Although this bacillus was never used as probiotic in the shrimp's tanks, B. licheniformis was found there. The bacterium could be transferred to the tanks by water, aerosols and/or nearby material. The appearance of re-emergent infections caused by this bacterium, led to the suspicion that a permanent contamination source might exist inside the culture tanks. Additionally, no effective method of treatment is known, and the search for alternatives is crucial. Over 99.9% of bacteria form biofilms (Donlan and Costerton, 2002) and it is common to find them in aqueous environments (Proal, 2008). Biofilms are sessile microbial communities attached to a surface, embedded within a self-produced matrix of extracellular polymeric substance (EPS) (Costerton et al., 2005). Bacteria can form these types of structures in any kind of environment they inhabit, being an essential and protective life strategy for them. The slow cell growth (that decreases the taking up of antimicrobials) and the matrix (which acts as a barrier) make them more resistant to antibiotics and chemical agents (Donlan and Costerton, 2002). The time of formation and complexity of the biofilm depend on different factors, like the type of microorganism, surface, light, quorum sensing, among others (Prescott et al., 2005). Biofilm formation involves three general steps: adherence, growth, and detachment. The first one initiates with a reversible adherence, which in time is converted into irreversible adherence once the exopolysacharide is secreted. The second one involves the formation of micro and macro - colonies, hydrated structures, where bacterial cells get embedded within the EPS and initiate its maturation. The last step, where the bacterial cells detached from the biofilm, enables the colonization of new surfaces, where the complete process starts again (Ghannoum and O'Toole, 2004; Sutherland et al., 2004). According to these, if B. licheniformis can form biofilms, the recurrent shrimps infections could be caused by the cell dispersion during the last stage of biofilm maturation. Due to the serious problem that bacteria cause to the shrimp industry, and the lack of an existing, efficient treatment, phage therapy has been proposed as a promising option. Phage therapy is based on the bactericidal effect of bacteriophages, or phages, viruses that only recognize and kill specific bacterial hosts. Phage therapy does not have negative side effects for humans, animals or plants as shown in several reports (Azeredo and Sutherland, 2008; Caplin, 2009; Frampton et al., 2014; Holguín et al., 2015; Martínez Díaz, 2010; Sulakvelidze et al., 2001). The use of phages has been also reported for the control of biofilms formed by pathogenic bacteria like Pseudomonas aeruginosa (Fu et al., 2010), Proteus mirabilis, Escherichia coli (Carson et al., 2010), and Listeria monocytogenes (Montañez Izquierdo et al., 2012), among others. The main goal of this study was to evaluate the biofilm control capabilities of the phage preparation FBL1, a novel native Colombian bacteriophage preparation. To achieve this goal, it was first necessary to evaluate the biofilm formation ability of B. licheniformis and
2. Materials and methods 2.1. Bacterial strains B. licheniformis strain 52 was used in this study for the biofilm assays. This is a virulent strain isolated from L. vannamei haemolymphs (Gálvez et al., 2016). Taxonomic confirmation of the bacterial isolate was done through the amplification and sequencing of the 16S rRNA gene (Vives Florez and Garnica, 2006). Host range test were performed using five additional bacterial strains of B. licheniformis, isolated by CENIACUA from L. vannamei, and nine other strains from the Bacillaceae family donated by A. Yousten (from Virginia Polytechnic Institute and State University) and A. Delecluse (from The Pasteur Institute, France) to J. Dussán, from Universidad de los Andes, and kindly provided by her for this work (supplemental e-component Table 1) (Cavados et al., 2001; HernandezSantana et al., 2016; Lee et al., 2001; Mahler et al., 1986; Stapleton et al., 2004). 2.2. Culture media and conditions Bacteria were grown at 30 °C on nutrient broth (13 g/L) under 250 rpm agitation. Incubation on nutrient agar (13 g/L nutrient broth and 15 g/L bacteriological agar) for 34–36 h allowed colonies observation. Culture media and components were obtained from Oxoid Ltd. 2.3. Bacteriophage isolation and characterization Phages were isolated from fresh water and sediment samples from shrimp ponds. Selective enrichment of the samples was done in 90 mL nutrient broth, where 1% B. licheniformis 52 was previously inoculated. The enriched culture was incubated over night at 30 °C without agitation. After this incubation period, chloroform was added at a proportion of 1% to the cultures followed by another incubation period (2 h, room temperature, 100 rpm). After this second incubation, the enriched culture plus chloroform was centrifuged (8500 rpm for 30 min at 4 °C), the supernatant (containing the phages) was filtered (0.22-μm pore size) and stored at 4 °C. This filtered supernatant was named phage preparation. Sixteen phage preparations were obtained. Due to the fact that these phages were difficult to concentrate and did not show isolated plaques in serial dilutions plating using the double layer agar method, they were plated by the spot-test method in double agar overlay plates to characterize them by host range test, against the fourteen strains of the Bacillaceae family (supplemental e-component Table 1). Briefly, 3–5 colonies of B. licheniformis 52 were inoculated on 3 mL nutrient broth and grown for 6 h at 30 °C and 250 rpm. 100 μL of each bacterial overnight culture were mixed with 3 mL of soft agar (TAE 1X, 13 g/L nutrient broth and 0.25% w/v agarose) and poured on top of nutrient agar plates. When the soft agar solidified, 10 μL of each phage preparation were spotted on the surface and let to absorb. The plates were incubated at 30 °C for 14 h. Spots that presented a plaque were categorized as a phage preparation with effect on the corresponding bacterial strain (positive, +), while those that resulted in no visible plaque were categorized as ineffective (negative, −). The structure of the viral particles from phage preparation FBL1 was characterized by transmission electron microscopy (TEM) according to the following protocol: drops of FBL1 were deposited on Parafilm®. Carbon-coated Formvar film (grid) were placed over the FBL1 drops for 5 min; the grid was air-dried for 3 min and then it was placed over the phosphotungstic acid solution for 2 min, in order to stain negatively the phage sample. When dried, FBL1 was observed using TEM in a microscope model JEOL 1400 plus. 161
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92 h and 116 h. The weight of the recovered biomasses was compared to controls without FBL1. All the experiments were conducted in triplicate.
2.4. B. licheniformis biofilm formation assay Biofilm formation was initially analyzed by the microtiter plate assay, the most commonly used method to evaluate biofilms. However, the biofilm obtained was thin and lacked adherence to the well walls, causing its loss during the procedure (data not shown). The same negative result was obtained in test tubes and flasks. Consequently, a novel method to evaluate the biofilm formation capacity in this Bacillus species was developed. B. licheniformis 52 biofilm formation was standardized using a microdrip solution administration set (BAXTER 60 drops/mL) (Chávez Escobar and Leidy, 2010) that simulated an aquatic flowing system. It consisted of two erlenmeyers, one of them located at a distance of 1.2 m below the other one, and connected by a 3 m microdrip hose. 5 mL of a 108 CFU/mL bacterial suspension were inoculated directly through the microdrip injection site; the culture was allowed to go down the hose for 10 s and then the system was kept closed for 3 h into the incubator at 30 °C. Afterwards, the flux was adjusted to 0.4–0.7 mL/min. Three different times for biofilm recovery after bacterial inoculation were tested: 24 h, 48 h, and 68 h. The formed biofilm was expelled from the plastic hose, centrifuged (8500 rpm for 30 min at 4 °C), and the pelleted biomass was lyophilized (Lyophilizer LABCONCO model freezone) for 24 h, and weighted. Experiments were conducted in triplicate. Biofilm surface structure was observed by scanning electron microscopy (SEM) (model JSM 6490-LV), using a sample of the recovered biofilm prior lyophilization and without sample preparation.
3. Results 3.1. Bacterial strain The sequence for the rRNA 16S gene obtained was compared with the NCBI gene bank database using BLAST search program (Altschul et al., 1990). The bacterium B. licheniformis 52 was identified as B. licheniformis with an E-value equal to 0.0, 100% query coverage and 99–100% maximum identity. 3.2. Bacteriophage isolation and characterization Sixteen phage preparations were obtained. We named them phage preparations instead of phages because it was not possible to guarantee that they contained only one type of phage. The phages were observed by plating using the spot method, and this method does not allow the definition of phage purity. Presence/absence of plaques in the host range test showed that the phage preparations were different (Table 1). Phage preparation FBL1 was chosen for the subsequent experiments due to its specificity, infecting, apart from the B. licheniformis, only one out of the other nine Bacillaceae strains. It was also effective against the most virulent strains of B. licheniformis, according to the information provided by CENIACUA, including the virulent B. licheniformis 52. TEM images were extremely difficult to obtain because it was not possible to concentrate the phages. Although the poor quality of the photographs, they show that phage preparation FBL1 contains tailed phages with heads of 45.03 nm to 68.41 nm. Due to the presence of long non-contractile tails they probably belong to the Siphoviridae family (Fig. 1).
2.5. Control of B. licheniformis biofilm by the phage preparation FBL1 Once the biofilm formation assay was standardized, experiments were set up to assess the effect of phages on the biofilm formed. Biofilms were treated with 1 mL of the phage suspension FBL1 at three different times: 0 h (simultaneous inoculation of bacteria and phage suspension), 24 h, and 48 h after bacterial inoculation. In all cases, once the phage suspension was added, the flux was suspended for 24 h and then resumed. In all treatments, biofilms were recovered at 68 h after bacterial inoculation. Dry weight of recovered biofilm was compared between treatments and controls (i.e. biofilms without FBL1 inoculation). Additionally, the surface of the 68 h-biofilms treated with FBL1 at 0 h was observed under SEM and compared to the appearance of the 24 and 68 h control biofilms. Each experimental assay was done in triplicate. In order to test for significance, the non-parametric data were analyzed by the Kruskal-Wallis test and the parametric data by the Tukey test using the R platform version 3.0.2, with a confidence level of p < 0.05. To elucidate the nature of FBL1 effect on the biofilm, it was predicted that either it would remove the formed biofilm, or prevent its formation by killing the planktonic cells. If the phages were removing the biofilm, FBL1 would induce the dispersion of planktonic cells and the material around them, which would cause a higher weight of the biomass in the disposal (where the flux from the lower Erlenmeyer was collected) compared to controls without phage treatment. On the other hand, if FBL1 were killing the planktonic cells, the biofilm formation cycle would take longer because of the reduced inoculum, with more time needed to form a mature biofilm. Therefore, the weight of the biomass in the disposal would be lower compared to controls, because the biofilm would not be in its mature stage, and the dispersion final step would not be reached at the same time. The experiment was conducted with and without phage treatment measuring the biomasses differences in the disposal, in order to evaluate which of the predictions was correct. FBL1 was inoculated at 0 h, 24 h, and 48 h after B. licheniformis strain 52.68 h after the bacterial inoculation in every assay, the accumulated disposal biomasses were recovered, freeze-dried and weighted. To assess the extended life cycle of the biofilm, the experiment was conducted as described previously, inoculating FBL1 simultaneously with the bacterium and recovering the biofilm at 68 h,
3.3. B. licheniformis biofilm formation The microdrip solution administration set allowed the formation, recovery (e-component Movie 1) and quantification of B. licheniformis 52 biofilm. To the best of our knowledge, this is the first report of B. licheniformis biofilm observation and quantification. The SEM images show the structure of the biofilms recovered at different times after bacterial inoculation, without FBL1 treatment (Fig. 2A–C). With longer times of incubation, the recovered biofilm increased its structural complexity, as well as its biomass (Fig. 3A). These results showed reproducible data, with low variability between replicates. At 68 h of incubation, a complex, continuous biofilm was observed in the SEM images (Fig. 2C). Thus, 68 h was chosen as the time for biofilm incubation in the following assays. 3.4. Control of B. licheniformis biofilm by the phage preparation FBL1 In order to evaluate the effect of the phage preparation FBL1 on the biofilm formed by B. licheniformis 52, the amount of formed biofilm was measured after it was treated with the phage preparation. The data showed on Fig. 3B evidenced that FBL1 caused a reduction on the dry weight of the biofilm, and the magnitude of these reductions varied according to the time of inoculation of FBL1: at 0 h, 24 h and 48 h, the reduction in the biofilm recovered was 53.33%, 30.16% and 0.79%, respectively. SEM images of the structure of biofilm treated with FBL1 at time 0 h (Fig. 2D), were almost identical to the structure of the non treated (control) biofilm at 24–48 h (Fig. 2A and B). Control biofilm at 68 h after bacterial inoculation (Fig. 2C), showed a hard and robust structure without holes. On the other hand, the phage-treated biofilm depicted a less complex structure, characteristic of earlier stages of biofilm 162
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formation (Fig. 2D). The experiments conducted to elucidate the mode of action of FBL1 on the biofilm identified interesting aspects. Fig. 4 shows that the biomass in the disposal was smaller in all experiments with FBL1, compared to controls. Significant difference (p < 0.05) were only found in the treatment where FBL1 was added at 48 h. This result would support the hypothesis that FBL1 killed the planktonic cells. It might also explain the lowest amount of biomass found in the disposal when FBL1 was inoculated at 48 h, since the phages could kill the dispersed cells from the formed, 48 h – old biofilm. On the other hand, in the control without FBL1, the biofilm continued its normal life cycle and the dispersed cells went to the disposal. Additionally, if the phage suspension killed the planktonic cells, the life cycle of the biofilm should also require a longer time. The results showed that in the normal life cycle of B. licheniformis biofilm (Fig. 3A), the dry weight increased regularly in ca. 5.5 mg every 24 h (Table 2). When the biofilm was treated with FBL1, injecting it simultaneously with the bacterium, the biofilm dry weight increased along time but at a significantly lower rate: from time 68 h to 92 h and 92 h to 116 h augmented in 0.43 mg and 1.94 mg, respectively (Table 2), showing a much lower growth rate compared to the control biofilm. The lower biofilm growth rate had the consequence of an extended biofilm life cycle, providing additional support to the hypothesis of FBL1 action at the planktonic cells stage.
+ − − + + + − −
+ − − − − + − + − − − − + − − − − − − − − + − − − − +
− + − + − + + + − + − − + − − − − − − − − − − − − − − − + − − +
− + − − − + − − − − +
Bacillus licheniformis 54(1)
The present work is the first study showing the standardization and quantification of the biofilm formed by a strain of B. licheniformis. Although there are no reports of B. licheniformis biofilms and it has been published that this bacterium was not known to form biofilms (Dhakal et al., 2013), this study demonstrated that B. licheniformis is capable of forming a robust, complex one, with abundant extracellular matrix produced within 24 h. This finding gives a plausible explanation for the recurrent outbreaks in L. vannamei tanks, since the biofilms can act as a source of new bacterial inocula. We used a new methodology, since the standard microtiter plate method was not useful for this bacterial strain. The new method utilizes a microdrip set, allowing the biofilm to form in a closed and controlled space, delivering reproducible data of biofilm dry weight, and also permitting the observation of its structural surface through SEM. Besides, the microdrip system resembles to the water pipelines used in aquaculture. Once the biofilm formation was standardized, it was possible to analyze the capacity of the phage preparation FBL1 to control it. The present study found that, depending on the time of the FLB1 treatment, the phages produced different outcomes in the amount of recovered biofilm (Fig. 3B). The best result (53.3% reduction) was obtained when the treatment was applied at the same time as the bacterial inoculation, and the worst (0.79%) with the more delayed treatment of 48 h postinoculation. These findings are likely due to the profuse extracellular matrix produced, which is known to provide impermeability and protection (Harper et al., 2014). Despite the resistant characteristics conferred, bacteriophages are excellent antibacterial agents that can overcome the biofilms barriers. However, it depends on the particular phage and biofilm and, in this case, FLB1 was not able to remove mature biofilms of B. licheniformis 52. Regarding the FBL1 mode of action, it was found that FBL1 caused a negative effect on the biofilm formation. The SEM image of the biofilm treated with FBL1 at time 0 h and recovered 68 h later (Fig. 2D), revealed a biofilm structure similar to the first stages of its formation without phage treatment (Fig. 2A and B). Bacteriophages are known to damage biofilms integrity, either killing EPS producing cells or producing enzymes that degrade the extracellular matrix (Harper et al., 2014). In this case, killing the planktonic cells probably caused the reduction of the biofilm.
+: phage preparation with effect on the corresponding bacterial strain; −: phage preparation without visible effect on the strain.
+ + + + + + − + + − + − − +
+
+ + + + + + − + − + + − + + − −
+ + + + + + − + − − − − + − − −
+ + + + + + + + − + + + + + + −
+ + + + + + + + − + + + + + − −
+ + + + + + + + − + + + + + − −
+ − + − + + − + − − − − + − − −
+ + + + + + − + − − − − + − − −
− − − − − − − − − − − − + − − −
4. Discussion
FBL16 FBL15 FBL14 FBL13 FBL12 FBL11 FBL10 FBL9 FBL8 FBL7 FBL6 FBL5 FBL4 FBL3 FBL2 FBL1
Lysinibacillus fusiformis Lysinibacillus sphaericus NRS1198 Lysinibacillus sphaericus LS 2362 Lysinibacillus sphaericus NRS400 Bacillus polymyxa
Bacteria strain Phage preparation
Table 1 Host range pattern of the phage preparations against representative strains of the Bacillaceae family.
Bacillus cereus RC607
Bacillus cereus 3501
Bacillus subtilis BGL
Bacillus thuringiensis
Bacillus licheniformis 52
Bacillus licheniformis 54(2)
Bacillus licheniformis 76
Bacillus licheniformis lume
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Fig. 1. TEM morphology of phage preparation FBL1. TEM morphology of the negatively stained phage preparation FBL1 showing long, non-contractile tails with heads of 45.03 nm to 68.41 nm. The morphology is compatible with Siphoviridaelike phages. (A) Original image. (B) Adjusted image: cutting the borders and getting a better contrast and brightness through the Fiji platform.
48 h. It is worth to mention that the amount of biofilm formed at 116 h was significantly lower (34.28%), compared to the controls, demonstrating the effect of FBL1. Additionally, the FBL1 treated biofilm grew at a lower rate (Table 2). These data also showed a gradual increase in the growth rate (0.017 mg/h to 0.080 mg/h); we expected that the biofilm would recover its normal growth rate (ca. 0.2 mg/h), as the effect of the phage suspension ends; however, we followed this experiment only up to the 116 h. These observations strongly suggest that FBL1 prevented the biofilm formation of B. licheniformis by killing
Analyzing the mode of action of FBL1, it was found that only its early addition was associated with a significant decrease in the dry weight of the collected biofilm, indicating that FBL1 killed the planktonic cells instead of removing the already formed biofilm. Thus, the B. licheniformis treated biofilm life cycle was longer than the untreated control. When the biofilm was treated with FBL1 and recovered at 92 h and 116 h, it did not even reach the amount of biofilm recovered at 68 h in the control assays (without FBL1). Moreover, the dry weigh of the biomass was similar to the biofilm biomasses of the control assays at
Fig. 2. SEM images of Bacillus licheniformis biofilm with and without phage preparation FBL1. B. licheniformis strain 52 biofilm recovered at (A) 24 h (B) 48 h and (C) 68 h. Fig. 2 (D) presents the B. licheniformis biofilm obtained after treatment with phage preparation FBL1; FBL1 was inoculated at time 0, and the biofilm formed was recovered 68 h later. Note the difference in complexity between the control and treated biofilms at 68 h.
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planktonic cells and delaying its development. The use of bacteriophages to control bacterial biofilms has been reported elsewhere, using the microtiter plate method (Carson et al., 2010; Holguín et al., 2015; Montañez Izquierdo et al., 2012). As far as we know, there are no reports on the use of phages to treat B. licheniformis planktonic cells or biofilms. In fact, not even the biofilm formed by Bacillus subtilis, the Bacillus species most studied, has been quantitatively assessed. Usually, biofilms of B. subtilis have been observed at the liquid-air interface of broth cultures, or by observing the colonies formed on the surface of agar plates, but the resulting data cannot be systematically quantified (Lemon et al., 2008; Vlamakis et al., 2013). Thus, this is the first time that the observation, standardization and quantification of B. licheniformis biofilm is reported. We were able to obtain the novel phage preparation FBL1 although the phages isolated in this work were difficult to titer and concentrate; we assayed a wide variety of methodologies (lysates, ultracentrifugation at 40,000 g, enrichment from isolated plaques, and lysates using agar) and did not obtain a concentrated suspension. This limitation prevented further characterization, like genome size estimation, sequencing and annotation. Additional work on these areas is needed to rule out the presence of integration sequences and other undesirable genes. A preliminary characterization based on phage structure allowed the putative classification of FBL1 in the Siphoviridae family, because of the binary morphology and the long non-contractile tail. According to the classification of tailed bacteriophages, phages from Bacillus species belong to the Myoviridae, Podoviridae (Kutter and Sulakvelidze, 2005) and Siphoviridae families (Yuan et al., 2012). Some reports have described B. licheniformis phages from Siphoviridae with a head diameter ranging from 41 nm to 66 nm (Hertel et al., 2015; Huang and Marmur, 1970); FBL1 phages showed a similar head size. In summary, the results obtained in this work give new information about the B. licheniformis ability to form biofilms, and offer a new method to study them. Also, phage therapy may represent a future alternative to control bacterial pathogens for the shrimp industry, provided that better characterized phages were available. Supplementary data to this article can be found online at https:// doi.org/10.1016/j.aquaculture.2017.11.007. Video showing the recovery of B. licheniformis biofilm formed inside the microdrip solution administration set.
Fig. 3. Bacillus licheniformis biofilm formation and effect of phage preparation FBL1. (A) Standardization of B. licheniformis strain 52 biofilm. The bars represent the amount of biofilm biomass recovered at five different times (24, 48, 68, 92 and 116 h) after the bacterial inoculation. All experiments were significantly different between each other (P < 0.05). (B) B. licheniformis strain 52 biofilm formed with and without FBL1 treatment, showing the effect of time addition of the phage suspension on the biofilm biomass recovered. FBL1 was inoculated at three different times after bacterial inoculation: 0 h, 24 h and 48 h. For all treatments, biofilm was recovered and the biomass was measured as dry weight 68 h after the bacterial inoculation. The asterisk (*) indicates that the difference was significant (P < 0.05). Controls (black bar) were measured independently for each treatment; the black bar represents the average of nine replicates (3 for each experiment). All experiments were conducted in triplicate.
Declaration of interest Catalina Prada-Peñaranda and Martha J. Vives-Florez are members of the spin off SciPhage S.A.S., which works for the development of phage therapy in Colombia.
Contributions Catalina Prada-Peñaranda: Conception and design, acquisition of data, biofilm formation assay, analysis and interpretation of data, drafting the article, revising it critically for important intellectual content, final approval of the version to be published critically for important intellectual content. Martha J. Vives-Florez: Conception and design, analysis and interpretation of data, drafting the article, revising it critically for important intellectual content, final approval of the version to be published critically for important intellectual content. Marcela Salazar, Linda Güiza: strains isolation and preliminary phage tests, revising the article, final approval of the version to be published critically for important intellectual content. Maria Isabel Pérez, Chad Leidy: biofilm formation assay, revising the article, final approval of the version to be published critically for important intellectual content.
Fig. 4. FBL1 prevents Bacillus licheniformis biofilm by killing the planktonic cells and delaying its formation. B. licheniformis strain 52 detached biomass with and without FBL1 treatment, when phage preparation was inoculated at 0 h, 24 h and 48 h after B. licheniformis strain 52. For all treatments, biofilm was recovered and the accumulated biomass was measured as dry weight 68 h after the bacterial inoculation. These results indicate that FBL1 kills the planktonic cells and lags the biofilm growth (see text). Controls (black bar) were measured independently for each treatment; the black bar represents the average of nine replicates (3 for each experiment). All experiments were conducted in triplicate. The asterisk (*) indicates that the difference was significant (P < 0.05).
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Table 2 Bacillus licheniformis strain 52 biofilm dry weight and growth rate, with and without phage suspension FBL1. B. licheniformis 52 (control) Time (h) 68 92 116
Dry weight (mg) 12.17 17.57 23.24
Difference – 5.40 5.67
B. licheniformis 52 + FBL1 (simultaneous inoculation) Biofilm growth rate (mg/h) – 0.225 0.236
Dry weight (mg) 5.60 6.03 7.97
Funding
Difference – 0.43 1.94
Biofilm growth rate (mg/h) – 0.017 0.080
% of biofilm formed compared to control 46.03 34.35 34.28
aeruginosa on catheters in an in vitro model system. Antimicrob. Agents Chemother. 54, 397–404. Gálvez, E.J.C., Carrillo-Castro, K., Zárate, L., Güiza, L., Pieper, D.H., García-Bonilla, E., Salazar, M., Junca, H., 2016. Draft genome sequence of Bacillus licheniformis CG-B 52, a highly virulent bacterium of Pacific white shrimp (Litopenaeus vannamei), isolated from a Colombian Caribbean aquaculture outbreak. Genome Announc. 4 (3), e00321–16. http://dx.doi.org/10.1128/genomeA.00321-1. Ghannoum, M., O'Toole, G.A. (Eds.), 2004. Microbial Biofilms. ASM Press, Washington. Harper, D.R., Parracho, H.M.R.T., Walker, J., Sharp, R., Hughes, G., Werthén, M., Lehman, S., Morales, S., 2014. Bacteriophages and biofilms. Antibiotics. 3, 270–284. Hernandez-Santana, A., Gómez-Garzón, C., Dussán, J., 2016. Complete genome sequence of Lysinibacillus sphaericus WHO reference strain 2362. Genome Announc. 4, e0054516. http://dx.doi.org/10.1128/genomeA.00545-16. Hertel, R., Rodríguez, D.P., Hollensteiner, J., Dietrich, S., Leimbach, A., Hoppert, M., Liesegang, H., Volland, S., 2015. Genome-based identification of active prophage regions by next generation sequencing in Bacillus licheniformis DSM13. PLoS One 10, e0120759. http://dx.doi.org/10.1371/journal.pone.0120759. Holguín, A.V., Rangel, G., Clavijo, V., Prada, C., Mantilla, M., Gomez, M.C., Kutter, E., Taylor, C., Fineran, P.C., Gonzalez Barrios, A.F., Vives, M.J., 2015. Phage _Pan70, a putative temperate phage, controls Pseudomonas aeruginosa in planktonic, biofilm and burn mouse model assays. Viruses. 7, 4602–4623. Huang, W.M., Marmur, J., 1970. Characterization of inducible bacteriophages in Bacillus licheniformis. J. Virol. 5, 237–246. ICA, 1962–2016. Productos medicamentos veterinarios con registro ICA vigente a ***20 de octubre 2015***. In: Colombia: Instituto Colombiano Agropecuario, . http:// www.ica.gov.co/Areas/Pecuaria/Servicios/Regulacion-y-Control-de-MedicamentosVeterinarios/Medicamentos/Listados/2015/03082015/Productos-MedicamentosVeterinarios-1.aspx (accessed 31.04.16). Jeyaram, K., Romi, W., Singh, T.A., Adewumi, G.A., Basanti, K., Oguntoyinbo, F.A., 2011. Distinct differentiation of closely related species of Bacillus subtilis group with industrial importance. J. Microbiol. Methods 87, 161–164. Kutter, E., Sulakvelidze, A. (Eds.), 2005. Bacteriophages: Biology and Applications. CRC Press, United States of America. Lee, I.H., Je, Y.H., Chang, J.H., Roh, J.Y., Oh, H.W., Lee, S.G., Shin, S.C., Boo, K.S., 2001. Isolation and characterization of a Bacillus thuringiensis ssp. kurstaki strain toxic to Spodoptera exigua and Culex pipiens. Curr. Microbiol. 43, 284–287. Lemon, K.P., Earl, A.M., Vlamakis, H.C., Aguilar, C., Kolter, R., 2008. Biofilm development with an emphasis on Bacillus subtilis. Curr. Top. Microbiol. Immunol. 322, 1–16. Mahler, I., Levinson, H.S., Wang, Y., Halvorson, H.O., 1986. Cadmium- and Mercuryresistant Bacillus strains from a salt marsh and from Boston Harbor. J Appl Environ Microbiol. 52, 1293–1298. Martínez Cruz, P., Ibañez, A.L., Monroy Hermosillo, O.A., Ramirez Saad, H.C., 2012. Use of probiotics in aquaculture. ISRN Microbiol. 2012, 13. Martínez Díaz, S.F., 2010. Phage therapy provides targeted bacteria treatment. In: Innovation, . http://pdf.gaalliance.org/pdf/GAA-Diaz-Jan10.pdf (accessed 20.05.15). MercoPress, 1997–2016. FAO Forecasts Shrimp World Production Could Double by 2030 [Internet]. South Atlantic News Agency, Montevideo. http://en.mercopress.com/ 2014/10/08/fao-forecasts-shrimp-world-production-could-double-by-2030 (accessed 20.01.16). Montañez Izquierdo, V.Y., Salas-Velazquez, D.I., Rodriguez-Jerez, J.J., 2012. Use of epifluorescence microscopy to assess the effectiveness of phage P100 in controlling Listeria monocytogenes biofilms on stainless steel surfaces. Food Control 23, 470–477. Newmark, U.F., Valverde, B.C., Humberto-Díaz, L.J.M., Parra, A.G., Bonilla, S.P., Salazar, V.M., 2009. Agenda prospectiva de investigación y desarrollo tecnológico para la cadena productiva de camarón de cultivo en Colombia. In: Bogotá (Colombia): Corporación Centro de Investigación de la Acuicultura de Colombia (CENIACUA). Universidad Nacional de, Colombia. Prescott, M.L., Harley, J.P., Klein, D.A., 2005. Microorganism interactions and microbial ecology microbiology. In: Prescott, M.L., Harley, J.P., Klein, D.A. (Eds.), Microbiology. McGraw-Hill, New York, pp. 577–613. Proal, A., 2008. Understanding Biofilms. [Internet]. Bacteriality Understanding Chronic Disease. http://bacteriality.com/2008/05/biofilm/ (accessed 22.03.16). Rey, M.W., Ramaiya, P., Nelson, B.A., Brody-Karpin, S.D., Zaretsky, E.J., Tang, M., Lopezde-Leon, A., Xiang, H., Gusti, V., Clausen, I.G., Olsen, P.B., Rasmussen, M.D., Andersen, J.T., Jorgensen, P.L., Larsen, T.S., Sorokin, A., Bolotin, A., Lapidus, A., Galleron, N., Ehlich, S.D., Berka, R.M., 2004. Complete genome sequence of the industrial bacterium Bacillus licheniformis and comparisons with closely related Bacillus species. Genome Biol. 5, 12. Salkinoja-Salonen, M.S., Vuorio, R., Andersson, M.A., Kämpfer, P., Andersson, M.C., Honkanen-Buzalski, T., Scoging, A.C., 1999. Toxigenic strains of Bacillus licheniformis related to food poisoning. J Appl Environ Microbiol. 65, 4637–4645. Sánchez-M, M., Romero-R, F., 2010. Producción de proteasas con Bacillus licheniformis ATCC 1259 y su acción hidrolítica sobre proteínas de arroz (Oryza sativa L.) y maíz
This work was supported by: Banco Santander, grant Premio Santander Emprendimiento Ciencia e Innovación 2010 awarded to MJV-F; the scholarship granted to CP-P by the Department of Biological Sciences, Universidad de los Andes, for her Master studies in Biological Sciences, area Microbiology; the Faculty of Sciences, Universidad de los Andes, project code OI160422001-BIO15; and the Office of the Vice president for Research, Universidad de los Andes, project code P16853122.003/09-01. Acknowledgments The authors thank Paula Esquinas from Fundación Santa Fé de Bogotá, Colombia, for the help with the TEM images; Dr. Jenny Dussán for lending the strains for the host range assays. We also thank Banco Santander for the financial support. Many thanks to the Faculty of Sciences Research Fund and the Department of Biological Sciences, Universidad de los Andes, that provided partial funding for the research and the CP-P Masters student scholarship, respectively. We acknowledge the Office of the Vicepresident for Research, Universidad de los Andes for the finantial support to CP-P. We thank Humberto Ibarra for the invaluable help with the TEM images. References Alibaba, 1999–2016. Leading Platform for Global Wholesale Trade. https://www.alibaba. com/trade/search?fsb=y&IndexArea=product_en&CatId=&SearchText= Aquaculture+Probiotics+Bacillus+Licheniformis (accessed 22.02.16). Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403–410. Azeredo, J., Sutherland, I.W., 2008. The use of phages for the removal of infectious biofilms. Curr. Pharm. Biotechnol. 9, 261–266. Caplin, J., 2009. Bacteriophage Therapy — Old Treatment, new Focus? http://eprints. brighton.ac.uk/6092/1/SfAM_article_BACTERIOPHAGE_THERAPY_-_OLD_ TREATMENT_NEW_FOCUS_LH.pdf (accessed 15.02.14). Carson, L., Gorman, S.P., Gilmore, B.F., 2010. The use of lytic bacteriophages in the preventionand eradication of biofilms of Proteus mirabilis and Escherichia coli. FEMS Immunol. Med. Microbiol. 59, 447–455. Cavados, C.F.G., Fonseca, R.N., Chaves, J.Q., Rabinovitch, L., Araújo-Coutinho, C.J.P.C., 2001. Identification of entomopathogenic Bacillus isolated from Simulium (Diptera, Simuliidae) larvae and adults. Mem. Inst. Oswaldo Cruz 96, 1017–1021. Chávez Escobar, J.P., Leidy, C., 2010. Regulación del nivel de fluidez membranal de bacterias patogénicas: formación de biopelículas y adaptación a la termorregulación del hospedero. dissertation. Universidad de los Andes, Colombia. Chong, C.M., Maha, A., 2014. Fish and Crustaceans: Immune System, Vaccine Development and Implications. In: Nurul Amin, S.M., Kamarudin, M.S., Arshad, A., Romano, N. (Eds.), Perspective of Fisheries and Aquaculture in Malaysia. Universiti Putra Malaysia Press, Selangor, pp. 188–207. Costerton, J.W., Montanaro, L., Arciola, C.R., 2005. Biofilm in implant infections: its production and regulation. Int J Artif Organs. 28, 1062–1068. Dhakal, R., Chauhan, K., Brent Seale, R., Deeth, H.C., Pillidge, C.J., Powell, I.B., Craven, H., Turner, M.S., 2013. Genotyping of dairy Bacillus licheniformis isolates by high resolution melt analysis of multiple variable number tandem repeat loci. Food Microbiol. 34, 344–351. Donlan, R.M., Costerton, J.W., 2002. Biofilms: survival mechanisms of clinically relevant microorganism. Clin. Microbiol. Rev. 15, 167–193. ElSersy, N.A., Mohamed, E.A.H., 2011. Biochemical and molecular characterization of hemolytic Bacillus licheniformis strains isolated from shrimp and clam aquacultures. Afr. J. Microbiol. Res. 5, 1830–1836. Frampton, R.A., Taylor, C., Holguín Moreno, A.V., Visnovsky, S.B., Petty, N.K., Pitman, A.R., Fineran, P.C., 2014. Identification of bacteriophages for biocontrol of the kiwifruit canker phytopathogen Pseudomonas syringae pv. actinidiae. J Appl Environ Microbiol. 80, 2216–2228. Fu, W., Forster, T., Mayer, O., Curtin, J.J., Lehman, S.M., Donlan, R.M., 2010. Bacteriophage cocktail for the prevention of biofilm formation by Pseudomonas
166
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C. Prada-Peñaranda et al.
environmental Pseudomonas aeruginosa isolates. Int. Microbiol. 9, 247–252. Vlamakis, H., Chai, Y., Beauregard, P., Losick, R., Kolter, R., 2013. Sticking together: building a biofilm the Bacillus subtilis way. Nat. Rev. Microbiol. 11, 157–168. WWF, 1961–2016. farmed shrimp. In: Washington: World Wildlife Fund, . http://www. worldwildlife.org/industries/farmed-shrimp (accessed 28.05.16). Yuan, Y., Gao, M., Wu, D., Liu, P., Wu, Y., 2012. Genome characteristics of a novel phage from Bacillus thuringiensis showing high similarity with phage from Bacillus cereus. PLoS One 7, e37557. http://dx.doi.org/10.1371/journal.pone.0037557. Zhang, Q., Tan, B., Mai, K., Zhang, W., Hongming, M., Ai, Q., Wang, X., Liufu, Z., 2011. Dietary administration of Bacillus (B. licheniformis and B. subtilis) and isomaltooligosaccharide influences the intestinal microflora, immunological parameters and resistance against Vibrio alginolyticus in shrimp, Penaeus japonicus (Decapoda: Penaeidae). Aquac. Res. 42, 943–952.
(Zea mays L.). dissertation. Universidad Nacional experimental Simón Rodríguez, Republica Bolivariana de Venezuela. Soto Rodríguez, S.A., Simoes, N., Roque, A., Gómez, G.B., 2006. Pathogenicity and colonization of Litopenaeus vannamei larvae by luminescent vibrios. Aquaculture 258, 109–115. Stapleton, P., Pike, R., Mullany, P., Lucas, V., Roberts, G., Rowbury, R., Wilson, M., Richards, H., 2004. Mercuric resistance genes in gram-positive oral bacteria. FEMS Microbiol. Lett. 236, 213–220. Sulakvelidze, A., Alavidze, Z., Morris, G., 2001. Bacteriophage therapy. Antimicrob. Agents Chemother. 45 (3), 649–659. Sutherland, I.W., Hughes, K.A., Skillman, L.C., Tait, K., 2004. The interaction of phage and biofilms. FEMS Microbiol. Lett. 232, 1–6. Vives Florez, M., Garnica, D., 2006. Comparison of virulence between clinical and
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Phage preparation FBL1 prevents Bacillus licheniformis biofilm, bacterium responsible for the mortality of the Pacific White Shrimp Litopenaeus vannamei
MARK
Catalina Prada-Peñarandaa, Marcela Salazarb, Linda Güizab, Maria Isabel Pérezc, Chad Leidyc, ⁎ Martha J. Vives-Floreza, a b c
Department of Biological Sciences, Universidad de los Andes, Carrera 1#18A-12 J Building, Bogotá 111711, Colombia Center for Aquaculture Research in Colombia (CENIACUA), Carrera 70F#78A-84, Bogotá 111711, Colombia Biophysics Laboratory, Physics Department, Universidad de los Andes, Cra 1#18A-12, Q Building, Bogotá 111711, Colombia
A R T I C L E I N F O
A B S T R A C T
Keywords: Bacillus licheniformis Biofilm Litopenaeus vannamei Phage therapy Biofilm prevention
Recurrent outbreaks of Bacillus licheniformis strain 52 caused high mortality on juvenile and reproducers of the Pacific White Shrimp Litopenaeus vannamei breeders, in a maturation facility in Colombia. This recurrence led to the suspicion of a permanent contamination source within the tanks. It was hypothesized that biofilms could be that source, but there were no previous reports on the ability of B. licheniformis to form biofilms. Consequently, we tested for the biofilm formation capacity of the bacterium. Using a novel method, a complex biofilm was obtained. Then, phage therapy was assayed as an alternative to control the biofilm. The best outcome was obtained when bacteria and phage preparation FBL1 were inoculated simultaneously, resulting in a 44.77% reduction of the biofilm. Our data indicated that the reduction was probably due to a diminished initial inoculum caused by the phage, instead of the removal of the formed biofilm. This work provides new information that contributes to the understanding of the outbreaks caused by the pathogenic strain 52 in aquaculture systems, and presents promising data supporting the future use of phage therapy.
1. Introduction
industry (Newmark et al., 2009). In the last years, new bacterial pathogens, such as Vibrio parahaemolyticus, responsible for AHPND (Acute Hepatopancreatic Necrosis Disease) have caused billions of dollars in losses. In Colombia, the Colombian Research Center of Aquaculture CENIACUA (Cartagena, Colombia), studied an outbreak of high mortality (up to 70%) of L. vannamei caused by Bacillus licheniformis. Histopathology analysis of moribund shrimps showed a total destruction of the lymphoid organ, and the hepatopancreas in some cases. Microbiological and molecular analyses made from L. vannamei juveniles' haemolymphs identified B. licheniformis as the bacterium responsible for the disease (Gálvez et al., 2016). In Colombia, shrimp farmers use control methods taking advantage of routine management practices, such as frequent water changes to lower the water salinity that could impair bacterial life conditions, decreasing the number of infected shrimps per square meter. Pseudomonads and vibrios are common pathogens in aquaculture productions worldwide but B. licheniformis is not. B. licheniformis is a gram positive, facultative anaerobe bacterium, belonging to the subtilis group, within the Bacillaceae family, closely related to B. subtilis, B.
Shrimp production is an important industry worldwide, valued at around USD 10.6 billion in 2005 (WWF, 1961–2016). According to FAO projections, in 2030 the world shrimp industry could reach 11–18 million tons, doubling its nowadays production (MercoPress, 1997–2016). However, as microbial infections cause mortalities over 90% in shrimp hatcheries (Soto Rodríguez et al., 2006), these represent an important drawback for shrimp farmers. Bacterial infections, the most prevalent during the shrimps' early life stages, are hard to eradicate. Most antibiotics (with the exception of tetracycline and florfenicol) are banned from the cultures with the result that bacteria have become resistant to them (ICA, 1962–2016). Vaccines are not a viable option due to the fact that shrimps do not seem to have a specific immune system; therefore no immunity is generated (Chong and Maha, 2014). Litopenaeus vannamei, the Pacific white shrimp, is the principal cultivated shrimp worldwide and the infectious diseases, caused by viruses and bacteria, are one of the main barriers to the growth of the ⁎
Corresponding author. E-mail addresses: [email protected] (C. Prada-Peñaranda), [email protected] (M. Salazar), [email protected] (L. Güiza), [email protected] (M.I. Pérez), [email protected] (C. Leidy), [email protected] (M.J. Vives-Florez). https://doi.org/10.1016/j.aquaculture.2017.11.007 Received 16 March 2017; Received in revised form 27 October 2017; Accepted 3 November 2017 Available online 07 November 2017 0044-8486/ © 2017 Published by Elsevier B.V.
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standardize a new method for its biofilm quantification.
amyloliquefaciens and B. pumilus. These species produce important enzymes, vitamins and fermented food (Jeyaram et al., 2011; Rey et al., 2004). B. licheniformis produces an assortment of extracellular enzymes, like proteases and amylases, with significant industrial importance (Rey et al., 2004; Sánchez-M and Romero-R, 2010). Several Bacillus species have been reported as capable to decrease pathogen population, being used as probiotics on the shrimp industries (Martínez Cruz et al., 2012); in particular, B. licheniformis is approved and used as probiotic in the aquaculture industries (Martínez Cruz et al., 2012; Sánchez-M and Romero-R, 2010). Probiotic products made from B. licheniformis, that assure inhibition of pathogenic microorganisms, water purification and improvement of health and growth on aquatic animals, poultry, swine, among others, are currently commercialized (Alibaba 1999–2014), as well as mixtures of B. licheniformis and B. subtilis, to control Vibrio alginolyticus (Zhang et al., 2011). Nevertheless, B. licheniformis have been associated to food poisoning and gastroenteritis (Salkinoja-Salonen et al., 1999). Moreover, according to ElSersy and Mohamed (2011) some B. licheniformis strains can produce exotoxins with strong hemolytic activity, causing a reduction in aquaculture products. Although this bacillus was never used as probiotic in the shrimp's tanks, B. licheniformis was found there. The bacterium could be transferred to the tanks by water, aerosols and/or nearby material. The appearance of re-emergent infections caused by this bacterium, led to the suspicion that a permanent contamination source might exist inside the culture tanks. Additionally, no effective method of treatment is known, and the search for alternatives is crucial. Over 99.9% of bacteria form biofilms (Donlan and Costerton, 2002) and it is common to find them in aqueous environments (Proal, 2008). Biofilms are sessile microbial communities attached to a surface, embedded within a self-produced matrix of extracellular polymeric substance (EPS) (Costerton et al., 2005). Bacteria can form these types of structures in any kind of environment they inhabit, being an essential and protective life strategy for them. The slow cell growth (that decreases the taking up of antimicrobials) and the matrix (which acts as a barrier) make them more resistant to antibiotics and chemical agents (Donlan and Costerton, 2002). The time of formation and complexity of the biofilm depend on different factors, like the type of microorganism, surface, light, quorum sensing, among others (Prescott et al., 2005). Biofilm formation involves three general steps: adherence, growth, and detachment. The first one initiates with a reversible adherence, which in time is converted into irreversible adherence once the exopolysacharide is secreted. The second one involves the formation of micro and macro - colonies, hydrated structures, where bacterial cells get embedded within the EPS and initiate its maturation. The last step, where the bacterial cells detached from the biofilm, enables the colonization of new surfaces, where the complete process starts again (Ghannoum and O'Toole, 2004; Sutherland et al., 2004). According to these, if B. licheniformis can form biofilms, the recurrent shrimps infections could be caused by the cell dispersion during the last stage of biofilm maturation. Due to the serious problem that bacteria cause to the shrimp industry, and the lack of an existing, efficient treatment, phage therapy has been proposed as a promising option. Phage therapy is based on the bactericidal effect of bacteriophages, or phages, viruses that only recognize and kill specific bacterial hosts. Phage therapy does not have negative side effects for humans, animals or plants as shown in several reports (Azeredo and Sutherland, 2008; Caplin, 2009; Frampton et al., 2014; Holguín et al., 2015; Martínez Díaz, 2010; Sulakvelidze et al., 2001). The use of phages has been also reported for the control of biofilms formed by pathogenic bacteria like Pseudomonas aeruginosa (Fu et al., 2010), Proteus mirabilis, Escherichia coli (Carson et al., 2010), and Listeria monocytogenes (Montañez Izquierdo et al., 2012), among others. The main goal of this study was to evaluate the biofilm control capabilities of the phage preparation FBL1, a novel native Colombian bacteriophage preparation. To achieve this goal, it was first necessary to evaluate the biofilm formation ability of B. licheniformis and
2. Materials and methods 2.1. Bacterial strains B. licheniformis strain 52 was used in this study for the biofilm assays. This is a virulent strain isolated from L. vannamei haemolymphs (Gálvez et al., 2016). Taxonomic confirmation of the bacterial isolate was done through the amplification and sequencing of the 16S rRNA gene (Vives Florez and Garnica, 2006). Host range test were performed using five additional bacterial strains of B. licheniformis, isolated by CENIACUA from L. vannamei, and nine other strains from the Bacillaceae family donated by A. Yousten (from Virginia Polytechnic Institute and State University) and A. Delecluse (from The Pasteur Institute, France) to J. Dussán, from Universidad de los Andes, and kindly provided by her for this work (supplemental e-component Table 1) (Cavados et al., 2001; HernandezSantana et al., 2016; Lee et al., 2001; Mahler et al., 1986; Stapleton et al., 2004). 2.2. Culture media and conditions Bacteria were grown at 30 °C on nutrient broth (13 g/L) under 250 rpm agitation. Incubation on nutrient agar (13 g/L nutrient broth and 15 g/L bacteriological agar) for 34–36 h allowed colonies observation. Culture media and components were obtained from Oxoid Ltd. 2.3. Bacteriophage isolation and characterization Phages were isolated from fresh water and sediment samples from shrimp ponds. Selective enrichment of the samples was done in 90 mL nutrient broth, where 1% B. licheniformis 52 was previously inoculated. The enriched culture was incubated over night at 30 °C without agitation. After this incubation period, chloroform was added at a proportion of 1% to the cultures followed by another incubation period (2 h, room temperature, 100 rpm). After this second incubation, the enriched culture plus chloroform was centrifuged (8500 rpm for 30 min at 4 °C), the supernatant (containing the phages) was filtered (0.22-μm pore size) and stored at 4 °C. This filtered supernatant was named phage preparation. Sixteen phage preparations were obtained. Due to the fact that these phages were difficult to concentrate and did not show isolated plaques in serial dilutions plating using the double layer agar method, they were plated by the spot-test method in double agar overlay plates to characterize them by host range test, against the fourteen strains of the Bacillaceae family (supplemental e-component Table 1). Briefly, 3–5 colonies of B. licheniformis 52 were inoculated on 3 mL nutrient broth and grown for 6 h at 30 °C and 250 rpm. 100 μL of each bacterial overnight culture were mixed with 3 mL of soft agar (TAE 1X, 13 g/L nutrient broth and 0.25% w/v agarose) and poured on top of nutrient agar plates. When the soft agar solidified, 10 μL of each phage preparation were spotted on the surface and let to absorb. The plates were incubated at 30 °C for 14 h. Spots that presented a plaque were categorized as a phage preparation with effect on the corresponding bacterial strain (positive, +), while those that resulted in no visible plaque were categorized as ineffective (negative, −). The structure of the viral particles from phage preparation FBL1 was characterized by transmission electron microscopy (TEM) according to the following protocol: drops of FBL1 were deposited on Parafilm®. Carbon-coated Formvar film (grid) were placed over the FBL1 drops for 5 min; the grid was air-dried for 3 min and then it was placed over the phosphotungstic acid solution for 2 min, in order to stain negatively the phage sample. When dried, FBL1 was observed using TEM in a microscope model JEOL 1400 plus. 161
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92 h and 116 h. The weight of the recovered biomasses was compared to controls without FBL1. All the experiments were conducted in triplicate.
2.4. B. licheniformis biofilm formation assay Biofilm formation was initially analyzed by the microtiter plate assay, the most commonly used method to evaluate biofilms. However, the biofilm obtained was thin and lacked adherence to the well walls, causing its loss during the procedure (data not shown). The same negative result was obtained in test tubes and flasks. Consequently, a novel method to evaluate the biofilm formation capacity in this Bacillus species was developed. B. licheniformis 52 biofilm formation was standardized using a microdrip solution administration set (BAXTER 60 drops/mL) (Chávez Escobar and Leidy, 2010) that simulated an aquatic flowing system. It consisted of two erlenmeyers, one of them located at a distance of 1.2 m below the other one, and connected by a 3 m microdrip hose. 5 mL of a 108 CFU/mL bacterial suspension were inoculated directly through the microdrip injection site; the culture was allowed to go down the hose for 10 s and then the system was kept closed for 3 h into the incubator at 30 °C. Afterwards, the flux was adjusted to 0.4–0.7 mL/min. Three different times for biofilm recovery after bacterial inoculation were tested: 24 h, 48 h, and 68 h. The formed biofilm was expelled from the plastic hose, centrifuged (8500 rpm for 30 min at 4 °C), and the pelleted biomass was lyophilized (Lyophilizer LABCONCO model freezone) for 24 h, and weighted. Experiments were conducted in triplicate. Biofilm surface structure was observed by scanning electron microscopy (SEM) (model JSM 6490-LV), using a sample of the recovered biofilm prior lyophilization and without sample preparation.
3. Results 3.1. Bacterial strain The sequence for the rRNA 16S gene obtained was compared with the NCBI gene bank database using BLAST search program (Altschul et al., 1990). The bacterium B. licheniformis 52 was identified as B. licheniformis with an E-value equal to 0.0, 100% query coverage and 99–100% maximum identity. 3.2. Bacteriophage isolation and characterization Sixteen phage preparations were obtained. We named them phage preparations instead of phages because it was not possible to guarantee that they contained only one type of phage. The phages were observed by plating using the spot method, and this method does not allow the definition of phage purity. Presence/absence of plaques in the host range test showed that the phage preparations were different (Table 1). Phage preparation FBL1 was chosen for the subsequent experiments due to its specificity, infecting, apart from the B. licheniformis, only one out of the other nine Bacillaceae strains. It was also effective against the most virulent strains of B. licheniformis, according to the information provided by CENIACUA, including the virulent B. licheniformis 52. TEM images were extremely difficult to obtain because it was not possible to concentrate the phages. Although the poor quality of the photographs, they show that phage preparation FBL1 contains tailed phages with heads of 45.03 nm to 68.41 nm. Due to the presence of long non-contractile tails they probably belong to the Siphoviridae family (Fig. 1).
2.5. Control of B. licheniformis biofilm by the phage preparation FBL1 Once the biofilm formation assay was standardized, experiments were set up to assess the effect of phages on the biofilm formed. Biofilms were treated with 1 mL of the phage suspension FBL1 at three different times: 0 h (simultaneous inoculation of bacteria and phage suspension), 24 h, and 48 h after bacterial inoculation. In all cases, once the phage suspension was added, the flux was suspended for 24 h and then resumed. In all treatments, biofilms were recovered at 68 h after bacterial inoculation. Dry weight of recovered biofilm was compared between treatments and controls (i.e. biofilms without FBL1 inoculation). Additionally, the surface of the 68 h-biofilms treated with FBL1 at 0 h was observed under SEM and compared to the appearance of the 24 and 68 h control biofilms. Each experimental assay was done in triplicate. In order to test for significance, the non-parametric data were analyzed by the Kruskal-Wallis test and the parametric data by the Tukey test using the R platform version 3.0.2, with a confidence level of p < 0.05. To elucidate the nature of FBL1 effect on the biofilm, it was predicted that either it would remove the formed biofilm, or prevent its formation by killing the planktonic cells. If the phages were removing the biofilm, FBL1 would induce the dispersion of planktonic cells and the material around them, which would cause a higher weight of the biomass in the disposal (where the flux from the lower Erlenmeyer was collected) compared to controls without phage treatment. On the other hand, if FBL1 were killing the planktonic cells, the biofilm formation cycle would take longer because of the reduced inoculum, with more time needed to form a mature biofilm. Therefore, the weight of the biomass in the disposal would be lower compared to controls, because the biofilm would not be in its mature stage, and the dispersion final step would not be reached at the same time. The experiment was conducted with and without phage treatment measuring the biomasses differences in the disposal, in order to evaluate which of the predictions was correct. FBL1 was inoculated at 0 h, 24 h, and 48 h after B. licheniformis strain 52.68 h after the bacterial inoculation in every assay, the accumulated disposal biomasses were recovered, freeze-dried and weighted. To assess the extended life cycle of the biofilm, the experiment was conducted as described previously, inoculating FBL1 simultaneously with the bacterium and recovering the biofilm at 68 h,
3.3. B. licheniformis biofilm formation The microdrip solution administration set allowed the formation, recovery (e-component Movie 1) and quantification of B. licheniformis 52 biofilm. To the best of our knowledge, this is the first report of B. licheniformis biofilm observation and quantification. The SEM images show the structure of the biofilms recovered at different times after bacterial inoculation, without FBL1 treatment (Fig. 2A–C). With longer times of incubation, the recovered biofilm increased its structural complexity, as well as its biomass (Fig. 3A). These results showed reproducible data, with low variability between replicates. At 68 h of incubation, a complex, continuous biofilm was observed in the SEM images (Fig. 2C). Thus, 68 h was chosen as the time for biofilm incubation in the following assays. 3.4. Control of B. licheniformis biofilm by the phage preparation FBL1 In order to evaluate the effect of the phage preparation FBL1 on the biofilm formed by B. licheniformis 52, the amount of formed biofilm was measured after it was treated with the phage preparation. The data showed on Fig. 3B evidenced that FBL1 caused a reduction on the dry weight of the biofilm, and the magnitude of these reductions varied according to the time of inoculation of FBL1: at 0 h, 24 h and 48 h, the reduction in the biofilm recovered was 53.33%, 30.16% and 0.79%, respectively. SEM images of the structure of biofilm treated with FBL1 at time 0 h (Fig. 2D), were almost identical to the structure of the non treated (control) biofilm at 24–48 h (Fig. 2A and B). Control biofilm at 68 h after bacterial inoculation (Fig. 2C), showed a hard and robust structure without holes. On the other hand, the phage-treated biofilm depicted a less complex structure, characteristic of earlier stages of biofilm 162
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formation (Fig. 2D). The experiments conducted to elucidate the mode of action of FBL1 on the biofilm identified interesting aspects. Fig. 4 shows that the biomass in the disposal was smaller in all experiments with FBL1, compared to controls. Significant difference (p < 0.05) were only found in the treatment where FBL1 was added at 48 h. This result would support the hypothesis that FBL1 killed the planktonic cells. It might also explain the lowest amount of biomass found in the disposal when FBL1 was inoculated at 48 h, since the phages could kill the dispersed cells from the formed, 48 h – old biofilm. On the other hand, in the control without FBL1, the biofilm continued its normal life cycle and the dispersed cells went to the disposal. Additionally, if the phage suspension killed the planktonic cells, the life cycle of the biofilm should also require a longer time. The results showed that in the normal life cycle of B. licheniformis biofilm (Fig. 3A), the dry weight increased regularly in ca. 5.5 mg every 24 h (Table 2). When the biofilm was treated with FBL1, injecting it simultaneously with the bacterium, the biofilm dry weight increased along time but at a significantly lower rate: from time 68 h to 92 h and 92 h to 116 h augmented in 0.43 mg and 1.94 mg, respectively (Table 2), showing a much lower growth rate compared to the control biofilm. The lower biofilm growth rate had the consequence of an extended biofilm life cycle, providing additional support to the hypothesis of FBL1 action at the planktonic cells stage.
+ − − + + + − −
+ − − − − + − + − − − − + − − − − − − − − + − − − − +
− + − + − + + + − + − − + − − − − − − − − − − − − − − − + − − +
− + − − − + − − − − +
Bacillus licheniformis 54(1)
The present work is the first study showing the standardization and quantification of the biofilm formed by a strain of B. licheniformis. Although there are no reports of B. licheniformis biofilms and it has been published that this bacterium was not known to form biofilms (Dhakal et al., 2013), this study demonstrated that B. licheniformis is capable of forming a robust, complex one, with abundant extracellular matrix produced within 24 h. This finding gives a plausible explanation for the recurrent outbreaks in L. vannamei tanks, since the biofilms can act as a source of new bacterial inocula. We used a new methodology, since the standard microtiter plate method was not useful for this bacterial strain. The new method utilizes a microdrip set, allowing the biofilm to form in a closed and controlled space, delivering reproducible data of biofilm dry weight, and also permitting the observation of its structural surface through SEM. Besides, the microdrip system resembles to the water pipelines used in aquaculture. Once the biofilm formation was standardized, it was possible to analyze the capacity of the phage preparation FBL1 to control it. The present study found that, depending on the time of the FLB1 treatment, the phages produced different outcomes in the amount of recovered biofilm (Fig. 3B). The best result (53.3% reduction) was obtained when the treatment was applied at the same time as the bacterial inoculation, and the worst (0.79%) with the more delayed treatment of 48 h postinoculation. These findings are likely due to the profuse extracellular matrix produced, which is known to provide impermeability and protection (Harper et al., 2014). Despite the resistant characteristics conferred, bacteriophages are excellent antibacterial agents that can overcome the biofilms barriers. However, it depends on the particular phage and biofilm and, in this case, FLB1 was not able to remove mature biofilms of B. licheniformis 52. Regarding the FBL1 mode of action, it was found that FBL1 caused a negative effect on the biofilm formation. The SEM image of the biofilm treated with FBL1 at time 0 h and recovered 68 h later (Fig. 2D), revealed a biofilm structure similar to the first stages of its formation without phage treatment (Fig. 2A and B). Bacteriophages are known to damage biofilms integrity, either killing EPS producing cells or producing enzymes that degrade the extracellular matrix (Harper et al., 2014). In this case, killing the planktonic cells probably caused the reduction of the biofilm.
+: phage preparation with effect on the corresponding bacterial strain; −: phage preparation without visible effect on the strain.
+ + + + + + − + + − + − − +
+
+ + + + + + − + − + + − + + − −
+ + + + + + − + − − − − + − − −
+ + + + + + + + − + + + + + + −
+ + + + + + + + − + + + + + − −
+ + + + + + + + − + + + + + − −
+ − + − + + − + − − − − + − − −
+ + + + + + − + − − − − + − − −
− − − − − − − − − − − − + − − −
4. Discussion
FBL16 FBL15 FBL14 FBL13 FBL12 FBL11 FBL10 FBL9 FBL8 FBL7 FBL6 FBL5 FBL4 FBL3 FBL2 FBL1
Lysinibacillus fusiformis Lysinibacillus sphaericus NRS1198 Lysinibacillus sphaericus LS 2362 Lysinibacillus sphaericus NRS400 Bacillus polymyxa
Bacteria strain Phage preparation
Table 1 Host range pattern of the phage preparations against representative strains of the Bacillaceae family.
Bacillus cereus RC607
Bacillus cereus 3501
Bacillus subtilis BGL
Bacillus thuringiensis
Bacillus licheniformis 52
Bacillus licheniformis 54(2)
Bacillus licheniformis 76
Bacillus licheniformis lume
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Fig. 1. TEM morphology of phage preparation FBL1. TEM morphology of the negatively stained phage preparation FBL1 showing long, non-contractile tails with heads of 45.03 nm to 68.41 nm. The morphology is compatible with Siphoviridaelike phages. (A) Original image. (B) Adjusted image: cutting the borders and getting a better contrast and brightness through the Fiji platform.
48 h. It is worth to mention that the amount of biofilm formed at 116 h was significantly lower (34.28%), compared to the controls, demonstrating the effect of FBL1. Additionally, the FBL1 treated biofilm grew at a lower rate (Table 2). These data also showed a gradual increase in the growth rate (0.017 mg/h to 0.080 mg/h); we expected that the biofilm would recover its normal growth rate (ca. 0.2 mg/h), as the effect of the phage suspension ends; however, we followed this experiment only up to the 116 h. These observations strongly suggest that FBL1 prevented the biofilm formation of B. licheniformis by killing
Analyzing the mode of action of FBL1, it was found that only its early addition was associated with a significant decrease in the dry weight of the collected biofilm, indicating that FBL1 killed the planktonic cells instead of removing the already formed biofilm. Thus, the B. licheniformis treated biofilm life cycle was longer than the untreated control. When the biofilm was treated with FBL1 and recovered at 92 h and 116 h, it did not even reach the amount of biofilm recovered at 68 h in the control assays (without FBL1). Moreover, the dry weigh of the biomass was similar to the biofilm biomasses of the control assays at
Fig. 2. SEM images of Bacillus licheniformis biofilm with and without phage preparation FBL1. B. licheniformis strain 52 biofilm recovered at (A) 24 h (B) 48 h and (C) 68 h. Fig. 2 (D) presents the B. licheniformis biofilm obtained after treatment with phage preparation FBL1; FBL1 was inoculated at time 0, and the biofilm formed was recovered 68 h later. Note the difference in complexity between the control and treated biofilms at 68 h.
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planktonic cells and delaying its development. The use of bacteriophages to control bacterial biofilms has been reported elsewhere, using the microtiter plate method (Carson et al., 2010; Holguín et al., 2015; Montañez Izquierdo et al., 2012). As far as we know, there are no reports on the use of phages to treat B. licheniformis planktonic cells or biofilms. In fact, not even the biofilm formed by Bacillus subtilis, the Bacillus species most studied, has been quantitatively assessed. Usually, biofilms of B. subtilis have been observed at the liquid-air interface of broth cultures, or by observing the colonies formed on the surface of agar plates, but the resulting data cannot be systematically quantified (Lemon et al., 2008; Vlamakis et al., 2013). Thus, this is the first time that the observation, standardization and quantification of B. licheniformis biofilm is reported. We were able to obtain the novel phage preparation FBL1 although the phages isolated in this work were difficult to titer and concentrate; we assayed a wide variety of methodologies (lysates, ultracentrifugation at 40,000 g, enrichment from isolated plaques, and lysates using agar) and did not obtain a concentrated suspension. This limitation prevented further characterization, like genome size estimation, sequencing and annotation. Additional work on these areas is needed to rule out the presence of integration sequences and other undesirable genes. A preliminary characterization based on phage structure allowed the putative classification of FBL1 in the Siphoviridae family, because of the binary morphology and the long non-contractile tail. According to the classification of tailed bacteriophages, phages from Bacillus species belong to the Myoviridae, Podoviridae (Kutter and Sulakvelidze, 2005) and Siphoviridae families (Yuan et al., 2012). Some reports have described B. licheniformis phages from Siphoviridae with a head diameter ranging from 41 nm to 66 nm (Hertel et al., 2015; Huang and Marmur, 1970); FBL1 phages showed a similar head size. In summary, the results obtained in this work give new information about the B. licheniformis ability to form biofilms, and offer a new method to study them. Also, phage therapy may represent a future alternative to control bacterial pathogens for the shrimp industry, provided that better characterized phages were available. Supplementary data to this article can be found online at https:// doi.org/10.1016/j.aquaculture.2017.11.007. Video showing the recovery of B. licheniformis biofilm formed inside the microdrip solution administration set.
Fig. 3. Bacillus licheniformis biofilm formation and effect of phage preparation FBL1. (A) Standardization of B. licheniformis strain 52 biofilm. The bars represent the amount of biofilm biomass recovered at five different times (24, 48, 68, 92 and 116 h) after the bacterial inoculation. All experiments were significantly different between each other (P < 0.05). (B) B. licheniformis strain 52 biofilm formed with and without FBL1 treatment, showing the effect of time addition of the phage suspension on the biofilm biomass recovered. FBL1 was inoculated at three different times after bacterial inoculation: 0 h, 24 h and 48 h. For all treatments, biofilm was recovered and the biomass was measured as dry weight 68 h after the bacterial inoculation. The asterisk (*) indicates that the difference was significant (P < 0.05). Controls (black bar) were measured independently for each treatment; the black bar represents the average of nine replicates (3 for each experiment). All experiments were conducted in triplicate.
Declaration of interest Catalina Prada-Peñaranda and Martha J. Vives-Florez are members of the spin off SciPhage S.A.S., which works for the development of phage therapy in Colombia.
Contributions Catalina Prada-Peñaranda: Conception and design, acquisition of data, biofilm formation assay, analysis and interpretation of data, drafting the article, revising it critically for important intellectual content, final approval of the version to be published critically for important intellectual content. Martha J. Vives-Florez: Conception and design, analysis and interpretation of data, drafting the article, revising it critically for important intellectual content, final approval of the version to be published critically for important intellectual content. Marcela Salazar, Linda Güiza: strains isolation and preliminary phage tests, revising the article, final approval of the version to be published critically for important intellectual content. Maria Isabel Pérez, Chad Leidy: biofilm formation assay, revising the article, final approval of the version to be published critically for important intellectual content.
Fig. 4. FBL1 prevents Bacillus licheniformis biofilm by killing the planktonic cells and delaying its formation. B. licheniformis strain 52 detached biomass with and without FBL1 treatment, when phage preparation was inoculated at 0 h, 24 h and 48 h after B. licheniformis strain 52. For all treatments, biofilm was recovered and the accumulated biomass was measured as dry weight 68 h after the bacterial inoculation. These results indicate that FBL1 kills the planktonic cells and lags the biofilm growth (see text). Controls (black bar) were measured independently for each treatment; the black bar represents the average of nine replicates (3 for each experiment). All experiments were conducted in triplicate. The asterisk (*) indicates that the difference was significant (P < 0.05).
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Table 2 Bacillus licheniformis strain 52 biofilm dry weight and growth rate, with and without phage suspension FBL1. B. licheniformis 52 (control) Time (h) 68 92 116
Dry weight (mg) 12.17 17.57 23.24
Difference – 5.40 5.67
B. licheniformis 52 + FBL1 (simultaneous inoculation) Biofilm growth rate (mg/h) – 0.225 0.236
Dry weight (mg) 5.60 6.03 7.97
Funding
Difference – 0.43 1.94
Biofilm growth rate (mg/h) – 0.017 0.080
% of biofilm formed compared to control 46.03 34.35 34.28
aeruginosa on catheters in an in vitro model system. Antimicrob. Agents Chemother. 54, 397–404. Gálvez, E.J.C., Carrillo-Castro, K., Zárate, L., Güiza, L., Pieper, D.H., García-Bonilla, E., Salazar, M., Junca, H., 2016. Draft genome sequence of Bacillus licheniformis CG-B 52, a highly virulent bacterium of Pacific white shrimp (Litopenaeus vannamei), isolated from a Colombian Caribbean aquaculture outbreak. Genome Announc. 4 (3), e00321–16. http://dx.doi.org/10.1128/genomeA.00321-1. Ghannoum, M., O'Toole, G.A. (Eds.), 2004. Microbial Biofilms. ASM Press, Washington. Harper, D.R., Parracho, H.M.R.T., Walker, J., Sharp, R., Hughes, G., Werthén, M., Lehman, S., Morales, S., 2014. Bacteriophages and biofilms. Antibiotics. 3, 270–284. Hernandez-Santana, A., Gómez-Garzón, C., Dussán, J., 2016. Complete genome sequence of Lysinibacillus sphaericus WHO reference strain 2362. Genome Announc. 4, e0054516. http://dx.doi.org/10.1128/genomeA.00545-16. Hertel, R., Rodríguez, D.P., Hollensteiner, J., Dietrich, S., Leimbach, A., Hoppert, M., Liesegang, H., Volland, S., 2015. Genome-based identification of active prophage regions by next generation sequencing in Bacillus licheniformis DSM13. PLoS One 10, e0120759. http://dx.doi.org/10.1371/journal.pone.0120759. Holguín, A.V., Rangel, G., Clavijo, V., Prada, C., Mantilla, M., Gomez, M.C., Kutter, E., Taylor, C., Fineran, P.C., Gonzalez Barrios, A.F., Vives, M.J., 2015. Phage _Pan70, a putative temperate phage, controls Pseudomonas aeruginosa in planktonic, biofilm and burn mouse model assays. Viruses. 7, 4602–4623. Huang, W.M., Marmur, J., 1970. Characterization of inducible bacteriophages in Bacillus licheniformis. J. Virol. 5, 237–246. ICA, 1962–2016. Productos medicamentos veterinarios con registro ICA vigente a ***20 de octubre 2015***. In: Colombia: Instituto Colombiano Agropecuario, . http:// www.ica.gov.co/Areas/Pecuaria/Servicios/Regulacion-y-Control-de-MedicamentosVeterinarios/Medicamentos/Listados/2015/03082015/Productos-MedicamentosVeterinarios-1.aspx (accessed 31.04.16). Jeyaram, K., Romi, W., Singh, T.A., Adewumi, G.A., Basanti, K., Oguntoyinbo, F.A., 2011. Distinct differentiation of closely related species of Bacillus subtilis group with industrial importance. J. Microbiol. Methods 87, 161–164. Kutter, E., Sulakvelidze, A. (Eds.), 2005. Bacteriophages: Biology and Applications. CRC Press, United States of America. Lee, I.H., Je, Y.H., Chang, J.H., Roh, J.Y., Oh, H.W., Lee, S.G., Shin, S.C., Boo, K.S., 2001. Isolation and characterization of a Bacillus thuringiensis ssp. kurstaki strain toxic to Spodoptera exigua and Culex pipiens. Curr. Microbiol. 43, 284–287. Lemon, K.P., Earl, A.M., Vlamakis, H.C., Aguilar, C., Kolter, R., 2008. Biofilm development with an emphasis on Bacillus subtilis. Curr. Top. Microbiol. Immunol. 322, 1–16. Mahler, I., Levinson, H.S., Wang, Y., Halvorson, H.O., 1986. Cadmium- and Mercuryresistant Bacillus strains from a salt marsh and from Boston Harbor. J Appl Environ Microbiol. 52, 1293–1298. Martínez Cruz, P., Ibañez, A.L., Monroy Hermosillo, O.A., Ramirez Saad, H.C., 2012. Use of probiotics in aquaculture. ISRN Microbiol. 2012, 13. Martínez Díaz, S.F., 2010. Phage therapy provides targeted bacteria treatment. In: Innovation, . http://pdf.gaalliance.org/pdf/GAA-Diaz-Jan10.pdf (accessed 20.05.15). MercoPress, 1997–2016. FAO Forecasts Shrimp World Production Could Double by 2030 [Internet]. South Atlantic News Agency, Montevideo. http://en.mercopress.com/ 2014/10/08/fao-forecasts-shrimp-world-production-could-double-by-2030 (accessed 20.01.16). Montañez Izquierdo, V.Y., Salas-Velazquez, D.I., Rodriguez-Jerez, J.J., 2012. Use of epifluorescence microscopy to assess the effectiveness of phage P100 in controlling Listeria monocytogenes biofilms on stainless steel surfaces. Food Control 23, 470–477. Newmark, U.F., Valverde, B.C., Humberto-Díaz, L.J.M., Parra, A.G., Bonilla, S.P., Salazar, V.M., 2009. Agenda prospectiva de investigación y desarrollo tecnológico para la cadena productiva de camarón de cultivo en Colombia. In: Bogotá (Colombia): Corporación Centro de Investigación de la Acuicultura de Colombia (CENIACUA). Universidad Nacional de, Colombia. Prescott, M.L., Harley, J.P., Klein, D.A., 2005. Microorganism interactions and microbial ecology microbiology. In: Prescott, M.L., Harley, J.P., Klein, D.A. (Eds.), Microbiology. McGraw-Hill, New York, pp. 577–613. Proal, A., 2008. Understanding Biofilms. [Internet]. Bacteriality Understanding Chronic Disease. http://bacteriality.com/2008/05/biofilm/ (accessed 22.03.16). Rey, M.W., Ramaiya, P., Nelson, B.A., Brody-Karpin, S.D., Zaretsky, E.J., Tang, M., Lopezde-Leon, A., Xiang, H., Gusti, V., Clausen, I.G., Olsen, P.B., Rasmussen, M.D., Andersen, J.T., Jorgensen, P.L., Larsen, T.S., Sorokin, A., Bolotin, A., Lapidus, A., Galleron, N., Ehlich, S.D., Berka, R.M., 2004. Complete genome sequence of the industrial bacterium Bacillus licheniformis and comparisons with closely related Bacillus species. Genome Biol. 5, 12. Salkinoja-Salonen, M.S., Vuorio, R., Andersson, M.A., Kämpfer, P., Andersson, M.C., Honkanen-Buzalski, T., Scoging, A.C., 1999. Toxigenic strains of Bacillus licheniformis related to food poisoning. J Appl Environ Microbiol. 65, 4637–4645. Sánchez-M, M., Romero-R, F., 2010. Producción de proteasas con Bacillus licheniformis ATCC 1259 y su acción hidrolítica sobre proteínas de arroz (Oryza sativa L.) y maíz
This work was supported by: Banco Santander, grant Premio Santander Emprendimiento Ciencia e Innovación 2010 awarded to MJV-F; the scholarship granted to CP-P by the Department of Biological Sciences, Universidad de los Andes, for her Master studies in Biological Sciences, area Microbiology; the Faculty of Sciences, Universidad de los Andes, project code OI160422001-BIO15; and the Office of the Vice president for Research, Universidad de los Andes, project code P16853122.003/09-01. Acknowledgments The authors thank Paula Esquinas from Fundación Santa Fé de Bogotá, Colombia, for the help with the TEM images; Dr. Jenny Dussán for lending the strains for the host range assays. We also thank Banco Santander for the financial support. Many thanks to the Faculty of Sciences Research Fund and the Department of Biological Sciences, Universidad de los Andes, that provided partial funding for the research and the CP-P Masters student scholarship, respectively. We acknowledge the Office of the Vicepresident for Research, Universidad de los Andes for the finantial support to CP-P. We thank Humberto Ibarra for the invaluable help with the TEM images. References Alibaba, 1999–2016. Leading Platform for Global Wholesale Trade. https://www.alibaba. com/trade/search?fsb=y&IndexArea=product_en&CatId=&SearchText= Aquaculture+Probiotics+Bacillus+Licheniformis (accessed 22.02.16). Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403–410. Azeredo, J., Sutherland, I.W., 2008. The use of phages for the removal of infectious biofilms. Curr. Pharm. Biotechnol. 9, 261–266. Caplin, J., 2009. Bacteriophage Therapy — Old Treatment, new Focus? http://eprints. brighton.ac.uk/6092/1/SfAM_article_BACTERIOPHAGE_THERAPY_-_OLD_ TREATMENT_NEW_FOCUS_LH.pdf (accessed 15.02.14). Carson, L., Gorman, S.P., Gilmore, B.F., 2010. The use of lytic bacteriophages in the preventionand eradication of biofilms of Proteus mirabilis and Escherichia coli. FEMS Immunol. Med. Microbiol. 59, 447–455. Cavados, C.F.G., Fonseca, R.N., Chaves, J.Q., Rabinovitch, L., Araújo-Coutinho, C.J.P.C., 2001. Identification of entomopathogenic Bacillus isolated from Simulium (Diptera, Simuliidae) larvae and adults. Mem. Inst. Oswaldo Cruz 96, 1017–1021. Chávez Escobar, J.P., Leidy, C., 2010. Regulación del nivel de fluidez membranal de bacterias patogénicas: formación de biopelículas y adaptación a la termorregulación del hospedero. dissertation. Universidad de los Andes, Colombia. Chong, C.M., Maha, A., 2014. Fish and Crustaceans: Immune System, Vaccine Development and Implications. In: Nurul Amin, S.M., Kamarudin, M.S., Arshad, A., Romano, N. (Eds.), Perspective of Fisheries and Aquaculture in Malaysia. Universiti Putra Malaysia Press, Selangor, pp. 188–207. Costerton, J.W., Montanaro, L., Arciola, C.R., 2005. Biofilm in implant infections: its production and regulation. Int J Artif Organs. 28, 1062–1068. Dhakal, R., Chauhan, K., Brent Seale, R., Deeth, H.C., Pillidge, C.J., Powell, I.B., Craven, H., Turner, M.S., 2013. Genotyping of dairy Bacillus licheniformis isolates by high resolution melt analysis of multiple variable number tandem repeat loci. Food Microbiol. 34, 344–351. Donlan, R.M., Costerton, J.W., 2002. Biofilms: survival mechanisms of clinically relevant microorganism. Clin. Microbiol. Rev. 15, 167–193. ElSersy, N.A., Mohamed, E.A.H., 2011. Biochemical and molecular characterization of hemolytic Bacillus licheniformis strains isolated from shrimp and clam aquacultures. Afr. J. Microbiol. Res. 5, 1830–1836. Frampton, R.A., Taylor, C., Holguín Moreno, A.V., Visnovsky, S.B., Petty, N.K., Pitman, A.R., Fineran, P.C., 2014. Identification of bacteriophages for biocontrol of the kiwifruit canker phytopathogen Pseudomonas syringae pv. actinidiae. J Appl Environ Microbiol. 80, 2216–2228. Fu, W., Forster, T., Mayer, O., Curtin, J.J., Lehman, S.M., Donlan, R.M., 2010. Bacteriophage cocktail for the prevention of biofilm formation by Pseudomonas
166
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C. Prada-Peñaranda et al.
environmental Pseudomonas aeruginosa isolates. Int. Microbiol. 9, 247–252. Vlamakis, H., Chai, Y., Beauregard, P., Losick, R., Kolter, R., 2013. Sticking together: building a biofilm the Bacillus subtilis way. Nat. Rev. Microbiol. 11, 157–168. WWF, 1961–2016. farmed shrimp. In: Washington: World Wildlife Fund, . http://www. worldwildlife.org/industries/farmed-shrimp (accessed 28.05.16). Yuan, Y., Gao, M., Wu, D., Liu, P., Wu, Y., 2012. Genome characteristics of a novel phage from Bacillus thuringiensis showing high similarity with phage from Bacillus cereus. PLoS One 7, e37557. http://dx.doi.org/10.1371/journal.pone.0037557. Zhang, Q., Tan, B., Mai, K., Zhang, W., Hongming, M., Ai, Q., Wang, X., Liufu, Z., 2011. Dietary administration of Bacillus (B. licheniformis and B. subtilis) and isomaltooligosaccharide influences the intestinal microflora, immunological parameters and resistance against Vibrio alginolyticus in shrimp, Penaeus japonicus (Decapoda: Penaeidae). Aquac. Res. 42, 943–952.
(Zea mays L.). dissertation. Universidad Nacional experimental Simón Rodríguez, Republica Bolivariana de Venezuela. Soto Rodríguez, S.A., Simoes, N., Roque, A., Gómez, G.B., 2006. Pathogenicity and colonization of Litopenaeus vannamei larvae by luminescent vibrios. Aquaculture 258, 109–115. Stapleton, P., Pike, R., Mullany, P., Lucas, V., Roberts, G., Rowbury, R., Wilson, M., Richards, H., 2004. Mercuric resistance genes in gram-positive oral bacteria. FEMS Microbiol. Lett. 236, 213–220. Sulakvelidze, A., Alavidze, Z., Morris, G., 2001. Bacteriophage therapy. Antimicrob. Agents Chemother. 45 (3), 649–659. Sutherland, I.W., Hughes, K.A., Skillman, L.C., Tait, K., 2004. The interaction of phage and biofilms. FEMS Microbiol. Lett. 232, 1–6. Vives Florez, M., Garnica, D., 2006. Comparison of virulence between clinical and
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