Background
Vibriosis is one of the most prevalent bacterial diseases that causes mortality of shrimp, fish and shellfish. It results from contamination with
Vibrio pathogens, such as
V. parahaemolyticus,
V. vulnificus,
V. alginolyticus and
V. harveyi [
1,
2]. With the rapid development of aquaculture and the rise in consumption of aquatic products, vibriosis in global aquaculture and
Vibrio-related food poisoning cases are increasing [
1,
3,
4].
V. parahaemolyticus is ubiquitous in seawater and seafood-associated environments [
5‐
7]. According to a comprehensive review and meta-analysis of studies between 2003 and 2015 on the occurrence and prevalence of
V. parahaemolyticus in seafood, it could be isolated from 47.5% of all seafood samples, and showed high overall prevalence rates in oysters (63.4%), clams (52.9%), fish (51.0%), shrimps (48.3%), and mussels, scallop and periwinkle (28.0%) [
8]. Eating any raw
V. parahaemolyticus-contaminated seafood can lead to gastroenteritis because the pathogen contains various virulence factors, including adhesin, heat-resistant direct hemolysin (TDH), TDH-associated hemolysin, and the type III secretory system [
9]. The reported incidence of
V. parahaemolyticus in fishery products in China was approximately 15% during the period 2008 to 2017 [
10]. The administration of antibiotics to quickly and effectively control pathogenic
Vibrio strains became common in aquaculture. However, prolonged use of antibiotics has resulted in an increase in the number of multidrug-resistant
Vibrio strains, presenting a substantial threat to the control of vibriosis [
11‐
13]. Calls have been made to decrease antibiotic use, and alternative measures to control bacterial pathogens are urgently required.
Phages infect and lyse host bacterial cells and plunder host cell resources for their own reproduction [
14‐
17]. Phages have been known as a potential antibacterial agent for over a century; they were used against human bacterial infection in the 1920s [
18]. Previous studies have reported on global efforts to prevent and control
Vibrio in aquaculture by using phages. Numerous in vitro and in vivo studies have shown that phage therapy was effective in controlling vibriosis caused by
Vibrio (e.g.,
V. parahaemolyticus and
V. harveyi) in shrimp, sea cucumber, abalone, oysters, and other species [
19‐
23]. By way of illustration, a 78.1% reduction in bacterial counts was observed within 1 h of phage application during an efficacy study of phage against
V. parahaemolyticus in shrimp [
23].
V. parahaemolyticus-infected shrimp larvae regained activity with no significant reduction in survival when they were treated with phage [
20]. The application of phages in a variety of aquaculture situations highlights the potential of phage therapy to decrease bacterial infections and diminish the significant economic losses to aquaculture and the harm to public health caused by contamination with
Vibrio.
The selection of appropriate phages is, however, a prerequisite, and important tests need to be performed before phage therapy can be applied in the field. In this study, a novel bacteriophage that infects V. parahaemolyticus 1.1997T was isolated from sewage from the Chigang seafood market in Guangdong, China. The phage was characterized based on morphological, host specificity, life cycle, genomic character and taxonomy, and evaluated for its bactericidal ability and possible use in phage therapy.
Methods
Phage isolation and purification
V. parahaemolyticus 1.1997
T brought from Guangdong marine pathogenic
Vibrio company in China was used as the host bacterium for phage isolation. It was incubated in rich organic (RO) medium (1 M peptone, 1 M yeast extract, and 1 M sodium acetate in artificial seawater, pH 7.5) at 30℃ with a shaking speed of 160 rpm min
− 1 [
24]. The sewage samples used for phage isolation were collected from the seafood markets in Guangzhou, China (23.10°N, 113.33°E), and filtered through a 0.22-µm membrane (Millipore, Massachusetts, USA) to remove bacteria and large particles. The sewage samples were added into an exponentially growing host bacterial culture to allow plaque formation using the double-layer agar method [
25]. A clear individual plaque was collected and suspended in storage medium (SM; 8 mM MgSO
4, 50 mM Tris-HCl, and 100 mM NaCl, pH 7.5). After purifying at least five times, the well-separated plaque was collected and stored in SM at 4 °C.
Phage amplification and enrichment
To obtain high-titer phage suspension, a purified phage plaque was inoculated into bacterial culture and amplified overnight, followed by centrifugation at 12,000 × g for 10 min. The supernatant was filtered through a 0.22-µm membrane to remove cell fragments. The filtrate was precipitated with polyethylene glycol 8000 (10% w/v) overnight. Then, phage pellets were obtained through centrifugation (10,000 × g, 60 min, 4 °C) and resuspended in SM. The phage particles were subjected to cesium chloride solutions (ρ = 1.3, 1.5, 1.7 g mL− 1) for purification and centrifuged at 200,000 × g at 4 °C for 24 h using an Optima L-100 XP ultracentrifuge (Beckman Coulter, CA, USA). The purified phage particles were dialyzed through 30-kDa superfilters (Millipore, Bedford, MA, USA).
Transmission electron microscopy (TEM)
Phage morphology was characterized by TEM. Briefly, approximately 20 µL of phage suspension was added onto the surface of a copper grid to adsorb in darkness for 30 min. Then, the phage sample was negatively stained with phosphotungstic acid (1%, pH 7.0) for 20 min and dried for 30 min. The phage sample was examined using a JEM-2100 transmission electron microscope (JEOL, Tokyo, Japan).
Determination of the host range
A total of 35
Vibrio strains, including 6 strains of
V. parahaemolyticus and 29 strains of other species (
V. alginolyticus,
V. antiquarius,
V. azureus,
V. campbellii,
V. caribbeanicus,
V. chagasii,
V. cholerae,
V. fortis,
V. harveyi,
V. inhibens,
V. neocaledonicus,
V. owensii,
V. rotiferianus,
V. tubiashii,
V. variabilis and
V. xuii), were used to determine the lytic host range of the phage isolated in this study with the double-layer agar method (Table
S1). The bacteria strains were grown on the RO agar medium plates and single colony was collected and then incubated in RO liquid medium at 30℃ with a shaking speed of 160 rpm min
− 1. The log-phase bacteria strains (incubated about 2–10 h) were mixed with the phage and kept in the dark for 20–30 min for infection. After infection, the phage-host mixtures were added with molten RO agar medium and poured onto solid agar plates, then incubated overnight at 28 °C. All bacteria strains have been tested at least three times. The presence of plaques on a bacterial lawn was checked to determine phage infection of the host bacterium.
Lipid test
To investigate whether the phage capsids contain lipid, phages were mixed with 0%, 0.2%, 2% or 20% chloroform and incubated in darkness at room temperature for 30 min. Then, the mixtures were centrifuged at 12,000 × g for 5 min and the phages were obtained from the upper suspension. The chloroform sensitivity of the phages was determined by the presence or absence of plaques on double-layer agar. The lipid test was carried out twice.
One-step growth curve
A one-step growth curve was determined to study the infectivity and replication ability of the phage. Briefly, phages were added to an early log-phase host culture (
V. parahaemolyticus 1.1997
T) at a multiplicity of infection of 0.03 and incubated for 20 min at room temperature in the dark. Then, phages that were not adsorbed were removed by centrifugation (8,000 ×
g, 4 °C, 5 min), and the cell pellets were washed and resuspended in 100 mL of RO medium. The phage suspension was incubated at 28 °C with a shaking of 160 rpm min
− 1. Subsamples were obtained at 10-min intervals and assayed by the double-layer agar method. The burst size was calculated as the ratio between the number of phages before and after the burst [
26]. The experiment of one-step growth curve has been conducted with three replications.
Determination of lytic activity
To understand the efficiency of bacterial inactivation, a lytic activity for the phage against V. parahaemolyticus were assessed at different multiplicity of infections (MOIs) by monitoring the OD600 (optical density measurements at a wave-length of 600 nm) using a microplate reader (Synergy H1, Bio-Tek, USA). Briefly, freshly prepared host culture was added to the a 96-well plate before infection with high-titer phage suspension at various MOIs (0.001, 0.01, 0.1, 1 and 10). Wells inoculated with RO medium or V. parahaemolyticus only was served as control groups. The plate was placed in the instrument and incubated at 30℃ with orbital shaking. The OD600 of the cultures were monitored in real time and recorded every 30 min. The assay was conducted with eight replications for each treatment.
DNA extraction and phage genome sequencing
Phage DNA was obtained by using the phenol–chloroform extraction method. Briefly, approximately 1 mL of high-titer phage suspension was treated with proteinase K, sodium dodecyl sulfate (10% w/v), and ethylenediaminetetraacetic acid (EDTA; pH 8.0) and incubated at 55 °C for 3 h. The digested sample was purified with phenol/chloroform/isoamyl alcohol (25:24:1 v:v:v) and chloroform/isoamyl alcohol (24:1 v:v) to remove any debris. Next, the DNA pellet was washed with pre-cooled 70% ethanol, air-dried at room temperature, dissolved in Tris-EDTA buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0), and stored at 4 °C. The phage genome was sequenced and assembled using the Illumina HiSeq 4000 platform with a 150-bp paired-end DNA library and Velvet software (v.1.2.03) [
27].
Genome annotation
The phage termini and packaging mechanism were identified by the online service PhageTerm on a Galaxy-based server (
https://galaxy.pasteur.fr) [
28]. The GeneMarkS online server (
http://topaz.gatech.edu/GeneMark/genemarks.cgi) was used to identify putative open reading frames (ORFs) [
29]. The functions of ORFs were annotated by BLASTP online against the non-redundant (nr) database of the National Center for Biotechnology Information (NCBI), with a cut-off
e-value of < 10
− 5, and the results were checked manually. Putative transfer RNA (tRNA) genes were detected with tRNAscan-SE (
http://lowelab.ucsc.edu/tRNAscan-SE/) [
30]. Virulence factors and antibiotic resistance-encoding genes were searched using the Virulence Factor Database (VFDB,
http://www.mgc.ac.cn/VFs/main.htm) and the Comprehensive Antibiotic Resistance Database (CARD,
https://card.mcmaster.ca/analyze/rgi) webservers, respectively [
31,
32]. In addition, genome-based life cycle classification was also carried out using an online platform called PhageAI (
https://phage.ai/). The complete genome data was deposited in the GenBank database with accession number OP793884.
Phylogenetic and taxonomic analysis
Viral CONTigs Automatic Clustering and Taxonomy v.2.0 (vConTACT2) was used to compare the isolated phage against the Prokaryotic Viral RefSeq 207 database using whole genome gene-sharing profiles, and related phages were identified by genome pairs with a similarity score > 1 [
33]. The whole phage sequence was also submitted to NCBI BLASTN to search forsimilar sequences. To analyze the evolutionary relationships of the phage, complete amino acid profiles of phages were submitted to the Virus Classification and Tree Building Online Resource (VICTOR,
https://ggdc.dsmz.de/victor.php) for phylogenetic tree construction [
34,
35]. All pairwise comparisons of the amino acid sequences were conducted using the Genome-BLAST Distance Phylogeny (GBDP) method with the settings recommended for prokaryotic viruses [
34,
36]. Branch support was inferred from 100 pseudo-bootstrap replicates each, and the tree was rooted at the midpoint and visualized with ggtree [
37,
38]. In addition, intergenomic nucleotide sequence similarity and aligned genome fractions within the imported phages were plotted with the Virus Inter-genomic Distance Calculator (VIRIDIC) using default parameters [
39].
Conclusions
This study isolated a lytic phage, vB_VpaM_R16F (R16F), active against Vibrio parahaemolyticus 1.1997T and investigated its biological properties. R16F is a myovirus with an icosahedral head and a contracted tail. R16F exhibits a narrow host spectrum, highly effective lytic activity, small burst size, and short latent period. A large portion of the genes of R16F are annotated as hypothetical proteins. However, serval functional genes that may improve the environmental competitiveness of the phage were found in the genome of R16F. Antibiotic resistance or virulence factor-related genes were not detected. R16F was most closely related to vibriophage qdvp001 with only 67% pairwise sequence identity and the highest value of intergenomic similarity was only 34.9%. R16F is considered a newly described vibriophage with a genome that differs from all previously described, belonging to a novel genus. The biological and genetic properties suggest that R16F has the potential to be used as a biocontrol agent for vibriosis induced by V. parahaemolyticus. However, further experiments still need to be conducted in the future to investigate the potential of R16F as a biological reagent for phage therapy, specifically with regards to lysogenic testing, pH and temperature stabilities, and so on. These follow-up studies will provide a deeper understanding of the practical applications of R16F in combating vibriosis caused by V. parahaemolyticus.
Open AccessThis article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit
http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (
http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.
Publisher’s Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.