Bacterial predators in the coral microbiome could be a type of top-down control, that directly alters the structure and function of the coral microbiome as demonstrated in other aquatic systems by bacterivorous predators (see reviews by Jürgens & Matz, 2002 ; Pernthaler, 2005 ; Matz & Kjelleberg, 2005 ). For example, we highlighted potential interactions of Halobacteriovorax and other members of the coral holobiont using co-occurrence network analysis of an in-field experimental time series of three coral genera, across three years, several treatments, and range of temperature conditions. These networks showed that Halobacteriovorax are core members of the coral microbiome, present in >78% of samples from three coral genera Porites, Agarica, and Siderastrea ( Welsh et al., 2016 ; Zaneveld et al., 2016 ). We also showed that isolated strains of coral-associated Halobacteriovorax prey upon known coral pathogens in cultured settings. Such antagonisms between predators and prey in the holobiont may have variable effects on the microbiome, such that they could be occlusive to pathogens or disruptive to the coral microbiome itself. Here we examine how a specific bacterial predator ( Halobacteriovorax ), a foreign bacterium ( V. coralliilyticus ), and a coral host ( M. cavernosa ) interact to affect the complex system of the coral microbiome in a laboratory-based system. While V. coralliilyticus is not associated with causing disease in M. cavernosa , our study still provides baseline experiments for predator and prey interactions on a coral host.

We have recently described how the predatory bacteria Halobacteriovorax , also likely influences the diversity and dynamics of the microbial community in the coral surface mucus layer through consumption of a broad range of bacterial prey ( Welsh et al., 2016 ). Halobacteriovorax spp. are small, highly motile predatory bacteria that exhibit a biphasic lifestyle and prey exclusively on gram negative bacteria, including known coral pathogens ( Williams, Falkler & Shay, 1980 ; Welsh et al., 2016 ). Halobacteriovorax are the marine component of a group of delta-proteobacteria known as Bdellovibrio and like organisms (BALOs). In free-living attack phase, BALOs actively seek out prey in order to attach, burrow inside, and restructure their host cell into a rounded bdelloplast. This kills their prey and provides BALOs with an osmotically stable structure free from competition to utilize prey resources for growth and replication. A new generation of attack-phase predators then bursts forth from the bdelloplast to seek new hosts.

Methods

Bacterial strains, growth conditions, and prey range assays V. coralliilyticus are known to be naturally abundant (Wilson et al., 2013) and infect corals (Ushijima et al., 2014; Tout et al., 2015). Although, V. coralliilyticus has not been shown to cause disease in the coral M. cavernosa, we believe that V. coralliilyticus serves as an interesting model to investigate the interactions and effects of bacterial predation on a host, given that it is a well understood bacterium and is consumed by our model predator, Halobacteriovorax, a member of the coral core microbiome (Welsh et al., 2016). We first evaluated if Vibrios can be preyed upon by coral-associated Halobacteriovorax, by conducting a series of predation assays in liquid and solid media. Bacterial strains Vibrio fortis PA1 and Vibrio coralliilyticus ATCC BAA450 (Accession # KT626460 and AJ440005, respectively), were grown on Marine 2216 Agar (MA) overnight. A single colony was re-suspended in 50 mL Marine 2216 Broth (MB) in a 250 mL flask at 30 °C and 250 rpm overnight. Cultures were diluted 1:100 in fresh media and incubated until late exponential growth before use in any experiment. Our predatory bacterial strain, Halobacteriovorax sp. PA1 (Accession # KR493097), was grown as previously reported (Welsh et al., 2016) in Pp20 media. Briefly, a single plaque from a double layer plate was resuspended in 3 mL of 109 V. fortis cells in filtered seawater in 15 mL test tubes. The culture was incubated at 28 °C and shaking at 250 rpm overnight. A 1:100 dilution of the overnight culture was prepared by adding 0.5 mL of 0.45 µm filtered culture to 50 mL of 109 V. fortis PA1 cells in filtered seawater. This new co-culture of predator and prey was grown in a 250 mL culture flask at 28 °C and 250 rpm and monitored until late exponential phase before use in any experiment. The double layer technique assayed whether Halobacteriovorax sp. PA1 was capable of preying on various bacteria (Table 1). One milliliter of the potential prey bacteria suspension (containing 109 cells/ml) and one milliliter of the appropriate predator dilution was mixed with 3 ml of molten agar (PP20 medium containing 1.1% Difco agar) held at 42 °C, for a final top layer agar concentration of 0.66%. The mixture was immediately spread over the surface of 1.8% agar PP20 Petri dish plates, and three replicates were plated for each predator–prey combination. Plaques were measured after 3–5 days of incubation (Fig. 1C). Prey taxon (accession number) 50% killing rate in liquid media (hours) Predation by Halobacteriovorax? (Double layer assay) V. coralliilyticus BAA 450 (AJ440005) 19.00 Yes V. coralliilyticus RE 98 (CP009617) 24.16 Yes V. coralliilyticus RE22 (PRJNA168268) No observable predation No V. tubiashii ATCC19106 (NZ_AFWI00000000.1) No observable predation No V. tubiashii ATCC19109 13.84 Yes V. fortis PA1 (KT626460) 11.67 Yes V. cholerae N1696 (AE003853) 8.96 Yes V. cholerae S10 (accession) 12.76 Yes DOI: 10.7717/peerj.3315/table-1 Figure 1: Halobacteriovorax predation of Vibrio spp. (A) Micrograph of the pathogen, Vibrio coralliilyticus BAA450 being attacked by Halobacteriovorax and rounded V. coralliilyticus bdelloplast (right) with Halobacteriovorax inside (B) double layer plate showing freshly lysed plaques on a lawn of V. coralliilyticus cells (C) Overnight liquid cultures of (1) a co-culture of Halobacteriovorax and V. fortis, (2) V. fortis and 0.2 µm filtrate from Halobacteriovorax culture, and (3) V. fortis alone. Triplicate biological replicates for each vibrio prey species were grown overnight in marine broth at 28 °C and shaking at 250 rpm, transferred using a 1:100 ratio into fresh MB, and monitored until late exponential phase. Prey were washed three times in 0.2 µm filtered and autoclaved seawater (FSW) and resuspended to a concentration of 109 cells/mL. Three biological replicates of overnight V. fortis and Halobacteriovorax sp. PA1 in FSW, which had lysed and cleared V. fortis prey, were 0.45 µm filtered to isolate predators. Filtered predators were then added to prey species at a 1:100 volume ratio. Predation was measured by OD 600 values using a microplate reader (Infiniti M200; Tecan Group Ltd, Männedorf, Switzerland). Tecan OD values were reported without conversion to a 1-cm path length. Halobacteriovorax in attack phase do not significantly alter the absorbance reading of the prey at 600 nm due to their small cell size. Predation rates in the liquid co-culture assay were measured by the host cell density reduction compared to the predator-free controls. Based on our observed predation rates and the biological relevance of the strain, we chose to conduct our predator–prey addition experiment using V. coralliilyticus BAA 450 (accession # AJ440005).

Collection and preparation of Montastraea cavernosa Montastraea cavernosa was selected as a model for this work as it is both a common reef-building Caribbean coral and is susceptible to a variety of coral diseases (Sutherland & Ritchie, 2002; Goodbody-Gringley, Woollacott & Giribet, 2012). The 33 × 32 cm M. cavernosa colony used in the main experiment was obtained from the Florida Keys National Marine Sanctuary (#FKNMS-2010-123) from the Key West (FL, USA), and was maintained for 10 weeks in a shaded flow through raceway tank at the University of Miami Experimental Hatchery. This single M. cavernosa colony was split into 3.5 cm diameter cores with skeleton trimmed to ∼2 cm. Coral cores were transferred back into a common garden experimental aquaria that only contained the M. cavernosa cores and were allowed to acclimate for an additional four weeks where they demonstrated signs of growth including lateral tissue extension over exposed skeleton and feeding behavior during recovery. Just prior to being subjected to the various experimental treatments as described below, the M. cavernosa cores were then transferred to a second common garden at FIU which was composed of a recirculating seawater tank that only contained the M. cavernosa cores. All coral cores were distributed randomly into new 40L recirculating treatment tanks and only used once in the experiment. Seawater for the experiment was obtained from the University of Miami Experimental Hatchery (sand and UV-filtered seawater pumped in from Biscayne Bay).

Montastraea cavernosa alien and predator additions Bacterial challenges were conducted in sterile beakers with water temperatures held at 31 °C to induce Vibrio coralliilyticus virulence as previously described (Kimes et al., 2012). To further encourage Vibrio coralliilyticus to affect the host and/or microbiome all coral cores (including the controls and Halobacteriovorax challenges) were taken from common garden tank at time zero, scored with a file to mimic tissue damage, and inoculated in the beakers by transferring the sterile media (control), V. coralliilyticus or Halobacteriovorax cells using a sterile q-tip (Adwin Scientific, Schaumburg, IL, USA). The 48 coral cores were divided into 4 treatments for the main experiment, providing 12 cores per treatment. These treatments were challenges of q-tips containing: (1) sterile media as a control, (2) a total of 109 Vibrio coralliilyticus, (3) a total of 106Halobacteriovorax, and (4) a total of 109 V. coralliilyticus and 106 Halobacteriovorax. Three replicate cores (one from each of the 3 replicate treatment tanks were collected at each of 4 time points per treatment (3 replicates × 4 timepoints = 12 per treatment; 12 × 4 treatments = 48 cores total)) (Fig. 2). Once challenged with the treatments, as described above, the fragments were placed in 40 liter tanks with natural seawater but constant recirculation, filtration, and temperature control for the 32 h experimental duration. Figure 2: Montastraea cavernosa microbiome manipulation experimental design detailing collection and inoculation of coral cores, treatment tanks and replication, sample preservation, tissue removal, DNA extraction, and microbiome sample processing. In each experimental challenge, late exponential phase bacteria cultures were pelleted, supernatant removed, and the cells washed three times with sterile artificial seawater (ASW) in 2 mL tubes by centrifugation at 10,000 × g for 10 min and gently re-suspension with sterile media. The late exponential Halobacteriovorax culture was passed through a 0.45 µm filter to remove prey and washed three times under the same condition as the V. coralliilyticus cells. The final cell pellet was resuspended in 100 µl of ASW and transferred using a dual q-tip approach (media control and media control, Vc and media control, Hbv and media control, and Vc and Hbv) to apply cells to freshly abraded corals in sterile beakers (Fig. 2). Q-tips were held on corals for 1 h in the inoculation beakers before transferring the challenged cores to recirculating seawater tanks. Laboratory experiments to quantify the number of cells that remain attached to the q-tip during transfer of the cell pellet were conducted by inoculating 1 ml of media for an hour, removing the q-tip, and performing direct counts of the ASW tubes using epifluorescent microscopy. At each time point (T = 4, 8, 24, and 32 h) one coral fragment from each replicate tank of each treatment (n = 3 per time per treatment) was removed, photographed, placed in a Whirlpak (Nasco, Salida, CA, USA), flash frozen in liquid nitrogen, and transferred to −80 °C freezer for microbial DNA analysis.

Montastraea cavernosa microbiome DNA extraction, sequencing, and quality control From each core one quadrant of the coral tissue layer was removed using a dental tool and transferred into separate microcentrifuge tubes (4 per core) containing 500 µl of TES Buffer (10 mM Tris–HCl [pH 7.5], 1 mM EDTA, and 100 mM NaCl). A 1.5 mL microtube pestle was used to homogenize the tissue before adding 400 µl of TES buffer with lysozyme (Epicentre; final: 10 U µl−1), followed by incubation at 37 °C for 30 min. A 200 µl aliquot of homogenized sample was used for DNA extraction with the Power Soil DNA extraction kit (MoBio Laboratories, Carlsbad, CA, USA); the remainder was stored at −20 °C. Microbial amplicon libraries were generated using 515F and 806R primers to the V4 region of the 16S rRNA gene with Schloss sequencing adapters (Kozich et al., 2013). AccuStart II PCR ToughMix (Gaithersburg, MD, USA) and the following thermocycling conditions were used for amplification:1 cycle of 94 °C for 3 min; 35 cycles of 94 °C for 30 s, 50 °C for 30 s, and 72 °C for 60 s; and 1 cycle of 72 °C for 10 min were used for amplification. Each sample underwent triplicate reactions that were pooled and cleaned using the Promega Wizard SV Gel and PCR Clean-Up System (Madison, WI, USA). The samples were then quantified using a Qubit dsDNA HS kit (Invitrogen, OR, USA) before being pooled in an equimolar ratio. The amplicon purity and length was checked on an Agilent Bioanalyzer 2100 prior to sequencing on a MiSeq Illumina sequencing platform at the Oregon State University’s Center for Genome Research and Biocomputing (CGRB) Core Laboratories. Quality control and selection of operational taxonomic units (OTUs) was performed using QIIME (v.1.8) (Caporaso et al., 2010). Sequences with quality scores less than a mean of 35 were removed. Sequences were clustered into (OTUs) at a 97% 16S rRNA gene identity threshold using USEARCH 6.1.54 (Edgar, 2010) and the subsampled open-reference OTU-picking protocol in QIIME v.1.8 (Rideout et al., 2014), using greengenes 13_8 as the reference (McDonald et al., 2012). Chimeric sequences were removed with QIIME’s wrapper of the UCHIME software (Edgar et al., 2011). Singleton OTUs were removed. The OTUs were assigned taxonomic classification using the QIIME wrapper to the UCLUST software package (Edgar, 2010). OTUs that were classified as chloroplast, eukaryotic or mitochondria were filtered out of the dataset.