Twenty four specimens of soft coral, representing 15 species (Supplementary Tables 1–3), were collected at a depth of 1–3 m near Orpheus Island (Great Barrier Reef, Australia; latitude 18°36.878′S; longitude 146°29.990′E). All specimens except Cespitularia sp. were photographed (Supplementary Figure 1) and sampled underwater, with samples placed directly into plastic bags filled with seawater. Samples were frozen (−80°C) within 1 h of collection and stored until freeze drying. Additional colonies of L. compactum, S. flexibilis, and Pachyclavularia violacea were collected for fraction analysis and isolation of pure compounds (Supplementary Tables 2,3). ¹H NMR spectra of the crude extracts of all soft coral samples were compared between collections and with reference spectra from the Bowden laboratory to ensure consistency in the metabolites and species tested.

Dried soft coral tissue was weighed and homogenized before extraction. Three extracts of different polarity were generated for each soft coral sample. Solvents used for extraction were, in order, dichloromethane (DCM), methanol (MeOH), and water (H₂O). Each extract was the result of three successive applications of solvent. Extracts were concentrated by rotary evaporation before being dried under a stream of nitrogen (N₂) and stored at −20°C until analysis. The DCM and MeOH extracts were dissolved in ethanol and the aqueous extracts in H₂O to a concentration of 20 mg/ml. All soft coral extracts were tested at two concentrations (4 μg/well and 40 μg/well) three times for the presence of QS induction and QS inhibition activity in A. tumefaciens A136 and C. violaceum CV026 in at least two independent experiments.

To further investigate the patterns of QS activity observed in crude extracts, nine soft coral species from five genera (three families) were chosen for further fractionation (Supplementary Table 2). The chosen species displayed five patterns of crude extract activity (Supplementary Table 4). In addition to four species with strong crude extract activities, five species were selected to investigate observed within genera variation in activity patterns or because they represent common genera on the Great Barrier Reef (GBR). Ten fractions of decreasing polarity were generated from the dichloromethane extracts of the chosen soft corals. Extracts were fractionated using flash column chromatography on RP-C18 silica cartridges (Phenomenex Strata C18-E 55 μm 70 Å, 1,000 mg) eluted with a stepwise 20–100% MeOH: H₂O gradient followed by a 1:1 DCM: MeOH wash. The resulting fractions were concentrated to dryness under a stream of N₂ gas and re-dissolved in ethanol for QS screening as described above. ¹H-NMR spectra (600 MHz) of active fractions were recorded with a Bruker 600 Avance spectrometer in deuterated chloroform (CDCl₃).

Ten cembranoid diterpenes with variable secondary ring structures were assessed in this study (Figure 1, Table 1). Isoneocembrene A (1) represents the base cembrene backbone, without an additional ring (Figure 1, Table 1). Compounds 2-4 are ɤ-lactones (5-membered rings). Two δ-lactones (6-membered rings) and one ε-lactone (7-membered ring) are also included in compounds 6-8 respectively (Figure 1, Table 1). The final two compounds are furanocembrenes (Figure 1, Table 1). Pure samples of isoneocembrene A (1), lobolide (4) and sarcophine (5) were acquired from the Bowden Laboratory, Townsville Australia. Isolobophytolide (2), and isolobophytolide monoacetates (3) were isolated from freshly collected and extracted L. compactum (see below). Flexibilide (6), dihydroflexibilide (7) and sinulariolide (8) were isolated from freshly collected and extracted S. flexibilis (see below). The furanocembrenes, pachyclavulariadiol (9) and pachyclavulariadiol diacetate (10), were isolated from freshly collected and extracted P. violacea. Extracts of each species were generated as described previously (section Soft Coral Extract Preparation).

For the L. compactum extract, vacuum liquid chromatography of active crude DCM extract (2 g) was performed over reverse phase C18 silica gel (Phenomenex Luna 10 mm C18 silica gel) and 10 fractions (200 ml) were collected using MeOH: H₂O 0–100% stepwise gradient for each extract. Activity was identified in the 80% MeOH fraction and this fraction was subjected to RP-HPLC 60–100% MeOH gradient over 30 min (Phenomenex Gemini 3 μm NX-C18 110 Å, LC Column 30 × 4.6 mm). Isolobophytolide (2) eluted at 15 min and the two isomers of isolobophytolide monoacetate (3) eluted at 17 min.

For the S. flexibilis extract, flexibilide (6) and dihydroflexibilide (7) were isolated as reported by Kazlauskas et al. (1978). In brief, the DCM extract was subjected to normal phase flash column chromatography with combinations of hexane, DCM and EtOAc. Flexibilide was eluted by 6:1 DCM: EtOAc, 4:1 DCM: EtOAc yielded a mixture of 6 and 7 and 3:1 DCM: EtOAc afforded pure dihydroflexibilide. Sinulariolide (8) was isolated separately by a method adapted from Tursch et al. (1975) by direct crystallization of the DCM extract after dissolution in diethyl ether. Purification of compounds was performed by HPLC as described above.

Due to instability of the furanocembrenes from P. violacea, the pure compounds were generated semi-synthetically as per Bowden et al. (1979). In brief, the DCM extract was prepared at 4°C then partitioned between hexane and 10% aqueous MeOH. After removal of the solvent, the aqueous MeOH fraction was subjected to normal phase flash chromatography with a hexane: EtOAc gradient. All fractions containing (by TLC and ¹H NMR) pachyclavulariadiol, diacetyl pachyclavulariadiol and the two monoacetyl pachyclavulariadiols were combined and hydrolyzed to pachyclavulariadiol (9) by incubation for 24 h at room temperature with MeOH containing 1% (w/v) potassium hydroxide. Methanol was removed under vacuum and the residue was partitioned between diethyl ether and water. The ether fraction was evaporated and the residue was dissolved in hexane. Diacetyl pachyclavulariadiol (10) was acquired by acetylation of half of the obtained pachyclavuriadiol (9). Acetylation was affected by incubation for 24 h with 1:1 acetic anhydride in pyridine before evaporation of the solvent and retrieval via liquid a partition between hexane and water. The semi-synthesis of monoacetyl pachyclavulariadiols was not performed as both 9 and 10 exhibited similar activity, so it was considered unlikely that the activities of monoacetyl pachyclavulariadiols would be different.

Structure and purity of each extracted compound was confirmed by 1D and 2D NMR and comparison with literature values. ¹H-NMR (600 MHz) and ¹³C -NMR (150 MHz) spectra were recorded with a Bruker 600 Avance spectrometer in CDCl₃, with tetramethylsilane (TMS) as internal standard. High resolution mass spectra were collected using an unmodified Bruker BioAPEX 47e mass spectrometer equipped with an Analytica model 103426 (Branford, CT) electrospray ionization (ESI) source. Analytical thin layer chromatography (TLC) was performed on Merck Kieselgel 60. Spots were visualized by UV light or by spraying with a 1% vanillin in acidified ethanol solution. Pure compounds were re-solubilized in ethanol and serially diluted to generate five different concentration solutions (1 × 10² mM to 1 × 10⁻² mM) for each compound.

Surface mucosal layer samples (SML) were collected from three healthy replicate colonies of S. flexibilis and L. compactum from the same depth and location. SML samples were collected underwater from the mid-capitulum region of the coral colony using 50 ml needleless sterile syringes. Samples were maintained at ambient temperatures and processed within 3 h of collection. At the same time as SML samples were retrieved, tissue samples of each replicate were collected and their metabolite ¹H NMR spectral profiles were compared with samples from S. flexibilis and L. compactum collected previously to ensure correct species identification.

SML samples were serially diluted (10⁻², 10⁻³, 10⁻⁴) using autoclaved artificial seawater (Instant Ocean; Spectrum Brands, Madison, WI, USA). One hundred microliters of each dilution were spread plated in triplicate on two types of media commonly used for studies of marine bacteria: 50% Marine Agar (50MA; BD) and Glycerol Artificial Seawater (GASW) agar (Smith and Hayasaka, 1982). Additionally, Thiosulfate Citrate Bile Salts (TCBS; BD) agar, which specifically selects for members of the family Vibrionaceae, was included. L. compactum SML dilutions were additionally plated onto 50MA and GASW agar supplemented with L. compactum's major secondary metabolite, isolobophytolide. All plates were incubated at 28°C and sampled after 48 h, 72 h, 1, and 2 weeks. Representatives of each colony morphotype from each plate were sub-cultured to purity for identification.

Where possible, two representatives of each morphotype were selected for QS activity screening. Where bacteria had initially been isolated using media embedded with isolobophytolide, growth was attempted on the equivalent medium without isolobophytolide. Strains that could not be cultured without isolobophytolide were not included in the screening. Screening was performed on acidified ethyl acetate (EtOAc) extracts of the cell free supernatant of soft coral isolates. These were acquired by transferring single colonies from 50MA plates to liquid culture (10 ml 50% Marine broth culture, 28°C at 170 rpm) and grown to late exponential phase. Cultures were centrifuged for 10 min at 4°C at 10,000 g to obtain the cell free supernatant (CFS). Each CFS was subjected to exhaustive extraction with acidified EtOAc (1% acetic acid) and concentrated to dryness under a stream of N₂ gas. Extracts were then dissolved and diluted to a concentration of 20 mg/l with ethanol.

The biosensor strains A. tumefaciens A136 (Fuqua and Winans, 1996) and C. violaceum CV026 (McClean et al., 1997) were used for detection of QS induction and inhibition in soft coral extracts. A. tumefaciens A136 was grown on ABt media (Clarrk and Maaloe, 1967) and C. violaceum CV026 was grown on Luria Bertani (LB) media (Bertani, 1951). In order to ensure that the QS plasmid was intact and functional, QS biosensor strains were grown in the presence of the appropriate antibiotic (Ravn et al., 2001). A. tumefaciens A136 was grown on media supplemented with 4.5 μg/ml of tetracycline and 50 μg/ml of spectinomycin, whereas, C. violaceum CV026 was grown on media supplemented with 20 μg/ml of kanamycin (Ravn et al., 2001).

The presence of AHL type QS induction activity in soft coral extracts, fractions and pure compounds was detected by performing an agar diffusion assay as described in detail by Ravn et al. (2001). The QS biosensor strain, either A. tumefaciens A136 or C. violaceum CV026, was embedded within the agar and the sample being tested was added to a well cut or formed in the agar. For induction of QS, N-hexanoyl homoserine lactone was used as a positive control and extraction solvents were used as negative controls. Positive results were read as a blue coloration surrounding the wells of A. tumefaciens A136 and a purple coloration surrounding the wells of C. violaceum CV026 (see above and Supplementary Figure 2). The intensity of the response was measured as the diameter of the colored zone and normalized to the response of the positive control.

The agar diffusion assays described above were modified in order to detect QS inhibition. Briefly, as A. tumefaciens A136 and C. violaceum CV026 are not able to QS without the exogenous addition of AHLs, 8.5 μmol n-hexanoyl homoserine lactone was added into the agar embedded with the biosensor strain in order to test for QS inhibition. Two positive controls were chosen based on their previously reported ability to inhibit QS: n-dodecanoyl-DL-homoserine lactone (McClean et al., 1997) and vanillin (Choo et al., 2006). These controls proved effective for both biosensors. The extraction solvents were once again used as negative controls. Positive results in the inhibition assay were read as inhibition of blue or purple coloration of the plates containing A. tumefaciens A136 and C. violaceum CV026, respectively (Supplementary Figure 2). The intensity of the response was measured as the width of the inhibition zone surrounding the well and normalized to the positive control.

To generate dose response curves for pure compounds, agar containing the respective biosensor was poured into custom built molds with 28 preformed wells 4 mm in diameter. After solidification of the agar, 20 μl of sample was added to each well. Pure compounds were only tested with the A. tumefaciens A136 strain as it was the only strain to have both QS induction and inhibition activity.

Genomic DNA of bacterial isolates was extracted using the Promega Wizard Genomic DNA Isolation kit (Promega, Madison WI USA) according to the manufacturer's directions. PCR amplification of 16S rRNA gene fragments was performed using the primers 27F and 1492R (Lane, 1991). The PCR reactions contained the following reagents: 0.4 mM of each primer, 1x MyTAQ buffer (Bioline, Australia), 1.25 U MyTAQ (Bioline, Australia), 1 μL DNA extract (final volume of 50 μL). Cycling conditions were an initial denaturing step of 94°C for 5 min, followed by 30 cycles at 95°C for 1 min, 56°C for 45 s, 72°C for 60 s, and a final elongation step at 72°C for 10 min. PCR products were verified by agarose gel electrophoresis and purified for sequencing using the Qiaquick PCR purification Kit (Qiagen, Valencia, CA) according to company supplied directions. Sanger sequencing was carried out at Macrogen (Seoul, South Korea) using both 27f and 1492R as sequencing primers.

Sequence fragments were assembled using Sequencher (Version 5, Gene Codes, Ann Arbour, USA). For each isolate, the 16S rRNA gene sequence was aligned with sequences in the nr and Ref_Seq database at the NCBI using the megablast tool (RRID:SCR_001598; Altschul et al., 1990) to identify closely related database sequences. Sequences of isolates and database matches were imported into MEGA6 (MEGA Software, RRID:SCR_000667) and aligned using ClustalW (Larkin et al., 2007). A Maximum Likelihood-based phylogenetic tree was constructed using the Maximum Parsimony algorithm for the starting tree, the Tamura-Nei model for nucleotide substitution, and 500 bootstrap replicates (Supplementary Figure 3). The 16S rRNA gene sequences for the 72 bacterial isolates were deposited into the NCBI Genbank database, under accession numbers KM360403-KM360473. Quantification and statistical analysis of CFUs and isolate morphotypes.

Colony forming unit (CFU) concentrations were estimated based on dilutions yielding between 30 and 300 colonies per plate. Differences in the number of CFUs between samples were determined based on three replicates for the corresponding dilution and media type. Statistical differences were determined using the non-parametric Kruskal-Wallis statistic, as the data violated both normality and homogeneity of variances required for ANOVA. Colony morphotype profile analysis was conducted on the variables color, size and texture and were compared using a nonmetric Multidimensional Scaling (nMDS) analysis in the vegan package (version 2.5-1; Oksanen et al., 2018) in R. The nMDS was chosen based on its suitability for spatial representation of complex data sets containing multiple variables, large numbers of zeroes and non-normal distributions (Rabinowitz, 1975). The statistical analyses were performed using Graphpad PRISM (GraphPad Prism, RRID:SCR_015807).

Crude soft coral extracts demonstrated the ability to both induce and inhibit QS in the biosensors tested. While both induction and inhibition of QS were observed for the biosensor A. tumefaciens A136, none of the extracts, regardless of source coral species, polarity, or concentration, were able to induce QS in C. violaceum CV026 (Figure 2A). Inhibition of QS was more prevalent than induction across all polarity extracts (Figure 2A). For both biosensors, the number of active soft coral extracts was reduced at lower extract concentration, with none of the aqueous extracts retaining their activity at the lower dosage (Figure 2B).

Crude extracts of five species induced QS in A. tumefaciens A136; two species from the family Alcyoniidae (L. compactum, Lobophytum sarcophytoides), one from the family Nephtheidae (Nephthea chabroli), one from the family Clavulariidae (P. violacea), and one from the family Xeniidae (Cespitularia sp.) (Figure 3). The highest incidence and strength of induction activity was seen for DCM crude extracts, with the largest haloes of coloration produced by DCM extracts of L. compactum and P. violacea (Figure 3).

Low level QS inhibition of A. tumefaciens A136 was demonstrated for DCM crude extracts of most species, with the only exceptions being L. microlobulatum, Sinularia polydactyla, P. violacea, Clavularia sp., and N. chabroli (Figure 4A). The same trend was seen for MeOH crude extracts, with the additional exception of Sarcophyton sp. 2 (Figure 4A). Inhibition of QS in C. violaceum CV026 was present in DCM crude extracts of all species except P. violacea and Clavularia sp. (Figure 4B). Of the five species that induced QS in A. tumefaciens A136, three (L. compactum, L. sarcophytoides and Cespitularia sp.) also inhibited QS in both biosensors (Figure 4). This contrasted with the other two species capable of QS induction; N. chabroli extracts inhibited QS only in C. violaceum CV026 (Figure 4) while P. violacea demonstrated no QS inhibitory activity with either biosensor (Figure 4).

For all fractionated species, at least two fractions induced QS in A. tumefaciens A136 (Figure 5), regardless of whether their corresponding crude extracts were active (Figure 4). All species produced fractions that induced QS and inhibited QS in at least one biosensor strain. The only exception was P. violacea, which did not inhibit QS in either strain (Figure 5) consistent with crude extract results (Figure 4). The two largest inductive haloes were observed for N. chabroli and L. compactum, which also retained their activity at the higher (1:10) dilution level (Figure 4 The two major bands of induction activity seen in the A. tumefaciens A136 bioassay occurred for fractions eluted at 60% MeOH and at 80–90% MeOH (Figure 5). Many of the latter fractions (80–90% MeOH) also showed activity in the corresponding QS inhibition bioassay (Figure 5). The distinct patterns of QS activity observed for A. tumefaciens A136 contrasted strongly with the broad C. violaceum CV026 inhibition activity (Figure 5). The presence of cembrene diterpenes correlated well with the QS active fractions (data not shown).

The response to pure compounds also depended on the biosensor strain utilized. None of the tested cembranoid diterpenes induced QS in C. violaceum CV026. In contrast, both QS induction and inhibition activity was observed against A. tumefaciens A136 over three to four orders of magnitude, with a loss of activity at higher concentrations (Figure 6). No QS interference was observed for isoneocembrene A (Compound 1; Figure 6). The strongest induction of QS in A. tumefaciens A136 was observed for pachyclavulariadiol and diacetyl pachyclavulariadiol (Compounds 9 and 10; Figure 6). Induction was also observed in isolobophytolide (Compounds 2 and 3), lobolide (Compound 4), and sarcophine (Compound 5). QS inhibition was strongest in the ε-lactone ring of sinulariolide (Figure 6). Peak QS interference for all compounds occurred at approximately 1 × 10⁻⁵ mM (or 3 ppm; Figure 6).

A significantly higher number of colony forming units (CFUs) were estimated for mucus of S. flexibilis as compared to L. compactum (Kruskal-Wallis, H = 7.200, 2 d.f., P = 0.0036; Supplementary Figure 4). Interestingly, when growth media for L. compactum were amended with isolobophytolide, the estimated number of CFUs increased for this species but remained lower than the estimates for S. flexibilis (Supplementary Figure 4). The number and type of colony morphotypes also differed between L. compactum and S. flexibilis (Figure 7). S. flexibilis showed little variation in the morphotype profiles of GASW or 50MA media, forming a tight cluster on the nMDS biplot (Figure 7). In comparison, the morphotype profiles generated from L. compactum showed higher variation in the same culture media. This trend was also consistent when isolobophytolide was added as a selection agent (Figure 7).

In total, 20 bacterial isolates from S. flexibilis were identified through 16S rRNA gene sequencing followed by BLAST searches and construction of phylogenetic trees (Supplementary Table 5, Figures 8, 9). The isolates were dominated by Gammaproteobacteria belonging to the family Vibrionaceae both for the non-Vibrionaceae targeted media (GASW and 50MA) as well as the Vibrionaceae targeted medium (TCBS). Other Gammaproteobacteria included two isolates whose closest relative was “Spongiobacter nickelotolerans” (hereafter referred to as Endozoicomonas, see below); and three Alteromonas-related strains. Finally, one isolate was identified with 99% sequence identity to Bacillus megaterium and Bacillus aryabhattai (phylum Firmicutes).

The potential of soft coral isolates from S. flexibilis to participate in AHL-type QS communication systems was investigated using the same reporter bioassays as used for coral extracts. Quorum sensing activity under the used test conditions was demonstrated for 52.4% of the tested S. flexibilis isolates. Both tested Alteromonas strains exhibited QS induction activity. The Alteromonas SFB10_2 strain triggered QS induction in both sensors strains, whereas, the Alteromonas YSF strain only triggered QS induction in C. violaceum CV026 (Figure 8). Both Endozoicomonas-related strains triggered QS induction in C. violaceum CV026 only. In addition, both strains triggered QS inhibition in A. tumefaciens A136, while QS inhibition in C. violaceum CV026 was only triggered by strain SF102 (Figure 8). We note that out of the 62 bacterial strains screened in this study, the two Endozoicomonas-related strains from S. flexibilis were the only strains with both induction and inhibition QS activity. Of the tested Vibrionaceae strains, 6 of 14 strains showed QS activity and this was evenly split between induction (3 strains) and inhibition (3 strains) activity.

The isolates cultured from L. compactum demonstrated a number of similarities to the bacteria isolated from S. flexibilis (Supplementary Table 5, Figures 10, 11). Firstly, the majority of L. compactum isolates were gammaproteobacteria of the genus Vibrionaceae (30/51). Secondly, strains related to the genera Endozoicomonas and Bacillus, and the order Alteromonadales, were isolated also from this soft coral species. In this instance, however, the diversity of Alteromonadales-related strains was higher with strains related not only to the genus Alteromonas (seven strains) but also to the genera Pseudoalteromonas (six strains), Paramoritella, Ferrimonas, and Shewanella. In contrast to the S. flexibilis isolates, the L. compactum isolates also included strains belonging to the genera Psychrobacter (class Gammaproteobacteria), Erythrobacter (class Alphaprotebacteria), and Micrococcus (class Actinobacteria).

The potential of soft coral isolates from L. compactum to participate in AHL-type QS communication systems was also investigated (Figure 10). Of the tested isolates from L. compactum, 47.5% demonstrated QS activity at the growth conditions tested and activity was mixed between induction and inhibition. Three of the tested Vibrio strains (LC111, LC103, and LC105) showed inhibitory activity against both biosensors. Of the strains that were initially isolated with media containing isolobophytolide, 41% were unable to be cultured in the absence of this compound and consequently were not tested for QS activity.

This study has demonstrated that QS interference extends across at least four soft coral families (Alcyoniidae, Clavulariidae, Nephtheidae, and Xeniidae). Further, it was shown that both induction and inhibition QS activity extends across both polar and non-polar fractions, indicating that QS interference capability is widespread in soft corals from the central Great Barrier Reef, Australia.

Widespread activity, across not only species that are known to contain different metabolite types but also across a range polarities (indicated by the activities of extracts obtained by use of solvents with different polarities), is probably indicative of active compounds of more than one structural type. This is further supported by the finding that most (except P. violacea) of the soft coral species that induced QS in A. tumefaciens A136, also inhibited both biosensors. The widespread prevalence of QS inhibition as well as the presence of QS induction in the soft corals screened here is consistent with QS activity found across a range of marine invertebrates (Taylor et al., 2004; Skindersoe et al., 2008; Hunt et al., 2012). The dual presence of induction and inhibition of QS is similar to that found previously in gorgonian coral extracts (Hunt et al., 2012) but contrasts with the sole QS inhibition activity that was identified in D. pulchra (Kjelleberg et al., 1997). QS induction was also established in extracts of marine sponges and sponge associated bacteria (Taylor et al., 2004). Results from this study highlight the need to examine both induction and inhibition of QS to generate a realistic understanding of the complexity of ecological interactions between a host organism and its associated bacteria.

In contrast with the initial crude extract testing, all the fractionated soft corals displayed at least one active fraction in the A. tumefaciens A136 QS induction assay. This may reflect an inherent increase in concentration of the active components or a decrease in complexity of the samples being tested. Soft coral extracts and fractions may be highly complex mixtures of compounds with contrasting QS regulatory activities. The potential for activity masking within extracts, a phenomenon previously observed in the QS screening of marine sponge extracts (Taylor et al., 2004), is high. This is particularly true if only one concentration or level of complexity is tested.

The QS induction pattern of the soft coral fractions was generally limited to one or two active fractions (80 and 90% methanol elution) for each species suggests that the inductive capability may be due to the presence of structurally similar compounds. Cembranoid diterpenes are well documented in eleven of the soft coral species tested (MarinLit Database, 2013) and correlated well to QS induction therefore could be responsible for the observed QS activity in these species. The same diterpene scaffolds, however, are not commonly known in the genera Nephthea (Amir et al., 2012a,b) Cespitularia (Elshamy et al., 2016) or in gorgonian corals (Changyun et al., 2008), so cembranoid diterpenes cannot fully explain the observed QS interference in these species. Isolation of cembranoid diterpenes was therefore required to understand the relative importance of these secondary metabolites to act as QS mimics.

A similarly discrete pattern was not reflected in the QS inhibition profile for these fractions. The broad QS inhibition profile of these fractions, might be due to multiple compounds, or, the compound(s) responsible may not be suited to the method of fractionation used and the same compounds could be spread across several fractions. The presence of multiple QS compounds within a single holobiont would potentially enable a larger number of interactions with different bacterial strains and / or trigger different QS responses. This complexity may also reflect the capability of some bacteria to possess multiple QS systems (Reshef et al., 2006), with each system regulating a different process or interaction.

The strength and type of QS interference by cembranoid diterpenes was observed to correlate with the size of the oxygenated ring. Those cembranoid diterpenes that contained either a five membered furan or lactone ring were capable of inducing QS in A. tumefaciens A136, whereas the cembranoid diterpenes with larger lactone rings (six or seven membered) were seen to inhibit QS in A. tumefaciens A136. The type of oxygenated functional group also appears to impact the strength these QS mimics, with the furans tested (Compounds 9 and 10) having higher activities than the ɤ lactones (Compounds 2-5). In keeping with our current understanding of the QS mechanism, QS interference was only observed for cembranoid diterpenes possessing secondary oxygen rings (Fuqua et al., 2001; Watson et al., 2002; Geske et al., 2008). The presence of other minor functional groups (epoxides, acetates or level of saturation) had minimal discernible impact on the strength of QS interference observed and no obvious effect on the type of activity observed with respect to A. tumefaciens A136. QS mimics have previously been isolated that possess a ɤ lactone, however, these mimics (such as the furanones of D. pulchra) are often associated with QS inhibition rather than the inductive activity demonstrated here (Givskov et al., 1996; Defoirdt et al., 2013). In the case of the furanones from D. pulchra, bromine substituents are also present and may be influencing the type of activity. The presence of these metabolites in soft corals is strongly correlated to their taxonomy and may represent different strategies of interaction between species.

A common feature of QS mimic compounds previously identified from eukaryotic extracts is multiple forms of biological activity (Davies, 2006; Yim et al., 2007; Defoirdt et al., 2013). Cembranoid diterpenes appear to be no different with a number, including those identified in the current study, having previously been reported to demonstrate antibiotic (Aceret et al., 1995), cytotoxic (Maida et al., 1993) and algacidal properties. The antimicrobial activity identified in flexibilide (7) by Aceret et al. (1995), however, was exhibited at concentrations at least one order of magnitude higher than those that produced QS interference in this study. The peak QS active concentration occurred 1 × 10⁻⁵ mM (or 3 ppm), reflecting the concentrations of flexibilide and sarcophytoxide in the mucous and water column surrounding S. flexibilis and S. crassocaule detected by Coll et al. (1982). Rather than being incompatible, the contrasting activities could be evidence of a hormetic response. Hormetic relationships have been previously observed in the QS mimics from garlic (Persson et al., 2005) and some antibiotic compounds, whereby growth stimulation or cell signaling properties are exhibited at concentrations below their minimum growth inhibitory concentration (Davies, 2006; Yim et al., 2007). A hormetic response could be relevant in soft corals with loosely packed sclerites, where uptake or release of water from the tissue can lead to large changes in volume over a matter of hours (Freckelton, 2015). As a result, associated metabolites will show a correspondingly dramatic change in concentration in the tissues on a volumetric basis over the same time period. More research is required to understand the potential of hormetic relationships in QS mimics and how such concentration changes could be manipulated in the defense of the coral.

Strong evidence for the ecological role of cembranolides and furanocembranoid diterpenes as QS mimics is further exhibited in the strong differences in the ability of the two biosensor strains to respond to QS mimics in the soft corals. The QS inductive compounds present within the soft corals were more readily detected by the A. tumefaciens A136 strain. In contrast, QS inhibition was observed more frequently for C. violaceum CV026. This could suggest that C. violaceum CV026 can be inhibited by a broader range of compounds or that it is more sensitive to a broader range of compound concentrations. This difference in sensitivity is despite an overlap of AHL acyl chain length detection by the two biosensor strains (Steindler and Venturi, 2007). A. tumefaciens A136 utilizes the TraR QS response regulator system and responds to a broad range of acyl chain lengths in AHL molecules (Steindler and Venturi, 2007). C. violaceum responds to a shorter range of acyl chain lengths in AHL molecules and utilizes the CivR QS response regulator (Steindler and Venturi, 2007). The differential responses of these two biosensors highlights the advantage of using multiple biosensors when screening for QS mimics where chain length sensitivities may have little applicability.

This study strongly suggests that isolobopytolide, the major secondary metabolite in L. compactum, is an important selection factor regulating the microbial community of this soft coral. Firstly, we demonstrated that isolobophytolide can interfere with the QS activity of sensor strains, and secondly we demonstrated that addition of isolobophytolide to culture media increased the number and morphological variation of colonies produced from L. compactum. Moreover, the latter result suggests that the inclusion of secondary metabolites in growth media can improve the success of culturing soft coral associated bacterial isolates.

Most isolates generated in this study had high sequence identity with bacterial sequences sourced from the marine environment, including marine invertebrate hosts (Supplementary Table 5). Many of the recovered genera have also previously been isolated from coral mucus samples including Alteromonas, Bacillus, Endozoicomonas, Erythrobacter, Micrococcus, Pseudoalteromonas, Shewanella, and Vibrio (Lampert et al., 2006; Nithyanand and Pandian, 2009; Pootakham et al., 2017). The isolates were dominated by gammaproteobacteria belonging to the family Vibrionaceae, a result that is consistent with previous observations in scleractinian corals (Kvennefors et al., 2010, 2012). For both coral species, several Vibrio strains were isolated whose sequences clustered together and separately from the most closely related database sequences and hence may represent novel species. Scleractinian corals have previously been recognized as harboring a number of novel bacterial taxa (Rohwer et al., 2002; Sunagawa et al., 2010). This situation still remains, with amplicon-based studies of coral microbiomes returning many unassigned OTUs (Blackall et al., 2015). The bacteria of alcyonacean corals are less well studied and it is reasonable to assume that a similar situation could exist.

Multiple strains capable of inducing and/or disrupting QS in the bacterial biosensors were isolated from L. compactum and S. flexibilis (57.5 and 57.8% of tested strains, respectively). The genera of all the bacteria isolated, regardless of activity detected in this study, have previously been reported to possess or interact with QS systems (Ansaldi et al., 2002; Long et al., 2003; Waters and Bassler, 2005; Case et al., 2008; Tait et al., 2009, 2010; Nithya et al., 2010; Albuquerque and Casadevall, 2012; Lade et al., 2014), providing support for the hypothesis that QS is one of the mechanisms regulating coral associated microbial communities.

There was no clear taxonomic pattern of QS activity within the Vibrionaceae, which is consistent with a previous study that assessed QS activity in 29 Vibrionaceae strains (Tait et al., 2010). It is well recognized that QS in Vibrio spp. is tightly regulated by environmental conditions including host-released cues and nutritional status (Waters and Bassler, 2005). Vibrio spp. are ubiquitous in the marine environment (Urakawa and Rivera, 2006), however many Vibrio strains have been implicated in disease either as primary or opportunistic pathogens (Urakawa and Rivera, 2006). Given that QS is involved in the regulation of a number of the genes involved in pathogenicity (de Kievit and Iglewski, 2000; LaSarre and Federle, 2013), the presence of a wide range of Vibrio spp. with QS capabilities in otherwise healthy corals warrants further investigation to elucidate which genes are under QS control in these species.

“Spongiobacter,” now recognized as belonging to the genus Endozoicomonas (Neave et al., 2016), was originally recovered from a marine sponge (Pike et al., 2013) but is also present in many gorgonian (Sunagawa et al., 2010; La Rivière et al., 2013) and scleractinian corals (Raina et al., 2009; Blackall et al., 2015; Bourne et al., 2016). “Spongiobacter” strains have been attributed a number of ecological roles; “Spongiobacter” strains from A. millepora demonstrated a dependence on DMSP and consequently a role in the biogeochemical sulfur cycle was postulated (Raina et al., 2009), whereas, “Spongiobacter” strains from the sponge Suberites carnosus demonstrated antibacterial activity (Flemer et al., 2012). Of greatest interest to this study is the QS activity detected in “Spongiobacter” strains from the sponges Mycale laxissima and Ircinia strobilina (Mohamed et al., 2008). The Endozoiocomonas-related strains SF102 and SF204 from S. flexibilis that were tested in this study induced QS activity in C. violaceum CV026 and not in A. tumefaciens A136, whereas Mohamed and coworkers found the opposite response (positive in A. tumefaciens and negative in C. violaceum).

In this study, QS activities were assessed for the culturable fraction of bacteria associated with the mucus of two soft coral species. In future, functional gene analysis and gene expression analysis may allow a more complete assessment of the genes that are responsible for and regulated by QS in these bacteria. Moreover, new –omics techniques will allow investigations of quorum sensing genes and their expression also in bacteria that cannot easily be cultured with standard methods. A combination of culture-independent studies and manipulative experiments using isolates holds great promise for further elucidation of QS mechanisms in soft coral holobiomes.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.