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Volcanic ash supports a diverse bacterial community in a marine mesocosm

Volcanic ash supports a diverse bacterial community in a marine mesocosm Sciences, Ludwig-Maximilians-Universität, Munich, Germany Shallow- water coral reef ecosystems, particularly those already impaired by anthropo- United States Geological Survey, Menlo Park, genic pressures, may be highly sensitive to disturbances from natural catastrophic CA, USA events, such as volcanic eruptions. Explosive volcanic eruptions expel large quantities 3 LMU GeoBio-Center , Munich, Germany 4 of silicate ash particles into the atmosphere, which can disperse across millions of SNSB-Bavarian State Collections of Palaeontology und Geology, Munich, Germany square kilometres and deposit into coral reef ecosystems. Following heavy ash deposi- MWM-Museum Witt München, Munich, tion, mass mortality of reef biota is expected, but little is known about the recovery of Germany post- burial reef ecosystems. Reef regeneration depends partly upon the capacity of Correspondence the ash deposit to be colonised by waterborne bacterial communities and may be in- V. Witt and C. Cimarelli, Department of fluenced to an unknown extent by the physiochemical properties of the ash substrate Earth and Environmental Sciences, Ludwig- Maximilians-Universität, Munich, Germany. itself. To determine the potential for volcanic ash to support pioneer bacterial coloni- Emails: wittverena@gmx.de and cimarelli@ sation, we exposed five well- characterised volcanic and coral reef substrates to a ma- min.uni-muenchen.de rine aquarium under low light conditions for 3 months: volcanic ash, synthetic volcanic glass, carbonate reef sand, calcite sand and quartz sand. Multivariate statistical analy- sis of Automated Ribosomal Intergenic Spacer Analysis (ARISA) fingerprinting data demonstrates clear segregation of volcanic substrates from the quartz and coral reef substrates over 3 months of bacterial colonisation. Overall bacterial diversity showed shared and substrate- specific bacterial communities; however, the volcanic ash sub- strate supported the most diverse bacterial community. These data suggest a signifi- cant influence of substrate properties (composition, granulometry and colour) on bacterial settlement. Our findings provide first insights into physicochemical controls on pioneer bacterial colonisation of volcanic ash and highlight the potential for vol- canic ash deposits to support bacterial diversity in the aftermath of reef burial, on timescales that could permit cascading effects on larval settlement. 1  |  INTR ODUCTION (Wilkinson 1999). The deterioration of water quality in coastal regions consequently favours macro- algal dominance (Fabricius, 2005; Coral reefs are unique, biodiverse ecosystems of high socio- economic Schaffelke, Mellors, & Duke, 2005) and increases the risk of disease importance on both global and local scales (Nicholls et al., 2007). for coral reef- building species, including sponges and corals (Haapkyla Anthropogenic disturbances, such as sedimentation and eutrophi- et al., 2011; Webster, Xavier, Freckelton, Motti, & Cobb, 2008), which cation, increasingly pressure fragile coral reef ecosystems worldwide further exacerbates coral reef vulnerability to catastrophic natural This is an open access article under the terms of the Creative Commons Attribution NonCommercial License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited and is not used for commercial purposes. © 2017 The Authors. Geobiology Published by John Wiley & Sons Ltd. Geobiology. 2017;15:453–463. wileyonlinelibrary.com/journal/gbi     453 WITT eT al . 454       disturbances, such as volcanic ash deposition (Vroom & Zgliczynski, Hoffmann, 2007; Munn, 2003), while others (e.g., Al, Cu) may be toxic 2011). After an explosive volcanic eruption, widespread dispersal and (Duggen et al., 2007b). deposition of volcanic ash over areas up to millions of square kilome- Investigating the propensity for volcanic ash to promote pioneer tres, in thickness of up to several centimetres, may be damaging to ash- bacterial colonisation in situ is hampered by the difficulties associated affected coral reef ecosystems; both Maniwavie, Rewald, Aitsi, Wagner, with substrate accessibility, geographic location and the dangers asso- and Munday (2001) and Vroom and Zgliczynski (2011) have reported ciated with sampling near active volcanoes. Laboratory experiments the destruction and mass mortality of reef biota following heavy ash conducted in simulated tropical coral reef aquaria, therefore, offer deposition. However, the capacity of reef ecosystems to recover after a viable method to approximate the properties of volcanic ash that burial by ash remains uncertain. Maniwavie et al. (2001) reported that may dictate its capacity to act as a colonisable substrate. In this study, 2 years after burial by volcanic ash corals had only re- colonised the sur- we incubate a selection of volcanogenic, terrigenic and biogenic sub- faces of protruding or unburied objects (e.g., boulders, tree stumps), strates in a coral reef- like marine aquarium system for 3 months and while the ash substrate itself remained barren; in contrast, Schils (2012) correlate differences in the bacterial colonising consortia with physi- noted that a period of frequent ash deposition into a tropical reef eco- cochemical properties (i.e., chemical composition, mineralogy, granu- system promoted a change in benthic microbial and macrofloral com- lometry, morphology, colour) of particulate substrates. munities on a similar timescale. These varying responses indicate a clear need to better understand the factors that may dictate the recovery of 2  |  MATERIALS AND METHODS vulnerable and valuable coral reef ecosystems after ash deposition. In the aftermath of large- scale burial, recovery of the reef eco- 2.1 | Substrate selection, preparation and system may depend on pioneer colonisation of the new substrate by characterisation free- living bacteria from the water column. After attachment to the surface, these bacteria produce an extracellular polymeric matrix The effect of physicochemical parameters on the composition of bac- that embeds further microbial organisms, forming so- called biofilms terial community colonisation was explored using five different sub- (Costerton, Lewandowski, Caldwell, Korber, & Lappin- Scott, 1995). strates in a marine aquaria under controlled conditions for bacterial Biofilm communities are highly abundant in coral reefs and are crucial colonisation: two volcanogenic materials (volcanic ash and synthetic in biogeochemical nutrient cycling and the degradation of anthropo- volcanic glass), two biogenic materials of reef origin (carbonate coral genic pollutants (reviewed in Davey & O’Toole, 2004). Further, they reef sand and calcite sand) and a terrigenic quartz sand (Table 1). Fresh, provide an essential settlement surface for larvae of important reef- crystal- bearing volcanic ash was obtained from Sakurajima volcano, building invertebrates (e.g., corals and sponges) and influence larval Japan, on 18 July 2013. Sakurajima was selected because it prevalently settlement cues and metamorphosis (Webster et al., 2004; Wieczorek erupts andesitic magma, which is of intermediate volcanic composi- & Todd, 1998). Accordingly, any changes in bacterial biofilm communi- tion and represents the dominant composition at island arc volcanoes, ties may influence invertebrate larval settlement, coral reef establish- and because the physicochemical properties of its eruptive products ment and further development. Therefore, the capacity of a volcanic are well characterised (Hillman et al., 2012). The synthetic volcanic ash substrate to support bacterial settlement, particularly compared to glass substrate was employed to provide a crystal- free system of com- the marine substrates it overlies, may play a crucial role in shaping the parable chemical composition to the volcanic ash substrate and was recovery of ash- affected reef ecosystems. prepared by subjecting the Sakurajima ash to five cycles of high tem- Previous studies on aquatic biofilm formation using an array of perature melting in a platinum crucible at 1500°C to produce a homo- natural and artificial substrates, including basaltic glasses and boro- geneous melt. This melt was quenched rapidly to produce a glass and silicate (Thorseth, Furnes, & Tumyr, 1995), biotite (Ward, 2013), gran- then mechanically crushed. The terrigenic quartz sand was selected to ite (Chung et al., 2010), coral skeletons and clay tiles (Witt, Wild, & provide a silicate reference material for comparison with the volcano- Uthicke, 2011), highlight the importance of physicochemical properties genic substrates and was obtained commercially (Sigma- Aldrich ID no. (e.g., granulometry, surface morphology, mineralogy and chemistry, 274739). To draw comparisons between the silicate substrates and bio- colour) in promoting initial substrate colonisation. Crucially, volcanic genic reef sands, we utilised two different carbonate materials, each ash materials are subject to a wide variation in all of these proper- with different origins. The coral reef sand was obtained from a shallow ties, which are the product of magma composition and eruption his- fringing reef in the north- eastern Gulf of Aqaba, Red Sea, located within tory (Dingwell, Lavallée, & Kueppers, 2012). Ash particles range in size a marine reserve close to the Marine Science Station in Aqaba, Jordan from the millimetre to submicron scale and vary in morphology from (29°27′N, 34°58′E), and was mechanically crushed before use. The cal- smooth, blocky particles, to rough- textured vesicular clasts (Heiken, cite sand was sourced from Longcliffe Quarries Ltd. (Brassington, UK). 1974). They commonly contain crystalline and amorphous silicates of The two volcanic substrates were subjected to a preliminary leach- various compositions, and range in colour from light to dark (Ayris & ing period of four hours in deionised water at an ash:water mass ratio Delmelle, 2012). Ash surfaces can be a source of variably extractable of 1:5 and dried at 105°C for six hours. The remaining three substrates elements, some of which (e.g., Al, Ca, Co, Cu, Fe, K, Mg, Mn, Ni, Mo, were washed in deionised water to remove very fine particulate P, S, Zn; Jones & Gislason, 2008) may be important macro- or micro- adhering to the surface of bigger clasts, and dried at 105°C for six nutrients for bacteria and phytoplankton (Duggen, Croot, Schacht, & hours. All substrates were then leached for 12 hr in sea water taken WITT eT al .       455 TABLE 1 Physicochemical data of the five substrates Carbonate reef a b b c Substrate Volcanic Glass Volcanic Ash sand Calcite Quartz Chemical composition (wt. %) SiO 58.2 58.5 0.2 0.9 >99.5 Al O 16.8 16 0.1 0.3 n.s. 2 3 Fe O 7.9 7.9 0 0 n.s. 2 3 MnO 0.2 0.2 0 0 n.s. MgO 3.4 3.6 1.3 0.4 n.s. CaO 7.4 7.2 52.2 54.4 n.s. Na O 3.4 3 0.3 0.1 n.s. K O 1.4 1.4 0 0 n.s. TiO 0.8 0.8 0 0 n.s. P O 0.2 0.2 0.1 0 n.s. 2 5 Total 99.6 100.1 99.0 100.0 >99.5 L.O.I −0.17 0.02 44.49 44.43 <0.5 Physical properties SSA (m /g) 0.02 0.1 0.49 0.22 0.11 Median diameter (μm) 234 221 224 230 263 Interquartile diameter 180–260 190–260 190–270 190–260 220–310 (μm) Colour Dark Dark White White White Morphology Angular Subangular Aggregate Aggregate Subangular Surface appearance Smooth Rough Rough Rough Rough Comprised of glass, plagioclase, pyroxenes, Fe–Ti oxides (Miwa et al., 2013). Comprised of CaCO . Compositional data of pure quartz from Sigma–Aldrich product specifications (n.s. not stated). from the aquarium system at a 1:5 ash:water mass ratio and dried at Brunauer–Emmett–Teller (BET) theory to argon adsorption measure- 105°C for six hours, spread out in large Petri dishes, UV- sterilised by ments conducted at −196°C using a Micrometrics Gemini surface area 2 × 30- min cycles and stirred in between the cycles. These treatments analyser. Samples were dried overnight at 105°C overnight prior to were intended to isolate the effect of the substrate by eliminating any analyses. Data are the mean of three repeated measurements. Particle potential contribution from mechanical crushing of the glass, calcite size distributions were determined by laser particle analysis using a and quartz substrates, any adsorbed chemical species (Ayris et al., Beckman- Coulter LS230. The particle morphology of the leached sub- 2014) that may be toxic to the other organisms in the aquarium water strates was examined by scanning electron microscopy (SEM; Leo 1430 system, and any pre- existing contamination by bacteria prior to or VP) with a maximum operating voltage of 15 kV. Samples were prepared between sample collection/synthesis. The absence of bacteria on the for imaging on SEM stubs and sputter coated with gold. Representative samples was confirmed at the onset of the experiment using the DNA images of single particles were taken for each sample to qualitatively fingerprinting tool of Automated Ribosomal Intergenic Spacer Analysis document differing particle morphologies amongst the substrates. (ARISA) (see details in Methods section below). As this study inten- tionally targets the physicochemical properties of substrates, the pre- 2.2 | Design of the marine colonisation experiment liminary leaching of volcanic ash was intended to remove any soluble salts emplaced on ash surfaces during transport through the volcanic The aquarium system consists of a main tank (330 L) containing a eruption plume. While the relevance of surface salts as a readily avail- 7- year established coral reef community, including corals, algae, able nutrient source in ocean surface waters has been previously con- diverse marine invertebrates and fish. The main tank supplies sea sidered (e.g., Duggen et al., 2007b), their capacity for rapid dissolution water to a series of 80- L subtanks. Those subtanks were used for on first contact with sea water likely limits their relevance to any sub- experiments and did not contain coral reef organisms. The subtank sequent influence once deposited as a substrate on the ocean floor. utilised in this study was configured for a 12:12- hr light:dark cycle and Bulk chemical compositions of the substrates were determined by was covered by shade cloth to ensure low light availability of <0.2 klux. X- ray fluorescence spectroscopy (Philips, MagiX Pro). The major and Light availability and temperature were recorded at 30- min intervals minor elements were measured using glass beads prepared by fusion over the course of the experiment using loggers (Onset HOBO TidBit). with a lithium borate flux in a Panalytical Eagon 2 furnace fusion Tank sea water was maintained at constant flow rate, as to not perturb system. Specific surface area was determined by application of the the substrates in the dishes (see below), constant average temperature WITT eT al . 456       (25.4°C ± 0.05), pH (8.13 ± 0.05) and conductivity (51.8 ± 1.14 mS), structure amongst different substrates. DNA was extracted from and was tested weekly to ensure that nutrient concentrations 0.25 g of each of the collected subsamples (three replicates and two remained low (<8 nmol/L NO , <0.2 nmol/L PO ; see Table S1). separate extractions from each) and water samples in separate tubes. 3 4 The duration of the experiment was 3 months, with sampling of Extraction utilised a PowerLyzer DNA Isolation Kit according to the all substrates after one (T1), two (T2) and three (T3) months. For each manufacturer’s instructions (MoBio, Carlsbad, CA, USA) with the fol- sampling time- point, each substrate was represented by three separate lowing alterations: bead beating cycles 2 × 30 s for higher yields of Petri dishes. Sterile glass Petri dishes of 50 mm diameter and 10 mm bacterial DNA and elution in 2 × 50 μL 1 × TE buffer to lower the pos- depth were filled with 1 g of substrate (a uniform depth of ~5 mm) and sibility of disturbing polymerase chain reaction (PCR) amplifications. positioned approximately 50 mm apart at the bottom of one of the 80- L DNA from all samples was stored in aliquots at −20°C until fur- subtanks, immersed to a depth of 250 mm in sea water. The granulom- ther processing. Extracted DNA from all replicates (three per sub- etry of the substrate was selected to minimise the possibility of any strate and time- point) was amplified by PCR using the ITSF-F AM disturbance by the water flow. To prevent loss of material during the and ITSReub primer pair (Cardinale et al., 2004) in three separate initial sample placement, the dishes were carefully filled with sea water reactions per replicate sample. The ARISA- PCR mixture (25 μL) con- via manual pipetting and loaded into the tank with their lids in place. tained 5 × PCR buffer (Promega, Madison, WI, USA); 2.5 mm MgCl After placement, the lids were removed. In total, 45 dishes were utilised (Promega); 0.25 mm deoxynucleoside triphosphate mix (Promega); in the experiment (5 substrates × 3 sampling points × 3 replicates). At 1.5 μg/μL bovine serum albumin (BSA); 400 nm universal primer each sampling time- point, 2 × 0.3 g of substrate was recovered from ITSF- FAM (5′- GTCGTAACAAGGTAGCCGTA- 3′), 5′- labelled with the each replicate dish using a sterilised spatula. DNA from the substrate phosphoramidite dye FAM and eubacterial ITSReub (5′- GCCAAGGC samples was either extracted immediately, or the sampled substrates ATCCACC- 3′); and 0.025 U of GoTaq polymerase (Promega). To each were directly snap- frozen in liquid nitrogen and stored at −80°C and reaction, approximately 20 ng of extracted DNA (quantified by a then extracted within 2 weeks after sampling. To ensure that observed ND- 1000 Nanodrop; Peqlab Biotechnology, Erlangen, Germany) was bacterial differences amongst substrates and time- points were not due added and the volume was adjusted to 25 μL using PCR water. Thermal to changes in water quality parameters (i.e., temperature, pH, conduc- cycling was carried out in an Eppendorf MasterCycler (Eppendorf, tivity, nutrient concentrations), 2 × 500 mL seawater samples were col- Hamburg, Germany) with an initial denaturation at 94°C for 3 min, lected at the time of substrate immersion (T0 beginning of the experi- followed by 30 cycles of 94°C for 45 s, 55°C for 45 s, 72°C for 90 s, ment) and at each sampling time- point (n = 2), filtered through 0.2 μm with a final extension at 72°C for 5 min and then cooled to 15°C. polycarbonate filters using a vacuum pump, and filters were stored and PCR products were purified utilising the Nucleospin PCR clean- up kit processed following the same protocols as the substrate samples. (Macherey- Nagel, Düren, Germany). Diluted PCR products (300–1200 bp) were prepared for analysis by ARISA using capillary electrophoresis as follows. A standardised 2.3 | Determination of differences in bacterial amount of DNA (100 ng) was added to a separation cocktail contain- community structures ing 0.5 μL of internal GeneScan 1200LIZ size standard (50–1000 bp) Differences in bacterial community structure amongst substrates and (Applied Biosystems, Foster City, CA, USA) and 14 μL of deionised time- points were determined by the DNA fingerprinting method of Hi- Di formamide. The preparation was denatured for 3 min at 95°C ARISA using capillary electrophoresis. The technique targets the ITS and kept on ice for at least 5 min before being further processed by region (Intergenic Transcribed Spacer) between the 16S and the 23S the sequencer. Separation of the PCR- amplified fragments via capil- rRNA gene regions, which is highly variable in nucleotide sequence lary electrophoresis was carried out on an 80 cm- capillary ABI PRISM and length (from 50 bp to 1500 bp). Different bacterial species exhibit 3730xl genetic analyzer (Applied Biosystems) with the following run specific nucleotide lengths relative to the ITS region and, hence, the parameters: 14.6 kV run voltage, 2.4 kV injection voltage, 20- s injec- bacterial community structure (community composition and relative tion time, and 60°C oven temperature. Raw profiles were checked abundance of dominant bacterial species) can be determined (Brown for stable baselines and voltage, and peak sizes and absolute areas & Fuhrman, 2005). While next- generation sequencing (NGS) enables were then determined using the GeneMapper software v4.0 (Applied more in- depth studies, Gobet, Boetius, and Ramette (2014) emphasise Biosystems) with minimum peak heights of 50 fluorescence units for that the DNA fingerprinting of ARISA retains value in determining dif- all dyes. The best- fit size- calling curves were built using a second- ferences in community structure. It is particularly beneficial for the order least- squares method (compensating for anomalously running current study, as it permits the identification of fundamental differ- fragments in the standard) and the Local Southern method as the size- ences in bacterial community composition amongst many different calling algorithm. A perfect fit for the calibration curves in the range samples. DNA fingerprinting is a cost- effective and rapid approach of 100–1000 bp was always checked before further processing the to identify taxonomic affiliations or diversity estimates and to sepa- samples. The fragments/peaks matrix generated with GeneMapper rate samples on local spatio- temporal scales, and correlate environ- was transferred into the T- align software (Smith et al., 2005). A differ- mental variables with bacterial community structure (van Dorst et al., ence of 1.5 bp between peaks was defined as a single operational tax- 2014), prior to embarking on more cost- intensive NGS campaigns. onomic unit (OTU) (Danovaro, Luna, Dell’Anno, & Pietrangeli, 2006). Hence, this tool was chosen to gain an initial overview of community The values of the relative fluorescence intensity of the peak area of the WITT eT al .       457 ARISA fingerprinting data were normalised, square- root transformed composed of micron- sized fragments of carbonate biogenic mate- and standardised prior to statistical analysis. rial (mainly shell and exoskeleton fragments of marine organisms) cemented by crystalline calcite. Calcite clasts are similarly blocky and equant and appear to consist of aggregates of micron- sized calcite 2.4 | Statistical analysis minerals. The specific surface area varies between 0.02 and 0.5 m /g, To explore the variation in the community structure and to determine with the highest surface area materials being the biogenic substrates 2 2 whether bacterial assemblages grouped by substrate, multivariate sta- (carbonate reef sand = 0.5 m /g, calcite = 0.2 m /g; Table 1). tistics were applied. Pairwise distance matrices were calculated from the relative abundance data (ARISA) using the Bray–Curtis dissimilar- 3.2 | Marine colonisation experiment ity index. Non- metric multidimensional scaling (nMDS) was applied to the distance matrices to explore the variation in community struc- Multivariate statistical analysis of the ARISA fingerprinting data of the ture. Differences in the bacterial community structure according to ITS region of the biofilm communities established upon the five sub- substrate were assessed for significance by applying the ANalysis Of strates repeatedly showed overall significantly different bacterial com- SIMilarity (ANOSIM) test based on permutation (9999 permutations) munity compositions over the 3- month period and was consistently procedures using the Bray–Curtis distance measure. The contribution distinct from the community within the water column, represented by of each single taxon to the overall dissimilarities of factors was deter- replicates from the beginning (T0) and additional data points as time mined using the SIMilarity PERcentage (SIMPER) routine. progressed. These differences are reflected by the variations in the total To visualise the community structure and relationships between number of OTUs and in the number of unique OTUs for each substrate substrate- specific and shared operational taxonomic units (OTUs) at each time- point. We summarise these trends in the following para- amongst the different substrates, a Venn diagram was constructed. graphs but, for brevity, we further interpret the data for month three To determine differences in the number of OTUs between substrate only, as this represents the most well- established bacterial community. groups and the water column, one- way Analysis of Variance (ANOVA) Total OTUs successively increased by over 2.5- fold from the first to was applied. Homogeneity of variance was tested using Levene’s test, the third month of the experiment (from 108 OTUs at T1 to 273 OTUs and the Tukey–Kramer test was used as a post hoc test. To determine at T3). At each time- point, most OTUs were shared amongst the sub- biodiversity differences from the fingerprinting data, the Shannon– strate communities, with few substrate- specific ones. Both shared and Wiener (H′) and Simpson’s (1- D) indices for bacterial OTUs in each substrate- specific OTUs increased with time from 99 shared and 10 substrate (average value for 6 replicates) were calculated. To test for substrate- specific OTUs at T1 to 259 and 13 substrate- specific OTUs significant differences between the bacterial communities on the dif- at T3. Amongst the five-substrates bacterial communities, volcanic ash ferent substrates, the Shannon index values (n = 6 per substrate) for showed the most substrate- specific OTUs, followed by glass, carbon- bacterial communities on the five substrates after 3 months of incu- ate and calcite, with quartz exhibiting no substrate- specific OTUs at all. bation were analysed using the t- test. All statistical analyses were per- The grouping patterns visualised in nMDS (Figure 2) showed sig- formed using the statistical program PAST (Hammer, Harper, & Paul, nificant differences amongst the bacterial communities on the five 2001). The level of significance was <0.05 for all statistical tests. substrates at T1 (ANOSIM R = .272, P = .0001), T2 (ANOSIM R = .557, P = .0002) and T3 (ANOSIM R = .507, P = .0001), apart from quartz, which was different to the volcanic substrates but very similar to the 3  |  RESULT S biogenic substrates carbonate and calcite at T2 (ANOSIM R = .191, P = .273) and T3 (ANOSIM R = .033, P = .065 and R = .209, P = .303, 3.1 | Particle substrate characterisation respectively, Table S2). At T1 and T2, Quartz did not exhibit any spe- Bulk chemical analysis by XRF confirms the volcanic ash, glass and cific bacterial community. The overall community for all five substrates quartz as silicates and the reef sand and calcite substrate as calcium showed distinct groupings of bacterial communities associated to the carbonates (Table 1). The volcanic ash and glass are both dark in col- substrates of volcanic origin and coral reef origin, while quartz and the our, whereas the quartz, reef sand and calcite are all white. Particle carbonate substrates both share bacterial OTUs. SIMPER analysis at T1 size data showed mono- modal Gaussian distributions for all tested revealed that the OTUs contributing the most to the community differ- substrates. Ash, glass, calcite and reef sand had median particle diam- ences were 545 bp (5%), 890 bp (4%) and 620 bp (4%). Of these, the eters of 220–234 μm, while the quartz sand had a median diameter of 545- and 890- bp OTUs were exclusive to, and dominant within, the 263 μm (Table 1). Particle morphology of the substrates was investi- volcanogenic substrate bacterial community, whereas 620- bp OTU was gated by SEM analyses (Figure 1). Ash comprised subangular blocky dominant in the volcanogenic substrates, occurred occasionally in the particles with rough surfaces, often with micron- sized vesicles. Glass biogenic substrates and was absent in the quartz substrate. In addition generated from ash melting and crushing shows smooth surfaces with to the groupings according to substrate, the bacterial assemblages on sharp edges (angular clasts) and conchoidal fractures imparted during each substrate differed significantly from those detected in the water preparation. Quartz particles display similar morphological character- column (ANOSIM R = .469, P = .0002). In the water column, 112 OTUs istics to the volcanic ash substrates, being blocky, subangular parti- were detected, of which 2 were specific to the water community. cles with rough surfaces. Reef sand particles are blocky aggregates, SIMPER analysis at T2 identified the fragments 530 bp (6%), 890 bp WITT eT al . 458       (a) (b) (c) (d) (e) FIGURE 1 Scanning electron microscopy (SEM) images of substrates: (a) volcanic ash, (b) synthetic volcanic glass, (c) quartz, (d) carbonate reef sand and (e) calcite. Images were collected at 15 kV (4%) and 721 bp (3%) contributing most to the overall community dif- as the bacterial assemblages on each substrate differed significantly ferences. We determined that the 530- bp fragment represents a domi- from those detected in the water column (ANOSIM R = .241, P = .049; nant OTU present in all substrates. Substrate- associated bacterial com- Table S2). SIMPER analysis revealed an overall dissimilarity pooled over munities differed significantly from those in the water column (ANOSIM all groups of 52.37% with the OTUs contributing most to community R = .294, P = .0002). In the bacterial community in the water column, structure differences being the 332- bp (4.6%), 422- bp (1.7%) and 481- 119 OTUs were detected, of which 3 OTUs were water- specific. bp (1.5%) fragments, but with unspecified taxonomic affiliations. These After 3 months (T3), a total of 273 OTUs could be detected in the OTUs were detected amongst all substrates, but were all higher in the overall bacterial community from the ARISA data. Of these, 272 OTUs volcanogenic than in the biogenic and terrigenic substrates; 481- bp were detected on the substrates, with 13 substrate- specific OTUs and OTU was a dominant OTU throughout all substrates. 259 shared ones (Figure 3). Of the shared OTUs, 144 were shared All five substrates and the water column varied significantly amongst all five substrates. The volcanic ash substrate showed the larg- from each other with respect to the number of associated OTUs at est bacterial community with the most substrate- specific OTUs (7), fol- the 3- month time- point (one-w ay ANOVA, degrees of freedom = 5, lowed by volcanic glass (3), while a further 30 shared OTUs were specific mean square = 13 844, F = 1044, P = <.0001, Tukey–Kramer post to the volcanogenic substrates only. The biogenic substrates showed hoc test P = .0001; Table S3). The volcanic substrates supported the very low substrate specificity with <2 substrate- specific OTUs each (reef highest number of total OTUs and, according to Shannon–Wiener and sand 1 OTU, calcite 2 OTUs). The bacterial community in the water col- Simpson’s diversity indices, also harboured the most diverse bacterial umn had a total of 127 OTUs, of which solely 1 OTU was water- specific. community compared to the quartz and biogenic substrates (Table in Multivariate statistical analysis of the ARISA fingerprinting data of Figure 2). Further, statistical t- tests confirmed significant differences in the ITS region of the biofilm communities established upon the five diversity of bacterial OTUs between the volcanogenic substrates and substrates after 3 months showed distinct groupings of bacterial com- the biogenic and quartz substrates, while calcite, carbonate reef sand munities associated with the substrates of volcanic origin and coral and quartz were not significantly different from each other (Table S4). reef origin (Figure 2), and of water. The two biogenic substrates had a higher variability amongst replicates than those of volcanic origin, but the greatest variability was detected amongst quartz substrate replicate 4  |  DISCUSSION samples. Bacterial communities on all substrates were significantly dif- ferent to each other (ANOSIM R = .507, P = .0001), apart from quartz, No previous in situ studies have investigated bacterial colonisation of which was different to the volcanic substrates but the same as the bio- volcanic ash as a substrate within a reef environment, and the only genic substrates (ANOSIM R = .209, P = .303 and R = .033, P = .065, in situ studies documenting the aftermath of ash deposition are both respectively, Table S2). Substrate specificity was further confirmed focused on macroflora and fauna and are contrasting in their results: WITT eT al .       459 FIGURE 2 Non- metric multidimensional scaling (nMDS) ordination (Bray–Curtis distance) of ARISA- derived bacterial community profiles at months 1, 2 and 3, with six replicates per substrate per time- point. Groupings are specified according to substrate. Data from an initial analysis (T0) of the bacterial community in the water column (n = 2) are plotted alongside the monthly water data. The water groupings are of cumulative data at each time- point, inclusive of T0, for reference. Points plotting close to each other show a more similar community than distant ones [Colour figure can be viewed at wileyonlinelibrary.com] FIGURE 3 Relationship between the substrate- specific and shared operational taxonomic units (OTUs) amongst all five substrates after 3 months. The diagram was constructed using the program VENN (http://bioinformatics.psb.ugent.be/webtools/Vennwileyonlinelibrary.com]). Table (left) shows the total number of OTUs per substrate after 3 months (T3), the Shannon–Wiener (H′) and Simpson’s (1- D) diversity indices. The values were calculated by taking the average of OTUs of the ARISA data of the replicate samples of each substrate (n = 6) [Colour figure can be viewed at wileyonlinelibrary.com] Maniwavie et al. (2001) implied that recolonisation of ash substrates community composition and settlement success rates increase with by reef flora did not occur, whereas Schils (2012) reported a sudden the age of the biofilm, and the timescale over which significant dif- change in benthic microbial and macrofloral communities of an ash- ferences in bacterial community structure were observed in the cur- affected reef. Therefore, while comparisons between the aquarium rent study are similar, and relevant, to the timescales required for the experiment of the current study and in situ reef settings should be settlement of invertebrate larva (days to weeks; Bao, Satuito, Yang, treated with caution, our study makes an important contribution to & Kitamura, 2007; Campbell et al., 2011). Consequently, the signifi- a highly uncertain subject by demonstrating that bacteria can colo- cant differences in pioneer bacterial colonisation between volcano- nise ash substrates and can establish a significantly different commu- genic substrates, and the terrigenic and biogenic substrates in the nity structure than those on co- existing substrates commonly found present study, even after 3 months, could impart further differences in reef habitats. Importantly, larval settlement is driven by bacterial in invertebrate larval settlement, and so have cascading effects on the WITT eT al . 460       subsequent reef formation and succession. This fast response time ash and carbonate reef sand compared to the glass and calcite sands. echoes recent studies of fresh basaltic lava flows in terrestrial envi- The former materials exhibit higher specific surface areas, reflecting ronments, where it was suggested that bacterial communities were higher surface roughness, porosity or the presence of smaller parti- rapidly established within days or months of lava flow emplacement cles adhering to larger particle surfaces; these properties are known (Kelly, Cockell, Thorsteinsson, Marteinsson, & Stevenson, 2014). to affect growth in other bacterial systems (Gerasimenko, Orleanskii, Karpov, & Ushantinskaya, 2013; Yamamoto & Lopez, 1985). We detected significantly different bacterial communities on white car- 4.1 | Differences in bacterial community structure bonate and calcite substrates, while quartz was the same as both of Evaluation of the bacterial communities colonising volcanogenic, bio- these biogenic substrates. In contrast, in a comparable study of sili- genic and terrigenic substrates in a coral reef environment mesocosm, cate and carbonate substrates in situ (Red Sea), Schöttner et al. (2011) as determined by DNA fingerprinting of the ITS region using Automated found that both sands showed similar bacterial density and diversity, Ribosomal Intergenic Spacer Analysis (ARISA), showed both substrate- although with differing bacterial community structure over seasonal specific and shared bacterial OTUs. Such differences in the establish- changes. However, the carbonate sand investigated by Schöttner et al. ment of biofilms on different substrates are compatible with previous (2011) was poorly sorted and significantly coarser (median particle investigations on biofilm formation on different silicate and carbonate size = 553 μm) than the quartz sand (median particle size = 326 μm). substrates incubated in tropical marine waters, which also noted the Yet, no evident influence of substrate chemistry between quartz and importance of the substrate for biofilm formation (Chung et al., 2010; carbonate materials in Schöttner et al. (2011) and the current study Dobretsov, Abed, & Voolstra, 2013; Witt et al., 2011). Further, DNA implies that differing substrate granulometry may have caused bacte- fingerprinting revealed distinct differences between bacterial biofilm rial niche- partitioning. communities on the substrates and the communities found in the water An effect of substrate colour could explain our observation that column over the 3- month experiment. Differences in the structure and the two dark volcanogenic substrates carried more diverse bacterial diversity between the attached and free- living bacterial communities communities than the three light substrates. The colour of a substrate are in line with in situ observations in different marine coastal systems has recently been observed to affect attraction of different microbial (Mohit, Archambault, Toupoint, & Lovejoy, 2014; Zhang, Liu, Lau, Ki, communities, whereby black substrates carry a higher bacterial den- & Qian, 2007) and have also previously been detected in coral reefs in sity than white substrates (Dobretsov et al., 2013). Notably, diverse situ (Santavy & Colwell, 1990; Schöttner et al., 2009). invertebrate settlement assays have shown that some larvae and Of the five substrates, we observed that the volcanogenic group algal spores prefer dark substrates over light- coloured ones, likely (ash, glass) carried a more diverse, and significantly different, bacte- due to diminished light reflection and greater heat retention of dark rial community compared to the communities associated with bio- substrates, conditions that are preferred by negatively photo- tactic genic (carbonate and calcite sand) and terrigenic (quartz) substrates. organisms (Svane & Dolmer, 1995). Further, invertebrates often pre- The SIMPER analysis revealed substrate- specific bacterial OTUs in fer dark substrates for colonisation to benefit from better protection the volcanogenic substrates contributing the most to the observed from grazers (Swain, Herpe, Ralston, & Tribou, 2006). These factors are differences. These were either OTUs found in all substrate commu- likely to apply to bacterial colonisation as well. Therefore, Dobretsov nities, which were more prevalent in the volcanogenic substrates, or et al. (2013) and the current study emphasise the effects of colour on were unique to the volcanogenic substrates, highlighting the substrate bacterial settlement, which should be tested further in future settle- specificity of the colonising bacterial community. Further, statistical ment assays. tests of the Shannon–Wiener diversity index confirmed significantly The chemistry and/or mineralogy of the substrates could equally higher diversity within the volcanogenic substrates when compared govern the observed differences in community structure, as substrate to the other three substrates, while, amongst the biogenic and terri- colour is strongly influenced by composition and the coordination genic substrates, no significant differences in diversity were detected. of atoms within a material (Nassau, 1978). These properties will, in However, within the volcanogenic and biogenic substrate groups, the turn, influence the distribution and availability of nutrients at the sub- number of total OTUs and diversity indices indicate that the ash and strate surfaces. Nutrients may be extracted from the substrate sur- carbonate reef sands carried a higher bacterial diversity than the cal- face by leaching and dissolution by organic compounds and by water cite and glass. The observed differences in bacterial abundance and within the established biofilm (Brehm, Gorbushina, & Mottershead, community differences amongst substrates indicate a substantive 2005). Pre- leaching of the materials in the present study was essen- control of substrate physicochemical properties on bacterial commu- tial to ensuring that nutrients were substrate- derived. However, the nity settlement and structure. response to available nutrients is likely to be species- dependent (Gerasimenko et al., 2013; Nies & Silver, 1989); thus, differences in substrate chemistry and/or mineralogy may promote or inhibit growth 4.2 | Physicochemical controls on bacterial of different bacterial species on the different substrates. communities Dependences of bacterial community structure on substrate Differences in substrate physical properties (surface morphology and mineralogy have been previously invoked by Kelly et al. (2011, granulometry) may account for the higher bacterial diversity on the 2009), Gleeson et al. (2006) and Hutchens, Gleeson, McDermott, WITT eT al .       461 Miranda-CasoLuengo, and Clipson (2010). The first two studies listed community structure is likely dictated by differences in substrate above note correlations between bacterial communities on weathered physicochemical properties. We identify greater diversity in sub- glasses and crystalline rocks, depending on their composition (basaltic strates with higher specific surface areas compared to those with to rhyolitic). The last two studies document significant differences in lower surface areas but are compositionally similar, which, coupled bacterial community structuring in biofilms growing on adjacent (on with comparisons to in situ studies, suggests possible controls associ- length scales of cm to m) silicate minerals (quartz, albite, K- feldspar, ated with particle physical properties (e.g., granulometry, surface mor- muscovite) at the surface of a terrestrial granite. Accordingly, there phology). Our findings also suggest a significant control of substrate may therefore be significant differences in bacterial community struc- composition (bulk chemistry and mineralogy), which could result ture both spatially across a single ash deposit, and in deposits of differ- from a direct influence of nutrient availability or an indirect influence ing composition produced by different volcanoes. through substrate colour, whereby compositionally diverse dark- Direct comparison of the fresh volcanic ash substrates used here and coloured volcanogenic substrates favoured development of a larger the weathered terrestrial rocks should be made carefully as weathering community structure relative to light- coloured quartz and biogenic alters nutrient availability and introduces mineral phases (e.g., palago- substrates. Identification of the bacterial community diversity using nite; Kelly et al., 2010) absent in the fresh material. It may even be dif- next- generation sequencing and an “omics” approach would provide ficult to compare fresh ash surfaces and those of fresh lava flows (e.g., additional information about the pioneer bacterial communities on the Kelly et al., 2014), as the surface of the latter is altered by crystallisation different substrates and help to understand their function. Critically, (Burkhard, 2002) and volatile degassing processes (White & Hochella, our findings indicate the potential for volcanic ash to promote bac- 1992) during and immediately after emplacement. However, if similar terial diversity in an immediate post- burial scenario in ash- affected differences in bacterial communities depending on the silicate mineral coral reefs, on timescales that could permit cascading effects on lar- substrate are found in our substrates as in terrestrial studies, our data val settlement and ultimately reef recovery. Further investigation of would imply a shared mineralogy between the ash and glass substrate. coral reef recovery and resilience following large- scale natural distur- As ash from Sakurajima volcano is predominantly glassy (70%–90%, bances, such as volcanic ash deposition, may contribute to predictions Miwa, Geshi, & Shinohara, 2013), otherwise comprising plagioclase and on ecosystem recovery and hazard management strategies. mafic minerals, this may suggest that the glass component is driving the bacterial community structure of both volcanogenic substrates. As nei- A CKNO WLEDGMENT S ther of the volcanic materials contain a significant quartz component, their different community structures relative to that of the terrigenic VW, PMA, CC and DED thank the AXA Research Grant “Risk from quartz would be consistent with the mineralogical dependence invoked volcanic ash in the Earth system” for supporting the research by Gleeson et al. (2006) and Hutchens et al. (2010). and DBD acknowledges the ERC Advanced Investigator Grant Volcanic ash is characterised by a wide range of physical (morphol- No. 247076 (EVOKES). We acknowledge R. Melzer for provid- ogy, particle size, surface area, colour) and chemical properties (min- ing access to the scanning electron microscope at the Zoologische eralogy, composition), and volcanic ash deposits can contain a diverse Staatssammlung München. Thanks to Larry Miller for providing an array of particles, including entirely glassy or crystalline particles from internal USGS review. fresh magma, remobilised older ejecta, and fragments of weathered lithic rocks. Furthermore, ash can deposit variably according to erup- REFERENCES tion dynamics as well as ash dispersal and sedimentation patterns Ayris, P. M., & Delmelle, P. (2012). The immediate environmental effects of with increasing distance from the volcano (Bonadonna, Costa, Folch, tephra emission. Bulletin of Volcanology, 74(9), 1905–1936. & Koyaguchi, 2015). 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Surface chemistry associated with https://doi.org/10.1111/gbi.12231 the cooling and subaerial weathering of recent basalt flows. Geochimica et Cosmochimica Acta, 56(10), 3711–3721. Wieczorek, S. K., & Todd, C. D. (1998). Inhibition and facilitation of settlement of epifaunal marine invertebrate larvae by microbial biofilm cues. Biofouling, 12(1–3), 81–118. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Geobiology Wiley

Volcanic ash supports a diverse bacterial community in a marine mesocosm

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Abstract

Sciences, Ludwig-Maximilians-Universität, Munich, Germany Shallow- water coral reef ecosystems, particularly those already impaired by anthropo- United States Geological Survey, Menlo Park, genic pressures, may be highly sensitive to disturbances from natural catastrophic CA, USA events, such as volcanic eruptions. Explosive volcanic eruptions expel large quantities 3 LMU GeoBio-Center , Munich, Germany 4 of silicate ash particles into the atmosphere, which can disperse across millions of SNSB-Bavarian State Collections of Palaeontology und Geology, Munich, Germany square kilometres and deposit into coral reef ecosystems. Following heavy ash deposi- MWM-Museum Witt München, Munich, tion, mass mortality of reef biota is expected, but little is known about the recovery of Germany post- burial reef ecosystems. Reef regeneration depends partly upon the capacity of Correspondence the ash deposit to be colonised by waterborne bacterial communities and may be in- V. Witt and C. Cimarelli, Department of fluenced to an unknown extent by the physiochemical properties of the ash substrate Earth and Environmental Sciences, Ludwig- Maximilians-Universität, Munich, Germany. itself. To determine the potential for volcanic ash to support pioneer bacterial coloni- Emails: wittverena@gmx.de and cimarelli@ sation, we exposed five well- characterised volcanic and coral reef substrates to a ma- min.uni-muenchen.de rine aquarium under low light conditions for 3 months: volcanic ash, synthetic volcanic glass, carbonate reef sand, calcite sand and quartz sand. Multivariate statistical analy- sis of Automated Ribosomal Intergenic Spacer Analysis (ARISA) fingerprinting data demonstrates clear segregation of volcanic substrates from the quartz and coral reef substrates over 3 months of bacterial colonisation. Overall bacterial diversity showed shared and substrate- specific bacterial communities; however, the volcanic ash sub- strate supported the most diverse bacterial community. These data suggest a signifi- cant influence of substrate properties (composition, granulometry and colour) on bacterial settlement. Our findings provide first insights into physicochemical controls on pioneer bacterial colonisation of volcanic ash and highlight the potential for vol- canic ash deposits to support bacterial diversity in the aftermath of reef burial, on timescales that could permit cascading effects on larval settlement. 1  |  INTR ODUCTION (Wilkinson 1999). The deterioration of water quality in coastal regions consequently favours macro- algal dominance (Fabricius, 2005; Coral reefs are unique, biodiverse ecosystems of high socio- economic Schaffelke, Mellors, & Duke, 2005) and increases the risk of disease importance on both global and local scales (Nicholls et al., 2007). for coral reef- building species, including sponges and corals (Haapkyla Anthropogenic disturbances, such as sedimentation and eutrophi- et al., 2011; Webster, Xavier, Freckelton, Motti, & Cobb, 2008), which cation, increasingly pressure fragile coral reef ecosystems worldwide further exacerbates coral reef vulnerability to catastrophic natural This is an open access article under the terms of the Creative Commons Attribution NonCommercial License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited and is not used for commercial purposes. © 2017 The Authors. Geobiology Published by John Wiley & Sons Ltd. Geobiology. 2017;15:453–463. wileyonlinelibrary.com/journal/gbi     453 WITT eT al . 454       disturbances, such as volcanic ash deposition (Vroom & Zgliczynski, Hoffmann, 2007; Munn, 2003), while others (e.g., Al, Cu) may be toxic 2011). After an explosive volcanic eruption, widespread dispersal and (Duggen et al., 2007b). deposition of volcanic ash over areas up to millions of square kilome- Investigating the propensity for volcanic ash to promote pioneer tres, in thickness of up to several centimetres, may be damaging to ash- bacterial colonisation in situ is hampered by the difficulties associated affected coral reef ecosystems; both Maniwavie, Rewald, Aitsi, Wagner, with substrate accessibility, geographic location and the dangers asso- and Munday (2001) and Vroom and Zgliczynski (2011) have reported ciated with sampling near active volcanoes. Laboratory experiments the destruction and mass mortality of reef biota following heavy ash conducted in simulated tropical coral reef aquaria, therefore, offer deposition. However, the capacity of reef ecosystems to recover after a viable method to approximate the properties of volcanic ash that burial by ash remains uncertain. Maniwavie et al. (2001) reported that may dictate its capacity to act as a colonisable substrate. In this study, 2 years after burial by volcanic ash corals had only re- colonised the sur- we incubate a selection of volcanogenic, terrigenic and biogenic sub- faces of protruding or unburied objects (e.g., boulders, tree stumps), strates in a coral reef- like marine aquarium system for 3 months and while the ash substrate itself remained barren; in contrast, Schils (2012) correlate differences in the bacterial colonising consortia with physi- noted that a period of frequent ash deposition into a tropical reef eco- cochemical properties (i.e., chemical composition, mineralogy, granu- system promoted a change in benthic microbial and macrofloral com- lometry, morphology, colour) of particulate substrates. munities on a similar timescale. These varying responses indicate a clear need to better understand the factors that may dictate the recovery of 2  |  MATERIALS AND METHODS vulnerable and valuable coral reef ecosystems after ash deposition. In the aftermath of large- scale burial, recovery of the reef eco- 2.1 | Substrate selection, preparation and system may depend on pioneer colonisation of the new substrate by characterisation free- living bacteria from the water column. After attachment to the surface, these bacteria produce an extracellular polymeric matrix The effect of physicochemical parameters on the composition of bac- that embeds further microbial organisms, forming so- called biofilms terial community colonisation was explored using five different sub- (Costerton, Lewandowski, Caldwell, Korber, & Lappin- Scott, 1995). strates in a marine aquaria under controlled conditions for bacterial Biofilm communities are highly abundant in coral reefs and are crucial colonisation: two volcanogenic materials (volcanic ash and synthetic in biogeochemical nutrient cycling and the degradation of anthropo- volcanic glass), two biogenic materials of reef origin (carbonate coral genic pollutants (reviewed in Davey & O’Toole, 2004). Further, they reef sand and calcite sand) and a terrigenic quartz sand (Table 1). Fresh, provide an essential settlement surface for larvae of important reef- crystal- bearing volcanic ash was obtained from Sakurajima volcano, building invertebrates (e.g., corals and sponges) and influence larval Japan, on 18 July 2013. Sakurajima was selected because it prevalently settlement cues and metamorphosis (Webster et al., 2004; Wieczorek erupts andesitic magma, which is of intermediate volcanic composi- & Todd, 1998). Accordingly, any changes in bacterial biofilm communi- tion and represents the dominant composition at island arc volcanoes, ties may influence invertebrate larval settlement, coral reef establish- and because the physicochemical properties of its eruptive products ment and further development. Therefore, the capacity of a volcanic are well characterised (Hillman et al., 2012). The synthetic volcanic ash substrate to support bacterial settlement, particularly compared to glass substrate was employed to provide a crystal- free system of com- the marine substrates it overlies, may play a crucial role in shaping the parable chemical composition to the volcanic ash substrate and was recovery of ash- affected reef ecosystems. prepared by subjecting the Sakurajima ash to five cycles of high tem- Previous studies on aquatic biofilm formation using an array of perature melting in a platinum crucible at 1500°C to produce a homo- natural and artificial substrates, including basaltic glasses and boro- geneous melt. This melt was quenched rapidly to produce a glass and silicate (Thorseth, Furnes, & Tumyr, 1995), biotite (Ward, 2013), gran- then mechanically crushed. The terrigenic quartz sand was selected to ite (Chung et al., 2010), coral skeletons and clay tiles (Witt, Wild, & provide a silicate reference material for comparison with the volcano- Uthicke, 2011), highlight the importance of physicochemical properties genic substrates and was obtained commercially (Sigma- Aldrich ID no. (e.g., granulometry, surface morphology, mineralogy and chemistry, 274739). To draw comparisons between the silicate substrates and bio- colour) in promoting initial substrate colonisation. Crucially, volcanic genic reef sands, we utilised two different carbonate materials, each ash materials are subject to a wide variation in all of these proper- with different origins. The coral reef sand was obtained from a shallow ties, which are the product of magma composition and eruption his- fringing reef in the north- eastern Gulf of Aqaba, Red Sea, located within tory (Dingwell, Lavallée, & Kueppers, 2012). Ash particles range in size a marine reserve close to the Marine Science Station in Aqaba, Jordan from the millimetre to submicron scale and vary in morphology from (29°27′N, 34°58′E), and was mechanically crushed before use. The cal- smooth, blocky particles, to rough- textured vesicular clasts (Heiken, cite sand was sourced from Longcliffe Quarries Ltd. (Brassington, UK). 1974). They commonly contain crystalline and amorphous silicates of The two volcanic substrates were subjected to a preliminary leach- various compositions, and range in colour from light to dark (Ayris & ing period of four hours in deionised water at an ash:water mass ratio Delmelle, 2012). Ash surfaces can be a source of variably extractable of 1:5 and dried at 105°C for six hours. The remaining three substrates elements, some of which (e.g., Al, Ca, Co, Cu, Fe, K, Mg, Mn, Ni, Mo, were washed in deionised water to remove very fine particulate P, S, Zn; Jones & Gislason, 2008) may be important macro- or micro- adhering to the surface of bigger clasts, and dried at 105°C for six nutrients for bacteria and phytoplankton (Duggen, Croot, Schacht, & hours. All substrates were then leached for 12 hr in sea water taken WITT eT al .       455 TABLE 1 Physicochemical data of the five substrates Carbonate reef a b b c Substrate Volcanic Glass Volcanic Ash sand Calcite Quartz Chemical composition (wt. %) SiO 58.2 58.5 0.2 0.9 >99.5 Al O 16.8 16 0.1 0.3 n.s. 2 3 Fe O 7.9 7.9 0 0 n.s. 2 3 MnO 0.2 0.2 0 0 n.s. MgO 3.4 3.6 1.3 0.4 n.s. CaO 7.4 7.2 52.2 54.4 n.s. Na O 3.4 3 0.3 0.1 n.s. K O 1.4 1.4 0 0 n.s. TiO 0.8 0.8 0 0 n.s. P O 0.2 0.2 0.1 0 n.s. 2 5 Total 99.6 100.1 99.0 100.0 >99.5 L.O.I −0.17 0.02 44.49 44.43 <0.5 Physical properties SSA (m /g) 0.02 0.1 0.49 0.22 0.11 Median diameter (μm) 234 221 224 230 263 Interquartile diameter 180–260 190–260 190–270 190–260 220–310 (μm) Colour Dark Dark White White White Morphology Angular Subangular Aggregate Aggregate Subangular Surface appearance Smooth Rough Rough Rough Rough Comprised of glass, plagioclase, pyroxenes, Fe–Ti oxides (Miwa et al., 2013). Comprised of CaCO . Compositional data of pure quartz from Sigma–Aldrich product specifications (n.s. not stated). from the aquarium system at a 1:5 ash:water mass ratio and dried at Brunauer–Emmett–Teller (BET) theory to argon adsorption measure- 105°C for six hours, spread out in large Petri dishes, UV- sterilised by ments conducted at −196°C using a Micrometrics Gemini surface area 2 × 30- min cycles and stirred in between the cycles. These treatments analyser. Samples were dried overnight at 105°C overnight prior to were intended to isolate the effect of the substrate by eliminating any analyses. Data are the mean of three repeated measurements. Particle potential contribution from mechanical crushing of the glass, calcite size distributions were determined by laser particle analysis using a and quartz substrates, any adsorbed chemical species (Ayris et al., Beckman- Coulter LS230. The particle morphology of the leached sub- 2014) that may be toxic to the other organisms in the aquarium water strates was examined by scanning electron microscopy (SEM; Leo 1430 system, and any pre- existing contamination by bacteria prior to or VP) with a maximum operating voltage of 15 kV. Samples were prepared between sample collection/synthesis. The absence of bacteria on the for imaging on SEM stubs and sputter coated with gold. Representative samples was confirmed at the onset of the experiment using the DNA images of single particles were taken for each sample to qualitatively fingerprinting tool of Automated Ribosomal Intergenic Spacer Analysis document differing particle morphologies amongst the substrates. (ARISA) (see details in Methods section below). As this study inten- tionally targets the physicochemical properties of substrates, the pre- 2.2 | Design of the marine colonisation experiment liminary leaching of volcanic ash was intended to remove any soluble salts emplaced on ash surfaces during transport through the volcanic The aquarium system consists of a main tank (330 L) containing a eruption plume. While the relevance of surface salts as a readily avail- 7- year established coral reef community, including corals, algae, able nutrient source in ocean surface waters has been previously con- diverse marine invertebrates and fish. The main tank supplies sea sidered (e.g., Duggen et al., 2007b), their capacity for rapid dissolution water to a series of 80- L subtanks. Those subtanks were used for on first contact with sea water likely limits their relevance to any sub- experiments and did not contain coral reef organisms. The subtank sequent influence once deposited as a substrate on the ocean floor. utilised in this study was configured for a 12:12- hr light:dark cycle and Bulk chemical compositions of the substrates were determined by was covered by shade cloth to ensure low light availability of <0.2 klux. X- ray fluorescence spectroscopy (Philips, MagiX Pro). The major and Light availability and temperature were recorded at 30- min intervals minor elements were measured using glass beads prepared by fusion over the course of the experiment using loggers (Onset HOBO TidBit). with a lithium borate flux in a Panalytical Eagon 2 furnace fusion Tank sea water was maintained at constant flow rate, as to not perturb system. Specific surface area was determined by application of the the substrates in the dishes (see below), constant average temperature WITT eT al . 456       (25.4°C ± 0.05), pH (8.13 ± 0.05) and conductivity (51.8 ± 1.14 mS), structure amongst different substrates. DNA was extracted from and was tested weekly to ensure that nutrient concentrations 0.25 g of each of the collected subsamples (three replicates and two remained low (<8 nmol/L NO , <0.2 nmol/L PO ; see Table S1). separate extractions from each) and water samples in separate tubes. 3 4 The duration of the experiment was 3 months, with sampling of Extraction utilised a PowerLyzer DNA Isolation Kit according to the all substrates after one (T1), two (T2) and three (T3) months. For each manufacturer’s instructions (MoBio, Carlsbad, CA, USA) with the fol- sampling time- point, each substrate was represented by three separate lowing alterations: bead beating cycles 2 × 30 s for higher yields of Petri dishes. Sterile glass Petri dishes of 50 mm diameter and 10 mm bacterial DNA and elution in 2 × 50 μL 1 × TE buffer to lower the pos- depth were filled with 1 g of substrate (a uniform depth of ~5 mm) and sibility of disturbing polymerase chain reaction (PCR) amplifications. positioned approximately 50 mm apart at the bottom of one of the 80- L DNA from all samples was stored in aliquots at −20°C until fur- subtanks, immersed to a depth of 250 mm in sea water. The granulom- ther processing. Extracted DNA from all replicates (three per sub- etry of the substrate was selected to minimise the possibility of any strate and time- point) was amplified by PCR using the ITSF-F AM disturbance by the water flow. To prevent loss of material during the and ITSReub primer pair (Cardinale et al., 2004) in three separate initial sample placement, the dishes were carefully filled with sea water reactions per replicate sample. The ARISA- PCR mixture (25 μL) con- via manual pipetting and loaded into the tank with their lids in place. tained 5 × PCR buffer (Promega, Madison, WI, USA); 2.5 mm MgCl After placement, the lids were removed. In total, 45 dishes were utilised (Promega); 0.25 mm deoxynucleoside triphosphate mix (Promega); in the experiment (5 substrates × 3 sampling points × 3 replicates). At 1.5 μg/μL bovine serum albumin (BSA); 400 nm universal primer each sampling time- point, 2 × 0.3 g of substrate was recovered from ITSF- FAM (5′- GTCGTAACAAGGTAGCCGTA- 3′), 5′- labelled with the each replicate dish using a sterilised spatula. DNA from the substrate phosphoramidite dye FAM and eubacterial ITSReub (5′- GCCAAGGC samples was either extracted immediately, or the sampled substrates ATCCACC- 3′); and 0.025 U of GoTaq polymerase (Promega). To each were directly snap- frozen in liquid nitrogen and stored at −80°C and reaction, approximately 20 ng of extracted DNA (quantified by a then extracted within 2 weeks after sampling. To ensure that observed ND- 1000 Nanodrop; Peqlab Biotechnology, Erlangen, Germany) was bacterial differences amongst substrates and time- points were not due added and the volume was adjusted to 25 μL using PCR water. Thermal to changes in water quality parameters (i.e., temperature, pH, conduc- cycling was carried out in an Eppendorf MasterCycler (Eppendorf, tivity, nutrient concentrations), 2 × 500 mL seawater samples were col- Hamburg, Germany) with an initial denaturation at 94°C for 3 min, lected at the time of substrate immersion (T0 beginning of the experi- followed by 30 cycles of 94°C for 45 s, 55°C for 45 s, 72°C for 90 s, ment) and at each sampling time- point (n = 2), filtered through 0.2 μm with a final extension at 72°C for 5 min and then cooled to 15°C. polycarbonate filters using a vacuum pump, and filters were stored and PCR products were purified utilising the Nucleospin PCR clean- up kit processed following the same protocols as the substrate samples. (Macherey- Nagel, Düren, Germany). Diluted PCR products (300–1200 bp) were prepared for analysis by ARISA using capillary electrophoresis as follows. A standardised 2.3 | Determination of differences in bacterial amount of DNA (100 ng) was added to a separation cocktail contain- community structures ing 0.5 μL of internal GeneScan 1200LIZ size standard (50–1000 bp) Differences in bacterial community structure amongst substrates and (Applied Biosystems, Foster City, CA, USA) and 14 μL of deionised time- points were determined by the DNA fingerprinting method of Hi- Di formamide. The preparation was denatured for 3 min at 95°C ARISA using capillary electrophoresis. The technique targets the ITS and kept on ice for at least 5 min before being further processed by region (Intergenic Transcribed Spacer) between the 16S and the 23S the sequencer. Separation of the PCR- amplified fragments via capil- rRNA gene regions, which is highly variable in nucleotide sequence lary electrophoresis was carried out on an 80 cm- capillary ABI PRISM and length (from 50 bp to 1500 bp). Different bacterial species exhibit 3730xl genetic analyzer (Applied Biosystems) with the following run specific nucleotide lengths relative to the ITS region and, hence, the parameters: 14.6 kV run voltage, 2.4 kV injection voltage, 20- s injec- bacterial community structure (community composition and relative tion time, and 60°C oven temperature. Raw profiles were checked abundance of dominant bacterial species) can be determined (Brown for stable baselines and voltage, and peak sizes and absolute areas & Fuhrman, 2005). While next- generation sequencing (NGS) enables were then determined using the GeneMapper software v4.0 (Applied more in- depth studies, Gobet, Boetius, and Ramette (2014) emphasise Biosystems) with minimum peak heights of 50 fluorescence units for that the DNA fingerprinting of ARISA retains value in determining dif- all dyes. The best- fit size- calling curves were built using a second- ferences in community structure. It is particularly beneficial for the order least- squares method (compensating for anomalously running current study, as it permits the identification of fundamental differ- fragments in the standard) and the Local Southern method as the size- ences in bacterial community composition amongst many different calling algorithm. A perfect fit for the calibration curves in the range samples. DNA fingerprinting is a cost- effective and rapid approach of 100–1000 bp was always checked before further processing the to identify taxonomic affiliations or diversity estimates and to sepa- samples. The fragments/peaks matrix generated with GeneMapper rate samples on local spatio- temporal scales, and correlate environ- was transferred into the T- align software (Smith et al., 2005). A differ- mental variables with bacterial community structure (van Dorst et al., ence of 1.5 bp between peaks was defined as a single operational tax- 2014), prior to embarking on more cost- intensive NGS campaigns. onomic unit (OTU) (Danovaro, Luna, Dell’Anno, & Pietrangeli, 2006). Hence, this tool was chosen to gain an initial overview of community The values of the relative fluorescence intensity of the peak area of the WITT eT al .       457 ARISA fingerprinting data were normalised, square- root transformed composed of micron- sized fragments of carbonate biogenic mate- and standardised prior to statistical analysis. rial (mainly shell and exoskeleton fragments of marine organisms) cemented by crystalline calcite. Calcite clasts are similarly blocky and equant and appear to consist of aggregates of micron- sized calcite 2.4 | Statistical analysis minerals. The specific surface area varies between 0.02 and 0.5 m /g, To explore the variation in the community structure and to determine with the highest surface area materials being the biogenic substrates 2 2 whether bacterial assemblages grouped by substrate, multivariate sta- (carbonate reef sand = 0.5 m /g, calcite = 0.2 m /g; Table 1). tistics were applied. Pairwise distance matrices were calculated from the relative abundance data (ARISA) using the Bray–Curtis dissimilar- 3.2 | Marine colonisation experiment ity index. Non- metric multidimensional scaling (nMDS) was applied to the distance matrices to explore the variation in community struc- Multivariate statistical analysis of the ARISA fingerprinting data of the ture. Differences in the bacterial community structure according to ITS region of the biofilm communities established upon the five sub- substrate were assessed for significance by applying the ANalysis Of strates repeatedly showed overall significantly different bacterial com- SIMilarity (ANOSIM) test based on permutation (9999 permutations) munity compositions over the 3- month period and was consistently procedures using the Bray–Curtis distance measure. The contribution distinct from the community within the water column, represented by of each single taxon to the overall dissimilarities of factors was deter- replicates from the beginning (T0) and additional data points as time mined using the SIMilarity PERcentage (SIMPER) routine. progressed. These differences are reflected by the variations in the total To visualise the community structure and relationships between number of OTUs and in the number of unique OTUs for each substrate substrate- specific and shared operational taxonomic units (OTUs) at each time- point. We summarise these trends in the following para- amongst the different substrates, a Venn diagram was constructed. graphs but, for brevity, we further interpret the data for month three To determine differences in the number of OTUs between substrate only, as this represents the most well- established bacterial community. groups and the water column, one- way Analysis of Variance (ANOVA) Total OTUs successively increased by over 2.5- fold from the first to was applied. Homogeneity of variance was tested using Levene’s test, the third month of the experiment (from 108 OTUs at T1 to 273 OTUs and the Tukey–Kramer test was used as a post hoc test. To determine at T3). At each time- point, most OTUs were shared amongst the sub- biodiversity differences from the fingerprinting data, the Shannon– strate communities, with few substrate- specific ones. Both shared and Wiener (H′) and Simpson’s (1- D) indices for bacterial OTUs in each substrate- specific OTUs increased with time from 99 shared and 10 substrate (average value for 6 replicates) were calculated. To test for substrate- specific OTUs at T1 to 259 and 13 substrate- specific OTUs significant differences between the bacterial communities on the dif- at T3. Amongst the five-substrates bacterial communities, volcanic ash ferent substrates, the Shannon index values (n = 6 per substrate) for showed the most substrate- specific OTUs, followed by glass, carbon- bacterial communities on the five substrates after 3 months of incu- ate and calcite, with quartz exhibiting no substrate- specific OTUs at all. bation were analysed using the t- test. All statistical analyses were per- The grouping patterns visualised in nMDS (Figure 2) showed sig- formed using the statistical program PAST (Hammer, Harper, & Paul, nificant differences amongst the bacterial communities on the five 2001). The level of significance was <0.05 for all statistical tests. substrates at T1 (ANOSIM R = .272, P = .0001), T2 (ANOSIM R = .557, P = .0002) and T3 (ANOSIM R = .507, P = .0001), apart from quartz, which was different to the volcanic substrates but very similar to the 3  |  RESULT S biogenic substrates carbonate and calcite at T2 (ANOSIM R = .191, P = .273) and T3 (ANOSIM R = .033, P = .065 and R = .209, P = .303, 3.1 | Particle substrate characterisation respectively, Table S2). At T1 and T2, Quartz did not exhibit any spe- Bulk chemical analysis by XRF confirms the volcanic ash, glass and cific bacterial community. The overall community for all five substrates quartz as silicates and the reef sand and calcite substrate as calcium showed distinct groupings of bacterial communities associated to the carbonates (Table 1). The volcanic ash and glass are both dark in col- substrates of volcanic origin and coral reef origin, while quartz and the our, whereas the quartz, reef sand and calcite are all white. Particle carbonate substrates both share bacterial OTUs. SIMPER analysis at T1 size data showed mono- modal Gaussian distributions for all tested revealed that the OTUs contributing the most to the community differ- substrates. Ash, glass, calcite and reef sand had median particle diam- ences were 545 bp (5%), 890 bp (4%) and 620 bp (4%). Of these, the eters of 220–234 μm, while the quartz sand had a median diameter of 545- and 890- bp OTUs were exclusive to, and dominant within, the 263 μm (Table 1). Particle morphology of the substrates was investi- volcanogenic substrate bacterial community, whereas 620- bp OTU was gated by SEM analyses (Figure 1). Ash comprised subangular blocky dominant in the volcanogenic substrates, occurred occasionally in the particles with rough surfaces, often with micron- sized vesicles. Glass biogenic substrates and was absent in the quartz substrate. In addition generated from ash melting and crushing shows smooth surfaces with to the groupings according to substrate, the bacterial assemblages on sharp edges (angular clasts) and conchoidal fractures imparted during each substrate differed significantly from those detected in the water preparation. Quartz particles display similar morphological character- column (ANOSIM R = .469, P = .0002). In the water column, 112 OTUs istics to the volcanic ash substrates, being blocky, subangular parti- were detected, of which 2 were specific to the water community. cles with rough surfaces. Reef sand particles are blocky aggregates, SIMPER analysis at T2 identified the fragments 530 bp (6%), 890 bp WITT eT al . 458       (a) (b) (c) (d) (e) FIGURE 1 Scanning electron microscopy (SEM) images of substrates: (a) volcanic ash, (b) synthetic volcanic glass, (c) quartz, (d) carbonate reef sand and (e) calcite. Images were collected at 15 kV (4%) and 721 bp (3%) contributing most to the overall community dif- as the bacterial assemblages on each substrate differed significantly ferences. We determined that the 530- bp fragment represents a domi- from those detected in the water column (ANOSIM R = .241, P = .049; nant OTU present in all substrates. Substrate- associated bacterial com- Table S2). SIMPER analysis revealed an overall dissimilarity pooled over munities differed significantly from those in the water column (ANOSIM all groups of 52.37% with the OTUs contributing most to community R = .294, P = .0002). In the bacterial community in the water column, structure differences being the 332- bp (4.6%), 422- bp (1.7%) and 481- 119 OTUs were detected, of which 3 OTUs were water- specific. bp (1.5%) fragments, but with unspecified taxonomic affiliations. These After 3 months (T3), a total of 273 OTUs could be detected in the OTUs were detected amongst all substrates, but were all higher in the overall bacterial community from the ARISA data. Of these, 272 OTUs volcanogenic than in the biogenic and terrigenic substrates; 481- bp were detected on the substrates, with 13 substrate- specific OTUs and OTU was a dominant OTU throughout all substrates. 259 shared ones (Figure 3). Of the shared OTUs, 144 were shared All five substrates and the water column varied significantly amongst all five substrates. The volcanic ash substrate showed the larg- from each other with respect to the number of associated OTUs at est bacterial community with the most substrate- specific OTUs (7), fol- the 3- month time- point (one-w ay ANOVA, degrees of freedom = 5, lowed by volcanic glass (3), while a further 30 shared OTUs were specific mean square = 13 844, F = 1044, P = <.0001, Tukey–Kramer post to the volcanogenic substrates only. The biogenic substrates showed hoc test P = .0001; Table S3). The volcanic substrates supported the very low substrate specificity with <2 substrate- specific OTUs each (reef highest number of total OTUs and, according to Shannon–Wiener and sand 1 OTU, calcite 2 OTUs). The bacterial community in the water col- Simpson’s diversity indices, also harboured the most diverse bacterial umn had a total of 127 OTUs, of which solely 1 OTU was water- specific. community compared to the quartz and biogenic substrates (Table in Multivariate statistical analysis of the ARISA fingerprinting data of Figure 2). Further, statistical t- tests confirmed significant differences in the ITS region of the biofilm communities established upon the five diversity of bacterial OTUs between the volcanogenic substrates and substrates after 3 months showed distinct groupings of bacterial com- the biogenic and quartz substrates, while calcite, carbonate reef sand munities associated with the substrates of volcanic origin and coral and quartz were not significantly different from each other (Table S4). reef origin (Figure 2), and of water. The two biogenic substrates had a higher variability amongst replicates than those of volcanic origin, but the greatest variability was detected amongst quartz substrate replicate 4  |  DISCUSSION samples. Bacterial communities on all substrates were significantly dif- ferent to each other (ANOSIM R = .507, P = .0001), apart from quartz, No previous in situ studies have investigated bacterial colonisation of which was different to the volcanic substrates but the same as the bio- volcanic ash as a substrate within a reef environment, and the only genic substrates (ANOSIM R = .209, P = .303 and R = .033, P = .065, in situ studies documenting the aftermath of ash deposition are both respectively, Table S2). Substrate specificity was further confirmed focused on macroflora and fauna and are contrasting in their results: WITT eT al .       459 FIGURE 2 Non- metric multidimensional scaling (nMDS) ordination (Bray–Curtis distance) of ARISA- derived bacterial community profiles at months 1, 2 and 3, with six replicates per substrate per time- point. Groupings are specified according to substrate. Data from an initial analysis (T0) of the bacterial community in the water column (n = 2) are plotted alongside the monthly water data. The water groupings are of cumulative data at each time- point, inclusive of T0, for reference. Points plotting close to each other show a more similar community than distant ones [Colour figure can be viewed at wileyonlinelibrary.com] FIGURE 3 Relationship between the substrate- specific and shared operational taxonomic units (OTUs) amongst all five substrates after 3 months. The diagram was constructed using the program VENN (http://bioinformatics.psb.ugent.be/webtools/Vennwileyonlinelibrary.com]). Table (left) shows the total number of OTUs per substrate after 3 months (T3), the Shannon–Wiener (H′) and Simpson’s (1- D) diversity indices. The values were calculated by taking the average of OTUs of the ARISA data of the replicate samples of each substrate (n = 6) [Colour figure can be viewed at wileyonlinelibrary.com] Maniwavie et al. (2001) implied that recolonisation of ash substrates community composition and settlement success rates increase with by reef flora did not occur, whereas Schils (2012) reported a sudden the age of the biofilm, and the timescale over which significant dif- change in benthic microbial and macrofloral communities of an ash- ferences in bacterial community structure were observed in the cur- affected reef. Therefore, while comparisons between the aquarium rent study are similar, and relevant, to the timescales required for the experiment of the current study and in situ reef settings should be settlement of invertebrate larva (days to weeks; Bao, Satuito, Yang, treated with caution, our study makes an important contribution to & Kitamura, 2007; Campbell et al., 2011). Consequently, the signifi- a highly uncertain subject by demonstrating that bacteria can colo- cant differences in pioneer bacterial colonisation between volcano- nise ash substrates and can establish a significantly different commu- genic substrates, and the terrigenic and biogenic substrates in the nity structure than those on co- existing substrates commonly found present study, even after 3 months, could impart further differences in reef habitats. Importantly, larval settlement is driven by bacterial in invertebrate larval settlement, and so have cascading effects on the WITT eT al . 460       subsequent reef formation and succession. This fast response time ash and carbonate reef sand compared to the glass and calcite sands. echoes recent studies of fresh basaltic lava flows in terrestrial envi- The former materials exhibit higher specific surface areas, reflecting ronments, where it was suggested that bacterial communities were higher surface roughness, porosity or the presence of smaller parti- rapidly established within days or months of lava flow emplacement cles adhering to larger particle surfaces; these properties are known (Kelly, Cockell, Thorsteinsson, Marteinsson, & Stevenson, 2014). to affect growth in other bacterial systems (Gerasimenko, Orleanskii, Karpov, & Ushantinskaya, 2013; Yamamoto & Lopez, 1985). We detected significantly different bacterial communities on white car- 4.1 | Differences in bacterial community structure bonate and calcite substrates, while quartz was the same as both of Evaluation of the bacterial communities colonising volcanogenic, bio- these biogenic substrates. In contrast, in a comparable study of sili- genic and terrigenic substrates in a coral reef environment mesocosm, cate and carbonate substrates in situ (Red Sea), Schöttner et al. (2011) as determined by DNA fingerprinting of the ITS region using Automated found that both sands showed similar bacterial density and diversity, Ribosomal Intergenic Spacer Analysis (ARISA), showed both substrate- although with differing bacterial community structure over seasonal specific and shared bacterial OTUs. Such differences in the establish- changes. However, the carbonate sand investigated by Schöttner et al. ment of biofilms on different substrates are compatible with previous (2011) was poorly sorted and significantly coarser (median particle investigations on biofilm formation on different silicate and carbonate size = 553 μm) than the quartz sand (median particle size = 326 μm). substrates incubated in tropical marine waters, which also noted the Yet, no evident influence of substrate chemistry between quartz and importance of the substrate for biofilm formation (Chung et al., 2010; carbonate materials in Schöttner et al. (2011) and the current study Dobretsov, Abed, & Voolstra, 2013; Witt et al., 2011). Further, DNA implies that differing substrate granulometry may have caused bacte- fingerprinting revealed distinct differences between bacterial biofilm rial niche- partitioning. communities on the substrates and the communities found in the water An effect of substrate colour could explain our observation that column over the 3- month experiment. Differences in the structure and the two dark volcanogenic substrates carried more diverse bacterial diversity between the attached and free- living bacterial communities communities than the three light substrates. The colour of a substrate are in line with in situ observations in different marine coastal systems has recently been observed to affect attraction of different microbial (Mohit, Archambault, Toupoint, & Lovejoy, 2014; Zhang, Liu, Lau, Ki, communities, whereby black substrates carry a higher bacterial den- & Qian, 2007) and have also previously been detected in coral reefs in sity than white substrates (Dobretsov et al., 2013). Notably, diverse situ (Santavy & Colwell, 1990; Schöttner et al., 2009). invertebrate settlement assays have shown that some larvae and Of the five substrates, we observed that the volcanogenic group algal spores prefer dark substrates over light- coloured ones, likely (ash, glass) carried a more diverse, and significantly different, bacte- due to diminished light reflection and greater heat retention of dark rial community compared to the communities associated with bio- substrates, conditions that are preferred by negatively photo- tactic genic (carbonate and calcite sand) and terrigenic (quartz) substrates. organisms (Svane & Dolmer, 1995). Further, invertebrates often pre- The SIMPER analysis revealed substrate- specific bacterial OTUs in fer dark substrates for colonisation to benefit from better protection the volcanogenic substrates contributing the most to the observed from grazers (Swain, Herpe, Ralston, & Tribou, 2006). These factors are differences. These were either OTUs found in all substrate commu- likely to apply to bacterial colonisation as well. Therefore, Dobretsov nities, which were more prevalent in the volcanogenic substrates, or et al. (2013) and the current study emphasise the effects of colour on were unique to the volcanogenic substrates, highlighting the substrate bacterial settlement, which should be tested further in future settle- specificity of the colonising bacterial community. Further, statistical ment assays. tests of the Shannon–Wiener diversity index confirmed significantly The chemistry and/or mineralogy of the substrates could equally higher diversity within the volcanogenic substrates when compared govern the observed differences in community structure, as substrate to the other three substrates, while, amongst the biogenic and terri- colour is strongly influenced by composition and the coordination genic substrates, no significant differences in diversity were detected. of atoms within a material (Nassau, 1978). These properties will, in However, within the volcanogenic and biogenic substrate groups, the turn, influence the distribution and availability of nutrients at the sub- number of total OTUs and diversity indices indicate that the ash and strate surfaces. Nutrients may be extracted from the substrate sur- carbonate reef sands carried a higher bacterial diversity than the cal- face by leaching and dissolution by organic compounds and by water cite and glass. The observed differences in bacterial abundance and within the established biofilm (Brehm, Gorbushina, & Mottershead, community differences amongst substrates indicate a substantive 2005). Pre- leaching of the materials in the present study was essen- control of substrate physicochemical properties on bacterial commu- tial to ensuring that nutrients were substrate- derived. However, the nity settlement and structure. response to available nutrients is likely to be species- dependent (Gerasimenko et al., 2013; Nies & Silver, 1989); thus, differences in substrate chemistry and/or mineralogy may promote or inhibit growth 4.2 | Physicochemical controls on bacterial of different bacterial species on the different substrates. communities Dependences of bacterial community structure on substrate Differences in substrate physical properties (surface morphology and mineralogy have been previously invoked by Kelly et al. (2011, granulometry) may account for the higher bacterial diversity on the 2009), Gleeson et al. (2006) and Hutchens, Gleeson, McDermott, WITT eT al .       461 Miranda-CasoLuengo, and Clipson (2010). The first two studies listed community structure is likely dictated by differences in substrate above note correlations between bacterial communities on weathered physicochemical properties. We identify greater diversity in sub- glasses and crystalline rocks, depending on their composition (basaltic strates with higher specific surface areas compared to those with to rhyolitic). The last two studies document significant differences in lower surface areas but are compositionally similar, which, coupled bacterial community structuring in biofilms growing on adjacent (on with comparisons to in situ studies, suggests possible controls associ- length scales of cm to m) silicate minerals (quartz, albite, K- feldspar, ated with particle physical properties (e.g., granulometry, surface mor- muscovite) at the surface of a terrestrial granite. Accordingly, there phology). Our findings also suggest a significant control of substrate may therefore be significant differences in bacterial community struc- composition (bulk chemistry and mineralogy), which could result ture both spatially across a single ash deposit, and in deposits of differ- from a direct influence of nutrient availability or an indirect influence ing composition produced by different volcanoes. through substrate colour, whereby compositionally diverse dark- Direct comparison of the fresh volcanic ash substrates used here and coloured volcanogenic substrates favoured development of a larger the weathered terrestrial rocks should be made carefully as weathering community structure relative to light- coloured quartz and biogenic alters nutrient availability and introduces mineral phases (e.g., palago- substrates. Identification of the bacterial community diversity using nite; Kelly et al., 2010) absent in the fresh material. It may even be dif- next- generation sequencing and an “omics” approach would provide ficult to compare fresh ash surfaces and those of fresh lava flows (e.g., additional information about the pioneer bacterial communities on the Kelly et al., 2014), as the surface of the latter is altered by crystallisation different substrates and help to understand their function. Critically, (Burkhard, 2002) and volatile degassing processes (White & Hochella, our findings indicate the potential for volcanic ash to promote bac- 1992) during and immediately after emplacement. However, if similar terial diversity in an immediate post- burial scenario in ash- affected differences in bacterial communities depending on the silicate mineral coral reefs, on timescales that could permit cascading effects on lar- substrate are found in our substrates as in terrestrial studies, our data val settlement and ultimately reef recovery. Further investigation of would imply a shared mineralogy between the ash and glass substrate. coral reef recovery and resilience following large- scale natural distur- As ash from Sakurajima volcano is predominantly glassy (70%–90%, bances, such as volcanic ash deposition, may contribute to predictions Miwa, Geshi, & Shinohara, 2013), otherwise comprising plagioclase and on ecosystem recovery and hazard management strategies. mafic minerals, this may suggest that the glass component is driving the bacterial community structure of both volcanogenic substrates. As nei- A CKNO WLEDGMENT S ther of the volcanic materials contain a significant quartz component, their different community structures relative to that of the terrigenic VW, PMA, CC and DED thank the AXA Research Grant “Risk from quartz would be consistent with the mineralogical dependence invoked volcanic ash in the Earth system” for supporting the research by Gleeson et al. (2006) and Hutchens et al. (2010). and DBD acknowledges the ERC Advanced Investigator Grant Volcanic ash is characterised by a wide range of physical (morphol- No. 247076 (EVOKES). We acknowledge R. Melzer for provid- ogy, particle size, surface area, colour) and chemical properties (min- ing access to the scanning electron microscope at the Zoologische eralogy, composition), and volcanic ash deposits can contain a diverse Staatssammlung München. Thanks to Larry Miller for providing an array of particles, including entirely glassy or crystalline particles from internal USGS review. fresh magma, remobilised older ejecta, and fragments of weathered lithic rocks. Furthermore, ash can deposit variably according to erup- REFERENCES tion dynamics as well as ash dispersal and sedimentation patterns Ayris, P. M., & Delmelle, P. (2012). The immediate environmental effects of with increasing distance from the volcano (Bonadonna, Costa, Folch, tephra emission. Bulletin of Volcanology, 74(9), 1905–1936. & Koyaguchi, 2015). 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Geobiology – Wiley

Published: May 1, 2017

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