# Acidification increases abundances of Vibrionales and Planctomycetia associated to a seaweed-grazer system: potential consequences for disease and prey digestion efficiency

Tânia Aires, Alexandra Serebryakova, Frédérique Viard, Ester A. Serrão, Aschwin H. Engelen

<figure><img src="image_1.png" /></figure>

## To cite this version:

Tânia Aires, Alexandra Serebryakova, Frédérique Viard, Ester A. Serrão, Aschwin H. Engelen. Acidification increases abundances of Vibrionales and Planctomycetia associated to a seaweed-grazer system: potential consequences for disease and prey digestion efficiency. PeerJ, 2018, 6, pp.e4377. 10.7717/peerj.4377. hal-01774858

HAL Id: hal-01774858

https://hal.sorbonne-universite.fr/hal-01774858

Submitted on 24 Apr 2018

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

<figure><img src="image_2.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# Acidification increases abundances of *Vibrionales* and *Planctomycetia* associated to a seaweed-grazer system: potential consequences for disease and prey digestion efficiency

Tania Aires<fnref n="1" />*, Alexandra Serebryakova<fnref n="1" />,<fnref n="2" />,* , Frédérique Viard<fnref n="2" />,<fnref n="3" /> , Ester A. Serrão<fnref n="1" /> and Aschwin H. Engelen<fnref n="1" />

## ABSTRACT

Ocean acidification significantly affects marine organisms in several ways, with complex interactions. Seaweeds might benefit from rising CO₂ through increased photosynthesis and carbon acquisition, with subsequent higher growth rates. However, changes in seaweed chemistry due to increased CO₂ may change the nutritional quality of tissue for grazers. In addition, organisms live in close association with a diverse microbiota, which can also be influenced by environmental changes, with feedback effects. As gut microbiomes are often linked to diet, changes in seaweed characteristics and associated microbiome can affect the gut microbiome of the grazer, with possible fitness consequences. In this study, we experimentally investigated the effects of acidification on the microbiome of the invasive brown seaweed *Sargassum muticum* and a native isopod consumer *Synisoma nadejda*. Both were exposed to ambient CO₂ conditions (380 ppm, pH 8.16) and an acidification treatment (1,000 ppm, pH 7.86) for three weeks. Microbiome diversity and composition were determined using high-throughput sequencing of the variable regions V5-7 of 16S rRNA. We anticipated that as a result of acidification, the seaweed-associated bacterial community would change, leading to further changes in the gut microbiome of grazers. However, no significant effects of elevated CO₂ on the overall bacterial community structure and composition were revealed in the seaweed. In contrast, significant changes were observed in the bacterial community of the grazer gut. Although the bacterial community of *S. muticum* as whole did not change, *Oceanospirillales* and *Vibrionales* (mainly *Pseudoalteromonas*) significantly increased their abundance in acidified conditions. The former, which uses organic matter compounds as its main source, may have opportunistically taken advantage of the possible increase of the C/N ratio in the seaweed under acidified conditions. *Pseudoalteromonas*, commonly associated to diseased seaweeds, suggesting that acidification may facilitate opportunistic/pathogenic bacteria. In the gut of *S. nadejda*, the bacterial genus *Planctomycetia* increased abundance under elevated CO₂. This shift might be associated to changes in food (*S. muticum*) quality under

<footnote n="1">¹Center for Marine Sciences (CCMAR), CIMAR, University of Algarve, Campus de Gambelas, Faro, Portugal</footnote>

<footnote n="2">²Sorbonne Université, CNRS, Lab Adaptation and Diversity in Marine Environments (UMR 7144 CNRS SU), Station Biologique de Roscoff, Roscoff, France</footnote>

<footnote n="3">³CNRS, UMR 7144, Divco Team, Station Biologique de Roscoff, Roscoff, France</footnote>

<footnote>*These authors contributed equally to this work.</footnote>
acidification. *Planctomycetia* are slow-acting decomposers of algal polymers that could be providing the isopod with an elevated algal digestion and availability of inorganic compounds to compensate the shifted C/N ratio under acidification in their food. In conclusion, our results indicate that even after only three weeks of acidified conditions, bacterial communities associated to ungrazed seaweed and to an isopod grazer show specific, differential shifts in associated bacterial community. These have potential consequences for seaweed health (as shown in corals) and isopod food digestion. The observed changes in the gut microbiome of the grazer seem to reflect changes in the seaweed chemistry rather than its microbial composition.

**Subjects** Ecology, Marine Biology, Microbiology, Climate Change Biology, Biological Oceanography

**Keywords** Invasive seaweeds, Ocean acidification, Grazer microbiomes, Algae microbiomes, Metabarcoding, *Sargassum muticum*, *Synisoma nadejda*

## BACKGROUND

Ocean acidification significantly affects marine organisms in diverse ways (Fabry et al., 2008; Kroeker et al., 2013). In the case of species interactions (e.g., predator–prey), the outcome of such effects can be difficult to predict as antagonistic or synergistic effects may be observed (Asnaghi et al., 2013; Branch et al., 2013; Poore et al., 2013). This is particularly true for non-calcifying seaweeds, which in contrast to most other organisms can benefit from rising CO₂ through increased photosynthesis and carbon acquisition, and subsequently acquire higher growth rates (Porzio, Buia & Hall-Spencer, 2011; Harley et al., 2012; Koch et al., 2013; Olischläger et al., 2013). However, changes in plant leaf chemistry in response to elevated carbon supply are expected to result in higher C:N and C:P ratios and, as such, reduce the nutritional quality of tissue for grazers (Urabe, Togari & Elser, 2003; Van De Waal et al., 2010) and the same is expected for seaweeds. Variations in the palatability of seaweeds may lead to changes in consumption rates by herbivores, which will have to absorb nutrients more efficiently or consume more to compensate for low concentrations of essential nutrients (Gutow et al., 2014). Thus, ocean acidification (OA) could have positive effects on seaweed growth rate, but may also induce behavioral changes on the herbivores and increased grazing rates. Therefore, it is important to understand the effects of ocean acidification on prey (bottom-up effects) and, as a top-down effect, on predation.

Interactions among organisms and their associated bacterial communities affect the holobiont physiology and health (Hollants et al., 2013; Egan et al., 2013), and play an important role in the functioning of hosts as, in the case of this study, seaweeds (Singh et al., 2011; Singh & Reddy, 2014). Seaweeds and marine organisms feeding on them live in a close association with diverse and abundant microbial communities (King et al., 2012; Hollants et al., 2013; Egan et al., 2013; Dudek et al., 2014). Seaweeds comprise dynamic species-specific bacterial communities (Aires et al., 2015; Aires, Serrão & Engelen, 2016; Vieira et al., 2016). The communities are recognized to have growth-promoting and nutritional effects (Head & Carpenter, 1975; Dimitrieva, Crawford & Yüksel, 2006), and to be involved
in the production of biologically active (Chojnacka et al., 2012) and defensive (Burgess et al., 1999) compounds. At a higher trophic level, symbiotic bacteria inhabiting the guts of marine herbivores are also known to support important physiological functions (Hacquard et al., 2015), including the mediation of the digestion of food components by producing critical digestive enzymes for breaking down complex molecular structures (Mackie et al., 2004). In addition to digestive functions, grazers depend on seaweed-associated microbiota for nutrients found in the algal biofilm (i.e., proteins, polysaccharides, lipids, etc.; Tietjen, 2014). As such, diet represents an important factor in shaping microbial diversity in the intestinal systems of grazers. So, any changes in bacterial composition of the seaweed may result in diet-induced changes in the gut microbiota of grazers that may eventually affect their metabolism, as well as its fitness and biology (Mattila et al., 2014; Tietjen, 2014). Because carbon acquisition is expected to be facilitated for seaweeds at elevated CO₂ levels, higher nutrient uptake is anticipated to help obtain other nutrients in the right balance with carbon. Part of these nutrients might be obtained through the microbiome and, therefore, the specific bacteria responsible for such acquisitions (e.g., phosphorous, nitrogen and iron) (Thomas et al., 2008; Burke et al., 2011b) might be positively selected and increase their abundance.

Because OA is expected to affect the interactions between marine herbivores and seaweeds through increased consumption of carbon enriched algal tissue (Gutow et al., 2014), the microbiomes of grazers might help with nutrient acquisition. While better understanding of the diversity and functions of associated symbiotic bacteria is needed, few studies have addressed the diversity and composition of gut microbiota of marine grazers (but see Hong et al., 2011; Devine, Pelletreau & Rumpho, 2012; Davis et al., 2013; Dudek et al., 2014).

To predict the responses of aquatic organisms to OA, it is necessary to understand responses of the host-associated microbiota to increasing CO₂ and reduced pH. Little is known about the responses of associated microbiota to changes in pCO₂ (partial pressure of carbon dioxide) including microbial metabolic capabilities or the ability to rapidly shift the host range (Morrow et al., 2015). Also, there is no consensus regarding whether a decrease in pH causes increase (Kerfahi et al., 2014), decrease (Taylor et al., 2014) or no changes (Hassenrück et al., 2016) in microbial richness and prevalence of dominant microbial taxa under acidification conditions. Furthermore, the current knowledge of acidification effects on the host-associated microbial communities is mostly based on the results of experiments conducted on corals. These experiments demonstrated that reduced pH initiates shifts in the coral microbiota towards microorganisms associated with stress and disease (Thurber et al., 2009; Meron et al., 2011; Webster et al., 2013). Therefore, there is a need for relevant studies on seaweeds, with a particular interest in species interactions as species may not respond similarly to OA and as effects can act synergistically. This response is particularly relevant to be examined in marine introduced seaweeds, because they are expected to benefit from future OA conditions, and thus increase their invasiveness.

In this study, we experimentally investigated the effects of acidification on the microbiomes of an emblematic invasive seaweed, the brown alga *Sargassum muticum*, and the gut microbiome of a native isopod consumer, *Synisoma nadejda*. This was done by
following a three-week mesocosms exposure to elevated pCO₂ followed by 16S amplicon sequencing in order to compare the bacterial community (hereafter microbiome) responses in these two hosts. Based on bacterial community characterization, our main hypotheses are that in acidified conditions (1) the seaweed-associated microbiome will have a different composition, (2) the grazers' gut microbiome will mirror the changes in food source (assuming that seaweed nutritional content will change with acidification), when compared with ambient conditions. Considering the existing evidence for corals responses to OA (Thurber et al., 2009; Meron et al., 2011; Webster et al., 2013) and seaweeds responses to other environmental stresses (e.g., temperature, Case et al., 2011; Mensch et al., 2016) we expect seaweed microbiome to shift towards a community composed by stress related bacteria and putative pathogens. Also, raw plant consumers' (e.g., fish, humans, etc.) gut microbiome and health is directly affected by environmental conditions (Sullam et al., 2012) or, following the “you are what you eat” premise, indirectly through changes in their food source/quality (Meziti et al., 2010; Berg et al., 2014), we predict that host's “normal” gut bacterial composition could be affected by OA. An increased abundance of bacteria potentially assisting digestion could provide the isopod with an elevated algal digestion and availability of inorganic compounds to compensate the shifted C/N ratio under acidification in their food. Furthermore, if S. muticum showed a shift under OA conditions, we will investigate the presence of possible bacterial taxa that could assist the seaweed obtaining nutrients (e.g., nitrogen fixing bacteria) under elevated pCO₂ conditions. These predictions, for both seaweed and gut microbiome, are limited to 16S taxonomic assignments and literature description and inherent reservations to this method will be considered.

# METHODOLOGY

## Experimental set-up

The experiment was performed at Centro de Ciencias do Mar (CCMAR) field station (Ramalhete) during the spring of 2014. Ambient (380 ppm CO₂, and pH 8.16—global levels of today's CO₂ conditions) and elevated pCO₂ (1,000) ppm CO₂, and pH 7.86—the year 2100 predictions by IPCC, A1FI scenario (Houghton et al., 2001) conditions were controlled by two separate CO₂ sensors systems. For acidified conditions, CO₂ was injected in seawater deposits that provided seawater for the experimental units. In both systems salinity was 36, alkalinity 2,550 µmol kg⁻¹ and seawater temperature 15 °C. Experimental units consisted of 3 L flowthrough mesocosms receiving each 30 L of seawater per hour. Experimental units were placed in one square meter tanks with 15 cm of the overflown seawater of the experimental units to stabilize temperature conditions in the units. *Sargassum muticum* and *Syniosoma nadejda* were sampled independently and kept isolated. Thus, the grazers were ‘naive’ and not previously on a diet containing *S. muticum*. After sampling, seaweeds and isopods were acclimated to ambient conditions in separate tanks for 1 week. A wet weight biomass of 1 g seaweed per experimental unit was used as a preliminary test and showed it did not affect the pH conditions in experimental units at the used volume and flow.

The two factors considered for the experiment were (1) CO₂ conditions (two levels: 380 and 1,000 ppm), and (2) host conditions (three levels: seaweed with grazers, seaweed
<figure><img src="image_3.png" /><figcaption>**Figure 1** Schematic representation of the mesocosms experiment. (A) Ambient (380 ppm) and (B) acidified (1,000 ppm) conditions each with four 3 L experimental units only containing 1 g wet weight (WW) *S. muticum* and four 3 L experimental units containing 1 g WW *S. muticum* along with the grazer *S. nadejda* were placed randomly in each CO₂ treatment. Each unit represented a replicate from which sample(s) (seaweed or seaweed and grazer) were taken.</figcaption></figure>

Full-size DOI: 10.7717/peerj.4377/fig-1

<figure><img src="image_4.png" /></figure>

without grazers, the grazers that fed on the seaweed). These were combined in a factorial design (all possible combinations of all levels of the two factors). The number of replicates within treatment combination was four. Therefore, in total, 24 samples (2 CO₂ × 3 hosts × 4 replicates) were collected (Fig. 1). The experiment ran for three weeks which was estimated sufficient for microbiomes to reflect the applied conditions considering the high turnover rate of food in the grazer gut and the multiplication rate of bacteria. During this period the units were cleaned, twice a week, to avoid epiphyte overgrowth on the experimental unit walls. In each experimental unit, seawater pH and the calibration of the automated CO₂ injection system was manually checked daily to make sure the pH was stable. At the end of the three weeks, the isopods and seaweeds were flash frozen, transported to the laboratory in liquid nitrogen and stored there at -80 °C until further processing.

## High-throughput sequencing of the microbiome

For both seaweeds and isopods's guts (also refered to as grazers, hereafter), DNA was extracted from all the 24 replicates using the Quick-gDNA kit (Zymo Research™, Irvine, CA, USA) according to the manufacturer protocol for “Solid Tissue Samples” (page 4 of the manual). Before extraction, isopods where dissected by removal of both ends and pulling out the intestinal tract. The total 16S rRNA was amplified using the universal primers 27F and 1492r with the following changes to the original protocol (Lane, 1991):
an initial denaturation at 95 °C for 2 min, 35 cycles of denaturation at 95 °C for 20 s, annealing at 55 °C for 20 s, and extension at 72 °C for 90 s, with a final extension was at 72 °C for 3 min. The 25 µl reaction mixture contained 250 µM dNTPs, 0.6 µM of each primer, 1 × 2 PCR buffer mix, 2 µl of template DNA (with a final concentration of about 10 ng µl⁻¹) and 0.3 µl of Taq polymerase (Advantage R2; Clontech, Mountain View, CA, USA). PCR products were cleaned using ExoFastAP enzyme following the Thermo Scientific™ protocol. Amplified DNA was sent to Molecular Research (MR DNA), Shallowater, Texas, where a nested-PCR was performed prior to sequencing. The modified 8 bp key-tagged primer 799F along with the reverse primer 1193R, covering the regions V5-V7 from 16S rRNA and amplifying a fragment of ~400 bp, were used to avoid chloroplast cross amplification (Bodenhausen, Horton & Bergelson, 2013). PCR conditions were as follow: 95 °C for 3 min, 10 cycles of 95 °C for 20 s, 50 °C for 30 s, 72 °C for 30 s, and a final elongation of 72 °C for 3 min. All amplifications and sample preparation procedures were the same for both seaweeds and isopodes. Samples were pooled together in equal proportions based on their molecular weight (calculated based on the size of the amplicon) and DNA concentrations (using Qubit™; Invitrogen®, Carlsbad, CA, USA) and purified using calibrated Agencourt® AMPure® XP beads. DNA libraries were prepared by following Illumina TruSeq DNA library preparation protocol and paired-end (2 × 250 bp) sequencing performed at MR DNA (http://www.mrdnalab.com; Shallowater, TX, USA) on a MiSeq following the manufacturer's guidelines.

Detailed protocols for sampling procedures, DNA extraction and PCR amplification mentioning all the important measures to avoid contamination, can be found in (Aires et al., 2018).

## Sequence analysis and bioinformatics

A total of 3,204,094 partial 16S rRNA gene sequences were obtained from the 24 samples (i.e., two CO₂ conditions X four replicates for seaweed in the presence and the absence of grazer and four grazer gut samples with two CO₂ conditions). The bacterial community analyses were performed using Quantitative Insights into Microbial Ecology (QIIME version 1.8.0) software (Caporaso et al., 2010). Sequences were screened and filtered for a minimum read length of 350 bp (after reads were paired) and less than two undetermined nucleotides. Selected high-quality sequences were clustered into Operational Taxonomic Units (OTUs) within reads using denovo OTU picking method. Representative sequences for each OTU were selected using the “most-abundant” method and OTU sequence alignment was carried out using PyNAST (Caporaso et al., 2010) and Greengenes v.13.8 (McDonald et al., 2012). Taxonomic assignments were done using the UCLUST (Edgar, 2010) method with a 97% confidence threshold. To assign each OTU to the closest matching described taxon, searches were performed against the Greengenes taxonomy database v.13.8 for 16S rRNA (McDonald et al., 2012), and sequences were putatively assigned to a described taxon with a minimum threshold of 0.001 (default value). Eukaryotes (i.e., chloroplasts and mitochondria) matching sequences were excluded from the OTU table in downstream analyses as well as rare OTUs (singletons and doubletons) and unassigned sequences (those sequences that did not match any of those from the Greengenes database, with a minimum threshold of 0.001).
Quality filtering resulted in 2,877,493 high-quality sequences, with an average of 119,896 ± 46,294 reads per sample, which were clustered into 42,730 unique operational taxonomic units (OTUs). The OTU table was rarefied to the minimum number of sequences (66,831). As a result, a total of 41,139 unique OTUs remained. Public access to the data can be done through: https://doi.org/10.6084/m9 figshare.5346316.v3.

All the statistical and diversity (alpha and beta) analysis were done using the filtered rarefied (to the minimum number of sequences—66,831) OTU table and considered significant at *P* < 0.05.

Alpha diversity indexes, including Chao *I* richness (Chao, 1984), observed number of species (OTUs) and Shannon diversity, were calculated using QIIME software. Bacterial community structure (beta diversity) was assessed by permutational multivariate analyses of variance (PERMANOVA) using Bray–Curtis dissimilarity matrices from square-root transformed data. PERMANOVA tested for differences among samples with different levels of *a priori* factors: Type of sample: Seaweed vs Grazer gut; for both CO₂ treatments: Ambient vs Acidified; for Seaweed: Grazed vs Non-grazed, and the interactions among these factors. The homogeneity of multivariate dispersions (based on mean distance to group centroid for all groups within each factor) was tested using a resemblance based permutation test (PERMDISP). To visualize differences and to assess dissimilarity between samples, Canonical Analysis of Principal coordinates (CAP) plots were constructed to test the assignment/clustering of treatments interaction (*S.muticum* XGrazingxAcidification and Grazer gutxAcidification) as *a priori* factor. Similarities and dissimilarities in bacterial communities between acidification treatments were explored using, similarity percentage analyses (SIMPER). For those bacterial taxonomic groups that displayed a high contribution (concerning their differential abundances in the treatments being compared) for the differences between the grazer gut and the seaweed and the two CO₂ levels, two-way analyses of variance (ANOVA) were performed (with the preliminary tests for normality and homogeneity of variances being implemented). Species (*S. muticum* and isopod) and acidification (CO₂ ambient and elevated) were tested as factors affecting the structure/composition bacterial communities. For bacterial OTUs for which significant interaction were detected, a *post-hoc t*-test was implemented using a Bonferroni correction and a conservative alpha (considering the comparisons made: CO₂ effect in *S. muticum*, CO₂ effect on isopod gut), effect of type of tissue (seaweed/gut) in ambient CO₂ and effect of type of tissue in elevated CO₂ (*P* (*T* ≤ *t*) two tail < 0.0125).

All bacterial community structure statistical analyses were performed using the software program PRIMER-E + PERMANOVA v.6 (Clarke & Warwick, 1994; Clarke & Gorley, 2006).

RESULTS

Bacterial communities associated with *S. muticum* and *S. nadejda's*
gut

Overall, 563 bacterial OTUs from 74 classes (apart from bacteria classified as ‘Other and non-ID’) distributed across 28 phyla (plus ‘Other and non-ID’) were identified. Among them, 551 bacterial OTUs distributed across 27 phyla (plus Other and non-ID) were
<figure><img src="image_5.png" /><figcaption>**Figure 2 Community structure.** Plot of canonical analysis of principal coordinates (CAP) based on Bray–Curtis distances calculated on square-root transformed bacterial abundances, showing the axes that best discriminate the bacterial assemblages across CO₂ levels (blue-ambient *versus* red-acidified), grazing by *S. nadejda* on *S. muticum* (open squares-grazed seaweed vs filled squares-non-grazed seaweed) and the gut of the isopod on a diet of *S. muticum* (circles).</figcaption></figure>

Full-size

<figure><img src="image_6.png" /></figure>

DOI: 10.7717/peerj.4377/fig-2

associated with *S. muticum* (over all samples and treatments), compared to 450 OTUs distributed across 22 phyla (plus Other and non-ID) for gut biome. Note that those sequences labelled as “others” were ambiguous assignments by the classifier and “no-ID” sequences result from a good match with a reference sequence but that reference sequence is poorly defined (not named at a certain taxonomic level and below). Public access to the data can be done through: https://doi.org/10.6084/m9.figshare.5346316.v3.

Alpha diversity of associated bacterial communities (Table S1) was similar or slightly higher in *S. muticum* than in the grazer gut for Shannon index (2-way ANOVA, *F* = 2.507, *P* = 0.139), OTU richness (2-way ANOVA, *F* = 3.361, *P* = 0.092) and Chao 1 (2-way ANOVA, *F* = 6.226, *P* = 0.028), respectively. Acidification did not affect bacterial diversity at OTU level, for both seaweeds and isopodes gut, as estimated by diversity indexes: Shannon index (2-way ANOVA, *F* = 0.048, *P* = 0.831), unique OTU richness (2-way ANOVA, *F* = 0.178, *P* = 0.681) and Chao 1 (2-way ANOVA, *F* = 0.003, *P* = 0.960).

Despite the overall PERMANOVA results showing that bacterial community composition was significantly different for the different type of samples (grazed and non-grazed *S. muticum* and grazer gut, *P* = 0.001, Fig. 2, Table S2), predation (grazing) did not significantly change bacterial community structure of *S. muticum* (*P* = 0.372, Table S3A). The main differences among *S. muticum* (grazed and non-grazed) and the isopode gut (Table S2) were due to a higher abundance of *Bacteroidetes* (73.7%) associated to *S. muticum* (contributing 15.11% to the dissimilarity; SIMPER analysis) and to more *Proteobacteria* (47.2%) and *Planctomycetes* (32.4%) in the gut of *S. nadejda*, contributing 11.76 and 14.98% to the dissimilarity, respectively (SIMPER analysis).
<figure><img src="image_7.png" /><figcaption>**Figure 3** Host and treatment effects on associated bacteria phyla. Relative abundance and distribution of the bacteria phyla associated to the brown seaweed *Sargassum muticum*, without (No grazing) and with (grazing) *Synisoma nadejda* isopods, and the gut of the isopod after three weeks on a *Sargassum muticum* diet, under ambient (380 ppm) and elevated/acidified (1,000 ppm) CO₂ conditions.</figcaption></figure>

Full-size

<figure><img src="image_8.png" /></figure>

DOI: 10.7717/peerj.4377/fig-3

Bacterial phyla-specific to host (referred in PERMANOVA analysis as “Type of sample”) and specific conditions are presented in Table S4.

As mentioned before, grazing did not show significant effects on *S. muticum* bacterial community structure so, most of further analyses focused on the comparison of grazed *S. muticum* vs grazer gut. Grazed *S. muticum* associated bacterial communities were dominated by *Bacteroidetes* (75.4%, Fig. 3; *Flavobacteriia* 75.2%) and *Proteobacteria* (16.2%, Fig. 3; *Alphaproteobacteria* 12.2%), while the isopod gut communities were dominated by *Proteobacteria* (47.2%, Fig. 3; *Alphaproteobacteria* 43.5%), *Planctomycetes* (32.4%, Fig. 3; *Planctomycetia* 32.1%), and *Bacteroidetes* (17.7%, Fig. 3; *Flavobacteriia* 16.8%). *Flavobacteriales* (73.3%) was the most common bacterial order associated with *S. muticum*, while *Rickettsiales* (38.4%), *Pirellulales* (30.9%) and *Flavobacteriales* (16.8%) were the most abundant grazer gut-associated orders. The distribution of the bacterial genera belonging to the main phyla found for the two different systems and described above (*S. muticum*—*Bacteroidetes* and *S. nadejda* gut—*Proteobacteria* and *Planctomycetes*) showed a clear higher relative abundance of genera assigned as Non-ID and “other” *Flavobacteraceae* (Fig. 4A) for the seaweed microbiome and a isopode gut dominated by *Neorickettsialles* (Fig. 4B) and non-identified *Pirellulaceae* (Fig. 4C).

Several bacterial orders were significantly more abundant within grazed *S. muticum* than in the isopod gut: *Flavobacteriales* from the phylum *Bacteroidetes* (2-way ANOVA, *F* = 49.124, *P* < 0.001); *Bdellovibrionales* from the class *Deltaproteobacteria* (2-way ANOVA, *F* = 13.221, *P* = 0.003); as well as *Acidithiobacillales* (2-way ANOVA, *F* = 22.589, *P* = 0.0005), *Alteromonadales* (2-way ANOVA, *F* = 15.285, *P* = 0.002), and *Oceanospirillales* (2-way ANOVA, *F* = 6.407, *P* = 0.026) from the class *Gammaproteobacteria* (Fig. 5). In contrast, the isopod gut microbiome had higher abundances of *Rickettsiales* from the class
<figure><img src="image_9.png" /><figcaption>**Figure 4** Relative abundance of the genera belonging to the main bacterial phyla. (A) Bacteroidetes, (B) Proteobacteria, and (C) Planctomycetes, associated with the brown seaweed *Sargassum muticum* grazed by *Synisoma nadejda* isopods (left side), and the gut microbiome of the isopod on a *Sargassum muticum* diet (right side), after three weeks under ambient (380 ppm; -CO₂) and elevated/acidified (1,000 ppm; +CO₂) CO₂ conditions.</figcaption></figure>

Full-size

<figure><img src="image_10.png" /></figure>

DOI: 10.7717/peerj.4377/fig-4

*Alphaproteobacteria* (2-way ANOVA, *F* = 18.574, *P* = 0.001); the low abundance bacteria from the phylum *TM6*-class *SJA-4* (2-way ANOVA, *F* = 12.491, *P* = 0.004); and the low abundance order *AKAU3564-Phycisphaerae* from the phylum *Planctomycetes* (2-way ANOVA, *F* = 7.325, *P* = 0.019) than *S. muticum* in the presence of grazer (Fig. 5).

## Bacterial diversity and composition under acidification conditions

Acidification did not affect the overall bacterial composition (PERMANOVA, *P* = 0.093, Table S2) and in particular that associated with *S. muticum* (*P* = 0.056, *P* = 0.584, Table S3B), but significantly affected the bacterial composition associated with the gut of *S. nadejda* (PERMANOVA, *p* = 0.022, Table S3B) (Fig. 2). However, under acidified conditions (and in the presence of grazers), the number of phyla in *S. muticum* bacterial community dropped from 26 to 17 (including one unidentified), compared to the ambient conditions (One-way ANOVA, *F* = 7.228, *P* = 0.009). In contrast, the number of phyla in the grazer gut-associated bacterial community under acidification treatment increased from 18 to 23 (including one unidentified), compared to the ambient conditions (One-way ANOVA, *F* = 1.923, *P* = 0.171).
<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /></figure>

**Figure 5** Mean relative abundances of bacterial classes, and respective orders (A, *Flavobacteriales* and Ricketsiales; B, *Bdellovibrionales* and *Oceanospirillales*; C, *Acidithiobacillales*; D, *Alteromonadales*; E, Non-ID SJA-4 and AKAU3564 *Phycisphaerae*), significantly more abundant in either grazed *Sargassum muticum* or the gut of *Synisoma nadejda*. After three weeks under ambient (380 ppm) and elevated/acidified (1,000 ppm) CO₂ conditions. Alpha = 0.05, error bars show standard error per treatment (n = 4).

Full-size

<figure><img src="image_16.png" /></figure>

DOI: 10.7717/peerj.4377/fig-5

The phyla specific to the microbiomes of *S. muticum* and gut of *S. nadejda*, under ambient vs acidified conditions, are presented in Table S4.

Some bacterial groups were unique to by their hosts under specific conditions (Fig. 6). The seaweed, in the presence of grazer and under the ambient conditions, had the highest
<figure><img src="image_17.png" /><figcaption>**Figure 6** Core communities. Venn diagram representing the number of bacterial genera (present in at least 75% of samples) shared between the different CO₂ treatments (ambient CO₂–↓CO₂; elevated/acidified CO₂–↑CO₂) and associated to grazed *Sargassum muticum* (Sm + G), and the gut microbiome of the isopod *Synisoma nadejda* on a *Sargassum muticum* diet. The bar plots show the distribution of Phyla of selected intersections.</figcaption></figure>

Full-size DOI: 10.7717/peerj.4377/fig-6

<figure><img src="image_18.png" /></figure>

number of unique bacterial OTUs (n = 28, 5%), while the grazer gut in the ambient conditions had the lowest number of unique bacterial OTUs (n = 2) (Fig. 6). *S. muticum* with the grazer present under the ambient conditions contained many OTUs belonging to *Planctomycetes* (3 OTUs, 24.8%), *Proteobacteria* (9 OTUs, 19.8%) and *Bacteroidetes* (9 OTUs, 19.5%) (Fig. 6). *S. muticum* with the grazer, but under acidification treatment, contained 13 unique OTUs, most of which belonged to *Proteobacteria* (7 OTUs, 74.9%) (Fig. 6). *Sphingobacterium* (*Bacteroidetes*) and *Caldicoprobacter* (*Firmicutes*) were unique to the gut of *S. nadejda* in the ambient conditions, compared to 23 OTUs unique to the grazer gut in the acidified conditions, a significant part of which belonged to *Planctomycetes* (2 OTUs, 50.7%) and *Proteobacteria* (8 OTUs, 15.8%) (Fig. 6).

A core bacterial community (present in at least 75% of samples) was composed of 181 bacterial OTUs (32.1%; including unidentified) (Fig. 6). Within this core bacterial
community, the highest number of OTUs belonged to *Bacteroidetes* (n = 29; 55.3%), *Proteobacteria* (n = 90; 26.7%) and *Planctomycetes* (n = 7; 13.2%) (Fig. 5). Shared bacterial communities within *S. muticum* in the presence of grazer between the ambient and acidified conditions (n = 29; 5.2%) had the highest number of bacterial OTUs belonging to *Proteobacteria* (n = 14; 47.2%), *Firmicutes* (n = 8; 19.9%), and *Bacteroidetes* (n = 5; 18.5%) (Fig. 6). Shared bacterial communities in the gut of *S. nadejda* between the ambient and acidified conditions (n = 9; 1.6%), belonged mostly to *Planctomycetes* (n = 3; 54.8%) and *Proteobacteria* (n = 4; 32.3%) (Fig. 6).

Under elevated CO₂ treatment, relative abundances of *Gammaproteobacteria* from the orders *Oceanospirillales* and *Vibrionales* (particularly *Pseudoalteromonas*), increased on *S. muticum* but only significantly in the absence of grazers (One-way ANOVA, *P* = 0.037 and *P* = 0.047, respectively; Figs. 7A and 7B). Under acidification, the abundance of *Planctomycetia* associated with the grazer gut was significantly higher than at the ambient CO₂ levels (Two-tailed *t*-test, *P* < 0.001), as was their abundance (at elevated CO₂) in the grazer gut than on *S. muticum* (Two-tailed *t*-test, *P* < 0.001) (Fig. 7C). Deeper analyses of bacterial abundance within *Planctomycetia* revealed that all bacterial orders detected within this class responded to acidification treatment (Figs. 7C–7E). A significant effect was confirmed within the grazer gut under different CO₂ conditions and between the seaweed and the grazer gut under increased CO₂. At elevated CO₂ levels, the abundance of *Pirellulales* (Two-tailed *t*-test, *P* < 0.001, Fig. 7C), *Planctomycetales* (Two-tailed *t*-test, *P* = 0.008, Fig. 7D) and *Gemmatales* (Two-tailed *t*-test, *P* = 0.009, Fig. 7D), were increased in the isopode gut. Alongside, under the acidification treatment, the abundance of the bacterial orders *Pirellulales* (Two-tailed *t*-test, *P* < 0.001, Fig. 7C), *Planctomycetales* (Two-tailed *t*-test, *P* < 0.001, Fig. 7D) and *Gemmatales* (Two-tailed *t*-test, *P* = 0.008, Fig. 7D), was significantly higher in the grazer gut than associated with the seaweed (in the presence of grazers). The class *C6-Planctomycetes* also exhibited a significant increase in abundance under elevated CO₂ conditions, but only within the grazer gut (Two-tailed *t*-test, *P* = 0.039) (Fig. 6). This was mostly due to the significant increase of bacteria from the order *d113* (two-tailed *t* test, *P* = 0.039) (Fig. 6). The abundance of *d113* under acidified conditions was significantly higher in the grazer gut than within *S. muticum* (two-tailed *t* test, *P* = 0.018) (Fig. 6). Overall, under acidification treatment, *Planctomycetes* increased in abundance (particularly within *Planctomycetia*) in the grazer gut from 20.9% to 43.8% (1-way ANOVA, *F* = 306.663, *P* < 0.001).

DISCUSSION

The results presented in this experimental study demonstrated that acidification affected specific bacterial groups, but hardly influenced the overall microbiome of the invasive brown seaweed *S. muticum*. In contrast, acidification caused significant changes in the gut microbiome of a native isopod consumer, *S. nadejda*. Interestingly, acidification increased abundances of *Planctomycetia* in the gut of *S. nadejda* and of *Oceanospirillales* and *Vibrionales* associated to *S. muticum*, raising hypotheses about their functional role under these conditions.
<figure><img src="image_19.png" /></figure>

<figure><img src="image_20.png" /></figure>

<figure><img src="image_21.png" /></figure>

<figure><img src="image_22.png" /></figure>

<figure><img src="image_23.png" /></figure>

**Figure 7** Mean relative abundances of associated bacterial orders. (A and B) *Sargassum muticum* under grazing/non grazing influence after three weeks under ambient (380 ppm) and elevated/acidified (1,000 ppm) CO₂ conditions and (C-E) grazed *Sargassum muticum* and the gut of *Synisoma nadejda* after three weeks under ambient (380 ppm) and elevated/acidified (1,000 ppm) CO₂ conditions, that responded to acidification, but for which a significant interaction between acidification and type of sample (seaweed or grazer gut) was observed. Alpha = 0.05, error bars show standard error per treatment (n = 4).

Full-size

<figure><img src="image_24.png" /></figure>

DOI: 10.7717/peerj.4377/fig-7
The guts of isopods are populated by symbiotic bacteria (e.g., Zimmer & Bartholmé, 2003; Wang, Brune & Zimmer, 2007; Fraune & Zimmer, 2008; Eberl, 2010) that assist in food utilization by the host (Zimmer, 2002; Zimmer et al., 2002; Zimmer & Bartholmé, 2003; Fraune & Zimmer, 2008). Because the diversity of gut bacterial communities is shaped by host diet (Tietjen, 2014), changes in seaweed characteristics and its microbiome, as a result of ocean acidification, were expected to affect the gut microbiome of the grazer with possible fitness consequences. Here, we documented that the most striking change resulting from acidification was a significantly increased abundance of *Planctomycetes* in the gut of *S. nadejda*, more specifically due to an increase of Non-ID *Pirellulaceae*. In this case, as seaweed microbiome was not overall affected by OA, these general changes in the grazer's gut cannot be directly attributed to changes in its food microbiome. Although the functions of *Pirellulaceae* are hardly known, its presence was documented in the gastrointestinal tract of fish (Parris et al., 2016) and as part of the resident microbiomes of marine copepods but in very low abundance in starved specimens (Moisander, Sexton & Daley, 2015). *Planctomycetes* constituted the second most prevalent phylum (after *Proteobacteria*) in the gut of *S. nadejda*. These bacteria are known to widely colonize aquatic and terrestrial ecosystems (Lage & Bondoso, 2014) and were, until recently, considered environmental organisms. However, *Planctomycetes* have been reported associated to the gut of, not only marine taxa (Singh & Reddy, 2014), but also terrestrial herbivores (Fuerst & Sagulenko, 2011), mammals (Frey et al., 2006) and even humans (Cayrou et al., 2013).

In this study, acidification strongly increased the abundance of *Planctomycetes* in the bacterial gut communities of *S. nadejda* compared to ambient conditions. An increase of this Phylum in response to simulated OA has also been shown in sandy sediments (Currie et al., 2017). Although it is clear that in our study this was a consequence of acidification, the lack of seawater samples does not allow us to determine whether the observed *Planctomycetes* increase was due to acidification of the seawater or a direct response of the grazer's gut to acidification. This phylum was also abundant (the third most abundant) in the bacterial community associated to *S. muticum*. These widespread bacteria have been often found associated to macroalgae (Lage & Bondoso, 2014), among which *Caulerpa taxifolia* (Meusnier et al., 2001) and the kelp *Laminaria hyperborea* (Bengtsson, Sjøtun & Øvreås, 2010). Bacteria from the phylum *Planctomycetes* are suggested to have potential benefits for their hosts, through their ability to mineralize organic molecules into inorganic compounds that match the nutritional requirements of macroalgae (Lage & Bondoso, 2014). *Planctomycetes* are also proposed to function as “slow-acting decomposers of organic matter” (i.e., algal polymer degradation; Bodelier & Dedysh, 2013) and important contributors to the global nitrogen cycle (i.e., anammox *Planctomycetes*, Fuerst & Sagulenko, 2011). So, the presence of these bacteria in the isopod gut might also be related to the ingestion of *S. muticum*. Its increased abundance (particularly those from the order *Pirellulales*) could also be a response to the change of the seaweed nutritional value under acidification, which likely increased its photosynthetic rate driven by high CO₂ and consequent C/N ratio shifts (Cornwall, Revill & Hurd, 2015; Briggs, 2017). An increase in *Planctomycetes* abundance could provide the isopod with an elevated algal digestion capacity to compensate the highly carbonated food. Again, with the lack of control samples
as starving isopods (empty guts) we can only raise new hypotheses about the direct (through its microbiome) or indirect (through its nutritional value/quality) influence of the food in the grazer's gut microbiome. Yet, there is no doubt that acidification resulted in a significant overall change in the isopod gut microbiome that is not directly related to bacterial community shifts of *S. muticum*, which did not occur for the seaweed at elevated CO₂.

Another important change resulting from acidification was a significant increase in *Oceanospirillales* (Non-ID *Oceanospirillaceae*) and *Vibrionales* (*Pseudoalteromonas*) associated with *S. muticum* in the absence of grazers. They all belong to *Gammaproteobacteria* which have been previously found in association with various seaweed species (e.g., Patel et al., 2003; Huggett et al., 2006). Members of the class *Gammaproteobacteria* are known to produce biologically active metabolites that mediate antifungal (Barbieri et al., 2001), antifouling (i.e., Alteromonas, Pseudomonas; Maki et al., 1988; Holmstrom, Rittschof & Kjelleberg, 1992; Avelin Mary et al., 1993; Holmström & Kjelleberg, 1999) and antibacterial activities (Hentschel et al., 2001). There are no studies documenting the effect of acidification on these bacteria in seaweeds, but environmental samples showed a high increase of *Gammaproteobacteria* under acidification, in particular from the order *Oceanospirillales* (Currie et al., 2017). Bacteria from the order *Oceanospirillales* are heterotrophic and capable of degrading complex organic compounds (Garrity et al., 2005; Goffredi, Johnson & Vrijenhoek, 2007). These bacteria use organic matter as food source and their increase in *S. muticum* under acidification might be opportunistic and related to the carbon content increase in the seaweed, and consequent seaweed enrichment, due to elevated CO₂ levels.

Increased abundance of *Vibrionales* has often been associated with stressed and diseased marine invertebrates and they are also known as coral pathogens (Bourne & Munn, 2005; Bourne et al., 2008; Sunagawa et al., 2009; Meron et al., 2011). *Vibrionales* were also responsible for a number of infections in humans and animals (Vezzulli et al., 2016), and identified as potential pathogens of sablefish larvae (Schulze et al., 2006) and bivalve mollusks (Asplund et al., 2014). Interestingly, it has been shown that under low pH, the coral-associated pathogen *Vibrio* sp. increased in abundance (Meron et al., 2011), while the blue mussel pathogen *Vibrio tubiashii* became more infectious (Asplund et al., 2014). In this study, among the *Vibrionales* that experienced a significant increase were predominantly *Pseudoalteromonas*. While certain members of the genus *Pseudoalteromonas* were reported to have antibacterial activity in corals, providing it with defense against potential pathogens (Shnit-Orland, Sivan & Kushmaro, 2012), multiple studies identify *Pseudoalteromonas* as opportunistic pathogens of marine organisms (Liu et al., 2010; Song et al., 2012; Wang et al., 2012). Bacteria affiliated to the genus *Pseudoalteromonas* were found associated to the kelp *Laminaria japonica* affected by two different diseases (holle-rotten disease and red spot disease) (Sawabe et al., 1998; Gachon et al., 2010) where its isolation and posterior reinfection resulted in observable symptoms (Wang et al., 2008). Nevertheless, particular bacteria that may be otherwise commensal, under stress of the seaweed host, can become saprophytic (Egan et al., 2013) and this could hypothetically be the case of the *Pseudoalteromonas* found in *S. muticum* under acidification. Ocean acidification could potentially result in shifts from healthy associated bacterial communities within seaweeds
towards a higher prevalence of pathogenic bacteria and/or increased vulnerability to disease. Unfortunately, our taxonomic assignments and our amplicon sequencing approach does not provide more detailed insights into possible bacterial pathogenicity.

*Flavobacteriales* from the *Bacteroidetes* phylum were among the bacterial groups that were significantly more abundant in association with *S. muticum* than in the isopod gut. *Bacteroidetes* colonize marine and freshwater environments widely (Thomas et al., 2011), populating a wide variety of surfaces, including macroalgae (Beleneva & Zhukova, 2006; Staufenberger et al., 2008) and marine sediments (Devine, Pelletreau & Rumpho, 2012). *Bacteroidetes* were isolated from *Caulerpa taxifolia* (Meusnier et al., 2001), *Ulva australis* and the red alga *Delisea pulchra* (Longford et al., 2007), suggesting that these typical marine bacteria are common seaweed associates (Tujula et al., 2010). This phylum represents some of the most abundant marine bacteria (Glöckner et al., 1999; Simon, Glöckner & Amann, 1999; Cottrell & Kirchman, 2000) and plays an important role as degraders of complex organic matter (Church, 2008). *Flavobacteriia*, the most prevalent class detected within the seaweed microbiome, are known to produce enzymes for polymer degradation (Fenchel, 2012). Most *Flavobacteriia* are able to degrade cellulose, chitin, proteins, and nucleic acids (Kirchman, 2002; Fenchel, 2012). However, *Bacteroidetes* are also related to stress conditions and often found in diseased corals (Barneah et al., 2007), as in the case of *Porites compressa* which when exposed to low pH showed an increase of disease-associated *Flavobacteriia* (Thurber et al., 2009). *Flavobacteriales*, the most widely represented *Flavobacteriia* order within *S. muticum*, are mostly associated with degradation of complex particle biomacromolecules, as well as algal debris (Kirchman, 2002), more specifically proteins, agars, xylan, fucoidan, cellulose, and chitin (Devine, Pelletreau & Rumpho, 2012). Bacteria from the genera *Aquimarina* and *Tenacibaculum* were among the most frequent OTUs and occur free-living in marine environments (Nedashkovskaya et al., 2006) and fixed to the surfaces of marine organisms (Suzuki et al., 2001). Bacteria from the genus *Tenacibaculum* are thought to induce morphogenesis in algae and possibly enhanced seaweed growth (Matsuo et al., 2003; Matsuo et al., 2005).

In this study, acidification resulted in a small (non-significant) decrease of seaweed-associated *Flavobacteriales* and just in the absence of grazer. This contrasts with a study conducted on biofilms from the Great Barrier Reef, which reported that with decreasing pH there was an increase in the relative abundance of *Flavobacteriales* (*Flavobacteriaceae*) (Witt et al., 2011). However, *Witt et al.* (2011) focused on natural biofilms on glass slides whereas biofilms from living organisms are the result of environment and host communication and may be differently influenced by any fluctuations that might have occurred in the seawater bacterial communities. Nevertheless, the lack of seawater bacterial community analysis in our study may limit the interpretation of some of these results. The decrease of these common seaweed associates (*Hollants et al., 2013*; *Egan et al., 2013*), although non significant, may result from the instability of environmental pH changes. Natural seaweed bacterial assemblages can be disrupted by environmental pressures (*Marzinelli et al., 2015*) and that could contribute to the initial response of *S. muticum* to acidification.

Seaweed consuming iguanas, which have adapted to use macroalgae as their primary resource, were found to host a large proportion of *Bacteroidetes* in their gut when compared
to terrestrial related species (Hong et al., 2011). Seaweed polysaccharides, many with
sulfated sugars that are absent in terrestrial plants, are easily hydrolized by *Bacteroides* spp.
present in the gastrointestinal tract (Shah & Gharbia, 1993). In particular, these bacteria
have been described as contributing to the degradation of brown algal polysaccharides
in the gastrointestinal tract of limpets (particularly *Flavobacteriia*; Dudek et al., 2014) and
other gastropods that are seaweed consumers (Cardoso et al., 2012). As discussed above,
*Bacteroidetes* were among the most abundant bacteria associated with *S. muticum* and,
as shown on copepods (Moisander, Sexton & Daley, 2015), these were the third most
abundant bacterial phylum in the intestinal tract of *S. nadejda*. *Bacteroidetes* are within
the most abundant phyla in three different copepod species, for both starving and full gut
specimens (Moisander, Sexton & Daley, 2015). These authors also found these bacteria to
be the most abundant in seawater samples which lead to the assumption that they were not
"gut permanent residents" but instead colonizers from the seawater. In our case, the lack of
seawater samples and starved isopodes (empty gut) limit the interpretation of our results,
so its relatively high abundance in *S. nadejda* gut might be both related with seawater
and/or food.

In contrast to *Bacteroidetes* and *Gammaproteobacteria* prevailing within *S. muticum*, *Alphaproteobacteria* (*Rickettsiales*), as well as bacteria from the phylum *TM6* (the class *SJA-4*) were more abundant in the grazer gut microbiome for both ambient and acidified conditions. *Alphaproteobacteria* were also reported as one of the most prevalent Classes in marine copepods (*Moisander, Sexton & Daley, 2015*). Bacteria from the order *Rickettsiales*, which dominated the gut of *S. nadejda*, are known as pathogenic to humans and animals (*Perlman, Hunter & Zchori-Fein, 2006*), and were found in the intestinal tract of infected isopod *Armadillidium vulgare* (*Dittmer et al., 2016*) and several other isopode species (*Wang, Brune & Zimmer, 2007*). *Neorickettsia* is known as a parasite to which the invertebrates usually serve as vectors (*Greiman et al., 2014*). This aspect is worth further investigation once the possibility of these herbivores to function as pathogen vectors may have consequences on human health if they are able to pass them along to some edible algae where they feed on.

The experimentally controlled mesocosm data allowed us to isolate variables to infer
their effects, even though they are not able to fully reconstruct the dynamic conditions of
nature, as documented for corals that had different bacterial communities in laboratory and
field (Kooperman et al., 2007; Meron et al., 2011). So, the cause of the observed bacterial
community shifts can be easily identified but may not mimick exactly the processes in
the field. Another limitation to realistic predictions is that global acidification may be
happening quicker than IPCC models predicted. So, pH and CO2 conditions used in this
and several other studies may not be realistic and underestimate the effects of OA that can
be more catastrophic than what is expected to be in the next 100 years (the maximum
IPCC prediction) (Thurber et al., 2009; Meron et al., 2011). However, those changes would
happen over a large period of time and not over the course of three weeks, as in our
experiment.

The results of this study demonstrate that bacterial communities associated within
an isopod—seaweed predator–prey system are dynamic and responsive to changes in
acidification. The observed changes in the associated bacterial communities of the seaweed
and the grazer gut might be a type of acclimation that facilitates tolerance and survival,
as suggested by Meron et al. (2011). The capacity of organisms to accommodate and
modify a highly diverse microbiota may influence the fitness and success of the host and
enhance its ability to survive changing environmental conditions (Morrow et al., 2015).
Further research is required to better understand the processes and conditions under
which different associated bacteria can increase tolerance of the host organisms to various
disturbances.

CONCLUSION

The responses of bacteria associated with *S. muticum* and the gut of *S. nadejda* revealed
in this study suggest that worst case acidification scenarios may not greatly affect overall
bacterial community composition and diversity, but might affect specific bacterial groups.

Contrarily to what was expected, acidification to expected levels seems to have less
significant impact on seaweed bacterial ecology than other environmental stressors, such
as increased temperature (Mensch et al., 2016). However, specific groups had significant
abundance shifts in seaweeds under acidification as was the case for Oceanospirillales
and Vibrionales (mainly Pseudomonadales). The former might be related to the possible
increase of the C/N ratio in the seaweed under acidified conditions and the latter, commonly
found associated with diseased seaweeds, could be an indicator that acidification enables
an increase of opportunistic/pathogenic bacteria. Unexpectedly, no particular bacteria
increased abundance that could assist the seaweed in obtaining nutrients (like nitrogen
fixing bacteria) under a higher carbon availability regime.

In contrast with the seaweed host, high CO₂ levels globally changed the bacterial
community associated to the isopod gut, particularly the abundance of members of the
Class *Planctomycetia*. However, as no significant changes occurred in the global seaweed
microbiome, the overall shift in the grazer gut bacterial community cannot be directly
attributed to bacterial changes in the food. Instead, we can hypothesize that the food
“quality” changed at elevated CO₂ triggering a shift in the isopod gut *Planctomycetia*
allowing it to better digest the seaweed and compensate the shifted C/N ratio as seaweed
becomes less nutritional under acidification, as previously hypothesized. The generality of
these findings must be addressed in further studies with other species.

*Bacteroidetes* (mainly *Flavobacteriia*), previously isolated and commonly found
associated with other seaweed species (*Huggett et al., 2006*; *Burke et al., 2011a*; *Lachnit
et al., 2011*), were the most dominant phyla, suggesting that they account for core roles
in the metabolism of *S. muticum*. Both in ambient and acidified conditions, the isopod
gut bacterial community was dominated by *Neorickettsia* (*Alphaproteobacteria*), for which
invertebrates are usually the vectors. The pathogenicity of these bacteria has already been
shown in other invertebrate species (*Greiman et al., 2014*) and the potential of isopod
species as possible vectors for these bacteria has already been suggested (*Yuksel, Thompson
& Adams, 2006*) and should be further investigated.

Concluding, our results show that after only three weeks of simulated acidification,
bacterial communities associated to a seaweed host, when ungrazed, and to an isopod
grazer gut, show shifts in composition. These bacterial community changes were particular and specific in the seaweed and only occurred when ungrazed, but were large and global community shifts in the isopod grazer gut. We hypothesized that these may have potential consequences for seaweed health and isopod food digestion. The observed changes in the gut microbiome of the grazer seem to be a reflection of changes in the seaweed chemistry rather than its microbial composition.

## ACKNOWLEDGEMENTS

We would like to thank Dario Nobre and João Reis for conducting the acidification experiment.

## ADDITIONAL INFORMATION AND DECLARATIONS

### Funding

This study was made possible by the Erasmus Mundus Doctoral Programme MARES on Marine Ecosystem Health & Conservation (MARES_13_08: Acclimation and adaptation of invasive seaweeds) for Alexandra Serebryakova, and funds from FCT (Foundation for Science and Technology, Portugal) fellowships SFRH/BPD/63703/2009 and SFRH/BPD/107878/2015 to Aschwin H. Engelen and SFRH/BPD/116774/2016 to Tania Aires, and the EU SEAS-ERA project INVASIVES (SEAS-ERA/0001/2012) and CCMAR/Multi/04326/2013. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

### Grant Disclosures

The following grant information was disclosed by the authors:
Erasmus Mundus Doctoral Programme MARES on Marine Ecosystem Health & Conservation.

FCT (Foundation for Science and Technology, Portugal): SFRH/BPD/63703/2009, SFRH/BPD/107878/2015, SFRH/BPD/116774/2016.

EU SEAS-ERA project INVASIVES: SEAS-ERA/0001/2012.
CCMAR/Multi/04326/2013.

### Competing Interests

The authors declare there are no competing interests.

### Author Contributions

* Tania Aires performed the experiments, analyzed the data, authored or reviewed drafts of the paper.

* Alexandra Serebryakova performed the experiments, analyzed the data, prepared figures and/or tables, authored or reviewed drafts of the paper.

* Frédérique Viard contributed reagents/materials/analysis tools, prepared figures and/or tables, authored or reviewed drafts of the paper, funding.

* Ester A. Serrão conceived and designed the experiments, contributed reagents/materials/analysis tools, authored or reviewed drafts of the paper.
* Aschwin H. Engelen conceived and designed the experiments, performed the experiments, analyzed the data, contributed reagents/materials/analysis tools, prepared figures and/or tables, authored or reviewed drafts of the paper, funding.

**Data Availability**

The following information was supplied regarding data availability:
Engelen, Aschwin (2018): Acidification + Grazing_Effects_Microbiome. figshare.
https://doi.org/10.6084/m9 figshare.5346316.v3.

**Supplemental Information**

Supplemental information for this article can be found online at http://dx.doi.org/10.7717/peerj.4377#supplemental-information.

**REFERENCES**

**Aires T, Moalic Y, Serrao EA, Arnaud-Haond S. 2015.** Hologenome theory supported by co-occurrence networks of species-specific bacterial communities in siphonous algae (*Caulerpa*). *FEMS Microbiology Ecology* **91**: fiv067 DOI 10.1093/femsec/fiv067.

**Aires T, Muyzer G, Serrão EA, Engelen AH. 2018.** Unraveling seaweeds bacteriomes—from field site to computer screen. In: Charrier B, Wichard T, Reddy CRK, eds. *Protocols for macroalgae research*. Oxfordshire: Taylor & Francis Group, 97–112.

**Aires T, Serrão EA, Engelen AH. 2016.** Host and environmental specificity in bacterial communities associated to two highly invasive marine species (*Genus Asparagopsis*). *Frontiers in Microbiology* **7**:559 DOI 10.3389/fmicb.2016.00559.

**Asnaghi V, Chiantore M, Mangialajo L, Gazeau F, Francour P, Alliouane S, Gattuso JP. 2013.** Cascading effects of ocean acidification in a rocky subtidal community. *PLOS ONE* **8:e61978** DOI 10.1371/journal.pone.0061978.

**Asplund ME, Baden SP, Russ S, Ellis RP, Gong N, Hernroth BE. 2014.** Ocean acidification and host-pathogen interactions: blue mussels, *Mytilus edulis*, encountering *Vibrio tubiashii*. *Environmental Microbiology* **16**:1029–1039 DOI 10.1111/1462-2920.12307.

**Avelin Mary S, Vitalina Mary S, Rittschof D, Nagabhushanam R. 1993.** Bacterial-barnacle interaction: potential of using juncellins and antibiotics to alter structure of bacterial communities. *Journal of Chemical Ecology* **19**:2155–2167 DOI 10.1007/BF00979654.

**Barbieri E, Paster BJ, Hughes D, Zurek L, Moser DP, Teske A, Sogin ML. 2001.** Phylogenetic characterization of epibiotic bacteria in the accessory nidamental gland and egg capsules of the squid *Loligo pealei* (Cephalopoda: Loliginidae). *Environmental Microbiology* **3**:151–167 DOI 10.1046/j.1462-2920.2001.00172.x.

**Barneah O, Ben-Dov E, Kramarsky-Winter E, Kushmaro A. 2007.** Characterization of black band disease in Red Sea stony corals. *Environmental Microbiology* **9**:1995–2006 DOI 10.1111/j.1462-2920.2007.01315.x.
**Beleneva IA, Zhukova NV. 2006.** Bacterial communities of some brown and red algae from Peter the Great Bay, the Sea of Japan. *Microbiology* **75**:348–357 DOI 10.1134/S0026261706030180.

**Bengtsson MM, Sjøtun K, Øvreås L. 2010.** Seasonal dynamics of bacterial biofilms on the kelp *Laminaria hyperborea*. *Aquatic Microbial Ecology* **60**:71–83 DOI 10.3354/ame01409.

**Berg G, Grube M, Schloter M, Smalla K. 2014.** The plant microbiome and its importance for plant and human health. *Frontiers in Microbiology* **5**:491 DOI 10.1007/10.

**Bodelier PLE, Dedysh SN. 2013.** Microbiology of wetlands. *Frontiers in Microbiology* **4**:Article 79 DOI 10.3389/fmicb.2013.00079.

**Bodenhausen N, Horton MW, Bergelson J. 2013.** Bacterial communities associated with the leaves and the roots of *Arabidopsis thaliana*. *PLOS ONE* **8**:e56329 DOI 10.1371/journal.pone.0056329.

**Bourne D, Iida Y, Uthicke S, Smith-Keune C. 2008.** Changes in coral-associated microbial communities during a bleaching event. *ISME Journal* **2**:350–363 DOI 10.1038/ismej.2007.112.

**Bourne DG, Munn CB. 2005.** Diversity of bacteria associated with the coral *Pocillopora damicornis* from the great barrier reef. *Environmental Microbiology* **7**:1162–1174 DOI 10.1111/j.1462-2920.2005.00793.x.

**Branch TA, DeJoseph BM, Ray LJ, Wagner CA. 2013.** Impacts of ocean acidification on marine seafood. *Trends in Ecology and Evolution* **28**:178–186 DOI 10.1016/j.tree.2012.10.001.

**Briggs LM. 2017.** *The effects of ocean warming and acidification on seawed growth and urchin grazing*. Pomona: California State Polytechnic University.

**Burgess JG, Jordan EM, Bregu M, Mearns-Spragg A, Boyd KG. 1999.** Microbial antagonism: a neglected avenue of natural products research. *Journal of Biotechnology* **70**:27–32 DOI 10.1016/S0168-1656(99)00054-1.

**Burke C, Steinberg P, Rusch D, Kjelleberg S, Thomas T. 2011a.** Bacterial community assembly based on functional genes rather than species. *Proceedings of the National Academy of Sciences of the United States of America* **108**:14288–14293 DOI 10.1073/pnas.1101591108.

**Burke C, Thomas T, Lewis M, Steinberg P, Kjelleberg S. 2011b.** Composition, uniqueness and variability of the epiphytic bacterial community of the green alga *Ulva australis*. *The ISME Journal* **5**:590–600 DOI 10.1038/ismej.2010.164.

**Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, Costello EK, Fierer N, Peña AG, Goodrich JK, Gordon JI, Huttley GA, Kelley ST, Knights D, Koenig JE, Ley RE, Lozupone CA, Mcdonald D, Muegge BD, Pirrung M, Reeder J, Sevinsky JR, Turnbaugh PJ, Walters WA, Widmann J, Yatsunenko T, Zaneveld J, Knight R. 2010.** QIIME allows analysis of high-throughput community sequencing data. *Nature Publishing Group* **7**:335–336 DOI 10.1038/nmeth0510-335.

**Cardoso AM, Cavalcante JJV, Vieira RP, Lima JL, Grieco MAB, Clementino MM, Vasconcelos ATR, Garcia ES, De Souza W, Albano RM, Martins OB. 2012.** Gut
bacterial communities in the giant land snail *Achatina fulica* and their modification by sugarcane-based diet. *PLOS ONE* 7:e33440 DOI 10.1371/journal.pone.0033440.

Case RJ, Longford SR, Campbell AH, Low A, Tujula N, Steinberg PD, Kjelleberg S. 2011. Temperature induced bacterial virulence and bleaching disease in a chemically defended marine macroalga. *Environmental Microbiology* 13:529–537 DOI 10.1111/j.1462-2920.2010.02356.x.

Cayrou C, Sambe B, Armougom F, Raoult D, Drancourt M. 2013. Molecular diversity of the *Planctomycetes* in the human gut microbiota in France and Senegal. *Apmis* 121:1082–1090 DOI 10.1111/apm.12087.

Chao A. 1984. Nonparametric estimation of the number of Classes in a population. *Scandinavian Journal of Statistics* 11:265–270.

Chojnacka K, Saeid A, Witkowska Z, Tuhy Ł. 2012. Biologically active compounds in seaweed extracts—the prospects for the application. *The Open Conference Proceedings Journal* 3:20–28 DOI 10.2174/1876326X01203020020.

Church MJ. 2008. Resource control of bacterial dynamics in the sea. In: Kirchman DL, ed. *Microbial ecology of the oceans*. Hoboken: John Wiley & Sons, 335–382.

Clarke K, Gorley RN. 2006. Primer v6: user manual/tutorial. Plymouth: Primer-E Ltd, 192p.

Clarke KR, Warwick RM. 1994. *Change in marine communities. An approach to statistical analysis and interpretation*. Plymouth: Natural Environment Research Council, 1–172.

Cornwall CE, Revill AT, Hurd CL. 2015. High prevalence of diffusive uptake of CO₂ by macroalgae in a temperate subtidal ecosystem. *Photosynthesis Research* 124:181–190 DOI 10.1007/s11120-015-0114-0.

Cottrell MT, Kirchman DL. 2000. Community composition of marine bacterioplankton determined by 16S rRNA gene clone libraries and fluorescence *in situ* hybridization. *Applied and Environmental Microbiology* 66:5116–5122 DOI 10.1128/aem.66.12.5116-5122.2000.

Currie AR, Tait K, Parry H, De Francisco-Mora B, Hicks N, Mark Osborn A, Wid-dicombe S, Stahl H. 2017. Marine microbial gene abundance and community composition in response to ocean acidification and elevated temperature in two contrasting coastal marine sediments. *Frontiers in Microbiology* 8:Article 1599 DOI 10.3389/fmicb.2017.01599.

Davis J, Fricke WF, Hamann MT, Esquenazi E, Dorrestein PC, Hill RT. 2013. Characterization of the bacterial community of the chemically defended Hawaiian sacoglossan *Elysia rufescens*. *Applied and Environmental Microbiology* 79:7073–7081 DOI 10.1128/AEM.01568-13.

Devine SP, Pelletreau KN, Rumpho ME. 2012. 16S rDNA-based metagenomic analysis of bacterial diversity associated with two populations of the kleptoplastic sea slug *Elysia chlorotica* and its algal prey *Vaucheria litorea*. *The Biological Bulletin* 223:138–154 DOI 10.1086/BBLv223n1p138.

Dimitrieva GY, Crawford RL, Yüksel GÜ. 2006. The nature of plant growth-promoting effects of a Pseudoalteromonad associated with the marine algae *Laminaria japonica*
and linked to catalase excretion. *Journal of Applied Microbiology* **100**:1159–1169
DOI 10.1111/j.1365-2672.2006.02831.x.

Dittmer J, Lesobre J, Moumen B, Bouchon D. 2016. Host origin and tissue microhabitat shaping the microbiota of the terrestrial isopod *Armadillidium vulgare*. *FEMS Microbiology Ecology* **92**:Article fiw063 DOI 10.1093/femsec/fiw063.

Dudek M, Adams J, Swain M, Hegarty M, Huws S, Gallagher J. 2014. Metaphylogen-
nomic and potential functionality of the limpet *Patella pellucida*’s gastrointestinal
tract microbiome. *International Journal of Molecular Sciences* **15**:18819–18839
DOI 10.3390/ijms151018819.

Eberl R. 2010. Sea-land transitions in isopods: pattern of symbiont distribution in two species of intertidal isopods *Ligia pallasii* and *Ligia occidentalis* in the Eastern Pacific. *Symbiosis* **51**:107–116 DOI 10.1007/s13199-010-0057-3.

Edgar RC. 2010. Search and clustering orders of magnitude faster than BLAST. *Bioinformatics* **26**:2460–2461 DOI 10.1093/bioinformatics/btq461.

Egan S, Harder T, Burke C, Steinberg P, Kjelleberg S, Thomas T. 2013. The seaweed holobiont: understanding seaweed-bacteria interactions. *FEMS Microbiology Reviews* **37**:462–476 DOI 10.1111/1574-6976.12011.

Fabry VJ, Seibel BA, Richard AF, Orr JC. 2008. Impacts of ocean acidification on marine fauna and ecosystem processes. *ICES Journal of Marine Science* **65**:414–432 DOI 10.1093/icesjms/fsn048.

Fenchel T. 2012. Chapter 3—degradation of organic polymers and hydrocarbons. Amsterdam: Elsevier DOI 10.1016/B978-0-12-415836-8.00003-7.

Fraune S, Zimmer M. 2008. Host-specificity of environmentally transmitted Mycoplasma-like isopod symbionts. *Environmental Microbiology* **10**:2497–2504 DOI 10.1111/j.1462-2920.2008.01672.x.

Frey JC, Rothman JM, Pell AN, Nizeyi B, Cranfield MR, Angert ER, Nizeyi JB. 2006. Fecal bacterial diversity in a wild gorilla. *Applied and Environmental Microbiology* **72**:3788–3792 DOI 10.1128/AEM.72.5.3788.

Fuerst JA, Sagulenko E. 2011. Beyond the bacterium: *Planctomycetes* challenge our concepts of microbial structure and function. *Nature Reviews Microbiology* **9**:403–413 DOI 10.1038/nrmicro2578.

Gachon CMM, Sime-Ngando T, Strittmatter M, Chambouvet A, Kim GH. 2010. Algal diseases: spotlight on a black box. *Trends in Plant Science* **15**:633–640 DOI 10.1016/j.tplants.2010.08.005.

Garrity GM, Bell JA, Lilburn T, Garrity GM, Bell JA, Lilburn T. 2005. Oceanospirillales ord. nov. In: Brenner D, Krieg N, Garrity G, Boone D, De Vos P, eds. *Bergey's manual of systematics of archaea and bacteria*. Hoboken: John Wiley & Sons, 270–323 DOI 10.1002/9781118960608.obm00100.

Glöckner FO, Fuchs BM, Fuchs BM, Glo FO, Amann R. 1999. Bacterioplankton compositions of lakes and oceans: a first comparison based on fluorescence *in situ* hybridization. *Applied and Environmental Microbiology* **65**:3721–3726.

Goffredi SK, Johnson SB, Vrijenhoek RC. 2007. Genetic diversity and potential function of microbial symbionts associated with newly discovered species of
*Osedax polychaete* worms. *Applied and Environmental Microbiology* 73:2314–2323
DOI 10.1128/AEM.01986-06.

Greiman SE, Tkach VV, Pulis E, Fayton TJ, Curran SS. 2014. Large scale screening of digeneans for *Neorickettsia* endosymbionts using real-time PCR reveals new *Neorickettsia* genotypes, host associations and geographic records. *PLOS ONE*
9:e98453 DOI 10.1371/journal.pone.0098453.

Gutow L, Rahman MM, Bartl K, Saborowski R, Bartsch I, Wiencke C. 2014. Ocean acidification affects growth but not nutritional quality of the seaweed *Fucus vesiculosus* (Phaeophyceae, Fucales). *Journal of Experimental Marine Biology and Ecology* 453:84–90 DOI 10.1016/j.jembe.2014.01.005.

Hacquard S, Garrido-Oter R, González A, Spaepen S, Ackermann G, Lebeis S, McHardy AC, Dangl JL, Knight R, Ley R, Schulze-Lefert P. 2015. Microbiota and host nutrition across plant and animal kingdoms. *Cell Host and Microbe* 17:603–616 DOI 10.1016/j.chom.2015.04.009.

Harley CDG, Anderson KM, Demes KW, Jorve JP, Kordas RL, Coyle TA, Graham MH. 2012. Effects of climate change on global seaweed communities. *Journal of Phycology* 48:1064–1078 DOI 10.1111/j.1529-8817.2012.01224.x.

Hassenrück C, Fink A, Lichtschlag A, Tegetmeyer HE, De Beer D, Ramette A. 2016. Quantification of the effects of ocean acidification on sediment microbial communities in the environment: the importance of ecosystem approaches. *FEMS Microbiology Ecology* 92:Article fiw027 DOI 10.1093/femsec/fiw027.

Head WD, Carpenter EJ. 1975. Nitrogen fixation associated with the marine macroalga *Codium fragile*. *Limnology and Oceanography* 20:815–823 DOI 10.1017/CBO9781107415324.004.

Hentschel U, Schmid M, Wagner M, Fieseler L, Gernert C, Hacker J. 2001. Isolation and phylogenetic analysis of bacteria with antimicrobial activities from the Mediterranean sponges *Aplysina aerophoba* and *Aplysina cavernicola*. *FEMS Microbiology Ecology* 35:305–312 DOI 10.1111/j.1574-6941.2001.tb00816.x.

Hollants J, Leliaert F, De Clerck O, Willems A. 2013. What we can learn from sushi: a review on seaweed-bacterial associations. *FEMS Microbiology Ecology* 83:1–16 DOI 10.1111/j.1574-6941.2012.01446.x.

Holmström C, Kjelleberg S. 1999. Marine *Pseudoalteromonas* species are associated with higher organisms and produce biologically active extracellular agents. *FEMS Microbiology Ecology* 30:285–293 DOI 10.1111/j.1574-6941.1999.tb00656.x.

Holmstrom C, Rittschof D, Kjelleberg S. 1992. Inhibition of settlement by larvae of *Balanus amphitrite* and *Ciona intestinalis* by a surface-colonizing marine bacterium. *Applied and Environmental Microbiology* 58:2111–2115.

Hong PY, Wheeler E, Cann IKO, Mackie RI. 2011. Phylogenetic analysis of the fecal microbial community in herbivorous land and marine iguanas of the Galápagos Islands using 16S rRNA-based pyrosequencing. *ISME Journal* 5:1461–1470 DOI 10.1038/ismej.2011.33.
Houghton JT, Ding Y, Griggs DJ, Noguer M, Van der Linden PJ, Dai X, Maskell K, Johnson C (eds.) 2001. IPCC, 2001: climate change 2001: the scientific basis, contribution of working group I to the third assessment report of the intergovernmental panel on climate change. In: *Climate Change 2001: the Scientific Basis*. Cambridge: Cambridge University Press, 881 DOI 10.1256/004316502320517344.

Huggett MJ, Williamson JE, De Nys R, Kjelleberg S, Steinberg PD. 2006. Larval settlement of the common Australian sea urchin *Heliocidaris erythrogramma* in response to bacteria from the surface of coralline algae. *Oecologia* 149:604–619 DOI 10.1007/s00442-006-0470-8.

Kerfahi D, Hall-Spencer JM, Tripathi BM, Milazzo M, Lee J, Adams JM. 2014. Shallow water marine sediment bacterial community shifts along a Natural CO2 gradient in the mediterranean sea off Vulcano, Italy. *Microbial Ecology* 67:819–828 DOI 10.1007/s00248-014-0368-7.

King GM, Judd C, Kuske CR, Smith C. 2012. Analysis of stomach and gut microbiomes of the Eastern Oyster (*Crassostrea virginica*) from Coastal Louisiana, USA. *PLOS ONE* 7:e51475 DOI 10.1371/journal.pone.0051475.

Kirchman DL. 2002. The ecology of Cytophaga-Flavobacteria in aquatic environments. *FEMS Microbiology Ecology* 39:91–100 DOI 10.1016/S0168-6496(01)00206-9.

Koch M, Bowes G, Ross C, Zhang XH. 2013. Climate change and ocean acidification effects on seagrasses and marine macroalgae. *Global Change Biology* 19:103–132 DOI 10.1111/j.1365-2486.2012.02791.x.

Kooperman N, Ben-Dov E, Kramarsky-Winter E, Barak Z, Kushmaro A. 2007. Coral mucus-associated bacterial communities from natural and aquarium environments. *FEMS Microbiology Letters* 276:106–113 DOI 10.1111/j.1574-6968.2007.00921.x.

Kroeker KJ, Kordas RL, Crim R, Hendriks IE, Ramajo L, Singh GS, Duarte CM, Gattuso JP. 2013. Impacts of ocean acidification on marine organisms: quantifying sensitivities and interaction with warming. *Global Change Biology* 19:1884–1896 DOI 10.1111/gcb.12179.

Lachnit T, Meske D, Wahl M, Harder T, Schmitz R. 2011. Epibacterial community patterns on marine macroalgae are host-specific but temporally variable. *Environmental Microbiology* 13:655–665 DOI 10.1111/j.1462-2920.2010.02371.x.

Lage OM, Bondoso J. 2014. *Planctomycetes* and macroalgae, a striking association. *Frontiers in Microbiology* 5:1–9 DOI 10.3389/fmicb.2014.00267.

Lane DJ. 1991. 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M, eds. *Nucleic acid techniques in bacterial systematics*. Chichester: John Wiley and Sons, 115–175.

Liu H, Zheng F, Sun X, Hong X, Dong S, Wang B, Tang X, Wang Y. 2010. Identification of the pathogens associated with skin ulceration and peristome tumescence in cultured sea cucumbers *Apostichopus japonicus* (Selenka). *Journal of Invertebrate Pathology* 105:236–242 DOI 10.1016/j.jip.2010.05.016.

Longford S, Tujula N, Crocetti G, Holmes A, Holmström C, Kjelleberg S, Steinberg P, Taylor M. 2007. Comparisons of diversity of bacterial communities associated
with three sessile marine eukaryotes. *Aquatic Microbial Ecology* **48**:217–229 DOI 10.3354/ame048217.

**Mackie RI, Rycyk M, Ruemmler RL, Aminov RI, Wikelski M. 2004.** Biochemical and microbiological evidence for fermentative digestion in free-living land iguanas (*Conolophus pallidus*) and marine iguanas (*Amblyrhynchus cristatus*) on the Galápagos archipelago. *Physiological and Biochemical Zoology* **77**:127–138 DOI 10.1086/383498.

**Maki JS, Rittschof D, Costlow JD, Mitchell R. 1988.** Inhibition of attachment of larval barnacles, *Balanus amphitrite*, by bacterial surface films. *Marine Biology* **97**:199–206 DOI 10.1007/BF00391303.

**Marzinelli EM, Campbell AH, Zozaya Valdes E, Vergés A, Nielsen S, Wernberg T, De Bettignies T, Bennett S, Caporaso JG, Thomas T, Steinberg PD. 2015.** Continental-scale variation in seaweed host-associated bacterial communities is a function of host condition, not geography. *Environmental Microbiology* **17**:4078–4088 DOI 10.1111/1462-2920.12972.

**Matsuo Y, Imagawa H, Nishizawa M, Shizuri Y. 2005.** Isolation of an algal morphogenesis inducer from a marine bacterium. *Science* **307**:1598 DOI 10.1126/science.1105486.

**Matsuo Y, Suzuki M, Kasai H, Shizuri Y, Harayama S. 2003.** Isolation and phylogenetic characterization of bacteria capable of inducing differentiation in the green alga *Monostroma oxyspermum*. *Environmental Microbiology* **5**:25–35 DOI 10.1046/j.1462-2920.2003.00382.x.

**Mattila JM, Zimmer M, Vesakoski O, Jormalainen V. 2014.** Habitat-specific gut microbiota of the marine herbivore *Idotea balthica* (Isopoda). *Journal of Experimental Marine Biology and Ecology* **455**:22–28 DOI 10.1016/j.jembe.2014.02.010.

**McDonald D, Price MN, Goodrich J, Nawrocki EP, DeSantis TZ, Probst A, Andersen GL, Knight R, Hugenholtz P. 2012.** An improved Greengenes taxonomy with explicit ranks for ecological and evolutionary analyses of bacteria and archaea. *The ISME Journal* **6**:610–618 DOI 10.1038/ismej.2011.139.

**Mensch B, Neulinger SC, Graiff A, Pansch A, Künzel S, Fischer MA, Schmitz RA. 2016.** Restructuring of epibacterial communities on *Fucus vesiculosus* forma *mytili* in response to elevated pCO₂ and increased temperature levels. *Frontiers in Microbiology* **7**:Article 434 DOI 10.3389/fmicb.2016.00434.

**Meron D, Atias E, Iasur Kruh L, Elifantz H, Minz D, Fine M, Banin E. 2011.** The impact of reduced pH on the microbial community of the coral *Acropora eurystoma*. *ISME Journal* **5**:51–60 DOI 10.1038/ismej.2010.102.

**Meusnier I, Olsen JL, Stam WT, Destombe C, Valero M. 2001.** Phylogenetic analyses of *Caulerpa taxifolia* (Chlorophyta) and of its associated bacterial microflora provide clues to the origin of the Mediterranean introduction. *Molecular Ecology* **10**:931–946 DOI 10.1046/j.1365-294X.2001.01245.x.

**Meziti A, Ramette A, Mente E, Kormas KA. 2010.** Temporal shifts of the Norway lobster (*Nephrops norvegicus*) gut bacterial communities. *FEMS Microbiology Ecology* **74**:472–484 DOI 10.1111/j.1574-6941.2010.00964.x.
**Moisander PH, Sexton AD, Daley MC. 2015.** Stable associations masked by temporal variability in the marine copepod microbiome. *PLOS ONE* 10:e0138967
DOI 10.1371/journal.pone.0138967.

**Morrow KM, Bourne DG, Humphrey C, Botté ES, Laffy P, Zaneveld J, Uthicke S, Fabricius KE, Webster NS. 2015.** Natural volcanic CO₂ seeps reveal future trajectories for host-microbial associations in corals and sponges. *The ISME Journal* 9:894–908
DOI 10.1038/ismej.2014.188.

**Nedashkovskaya OI, Vancanneyt M, Christiaens L, Kalinovskaya NI, Mikhailov VV, Swings J. 2006.** *Aquimarina intermedia* sp. nov., reclassification of *Stanierella latercula* (Lewin 1969) as *Aquimarina latercula* comb. nov and *Gaetbulimicrobium brevivitae* Yoon, et al. 2006 as *Aquimarina brevivitae* comb. nov., and amended description of the genus *Aquimarina*. *International Journal of Systematic and Evolutionary Microbiology* 56:2037–2041 DOI 10.1099/ijs.0.64155-0.

**Olischläger M, Bartsch I, Gutow L, Wiencke C. 2013.** Effects of ocean acidification on growth and physiology of *Ulva lactuca* (Chlorophyta) in a rockpool-scenario. *Phycological Research* 61:180–190 DOI 10.1111/pre.12006.

**Parris DJ, Brooker RM, Morgan MA, Dixson DL, Stewart FJ. 2016.** Whole gut microbiome composition of damselfish and cardinalfish before and after reef settlement. *PeerJ* 4:e2412 DOI 10.7717/peerj.2412.

**Patel P, Callow ME, Joint I, Callow JA. 2003.** Specificity in the settlement—modifying response of bacterial biofilms towards zoospores of the marine alga *Enteromorpha*. *Environmental Microbiology* 5:338–349 DOI 10.1046/j.1462-2920.2003.00407.x.

**Perlman SJ, Hunter MS, Zchori-Fein E. 2006.** The emerging diversity of *Rickettsia*. *Proceedings of the Royal Society B: Biological Sciences* 273:2097–2106
DOI 10.1098/rspb.2006.3541.

**Poore AGB, Graba-Landry A, Favret M, Sheppard Brennand H, Byrne M, Dworjanyn SA. 2013.** Direct and indirect effects of ocean acidification and warming on a marine plant-herbivore interaction. *Oecologia* 173:1113–1124
DOI 10.1007/s00442-013-2683-y.

**Porzio L, Buia MC, Hall-Spencer JM. 2011.** Effects of ocean acidification on macroalgal communities. *Journal of Experimental Marine Biology and Ecology* 400:278–287
DOI 10.1016/j.jembe.2011.02.011.

**Sawabe T, Makino H, Tatsumi M, Nakano K, Tajima K, Iqbal MM, Yumoto I, Ezura Y, Christen R. 1998.** *Pseudoalteromonas bacteriolytica* sp. nov., a marine bacterium that is the causative agent of red spot disease of *Laminaria japonica*. *International Journal of Systematic Bacteriology* 3:769–774.

**Schulze AD, Alabi AO, Tattersall-Sheldrake AR, Miller KM. 2006.** Bacterial diversity in a marine hatchery: balance between pathogenic and potentially probiotic bacterial strains. *Aquaculture* 256:50–73 DOI 10.1016/j.aquaculture.2006.02.008.

**Shah HN, Gharbia SE. 1993.** Ecophysiology and taxonomy of bacteroides and related taxa. *Clinical Infectious Diseases* 16:S160–S167
DOI 10.1093/clinids/16.Supplement_4.S160.
**Shnit-Orland M, Sivan A, Kushmaro A. 2012.** Antibacterial activity of pseudoalteromonas in the coral holobiont. *Microbial Ecology* **64**:851–859
DOI 10.1007/s00248-012-0086-y.

**Simon M, Glöckner FO, Amann R. 1999.** Different community structure and temperature optima of heterotrophic picoplankton in various regions of the Southern Ocean. *Aquatic Microbial Ecology* **18**:275–284 DOI 10.3354/ame018275.

**Singh RP, Mantri VA, Reddy CRK, Jha B. 2011.** Isolation of seaweed-associated bacteria and their morphogenesis-inducing capability in axenic cultures of the green alga *Ulva fasciata*. *Aquatic Biology* **12**:13–21 DOI 10.3354/ab00312.

**Singh RP, Reddy CRKRK. 2014.** Seaweed-microbial interactions: key functions of seaweed-associated bacteria. *FEMS Microbiology Ecology* **88**:213–230
DOI 10.1111/1574-6941.12297.

**Song JK, Kim D, Eun JB, Choi BD, Oh MJ, Jung SJ. 2012.** Identification of cellulolytic bacteria associated with tunic softness syndrome in the sea squirt, *Halocynthia roretzi*. *Food Science and Biotechnology* **21**:1405–1411
DOI 10.1007/s10068-012-0185-z.

**Staufenberger T, Thiel V, Wiese J, Imhoff JF. 2008.** Phylogenetic analysis of bacteria associated with *Laminaria saccharina*. *FEMS Microbiology Ecology* **64**:65–77
DOI 10.1111/j.1574-6941.2008.00445.x.

**Sullam KE, Essinger SD, Lozupone CA, O'Connor MP, Rosen GL, Knight R, Kilham SS, Russell JA. 2012.** Environmental and ecological factors that shape the gut bacterial communities of fish: a meta-analysis. *Molecular Ecology* **21**:3363–3378
DOI 10.1111/j.1365-294X.2012.05552.x.

**Sunagawa S, Desantis TZ, Piceno YM, Brodie EL, Desalvo MK, Voolstra CR, Weil E, Andersen GL, Medina M. 2009.** Bacterial diversity and white Plague disease-associated community changes in the caribbean coral *Montastraea faveolata*. *ISME Journal* **3**:512–521 DOI 10.1038/ismej.2008.131.

**Suzuki M, Nakagawa Y, Harayama S, Yamamoto S. 2001.** Phylogenetic analysis and taxonomic study of marine Cytophaga-like bacteria: proposal for *Tenacibaculum* gen. nov. with *Tenacibaculum maritimum* comb. nov. and *Tenacibaculum ovolyticum* comb. nov. and description of *Tenacibaculum mesophilum* sp. nov. and Ten. *International Journal of Systematic and Evolutionary Microbiology* **51**:1639–1652
DOI 10.1099/00207713-51-5-1639.

**Taylor JD, Ellis R, Milazzo M, Hall-Spencer JM, Cunliffe M. 2014.** Intertidal epilithic bacteria diversity changes along a naturally occurring carbon dioxide and pH gradient. *FEMS Microbiology Ecology* **89**:670–678 DOI 10.1111/1574-6941.12368.

**Thomas T, Evans FF, Schleheck D, Mai-Prochnow A, Burke C, Penesyan A, Dalisay DS, Stelzer-Braid S, Saunders N, Johnson J, Ferriera S, Kjelleberg S, Egan S. 2008.** Analysis of the *Pseudoalteromonas tunicata* genome reveals properties of a surface-associated life style in the marine environment. *PLOS ONE* **3**:e3252
DOI 10.1371/journal.pone.0003252.
**Thomas F, Hehemann JH, Rebuffet E, Czjzek M, Michel G. 2011.** Environmental and gut Bacteroidetes: the food connection. *Frontiers in Microbiology* 2:Article 93 DOI 10.3389/fmicb.2011.00093.

**Thurber RV, Willner-Hall D, Rodriguez-Mueller B, Desnues C, Edwards RA, Angly F, Dinsdale E, Kelly L, Rohwer F. 2009.** Metagenomic analysis of stressed coral holobionts. *Environmental Microbiology* 11:2148–2163 DOI 10.1111/j.1462-2920.2009.01935.x.

**Tietjen M. 2014.** "You are what you eat": how diet can influence the gut microbiota of marine invertebrates. *The Plymouth Student Scientist* 7:203–211.

**Tujula NA, Crocetti GR, Burke C, Thomas T, Holmström C, Kjelleberg S. 2010.** Variability and abundance of the epiphytic bacterial community associated with a green marine Ulvacean alga. *The ISME Journal* 4:301–311 DOI 10.1038/ismej.2009.107.

**Urabe J, Togari J, Elser JJ. 2003.** Stoichiometric impacts of increased carbon dioxide on a planktonic herbivore. *Global Change Biology* 9:818–825 DOI 10.1046/j.1365-2486.2003.00634.x.

**Van De Waal DB, Verschoor AM, Verspagen JMH, Van Donk E, Huisman J. 2010.** Climate-driven changes in the ecological stoichiometry of aquatic ecosystems. *Frontiers in Ecology and the Environment* 8:145–152 DOI 10.1890/080178.

**Vezzulli L, Grande C, Reid PC, Hélaouët P, Edwards M, Höfle MG, Brettar I, Colwell RR, Pruzzo C. 2016.** Climate influence on *Vibrio* and associated human diseases during the past half-century in the coastal North Atlantic. *Proceedings of the National Academy of Sciences of the United States of America* 113:E5062–E5071 DOI 10.1073/pnas.1609157113.

**Vieira C, Engelen AH, Guentas L, Aires T, Houlbreque F, Gaubert J, Serrão EA, Clerck O De, Payri CE. 2016.** Species specificity of bacteria associated to the brown seaweeds *Lobophora* (Dictyotales, Phaeophyceae) and their potential for induction of rapid coral bleaching in *Acropora muricata*. *Frontiers in Microbiology* 7:Article 316 DOI 10.3389/fmicb.2016.00316.

**Wang Y, Brune A, Zimmer M. 2007.** Bacterial symbionts in the hepatopancreas of isopods: diversity and environmental transmission. *FEMS Microbiology Ecology* 61:141–152 DOI 10.1111/j.1574-6941.2007.00329.x.

**Wang Y, Feng N, Li Q, Ding J, Zhan Y, Chang Y. 2012.** Isolation and characterization of bacteria associated with a syndrome disease of sea urchin *Strongylocentrotus intermedius* in North China. *Aquaculture Research* 44:691–700 DOI 10.1111/j.1365-2109.2011.03073.x.

**Wang G, Shuai L, Li Y, Lin W, Zhao X, Duan D. 2008.** Phylogenetic analysis of epiphytic marine bacteria on Hole-Rotten diseased sporophytes of *Laminaria japonica*. *Journal of Applied Phycology* 20:403–409 DOI 10.1007/s10811-007-9274-4.

**Webster NS, Negri AP, Flores F, Humphrey C, Soo R, Botté ES, Vogel N, Uthicke S. 2013.** Near-future ocean acidification causes differences in microbial associations within diverse coral reef taxa. *Environmental Microbiology Reports* 5:243–251 DOI 10.1111/1758-2229.12006.
Witt V, Wild C, Anthony KRN, Diaz-Pulido G, Uthicke S. 2011. Effects of ocean acidification on microbial community composition of, and oxygen fluxes through, biofilms from the Great Barrier Reef. *Environmental Microbiology* 13:2976–2989 DOI 10.1111/j.1462-2920.2011.02571.x.

Yuksel SA, Thompson KD, Adams A. 2006. Rickettsial infections of fish. *Turkish Journal of Fisheries and Aquatic Sciences* 6:63–78 DOI 10.1016/S0959-8030(96)00001-4.

Zimmer M. 2002. Nutrition in terrestrial isopods (Isopoda: Oniscidea): an evolutionary-ecological approach. *Biological Reviews of the Cambridge Philosophical Society* 77:455–493 DOI 10.1017/S1464793102005912.

Zimmer M, Bartholmé S. 2003. Bacterial endosymbionts in *Asellus aquaticus* (Isopoda) and *Gammarus pulex* (Amphipoda), and their contribution to digestion. *Limnology and Oceanography* 48:2208–2213 DOI 10.4319/lo.2003.48.6.2208.

Zimmer M, Danko JP, Pennings SC, Danford AR, Carefoot TH, Ziegler A, Uglow RF. 2002. Cellulose digestion and phenol oxidation in coastal isopods (Crustacea: Isopoda). *Marine Biology* 140:1207–1213 DOI 10.1007/s00227-002-0800-2.