*4.2. Field Sampling*

To assess the composition of the fouling organism communities, oyster clusters growing on scallop shells were collected from four oyster farms in the inner part of the bay in August 2014 (Figure 5). The seawater temperature at a depth of 2 m in farm A was 20–21 ◦C. At all sampling farms, two-year-old oysters were cultured (the oysters were hatched in summer 2012). One oyster cluster (Figure 6) was collected from ~2 m depth from each of the three ropes that were randomly selected at each farm. Since vertical distribution of fouling organisms are expected to be different [23], three clusters were collected from ~8 m depth at farm A. Thus, 15 oyster clusters were sampled in total. Immediately after sample collection, the oyster clusters, including oysters, mussels, sponges and algae, and others, were disassembled by hand and sorted into four groups. Because sponges and macroalgae were attached together tightly and intricately, these two groups could not be separated. Thus, sponges and macroalgae were treated as one group. The macroalgae were mainly composed of *Ulva* and brown algae. Each group of sessile fouling organisms was measured for abundance and wet biomass. For the wet weight measurement, the samples were carefully dried with paper towels to minimise errors and then weighed using an electronic scale. For the sampled oysters, all individuals were measured for length, width, shell height, and whole-body wet weight. The soft tissues were then obtained by dissection and measured for wet weight. The shells were weighed after air drying. For each sampled cluster, to remove the effect of individual size on the fatty acid analysis, tissue samples were selected from five individuals with similar shell lengths (103.3 ± 12.3 mm, mean ± SD) and preserved in a freezer at −30 ◦C for later fatty acid analysis. Similarly, for the *M. galloprovincialis* mussels, which were predominant among the sessile fouling organisms, five individuals were randomly selected from each cluster sample with similar shell lengths (46.8 ± 6.8 mm, mean ± SD), and the shell size, wet weight of the soft tissue, and dry weight of the shell were measured. The soft tissue samples of the mussels were preserved in a freezer for later fatty acid analysis.

**Figure 6.** Two oyster clusters fouled by sponges, macroalgae, *M. galloprovincialis*, and other organisms. This photo was taken at farm B in February 2017.

#### *4.3. Fatty Acid Analysis*

Since the whole body of the oyster is eaten by humans, we evaluated the fatty acid contents of oysters from the whole body. First, the soft tissue samples of the oysters and mussels were freeze-dried. Then, the whole body was powdered and homogenised in a blender. The 'one-step method' described by Abdulkadir and Tsuchiya [57] was applied for lipid extraction and derivatisation from the freeze-dried samples. Approximately 100 mg of freeze-dried sample was moved to a 50 mL glass tube. One millilitre of internal standard (1 mg tricosanoic acid per ml hexane), 4 mL hexane, and 2 mL 14% boron trifluoride methanol were added to the test tube, and nitrogen gas was added to fill the head space. The glass tubes were heated at 100 ◦C in a water bath for 2 h, then cooled to room temperature, and 1 mL hexane and 2 mL ultrapure water were added. The samples were shaken vigorously and centrifuged for 3 min at 2500 rpm (M-4000, KUBOTA Corp., Tokyo, Japan). The upper layer of hexane, which contained fatty acid methyl esters (FAME), was then placed in a 1.5 mL GC vial.

For quantification of the fatty acids, 1 μL FAME solution was injected into a gas chromatograph with an FID detector (GC-2014, Shimadzu, Kyoto, Japan) equipped with a capillary column (Select FAME, 100 m × 0.25 mm i.d., Agilent Technologies, Tokyo, Japan). The analytical conditions followed those outlined by Fujibayashi et al. [58]. The peak of each fatty acid was identified by comparison with the retention time of commercial standard mixtures (Supelco37, PUFA No.3, Bacterial FAME, Supelco®, Darmstadt, Germany). The amount of each fatty acid (milligram fatty acid per dry weight of animal) was calculated by following the method of Abdulkadir and Tsuchiya [57], with the internal standard (i.e., tricosanoic acid).

#### *4.4. Data Analysis*

We applied two condition indices (CI1 and CI2) in this study. CI1 and CI2 have been generally applied for oysters and other bivalve species, including mussels [24,59,60].

The oyster body condition index (CI1), which was developed by Lawrence and Scott [50], was calculated by

$$\text{CI1} = \text{(Dfrw (g))} / \text{(Ww (g) -- Shw (g))} \times 100$$

where Dfw is the dry weight of soft tissue, which was measured after freeze drying, Ww is the total wet weight of the shell and soft tissue without any fouling organisms, and Shw is the dried shell weight. CI1 expresses the ratio of the dry weight of soft tissue to the internal shell volume, with the assumption that the density of soft tissue is almost the same as that of seawater. This assumption has been validated in oysters [24]. However, there has been no attempt to verify this for *M. galloprovincialis,* and Lucas and Beninger [59] pointed out that it is unlikely that the underlying assumptions are applicable to all bivalves. Therefore, we considered CI2 more appropriate for *M. galloprovincialis* in this study.

For mussels, the following condition index was applied [60]:

$$\text{CI2} = \text{(Dfw (g))} / \text{(Shw (g))}$$

To examine the effects of fouling organisms on the CI and content of EPA and DHA in the cultivated oysters, correlation analysis was conducted by SPSS software (IBM, ver.20). All data were explored for normality using a Kolmogorov–Smirnov test and normality was not supported for the wet weight and relative weight of *M. galloprovincialis*. Then, Spearman rank correlation analysis was applied for *M. galloprovincialis*, and Pearson's correlation analysis was applied for other fouling organisms. Fouling organisms were expressed as the total wet weight (g cluster−1). Furthermore, relative weight to oysters (g g−1) was also calculated since relative weight of fouling organisms can be expected to affect *C. gigas*. For the content of EPA and DHA, the concentration in each individual (mg g−1) and total amount in a cluster (g cluster−1) were evaluated and applied in the correlation analysis.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/md19070369/s1, Table S1: Composition of phytoplankton taxa collected near farm A in July 2017 (cell L<sup>−</sup>1).

**Author Contributions:** Conceptualisation, M.F., O.N. and T.S.; methodology, M.F. and T.S.; investigation, M.F. and T.S.; chemical analysis, M.F. and T.S.; writing—original draft preparation, M.F. and T.S.; writing—review and editing, M.F., O.N. and T.S.; visualisation, M.F.; supervision, O.N.; project administration, O.N.; funding acquisition, O.N. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was financially supported by the Environment Research and Technology Development Fund of the Ministry of the Environment, Japan (grant number S-13), the Japanese Institute of Fisheries Infrastructure and Communities, and KAKENHI (grant number JP17H01885, 19KT0006).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data are included in the manuscript.

**Acknowledgments:** We thank K. Goto, T. Kudo, N. Chiba, A. Kato, C. Maruo, H. Kanzaki, and Y. Zheng for assistance with the field and laboratory work. We thank R. Filgueira for their valuable comments on the manuscript.

**Conflicts of Interest:** The authors declare no conflict of interest.
