Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae.

We investigated the succession process of intestinal bacteria during seed production in full-cycle aquaculture of Pacific bluefin tuna (Thunnus orientalis). During the seed production, eggs, healthy fish, rearing water, and feeds from three experimental rounds in 2012 and 2013 were collected before transferring to offshore net cages and subjected to a fragment analysis of the bacterial community structure. We identified a clear succession of intestinal bacteria in bluefin tuna during seed production. While community structures of intestinal bacteria in the early stage of larvae were relatively similar to those of rearing water and feed, the bacterial community structures seen 17 days after hatching were different. Moreover, although intestinal bacteria in the late stage of larvae were less diverse than those in the early stage of larvae, the specific bacteria were predominant, suggesting that the developed intestinal environment of the host puts selection pressure on the bacteria in the late stage. The specific bacteria in the late stage of larvae, which likely composed 'core microbiota', were also found on the egg surface. The present study highlights that proper management of the seed production process, including the preparation of rearing water, feeds, and fish eggs, is important for the aquaculture of healthy fish.

Introduction

The intestinal microbiome of mammals has been well studied, and various findings have been obtained. In mammals, infancy and early childhood are the key periods for shaping the gut microbiome [1], which supports host functions such as indirect (immune system-mediated) and direct protection against pathogens [2] and nutrient absorption [3]. Hviid et al. [4] reported that the administration of antibiotics during this key period increases the incidence of other diseases, due to interference with the still unstable gut microbiome. Environmental factors play an important role in shaping the gut microbiome of mammals. The effects of oral acquisition are also important. For example, Ferretti et al. [5] highlighted that oral acquisition of bacteria from maternal skin, breast milk, feces, vagina, and oral cavity is crucial for the development of infant microbiome. Furthermore, it has been shown that the gut microbiome of Japanese individuals exhibits a higher abundance of unique bacteria (Bifidobacterium) than that of individuals in other nations, indicating the influence of unique Japanese traditional foods [6].

The intestinal microbiome of fish has also been well studied. For example, Sugita et al. [7] using a direct counting method showed that the intestinal bacteria in eight marine fish species comprised approximately 109–1010 cells per g of intestinal contents. Such abundance of bacteria likely benefit the host fish as they do for mammals, improving digestion [8], production of vitamins [9], and exhibiting antibacterial activity against fish pathogens [10]. Environmental factors, among which food resources and habitat are important, shape the intestinal microbiome of fish [11-13]. Uchii et al. [11] reported that different feeding habitats among the same fish species led to the formation of different intestinal microbiota. The number of studies on fish intestinal bacteria has been increasing; however, their succession process during fish development from the egg is still unknown. This is due to the complex bacterial community in fish intestines and the surrounding environments such as water and feeds, which makes it difficult to trace the intestinal bacteria.

Pacific bluefin tuna Thunnus orientalis is one of the most popular and important fishery species, not only in Japan but also worldwide. Increasing global demand of bluefin tuna is threatening its natural population towards extinction. Therefore, various committees such as the Western and Central Pacific Fisheries Commission (WCPFC) have been discussing conservation and management of tuna resources every year. The Pacific bluefin tuna was reclassified from Vulnerable to Near Threatened in 2021 according to the International Union for Conservation of Nature (IUCN), but it is still under critical conditions. To remedy this alarming situation, Kindai University conducted full cycle aquaculture of bluefin tuna in 2002 [14]. Full-cycle aquaculture is an important technology that does not rely on natural resources and enables a stable supply of bluefin tuna, without threatening the natural population. However, the full cycle aquaculture of bluefin tuna also has various problems, such as disease and feeding regime during larval production [14,15].

The intestinal microbiota is important for the health and development of bluefin tuna, however, studies on this topic are limited [16,17]. Minich et al. [17] investigated the microbial diversity associated with mucosal membranes, including the gut farmed southern bluefin tuna (Thunnus maccoyii) and showed that the microbiota structure was affected by pontoon location and treatment with an antihelminthic drug. Gatesoupe et al. [16] investigated the change in intestinal microbiota in Atlantic bluefin tuna Thunnus thynnus larvae and showed that the microbiota varied greatly among individuals. Pathogenic bacteria such as Photobacterium and Vibrio species have been shown to cause disease in tuna [18-21]. Given that blood flukes often cause serious problems in bluefin tuna aquaculture [22] and also the ‘pathobiome’ concept [23], it is important to understand how intestinal bacteria directly and indirectly affect the health of the host.

In this study, we investigated the succession process of intestinal bacteria during the seed production in full cycle aquaculture of bluefin tuna. We hypothesized that bacteria derived from rearing water and feeds would colonize the intestinal tract of bluefin tuna, thereby shaping the gut bacterial community. In this study, we used healthy bluefin tuna larvae produced commercially at Susami Hatchery, Aquaculture Technology and Production Center, Kindai University.

Materials and methods

Sample collection

All experimental procedure was approved by the Institutional Animal Care and Use Committee and was conducted in accordance with the Kindai University Animal Experimentation Regulations (2021-A-00118). This study was performed in accordance with ARRIVE guidelines. Informed consent was not required for this study.

Samples were collected three times: once in 2012 (expressed as Exp12) and twice in 2013 (expressed as Exp13r1 and Exp13r2). We used a full-cycle aquaculture of Pacific bluefin tuna Thunnus orientalis, rearing water, and feeds. The larval fish density was 150,000 fish/30 t tank in Exp12, and 250,000 and 500,000 fishes/50 t tank for Exp13r1 and Exp13r2, respectively. The periods were set from the egg to offshore for the analysis, and the feeding schedule was performed according to Sawada et al. [14] (S1 Fig and S1 Table).

The rearing water and feed were sampled immediately after feeding to the fish, and fish samples were taken immediately before feeding on the next day to investigate the relationship between intestinal and environmental bacteria. The fish samples were collected with a beaker or fry net, depending on the swimming capability. For the feed samples, rotifers (Brachionus plicatilis), Artemia (Artemia salina) nauplii, and feeder larvae (striped beakfish) were placed in a beaker, followed by collection onto a 60-μm-opening nylon membrane, and the commercial pellets were placed in a plastic bag. The rearing water pre-filtered, to remove feeds using a 60-μm-opening nylon membrane, was collected into a 50 mL centrifuge tube. All samples were immediately stored at −60°C until further analysis. The beaker, net, and nylon membranes were washed with hypochlorite.

For the egg and fish samples, we first performed a washing step to remove the bacteria from the samples. Water around the egg (71–120 eggs) and larval (16–44 fish; total length (TL) 2.0–4.6 mm) samples on the nylon membrane were removed with a 47 mm diameter GF/C filter (1822–047, Whatman, Kent, UK; pre-combusted at 450°C for 2 h) and washed with 5 mL of 0.22-μm-filtered seawater. This washing step was repeated seven times to prevent bacterial contamination from the rearing water. The whole fish body was subjected to DNA extraction because it was difficult to remove only the intestine. For experimental convenience, the bacteria of eggs and small fish were treated as intestinal bacteria. As for the larger larvae and fry (5–44 fish, TL 7.2–65.6 mm), the water around each individual fish was removed in the same manner as described above and washed with 1 mL of 0.22-μm-filtered seawater. This washing step was repeated seven times. The intestines of larvae and fry were removed with a sterilized razor blade and placed into a ZircoPrep Mini tube (FGM50M, NIPPON Genetics Co. Ltd., Tokyo, Japan) containing 400 μL of TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0).

For the feed samples, rotifers, Artemia, and the feeder larvae of striped beakfish were washed in the same manner as the egg and fish samples. The weights of rotifer, Artemia, and feeder larvae were 33.2–53.5, 40.5–101.2, and 35.4–82.7 mg, respectively. The commercial pellets were weighed to approximately 50 mg. Each sample was placed in a ZircoPrep Mini tube containing 400 μL of TE buffer. For the water sample, 10 mL of water was filtered through a 0.2-μm-pore size polycarbonate membrane filter (16020002, Advantec Toyo Co. Ltd., Tokyo, Japan) under vacuum < 0.02 MPa. A 0.45-μm-pore size nitrocellulose membrane filter (HAWP02500, Millipore, MA, USA) was used as a base filter to prevent the contamination of filtered seawater. The polycarbonate filter was placed in a ZircoPrep Mini tube containing 400 μL of TE buffer. Filtered seawater was also subjected to filtration in the same manner to check for bacterial contamination.

DNA extraction

The ZircoPrep Mini tubes containing the samples were vortexed for 15 min using a Vortex-Genie 2 (G-560, Scientific Industries, Inc., NY, USA). Lysozyme (final concentration 15 mg/mL) was added to the tubes and incubated for 60 min at 37°C with gentle mixing. After incubation, proteinase K (2 mg/mL) and SDS (1.8%) were added and incubated for 60 min at 55°C with gentle mixing. After incubation, equal amounts of phenol were added to the supernatant and centrifuged at 20,630 × g for 5 min at 20°C. This phenol step was repeated three times. An equal amount of phenol/chloroform supernatant was added and centrifuged at 20,630 × g for 5 min at 20°C. Ethanol precipitation was subsequently performed. The DNA was dissolved in 30 μL of TE buffer, and the concentration was measured at 260 nm using a NanoDrop spectrophotometer (ND-1000, Thermo Fisher Scientific Inc., MA, USA).

Automated ribosomal intergenic spacer analysis (ARISA)

PCR amplification was performed in a 20 μL reaction mixture using TaKaRa Ex Taq polymerase (Takara Bio Inc., Shiga, Japan) in a C1000 Thermal Cycler (Bio-Rad, Richmond, CA, USA). The mixture contained 1 × PCR buffer, 2 mM MgCl2, 0.2 mM of each dNTP, primers, 0.8 mg/mL bovine serum albumin, and 0.025 units TaKaRa Ex Taq polymerase. The primers used in this study were 16S-1392F (5ʹ-G[C/T]A CAC ACC GCC CA-3ʹ) and 23S-125R (5ʹ-GGG TT[C/G/T] CCC CAT TC[A/G] G-3ʹ) labeled with 6-FAM at the 5ʹ end [24]. The following PCR cycling conditions were used: initial denaturation step at 95°C for 3 min, 30 cycles of 30 s at 95°C, 30 s at 56°C, and 45 s at 72°C, followed by a final extension step at 72°C for 7 min. PCR products were analyzed using 2.0% agarose gel electrophoresis.

PCR products were purified using a PCR Clean-Up Mini Kit (Favorgen, Ring-Tung, Taiwan), and the concentration was measured using a NanoDrop spectrophotometer. Purified products were diluted to 25 ng/μL with TE buffer, and 100 ng of a standardized amount were loaded into the fragment analysis. Products were then run for 3 h on an ABI 3130xl Genetic Analyzer (Applied Biosystems, Waltham, MA, USA) with a GeneScan 1200 LIZ dye Size Standard (Applied Biosystems, Waltham, MA, USA). The ARISA peak patterns were analyzed using PeakStudio version 2.2 [25]. According to Chow et al. [26], ARISA peaks were manually binned with maximum bin sizes of 1, 2, 3, and 5 bp for 390–450, 450–650, 650–900, and 900–1200 bp, respectively. Each ARISA peak was regarded as an operational taxonomic unit (OTU), and the relative abundance (peak height divided by the cumulative height of all peaks in the sample) was subjected to community analysis.

Community analysis

All analyses were performed using R software [27]. Diversity indices (OTU number, logarithm of inverse Simpson and Shannon-Weiner indices) were calculated, using the ’Vegan’ package [28]. Scatter plots of OTU abundance numbers and diversity indices during experiments were generated and the Pearson’s coefficients were calculated using the ’ggpubr’ package [29]. Bray-Curtis dissimilarities were calculated, and the distance matrix was analyzed using the between-group average linkage method for clustering and the ’Vegan’ package. We performed a similarity profile (SIMPROF) test (9999 permutations with a p < 0.0001) to determine significant clusters among samples, using the ’clustsig’ package [30]. Also, a principal coordinates analysis (PCoA) and an Adonis test (9999 permutations) were used to show OTU variations among samples, using ’ape’ [31] and ’Vegan’ [28] packages. Venn analysis was performed between the early and late stages of larvae as the result of cluster analysis, using the ’VennDiagram’ package [32], to reveal characteristic OTUs. The derived OTUs and abundance were visualized by a heatmap using the ’tidyr’ package [33].

Results

Bacterial diversity

During the experiments, bacterial OTU numbers of rearing water, feeds, and intestinal samples ranged from 7 to 81 OTUs, 2 to 80 OTUs, and 1 to 49 OTUs, respectively (Tables 1–3); however, no PCR products were obtained from some intestinal samples (S1 Table). The number of OTUs in the rearing water sample tended to be smaller before the beginning of feeding, and those of feed samples varied, even though the feed type was the same. The OTU numbers of intestinal samples decreased with the progression of larval growth (Pearson’s correlation coefficient r = −0.498, n = 27, p < 0.01), unlike those of seawater (r = 0.394, n = 31, p < 0.05) (S2 Fig). Simpson and Shannon-Wiener indices of rearing water sample ranged from 1.42 to 4.47 and 1.88 to 5.17, respectively, and those of feed samples ranged from 0.27 to 4.50 and 0.46 to 5.24, respectively. In intestinal samples, Simpson and Shannon-Wiener indices ranged from 0.08 to 4.04 and 0.23 to 4.68, respectively. No correlation of the indices with the progression of larval growth was observed for all samples of rearing water, feed, or intestine.

Table 1

Diversity indices of bacterial communities in Exp12.

	Rearing water												Feed											Intestine
	W01	W02	W05	W08	W11	W13	W16	W19	W22	W25	W28	W31	R02	R05	R08	R11	A13	AL16	L19	P19	L22	P25	P28	G-1	G01	G03	G06	G09	G12	G14	G17	G20	G26	G29	G31
OTU number	11	42	36	42	28	40	41	40	36	32	19	39	57	80	33	17	18	29	36	38	45	34	31	10	8	34	10	25	23	15	9	7	6	6	6
H’	3.02	4.10	4.48	4.11	3.97	4.36	4.29	4.24	3.94	3.91	3.24	3.06	5.05	5.24	4.04	1.94	3.30	3.21	4.28	3.41	4.38	3.49	2.97	3.07	2.29	3.16	3.28	4.26	3.62	3.52	2.98	2.57	2.50	2.20	2.41
log(1/D)	2.73	3.43	4.08	3.34	3.46	3.78	3.75	3.43	2.98	2.84	2.50	1.89	4.47	4.42	3.44	1.23	2.55	2.47	3.74	2.39	3.81	2.47	1.76	2.85	1.92	1.74	3.25	3.98	3.00	3.16	2.82	2.42	2.42	1.99	2.30

W, water; R, rotifer; A, Artemia; L, feeder larvae; P, commercial pellet; G, intestine. The number after letters indicates the day after hatching of bluefin tuna. Day 0 is the day of hatching and Day -1 represents the unhatched egg.

Table 3

Diversity indices of bacterial communities in Exp13r2.

	Rearing water										Feed								Intestine
	W-1	W01	W02	W05	W08	W12	W16	W20	W24	W29	R01	R02	R05	R08	R12	L13	L16	L20	G-1	G00	G03	G06	G21	G25	G29
OTU number	7	9	9	15	10	44	56	58	58	81	10	12	50	28	15	2	20	18	33	13	4	2	3	7	8
H’	1.88	2.07	2.65	2.36	2.28	3.94	4.18	3.91	4.18	5.17	2.27	2.75	3.94	2.81	3.56	0.46	3.39	3.62	4.39	2.72	0.23	0.98	1.58	2.41	2.57
log(1/D)	1.42	1.57	2.41	1.47	1.63	3.12	3.32	3.10	3.40	4.47	1.74	2.33	3.16	2.08	3.30	0.27	2.95	3.29	3.78	2.11	0.08	0.96	1.57	2.10	2.34

W, water; R, rotifer; A, Artemia; L, feeder larvae; P, commercial pellet; G, intestine. The number after letters indicates the day after hatching of bluefin tuna. Day 0 is the day of hatching and Day -1 represents the unhatched egg.

Bacterial community structure

ARISA peak patterns were separated into two clusters, communities of rearing water and feed, regardless of the sampling year (Figs 1A and S3A). The early and late stages of intestinal communities appeared to resemble communities of rearing water and feed, respectively. Examining the communities of intestinal samples in detail, 17 days after hatching (DAH) formed the same cluster (p < 0.0001 SIMPROF and Adonis tests, Cluster-L, Figs 1B and S3B). Venn analysis between Cluster-L and the other clusters (Cluster-E) from the cluster analysis revealed characteristic OTUs: 130 in Cluster-E, 4 in Cluster-L, and 13 in both clusters (Fig 2 and S2 Table in S1 File). Although most of the OTUs were observed in the early stage of larvae (Cluster-E), the OTUs of intestinal samples in the late stage (Cluster-L) consisted of a small number of OTUs (Cluster-L ∩ Cluster-E). In response to this result, the occurrence pattern and abundance variation of characteristic OTUs in both clusters (Cluster-L ∩ Cluster-E) and in only Cluster-L (Cluster-L ∖ Cluster-E) were visualized by heatmap analysis (Fig 3). Regardless of the year of the experiments, six OTUs (OTU507.9, OTU553.3, OTU591.1, OTU653.1, OTU789.5, and OTU814.4) were particularly predominant and frequently observed in intestinal samples. In some cases (e.g., 12_G26 in Fig 3), intestinal samples contained only these six OTUs. Among them, OTU507.9, OTU591.1, OTU653.1, and OTU789.5 were detected throughout the experiment and became dominant after 17 DAH. Notably, these OTUs were also detected in the egg samples (12_G-1 and 13r2_G-1 in Fig 3). OTU553.3 and OTU814.4 were less dominant in the early stage, but more frequently observed after 17 DAH. Most of the other OTUs observed in the rearing water and feed samples were not predominant in the intestinal samples. The OTU421.9 was observed frequently and predominantly in the samples of rearing water and feeds and the intestinal samples before 17 DAH. However, it was seldom observed in the intestinal samples after 17 DAH. Four OTUs (OTU420.5, OTU555.0, OTU622.9, and OTU669.4) in Cluster-L (Cluster-L ∖ Cluster-E) were detected but were not predominant throughout the experiment in the intestinal samples as well as in the rearing water and feed samples. The distribution of the other OTUs (Cluster-E ∖ Cluster-L) is shown in S4 Fig.

Fig 1

Cluster analysis of bacterial community structures.

(a) Cluster constructed using all samples, rearing water (blue letters), feed (green letters) and intestine (brown letters). (b) Cluster constructed using only intestinal samples. Sample names are represented by experimental year, sample type, and DAH of bluefin tuna; 0 DAH is the day of hatching and Day -1 represents the unhatched egg. For example, 12_G17 represents intestinal samples at 17 DAH in 2012. W, water; G, intestine; R, rotifer, A, Artemia; L, feeder larvae (striped beakfish); P, commercial pellet.

Fig 2

Venn diagram of OTUs in Cluster-E and Cluster-L from cluster analysis.

The name of clusters corresponds to descriptions in Figs 1B and S3B.

Fig 3

Heatmap analysis of characteristic OTUs derived from Venn analysis.

The heatmap color corresponds to the square root of transformed relative abundance of each OTU in each sample. Sample names are represented by experimental year, sample type (rearing water, blue letters; feeds, green letters; intestine, brown letters), and DAH of bluefin tuna; 0 DAH is the day of hatching and Day -1 represents the unhatched egg. For example, 12_G17 represents intestinal samples of 17 DAH in 2012. The order of samples is based on the hatching day of larvae. W, water; G, intestine; R, rotifer, A, Artemia; L, feeder larvae (striped beakfish); P, commercial pellet.

Discussion

In the present study, we demonstrated the succession process of intestinal bacteria in bluefin tuna, in full-cycle aquaculture for the first time. For the analysis, we used a PCR-based fingerprinting analysis, ARISA, which seemed to detect particularly the dominant bacteria in a sample, with some bias [34]. Kashinskaya et al. [35] used several molecular methods to compare differences in the intestinal bacterial communities of Prussian carp and reported that the PCR-based cloning method successfully appeared to detect dominant bacteria in the samples. It is also reported that there is ecological coherence of bacterial diversity pattern between ARISA and deep sequencing techniques [36-38]. Therefore, we intended to see a similar trend using the PCR-based method, ARISA, in this study. The dominant bacteria may have a great impact on host development, as can be seen from the probiotic strategy. The simplification in the present study would be key to revealing the succession process of intestinal microbiota. In addition, bacterial contamination from the body surface might be observed in samples of the smaller larvae due to the handling of the whole fish body. Yoshimizu et al. [39] detected bacterial counts in the range of 102−103 colony forming units/g of body weight from the body surface of the sac fry. However, the contamination risk could be significantly low in this study as our washing steps were notably more frequent (at least 107 times higher) than those of Yoshimizu et al. [39]. Our experiments yield three major findings: 1) the intestinal bacteria in the early stage of larvae were affected by rearing water and feed, 2) the specific intestinal bacteria in the late stage of larvae became dominant regardless of the diversity of rearing water and feed, and 3) the specific intestinal bacteria in the late stage of larvae, especially after 17 DAH, originated from the egg surface.

This study suggests that the management of rearing water and live feeds, such as rotifers, is important for controlling the bacterial communities in seed production because they affect the intestinal bacteria in the early stage of larvae (Figs 1A and S3A). Other studies have also shown that fish intestinal bacteria are mainly affected by feed [12,40-43], water [44,45], and both [39,46-48]. Various types of water used for seed production have different bacterial communities [49]. They mentioned that the management of various waters in seed production was a key point because pathogenic microbes would increase during larval development even though the initial rearing water was free of pathogens. Effective management of rearing water has been shown to be ‘green-water’, a type of rearing water containing a large amount of microalga [50-53]. Studies have shown that live microalgae such as Nannochloropsis oculata and Chlorella vulgaris could eradicate the pathogenic bacterium Vibrio anguillarum by collaborating with the indigenous probiotic bacterium Sulfitobacter in rearing water. This ‘green-water’ technique is necessary to maintain the bacterial communities in desirable condition in the early stage of larvae.

Specific bacteria in the late stage of larvae in this study would colonize the favorable intestinal environment of bluefin tuna (Figs 1B and S3B) by the host and/or interactions of host-microbe and microbe-microbe [39,46-48,54-56]. A pioneering study of the succession process of intestinal bacteria by Yoshimizu et al. [39] indicated the possibility of regulation by the host. They reported that intestinal bacteria in the early stage of salmonid larvae were affected by the rearing water and diet, and then the developed intestinal environment by the activation of digestive tract would establish the characteristic intestinal microbiota. Ontogenetic development of the digestive tract and enzyme activity in bluefin tuna has been reported to develop relatively fast [57,58]. In bluefin tuna farmed at Kindai University, Miyashita et al. [59] showed that activities of pepsin-like and trypsin-like enzymes increased simultaneously with the development of stomach and pyloric caecum functions during the transitional period of juvenile tuna (17–25 DAH) as the rate of percentage of preanal length to standard length increased. The selection pressure would be due to host regulation as well as host-microbe and microbe-microbe interactions [55]. Changes in microbiota contribute to larval health and growth by improving host abilities.

The possible source of characteristic intestinal bacteria in the late stage of larvae would be the egg surface where these characteristic bacteria were also found (Fig 3), and the bacteria should compose of ‘core microbiota’ [60,61] in bluefin tuna. Fish eggs contain many bacteria on their surfaces [39,41,62], although sometimes the associated bacteria can cause mortality in marine fish [63]. Lauzon et al. [64] demonstrated that probiotic bathing treatment of eggs resulted in the establishment of added bacteria in the larval intestine, leading to increased survival, stress tolerance, and growth of host larvae. Notably, they highlighted that bacterial control was mainly evidenced prior to larval feeding, suggesting the importance of bacterial establishment on the egg surface. The mechanism of transfer from egg surface to larva is still unknown in bluefin tuna, but one possibility is ingestion of egg surface bacteria via grazing on egg debris and/or drinking seawater [62,65]. In the present study, we demonstrated that bacteria on the egg surface would be an important key species for determining ‘core microbiota’ of intestine in bluefin tuna. As bacteria associated with fish egg surface are likely to be opportunistic species [63], the management of eggs shortly after release, including disinfection [15] and preparation of rearing water, would be an important process. Further investigation is needed to determine whether the key species are still colonized in the adult bluefin tuna.

In conclusion, a clear succession process of intestinal bacteria in full-cycle aquaculture of Pacific bluefin tuna was shown for a total of three times of seed production. While bacteria in the early stage of larvae were affected by bacteria of rearing water and feeds, in the late stage (especially, after 17 DAH when the intestinal environment was developed), they changed to specific bacteria originating from the egg surface, which are likely to be composed of the ‘core microbiota’. Thus, we highlighted the importance of proper management in the seed production process, including egg management for the aquaculture of healthy fish. Further research is needed to determine the bacterial function and whether the bacteria are well colonized in adult fish.

Growth curve of Pacific bluefin tuna and feeding schedules.

The growth of bluefin tuna is indicated as a plot graph, and the feeding schedules are shown as a bar graph at the top of the plot graph. Exp12, Exp13r1, and Exp13r2 indicate the year (2012 or 2013) and round (r1 or r2) in which the experiment was performed.

(TIF)

Click here for additional data file.

Variation of OTU abundance and diversity indices during experiments.

a)–c) Show the number of OTUs in intestine, feed, and water samples, respectively. d)–f) Show indices of the Shannon-Wiener (H’) (open circle) and Simpson (log(1/D)) (filled orange circle) in intestine, feed, and water samples, respectively. r and p values describe Pearson’s correlation.

(TIF)

Click here for additional data file.

Principal coordinate analysis (PCoA) of bacterial community structures.

(a) PCoA constructed using all samples, rearing water (blue letters), feed (green letters) and intestine (brown letters). (b) PCoA constructed using only intestinal samples, the early (yellow group) and late (brown group) stages based on Adonis test (p = 0.0001). Sample names are represented by experimental year, sample type, and DAH of bluefin tuna; 0 DAH is the day of hatching and Day -1 represents the unhatched egg. For example, 12_G17 represents intestinal samples of 17 DAH in 2012. W, water; G, intestine; R, rotifer, A, Artemia; L, feeder larvae (striped beakfish); P, commercial pellet.

(TIF)

Click here for additional data file.

Heatmap analysis of all intestinal OTUs.

The heatmap color corresponds to the square root of transformed relative abundance of each OTU in each sample. Sample names are represented by experimental year, sample type (rearing water, blue letters; feeds, green letters; intestine, brown letters), and DAH of bluefin tuna; 0 DAH is the day of hatching and Day -1 represents the unhatched egg. For example, 12_G17 represents intestinal samples of 17 DAH in 2012. The order of sample is based on the hatching day of larvae. W, water; G, intestine; R, rotifer, A, Artemia; L, feeder larvae (striped beakfish); P, commercial pellet.

(TIF)

Click here for additional data file.

Sample list in this study.

(XLSX)

Click here for additional data file.

Relative abundance of all OTUs in this study.

(XLSX)

Click here for additional data file.

13 May 2022

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Reviewer #1: Partly

Reviewer #2: Partly

**********

2. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: N/A

Reviewer #2: I Don't Know

**********

3. Have the authors made all data underlying the findings in their manuscript fully available?

The PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.

Reviewer #1: Yes

Reviewer #2: Yes

**********

4. Is the manuscript presented in an intelligible fashion and written in standard English?

PLOS ONE does not copyedit accepted manuscripts, so the language in submitted articles must be clear, correct, and unambiguous. Any typographical or grammatical errors should be corrected at revision, so please note any specific errors here.

Reviewer #1: Yes

Reviewer #2: Yes

**********

5. Review Comments to the Author

Please use the space provided to explain your answers to the questions above. You may also include additional comments for the author, including concerns about dual publication, research ethics, or publication ethics. (Please upload your review as an attachment if it exceeds 20,000 characters)

Reviewer #1: The author examined the gut microbiota of blue fin tuna from the offshore cage rearing system by Automated approach for ribosomal intergenic spacer (ARISA) analysis. However, even with dissimilarity tree analysis, this method can give only preliminary dominant microbiota change but not enough to state the solid succession limited by the lack of bacteria annotation and the multi-analysis of the microbiota change of feeds and rearing water due to seemingly technical issue of DNA extraction and the uneven distribution of bacteria abundance.

Major issue:

1. The biggest issue with this work is still the accuracy of ARISA. Have author(s) done any single 16s for getting idea on whether How different OTU number or major OTU number difference? Even though the author(s) claimed ARISA can represent major dominant bacteria?

2. Figure 1. Can author(s) use PCoA to represent the beta-diversity of dominant bacteria from ARISA? PCoA can represent better than tree in terms of distinguishing the different microbiota group along with statistical analysis such as ANOSIM/ADONIS.

3. Venn diagram: how were the 117 and 4 distinguished bacteria distributed in the cluster E and L samples respectively?

4. It seems the type of feeding organisms changed during the seedling (Table. S1) and their microbiota could change (Fig. S2e). Can authors show how different the microbiota is between different feeding organisms? This will tell us how if the feeds can really impact the intestinal microbiota.

5. Figure 3. It seems a lot of OTUs (from 789.5- 774.7) were missing in the most of the rearing water and feed samples but presenting in either egg or larvae intestine. If so, then host factor can be more critical than feed/rearing water in microbiota succession?

6. Line179-180. however, no PCR products were obtained from some intestinal samples. Could it be the issue of bacteria DNA extraction? if so, then this will also affect the diversity and total abundance of the microbiota.

Minor issue:

1. Figure S2. – Y-axis. Number of OTU instead of Number if OTU. P value of R2?

Reviewer #2: The manuscript entitled “Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae” is try to investigated the succession process of intestinal bacteria during larvae rearing and their relationship between rearing water and feed (rotifer, artemia, feeder larvae and commercial pellets. The bacterial community was detected by an “automated ribosomal intergenic spacer analysis (ARISA)” methods and the operational taxonomic unit (OTUs), logarithm of inverse Simpson and Shannon-Weiner indices were used as the diversity indices. I have to say that this experiment is difficult to perform due to the sample collection was difficult. This manuscript also may give us very different results. However, there are some issues still need to be clarified before this draft can be published.

1. Since the fish samples were collected individually, how to define the data collected from these fish is succession?

2. I’m curious what is the OTUs number variation between each pooled sample?

3. In the manuscript, the authors mentioned the bacterial community in fish larvae have great different in the late larvae stage (after 17 dph). Even the number of OTUs seems reduce after this stage. But how to define this observation.

4. As I understand, the next generation sequencing (NGS) methods had also can using in analyse bacterial community composition in gut or environment for some years. Not only give you the different OTUs reactions, but also can give you their composition. How you compare the ARISA and NGS methods using in bacterial community ananlysis in your research.

**********

6. PLOS authors have the option to publish the peer review history of their article (what does this mean?). If published, this will include your full peer review and any attached files.

If you choose “no”, your identity will remain anonymous but your review may still be made public.

Do you want your identity to be public for this peer review? For information about this choice, including consent withdrawal, please see our Privacy Policy.

Reviewer #1: No

Reviewer #2: No

[NOTE: If reviewer comments were submitted as an attachment file, they will be attached to this email and accessible via the submission site. Please log into your account, locate the manuscript record, and check for the action link "View Attachments". If this link does not appear, there are no attachment files.]

While revising your submission, please upload your figure files to the Preflight Analysis and Conversion Engine (PACE) digital diagnostic tool, https://pacev2.apexcovantage.com/. PACE helps ensure that figures meet PLOS requirements. To use PACE, you must first register as a user. Registration is free. Then, login and navigate to the UPLOAD tab, where you will find detailed instructions on how to use the tool. If you encounter any issues or have any questions when using PACE, please email PLOS at figures@plos.org. Please note that Supporting Information files do not need this step.

25 Jun 2022

PONE-D-21-35860

Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae

PLOS ONE

1. The data cannot support the conclusions. PLOS ONE is designed to communicate primary scientific research, and welcome submissions in any applied discipline that will contribute to the base of scientific knowledge. But the data of this manuscript cannot support the conclusions.

> This paper presents a molecular ecological approach to the succession pattern of intestinal bacteria. We did not perform species identification because the main purpose of the present manuscript is to understand how environmental factors such as feed and rearing water influence intestinal microbiota. We hope this meets the aim of your journal.

2. This manuscript has the statistical analysis problem.

> We have addressed this problem.

3. The revised manuscript needs to address each of the comments of the reviewers.

> We have responded to the reviewers’ comments individually.

Reviewers' comments:

Reviewer's Responses to Questions

Comments to the Author

1. Is the manuscript technically sound, and do the data support the conclusions?

The manuscript must describe a technically sound piece of scientific research with data that supports the conclusions. Experiments must have been conducted rigorously, with appropriate controls, replication, and sample sizes. The conclusions must be drawn appropriately based on the data presented.

Reviewer #1: Partly

Reviewer #2: Partly

2. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: N/A

Reviewer #2: I Don't Know

3. Have the authors made all data underlying the findings in their manuscript fully available?

The PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.

Reviewer #1: Yes

Reviewer #2: Yes

4. Is the manuscript presented in an intelligible fashion and written in standard English?

PLOS ONE does not copyedit accepted manuscripts, so the language in submitted articles must be clear, correct, and unambiguous. Any typographical or grammatical errors should be corrected at revision, so please note any specific errors here.

Reviewer #1: Yes

Reviewer #2: Yes

5. Review Comments to the Author

Please use the space provided to explain your answers to the questions above. You may also include additional comments for the author, including concerns about dual publication, research ethics, or publication ethics. (Please upload your review as an attachment if it exceeds 20,000 characters)

Reviewer #1: The author examined the gut microbiota of blue fin tuna from the offshore cage rearing system by Automated approach for ribosomal intergenic spacer (ARISA) analysis. However, even with dissimilarity tree analysis, this method can give only preliminary dominant microbiota change but not enough to state the solid succession limited by the lack of bacteria annotation and the multi-analysis of the microbiota change of feeds and rearing water due to seemingly technical issue of DNA extraction and the uneven distribution of bacteria abundance.

Major issue:

1. The biggest issue with this work is still the accuracy of ARISA. Have author(s) done any single 16s for getting idea on whether How different OTU number or major OTU number difference? Even though the author(s) claimed ARISA can represent major dominant bacteria?

> In this manuscript, we aimed to investigate how intestinal microbiota communities are formed with respect to microbes found in food and water. Therefore, we used ARISA, which is reported to be able to estimate microbial diversity and community composition in terms of OTUs (Fisher and Triplett, 1999). It is also a semi-quantitative technique (Brown et al, 2005) and has been utilized in many studies. In humans, it has been reported that the predominant bacteria have a significant impact on the host (Gerritsen et al, 2011), so understanding the microbial dynamics detected by ARISA will be very meaningful. However, as you pointed out, because species composition needs to be elucidated to utilize the data, a deep sequencing approach should be used in the future.

2. Figure 1. Can author(s) use PCoA to represent the beta-diversity of dominant bacteria from ARISA? PCoA can represent better than tree in terms of distinguishing the different microbiota group along with statistical analysis such as ANOSIM/ADONIS.

> Thank you for your suggestions. We performed PCoA and ADONIS (9999 permutations). These results are shown in supplementary Fig S3 because the eigenvalues of axes 1 & 2 were low, especially in PCoA of all samples (S3a Fig). The ADONIS results are also shown in Fig S3, which shows the adjusted R2 of 0.117 with a p value of 0.0001 among sample types, and an adjusted R2 of 0.213 with a p value of 0.0001 between Clusters E and L, indicating significant differences. Additionally, for the significance in the cluster analysis (Fig. 1), we performed SIMPROF analysis (9999 permutations with a p < 0.0001 according to ADONIS result). The results are shown in Fig. 1 as bars below each cluster. These results were similar to those yielded by PCoA and ADONIS.

3. Venn diagram: how were the 117 and 4 distinguished bacteria distributed in the cluster E and L samples respectively?

> Thank you for pointing this out. First, after rechecking all OTUs, we re-drew the Venn diagram (Fig. 2) because the OTU number detected from Cluster-E was wrong in the previous manuscript; the correct number was 130 OTUs, not 117 OTUs. The distribution of the 130 and 4 bacterial OTUs is shown in Fig S4.

4. It seems the type of feeding organisms changed during the seedling (Table. S1) and their microbiota could change (Fig. S2e). Can authors show how different the microbiota is between different feeding organisms? This will tell us how if the feeds can really impact the intestinal microbiota.

> Thank you for your comment. We showed the distribution in S4 Fig. Bacteria in feed sample are highly variable, but similar for the same feed items. Interestingly, especially during the feeding period of rotifers and Artemia, bacteria from them were detected as intestinal bacteria, but most of the bacteria were no longer detected after the period.

5. Figure 3. It seems a lot of OTUs (from 789.5- 774.7) were missing in the most of the rearing water and feed samples but presenting in either egg or larvae intestine. If so, then host factor can be more critical than feed/rearing water in microbiota succession?

> As you mentioned, we speculate that host factors, such as digestive enzymatic activity, are among the critical factors for determining microbiota composition. In this manuscript, we aimed to identify the general process of intestinal microbiota development, that is, to determine whether the microbiota come from water, feed items, or another source.

6. Line179-180. however, no PCR products were obtained from some intestinal samples. Could it be the issue of bacteria DNA extraction? if so, then this will also affect the diversity and total abundance of the microbiota.

> Unfortunately, DNA extraction did not work for some samples. However, the method in this manuscript (physical disruption by bead beating; chemical lysis by lysozyme, proteinase K and SDS; and phenol/chloroform purification) has been used for DNA extraction in many previous papers. Every step commonly reported for effective DNA extraction from intestinal samples (e.g., Daly et al., 2012; Yuan et al., 2012; Eun et al., 2014; Dubin et al., 2016; Han et al., 2018). One of the reasons why no PCR products were obtained from some intestinal samples could be low abundance of intestinal microbes during development of host digestive tract. In any case, as you mentioned, care should be taken in interpreting the results because DNA extraction could introduce bias with regard to microbial diversity.

Minor issue:

1. Figure S2. – Y-axis. Number of OTU instead of Number if OTU. P value of R2?

> Thank you for pointing this out. We have corrected “if” to “of” and showed P- values in S2 Fig.

Reviewer #2: The manuscript entitled “Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae” is try to investigated the succession process of intestinal bacteria during larvae rearing and their relationship between rearing water and feed (rotifer, artemia, feeder larvae and commercial pellets. The bacterial community was detected by an “automated ribosomal intergenic spacer analysis (ARISA)” methods and the operational taxonomic unit (OTUs), logarithm of inverse Simpson and Shannon-Weiner indices were used as the diversity indices. I have to say that this experiment is difficult to perform due to the sample collection was difficult. This manuscript also may give us very different results. However, there are some issues still need to be clarified before this draft can be published.

> Thank you for your comments.

1. Since the fish samples were collected individually, how to define the data collected from these fish is succession?

> The fish were reared under proper management, such as water quality, feed quality and quantity, and the number of reared fish from hatching to net-cage transfer. Fish were maintained under the same physiological conditions in each trial. In this manuscript, the formation of similar intestinal microbiota in the three rounds was considered evidence of this. We also collected multiple fish for each sample to decrease the effect of individual differences. Therefore, although fish samples were collected individually, we feel that our data are sufficient to indicate succession.

2. I’m curious what is the OTUs number variation between each pooled sample?

> “Bacterial abundance in rearing water, feed, and intestine samples ranged from 7–81, 2 –80, and 1– 49 OTUs, respectively (Tables 1–3)” lines 195–196 in “Revised Manuscript with Tracked Change”. This variation likely occurred because the predominant bacteria changed during the development of the host gastrointestinal tract. In this manuscript, we standardized the DNA amount loaded (100 ng) into the ARISA instrument to reduce the variation between samples (line 168).

3. In the manuscript, the authors mentioned the bacterial community in fish larvae have great different in the late larvae stage (after 17 dph). Even the number of OTUs seems reduce after this stage. But how to define this observation.

> This probably occurs due to the development of the host gastrointestinal tract, particularly the increased activities of digestive enzymes. Only bacteria that can tolerate the intestinal environment can survive.

4. As I understand, the next generation sequencing (NGS) methods had also can using in analyse bacterial community composition in gut or environment for some years. Not only give you the different OTUs reactions, but also can give you their composition. How you compare the ARISA and NGS methods using in bacterial community ananlysis in your research.

> As you mentioned, the NGS method is one of the strongest approaches for analysis of intestinal microbiota. However, in this manuscript, we focused on the relationship between the intestinal bacteria and the bacteria found in the feed and rearing water, which does not necessarily require identification of bacterial species composition. In fact, using ARISA, which is more economical than NGS, it is easy to compare multiple samples. We identified the importance of egg management for the formation of intestinal microbiota in the late stage. When compared to NGS data, the trend in major species may not change because ARISA detects predominant bacteria. However, because NGS can detect rare species, it may provide different findings in this respect.

6. PLOS authors have the option to publish the peer review history of their article (what does this mean?). If published, this will include your full peer review and any attached files.

If you choose “no”, your identity will remain anonymous but your review may still be made public.

Do you want your identity to be public for this peer review? For information about this choice, including consent withdrawal, please see our Privacy Policy.

Reviewer #1: No

Reviewer #2: No

Submitted filename: Response to Reiewers.docx

Click here for additional data file.

9 Aug 2022

A rebuttal letter that responds to each point raised by the academic editor and reviewer(s). You should upload this letter as a separate file labeled 'Response to Reviewers'.

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If you would like to make changes to your financial disclosure, please include your updated statement in your cover letter. Guidelines for resubmitting your figure files are available below the reviewer comments at the end of this letter.

If applicable, we recommend that you deposit your laboratory protocols in protocols.io to enhance the reproducibility of your results. Protocols.io assigns your protocol its own identifier (DOI) so that it can be cited independently in the future. For instructions see: https://journals.plos.org/plosone/s/submission-guidelines#loc-laboratory-protocols. Additionally, PLOS ONE offers an option for publishing peer-reviewed Lab Protocol articles, which describe protocols hosted on protocols.io. Read more information on sharing protocols at https://plos.org/protocols?utm_medium=editorial-email&utm_source=authorletters&utm_campaign=protocols.

We look forward to receiving your revised manuscript.

Kind regards,

Tzong-Yueh Chen, Ph.D.

Academic Editor

PLOS ONE

[Note: HTML markup is below. Please do not edit.]

Reviewers' comments:

Reviewer's Responses to Questions

Comments to the Author

1. If the authors have adequately addressed your comments raised in a previous round of review and you feel that this manuscript is now acceptable for publication, you may indicate that here to bypass the “Comments to the Author” section, enter your conflict of interest statement in the “Confidential to Editor” section, and submit your "Accept" recommendation.

Reviewer #1: (No Response)

Reviewer #2: All comments have been addressed

**********

2. Is the manuscript technically sound, and do the data support the conclusions?

The manuscript must describe a technically sound piece of scientific research with data that supports the conclusions. Experiments must have been conducted rigorously, with appropriate controls, replication, and sample sizes. The conclusions must be drawn appropriately based on the data presented.

Reviewer #1: Partly

Reviewer #2: Yes

**********

3. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: I Don't Know

Reviewer #2: Yes

**********

4. Have the authors made all data underlying the findings in their manuscript fully available?

The PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.

Reviewer #1: Yes

Reviewer #2: (No Response)

**********

5. Is the manuscript presented in an intelligible fashion and written in standard English?

PLOS ONE does not copyedit accepted manuscripts, so the language in submitted articles must be clear, correct, and unambiguous. Any typographical or grammatical errors should be corrected at revision, so please note any specific errors here.

Reviewer #1: Yes

Reviewer #2: Yes

**********

6. Review Comments to the Author

Please use the space provided to explain your answers to the questions above. You may also include additional comments for the author, including concerns about dual publication, research ethics, or publication ethics. (Please upload your review as an attachment if it exceeds 20,000 characters)

Reviewer #1: The revision has improved and given more clues in the potential bacterial community change. However, come technical issues have to be resolved or addressed in the context.

1. ARISA: I thank the author for his/her detailed reply on how ARISA was used in previous studies. Nonetheless, most of the studies using ARISA were back in the time NGS was not widely used. The current bacterial studies would need NGS (16S rDNA) analysis to really point out the bacterial community change. It would be more technical sound if the author can show a data to indicate that each ARISA peak really represents one bacterial species/genera in order to reach NGS resolution. (Is the binning method good enough to separate single bacteria type?)

2. Can the author explain how this is happening in my previous comment#4 regarding the highly variable bacterial community in feeds but later disappeared in the intestine?

3. Have author compared the ARISA peaks to parental fish from the early- and late-stage larva? I'm asking because part of the bacterial community may be acquired from the parents.

4. Can the author discuss the error by DNA extraction and the resulting potential error in bacterial community diversity/richness regarding to my previous comment# 6?

5. It would be good to put a rough estimate of bacterial change in phyla based on ARISA peak change (eg. from Gamma-proteobacteria to alpha-proteobacteria based on length) for NGS substitution.

6. As the author mentioned. OTU 429.1 is of my interest. Is there any possible way to know what bacteria it is? It would be powerful to show this bacteria genera/species to consolidate your conclusion in succession of bacterial community.

7. Fig S3 (b) : Please indicate color difference (What is yellow and brown group?).

8. Line 266-267 : The sentence "the dominant bacteria are likely to have a great impact on host development, as can be seen from probiotic strategy" is overstated here without evidence based data.

Reviewer #2: The manuscripte had been reviced, all commands have been addressed, and seems there are no more questions from me.

**********

7. PLOS authors have the option to publish the peer review history of their article (what does this mean?). If published, this will include your full peer review and any attached files.

If you choose “no”, your identity will remain anonymous but your review may still be made public.

Do you want your identity to be public for this peer review? For information about this choice, including consent withdrawal, please see our Privacy Policy.

Reviewer #1: No

Reviewer #2: No

**********

[NOTE: If reviewer comments were submitted as an attachment file, they will be attached to this email and accessible via the submission site. Please log into your account, locate the manuscript record, and check for the action link "View Attachments". If this link does not appear, there are no attachment files.]

While revising your submission, please upload your figure files to the Preflight Analysis and Conversion Engine (PACE) digital diagnostic tool, https://pacev2.apexcovantage.com/. PACE helps ensure that figures meet PLOS requirements. To use PACE, you must first register as a user. Registration is free. Then, login and navigate to the UPLOAD tab, where you will find detailed instructions on how to use the tool. If you encounter any issues or have any questions when using PACE, please email PLOS at figures@plos.org. Please note that Supporting Information files do not need this step.

26 Aug 2022

PONE-D-21-35860R1

Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae

PLOS ONE

1. The data cannot support the conclusions. PLOS ONE is designed to communicate primary scientific research, and welcome submissions in any applied discipline that will contribute to the base of scientific knowledge. But the data of this manuscript cannot support the conclusions. Some technical issues have to be resolved in the manuscript.

> In this manuscript, we aimed to know how the pattern of intestinal bacteria change as the host fish grow. Hence, we think that the method ARISA and obtained data are enough to address it and support the conclusions.

2. This manuscript has the statistical analysis problem.

> We have already addressed statistical problems pointed out by two reviewers. Reviewer #1, who initially indicated that there was a problem, no longer indicate it, and Reviewer's Responses to Questions #3 in this time is also “no problem”.

3. The revised manuscript needs to address each of the comments of the reviewers.

> We have responded to the reviewers’ comments individually.

Reviewer #1: The revision has improved and given more clues in the potential bacterial community change. However, come technical issues have to be resolved or addressed in the context.

> We would like to thank the Reviewer for the careful review of this manuscript.

1. ARISA: I thank the author for his/her detailed reply on how ARISA was used in previous studies. Nonetheless, most of the studies using ARISA were back in the time NGS was not widely used. The current bacterial studies would need NGS (16S rDNA) analysis to really point out the bacterial community change. It would be more technical sound if the author can show a data to indicate that each ARISA peak really represents one bacterial species/genera in order to reach NGS resolution. (Is the binning method good enough to separate single bacteria type?)

> In this manuscript, we aimed to investigate how intestinal microbiota communities are formed with respect to microbes found in food and water. We know your point, however ARISA allows us for the rapid comparison with high-throughput and cost-effective, and can also yield meaningful and valuable results. Previous studies show that each ARISA peak represents one bacterial species or strain and the binning method good enough to separate single species (e.g., Brown et al. 2005, Kovacs et al. 2010). Also, several reports show ecological coherence of bacterial diversity pattern between ARISA and NGS techniques (e.g., Bienhold et al. 2011, Gobet et al. 2014, Jami et al. 2014).

Thus, as you suggested, we added one sentence on the ecological coherence between the techniques: lines 253–255 in Revised_Manuscript “It is also reported that there is ecological coherence of bacterial diversity pattern between ARISA and deep sequencing techniques [36-38]”.

2. Can the author explain how this is happening in my previous comment#4 regarding the highly variable bacterial community in feeds but later disappeared in the intestine?

> We think this probably occurs due to the development of the host gastrointestinal tract, particularly the increased activities of digestive enzymes. Only bacteria that can tolerate the intestinal environment can survive and so the most of highly variable bacterial community in feeds cannot survive and colonize in the intestine (lines 281–294 in Revised_Manuscript).

3. Have author compared the ARISA peaks to parental fish from the early- and late-stage larva? I'm asking because part of the bacterial community may be acquired from the parents.

> No, we haven’t. The specific bacteria in the late stage larvae, 17 days after hatching, were also found on egg surface, egg shell. The egg shells were floating in the rearing water for long time after hatching. If the vertical transmission of bacteria from the parent fish occurred, the bacteria from the parents might give an influence on the early stage of larvae. Also, to our knowledge, a bacterial vertical transmission has not yet been reported as for bluefin tuna and there is only one paper on betanodaviruses in bluefin tuna (Sugaya et al., 2009). Anyway, as we have no data about the intestinal bacteria of adult bluefin tuna, this point, the vertical transmission from the parents, should be discussed in the future study. Thank you.

4. Can the author discuss the error by DNA extraction and the resulting potential error in bacterial community diversity/richness regarding to my previous comment# 6?

> This was the challenge in the past as we commented previously (e.g., Daly et al., 2012; Yuan et al., 2012; Eun et al., 2014; Dubin et al., 2016; Han et al., 2018). The results of DNA extraction are controversial even now as you know, and there may or may not be some errors. Therefore, readers interested in such paper should know this possibility, as all DNA extraction methods contain some errors.

5. It would be good to put a rough estimate of bacterial change in phyla based on ARISA peak change (eg. from Gamma-proteobacteria to alpha-proteobacteria based on length) for NGS substitution.

> Unfortunately, we cannot estimate this bacterial change. As you suggested, we are now planning a new experiment to compare the bacterial communities in not only the larvae but also the adult fish. This is because the present study showed us the important period of the formation of intestinal bacteria.

6. As the author mentioned. OTU 429.1 is of my interest. Is there any possible way to know what bacteria it is? It would be powerful to show this bacteria genera/species to consolidate your conclusion in succession of bacterial community.

> Unfortunately, we cannot know this bacterial species. However, one of the important results in this manuscript is that bacteria in feeds, controlled by the host fish, do not necessarily establish intestinal bacterial community. We think that the findings obtained in this manuscript are also valuable and appropriate for this journal.

7. Fig S3 (b) : Please indicate color difference (What is yellow and brown group?).

> We have addressed it. Thank you. “(b) PCoA constructed using only intestinal samples, the early (yellow group) and late (brown group) stages based on Adonis test (p = 0.0001).”

8. Line 266-267 : The sentence "the dominant bacteria are likely to have a great impact on host development, as can be seen from probiotic strategy" is overstated here without evidence based data.

> We have toned down this statement as follows: lines 254–255 "The dominant bacteria may have a great impact on host development, as can be seen from probiotic strategy"

Submitted filename: Response to Reiewers.docx

Click here for additional data file.

13 Sep 2022

Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae

PONE-D-21-35860R2

Dear Dr. Eguchi,

We’re pleased to inform you that your manuscript has been judged scientifically suitable for publication and will be formally accepted for publication once it meets all outstanding technical requirements.

Within one week, you’ll receive an e-mail detailing the required amendments. When these have been addressed, you’ll receive a formal acceptance letter and your manuscript will be scheduled for publication.

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Kind regards,

Tzong-Yueh Chen, Ph.D.

Academic Editor

PLOS ONE

Additional Editor Comments (optional):

Reviewers' comments:

Reviewer's Responses to Questions

Comments to the Author

1. If the authors have adequately addressed your comments raised in a previous round of review and you feel that this manuscript is now acceptable for publication, you may indicate that here to bypass the “Comments to the Author” section, enter your conflict of interest statement in the “Confidential to Editor” section, and submit your "Accept" recommendation.

Reviewer #1: All comments have been addressed

**********

2. Is the manuscript technically sound, and do the data support the conclusions?

The manuscript must describe a technically sound piece of scientific research with data that supports the conclusions. Experiments must have been conducted rigorously, with appropriate controls, replication, and sample sizes. The conclusions must be drawn appropriately based on the data presented.

Reviewer #1: Yes

**********

3. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: Yes

**********

4. Have the authors made all data underlying the findings in their manuscript fully available?

The PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.

Reviewer #1: Yes

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19 Sep 2022

PONE-D-21-35860R2

Succession of the intestinal bacterial community in Pacific bluefin tuna (Thunnus orientalis) larvae

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Table 2

Diversity indices of bacterial communities in Exp13r1.

	Rearing water									Feed							Intestine
	W-1	W01	W03	W07	W11	W13	W15	W19	W25	R03	A07	A11	L14	AL15	L16	L19	G01	G03	G04	G08	G12	G16	G25	G35
OTU number	61	7	29	27	60	18	22	27	14	5	42	21	63	64	25	7	49	45	17	19	1	1	6	9
H’	5.17	2.57	3.70	3.40	4.17	3.31	3.34	3.67	2.93	1.46	3.62	3.45	4.32	5.04	3.64	1.32	3.54	4.68	2.85	2.71	-	-	2.21	2.69
log(1/D)	4.41	2.40	2.99	2.48	3.25	2.98	2.76	3.16	2.36	0.95	2.48	2.85	3.35	4.50	2.83	0.78	2.58	4.04	2.28	2.13	-	-	2.03	2.41

W, water; R, rotifer; A, Artemia; L, feeder larvae; P, commercial pellet; G, intestine. The number after letters indicates the day after hatching of bluefin tuna. Day 0 is the day of hatching and Day -1 represents the unhatched egg.