Dietary lipid levels could improve growth and intestinal microbiota of juvenile swimming crab, Portunus trituberculatus

Accepted Manuscript Dietary lipid levels could improve growth and intestinal microbiota of juvenile swimming crab, Portunus trituberculatus

Peng Sun, Min Jin, Liyun Ding, You Lu, Hongna Ma, Ye Yuan, Qicun Zhou PII: DOI: Reference:

S0044-8486(17)31521-1 doi:10.1016/j.aquaculture.2018.02.018 AQUA 633066

To appear in:

aquaculture

Received date: Revised date: Accepted date:

1 August 2017 6 February 2018 12 February 2018

Please cite this article as: Peng Sun, Min Jin, Liyun Ding, You Lu, Hongna Ma, Ye Yuan, Qicun Zhou , Dietary lipid levels could improve growth and intestinal microbiota of juvenile swimming crab, Portunus trituberculatus. The address for the corresponding author was captured as affiliation for all authors. Please check if appropriate. Aqua(2017), doi:10.1016/j.aquaculture.2018.02.018

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ACCEPTED MANUSCRIPT Dietary lipid levels could improve growth and intestinal microbiota of juvenile swimming crab, Portunus trituberculatus

Laboratory of Fish Nutrition, School of Marine Sciences, Ningbo University, Ningbo 315211,

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Peng Suna,b, Min Jina, Liyun Dinga, You Lua, Hongna Maa, Ye Yuana, Qicun Zhoua,b,*

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China. b

Collaborative Innovation Center for Zhejiang Marine High-efficiency and Healthy Aquaculture,

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Ningbo University, Ningbo, China

* Corresponding author. Tel/Fax: +86-574-876-09878.

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E-mail address: [email protected] (Q. -C. Zhou)

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Abstract An 8 week feeding trial was conducted to evaluate the effects of dietary lipid levels on growth and intestinal microbiota of juvenile swimming crab. Three isonitrogenous (47% crude protein) diets were formulated to contain 5.8%, 9.9% and 15.1% crude lipid levels, respectively. Three groups of swimming

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crab (n = 60 per group) were randomly assigned to each diet. An Illumina-based sequencing method was

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used to analyze the intestinal bacterial composition of the crabs. The results indicated that swimming

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crab fed the diets containing 5.8% and 9.9% lipid had significantly higher weight gain and survival than those fed the 15.1% lipid diet. Proteobacteria, Fusobacteria and Tenericutes were dominant in the

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intestines of swimming crab regardless of the diet; the relative abundance of Fusobacteria

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decreased while Proteobacteria increased with increase of dietary lipid levels. Swimming crab fed the diets containing 9.9% and 15.1% lipid had a higher abundance of pathogenic bacteria than

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those fed the 5.8% lipid diet. These results indicate that dietary lipid levels could affect the

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composition of the intestinal microbiota; also, higher dietary lipid levels have a negative impact and increase the potential risk of disease in swimming crab.

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Key words: Portunus trituberculatus; Lipid levels; Growth performance; Intestinal bacterial

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composition; Illumina sequencing.

1. Introduction

Lipid, one of the macronutrients of formulated feed, plays a vital role as a principal source of energy and provides essential fatty acids for such important functions as forming the main components of biomembranes, acting as carriers of fat-soluble vitamins and playing a part in precursors of eicosanoids, hormones and enzyme cofactors in crustaceans (Watanabe, 1982; Higgs

ACCEPTED MANUSCRIPT and Dong, 2000). Optimal dietary lipid levels can also help spare protein (Helland et al., 1998; Torstensen et al., 2001). Meanwhile, the optimum dietary lipid level can reduce feed costs and increase economic benefits of farming (Peres and Olivia-Teres, 1999; Yoshii et al., 2010). However, excessive lipid in the diet can retard growth, decrease intake and reduce feed utilization

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(NRC, 2011). Furthermore, high-fat diets can increase fat deposition in organisms (Shiau and

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Huang, 1990). The requirements for lipid have been demonstrated in some crustaceans, ranging

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from 2 to 14 % (Sheen and D'Abramo, 1991; Sheen, 1997; Sheen and Wu, 1999; Catacutan, 2002; Cortés-Jacinto et al., 2005; Duan et al., 2011; Han et al., 2013a; Xu et al., 2013; Zhang et al., 2013,

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Zhao et al., 2015). The wide variation observed in dietary lipid requirement among crustacean

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species may be due to species, life stages, lipid sources, and culture environment (D'Abramo, 1997).

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The intestinal tract of crabs is inhabited by trillions of bacteria, and they play an important

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role in host health (Zhang et al., 2014a). Moreover, gut microbiota can perform many beneficial functions for the host, such as absorbing nutrients, improving energy production, and balancing

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the immune response (Hooper et al., 2002; Bates et al., 2006; Li et al., 2008; Hooper and

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Macpherson, 2010; Semova et al., 2012). Recent studies have shown that the gut microbial communities of several crustaceans are influenced by the nutritional habits of their hosts, as they metabolize part of the ingested food (e.g. cellulose digestion) and provide the host with important nutrients (Meziti et al., 2012). The intestinal bacterial are also involved in lipid metabolism, consisting of lipid storage, energy balance, bile acid synthesis and choline bio-utilization in mice (Backhed et al., 2004; Dumas et al., 2006) and human (Ridlon et al., 2006; Fukiya et al., 2009). A previous study had demonstrated that high-fat diets can affect bacterial diversity and composition

ACCEPTED MANUSCRIPT in the intestinal tract of mice (Daniel et al., 2014). Also, Zhang et al. (2014a) indicated that lipid sources with different fatty acid compositions could affect the composition of the intestinal microbiota of Litopenaeus vannamei. The intestinal bacteria in crustaceans have been studied in Eriocheir sinensis (Li et al., 2007; Chen et al., 2015; Zhang et al., 2016a), Fenneropenaeus

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chinensis (Liu et al., 2011), L. vannamei (Moss et al., 2000; Zhou et al., 2007a, b; Johnson et al.,

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2008; Luis-Villasenor et al., 2013; Zhang et al., 2014a; Zhang et al., 2016b; Suo et al., 2017), L.

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stylirostris (Cardona et al., 2016), Macrobrachium rosenbergii (Mente et al., 2016), Nephrops norvegicus (Meziti et al., 2010; 2012), Penaeus monodon (Chaiyapechara et al., 2012;

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Rungrassamee et al., 2013, 2014), and Scylla paramamosain (Li et al., 2012). In the only study

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performed in P. trituberculatus, it was demonstrated that the gut was dominated by Proteobacteria, Bacteroidetes, Fusobacteria, Tenericutes, and Acidobacteria at the phylum level (Zeng et al., 2016).

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However, no group has yet studied the effects of dietary lipid levels on intestinal microbiota in

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crabs.

The swimming crab is an important mariculture crustacean species and is widely distributed

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along the coastal waters of China, Japan, Korea, and Malaysia (Jin et al., 2015; Ding et al., 2017).

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Owing to its economic value, the P. trituberculatus industry is now expanding rapidly in China, and the aquaculture production of this crab reached approximately 117,772 metric tonnes in 2015 (China Fishery Statistical Yearbook, 2016). It has become one of the most important aquaculture crustaceans in China (Chen et al., 2006). To date, studies have primarily focused on the optimal requirements of lipid (Duan et al., 2011; Han et al., 2013a; Huo et al., 2014), essential fatty acids (Takeuchi et al., 1999a, b; Wang et al., 2012; Zhang et al., 2014d; Han et al., 2015), phospholipid (Li et al., 2014) and cholesterol (Han et al., 2013b). However, the relationship between lipid levels

ACCEPTED MANUSCRIPT and the intestinal microbiota has not been elucidated in P. trituberculatus. Therefore, the objectives of the present study were to analyze the biodiversity and composition of the intestinal microbiota with an Illumina HiSeq2500 PE250 sequencing of 16S rRNA gene, and evaluate the relationship between dietary lipid levels and intestinal bacterial profiles of juveniles swimming

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crabs.

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2. Materials and Methods 2.1. Diet preparation

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Three isonitrogenous (47% crude protein) diets were formulated to contain 5.8%, 9.9% and

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15.1% (low, medium, and high fat) crude lipid levels by gradually adding fish oil; and cellulose was used to adjust the different dietary lipid levels to keep isonitrogenous among all treatments.

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The formulation and approximate composition of the diets are presented in Table 1, and fatty acid

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composition of the experimental diets is shown in Table 2. Fish meal, wheat gluten meal, soybean protein concentrate and krill meal were used as protein sources; fish oil and soy lecithin were used

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as lipid sources; and dextrin was used as the carbohydrate source. All ingredients were ground into

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fine powder with a particle size less than 177 microns. The micro-components such as vitamin and mineral premixes, were then mixed thoroughly by the progressive enlargement method. Finally, lipids and distilled water (40% by w/w) were added to the premixed ingredients and mixed until homogenous in a Hobart-type mixer. Cold-extruded pellets were produced, and the pellet strands were diced into two uniform pellet sizes (pellet sizes: 3.0 mm in diameter and 5.0 mm in length and 4.0 mm in diameter and 7.0 mm in length) using a granulating machine (G-250, machine factory of South China University of Technology, Guangzhou, China), they were steamed for 30

ACCEPTED MANUSCRIPT min at 90 °C, and then air-dried to approximately 10% moisture. The diets were stored frozen in sealed plastic bags until their use in the feeding trial (stored at -20 °C). Insert Table 1 here. Insert Table 2 here.

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2.2. Crab rearing and experimental conditions The juvenile P. trituberculatus were obtained from the Lai-Fa nursery farm (Ningbo, China).

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Prior to the start of the feeding trial, the juvenile crabs were acclimated and fed trash fish for 1

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week. Juvenile crabs (approximate initial weight 21.1±1.1 g) were randomly sorted into 180

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rectangular plastic baskets (36 cm × 21 cm × 35 cm; length × width × height) in two cement pools (7.0 m × 5.0 m × 1.8 m; length × width × depth) and fed twice daily at 8:00 h and 18:00 h. Each

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experimental diet was randomly assigned to three replicates, twenty juvenile swimming crabs in each replicate, with one crab in each basket. Each basket was divided into two equal parts by a

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nylon screen; one partitioned area was filled with sand and served as the crab habitat, and the other area was filled with seawater and served as the feeding area. All groups of crabs were fed at

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the same rate, and the amount of feed was 6-8% of wet body weight. The feeding procedure has been previously described in Jin et al. (2015).

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All baskets were put in a cement pool, and the seawater of the pools was continuously aerated through multiple air tubes to maintain high dissolved oxygen levels. During the experimental period, water temperature in the pool was 26-30 °C, pH was 7.7-8.3, salinity was 26-28 ppt, ammonia nitrogen was lower than 0.05 mg L-1, and dissolved oxygen was 6.7-7.1 mg L−1. The pH, salinity, ammonia nitrogen and dissolved oxygen were measured using a YSI Proplus instrument (YSI, Yellow Springs, OH, USA). The feeding trial lasted for 8 weeks. 2.3. Sample collection and DNA extraction

ACCEPTED MANUSCRIPT At the termination of the feeding trial, the individual crab from each plastic basket were anaesthetized and then weighed. The whole intestines (including contents) of five crabs from each replicate were collected into autoclaved tubes for DNA extraction. Total bacterial community DNA was isolated with a TIANamp Micro DNA Purification Kit (Tiangen, Beijing, China). DNA

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concentration and purity were checked on 1% agarose gels. According to the concentration, DNA

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was diluted to 1 ng/μL; DNA specimens from five crabs in each replicate were mixed in equal

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concentrations, and the mixed samples were sent to Novogene Biological Information Technology Co. (Beijing, China) for analyses by high-throughput sequencing.

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2.4. Illumina high-throughput sequencing of barcoded 16S rRNA genes

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Bacterial DNA was used as the template for 16S rRNA gene V4-V5 region amplifica tion (Sun et al., 2013). The forward and reverse primers were 515F: 5'-GTGCCAGCMGC

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CGCGGTAA-3' and 907R: 5'-CCGTCAATTCCTTTGAGTTT-3' respectively (Zhang et al.,

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2016a, b). All PCR reactions were executed in a 30 μL system with 15 μL of Phusion ® High-Fidelity PCR Master Mix (New England Biolabs), 0.2 μM of forward and reverse pr

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imers, and approximately 10 ng template DNA. Thermal cycling conditions were: initial de

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naturation at 98 ℃ for 1 min, followed by 30 cycles of denaturation at 98 ℃ for 10 s, a nnealing at 50 ℃ for 30 s, extension at 72 ℃ for 30 s, then a final 6-min extension at 7 2 ℃ (Gunimaladevi et al., 2006). The PCR products were purified using a Gene JET Gel Extraction Kit (Thermo Scientific, Carlsbad, CA, USA). Sequencing libraries were generat ed by Illumina TruSeq DNA PCR-Free Library Preparation Kit (Illumina, USA) following manufacturer’s recommendations and index codes were added. The library quantity was ass essed on a Qubit 2.0 Fluorometer (Thermo Fisher Scientific, Carlsbad, CA, USA) and qua

ACCEPTED MANUSCRIPT lity in an Agilent Bioanalyzer 2100 system. Lastly, the library was sequenced on an Illum ina HiSeq 2500 platform and 250 bp paired-end reads were generated. The sequences obta ined in this paper are available in the GenBank with the accession number PRJNA389218. 2.5. Processing of Illumina Sequencing Data

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Paired-end reads were assigned to samples based on their unique barcode and truncate

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d by cutting off the barcode and primer sequence. We obtained the merged reads with FL

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ASH v1.2.7 (http://ccb.jhu.edu/software/FLASH/) based on overlapping regions between pair ed-end reads (Magoč and Salzberg, 2011), and the splicing sequences were called raw tags.

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Qualities filtering on the raw tags were performed according to the QIIME V1.7.0 (http://

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qiime.org/index.html) quality control process (Caporaso et al., 2010). The reads with ambig uous bases or truncated at any site of more than three consecutive bases receiving a Phre

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d quality score (Q) < 20 were discarded, as were truncated reads that had < 75% of their

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original length and a minimum length > 210 bp were selected in further analysis. Chime ra sequences were checked using an UCHIME algorithm (http://www.drive5.com/usearch /m

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anual/uchime_algo.html), and were then removed from the raw data (Edgar et al., 2011; H

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aas et al., 2011). Lastly, we obtained the high-quality effective tags. These sequences were classified into the same OTUs (operational taxonomic units) at an identity threshold of 9 7% similarity using UPARSE v7.0.1001 (http://drive5. com/uparse/) (Edgar, 2013). For eac h representative sequence, the Greengenes Database (http://greengenes.lbl.gov/cgi-bin/ nph-in dex.cgi) was used based on a RDP classifier (Version 2.2, http://sourceforge.net/projects/ rd p-classifier/) algorithm to annotate taxonomic information (DeSantis et al., 2006; Wang et al., 2007). After taxonomies had been performed, any OTUs identified as belonging to arc

ACCEPTED MANUSCRIPT haea, chloroplasts, eukarya, and those unassigned at the bacteria domain level were remov ed from the dataset. In order to study the phylogenetic relationship of different OTUs, and the difference in the dominant species among different samples (groups), multiple sequenc e alignment were conducted using the MUSCLE software (Version 3.8.31, http://www. driv

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e5.com/muscle/) (Edgar, 2004). To correct for unequal sequencing depth, we used 64,002 s

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equences per sample to calculate the diversity and distance among samples.

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Taxonomic richness and diversity estimators were determined for each library in Mothur (Schloss, 2009). The mean of the estimated richness was used for comparisons among samples.

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The taxa relative abundances were generated into phylum, class, order, family, and genus levels

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for each sample. The analyses from clustering to alpha (within sample) and beta diversity (between samples) were performed with QIIME (Version 1.7.0) and displayed with R software

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(Version 2.15.3) (Zhang et al., 2016a, b). Alpha diversity was estimated using five phylogenetic

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diversity metrics: Chao1and ACE indices, Observed OTUs, and Simpson and Shannon indices (Zhang et al., 2016a, b). Rarefaction curves were generated based on observed species. Beta

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diversity between bacterial communities was evaluated using weighted Unifrac distances (Zhang

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et al., 2016a, b). Hierarchical clustering of samples was completed using UPGMA. PCoA on the Unifrac distances of the weighted distance matrices was performed to visualize differences in bacterial community composition and structure. 2.6. Statistical Analysis Data in this study were expressed as mean ± S.E.M.. Proportional data were arcsine square root transformed and analyzed by a one-way ANOVA, followed by a Tukey's multiple comparison

ACCEPTED MANUSCRIPT tests was used to determine significant differences between treatments (SPSS 17.0, IL, USA). The level of significant difference was set at P < 0.05.

3. Results

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3.1. Growth performance and survival

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Effect of dietary lipid levels on growth and survival of swimming crab are presented in Table

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3. The average survival ranged from 63.3 to 83.3%, and swimming crab fed the high-fat diet (HF) (15.1%) had significantly lower survival than those fed the low-fat (5.8%, LF) and medium-fat

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(9.9%, MF) diets. Swimming crab fed the diets containing the LF and MF diets had higher final

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weight and weight gain than those fed the HF diet.

Insert Table 3 here.

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3.2. Illumina sequencing and microbial complexity in crab gut

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After quality control, we obtained a total of 677,734 high quality sequences and 64,507-84,625 sequences per sample. The average length of the sequences was 369 bp. To explore

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the bacterial community diversity among the three groups, we estimated a series of alpha diversity

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indices (Table 4). Sequences with more than 97% similarity were clustered into OTUs. Rarefaction analysis indicated that sufficient sampling depth was achieved for each sample (Fig. 1).

The richness indices of Chao1 and ACE varied from 97.9 to 107.6, and from 103.0 to 112.4, respectively. Bacterial diversity was estimated by the Shannon and Simpson indices, which varied from 1.2 to 1.7, and from 0.4 to 0.6, respectively. Based on the ACE, Chao1, Shannon, and

ACCEPTED MANUSCRIPT Simpson indices, different lipid levels did not influence the intestinal bacterial richness and diversity in swimming crab (Table 4). Insert Table 4 here. Insert Figure 1 here.

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3.3. Microbial community composition

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Proteobacteria, Fusobacteria, and Tenericutes were the dominant phyla (relative

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abundance>1%) in the P. trituberculatus intestinal microbiota (Fig. 2). The relative abundances of Proteobacteria and Fusobacteria were significantly influenced by dietary lipid levels (P<0.05).

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The UPGMA cluster analysis revealed that the three LF diet-associated gut communities were

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more similar to each other than was the case for the MF and HF diets gut microbial communities. Proteobacteria were dominant in the MF (68.7±13.7%) and HF (78.1±15.3%) groups, but they

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were much lower in the LF group (9.4±5.2%). In contrast, the relative abundance of Fusobacteria

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in the LF group (60.9±22.4%) was higher than that in the MF (9.5±7.4%) and HF (1.2±0.5%) groups (P<0.05). The Tenericutes phylum exhibited the same change pattern among all treatments

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(29.3±17.6, 20.0±10.5 and 19.6±15.8 in LF, MF and HF groups, respectively) as did Fusobacteria. Insert Figure 2 here.

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The intestinal bacterial composition which had the best growth conditions and the worst growth conditions in the three treatments was studied further. The main families among the three treatment groups were Vibrionaceae, Leptotrichiaceae, Mycoplasmataceae and unidentified Rickettsiales (proportions>1%; Fig. 3). The relative abundances of Vibrionaceae and Leptotrichiaceae were influenced by lipid levels (P<0.05). Vibrionaceae was dominant in the MF (67.4±13.7%) and HF (41.8±4.4%) groups, but was much lower in the LF group (7.0±3.4%). In contrast, the relative abundance of Leptotrichiaceae in the LF group (60.8±22.5%) was higher than

ACCEPTED MANUSCRIPT that in the MF (9.5±7.4%) and HF (1.1±0.5%) groups (P<0.05). The prevalence of unidentified Rickettsiales increased in the HF group while it decreased in the LF and MF groups, but no significant differences were detected, which may be attributed to the intra-group variation and the limited number of samples. For the remaining family, Mycoplasmataceae (29.3 ± 17.6%,

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20.0±10.5%, 19.6±15.8% in LF, MF and HF groups, respectively) was the most dominant

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Insert Figure 3 here.

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members, followed by Vibrionaceae, Leptotrichiaceae and unidentified Rickettsiales.

3.4. Bacterial phylotype distribution

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A PCoA plot was used to show the microbial community compositions for swimming crab

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fed with different lipid level diets based on the Unifrac distance (Fig. 4). This figure demonstrated that the lipid was important determinants of bacterial community composition. Bacterial

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communities from the different treatment groups were divided into two major clusters (LF group

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and other groups) along PC1, which could describe 72.6% of total variation. For swimming crab intestines, the LF group was distinctly separated from the MF and HF groups. Moreover, HF

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groups also showed somewhat distinct patterns in community composition along PC2, which could describe 14.76% of total variation.

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Insert Figure 4 here.

The heat map was used to show genera whose abundances were changed by dietary lipid level. Top thirty-five genera in the swimming crab were selected for the construction of a heat map (Fig. 5). Four out of thirty-five genera were specifically detected in the intestines of the LF group, and

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were

affiliated

to

Tenericutes

(Candidatus_Bacilloplasma),

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(Propionigenium), Proteobacteria (Acetobacter) and Firmicutes (Defluviitaleaceae_UCG-011). Four out of thirty-five genera were higher in the intestines of the MF group, and they were related

ACCEPTED MANUSCRIPT to Proteobacteria (Photobacterium, Brevundimonas, Stenotrophomonas and Bordetella) and Firmicutes (Fusibacter). Eighteen out of thirty-five genera were detected in the intestines of the HF group, and they were related to Proteobacteria (Labrenzia, Nesiotobacter, Psychrobacter, Pelagibacterium, Maritalea, Candidatus_Thiobios, Ruegeria, Pseudahrensia, Desulfovibrio,

and

Bergeyella),

Verrucomicrobia

(Rubritalea),

Cyanobacteria

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(Draconibacterium

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Desulforhopalus, Candidatus_Hepatincola, Cohaesibacter and Raoultella), Bacteroidetes

observed microbial assembly is at the genus level.

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(Chloroplast_unidentified) and Firmicutes (Staphylococcus). These results indicated that the

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Insert Figure 5 here.

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In order to further study the phylogenetic relationships at the genus level, the phylogenetic relationships of Top100 genera were obtained by multiple sequence alignment. Combined with the

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relative abundance of each genus, we observed the phylogenetic relationship of these genera in the

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three lipid level diets (Fig. 6). The results of the Phylogenetic tree revealed that the MF and HF groups were observed to be concentrated in the same phyla and separated from the LF group. Four

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genera, Photobacterium, Candidatus_Bacilloplasma, Candidatus_Hepatincola, and Vibrio, were detected in all intestinal bacterial communities (proportions>1%). The relative abundances of

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Candidatus_Hepatincola was most in the HF group, but was much lower in the LF and MF groups. Meanwhile, the relative abundances of Photobacterium and Vibrio in the MF and HF groups were higher than those in the LF group. For the remaining one genus, the relative abundance of Candidatus_Bacilloplasma in the LF group was slightly higher than those in the MF and HF groups. Insert Figure 6 here.

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4. Discussion A true lipid requirement for fish or crustacean species cannot be specifically defined because it is influenced by a variety of nutritional factors (NRC, 2011). As a macronutrient, lipids are principal sources of energy and essential fatty acids for the aquatic animal (Watanabe, 1982, NRC,

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2011). In crustaceans, such as crabs or shrimp, weight gain responses to different dietary lipid

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levels, either alone or in combination, indicate that highest weight gain are generally achieved at

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inclusion levels of 5-6% in the diet (NRC, 2011). In the present study, the results indicated that swimming crab in the LF and MF groups showed better growth performance and survival than

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those in the HF group. Huo et al. (2014) found that a diet containing 5% lipid level with a

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protein/energy of 26.9 mg protein kJ-1 is optimal for juvenile swimming crab. However, Han et al. (2013a) reported that swimming crab fed diets containing 4.2 to 13.8% lipid appears to meet the

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lipid requirement. Previous studies demonstrated a negative relationship between growth and

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dietary lipid levels, indicating no distinct dietary lipid level required by crustacean species (Sheen and D'Abramo, 1991; Sheen, 1997; Sheen and Wu, 1999). Moreover, higher lipid levels (>10%) in

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diets are known to retard growth and survival of shrimp, most probably due to a reduction in

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consumption caused by high caloric content or an inability to metabolize high levels efficiently (i.e., reduced digestibility) (Chuntapa et al., 1999, NRC, 2011). Cuzon and Guillaume (1997) also observed that crustaceans are not able to tolerate more than 10% lipid in the diet. Reduced growth has been shown to be associated with accumulation of lipid in tissue (González-Felix et al., 2002). Certain factors such as genetic background, diet, and living environments, can affect intestinal microbiota (Zhang et al., 2014a). Intestinal bacteria are important in nutrient absorption and utilization (Roeselers et al., 2011; Sullam et al., 2012; Daniel et al., 2014; Zhang et al., 2014a;

ACCEPTED MANUSCRIPT Qiao et al., 2016; Xiong et al., 2016). In recent years, some studies in invertebrates have focused on the relationship between intestinal microbiota and nutrient absorption and degradation (Johnson et al., 2008; Zhang et al., 2014a, b; Zhang et al., 2016a, b). In the present study, the intestinal microbiota of swimming crab fed the MF and HF diets were similar to each other, and

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different from those fed the LF diet. The relative abundance of Fusobacteria decreased, while

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Proteobacteria increased in the intestine of the crabs fed the LF diet, suggesting that the abundance

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of these two phyla respond to dietary lipid levels. These data also suggested that the alteration of intestinal microbiota in the HF group may account for the poorer growth performance. Close

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relationship between intestinal bacteria composition and lipid metabolism have been seen before

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(Musso et al., 2011; Semova et al., 2012; Ghosh et al., 2013; Druart et al., 2014), although the role of intestinal bacteria in lipid metabolism remains unknown.

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Illumina high-throughput sequencing data revealed that the dominant phyla in P.

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trituberculatus were within Proteobacteria, followed by Fusobacteria and Tenericutes. Proteobacteria were the most prevalent members in each sample, which is consistent with previous

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studies in other aquatic animals, such as Brachydanio rerio (Roeselers et al., 2011),

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Ctenopharyngodon idellus (Wu et al., 2012), L. vannamei (Zhang et al., 2014a; Xiong et al., 2015; Qiao et al., 2016; Zhang et al., 2016b), N. norvegicus (Meziti et al., 2010; 2012), Oreochromis niloticus (Zhang et al., 2016b), P. monodon (Rungrassamee et al., 2014), and Pelteobagrus fulvidraco (Wu et al., 2010). Also, Fusobacteria have been detected in some aquatic animal species, such as Hypophthalmichthys molitrix and Dorosoma cepedianum (Sullam et al., 2012; Ye et al., 2014), C. idellus (Ni et al., 2014), O. niloticus (Zhang et al., 2016b), and N. norvegicus (Meziti and Kormas, 2013). Although Fusobacteria were detected in P. monodon, the relative

ACCEPTED MANUSCRIPT abundance was about one fifth of that observed in fish gut, and they were almost absent from L. vannamei (Ni et al., 2014; Rungrassamee et al., 2014). Tenericutes have been detected in the gut of L. vannamei and P. monodon, but were barely detectable in that of O. niloticus (Chaiyapechara et al., 2012; Zhang et al., 2016b). The function of these phyla in crab intestines remains unknown

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due to the limited number of studies with gut microbiota in crustaceans. In contrast, Firmicutes

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and Bacteroidetes are the dominant phyla in human and mice intestines (Zhang et al., 2014a).

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However, the relative abundance of Firmicutes and Bacteroidetes in the swimming crab was relatively low. Consistent with previous studies, Firmicutes or Bacteroidetes comprised a small

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proportion of bacterial sequences retrieved from some shrimp intestines (Zhang et al., 2014a;

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Rungrassamee et al., 2013, 2014; Xiong et al., 2017), while Firmicutes were dominant in P. monodon (Rungrassamee et al., 2014). Also, the relative abundance of Bacteroidetes was similar

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to that of Proteobacteria and Tenericutes in the intestines of L. vannamei (Yan et al., 2012). This

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discrepancy between the present work and previous studies may be attributed to differences in the phylogeny, culture condition or diets. Thus, further work is needed to gather more information on

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diets.

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how the intestinal bacterial structure and composition of swimming crab is affected by various

A clear shift in microbial community structure was observed based on the PCoA plot, suggesting that lipid was a critical factor in shaping the gut microbiota of swimming crab. The dominant microbial populations accounted for a significant portion of the total populations and these were substantially altered by different lipid levels. In this study, swimming crab fed the medium- and high-lipid diets had less Leptotrichiaceae than those fed the low-lipid diet, but the role of this family in the crustacean intestine remains unclear. Moreover, higher Vibrionaceae

ACCEPTED MANUSCRIPT were detected in the swimming crab fed the medium- and high-lipid diets than in those fed the low-lipid diet. The Vbrionaceae family is one of the most important bacterial groups in marine environments (Kita-Tsukamoto et al., 1993). Members of the Vibrionaceae are the main organisms present in the intestinal flora of marine fish (Simidu et al., 1977). In addition, some members of

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the Vibrionaceae are important pathogens for humans and animals. Members of the family

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Vibrionaceae associated with the larvae of M. rosenbergii were shown to be pathogenic under

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laboratory conditions (Bhat et al., 1999). Our results suggested that the Vibrionaceae would prefer to live in rich lipid surroundings and this may be a primary reason for the reduced growth

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observed in the high-lipid group. Both Photobacterium and Vibrio belong to the Vibrionaceae

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family. Several studies have reported symbiotic associations between Photobacterium species and marine animals, but studies have also shown that Photobacterium may be an important pathogen

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in the marine ecosystem (Reichelt et al., 1977; Prayitno and Latchford, 1995; Meziti et al., 2010).

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The members of Vibrio are ubiquitous and are associated with many marine or freshwater crustaceans. However, the pathological and physiological effects of Vibrio infections are reported

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in marine crabs where Vibrio infections commonly cause or produce bacteremias and shell disease

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(Wang, 2011). Under conditions of Vibrio-caused bacteremias, a marked reduction in hemocyte numbers and intravascular clotting is observed; the apparent result of an endotoxin in the cell wall of the bacterium (Sritunyalucksana and Söderhäll, 2000). Vibrio-caused bacteremias have been reported for the blue crab, Callinectes sapidus (Davis and Sizemore, 1982; Sizemore and Davis, 1985; Welsh and Sizemore, 1985), Callinectes bocourti, (Rivera et al., 1999), and rock crab, Cancer irroratus (Newman and Feng, 1982). Affected crabs were weak and their haemolymph contained many bacteria when examined by phase microscopy. Vibrio-caused shell disease has

ACCEPTED MANUSCRIPT been reported for Callinectes sapidus (Tubiash and Krantz, 1970). Also, common Vibrio human pathogens, such as V. cholerae and V. vulnificus, have been reported to regularly occur in blue crab haemolymph (Sizemore et al., 1975; Davis and Sizemore, 1982). In the present study, more Rickettsiales were found in swimming crab fed the HF diet than those fed the LF and MF diets.

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Many members of the Rickettsiales are intracellular symbionts (Fredricks, 2006). Orders of

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Rickettsiales are well-known parasites and pathogens of plants, animals and human (Wang et al.,

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2004). The large number of Rickettsiales may explain the low survival rate in swimming crab fed the HF diet. However, the exact role of these families in the crab intestine still needs to be

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revealed. As mentioned before, Proteobacteria, Fusobacteria and Tenericute were the most

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abundant members found in the intestines of swimming crab; moreover, Vibrionaceae and unidentified Rickettsiales adapt to the swimming crab intestinal environment when fed the MF or

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HF diets, demonstrating that the intestinal environment of the host can exert selective pressure on

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the establishment of microbial communities (Rungrassamee et al., 2014; Zhu et al., 2016; Xiong et al., 2017).

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In conclusion, the findings of the present study revealed that swimming crab harbor specific

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gut microbiota and that dietary lipid levels impose selective pressure on these intestinal bacterial communities. Indeed, the composition of intestinal microbiota could be an indicator of the nutritional and growth status of swimming crab.

Acknowledgments This study was supported by China Agriculture Research System-46 (CARS-46), the National Natural Science Foundation of China (41476125), the Nature Science Foundation of

ACCEPTED MANUSCRIPT Zhejiang Province (LY17C190002), Key Research Program of Zhejiang Province of China (2017C02G1460834). This research was also sponsored by the K. C. Wong Magna Fund and the K. C. Wong Education Foundation at Ningbo University. We are grateful to Y. Li, H. Qiu and Y.

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M. Hou for their valuable help during the feeding trial, sampling and chemical analyses.

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ACCEPTED MANUSCRIPT metagenome in Litopenaeus vannamei. Marine Sci. 36, 9-14. Ye, L., Amberg, J., Chapman, D., Gaikowski, M., Liu, W.T., 2014. Fish gut microbiota analysis differentiates physiology and behavior of invasive Asian carp and indigenous American fish. ISME J. 8 (3), 541-551.

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Yoshii, K., Takakuwa, F., Nguyen, H.P., Masumoto, T., Fukada, H., 2010. Effect of dietary lipid

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Zeng, T.L., Ye, Y.F., Mu, C.K., Wang, K., Li, R.H., Wang, C.L., 2016. Gut Microbiota and

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Zhang, M.L., Sun, Y.H., Liu Y.K., Qiao, F., Chen, L.Q., Liu, W.T., Du, Z.Y., Li, E.C., 2016b.

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Response of gut microbiota to salinity change in two euryhaline aquatic animals with reverse

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salinity preference. Aquaculture 454, 72-80. Zhang, W, Xie, F.J, Jin, M, Zhou, Q.C., Li, M., Han, C.M., 2014d. Effects of dietary n-3 highly

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swimming crab (Portunus trituberculatus). Chinese J. Anim. Nutr. 5, 1254-1264. Zhang, M.L., Chekan, J. R., Dodd, D., Hong, P. Y., Radlinski, L., Revindran, V., Nair, S. K., Mackie, R.I., Cann, I., 2014c. Xylan utilization in human gut commensal bacteria is orchestrated by unique modular organization of polysaccharide-degrading enzymes. Proc. Natl. Acad. Sci. U. S. A. 111(35), 3708-17. Zhang, M.L., Sun, Y..H., Chen, K., Yu, N., Zhou, Z.G., Chen, L.Q., Du, Z.Y., Li, E.C., 2014a. Characterization of the intestinal microbiota in Pacific white shrimp, Litopenaeus vannamei,

ACCEPTED MANUSCRIPT fed diets with different lipid sources. Aquaculture 434, 449-455. Zhang, M.L., Sun, Y.H., Chen, L.Q., Cai, C., Qiao, F., Du, Z.Y., Li, E.C., 2016a. Symbiotic bacteria in gills and guts of Chinese mitten crab (Eriocheir sinensis) differ from the free-living bacteria in water. PlOS ONE 11(1), e0148135.

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Zhang, M.L., Liu, N., Qian, C.L., Wang, Q.F., Wang, Q., Long, Y.H., Huang, Y.P., Zhou, Z.H., Yan,

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X., 2014b. Phylogenetic and functional analysis of gut microbiota of a fungus-growing

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higher termite: Bacteroidetes from higher termites are a rich source of B-glucosidase genes. Microb. Ecol. 68, 416-425.

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Zhang, S., Li, J., Wu, X., Zhong, W., Xian, J., Liao, S., Miao, Y., Wang, A., 2013. Effects of

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different dietary lipid level on the growth, survival and immune-relating genes expression in Pacific white shrimp, Litopenaeus vannamei. Fish Shellfish Immunol. 34, 1131-1138.

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Zhao, J., Wen, X., Li, S., Zhu, D., Li, Y., 2015. Effects of dietary lipid levels on growth, feed

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Zhou, X.X., Pan, Y.J., Wang, Y.B., Li, W.F., 2007. In vitro assessment of gastrointestinal viability

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shrimp. Microb. Ecol.72, 975-985.

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Table 1 Formulation and proximate composition of experimental diets. Dietary lipid levels (%) Ingredient (%) 9.9

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White fish meal

25.0

25.0

25.0

Wheat gluten meal

12.0

12.0

12.0

Soybean protein concentrate

19.0

19.0

19.0

Krill meal

5.0

5.0

5.0

Dextrin

19.7

19.7

Fish oil

2.0

Soy lecithin

1.0

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12.0

1.0

1.0

1.0

1.0

1.0

1.5

1.5

1.5

0.3

0.3

0.3

1.5

1.5

1.5

10.0

5.0

0.0

2.0

2.0

2.0

90.8

91.5

91.6

47.3

47.3

47.6

5.8

9.9

15.1

9.5

9.5

9.5

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19.7

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Vitamin premixa

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Mineral premixb Choline chloride

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Ca(H2PO4)2

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Cellulose Sodium alginate

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Proximate composition (dry matter %)

Crude protein

Ash a,b

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Crude lipid

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Dry matter

Same as Jin et al. (2015).

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Table 2 Fatty acid composition of the experimental diets (% of total fatty acids). Dietary lipid level (%) Fatty acid 9.9

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C14:0

4.6

6.1

7.0

C16:0

23.7

24.6

24.1

C18:0

4.7

5.0

C20:0

0.3

0.6

C22:0

0.2

0.2

∑SFA1

33.6

C16:1n-7

5.3

C18:1n-9

17.2

C20:1n-9

4.4

C22:1n-9

0.9

∑MUFA2

27.8

C18:3n-3

1.6

C20:5n-3

7.3

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5.8

0.7 0.2 36.7

6.7

7.5

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36.5

16.0

4.1

3.8

0.9

0.8

28.7

28.1

1.4

1.4

8.2

9.5

10.1

10.7

11.5

19.1

20.3

22.4

16.5

10.2

7.9

0.6

0.9

0.8

17.1

11.1

8.7

DHA/EPA

1.4

1.3

1.2

∑PUFA3

18.1

11.6

9.3

∑HUFA4

18.9

20.7

23.0

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∑n-3PUFA5

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∑n-6PUFA6

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C18:2n-6 C20:4n-6

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C22:6n-3

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∑SFA, saturated fatty acids: C14:0, C16:0, C18:0, C20:0, C22:0.

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∑MUFA, monounsaturated fatty acids: C16:1n-7, C18:1n-9, C20:1n-9.

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∑PUFA, polyunsaturated fatty acids: C18:2n-6, C18:3n-3.

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∑HUFA, highly unsaturated fatty acids: C20:4n-6, C20:5n-3, C22:5n-3, C22:6n-3.

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∑n-3PUFA: C18:3n-3, C20:5n-3, C22:6n-3.

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∑n-6PUFA: C18:2n-6, C20:4n-6.

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Dietary lipid level (%) 9.9

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Initial weight (g)

19.9±1.2

21.4±1.1

22.0±0.3

Final weight (g)

83.9±5.1a

84.1±6.5 a

76.6±4.7 b

Weight gain (%)

322.9±10.6a

292.2±14.7a

248.0±21.7b

80.0±4.1a

83.3±4.9a

63.3±1.4b

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Survival (%)

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Data represent means ± S.E.M. from three repetitions. The values in the same row with different

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superscripts are different (P<0.05).

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Table 4 Illumina high-throughput data, bacterial diversity richness (OTUs), diversity index (Shannon & Simpson), and estimated OTU richness (Chao & ACE) for intestinal bacterial diversity analysis of P. trituberculatus fed with different dietary lipid levels for 8 weeks. Dietary lipid levels (%)

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Parameters

5.8

9.9

Mean sequences

74,189

76,025

Observed OTUs

78.0±13.8

88.3±15.3

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Richness estimators

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Sampling depth

15.1

75,697 100.0±19.3

97.9±12.0

101.2±14.0

107.6±17.8

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103.0±10.8

103.6±14.1

112.4±17.3

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Chao1

Diversity estimators

1.2±0.3

1.7±0.5

1.7±0.0

Simpson

0.4±0.1

0.6±0.2

0.6±0.0

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Data represent means ± S.E.M from three repetitions. The values in the same row with different

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superscripts are different (P<0.05).

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Fig. 1. Rarefaction analysis of the intestine bacterial community of P. trituberculatus fed with different dietary lipid levels for 8 weeks. Operational taxonomic units (OTUs) were clustered

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based on 97% sequence similarity. The number of sequences sampled indicates the number of

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sequencing reads. Each point made 10 iterations for each sample. LF: low-fat; MF: medium-fat;

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HF: high-fat.

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Dietary lipid levels (%) Phylum (%)

5.8

9.9

15.1

9.4±5.2b

68.7±13.7a

78.1±15.3a

60.9±22.4a

9.5±7.4b

1.2±0.5b

29.3±17.6

20.0±10.5

19.6±15.8

0.07±0.03

0.15±0.05

0.18±0.11

0.01±0.00

0.03±0.01

0.06±0.06

0.01±0.00

0.01±0.01

0.05±0.04

Firmicutes

0.03±0.01

0.06±0.01

0.06±0.02

Other bacteria

0.33±0.12

1.51±0.99

0.76±0.45

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Proteobacteria

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Fusobacteria Tenericutes

Chlorobi

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Bacteroidetes

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Verrucomicrobia

Fig. 2. Cluster analysis of the intestinal bacterial phyla of P. trituberculatus fed different dietary lipid levels for 8 weeks. The phyla with relative abundance higher than 1% in at least one individual are shown in the schematic, and the relative abundance of main phyla are indicated in the table section. LF: low-fat; MF: medium-fat; HF: high-fat.

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Dietary lipid levels (%)

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Family (%)

5.8

9.9

15.1

7.0±3.4b

67.4±13.7a

41.8±4.4a

60.8±22.5a

9.5±7.4b

1.1±0.5b

29.3±17.6

20.0±10.5

19.6±15.8

2.1±2.0

0.8±0.3

35.4±17.9

0.13±0.05

0.18±0.07

0.4±0.16

Gammaproteobacteria_unidentified

0.04±0.02

0.2±0.1

0.16±0.09

Fusobacteriaceae

0.11±0.1

0.01±0

0.07±0.03

0.02±0.01

0.05±0.02

0.09±0.05

0.01±0

0.03±0.01

0.06±0.05

Cytophagaceae

0.03±0.02

0.08±0.03

0.08±0.05

others

0.48±0.17

1.77±1.05

1.23±0.6

Vibrionaceae

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Mycoplasmataceae

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Leptotrichiaceae

Rickettsiales_unidentified

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Rhodobacteraceae

Flavobacteriaceae IheB3-7

Fig. 3. Relative abundances of the main bacterial families in the low lipid, middle lipid and high lipid groups. The families with abundance higher than 1% in at least one individual are shown in the schematic. LF: low-fat; MF: medium-fat; HF: high-fat.

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Fig. 4. Principal co-ordinate analysis (PCoA), based on weighted-Unifrac distance, of the

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intestinal bacterial communities of P. trituberculatus fed low-fat (LF; red), medium-fat (MF; green)

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or high-fat (HL; blue). LF: low-fat; MF: medium-fat; HF: high-fat.

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Fig. 5. A heatmap showing the relative abundances of the bacterial genera in the intestine of P.

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trituberculatus fed different dietary lipid levels for 8 weeks. At the genus level, we focused on variations in the top 35 genera among the three groups.The relative values of the genus are depicted by color intensity. LF: low-fat; MF: medium-fat; HF: high-fat.

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Fig. 6. Phylogenetic tree of bacterial genera in the intestine of P. trituberculatus fed three different

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dietary lipid levels for 8 weeks. The inner circle is a phylogenetic tree constructed from a representative sequence (genus level). The color of the branch represents its corresponding

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phylum, while each color represents a phylum. The outer circle is the relative abundance of each genus in each treatment group. The color of the relative abundance represents the different treatment groups. LF: low-fat; MF: medium-fat; HF: high-fat.

ACCEPTED MANUSCRIPT Highlights

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 

Dietary lipid levels being not more than 9.91% could improve growth of swimming crab. Dietary lipid levels influenced the intestinal bacterial composition. This study characterized the intestinal microbiota in swimming crab.

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