Meziti etal12

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Authors requiring further information regarding Elsevier’s archiving and manuscript policies are encouraged to visit: http://www.elsevier.com/copyright Author's personal copy Systematic and Applied Microbiology 35 (2012) 473–482 Contents lists available at SciVerse ScienceDirect Systematic and Applied Microbiology journal homepage: www.elsevier.de/syapm Gut bacteria associated with different diets in reared Nephrops norvegicus Alexandra Meziti, Eleni Mente, Konstantinos Ar. Kormas ∗ Department of Ichthyology & Aquatic Environment, School of Agricultural Sciences, University of Thessaly, 384 46 Volos, Greece a r t i c l e i n f o Article history: Received 7 June 2012 Received in revised form 26 July 2012 Accepted 27 July 2012 Keywords: Bacteria Gut Diet Nephrops norvegicus Aquaculture a b s t r a c t The impact of different diets on the gut microbiota of reared Nephrops norvegicus was investigated based on bacterial 16S rRNA gene diversity. Specimens were collected from Pagasitikos Gulf (Greece) and kept in experimental rearing tanks, under in situ conditions, for 6 months. Treatments included three diets: frozen natural (mussel) food (M), dry formulated pellet (P) and starvation (S). Gut samples were collected at the initiation of the experiment, and after 3 and 6 months. Tank water and diet samples were also analyzed for bacterial 16S rRNA gene diversity. Statistical analysis separated the two groups fed or starved (M and P vs. S samples). Most gut bacteria were not related to the water or diet bacteria, while bacterial diversity was higher in the starvation samples. M and P samples were dominated by Gammaproteobacteria, Epsilonproteobacteria and Tenericutes. Phylotypes clustering in Photobacterium leiognathi, Shewanella sp. and Entomoplasmatales had high frequencies in the M and P samples but low sequence frequencies in S samples. The study showed that feeding resulted in the selection of specific species, which also occurs in the natural population, and might be associated with the animal’s nutrition. © 2012 Elsevier GmbH. All rights reserved. Introduction The Norway lobster Nephrops norvegicus is a decapod crustacean living at depths of 20–800 m in the Mediterranean, the North Sea and the North East Atlantic Ocean. It is a commercially important species [4], but overexploitation and inappropriate management strategies have led to possible depletion of the existing stocks. Several studies of N. norvegicus have been performed in the past on their biology [4], population abundance and structure [1], molting and growth [17], feeding ecology and behavior [9] and reproductive biology [35,46,47]. However, the lack of knowledge concerning their nutritional requirements and rearing under laboratory conditions [45] is the constraint limiting the successful culture of the Norway lobster, either for restocking or for commercial size production. Recent studies [34] have provided valuable data on the survival, growth and feeding behavior of N. norvegicus kept in controlled laboratory conditions, and they have shown, among others, the need for high-quality dry food. N. norvegicus mainly feeds on fish, mollusks, crustaceans, polychaetes, echinoderms and foraminifers [9]. However, the synthetic feed provided to N. norvegicus in past studies [34] was mostly pellets used in fish aquaculture that consisted mainly of fishmeal and soy. The use of probiotics, although very important in the aquaculture of crustaceans [15], has never been attempted in the rearing of N. norvegicus. The probiotics used in crustacean aquaculture have ∗ Corresponding author. Tel.: +30 242 109 3082; fax: +30 242 109 3157. E-mail address: kkormas@uth.gr (K.Ar. Kormas). 0723-2020/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.syapm.2012.07.004 been shown to increase the growth and survival of the species ([15] and references therein) without the use of antibiotics. The probiotics used for each species are determined by several factors, such as the non-pathogenicity and non-toxicity of the strain used, as well as its ability to survive in and adhere to the gut [15]. Additionally, probiotics should benefit their host in certain ways, such as promoting growth or protecting against pathogens [2,25]. Recent studies have shown that the gut microbial communities of several animals are influenced by the nutritional habits of their hosts [29] and, at the same time, they metabolize part of the ingested food and provide the host with important nutrients (e.g. cellulose digestion). Resident gut microbes that are able to metabolize complex compounds are very good candidates as probiotics since they fulfill all the above-mentioned criteria. In the case of the gut microorganisms of N. norvegicus, the only study that has been performed in wild specimens showed that a seasonal variation of mid-gut bacterial communities was mostly related to differences in food supply from the overlying water column [36]. During the last 20 years, N. norvegicus catches per unit of effort have steadily declined within the Mediterranean Sea and, thus, support the need for measures to conserve this crustacean species. Aquaculture has been a very useful tool for restocking and stock enhancement programs for a number of fish and shellfish species. Although commercial cultivation of Norway lobsters might be hindered by the slow growth rate exhibited by juveniles of this species in nature [38] and in captivity [10], there is a lot of interest in the intensive cultivation of these animals due to their high nutritional and commercial value. Although there are studies on husbandry and rearing conditions for the Norway lobster [34,45], knowledge Author's personal copy 474 A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 Table 1 Codes of samples collected and analyzed. Samples from the 2nd batch (April 2009) are indicated in boldface. Samples t0 (start) Natural population Starvation Mussels Pellets Nat1/Nat2 t1 (3 months) t2 (6 months) S3m1 M3m1 S6m1/S6m2 M6m1/M6m2 P6m1/P6m2 on the nutritional physiology of the species remains limited. Therefore, the present study contributes to our knowledge of the possible role of its gut microbiota in nutrition. Such information can provide the basis for diet formulation in commercial scale culturing of the Norway lobster. In particular, this study aimed at investigating the factors that affected the dominant gut bacterial communities of N. norvegicus grown under stable laboratory conditions. For this purpose, N. norvegicus individuals were reared in three different groups where mussels (natural feed) and pellets were provided, respectively, while the third group was starved. Mid-gut samples were collected at the beginning of the experiment and 3 and 6 months afterwards, and they were investigated by 16S rRNA gene diversity analysis. Materials and methods Collection of N. norvegicus individuals Individuals were collected from Pagasitikos Gulf (Greece) in March (1st batch) and April 2009 (2nd batch). Sampling depth varied from 60 to 88 m and only male individuals were kept for further analysis, since it has been shown from previous studies that males have better survival rates than females under rearing conditions [41], and that sex is not a significant factor for gut bacterial diversity [36]. After collection, N. norvegicus individuals were immediately transferred to the laboratory in aerated seawater, and animal weight and carapace length were measured. One individual from each batch (Nat1–Nat2) was immediately sacrificed, while the rest were kept for rearing (Table 1). when sacrificed. During the rearing period, molting had been observed only in one individual from group P (P6m2) 2 days before the animal was sacrificed (post-molt stage). Water samples were collected from the tanks 2–3 days before the sampling of the animals. Water samples were collected in sterile 1 L bottles and 1 L, 500 mL and 800 mL were filtered from samples wt2 (t1), wt3 (t2, 1st batch) and wt4 (t2, 2nd batch), respectively. Filtering was performed under vacuum using 0.2 ␮m filters (GTTP, Millipore, USA) and filters were kept at −20 ◦ C until further processing. Samples from mussels and pellets from the batches provided for feeding were also kept. Mid-gut isolation The animals were dissected using sterile lancets and the intestine was extracted using sterile forceps, as described in Meziti et al. [36]. Since the bacterial communities established on or in the gut tissue were required, the intestine was emptied by applying mechanical force and by rinsing three times in autoclaved particlefree seawater in order to remove all gut content. The posterior part of the mid-gut was used for further analysis. All dissecting tools were alcohol-flame sterilized between each individual sample. DNA extraction DNA extraction was performed on 10 N. norvegicus gut tissues using the QIAamp DNA Mini Kit (Qiagen Inc., USA) and following the manufacturer’s standard protocol. At the final step, DNA was diluted in 100 ␮L of the elution buffer provided with the kit and it was stored at −20 ◦ C. DNA extraction from the water samples was performed using the UltraClean Soil DNA Kit (MoBio Laboratories Inc. USA), following the manufacturer’s protocol. DNA was finally eluted in the 50 ␮L elution buffer provided by the manufacturer. DNA was extracted from pooled foot and mantle tissues of three mussels from the batch used for feeding (frozen Mytilus edulis) following the same procedure as for the mid-gut samples. For the pellets, DNA was extracted from 10 pooled pellets using the UltraClean Soil DNA kit (MoBio Laboratories Inc.), following the manufacturer’s standard protocol. Rearing procedure Cloning and sequencing of 16S rRNA genes Individuals for rearing were placed in 100 L glass tanks in the laboratory. Water from Pagasitikos Gulf was transported in the tanks before N. norvegicus transport. In order to establish the bacterial nitrifying communities on the filters of the tanks, bacteria from commercial solutions (Stability, Seachem Inc., USA) were added to the tanks. Water was constantly recycled through carbon and biological filters. Water temperature, salinity and photoperiod were maintained at 11.9 ± 0.8 ◦ C, 374 ± 0.2 ppm and 24 h darkness, respectively, reflecting in situ conditions. After a 15-day acclimatization period, the animals were divided into three groups (M, P, S), and each animal was placed in a separate compartment in tanks made from Plexiglas and plastic netting. Groups M and P were supplied with 1–4 g frozen mussels (natural feed) and approximately 1 g fish pellets (synthetic feed), respectively, three times per week, while group S was starved. The diet’s chemical composition was 69% protein, 7.5% lipid and 23.5% carbohydrate for dry mussels and 42% protein, 11.1% lipid and 46.9% carbohydrate for pellets (Rotllant, unpublished data). Collection of samples Three and 6 months after the initiation of the experiment, animals from each group (Table 1) were sacrificed and their morphometric characteristics were measured. All animals were healthy Part of the bacterial 16S rRNA gene was amplified from all mid-gut, water, mussel and pellet samples using the primers 27f BAC (5′ -AGAGTTTGATCMTGGCTCAG-3′ ) [27] and 907R (5′ CCGTCAATTCCTTTRAGTTT-3′ ) [37]. PCR conditions were 5 min at 94 ◦ C followed by 22–27 cycles of 1 min at 94 ◦ C, 1 min at 52.5 ◦ C and 1 min at 72 ◦ C and a final step of 7 min at 72 ◦ C. The number of PCR cycles was adjusted when needed in order to decrease non-specific products. The total number of cycles for all samples varied from 23 (sample S6m1) to 27 cycles (sample P6m1). PCR products were purified with the Montage Purification Kit (Millipore, USA) and were cloned directly using the TOPO TA Kit for sequencing (Invitrogen Inc., USA) with electrocompetent cells. The insert size was checked using PCR with M13f–M13R vectorbinding primers. Positive clones were grown overnight in 1.5 mL of Luria–Bertani medium containing kanamycin (50 ␮g mL−1 ), and plasmids were purified from the pelleted cells using the Nucleospin Plasmid QuickPure Kit (Macherey-Nagel GmbH and Co., KG, Germany). Plasmids were partially sequenced with primer M13f (5′ -GTAAAACGACGGCCAG-3′ ). After alignment with ClustalW [28], manual correction, elimination of chimeras using the Pintail software [3] and visual examination of the alignments, clones were grouped based on a 16S rRNA similarity cut-off level of 98% and representatives from each group were sequenced using primer M13R Author's personal copy A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 475 Table 2 Dominant phylotypes (>15%) in the studied samples. Phylotype Frequency (%) Closest relative (GenBank No) (% similarity) Phylogeny Publication Nat1-12 Nat2-8 S3m1-3 S3m1-6 S3m1-7 M3m1-15 M3m1-41 M6m1-4 M6m1-56 M6m2-6 M6m2-3 P6m1-12 P6m1-11 P6m2-2 66.6 48.0 15.0 15.0 15.0 15.8 15.8 20.5 17.9 28.9 15.8 33.3 20.8 73.5 Uncultured bacterium clone My46-424 (GQ866072) (99) Uncultured clone Ag31-3 (GQ866101) (99) Marine sponge bacterium plate OTU18 (EU346505) (99) Marinicella litoralis (AB500095) (96) Litoreibacter albidus (AB518881) (99) Uncultured bacterium clone C2E (DQ856531) (96) Epsilonproteobacterium Oy-M7 clone 465.4 (DQ357825) (96) Uncultured clone Ag31-3 (GQ866101) (99) Uncultured bacterium clone TIGU1075 (HM558927) (99) Uncultured clone Ag31-3 (GQ866101) (99) Uncultured Mycoplasmataceae clone Lo Hep1.15 (EU646198) (88) Uncultured bacterium clone D1-674 (GQ866083) (99) Uncultured clone Ag31-3 (GQ866101) (99) Uncultured clone Ag31-3 (GQ866101) (99) Gammaproteobacteria Gammaproteobacteria Alphaproteobacteria Gammaproteobacteria Alphaproteobacteria Alphaproteobacteria Epsilonproteobacteria Gammaproteobacteria Gammaproteobacteria Gammaproteobacteria Tenericutes Gammaproteobacteria Gammaproteobacteria Gammaproteobacteria [36] [36] [52] [42] [43] [30] [49] [36] [54] [36] [16] [36] [36] [36] (5′ -CAGGAAACAGCTATGAC-3′ ). Sequence data were obtained by capillary electrophoresis (Macrogen Inc., Korea) using the Big Dye Terminator Kit (Applied Biosystems Inc., USA). Sequences were checked for closest relatives using the BLAST application and all sequences were checked for chimeras using Pintail. 16S rRNA sequences were aligned using the ARB software [31] and the SILVA aligner application [40]. 16S rRNA distance matrices were calculated with the Jukes–Cantor formula and they were clustered with the neighbor-joining method. Bootstrap values were obtained from 1000 replicates using similar parameters. All 16S rRNA sequences from this study were deposited in GenBank under numbers JN092133–JN092292 (mid-gut samples), JN639288–JN639332 (water samples) and JN858926–JN858954 (mussel and pellet samples). NMDS analysis Unconstrained ordinations, based on the frequencies of the phylotypes, were performed in order to illustrate the relationships between gut and water samples graphically using three-dimensional non-metric multidimensional scaling (NMDS) [26], implemented in R (version 2.9.1). NMDS ordination attempts to place all samples in a three-dimensional space such that their ordering relationships (here based on a Bray–Curtis similarity matrix) can be preserved. Hence, the closer the samples are in the resulting ordination, the more similar the bacterial communities are. Kruskal’s stress value reflects the difficulty involved in fitting the relationships of the samples into a threedimensional ordination space. The hypothesis that gut microbial communities differed depending on whether food was provided or not was tested with the use of the non-parametric analysis of similarities (ANOSIM) [6]. ANOSIM generates a test statistic, R, that ranges from −1 to 1. The magnitude of R is indicative of the degree of separation between groups, with a score of 1 indicating complete separation and 0 indicating no separation [5]. Diversity and similarity analysis The indices of Shannon–Wiener (H) [50], Simpson (D) [51] and Margalef [33] were used for diversity estimates and were calculated using the PAST software. Morisita similarity indices on the phylotypes of the samples were calculated with the SPADE software (http://chao.stat.nthu.edu.tw/softwareCE.html). Cluster analysis was applied to Morisita similarity indices using the PAST program [19]. ANOSIM between the phylotype frequencies of the potential groups was also performed using the PAST program. Results Phylogenetic analysis A total of 520 partial 16S rRNA sequences were analyzed for samples Nat1 (30), Nat2 (25), S6m1 (39), S6m2 (43), M6m1 (39), M6m2 (38), P6m1 (24), P6m2 (34), M3m1 (38) and S3m1 (40), wt2 (34), wt3 (44), wt4 (34), Mus (30) and Pl (28). Each clone library had 6–30 different phylotypes based on a 98% cutoff similarity. Good’s coverage was calculated using the formula C = 1 − (ni /N), where ni is the number of singleton phylotypes and N is the total number of clones analyzed. It ranged from 52% (S6m2) to 91% (P6m2) (Table S1), showing that at least 50% of the total mid-gut bacterial species richness was revealed in all samples. The slope of the collector’s curves, plotting the number of total clones sampled against the number of different phylotypes showed that sampling was incomplete and rare species probably remained undetected (Fig. S1). However, almost all libraries from the rearing samples (apart from the starved ones) had one to two dominant phylotypes (sequence frequencies >15%) implying that the dominant phylotypes were detected (Table 2). Mid-gut samples In five samples (Nat2, M6m1, M6m2, P6m1 and P6m2) the dominant phylotypes (Table 2) were 98–99% similar and were closely related (98–99%) to Photobacterium leiognathi strain RM1 (AY292947) [39] and to phylotypes Jl1-1 (GQ866087), O21 (GQ866108) and Ag31-3 (GQ866101), previously detected in the gut of the Pagasitikos Gulf Norway lobster population [36] (Figs. 1 and 2). In sample P6m1, the dominant phylotype (33.3%) was 99% similar to Shewanella sp. E5050-7 (FJ169983), a protease producing bacterium from the South China Sea [59], and to phylotype D1-674 (GQ866083) detected in the gut of wild N. norvegicus [36]. Closely related (98%) phylotypes were also found in lower frequencies (Figs. 1 and 2) in samples M6m2, S6m1 and S6m2. Phylotypes showing high frequencies in the 6-month musselfed samples (M6m1-2, 12.8% and M6m2-3, 15.7%) clustered within the order of Entomoplasmatales (Fig. 3) but they were distantly related (91%) to all other members of the order. Their closest relatives were the uncultured Candidatus Hepatoplasma clones TyHep1.19 and Lo-Hep 11.5 detected in the hepatopancreas of the isopods Tylos europaeus and Ligia oceanica, respectively [16]. Phylotypes clustering in the same group were also detected in other samples (S6m2 and Nat2) but in lower frequencies (Figs. 2 and 3). Author's personal copy 476 A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 Fig. 1. Neighbor-joining tree based on gammaproteobacterial 16S rRNA gene sequences from wild and reared Nephrops norvegicus. Arcobacter nitrofigilis was set as an outgroup. The bar corresponds to a 10% nucleotide difference and bootstrap values were calculated from 1000 replicate trees. Phylotypes from previous N. norvegicus studies are designated with *. Frequencies of retrieved phylotypes in each clone library are shown in parentheses. Similarly, the dominant phylotype in the 3-month mussel-fed sample (M3m1-41, 15.8%), was closely related (99%) to phylotypes from the rest of the mussel-fed samples and from sample P6m1 (Figs. 2 and 3). These phylotypes were affiliated to the uncultured bacterium Oy-M7 clone 465.4 (DQ357825) from oyster hatcheries [49], clustering within the genus Arcobacter according to the SILVA database 108 [40]. The other dominant phylotype from the 3month mussel-fed sample (M3m1-15, 15.8%) was affiliated (96%) to the alphaproteobacterial clone C2E (DQ856531) detected in the intestine of the Chinese mitten crab Eriocheir sinensis [30], which clusters in the genus Defluviicoccus according to the SILVA database. The rest of the phylotypes clustered within the Alpha-, Betaand Gammaproteobacteria, in the phyla Bacteroidetes, Fibrobacteres, Firmicutes and Actinobacteria, and in the candidate divisions of OD1, OP11 and TM6 (Table S2). Water samples The dominant phylotype (23.5%) in sample wt2 was classified as Flavobacteria (Fig. 4). Its closest relative was strain Kordia algicida OT-1 isolated from the marine environment and it was able to decompose the diatom Skeletonema costatum, which is responsible for the formation of red tides [53]. In sample wt3, the dominant phylotype (47.7%) was classified in Rhodobacteraceae (Fig. 3). Its closest relative was the marine bacterium strain ATAM407 56 (AF359535) belonging to the genus Phaeobacter, that has been isolated from cultures of the slightly toxic dinoflagellate Alexandrium affine NEPCC 607 [22]. Similar phylotypes (>98%) were detected in samples S6m1 and M6m1 (Fig. 2). The dominant phylotype (47%) of sample wt4 was grouped in the family Rhodobacteraceae of Alphaproteobacteria. Its closest relative was Author's personal copy A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 477 Fig. 2. Dominant phylotypes in the mid-gut of Nephrops norvegicus-reared populations. Different colors correspond to different groups of phylotypes, named by the genus of their closest relative. Grouping is based on a cut-off similarity of 97%, except for the relatives of Candidatus Hepatoplasma (<97%). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) strain Marivita cryptomonadis CL-SK44 (EU512919) that has been isolated from the marine species Cryptophyta sp. CR-MAL01 [24]. A similar phylotype was detected in sample S6m1. Generally, several phylotypes from the water samples were closely related to phylotypes detected in the gut samples and mostly in the ones from the starvation group (Fig. 4). Mussel and pellet samples The phylotypes detected in the mussels clustered within the Alpha-, Gamma-, Epsilonproteobacteria, Bacteroidetes and Fibrobacteres, while the ones detected in the pellets clustered mostly within the Firmicutes (data not shown). No common phylotypes were detected between the samples of the feed provided and the mid-gut samples. NMDS analysis The three-dimensional NMDS analysis performed on all water and mid-gut samples revealed the grouping of all mid-gut samples that had been fed with mussels or pellets for 6 months with one sample from the natural populations (Nat2), while samples from starved animals and from the animal that had been fed only for 3 months (M3m1) were grouped with the water samples (Fig. 5). This grouping was statistically significant, as shown by the ANOSIM analysis (R = 0.613, p = 0.004). Diversity and similarity of bacterial communities The three diversity indices gave similar results regarding the bacterial diversity of the gut samples (Table S3). They all showed their lowest values in sample P6m2 and the highest Author's personal copy 478 A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 Fig. 3. Neighbor-joining tree based on 16S rRNA gene sequences from wild and reared Nephrops norvegicus. Aquifex pyrophilus was set as an outgroup. The bar corresponds to a 10% nucleotide difference and bootstrap values were calculated from 1000 replicate trees. Frequencies of retrieved phylotypes in each clone library are shown in parentheses. values in sample S6m2. Overall, the indices indicated higher diversity in the starvation and the M3m1 samples. Neighborjoining analysis of the samples based on the frequencies of the phylotypes showed similar results to the Morisita similarities cluster analysis and NMDS analysis (Fig. S2). Neighbor-joining analysis exhibited a differentiation between Group I (G1: M6m1, M6m2, P6m1, P6m2 and Nat2) and Group II (G2: M3m1, S3m1, S6m1 and S6m2) with Nat1 as an outgroup. The Morisita similarities cluster analysis exhibited similarities of >0.7 between the members of G1, while the members of G2 were highly differentiated with similarities of <0.4 (Fig. S2). This grouping also proved to be statistically significant after ANOSIM analysis (R = 0.665, p < 0.001). Discussion This study analyzed the differences of the gut bacterial communities in experimentally reared N. norvegicus individuals when different food sources were provided. Statistical analysis of the gut bacterial diversity showed the presence of two groups depending primarily on whether food was provided or not. The G1 samples (Nat2, M6m1, M6m2, P6m1 and P6m2) had lower bacterial diversity than the G2 samples (S3m1, S6m1, S6m2 and M3m) (Table S3). All starvation samples were grouped together with M3m1 while some of the phylotypes detected in G2 were similar to phylotypes detected in the water of the tanks (Fig. 4). After 3 months feeding, the bacterial diversity in mussel-fed animals was higher than at the beginning of the experiment, although specific bacterial communities had still not been established and resembled the starvation samples more. Bacterial diversity was lower in the members of G1 and this difference was attributed to food provision that helped the establishment of more stable gut bacterial communities. The high gut bacterial diversity in the members of G2 could be attributed either to starvation or to the short time (3 months) between the initiation of the experiment and the first sampling. An increase in gut bacterial diversity in starved animals has also been observed in the locust Schistocerca gregaria [12]. However, the exact reasons for this increase in bacterial diversity have not been studied in either of the studies. Apart from samples Nat1, Nat2 and P6m2, frequencies of specific bacterial phylotypes never exceeded 33.3% in reared populations (P6m1-12). This is different to previous results [36] where gut bacterial communities in N. norvegicus natural populations were dominated (≥58%) by a single bacterial phylotype. Thus, the gut bacterial communities of reared samples after 3 and 6 months showed higher diversity than the communities of the wild ones (Table S3). In the case of the Norway lobster, dominant bacterial diversities are considered to indicate the performance of specific digestive functions that assist the host. The decrease of the bacterial diversity in reared animals could be attributed to the slow establishment of dominant bacterial communities resulting from the low food consumption (0.049–0.069 mg/gdry body weight /day) of mussels and pellets (Mente and Karapanagiotidis, unpublished data) compared to previous studies (0.025 mg/gbody weight /day) [47]. The phylotypes related to P. leiognathi were practically identical to those previously detected in the gut of wild Norway lobsters [36]. They were present in all the gut bacterial communities of G1 samples where mussels and pellets were consumed. Microbiological studies have proved that most P. leiognathi strains Author's personal copy A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 479 Fig. 4. Neighbor-joining tree based on 16S rRNA gene sequences from the water of the rearing tanks. Aquifex pyrophilus was set as an outgroup. The bar corresponds to a 10% nucleotide difference and bootstrap values were calculated from 1000 replicate trees. Frequencies of retrieved phylotypes in each clone library are shown in parentheses. are chitinolytic (96%) and lipolytic (82%) [14]. Recently, the first completed genomic study of the species (NZ BACE00000000), showed the presence of multiple genes coding for lipases, proteases and chitinases (Microbial Genome Resources, 2011; http://www.ncbi.nlm.nih.gov/genomes/MICROBES/microbial taxt ree.html). Although no lipase, chitinase or protease tests were performed in this study, data showing the high similarities between phylotypes, the dominance of the phylotypes in all G1 and wild samples (from this and from previous studies), and the results from previous microbiologic and genomic studies on the species indicated the presence of a specific bacterial community with potential positive effects on N. norvegicus digestive function. Author's personal copy 480 A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 Similar P. leiognathi clustering phylotypes were dominant (73%) in the P6m2 sample that had molted 2–3 days before sampling. During molting, the chitinase lining of the hind-gut cuticle is shed together with the rest of the exoskeleton [7] and, as a result, the bacterial abundance of the hind-gut decreases [8]. However, both the role of gut bacteria in the molting procedure and their fate after molting is as yet unknown. Studies have shown that chitinase activity in the gut is increased during the post-molt period (the first 2–3 days after molting) [18], since this has been related to the release of the molt [32,56]. In this case, the inferred chitinolytic activity of P. leiognathi-related phylotypes could explain their dominance in the post-molt sample P6m2. Arcobacter-related phylotypes were frequent and recurring in the gut samples and mostly in the ones fed with mussels (Figs. 2 and 3). Their closest relative was isolated from oyster mantle [49] clustering in the genus Arcobacter. The genus Arcobacter oxidizes hydrogen sulfide and produces sulfur [55], while some of the strains of this species are capable of denitrification [21]. Arcobacterlike bacteria have been found in the intestinal tract of humans and animals [55], the deep sea [23], lake water columns [48], activated sludge [21] and marine oysters [44,51]. In this study, the presence of Arcobacter-like phylotypes was mostly associated with the protein-rich mussels that were used for feeding, and they might be connected to nitrogen metabolism and the sulfur cycle. Similarly, high rates of denitrification have been detected in previous studies [20] in the gut of the aquacultured shrimp Litopenaeus vannamei and were mainly attributed to the activity of denitrifying bacteria. The Shewanella-like phylotypes (Fig. 1 and Table 2) were closely related to a protease producing strain. Apart from the potential proteolytic function in the gut of the Norway lobster these phylotypes were similar to the one previously detected (D1-674) in the gut of N. norvegicus [36] and, thus, may belong to the resident bacterial community of the gut. Their presence is reinforced by their ability to grow solely on leucine, which was abundant on the pellets provided as synthetic feed in this experiment [34]. Entomoplasmatales phylotypes were dominant in the 6-month mussel-fed samples (M6m) and appeared with lower frequencies in other samples (Figs. 2 and 3). Phylotypes from the same cluster have appeared in the intestine [11,12,13,58] and the hepatopancreas [16] of other invertebrates with functions that are still unknown. In the case of the uncultured Candidatus Hepatoplasma clones, which were detected as closest relatives in our study, they had been related to higher survival rates of their hosts when low quality food was provided [16]. The phylotypes detected in the gut of N. norvegicus were distantly related to the Candidatus Hepatoplasma clones (<93%) but clearly clustered in different genera, as has been described recently from Yarza et al. [57] when setting genus boundaries at 94.5% SSU similarities. Thus, a similar relationship between higher survival rates and food quality cannot be assumed. Regarding the phylotypes detected in the starvation samples, there was no pattern that was present in all of them, according to sampling time or to specific bacterial communities. The starvation phylotypes mostly clustered in the Alphaproteobacteria and the Bacteroidetes (Figs. 3 and 4, Table S2) and some of them were closely related to phylotypes detected in the water samples (Fig. 4), suggesting an influence of the gut bacterial communities from the surrounding environment. The reverse hypothesis has been excluded since all the dominant common phylotypes were related to bacteria detected in the marine environment that were not associated with the digestive tract. Apart from that, some rare S phylotypes (Figs. 1 and 2) were closely related to the dominant Photobacterium-like and Shewanella-like phylotypes detected in the M6m and P6m samples. However, their occurrence in S samples was low (<7%) and occasional. Although no clear differences between mussel- and pellet-fed animals were observed, the presence of food seemed to fuel the Fig. 5. NMDS ordination plot (Bray–Curtis distance matrix) of the phylotype frequencies from the Nephrops norvegicus samples (ordination stress = 0.04). Each gut sample is indicated by a dot with different colors (light red: wild populations t0; dark-red: pellet-fed samples; orange: mussel-fed samples; blue: starvation samples; cyan: tank water samples). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) establishment of dominant microbial communities. Bacterial communities seemed to be agitated after transport in the rearing tanks, since starvation samples and the 3-month mussel-fed sample were more diverse compared to that known concerning gut bacterial diversity of wild Nephrops from past studies [36] and from this study (Table S3). The dominance of Photobacterium sp., Shewanella sp. and Mycoplasmataceae clustering phylotypes in G1 samples (6 months) seemed to be a result of feeding, since these phylotypes, although potentially resident (as assumed from their concurrence in wild samples), showed a low and random presence in G2 samples and formed more stable communities after 6 months feeding. From our findings, combined with previous studies [36], it seems that among all the dominant phylotypes detected in this study, P. leiognathi fulfills most of the known criteria [15] for the selection of a probiotic microorganism. It is non-pathogenic for N. norvegicus, forms dominant communities in the gut of wild and reared N. norvegicus populations, and has potentially positive effects on N. norvegicus digestive function. Thus, it is a very promising candidate for future use as a probiotic. This is the first study of gut bacterial communities in reared N. norvegicus. By analyzing the community changes under different diets for a rearing period of 6 months, it was shown that food intake promoted the establishment of specific bacterial communities, which were dominated by species that have been found to occur previously in natural populations of N. norvegicus, rendering them possible resident symbionts. From these bacterial species, P. leiognathi appeared as the most promising candidate to be used as a probiotic in future N. norvegicus rearing efforts. Acknowledgments We thank Ioannis Karapanagiotidis for his contribution to the rearing experiments and Zisis Petmezas, Konstantinos Kroupis, Vaso Kefeke and Maria Sakkomitrou for helping in the rearing experiments, as well as the histological and physiological analyses. The two fishermen from Volos are fully acknowledged for their assistance in the collection of the lobsters. AM would also like to thank the International Max Planck Research School of Marine Author's personal copy A. Meziti et al. / Systematic and Applied Microbiology 35 (2012) 473–482 Microbiology (MarMic), Bremen, Germany program for supporting part of this work, and Elmar Pruesse and Tryfonas Farmakakis for their assistance in the bioinformatics part of this work. Appendix A. 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