Changes in the gut microbial assemblages associated with dietary changes are still poorly explored, but may have profound effects in the life-history and physiology of fishes. Here, we have investigated the influence of the diet type on the autochthonous gut microbiomes of gilthead seabream (Sparus aurata). We conducted a 60-day assay in which a dietary shift from commercial to a diet based only on shrimp, followed by determination of gut microbial assemblages using 16S rRNA gene amplicon sequencing. The most prevalent species belonged to the genus Ralstonia, present in all samples, but was enhanced after the switch to shrimp diet. This seems to be a key genus whose functional relevance needs still to be unveiled given the wide distribution in samples with marine origin. Our results also pointed that nor the diet or the environmental microbiota (i.e. present in the water tanks) played a paramount role in the structure of the gut microbiome at the long-term. After 40 days, the gut microbial communities under both dietary treatments were very similar, being some species of the genera Ralstonia, Paraburkholderia, Fulvimonas, Pseudomonas and Cutibacteriumpart of the core-microbiome conserved at similar abundances over time. These results may challenge the common wisdom that diet is the primary determinant of gut microbiome composition in the long-term. In the era of the search for alternatives to fish-based ingredients in the aquafeed sector, and considering the paramount role of the gut microbiota in the fish health, this work offers valuable information for feed manufacturers to explore different nutritional strategies for aquaculture production, as long as the impact of the feeds on the physiology and quality of the fish are also tested.
Article
Resilience of gut microbial assemblages in farmed fish revealed by long-term dietary changes
https://doi.org/10.21203/rs.3.rs-3778254/v1
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Fish gut
microbial diversity
16S amplicon
aquaculture
Sparus aurata
The fish gut microbiome plays an important role in numerous aspects of fish life-history and physiology, including digestion, nutrient absorption, immunity, and even behavior or stress response [1]. In recent years, understanding the microbial diversity and their metabolic function has garnered increasing attention due to its profound implications for fish condition and health, aquaculture practices, and ecosystem sustainability. The composition and diversity of the fish gut microbiome are influenced by various factors, such as habitat, host genetics, environmental conditions, and particularly, the diet-associated factors [2, 3]. Different dietary components, such as carbohydrates, proteins, lipids, and fiber, provide distinct substrates for microbial metabolism, that together with the accompanying microbes may have an influence in the microbial species diversity [4, 5]. The growing interest in optimizing the utilization of fish meal and fish oil in aquafeeds has prompted the exploration of sustainable and economically viable alternative ingredients. This exploration aims to identify substitutes that not only align with environmental sustainability but also contribute positively to fish health, with a particular focus on understanding the potential influence of dietary components on the gut microbiome [6–8]. In addition, the importance of studying the diet-microbiome modulation also lies in the fact that dietary alternatives cannot only have health-promoting effects, but may also cause dysbiosis, impairing the composition and/or functions of the gut microbiota, which subsequently can have a physiological effect, such as the generation of an intestinal injury, inflammation or a reduced immune response [9]. Hence, examining the alterations in microbial diversity arising from dietary changes can offer valuable insights into the formulation of sustainable aquaculture practices.
In our previous study [5], we differentiated between a transient microbiota, probably accompanying feed and not clearly active in the gut, and the autochthonous microbiome that establishes close to the mucosa and seems to be permanently associated with the fish. In addition, the autochthonous microbiome can be divided in core that is stable and persistent regardless changing factors and normally composed of highly abundant members, and the non-core, which is more modulable and transient [10], and as demonstrated for mammals, fish also have a core gut microbiome [11–13]. In the assessment of the gut microbial structures in wild pearly razorfish (Xyrichtys novacula), and wild and farmed gilthead seabream (Sparus aurata), we could reveal how the global microbiomes changed in relation to different environmental conditions [5]. In the context of aquaculture species, variations in feeding habits between wild and farmed fish are apparent. Our findings revealed that the ecological context and feeding behaviors may exert a more pronounced influence on the structure of intestinal microbiomes compared to host species. Notably, two distinct wild fish species exhibited more similar microbiomes than the pair consisting of a wild fish and a farmed S. aurata, suggesting that the environment and dietary practices play significant roles in shaping the microbial composition [5]. In the same study, we also showed that fasting for 86 h the captivity individuals led to a decrease in species diversity, which reached similar values to the wild fish. Thus, we pointed out to that if the fish is not previously fasted, most of the microbiome detected is transient, non-stable and ephemerous, and to reveal the autochthonous (mucosa-adhered) bacteria, a previous fasting period is necessary.
Gilthead seabream (S. aurata) is the second most farmed fish in the European Union (first being rainbow trout), with an estimated aquaculture production which reached 320,630 tones in 2021 [14]. This sparid species is predominantly carnivorous and eventually herbivorous. Consequently, within the context of European aquaculture, extensive endeavors have been undertaken to investigate the impact of sustainable, readily available, and economically viable alternative ingredients using gilthead seabream as a model species [15, 16]. While current studies are focusing on the effect of the alternative ingredient and/or feed formulation on the growth, feeding performance, and nutritional quality of the fish [15, 16], their microbiomes are still poorly studied and unknown despite to their close connection with the animal’s health and welfare. In the current study, we aim to evaluate the dynamics of gut resident microbial communities in farmed S. aurata fed with commercial feed and assess the changes upon a shift in their diet to deep-water pink shrimp (Parapenaeus longirostris). Here, we applied 16S rRNA gene amplicon sequencing followed by the operational phylogenetic unit (OPU) approach for gut microbial assemblages determination along a 60-day assay [17, 18].
Experimental setup
The experiment was conducted at Insitute of Agrefood Research and Technology (IRTA) facilities, in La Ràpita (Tarragona, Spain). Sparus aurata juveniles (1.6 g in body weight, BW) were purchased from a commercial fish hatchery in the Est coast of Spain and transported by road to IRTA. A subset of these fish was used in our previous assay, so details about their acclimation may be found there [5]. Fish with an average initial body weight (BWi) of 25.29 ± 4.90 g (standard deviation, SD) were randomly distributed in two tanks, each with a volume of 400 L (n = 30 per tank), connected to an IRTAmar™ water recirculation system, where they were maintained for 4 months. During three months the fish were fed with the commercial feed D-2 Optibream AE 1P (2.2 mm pellet size, Skretting; Nutreco N.V.), and in the fourth month, the commercial feed was progressively switched to D-4 Optibream AE 3P (4 mm pellet size, Skretting) in order to adjust pellet size to fish growth. The macronutrient proximate composition of D-2 Optibream AE 1P was 48.5% crude protein, 18% lipids and 18.5 MJ kg− 1 digestible energy, while D-4 Optibream AE 3P contained 44% crude protein, 20% lipids and 18 MJ kg− 1 digestible energy (Serie D2-D5 Fishmeal composition is: fish oil, vegetable oils, cereal products and by-products, legume products and by-products, oilseed products and by-products, vitamins and minerals). After 4 months, fish from one tank continued being fed with D-4 Optibream AE 3P (Skretting), while those from the second tank were switched to a diet exclusively based on deep-water pink shrimp (P. longirostris). For the diet shift assay, 25 kg of entire individuals of P. longirostris (total length: 4–6 cm) which had been frozen only once just after capture with no further manipulation, were purchased from a local fishing company. However, commercial crustaceans normally contain sodium metabisulfite (E223), which is a synthetic antioxidant preservative added to avoid melanosis after removal from the water with a slight antimicrobial activity [19]. To avoid the potential penetration of sodium metabisulfite into the shrimp tissues, before thawing, shrimps were cleaned with abundant sterile seawater. Afterwards, shrimps were cut into pieces of between 2–3 cm length to facilitate their ingestion. Fish from both tanks were manually fed ad libitum three times per day for a period of 60 days. Specimens from both tanks were collected at 20, 40, and 60 days, which we refer to as T20, T40, and T60 datasets, respectively. In each sampling time, eight specimens were collected from each condition (Sup. Figure S1 and Sup. Table S1). To monitor the tanks’ water quality at the microbiological level during the experiment, each week an aliquot of 100 µL was spread onto TSA (Trypticase Soy Agar) + 0.6% NaCl, and TCBS (Thiosulphate Citrate Bile Salts Sucrose) Agar medium and incubated at 22 ºC for 48 h to obtain the Total Viable Counts (CFU) of heterotrophic microorganism and Vibrionaceae, respectively.
The water temperature (18–22 ºC) and dissolved oxygen (6.4 ± 0.6 mg L− 1; OXI330, Crison Instruments) were daily measured and controlled, and pH (7.4 ± 0.2; pHmeter 507, Crison Instruments), salinity (36‰; MASTER-20 T, ATAGO Co. Ltd), ammonia (0.14 ± 0.1 mg NH4+ L− 1) and nitrite (0.2 ± 0.1 mg NO2− L− 1) levels (HACH DR9000 Colorimeter, Hach, Spain) were weekly monitored. Just after T20, the fish were transferred to two other 400 L tanks for the cleaning and maintenance of the tanks in this module, but the water quality and environmental conditions among modules were maintained.
Sample processing
For each sampling point, fish were manipulated as detailed previously [5]. Briefly, after a fasting period of 86 h for collecting only the autochthonous microbiota [20, 21], fish were euthanized with an overdose of the buffered anesthetic (tricaine methanesulfonate, MS-222, Sigma-Aldrich; 300 mg L− 1). The entire intestinal tract was aseptically extracted and the perivisceral fat surrounding the intestine was gently removed with a scalpel; then, a 3 cm section of the anterior intestine and another of the posterior intestine with the same length, were dissected using sterile scissors, and opened lengthwise. The mucosal content was gently scraped with a round edge spatula and immediately frozen at -80 ºC for further microbial analysis.
Microbial composition of commercial feed and shrimp
Commercial feed pellets (D-4 OptiBream AE 3P) and samples of shrimps were stored at -80 ºC at each sampling time. For processing purposes, three feed pellets and three shrimps were crushed with an autoclaved mortar and cut into very small pieces with sterile scissors, respectively. A total amount of ca. 150 mg in wet weight from feed or shrimps were collected and used for DNA extraction by means of the FastDNA™ Spin Kit for Feces (MP Biomedicals) following the manufacturer’s protocol. Three extractions for feed and shrimps were performed from each sampling time.
DNA isolation, amplification, and sequencing
The FastDNA™ Spin Kit for Feces (MP Biomedicals) was used to perform DNA extraction from the scraped mucosa content (up to 150 mg), following the manufacturer’s instructions. Illumina amplicon sequencing was performed using the set of primers for Bacteria forward (5’ - CCTACGGGNGGCWGCAG − 3’) and reverse (5’ - GACTACHVGGGTATCTAATCC − 3’) [22] containing the forward (5’ - TCGTCGGCAGCGTCAGATGTGTATAAGAGACAG − 3’) and reverse (5’ - GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAG − 3’) Illumina sequencing adapters. Sequencing was performed using an Illumina MiSeq instrument (2x250 bp) (FISABIO, Valencia, Spain).
Amplicon sequence variant (ASV) and operational phylogenetic unit (OPU) approaches
Raw sequencing data was analyzed using the Qiime2 bioinformatic platform [23] with the parameters --p-trunc-len-f 280, --p-trunc-len-r 220, --p-trim-left-f 19 and --p-trim-left-r 22. Amplicon sequence variant analyses (ASV) were obtained with DADA2 software implemented in Qiime2. The longest sequence of each ASV was selected as representative, and sequences of the bacterial and archaeal ASVs were aligned using the non-redundant SILVA REF 138.1 [24] database and the ARB package [25]. Sequence alignment was performed using the SINA tool implemented in the ARB program [26]. The aligned sequences were inserted in the SILVA REF 138.1 database containing the pre-stablished OPUs detected in Viver et al. [5] for taxonomic classification, using the parsimony tool available in the ARB package. The non-affiliated sequences to the pre-stablished OPUs were inserted again in the SILVA REF 138.1 database to select the close relative sequences for phylogenetic analysis and denovo OPUs identification.
Statistical analyses
After verifying normal distribution (Shapiro-Wilk test) and homoscedasticity (Levene's test) of data, a Student's t-test was performed for testing significant differences in BW and standard length (SL) between both dietary groups (P ≤ 0.05). To analyze the alpha and beta diversity indices and for statistical purposes, the dataset was rarified to the minimum sample depth (21,494 sequencing reads per sample) with the vegan R package [27]. To assess the phylogenetic complexity of the samples, rarefaction curves, Shannon and Dominance indices were calculated using the PAST statistic tool [28] based on the diversity and abundance of 16S-based OPUs. The microeco R package [29] was used to calculate the Bray-Curtis dissimilarity values between the different community profiles of samples and periods of the assay based on OPUs. Non-metric multidimensional scaling (NMDS) plot based on Bray-Curtis dissimilarity was constructed using the vegan package [27] and plotted using the R package ggplot2 [30] in R tool version 3.6.3 [31]. A divergence test between samples based on OPU abundance and composition was performed using non-parametric two-sample Kolmogorov-Smirnov tests [32], since the data did not meet the assumptions of normality and homoscedasticity.
Data availability statement
16S rRNA gene amplicon sequences were submitted to the European Nucleotide Archive (ENA) under the study number PRJEB64773, and accession numbers from ERS16220008 to ERS16220143.
ARRIVE guidelines
This work was performed in accordance with the ARRIVE guidelines [33].
Ethics statement
All animal care procedures were approved by the Ethical Committee of Animal Experimentation of the Institute of Agrifood Research and Technology (Spain) and the Generalitat of Catalunya (Ref. CEEA 219/2020) and were carried out by trained competent personnel, in accordance with European Directive 2010/63/UE and Spanish Royal Decree RD53/2013 to ensure good practices for animal care, health, and welfare. The IRTA facilities are certified and have obtained the necessary authorization for the breeding and husbandry of animals for scientific purposes. Experimental procedures complied with the Guiding Principles for Biomedical Research Involving Animals (EU2010/63) and with the ongoing Spanish legislation (Law 32/ 2007 and RD 1201/2015).
Fish growth
Along the 60-day experimental trial, we observed a consistent BW pattern in both dietary groups. At the initial sampling day in which a group was submitted to the diet change (T0), the fish weighed an average of 158 ± 32.9 grams. During T20 and T40, the fish feed with shrimp showed, a significantly higher BW (211.2 ± 23.8 and 220.3 ± 31.9 g, respectively) than the fish feed with commercial feed (190.0 ± 25.1 and 202.1 ± 34.1 g, respectively; P < 0.05). At the end of the trial (T60), fish feed with both diets exhibited a similar BW (Wilcoxon-test; P = 0.59): an average BW of 259 ± 48 g for those fed with commercial feed and 252 ± 31.8 g for those fed with deep-water pink shrimp (P > 0.05; Table 1 and Sup. Table S1).
Samples phylogenetic diversity based on OPUs
After trimming and chimera removal, all samples included in this study rendered a total of 10,957,204 sequenced reads, and the number of sequences per sample ranged between 21,494 and 160,269. All sequences were clustered in 8,072 ASVs and after phylogenetic inference a total of 1,256 OPUs were detected (ranging between 22 and 428 OPUs per sample) (Sup. Table S2). To ensure statistical accuracy, the dataset was rarified at the lowest number of sequences detected in a single sample (~21,000; Sup. Table S3), and the rarefaction curves demonstrated that diversity had reached saturation level in all of the rarified datasets (Sup. Figure S2).
Taking into consideration that an OPU is the smallest cluster of query sequences that affiliates with at least one reference sequence and considering that the size of each cluster typically falls within a sequence distance of less than <97 % (considering the relatively short length of ~400 pb for partial sequences), we can assert with confidence that an OPU represents a single species [18]. Each OPUs is recognized as a distinct species within genera whenever possible. Here, we used the commonly applied terminology for the higher taxa as in the past. We are aware that certain phyla names have been officially proposed by Oren and Garrity [34] as Bacillota (= Firmicutes), Bacteroidota (= Bacteroidetes), Pseudomonadota (= Proteobacteria), Actinomycetota (= Actinobacteria), Planctomycetota (= Planctomycetes), Chloroflexota (= Chloroflexi), Fusobacteriota (= Fusobacteria) and Chlamydiota (= Chlamydia), but for pragmatic reasons and comparative purposes with available published literarature, we still prefer retaining the former nomenclature.
The phylogenetic analysis showed that the 1,256 OPUs were classified in 620 genera and 314 families, affiliated with 38 different phyla (Sup. Table S4 and S5). In agreement with our previous assay [5], the phylum Proteobacteria was the most prevalent phylum, accounting for 42.9% of the total OPUs. Within this phylum, the majority of OPUs were classified as Gamma- and Alphaproteobacteria, representing 20.7% and 18.9% of the OPUs, respectively. Firmicutes (22.9% of the OPUs), Actinobacteriota (12.7%), Bacteroidota (7.0%), and Planctomycetota (2.2%) were also predominant in the examined samples. Furthermore, the study identified a total of 620 genera, from which 514 were represented by only one OPU. However, there was a considerable OPU diversity within some genera, such as Vibrio (32 distinct OPUs), Bacillus (29 OPUs), Lactobacillus (27 OPUs), Pseudomonas (22 OPUs), and Paenibacillus (15 OPUs).
Microbial composition of the feed pellets and shrimp
The microbial diversity patterns showed that the commercial feed and shrimps presented the highest diversity values, as measured by the Shannon index (average of 2.09 ± 0.22 in the commercial feed and 3.35 ± 0.58 in pink shrimps), and the lowest dominance values (average of 0.22 ± 0.04 in the commercial feed and 0.13 ± 0.1 in pink shrimps), indicating a more even distribution of species compared to the fish gut microbiomes (Figure 3 and Sup. Table S6). Shannon diversity values were positively correlated with the detection of a higher number of specific OPUs from commercial feed datasets. Out of the 424 OPUs detected in commercial feed and 810 OPUs detected in pink shrimp, a substantial proportion (30% and 35.7%, respectively) were exclusive to each set of samples. In contrast, the exclusive detected OPUs in the fish microbiomes ranged from 6.1% to 2.0% (Sup. Table S4). The microbial diversity of the commercial feed remained relatively consistent throughout the experiment, with the lowest range of Bray-Curtis values (average value of 0.12 ± 0.06) (Sup, Table S7) suggesting that storage did not significantly alter the microbial assemblage of feed pellets for at least 60 days. The OPU analysis revealed that 12 microbial species were consistently abundant (³ 1% in all samples), and their combined abundance ranged from 89.8% to 95.02%. Among them, six of these species affiliated with phylum Firmicutes, specifically the class Bacilli, where the OPU0015, OPU0011, OPU0036 and OPU0037 affiliated with Lactobacillus (average abundance of 36.02 ± 5.8%, 9.02 ± 1.4%, 1.75 ± 0.4% and 1.67 ± 0.3%, respectively), Bacillus OPU0145 (2.61 ± 0.4%) and Staphylococcus OPU0076 (1.49 ± 0.3%). The other highly abundant OPUs affiliated with Ralstonia OPU0522 (4.4 ± 3.6%), Photobacterium OPU0624 (3.47 ± 0.9%) and Leptospira (0.35 ± 0.4%). Moreover, a high abundance of chloroplasts (OPU1059) and mitochondria (OPU1038) sequences were detected in all samples (Figure 1 and Sup. Table S8). Ralstonia OPU0522 was found to be shared among the commercial feed (4.39 ± 3.65%), deep-water pink shrimp (18.84 ± 19.51%) and all fish gut microbiome samples (37.81 ± 29.37%) (Figure 1)
The microbial composition of shrimp exhibited a slightly higher level of heterogeneity (average Bray-Curtis value of 0.54 ± 0.2) than the compound feed, but all samples were closely clustered in the NMDS plot (Figure 2A). This variability could be associated to the batch selected for DNA extraction and sequencing. The high abundant OPUs detected in shrimp and absent or poorly detected in fish microbiomes affiliated with the Flavobacterium OPU1209 (relative abundance of 14.4 ± 11.9%) and Chryseobacterium OPU_R402 (2.1 ± 1.5%) of the Bacteroidia class; Glutamicibacter OPU0420 (2.4 ± 1.4%) and Psychromicrobium OPU0427 (2.07 ± 1.4%) of the Actinobacteria class; the Psychrobacter OPU0705 (11.9 ± 7.3%), the Oceanimonas OPU_P06 (1.9 ± 1.9%) and Marinomonas OPU0684 (3.5 ± 1.78%) of the Gammaproteobacteria class.
The microbial gut content of specimens fed with commercial diet
All S. aurata juveniles were fed with commercial pellets upon arrival to the experimental facilities and during the 4 months preceding the start of the experiment on the shift of food items. At time zero (T0), the control group continued being fed with the commercial diet over a period of 60 days and we did not observe any differences in species diversity and abundance between distal and proximal intestine for 80% of the specimens studied (n = 16 intestinal samples per diet) as indicated by the Kolmogorov-Smirnov test (Sup. Table S9). At T0, all samples exhibited a similar dominance and Shannon indices with T20 (with an average dominance value ranging from 0.47 ± 0.15 to 0.47 ± 0.22 and Shannon index ranging from 1.22 ± 0.46 to 1.33 ± 0.53 in T0 and T20, respectively). We detected a slight increase in diversity with a decrease in the dominance after 40 and 60 days (average dominance value ranging from 0.37 ± 0.14 to 0.39 ± 0.16 and a Shannon index ranging from 1.44 ± 0.43 to 1.46 ± 0.55 in T40 and T60, respectively) (Figure 3 and Sup. Table S6). No significant differences were observed in dominance and Shannon indices over time (Wilcoxon-test; P > 0.05). The NMDS plot, based on the Bray-Curtis index, indicated that the dissimilarity between the control samples from the T0 and T20 datasets was higher (averaging 0.67 ± 0.24 and 0.7 ± 0.24, respectively) compared to that within the T40 and T60 datasets (averaging 0.46 ± 0.23 and 0.53 ± 0.29, respectively), suggesting a higher species homogeneity among individuals in the final stages of the experiment (Figure 2B and Sup. Table S7). Comparisons between T0 and the rest of sampling times indicated that T60 showed the most significant microbial diversity changes (average Bray-Curtis dissimilarity of 0.85 ± 0.12 between T0 and T60 samples) (Sup. Table S7).
Commercial feed and control fish in all timepoints shared 206 OPUs, confirming that commercial feed was the source of these species in fish gut (Sup. Table S8). Notably, the species Ralstonia OPU0522, Paraburkholderia OPU0542, Leptospira OPU1052, Fulvimonas OPU0572, Pseudomonas OPU0713 and Cutibacterium OPU0409 were consistently detected in both the feed samples and fish collected throughout the experiment (Sup. Table S8). There were microbial species exclusive in each dataset or sampling day. Out of the 319 total species detected, 5.6% were exclusive of the initial time (T0). At 20 days (T20), the percentage of exclusive species decreased to 4.6%, and at T40 and T60 the values were of 2.2% and 3.6%, respectively (Sup. Table S4). The genus Vibrio exhibited the highest richness with 32 OPUs identified. In the fish, ten Vibrio species were identified as highly abundant (Figure 1). However, the prevalence of these species varied significantly over time. Specifically, Vibrio OPU0604, OPU0606, OPU0599, OPU0581 and OPU0594 were highly abundant in T0 and T20 days, and Vibrio OPU0600 and OPU0588 showed higher abundance in samples T20, T40 and T60. Aliivibrio OPU0614, OPU0615 and OPU0616 were abundant in T20. Candidatus Kaiserbacteria OPU1085 and Leptospira OPU1052 were highly abundant at T0 and T20 days, while Psychrobacter OPU0705, Corynebacterium OPU0366, Acinetobacter OPU0701 and Subdoligranulum OPU0242 were highly abundant in T40 and/or T60 days.
The microbial gut content of specimens fed with shrimp
The feed of the second group was replaced by shrimp at T0 and was maintained during 60 days. In this case, the alpha diversity values showed that the switch to a shrimp diet resulted in a reduction in microbial diversity along the first 20 days, as indicated by the non-significant, but numeral decrease in Shannon values (average from 1.22 ± 0.46 at T0 to 1.04 ± 0.25 at T20 (Wilcoxon-test; P = 0.42)), accompanied by a significant increase in dominance (average from 0.47 ± 0.15 at T0 to 0.63 ± 0.12 at T20 (Wilcoxon-test; P = 0.008)). However, after a gradual increase in diversity and a decrease in dominance occurred, reaching similar levels to the control at T60 (average Shannon value of 1.6 ± 0.27 (Wilcoxon-test; P = 0.26) and dominance of 0.33 ± 0.1 at T60 (Wilcoxon-test; P = 0.28)) (Figure 3 and Sup. Table S6).
The NMDS plot shows that the samples became more homogeneous after 20 days reflected by decrease in the average Bray-Curtis value from 0.67 ± 0.24 at T0 to 0.3 ± 0.25 at T20. This shift coincided with a significant increase in the dominance of the Ralstonia OPU0522, which exhibited an average abundance value of 73.59 ± 20.49% at T20. A temporal shift in beta diversity was observed during the experiment, with the most significant changes occurring in the samples collected at T60 (Figure 2C). We observed a similar pattern for the control dataset in the percentage of exclusive OPUs, with 6.1%, 6%, and 2% of OPUs being unique to T20, T40, and T60, respectively (Sup. Table S4). We also observed a shared number of 324 OPUs between the shrimps and the gut microbiome of the fish fed with them. Similar to what was observed with the control, the most abundant OPUs from shrimp did not establish in the gut microbiomes, except for Paraburkholderia OPU0542, Leptospira OPU1052, Pseudomonas OPU0713, Psychrobacter OPU0705 and Acinetobacter OPU0701 (Figure 1). The Candidatus Kaiserbacteria OPU1085 was manly present in samples T20 and Vibrio OPU0600 and OPU0573 were mainly detected in specimens collected at T40 and T60. On the other hand, Acidovorax OPU0530 and Aliivibrio OPU0614, OPU0615 and OPU0616 were most abundant in T60 (Figure 1).
The NMDS based on Bray-Curtis dissimilarity indicates that despite the different diets the fish communities clustered together at the final stage of the experiment (T60) (Figure 2A). Moreover, when evaluating dominance and Shannon index values at T60, no statistically significant differences were observed between fish fed with commercial feed and shrimp (Wilcoxon-test; P = 0.05).
In this study, we examined the impact of a dietary shift (from a compound diet to a shrimp-based diet) on the resident (autochthonous) gut microbiota of a commercially significant fish species. In all cases, bacteria accompanying the feed had a limited influence on the diversity in the gut, which was evidenced by the independent clustering of the diets with respect to the fish as well as by the different Shannon diversity and dominance indices. For instance, highly abundant species in the commercial feed or in shrimps did not establish in fish gut. Other studies also described different bacterial compositions between feed and intestinal autochthonous microbiota of many teleosts as gilthead seabream [35], European sea bass (Dicentrarchus labrax) [36] or rainbow trout (Oncorhynchus mykiss) [37]. Our findings are also supported by similar studies using experimental diets for gilthead seabream with increasing levels of Firmicutes that not only did not establish, but rather decreased in abundance over time [38], or the particular case of the European sea bass [36], in where the feed pellets were dominated by Lactobacillus that did not either establish in their guts [36].
Contrarily to our findings, there are other studies that have shown a direct establishment of the feed-associated bacteria in fish species like yellowtail kingfish (Seriola lalandi) [39] and rainbow trout [40]. However, some of these studies may have some caveats as e.g. for the yellowtail kingfish [39], no previous fasting was reported and were probably detecting both indigenous and transient microbiota as we demonstrated in our previous study [5]; or for the rainbow trout [40], where the lumen-associated bacteria was collected but not the mucosal, which explains the differences with our results. Commercial feeds are submitted to an extrusion process reaching high temperatures (> 100 ºC) that would washout the vast majority of viable bacteria [41, 42], but some species, as the observed Lactobacillus sp. may develop as a consequence of fermentative conditions during feed processing, manipulation or storage. These organisms or their residual DNA can be identified through amplicon sequencing, even in cases where they are non-viable, signifying their transient presence in the fish gut.
One of the most relevant species detected was Ralstonia sp. 1, which was found already with remarkable abundance in our previous study [5]. Members of this genus are normally found as part of the autochthonous gut microbiota in several fish species, such as rainbow trout [43], yellow catfish (Pelteobagrus fulvidraco) [44], European sea bass [45], yellowtail kingfish [46], large yellow croaker (Larimichthys crocea) [47], pearly razorfish (Xyrichtys novacula) and gilthead seabream [5]. Here, Ralstonia was present in all samples, but its abundance was clearly enhanced in fish fed with shrimps. Contrarily to the commercial feed, beyond freezing, shrimps were not submitted to a pre-processing to destroy or minimize the accompanying bacteria. Thus, the increased amounts of Ralstonia in the fish fed with shrimps may partially respond to the colonization of this genus as part of the allochthonous microbiota. Despite the commercial feed also contained Ralstonia, probably originated from fish meal, these bacteria may be not viable and this fact would explain the lack of their growth in fish guts. Although further research is needed to unravel the role of the member of this aerobic genus in the fish intestine, we believe that this species may play an important role in marine animals, and the significant presence in the guts of fish may respond to a symbiotic or mutualistic relation. Indeed, some Ralstonia species have been suggested to have antimicrobial activity or biosynthesize bioactive compounds, and could produce beneficial secondary metabolites for the host fish [48].
Altogether, here the type of feed does not seem to be a key factor as a source of bacteria, as no remarkable changes were found between the two groups for the rest of represented species (> 2%). However, the diet ingredients could modulate the gut autochthonous microbiomes. Utilizing diverse protein, lipid, and carbohydrate sources, or incorporating varying levels of dietary supplements, can induce a spectrum of pronounced effects on the microbial profile of the fish. These effects may be directly and/or indirectly initiated by alterations in host physiology [49]. For example, here, the effect of the diet was reflected in the differential relative abundances of some Aliivibrio species in the group fed shrimps, especially at T60, that were not detected when using the commercial diet. These species related to A. finisterrensis, A. fischeri, and A. logei, seem to be established in the gut of marine teleosts [50-52]. Some Aliivibrio species in the gut of the Atlantic cod (Gadus morhua) has been reported to be stimulated by the supplementation with chitin [53], one of the main components of exoskeletal shells from crustaceans [54]. In aquafeed, chitin is considered as a prebiotic, able to increase bacterial intestinal richness by favoring the growth of beneficial and chitin-degrading bacteria [55, 56]. In this sense, many Aliivibrio species have chitinase activity [57], which may aid in the digestion of this ingredient since many marine fish, such as gilthead seabream, have shown a low chitin digestibility [58-60]. Then, it seems plausible that the higher growth of Aliivibrio in fish fed with shrimp with respect to their congeners fed with commercial feed was due to the probiotic effect of chitin. In addition, we cannot discard that the transient feed-associated microbes that do not establish in the gut, if viable, may also influence the gut autochthonous microbiota, e.g., the probiotic effects in modulating microbial functionality and exerting a beneficial effect on the fish health [37, 61, 62].
Beyond dietary factors, the gut microbiome is also influenced by the environmental conditions such as salinity and/or temperature that are paramount drivers determining fish microbiomes [12, 63, 64]. In this context, we observed and increase in abundance of Staphylococcus, Acidovorax, Vibrio, Corynebacterium, and Psychrobacter in both groups regardless the diet provided just after T20 when we relocated the fish in the new tanks for the cleaning and maintenance. Although we maintained the environmental conditions between tanks after the change, the differential microbiota in the water and on the tanks’ wall biofilms might have been the source of these taxonomic changes [65, 66]. Unfortunately, we did not collect samples from water or biofilms from the tanks to compare their taxonomic profiles, and therefore, we can just hypothesize that these bacteria were present in the new tanks.
Some of the changes induced by the relocation were more marked in gilthead seabream fed with the commercial diet, such as the differential abundances of some Vibrio species and Clostridium sp. 22 before and after T20. The reason why the impact of the water change was more pronounced in fish fed with the commercial diet than in those fed with shrimps is that at the moment of the relocation fish fed the commercial feed had been already adapting to this diet type for 7 weeks plus other 3 previous months being fed with a similar commercial diet after T20. In this sense, a long-term microbial adaptation of many weeks will be expected in response to any external changing factor, such as a shift in the diet [67, 68] or in the feeding regime [69], an antibiotic administration [70], or an induced stress [71].
Under current experimental conditions, the most striking observation was to find that there were some bacteria which were present in all groups of individuals at very similar average abundances regardless the sampling time and diet provided, and consequently, Ralstonia sp. 1, Paraburkholderia sp. 1, Fulvimonas sp. 1, Pseudomonas sp. 1 and Cutibacterium sp. 1, could be considered as part of core microbiome of this farmed species [10-13]. These similarities in terms of species composition and abundance were reflected in a similar diversity (Shannon diversity and Dominance indices) and structure (Bray-Curtis distances) of both dietary groups at the later stages of the experiment (T40 and T60). It seems that the core microbiome prevalence is not affected by the environmental changes after the adaptation processes in the tanks. In contraposition, studies in juveniles of rainbow trout [67] and brown trout (Salmo trutta) [68] have shown that the microbiome composition is directly dependent in the actual diet provided after a dietary shift, but barely influenced by the initial diet provided before the change. However, these juveniles were younger and with much lesser body weight (around two orders of magnitude smaller) than ours at the end of the assay, and this may indicate that the differential plasticity of the fish gut microbiota to adapt to diet changes depends on the animal age as it was previously postulated in gilthead seabream [72] as well as in mammals gut [73, 74] and rumen [75].
Previous results have already showed that there are host-inherent factors which act as paramount modulators of the fish gut microbiota, such as the species, sex, age, and genetic background [72, 76]. Overall, the present results suggested that a diet change have much less weight in the modulation of the gut microbiota than these host-inherent factors. This is consistent with the evidence of a core microbiome in all fish regardless the diet provided, leading to gut bacterial communities with a similar diversity, structure, and composition in the long-term. As long as the impact of the feeds on the health and quality of the fish are also tested, the findings herein could be used as an advantage in the aquaculture industry to explore novel feeds formulated with sustainable, cost-effective, and readily available ingredients with the security that in the long-term most part of the gut microbiota will adapt and persist to the diet influence and to the adjustments of feed formulae linked to the nutritional requirements of the fish for an optimal growth over the production cycle.
Author Contributions Statement
TV, AR, JA and DF designed the experiment. AR collected and processed the farmed fish, and isolated, amplified and sequenced the DNAs. TV analyzed the sequenced data and prepared figures and tables. TV and AR wrote the manuscript. EG and DF, resources, supervision and validation. All authors reviewed, discussed, and edited the manuscript.
Acknowledgements:
We acknowledge Ramon Rossello-Mora for critical reading and recommendations, and also for participating in funding the amplicon sequencing of the samples included in this study by the projects CTM2017-91490-EXP and PID2021-126114NB-C42 of the Spanish Ministry of Science, Innovation and Universities which were also supported by the European Regional Development Fund (FEDER). TV acknowledges the “Margarita Salas” postdoctoral grant, funded by the Spanish Ministry of Universities, within the framework of the Recovery, Transformation and Resilience Plan funded by the European Union (NextGenerationEU), with the participation of the University of the Balearic Islands (UIB). AR was supported by a pre-doctoral grant (PRE2019-091259) linked to the ADIPOQUIZ project (RTI2018-095653-R-I00), funded by the Spanish Ministry of Science and Innovation. JA received funding by the METARAOR Project (Grant num. PID2022-139349OB-I00) funded by MCIN/AEI/10.13039/501100011033/FEDER, UE. The research was carried out within the framework of the activities of the Spanish Government through the "Maria de Maeztu Centre of Excellence" accreditation to IMEDEA (CSIC-UIB) (CEX2021-001198).
Animal welfare statement:
The IRTA facilities are certified and have obtained the necessary authorization for the breeding and husbandry of animals for scientific purposes. Experimental procedures were conducted following the Guiding Principles from Biomedical Research Involving Animals (EU2010/63) and the guidelines of the relevant Spanish laws (Law 32/2007 and RD 1201/2015).
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Table 1. Body weight (BW) of gilthead seabream (Sparus aurata) at the different sampling times: at their arrival to the facilities where the experiment was conducted; at the beginning of the trial, when they started to be fed with a commercial feed; at T0, when a group of fish was submitted to a dietary shift to deep-water pink shrimp; and after 20, 40 and 60 days from the diet shift (T20, T40 and T60, respectively). Values are represented as mean ± SD. There were no significant differences among fish fed the commercial feed and shrimp at any sampling time (Student's t-test).
No competing interests reported.