Jurnal Internasional 5v5211

Aquaculture 416–417 (2013) 396–406

Contents lists available at ScienceDirect

Aquaculture journal homepage: www.elsevier.com/locate/aqua-online

Probiotic, prebiotic and synbiotic applications for the improvement of larval European lobster (Homarus gammarus) culture Carly L. Daniels a,b,⁎, Daniel L. Merrifield b, Einar Ringø c, Simon J. Davies b a b c

The National Lobster Hatchery, Padstow, Cornwall, UK Aquaculture and Fish Nutrition Research Group, School of Biomedical and Biological Sciences, Plymouth University, Devon, UK Norwegian College of Fishery Science, Faculty of Biosciences, Fisheries and Economics, University of Tromsø, Tromsø, Norway

a r t i c l e

i n f o

Available online 13 August 2013 Keywords: Health Stress Microalgae Larval development Gastrointestinal microbiota

a b s t r a c t The effects of dietary applications of a commercial probiotic (Bacillus spp.) and prebiotic (mannan oligosaccharides [MOS]), used singularly and combined (i.e. synbiotic), on larval survival, growth, intestinal microbial communities and stress resistance of larval European lobster, Homarus gammarus, were assessed. Larvae were reared in green water culture for 12 days from hatch until metamorphosis to zoea III. Un-supplemented Artemia nauplii (control) or Artemia nauplii enriched with probiotics (Bacillus spp. 100 mg l−1), prebiotic (MOS 12 mg l−1) or synbiotics (100 mg l−1 Bacillus spp. + MOS 12 mg l−1) were each fed to 3 replicate groups of zoea I lobsters for 12 days. The effects on gut microbiota were assessed using culture-dependent methods at 1, 7 and 12 days post-hatch (dph) and PCR-DGGE at 1 and 12 dph. PCR-DGGE was also used to assess microbial communities of the live feeds. Carapace length and weight of five H. gammarus from each replicate was recorded on 1, 7, and 12 dph and survival to zoea III was recorded. A low salinity stress test was used as a measure of organism fitness at day 12. After 12 dph, H. gammarus larvae fed experimental treatments had significantly (P b 0.02) improved weight, carapace length and weight gain, compared to larvae fed control treatments. Survival to 12 dph was elevated by all treatments and was significantly (P b 0.001) increased for the Bacillus and MOS fed larvae. Salinity stress tolerance was greatest in larvae fed Bacillus although all experimental treatments produced enhanced tolerance to salinity stress in comparison to larvae fed control treatment. Culture-dependent analysis of the gut microbiota of larval lobsters demonstrated the colonisation of Bacillus spp. in larvae fed probiotic or synbiotic enriched live feeds. There was also a reduction in Vibrio levels in certain biotic fed larvae and live feed treatments. PCR-DGGE revealed that the number of observed taxonomical units (OTUs), species richness and species diversity increased in zoea III lobsters fed probiotic, prebiotic and synbiotic. Subsequently, the microbial profiles were dissimilar to the control group with the synbiotic group showing the greatest dissimilarity to the control (36.54 ± 2.54%). The similarity between bacterial communities associated with Artemia and zoea III larvae was highest in the Bacillus treatments (53.86%). The present study demonstrates the benefit of applying dietary supplementation of Bacillus, MOS and Bacillus + MOS on the GI microbiota of lobster larvae which subsequently improved growth performance and stress tolerance. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Probiotics, live microbial feed supplements which modulate gastrointestinal microbial communities, and prebiotics, non-digestible feed additives which stimulate the abundance or activity of beneficial gastrointestinal microbes, have received extensive attention showing improved production, health and disease resistance of aquatic animals (Dimitroglou et al., 2011; Merrifield et al., 2010). Furthermore, these applications have shown potential to enhance bacterial assemblages of live feeds used for aquaculture purposes, thus reducing the introduction ⁎ Corresponding author at: The National Lobster Hatchery, Padstow, Cornwall, UK. Tel.: + 44 1841 533877; fax: +44 8707 060299. E-mail address: [email protected] (C.L. Daniels). 0044-8486/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.aquaculture.2013.08.001

of opportunistic pathogens to the targeted cultured organism (Gatesoupe, 2002; Makridis et al., 2000; Verschuere et al., 2000). The bacteria present in the intestinal tract of aquatic organisms appear to represent those from both the environment and diets (Cahill, 1990; Verner-Jeffreys et al., 2003). This is particularly true for filter feeders, including many live feed species, such as crustacean larvae, including Artemia, which are affected by the environment due to the large flow of water into the body cavity during feeding (Gatesoupe, 1999; Olafsen, 2001). Studies examining the use of probiotic and prebiotic supplements in aquaculture have typically relied on recirculation systems (Dimitroglou et al., 2011). Media exchange must be considered in these situations, with the potential for bacterial contamination between experimental dietary treatments due to the environment. This is especially true for

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studies of live probiotic microorganisms, because potential intertreatment contamination in these situations may affect studies focussed on changes in bacterial abundances and possibly distort overall culture success, as well as morphological and physical outcomes. Isolated culture environments may yield better information to fully understand the use of probiotics in culture situations. In larval European lobster culture the use of isolated vessels and green water culture methods have shown success (Beal and Chapman, 2001; Browne et al., 2009) in comparison to the more traditional larval rearing techniques using recirculation systems (Beard et al., 1985; Daniels et al., 2010). Green water culture involves the supplementation of live microalgae to rearing vessels which has shown positive effects on general water quality; including reductions in Vibrio abundance (Makridis et al., 2009; Ritar et al., 2004), and can also provide nutritional and physiological benefits for the live feed (Artemia) subsequently delivered to lobster larvae (Marques et al., 2006; Ritar et al., 2004). European lobster larvae are unable to consume small algal cells (phytoplankton), this system therefore relies on the encapsulation of microalgae into live feeds known to be of suitable prey size for decapod crustacean, such as Artemia (Browne et al., 2009; Marques et al., 2006). Microalgae vary in their nutritional properties (Ritar et al., 2004) and several species, such as Isochrysis galbana and Chaetoceros muelleri, have been used for homarid lobster culture due to their rich fatty acid content (Browne et al., 2009). Various larval crustaceans are unable to produce specific fatty acids essential for growth, for example, the n−3 (linolenic) and n−6 (linoleic) fatty acids (Tamaru et al, 2003). Thus, microalgal species such as C. muelleri, Chaetoceros gracilis and I. galbana which are high in some essential fatty acids, such as EPA (20:5n−3), DHA (22:6n−3) and ARA (20:4n −6) (Ritar et al., 2004) are used routinely in the green water culture of Homarus spp. (Beal and Chapman, 2001; Uglem et al., 2006; Browne at al., 2009). Dietary probiotic and prebiotic supplements have been reported to improve the growth and survival of the larval stages of many aquaculture species (Gatesoupe, 1994, 2002; Rojas-Garcia et al., 2008; Salze et al., 2008; Skjermo et al., 2006; Suzer et al., 2008), including H. gammarus (Daniels et al., 2010). Improvements in animal quality have also been determined using stress tests (Dimitroglou et al., 2010; Salze et al., 2008). Enhancements in the culture success and quality of aquatic animals fed probiotics and prebiotics are thought to be due to immunomodulation (Sang et al., 2011; Staykov et al., 2005, 2007; Torrecillas et al., 2007; Zhou and Li, 2004), improved digestive morphology (Daniels et al., 2010; De Rodriganez et al., 2009; Dimitroglou et al., 2009; Salze et al., 2008; Torrecillas et al., 2007) and modulated intestinal bacterial populations (Dimitroglou et al., 2009). Therefore, the aim of the present study was to investigate the effect of the individual and combined use of a probiotic, Bacillus spp. (SANOLIFE® - INVE Aquaculture, Belgium), a prebiotic, MOS (BIOMOS® - Alltech Inc., KY USA) and the combined use of Bacillus spp. with MOS (i.e. synbiotic treatment) on the growth performance, survival, quality and gut microbiota of cultured H. gammarus larvae using isolated green water techniques.

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NO2 b 0.1 mg l−1. Every 48 h, 100% water changes were performed with vessels left to acclimate to the desired temperature for 2 h before larvae were transferred and then feed and microalgae were added. Two species of live microalgae (I. galbana and C. muelleri) were added to each rearing vessel at a density of 150 cells μl−1 calculated using an algal cell counting chamber (Improved Neubauer, Weber Scientific International Ltd. Sussex, UK). Microalgae density was monitored and replenished daily where appropriate to keep density constant at 300 cells μl−1. The cones were vigorously aerated (14 l min−1) from the base using a Koi Air 65 air compressor (Blagdon, Surrey, UK). All cones were covered with clear polythene lids to prevent cross contamination by micro-organisms. Newly hatched larvae were added to 80 l conical vessels across a maximum period of two consecutive days to a maximum density of 12–14 larvae l−1 (replicate: 1 = 12.1; replicate 2 = 12.7; replicate 3 = 13.75 larvae l−1). 2.2. Experimental diets and feeding H. gammarus larvae were fed with enriched Artemia (Salt Creek Inc., South Salt Lake City, UT, USA) produced in accordance with the 48 h protocol described by Daniels et al. (2010) (24 h hatch period followed by a 24 h enrichment period). Pyceze was added at 0.3 g l−1 to reduce bacterial and fungal concentration during the hatching process. Artemia were then enriched with a solution of the designated enrichment emulsion as specified in Table 1. Larvae were fed at a density of 5 Artemia ml−1, as specified by Fitzsimmons et al. (2004), equivalent to 1.25 g of dried decapsulated Artemia cysts per cone. Live Artemia and algal counts were monitored daily and replenished where necessary. 2.3. Live feed microbiology Both culture water and Artemia were sampled after the 24 h enrichment period. Culture water samples (1 ml) were taken directly from the Artemia culture vessels. Artemia samples were then collected by draining Artemia onto 50 μm mesh and rinsing with sterile seawater to remove non-adherent external bacteria prior to further analyses. 2.3.1. Culture-dependent analyses Levels of total culturable aerobic marine heterotrophic bacteria, Vibrio spp. and Bacillus spp. were analysed following the techniques employed by Daniels et al. (2010), using marine salts agar, thiosulphate citrate bile salts (TCBS) agar (Oxoid Ltd, UK) and Bacillus selective agar (agar base; 5 g l−1 special peptone, 30 g l−1 soluble starch, 5 g l−1 granulated yeast extract and 12 g l−1 bacteriological grade agar) supplemented with Polymyxin B (Oxoid Limited, Hampshire, UK) at 100,000 IU l−1, respectively. Five samples, of Artemia (n = 5 treatment−1) and Artemia culture water (n = 5 treatment−1), for each enrichment treatment were collected in sterile microcentrifuge tubes from five different batches of the enrichment. Artemia samples were weighed, homogenised, serially diluted with phosphate buffer saline (PBS) (to 10−4), and spread

2. Materials and methods 2.1. Experimental system and animals Twelve day feeding trials were conducted at The National Lobster Hatchery (NLH), Padstow, Cornwall, UK, between June and August 2009. All larvae were hatched and collected according to Daniels et al. (2010). Larvae were reared by green water technique (Browne et al., 2009), in isolated 80 l conical vessels using pre-treated seawater and subject to natural photoperiod. The culture water in the rearing vessels was circulated by a bottom feed of air, water temperature was maintained at 19 ± 1 °C using a GET Portable 12 k BTU air conditioner (model No. GPACU12HR, GET plc, Birmingham, UK) and water chemistry sustained at a salinity of 35 g l−1, pH at 8.2–8.3, NH3 b 0.1 mg l−1 and

Table 1 Formulations of diet enrichments (g l−1 Artemia culture water) added daily to the hatched Artemia. Diet

a

Easy DHA Selco™ b Sanolife® c Bio-Mos® Water a

Control

Bacillus

MOS

Bacillus + MOS

0.6 0 0 0.3

0.6 0.1 0 0.3

0.6 0 0.012 0.3

0.6 0.1 0.012 0.3

INVE Aquaculture, Belgium. INVE Aquaculture, Belgium; containing Bacillus subtilis, Bacillus licheniformis and Bacillus pumulis. c Alltech Inc., Lexington, KY. b

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(100 μl) onto the three selected media in triplicate and incubated at 18 °C for 48 h. Colony forming units (CFU) were counted post incubation and values per g or ml were calculated. 2.3.2. PCR-DGGE Three further Artemia samples (n = 3 treatment−1) were collected from separate batches of Artemia. These were then homogenised prior to DNA extraction, PCR amplification of V3 16S rRNA genes and DGGE (as described elsewhere [Merrifield et al., 2009]). Selected dominant bands were then excised and DNA was eluted in TE buffer at 4 °C overnight before re-PCR. The PCR products were cleaned (SureClean, Bioline USA Inc.) and sequenced as described elsewhere (Ringø et al., 2006). The resultant nucleotide sequences were submitted to a BLAST search in GenBank (http://blast.ncbi.nlm.nih.gov/Blast.cgi) to retrieve the closest known alignment identities for the partial 16S rRNA sequences. 2.4. Larval gastrointestinal microbiology 2.4.1. Culture-dependent analyses On 1, 7 and 12 days post hatch (dph), 15 larvae treatment−1 (i.e. three samples per dietary treatment, each containing 5 pooled larvae) were surface sterilised and rinsed as described by Daniels et al. (2010) using 0.1% benzalkonium chloride (BKC). These samples were homogenised, serially diluted, spread onto the three agar types, incubated and CFU counted, as previously described. 2.4.2. PCR-DGGE On 1 and 12 dph, 15 larvae treatment−1 (i.e., three samples per dietary treatment, each consisting of 5 pooled larvae) were surface sterilised and rinsed as previously described and placed into molecular grade 1.5 ml microcentrifuge tubes. At day 1 and 12 the three samples per treatment were stored at −20 °C before further analysis, which consisted of DNA extraction, PCR amplification, DGGE, band extraction and sequencing, as previously described. 2.5. Larval growth and survival At 1 (initial measurements, zoea I), 7 and 12 dph (final measurements, zoea III), five larvae were removed at random from each of the three replicate cones per treatment (n = 15 treatment−1), weighed and carapace length (CL) measured as described by Daniels et al. (2010). At 12 dph the numbers of zoea III in each cone were counted to calculate survival. The following variables were calculated for each replicate tank within each of the four treatments; survival (%) = (FN/IN) × 100; live weight gain (LWG) = Wt − W0; and carapace length gain (CLG) = (CLt/CL0). Where IN is initial number of larvae, FN is the final number of larvae, W0 is initial wet weight (mg), Wt is final wet weight (mg), CL0 is initial carapace length (mm), and CLt is final carapace length (mm). 2.6. Salinity stress tests At 12 dph, 120 larval lobsters were randomly removed from each treatment (n = 40 replicate−1) and subjected to a salinity challenge stress test. The stress tests were undertaken using 1 l aerated beakers (n = 10 lobsters beaker−1). Low salinity stressors were set at 15 g l−1 at 19 °C (lethal time for 50% mortality [LT50] after 1 h and lethal time for 100% mortality [LT100] after 12 h). Larvae were evaluated at 5 min intervals to record cumulative mortality over the 1 h time interval then every hour until 12 h. The larvae presenting no movement of pleopods and giving no reaction to physical stimuli were recorded as dead. The sensitivity of larvae to low salinity stress was expressed by the cumulative stress index (CSI); CSI = CM/T where CM is the cumulative mortality observed and T is the time of exposure to the stressor. The higher the value of the CSI index, the earlier and/or higher the

mortalities, thus the greater the sensitivity to osmotic stress (Dhert et al., 1992). 3. Statistical analyses To determine if there was significant differences between treatments for growth performance (Wt, LWG, CLt, CLG), survival, bacterial abundance and CSI, data were subject to one way analysis of variance (ANOVA). Kruskal–Wallis non-parametric tests were applied to nonnormal distributed data (Vibrio and Bacillus CFU counts from 12 dph larvae and Artemia samples). To determine whether significant differences in growth parameters (W, CL) and gastrointestinal bacterial counts existed between treatments over time a series of one-way repeated measures ANOVA's were conducted, the repeated measure being dph (1, 7 and 12 dph). Primer V6 was used to conduct nonmetric multidimensional scaling analysis (nMDS) on data from the DGGE to represent the relative similarities between treatments (Powell et al., 2003) and cluster analysis was employed to check the groupings by the nMDS procedure. Stress level was low (0.02–0.09 for Artemia and larval DGGEs, respectively). Similarity percentages (SIMPER) were also performed on the observed clusters from the nMDS plots and for pairwise comparisons a one way analysis of similarity (ANOSIM) was used to determine differences between DGGE banding profiles where appropriate (Abell and Bowman, 2005). One-way ANOVA was applied where applicable to the number of observed taxonomical units (OTUs), species richness, evenness, diversity, similarity and dissimilarity data to determine differences between treatments. 4. Results 4.1. Live feed microbiology 4.1.1. Culture-dependent analyses Bacillus levels within the Artemia culture water differed (P b 0.001) among treatments (Table 2). Bacillus CFU per ml of Artemia culture water after 24 h of enrichment was significantly higher than in the control for Bacillus (P b 0.001) and Bacillus + MOS (P b 0.001) treatments (log 4.4 and log 4.6 CFU ml−1, respectively). Control and MOS treatments both contained Bacillus spp. at levels which were too few to enumerate accurately (i.e., less than 20 CFU at the lowest dilution). Bacterial levels in the culture medium from the Bacillus treatment either alone or in combination with MOS showed significantly (P b 0.001 and P = 0.006, respectively) lower Vibrio levels in comparison with the control. However, Vibrio levels were not lower than the control for the MOS treatment (P N 0.05). The total viable count (TVC) was not affected (P N 0.05) by the probiotic, prebiotic or synbiotic supplementation to the live feed culture medium. Table 2 Summary of culturable bacterial levels within Artemia and Artemia culture water samples enriched in four ways. Levels (log CFU g−1 Artemia or log CFU ml−1 culture water) of total culturable viable counts (TVC), Vibrio and Bacillus. n = 5. Source

Bacterial log CFU g−1 or ml−1 group Control Bacillus

MOS

Bacillus + MOS

TVC Artemia culture Vibrio water Bacillus

6.89 ± 0.36a 7.11 ± 0.46a 6.83 ± 0.33a 7.24 ± 0.49a 5.28 ± 0.33c 2.82 ± 0.29a 4.89 ± 0.28c,b 4.00 ± 0.15b TFTCa 4.40 ± 0.11b TFTCa 4.60 ± 0.13b

Artemia

7.62 ± 0.10a 7.85 ± 0.17a 8.24 ± 0.35a 5.35 ± 0.84a 6.28 ± 0.30a 6.54 ± 0.18a TFTCa 5.77 ± 0.24b TFTCa

TVC Vibrio Bacillus

7.93 ± 0.14a 5.93 ± 0.32a 5.56 ± 0.02b

Values expressed as means ± standard error where applicable. TFTC = too few colonies to reliably enumerate (i.e. less than 20 colonies at 10−1 dilution). a,b,c Values in the same row with the different superscripts denotes significant differences (P b 0.05).

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Significant differences in Bacillus abundances were observed among the treatments for the bacterial populations in the Artemia after 24 h of enrichment (P b 0.001). Bacillus levels in Artemia were higher than that of the culture water and had accumulated levels of log 5.77 and 5.56 CFU g Artemia−1 in the Bacillus and Bacillus + MOS treatments, respectively. However, Vibrio levels in Artemia were not affected by treatment (P = 0.385). There were no differences in Artemia TVC levels among samples from the four different treatments (P N 0.05). 4.1.2. PCR-DGGE Artemia DGGE profiles were quite dissimilar from larvae DGGE profiles within experimental treatments (46–64%). The similarity between Artemia and larvae of the same treatment was highest in the Bacillus (53.86%) and decreased to 40.59% and 44.27% in the MOS and control samples, respectively, with the lowest similarity between the Bacillus + MOS Artemia and Bacillus + MOS larvae (Table 3). The similarity between stage I larval samples was extremely high, 91.8 ± 2.09%. This relationship is shown in the Bray Curtis similarity 3-dimensional scaling plot (Fig. 1). Pielou's evenness (values close to 1 represent low variation in communities between the species) shows that variation in the communities within the respective treatments was low. The OTUs, species richness and Shannon's diversity index were lowest in the Artemia samples in comparison to larval samples and this was particularly true in the MOS treatment. Not all selected DGGE bands (Table 4) could be identified to species level due to equal similarity to a number of sequences in GenBank, however, phylotypes were most closely identified as being: Lactococcus spp., Weissella spp., Chlorogloeopsis spp. and Varibaculum spp. previously isolated from marine environments including from plankton, and other aquatic environments including the gastrointestinal tracts of yellow catfish, Pelteobagrus fulvidraco. Uncultured bacteria from the orders Mycoplasmatales, Bacillales, Flavobacteriales, Actinomycetales and Rhizobiales were also identified. 4.2. Larval microbiology 4.2.1. Culture-dependent analyses Zoea I, at one dph showed no difference (P N 0.05) in larval gastric bacterial abundances among treatments (Fig. 2A) with TVC and Vibrio counts on average 5.14 ± 0.07 and 3.81 ± 0.33 log CFU g larvae−1, respectively. No significant differences (P N 0.05) were observed in TVC or Vibrio counts at 7 dph (Fig. 2B). At 12 dph TVC showed no difference among treatments (P N 0.05) (Fig. 2C). TVC did not change from 7 to 12 dph (P N 0.05). However, at 12 dph, gastric Vibrio counts in zoea III larvae were significantly lower (P b 0.005) in MOS fed larvae compared to the control larvae, however, this reduction was not observed in Bacillus or Bacillus + MOS fed larvae. Vibrio counts reduced over

399

time from 7 to 12 dph (P b 0.001). GI Bacillus levels were different between treatments (P b 0.001). Bacillus levels were below detection levels the in gut of larvae fed MOS or control diets and too few to enumerate accurately in all treatments at one dph (Fig. 2). Bacillus bacteria appeared to accumulate over time in those larvae fed Bacillus and Bacillus + MOS enriched Artemia, with levels increasing by ~1 log CFU g larvae−1 from 7 to 12 dph (P b 0.001). 4.2.2. PCR-DGGE DGGE profiles of the experimental larval groups were dissimilar to the control (33–37%) and relatively similar to each other (72–76% similarity) (Table 5). Of the experimental treatments, MOS or Bacillus fed larvae contained intestinal microbial communities most similar to control fed larvae, both displaying 33% dissimilarity to the control (Fig. 3). The ANOSIM R statistic shows that replicates within all treatments were more similar to each other compared to replicates of other treatments. The lowest value was recorded when comparing the control with MOS replicates (0.37). The highest R statistics were found when comparing Bacillus + MOS with the Bacillus group with values as high as 0.96 which was approaching significance (P = 0.10). The similarity among in-treatment replicates was highest in the Bacillus fed larvae (88.71 ± 1.10%) and lowest in the control fed larvae (72.45 ± 3.65%); this difference was approaching significance (P = 0.067) (Table 5). In larval lobsters the number of OTUs, species richness and Shannon's diversity index was generally higher in all zoea III larval groups (Table 5) than that of the zoea I larvae and Artemia samples (Table 3). In addition, the number of OTUs, species richness and diversity were significantly higher (P b 0.01) in all of the experimental groups than the control, with the highest number of OTUs (P = 0.006), richness (P = 0.007) and diversity (P = 0.009) observed in Bacillus and Bacillus + MOS fed larvae (Table 5). Selected phylotypes, identified from DGGE bands, were most closely related to Weissella spp., Bacillus spp., Lysinibacillus spp., Pseudomonas spp., Scytonema spp. and Colwellia spp., many of which were previously isolated from marine environments including marine biofilms, deep hydrothermal plumes, and other aquatic environments including the GI tract of yellow catfish. Five bands were also closely related to uncultured bacterial species from the orders Bacillales, Mycoplasmatales, Clostridiales, Sphingobacteriales and Flavobacteriales (Table 6). The bands identified as the Bacillus probionts were present only in the Bacillus and Bacillus + MOS fed larvae. 4.3. Larval growth and survival Larval weight at 1 and 7 dph was not different among treatment groups (P N 0.05). However, by 12 dph the larvae fed Bacillus, MOS and

Table 3 Summary of microbial community parameters obtained from DGGE fingerprints of microbiota from Artemia (A) and H. gammarus zoea III larvae (L) for each of four dietary treatmentsa and for the zoea I larval lobsters at the outset of the experiment. Groupa

OTUsb

Species richnessc

Evennessd

Diversitye

SIMPERf (similarity) %

Dissimilarity %

Control A Control L MOS A MOS L Bacillus A Bacillus L Bacillus + MOS A Bacillus + MOS L Day 1 larvae

11 15 8 12 9 18 12 14 13.00 ± 0.00

0.99 1.34 0.72 1.05 0.81 1.59 1.07 1.25 1.18 ± 0.01

0.994 0.995 0.992 0.991 0.993 0.993 0.991 0.990 0.984 ± 0.00

2.38 2.69 2.06 2.46 2.18 2.87 2.46 2.61 2.52 ± 0.00

44.27

55.73

40.59

59.41

53.86

46.14

35.64

64.36

91.85 ± 2.09

8.15 ± 2.09

Values expressed as means ± standard error where applicable. a Feeding groups: control, MOS, Bacillus, Bacillus + MOS. b Average number of observed taxonomical units (OTUs). c Margalef species richness: d = (S−1)/log(N). d Pielou's evenness: J’ = H’/log(S). e Shannon's diversity index: H’ = -Σ(pi(lnpi)). f SIMPER = similarity of replicates within group.

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Fig. 1. Nonmetric 3D multidimensional scaling analysis plot of DGGE fingerprints of H. gammarus larvae and Artemia showing similarities between Artemia (A) and larvae (L) of treatments (Control, MOS, Bacillus and Bacillus + MOS) and changes from initial stage I larval samples. Clustering indicates that at stage I larvae commence life with similar bacterial profiles. The effect feed has on changing these microbial community profiles is highlighted by clustering between Artemia and larval samples namely MOS and Bacillus, although Bacillus + MOS and control treatments appear to be most variable between Artemia and larval samples. All treatments cause a shift from the bacterial community structure seen in stage I larvae.

Bacillus + MOS were 26, 32 and 20% heavier, respectively, (P =0.002, P b 0.001, P = 0.02, respectively) than control fed lobsters. For LWG there was significant differences (P =0.009) among treatments, with Bacillus only or MOS only fed lobsters producing the greatest weight gains (P = 0.015, P = 0.001, respectively) at 12dph in comparison to the control. Whereas LWG was not significantly different (P = 0.165) from the control treatment in Bacillus + MOS fed lobsters. CL at 1 and 7 dph did not vary between larvae fed different treatments (P N 0.05). However, by 12 dph, CL was significantly greater in Bacillus, MOS and Bacillus + MOS fed larvae (P b 0.001, P b 0.001, P = 0.016, respectively) than control fed larvae (15, 14 and 9% larger, respectively). There was no interaction effect between treatment and time (P = 0.121, Fig. 4B). Survival rate to 12 dph (zoeal III) significantly differed among treatments (P b 0.001). Larvae fed Bacillus, MOS or Bacillus + MOS showed significantly higher levels of survival (P b 0.004) than control fed larvae, with the highest survival observed in larvae fed Bacillus enriched Artemia (35%) compared to all other treatments (vs control P b 0.001, vs MOS P b 0.001, vs Bacillus + MOS P b 0.001). Control fed larvae displayed the lowest survival rate (11%, Table 7).

4.4. Salinity stress test At the end of 1 and 12 h the CSI was significantly different (P b 0.001) among treatments (Table 8). Higher mortalities were observed for control fed lobsters in the salinity stress test than experimental treatments, as shown by the significantly lower CSI for Bacillus, MOS and Bacillus + MOS fed lobsters (P b 0.001) (Fig. 5). The lowest CSI's in comparison to all other treatments were for Bacillus fed lobsters after 1 and 12 h (both P b 0.001) of low salinity stress.

5. Discussion The use of green water systems for lobster culture is still under examination and high variability in success rates between batches has been reported (Browne et al., 2009). In the present study, major losses of larvae occurred after 12 dph (stage III) across all treatments and replicates, these crashes have previously been accredited to the accumulation of Vibrio levels (Uglem et al., 2006). This would explain the high mortalities experienced at stage III in the control cones in the present study, which had the highest Vibrio counts at 12 dph. These Vibrio levels may originate from the high Vibrio counts present in the live feed and their culture media, the latter of which, was again highest in the control. The present study shows the benefits of adding probiotics and prebiotics during the culture of live feeds on the bacterial quality of the feed and on subsequent survival and fitness of larvae. Many Vibrio spp. are known to be pathogenic to crustaceans (for review see Soto-Rodriguez et al., 2003); thus, exclusion or reduction of Vibrio from the feed source is potentially beneficial to the host species. Especially when considering that Artemia enriched with Selco™ have previously been reported to contain these potential pathogens (Hoj et al., 2009; Villamil et al., 2003). Reductions in presumptive Vibrio spp. associated with Artemia culture water by the presence of Bacillus or Bacillus + MOS were observed in the present study. Similar reductions have previously been noted with the use of probiotics in live feeds (Verschuere et al., 2000; Villamil et al., 2003). In the present study, no differences in culturable TVC were observed in relation to the enrichment of Artemia with the probiotic, prebiotic or synbiotic, which concurs with Gatesoupe (2002) who also found no difference in Artemia TVC when using probiotics. Pseudomonas spp., Aeromonas spp., Bacillus spp. and Vibrio spp. have previously been reported to dominate the bacterial assemblages associated

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Table 4 Isolated bacterial bands and their closest relatives (BLAST) from PCR-DGGE of the intestinal communities of Artemia samples and zoea I and III larval lobsters. Band number

Groups (Artemia or Larvae) and number of lanes present in Control

MOS

Bacillus

Bacillus + MOS

A

L

A

L

A

L

A

L

0 1 0 0 1 0 1 1 1

1 0 0 1 1 0 1 1 1

0 1 0 0 1 1 1 1 1

1 0 1 1 1 1 0 1 1

0 0 1 0 1 0 1 1 1

1 0 1 1 1 1 1 1 1

1 0 1 0 1 0 0 1 1

1 0 0 1 1 1 1 1 1

3 0 0 0 3 3 3 3 3

18

0

0

1

0

1

0

1

0

0

b

Similarity to nearest neighbour

Accession number of nearest neighbour

Uncultured bacterium clone SC57 Uncultured bacterium clone ncd960h01c1 Lactococcus sp. TP2MJ Uncultured bacterium clone WLB13-202 Weissella cibaria strain MGD4-4 Chlorogloeopsis fritschii PCC 6718 Uncultured bacterium clone ncd307e06cl Uncultured bacterium clone FRC-AI_667 Uncultured Varibaculum sp. Clone W5_maj-078 Uncultured bacterium clone TI-154

93% 98% 91% 91% 98% 94% 95% 88% 88%

GU293212b HM331027 GU272382b DQ015842b HM058481 AF132777 HM316521 DQ015842b DQ976120

96%

FN821846b

Day 1 larvaea

6 10 12 17 4 20 21 24 25

a

Nearest neighbour

Number of replicate lanes present in. Previously isolated from an aquatic based source.

with Artemia (Kennedy et al., 2006). However, the present study identified Lactococcus spp., Weissella spp., Chlorogloeopsis spp. and many uncultured bacterial strains closely related to bacteria from the orders

Fig. 2. Gastric bacterial levels (total culturable viable, Vibrio and Bacillus counts in log — CFU g larvae−1) from larval H. gammarus, A) 1 dph (zoea I), B) 7 dph (zoea II), C) 11 dph (zoea III), fed live Artemia diets supplemented with Bacillus and/or MOS or without supplementation (control). Different letters from the same bacterial classification (e.g. total, Vibrio or Bacillus) indicated significant differences between dietary treatments (Control, Bacillus, MOS or Bacillus + MOS) (P b 0.05) values represent mean ± standard error.

Mycoplasmatales, Bacillales, Flavobacteriales, and Rhizobiales, as well as Varibaculum spp., which indicate that the microbial communities of Artemia cultures may be more complex and diverse than previously believed. Bacillus levels established within Artemia samples in the present study are presumed to be those accumulated in the gut and/or on the surface of the organism. Elevated Bacillus levels found in larvae fed Bacillus and Bacillus + MOS diets are likely to have been introduced from the feed. When comparing bacterial levels in the Artemia to that obtained in the larvae at 12 dph (i.e. translocation from live feed into the digestive tract of larvae) all culturable bacterial levels were less than that in the Artemia. The only exception was the Vibrio levels in control fed larvae which increased by 1.85 CFU g−1. This indicates the accumulation of Vibrio levels in the gut of larvae. PCR-DGGE analysis revealed increased number of OTUs, species richness and diversity in the larvae compared to their Artemia feed, irrespective of the experimental treatments. It has been reported that the bacterial community structure of live prey are only partly reflected in the host (Bjornsdottir et al., 2010). This was also the case in the present study when comparing Artemia with 12 dph H. gammarus larvae; bacteria commonly found across treatments and sample types were Weissella spp. (order Lactobacillales), Lactococcus spp. (order Lactobacillales), Chlorogloeopsis spp. (order Stigonematales) and uncultured bacterial clones from the orders Mycoplasmatales, Actinomycetales and Flavobacteriales. However, many of the OTUs observed by DGGE were not successfully identified and most species were found inconsistently in both Artemia and larvae from the same treatment. Culture-based analysis of bacterial communities in larval samples showed that newly-hatched larvae had low gastric bacterial levels which increased as the larvae developed. This is in agreement with studies from other aquatic species (Ringø and Birkbeck, 1999; Skjermo et al., 2006). By 7 dph bacterial levels similar to those from the feed were obtained from larvae, however, it was only after 12 dph that significant differences between the treatments were observed. This observation is in accordance with that of Qi et al. (2009) who reported that probionts were unable to influence microbial community composition associated with cultured rotifers after short periods of feeding (3 days). In the present study, reduced Vibrio levels were found in zoea III (12 dph) larvae fed MOS supplemented Artemia in comparison to the control. Similar findings were reported with the application of dietary MOS in rainbow trout, Oncorhynchus mykiss (Dimitroglou et al., 2009). Despite apparent trends for decreased larval gastric presumptive Vibrio levels in the present study, not all treatments produced significant reductions. As with bacterial abundances in Artemia, no change in gastric culturable TVC was observed between treatments.

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Table 5 Summary of microbial community parameters and pairwise comparisons obtained from DGGE fingerprints of gut microbiota from zoea III larval lobsters of each dietary group a. ANOSIMg Group a Control MOS Bacillus Bacillus + MOS Significance

OTUsb

Species richnessc a

13.00 ± 1.15 14.00 ± 0.88b 15.33 ± 0.33c 16.33 ± 1.15b,c P = 0.006

a

1.18 ± 0.10 1.25 ± 0.08b 1.37 ± 0.03c 1.46 ± 0.10b,c P = 0.007

Evennessd

Diversitye a

0.981 ± 0.00 0.978 ± 0.00a 0.978 ± 0.00a 0.979 ± 0.00a P = 0.169

SIMPERf (similarity) % a

2.51 ± 0.09 2.57 ± 0.05b 2.67 ± 0.01c 2.73 ± 0.07b,c P = 0.009

R-value

P-value

a

72.45 ± 3.65 75.02 ± 6.68a 88.71 ± 1.10a 86.47 ± 3.82a P = 0.067

Pairwise comparisons Control v MOS Control v Bacillus Control v Bacillus + MOS MOS v Bacillus MOS v Bacillus + MOS Bacillus v Bacillus + MOS

Dissimilarity % 27.55 ± 3.35 24.98 ± 6.68 11.29 ± 1.10 13.53 ± 3.82 P = 0.067

0.37 0.59 0.82 0.52 0.48 0.96

0.20 0.10 0.10 0.10 0.10 0.10

33.16 32.77 36.54 24.01 26.90 27.98

± ± ± ± ± ±

2.97 3.72 2.54 3.65 1.59 2.65

Values in the same column with different subscripts denotes significant differences. Values expressed as means ± standard error. a Feeding groups: control, MOS, Bacillus, Bacillus + MOS. b Average number of observed taxonomical units. c Margalef species richness: d = (S-1)/log(N). d Pielou's evenness: J’ = H’/log(S). e Shannons diversity index: H’ = -Σ(pi(lnpi)). f SIMPER = similarity of replicates within group. g Analysis of similarity (ANOSIM).

This lack in modification of culturable bacterial levels has also been reported in early stages of fish fed probiotics (Gatesoupe, 2002; Makridis et al., 2001). Bacillus probionts were detected in Bacillus and Bacillus + MOS fed larvae but were not detected in the control or MOS fed larvae using culture-dependent or PCR-DGGE analyses. An uncultured Bacillus spp. was also detected by PCR-DGGE in Bacillus and control fed larvae, though Bacillus spp. have previously been documented as normal components of the gut microbiota of lobsters (Battison et al., 2008). Vibrio spp. and Pseudomonas spp. are thought to be the most abundant genera in the digestive tract of crustaceans (Moriarty, 1990) and in the present study Pseudomonas spp., Weissella spp., Bacillus spp. Lysinibacillus spp., Scytonema spp. and Colwellia spp. were also found to be among the dominant bacterial genera. DGGE revealed that microbial species

richness and diversity was significantly increased in Bacillus, MOS and Bacillus + MOS fed zoea III larval lobsters in comparison to the control, with the largest increases in Bacillus fed larvae. Subsequently the microbial profiles of the experimental groups were dissimilar to the control group; with Bacillus + MOS showing the greatest dissimilarity (36.54 ± 2.54%) to the control. The present study demonstrates the benefits of supplementing larval live feeds with probiotic, prebiotic and synbiotic on the growth of green water cultured H. gammarus which is similar to the findings reported by Daniels et al. (2010) in lobsters reared in re-circulatory systems. Although no additive relationship (i.e., greater improvements beyond the individual applications) was found on growth parameters in the present study with the application of Bacillus + MOS, which was in contrast to the findings of Daniels et al. (2010). Larval growth

Fig. 3. 2D Nonmetric multidimensional scaling analysis plot of DGGE fingerprints of bacteria from the gut of H. gammarus larvae showing similarities among treatments; Control, MOS, Bacillus, and Bacillus + MOS. Clustering indicates that biotic supplementation of larval diets causes a shift in the bacterial community structure from the control, with Bacillus tending to decrease variability bacterial community structure. MOS supplemented lobsters are not as closely clustered, but MOS supplementation appears to cause a shift toward the Bacillus and Bacillus + MOS larval groups.

C.L. Daniels et al. / Aquaculture 416–417 (2013) 396–406

403

Table 6 Isolated bacterial bands and their closest relatives (BLAST) from PCR-DGGE of the intestinal communities of stage III larval lobsters. Band number

3 5 7 6 8 9 19 17 27 4 20 18 a

Groups and number of lanes present in Control

MOS

Bacillus

Bacillus + MOS

3 1 3 3 1 3 3 3 3 2 3 0

3 0 3 3 0 3 3 3 3 1 3 2

3 1 3 3 2 3 3 3 3 3 3 3

3 0 3 3 0 3 3 3 3 1 3 3

Nearest neighbour

Similarity to nearest neighbour

Accession number of nearest neighbour

Weissella confusa Uncultured Bacillus spp. Uncultured bacterium clone SC57 Uncultured bacterium clone SC57 Lysinibacillus spp. P-001 Uncultured bacterium Uncultured bacterium clone 80 Uncultured bacterium clone 1E-059 Pseudomonas sp. INBio2926B Weissella cibaria strain MGD4-4 Scytonema sp. R01 Colwellia sp. ‘Disko Bay’

99% 90% 93% 92% 94% 88% 96% 94% 95% 99% 87% 88%

AB494723 FJ227518 GU293212a GU293212a GU288531a GQ351438 GU066450a FJ980811a GU827529 HM058481 EU818963 FJ581925

Previously isolated from an aquatic based source.

Fig. 4. Growth measurements of larval H. gammarus fed with and without the addition of probiotics, prebiotics or synbiotics to the diet over a 12 day feeding trial. A) wet weight, B) carapace length (CL). Data represents mean ± SEM. Different letters represent significant differences between treatments (P b 0.05).

in all treatments in the present study was greatly improved compared to the control. Similar improvements in growth have previously been reported when supplementing Bacillus probionts at: 0.3 g kg−1 in black tiger shrimp (Penaeus monodon) (Rengpipat et al., 2003); at 1 and 5 g kg−1 in shrimp (Penaeus vannamei) (Wang, 2007); and at various concentrations in white shrimp (Litopenaeus vannamei) (Liu et al., 2009). The dietary application of MOS has also been reported to improve growth of green tiger prawn (Penaeus semisulcatus) when fed at 3 g kg−1 (Genc et al., 2007). Improved daily weight gain (DWG) and final body weight, as observed in the present study, is in agreement with previous studies on shrimp reported by Wang (2007) and Liu et al. (2009) with the dietary addition of different concentrations of probiotics. The present study demonstrates that the dietary probiotic, prebiotic and synbiotic application had beneficial effects on the survival of lobster larvae with increases of ~24% with the probiotic treatment, ~10% with the prebiotic treatment and ~6% with the synbiotic treatment. Similar findings have been reported in other crustacean species including shrimp, L. vannamei, L. stylirostris and prawn, P. monodon with the dietary supplementation of SANOLIFE® and other Bacillus spp. (Decamp and Moriarty, 2007). Increased survival of ~21% has also been reported in green tiger prawn fed MOS dietary supplementation (Genc et al., 2007). Many studies use stress challenges to relay information on an animal's ability to tolerate change as an indication of fitness or the quality of cultured individuals (Dhert et al., 1992; Palacios and Racotta, 2005; Salze et al., 2008; Taoka et al., 2006) and salinity stress tests have been used as a final indicator of animal quality at the end of nutritional/experimental studies (Palacios and Racotta, 2005; Salze et al., 2008; Smith et al., 2004; Taoka et al., 2006). In the present study, larvae fed the experimental diets appeared more capable of withstanding hyposaline changes than control fed larvae, with the best tolerance shown in the Bacillus group. Similar improvements in hyposaline stress tolerance were seen in larval cobia (Salze et al., 2008) and white sea bream (Diplodus sargus) (Dimitroglou et al., 2010) fed with MOS. Dietary changes can affect nutritional reserves which in turn can affect the tolerance of an organism to a salinity stress (Palacios and Racotta, 2007). Carbohydrates and other nutrient reserves such as proteins and fatty acids can be used in osmoregulation therefore larvae with better reserves are more likely to survive in low salinities (Palacios and Racotta, 2007). Bacteria are also known to offer an extra source of nutrients for the live feed (Gorospe et al, 1996; Marques et al., 2005), thus in the present study the feed additive enrichments of Artemia may have acted as either nutrient sources for Artemia and/or caused improved feed conversion (via the production or stimulation of enzymes which aid digestion (Castex et al., 2008; Ghosh et al., 2008; Zhou et al., 2009) and/or allowing for improved GI structure (Verschuere et al., 2000;

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Table 7 Growth response and survival of H. gammarus larvae fed on diets supplemented with and without probiotics, prebiotics or synbiotics until metamorphosis to zoeal stage III. Control Initial weight (mg) Final weight (mg) Live weight gain (mg) Initial CL (mm) Final CL (mm) CL gain (mm) Survival (%)

12.53 32.1 19.6 2.98 4.21 1.22 11.39

± ± ± ± ± ± ±

Bacillus 0.48a 3.04a 3.34a 0.12a 0.15a 0.12a 1.18a

13.47 40.6 27.13 3.18 4.82 1.64 34.88

± ± ± ± ± ± ±

MOS 0.46a 1.00b 1.21b 0.18a 0.06b 0.12a 2.68c

12.80 42.5 29.67 3.30 4.79 1.49 20.60

Bacillus + MOS ± ± ± ± ± ± ±

0.66a 0.98b 1.42b 0.18a 0.09b 1.14a 0.68b

14.6 38.4 23.80 3.11 4.59 1.42 16.54

± ± ± ± ± ± ±

0.62a 1.56b 1.79a,b 0.15a 0.12b 0.16a 1.11a,b

Significance P P P P P P P

= 0.053 b 0.001 b 0.01 = 0.167 b 0.001 = 0.210 b 0.001

Data represented as mean ± SE mean. Data subject to one way ANOVA. Mean values in rows with different superscripts are significantly different (P b 0.05).

Table 8 Cumulative sensitivity index (CSI) at 1 and 12 h, for 12 dph larval H. gammarus subject to low salinity stress. Lobster larvae pre-fed live Artemia diets supplemented with Bacillus and/or MOS or without supplementation (control). Time (h)

Measure

Control

Bacillus

MOS

Bacillus + MOS

Significance

1 12

CSI CSI

42.75 ± 0.63a 131.00 ± 1.47a

16.00 ± 0.91d 77.50 ± 2.75d

19.25 ± 1.18c 85.50 ± 2.66c

23.00 ± 0.41b 99.25 ± 2.56b

P b 0.001 P b 0.001

Data represented as mean ± SE mean. Data subject to one way ANOVA. Mean values in rows with different superscripts are significantly different (P b 0.05).

Daniels et al., 2010) thus improving feed quality for the larvae. Therefore, improvements in the nutritional quality of feed or host feed conversion may explain the improved tolerance to hyposalinity observed in probiotic, prebiotic and synbiotic fed larvae. Improved tolerance to stress has also been accredited to probiotic supplements mediating stress responses which can cause immune suppression following stress; for example, probiotics can reduce cortisol production (Avella et al., 2010) and heat shock protein (HSP70) production and gene expression (Avella et al., 2010; Rollo et al., 2006). 6. Conclusion The use of the green water technique for rearing larvae H. gammarus proved beneficial for analysing the application of feed additives. This study suggests that the dietary supplementation of prebiotics, probiotics and synbiotics to larval H. gammarus reared with microalgae

improves growth, survival and microbial parameters. The individual supplementation of Bacillus to diets produced greater enhancements in resistance to stress, gastrointestinal bacterial communities and survival in comparison to all other treatments.

Acknowledgements The authors would like to thank G. Scofield for all her hard work and commitment towards the practical research and D. Boothroyd, C. Wells and C. Ellis of the National Lobster Hatchery, Cornwall, UK for their critical assistance throughout these experiments. We also thank Plymouth University staff for their technical assistance. This work was funded by the Great Western Research Fund and the National Lobster Hatchery with additional financial from the University of Plymouth and the Worshipful Company of Fishmongers. We kindly thank INVE

Fig. 5. Cumulative sensitivity stress index (CSI) for 12 dph H. gammarus larvae subject to low salinity stress over 12 h. Larval lobsters were pre fed live Artemia diets supplemented with Bacillus and/or MOS or without supplementation (control). Data represented as mean ± SE mean. Different letters represent significant differences between treatments (P b 0.05).

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