Gut microbial communities associated with the molting stages of the giant freshwater prawn Macrobrachium rosenbergii

    Gut microbial communities associated with the molting stages of the giant freshwater prawn Macrobrachium rosenbergii Eleni Mente, Andrew T. Gannon, Eleni Nikouli, Hugh Hammer, Konstantinos A. Kormas PII: DOI: Reference:

S0044-8486(16)30302-7 doi: 10.1016/j.aquaculture.2016.05.045 AQUA 632173

To appear in:

Aquaculture

Received date: Revised date: Accepted date:

4 January 2016 15 May 2016 31 May 2016

Please cite this article as: Mente, Eleni, Gannon, Andrew T., Nikouli, Eleni, Hammer, Hugh, Kormas, Konstantinos A., Gut microbial communities associated with the molting stages of the giant freshwater prawn Macrobrachium rosenbergii, Aquaculture (2016), doi: 10.1016/j.aquaculture.2016.05.045

This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

ACCEPTED MANUSCRIPT 1

Gut microbial communities associated with the molting stages of the giant freshwater

PT

prawn Macrobrachium rosenbergii

SC

a

RI

Eleni Mentea*, Andrew T. Gannonb, Eleni Nikoulia, Hugh Hammerc, Konstantinos A. Kormasa

Department of Ichthyology and Aquatic Environment, School of Agricultural Sciences,

Department of Biology, Birmingham-Southern College, 900 Arkadelphia Road, Birmingham,

MA

b

NU

University of Thessaly, 384 46 Volos, Greece

c

TE

D

AL 35254, USA

Aquaculture Education and Development Center, Gadsden State Community College, 1001

AC CE P

George Wallace Drive, Gadsden, AL 35903 USA

*Corresponding author: Dr. E. Mente, Department of Ichthyology and Aquatic Environment, School of Agricultural Sciences, University of Thessaly, 384 46 Volos, Greece. Tel: +302421093176, Fax: +302421093157 E-mail: [email protected] and [email protected]

ACCEPTED MANUSCRIPT 2

Abstract The reciprocal interaction between host organisms’ physiology and their gut microorganism

PT

community is of great interest in aquatic animal biology and aquaculture but for crustaceans, it

RI

remains understudied. This study enhances our understanding of this community of

SC

microorganisms as it changes during the molt cycle. Because crustaceans shed a major component of their gut, and the associated microbiome, with each molt this adds a level of

NU

complexity heretofore unexamined. We have identified the bacterial communities that are

MA

affected by the changing gut environment and that may in turn, exert some control over aspects of the molt cycle. We investigated the structural changes of the resident gut bacterial

D

communities, using the diversity of the 16S rRNA gene by 454 pyrosequencing, in the freshwater

TE

prawn Macrobrachium rosenbergii during its four-stage molt cycle. The number of bacterial

AC CE P

operational taxonomic units (OTUs) increased from stages A to C. Stage C the intermolt and longest lasting stage was different in the gut bacterial community structure having the (a) highest number of total OTUs, (b) highest number of unique and newly introduced OTUs, (c) highest percentage of estimated specialists OTUs, i.e. that are more ecologically restricted. Moreover, stage C was characterized by greater contribution of Actinobacteria-related and unaffiliated OTUs. The most dominant OTUs found in stage C of the gut of M. rosenbergii were related to microorganisms involved in fermentation and food material processing originating from similar, i.e. gut, or habitats of terrestrial and freshwater animals. Thus, the distinct gut bacterial communities found in molting stage C corroborate with the physiological significance of this molting stage. The abiotic factors and the exact role of the corresponding specific bacterial communities in the animal’s physiology and growth are areas that remain to be elucidated. Keywords: prawn, molting, gut, microbial.

ACCEPTED MANUSCRIPT 3

Introduction

PT

Giant freshwater prawn Macrobrachium rosenbergii de Man, (1879) is commercially

RI

cultured in inland aquaculture in India, China, Taiwan, Bangladesh, Vietnam, Thailand and in South America (New and Nair, 2012). It is one of the most important cultured crustacean species

SC

being farmed to promote rural livelihood and contribute to food security. Freshwater prawns are

NU

suitable candidates for inclusion in polyculture and integrated crop production systems providing the opportunity for fish farmers to increase their production and also promote sustainable rural

MA

aquaculture (New et al., 2009, FAO, 2013). Prawn farming is preferable to marine shrimp farming in inland areas because saline water is not required in the grow-out phase, once prawns

D

have metamorphosed from the larval stages, therefore there is no environmental concern for

TE

making the agriculture land saline which might affect the suitability of these areas for farming.

AC CE P

Mating and spawning can easily be achieved in captivity and management is less labour intensive (FAO, 2013). Furthermore, it is a product that can diversify crustacean aquaculture from marine shrimp due to its own culinary characteristics and new market trends. The culture of Macrobrachium can supply a good quality product to the consumer by using socially and environmentally acceptable aquaculture practices. In total, the output of M. rosenbergii from aquaculture expanded during the decade 19932002 from 17 000 tonnes to 195 000 tonnes (New et al., 2009). In 2004, the total world farmed M. rosenbergii volume reached more than 194,000 tons and in 2013 it was 203,299 (FAO, 2013). Research on growth performance and selective breeding was suggested and initiated to help the industry to increase productivity (Aflato et al., 2012). Productivity stagnation was attributed to declines in growth performance of the species due to repeated culture of unimproved stocks, thus somatic tissue growth information about the species became one of the greatest challenge for

ACCEPTED MANUSCRIPT 4

future research directed towards understanding better the physiological events of its molt cycle which is essential for growth and reproduction. A thorough understanding of M. rosenbergii

PT

changes in biochemistry and physiology during molting is essential to achieve better growth

RI

performance when cultured. Moreover, research on the microbial communities in the gut of prawns together with a demonstration of their relevance in terms of physiological implications is

SC

of great importance. By comparing different molt stages where we know that changes in the

NU

biochemistry and physiology during molt occur, we compare the responsive change of the gut microorganism communities in relation to the molt stage. Such information will be useful in

MA

order to design the best strategy to manipulate these communities in order to optimize practices to determine the physiological comfort zone for prawns and to increase their growth, health and

D

farmed production.

TE

The crustacean exoskeleton is composed of the polysaccharide chitin, structural proteins

AC CE P

and mineral deposits and its construction is an energy demanding process. The relative substrates and metabolic pathways for this are not fully understood and involvement of microbes cannot be ruled out. All freshwater prawns (like other crustaceans) have to regularly cast their ‘exoskeleton’ or shell to enable growth. This process is referred to as “molting” and is accompanied by a sudden increase in size and weight (Hartnoll, 2001). Crustacean molting is a complex process involving many regulatory pathways. Knowledge of the progression of the molting cycle stage is highly important for the understanding of various aspects of crustacean biology, including physiology and biochemistry (Hartnoll, 2001; Hayd et al. 2008). Drach, (1939) and Drach and Tehernigovtzeff (1967) classified the decapod moult cycle. Peebles, (1977) defined the molt cycle in prawns and provided a framework of terms by which to describe the various stages. There are two post-molt stages A or B, an intermolt stage C, and D is the premolt phase of the cycle. During post-molt, stages A and B, the new exoskeleton is gradually hardened by

ACCEPTED MANUSCRIPT 5

sclerotization and calcification and there is a gradual replacement of the absorbed water by tissues. This phase is followed by the intermolt stage; stage C during which the animal

PT

accumulates resources. Stage D, proecdysis, the premolt stage is the period preceding the molt

RI

event, is characterized by withdrawal of the underlying epidermis from the old cuticle, digestion and demineralization of the existing cuticular chitin and proteins and development of a new

SC

cuticle layer. Ecdysis, the molting event, is driven by active absorption of water to increase the

NU

body’s volume, leading to rupture of the partially degraded old cuticle and stretching of the new

1979) is used to determine molt stage.

MA

soft one, enveloping the now larger individual. In M. rosenbergii an index by Peebles (1977;

The structure and activity of gut microbiome as well as the effects on the host itself are

D

useful to support intestine microbial communities and maintain microbial balance with the

TE

surrounding environment. Additionally, microbial communities can be provided via the diet or

AC CE P

the rearing water to the prawn to improve growth, health and welfare of the farmed prawn. It can be a tool implemented by farmers as a best management practice in prawn aquaculture. A limited number of studies have assessed the gut microorganisms of crustaceans (Merrifield and Ringo, 2014). Even less is known about prawns, but Kennedy et al. (2006) did find in M. rosenbergii larvae a small proportion of lactobacilli, accounting for approximately 4.5% of the culturable microbiota. Tzeng et al. (2015) found proteobacteria as the major phylum in oriental river prawn Macrobrachium nipponense. Further information could be useful for developing effective strategies to manipulate gut microbial communities to promote animal growth and health and improve aquaculture productivity. However, to achieve this goal, factors that influence microbiota composition should be topics for further research. To our knowledge this is the first study that investigates the dynamics of the gut microbe community during the prawn molt cycle. Molting is a complex growth process in crustaceans that is affected by a range of external and

ACCEPTED MANUSCRIPT 6

internal factors including temperature, photoperiod, and nutritional state and eyestalk integrity. This is a step towards understanding the interactions between the gut microbiota and the host

PT

physiology and metabolism as they impact and are affected by the molt cycle. The implication of

RI

gut microbiota on the molt cycle in prawns has not been considered so far although molting causes an extreme change in the physiological programming of the animal and metabolism and

SC

growth in crustacean depends on the molt stage. M. rosenbergii was used as a species to study

NU

molting as it is an important freshwater farmed species and there is an increasing need to enhance its growth performance and improve aquaculture productivity to support the growing demand for

MA

prawns.

This study investigates the structural changes of gut microbial communities throughout

D

different molt stages of M. rosenbergii individuals by comparing the 16S rRNA gene diversity by

TE

454 pyrosequencing in order to reveal the dominant bacteria and their ecophysiological role

AC CE P

during molting with potential impact on developing conditions that prompt ecdysis and growth of the prawn. Originally we hypothesized that since the prawns habitat and presumably diet, was stable there would be little change in the gut microbiome community between molt stages. By comparing different molting stages, the dominant bacterial taxa were identified in the gut microflora with potential involvement in the molting process, including those being independent from the molting process and those being unique to each molting stage. This has enabled the assessment of gut microbial communities that play a role in the molt cycle of crustaceans.

Material and methods

Animals rearing and sampling. Day 30 postlarval, freshwater prawns, Macrobrachium

ACCEPTED MANUSCRIPT 7

rosenbergii, were obtained from a commercial hatchery (Aquaculture of Texas, Weatherford, TX, USA) in early June 2013 and stocked in a 0.10 hectare outdoor freshwater pond, at a 20-24000

PT

prawns per acre that qualifies a semi-intensive production system, at the Gadsden-State

RI

Community College Aquaculture facility, Gadsden AL, for grow-out. The prawns were raised in outdoor ponds from postlarva to adult at the growing phase. They were fed a commercially-

SC

formulated food (Sinking Catfish pellets), with a 32% crude protein which is greater than their

NU

nutritional protein requirement of the 20% crude protein (D’Abramo). However, in grow-out ponds they are thought to feed as much or more on naturally occurring pond organisms whose

MA

population growth is stimulated by the feed provided (New et al. 2009). After 120 days, on October 2013, the full-grown, adult prawns (25 – 75 g) were harvested by draining the pond and

D

hand collection. A seasonal drop in pond water temperature to 21.5°C determined harvest time.

TE

After harvest they were held overnight in an indoor facility with a recirculating freshwater system

AC CE P

without any food provided before they were sorted into molt stage A (early stage A), B, C, or D (late stage D) by physical characteristics and microscopic examination of a clipped piece of uropod using staging characteristics for M. rosenbergii from Peebles (1977). During their molt they do not eat their shed exoskeleton, unlike crayfish, which usually do. For near full grown adults, stage A lasts for about 2 days, stage B lasts about 2 days, stage C is the most variable, lasting 29-79 days (New et al. 2009) and stage D lasts 5-7 days, in total from stage A to D usually takes 38 days. Later the same day, staged prawns were transported to Birmingham-Southern College in Birmingham, AL for sterile removal of the midgut and hindgut, hereafter mentioned as gut. Prawns were weighed and morphometric data (carapace width at widest point, abdominal width at widest point, carapace length from caudal end to tip of rostrum, and total length from uropod to rostrum) was measured with calipers. Prawns were subjected to cryoanesthesia by being

ACCEPTED MANUSCRIPT 8

placed in a freezer (0°C) for 5 – 10 min before the gut was removed using sterile technique. The animals were dissected using sterile lancets and the midgut was extracted using

PT

sterile forceps. The midgut started where the digestive gland joins the main tract and ended

RI

before the hindgut. All dissecting tools were alcohol flame sterilized between each individual sample. Gut contents were emptied since we were not interested in revealing the exhaustive

SC

bacterial diversity of the gut, i.e. food particles and associated bacteria+peritrophic

NU

membrane+gut tissue epi- and endobionts, but rather in the “true” gut symbionts, i.e. epi- and endobionts of gut cell tissue (enterocytes), which are the most likely to have a more constant role

MA

in nutrient assimilation. In order to exclude as many as possible transient bacterial cells ingested with food particles, the extracted midgut was emptied into sterile diH2O by applying gently

D

mechanical force to evacuate it and by rinsing it in distilled water in the pedri dish. Evacuated

TE

guts were immediately placed in 1.5 ml. Eppendorf snap cap vials on dry ice and then stored in a

AC CE P

-80°C freezer until further analysis. Animals were selected from a larger harvest, with relatively equal representation of each molt stage. They were sampled according to the molt stage and the number of sampled animals was 21 (Table 1). The males were a mix of blue claw, orange claw, and small male. The females were smaller than the males (Table 1) although there was some variance and overlap between the sexes as has been demonstrated in numerous studies cited in Karplus and Sagi, 2010. Molecular analysis. Bulk DNA from individual gut samples (ca. 0.2 - 0.5 g) was extracted using the PowerSoil DNA isolation kit (MoBio Laboratories, USA) according to the manufacturer’s protocol. DNA concentrations ranged from 4.7 – 17.8 ng μl-1. DNA was amplified with the bacterial primers S-DBact-0341-b-S-17 (5’-CCTACGGGNGGCWGCAG-3’) and S-D-Bact-0785-a-A-21 (5’-GACTACHVGGGTATCTAATCC-3’) (Klindworth et al. 2012)

ACCEPTED MANUSCRIPT 9

targeting the V3 – V4 regions of the 16S rRNA (Claesson et al. 2010) gene at the sequencing facilities of MRDNA Ltd. (Shallowater, TX, USA). Amplicon pyrosequencing (bTEFAP) was

PT

performed as described in Dowd et al. (2008). In brief, a one-step 30 cycle PCR was applied using HotStarTaq Plus Master Mix Kit (Qiagen, Valencia, CA). PCR conditions included: 94°C

RI

for 3 minutes, then followed by 28 cycles of 94°C for 30 seconds; 53°C for 40 seconds and 72°C

SC

for 1 minute; and a final elongation step at 72°C for 5 minutes. Following PCR, all amplicon

NU

products from different samples were mixed in equal concentrations and purified using Agencourt Ampure beads (Agencourt Bioscience Corporation, MA, USA). Samples were

MA

sequenced utilizing Roche 454 FLX titanium instruments and reagents after following manufacturer’s guidelines.

D

Data processing and analysis. Samples were analysed from individuals and processed

TE

with the average OTUs abundance per molting stage. Sequencing data were analysed using the

AC CE P

MOTHUR 1.28.0 software (Schloss et al. 2009). In brief, flowgrams from the individual samples were separated according to their TAG and then were denoised using PyroNoise software (Quince et al. 2009). After removing primer sequences, TAG, and key fragments, only sequences with ≥200 bp long with homopolymers shorter than 8bp were considered for further analysis. Chimeric sequences were recognised and removed using the UCHIME software (Edgar 2010). A 97% similarity cut-off limit was used for clustering the remaining sequences. Singletons, i.e. sequences that occurred only once in the whole dataset, were removed from downstream analyses, as considered most likely sequencing artifacts (Kunin et al. 2010). The data were standardized based on the smallest sample with permutation in order to minimize differences in sequencing depths between samples. Taxonomic affiliation was assigned according to the SILVA 111 SSU RNA database (Pruesse et al. 2007). The batch of sequences from this study has been submitted to the Short Reads Archive (http://www.ncbi.nlm.nih.gov/sra) with accession number

ACCEPTED MANUSCRIPT 10

SRR1502207. We used Levins’ niche width (B) index in order to calculate habitat, i.e. molting stage,

RI

PT

specialization according to Pandit et al. (2009):

SC

where pij is the proportion of OTU j in molting stage i, and N is the total number of molting

NU

stages. Index B is irrelevant to spatial locale and abiotic conditions of a specific community, describing the extent of habitat specialization based on the distribution of species abundances in

MA

any meta-community. Based on this, OTUs with higher B scores are assigned as habitat generalists, while these with lower B scores are considered to have unevenly distributed

D

abundances among the four molting stages and are considered as specialists. According to the

TE

above equation, the lowest possible B value is 1. For this paper, and according to the values we

AC CE P

found, we set the following arbitrary categories: generalists, G (B>2.001), intermediate specialists, I, (2.0001.501) and specialists, S (1.5001.000).

Results

No statistically significant differences (p>0.05) were found between the four molt stages for carapace lengths and widths of Macrobrachium rosenbergii. However, there was a significant wet weight loss during molting from stage A to stage C (p<0.05) (Table 1). Weight had fallen at the time of molting due to the loss of the cuticle and some loss of water that is not replaced because the animal is not feeding. The OTUs communities were significantly affected by the molting stage. Tukey comparisons indicated that the OTU profiles were similar between molting stages A, B and D. However, molting stage C had significantly different OTUs communities

ACCEPTED MANUSCRIPT 11

compared to the three other molting stages (P= 0.0002). Pearson’s r ranged from 0.53-0.57, thus, there is a relationship between the molting stages A, B and D but not a strong correlation with

PT

any changes in the molting stages of A, B and D.

RI

In total, 129 unique bacterial OTUs were found in all four molt stages. The highest number of OTUs occurred in stage C (93 OTUs) and the lowest in stage A (29 OTUs) (Table 2).

SC

In each molting stage, the most dominant OTU consisted of between 34.8% (Stage C) and 63.6%

NU

(Stage B) of relative abundance, while the number of OTUs that consisted ≥90% of relative abundance ranged between 6 (Stage B) and 9 (Stage C) (Table 2). The majority of the dominant

MA

OTUs were shared among the molt stages, however their relative abundance varied among the stages (Fig. 1). Most of the dominant OTUs were affiliated to the γ-Proteobacteria, followed by

D

Firmicutes, while one dominant OUT was assigned to each of Tenericutes, Fusobacteria, β-

TE

Proteobacteria and one unaffiliated group (Table S1).

AC CE P

The four molting stages shared 13 OTUs (Fig. 2), with seven of them being among the most dominant ones (Fig. 1). The number of unique OTUs was 7, 10, 56 and 13 for molting stages A, B, C and D, respectively (Fig. 2). A similar increase in the number of novel OTUs per molting stage was also observed from stage A to C, followed by a decline in novel OTUs in stage D (Fig. 3). As most of the shared OTUs were among the most abundant OTUs, these OTUs belonged to the γ-Proteobacteria, Firmicutes, Tenericutes, Actinobacteria and one unaffiliated group (Table S1). The phylogenetic affiliation of the unique OTUs in each molting stage was different from that of the most abundant and shared OTUs (Table S1, Fig. S2). Molting stage A unique OTUs were related to the γ-, α-Proteobacteria and three unaffiliated groups, while B had more of its unique OTUs falling into unaffiliated groups followed by Cyanobacteria, γ-Proteobacteria and Bacteroidetes. Molting stage C had the highest (56) number of unique OTUs. More than half of

ACCEPTED MANUSCRIPT 12

these OTUs were associated with the intestinal tract or had symbiotic associations with various animals, while several of the OTUs were related to habitats with high organic matter degradation

PT

processes. Most of these OTUs (24) could not be affiliated to any of the known to date major bacterial phyla, while the most abundant group was the Actinobacteria (12 OTUs) followed by

RI

only a few OTUs belonging to the Firmicutes, α-, β-, γ- δ-Proteobacteria, Bacteroidetes and

SC

Cyanobacteria. Finally, molting stage D had only 13 unique OTUs belonging to the α-, β-, γ-

NU

Proteobacteria, Bacteroidetes, Firmicutes, Cyanobacteria and two unaffiliated groups. The habitat specialization of the OTUs found in each of the molting stages A, B and D

MA

had comparable numbers of generalists, intermediate and specialists (Fig. 4). Molting stage C, however, was characterized by a marked increase in the contribution of specialists OTUs (from

AC CE P

DISCUSSION

TE

D

13 to 59%).

The molt cycle in decapod crustaceans has been widely studied from the endocrinal basis to applied studies of digestive physiology for many species (Hayd et al. 2008). Resolving the species-specific molt cycle is critical in investigating their biology, or culturing them successfully. The gut microbiota of crustaceans contributes positively to their physiology mainly by participating in molting, food digestion, protecting against pathogens or provisioning vitamins, amino acids and metals (Merrifield and Ringo 2014). However, the dynamics of the gut microbe community in the molting cycle is not well understood. The effect of the gut microorganisms on the host health, development and nutrition is clearly of great interest in aquatic animal biology and aquaculture, recently. Gut microorganisms influence various host functions including development, digestion, nutrition, disease resistance and immunity

ACCEPTED MANUSCRIPT 13

(Chaiyapechara et al. 2011). Gut microorganisms in farmed aquatic animals are influenced by many host and non-host related factors such as environmental conditions, nutrition, farming

PT

practices, stocking density, diet, life history, physiochemical aspects of the gut (Merrifield and

RI

Ringo 2014). The limited number of relevant studies of crustaceans include shrimps (Chaiyapechara et al. 2011), prawns (Tzeng et al. 2015), crabs (Chen et al. 2015) lobsters (Meziti

SC

et al. 2010) and in most cases are focused on nutrition, growth and immunity.

NU

In this study, we investigated the changes of the resident midgut bacterial communities structure throughout the four molting stages of pond-reared Macrobrachium rosenbergii

MA

individuals by analyzing the bacterial community structure as revealed with the diversity of the 16S rRNA gene by using 454 pyrosequencing. By comparing different molting stages, the

D

dominant bacterial taxa were identified in the gut microflora with potential involvement in the

TE

molting process, including those being independent from the molting procedure and those being

AC CE P

unique to each molting stage. This has enabled the assessment of gut microbial communities that play a role in molt cycle of crustaceans. Besides affecting anatomy and morphology, the molting cycle also creates changes in behaviour, physiology and biochemistry. In mature M. rosenbergii the intermoult period is relatively short (29–79 days) (Peebles 1977) but longer than all the other stages. Molting stage C is the stage where the animal must build reserves of glycogen and lipid. It is the metabolically and morphologically most stable period within the moulting cycle. It seems to be a period without dramatic structural and metabolic changes, characterized only by substantial tissue growth. Similarly, this study showed that the gut bacterial community at stage C was different compared to the other three stages. Stage C is significantly different from the other three molting stages regarding major morphometric features (Table 1). The difference of stage C gut bacteria communities compared to the other three stages lies not only at its highest number of bacterial OTUs (Table 2) but also at

ACCEPTED MANUSCRIPT 14

its high number of unique (Fig. 2) and newly introduced -for the gut environment- OTUs (Fig. 3) with most of them being characterized as specialists (Fig. 4).

PT

The high number of OTUs found in the gut seems to be a beneficial feature for the host.

RI

For example, high OTUs richness –determined by 454 pyrosequencing– has been found in the gut of wild and biological reared sea bream (Sparus aurata) compared to the conventional reared

SC

ones (Kormas et al. 2014) and in wild-caught vs. domesticated black tiger shrimps (Penaus

NU

monodon) (Rungrassamee et al. 2014). Moreover, in humans, a high number of OTUs has recently been associated with increased health (Sankar et al. 2015) while gut bacterial diversity

MA

increases from neonates and after 1-3 years the gut microbiome closely resembles that of adults (Costello et al. 2013). In our study, we observed a drop in OTUs numbers from stage C to D,

D

which suggests a change in the gut environment and, subsequently, function. Since it is known

TE

that environmental complexity and resource partitioning favors rich bacterial diversity (Azam and

AC CE P

Malfatti 2007) it is possible that the increased number of OTUs at stage C is related to the M. rosenbergii’s building energy reserves since shortly after molting true tissue growth occurs, as muscle and tissues replace the water absorbed for inflating the new carapace. Stern and Cohen (1982) suggest that oxygen consumption was lowest during the intermolt (C) stage and ammonia excretion increased during the premolt (D) stage to molt which accurately reflect the physiological condition of M. rosenbergii regarding its molt cycle. The increase in oxygen consumption preceding the molt has been attributed to the cumulative energy expense required for the several-fold increase in water uptake and the increase in catabolic as well as anabolic metabolism. Nitrogen is being assimilated for growth during the intermolt stage while the premolt and molt stage reflect an increase in protein metabolism. As amino acids are being deaminated, ammonia is being excreted as the catabolic by-product. Protein levels were correlated with successive stages of the molt cycle for brachyurans and, after allowing for normal

ACCEPTED MANUSCRIPT 15

tissue growth nearly 70% of the protein nitrogen at stage D is calculated to be utilized in the succeeding molt (New et al. 2009). The variation in muscle protein content is reflected in

PT

differences in the levels of amino acids incorporation into protein during the course of the molt

RI

cycle (New et al. 2009).

Another reason for the relatively higher number of bacterial OTUs in stage C could be the

SC

shedding of the hindgut during ecdysis and the subsequent loss of the intestinal microbial biome

NU

followed by recolonization in the postmolt stages. However, a contributing factor could be host suppression of bacterial symbionts in the premolt due to host antimicrobial activity as shown in

MA

an insect (Kim et al. 2014). Additionally, the changes that occur in the gut milieu during the molt, such as altered proteolytic enzyme activity (Muhlia-Almazán and García-Carreño 2002) may

D

reduce its suitability for some symbionts.

TE

The four molting stages hosted very different unique OTUs even at the phylum level

AC CE P

(Table S1. Fig. S2). Again, stage C had the highest number of phyla, while Actinobacteria appeared only at this stage. Actinobacteria is a phylum related mostly to freshwater/terrestrial habitats (Barberán and Casamayor 2010) but OTUs from this phylum have been found in low relative abundances in the gut of the Norway lobster (Meziti et al. 2010) and the Artemia brine shrimp (Quiroz et al. 2015). Likewise, cultivable Actinobacteria have been isolated from the gut of freshwater animals like fish (Jami et al. 2015) and copepods (Homonnay et al. 2012). The Actinobacteria is a large phylum containing microorganisms with a wide array of metabolic features due to its large number of unaffiliated and yet-uncultivated bacteria (Newton et al. 2011). Moreover, the short sequences’ length produced by the 454 pyrosequencing which does not allow secure species identification (Yarza et al. 2014), despite that these short reads are satisfactory proxies of the whole 16S rRNA gene phylogeny (Logares et al. 2014), are precautious on the inferred ecophysiological role of these OTUs. The different environmental

ACCEPTED MANUSCRIPT 16

setting of the gut at stage C is also evidenced by the high number of novel, yet-unaffiliated OTUs (Fig. S2) with currently unknown metabolic features.

PT

A small percentage (10.0%) of the total OTUs number was found to be shared among the

RI

four molting stages (Fig. 2). The presence of these 13 OTUs (OTU1, 2, 3, 4, 5, 6, 9, 10, 12, 20, 22, 25, 29) regardless of the molting stage, renders them possible members of the permanent M.

SC

rosenbergii gut microbiome or at least these OTUs are not affected by the molting procedure.

NU

Following the sensu Hamady and Knight (2009) definition for “core microbiome, this study is statistically secure to claim the realistic presence of each of the core OTUs by using the term

MA

“core”. Since we used normalized data, i.e. all samples had similar total OTUs abundance we are safe to assign the OTUs, which have been found in all four stages as core OTUs in relation to the

D

molting process and are not affected by the changes caused by the molting process. Moreover,

TE

eight of these 13 OTUs are among the 15 most abundant ones (OTU1, 2, 3, 4, 5, 6, 9, 12; Fig. 1)

AC CE P

with the majority of them being characterized as generalists (OTU1, 4, 5, 6, 9) while only one (OTU3) is considered a specialist (Fig. 4). The OTUs characterized as generalists, i.e. occupying a broad range of various habitats (Pandit et al. 2009), have been found to be related to bacteria from dairy products fermentation processes, earthworm gut or endophytic bacteria (Table S1). Consequently, although in this study the exact ecophysiological role of these bacteria remains unresolved, the ecological characterization of their molecular fingerprints relates them to habitats where food fermenters/degraders dominate, assigning to them a possibly similar role for the M. rosenbergii gut. Additionally, the rest of the shared OTUs among all four molting stages, which are not included in the most dominant ones (OTU10, 20, 22, 25, 29), were also related to gut bacteria found in terrestrial and freshwater invertebrates (Table S1). Finally, OTU3 which was the only specialist being shared among all four molting stages and being among the dominant OTUs, was closely related to Aeromonas veronii (Table S1), a

ACCEPTED MANUSCRIPT 17

potential pathogen of humans (Hickman-Brenner et al. 1987) but also of M. rosenbergii (Sung et al. 2000), indicating its specific ecological role as a specialist, i.e. a more ecologically restricted

PT

OTU compared to generalists in terms of habitat range, having very specific environmental

RI

requirements for its growth (Pandit et al. 2009).

Apart from the shared OTUs, of special interest are the rest of the dominant ones. Since in

SC

each stage between 6 and 9 OTUs comprised more than 90% of the cumulative relative OTU

NU

abundance (Table 2), it is reasonable to hypothesize that these bacteria (Fig. 1) pose a central role in the gut function or at least are the best adapted in this environment (Konopka 2009). The

MA

majority of the rest of the dominant OTUs (OTU7, 8, 11, 14, 16, 19, 21) belong to the γProteobacteria and Firmicutes and are related to animal faeces, gastrointestinal tracts, dairy

D

products or fermentation processes (Table S1), enhancing their potential involvement in food

TE

processing in the gut. These phyla are among the dominant ones in other decapod crustaceans’

AC CE P

gut environments (Meziti et al. 2010, Rungrassamee et al. 2014) but also of fish (e.g. CardaDiéguez et al. 2014), insects (Engel and Moran 2013) and even humans’ (Sankar et al. 2015). In conclusion, in this study we investigated whether there is a succession in the resident bacterial community structure along the molting process; by recognizing the bacteria that dominate in each stage, i.e. are most likely better adapted to the gut conditions during that stage, one can depict bacteria with specific potential metabolic repertoires, potential pathogens and candidate probiotic species –specific to each molting stage- based on their inferred ecophysiological role. This study presented for the first time that major structural changes in the Macrobrachium rosenbregii freshwater prawn gut bacterial community coincide with the molting stage that is the longest and with the most significant differences in animal morphometry (stage C). These gut bacterial community changes have to do with increased number of OTUs, most of them being unique, newly introduced in the gut habitat and characterized as specialists, with the majority of them belonging to the

ACCEPTED MANUSCRIPT 18

Actinobacteria and unaffiliated groups, indicating the unique gut environmental conditions that prevail at stage C. Moreover, the inferred ecophysiological role of these OTsUs is related to food

PT

material processing and/or originate from other aquatic animals’ gut environments. The presence of 13 OTUs, with most of them being related to the γ-Proteobacteria and Firmicutes, was found to be

RI

unaffected by the molting stage, indicating these bacteria’s role in the general gut ecophysiological

SC

state, unrelated to the animal’s growth. The establishment and succession of gut microbiota is

NU

controlled by the ingested food, which serves as the bacteria’s growth medium, and the gut’s prevailing conditions, i.e. the “incubation” conditions. Thus, the structure and activity of the gut

MA

microbiota as well as the effects on the host itself are useful to support intestine microbial communities and maintain microbial balance with the surrounding environment. Additionally,

D

microbial communities can be provided via the diet or the rearing water to the prawn to improve

TE

growth, health and welfare of the farmed prawn. It can be a tool implemented by farmers as a best

AC CE P

management practice in prawn aquaculture. Further research should attempt to associate the importance of these bacterial groups with specific functions of the host.

Acknowledgments

We would like to thank the Gadsden State Community College Aquaculture Program for providing the animals used in this study and the technical expertise of Tim Adams. GSCC Aquaculture students assisted in prawn harvest and animal maintenance.

References

ACCEPTED MANUSCRIPT 19

Aflalo, E.D, Raju, D.V.S.N, Bommi, N.A, Verghese, J.T, Samraj, T.Y.C., Hulata, G., Ovadia, O., Sagi, A. 2012. Toward a sustainable production of genetically improved all-male prawn

PT

(Macrobrachium rosenbergii): Evaluation of production traits and obtaining neo-females in

RI

three Indian strains. Aquaculture, 338-341, 197-207.

Azam, F, Malfatti, F. 2007. Microbial structuring of marine ecosystems. Nat Rev Microbiol.,

SC

5, 782–791.

NU

Barberán, A., Casamayor, E. 2010. Global phylogenetic community structure and β-diversity patterns in surface bacterioplankton metacommunities. Aquatic Microbial Ecology, 59, 1-

MA

10.

Carda-Diéguez, M., Mira, A., Fouz, B. 2014. Pyrosequencing survey of intestinal microbiota

D

diversity in cultured sea bass (Dicentrarchus labrax) fed functional diets. FEMS

TE

Microbiology Ecology, 87, 451-459.

AC CE P

Chaiyapechara, S., Rungrassamee, W., Suriyachay, I., Kuncharin, Y., Klanchui, A, et al. 2011. Bacterial community associated with the intestinal tract of P. monodon in commercial farms. Microb Ecol, 63, 938–953. Chen, X., Di, P., Wang, H., Li, B., Pan, Y., Yan, S., Wang, Y. 2015. Bacterial Community Associated with the Intestinal Tract of Chinese Mitten Crab (Eriocheir sinensis) Farmed in Lake Tai, China. PLoS ONE, 10, e0123990. Claesson, M.J., Wang, Q., O'Sullivan, O., Greene-Diniz, R., Cole, J.R., Ross, R.P., O'Toole P.W. 2010. Comparison of two next-generation sequencing technologies for resolving highly complex microbiota composition using tandem variable 16S rRNA gene regions. Nucleic Acids Research, 38(22), e200. Dowd, S., Callaway, T., Wolcott, R., Sun, Y., McKeehan, T., Hagevoort, R., Edrington, T.S. 2008. Evaluation of the bacterial diversity in the feces of cattle using 16S rDNA bacterial

ACCEPTED MANUSCRIPT 20

tag-encoded FLX amplicon pyrosequencing (bTEFAP). BMC Microbiology, 8:125 D’Abramo L.R. New MB. 2010 Nutrition, feeds and feeding. pp. 218 – 238 in Freshwater

RI

M.N. Kutty, eds. John Wiley and Sons. West Sussex, UK.

PT

Prawns: Biology and Farming. M.B. New, W.C. Valenti, J.H Tidwell, L.R. D’Abramo, and

Drach, P. 1939 Mue et cycle d'intermue chez les Crustacés Décapodes. (Molting and the

SC

intermolt cycle in Decapod Crustaceans). Ann. Inst. océanogr., Paris, 19, 103-391, 6 pl.

NU

Drach, P. Tchernigovtzeff, C. 1967. Sur la methode de determination des stades d'intermue et son application generale aux crustaces. Vie et Milieu. Series A, 18, 595-610.

MA

Edgar, R. 2010. Search and clustering orders of magnitude faster than BLAST. Bioinformatics 26, 2460-2461.

D

Engel, P., Moran, N.A. 2013. The gut microbiota of insects – diversity in structure and

TE

function. FEMS Microbiology Reviews, 37, 699-735.

AC CE P

FAO (2013). Fisheries and Aquaculture Department. Cultured aquatic species information programme data collection (FAO 2004-2016). Aquaculture management and conservation service. FAO-FIMA.

http://www.fao.org/fishery/culturedspecies/Macrobrachium_rosenbergii/en Hamady, M. 2009. Knight R. Microbial community profiling for human microbiome projects: Tools, techniques, and challenges. Genome Research, 19, 1141-1152. Hartnoll, R.G. 2001. Growth in Crustacea – twenty years on. Hydrobiologia, 449, 111-22. Hayd, L.A., Anger, K., Valenti, W.C. 2008. The moulting cycle of larval Amazon river prawn Macrobrachium amazonicum reared in the laboratory. Nauplius, 16, 55-63. Hickman-Brenner, F.W., MacDonald, K.L., Steigerwalt, A.G., Fanning, G.R., Brenner, D.J., Farmer, J.J. 1987. Aeromonas veronii, a new ornithine decarboxylase-positive species that

ACCEPTED MANUSCRIPT 21

may cause diarrhea. Journal of Clinical Microbiology, 25, 900-906. Homonnay, Z.G., Kéki, Z., Márialigeti, K., Tóth, E.M. 2012. Bacterial communities in the gut

PT

of the freshwater copepod Eudiaptomus gracilis. Journal of Basic Microbiology, 52, 86-90.

RI

Jami, M., Ghanbari, M,. Kneifel, W., Domig, K.J. 2015. Phylogenetic diversity and biological activity of culturable Actinobacteria isolated from freshwater fish gut microbiota.

SC

Microbiol Res., 175, 6-15.

NU

Karplus, I. and A. Sagi. 2010. The Biology and Management of Size Variation p. 316 - 345 in Freshwater Prawns: Biology and Farming edited by M.B. New et al. Blackwell Publishing,

MA

West Sussex UK)

Kennedy, B., Venugopal, M.N., Karunasagar, I., Karunasagar, I. 2006. Bacterial flora

D

associated with the giant freshwater prawn Macrobrachium rosenbergii, in the hatchery

TE

system. Aquaculture, 261, 1156-116.

AC CE P

Kim, J.K., Han, S.H., Kim, C-H, Jo, Y.H., Futahashi, R., Kikuchi, Y., Fukatsu, T., Lee, B.L. 2014. Molting-associated suppression of symbiont population and up-regulation of antimicrobial activity in the midgut symbiotic organ of the Riptortus– Burkholderia symbiosis. Developmental and Comparative Immunology, 43(1), 10-14. Klindworth, A., Pruesse, E., Schweer, T., Peplies, Jr., Quast, C., Horn, M., Glöckner, F.O. 2012. Evaluation of general 16S ribosomal RNA gene PCR primers for classical and nextgeneration sequencing-based diversity studies. Nucleic Acids Research, 41(1), e1 Konopka, A. 2009. What is microbial community ecology? ISME Journal, 3, 1223-1230. Kormas, K.A., Meziti, A., Mente, E., Frentzos, A. 2014. Dietary differences are reflected on the gut prokaryotic community structure of wild and commercially reared sea bream (Sparus aurata). MicrobiologyOpen, 3, 718-728. Kunin, V., Engelbrektson, A., Ochman, H., Hugenholtz, P. 2010. Wrinkles in the rare

ACCEPTED MANUSCRIPT 22

biosphere: pyrosequencing errors can lead to artificial inflation of diversity estimates. Environmental Microbiology, 12, 118-123.

PT

Logares, R., Sunagawa, S., Salazar, G., et al. 2014. Metagenomic 16S rDNA Illumina tags are

RI

a powerful alternative to amplicon sequencing to explore diversity and structure of microbial communities. Environmental Microbiology, 16, 2659-2671.

SC

Meziti, A., Ramette, A., Mente, E., Kormas, K.A. 2010. Temporal shifts of the Norway lobster

NU

(Nephrops norvegicus) gut bacterial communities. FEMS Microbiology Ecology, 74, 472484.

MA

Merrifield, D., Ringø, E. 2014. Aquaculture nutrition: gut health, probiotics and prebiotics. Oxford, John Wiley and Sons, Ltd. pp. 480.

D

Muhlia-Almazán, A., García Carreño, F. 2002. Influence of molting and starvation on the

TE

synthesis of proteolytic enzymes in the midgut gland of the white shrimp, Penaeus

AC CE P

vannamei. Comparative Biochemistry and Physiology Part B, 133, 383–394 New, M., Nair, C.M. 2012. Global scale of freshwater prawn farming. Aquaculture Research, 43(7), 960-969.

New, M.B., Valenti, W.C., Tidwell, J.H., D’Abramo, L.R., Kutty, M.N. 2009. Freshwater prawns: biology and farming. Wiley-Blackwell Publishing, Oxford, UK 560p. Palmer, C., Bik, E.M., DiGiulio, D.B., Relman, D.A., Brown, P.O. 2007. Development of the human infant intestinal microbiota. PLoS Biol , 5, e177. Pandit, S.N., Kolasa, J., Cottenie, K.2009. Contrasts between habitat generalists and specialists: an empirical extension to the basic metacommunity framework. Ecology, 90, 2253-2262. Peebles, J.B. 1977. A rapid technique for molt staging in live Macrobrachium rosenbergii. Aquaculture, 12, 173-180.

ACCEPTED MANUSCRIPT 23

Peebles, J.B.1979. The role of prior residence and relative size in competition for shelter by the Lamaysian prawn Macrobrachium rosenbergii. Fisheries Bulletin, 76(4), 905-911.

PT

Pruesse, E., Quast, C., Knittel, K., Fuchs, B., Ludwig, W., Peplies, J. Glöckner, F. 2007.

RI

SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Research, 35, 7188-7196.

SC

Quince, C., Lanzen, A., Curtis, T.P., Davenport, R.J., Hall, N., Head, I.M., Read, L.F., Sloan

NU

WT 2009. Accurate determination of microbial diversity from 454 pyrosequencing data. Nat Meth, 6, 639-641.

MA

Quiroz, M., Triadó-Margarit, X., Casamayor, E., Gajardo, G. 2015. Comparison of artemia– bacteria associations in brines, laboratory cultures and the gut environment: a study based

D

on Chilean hypersaline environments. Extremophiles, 19, 135-147.

TE

Rungrassamee, W., Klanchui, A., Maibunkaew, S., Chaiyapechara, S., Jiravanichpaisal, P.,

AC CE P

Karoonuthaisiri, N. 2014. Characterization of Intestinal Bacteria in Wild and Domesticated Adult Black Tiger Shrimp (Penaeus monodon). PLoS ONE, 9, e91853. Sankar, S.A., Lagier, J-C., Pontarotti, P., Raoult, D., Fournier, P-E. 2015. The human gut microbiome, a taxonomic conundrum. Systematic and Applied Microbiology, 38, 276-286. Schloss, P.D., Westcott, S.L., Ryabin, T., Hall, J.R., Hartmann, M., Hollister, E.B., Lesniewski, R.A., Oakley, B.B., Parks, D.H., Robinson, C.J., Sahl, J.W., Stres, B., Thallinger, G.G., Van Horn, D.J., Weber, C.F. 2009. Introducing mothur: Open-source, platform-independent, community-supported software for describing and comparing microbial communities. Applied and Environmental Microbiology, 75, 7537-7541. Stern, S., Cohen, D. 1982. Oxygen consumption and ammonia excretion during the molt cycle of the freshwater prawn Macrobrachium rosenbergii (De Man). Comparative Biochemistry and Physiology, 73A, 417-419.

ACCEPTED MANUSCRIPT 24

Sung, H-H., Hwang, S-F., Tasi, F-M. 2000. Responses of giant freshwater prawn (Macrobrachium rosenbergii) to challenge by two strains of Aeromonas spp. Journal of

PT

Invertebrate Pathology, 76, 278-284.

RI

Tzeng, T-D., Pao, Y-Y., Chen, P-C., Weng, FC-H., Jean, WD., Wang, D (2015). Effects of host phylogeny and habitats on gut microbiomes of oriental river prawn (Macrobrachium

SC

nipponense). PLoS ONE 10(7):e0132860. Doi10.1371/journal. Pone.0132860.

NU

Yarza, P., Yilmaz, P., Pruesse E et al. 2014. Uniting the classification of cultured and uncultured bacteria and archaea using 16S rRNA gene sequences. Nat Rev Micro., 12, 635-

AC CE P

TE

D

MA

45.

ACCEPTED MANUSCRIPT 25

Legends to figures

PT

Figure 1. Dominant (cumulative relative abundance of ≥90%) operational taxonomic units (OTUs) of gut Bacteria in each of the four (A, B, C, D) molting stages of Macrobrachium

RI

rosenbergii. G: generalist, I: intermediate, S: specialist OTUs according to Levin’s niche width

NU

SC

(B) index (see text for further information).

MA

Figure 2. Venn diagrams showing the number of unique and shared operational taxonomic (OTUs) of gut Bacteria between the four molting stages (A, B, C, D) in Macrobrachium

TE

D

rosenbergii.

AC CE P

Figure 3. Number of total (red line) and newly introduced operational taxonomic (OTUs) of Bacteria in each of the four (A, B, C, D) molting stages of Macrobrachium rosenbergii.

Figure 4. The contribution of generalists (G), intermediate (I) and specialists (S) operational taxonomic (OTUs) of Bacteria during the four (A, B, C, D) molting stages of Macrobrachium rosenbergii according to Levin’s niche width (B) index (see text for further information).

ACCEPTED MANUSCRIPT

MA

NU

SC

RI

PT

26

AC CE P

TE

D

Fig. 1

ACCEPTED MANUSCRIPT

Fig. 2

AC CE P

TE

D

MA

NU

SC

RI

PT

27

ACCEPTED MANUSCRIPT

TE AC CE P

Fig. 3

D

MA

NU

SC

RI

PT

28

ACCEPTED MANUSCRIPT

AC CE P

Fig. 4

TE

D

MA

NU

SC

RI

PT

29

ACCEPTED MANUSCRIPT 30

Table 1. Morphometric data of Macrobrachium rosenbergii measured in the present study. M: male, F: female.

A

M

28.50

57.0

20.5

3

A

M

46.48

68.6

25.3

4

A

M

40.86

36.4

5

A

M

56.39

60.8

6

B

M

47.64

79.9

7

B

F

39.56

71.5

8

B

M

60.51

9

B

M

38.44

10

B

M

42.00

11

C

M

25.16

12

C

13

C

14

C

15

C

16

C

17

21.6 20.6

150.9

20.5

25.3

153.5

22.4

27.6

148.3

20.9

21.6

154.2

21.9

25.4

152.2

21.3

85.4

28.2

169.9

24.4

73.0

23.8

148.8

19.2

80.7

24.5

157.3

21.1

63.4

20.9

125.0

18.1

SC NU

MA D

TE

AC CE P

Abdominal width (mm)

125.4

RI

2

PT

Sample Molting Gender Wet body mass Carapace length Carapace width Rostrum to stage (g) (mm) (mm) uropod length (mm) 1 A M 75.90 88.0 30.5 182.0

F

24.91

62.6

19.7

124.9

17.3

M

41.93

71.0

23.8

146.2

20.0

F

30.21

68.4

23.0

133.1

20.0

F

27.23

66.3

20.5

129.0

19.8

F

29.45

62.5

23.4

141.1

21.7

D

F

34.00

70.4

22.1

150.9

19.6

18

D

F

46.04

71.6

25.2

166.8

20.1

19

D

M

42.30

73.7

25.0

152.8

21.1

20

D

F

27.78

64.6

20.5

141.1

18.7

21

D

F

27.98

64.6

20.9

140.7

19.2

ACCEPTED MANUSCRIPT 31

Table 2. Normalised pyrosequencing results of the 16S rRNA gene in the gut of the four molting

Dominance of the most

(cumulative relative dominance)

2408

(29)

39.8%

7 (90.8%)

B

2413

(39)

63.6%

6 (92.7%)

C

2401

(93)

34.8%

9 (90.8%)

D

2410

(43)

39.7%

7 (90.4%)

D

MA

A

NU

SC

abundant OTU

No. of most dominant OTUs

RI

Molting stage Reads (OTUs)

PT

stages of Macrobrachium rosenbergii. OTU: operational taxonomic unit.

AC CE P

TE

Levin’s niche width (B) index (see text for further information).

ACCEPTED MANUSCRIPT 32

Highlights To our knowledge this is the first study that investigates the dynamics of the gut microbe community during the prawn molt cycle.



Major changes in the M. rosenbergii freshwater prawn gut bacterial community structure coincide with molting stage C



The distinct gut bacterial communities found in molting stage C corroborates with the physiological significance of this molting stage.

AC CE P

TE

D

MA

NU

SC

RI

PT

