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Boletín de Investigaciones Marinas y Costeras - INVEMAR

Print version ISSN 0122-9761

Bol. Invest. Mar. Cost. vol.47 no.2 Santa Marta July/Dec. 2018

http://dx.doi.org/10.25268/bimc.invemar.2018.47.2.746 

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Bacterial community structure in different tissues of the wild Lobatus gigas (Linnaeus, 1758) from the Caribbean Seaflower Biosphera Reserve

Mónica Marcela Higuita-Valencia 0000-0002-4031-49311  *  , Olga Inés Montoya-Campuzano 0000-0002-4820-03811  **  , Edna Judith Márquez-Fernández 0000-0003-0760-37471  ***  , Claudia Ximena Moreno-Herrera 0000-0002-8132-52231  **** 

1 Facultad de Ciencias, Universidad Nacional de Colombia, sede Medellín, Colombia. Grupo de Microbiodiversidad y Bioprospección (Microbiop), Universidad Nacional de Colombia-Sede Medellín-Facultad de Ciencias-Escuela de Biociencias-Laboratorio de Biología Celular y Molecular, Calle 59A No 63 - 20 Bloque 19 A Laboratorio 310, Medellín, 050034 Colombia.

ABSTRACT

The microbial diversity of Lobatus gigas has not been thoroughly studied despite of them is a specie endangered. Knowledge of microbiota may help to improve the conservation and cultivation of this species. The objective of this study was to evaluate the bacterial populations associated with the gonad and the gut compartments of the wild endangered L. gigas from the Caribbean Seaflower Biosphere Reserve, using microbiological methods and culture-independent molecular tools. The genetic profiles of the bacterial populations were generated and Temporal Temperature Gradient Electrophoresis (TTGE) was used to compare them with total DNA. A genetic and statistical analysis of the bacterial communities revealed a low level of diversity in gonad tissue based on the number of bands detected using TTGE. In addition, statistical differences in bacterial community structure were found between the foregut and hindgut tissue of L. gigas. The dominant phylogenetic affiliations of the gonad bacteria, as determined using 16S rRNA gene sequencing, belong to Ralstonia (50%). The possible involvement of this genus in the reproduction and development of the conch is discussed. On the other hand, the bacterial phylotypes from foregut and hindgut included members of Alphaproteobactera (12.5%), Betaproteobacteria (12.5%), Gammaproteobacteria (12.5%), Bacilli (31.25%), Clostridia (6.25%), Actinobacteria (6.25%), Mollicutes (6.25%) and Deinococci (6.25%) classes. Knowing the composition of the gonad and foregut and hindgut bacteria of L. gigas is the first step toward exploring the proper management of this species, as well as provides useful information to future researches that allow a better understanding of the role of these bacterial populations in the health and reproductive rate of L. gigas.

Key words: Lobatus gigas, 16S rRNA gene; Bacterial Community; Gonad; gut

INTRODUCTION

The Queen conch, Lobatus gigas, is a species of ecological and economic importance throughout its distribution area (Catarci, 2004). This species plays an important role in the ecological interactions of the near shore marine populations, as one of the main herbivores of seagrass, epiphyte algae and detritus. Additionally, it is a commensal organism, serves as a food source for a wide variety of marine organisms, and competes with them for resources (Stoner and Waite, 1991; Tewfik, 1997; Catarci, 2004). Furthermore, it is a fishery resource with economic value in the marine food trade. Thus, many countries are required to establish policies to regulate its exploitation by commercializing it in a sustainable way without jeopardizing its conservation. Nonetheless, despite the design and implementation of national and international management measures to control the overexploitation and access to this resource (Prada et al., 2008), their population density continues to decline (Ballesteros et al., 2005; Stoner et al., 2012). This condition hinders the proper maintenance of the population, affecting its self-healing abilities and rendering its commercialization unsustainable (Stoner and Ray-Culp, 2000). As a consequence, it has been included in appendix II of the Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES) (Daves and Fields, 2004). In order to overcome this situation and to recover the overexploited populations of L. gigas, several alternatives have been established, such as the creation of marine protected areas (MPA’s) (Appeldoorn, 1994 ; Stoner, 1996; Anon, 1999) which allow the preservation of spawning stock at high densities and maintain a haven for adults with higher reproducibility (Anon, 1999). Another alternative is aquaculture with techniques directed to produce juveniles (Brownell, 1977; Creswell, 1994).

In the Seaflower Biosphere Reserve, several conservation measures have been stablished to regulate L. gigas fishing and to improve understanding of its biology (Castro et al., 2007). More specifically, studies have focused mainly on the population genetics (Landínez- García et al., 2011; Márquez et al., 2012), dynamics, and reproductive biology of this mollusk (Aranda et al., 2001; Aranda et al., 2003a, b; Delgado et al., 2004; Castro et al., 2007). But, these conservation alternatives did not consider studies on microbiota, which is a fundamental aspect of successful culture and restoration of marine organisms. For instance, it has been demonstrated in several species of fish and crustaceans that the microbial community influences various host functions including development, digestion, nutrition, disease resistance and immunity (Merrifield and Ringø, 2014), aspects that are fundamental to the health of any organism. Despite this, to date there is very little known about how the microbiota influence the growth and development of the queen conch (Kirjavainen and Gibson, 1999; Kowalik et al., 2006, Pérez et al., 2014). Recently, others have analyzed the queen conch microbiota through culture-dependent tools, genotypic identification (Acosta et al., 2009, Pérez et al., 2014) and culture-independent approaches (Pérez et al., 2014) using gut and secretion samples from wild conch collected near the Isla del Rosario and the Isla Tortugas in Colombia. These studies demonstrated the presence of Pseudoalteromonas sp., Halomonas sp., Psychrobacter sp., Cobetia sp., Pseudomonas sp., Vibrio sp. and Burkholderia sp. (Acosta et al., 2009; Pérez et al., 2014). Cuartas et al., (2018) using histological analysis, 454 pyrosequencing and a combination of PCR amplification and automated Sanger sequencing indicated that the etiological agent of the muscle protrusions is a parasite belonging to the subclass Digenea and is associated with bacterial and fungi clades. Nevertheless, currently, there are no reports about the molecular characterization of bacterial microbiota associated with the compartments of the gonad and the digestive tract of L. gigas.

The digestive tract is a compound ecosystem that contains a dynamic and complex consortium of microorganisms, which play a key role in the nutrition and health of the host (Bäckhed et al., 2004). The gut microbiota is involved in important processes, such as stimulation of epithelial proliferation, development of nutrient metabolism and innate immune response (Rawls et al., 2004; Sommer and Bäckhed, 2013; Thaiss et al., 2016). Thus, knowledge about the bacteria associated with the different gastrointestinal compartments could be useful to control microbiota as a strategy to improve nutrition, prevent pathogenic infections and to develop new methodologies that contribute to the successful culture of this species. To date, there are no studies aimed at identifying the bacterial communities that inhabit the conch gonad and the potential role on the health and the development of this species. Nonetheless, the presence of bacteria in the gonad has been reported in other mollusks, such as, Argopecten purpuratus where the permanent presence of the genera Vibrio, Acinetobacter, Moraxella, Pseudomonas and Cytophaga in the reproductive organs was found (Chavez and Riquelme, 1994; Riquelme et al., 1995a). In the mollusk Dendrodoris nigra the presence of bacteria in the gonad was determined by means of electron microscopy, but these bacteria were not classified (Klussmann-Kolb and Brodie, 1999). Although the gonad microbiota is poorly understood in marine organisms, previous studies using mammals suggests that the reproductive system microbiome is an important site of bacterial-mammalian symbiosis. These bacteria play important roles, such as, providing protection against pathobiont colonization and producing antimicrobial compounds (Reid et al., 2011; Miller et al., 2017; Kindinger et al., 2017).

The main objective of the present work was to estimate the bacterial community structure associated with gonad, and throughout the different sections of the gut of the wild L. gigas from the Caribbean Seaflower Biosphere Reserve, using culture-dependent and culture- independent analysis. Also, we analyzed microbiota of pool and individuals to study inter-individual variation. To our knowledge, the present work is the first examination of bacterial microbiota of the gonad and throughout the gut compartments of queen conch. This approach enables the detection of dominant bacterial communities that could be of importance in the reproduction and development of the queen conch.

MATERIALS AND METHODS

Sample Collection and Processing

Twenty-two samples of L. gigas (queen conch) juvenile specimens of approximately 350 g each, were collected by fishermen in the Colombian Caribbean Sea, San Andres archipelago (12° -16° N and between 78° - 82°O), these samples were provided by the Gobernación del Archipielago de San Andrés, Providencia y Santa Catalina, through the scientific cooperation agreement #083/2012 (Cuartas et al., 2018). The conchs were transported to the laboratory on dry ice, where the gonad (G), foregut (F), and hindgut (H) of each conch was aseptically extracted under cold conditions. Tissues removed from 15 conchs with similar weight were separatedintofivegroupsandeachgroupwashomogenized and referred to as a pool (P) of each tissue. Pooling samples is a common practice to study the gut microbiota in fish (Hovda et al., 2007; Andlid et al., 1998; Romero and Navarrete, 2006). The tissues corresponding to the seven (7) remaining conchs were processed individually (I), to study inter-individual variation (Romero and Navarrete, 2006). Finally, the samples obtained from tissues (P and I) were immediately macerated with liquid nitrogen and then stored at -80ºC until their processing. Since bacterial communities were obtained from each gut compartment that contained epithelium and digested food, the microbiota analyzed was a combination of both autochthonous (able to colonize the epithelial surface or mucus of the host gut) and allochthonous (transient or associated with digestion) bacteria (Figure 1).

Figure 1 (a). Lobatus gigas anatomy scheme taken from Reed (1996). (b). Dissection and processing of gonad (G), foregut (F) and hindgut of L. gigas

For the culture-dependent analysis, serial dilutions of homogenates from either P or I conches were plated in marine agar (Difco) and Thiosulfate-citrate-bile salts-sucrose agar-TCBS (Merck), and then the plates were incubated aerobically at 20ºC for 2 days. Colony forming units (CFUs) recovered from the different media were preserved in 20% glycerol at -80ºC and referenced as cultivable fraction (C). The direct fraction (D) refers to the macerated tissue samples frozen at -80ºC without being cultured (G, F, and H). Nomenclature of the samples is summarized in table 1.

Table 1 Nomenclature of samples obtained from the gut compartments and gonad of Lobatus gigas from the biosphere reserve (SEAFLOWER). D= Total DNA from direct fraction, C= DNA from cultivable fraction, I= individual samples, P= pools, G= Gonad, F= foregut and H= hindgut. 

Nomenclatura Nomenclature Extracción de ADN DNA Extraction Orígenes Origen Proceso Process Muestras Samples
CIDI / CFI Fracción cultivable "C" / Cultivable fraction “C” Intestino delgado "ID" Foregut “F” Individual “I” CIDI2, CIDI3, CIDI4, CIDI6, CIDI16 CFI2, CFI3, CFI4, CFI6, CFI16
CIDP / CFP Pool "P" CIDP1, CIDP2, CIDP4, CIDP5 CFP1, CFP2, CFP4, CFP5
CIGI / CHI Intestino grueso "IG" Hindgut “H” Individual “I” CIGI2, CIGI3, CIGI6, CIGI16 CHI2, CHI3, CHI6, CHI16
CIGP / CHP Pool "P" CIGP1, CIGP2, CIGP4, CIGP5 CHP1, CHP2, CHP4, CHP5
CGI / CGI Gónada "G" Gonad “G” Individual “I” CGI3, CGI4, CGI6, CGI16
CGP / CGP Pool "P" CGP1, CGP2, CGP4, CGP5
DIDI / DFI Fracción directa "D" / Direct fraction “D” Intestino delgado "ID" Foregut “F” Individual “I” DIDI2, DIDI3, DIDI6, DIDI16 DFI2, DFI3, DFI6, DFI16
DIDP / DFP Pool "P" DIDP1, DIDP2, DIDP3, DIDP4, DIDP5 DFP1, DFP2, DFP3, DFP4, DFP5
DIGI / DHI Intestino grueso "IG" Hindgut “H” Individual “I” DIGI2, DIGI3, DIGI6, DIGI16 DHI2, DHI3, DHI6, DHI16
DIGP / DHP Pool "P" DIGP1, DIGP2, DIGP3, DIGP4, DIGP5 DHP1, DHP2, DHP3, DHP4, DHP5
DGI / DGI Gónada "G" Gonad “G” Individual “I” DGI3, DGI4, DGI6, DGI16
DGP / DGP Pool "P" DGP1, DGP2, DGP3, DGP4, DGP5

DNA Extraction

The DNA from the cultivable fraction (F) of each sample was extracted with a DNEasy tissue kit (Qiagen, Duesseldford, Germany) using a modified DNEasy DNA extraction protocol to ensure lysis of Gram positive and Gram negative bacteria, as described by Li et al. (2009).

The total genomic DNA from the direct fraction of each tissue was extracted using an Ultraclean soil DNA extraction kit with a prior tissue lysis step, as described by Ó Cuív et al. (2011). Briefly, each dissected tissue from L. gigas was obtained as described above and 300 mg were homogenized with 800 μl of lysis buffer (6 mM Tris-HCl [pH 8], 100 mM EDTA, 1M NaCl) and incubated at 75ºC for 10 minutes to inactivate nucleases. 20 μl of lysozyme were added (200 mg/ml) and incubated overnight at 37ºC. Subsequently, a digestion with 20 μl of proteinase K (20 mg/ml) at 56ºC was performed until complete lysis of each tissue.

PCR amplification

In order to obtain molecular fingerprinting of the bacterial community from the foregut, hindgut and gonad cultured on marine agar (C), the 16S rRNA gene between V3 and V6 region was amplified with specific primers 341F with an extra GC tail (5 ′GCCTACGGGAGGCAGCAG 3′) and 907R (5′CCGTCAATTCMTTTGAGTTT 3′) (McCracken et al., 2001). The PCR was carried out using the conditions described by García et al. (2016).

Amplification of the hypervariable region between V3 and V6 regions of 16S rRNA from the D was performed with a nested PCR. The first PCR was carried out to amplify nearly full-length 16S rRNA gene sequences with universal primers 27F (5`AGAGTTTGATCMTGGCTCAG 3`) and 1492R (5` GTTACCTTGTTACGACTT 3`) following the conditions given by Espejo and Romero (1997). Then, 2.5 μl from the product of the first PCR were used as template DNA in the second reaction, which was performed with the specific primers 341-GC F and 907R (as described previously for cultivable samples) and under the conditions reported by Espejo et al. (1998).

Temporal Temperature Gradient Electrophoresis (TTGE) analysis

The structure of bacterial communities associated with the foregut, hindgut and gonad was revealed using the TTGE technique, which allowed separating fragments of the 16S rRNA gene of approximately 585 bp, with the assumption that each band observed in the banding patterns represents a bacterial species.

The PCR products obtained from both fractions, using the 341F-GC and 907R primers, were separated by TTGE using the DCode Universal Mutation Detection System (BioRad, USA) on 7% (w/v) polyacrylamide, 7M Urea gels in 1.25X TAE buffer for 15h at 55V. The initial and final temperatures were 66ºC and 69º respectively, with a temperature ramp of 0.1ºC per hour. The gels were stained with AgNO3 (AMRESCO, OH, USA) (Sanguinetti et al., 1994). The band patterns obtained for each sample studied were analyzed using GelCompar II (Applied Maths TX, USA) (Rademaker and De Bruijn, 2008, Garcia et al., 2016). The reference lanes from each sample were aligned according to a 100 bp ladder, and a cluster analysis was performed using DICE coefficient (Nei and Li, 1979) and the Unweighted Pair Group Method with Arithmetic Average (UPGMA) (Mohammadi and Prasanna, 2003).

Bacterial community analysis

GelCompar II software (Applied Biosystems, Belgium) was used to analyze TTGE profile patterns from the DNA samples. For this analysis, the presence and absence matrix was used to generate an analysis of similarity (ANOSIM) based on the Bray-Curtis index using PAST software version 2.0 (Hammer et al., 2001). Significant differences in bacterial diversity were assessed with the “Diversity t-test” option of the PAST program. For this analysis, the band number was considered to represent the species number or Taxa (S) and the band intensity was considered to represent the relative abundance or number of individuals of each bacterial species. The “Principal Component Analysis” option was used to view overall similarities and dissimilarities among sampling of the gonad, foregut and hindgut and between the cultivable (C) and direct (D) fraction from each tissue (Chaiyapechara et al., 2012; Pérez et al., 2014, Boon, 2002). This clustering distribution was confirmed by the analysis carried out with the two gel replicates. Finally, the Shannon index (H`) which reflects the diversity of the bacterial community of F, H and G was calculated following the formula: H` = - SPi ln Pi, where Pi is defined as (ni ⁄N), ni is each band present, and N is the sum of all bands (Sun et al., 2011).

TTGE bands with unique migration patterns according to the GenRulerTM 100bp DNA ladder (Thermo Fisher Scientific) and bands with higher intensity in each TTGE, were cleaved and DNA was eluted following the Soak and Crush method described by Sambrook and Russell (2002). Then 5μl from the final elution were taken and re-amplified with primers 341F without GC tail and 907R. Re-amplified products from the C fraction was verified by gel electrophoresis on a 1% agarose gel. The re-amplified products from D were purified using the QIAquick PCR purification kit (QIAGEN CA, U.S.A) and cloned into E. coli DH5α competent cells using a Clone JET TM kit (Thermo Scientific, U.S.A). Ligation was performed following the manufacturer’s indications and transformation was completed as described by Sambrook and Russell (2002). The insert was amplified following the QIAquick purification kit (QIAGEN CA, U.S.A) recommendations with pJET1.1 and pJET 1.2 primers. Amplicons from D and C were sequenced in on an ABI PrimR 3100 Genetic Analyser (Applied Biosystems, CA, USA).

Sequences of 16S rDNA obtained from both fractions were edited using BioEdit ® (Hall, 1999) and the presence of chimeras was assessed with the online software DECIPHER v2.0 described by Wright et al. (2012). The edited sequences were then compared with reference sequences from The National Center for Biotechnology information (NCBI) and Ribosomal Database Project (RDP). To determine the phylogenetic affiliation of the microbial isolates and the excised bands, similarity searches and phylogenetic analysis were performed using the BLASTN and MEGA 7 software (Tamura et al., 2007). Phylogenetic analysis was performed using the Neighbor-Joining method (Saitou and Nei, 1987) with 1000 bootstrap replicates. It was rooted with a 16S rRNA gene sequence from Crenarchaeote symbiont of Axinella sp., in order to improve the topology. The sequences obtained in this study were deposited in GenBank under accession numbers KX891431-KX891450 and KX886796-KAX886797.

RESULTS

Bacterial count

The average cultivable bacterial abundance of gut tract compartments and gonad in each medium is observed in Table 2 and expressed as CFUs per gram of tissue. The averages of bacterial abundance recovered from the gonad and cultured in marine and TCBS agar were 1,21x107 y 2,83x 107 CFUs per gram of tissue, respectively. The TCBS agar counts indicated a similar abundance in both gut compartments (1.01 x 107CFUs per gram of tissue), while on the other hand, the counts from the foregut bacteria cultured in marine agar revealed a concentration of heterotrophic bacteria five times greater than the hindgut.

Table 2 Average bacterial abundance (CFUs/g) from gonad, foregut and hindgut cultured in marine and TCBS agar. 

Tejido Tissue Agar marino (UFC/g) Marine Agar (CFU/g) Agar TCBS (UFC/g) Agar TCBS (CFU/g)T
Intestino delgado Foregut 5.57*107 1.01*107
Intestino grueso Hindgut 1.10*107 1.00*107
Gónada Gonad 1.21*107 2.83*107

Analysis of PCR-TTGE profiles

The estimation of the bacterial community that were associated with gonad and gut tissues and between cultivable “C” and direct “D” fraction were evaluated through TTGE profiling of partial 16S rRNAgenes (Figure 2A and Figure 2B). The results of the banding profiles, evaluated using the program GelCompar II, revealed the presence of several migration patterns, these profiles revealed differences between the bacterial communities associated with each tissue and the “D” fraction samples exhibited more complex patterns and more diverse bands compared with those of the cultivable “C” fraction.

Figure 2 TTGE profile of V3-V6 region of 16S rRNA gene from cultivable and direct fraction gonad (A) and foregut, hindgut (B). The labels above the image indicate the samples origin. “G” for gonad, “F” for foregut and “H” for hindgut, “P” for pool and “I” for individual. The lanes labeled with “L” correspond to markers of 100 bp used as references. The codes above each lane indicate the bands selected for sequencing. 

From this gel analysis (Figure 2), 105 (69 of gut and 36 gonad) unique and common DNA bands were selected based on the band intensity. Referring to the gonad of Lobatus gigas, the G38 band showed a predominant population associated with individual and pooled samples from the direct fraction. The cultivable fraction was characterized by the presence of bands G1 and G8 in three pools and four individual samples (Figure 2A). The results of the DNA bands sequences are provided in Table 4.

The genetic profiles and the clustering analysis performed from direct and cultivable fraction of the gonad revealed two different groups according to the fraction, with 22.2% similarity for group A and 27.9% for group B (Figure 3A). Two subgroups were formed in group A from cultivable fraction samples, one of them with a higher than 60% similarity and the other with a similarity level of 50%. Group B contained the samples from the direct fractions (Figure 3A).

Figure 3 TTGE banding patterns cluster analysis from cultivable “C” and direct “D” fraction of individual “I” and pools “P” samples of gonad “G” (A) and foregut “F” and hindgut “H” (B) of L. gigas. The cluster analysis was performed with Gel compare II ® software using Dice coefficient and Unweighted - Pair Group Method with Arithmetic Mean (UPGMA) (Mohammadi and Prasanna, 2003). The letters from A to F and the numbers represent the groups and the sub-groups generated, respectively. 

On the other hand, the genetic profiles with the UPGMA method for the clusters analysis of foregut and hindgut showed six groups A, B, C, D, E and F with similarity percentages of 20.7, 45, 38.9, 33.1 and 44.6 %, respectively (Figure 3B). Group A is comprised of hindgut and foregut samples from both cultivable and direct fraction, however, two subgroups were also formed according to the fraction where the samples came from (Figure 3B, subgroup 1 and 2). Group B clustered two individual samples (DFI2 and DFI16) and one pool (DFP5) from the direct fraction of the foregut with a similarity percentage of 45%. Group C contained all samples from the direct fraction of the hindgut, with a similarity percentage higher than 50%. In cluster D the samples from cultivable fraction were grouped with similarity percentages of 38.9% and 44.6% respectively, and a high similarity percentage was observed in the bacterial population from the foregut. Furthermore, the samples from direct fractions were clustered mainly in the groups B, C and E, with a similarity of 45, 50 and 33.1%, respectively. Group E contained hindgut and foregut samples from direct fractions, with similarity levels higher than 33.1%.

Variations in bacterial communities on different tissue (G, F, and H) observed in the UPGMA analyses, were verified through a principal component analyses PCA (Figure 4).

Figure 4 (A) Principal Components Analysis (PCA) of individual samples and pools of the gonad “G” obtained from “C” cultivable and “D” direct fraction. (B) PCA of “G” gonad samples obtained from “C” cultivable and “D” direct fraction. (C) PCA to evaluate the differences between bacterial communities associated with “F” foregut and “H” hindgut of Lobatus gigas. (D). PCA of gut compartments obtained from “C” cultivable and “D” direct fraction. 

According to the binary matrix of data on the presence (1) and absence (0) of bands, 36 bands were detected on the gels of the gonad. A band-based binary presence/absence matrix was calculated by applying the Dice similarity coefficient and used for an analysis of similarity (ANOSIM) based on the Bray-Curtis coefficient, which enables significance testing of the data groups (Hammer et al., 2001). Significant differences were found between the samples associated with “C” cultivable and “D” direct fraction [R = 0.868 (Bray Curtis) and (Jaccard), (P < 0.004 in both analyses)]. Moreover, differences between bacterial community compositions of the cultivable fraction of individual samples and pools were demonstrated (Figure 4A). The Shannon Index (H`) analysis showed a difference between the bacterial communities associated with “C” cultivable and “D” direct fraction (Table 3). Also, the analysis showed that bacterial populations associated with individual gonad tissue samples are less related than those obtained from pools of gonad tissue and that this is true for both fractions.

Table 3 Shannon index (H`) calculated from the TTGE profiles of V3-V6 region of 16S rRNA for each tissue, each fraction. 

Índice de Shannon (H`) Shannon Index (H`)
Tejidos Tissues Fracción C C Fraction Fracción D D Fraction Fracciones C y D C and D Fractions
Intestino delgado (ID) / Foregut (F) 1.4 2.5 2.5
Intestino grueso (IG) / Hindgut (H) 1.7 1.7 2.4
ID e IG / F and H 2.2 2.7 2.9
Gónada / Gonad 1.6 1.9 2.3

Furthermore, the results of the analysis shownthat 69 bands were detected on the gels of the gut tissue. PCA observed in figure 4C and the analysis of similarity (ANOSIM), showed significant differences in the bacterial community associated with the foregut (F) and hindgut (H) [R=0.1406 and (Bray Curtis) and (Jaccard), (P < 0.0018 in both analyses)]. The differences among bacterial communities related to cultivable and direct fraction from each compartment is observed in the PCA (Figure 4D). Significant differences were found between the samples associated with “C” cultivable and “D” direct fraction of the foregut (F) [R = 0.3479 (Jaccard) and R= 0.3479 (Bray-Curtis), (P value < 0.0024 in both analyses)], and hindgut (H)[R = 0.4643 (Jaccard) and R= 0.4643 (Bray Curtis), (P value < 0.0003) ].

The H` values calculated from G, F, and H (taking into account C and D fraction) revealed that the foregut has the most diverse bacterial community while the gonad has the least diverse. When H` was calculated for each tissue and for C and D separately, it was possible observe that the diversity is higher in D fraction than the C fraction, except for hindgut tissue which kept its diversity for both approaches (Table 3).

Sequencing and phylogenetic analysis of the bands (≈400 bp) excised from TTGE gel revealed several different phylogenetic groups for each sampling tissue (gonad, foregut and hindgut) according to their similarity with 16S rRNA gene sequences held in public databases (Table 4, Figure 5). These belonged to Alphaproteobacteria, Gammaproteobacteria and Bacilli (Table 4, Figure 5A) for the gonad, and the most common species was Ralstonia pikettii, which was observed in both cultivable and direct fraction (Figure 4A). In the gut compartments, the results of partial sequences of the 16S rRNA gene revealed high similarity with bacteria belonging to Alphaproteobacteria (12.5%), Betaproteobacteria (12.5%), Gammaproteobacteria (12.5%), Bacilli (31.25%), Clostridia (6.25%), Actinobacteria (6.25%), Mollicutes (6.25%) and Deinococci (6.25%) and unclassified bacteria (Table 4, Figure 5B).

Table 4 Taxonomic affiliations of DNA partial sequences obtained from TTGE bands of L. gigas gonad, foregut, and hindgut. 

Origen Origin Banda TTGE TTGE Band Nº de acceso Accession N° Porcentaje de similitud (BlastN) Percent Similarity (BlastN) Filo Affiliation Phylum Clase Class Secuencia más cercana Closest sequence
DIDI6 / DFI6 4F KX891442 93% α-Proteobacteria Oceanicola marinus. NR_043969.1
DIDP1 / DFP1 9F KX891443 98% β-Proteobacteria Undibacterium oligocarboniphilum. NR_117348.1
DIDI3 / DFI3 3F KX891440 97% Proteobacteria
DIGP3 / DPH3 Clon 38H KX891439 98%
CIGP5 / CHP5 AH KX891445 99% γ-Proteobacteria Pseudomonas azotoformans. NR_113600.1
DIGI16 / DHI16 Clon 31H KX891438 99% Pantoea stewartii. NR_104928.1
DIDP3 / DFP3 Clon 18F KX891432 95% Firmicutes Clostridia Clostridium straminisolvens. NR_024829.1
DIDP4 / DFP4 21F KX891442 100% Bacilli Streptococcus sanguinis. NR_113260.1
DIGI6 / DHI6 29H KX891431 100% Streptococcus sanguinis. NR_074974.1
DIGI3 / DHI3 Clon 28H KX891436 100% Streptococcus mitis. NR_116207.1
CIDI2 / CFI2 ZF KX891444 96% Anoxibacillus amylolyticus. NR_042225.1
DIDI2 / DFI2 2F KX891437 100% Bacillus cereus. NR_074540.1
DIGI2 / DHI2 Clon 24H KX891434 99% Actinobacteria Actinobacteria Propionibacterium acnes. NR_040847.1
26H KX891435 88% Deinococcus- Thermus Deinococci Deinococcus geothermalis. NR_074342.1
DIGP5 / DHP5 42H KX891441 88% Tenericutes Mollicutes Mycoplasma neophronis. NR_108494.1
DIDP1 / DFP1 Clon 10F Secuencia de la quimera 93% Spirochaetae Spirochaetia Spirochaeta litoralis. NR_104732.1
CGI4 / CGI4 G1 KX891446 96% Ralstonia pickettii. NR_043152.1
96% Ralstonia pickettii. NR_102967.1
G8 KX891447 96% β-Proteobacteria Ralstonia pickettii. NR_043152.1
96% Ralstonia mannitolilytica. NR_025385.1
DGP5 / DGP5 Clon G38 KX886796 99% Proteobacteria Ralstonia pickettii. NR_043152.1
98% Ralstonia mannitolilytica. NR_025385.1
CGP5 / CGP5 G22 KX891450 95% α-Proteobacteria Roseomonas aquatica. NR_042501.1
94% Roseomonas vinacea. NR_044191.1
CGP4 / CGP4 G21 KX886797 83% Firmicutes Bacilli Bacillus gotheilli. NR_108491.1
83% Bacillus foraminis. NR_042274.1
DGP2 / DGP2 G32 KX891449 94% Bacillus lichenoformis. NR_074923.1
94% Bacillus shackletonii. NR_025373.1

Figure 5 16S rRNA gene-based dendogram showing the phylogenetic relationships of TTGE bands obtained from the total DNA samples from the “G” gonad (A) and “F” foregut, “H” hindgut (B) of L. gigas. 

DISCUSSION

L. gigas microbiota from the Colombian Caribbean has been characterized by recent studies, with the goal of study the bacterial diversity associated with both wild and captive conchs (Pérez et al., 2014; Acosta et al., 2009; Rodriguez et al., 2011). In this study we conduct one of the first exploratory investigations into consideration the different tissues to observe the dominant bacterial community in of L. gigas. We analyzed the bacterial composition of the gonad and the gut (foregut and hindgut) of the wild queen conch from the Seaflower biosphere reserve by using different microbiological and molecular methods. When we compared the cultured bacteria and a molecular approach based on analysis of DNA extract direct from the sample seems to be a complementary strategy to determine the principal components in the microbial community of queen conch.

The analysis performed using conventional culture techniques, such as counting CFUs in marine and TCBS agar, allowed us to establish the presence of heterotrophic bacteria and bacteria belonging to the family Vibrionacea in the gonad and in the gut compartments of the queen conch. The TCBS agar count suggests an abundance of vibrios with a total of 2.83 x 107 CFUs per gram in gonad tissue and 1.01 x 107 CFUs per gram of tissue for the gut compartments analyzed; nevertheless, to date no studies have described the gonad microbiota of the conch. Previously, Avendaño-Herrera et al. (2001) reported a load of heterotrophic bacteria in the bivalve Argopecten purpuratus gonad in captivity; they counted 4 x 104 CFUs per gram and a Vibrionacea count of 3 x 102 CFUs per gram. Our results contrast with those describe by Avendaño-Herrera et al. (2001) and could be due to the captivity conditions under which the A. purpuratus studies were performed. The affinity of vibrios to the gonad tissue has been previously described in the mollusk Crassostrea gigas and it has been related to episodes of mortality of this species (De Decker et al., 2011). Studies aimed at establishing the identity of vibrios associated with the gonad, foregut and hindgut and its possible effects on mollusk development are necessary. Also, the bacteria belonging to the family Enterobacteriaceae and the genera Aeromonas and Pseudomonas can develop in the TCBS medium, and they could interfere with bacterial counts, generating an overestimation in the total vibrio abundance present in the tissue analyzed.

The analysis of TTGE profiles showed similar bacterial communities among the individual and pool samples in the gonad tissue (Figure 2A), with dominance of two or three bands per profile. These results could be related with the autochthonous bacteria of this mollusk tissue. The differences in the banding profiles observed between individual and pooled samples from gonad, foregut and hindgut were verified with a PCA. The analysis highly suggests that there is a greater difference in bacterial populations associated with individual samples than those associated with pooled samples from both fractions (C and D) (Figure 3). Similar results have been reported in different studies with marine organisms. In these studies individual variation was observed even among fish raised under similar environmental conditions, with similar genetic background, and fed the same diet, emphasizing the influence of the host on bacterial diversity (Spor et al., 2011; McKnite et al., 2012; Navarrete et al., 2012). Besides this, banding patterns from pools were similar in some samples, suggesting the presence of a dominant bacterial population in these tissues. However, these results could lead to biased conclusions, due to the possibility that the dominant bacteria present in one individual could be interpreted as common bacteria for all the individuals analyzed in the sample (Reveco et al., 2014).

Meanwhile, the results revealed the presence of a more complex and diverse community associated with the conch foregut when compared to the hindgut. Important shifts in the bacterial community structure were also observed between cultivable fraction (C) and direct fraction (D). Significant differences between both gut compartments were found with an average percentage of dissimilarity of 94.41%. These results were verified with the principal components analysis presented in figure 4C, which suggests that the foregut and hindgut have been colonized by different bacterial communities, this fact was supported with diversity index (H`), which showed a higher diversity in foregut than in hindgut (Table 3). These differences could be associated with metabolic processes carried out specifically in each compartment. Studies on fish suggest that variation in microbial diversity among gut compartments could be due to differences in pH and protease activity in the lumen (Yu et al., 2007; Yúfera and Darías, 2007). Furthermore, differences in the pH of the Epinephelus coioides fish stomach, pyloric caeca and intestine have been reported (Yu et al., 2007). These physicochemical parameters might be associated with the changes observed in the microbiota of different gut compartments, which could be related to specific functions in each gut and the ecological conditions of the habitat where L. gigas feeds.

According to the phylogenetic analysis, sequences from the gonad, indicated that the bacterial community has been colonized mainly by bacteria belonging to the genus Ralstonia, which were represented by several TTGE bands in the direct and cultivable fraction of pooled and individual samples (Figure 2A, Table 4). This finding suggests that the gonad could be a site of bacteria symbiosis in the queen conch. Although little is known about gonad microbiota in marine organisms and its interaction with the host, previous studies of the microbiome in mammals have demonstrated that in the reproductive system there is a mammal-bacteria symbiosis and this interaction may help defend the host against infectious diseases and improve its reproductive health (Miller et al., 2017; Kindinger et al., 2017).

In the case of the foregut and hindgut, phylogenetic analysis revealed differences between the two compartments (Table 4 and Figure 5B). The species found in both compartments belong to the phylum Firmicutes and Actinobacteria, while the phylum Proteobacteria, Deinococcus - Thermus, and Tenericutes were only found in the conch hindgut. These results are consistent with those reported in gut samples of the queen conch from Islas del Rosario (Acosta et al., 2009; Pérez et al., 2014) and other marine organisms (Sun et al., 2011; Trabal et al., 2012; Chaiyapechara et al., 2012). Although there are differences in the methodologies employed, the results are related to a high bacterial diversity associated with this mollusk.

Bacteria of the genus Bacillus and Propionibacterium have been characterized as probiotics in fish and marine crustaceans (Nakagawa et al., 2007; Lan et al et al., 2007; Cousin et al., 2010). More specifically, the genus Bacillus has been established as a biological agent against Aeromonas hydrophila (Lalloo et al., 2007, 2008) and the members of this genus produce a wide range of antagonistic compounds that have been found in the gut tract of crustaceans and fish (Balcázar et al., 2006). The presence of Pseudomonas in the foregut of the queen conch coincides with the results reported in other mollusks and in different marine organisms such as crustaceans and fish (Sfanos et al., 2005; Romero and Navarrete, 2006; Trabal et al., 2012). It is important to notice that members of this genus isolated from marine organisms have been recognized for their antibiotic activity against a broad spectrum of microorganisms (Gram, 1993; Jayatilake et al., 1996; Zheng et al., 2005). These results suggest that the presence of Pseudomonas libanensis could be related to beneficial effects and it could contribute to the survival of L. gigas by defending it against potential pathogens that may affect its health and development. Meanwhile, the genus Streptococcus (Table 4 and Figure 5B) has been associated with several diseases in fish and crustaceans, and it has been identified as the causative agent of the slow development disease in the crustacean Carcinus mediterraneus (Pappalardo and Boemare, 1982). The genus Undibacterium and Oceanicola found in the foregut and hindgut of the queen conch (Table 4) has also been recently reported in the gut of wild crustaceans Penaeus monodon samples (Rungrassamee et al., 2014). It is remarkable that Oceanicola marinus is a common specie in marine environments (Lin et al., 2007; Chunyan et al., 2012).

Differences between the results obtained from cultivable and direct fraction analysis were observed in the conch gonad, foregut and hindgut. It is possible that these differences are due to the culture dependent techniques which favor the growth of bacteria able to grow on synthetic media, regardless whether these bacteria are dominant in the sample. It has been shown that these bacteria represent only 1% of the total population, generating an underestimation of the actual microbial population diversity (Hansen and Olafsen, 1999; Suau et al., 1999). Further studies based on culture-independent methods have demonstrated a less biased picture of the bacterial population present in environmental samples than culture-dependent methods (Amann et al., 1995). Nonetheless, it is necessary to take into account that techniques based on ribosomal genes may be affected by low concentrations of DNA from different species or the presence of high concentrations of DNA that are competing during PCR reactions (Ogier et al., 2002), which can affect the detection of some species in the community (Kisand and Wikner, 2003). That is why; it is necessary to implement both approaches (culture-dependent and culture-independent) in the study of microbiota from natural samples, in order to recover as much information as possible about the bacterial species present in the samples analyzed (Kisand and Wikner, 2003).

This research established the bacterial community associated with gonad and foregut and hindgut of L. gigas. Regarding the gonad, the results suggest that this organ has a stable microbiota that is mainly dominated by members of the genus Ralstonia that could be endosymbionts of this tissue. Additionally, the presence of vibrios should be verified and it is necessary to determine their effect on the gonad. But, more comprehensive studies are required to determine the role played by this bacterial community in the health and the reproductive cycle of the L. gigas. In the case of the foregut and hindgut, phylogenetic analysis revealed differences between the two compartments. However, to establish the effect of these communities on the different compartments of the conch, further investigations are necessary. Overall, the findings reported here could be used for the development of strategies to increase nutrition, prevent pathogens and contribute to the conservation of this vulnerable species from the Seaflower Biosphere Reserve in the Colombian Caribbean.

ACKNOWLEDGMENTS

We thank to the Universidad Nacional de Colombia and Secretaría de Agricultura y Pesca del Archipiélago de San Andrés, Providencia y Santa Catalina, Isla de San Andrés, through the projects 20101009144 and 201010011106, for their financial support. According to the Colombian legislative framework of the Environmental Ministry of Colombia, this project is subscribed to “Permiso Marco No. 0255” granted by the Autoridad Nacional de Licencias Ambientales (ANLA) to the Universidad Nacional de Colombia on March 14, 2014. Special thanks to Erick Castro, Jaisón Cuartas and Luz Pineda for all technical support provided and PhD Gloria Cadavid for her unconditional support.

AGRADECIMIENTOS

Agradecimientos a la Universidad Nacional de Colombia y a la Secretaría de Agricultura y Pesca del Archipiélago de San Andrés, Providencia y Santa Catalina, Isla de San Andrés, a través de los proyectos 20101009144 y 201010011106, por su apoyo financiero. De acuerdo con el marco legislativo colombiano del Ministerio del Ambiente de Colombia, este proyecto está suscrito a "Permiso Macro No. 0255" otorgado por la Autoridad Nacional de Licencias Ambientales (ANLA) a la Universidad Nacional de Colombia el 14 de marzo de 2014. Agradecimiento especial a Erick Castro, Jaisón Cuartas y Luz Pineda por todo el apoyo técnico proporcionado y la doctora Gloria Cadavid por su apoyo incondicional.

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Received: October 05, 2017; Accepted: July 09, 2018

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