Download Microbial Ecology

Microb Ecol (2012) 64:555–569
DOI 10.1007/s00248-012-0039-5
Molecular Analysis of Bacterial Microbiota Associated
with Oysters (Crassostrea gigas and Crassostrea corteziensis)
in Different Growth Phases at Two Cultivation Sites
Natalia Trabal & José M. Mazón-Suástegui &
Ricardo Vázquez-Juárez & Felipe Asencio-Valle &
Enrique Morales-Bojórquez & Jaime Romero
Received: 6 December 2011 / Accepted: 6 March 2012 / Published online: 27 March 2012
# Springer Science+Business Media, LLC 2012
Abstract Microbiota presumably plays an essential role in
inhibiting pathogen colonization and in the maintenance of
health in oysters, but limited data exist concerning their different growth phases and conditions. We analyzed the bacterial microbiota composition of two commercial oysters:
Crassostrea gigas and Crassostrea corteziensis. Differences
in microbiota were assayed in three growth phases: postlarvae at the hatchery, juvenile, and adult at two grow-out
cultivation sites. Variations in the microbiota were assessed by
PCR analysis of the 16S rRNA gene in DNA extracted from
depurated oysters. Restriction fragment length polymorphism
(RFLP) profiles were studied using Dice’s similarity coefficient (Cs) and statistical principal component analysis (PCA).
The microbiota composition was determined by sequencing
temperature gradient gel electrophoresis (TGGE) bands. The
RFLP analysis of post-larvae revealed homology in the microbiota of both oyster species (Cs>88 %). Dice and PCA
analyses of C. corteziensis but not C. gigas showed differences in the microbiota according to the cultivation sites. The
sequencing analysis revealed low bacterial diversity (primarily β-Proteobacteria, Firmicutes, and Spirochaetes), with
N. Trabal : J. M. Mazón-Suástegui : R. Vázquez-Juárez (*) :
F. Asencio-Valle : E. Morales-Bojórquez
Centro de Investigaciones Biológicas del Noroeste, (CIBNOR),
Mar Bermejo 195, Col. Playa Palo de Santa Rita,
La Paz, Baja California Sur 23095, Mexico
e-mail: [email protected]
N. Trabal
e-mail: [email protected]
J. Romero
Laboratorio de Biotecnología,
Instituto de Nutrición y Tecnología de los Alimentos,
Universidad de Chile,
El Líbano 5524,
Macul, Santiago, Chile
Burkholderia cepacia being the most abundant bacteria in
both oyster species. This study provides the first description
of the microbiota in C. corteziensis, which was shown to be
influenced by cultivation site conditions. During early growth,
we observed that B. cepacia colonized and remained strongly
associated with the two oysters, probably in a symbiotic host–
bacteria relationship. This association was maintained in the
three growth phases and was not altered by environmental
conditions or the management of the oysters at the grow-out
site.
Introduction
Oysters are the most produced and most valuable bivalve
mollusks [23]. However, one of the main problems in the
aquaculture of oysters is the repetitive episodes of mortality,
which seriously reduces production. These outbreaks of
disease affect the larval and post-larval stages in hatcheries
and also affect juveniles and adults cultured in the natural
environment. In some cases, studies have demonstrated diseases of bacterial etiology, including vibrioses nocardosis
and infections by Chlamydia-, Mycoplasma-, and Rickettsia-like organisms [7, 39, 59, 65, 68, 75]. Because oysters
may concentrate pathogenic microorganisms, the microbiota
of oysters have been examined mainly from the public
health perspective, and most of the studies have focused
on fecal contamination, enteric pathogens, and pathogenic
species of Vibrio [17, 34, 39, 54, 67, 75–77].
The association between aquatic invertebrates and gut
microbes is typically attributed to the ingestion of bacteria
[31, 62]. Research with aquatic organisms has suggested a
variety of beneficial roles for microbiota, including the
development of the host gastrointestinal tract, nutrition
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(providing vitamins, enzymes, and essential fatty acids for
the host), immune responses, and disease resistance [3, 19,
26, 31, 46, 56, 62, 64]. An increased susceptibility to infections may be related to the lack of the barrier provided by
microbiota, which compete with pathogenic microorganisms for nutrients and space in the intestinal tract [5, 46,
51, 52, 61] or produce substances that inhibit pathogens [26,
28, 61].
In recent years, the natural bacterial microbiota of different oysters (Crassostrea gigas [33], Crassostrea virginica
[40], Saccostrea glomerata [29], Ostrea chilensis [69], and
Crassostrea iredalei [54]) have been described. However,
these studies do not focus on the changes in microbiota that
may occur during the growth of the oysters. With the
exception of the study by Romero et al. [69], these reports
do not distinguish between resident and transient bacteria,
which lead to an overestimation of bacterial diversity within
the microbiota. Because oysters are filter feeders, their gills
are covered with mucus and vibrating cilia that facilitate gas
exchange for respiration and simultaneously trap suspended
particles, including bacteria and viruses [62]. In oysters, two
categories of normal bacterial microbiota can be described:
(1) indigenous or resident bacteria, including relatively
fixed types of organisms that remain relatively stable over
time and do not change with the particles ingested by the
host (even if the bacteria are disturbed, they tend to spontaneously recover), and (2) transient or non-indigenous microbiota, which are composed of potentially pathogenic
microorganisms that are ingested with the food, survive
passage through the gut (possibly proliferating in the gut),
and are voided with the feces [6, 31, 52]. During larval
development, transient microbiota rapidly become residents
of the oyster microbiota [8, 37, 51, 52, 62], although little is
known about the dynamics and stability of microbiota in the
juvenile and adult growth stages. It is, however, recognized
that the physiology of the invertebrate marine organism can
influence the gut microbiota [31, 62]. In bivalve mollusks,
bacteria have been reported in the style, gland of Deshayes,
esophagus, stomach, and intestine [31, 37, 51, 62]; however,
many other studies on the microbiota were performed on the
whole-oyster homogenate [31, 40, 63, 69]. Colonization by
bacteria in the oyster gastrointestinal tract has a particular
dependence on the external environment because of the flow
of water passing through the digestive tract [26, 31, 62].
Considering that ambient conditions and seasonality have
been shown to alter the gut bacteria community in some
aquatic invertebrates, it is probable that the habitat can influence the conditions in the gut and may therefore be an important factor in the composition of gut microbiota [26, 31, 62].
However, studies of the changes in microbiota under different
environmental conditions are lacking [40, 62, 63].
One of the most critical steps in the cultivation of these
oysters is the planting of post-larvae (spat) from production
N. Trabal et al.
at hatcheries to the field, where large numbers die [7, 11,
18]. Additionally, at high stocking densities in the field, the
identification of microbiota is important because the resident microbiota can be disturbed, and transient microorganisms subsequently exploit a temporary advantage [51, 52].
Productivity is related to larval transfer to the field and
environmental conditions that can alter the bacterial community associated with oysters.
During the past 20 years, oyster cultivation in Mexico has
improved; commercially important mollusks, such as the
Pacific oyster C. gigas (Thunberg, 1793) and the Cortez
oyster C. corteziensis (Hertlein, 1951), have been used. C.
gigas is the most cultivated and widely studied oyster
worldwide [30] and is the most prevalent species along the
Pacific coast; however, massive deaths have led to many
studies of the regional species C. corteziensis [11, 13,
47–49]. Presently, the microbiota composition of Cortez
oyster is unknown. In this study, using molecular techniques, we analyzed the composition of the bacterial microbiota of C. gigas and C. corteziensis. Variations and
differences in the bacterial microbiota composition were
assayed in three growth phases: post-larvae at the hatchery,
juvenile, and adult at two grow-out cultivation sites in the
field. In particular, we studied the variation of the microbiota composition when the post-larvae were planted at
different cultivation sites.
Methods
A synthetic diagram of the experimental design and methodology used in this study is shown in Fig. 1.
Oyster Collection and Experimental Design
Larvae of C. gigas and C. corteziensis were collected from
certified breeding adults following Office International des
Epizooties (OIE) standards. Hatchery procedures were followed as described by Mazón-Suástegui et al. [47–49]. To
obtain and culture embryos and larvae at Playa Eréndira
(PE) (24°14′09″N, 110°18′24″W), broodstock were conditioned for maturation under controlled temperature conditions and food supply in the hatchery prior to being induced
to spawn by thermal stimulation (Fig. 1). Swimming larvae
were fed with increasing concentrations of cultured Isochrysis galbana and Chaetoceros calcitrans microalgae
(25,000–60,000 cells ml–1) while decreasing the culture
density (10−1 larvae ml–1). Post-larvae (small and delicate
young oyster) were cultivated in upwelling recirculation
systems using the same microalgae diet mentioned above;
until they reach the optimal post-larvae planting size, these
post-larvae are known as spats (mean shell height 3–5 mm;
6 weeks of age after settling). In August 2009, a set of post-
Bacterial Microbiota in Two Commercial Oysters
557
Figure 1 Diagram of the experimental design used in this study
larvae was transferred (planting) to two grow-out cultivation
sites (Fig. 1). An initial culture of spats from the oysters in
the cultured site was performed in NestierTM-type trays
suspended from a floating long-line. The juvenile oysters
(3–5 cm in shell height; age 5 months after planting; Fig. 1)
were cultured in cages suspended on rigid structures near the
bottom. The adult oysters (6–12 cm in shell height; age
6–10 months after planting, Fig. 1) were cultured exclusively
at the bottom of the cages for improved stability and food
supply.
Two different grow-out cultivation locations were used:
Punta Botella (PB; 25°18′14.52″N, 112°04′35″W) at Bahía
Magdalena and Bahía de Topolobampo (BT; 25°54′N, 108°
33′W) (Fig. 2). Thirty samples of juveniles of C. gigas and
C. corteziensis were collected at both grow-out cultivation
areas in December 2009 and February 2010, respectively.
The adult oysters (n030 of each species) were collected in
April 2010 (Fig. 1). At this time, at both sites, gonad
maturation was not detected in the oysters. The samples
were transported on ice to the laboratory. On arrival, the
surface of the shells was scrubbed with filtered and
UV-sterilized seawater for epifauna elimination. Next, the
shells were washed with 70 % ethanol and were finally
depurated for 72 h in filtered and UV-sterilized seawater
with constant aeration according to the Scheduled Depuration Process [24]. Transient microbiota was removed from
the oysters following methods previously reported by Son
and Fleet [72] and Lee et al. [41]. After cleaning, the
juvenile and adult oysters were dissected in a sterile Petri
dish using one sterile scalpel blade for each oyster. After
discarding the intervalve liquid and adductor muscles, we
proceeded to dissect the organisms (juveniles and adults).
The tissues of the gastrointestinal tract, gills, and mantle
Figure 2 Collection sites for C. gigas and C. corteziensis specimens.
p-L0post-larvae; J0juvenile; A0adults. PE0hatchery production of
post-larvae at Playa Eréndira; PB0grow-out cultivation site at Punta
Botella; BT0grow-out cultivation site at Bahía Topolobampo
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N. Trabal et al.
were separated, and the gastrointestinal tract was frozen
at −20°C. Post-larval oysters (n030, for each species) were
vigorously rinsed in sterile seawater, and after the intervalve
liquid and adductor muscles were eliminated, the organisms
were frozen at −20°C. In all cases, at the time of dissection,
the oysters were alive, and the appearance of the bodies was
normal with no evidence of infection.
temperate Pacific environmental conditions that are
more suitable for C. gigas, which is a temperate water
species, whereas the subtropical conditions in parts of
the Gulf of California are more suitable for C. corteziensis, which is a subtropical species [13, 47, 48].
Environmental Conditions of the Grow-Out Cultivation
Area
The total tissue of the post-larvae was homogenized by lysis
using a buffer containing Tris–EDTA–SDS (100 mM NaCl,
50 mM Tris [pH 8], 100 mM EDTA [pH 8], sodium dodecyl
sulfate [SDS 1 %]) and 20 μl proteinase K (20 mg ml−1)
(Sigma, St. Louis, MO) and was incubated for 1 h at 65°C.
The DNA was subsequently extracted with phenol/chloroform and precipitated with ethanol, as previously described
[69].
The gastrointestinal tissues (cut into pieces) of juvenile
and adult oysters were lysed with 50 ml of Tris–EDTA–SDS
buffer (as previously described), and the DNA was extracted
using a QIAmp® DNA Mini kit (Qiagen, Valencia, CA)
following the manufacturer's instructions. A final purification was performed using the Wizard® SV Gel and PCR
Clean-up System (Promega, Madison, WI). The concentration and quality of the DNA were determined at A260 and
A280 nm using an ND-1000 spectrophotometer (NanoDrop
Technologies, Wilmington, DE).
The Punta Botella grow-out cultivation site (PB) is located
in Bahía Magdalena, which has one of the most diverse
ecosystems in the state of Baja California Sur. This location
has low pollution and minor human activity, and it is influenced by oceanic currents of temperate seawater from further south. In Bahía Magdalena, the large lagoon has an
irregularly shaped barrier island. There are numerous small
estuaries and shallow channels bordered by mangroves that
provide organic matter deposited as sediment. Mangroves
are the common feeding habitat for communities of many
vertebrate and invertebrate species and possess commercial
and ecological value [1, 12].
The Bahía Topolobampo grow-out site (BT) is a coastal
lagoon complex with relatively large mouths connecting to
the Gulf of California that is influenced by oceanic currents.
In the last several decades, BT has been supporting shrimp
and oyster aquaculture projects. However, the lagoon water
is contaminated by agricultural runoff and urban and industrial sewage effluents [57, 58].
The salinity at both sites ranged from 35 to 40 psu with
an average of 37 psu.
The weekly sea surface temperature (SST) data for 2009–
2010 were derived from the Moderate Resolution Imaging
Spectroradiometer-Aqua (MODIS-Aqua) satellite. The SST
data were downloaded from the National Aeronautics and
Space Administration web site (http://modis.gsfc.nasa.gov/
data/). The monthly SST was estimated along a straight line
centered at each one of the study areas, and the
corresponding mean was calculated. The following data
show the SST during grow-out of juveniles and adults at
both sites: Punta Botella (August, 26.6°C; September, 28.0°C;
October, 26.2°C; November, 25.5°C; December, 23.4°C;
January, 21.7°C; February, 20.6°C; March, 20.4°C; and April,
19.9°C) and Bahía Topolobampo (August, 27.9°C; September,
29.1°C; October, 26.7°C; November, 25.9°C; December,
24.6°C; January, 21.2°C; February, 19.7°C; March, 23.8°C;
and April, 25.5°C). Temperate temperatures were found at the
PB site, whereas the temperatures at the BT site were
subtropical. A temperature drop was also noted in
October, which indicated the beginning of autumn at
both sites. C. gigas grew faster at PB, and C. corteziensis grew faster at BT. The difference is based on
DNA Extraction and Purification
PCR Amplification of the 16S rRNA Gene and Analysis
of the Products
The total DNA was extracted from each oyster (total tissue
post-larvae and gastrointestinal tissues of juveniles and
adults), diluted in nuclease-free water to obtain a concentration of 50 ng μl−1, and used as a template in PCR. Nested
PCR of the V3–V5 region of the 16S rRNA gene was used
to obtain profiles of the bacterial communities present in
three different growth stages in both oyster species and at
the two grow-out cultivation sites. Eubacterial 16S rRNA
genes were amplified with the following primers: EUB 27 F
(5′-AGAGTTTGATCCTGGCTCAG-3′) and EUB1492R
(5′-GTTACCTTGTTACGACTT-3′) [43] in the first amplification and 341 F (5′-GCCTACGGGAGGCAGCAG-3′
with GC clamps at the 5′ end for temperature gradient gel
electrophoresis analysis, TGGE) [53] and 939R
(5′-CTTGTGCGGGCCCCCGTCAA TTC-3′) [70] for the
second phase. The primers used in the specific amplification
of the eubacterial 16S rRNA gene variable region were
selected from a set of primers available in the literature
and aligned in silico using the Sequence Match software
with reference sequences (1,921,179 16S rRNAs) from the
Ribosomal Database Project II Database (RDP II). The
criteria used to choose the best primers were as follows:
greater alignment with eubacteria domain sequences,
Bacterial Microbiota in Two Commercial Oysters
especially with the Proteobacteria group; lack of alignment
with Archaea domain sequences; and no cross-amplification
with the oyster genome. PCR was performed with a reaction
mixture (25 μl) containing 0.2 mM of each deoxynucleoside
triphosphate, 0.05 Uml−1 Platinum Taq DNA polymerase
(Invitrogen, San Diego, CA), 1× polymerase reaction buffer,
2 mM MgCl2, and 0.25 pmol/ml−1 of each primer. The
reaction mixtures were incubated in an Eppendorf Mastercycler (Eppendorf, Hamburg, Germany) with an initial
denaturation at 95°C for 10 min followed by 15 cycles at
95°C for 1 min 30 s, 55°C for 1 min 30 s, 72°C for 1 min
30 s, and a final elongation at 72°C for 10 min for the first
phase of the nested PCR. Subsequently, 1 μl of the amplified DNA was re-amplified with an initial denaturation at
94°C for 10 min followed by 20 cycles at 97°C for 1 min,
54°C for 1 min, 72°C for 1 min 30 s, and a final elongation
at 72°C for 10 min for second phase of the nested PCR. The
PCR products were analyzed using polyacrylamide gel electrophoresis and silver nitrate staining [20].
Restriction Fragment Length Polymorphism Analysis
of the 16S rRNA Gene
By applying restriction fragment length polymorphism
(RFLP) analysis to the PCR amplification products of the
16S rRNA gene, we sought to observe variations of the
resident microbiota in C. gigas and C. corteziensis in the
three growth stages at two different grow-out sites. The
products of the 16S rRNA gene amplification (10 μl) from
the DNA of post-larval and gastrointestinal tissues of juveniles and adults were digested for 2 h at 37°C with 1.5 U of
Alu1 restriction endonuclease (Invitrogen, San Diego, CA).
The resulting fragments were subsequently analyzed by
polyacrylamide gel electrophoresis and silver nitrate staining [20]. Each gel included BenchTop 100 bp DNA Ladder
markers (Promega) and standard bacterial RFLP controls
(Vibrio harveyi ATCC 14126, Listonella pelagia ATCC
25916) to validate the comparisons between the gels.
Analysis of the Variation of Microbiota Using RFLP
Patterns
The RFLP profiles were analyzed using software (GelCompar II® 5.2, Applied Maths, Austin, TX) and by applying the
Dice similarity index (Cs). The pairwise Cs is a similarity
index used to compare microbiotal variation in different
samples based on the presence or absence of bands in each
lane in the gels [50]. Thus, two identical profiles create a Cs
value of 100 %, whereas completely different profiles result
in a Cs value of 0 %. Each lane in the gels (samples from
individual oysters) can be compared with every other sample; therefore, the mean percent similarities (Cs values) were
used to compare the similarity of microbiota profiles in
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post-larvae, juveniles, and adults in the two grow-out sites
[50]. Using this index, microbiota profiles derived from
different species (C. gigas vs. C. corteziensis) and from
each individual of the same species were compared. The
cluster analysis results were displayed as a dendrogram.
UPGMA was applied with GelComparII software. The reliability was tested by calculating the co-phenetic correlations
of the nodes with tools available in the GelCompar program.
Analysis of Microbiota Composition Using 16S rRNA Gene
Temperature Gradient Gel Electrophoresis
Using TGGE analysis, we determined the composition of
the microbiota associated with the oysters. We chose individual samples that showed different RFLP profiles and
triplicate samples of organisms having the same pattern
from each group analyzed (the two species of post-larvae,
juveniles, and adults from Punta Botella and Bahía
Topolobampo). Products obtained from the nested PCR of
the 16S rRNA gene were separated by TGGE and analyzed
using the DCode System (Bio-Rad, Hercules, CA, USA) in
0.8 mm gels composed of 6 % [w/v] acrylamide, 0.1 %
bisacrylamide, 8 M urea, 20 % [v/v] formamide, and 2 %
[v/v] glycerol with 1× Tris–acetate (Tris 0.04 M, acetate
0.002 M, EDTA 0.001 M, pH 8.5) as the electrophoresis
buffer. The polymerization was catalyzed by 110 μl of N-NN′-N′-tetramethylethylenediamine and 80 μl of 10 %
ammonium persulfate solution added to 50 ml of the gel
solution. The gel was loaded with 6 μl of 341f-939r-GC
clamp PCR products and 1 μl of dye solution and run for
18 h 30 min at a fixed voltage of 65 V (±9 mA) with a
parallel temperature gradient ranging from 66 to 70°C. After
electrophoresis, the gels were stained for 1 h by incubation
with SYBR Green at 25±2°C (room temperature). Each gel
included standards containing PCR amplicons of known
bacterial sequences ( % GC) to validate comparisons
between gels [55]. The standard was composed of amplicons from the following bacteria: Pseudomona aeruginosa
P1 (%GC: 51.8), V. harveyi ATCC 14126 (%GC: 53.1),
Bacillus subtilus B4 (% GC: 54.6) and Citrobacter gillenii
(%GC: 56).
Sequence Analysis of the 16S rRNA Gene
The dominant bands were identified by their corresponding
intensity in each TGGE pattern. The bands were excised
from the gel and eluted for 12 h at 25°C in 50 μl MilliQ
water. After elution, 1 μl of DNA was used for
re-amplification, using the same conditions described
above. To test for the presence of similar amplicons, bands
showing the same migration in different lanes were checked
by RFLP using AluI (an enzyme that cuts frequently) [55].
Next, 16S rDNA from the re-amplified bands or from the
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bacterial isolates was purified using the Wizard® SV Gel
and PCR Clean-up System and sequenced (by Macrogen,
Rockville, MD) with an automatic sequencer (Applied Biosystems ABI3730XL, Carlsbad, CA).
The sequences were initially compared to the available databases using the basic Local Alignment Search
Tool network service [2] and aligned with reference
sequences using Sequence Match software from the
Ribosomal Database Project II (RDP II) website [15]
to determine the approximate phylogenetic affiliations.
The sequences were deposited in GenBank (accession
numbers JF522195–JF522232).
Statistical Analysis
We analyzed the differences in the bacterial communities
associated with C. gigas and C. corteziensis in three growth
phases (post-larvae, juvenile, and adult) between two growout cultivation sites using orthogonal empirical functions.
The differences were estimated using a multivariate analysis, namely, the principal component analysis (PCA). The
vectors computed from PCA are orthogonal and are uncorrelated, which prevents the regression on residuals [36]. The
PCA permitted the ranking and simplification of the new
variables, called principal components, and determination of
the total variation of the data Xj, thereby explaining the
variation with a few principal components (factor loadings≥0.7). Only the principal component was considered
to be statistically significant for eigenvalues (l)>1.0 [36].
PCA was performed from correlation matrices generated
from a binary matrix considering the presence/absence of
the RFLP patterns bands, which were expressed as a value
of the Dice similarity coefficient [25]. The PCA was conducted using the Statistica 8.0 software package (StatSoft
Inc., Tulsa, OK).
Results
Similarity of Bacterial Microbiota in the Post-larvae
of C. gigas and C. corteziensis
The similar RFLP profiles obtained from the cluster of postlarvae of C. corteziensis revealed a homology in the microbiota of this oyster with Cs>92 % (Table 1, Fig. 3a). C.
gigas, however, exhibited a lower similarity than did C.
corteziensis with Cs>72 % (Fig. 3a, Table 1). A comparison
of the patterns obtained from the post-larvae of both species
showed uniformity in the composition of the microbiota and
clustering with high similarity (cluster I, Cs>88 %). However, cluster II was observed with high similarity (Cs>
83 %), which partly corresponded to the post-larvae of C.
gigas (Fig. 3a).
N. Trabal et al.
Table 1 Summary of Dice similarity coefficients (percent) of the
RFLP profiles of the bacterial microbiota associated with oysters,
depending on the growth stage and two grow-out cultivation sites
Species
Growth stage
Site
H
C. corteziensis
PB
BT
87
75
81
55
Juvenile
53
57
Adult
Post-larvae
56
68
80
40
55
40
Post-larvae
92
Juvenile
Adult
C. gigas
Both speciesa
Post-larvae
Juvenile
Adult
72
88
H hatchery production of post-larvae at Playa Eréndira, PB grow-out
cultivation at Punta Botella, BT grow-out cultivation at Bahía
Topolobampo
a
Comparison of RFLP profiles of C. gigas and C. corteziensis
The PCA statistical method was used to compare the
bacterial microbiota variation in the three growth phases
(post-larvae, juvenile, and adult) of C. gigas and C. corteziensis at different grow-out cultivation sites, and the results
are presented as a tridimensional scatter plot in Fig. 4. The
results for the microbiota associated with the post-larvae in
both species of oyster are shown in Fig. 4a. The observed
post-larval cluster of C. gigas and C. corteziensis was
explained by three principal components (PC1 042.4 %
and l036.9, PC2019.8 % and l017.1, PC3010.2 % and
l08.8) with a 72.4 % cumulative variance. However, a
separated group of C. gigas post-larvae was also observed,
which agrees with the results of the overall cluster analysis
of RFLP patterns of post-larvae (Fig. 3a).
Variation in the Bacterial Microbiota of Juveniles
and Adults in Different Grow-Out Cultivation Sites
The microbiota composition in juveniles and adults of both
species at the two sites was studied by analyzing the patterns
obtained by RFLP (Table 1). The profiles obtained from
juvenile C. corteziensis indicated differences in the microbiota
composition when these oysters were raised at different sites
(Fig. 3b). Two groups were defined on the basis of the RFLP
profiles, which correlated with the C. corteziensis grow-out
site (Punta Botella cluster and Bahía Topolobampo cluster).
For the population at Bahía Topolobampo, the Cs was >82 %,
whereas at Punta Botella, the Cs was >87 %, which indicates
uniformity of the microbiota associated with each locality
with the exception of oyster JC6 (Fig. 3b, Table 1). For the
C. corteziensis adults, the similarity between samples was
higher at PB (Cs>75 %) than at BT (Cs>55 %) (Table 1).
Bacterial Microbiota in Two Commercial Oysters
561
Figure 3 Dendrogram
representing the similarity
(Dice coefficient) observed in
the RFLP profiles of rRNA
genes amplified from bacterial
microbiota associated with
oysters. Only representatives of
each cluster are shown. a
Comparisons between postlarvae C. gigas and C. corteziensis, and b comparisons
between juvenile samples of C.
corteziensis at two grow-out
cultivation sites. pL0postlarvae; J0juvenile; G0C.
gigas; C0C. corteziensis;
PB0cultivation at Punta
Botella; BT0cultivation at
Bahía Topolobampo
When this analysis was performed for juveniles and
adults of C. gigas, differences in the microbiota profiles of
individual samples were observed. In contrast to the results
for C. corteziensis, the dendrogram based on the Dice coefficient did not reveal an association between the microbiota
profile and the grow-out cultivation location (data not
shown). Juveniles and adults of C. gigas had lower Cs
values than C. corteziensis, which indicates variation within
the microbiota among individual oysters of C. gigas
(Table 1).The similarity in the microbiota associated with
juvenile or adult C. gigas and C. corteziensis was always
higher at PB than at BT (Table 1).
The PCA of juveniles and adults of C. corteziensis (Fig. 4b)
generated two clusters in relation to the grow-out cultivation
sites with a 73.6 % cumulative variance (PC1035.0 % and
l030.4, PC2023.2 % l020.1, PC3015.4 % l013.4). These
results confirmed the pattern observed in the dendrogram
estimated from the same samples (Fig. 3b) and the results
found by Dice coefficient analysis (Table 1). In contrast, the
PCA for juveniles and adults of C. gigas showed minor
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N. Trabal et al.
Figure 4 A 3D scatter plot of the principal component (PC) analysis
based on the RFLP of the rRNA gene fragments of C. gigas and C.
corteziensis in three growth stages and two grow-out cultivation sites. a
RFLP banding patterns of post-larvae C. corteziensis (filled circle) vs. C.
gigas (empty circle). b RFLP banding patterns of juveniles and adults of C.
corteziensis in two grow-out cultivation sites. Punta Botella (filled triangle)
and Bahia Topolobampo (empty triangle). c RFLP banding patterns of
juveniles and adults of C. gigas in two grow-out cultivation sites. Punta
Botella (filled triangle) and Bahia Topolobampo (empty triangle). d RFLP
banding patterns of juveniles of C. corteziensis (filled triangle) vs. C. gigas
from Punta Botella (filled diamond), juveniles of C. corteziensis (empty
triangle) vs. C. gigas (empty diamond) from Bahia Topolobampo
correlations between the RFLP profiles at the two different
grow-out cultivation sites (Fig. 4c). The results were similar to
those obtained with the Dice coefficient (Table 1).
The PCA for juveniles of C. gigas and C. corteziensis in the
same cultivation site (Fig. 4d) showed a clear dispersion of the
data, thereby supporting the results obtained with the Dice
coefficient analysis (Table 1), which did not suggest any similarities in the microbiota associated with these oyster species.
Analysis of the Stability of the Microbiota Associated
with the Oysters When the Post-larvae are Planted
at the Grow-Out Cultivation Sites
Using RFLP profiles, the effect of cultivation site transfer
upon microbiota associated with the post-larvae was studied
(Table 2). Strong similarities (Cs >80 %) in the RFLP profiles of post-larvae and juveniles of C. corteziensis were
Bacterial Microbiota in Two Commercial Oysters
563
Table 2 Summary of Dice similarity coefficients (percent) of RFLP
profiles of microbiota associated with post-larvae during planting
versus juveniles at two grow-out cultivation sites
Site
Species
Growth stages
C. corteziensis
Post-larvaea
C. gigas
Post-larvaea
a
PB
Juvenile
BT
Juvenile
80
40
54
65
Post-larvae planting grow-out sites
larvae at the grow-out site (Fig. 5). The PCA showed significant correlations between the RFLP profiles for postlarvae and the juveniles of C. corteziensis at Punta
Botella (PC1037.2 %, l032.4, PC2019.1 % l016.6,
PC3016.2 % l014.1); this correlation did not exist in
post-larvae planted at Bahía Topolobampo (Fig. 5a).
These results confirm the findings from the dendrogram
analysis using the Dice coefficient (Table 2). The PCA
for C. gigas showed considerable variation between the
profiles obtained for the post-larvae and the juveniles at
both cultivation sites (Fig. 5b).
PB cultivation at Punta Botella, BT cultivation at Bahía Topolobampo
Identification of Dominant Bacterial Composition
Based on TGGE Band Sequencing
found at Punta Botella, which suggests that the microbiota
composition was stable with little environmental influence
(Table 2). The microbiota of C. corteziensis in the Punta
Botella cultivation site remained stable, even throughout the
fattening process, as the RFLP profiles between juveniles
and adults was also similar (Cs>84 %, data not shown).
However, upon introduction into Bahía Topolobampo, the
microbiota of C. corteziensis was not observed to be stable
(Table 2).
Similar Dice values (Cs >40 % at PB, Cs >65 % at BT)
showed that the microbiota of C. gigas undergo variations
during the planting of the post-larvae (Table 2).
Using PCA, we studied the variation in the microbiota
associated with the oysters when we introduced the post-
The sequences of the bands detected visually by TGGE
were identified for both oyster species in different growth
stages and at the two cultivation sites. The partial sequences
of the 16S rRNA gene (400–600 bp) were compared with
sequences available in RDP II and GenBank (Table 3). Our
results indicate that the microorganisms represented by the
main bands (OTUs) belong to the phyla Proteobacteria and
Firmicutes. Specifically, these OTUs were related to gramnegative bacteria from the genera Burkhordelia and Borrelia
and to gram-positive bacteria such as Bacillus and Propionibacterium. Unique bands corresponding to genera Ruegeria, Paracoccus, Methylobacterium, and Vibrio were also
observed. The most frequent OTU in both species was
Figure 5 A 3D scatter plot of principal component (PC) analysis
based on the RFLP analysis of rRNA gene fragments of C. gigas and
C. corteziensis of post-larvae vs. juveniles in two grow-out cultivation
sites. a RFLP banding patterns of C. corteziensis post-larvae (filled
circle) vs. juveniles from Punta Botella (filled triangle) and, Bahía
Topolobampo (empty triangle). b RFLP banding patterns of C. gigas
post-larvae (empty circle) vs. juveniles from Punta Botella (filled
diamond) and Bahía Topolobampo (empty diamond)
564
N. Trabal et al.
Table 3 Nearest-match identification of 16S rRNA gene sequences obtained by PCR-TGGE from known bacterial sequences in the RDP II and
GenBank databases
Accession number
OTU
Class affiliation
Identity (%)
Closest relative
Growth stage
N/Total Sample
Site
Burkholderia cepacia
Post-larvae
8/8
H
Borrelia sp.
Juvenile
4/7
PB
Crassostrea corteziensis
β-Proteobacteria
JF522209
B2
JF522195
JF522210
A1
B1
JF522206
JC13E
JF522205
JF522208
JC13F
JC6A
JF522207
JC13A
β-Proteobacteria
100
Burkholderia cepacia
Juvenile
7/7
PB
JF522232
AC9B
β-Proteobacteria
100
Burkholderia cepacia
Adult
6/6
PB
JF522231
JF522203
AC9F
JC24C
Actinobacteria
99
Propionibacterium sp.
Juvenile
2/7
BT
JF522204
JC24 A
JF522202
JC31A
JF522201
JC33 A
β-Proteobacteria
100
Burkholderia cepacia
Juvenile
7/7
BT
JF522200
JF522229
JF522230
JC33B
AC28B
AC28C
β-Proteobacteria
100
Burkholderia cepacia
Adult
7/7
BT
JF522228
Crasssotrea gigas
JF522197
Spirochaetes
100
85.4
AC29A
SG2D
Bacilli
Bacillus subtilis
Post-larvae
3/8
H
JF522213
JF522196
SG22B
SG30G
β-Proteobacteria
α-Proteobacteria
100
98
89
Burkholderia cepacia
Ruegeria sp.
Post-larvae
Post-larvae
8/8
1/8
H
H
JF522199
JF522198
JF522227
JG5C
JG6F
JG9A
Bacilli
α-Proteobacteria
β-Proteobacteria
96
98
100
Bacillus sp.
Paracoccus sp.
Burkholderia cepacia
Juvenile
Juvenile
Juvenile
4/7
1/7
7/7
PB
PB
PB
JF522226
JF522223
JF522220
AG9B
AG9F
AG10B
JF522218
JF522217
JF522216
AG12A
AG12P
AG14P
β-Proteobacteria
100
Burkholderia cepacia
Adult
6/6
PB
JF522225
JF522222
JF522219
AG9D
AG9I
AG11I
α-Proteobacteria
100
Methylobacterium sp.
Adult
1/6
PB
Bacilli
100
Bacillus subtilis
Adult
2/6
PB
JF522215
JF522214
JF522211
JF522212
JG22A
AG22C
AG30B
AG26H
β-Proteobacteria
100
Burkholderia cepacia
Juvenile
6/6
BT
β-Proteobacteria
γ-Proteobacteria
100
100
Burkholderia cepacia
Vibrio sp.
Adult
Adult
6/6
1/6
BT
BT
N Number of oysters presenting the closest relative
OTU operational taxonomic unit/number total of oyster samples, H hatchery production of post-larvae at Playa Eréndira, PB cultivation at Punta
Botella, BT cultivation at Bahía Topolobampo
Burkholderia cepacia, which was found during all growth
stages and at both sites. A particular bacterium (probably
Bacillus subtilis or another in the genus Bacillus) was a
component of C. gigas throughout its growth at Punta Botella.
In most cases, the bacterial diversity was higher in C. gigas
than in C. corteziensis, especially in adults (Table y3).
Discussion
The composition of microbiota associated with oysters can
be influenced by numerous factors, including particular
characteristics of the host, diet, and environmental conditions. In this study, bacterial microbiota of depurated C.
Bacterial Microbiota in Two Commercial Oysters
gigas and C. corteziensis oysters were compared using the
RFLP molecular technique to analyze the microbiota variation in three growth stages of the oysters and at two growout cultivation sites. In parallel, the microbiota composition
in these two species of oysters was also determined using
the TGGE method.
At the post-larval stage, the microbiota presented a uniform composition that was independent of the host (C. gigas
or C. corteziensis) with similar RFLP profiles and formed a
group when analyzed by PCA. During the cultivation of the
larvae at the hatchery, the bacteria associated with the food,
which influence the composition of the microbiota, were
controlled in the laboratory [4, 5, 26, 31, 74]. All postlarvae were produced in the same hatchery, where they were
fed a mixture of microalgae. Generally, the microalgae
cultivated in shellfish hatcheries are not free of bacteria
and are a highly nutritive product, that is, notably useful as
oyster food [38, 39]. Thus, when oysters are fed microalgae,
the associated bacteria may colonize the gastrointestinal
tract and become part of the microbiota of the oyster species.
It has been reported that bivalve mollusks, which have
similar diets, provide similar gastrointestinal environments
for bacterial colonization [4, 27, 31, 66]; this condition may
explain the similarity found in the composition of the microbiota in C. gigas and C. corteziensis.
There is evidence that certain aquatic invertebrates, particularly oysters, maintain a permanent microbiota in the
gastrointestinal tract [31, 33, 40, 62, 69, 74]. Several studies
have revealed that the bacterial populations isolated from
the gut differ in species composition from those isolated
from the habitat [31, 40, 62, 63]. Other studies have indicated that ambient conditions significantly affect the microbiota in marine invertebrates; oysters of the same species
caught in the wild and laboratory-reared animals may have
different bacterial communities in the gastrointestinal tract
[31, 40, 62, 75]. During the cultivation of marine organisms,
changes in environmental parameters, such as salinity, temperature [17, 34], nutrients, availability of food, and operational design in grow-out and management methods, may
induce stress in the animals [38] and influence the composition of the gastrointestinal microbiota. Our results show
that C. corteziensis microbiota composition was influenced
by the conditions presented at the cultivation site. The
results obtained by RFLP profile analysis and PCA show
that the associated microbiota vary depending on where C.
corteziensis was fattened (differences were observed between Punta Botella and Bahía Topolobampo). It is noteworthy that the microbiota composition of C. corteziensis
remained stable from the post-larvae stage to the growth of
the oyster when the oysters were planted at Punta Botella,
which was not observed at Bahía Topolobampo. This result
was somewhat surprising because the subtropical temperature observed at Bahía Topolobampo promotes better
565
growth in C. corteziensis [13, 47, 48]. There is limited data
about the environmental parameters of the field site used in
this study. Thus, we opted to report the temperature ranges,
but these data are not sufficient to establish a relationship
with the microbiota composition. It will be interesting to
evaluate the impact of environmental parameters on microbiota composition. The differences found in the stability of
the microbiota in C. corteziensis at Punta Botella and Bahía
Topolobampo may be the result of the characteristics of each
cultivation site. In particular, the environmental conditions
and grow-out management at Punta Botella reflect low
pollution and little anthropogenic activity, which support
the stability of the associated microbiota. In contrast, Bahía
Topolobampo is a highly impacted site, where the lagoon
water is contaminated by agricultural runoff and urban and
industrial sewage effluents [57, 58].
The difference in the filtering capacity of the two species
is another factor that may explain the differences in microbiota [31, 62]. C. gigas has a higher filtration capacity than
native oysters, and the capacity increases further when food
is abundant; thus these oysters are typically larger in size
[11, 22]. A difference in filtration capacity might explain the
greater diversity of bacteria in C. gigas because this oyster
has a greater capacity to ingest bacteria during respiration
and feeding.
Despite the differences in behavior among microbiota
associated with oysters in different grow-out sites, we found
a common bacterial component associated with C. corteziensis and C. gigas. This bacterium was a member of the
Burkholderia genus, was acquired in the post-larval stage,
and remained associated with the gastrointestinal tract in
juvenile and adult oysters at both cultivation sites. Presumably, the changes during the different growth stages of the
oysters result in changes in the environment of the microbiota associated with the gastrointestinal tract [8, 31, 37,
62]. Changes in the composition of the microbiota were
mainly observed in C. gigas in the three growing phases
included in this study and at the two cultivation sites. These
changes are probably related to a particular feature of the
host [31]. However, our results show that at least part of the
microbiota remains strongly associated with the gastrointestinal tracts of C. corteziensis and C. gigas, presumably
because of attributes of the bacteria (such as adhesion to
the gut wall) that prevent expulsion from the gut. In a
previous report, B. cepacia was found in the three growth
stages and was not influenced by the conditions at the
cultivation site, which could indicate a symbiotic association between this bacterium and the host (i.e., the two oyster
species). It would be interesting to demonstrate positive
relationships between these host oysters and B. cepacia
and to determine whether the bacteria play an important role
in the physiological processes of C. gigas and C. corteziensis. The genus Burkholderia was originally classified with
566
Pseudomonas [44, 47, 78]. In the past two decades, Burkholderia has been found in a variety of ecological niches,
including soil [21], plants [14, 44, 71], and freshwater and
marine habitats [16]. Recently, a strain was isolated from C.
corteziensis and identified as B. cepacia. This strain had
been isolated from the digestive gland of native C. corteziensis from the State of Nayarit, Mexico [9, 10].
Another common bacterium in the post-larvae of C. gigas
was Bacillus sp., which was also present in juveniles and
adults at Punta Botella. Bacillus spp. have been reported in
oysters, especially in Saccrostrea glomerata [29] and Ostrea
edulis [63]. Hernández and Olmos [33] reported findings
similar to ours, and they explained that Bacillus spp. adheres
to the gills of C. gigas at the cultivation site in the west coast
of Mexico. It has been reported that some species of Bacillus may have potential as a probiotic in cultivated marine
animals because it can colonize marine organisms and have
an antagonistic effect on pathogenic microorganisms. In
addition, the spores of these bacteria can be easily introduced into the food supply of the oysters [26, 52].
Several OTUs were found in C. corteziensis sampled at
the Punta Botella cultivation site and were recognized as
bacteria of the family Spirochaetaceae. The biodiversity of
spirochetes is high, and they are present in large numbers in
the gastrointestinal tract of humans, mammals, insects, and
bivalves (especially oysters), as well as in aquatic and marine environments [45, 60, 73]. These bacteria seem to be
limited to the crystalline styles (mucoproteinaceous rod of
the digestive gland) [45, 73], but the hypothesis that styleassociated spirochetes play a major role in the extracellular
enzymatic digestion processes of bivalves has not been
tested [35]. Recently, Husmann et al. [35] investigated the
phylogeny of spirochete groups present in crystalline styles
from select bivalves at different habitats and found that
spirochetes are not randomly distributed, which implies an
association between oyster species and specific spirochete
clusters. However, the spirochetes were present in only a
portion of the crystalline styles investigated; therefore, the
authors concluded that spirochetes are not essential symbionts for these bivalves. All of the identified spirochetes
were grouped into two families: Spirochaetaceae, which
included two genera (Cristispira and Spirochaeta), and
Brachyspiraceae, with one genus (Brachyspira) [35]. Similarity analyses of the OTU found in this study using databases and GenBank RPDII revealed an 85 % sequence
similarity with Borrelia spp. This result is similar to the
results of Green & Barnes [29] for microbiota in the digestive gland of the Sydney rock oyster. In this study, we
demonstrate that molecular biology techniques may clarify
differences in bacterial species associated with an oyster
species, as most spirochetes cannot be cultured.
The α-, β-, and γ-Proteobacteria, Firmicutes, Spirochaetes, and Actinobacteria species that we identified
N. Trabal et al.
among the microbiota associated with C. gigas or C. corteziensis have been reported in other oysters using cultureindependent techniques [33, 40, 54, 63, 69]. However, our
results showed low bacterial diversity compared with other
studies. We used TGGE methodology to analyze the bacterial community composition in the oysters. To rule out the
possibility that the observed lack of diversity may be the
result of primer bias, the 16S rRNA primers were analyzed
using the informatics tool Probe Match in RDP II, which
contains almost 2 million sequences that are distributed in
33 phyla. Each primer matched with over 80 % of the total
sequences deposited in RDP II, recognizing all phyla. More
than 70 % of the sequences assigned in each phylum were
recognized by these primers. Therefore, no specific preference during amplification should be expected, and the narrow diversity is therefore not a consequence of primer bias.
To reduce the nested PCR bias, the first amplification step
was limited to 15 cycles. For TGGE analysis, we used a
previously established protocol [55]. This protocol includes
the use of known sequences (%GC content) as a control for
each run and checks the separation of bands with different
GC contents.
Regardless of the growth stage or the cultivation site of
oysters, we found low bacterial diversity and this may be
due to the use of depurated oysters. With the exception of
the study by Romero et al. [69], the reports on microbiota
associated with other oysters, do not distinguish between
resident and transient bacteria. This could lead to an overestimation of bacterial diversity within the microbiota.
The Vibrio and Pseudomonas species are commonly isolated from oyster tissues using traditional methods [31, 54,
62, 63, 75, 77] and are recognized as symbionts of diverse
marine invertebrates [32]. In general, the Vibrios species
have been reported in sick oysters and have been detected
using selective culturing [17, 59, 75, 76]. However, we did
not find Vibrios in this study (only one OTU in an adult C.
gigas). It has been demonstrated that the distribution of
Vibrios is typically seasonal with peaks of abundance that
depend on environmental conditions, similar to mollusks
[17, 32, 75, 76]. However, although the juveniles and adults
of C. corteziensis and C. gigas were collected in warm
water, which has a positive effect on the abundance of
Vibrios, these bacteria were not found. In recent years, some
authors have suggested that Vibrio is not necessarily predominant in the microbiota of healthy oysters [29, 42]. We
analyzed samples from healthy and depurated oysters,
which may explain why we did not find Vibrios to be a
component of the microbiota of C. gigas and C. corteziensis. It would be interesting in a future study to use Vibriospecific primers to verify our results.
This study provides the first description of the bacterial
community in Crassostrea corteziensis. Additionally, this
study is the first attempt to describe what happens to the
Bacterial Microbiota in Two Commercial Oysters
stability of the microbiota when the post-larvae are planted
at grow-out cultivation sites. Considering that one of the
most critical steps in the cultivation of these oysters is the
planting of post-larvae (spat) at the cultivation site in the
natural environment in which large numbers of oyster die [7,
11, 18], the studies described here are probably important.
We observed that during the early growth stages, certain
bacteria (especially B. cepacia) colonize oysters and remain
strongly associated with their host, probably in a symbiotic
host–bacteria relationship. This association is maintained
during the growth phases of the oysters and is not altered
by environmental conditions or the management of the
organisms at the grow-out site. This finding is important
because probiotics have been used in aquaculture to control
pathogenic microorganisms by competitive exclusion and
may help to reduce and prevent outbreaks in several marine
species [26–28, 51, 61]. Resident bacteria with potential
probiotic characteristics that strongly associate with the
oysters seem to be stable in the host because the community
was not affected by the conditions of the grow-out site,
which suggests a competitive advantage for the sustainable
development of aquaculture.
The main findings of the present study were (1) the bacterial communities associated with the oysters, especially C.
corteziensis, at each grow-out site were similar, if not identical, and were influenced by the environment. (2) The microbiota associated with the oysters underwent certain changes
when the post-larvae were planted at grow-out sites and
during their growth from juvenile to adult. (3) However, B.
cepacia was established in the post-larval phase and remained
associated with the oysters throughout their growth, regardless
of the grow-out site. (4) There were certain differences among
the microbiota associated with C. gigas and C. corteziensis,
even when grown at the same site. (5) C. gigas has more
intraspecific differences in its composition of microbiota.
567
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
Acknowledgments We thank Acuícola Robles and Acuícola CuateMachado for providing the oysters used in this research. We also thank
Victoria Urzúa (INTA), Alejandra Givovich (INTA), Angel Carrillo
García (CIBNOR), and Hever Latisnere-Barragan of Marine Biotechnology Laboratory (CIBNOR) for technical support. Funding was
provided by Consejo Nacional de Ciencia y Tecnología of Mexico
(SEP-CONACYT grants 129025 and 106887). N. A. is a recipient of a
CONACYT doctoral fellowship and also an internship grant at the
Instituto Nacional de Tecnología de los Alimentos (Universidad de
Chile).
14.
15.
16.
17.
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