Introduction

Crassostrea species are responsible for almost all of the worldwide oyster production [1]. Oyster production in the South Atlantic is dominated by three species: the native Crassostrea gasar (Adanson, 1757; sin. C. brasiliana, [2]) and C. rhizophorae (Guilding, 1828), which are extracted from natural banks, and the cultivated C. gigas (Thunberg, 1793), responsible for 91 % of the total Brazilian oyster production [3] and recently becoming invasive in natural banks [4]. Native oyster species have a wide distribution along the Atlantic coast of Latin America and Africa [5], but their cultivation is hampered by the uncertainty about the existing stocks and their limits. To date, the few studies that have analysed the genetic structure of C. gasar and C. rhizophorae used mitochondrial (cytochrome oxidase I) and allozyme markers [2, 4, 6]. Recently, ten microsatellite loci were developed for C. gasar [7] and will allow new genetic structuring studies with more polymorphic markers. In contrast, microsatellite loci for C. rhizophorae are still lacking.

According to Beck et al. [8], over 100 years of intense and unregulated exploitation has eliminated more than 99 % of the oyster reefs of many bays around the world, and thus, the worldwide oyster populations can be considered functionally extinct. In some cases, oyster populations have vanished altogether, such as for C. rhizophorae, which has not been found in Ciénaga Grande de Santa Marta (Colombia) since 1996, most likely due to the combined effects of uncontrolled extraction and alterations in freshwater flow [9]. Aquaculture is accepted as an alternative to this uncontrolled oyster extraction; however, to avoid the risks of outbreeding depression and loss of locally adapted gene traits, it relies on knowledge of the number and limits of genetic stocks. This knowledge can be greatly improved by population genetics studies based on highly polymorphic loci. Here, we describe the isolation and characterisation of microsatellite loci for C. gasar and C. rhizophorae and test their cross amplification in the cultivated C. gigas and the native C. virginica (Gmelin, 1791).

Materials and methods

DNA was extracted from the adductor muscle according to the modified cetyltrimethylammonium bromide (CTAB) protocol, as previously described [10]. Microsatellite DNA was isolated from an enriched partial genomic library following the protocol of Bloor et al. [11]. A pool of high-quality genomic DNA from four individuals (10 μg) was digested with SauIIIA and ligated to phosphorylated double-stranded linkers; the fragments (between 500 and 1,000 bp) were excised from a 2 % agarose gel. The DNA fragments were hybridised with biotinylated (CA)12 and (CAA)8 probes and isolated using streptavidin-coated magnetic beads. A PCR primed with the forward linker oligonucleotide was used for enrichment of the DNA containing microsatellites. The enriched fragments were cloned using the pGEM-T vector (Promega) for posterior insertion in One Shot TOP10 competent cells (Invitrogen). White colonies were selected during screening on S-gal (Sigma)/agar/ampicillin plates. The presence of a microsatellite insert was confirmed by two or more PCR products after amplification using the forward linker oligonucleotide and (non-biotinylated) microsatellite oligonucleotides as primers. 48 positive clones from each species were sequenced in both directions using an ABI3500 automated sequencer (Applied Biosystems). The sequences were edited using SEQMAN (Lasergene Inc.). 19 and 47 sequences from C. gasar and C. rhizophorae, respectively, were selected for the design of primer pairs flanking the microsatellite regions using WEBSAT [12]. Primers were initially screened for quality of PCR amplification on a Veriti® Cycler (Applied Biosystems), using eight individuals of each species. Those primers that did not amplify, or that produced multiple bands on high-resolution agarose gels were discarded. The 22 loci selected were tested for polymorphism through genotyping on an ABI3500 automated sequencer using the tailed primer method [13], whereby all the forward primers were synthesised with a M13 tail at their 5 end to which oligonucleotides labelled with different dyes (6-FAM, VIC, NED, or PET) could anneal. The best annealing temperature (Ta) for each set of primers was estimated in a 54 to 64 °C temperature gradient PCR. Subsequently, thermocycling was performed using an initial denaturing at 94 °C for 3 min, 30 cycles of 45 s at 93 °C, 45 s at the specific Ta, and 1 min at 72 °C, followed by 8 cycles identical to the 30 initial cycles but with the Ta fixed at 53 °C (as in [13]). The cycling was finalised with a 15 min extension at 72 °C. The PCR reactions included 20 ng template DNA, 1 U GoTaq® DNA Polymerase (Promega), 200 μM dNTPs, 0.2 μM tailed forward primer, 0.4 μM labelled primer, 0.8 μM reverse primer, 2.5 mM MgCl2, and 15 μg BSA in a 15 μL total volume. The PCR products were pooled with a GSLIZ-600 size standard (Applied Biosystems) and were denatured in formamide prior to capillary electrophoresis. Allele sizing and genotype confirmation were performed using the GENEMARKER 1.97 programme (SoftGenetics—free trial version). After the analyses, 5 monomorphic loci were discarded, so 17 polymorphic loci (6 for C. gasar and 11 for C. rhizophorae) were selected. Binning of the alleles was performed using FLEXIBIN software [14]. The number of alleles was determined, and tests of linkage disequilibria were performed using FSTAT [15]. The observed and expected heterozygosities and tests of Hardy–Weinberg equilibrium were calculated using ARLEQUIN 3.5 [16]. The presence of null alleles was investigated with the MICRO-CHECKER programme [17].

Results and discussion

Twenty-four individuals of C. rhizophorae, from Southeast Brazil (São Paulo: 25°01S, 47°55W) and 23 individuals of C. gasar from North Brazil (Maranhão: 02°41S, 41°58W) were genotyped. Linkage disequilibrium was not found for any pair of loci after Bonferroni correction for either species, but four loci (GASA5, RHIZ7, RHIZ12, and RHIZ18) exhibited significant heterozygote deficiencies, possibly due to the presence of null alleles (Table 1). A high frequency of null alleles has been previously verified for microsatellite markers in molluscs [18]. The analysed loci demonstrated high polymorphism, ranging from 5 to 27 alleles, and the observed and expected heterozygosities ranged between 0.609–0.958 and 0.582–0.972 in C. rhizophorae and C. gasar, respectively (Table 1). An exception was GASA15, which had only three alleles and observed and expected heterozygosities of 0.174 and 0.237, respectively.

Table 1 Microsatellite loci developed for C. gasar and C. rhizophorae
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Cross-amplification was tested for all the loci in two other species of Crassostrea: C. virginica and C. gigas. Seven C. virginica individuals from Panamá (8°58′N, 79°31′W) and eight C. gigas individuals from an oyster farm in South Brazil (Santa Catarina: 26°15′S, 48°41′W) were analysed. 11 loci amplified well for C. virginica, while just two loci worked for C. gigas. No pairwise linkage disequilibrium was found between loci of either species, nor did they present any significant deviations (after Bonferroni correction) from Hardy–Weinberg equilibrium. Values of annealing temperature, heterozygosity and number of alleles for cross-amplification are shown in Table 2. All loci exhibited high levels of polymorphism, except for RHIZ8 in C. virginica (H = 0.14). Surprisingly, none of the loci from C. rhizophorae could be used in C. gasar and vice versa: some heterologous loci did amplify between the two species, but they did not pass the stringent conditions set for the selection of loci for the two species that were the main objective of this work.

Table 2 Cross-amplification of the developed loci in C. virginica and C. gigas
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The new markers developed here will be useful for population and aquaculture studies. More specifically, they will help with the delimitation of the genetic stocks along the coast, which will provide critical information for the cultivation of native species, contributing both to their protection and to habitat restoration.