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Fast, cost-effective development of species-specific microsatellite markers by genomic sequencing
Jawad Abdelkrim1,2, Bruce C. Robertson1,3, Jo-Ann L. Stanton4, and Neil J. Gemmell1,2
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Supplementary Material

Using a 1/16th run, which costs <$1500 USD at the facility used for this work (Department of Anatomy and Structural Biology, University of Otago, Dunedin, NZ), this approach cuts the cost of microsatellite marker development by 3–5 times, once the cost of the primer synthesis is included, versus the cost cited for most commercial microsatellite marker development. If more markers were desired using our approach, the same ligation library could be used again as well. A second 1/16th run would cost ∼$1060 USD (or ∼$81 USD per marker assuming the same return). If a large amount of markers are needed, the cost is further decreased by running a whole plate, which also takes less time than running 1/16th. In this case, the cost would be $11,000 USD. If the average yield is the same, 455 polymorphic markers would be expected and would represent a price of $24 USD per polymorphic marker. Of course, the number of markers could also be easily increased if needed just by relaxing some parameters of the microsatellite mining, such as reducing the minimum number of repeats or a search for interrupted and compound repeats could increase the number of loci returned. In our case, by simply decreasing the minimum number of repeats from 8 to 5, the number of microsatellites detected increased 3-fold. Finally, a popular approach to increase the yield of usable microsatellite loci is enrichment (2). We have not yet tested the efficiency of new genomic sequencing technologies for isolating microsatellites following an enrichment step, but this is an obvious extension of our work and we see no reason why it would not be successful.

The method described here reduces lab time; in our case, the entire process was completed in 2 weeks, and has the potential to be faster. The sequencing process—from the construction of the library to obtaining sequences—only took 4 days in this case. No cloning or library screening is necessary, and if the sequencing step is sourced at an external facility, no molecular biology work is necessary for the researcher, except for the initial DNA extraction. The bioinformatics tools we used for repeat finding and primer design are freely available and run on a desktop PC quickly (Intel Core 2 Duo, 2.66GHz, 3.24 GB of RAM). This new method also targets all microsatellite repeat types (e.g., di-, tri-, tetra-, and penta-nucleotides) and motifs (i.e., nucleotide composition of the repeated sequence). In contrast, traditional methods require the genomic library to be enriched for a limited number of specific microsatellite motifs, and it is known that the choice of motif can have an effect on the variability detected (11). Another advantage of this approach is that in addition to the possibility of designing microsatellite primers, millions of base pairs of genomic sequence are available, potentially providing a framework for further genomic analyses and a useful resource for comparative research, as more and more data will be generated on non-model organisms.

In this study, we focused on a single bird species, the blue duck, though recent application of this approach to develop microsatellite markers for other species in our hands show that this method is promising in other phylum as well (Table 2). Indeed, one insect, one gastropod, and one trematode species have also been run through the same process and in all cases, a large number of microsatellite-containing sequences were detected (472–2541, depending on the species). The important number of potential primer pairs, following our parameter settings above, range from 46 to 170, making it likely that enough polymorphic markers will be available to conduct very detailed population genetic studies in each of these species.

In summary, we describe a reliable method based on the combined use of high-throughput sequencing technologies and bioinformatics tools by which microsatellite isolation can be performed at a fraction of the cost and timeframe of current methodologies. The one drawback with this approach is the current technical limitation of the sequencing technology, which presently only produces reads of an average size of 250 bp. The size constraint of these short reads could have two implications. First, one of the main determinants of microsatellite variability, mutation rate, is dependent on the number of repeats as well as other factors (12). Thus, as it stands, this approach may limit the isolation of longer, potentially more polymorphic microsatellites. However, this drawback also applies to traditional library construction, due to the necessity to size-select prior to bacterial cloning (2). Second, the short fragment length frame could make it more difficult to design large sets of variable-length markers that could be multiplexed for genotyping without needing to resort to primer tagging to create different-length products. Nevertheless, the results we obtained in our test species highlights that the final markers exhibit a decent level of polymorphism, similar to many traditionally designed markers and can be easily pooled into two sets for genotyping based on allele sizes (Table 1). Moreover, technical limitations introduced by sequence read lengths have been overcome in the past, and a recent announcement from Roche indicates that an increase in sequence read length to 400 bp and a 5-fold increase in throughput have been achieved solely through improvements in the reagents used (

In conclusion, the genomic shotgun sequencing approach using high-throughput technology offers a cost-effective, simple, and fast alternative to traditional protocols or commercial services for microsatellite isolation. This approach should be of particular advantage to research groups conducting occasional genotyping projects on a limited number of species, as well as larger research groups who regularly develop microsatellite markers. Under this new paradigm, microsatellite marker development becomes as simple and routine as traditional Sanger sequencing has become and the cost per sample—and cost per marker—will only reduce further.


Three anonymous referees kindly provided helpful comments on the manuscript. We thank the New Zealand Department of Conservation and the blue duck recovery group for collecting and providing the samples under permits ECHB-23214-RES and TT-23326-FAU. This work was funded as part of the Sustaining and Restoring Biodiversity OBI FRST contract C09X0503. We also thank Yuri Springer (University of Otago) and Thomas Buckley (Landcare Research) for providing results from their studies.

The authors declare no competing interests.

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