The duckweed family encompasses 38 monocotyledonous species in four genera (Landolt, 1986; Les et al., 2002). Duckweeds had long been classified in their own family, the Lemnaceae, but are now considered to be members of the arum or aroid family (Araceae); the name Lemnaceae is therefore rapidly falling out of favor among taxonomists, who treat it as the subfamily Lemnoideae (Cabrera et al., 2008). Lemna minor L., which is known as common duckweed, is a small, free-floating, and fast-growing aquatic plant with a chromosome count of 40 (Blackburn, 1933). The species is distributed worldwide and often grows as blanket-like mats on the surface of still or slow-moving, nutrient-rich, fresh and brackish waters. Lemna minor represents a potential source of bioethanol (Xu et al., 2011) and can extract organic pollutants and toxic metals from waters, which makes it useful in remediation efforts (Alvarado et al., 2008; Wang et al., 2010). The species reproduces quickly through vegetative budding and doubles its biomass in two to seven days, depending upon culture conditions. Conditions affecting growth include the availability of nutrients and water temperature (Landolt, 1986; Brain and Solomon, 2007; Kanoun-Boulé et al., 2009). Notwithstanding its wide geographic range, L. minor displays a characteristically sporadic local distribution (Savile, 1956). In some parts of the world, such as the Kashmir Himalayas, this species has tended to become more invasive (Shah and Reshi, 2014).

Because of its widespread distribution, a monographic account of the Lemnaceae by Daubs (1965) puts L. minor in a “catch-all” category, as many herbarium specimens have been inadvertently labeled with this binomial but are actually other species. Development of appropriate molecular markers has therefore attained a special significance for correct taxonomic delineation of the species. Moreover, a lack of genetic markers impedes our understanding of the population biology and dynamics of L. minor. The development of such markers promises to yield important insights into the biology and biogeography of this species, with useful implications for understanding its invasiveness. Although Wang and Messing (2011) recently sequenced the chloroplast genomes from species in three different genera within the Lemnoideae (i.e., Spirodela polyrhiza (L.) Schleid., Wolffiella lingulata Hegelm., and Wolffia australiana (Benth.) Hartog & Plas) for systematic analysis, there are no studies so far on duckweed species using simple sequence repeat (SSR) or microsatellite markers. Thus, our objective was to develop cpDNA-based SSR markers for L. minor because such markers could provide a wealth of information for evolutionary and population genetic studies.

Table 1.

Nine polymorphic microsatellite markers used for optimization on Lemna minor.

METHODS AND RESULTS

Individuals of L. minor were collected from five populations in the Kashmir Valley, India, and from three populations in Quebec, Canada, using a panel of five to seven individuals per population. The date and site of collection within each region, together with geographic coordinates of the sites, are given in Appendix 1; voucher specimens could not be collected due to lack of availability of suitable specimens. To develop SSR markers for L. minor, the chloroplast genome of L. minor was downloaded from the National Center for Biotechnology Information (NCBI) database. The PerlScript MIcroSAtelitte (MISA;  http://pgrc.ipk-gatersleben.de/misa/) was used to identify microsatellites in the L. minor chloroplast genome. The SSR information that was generated by MISA was used for designing primers flanking the repeats. To design primers that flanked the microsatellite locus, two PerlScripts were used as interface modules for the program-to-program data interchange between MISA and the primer-designing software Prim er 3 (Rozen and Skaletsky, 2000). Primer pairs were designed from the flanking sequences of SSRs using primer3_core in batch mode via the p3_in.pl and p3_out.pl PerlScripts (Sonah et al., 2011). The primer-designing conditions were: 100–300 bp amplicon size, 60°C optimal annealing temperature, 20 bp optimal primer length, and 50% optimal GC content (Sonah et al., 2011). Three sets of primer pairs were designed for each SSR to provide alternatives if amplification was unsuccessful.

Genomic DNA was extracted by grinding 0.25 g of fresh leaf tissue in liquid nitrogen and by using a prewarmed cetyltrimethylammonium bromide (CTAB) extraction protocol (Doyle and Doyle, 1987). Thirty-three primer pairs were designed initially, synthesized, and tested on seven individuals from Kashmir and Quebec by running the PCR products in 1.5% agarose gel in l× Tris-acetate/EDTA (TAE). PCR amplifications were carried out in total reaction volumes of 15 µL containing 50 ng of template DNA, 0.2 µM forward primer, 0.5 µM reverse primer, 1.5 mM dNTPs (Applied Biosystems/ Life Technologies, Grand Island, New York, USA), 1× PCR buffer including MgCl₂ (10 mM Tris [pH 8.0], 50 mM KCl, and 50 mM ammonium sulphate; Sigma Aldrich, St. Louis, Missouri, USA), 0.5 µM fluorochrome (Applied Biosystems/Life Technologies), and 1 unit of Taq DNA polymerase (Sigma Aldrich). The thermal cycling profile was 4 min at 94°C; followed by 35 cycles of 94°C for 1 min, 51°C annealing for 1 min, and 72°C for 1 min; followed by a final extension of 72°C for 10 min. The PCR products were separated by electrophoresis in 1.5% agarose gels in 1× Tris-borate/EDTA (TBE) buffer and visualized by ethidium bromide staining. To check for variability in L. minor, five to seven individuals from each of the different populations were amplified for each primer set. Amplicons were aligned using BioEdit Sequence Alignment Editor (Ibis Biosciences, Carlsbad, California, USA) to determine the possible identity of haplotypes, and fragments were measured using an ABI PRISM 3130xL Analyzer (Applied Biosystems, Carlsbad, California, USA) and scored using Peak Scanner version 1.0 software (Applied Biosystems).

DNA samples that were obtained from 26 individuals of five L. minor populations in Kashmir and from 17 individuals of three populations in Quebec were screened against 33 primer pairs. We found nine polymorphic loci (Table 1) and 24 monomorphic loci ( Appendix S1 (APPSD1300099_AppendixS1.docx)), which allowed the identification of 11 haplotypes in Kashmir and one haplotype in Quebec (Table 2). Of these 11 haplotypes, one occurs in 56% of the genotypes, one in 8%, and nine in 4% each. The number of alleles and unbiased estimates of haploid diversity are shown in Table 3. One intraspecific diagnostic locus (L16^*) showed discriminating alleles between Kashmir and Quebec, and could be useful to determine whether individuals introduced outside of the native range are from similar or different source populations in the native range.

CONCLUSIONS

For the first time, we have developed and characterized nine polymorphic and 24 monomorphic cpDNA microsatellite markers for L. minor. We expect these markers to be useful for population genetic studies and the reconstruction of introduction history, as well as to facilitate the understanding of other life history questions regarding Lemna and related species.

Table 2.

Haplotypes of cpSSRs at nine polymorphic loci of Lemna minor.^a

Table 3.

Chloroplast microsatellite genetic diversity values for nine polymorphic loci of Lemna minor.