Open Access
28 September 2017 Isolation of Polymorphic Microsatellite Loci in the New Zealand Endemic Sand-Binder, Ficinia spiralis (Cyperaceae)
Rachel A. van Heugten, Marie L. Hale, Stacey Bryan, Bart van Griensven, Sophia Satter, Lily Brailsford, Hannah L. Buckley
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Ficinia spiralis (A. Rich.) Muasya & de Lange (Cyperaceae), otherwise known as pīngao, pikao, or golden sand sedge, is a sand-binding sedge that occurs on the foredunes of New Zealand sandy beaches (Bergin and Herbert, 1998). Small remnants and numerous human-planted restoration populations are all that remain of this previously widespread plant (Courtney, 1983; Bergin, 2011). Despite significant effort being put into conservation and restoration activities, this species is currently ranked as “Declining” in the New Zealand Threat Classification System (de Lange et al., 2013). Pīngao is of great cultural and ecological value and significance for New Zealand because it provides habitat for other native sand dune specialists and is an important weaving fiber, being highly valued for its beautiful golden color when dry. It exhibits pronounced morphological and physiological variation throughout its geographic range (Bergin and Herbert, 1998); however, the causes and consequences of this variation are not understood.

Microsatellite markers have not previously been successfully developed for this species, but are essential for characterizing the genetic variation within F. spiralis. In this paper, we present the development of microsatellite markers that will enable future genotyping of individuals from F. spiralis populations to correlate genetic, morphological, physiological, and ecological variation within this species. Our ultimate goal is to determine (1) what genetic variation is being lost as this species continues to decline and (2) what potential there is to use knowledge of the spatial genetic variation in this species to make better-informed conservation and restoration management decisions to reverse its decline.


Whole genomic DNA was extracted from 100-mg leaf samples of two F. spiralis individuals (geographic coordinates: −43.435°S, 172.713°E and −43.852°S, 172.771°E) using a modified cetyltrimethylammonium bromide (CTAB) method (Weising et al., 1995). Voucher specimens could not be collected due to permit restrictions (New Zealand Department of Conservation Research and Collection Permit 47972); a photograph of F. spiralis is provided in  Appendix S1 (apps.1700039_s1.pdf). The samples were combined before microsatellite regions were identified using a 454 sequencing approach. The 454 sequencing was conducted by Macrogen (Seoul, Korea) as follows. One microgram of genomic DNA was run on a GS FLX Titanium platform (1/8 region) after GS FLX Rapid Library shotgun library construction (454 Life Sciences, a Roche Company, Branford, Connecticut, USA). This generated 157,274 raw reads (National Center for Biotechnology Information [NCBI] Sequence Read Archive [SRA] Bioproject number PRJNA378666), which were then screened for microsatellite regions using MSATCOMMANDER version 0.8.2 (Faircloth, 2008). The search conditions were: dinucleotide repeats >8, trinucleotide repeats >7, tetranucleotide repeats >7, pentanucleotide repeats >6, and hexanucleotide repeats >6. The number of reads containing microsatellites was 536, and Primer3 (Koressaar and Remm, 2007; Untergasser et al., 2012) was used to design primers for all reads containing microsatellites with sufficient flanking sequence for primer design. Forty loci were then chosen for testing of amplification and variation based on the number of repeats present in the raw sequence. Reads with 15 to 25 repeats were preferred as they will likely maximize the chance of polymorphism while minimizing the likelihood of excessive stutter. Fourteen di-, 15 tri-, six tetra-, two penta-, and three hexanucleotide repeat regions were chosen for testing. The forward primer for each locus was appended with an M13 sequence (5′-TGTAAAACGACGGCCAGT-3′) at the 5′ end to allow the use of universal labeled primers (Schuelke, 2000). and PIG-tails (5′-GTTTCTT-3′) were added to the 5′ end of the reverse primers to reduce incomplete A-addition during PCR (Brownstein et al. 1996).

Table 1.

Characteristics of 15 nuclear microsatellite loci isolated from Ficinia spiralis.a,b


The primers (Invitrogen, Carlsbad, California, USA) were initially tested for amplification against four F. spiralis individuals from two sample sites (geographic coordinates: −43.435°S, 172.713°E and −43.852°S, 172.771°E). Primers showing evidence of amplification were screened for polymorphism with a fluorescent M13 tag (Applied Biosystems, Carlsbad, California, USA) using an additional six F. spiralis individuals spanning the same two sample sites. PCRs used the following protocol: 15-µL reactions containing 1 µL of template DNA, 1× PCR buffer (Bioline, London, United Kingdom), 2 mM MgCl2, 0.08 mM dNTPs, 0.33 µM reverse primer, 0.33 µM fluorescent M13 tag (6-FAM, VIC, NED, PET), 0.08 µM forward primer, and 0.6 unit BIOTAQ (Bioline). PCR cycles for loci FSP2, FSP4, FSP6, FSP11, FSP30, and FSP50 ran on a touchdown protocol (TD) as follows: 95°C for 15 min; 10 cycles of 94°C for 15 s, 62–53°C (decreasing 1°C per cycle) for 30 s, 72°C for 1 min; 25 cycles of 94°C for 15 s, 52°C for 30 s, 72°C for 1 min; and a final extension step of 15 min at 72°C. The remaining loci were amplified with the following PCR cycle: 95°C for 12 min; 10 cycles of 94°C for 15 s, annealing temperature (Table 1) for 30 s, 72°C for 30 s; 30 cycles of 89°C for 15 s, annealing temperature for 30 s, 72°C for 30 s; and a final extension of 72°C for 10 min. DNA fragments were detected and sized using the GeneScan 500 LIZ Internal Size Standard (Applied Biosystems) and an ABI 3130xl Genetic Analyzer (Applied Biosystems). Loci FSP19, FSP44, and FSP45 were genotyped individually. The remaining loci were combined in the following pairs: FSP6 and FSP16, FSP30 and FSP29,FsP7 and FSP2, FSP4 and FSP21 FSP51 and FSP50, and FSP11 and FSP1. Fragments were scored manually using GeneMarker software version 6.41 (SoftGenetics, State College, Pennsylvania, USA).

Of the 40 loci tested, 15 amplified cleanly. Of the 15 loci, one locus (FSP19) was monomorphic in F. spiralis, while 14 were polymorphic (see  Appendix S2 (apps.1700039_s2.docx) for raw sequences for these loci). The 14 polymorphic loci were then used to genotype 100 F. spiralis individuals from across four sample sites (each with at least 20 samples) to gain a better understanding of the variation present at these loci (Table 2). All 15 loci were tested for cross-amplification in six specimens (from two different locations: Christchurch [−43.486°S, 172.693°E] and Auckland [−36.956°S, 174.468°E]) of F. nodosa (Rottb.) Goetgh., Muasya & D. A. Simpson, the only other Ficinia Schrad. species recorded in New Zealand (a photograph of F. nodosa is provided in  Appendix S1 (apps.1700039_s1.pdf)). Ten of the 15 loci amplified in F. nodosa, and two loci (FSP19 and FSP51) were polymorphic (Table 3). Analysis with POLYSAT version 1.7-1 (Clark and Jasieniuk, 2011; Clark and Schreier, 2017) in R version 3.0.2 (R Core Team, 2013) indicated that F. spiralis individuals are either triploid or tetraploid. However, there is no record in the literature to indicate whether F. spiralis is allopolyploid or autopolyploid. Therefore, F. spiralis alleles were scored as either present or absent, and it was not possible to calculate Hardy–Weinberg equilibrium and linkage disequilibrium.


Ficinia spiralis (Cyperaceae) is a declining, endemic species of great cultural and ecological value to New Zealand. Here, we describe one monomorphic and 14 polymorphic microsatellite loci that will enable us to assess genetic variation across the geographic range of this species, and to determine whether this variation correlates with morphological and/or physiological variation. Ten of these loci also amplified in F. nodosa, suggesting these loci may be useful for assessing genetic variation and population structure in other species within this genus. Although the polyploidy of F. spiralis limits the range of population genetic analyses available to those that can accommodate dominant data, numerous other studies have used similar data to investigate population structure and how it relates to other factors such as the environment (i.e., Brito et al., 2016; Wu et al., 2016). The high number of alleles at many of these loci should provide sufficient information for the assessment of fine-scale genetic structure throughout the species' range.

Table 2.

Genetic diversity statistics for 14 polymorphic microsatellite loci in four populations of Ficinia spiralis.


Table 3.

Cross-amplification of 15 nuclear microsatellite loci developed for Ficinia spiralis in F. nodosa (n = 6).a



Funding for this research was provided by the Brian Mason Scientific and Technical Trust.



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Rachel A. van Heugten, Marie L. Hale, Stacey Bryan, Bart van Griensven, Sophia Satter, Lily Brailsford, and Hannah L. Buckley "Isolation of Polymorphic Microsatellite Loci in the New Zealand Endemic Sand-Binder, Ficinia spiralis (Cyperaceae)," Applications in Plant Sciences 5(9), (28 September 2017).
Received: 17 April 2017; Accepted: 1 July 2017; Published: 28 September 2017
Ficinia nodosa
Ficinia spiralis
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