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24 August 2017 Characterization and Transferability of Microsatellites for the Kangaroo Paw, Anigozanthos manglesii (Haemodoraceae)
Bronwyn M. Ayre, Janet M. Anthony, David G. Roberts, Richard J. N. Allcock, Siegfried L. Krauss
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Anigozanthos manglesii D. Don (Haemodoraceae), the Red and Green Kangaroo Paw, is a perennial wildflower endemic to the Southwest Australian Floristic Region. Flowering occurs between July and November, with large inflorescences of red and green tubular flowers on stems up to a meter tall. These flowers are visited by nectar-feeding birds and invertebrates seeking nectar and pollen (Hopper, 1993). Differences in the foraging behavior of vertebrates and invertebrates are predicted to have a significant impact on pollen dispersal patterns, multiple paternity, genetic diversity, and fitness of offspring (Krauss et al., 2017). Manipulation of pollinator access to inflorescences and paternity assignment of the resulting seeds allows for the quantification of pollen dispersal patterns by specific pollinators. Here, we describe the development of microsatellite markers that will facilitate future research on the genetic consequences of pollen dispersal by bird and invertebrate pollinators of A. manglesii. In particular, we will use these markers for mating system and paternity assignment following pollinator manipulation studies to test hypotheses of high paternal diversity for plants pollinated by nectar-feeding birds (Krauss et al., 2017). The degree of congeneric cross-transferability of the markers was also assessed in eight other species, covering over 80% of the genus.

METHODS AND RESULTS

DNA was extracted from a leaf sample collected in Kings Park, Perth, Western Australia (Appendix 1), using the extraction method of Carlson et al. (1991), modified with the addition of potassium acetate after lysis incubation, a 5 M NaCl step, and an additional ethanol precipitation after the isopropanol precipitation. One hundred grams of DNA was sheared to approximately 300–400 bp using an S2 sonicator (Covaris, Woburn, Massachusetts, USA), and a single barcoded library was prepared using a NEBNext Ultra DNA Library Prep Kit (New England Biolabs, Ipswich, Massachusetts, USA). Inserts sized 330–360 bp were selected by gel excision (E-Gel, Invitrogen/Thermo Fisher Scientific, Waltham, Massachusetts, USA), and the libraries were produced, assessed, and quantified using a Bioanalyzer 2100 (Agilent Technologies, Santa Clara, California, USA). The final library was diluted to 9 pM using a OneTouch 2 Template 400 kit (Life Technologies, Carlsbad, California, USA) and enriched. A Personal Genome Machine (PGM) semiconductor sequencer (Life Technologies) using 850 flows on a 316 sequencing chip produced approximately 350–400 bp read lengths. Signal processing, base-calling, and quality trimming were conducted using the default settings on Torrent Suite 4.0 (Thermo Fisher Scientific), and library-specific FASTQ files were generated. This resulted in 4.03 million reads with a modal read length of 354 bp and 2.4 Gb of data (National Center for Biotechnology Information [NCBI] Sequence Read Archive Bioproject no. PRJNA390010).

Table 1.

Characteristics of 15 polymorphic microsatellite loci developed for Anigozanthos manglesii.

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Using QDD 3.1 software, all reads were screened for microsatellite-containing regions (Meglecz et al., 2014). A total of 190,000 were identified. Thirty primer pairs were chosen for screening at a time. Primers chosen were all categorized as design A (no homopolymers, no other target microsatellites in flanking region, no nanosatellite in primer or flanking regions, pure not compound microsatellites), were unable to form a hairpin, had a low PCR align score, had a >20-bp distance between primer and microsatellite, had higher microsatellite repeats, and had similar annealing temperatures, but had a variety of PCR product lengths (Meglecz et al., 2014;  http://net.imbe.fr/∼emeglecz/qdd.html#choice). Each assay had a final volume of 10 µL and contained 5 µL of SsoAdvanced SYBR Green Supermix (Bio-Rad Laboratories, Hercules, California, USA), 0.3 µM of forward and reverse primers, and 5–10 ng of genomic DNA. PCR was conducted on a CFX96 Touch Real-Time PCR Detection System (Bio-Rad Laboratories). Using a single sample, DNA was amplified across a range of temperatures to determine an appropriate annealing temperature. To test for polymorphism, eight individuals were amplified at the chosen best temperature and analyzed using Precision Melt Analysis (Bio-Rad Laboratories). The forward primer of primer pairs that amplified consistently across all eight individuals were each tagged with a fluorescent label (6-FAM, NED, VIC, or PET) compatible with the ABI 3500 sequencer (Life Technologies). This process was repeated three times, until 15 reliable primer pairs were produced. All other primer pairs failed to amplify consistently and/or cleanly (i.e., they displayed stuttering and allelic patterns were difficult to distinguish) across different DNA samples.

Table 2.

Genetic properties of 15 polymorphic microsatellite loci for three populations of Anigozanthos manglesii.a

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Table 3.

Results of cross-amplification (allele size ranges) of microsatellite loci isolated in Anigozanthos manglesii and tested in five individuals across eight congeneric taxa. Anigozanthos manglesii is included for comparison.

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To amplify microsatellite regions, PCR was performed on a Veriti Thermocycler (Life Technologies), either individually or in multiplex. Individual microsatellite loci (Am1, Am2, Am8, Am23, Am28, Am56, Am60, and Am79) were amplified using 10–20 ng of DNA with 2 µL of 5× buffer containing dNTPs (Fisher Biotec, Wembley, Western Australia, Australia), 2 mM MgCl2, 0.16 µM of both reverse and fluorescently labeled forward primers, and 0.05 µL of 5.5 units/µL Taq polymerase (Fisher Biotec) in a 10-µL reaction. The amplification cycle began with a 1-min denaturation at 95°C; followed by 35 cycles of denaturation at 95°C for 10 s, annealing (at variable temperatures, see Table 1) for 30 s, and extension at 72°C for 45 s; and a final extension of 15 min at 72°C. The remaining seven primer pairs were amplified across two multiplex mixes (primer mix 1 contained Am11, Am13, Am20, and Am29, all at 0.2 µM; primer mix 2 contained Am2 [0.1 µM], Am75 [0.4 µM], and Am82 [0.3 µM]). All multiplex reactions used 6 µL of 2× Multimix (QIAGEN, Hilden, Germany), 2 µL of 5× Q-solution (QIAGEN), 1.25 µL of primer mix, and 2.75 µL of 10–20 ng DNA in a final 12-µL reaction. The amplification cycle began with 15-min denaturation at 95°C; followed by 30 cycles of denaturation at 94°C for 30 s, annealing (at variable temperatures, see Table 1) for 90 s, and extension at 72°C for 90 s; and a final extension of 30 min at 60°C. PCR products were separated by capillary electrophoresis on an ABI 3500 Genetic Analyzer (Life Technologies), and allele sizes scored using Geneious version 7.1 (Biomatters Ltd., Auckland, New Zealand;  http://www.geneious.com/).

Primers were tested on leaf samples collected from three populations of A. manglesii (Appendix 1, Table 2). All 15 markers were polymorphic in at least one population. Analysis for observed heterozygosity, expected heterozygosity, and Hardy–Weinberg equilibrium was completed with GenAlEx (Peakall and Smouse, 2006, 2012). Observed and expected heterozygosities ranged from 0.182 to 0.950 and 0.133 to 0.931, respectively. A significant departure from Hardy–Weinberg equilibrium was recorded in different loci across the three populations (Table 2). MICRO-CHECKER (van Oosterhout et al., 2004) identified the possibility of null alleles in some loci, but not consistently across populations. No stuttering or large allele dropouts were identified.

Using the same extraction and amplification methods as above, the primers were tested on DNA extracted from five individuals from each of A. bicolor Endl., A. flavidus DC., A. gabrielae Domin, A. humilis Lindl., A. preissii Endl., A. pulcherrimus Hook., A. rufus Labill., and A. viridis Endl. Success varied, with four to eight markers successfully amplified across different species (Table 3).

CONCLUSIONS

Fifteen microsatellite markers have been developed for A. manglesii. Without changing any of the amplification conditions, between four and eight of these markers successfully amplified in each of eight congeneric species. This suggests that with further species-specific refinement, these markers will provide a valuable resource for population genetic studies of the genus.

ACKNOWLEDGMENTS

The authors thank M. Jones and R. Phillips for their help sourcing leaf samples from additional Anigozanthos species, and an anonymous reviewer for valuable feedback. This work was supported by an Australian Research Council Discovery Grant to S. Hopper, S.L.K., and R. Phillips (DP 140103357) and an Australian Government Research Training Program Scholarship and a Holsworth Wildlife Research Endowment Equity Trustees Charitable Foundation grant to B.M.A.

LITERATURE CITED

1.

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Meglécz, E., N. Pech, A. Gilles, V. Dubut, P. Hingamp, A. Trilles, R. Grenier, and J. F. Martin. 2014. QDD version 3.1: A user-friendly computer program for microsatellite selection and primer design revisited: Experimental validation of variables determining genotyping success rate. Molecular Ecology Resources 14: 1302–1313. Google Scholar

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Appendices

Appendix 1.

Voucher information for Anigozanthos species used in this study.a

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Bronwyn M. Ayre, Janet M. Anthony, David G. Roberts, Richard J. N. Allcock, and Siegfried L. Krauss "Characterization and Transferability of Microsatellites for the Kangaroo Paw, Anigozanthos manglesii (Haemodoraceae)," Applications in Plant Sciences 5(8), (24 August 2017). https://doi.org/10.3732/apps.1700055
Received: 21 May 2017; Accepted: 1 June 2017; Published: 24 August 2017
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KEYWORDS
Anigozanthos
Catspaw
Haemodoraceae
Kangaroo Paw
microsatellite primers
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