See discussions, stats, and author profiles for this publication at: https://www.researchgate.net/publication/12204512 Characterization of microsatellite loci in King George Whiting Sillaginodes punctata Cuvier and Valenciennes... Article in Molecular Ecology · January 2001 DOI: 10.1046/j.1365-294X.2000.105331.x · Source: PubMed CITATIONS READS 0 27 2 authors, including: Steve C. Donnellan South Australian Museum 245 PUBLICATIONS 5,002 CITATIONS SEE PROFILE All content following this page was uploaded by Steve C. Donnellan on 30 March 2015. The user has requested enhancement of the downloaded file. All in-text references underlined in blue are linked to publications on ResearchGate, letting you access and read them immediately. Molecular Ecology (2000) 9, 2155–2234 PRIMER NOTES Blackwell Science, Ltd Microsatellite loci for the social wasp Polistes dominulus and their application in other polistine wasps Graphicraft 00 Limited, Hong Kong M I C H A E L T. H E N S H AW * Department of Ecology and Evolutionary Biology, Rice University, PO Box 1892, Houston, TX 77251–1892, USA Keywords: Polistinae, social wasps, Vespidae Received 21 March 2000; revision accepted 29 May 2000 Correspondence: Michael T. Henshaw. *Present address: Department of Entomology, 102 Fernald Hall, University of Massachusetts, Amherst, MA 01003–2410 USA. Fax: (413) 545–0231; E-mail: henshawm@ent.umass.edu The social wasps of the genus Polistes are an important model system for understanding the evolution of cooperation. Their relatively simple societies lack the distinct morphological castes which characterize many of the social insects, and newly emerged females possess a variety of reproductive options (Reeve 1991). A female may remain on her natal nest as a helper gaining indirect fitness; usurp a foreign nest and become reproductively dominant; initiate a new nest independently; reproduce on a satellite nest; or initiate a new nest in cooperation with other wasps (Strassmann 1981; Reeve 1991; Mead et al. 1995; Cervo & Lorenzi 1996; Queller et al. 2000). By characterizing the reproductive payoffs associated with different reproductive strategies, we are better able to understand how cooperative societies are maintained. Recently, microsatellite genetic loci have greatly extended our ability to characterize the reproductive strategies used by social wasps (Hughes 1998; Queller et al. 1993a). Using microsatellite loci we can reconstruct pedigrees, and estimate relatedness. Using this information, unobserved events such as queen death, nest usurpation or past reproductive dominance can be inferred (Queller et al. 1993a,b; Field et al. 1998; Hughes 1998). In this paper, I describe microsatellite loci isolated from the social wasp Polistes dominulus, one of the best studied Polistes species. We followed published protocols for the isolation of microsatellite loci (Strassmann et al. 1996) with clarifications and modifications to those protocols as noted below. DNA was extracted from 1 to 1.5 g of pupal thoraces ground in a mortar and pestle which had been chilled in liquid nitrogen. The ground tissue was suspended in grinding buffer (0.1 m NaCl; 0.1 m Tris-HCl, pH 9.1; 0.05 m EDTA; 0.05% SDS), and purified three times with phenol:chloroform:isoamyl alcohol (25:24:1), and then three times with chloroform:isoamyl alcohol (24:1). The purified genomic DNA was then ethanol precipitated, and resuspended in distilled water. Genomic DNA was digested with Sau3aI, and 300 –1000 bp inserts were ligated into the pZErO –2 plasmid (Zero Background cloning kit, Invitrogen) digested with BamHI. We transformed TOP10 cells (Invitrogen) to obtain approximately © 2000 Blackwell Science Ltd 5000 – 6000 clones. Nylon replicates of the genomic library were probed with five oligonucleotides (AAT10, AAG10, AAC10, TAG10, and CAT10) which were end-labelled with [γ-33P]-dATP. Probes of the nylon replicates yielded 151 positives and subsequent probing of plasmid DNA on the southern blot confirmed 34 unique positives. Clones which were positive on the southern blot were sequenced on an ABI 377 automated sequencer (Perkin-Elmer), and 19 sets of polymerase chain reaction (PCR) primers were designed from the 28 resulting sequences using Mac Ventor 5.0 (Kodak Scientific Imaging Systems). We optimized the PCR primers on an MJ Research PTC100 thermocycler using 10 µL reactions (Peters et al. 1998), and assessed within-species polymorphisms for eight species of polistine wasps, using from one to eight unrelated females for each species (Table 1). PCR products were visualized on 6% polyacrylamide/8 m Urea sequencing gels. Twelve of the 19 loci tested were polymorphic within our P. dominulus population and had a mean observed heterozygosity (HO ) of 0.76. Loci with a minimum of five uninterrupted repeats were polymorphic, and heterozygosity increased logarithmically with the number of uninterrupted repeats (Fig. 1; logarithmic regression, R2 = 0.454, P = 0.0016). The loci retained much of their polymorphism in other species of Polistes with six polymorphic loci for P. fuscatus and P. apachus which had a mean HO of 0.48. No polymorphisms were detected outside of the Polistes genus, however, it is likely that some polymorphisms went undetected due to the small number of individuals screened in the other species (Table 1). Acknowledgements This work was supported by a National Science Foundation (NSF) grant DEB-9510126 to Joan Strassmann and David Queller, Fig. 1 The relationship between the observed heterozygosity and the number of uninterrupted repeats for 19 microsatellite loci isolated from Polistes dominulus. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Locus Size (bp) Ta (°C) Pdom 1 209 55 Pdom 2 184 (CAG)9TAG(CAG)5 (CAT)5GGCAC(CAG)3 51, 48 (AAG)8CG(AAG)2 Pdom 7 160 54 Pdom 20 236 55, 52 (CAT)18 Pdom 25 157 50, 45 (AAG)11 Pdom 93 131 55 Pdom 117 260 51, 48 Pdom 121 218 54, 50 Pdom 122 172 46, 48 Repeat (AAG)CAG(AAG)9 (AAG)2ACG(AAG)2 ACG(AAG)5 (AAG)4AGG(AAG)2 AGG(AAG)14 (AAG)8AGGAAC (AAG)2AAC(AAG)2 (AAT)10GAAAAT (AAT)2GAAAAT (AAT)8 (AAT)13...(AAT)6 AA (AAT)4AAC(AAT) (AAC)7(AAT)2(AAC) (AAT)2(AAC)2 (TAG)9 Pdom 127b 119 48 Pdom 139 186 48, 45 Pdom 140 192 55 Pdom 151 115 52, 50 (CAT)2AA(CAT)CAAT (CAT)3 Ta, annealing temperature. Polistes dominulus (n = 8) Polistes fuscatus (n = 4) Polistes apachus (n = 4) Protopolybia Brachgastera Polybia Ropalidia Miscocyterus exigua mellifica occidentallis excavata alfkenii (n = 2) (n = 2) (n = 1) (n = 1) (n = 1) Primers (5′– 3′) HO = 0.38 (3) HE = 0.41 HO = 0.75 (4) HE = 0.63 HO = 0.75 (5) HE = 0.73 HO = 0.88 (4) HE = 0.63 HO = 0.50 (3) HE = 0.53 HO = 0.63 (2) HE = 0.43 HO = 1.00 (9) HE = 0.83 HO = 0.63 (6) HE = 0.78 HO = 1.00 (9) HE = 0.85 HO = 0.88 (9) HE = 0.80 HO = 0.88 (6) HE = 0.72 HO = 0.88 (9) HE = 0.85 HO = 0.00 (1) HE = 0.00 0.00 (1) –(2) n=1 0.33 (3) – (1) –(1) –(1) – (1) – (1) NP NP NP NP NP –(1) n=1 NP –(1) NP – (1) NP NP NP 0.50 (4) – (1) n=2 0.75 (6) 1.00 (5) – (1) n=1 NP NP NP NP NP NP NP NP 0.25 (4) 0.50 (5) – (1) –(1) NP – (1) NP 0.25 (2) –(2) n=2 0.00 (1) – (1) –(1) NP NP NP NP NP NP NP – (1) NP NP NP NP – (1) 0.00 (1) –(2) n=1 0.00 (1) – (1) –(1) –(1) – (1) – (1) 0.00 (1) 0.00 (1) NP NP NP NP – (1) 0.00 (1) 0.00 (1) – (1) –(1) –(1) NP NP 0.25 (2) –(1) – (1) –(1) NP – (2) – (1) 0.00 (1) 0.50 (2) 0.00 (1) F:GGACGCTCGGCTGATTTGTC R:AAGGGATTTTTCCTGAGACTATTCG F:CGTCTCTCGAAATATGCTAAAC R:AGAACGGTAAACATTCTTCTATC F:CACTGTATTGTCCTACGGTGGTCC R:GCGAGAACCTGTACTCAAAACAAAC F:TTCTCTGGCGAGCTGCACTC R:AGATGGCATCGTTTGAAAGAGC F:CATTATAAACGCCGCG R:ACGATGGAAACGTAAGTCC F:CCATCAGCTGTCCCATTCGC R:AATCGGTTTCGCTCGTCCACCTCC F:AAGAAAACCTACTACGTTGTGTGAG R:TTTCAACATTCCATAGGGACAG F:GAGTGGGTATGACGAAGATGATGG R:TGATTATAGCCTGCCGAAACTCTG F:CCGAAGAATGATAGTAGGTCC R:AGACCATCTCTCGCACGC F:TCCCCCGTTTTTGGTCCTTG R:GGGAGAGAATCGTGCCTTTTC F:TGACAAAAGACAACAAAATATG R:AGCTTCGGTAGGGCTTCG F:GCTTTTCCCTTATTTTCCCG R:CGTGTTCGTATATTCCTGTAACG F:TGATGTTACCACTGCTTTGAGCG R:TTCAGCACCGTCGTCGTTGTTG 2156 P R I M E R N O T E S Table 1 A description of polymorphic microsatellite loci isolated from Polistes dominulus, including their utility in related polistine taxa. The sample size (n) for each species is given in the column heading with exceptions for certain primers noted in the table. Where n ≥ 3, we report the observed heterozygosity for all species, as well as the expected heterozygosity for P. dominulus. In all cases we report the observed number of alleles in parentheses. The product size and repeat region data are based on the sequenced allele. NP = no scorable product. GenBank accession nos are AF155596 to AF155623 and include 16 additional loci not summarized in the table P R I M E R N O T E S 2157 and by a NSF predoctoral fellowship to MT Henshaw. I thank JE Strassmann and DC Queller for comments on the manuscript, Steffano Turillazzi and Rita Cervo for their help collecting wasps in Italy, and Aviva Liebert for help screening the loci in other species. References Cervo R, Lorenzi MC (1996) Behavior in usurpers and late joiners of Polistes biglumis bimaculatus (Hymenoptera: Vespidae). Insectes Sociaux, 43 (3), 255– 266. Field J, Solis CR, Queller DC, Strassmann JE (1998) Social and genetic structure of Papers Wasp Cofoundress Associations: tests of reproductive skew models. The American Naturalist, 151 (6), 545 – 563. Hughes CR (1998) Integrating molecular techniques with field methods in studies of social behavior: a revolution results. Ecology, 79, 383– 399. Mead F, Gabouriaut D, Habersetzer C (1995) Nest-founding behavior induced in the first descendants of Polistes dominulus Christ (Hymenoptera: Vespidae) colonies. Insectes Sociaux, 42 (4), 385 – 396. Peters JM, Queller DC, Imperatriz Fonseca VL, Strassmann JE (1998) Microsatellite loci for stingless bees. Molecular Ecology, 7, 783–792. Queller DC, Strassmann JE, Hughes CR (1993a) Microsatellites and kinship. Trends in Ecology and Evolution, 8 (8), 285–288. Queller DC, Strassmann JE, Solís CR, Hughes CR, DeLoach DM (1993b) A selfish strategy of social insect workers that promotes social cohesion. Nature, 365, 639–641. Queller DC, Zacchi F, Cervo R, et al. (2000) Unrelated helpers in a social insect. Nature, 405, 784–787. Reeve HK (1991) Polistes. In: The Social Biology of Wasps (eds Ross KG, Matthews RW), pp. 99–148. Cornell University Press, Ithaca. Strassmann JE (1981) Evolutionary implications of early male and satellite nest production in Polistes exclamans colony cycles. Behavioral Ecology and Sociobiology, 8, 55 –64. Strassmann JE, Solís CR, Peters JM, Queller DC (1996) Strategies for finding and using highly polymorphic DNA microsatellite loci for studies of genetic relatedness and pedigrees. In: Molecular Zoology: Advances, Strategies and Protocols (eds Ferraris JD, Palumbi SR), pp. 163–180, 528 –549. Wiley-Liss, Inc., New York. 2000 Graphicraft PRIMER 9PRIMER 101112 02 NOTEs NOTEs Limited, Hong Kong Characterization of nuclear microsatellites in Pinus halepensis Mill. and their inheritance in P. halepensis and Pinus brutia Ten. R . N . K E Y S , * A . A U T I N O , † K . J . E D WA R D S , ‡ B . FA D Y , * C . P I C H O T * and G. G. VENDRAMIN† *Institut National de la Recherche Agronomique, Unité des Recherches Forestières Méditerranéennes, Avenue Vivaldi, 84000 Avignon, France, †Istituto Miglioramento Genetico Piante Forestali, Consiglio Nazionale delle Ricerche, via Atto Vanucci 13, 50134 Firenze, Italy, ‡IACR-Long Ashton Research Station, University of Bristol, Bristol BS41 9AF, UK © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Keywords: microsatellite primers, Pinus brutia, Pinus halepensis Received 25 May 2000; revision received 29 June 2000; accepted 24 July 2000 Correspondence: B. Fady. Fax: +33 4 90 13 59 59; E-mail: fady@avignon.inra.fr Nuclear microsatellites, or single sequence repeats (nSSRs), have been characterized in many tree species and are powerful markers for genetic diversity studies in natural populations (e.g. Echt et al. 1996; Pfeiffer et al. 1997). Although nSSR enrichment protocols have successfully been applied to conifers (Edwards et al. 1996), identification of single-locus, reproducible markers is difficult, probably because of their large genome size and complexity (Pfeiffer et al. 1997; Soranzo et al. 1998). In this study, we report the successful isolation of nSSRs in Pinus halepensis Mill. and their Mendelian segregation in both P. halepensis and P. brutia, two closely related Mediterranean pines. A microsatellite library enriched for di- (GC, CT, CA), tri- (CAA, GCC) and tetra-nucleotide (GATA, CATA) repeats was constructed for Pinus halepensis, following the method described by Edwards et al. (1996). A total of 43 clones containing a microsatellite were detected from 47 clones randomly chosen from the library: 16% were repetitions of a single nucleotide (A/T), 77% were repetitions of dinucleotides (CA, CT or compounds CA–TA, CA–GA) and 7% were repetitions of trinucleotides (TAA, GCC). Sequencing reactions were performed using the Pharmacia AutoRead Sequencing Kit, and run on a 6% polyacrylamide gel containing 7 m urea using an ALF Pharmacia automatic sequencer. Primers were designed for the amplification of 25 dinucleotide nSSRs using the computer program Primer (http://wwwgenome.wi.mit.edu/genome_software/other/primer3.html). Total genomic DNA extracted from leaf and megagametophyte tissue was used for testing the primer pairs. The procedure described by Doyle and Doyle (1990) and the Nucleon Phytopur DNA extraction kit were used for leaf tissue and mega-gametophytes, respectively. Polymerase chain reaction (PCR) was carried out using a Gradient 96 Stratagene Robocycler: the reaction solution (25 µL) contained four dNTPs (each 0.2 mm), 0.25 µm of each primer, 2.5 µL reaction buffer (100 mm Tris–HCl pH 9.0, 15 mm MgCl2, 500 mm KCl), 25 ng of template DNA and 1 unit of Taq polymerase (Pharmacia). After a preliminary denaturing step at 95 °C for 1.5 min, PCR amplification was performed for 35 cycles: 1.5 min denaturing at 94 °C, 1.5 min at annealing temperature (Table 1) and 1.5 min extension at 72 °C, with a final 5 min step at 72 °C. After amplification, PCR products were mixed with a loading buffer (98% formamide, 10 mm EDTA pH 8.0, 0.1% bromophenol blue, 0.1% xylene cyanol and 10 mm NaOH), heated for 5 min at 95 °C, and then set on ice. Fragments were electrophoretically separated on a 6% polyacrylamide gel and stained using silver nitrate (Rajora et al. 2000). Out of 25 primer pairs, nine (36%) either gave no amplification (n = 4) or produced multi-band patterns (n = 5). Sixteen produced fragment amplification in the expected size range, of which eight were polymorphic within one or the other species (Table 1). This proportion of functional markers is comparable to what is generally observed in conifers (e.g. Echt 2158 P R I M E R N O T E S Table 1 Primers and characteristics of seven microsatellite loci that were polymorphic either within Pinus halepensis or within Pinus brutia* © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Locus† Repeat sequence PHAF01 (CA)18 PHAF02 (CA)15 PHAF05 (CA)17 PHAF07 (CT)16 PHAF08 (CT)25 PHAF09 (CT)18 PHAF10 (CA)17(TA)3 Primer sequence (5′ → 3′) F: TTCAGATATGGTCCATGGATG R: GATCACAATGTCATTATCGGG F: TGGCAATGGAAACCTGATAC R: GCCCCACCATCATATCTCTTTAG F: TCATAAGCCCTTTGTTTCTTTTC R: TTTTTCGCCCTGTATTTTCTG F: ATCAGCTTAGTAGGTCTCGCC R: AGACACTAAAGGGGAGTCCG F: TTCCACATTGTATTTTGATGCT R: AACTTTGGAAGTGACCAAATGT F: ACTAAGAAACGGTGTGATGCTG R: CTTCGCATAGGCATGCATAC F: TCCTTTCTTGTTCTTGGTAACTG R: ACCGCGGATTATAACCTGTG Annealing temp. (°C) MgCl2 (mm) Expected size (bp) Number of alleles‡ Heterozygosity (HO/HE)§ Number of megagametophytes per bi-allelic combination 54 2.5 194 3/3 0.611/0.538 15 0.795 AF195535 54 2.5 149 3/3 0.550/0.609 15 0.795 AF195536 56 3.5 125 4/4 0.611/0.624 20, 15, 8 0.655, 0.197, 1 AF195540 54 2.5 123 3/3 0.700/0.676 13, 9 0.782, 0.739 AF195541 53 4.5 150 2/1 0.500/0.479 19 0.251 AF195542 59 2.5 198 2/1 0.600/0.505 19 0.819 AF195538 53 2.5 129 4/4 0.529/0.665 19, 16 0.108, 1 AF195543 χ2 test (P value) Accession no. *An eighth locus, ITPF4516 (accession AJ012087) tested in P. pinaster (Mariette et al. 2000), is polymorphic in P. halepensis and P. brutia (four common alleles in both species). †PHAF, Pinus halepensis Avignon Firenze. ‡Values are for P. halapensis/P. brutia. In loci PHAF08 and PHAF09, P. halepensis and P. brutia do not share common alleles (sizes 205 and 155 bp respectively). §HO is the frequency of heterozygotes in the sample and HE is the unbiased expected heterozygosity (Nei 1978), where HE = (2n/2n – 1) (1 – Σpi2). P R I M E R N O T E S 2159 et al. 1996; Pfeiffer et al. 1997). A single marker was found to be polymorphic in Pinus pinaster when the same 25 primer pairs were tested (Mariette et al. 2000). Transfer of nSSR markers across species of the same genus is generally difficult in conifers (e.g. Echt & May-Marquardt 1997), and the results thus confirm the close taxonomic relatedness between P. halepensis and P. brutia. nSSR polymorphism was screened at population level using 50 P. brutia individuals (two populations) and 47 P. halepensis individuals (three populations). The maximum number of alleles per locus was four, and the expected heterozygosity per locus was between 0.479 and 0.676 (Table 1), which is lower than observed for other conifers, e.g. Pinus sylvestris (Soranzo et al. 1998) or Picea abies (Pfeiffer et al. 1997), but higher than found using isozymes (Schiller et al. 1986; Teisseire et al. 1995). Mendelian segregation was tested on 1– 3 bi-allelic combinations in all polymorphic loci (Table 1). No significant deviation from the expected 1:1 ratio was observed. nSSRs are thus potentially helpful markers for studying population diversity in P. halepensis and P. brutia. Acknowledgements This study was supported by the European Union, contract FAIR CT95-0097 ‘Mediterranean Pinus and Cedrus’. Many thanks to B. Jouaud for technical assistance. References Doyle JJ, Doyle JL (1990) Isolation of plant DNA from fresh tissue. Focus, 12, 13 –15. Echt CS, May-Marquardt P (1997) Survey of microsatellite DNA in pine. Genome, 40, 9–17. Echt CS, May-Marquardt P, Hseih M, Zahorchak R (1996) Characterization of microsatellite markers in eastern white pine. Genome, 39, 1102–1108. Edwards KJ, Barker JHA, Daly A, Jones C, Karp A (1996) Microsatellite libraries enriched for several microsatellite sequences in plants. Biotechniques, 20, 758–760. Mariette S, Chagne D, Decroocq S, Vendramin GG, Lalanne C, Madur D, Plomion C (2000) Microsatellite markers for Pinus pinaster Ait. Annals of Forest Science, in press. Nei M (1978) Estimation of average heterozygosity and genetic distance from a small number of individuals. Genetics, 89, 583– 590. Pfeiffer A, Olivieri AM, Morgante M (1997) Identification and characterization of microsatellites in Norway spruce (Picea abies K.). Genome, 40, 411– 419. Rajora OP, Rahman MH, Buchert GP, Dancik BP (2000) Microsatellite DNA analysis of genetic effects of harvesting in old-growth eastern white pine (Pinus strobus) in Ontario, Canada. Molecular Ecology, 9, 339–348. Schiller G, Conkle MT, Grunwald C (1986) Local differentiation among Mediterranean populations of Aleppo pine in their isoenzymes. Silvae Genetica, 35, 11–18. Soranzo N, Provan J, Powell W (1998) Characterisation of microsatellite loci in Pinus sylvestris L. Molecular Ecology, 7, 1247 –1248. Teisseire H, Fady B, Pichot C (1995) Allozyme variation in five French populations of Aleppo pine (Pinus halepensis Mill.). Forest Genetics, 2, 225–236. 2000 Graphicraft PRIMER 9PRIMER 11113 32 NOTEs NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Microsatellite markers for behavioural studies in a semi-fossorial shrew (Soricidae: Anourosorex squamipes) H O N - T S E N Y U and Y U - Y I N G L I A O Department of Zoology, National Taiwan University, Taipei, Taiwan, ROC 106, Republic of China Keywords: Anourosorex squamipes, behavioural genetics, fossorial, microsatellite, Soricidae Received 23 June 2000; revision accepted 24 July 2000 Correspondence: Alex Hon-Tsen Yu. Fax: +886 2 23638179; E-mail: ayu@ccms.ntu.edu.tw Genetic information revealed by microsatellite markers is useful for inferring social behaviours in animals (Garza et al. 1997), particularly for species that lead a secretive life style. The mole shrews (Anourosorex squamipes) are semi-fossorial, living underground and digging burrows but also coming to the forest floor to search for food (Hutterer 1985). Yu (1994) suggested that several mole shrews might share the same burrow system, as three or four mole shrews were often caught successively by one trap placed on the same spot. Thus, the mole shrew may have the peculiar social structure and behaviour common to some other subterranean mammals (Nevo 1979). As a preparatory step for studying behavioural genetics, we have characterized 11 microsatellite loci that are polymorphic and suitable for use to address questions regarding social structure in Anourosorex squamipes. Genomic DNA for constructing the partial libraries was prepared according to procedures described by Sambrook et al. (1989). Genomic DNA was digested with Sau3A and fractioned in a 2.5% NuSieve™ GTG gel (FMC, Rockland, ME, USA). DNA of size range of 300–700 bp was isolated, purified with a GeneClean III kit (Bio101 Inc.) and ligated into plasmid PUC18/ BamHI/BAP (Pharmacia, Vista, CA, USA) according manufacturer’s protocols. Ligated plasmids were transformed into competent SURE cells or XL-2 Blue ultracompetent cells (Stratagene). Recombinant clones containing inserts were transferred to Hybond N+ nylon membranes (Amersham), which were hybridized to a set of six oligonucleotide probes: (AC)10, (TC)10, (CAC)5CA, CT(ATCT)6, (TGTA)6TG and CT(CCT)5. Probes were labelled with a DIG Oligonucleotide 3′-End Labelling Kit (Boehringer Mannheim). Hybridization was performed at 45 °C for 16 h in a standard hybridization buffer consisting of 5 × SSC, 0.1% N-lauroylsarcosine, 0.2% SDS and 1% blocking reagent (Boehringer Mannheim). The membranes were washed twice for 5 min at 45 °C, with a solution of 2 × SSC, 0.1% SDS, and then twice for 15 min at 65 °C with a solution of 0.1 × SSC, 0.1% SDS. Chemi-luminescent detection was performed with a DIG Luminescent Detection Kit (Boehringer Mannheim). The exposure time ranged from 15 to 30 min. Positive clones were chosen for sequencing to confirm suitable length and base composition. The sequencing reactions were performed with a Big Dye dye-terminator kit, following the manufacturer’s protocols, and analysed on polyacrylamide gels with an ABI 377 automated sequencer (Perkin-Elmer Applied Biosystems). The online program 2160 P R I M E R N O T E S Table 1 Characteristics of 11 polymorphic microsatellite loci in Anourosorex squamipes, including repeat motif, primer sequences, annealing temperature, allele size range, number of alleles, observed heterozygosity (HO) and expected heterozygosity (HE) Locus* Repeat motif Primer sequences (5′ → 3′) AS1 (AC)15 AS2 (TC)9(TG)6 AS3 (TG)3TA(TG)18 AS4 (TGTC)5(TC)11(AC)6 AS5 (CA)17 AS6 (AC)13 AS7 (TG)14 AS8 (TG)12 AS9 (TG)12 AS10 (CA)26 AS11 (CCA)6CCG(CCA)8 GGATTCTATTTCATTCTTGAGTCAC GTAAAACTCTGGCTGGTGCC CCTGGTTTGACCTCATGTTTGG GACAGAGAGAGATGGGTGGGG TTCCGCCTTGTACTTTGCTG CCCCGGGGATCCAGTGTCTTAC GGATCCTTCCAGCGTTCTCTCTC GCAGCATGTTTCCCCAGTGTC AGGCAAACGCTTTACCCTTG TGTAGAAGGCTGGAGAGACAGTG GGTATGGAGGCACACAACGG TGCTTGCCAGTCTTCTCTGCG CGCATGCGTGTGTGTGAATC CCAGGTGTGCCCTTGAAACC TGCTCAAAAGCAATGCTAGCTG GTTCCAAGGACAATGCACGG CGCACTTTTGTTGTTGTATGCG TTCCTGGCGCCCCATAATAG GGGGCCTATTCCCCTGTTTC GGATGAGGGAATCCAGAAGACG AGCCACAGGTTTCCACCCAC TTCCGCCTGTCTGCTTCTCC Annealing temp. (°C) Allele size range (bp) Number of alleles HO HE 53 129–155 10 0.75 0.88 58 136–166 15 0.56 0.89 56 118–138 20 0.67 0.93 53 140–164 11 0.78 0.89 56 94 –112 17 0.58 0.89 56 96 –126 13 0.56 0.86 53 120–150 13 0.36 0.82 52 112–138 13 0.53 0.88 50 126–148 19 0.58 0.93 56 79 –111 20 0.92 0.96 56 80 –119 19 0.33 0.88 *GenBank accession nos (order listed in table): AF261959–AF261969. Primer 3.0 (http://www.genome.wi.mit.edu) was used to design primers from flanking regions of microsatellite DNA loci that contain more than 10 repeat units. Individual genotypes were determined by polymerase chain reaction (PCR). PCR reactions were performed either with non-radioactive primers or radioactive primers. For nonradioactive PCR, 25 µL reactions were performed, containing 200 ng template DNA, 10 mm Tris–HCl, 50 mm KCl, 0.1% Triton X-100, 0.75 mm Mg2+, 0.15 mm dNTP, 0.5 µm of each primer and 2 units Taq DNA polymerase (Promega). Amplification was carried out according to the thermal profile: 95 °C for 4 min, followed by 25 cycles of 94 °C for 30 s, optimal annealing temperature (Table 1) for 30 s and 72 °C for 30 s, with a final extension step at 72 °C for 7 min. PCR products were run on 6% native polyacrylamide gel, stained by ethidium bromide and visualized on a UV light box. The non-radioactive PCR was used to screen for polymorphic loci and the initial round of genotyping. For radioactive PCR, one primer from each pair was 5′ end-labelled with [γ 32P]-ATP (NEN) and T4 polynucleotide kinase (Promega, Boston, MA, USA), following the manufacturer’s protocols. Each PCR reaction totalled 10 µL, containing 200 ng template DNA, 10 mm Tris–HCl, 50 mm KCl, 0.1% Triton X-100, 0.25 mm dNTP, 0.2 µm of each unlabelled primer, 0.6 mm Mg2+, 0.25 units Taq DNA polymerase (Promega) and 0.5 pmol [γ 32P]-ATP labelled primer. Amplification was carried out according to the thermal profile: 95 °C for 3 min, followed by 25 cycles of 95 °C for 15 s, optimal annealing temperature (Table 1) for 2 min and 72 °C for 2 min, with a final extension step at 72 °C for 7 min. PCR products were run on a regular denaturing 6% polyacrylamide sequencing gel. The sizes of alleles were estimated by using control DNA (PUC18) from a Thermo Sequenase Cycle Sequencing Kit (Amersham) as markers. The radioactive PCR was used for a second round of screening: all the alleles of different sizes detected in the first round of screening were run on comparison gels to accurately determine their sizes. Running radioactive PCR products on denaturing gels also helps reduce the confusion caused by the heteroduplex bands that sometimes appeared in the first round of screening. Eleven clones were confirmed to be polymorphic (Table 1) by typing 36 mole shrews collected from Taiwan. The number of alleles per locus ranged from 10 to 20, and the observed and expected heterozygosities ranged from 0.33 to 0.92 and from 0.82 to 0.96, respectively. The observed genotypes deviated from Hardy–Weinberg expectation at the 11 loci (all P < 0.05), resulting from heterozygote deficiency, which may be caused by combining samples from various disparate localities (Wahlund’s effect). Acknowledgements Chu-Fong Lo and members of her laboratory offered technical support for molecular cloning. Financial aid was granted to HTY by the National Science Council (89-2311-B-002-029, 88-2311-B002-051). © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2161 References Garza JC, Dallas J, Duryadi D, Gerasimov S, Croset H, Boursot P (1997) Social structure of the mound-building mouse Mus spicilegus revealed by genetic analysis with microsatellite. Molecular Ecology, 6, 1009–1017. Hutterer R (1985) Anatomical adaptations of shrews. Mammal Review, 15, 43 – 55. Nevo E (1979) Adaptive convergence and divergence of subterranean mammals. Annual Review of Ecology and Systematics, 10, 269– 308. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning. A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Yu HT (1994) Distribution and abundance of small mammals along a subtropical elevational gradient in central Taiwan. Journal of Zoology, London, 234, 577–600. 2000 Graphicraft Primer 932 11115 notesLimited, Hong Kong Isolation and characterization of microsatellite DNA markers in the Florida manatee (Trichechus manatus latirostris) and their application in selected Sirenian species A. I. GARCIA-RODRIGUEZ,* D . M O R A G A - A M A D O R , † W. FA R M E R I E , ‡ P. M C G U I R E § and T. L . K I N G ¶ *United States Geological Survey, Biological Resources Division, Sirenia Project, Gainesville, FL 32601, USA, †Education and Training Core, Interdisciplinary Center for Biotechnology Research, University of Florida, Gainesville, FL 32610, USA, ‡ Molecular Services Core, Interdisciplinary Center for Biotechnology Research, University of Florida, Gainesville, FL 32610, USA, §Department of Biochemistry and Molecular Biology, University of Florida, Gainesville, FL 32610, USA, ¶United States Geological Survey, Biological Resources Division, Aquatic Ecology Laboratory, Leetown Science Center, 1700 Leetown Road, Kearneysville, WV 25430, USA Keywords: Dugong dugong, microsatellite DNA, Trichechus inunguis, Trichechus manatus Received 3 February 2000; revision received 2 July 2000; accepted 27 July 2000 Correspondence: T.L. King. Fax: 304 724 4498; E-mail: tim_king@usgs.gov The West Indian manatee (Trichechus manatus) inhabits subtropical and tropical waters of the Caribbean Sea from the southern USA to Brazil’s north-east coast. Two sub-species are recognized, the Florida manatee (T. m. latirostris) and the Antillean manatee (T. m. manatus) (Domning & Hayek 1986). Abundant biological and ecological data for the Florida manatee have been collected, and the information has formed the basis for management and conservation programmes. However, to plan and implement biologically sound management programmes for this marine mammal, knowledge of the amount of genetic diversity present and a thorough understanding of the evolutionary relationships among geographical populations are essential. Genetic studies employing allozymes (McClenaghan & O’Shea 1988) and mitochondrial DNA (Bradley et al. 1993; Garcia-Rodriguez et al. 1998) have identified © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 low levels of genetic diversity, and failed to resolve population structure for the Florida manatee. A technique with a higher resolution of genetic population structure and pedigree analysis is needed. We report the development and characterization of microsatellite DNA markers in the Florida manatee and test the utility of these markers in three closely related Sirenian species. Two methodologies were used to generate microsatelliteenriched libraries for T. m. latirostris. Four enriched libraries were produced by Genetic Identification Services (Chatworth, California, USA) using proprietary magnetic bead capture technology. An additional library was constructed and screened for polymorphic loci following a protocol modified from Armour et al. (1994). For this protocol, approximately 50 µg of manatee genomic DNA were digested with Sau3AI (Life Technologies, Rockville, Maryland, USA), gel-fractionated to isolate 0.4 –1.0 kbp fragments, and ligated to Sau3AI linkers. Polymerase chain reaction (PCR) amplifications were performed in a 100 µL volume containing 15 ng of purified DNA, 50 mm KCl, 10 mm Tris–HCl (pH 9.0), 0.1% Triton X-100, 0.25 mm MgCl2, 0.2 mm dNTPs (Applied Biosystems, Foster City, California, USA), 1 µm Sau-L-A primer and 2.5 units of Taq DNA polymerase (Promega, Madison, Wisconsin, USA). The following amplification programme was used: 94 °C for 3 min, 30 cycles of 94 °C for 45 s, 68 °C for 45 s and 72 °C for 1.5 min, followed by 72 °C for 10 min. Purified PCR products were denatured by alkali treatment and hybridized to nylon filters containing (CA)n oligonucleotide repeats. Hybridization was performed overnight at 65 °C, and 5 µL of the recovered hybridized molecules were used for a 100 µL PCR amplification of microsatellite-enriched genomic DNA fragments following the amplification and PCR conditions described above. PCR products were directly ligated to pCR®2.1 (Invitrogen, Carlsbad, California, USA) followed by transformation into INVαF′ One Shot™ competent cells (Invitrogen). A total of 186 colonies were screened for (CA)ncontaining inserts using alkaline phosphatase-conjugated (TG)n oligomer and a chemi-luminescent detection system (FMC BioProducts Corp., Rockland, Maine, USA). Following a secondary screening, 60 positive colonies were sequenced using the ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems) employing M13 forward and reverse primers. Sequencing reactions were electrophoresed on an ABI 377 automated sequencer (Applied Biosystems). From the two sets of libraries, primers were designed in the flanking regions of 61 microsatellite-bearing clones using oligo 5.1 (National Biosciences, Molecular Biology Insights Inc., Cascade, Colorado, USA). Microsatellite DNA amplification reactions consisted of 200 ng DNA, 10 mm Tris –HCl (pH 8.3), 50 mm KCl, 1.5 mm MgCl2, 0.20 mm dNTP, 5 pmol of forward and reverse primer and 1.0 U Taq DNA polymerase (Promega) in a total volume of 20 µL. The forward primer was 5′ modified with either TET, FAM or HEX fluorescent labels (Applied Biosystems). Amplification was performed in a Biometra® UNO II thermal cycler using the following conditions: 94 °C for 2 min, 34 cycles of 94 °C for 30 s, 54 °C for 30 s and 72 °C for 30 s, and a final extension at 72 °C for 10 min. Amplified fragments were subjected to fragment analysis on an Manatee species Florida © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Locus Size Repeat type (and length) N HO HE Primer sequences (5′ → 3′) TmaA01 107 (TA)3(CA)3CG(CA)7 1 0.00 0.00 TmaA02 247– 251 (CACT)2(CA)16 3 0.51 0.54 TmaA03 163–183 (GACA)4 2 0.30 0.40 TmaA04 204 (CT)2(GT)12AT(GT)7AT(GT)2 1 0.00 0.00 TmaA09 150 (GT)15 1 0.00 0.00 TmaE02 172–174 (GT)13 2 0.44 0.46 TmaE08 149–165 (CA)13TA(CA)5 3 0.47 0.55 TmaE11 177 –197 (CA)13 6 0.58 0.63 TmaE26 199– 201 (CA)8C(CA)17 2 0.24 0.26 TmaF14 204– 206 (TC)6(TG)2TA(TC)5TG(TC)3 2 0.24 0.32 TmaF34 271 1 0.00 0.00 TmaH1I 298 1 0.00 0.00 TmaM61 176 (TCTCTCTCTTTCTG)2(TC)4 TT(TC)3 (AC)8AT(AC)9 (TCTG)4(TCTA)5CCTGTCTATCCA (TCTA)3CCTG(TCTA)8CCTG(TCTA)5 (TG)3(GT)17 1 0.00 0.00 TmaM79 154–156 (GT)15 2 0.56 0.54 F-CAGAAGGGATACATATACA R-CAGCCCCTGGCTGTCTCTTGTC F-CTCAGTCCAAACAGCTAATG R-TAGTCATTTGTGCAGAGTGC F-ACATGTGTTCCCTGCTGTAT R-GATTTTTGGAGCAGTTGTCA F-GAACACAAGACCGCAATAAC R-TGGTGTATCACTCAGGGTTC F-GATGGGATACTGGGTTATGC R-ATGCAGACACTGGACATAGG F-GTCTCTACGGCCTAGAATTGTG R-TTTCTCTACCTCTCCTCACACG F-GAATAGAGACTGGGCTAGAATCC R-GCCTTTTGGAGGGATAGAAGTAG F-ACACACAACATCACTCATCCAC R-AAGCTGCGTTCTACTTCATATAATC F-CATTCCTGATCCACAAAATC R-CCTGTCTTCTCTCTGTTTCTCC F-CTAAGACATTGCTCCAAAAGC R-GGGCAGTGGGATTTGAGATG F-CATGAGAGACTATGCTCCCTTC R-CAGGTAGGAAGATGATGAGGAC F-AGCAGATAGACACACTGGGAAG R-GAGTCTGAATGAATGAATTACTGC F-TTGAGGTGTAATCTGTGTG R-GGTAATCGGAGTTGGTGTA F-CCAATCATGTCCCAAACT R-CAATAGAAGAAGCAGCAG Antillean N Amazonian N Dugong N GenBank accession no. 2 2 2 AF223649 3 5 2 AF223650 4 3 1 AF223651 3 1 3 AF223652 1 4 3 AF223653 2 1 3 AF223656 4 2 3 AF223657 8 1 3 AF223658 5 3 2 AF223659 2 2 1 AF223660 1 2 — AF223661 1 3 1 AF223662 2 2 1 AF223655 3 3 2 AF223654 Tests for goodness of fit to Hardy–Weinberg expectations suggested that there were no significant differences between observed and expected values (Raymond & Rousset 1995). The results of cross-species amplification of these markers in three other Sirenian taxa are also provided: Antillean manatee (Trichechus manatus manatus), n = 21 animals surveyed; Amazonian manatee (Trichechus inunguis), n = 7; and the dugong (Dugong dugong), n = 3. ‘—’ indicates no or sub-optimal amplification products in cross-species tests. N, number of alleles observed. 2162 P R I M E R N O T E S Table 1 Expected size of fragment (bp), repeat type, number of alleles detected, observed and expected levels of heterozygosity, primer sequence, and GenBank accession nos for 14 Trichechus manatus latirostris microsatellite DNA markers surveyed in 50 animals collected throughout Florida, and the results of cross-species amplification of these markers in three other Sirenian taxa P R I M E R N O T E S 2163 ABI PRISM™ 310 Genetic Analyser (Applied Biosystems). Genescan™ 2.1 and Genotyper™ 2.1 Fragment Analysis software (Applied Biosystems) were used to score, bin and output allelic (and genotypic) data. Fourteen sets of primers amplified fragments of expected size from Florida manatee genomic DNA (Table 1). These markers were screened in 50 manatees collected throughout the Florida peninsula. Eight of the 14 loci were polymorphic in this initial survey, and overall levels of heterozygosity averaged 41%. Low levels of allelic diversity were observed in the Florida manatee. The maximum number of alleles identified was six (TmaE11), and the average number of alleles observed at polymorphic loci was 2.9. This paucity of genetic diversity suggests a founder effect or major population bottleneck of evolutionary significance (see GarciaRodriguez et al. 1998). In addition, this study reports one of the lowest levels of genetic diversity observed in species-specific microsatellite DNA markers [see Nyakaana & Arctander 1999 (African elephant); Waldick et al. 1999 (right whale) ]. Cross-species amplification was tested in three Sirenian taxa: the Antillean manatee (T. m. manatus), the Amazonian manatee (T. inunguis) and the dugong (Dugong dugong). Eleven of 14 markers were polymorphic for the Antillean and the Amazonian manatee (Table 1). At least nine markers were polymorphic in the dugong; the polymorphism is likely to be under-estimated due to the small sample size (n = 3). This suite of markers appears to be ideal for the identification of population structure and possibly pedigree analysis in all four Sirenian species, and provides a nuclear DNA-based approach to complement existing mitochondrial DNA genetic information for these vulnerable species. References Armour JAL, Neumann R, Gobert S, Jeffreys AJ (1994) Isolation of human simple repeat loci by hybridization selection. Human Molecular Genetics, 3, 599–605. Bradley JL, Wright SD, McGuire PM (1993) The Florida manatee: cytochrome b DNA sequence. Marine Mammal Science, 9, 197–202. Domning DP, Hayek LC (1986) Interspecific and intraspecific morphological variation in manatees (Sirenia: Trichechus). Marine Mammal Science, 2, 87–144. Garcia-Rodriguez AI, Bowen BW, Domning D, et al. (1998) Phylogeography of the West Indian manatee (Trichechus manatus): how many populations and how many taxa? Molecular Ecology, 7, 1137–1149. McClenaghan LR Jr, O’Shea TJ (1988) Genetic variability in the Florida manatee (Trichechus manatus). Journal of Mammalogy, 69, 481– 488. Nyakaana S, Arctander P (1999) Population genetic structure of the African elephant in Uganda based on variation at mitochondrial and nuclear loci: evidence for male-biased gene flow. Molecular Ecology, 8, 1105–1115. Raymond M, Rousset F (1995) genepop (version 1.2): population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248– 249. Waldick RC, Brown MW, White BN (1999) Characterization and isolation of microsatellite loci from the endangered North Atlantic right whale. Molecular Ecology, 8, 1753–1768. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Microsatellite loci for two European sciurid species (Marmota marmota, Spermophilus citellus) 2000 Graphicraft 932 Primer 11116 notesLimited, Hong Kong S . H A N S L I K * and L . K R U C K E N H A U S E R † *Department of Animal Breeding and Genetics, University of Veterinary Medicine Vienna, A-1210 Vienna, Austria, †Museum of Natural History Vienna, 1st Zoology Department, Burgring 7, A-1014 Vienna, Austria Keywords: Marmota marmota, microsatellite, primer, population genetics, Spermophilus citellus Received 10 May 2000; revision received 19 June 2000; accepted 29 July 2000 Correspondence: L. Kruckenhauser. Fax: +43 15235254; E-mail: Luise.Kruckenhauser@univie.ac.at Two species of European sciurid rodents are of particular interest for behavioural ecology and population genetics: Marmota marmota and Spermophilus citellus. The Alpine marmot (M. marmota) inhabits higher elevations of the European Alps and some isolated mountain massifs. Autochthonous populations occur only in the Alpine core area and in a small area near Berchtesgaden. The distribution of the European groundsquirrel (S. citellus) comprises the grassland of the Pannonian plain ranging from eastern Europe to the foothills of the Alps. It is presently listed as endangered (Berner Convention 1999). We isolated six new microsatellite markers for each of the two species (M. marmota: L. Kruckenhauser; S. citellus: S. Hanslik). Genomic DNA was extracted from frozen liver (M. marmota) or ethanol-stored tissue samples from the tail (S. citellus) using a standard phenol– chloroform extraction method (Sambrook et al. 1989). Following the protocol of Rassmann et al. (1991), partial genomic libraries were established for M. marmota and S. citellus and around 1400 clones from each species were screened for the presence of microsatellite sequences using a dinucleotide simple sequence polymer probe AC/GT. Fifty-eight marmot clones showed a positive signal. Twentytwo were sequenced using the SequiTherm EXCEL™ II DNA Sequencing Kit (Epicentre Technologies) with biotinylated primers and the SAAP/CSPD detection system (US Biochemicals, Inc.). Primer pairs were synthesized for 11 loci; six of these microsatellite loci showed unambiguous allelic patterns in M. marmota (Table 1). Polymerase chain reaction (PCR) amplifications were performed on a HYBAID Omnigene thermocycler in a volume of 12.5 µL containing 10 mm Tris –HCl (pH 8.8), 1.5 mm MgCl2, 150 mm KCl, 0.1% Triton X-100, 0.25 U DynaZyme DNA polymerase (Finnzymes OY), 2 pmol of each primer (forward primer labelled with IRD-800), 200 µmol of each dNTP, 0.25 µL DMSO and 50 ng template DNA. The amplification protocols were as follows: 94 °C for 5 min, then two cycles of 94 °C for 20 s, annealing temperature plus 6 °C for 20 s, 70 °C for 20 s, then 30 cycles of 94 °C for 30 s, annealing temperature for 20 s, 70 °C for 20 s, and finally 72 °C for 2 min. PCR products were separated on 6% denaturating polyacrylamide gels in a Li-Cor automatic sequencer. Analysis of PCR fragments was carried out using RFLPscan (Scanalytics). The six loci were tested in 19 individuals of M. marmota from the Austrian allochthonous population Turracher Nockberge. In addition, 10 individuals of S. citellus were cross-tested with the same primer sets. 2164 P R I M E R N O T E S Table 1 Primer sequences (5′ → 3′) of microsatellites from Marmota marmota (MS6, MS41, MS45, MS57, MS53, MS56) and Spermophilus citellus (ST7, ST10, SB10, SC2, SC4, SX), GenBank accession nos, repeat motifs and annealing temperatures Locus Repeat motif Primer Accession no. Annealing temp. (°C) MS6 (GT)20 AF259372 53 MS41 (GT)11 AF259373 53 MS45 (GT)13 AF259374 53 MS47 AF259375 50 MS53 (GT)4TC(GT)3AT(GT)7GAGG (GA)4TT(GA)3AA(GA)11 (GT)18 AF259376 53 MS56 (CA)14 AF259377 53 SB10 (GA)12(TG)18 AF254435 50 SC2 (GA)31 AF254438 56 SC4 (GT)20 AF254437 56 ST7 AF254439 50 ST10 (TGG)7T(GT)2 AT(GT)7AT(TG)8 (CA)12 AF254436 52 SX (GA)25 F: CTGATGGGGTTAAGATTGCC R: CCCCACTGACCCACCTCC F: GGTGTATATGGGAATAGGGGG R: GCCTTCAAATCAAAGCAGGTTG F: CTGTCTCTTTGTCCCTGCC R: CTCCTTACCATCATCTTTCCG F: CCTGATGTAGTCAGTCAG R: TGTGGGAAATGGCACATC F: ATTGAGGAGCAGCATCTAGG R: TCAGGGAAAGGCAGACCTG F: CAGACTCCCACCAGTGACC R: CCTGATCTATGTAGGTTCCAT F: TCTGTTTAGTTCATTTGCCATTT R: TCAAGAGAGGTCCTACAGAATGA F: CATCATGGCAGAAGATGTGG R: TTGACTGGAAGTGGGACTCTC F: AAAAGCGTGCATTGCCTTAC R: CCTCTCAAGACGGGCAGA F: GAATCTTGACTCCTGAGATA R: CCATCTCCTGACATTTAATA F: TTGTGATCCTCCAGGGAGTT R: GTGATTTCCAAACCCCATTC F: TTTTCCTCTCCTGAATGCTTTT R: CAAAGATGTTGTGTCCGACG AF254440 56 Thirty-six positive ground-squirrel clones were sequenced with the M13-40 forward primer using Sequenase version 2.0 (Amersham Life Sciences). Sequencing products were separated on a 4% denaturing polyacrylamide gel and visualized autoradiographically. Primers were designed with the oligo software package (National Biosciences Inc., version 5.0). PCR amplification was carried out on a HYBAID Omnigene thermocycler in 10 µL reaction volume with 10 mm Tris –HCl (pH 9.0), 50 mm KCl, 1.5 mm MgCl, 0.1% Triton X-100, 0.2 mg/mL BSA, 200 µm dNTPs, 1 µmol of each primer (0.02 pmol forward primer end-labelled with γ 32P), 50 –100 ng template DNA, and 0.5 units Taq DNA polymerase. A 4 min initial denaturation at 94 °C was followed by 30 cycles of 1 min at 94 °C, 1 min at 47– 61 °C (depending on the primer combination), 1 min at 72 °C, and a final extension at 72 °C for 45 min. PCR products were separated on a 7% denaturing polyacrylamide gel. Alleles were sized by running a sequencing reaction of M13 next to the amplified microsatellites. Six primer pairs yielded clear amplification products in S. citellus (Table 1). The six loci were analysed in 54 ground-squirrel and 10 marmot individuals. Observed and expected heterozygosities were calculated using genepop (version 1.2; Raymond & Rousset 1995). Altogether 12 microsatellite loci were tested in both species, the results for these are shown in Table 2. Ten loci amplified in both species, two amplified in M. marmota only. All loci were polymorphic in at least one of the two species, and up to seven different alleles were observed in one species. Significant deviations from the Hardy–Weinberg expectations as calculated with the program genepop (version 1.2; Raymond & Rousset 1995) were found for the loci SB10 (P = 0.0038) and SX (P = 0.0098) in the ground-squirrel population and MS56 (P = 0.0001) in the marmot population. These deviations might be due to null alleles in the respective populations. So far, only a small number of microsatellite loci have been identified for M. marmota (Klinkicht 1993), and no markers have been isolated for S. citellus. The primer sets for 12 loci compiled here should provide sufficient information for genetic investigations not only in M. marmota and S. citellus but also over a larger species range within the two genera. Acknowledgements We are very much indebted to C. Schlötterer and W. Pinsker for useful comments on the manuscript. We thank I. Hoffman and S. Huber for field work assistance and P. Taberlet and M. Preleuthner for providing marmot samples. The work was supported by a Jubiläumsfonds der Österreichischen National bank grant (project 6590) to Eva Millesi and Fords zur Förderung der wissenschaftlichen Forschung grants to WP (project P-11840-GEN) and C.S. (P-11628, S-8207, S-8213). References Berner Convention (1999) EU Habitats & Species Directive Annex II & Annex IV. Klinkicht M (1993) Untersuchungen zum Paarungssystem des Alpenmurmeltieres, Marmota m. marmota (Linné, 1758), mittels DNAFingerprinting. PhD Thesis, Ludwig-Maximilians-Universität, München, Germany. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2165 Table 2 Microsatellite loci tested in Marmota marmota and Spermophilus citellus: number of detected alleles, size range of alleles, expected and observed heterozygosities (HE, HO ) and number of individuals analysed (n) Marmota marmota Spermophilus citellus Locus n Number of alleles Size range HE HO n Number of alleles Size range HE HO MS6 MS41 MS45 MS47 MS53 MS56 19 19 19 19 19 19 5 3 3 7 5 3 142–164 186–190 109–113 163–191 141–149 111–115 0.67 0.42 0.68 0.87 0.71 0.58 0.67 0.41 0.69 0.81 0.78 0.16 10 10 10 10 10 10 — 3 2 — 4 3 — 195–201 127–129 — 147–153 113 –121 — 0.43 0.16 — 0.58 0.49 — 0.55 0.17 — 0.46 0.62 SC2 SC4 ST7 SB10 ST10 SX 10 10 10 10 10 10 2 2 5 1 4 2 128–130 134–145 135–154 154 124–130 142–146 0.50 0.53 0.60 0.00 0.63 0.50 0.50 0.00 0.87 0.00 0.83 0.50 54 54 54 54 54 54 1 1 3 4 3 3 146 102 151–156 150–162 127–134 142–146 0.00 0.00 0.55 0.73 0.51 0.60 0.00 0.00 0.44 0.65 0.62 0.61 Rassmann K, Schlötterer C, Tautz D (1991) Isolation of simplesequence loci for use in polymerase chain reaction-based DNA fingerprinting. Electrophoresis, 12, 113–118. Raymond M, Rousset F (1995) genepop (version 1.2): a population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248– 249. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning: A Laboratory Manual. 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 2000 Graphicraft PRIMER 902 101117 NOTEs Limited, Hong Kong Microsatellite loci in the Eurasian red squirrel, Sciurus vulgaris L. REBECCA TODD Division of Genetics, University of Nottingham, Queen’s Medical Centre, Nottingham, NG7 2UH, UK Keywords: microsatellites, primers, red squirrels, Sciurus vulgaris Received 18 May 2000; revision received 12 July 2000; accepted 29 July 2000 Correspondence: Rebecca Todd. E-mail: bectodd@yahoo.com Ever since microsatellites were first amplified using the polymerase chain reaction (PCR) and shown to be variable, they have been enthusiastically adopted by population geneticists. Microsatellites quickly became the molecular marker of choice during the 1990s because of the speed and ease with which they can be applied to large samples, and the possibility of their amplification from poor-quality samples collected by non-invasive methods. The level of variability found at microsatellite loci has meant that they can be used to answer phylogenetic questions on many levels (McDonald & Potts 1997). However, the main disadvantage in the use of microsatellites is the frequent need to develop a set of markers for each species under investigation; this limitation will diminish as more markers are isolated for different species. This paper reports the development of five polymorphic microsatellite loci from the genome of Sciurus vulgaris L., the Eurasian red squirrel. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 The loci were isolated using the enrichment method of Armour et al. (1994). Three partial genomic libraries were constructed using DNA extracted from Sciurus vulgaris tissue and digested with the enzyme Mbo1 (Gibco BRL). SAU linkers were ligated to a size-selected fragment (400 –1300 bp), as described in Armour et al. (1994), and used to prime a whole-genome PCR reaction. The product of this reaction was further size-selected before hybridization selection was carried out. Each library was constructed using genomic fractions selected by hybridization to a different set of tetra-, tri- and dinucleotide target repeat sequences taken from (GATA)n, (GACA)n, (CCAT)n, (ACCT)n, (TTGG)n, (GGAA)n, (TTTG)n, (TTTC)n, (GTA)n, (GAT)n, (GCT)n, (CGT)n, (TCC)n, (CAC)n, (GTT)n, (AAG)n and (GT)n. The hybridization selection reactions were carried out as described by Armour et al. (1994). The selected fraction was re-amplified in a whole-genome PCR and ligated directly into the pGEM-T vector (Promega) or the pNoTA/T7 shuttle vector of the Prime PCR Cloner Cloning System (5 Prime → 3 Prime, Inc.). These ligations were used to transform Epicurian coli XL2-Blue MRF ultracompetent cells (Stratagene). Positive colonies were cultured and stored as glycerol stocks in microtitre plates; the contents of each plate were replicated onto nylon filters and probed with labelled target oligonucleotide repeat sequences. Positive colonies were identified and sequenced manually using either isolated plasmid DNA or amplified PCR products as template (the PCR products were generated using the primers M13for (Gibco BRL) and M13rev (Promega) which flank the insertion site). Sequencing was carried out using the T7 sequencing mixes and the T7 polymerase enzyme (Pharmacia Biotech) following a protocol based on protocol 11 described in Hoelzel & Green (1998). Primers for all useful repeat sequences were designed (with the aid of the computer program oligo™; National Bioscience) and tested for variability on a panel of 10 DNA samples. PCR amplification of variable loci was optimized using the method described by Cobb & Clarkson (1994). The forward reaction primer in each case was end-labelled with 32P γ-dATP 2166 P R I M E R N O T E S Table 1 The characteristics of five Eurasian red squirrel microsatellites Locus name GenBank accession no. Repeat structure Primer sequences (5′ → 3′) RSµ1 AF285149 [GGAT]13 RSµ3 AF285150 [GA]9[GACA]9 RSµ4 AF285151 [ATCC]12 RSµ5 AF285152 [GT]10 RSµ6 AF285153 [GTT]10 F 5′-CTGGGTTCACTGACTTCTCC-3′ R 5′-CACTCTCAGAGGCCAAAGTC-3′ F 5′-GCCAAAATCTAGCCCAAGAAG-3′ R 5′-CTCAGGTGTGGGAAAGAAGC-3′ F 5′-CAATCCTCCCATCCTGCTGC-3′ R 5′-TAGGCAGTCAGATAGGTGGG-3′ F 5′-CCCAGTCTACATTAAAGGGC-3′ R 5′-GCCTATACACTATAATTGACTG-3′ F 5′-GGCATAGGGCACGTGAAG-3′ R 5′-GGGCCAATCTCATACCAAG-3′ Allele size range (bp) Number of alleles HO (%) HE (%) 172–196 7 71.9 73 161–173 7 52.2 57.4 256–284 8 78.1 72.3 123–143 7 39.3 45.5 122–131 4 27.3 36.5 HO, observed average heterozygosity; HE, expected average heterozygosity. F, forward primer; R, reverse primer. using T4 polynucleotide kinase (Gibco BRL). Amplification reactions were carried out on a PTC-200 thermocycler (MJ Research, Inc.) with 25 µL reactions containing dNTPs (0.15 mm for RSµ1, 0.1 mm for RSµ3, 0.2 mm for RSµ4 and 0.05 mm for RSµ5 and 6), 1 mm MgCl2 (1.5 mm for RSµ1), 10 pmol of each primer (5 pmol for RSµ1 and 3) including 1 pmol of labelled primer (2 pmol for RSµ1), 1 unit of ‘red hot’ Taq DNA polymerase (Advanced Biotechnologies) with Taq buffer (final concentration 0.75 m Tris–HCl, pH 9.0, 20 mm (NH4)2SO4, 0.01% w/v Tween; Advanced Biotechnologies) and approximately 0.8 ng of template DNA. The reactions were denatured at 94 °C for 3 min, and then subjected to 30 cycles of 94 °C for 1 min, 54 °C for 1 min and 72 °C for 90 s. A final extension step was carried out for 5 min at 72 °C. The PCR products were visualized by electrophoresis through 6% polyacrylamide gels using Sequi-GenII GT gel rigs (BioRad) and exposure to X-ray film. Allele sizes were determined by comparison to a known sequence ladder. Five loci, named RSµ1, RSµ3, RSµ4, RSµ5 and RSµ6, were found to be polymorphic in the Eurasian red squirrel (Table 1). These loci were amplified from 163 samples of red squirrel DNA in individuals taken from 11 populations in Belgium and Germany (Todd 2000); the proportion of individuals found to be heterozygous at each locus is also given in Table 1. The study included samples of more than 20 individuals from three large populations, and these were used to test for null alleles. Fisher’s exact test was carried out using Biomstat (version 3.2) (Applied Biostatistics Inc.) on the observed and expected number of heterozygotes at each locus in the three populations, and no evidence to indicate the presence of null alleles was found (P > 0.2). Acknowledgements This work was carried out under the supervision of Professor David Parkin with financial support from the University of Nottingham. Red squirrel tissue samples were collected by Goedele Verbeyen, University of Antwerp, Belgium. The assistance of Dr Jon Wetton is gratefully acknowledged. References Armour JAL, Neumann R, Gobert S, Jeffreys AJ (1994) Isolation of human simple repeat loci by hybridization selection. Human Molecular Genetics, 3, 599–605. Cobb BD, Clarkson JM (1994) A simple procedure for optimising the polymerase chain reaction (PCR) using modified Taguchi methods. Nucleic Acids Research, 22, 3801– 3805. Hoelzel AR, Green A (1998) PCR protocols and population analysis by direct DNA sequencing and PCR-based DNA fingerprinting. In: Molecular Genetic Analysis of Populations: A Practical Approach (ed. Hoelzel AR), pp. 201–235. Oxford University Press, Oxford. McDonald DB, Potts WK (1997) DNA microsatellites as genetic markers at several scales. In: Avian Molecular Evolution and Systematics (ed. Mindell DP), pp. 29 –48. Academic Press, San Diego. Todd RT (2000) The population genetics of red squirrels in a fragmented habitat. PhD Thesis, University of Nottingham, Nottingham, UK. 2000 Graphicraft 9PRIMER 101118 02 NOTEs Limited, Hong Kong Isolation and characterization of microsatellite loci from the ocellated wrasse Symphodus ocellatus (Perciformes: Labridae) and their applicability to related taxa S . A R I G O N I * † ‡ and C . R . L A R G I A D È R * *Division of Population Biology, Institute of Zoology, University of Berne, Baltzerstrasse 3, CH-3012 Berne, Switzerland, †Department of Zoology and Animal Biology, University of Geneva, 13, rue des Maraîchers, CH-1211 Geneva, Switzerland, ‡Station Marine d’Endoume, University of the Mediterranean, rue de la Batterie des Lions, F-13007 Marseille, France Keywords: Labridae, microsatellites, ocellated wrasse, Symphodus ocellatus Received 30 June 2000; revision accepted 27 July 2000 Correspondence: S. Arigoni. Fax: +41 31 6314888; E-mail: arigoni@zoo.unibe.ch © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2167 Table 1 Characterization of seven Symphodus ocellatus microsatellite loci based on five samples (10 individuals each) Locus Repeat array Soc1017PBBE (AC)20GC(AC)2 Soc1063PBBE (GA)2(GT)8 AT(GT)4 Soc1093PBBE (AC)26 Soc1109PBBE (GT)10 Soc1198PBBE (TG)5TA (TG)13 Soc3121PBBE (GT)18 Soc3200PBBE (GT)15 Annealing MgCl2 Number temp. (°C) (mm) of alleles Primer sequences (5′ → 3′) *TCC GTG *CCC AAG *CCT CTG *AGG TGC *CTC GAC *ACG CCA *AGT CAT TGT ATT TTC CCT CCA ACC ATT GGT TTT TTC ACA GTA GCC GGA CAG GAT TTG CAC ATT ACT TAG GAA CTG ATT AGC ATT AGA CGC TCT TAG TGT TTG CCC GGC CCT TGG CCT GGA TGC CTG TGT ATT CCC GCG CAT ATA AAA ACA GCC CTG GCA CAG ACG ACT ATA TGT TTC ATG TCC TGT ACA CTC CAG TAG CTC CAC AAC CCA TGG AGC A AG Size range (bp)** HO HE 63 0.8 20 (9 –15) 77–123 (101) 0.86 (0.6 –1.0) 0.92 (0.87 –0.96) 56 0.8 14 (6 –9) 92–134 (98) 0.76 (0.4 –1.0) 0.86 (0.83 –0.91) 96–294 (132) 0.96 (0.9 –1.0) 0.95 (0.91–0.97) 133–167 (137) 0.84 (0.7–1.0) 0.87 (0.80 –0.89) CC AC AT GA GT 63 0.8 30 (12 –15) 57 1.0 13 (7 –9) AC 57 1.2 11 (5 –9) 89–113 (109) 0.76 (0.6 –0.9) 0.80 (0.67 –0.89) CCC 56 G 51 0.9 28 (12 –14) 82–205 (102) 0.90 (0.8 –1.0) 0.95 (0.94 –0.96) 1.0 27 (12–14) 120–188 (134) 0.88 (0.8 –1.0) 0.94 (0.91–0.96) The sequences of cloned fragments have GenBank accession nos AJ278566–AJ278572; HO, observed heterozygosity; HE, expected heterozygosity; given are the mean values across the five populations with range in parentheses. *Primer used for end-labelling. **Cloned insert size in parentheses. The ocellated wrasse, Symphodus ocellatus, is a common Mediterranean labrid fish of shallow coastal waters. Its geographical distribution also includes the Black Sea, the Azov Sea and the North-Eastern Atlantic (Whitehead et al. 1984). This species is abundant in the Mediterranean and inhabits various biotopes such as shallow rocky areas and seagrass beds (Michel et al. 1987; Francour 1997). The ocellated wrasse is a partially sedentary fish, with territorial males, exclusive male parental care and conspicuous male nuptial coloration and courtship (Warner & Lejeune 1985), and thus constitutes an interesting species for investigating various aspects of population genetics and behavioural ecology of marine fishes. Here we report seven microsatellite loci of the labrid fish S. ocellatus and their amplification in five related taxa. Ocellated wrasse microsatellite loci were cloned as described by Estoup et al. (1993) and in detailed protocols by A. Estoup and J. Turgeon available at http://www.inapg.inra.fr/dsa/ microsat/microsat.htm. The genomic library was constructed with about 10 µg of DNA isolated from muscle tissue of a single ocellated wrasse from a population near Marseille (France). Approximately 1500 colonies were screened for microsatellites using a mixture of six probes (TC)10, (TG)10, (CAC)5CA, CT(CCT)5, CT(ATCT)6 and (TGTA)6TG, yielding 176 positively hybridizing clones. Plasmid DNA of positive clones was purified using a QIAprep Spin Miniprep Kit™ (Qiagen). Both strands of the wrasse DNA inserts were sequenced using a Thermo sequenase cycle sequencing kit™ (Amersham) and M13 forward and reverse primers end-labelled with fluorescent dye (IRD800™; Li-Cor). Miniprep preparation and sequencing reactions were carried out according to the recommendations of the manufacturers, and sequence reaction products were resolved on an automated DNA sequencer (model 4200™; Li-Cor). Here we report the seven microsatellite loci (Table 1) for which we so far have successfully designed primer pairs. The genomic DNA for genotyping was prepared either using © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 a phenol–ethanol extraction method or a rapid BIO RAD (Celex 100 resin) extraction protocol as described by Estoup et al. (1996). Polymerase chain reaction (PCR) amplifications were carried out in 10 µL volumes using a PTC100™ machine (MJ Research, USA). Each reaction contained 20 ng genomic DNA, 2 pmol of each primer, one of which was end-labelled with an infra-red fluorescent dye (IRD800™), MgCl2 (concentration in Table 1), 0.06 mm of each dNTP, 1 × PCR buffer (Qiagen) and 0.25 U Taq DNA polymerase (Qiagen). Reaction conditions were as follows: an initial denaturation step of 5 min at 95 °C, five cycles consisting of 30 s at 95 °C, 30 s at annealing temperature (see Table 1) and 75 s at 72 °C, 25 cycles consisting of 30 s at 94 °C, 30 s at annealing temperature and 75 s at 72 °C, followed by a final 5 min extension at 72 °C. PCR products were analysed on an automated DNA sequencer (model 4200™), and amplified fragments of cloned alleles were used for size determination at the respective loci. Variability of the loci was tested in five populations of S. ocellatus from the French Mediterranean coast (Cap Martin, St Jean Cap Ferrat, Antibes, Cannes and Marseille). Ten individuals from each population were analysed. The number of alleles per locus and the observed and expected heterozygosities are listed in Table 1. All loci were polymorphic, with the number of alleles per locus ranging between 11 and 30 and the observed heterozygosity between 0.76 and 0.96. Additionally, we tested the amplification of these primers in the labrid Coris julis and four other species of the genus Symphodus: S. tinca, S. roissali, S. rostratus and S. cinereus (Table 2). All specimens were sampled along the French Mediterranean coast between Cap Martin and Cannes. Acknowledgements We thank Alex Kohler and Susanne Wüthrich for their laboratory assistance and the Swiss Federal Office of Education and Science for financial support. S.A. thanks L. Zaninetti, 2168 P R I M E R N O T E S Species Annealing temp. (°C) Mg concentration (mm) Number of alleles Size range (bp) n Soc1017PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 63 63 61 61 61 0.8 0.8 0.8 0.8 0.8 5 7 6 2 1 5 9 5 4 2 75 –85 83 –109 93 –101 81–97 93 –97 Soc1093PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 63 63 63 63 63 0.8 0.8 0.8 0.8 0.8 3 5 6 2 1 5 1 5 4 1 114–206 112 86 –96 106–120 104 Soc1198PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 57 57 57 57 57 1.2 1.2 1.2 1.2 1.2 5 8 6 2 1 7 9 2 4 1 89 –105 95 –133 85 –103 93 –109 85 Soc3200PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 61 61 61 61 61 0.9 0.9 0.9 0.9 0.9 5 8 6 2 1 3 11 5 3 — 142–148 117 –154 122–134 132–150 — Soc1063PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 56 56 56 56 54 0.8 0.8 0.8 0.8 0.8 5 6 6 2 1 4 12 7 4 2 96 –102 98–164 94 –112 92 –120 103–113 Soc1109PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 57 57 57 57 57 1.0 1.0 1.0 1.0 1.0 5 8 6 2 1 6 12 4 3 1 141–155 133–167 127–147 143–159 125 Soc3121PBBE S. tinca S. roissali S. rostratus S. cinereus Coris julis 56 56 56 56 56 0.9 0.9 0.9 0.9 0.9 5 8 6 2 1 5 13 2 4 1 91–125 86–147 87 –95 87 –97 87 Table 2 Amplification results of seven Symphodus ocellatus microsatellite loci in related taxa n, number of analysed specimens; —, no alleles obtained. M. Harmelin-Vivien and P. Francour for scientific support, and F. Palluy and C. Marschal for assistance in obtaining samples. References Estoup A, Largiadèr CR, Perrot E, Chourrout D (1996) Rapid one tube DNA extraction for reliable PCR detection of fish polymorphic markers and transgenes. Molecular Marine Biology and Biotechnology, 5, 295–298. Estoup A, Solingnac M, Harry M, Cornuet JM (1993) Characterisation of (GT)n and (CT)n microsatellites in two insect species: Apis mellifera and Bombus terrestris. Nucleic Acids Research, 21, 1427 –1431. Francour P (1997) Fish assemblages of Posidonia oceanica beds at Port-Cros (France, NW Mediterranean): assessment of composition and long-term fluctuations by visual census. Publicazioni della Stazione Zoologica di Napoli Halia: Marine Ecology, 18, 157–173. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2169 Michel Ch, Lejeune P, Voss J (1987) Biologie et comportement des Labres européens. Revue Française d’Aquariologie Herpétologie, 14, 1– 80. Warner RR, Lejeune P (1985) Sex change limited by paternal care: a test using four Mediterranean labrid fishes, genus Symphodus. Marine Biology, 87, 89 –99. Whitehead PJP, Bauchot ML, Hureau JC, Nielsen J, Tortonese E (1984) Fishes of the North-Eastern Atlantic and Mediterranean. UNESCO Publications, Paris. PRIMER 1126 2000 Graphicraft 1932 NOTEs Limited, Hong Kong Characterization of microsatellite loci in the primitive ant Nothomyrmecia macrops Clark M AT T H I A S S A N E T R A * and ROSS H. CROZIER* School of Biochemistry and Genetics, La Trobe University, Bundoora, Victoria 3083, Australia Keywords: microsatellites, Myrmecia, Nothomyrmecia macrops, primitive ants Received 14 June 2000; revision accepted 7 August 2000 Correspondence: M. Sanetra. Fax: + 61 7 4725 1570; E-mail: matthias.sanetra@jcu.edu.au *Present address: School of Tropical Biology, James Cook University, Townsville 4811, Queensland, Australia. Although a number of microsatellite loci have been isolated for some species of ‘primitive ants’ in the subfamily Ponerinae [e.g. Diacamma (Doums 1999), Gnamptogenys (Giraut et al. 1999) ], the availability of genetic markers for the unique Australian ant Nothomyrmecia macrops has been poor. Of 16 allozyme loci studied by Ward & Taylor (1981) only one locus was polymorphic. Colonies appear to have low nestmate relatedness (Ward & Taylor 1981) but these estimates must be interpreted with caution because of limited sample size. N. macrops has great significance in evolutionary sociobiology because it possesses a relatively large proportion of ancestral characters (e.g. Taylor 1978). Thus, a more detailed knowledge of the genetics of this ant is desirable and will perhaps shed new light on a number of issues related to the evolution of eusociality in the Hymenoptera. In this paper we describe the isolation of variable microsatellite loci that can be used for precise colony- and population-level genetic analyses in Nothomyrmecia, and in the most closely related subfamily, the Myrmeciinae. Genomic DNA was extracted from five worker pupae as described by Baur et al. (1993). The DNA was digested with the restriction enzymes Sau3AI and RsaI, and size-selected fragments (300 – 600 bp) were ligated into the BamHI/HincII site of the vector pUC19. Electrocompetent Escherichia coli JM109 strains were transformed by electroporation using a BioRad genepulser and colonies were hybridized onto Hybond N + (Amersham) nylon membranes. Approximately 9000 recombinant colonies were screened with a radiolabelled (GA)10 oligonucleotide probe and 117 positive clones identified. Thirty clones were sequenced either manually using the fmol® cycle sequencing kit (Promega) or using the Big Dye Terminator cycle sequencing ready reaction kit (Perkin Elmer) with an © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 ABI Prism 377 DNA auto-sequencer. Primers were designed for 18 loci using the computer program oligoTM (Macintosh version 4.0, National Biosciences Inc.). DNA for microsatellite analysis was prepared from gasters of single Nothomyrmecia workers using a modification of the Chelex® 100 Resin extraction protocol (Walsh et al. 1991). Polymerase chain reactions contained 10 mm Tris-HCl (pH 9.0), 50 mm KCl, 1.5 mm MgCl2, 0.1% Triton® X-100, 165 µm dNTPs, 0.1 µm of forward primer, 0.03– 0.06 µm of forward primer end-labelled with [γ 33P]-ATP, 0.4 µm of reverse primer, 0.5 µg/µL bovine serum albumin, 0.4 U of Taq DNA polymerase (Promega) and 2 µL of template DNA in a total volume of 10 µL. Amplifications were conducted in a Corbett thermal cycler using the following temperature profile: 2 min at 94 °C followed by 35 cycles of 30 s at 93 °C, 30 s at 50 or 55 °C for annealing (see Table 1) and 30 s at 72 °C, and a final elongation step of 10 min at 72 °C. The amplified products were electrophoresed on 5% polyacrylamide sequencing gels and visualized by autoradiography. Of the 18 sets of primers, two failed to amplify and one gave a banding pattern that was difficult to interpret. The other 15 loci yielded repeatable and scorable results. A sample of 36 workers from Poochera, South Australia (taken from trees in an area of approximately 200 × 20 m) was used to assess the variability of these markers. We found that all but one of the 15 loci showed considerable polymorphism (Table 1), which is surprising given the small and geographically restricted sample analysed. Each polymorphic locus had between three and 12 alleles. The expected heterozygosity based on allele frequencies ranged from 0.53 – 0.90 with a mean of 0.70 across all loci. We tested for heterozygote deficiency using the computer program genepop (web version 3.1c) in order to detect the presence of null alleles. Except for the significant excess of homozygotes at locus Nmac 115, no deviations from expected heterozygosities were discovered. We investigated cross-species amplification in two species of Myrmecia and found that a large proportion of the loci could be amplified in M. forficata (see Table 1). Despite being highly polymorphic in Nothomyrmecia, the loci Nmac 11, 28 and 45 turned out to be monomorphic in M. pyriformis (three individuals from each of five colonies examined). The relatively high success rate of cross-species amplification may support the general contention that the two subfamilies Nothomyrmeciinae and Myrmeciinae are closely related (Baroni Urbani et al. 1992). In other groups of ants, successful cross-species amplifications have been reported most frequently among genera within the same subfamily (e.g. Doums 1999) suggesting a relatively low level of conservation of microsatellites across ant taxa. Acknowledgements We thank Seigo Higashi and Hiroki Miyata for field assistance, and Ching Crozier, Melissa Carew, Maria Chiotis and Vanessa Fraser for their help during the laboratory work. Lynn Atkinson and Michael Goodisman made comments on the manuscript. This work was supported by the Deutsche Akademie der Naturforscher Leopoldina (grant no. LPD 9701–6 to M.S.) and by the Australian Research Council (grant no. A19925028 to R.H.C.). 2170 P R I M E R N O T E S Table 1 Microsatellite loci and their characteristics developed in the ant Nothomyrrmecia macrops. The number of alleles (Na), frequency of the most common allele ( f ) and the estimates of observed (HO) and expected heterozygosity (HE) are based on a worker sample of 36 individuals collected near Poochera. Amplification success in Myrmecia forficata (Mf — three individuals examined) is indicated by ± but with the annealing temperature (Ta) set to 50 °C for all runs Locus Core repeat Size (bp) Na f HO HE Ta (°C) Primers (5′−3′) Nmac 1 (AG)13 186–209 8 0.32 0.94 0.82 50 Nmac 11 12 0.21 0.97 0.90 55 Nmac 13 (GA)3G(GA)2CG(GA)18(G)5(GA)2AA (GA)3 187–210 (GGA)2(GA)3TA(GA)2 (AG)15 108–126 5 0.65 0.50 0.53 55 Nmac 14 (T)3(CT)10(CCCT)2(CT)4TTCA(TC)2 151–165 5 0.47 0.49 0.62 50 Nmac 18 282–302 7 0.54 0.78 0.66 50 Nmac 20 (TC)4TTTG (TC)2(N)9(TC)11(AC)4 TCTT(TC)2 CC(CCCT)2(CT)3A(C)6TC(CT)8CA(CT)2 198–204 3 0.85 0.31 0.27 55 Nmac 23 (AG)15 286–292 4 0.32 0.78 0.74 55 Nmac 28 (C)9(TC)11TT (TC)5 156–186 10 0.60 0.64 0.63 55 Nmac 39 (GA)14 206–222 12 0.21 0.94 0.89 55 Nmac 43 (CT)23 105–139 11 0.49 0.67 0.73 50 Nmac 45 (GA)15GG(GA)10 133–173 10 0.33 0.83 0.83 50 Nmac 47 (GA)24 291– 321 11 0.22 0.82 0.88 50 Nmac 53 (GA)18 308–328 6 0.42 0.67 0.76 55 309–313 3 0.56 0.22 0.53 55 Nmac 115 (CT)3(AT)2(CA)10(CT)10TTCT (CTTT)4 F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: CGT CAG ATT AAT TGC TAG TAT TGT CCA GGC TGG AGA TCG CTC AGA ATA GGC GTC GTT CTC CGC GCC GAT GAA ACA CCC GCC CGG TTC GAT ACA CCC TCG AAC AAG AAT ATT GAG TAA CTG GCA CCA CCG AAT TCT CCG CGT CGT TTT ATA GTC ACT CAA CCT ATT GGG CGA GAC ACA TGC CCG ACC ATT TCT CGT GGT AGC GAA AAG CTG TAT CCT CCA AGA GGC GCT CAA CCC GTT TCG GGC TTC TAG TGA Mf TAT AGC TAG TGC CTT AGA GAG TAG GCG TAT AAA GGT TGC CTG AAA CGA TTC CAT AGC TTC ACC TTT GGG GCA GAG CTC TAT GTG TCG CGG ACG GGC ATC TGC AAT CTC TCC TTC TGT GTG GGT AGT ATT AGG TGA GCA AGT CAG TGC TAG TTC GGG CCA AAC CGC ATT AGC TGA GCA TTA CTT GTC GTA TCG CCA TTA AAA CTC TGA TGG CGT TGG CGG CAT CGG AAC TTC AGA GTA ACT AAC TAC CGT AAG AG G AGA G C GT TCG CAA T CG GCC G GC TA TGA CGA TG AC + T + – CT + C + G – + G + – + G TG TAA C TC C G C GTG CG + + + + GenBank Accession numbers AF264862 –264874, 264876. References Baroni Urbani C, Bolton B, Ward PS (1992) The internal phylogeny of ants (Hymenoptera: Formicidae). Systematic Entomology, 17, 301– 329. Baur A, Buschinger A, Zimmermann FK (1993) Molecular cloning and sequencing of 18S rDNA gene fragments from six different ant species. Insectes Sociaux, 40, 325–335. Doums C (1999) Characterization of microsatellite loci in the queenless Ponerine ant Diacamma cyaneiventre. Molecular Ecology, 8, 1957–1959. Giraud T, Blatrix R, Solignac M, Jaisson P (1999) Polymorphic microsatellite DNA markers in the ant Gnamptogenys striatula. Molecular Ecology, 8, 2143–2145. Taylor RW (1978) Nothomyrmecia macrops: a living-fossil ant rediscovered. Science, 201, 979–985. Walsh PS, Metzger DA, Higuchi R (1991) Chelex® 100 as a medium for simple extraction of DNA for PCR-based typing from forensic material. Biotechniques, 10, 506–513. Ward PS, Taylor RW (1981) Allozyme variation, colony structure and genetic relatedness in the primitive ant Nothomyrmecia macrops Clark (Hymenoptera: Formicidae). Journal of the Australian Entomological Society, 20, 177–183. Characterization of microsatellite loci in the aflatoxigenic fungi Aspergillus flavus and Aspergillus parasiticus N A I T R A N - D I N H and D E E C A RT E R Department of Microbiology, Building G08, University of Sydney, NSW 2006, Australia Keywords: Aspergillus flavus, Aspergillus parasiticus, microsatellite, PCR Received 30 June 2000; revision accepted 7 August 2000 Correspondence: D. Carter. Fax: + 612 93514571; E-mail: d.carter@microbio.usyd.edu.au Aspergillus flavus and Aspergillus parasiticus are closely related, morphologically similar species belonging to Aspergillus section Flavi. Both A. flavus and A. parasiticus have a worldwide impact on agriculture due to their ability to produce aflatoxin. Contamination of crops poses a serious health risk, as aflatoxins are extremely potent hepatocarcinogens (Diener et al. 1987). The need to monitor and control aflatoxin levels 000 PRIMER Graphicraft 2000 1127 912 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2171 Table 1 PCR primer sequences, number of alleles, size range, discriminatory index, and observed heterozygosity for microsatellite loci HO¶ No. of alleles D§ A.f † A.p‡ A.f A.p A.f A.p GenBank accession no. Locus Repeat motif Primer (5′−3′) Size Range AFPM1 (CCA)3(CTA)4(CCA)4 117–120 2 1 0.51 0 0.48 0 AFPM2 (ACT)5T(CTC)4 206–266 7 6 0.85 0.79 0.81 0.74 APU52151 AFPM3 (AT)6AAGGGCG(GA)8 199–217 7 4 0.79 0.71 0.75 0.67 APU76621 AFPM4 (CA)13 179–206 5 2 0.73 0.26 0.70 0.24 AB010432 AFPM5 (AG)5AC(AG)2 210–338 10 7 0.86 0.88 0.82 0.82 AF098293 AFPM6 (GT)6 341– 355 4 4 0.62 0.37 0.59 0.35 ASNAMDR AFPM7 (AC)35 CCCAGTCACGACCATTAC *GGTTCGTAGGTGGATAGAG CCACGCTCCTCAAATACG *CTGGACGGAGATCACGAC CACCACCAGTGATGAGGG *CCTTTCGCACTCCGAGAC TCTTGCTATACATATCTTCACC *AGCGATACAGTTTTAACACC CCATTATGACATGTGGTTAAGAG *TCCTACCCGAGAGAGTCTG CTCAACGCAAGTCAGGTACGC *CGAAAGGCAGTTGTGAAGGC CAAATACCAATTACGTCCAACAAGGG *TTGAGGCTGCTGTGGAACGC 215–276 11 9 0.89 0.90 0.85 0.84 AF152374 *Fluorophor-labelled primer. †Aspergillus flavus; ‡Aspergillus parasiticus; §numerical index of discriminatory power; ¶observed heterozygosity. in food and feed means contamination is also a major economic concern in many countries. Currently the methods used to control aflatoxin contamination are expensive and cannot guarantee the total elimination of toxins. A possible solution to the problem is the use of nontoxigenic isolates of Aspergillus to competitively exclude their toxigenic counterparts: a biological control strategy. Knowledge of genetic diversity, dispersal and potential for genetic exchange are essential for predicting the likely success of such a strategy. Small-scale studies of A. flavus and A. parasiticus populations carried out to date have used random amplified polymorphic DNA (RAPD) markers and DNA sequence data (Geiser et al. 1998; Tran-Dinh et al. 1999). Studies on a larger scale will require markers that are inexpensive and easy to apply, but also highly discriminatory and reproducible. We, therefore, set out to develop microsatellite markers that could be amplified from the genomes of A. flavus and A. parasiticus. In this note, we characterized seven polymorphic microsatellite markers for A. flavus and A. parasiticus. To our knowledge, no microsatellite markers have been reported for A. flavus or A. parasiticus. We also examined the ability to amplify these loci from six other species of Aspergillus. Microsatellites were found in GenBank using microsatellite motifs as queries for searches. Searches were performed on submitted sequences from A. flavus and A. parasiticus, and the two closely related species, A. oryzae and A. sojae. Six markers were found in this way (AFPM2-AFPM7). GenBank accession numbers are shown in Table 1. One microsatellite marker (AFPM1) was found by hybridizing DNA amplified from A. flavus and A. parasiticus by RAPD–PCR with chemiluminescent microsatellite probes (Carter et al. 1996). Hybridizing bands were identified, reamplified and sequenced. Primers for polymerase chain reaction (PCR) amplification were designed from sequences flanking the microsatellites © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 using oligo™ version 4.0 (National Biosciences) software. One primer from each pair was 5′-labelled with a fluorophor (PE Biosystems) (Table 1). All primer pairs were designed with an annealing temperature of 54 °C and amplified fragment sizes were optimized to allow analysis of all the microsatellite markers in a single lane. PCR amplifications were carried out in 25 µL reaction volumes using a Perkin-Elmer 2400 thermocycler. Reaction mixtures contained 1× PCR buffer (10 mm Tris-HCl pH 8.3, 50 mm KCl, 1.5 mm MgCl2, 0.001% gelatin), 200 µm each dNTP, 10 pmol of each primer, and 1 U of AmpliTaq DNA polymerase (Perkin Elmer). Each reaction contained approximately 20–40 ng of DNA, which was isolated using small-scale extraction protocol (Lee & Taylor 1990). Reactions were subjected to an initial denaturing step of 5 min at 94 °C, 30 cycles of 1 min at 94 °C, 1 min at 54 °C, 1 min at 72 °C, followed by a final elongation step of 10 min at 72 °C. Electrophoresis was conducted using an ABI 373 XL sequencer with genescan version 3.0 (PE Biosystems) software. Fragment sizes were determined with reference to a TAMRA 500 (PE Biosystems) internal standard. Microsatellite variability was analysed using 20 isolates of A. flavus and 15 isolates of A. parasiticus that have previously been found to be genetically diverse (Tran-Dinh et al. 1999). All of the microsatellite markers were reliably amplified from each isolate. The number of alleles, range of allele sizes, numerical index of discriminatory power (Hunter 1991) and observed heterozygosities for the two species are detailed in Table 1. All loci were polymorphic, with some showing higher degrees of polymorphism. Greater variation was seen within A. flavus than in A. parasiticus, which was consistent with our previous analysis using RAPD markers (Tran-Dinh et al. 1999). Amplification of the microsatellites was also attempted with DNA from A. niger, A. carbonarius, A. tamarii, A. nomius, A. oryzae and A. sojae. Only A. oryzae and A. sojae produced clear amplification products. These species are thought to 2172 P R I M E R N O T E S be domesticated variants of A. flavus and A. parasiticus, respectively. The microsatellite alleles amplified were consistent with this assumption (results not shown). In conclusion, given the levels of polymorphism and the ease of amplification and analysis, the microsatellite markers presented here will be very useful for investigating the diversity and population structure of A. flavus and A. parasiticus. Acknowledgements We thank John Pitt of Food Science Australia for providing the isolates used in this study. References Carter DA, Reynolds R, Fildes N, White TJ (1996) Future applications of PCR to conservation biology. In: Molecular Genetic Approaches in Conservation (eds Smith TB, Wayne RK), pp. 314– 326. Oxford University Press, New York. Diener UL, Cole RJ, Sanders TH et al. (1987) Epidemiology of aflatoxin formation by Aspergillus flavus. Annual Review of Phytopathology, 25, 249–270. Geiser DM, Pitt JI, Taylor JW (1998) Cryptic speciation and recombination in the aflatoxin-producing fungus Aspergillus flavus. Proceedings of the National Academy of Sciences of the USA, 95, 388– 393. Hunter PR (1991) A critical review of typing methods for Candida albicans and their applications. Critical Reviews in Microbiology, 17, 417– 434. Lee SB, Taylor JW (1990) Isolation of DNA from fungal mycelia and single spores. In: PCR Protocols (eds Innis MA, Gelfand DH, Sninsky JJ, White TJ), pp. 282 –287. Academic Press, San Diego. Tran-Dinh N, Pitt JI, Carter DA (1999) Molecular genotype analysis of natural toxigenic and nontoxigenic isolate of Aspergillus flavus and A. parasiticus. Mycological Research, 103, 1485–1490. PRIMER 2000 Graphicraft 1128 1932 NOTEs Limited, Hong Kong Identification of microsatellite loci in the water-rat Nectomys squamipes (Rodentia, Sigmodontinae) F. C . A L M E I D A , * L . S . M A R O J A , * H . N . S E U Á N E Z , * † R . C E R Q U E I R A ‡ and M. A. M. MOREIRA† *Genetics Department, Universidade Federal do Rio de Janeiro, Rio de Janeiro, RJ, Brazil, †Genetics Division, Instituto Nacional de Cancer, Praça da Cruz Vermelha 23, 6° andar, Rio de Janeiro, RJ 20230-130, Brazil, ‡Ecology Department, Universidade Federal do Rio de Janeiro. Rio de Janeiro, RJ, Brazil Keywords: Cricetidae, DNA marker, heterozygosity, microsatellite isolation, primers Received 10 July 2000; revision accepted 7 August 2000 Correspondence: Dr M. A. M. Moreira. Fax: + 55 21 224 41 48; E-mail: genetics@inca.org.br The water-rat, Nectomys squamipes (Cricetidae, Sigmodontinae), is a South American semiaquatic rodent that is well adapted to peridomiciliar habitats. This species is a primary host of Schistosoma mansoni, the prevalence of which may be as high as 90% in some natural rodent populations (Rey 1993). Because rodent fitness is not reduced by infection (D’Andrea et al. 2000), migrating rats, once infected, will carry the parasite and establish new infective foci in sites where secondary parasite hosts (Biomphalaria species) are present. These characteristics make the presence of water-rat populations a complicated factor for schistosomiasis control. Several populations of N. squamipes studied with random amplified polymorphic DNA (RAPD) showed limited differentiation indicating effects of migration or recent range expansion (Almeida et al., in press). Microsatellites were developed as a tool for studying migration patterns of N. squamipes and for evaluating its potential in spreading infection. Genomic DNA was digested with AluI, and 200 – 700 bp fragments were excised and purified (QIAquick Gel Extraction, QIAGEN) following separation in low melting agarose gel electrophoresis. Size-selected fragments were ligated to a SmaI-digested and dephosphorilated pUC18 vector (Pharmacia) and transferred to Escherichia coli DH5α competent cells. Some 18 210 recombinant colonies were transferred to nylon membranes (NEN) following Sambrook et al. (1989). Nylon filters were hybridized with [γ 32P]-ATP labelled (GT)10, (CT)10, (AGG)7, (GAA)7 and (GATA)5 oligonucleotides. A total of 11 224 colonies were hybridized with (GT)10, 12 984 with (CT)10, 8878 with (AGG)7, 5981 with (GAA)7, and 5981 with (GATA)5. One hundred and eleven positive colonies were detected and 53 were sequenced with an ABI PRISM 377 automated sequencer using BigDye terminator labelling (Applied Biosystems). Thirty-three sequenced clones showed microsatellite repeats. Of the 28 well resolved sequences, all had (CA)n microsatellites motifs except for two, indicating that this was the most abundant in N. squamipes. Eight microsatellites were characterized of which only five were polymorphic (Table 1). Genomic DNA of N. squamipes was extracted from blood or liver tissue of several water-rat populations by the standard proteinase-K/phenol– chloroform procedure (Sambrook et al. 1989). Polymerase chain reactions (PCR) were carried out in final volumes of 15 µL with ~10 – 40 ng of genomic DNA, 10 mm Tris-HCl pH 9.0, 50 mm KCl2, 2.5 mm MgCl2, 7 pmol of fluorescence labelled forward primer, 10 pmol of reverse primer, 300 µm of each dNTP and 1 U of Taq DNA polymerase (Pharmacia). PCR amplifications were performed using a thermal cycler (GeneAmp PCR System 9700 – PE Applied Biosystems) under the following conditions: an initial denaturation at 94 °C for 5 min followed by 30 cycles (except for Nec15 and Nec18 with 34 cycles) of 15 s at 94 °C, 30 s at Ta °C (Table 1), 30 s at 72 °C and a final extension period of 4 min at 72 °C. Fragment analyses were conducted with an ABI PRISM 377 with standard loading and electrophoresis conditions. Alleles were sized relatively to an internal size standard (ROX GS 500; Applied Biosystems) and analysed with genescan version 2.1 (Applied Biosystems). A maximum of 110 N. squamipes individuals was analysed for each locus (Table 2). Five microsatellites were polymorphic (Table 2) with the number of alleles ranging from 12– 26. Linkage disequilibrium between all pairs of loci was not detected (P > 0.5 Fisher’s exact test) when tested using genepop version © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2173 Table 1 Motifs and primer sequences of eight microsatellite loci of Nectomys squamipes. F, Forward primer; R, Reverse primer. Ta °C, annealing temperature Locus Repeat motif Primer sequences (5′ → 3′) Ta °C GenBank accession no. Nec12 (CA)4T(CA)19 61 AF283417 Nec14 (CA)24 57 Nec15 (AC)24T(CA)6 Nec18 (CA)34 Nec28 (CA)19 F: CTCCCTTCCCTCAATTTGCTGAGT R: ACATGTGCAAAGCATGAAAATGGA F: CAGGCGATTTACACAAAAGAAT R: CACTGAGCCATCTATCCAGTTC F: AGGAAATGCTTATCTGGAGGAG R: GACTCCTGATGTTGAACTGACC F: CTCTTTGAGGCCACTTCATTAA R: GAACTAACATTTGCATCCTCCAG F: AGGAGAAAACCTGTATGCCATG R: GTTTCTTCTTGCTGACCATGAGG AF283420 AF283419 AF283422 AF283421 AF283426 AF283424 AF283428 Table 2 Genetic variation of eight microsatellite loci in Nectomys squamipes. N, number of examined animals; A, number of alleles per loci; Freq., frequency of the most common allele; HO, observed heterozygosity; HE, expected heterozygosity Locus N A Freq. HE HO Allele range (bp) Nec12 Nec14 Nec15 Nec18 Nec28 110 110 100 109 110 26 16 19 21 12 0.15 0.16 0.17 0.11 0.24 0.93 0.90 0.90 0.93 0.85 0.72* 0.73*** 0.68*** 0.80** 0.82 206–242 204–236 171– 213 128–170 133–155 *P < 0.01, **P < 0.001, ***P < 0.0001. P-values obtained with Fisher’s exact test for difference between HE and HO considering the null hypothesis of heterozygote deficiency. 3.2 (Raymond & Rousset 1995). Expected heterozygozity was significantly higher than observed heterozygosity for all but one locus (Table 2). Although this was probably a result of the Wahlund effect (Hartl & Clark 1997), and since samples were collected in eight different localities, the existence of null alleles cannot be ruled out until a more detailed population study can be performed. The five polymorphic microsatellites loci, the first known for Nectomys, will be useful for assessing genetic variability within and among water-rat populations as well as for detecting differentiation and migration. 58 58 59 D’Andrea PS, Maroja LS, Gentile R, Cerqueira R, Maldonado A, Jr, Rey L (2000) The influence of Schistosoma mansoni on a naturally infected population of water-rats in Brazil. Parasitology, 120, 573–582. Hartl DL, Clark AG (1997) Principles of Population Genetics. 3rd edn. Sinauer Associates Inc., Sunderland, MA. Raymond M, Rousset F (1995) genepop (Version 1.2): a population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. Rey L (1993) Non-human vertebrate hosts of Schistosoma mansoni and Schistosomiasis transmission in Brazil. Research and Reviews in Parasitology, 53, 13 –25. Sambrook J, Fritcsh E, Maniatis T (1989) Molecular Cloning a Laboratory Manual. 2nd edn. Cold Spring Harbour Laboratory Press, New York. 2000 Graphicraft 1129 109PRIMER 02 NOTEs Limited, Hong Kong A set of CA repeat microsatellite markers derived from the pool frog, Rana lessonae T. W. J . G A R N E R , * B . G A U T S C H I , † S . R Ö T H L I S B E R G E R , * and H . - U . R E Y E R * *Zoologisches Institut, Universität Zürich-Irchel, Winterthurerstrasse 190, CH-8057 Zürich, Switzerland, †Institut für Umweltwissenschaften, Universität Zürich-Irchel, Winterthurerstrasse 190, CH-8057 Zürich, Switzerland Keywords: hybridogenesis, microsatellites, primers, Rana esculenta, Rana lessonae, water frogs Received 21 July 2000; revision accepted 7 August 2000 Acknowledgements This work was funded by CNPq-PRONEX 100/98, PROBIO/ MMA, FIOCRUZ (to L. Rey and P. S. D’Andrea), FUJB, INCa/ FAF and CAPES (graduate grants). We are grateful to P. S. D’Andrea and C. R. Bonvicino for providing part of the analysed samples. References Almeida FC, Moreira MAM, Bonvicino CR, Cerqueira R (in press) RAPD analysis of Nectomys squamipes (Rodentia, Sigmodontinae) populations. Genetics and Molecular Biology, in press. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Correspondence: T. W. J. Garner. Fax: + 41 635 68 21; E-mail: twjg@zool.unizh.ch The pool frog, Rana lessonae, is broadly distributed in central Europe and often forms hybridogenetic, hemiclonal hybrids with the lake frog, Rana ridibunda (Blankenhorn 1977). These hybrids, known as Rana klepton esculenta, sexually parasitize either one or the other of the parental species, the most common form of which is the L-E system (R. lessonae LL × R. esculenta LR) (Graf & Polls-Pelaz 1989). In this system, hybrids transmit a clonal R. ridibunda haplotype by mating with a syntopic R. lessonae, while hybrid by hybrid crosses result in inviable offspring (Graf & Müller 1979; Uzzell et al. 2174 P R I M E R N O T E S 1980). Hybrid lineages, therefore, represent frozen lineages that are assumed to be subject to an accumulation of deleterious mutations; mutations that are expressed when a hybrid by hybrid cross occurs and are suppressed when backcrosses with the parental species occur (Uzzell et al. 1980). The obvious lack of fitness benefits for R. lessonae individuals involved in LL × LR pairings make investigations of mate choice and sexual selection in this complex of great interest (Abt & Reyer 1993; Reyer et al. 1999). As well, pure R. esculenta populations have been detected, while theoretical investigations show that such pure populations cannot persist in isolation (Som et al. 2000). Even when a few R. lessonae are present, these generally are involved in hybrid matings due to the predominance of hybrids, which suggests that immigration by R. lessonae into such populations is required for population maintenance (Som et al. 2000; Hellriegel & Reyer in press). In these cases, management of pure or almost pure hybrid populations also requires identifying and managing R. lessonae source ponds. With these and other applications in mind, we identified and characterized a suite of CA repeat microsatellite loci derived from R. lessonae. We constructed a highly enriched subgenomic library following standard protocols (Tenzer et al. 1999). A brief outline follows: genomic DNA isolated from a single male R. lessonae was digested to completion with Tsp509I (New England Biolabs) and the 500 –1000 bp size fraction was isolated from LM-MP agarose (Boehringer Mannheim) using freezer phenol extraction. This size fraction was ligated to TSPADSHORT/TSPADLONG linkers (Tenzer et al. 1999) and amplified using TSPADSHORT and the polymerase chain reaction (PCR) as follows; total reaction volume was 25 µL and included 100 ng DNA, 1 U Taq polymerase (Quantum-Appligene), 10 mm Tris-HCl, pH 9.0, 50 mm KCl, 1.5 mm MgCl2, 0.01% TritonX100, 0.2 mg BSA (QuantumAppligene), 100 µm of each dNTP (Promega), and 1 µm of TSPADSHORT. PCR was performed on a Techne Genius thermocycler (Techne Ltd) using the following thermotreatment: 2 min at 72 °C, followed by 25 cycles of 1 min at 94 °C, 1 min at 55 °C, and 1 min at 72 °. A total of 32 PCRs were carried out, pooled, cleaned and concentrated to minimize the likelihood of redundant products being detected during screening for positive clones (B. Gautschi et al. submitted). PCR products were hybridized to biotinylated CA(20) probes bonded to streptavidin-coated magnetic beads (Dynabeads M-280 Streptavadin, DYNAL, France) and amplified again. These final PCR products were cloned following the Original TA Cloning® Kit (Invitrogen) protocol. White colonies were dot-blotted onto nylon membranes (Hybond™-N+, Amersham Pharmacia) and screened for CA repeats using the ECL 3′oligolabelling and detection system (Amersham Pharmacia) and a 40mer CA oligonucleotide. All positive clones were sequenced using M13 forward and reverse primers, following the ABI Prism® BigDye™ Terminator Cycle Sequencing Ready Reaction Kit protocol, version 2.0 (PE Biosystems) and using the ABI 377 automated sequencing system (PE Biosystems). Primer design was carried out using primer 3 software (Rozen & Skaletsky 1998) and oligonucleotides were synthesized by Microsynth GmbH (Switzerland). Initial tests for amplification and polymorphism were done at 55 °C and electrophoresed on 8%, nondenaturing, 14.5 cm by 17 cm acrylamide gels at 80 V overnight. Those primers amplifying polymorphic products using five test templates (Table 1) were used for subsequent analyses reported below. PCR amplification of frog DNA isolated from a sample of R. lessonae and R. esculenta adults captured and toe-clipped at a pond near Hellberg, north of Zürich, Switzerland was performed as follows. Reactions were 10 µL total volume and contained 50 –100 ng template DNA, 0.5 U Taq polymerase (Quantum-Appligene), buffer components and dNTPs as listed above, and 0.5 µm of both forward and reverse primer. All PCR was performed using the following conditions: 3 min at 94 °C, followed by 25 cycles of 30 s at 94 °C, 30 s at 57 or 58 °C, and 30 s at 72 °C, followed by a final step of 2 min at 72 °C. Products were electrophoresed on Spreadex™ gels, either EL-300 or EL-500 (Elchrom Scientific AG, Switzerland), depending on the size of the alleles generated. All electrophoresis was performed using the SEA 2000™ advanced submerged gel electrophoresis apparatus (Elchrom Scientific AG, Switzerland) at 100 V for 60 –120 min, depending on allele size, then scored against the M3 Marker ladder (Elchrom Scientific AG, Switzerland) and a 20-bp ladder (Bio-Rad). Expected and observed counts for homozygotes/heterozygotes were determined using genepop, version 3x (Raymond & Rousset 1995) and tested for significant deviations using Chi-square analysis (null hypothesis rejected at P < 0.05). All 10 loci were variable in R. lessonae and as well in R. esculenta (data not shown). Locus RlCA1b5 amplified an allele 137 bp in length only in R. esculenta individuals and is most likely the clonally transmitted R. ridibunda allele (data not shown). Loci RlCA1b17, RlCA1b20, RlCA1b27, RlCA18, RlCA19 and RlCA31 all appear to amplify only a single allele in a sample of R. esculenta tested in two other populations not reported here. Only R. lessonae frogs were used to test for homozygote excess, for obvious reasons. Loci RlCA5, RlCA1b17, RlCA1b20 and RlCA2a49 all exhibited homozygote excess (P ≥ 0.05), which may indicate the presence of at least one null allele at each of these loci. Considering the bizarre nature of the LE complex, these homozygote excesses may instead be indicative of a departure from Hardy–Weinberg due to a violation of the assumption of random mating. Acknowledgements P. Zipperlen, J. Berger and M. Spörri provided invaluable assistance with sequencing. This work was supported through a grant to H.-U. Reyer from the Swiss National Foundation (SNF 31–40688.94). References Abt G, Reyer H-U (1993) Mate choice and fitness in a hybrid frog: Rana esculenta females prefer Rana lessonae males over their own. Behavioral Ecology and Sociobiology, 32, 221–228. Blankenhorn H (1977) Reproduction and mating behavior in Rana lessonae-Rana esculenta mixed populations. In: The Reproductive Biology of Amphibians (eds Taylor DH, Guttman SI), pp. 389 –410. Plenum Press, New York. Graf J-D, Müller WP (1979) Experimental gynogenesis provides evidence of hybridogenetic reproduction in the Rana esculenta complex. Experentia, 35, 1574–1576. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2175 Table 1 10 CA repeat microsatellite loci developed for Rana lessonae. All data based on PCR analysis of 25 R. lessonae individuals. Ta, annealing temperature; HO, observed number of homozygotes; HE, unbiased average heterozygosity estimate (Nei 1978). Both size and repeat motif are based on that detected in the original sequenced clone (GenBank Accession nos: AF286384 –93) Locus Primer Sequences (5′– 3′) Repeat motif Ta (°C) No. alleles Size (bp) HO HE RlCA1 AAATGCAAGCGTCCCAATAC GGACGCAGTTTCTGGATTTG CTTCCACTTTGCCCATCAAG ATGTGTCGGCAGCTATGTTC CTCTGCTCCCTCAGCTATGC AAAAAGTGGTCCTTTCATTTTGAG GTCTGTCCGTGTGCAGAGAG CAAGTGATTGAGAGCCTCAGC CCCAGTGACAGTGAGTACCG CCCAACTGGAGGACCAAAAG TAAACCTTAAAAGTGGTTATAAAAACC GTAAGTGTTAGGGATGCTGAGG GGGCAGGTATTGTACTCAATATCAC CAACACAAGGACTCCACTGC TGTCCACATTAAGGAACTTTTGC TTCAGAGATCAGGGGTCTCC GTAAGTGTTAGGGATGCTGAGG TAAACCTTAAAAGTGGTTATAAAAAGG GAAGCTTAAACCACTTGACCAAC TCCCTTTTTCAGGTCTTTGG (CA)16 58 10 110 0.20 0.832 (CA)17 58 6 250 0.52 0.678 (CA)22 57 5 177 0.48 0.573 (CA)15 57 2 129 0.52 0.490 (CA)17 58 3* 145 0.56 0.476 (A)8(CAAA)2(CA)16 57 8 134 0.44 0.742 (CA)8(C)13 57 4 87 0.72 0.506 (C)8(A)2(CA)15CG(CA)4 57 5 200 0.48 0.710 (CA)15(CAAA)3(A)5 58 6 134 0.48 0.644 (C)4A(C)5GACAAA CATA(CA)6TA(CA)5 58 3 98 0.44 0.640 RlCA5 RlCA18 RlCA19 RlCA1b5 RlCA1b17 RlCA1b20 RlCA2a49 RlCA1b27 RlCA31 *Third allele detected at this locus only amplifies in R. esculenta and is not included in enumerations of HO, and calculations of HE and homozygote excess (see text for last). Graf J-D, Polls-Pelaz M (1989) Evolutionary genetics of the Rana esculenta complex. In: Evolution and Ecology of Unisexual Vertebrates (eds Dawley RM, Bogert JP), pp. 289 –302. Bulletin 466, New York State Museum, Albany, New York. Hellriegel B, Reyer H-U (in press) Factors influencing the composition of mixed populations of a hemiclonal hybrid and its sexual host. Journal of Evolutionary Biology, in press. Nei M (1978) Estimation of average heterozygosity and genetic distance from a small number of individuals. Genetics, 89, 583– 590. Raymond M, Rousset F (1995) genepop (Version 1.2): Population genetics software for exact tests and ecumenism. Journal of Heredity, 86, 248– 249. Reyer H-U, Frei G, Som C (1999) Cryptic female choice: frogs reduce clutch size when amplexed by undesired males. Proceedings of the Royal Society, London, Series B, 266, 2101– 2107. Rozen S, Skaletsky HJ (1998) Primer 3. Code available at http://www-genome.wi.mit.edu/genome_software/other/ primer3.html. Som C, Anholt BR, Reyer H-U (2000) The effect of assortative mating on the coexistence of a hybridogenetic waterfrog and its sexual host. American Naturalist, 156, 34 –46. Tenzer I, degli Ivanissevich S, Morgante M, Gessler C (1999) Identification of microsatellite markers and their application to population genetics of Venturia inaequalis. Phytopathology, 89, 748– 753. Uzzell T, Hotz H, Berger L (1980) Genome exclusion in gametogenesis by an interspecific Rana hybrid: evidence from electrophoresis of individual oocytes. Journal of Experimental Zoology, 214, 251– 259. PRIMER 1130 2000 Graphicraft 10902 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Characterization of microsatellite and minisatellite loci in Atlantic salmon (Salmo salar L.) and cross-species amplification in other salmonids M . C A I R N E Y , * J . B . TA G G A RT * and B. HØYHEIM† *Institute of Aquaculture, University of Stirling, Stirling, FK9 4LA, UK, †Norwegian School of Veterinary Science, MGA-Genetics, PO Box 8146 DEP, N-0033 Oslo, Norway Keywords: linkage, microsatellite enrichment, minisatellite, Salmo salar Received 21 July 2000; revision accepted 7 August 2000 Correspondence: Margaret Cairney. Fax: 01786 472133; E-mail: margaret.cairney@stir.ac.uk The Atlantic salmon (Salmo salar) is a salmonid fish species which naturally inhabits cool rivers and oceans of the Northern hemisphere. It is of considerable economic importance, both for recreational fishing and as a major aquaculture species. Novel polymorphic genetic markers are in continual demand to extend familial and population genetic studies in this species. We report here on the identification of ‘higher order’ (tri- and tetranucleotide) Atlantic salmon microsatellites. A number of different size-selected Atlantic salmon genomic DNA libraries were constructed, employing a microsatellite enrichment methodology (comprehensively described by Kijas et al. 1994). This protocol uses biotinylated microsatellite motif PCR Conditions No. of alleles Heterozygosity§ HO HE H–W EMBL Accession no. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Locus Repeat motif of original clone† Primer Sequence (5′– 3′)‡ Ta (°C) MgCl2 (mm) Allele size range (bp) Ssa401UOS (GACA)38 64 1.5 230– 340 20 0.86 0.92 ns AJ402718 Ssa402/1UOS¶ /2UOS Ssa403UOS (GA)55 64 0.9 1.0 10 5 18 0.95 0.57 1.00 0.84 0.70 0.94 ns ns ns AJ402719 62.5 206– 246 154 –172 152– 252 Ssa404UOS (GACA)27 59 0.9 194– 314 20 0.95 0.94 ns AJ402721 Ssa405UOS (GACA)34 62 1.0 302– 405 16 0.95 0.95 ns AJ402722 Ssa406UOS (GA)18C(GGAC)5A(GACA)4 62 1.0 322– 520 12 0.71 0.82 ns AJ402723 Ssa407UOS (GACA)37 61 1.0 176– 304 15 0.95 0.93 ns AJ402724 Ssa408UOS (GACA)27 62 1.5 248–340 16 0.90 0.93 ns AJ402725 Ssa410UOS (GACA)22 58 1.0 198– 324 25 0.90 0.97 ns AJ402727 Ssa411UOS (CT)70 inc. interspersed (GT)1 62 1.0 290– 294 2 0.20 0.18 ns AJ402728 Ssa412UOS (GA)7GG(GA)10GG(GA)20 62 1.0 246– 252 3 0.56 0.51 ns AJ402729 Ssa413/1UOS¶ /2UOS Ssa416UOS (ATT)2G(TTA)4(GTA)3(N)65(ATT)7 64 0.9 1 5 6 0.00 0.71 0.67 0.00 0.75 0.66 ns ns AJ402733 Ssa417UOS (GATA)114 Ssa418UOS (GATA)59 Ssa419UOS [86 bp MS]3 Ssa420/1UOS¶ /2UOS Ssa421UOS (CA)5T(GACA)21 Ssa422UOS (GA)3(GT)2(GA)6GGG(GA)20 * f: ACTGGTTGTTGCAGAGTTTGATGC r: AAACATACCTGATTCCCGAACCAG * f: GCTTTGGCAATGCATGTGGTAAT r: CCTATCCCTGTTGTTGCTGAC * f: CTTTAGAAGACGGCTCACCCTGTA r: GCTACTTCGTACTGACTGCCTCA * f: ATGCAGTGTAAGAGGGGTAAAAAC r: CTCTGCTCTCCTCTGACTCTC * f: CTGAGTGGGAATGGACCAGACA r: ACTCGGGAGGCCCAGACTTGAT * f: ACCAACCTGCACATGTCTTCTATG r: GCTGCCGCCTGTTGTCTCTTT * f: TGTGTAGGCAGGTGTGGAC r: CACTGCTGTTACTTTGGTGATTC * f: AATGGATTACGGGTACGTTAGACA r: CTCTTGTGCAGGTTCTTCATCTGT * f: GGAAAATAATCAATGCTGCTGGTT r: CTACAATCTGGACTATCTTCTTCA * f: TCCGCACAGACCAGAAGAACG r: AGGGGAGACCGCGAGTGAGA * f: GTGGAGATACACAGCACTTA r: CACCCCTCCGTTTTATCAC * f: GTAGACGCCATCGGTATTGTG r: CGTGATGCCGCTGTAGACTTG * f: TGACCAACAACAAACGCACAT r: CCCACCCATTAACACAACTAT * f: AGACAGGTCCAGACAAGCACTCA r: ATCAAATCCACTGGGGTTATACTG * f: CACACCTCAACCTGGACACT r: GACATCAACAACCTCAAGACTG * f: GGTCGTATCGCGTTTCAGGA r: TGCTGCAATAAAGAGATGCTTGTT * f: GCAGGAGAGTCGCTACAG r: GATCTATGCCCACAAACAG f: CAGGGTCTGTGGTGGACTGTTC * r: CGTTTGCACATTGTGAGGTGTC f: TTATGGGCGTCCACCTCTGACA * r: CACCCCAGCCTCCTCAACCTTC (GACA)28(N)39(GT)56 [90 bp MS]5 (GACA)24(GAGACA)10(GA)4 AJ402720 63 0.9 234 214– 234 214– 400 AJ402730 60 1.0 265– 424 24 0.95 0.96 ns AJ402734 64 1.0 328– 570+ 22 1.00 0.96 ns AJ402735 64 1.0 314– 406 3 0.19 0.18 ns AJ402736 63 1.0 0.48 0.00 0.95 0.96 0.00 0.94 AJ402737 1.3 22 1 18 *** 60 164– 700 142 282– 370 ns AJ402738 60 1.1 194– 220 10 0.86 0.87 ns AJ402739 †N, any nucleotide; [25 bp MS], minisatellite with 25 bp repeat motif. ‡f & r are forward and reverse primers, respectively; *indicates isotopically labelled primer. §HO, observed heterozygosity; HE, expected heterozygosity (Nei’s unbiased); H–W, test for conformance to Hardy–Weinberg equilibrium (ns, not significant, P > 0.05; ***P < 0.001). Pseudo-exact tests performed using genepop v3.1 population genetics software (Raymond & Rousset 1995). ¶Two loci detected — presumed duplicate pair reflecting the tetraploid origin of the salmonid genome (Ohno 1970). 2176 P R I M E R N O T E S Table 1 Repeat motif, PCR primer sequences, optimal annealing temperature (Ta), MgCl2 concentration for amplification and preliminary population characteristics (based on 21 individuals) for 20 polymorphic Atlantic salmon microsatellite and minisatellite loci P R I M E R N O T E S 2177 Table 2 Cross amplification of 19 Atlantic salmon derived primer sets in seven other salmonid species, as determined by agarose gel electrophoresis. + or (+) indicates detection of a discrete fragment of appropriate size at 1 °C or 5 °C below Atlantic salmon optimum annealing temperature, respectively; – indicates absence or smeared product. Additionally, P indicates polymorphism confirmed by isotopic screening (Salmo trutta and Oncorhynchus mykiss individuals only); see text for details Primer set Salmo trutta Oncorhynchus mykiss Oncorhynchus clarki Oncorhynchus nerka Salvelinus alpinus Coregonus lavaretus Thymallus thymallus Ssa401UOS Ssa402UOS Ssa403UOS Ssa404UOS Ssa405UOS Ssa406UOS Ssa407UOS Ssa408UOS Ssa410UOS Ssa411UOS Ssa412UOS Ssa413UOS Ssa416UOS Ssa417UOS Ssa418UOS Ssa419UOS Ssa420UOS Ssa421UOS Ssa422UOS (+) +P +P + + +P +P +P +P – + +P +P +P + P* +P +P – +P – +P +P – + + +P +P +P – + +P + +P + P* – – – (+) – + + – + + + + + – + + + + + + + – + – + – – – + + + + – + (+) – + + (+) (+) – – – + + + + + + + + – + + + + + + + – + – + + – – + – + + – + + – + + – + – + – + – – – + + (+) – – – (+) – + + – – – + *Two loci detected. sequences bound to streptavidin-coated magnetic particles as the basis for enrichment. The libraries were constructed using dephosphorylated pBluescript II KS(–) phagemid vector (BamHI or EcoRV digested), Epicurian Coli XL2-Blue ultracompetent host cells (Stratagene) and size selected (≈200– 500 bp) restriction-digested Atlantic salmon genomic DNA. One library was constructed using MboI digested DNA fragments which was enriched for (GACA) n sequences. Additional libraries were made using blunt-ended restriction digested DNA (pooled from separate AluI, HaeIII and RsaI digests). These libraries were potentially enriched for (GACA)n, (GATA)n, (TAA)n, and (AAGG)n motifs. Microsatellite containing recombinant clones were identified by ordered array screening of the libraries (Armour et al. 1994). Recombinant DNA was fixed onto Hybond-N membrane (Amersham) and hybridized with isotopically [γ 32P]-ATP end-labelled target oligonucleotide [(TAA)6, (GACA)4, (GATA)4 or (AAGG)4]. Hybridization was performed overnight in 6× SSC (20× SSC is 3 m NaCl, 0.3 m Na3 citrate, pH 7.0), 0.1% SDS (sodium dodecyl sulphate), 42 °C with subsequent washes to a stringency of 5× SSC, 0.1% SDS, 42 °C for 30 min. Following autoradiography, clones exhibiting strong signal were sequenced (ABI PRISM Dye Primer Cycle Sequencing Ready Reaction Kit and ABI 377 mediated automated detection) and, where possible, appropriate polymerase chain reaction (PCR) primers were designed (assisted by Primer Select software, DNASTAR Inc.). Characterization of primer sets involved both nonisotopic and isotopic screening. Atlantic salmon DNA was extracted from liver tissue by a rapid phenol-based method (Taggart et al. 1992). Common components in both assays (10 µL final © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 volume) were: 50 ng template DNA, 50 mm KCl, 10 mm TrisHCl pH 9.0, 0.1% Triton X-100, 0.9–1.5 mm MgCl2, 130 µm each nucleotide and 0.5 U Taq DNA polymerase (Promega). For nonisotopic PCRs 1 µm of each primer was included. For isotopic assays 0.1 µm of each primer was added with 10% of one primer being end-labelled with [γ32P]-ATP (4500 Ci/mMol). Cycling parameters, using a Hybaid TouchDown thermocycler, were: 96 °C for 3 min, four cycles of 95 °C for 50 s, xx °C annealing for 50 s, 72 °C for 50 s and n cycles of 94 °C for 50 s, xx °C annealing for 50 s, 72 °C for 50 s, where xx is locus specific annealing temperature (Table 1) and n = 28 for nonisotopic or 25 for isotopic reactions. Non-isotopic products were resolved on ethidium bromide stained 1.4% agarose gels while isotopic products were separated on 50 cm long denaturing polyacrylamide gels (SequaGel XR; National Diagnostics) followed by autoradiography. Allele sizes were determined relative to pBluescript II KS(–) sequence reactions run on each gel. Level of variability at each identified locus was assessed in 21 wild adult salmon. Polymorphic loci were also screened in two Atlantic salmon families (each consisting of two parents + 46 progeny). Cross-species amplification was assessed in seven other salmonid species (Table 2). Two individuals of each species were screened (nonisotopically) using two different annealing temperatures (1 °C and 4 °C below optimum for Atlantic salmon) and 1.5 mm MgCl2. Additionally four brown trout (Salmo trutta) and two rainbow trout (Oncorhynchus mykiss) were screened for polymorphism using identical isotopic conditions to those employed for Atlantic salmon. Of 164 clones sequenced, 144 had identifiable repeat motifs. While most (87%) were the expected target repeats both 2178 P R I M E R N O T E S dinucleotide microsatellites (5%) and minisatellites (8%) were also identified. Clones containing (AAGG) n showed few consecutive repeat units (2 – 3) and these were invariably found within larger minisatellite motifs. Identified (TAA)n microsatellites comprised relatively low numbers of repeats (n = 4 –15) while both (GACA)n and (GATA)n repeats were much larger (n = 10 –100+ ; Table 1). Forty-two primer sets could be designed that flanked micro- or minisatellite sequences. Twenty-two sets gave discrete products on nonisotopic testing with Atlantic salmon samples and were further optimized for isotopic screening. Of 25 loci amplified, 20 were detected as being polymorphic (Table 1). Inheritance studies confirmed disomic segregation of alleles at each locus. Length mutations were observed for two loci (Ssa404UOS, one allele; Ssa417UOS, three alleles; out of 184 progeny alleles assayed). Presence of a high frequency null allele at Ssa420UOS, suggested from population data (Table 1), was confirmed in both pedigrees screened. Furthermore, joint segregation statistics identified four significant linkage associations (P < 0.01): Ssa402/1UOS with Ssa403UOS; Ssa402/2UOS with Ssa404UOS; Ssa407UOS with Ssa422UOS; and Ssa408UOS with Ssa413UOS. Many of the primer sets are potentially informative for other salmonid species (Table 2). of microsatellites from the citrus genome using biotinylated oligonucleotide sequences bound to streptavidin-coated magnetic particles. Biotechniques, 16, 657–662. Ohno S (1970) Evolution by Gene Duplication. Allen & Unwin, London. Raymond M, Rousset F (1995) genepop (version 1.2): population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. Taggart JB, Hynes RA, Prodöhl PA, Ferguson A (1992) A simplified protocol for routine total DNA isolation from salmonid fishes. Journal of Fish Biology, 40, 963–965. 2000 Graphicraft 1131 109PRIMER 02 NOTEs Limited, Hong Kong Isolation and characterization of microsatellite loci in the orchid Ophrys araneola (Orchidaceae) and a test of cross-species amplification M. SOLIVA,* B. GAUTSC HI, † C. SA LZMA NN,* I. TEN Z ER‡ and A. WIDMER* *Geobotanisches Institut, ETH Zürich, Zollikerstrasse 107, CH-8008 Zürich, Switzerland, †Institut für Umweltwissenschaften, Universität Zürich-Irchel, Winterthurerstrasse 190, CH-8057 Zürich, Switzerland, ‡Institut für Pflanzenwissenschaften, ETH Zürich, Universitätstrasse 2, CH-8092 Zürich, Switzerland Acknowledgements We thank Paulo Prodöhl for advice on enrichment strategies and Jens Carlsson, Karim Gharbi and Kate Lindner for salmonid DNA samples. The project was supported by European Commission contract FAIR CT96 1591 (SALMAP). Keywords: cross-species amplification, microsatellites, Ophrys, Orchidaceae, pollination References The orchid genus Ophrys shows a highly specialized pollination system in which flowers deceive male hymenopterans by imitating females. Pollination occurs when males attempt to mate with the labellum of the flowers. This interaction, known as sexual deception, is assumed to be highly specific Received 21 July 2000; revision accepted 24 July 2000 Correspondence: Marco Soliva. Fax: + 41 1632 14 63; E-mail: soliva@geobot.umnw.ethz.ch Armour JAL, Neumann R, Gobert S, Jeffreys AJJ (1994) Isolation of human simple repeat loci by hybridization selection. Human Molecular Genetics, 3, 599–605. Kijas JMH, Fowler JCS, Garbett CA, Thomas MR (1994) Enrichment Table 1 Characteristics of seven microsatellite loci of Ophrys araneola. Data are based on two populations, one from Switzerland (CH), and one from France (FR). †labelled primer; ‡labelled dNTPs; Ta, locus specific annealing temperature; HO, observed heterozygosity; HE, expected heterozygosity; *significant heterozygote deficiency (P < 0.05). Repeat motifs and PCR-product lengths are derived from the sequenced clone Locus Primer sequence (5′–3′) Repeat motif Ta Size (°C) (bp) OaCT1 F: TCGTGCTACATAGGAAGGCAAATC† R: AGTCTCCAAACGGCACCCAG F: GCCAACCCCTTGGAGAAAGC† R: CAAGCTCGCTCCTTTAACTCGC F: ATAGAGGCGGTCTCCTTCAAGTCG† R: CAGTGACGAACTCATGCTCTCCAG F: CACGTCGGTGCCTCATTTAC† R: TGAGTCGATATGAATAACCTGCC F: AGCATTGGAGGCATATCCGAC R: CGTGCTTTGTGATTTTTGGCG F: GGTTTGTGGTTGTTGTTTGCG† R: AAGCTCCTCCAATGGAACCTTC F: GCACTGAGGTTGTATGCTGAGAGG† R: GCTCGGATTGTGATTCCAAGC (CT)20 50 168 7 (CT)31TT(CT)5 58 201 15 (CT)19 58 171 7 (CT)16AT(CT)5(ATCT)5 50 158 14 (CT)28AA(CT)8AA(CT)4 49 164 16 (CT)28 58 186 14 (CT)25 58 195 22 OaCT2 OaCT3 OaCT4 OaCT5‡ OaCT6 OaCT7 Total no. of alleles Population (no. of individuals) HO HE CH (24) FR (19) CH (23) FR (19) CH (24) FR (17) CH (24) FR (19) CH (24) FR (19) CH (23) FR (19) CH (24) FR (19) 0.370 0.634 0.793* 0.910 0.714 0.745 0.872 0.872 0.772 0.899* 0.769* 0.765* 0.775 0.940 0.375 0.684 0.700 0.789 0.750 0.529 0.917 0.737 0.833 0.789 0.478 0.368 0.667 0.895 Accession number AF277788 AF277789 AF277790 AF277791 AF277792 AF277793 AF277794 © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2179 Locus Species OaCT1 OaCT2 OaCT3 OaCT4 OaCT5 OaCT6 OaCT7 Ophrys aveyronensis Ophrys fuciflora Ophrys insectifera Ophrys lutea Ophrys sphegodes Ophrys tenthredinifera + + + – – – + + + – + + + – – – + – + + + – + – + + + – + – + + – – + + + + + – + + due to the imitation of sexual pheromones (Schiestl et al. 1999). To study the influence of this specialized pollination system on genetic population structure and to estimate gene flow among morphologically similar, coflowering Ophrys species with presumably different pollinators, variable and codominant genetic markers are needed. We, therefore, isolated and characterized microsatellite loci from Ophrys araneola and tested their variability in two O. araneola populations. Furthermore, we assessed whether these loci can be amplified in other Ophrys species. Genomic DNA was extracted from leaf material stored in silica gel using the CTAB protocol (Doyle & Doyle 1990). Microsatellite loci were isolated and identified from a partial genomic library enriched for GA/CT repeats, following Tenzer et al. (1999). Enriched DNA was ligated into pGEM®-T vector and cloned using JM109 high efficiency competent cells (Promega). The 672 colonies with inserts were blotted onto nylon filters (Hybond-N+, Amersham Pharmacia Biotech) and screened for GA/CT repeats using the ECL 3′-oligolabelling and detection systems (Amersham Pharmacia Biotech). Plasmid DNA of 74 positive clones was purified with the GFXTM Micro Plasmid Prep Kit (Amersham Pharmacia Biotech). Cycle-sequencing reactions were performed with BigDye terminator chemistry (PE Biosystems) using the universal primers pUC/M13 forward and pUC/M13 reverse, and run on an ABI Prism 310 Genetic Analyser. Microsatellite motifs were found in 51 clones. Primers annealing to flanking regions were designed for 21 loci using MacVector™ 6.0 (Oxford Molecular LTD). Polymerase chain reactions (PCRs) were performed in 10 µL reaction volumes containing 4.5 µL ddH2O, 10 mm Tris-HCl, 50 mm KCl, 0.4 µm of each forward and reverse primer, 200 µm of each dNTP, 0.5 U AmpliTaq Gold® DNA Polymerase (PE Biosystems), 1.5 mm MgCl2 and 10 – 50 ng of DNA. PCRs were run on a Perkin-Elmer GeneAmp PCR System 9700 thermocycler. An initial denaturation step (95 °C, 10 min) was followed by 35 cycles of 30 s at 95 °C, 30 s at the locus specific annealing temperature (see Table 1), 30 s at 72 °C; a final extension step for 5 min at 72 °C was performed at the end. Products were visualized on an ABI PRISM 310 Genetic Analyser (PE Biosystems) using either labelled primers or labelled nucleotides (Table 1). Allele sizes were scored against the internal GeneScan-500 (ROX) size standard (PE Biosystems) and individuals were genotyped using GeneScan Analysis® 3.1 and Genotyper® 2.1 software (PE Biosystems). Seven out of 21 primer pairs amplified fragments of the expected size. Levels of variability detected at these seven loci are high, with numbers of alleles ranging from 7–22 and observed heterozygosities ranging from 37– 92% (Table 1). Using genepop 3.1c (Rousset & Raymond 1995) we found a significant heterozygote deficiency (P < 0.01) for loci OaCT2 and OaCT6 in the Swiss O. araneola population, and for OaCT5 and OaCT6 in the French population. Loci OaCT1 and OaCT7 exhibit significant linkage disequilibrium across both populations (P < 0.01). We tested for cross-species amplification of O. araneola primers with six other Ophrys species of increasing phylogenetic distance to O. araneola (M. Soliva et al., unpublished results), using two individuals per species. PCR conditions were the same as described above. Qualities of PCR products were classified according to Smulders et al. (1997). Amplification products of qualities 1– 3 were regarded as successful crossspecies amplifications, whereas products of qualities 4 and 5 were treated as failure. Microsatellite loci were successfully cross-amplified with Ophrys aveyronensis, partially with O. sphegodes, O. fuciflora, O. insectifera, and O. tenthredinifera and failed to cross-amplify with the more distantly related O. lutea (Table 2). Acknowledgements The authors thank Susanne Graf and Elke Karaus for technical assistance. This study was supported by Swiss Federal Institute of Technology (ETH) internal grants (no. 0–20–477–98 and no. 0–20–600–99). References Doyle JJ, Doyle JL (1990) Isolation of plant DNA from fresh tissue. Focus, 12, 13 –15. Rousset F, Raymond M (1995) genepop (version 1.2): population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. Schiestl FP, Ayasse M, Paulus HFL, Löfstedt C, Hansson BS, Ibarra F, Francke W (1999) Orchid pollination by sexual swindle. Nature, 399, 421–422. Smulders MJM, Bredemeijer G, Rus-Kortekaas W, Arens P, Vosman B (1997) Use of short microsatellites from database sequences to generate polymorphisms among Lycopersicon esculentum cultivars and accessions of other Lycopersicon species. Theoretical and Applied Genetics, 97, 264–272. Tenzer I, degli Ivanissevich S, Morgante M, Gessler C (1999) Identification of microsatellite markers and their application to population genetics of Venturia inaequalis. Phytopathology, 89, 748 – 753. Graphicraft 2000 1133 00 912 primer PRIMER notes NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Table 2 Cross-species amplification with Ophrys araneola microsatellite primers. +, successful amplification (qualities 1–3 of Smulders et al. 1997); –, no amplification (qualities 4 and 5 of Smulders et al. (1997) 2180 P R I M E R N O T E S Microsatellite DNA loci suitable for parentage analysis in the yellow-pine chipmunk (Tamias amoenus) A L B R E C H T I . S C H U LT E - H O S T E D D E , * H . L I S L E G I B B S † and J O H N S . M I L L A R * *Ecology and Evolution Group, Department of Zoology, University of Western Ontario, London, Ontario, Canada, N6A 5B7, †Department of Biology, McMaster University, Hamilton, Ontario, Canada. L8S 4K1 Keywords: microsatellite DNA loci, parentage analysis, Tamias amoenus, yellow-pine chipmunk Received 20 April 2000; revision accepted 16 June 2000 Correspondence: Albrecht I. Schulte-Hostedde. Fax: (519) 661–2014; E-mail: aischult@julian.uwo.ca The yellow-pine chipmunk (Tamias amoenus) exhibits femalebiased sexual size dimorphism (Schulte-Hostedde & Millar 2000), and an understanding of the evolution and/or maintenance of this dimorphism requires the determination of individual reproductive success. DNA-based genetic markers are necessary for assigning parentage to quantify reproductive success in promiscuous mating systems, such as chipmunks (Callahan 1981). Here, we: (i) characterize primers for 11 microsatellite loci suitable for parentage studies of yellow-pine chipmunks; and (ii) assess variation in loci derived from Columbian ground squirrels (Spermophilus columbianus) which produce amplification products in the least chipmunk (Tamias minimus) (Stevens et al. 1997). We extracted DNA from the kidney of a yellow-pine chipmunk taken from the Kananaskis Valley, Alberta and constructed a plasmid library consisting of 250 – 400 bp fragments using the method described by Dawson et al. (1997). Briefly, approximately 10 µg of DNA was digested and fragments containing 250– 400 bp were purified from an agarose gel and cloned into a plasmid vector. The library was transformed into XL1-Blue (Stratagene) competent cells and plate lifts made using HybondN (Amersham-Pharmacia) nylon membranes. Approximately 50 000 colonies were screened using two dinucleotide polymers [ (TG)n and (TC)n (Amersham-Pharmacia) labelled with 32P-dCTP] and 165 positive clones were identified. Twenty-five clones, each containing a single insert, were sequenced by MOBIX Central Facility, McMaster University, using dye-terminator chemistry on an ABI 373 A Stretch DNA sequencer. Primers to amplify regions containing repeats were designed from 17 clones using primer (version 0.5; Lincoln et al. 1991); however, only 11 of these primer pairs were sufficiently variable for parentage studies (i.e. ≥ 3 alleles). To assess variability of these 11 loci, we used DNA from ear tissue collected from 76 chipmunks (43 adults, 33 juveniles) in the Kananaskis Valley in 1999. DNA was extracted using QIAGEN® QIAmp tissue kits. Polymerase chain reaction (PCR) was performed on the samples generally following Dawson et al. (1997) on a 480 Perkin-Elmer DNA Thermal Cycler, with the following changes: after the Table 1 Primer sequences, repeat motif, PCR product size for clone, annealing temperature (Ta), number of alleles among 43 adults surveyd, observed (HO) and expected (HE) heterozygosity, and GenBank accession nos for microsatellite loci of the yellow-pine chipmunk. F and R are the forward and reverse primer, respectively No. of Alleles HO Primer Primer sequence Repeat Clone size Ta EuAmMS 26 F 5′ ACA GGA ACA GCA GAT TGT TGT 3′ R 5′ CAC TGT TTG CCT GTG AAG AG 3′ F 5′ ATC CGT TTA GTC TGT TAT GTC TCA 3′ R 5′ TTT AAT CTA AAG GAC AAC AAT TGC 3′ F 5′ CCT GGG AGA AAA TAC TTG GAT G 3′ R 5′ AGA AAT GAG GGC AGG GAT AAT T 3′ F 5′ ATT CAG GCT CCA GAA AAA CAA A 3′ R 5′ TCT GCC CCA GAG ATA TTG ATC T 3′ F 5′ AAA GAA TGT GCA GCA AAC CTG 3′ R 5′ TTC AAT CCT TTC TAG TGC TCT TCC 5′ F 5′ TGG CTC AGT TTT TCA GTT TTT 3′ R 5′ ATC TCA AAG CCA TCA AGA GTT T 5′ F 5′ TCC CAA CAA CCT CTC TTG ATG 3′ R 3′ AAC TTG AAA ATT TTC TTC TGG GC 3′ F 5′ CTC AGT CTC CCC AAA CAT TG 3′ R 5′ TAG TTC AGT GGT AGG GCA TTC 3′ F 5′ AAT GTA TGC TAG AGT GCC CAC C 3′ R 5′ TTT TCT AGA GAC ACA AAA ATT TAG CA 3′ F 5′ CTG TGG CGG TCT TAT CTG TAT G 3′ R 5′ CCA GTT ACA GCC AGA ACC ACT T 3′ F 5′ GCC CAT CAA TAG TTG AAT GGA TA 3′ R 5′ CCT GGA AAT GCC ATA ATT TTA TTC 3′ (CA)20 181 bp 55 °C 4 0.605 0.551 AF255957 (TG)12 139 bp 55 °C 5 0.674 0.657 AF255958 (GA)17 134 bp 55 °C 3 0.488 0.506 AF255959 (GT)16 143 bp 54 °C 5 0.721 0.715 AF255960 (AC)21 159 bp 55 °C 5 0.465 0.533 AF255961 (GT)14 104 bp 51 °C 4 0.279 0.282 AF255962 (GT)10 182 bp 53 °C 4 0.651 0.634 AF255963 (CT)21 159 bp 53 °C 8 0.860 0.745* AF255964 (AC)19 128 bp 54 °C 5 0.581 0.694 AF255965 (CT)14(CA)14 120 bp 53 °C 4 0.814 0.698 AF255966 (TC)6G(TC)5G(TC)9(AC)20 169 bp 60 °C 9 0.710 0.642 AF255967 EuAmMS 35 EuAmMS 37 EuAmMS 41 EuAmMS 86 EuAmMS 94 EuAmMS 108 EuAmMS 114 EuAmMS 138 EuAmMS 142 EuAmMS 163 HE GenBank *indicates a significant deviation from Hardy–Weinberg equilibrium (P < 0.05). © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2181 initial denaturing step at 94 °C for 3 min, 32 PCR cycles were performed consisting of 45 s at 94 °C, 45 s at the appropriate annealing temperature, and 45 s at 72 °C. Amplification products were resolved on polyacrylamide gels, as described in Dawson et al. (1997) except gels were run at 70 W. PCR reactions (1 µL volume) consisted of the following reagents; 2.5 mm of MgCl2 (MBI Fermentas), PCR buffer [75 mm Tris-HCl (pH 8.8), 20 mm (NH4)2SO4, 0.01% Tween (MBI Fermentas) ] 1 µg/µL BSA (Amersham-Pharmacia), 200 µm dNTP’s, 0.25 U Taq DNA polymerase (MBI Fermentas), 0.2 pmol of the forward primer labelled with [γ-33P]-ATP (Amersham-Pharmacia), 0.3 pmol of the unlabelled forward primer, and 0.5 pmol unlabelled reverse primer. Table 1 describes the primer sequence, size of clone product, annealing temperature, and number of observed alleles for each locus. We determined whether there were deviations from Hardy–Weinberg equilibrium for each locus from adult chipmunks using genepop (Raymond & Rousset 1995). Only EuAmMS 114 was found to deviate from Hardy–Weinberg expectation due to heterozygote excess (Table 1). To assess the utility of these microsatellite loci for parentage analysis, we used the likelihood-based approach and simulation procedures of cervus 1.0 (Marshall et al. 1998). Using this program, we were able to assign maternity to all 33 juveniles (100%) with 80% confidence, 18 (54.5%) of these with 95% confidence. Using known maternity data, we were able to assign paternity to 30 juveniles (90.9%) with 80% confidence, 20 (60.6%) of these with 95% confidence. The microsatellite loci presented here provide adequate information to assess parentage in yellow-pine chipmunks. We also attempted to amplify samples of yellow-pine chipmunk DNA using four primers which amplify DNA from Columbian ground squirrels and least chipmunks (Loci: GS3, GS17, GS20, and GS34) (Stevens et al. 1997). Only two alleles were observed among samples from 22 chipmunks for GS22. At a low-stringency annealing temperature (50 °C) we found only GS20 to amplify, producing one allele. These primers were not considered to be appropriate for further parentage analysis. Acknowledgements We thank Liliana De Sousa for superb technical assistance. This study was supported by a grant-in-aid of research from the American Society of Mammalogists, and postgraduate scholarship from the Natural Sciences and Engineering Research Council of Canada (NSERC) to AISH, and NSERC operating grants to HLG and JSM. References Callahan JR (1981) Vocal solicitation and parental investment in female Eutamias. American Naturalist, 118, 872–875. Dawson RJG, Gibbs HL, Hobson KA, Yezerinac SM (1997) Isolation of microsatellite DNA markers from a passerine bird, Dendroica petechia (the yellow warbler), and their use in population studies. Heredity, 79, 506–514. Lincoln SE, Daly MJ, Lander ES (1991) Primer: A computer program for automatically selecting PCR primers, Version 0.5. Whitehead Institute for Biomedical Research, Cambridge, MA. Marshall TC, Slate J, Kruuk LEB, Pemberton JM (1998) Statistical confidence for likelihood-based paternity inference in natural populations. Molecular Ecology, 7, 639–655. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Raymond M, Rousset F (1995) genepop (Version 1.2): a population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. Schulte-Hostedde AI, Millar JS (2000) Measuring sexual size dimorphism in the yellow-pine chipmunk (Tamias amoenus). Canadian Journal of Zoology, 78, 728–733. Stevens S, Coffin J, Strobeck C (1997) Microsatellite loci in Columbian ground squirrels Spermophilus columbianus. Molecular Ecology, 6, 493–495. 2000 Graphicraft 1132 109PRIMER 02 NOTEs Limited, Hong Kong Polymorphic di-nucleotide microsatellite loci isolated from the humpback whale, Megaptera novaeangliae MARTIN E BÉRUBÉ,* † HAN NE J Ø RG ENSEN,† ROSS MC EWIN G* and P ER J. PA LSBØ LL*† *School of Biological Sciences, University of Wales, Deiniol Road, Bangor, Gwynedd LL57 2UW, UK, †Department of Evolutionary Biology, University of Copenhagen, Universitetsparken 15, DK-2100 Copenhagen Ø, Denmark Keywords: baleen whale, kinship, Mysticeti, STR loci Received 10 August 2000; revision accepted 11 August 2000 Correspondence: Martine Bérubé. School of Biological Sciences, University of Wales, Deiniol Road, Bangor, Gwynedd LL57 2UW, UK. Fax: +44 1248 38 28 25; E-mail: martine@sbs.bangor.ac.uk The study of cetaceans by genetic methods is moving increasingly towards the estimation of kinship among individuals using genetic data. Microsatellite loci are ideal for this kind of study given their high mutation rates and co-dominant inheritance. However, a large number of loci need to be genotyped in order to ensure reliable estimation of kinship. Even for relatively small sample sizes, reliable identification of parent– offspring pairs is likely to require more than 17 loci genotyped in each individual (Palsbøll 1999). Towards this end, we isolated an additional nine polymorphic microsatellite loci from genomic DNA of the humpback whale, Megaptera novaeangliae, which are presented here. The loci originate from the same partial genomic library from which we previously presented tri- and tetra-nucleotide microsatellite loci (Palsbøll et al. 1997). In this paper, we present additional di-nucleotide loci identified among the positive clones in the above-mentioned genomic library. The isolation and sequencing of clones containing inserts has been described previously (Palsbøll et al. 1997). The data presented here are based upon genotypes obtained from up to 353 individual humpback whales, 65 individual fin whales (Balaenoptera physalus), 169 individual minke whales (B. acutorostrata) and 92 individual blue whales (B. musculus). Total-cell DNA was extracted from skin biopsies by standard phenol and chloroform extractions (Sambrook et al. 1989) and the DNA re-suspended in 1 × TE (Sambrook et al. 1989). The nucleotide sequence at each locus was amplified by polymerase chain reaction (PCR) (Mullis & Faloona 1987) using 10 µL reaction volumes, each containing 10 ng of genomic DNA, 67 mm Tris –HCl, pH 8.8, 2 mm MgCl2, 16.6 mm (NH4)2SO4, 10 mm β-mercaptoethanol, 0.2 mm dNTPs, 1 mm unlabelled oligo-nucleotide primer, 40 µm end-labelled oligo-nucleotide 2182 P R I M E R N O T E S Table 1 Summary of the experimental conditions for amplification of the microsatellite loci Thermocycling profile Locus GT023 GT101 GT195 GT211 GT271 GT307 GT310 GT509 GT575 Oligo designation* Oligonuceotide primer sequence (5′ → 3′) GT023R GT023F GT101R GT101F GT195R GT195F GT211R2 GT211F2 GT271F GT271R GT307F GT307R GT310R GT310F GT509F GT509R GT575F GT575R CAT GTT CTT CTG TGA TGA CAT GGC GCT CCC ATA TTA TAA GAA CAG GTA TAT ACC TTC CCC TCT TGC GAA AGT CTG ACA CAC TAG TAG GCG CTT TAC CTG AAA AAG ATC CTA AGG CCT TGG AGA AAC TGC AGT ACT GAA TTA AGT GTG TCC CAA TGT TGA AAC CCC CTC AGT TAT TGA AGT TTC CAG GGT GGA TAT CAT GAA CAG AAC TTC ATA TGG ACC TGC GCT ATG CTA TAA CAC TAA AAT TAG CTG ATT GAT TAG CTT CAG CAA AAG TGT ACT CCC CTA TGA TAT AAG GGT CTG ACA TTG ATA GCC TTT GAC TGC AGA TCT CAT CTG CGC TCC CTC ACC CCC AGG TGG TAG CTC AAG AAC CTC ATT ATC CCC TTC GenBank accession no. Annealing temperature (°C) Cycling times† Number of cycles Thermocycler AF309690 62 15/15/15 28 MJR PCT100‡ AF309691 60 30/30/30 30 RoboCycler§ AF309692 54 15/15/15 30 MJR PCT100 AF309693 60 30/30/30 28 RoboCycler AF309694 62 15/15/15 28 MJR PCT100 AF309695 49 15/15/15 33 MJR PCT100 AF309696 62 15/15/15 28 MJR PCT100 AF309697 58 15/15/15 28 MJR PCT100 AF309698 60 20/45/60 30 MJR PCT100 *The upper oligonucleotide primer was end-labelled. †Times are given in seconds, starting with time at denaturing temperature (94 °C), then time at annealing temperature, followed by time at extension temperature (72 °C). ‡MJ Research model PCT100. §Stratagene RoboCycler model 96. primer, and 0.4 units of Taq DNA polymerase (Life Technologies Inc.). The end-labelled oligo-nucleotide primer was labelled with [γ -32P]ATP using T4 polynucleotide kinase (Sambrook et al. 1989). The thermo-cycling profiles and GenBank accession numbers are listed in Table 1. The amplification products were separated by electrophoresis through a denaturing 5% polyacrylamide gel. After electrophoresis, the gel was fixed in 5% ethanol: 5% acetic acid for 40 min, followed by a 15 min rinse in tap water. The fixed polyacrylamide gel was dried at 80 °C for 45 min and autoradiography performed with Kodak BioMax™ film for 5 – 48 h depending on the intensity of radioactive signal. The size of the amplification products was estimated from λM13 sequences and multiple positive control samples (of known genotype) included in each amplification and detection. Our Gulf of Maine sample contained 73 known mother and calf pairs in which we detected no indications of null alleles (i.e. a mother and calf both homozygous but for different alleles; Pemberton et al. 1995) at the loci analysed in these samples. Neither did we detect any significant deviation between the observed and expected levels of heterozygosity for any of the remaining locus and species combinations (Table 2). Acknowledgements Samples were kindly supplied by the Center for Coastal Studies, Greenland Institute of Natural Resources, Mingan Island Cetacean Study Inc., and the US National Marine Fisheries Service South-West Fisheries Science Center. This work was in part supported by the International Whaling Commission the Greenland Home Rule, and the Commission for Scientific Research in Greenland, as well as the Danish Natural Science Research Council. References Bérubé M, Aguilar A, Dendanto D, et al. (1998) Population genetic structure of North Atlantic, Mediterranean Sea and Sea of Cortez fin whales, Balaenoptera physalus (Linnaeus, 1758): analysis of mitochondrial and nuclear loci. Molecular Ecology, 7, 585–600. Mullis KB, Faloona F (1987) Specific synthesis of DNA in vitro via a polymerase-catalyzed chain reaction. Methods in Enzymology, 155, 335–350. Palsbøll PJ (1999) Genetic tagging: contemporary molecular ecology. Biological Journal of the Linnean Society, 68, 3–22. Palsbøll PJ, Bérubé M, Larsen AH, Jørgensen H (1997) Primers for the amplification of tri- and tetramer microsatellite loci in cetaceans. Molecular Ecology, 6, 893–895. Pemberton JM, Slate J, Bancroft DR, Barrett JA (1995) Nonamplifying alleles at microsatellite loci: a caution for parentage and population studies. Molecular Ecology, 4, 249–252. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning. A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 2000 1137 109PRIMER Graphicraft 2 00 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2183 Number of alleles Size range (bp) HO† HE‡ 0.82 0.86 0.85 0.66 0.80 0.81 0.83 0.64 Locus Species n* GT023 M. novaeangliae B. acutorostrata B. musculus B. physalus 353 91 92 65 8 9 8 7 114 –128 100–116 122–136 112 –138 GT101 M. novaeangliae B. acutorostrata B. musculus B. physalus 4 — 92 4 2 92 – 94 — — 9 5 85–101 94 –112 0.65 — 0.67 — GT195 M. novaeangliae B. acutorostrata B. musculus B. physalus 353 3 4 65 5 2 2 8 151–163 162–166 146–148 158–176 0.65 — — 0.74 0.65 — — 0.70 GT211 M. novaeangliae B. acutorostrata B. musculus B. physalus 353 21 — 73 7 7 196–208 185–203 0.80 0.85 0.82 0.75 6 193–213 0.60 0.55 GT271 M. novaeangliae B. acutorostrata B. musculus B. physalus 353 2 4 65 10 3 3 6 97 –123 101–104 101–105 112 –128 0.57 — — 0.45 0.59 — — 0.43 GT307 M. novaeangliae B. acutorostrata B. musculus B. physalus 353 2 4 65 7 3 3 7 127–139 135–141 127–133 121–139 0.67 — — 0.70 0.68 — — 0.64 GT310 M. novaeangliae B. acutorostrata B. musculus B. physalus 4 21 4 65 2 6 3 2 102–106 112 –122 110–116 104–130 — 0.60 — 0.54 — 0.70 — 0.50 GT509 M. novaeangliae B. acutorostrata B. musculus B. physalus 6 169 — — 1 11 195 195–217 — 0.81 — 0.81 GT575 M. novaeangliae B. acutorostrata B. musculus B. physalus 5 21 — 5 6 5 140–154 195–211 — 0.80 — 0.85 5 140–154 — — Table 2 Levels of genetic variation estimated in selected baleen whale species *Number of individual whales genotyped. Estimated †observed and ‡expected heterozygosity. Megaptera novaeangliae was sampled in the Gulf of Maine, Balaenoptera acutorostrata across the North Atlantic, B. musculus in the Gulf of S. Lawrence as well as off West Greenland, and B. physalus in the Sea of Cortez (a small population with low levels of genetic variation, Bérubé et al. 1998). Novel chloroplast microsatellites reveal cytoplasmic variation in Arabidopsis thaliana J . P R O VA N School of Biology and Biochemistry, The Queen’s University of Belfast, Medical Biology Centre, 97 Lisburn Road, Belfast BT9 7BL, Northern Ireland Keywords: Arabidopsis thaliana, Brassicaceae, cytoplasm, chloroplast, microsatellites, simple sequence repeats Received 29 July 2000; revision accepted 15 August 2000 © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Correspondence: Dr Jim Provan. Fax: + 44 028 90 236505; E-mail: J.Provan@qub.ac.uk The analysis of levels and patterns of cytoplasmic variation in plants is now widely recognized as providing important and complementary information to that obtained using nuclear markers. In particular, the uniparentally inherited, nonrecombining chloroplast genome has been utilized in many studies in plant population and evolutionary genetics (Soltis et al. 1992; Ennos et al. 1999). Until recently, however, the typically low substitution rates associated with the chloroplast genome meant that detecting sufficient levels of polymorphism 2184 P R I M E R N O T E S Table 1 Arabidopsis chloroplast microsatellite primers Locus Repeat Location Primers (5 – 3′) Alleles ATCP112 (A)15 trnH(GUG)/psbA intergenic 5 96–100 ATCP7905 (A)13 trnS(GCU)/trn G(UCC) intergenic 2 140–141 ATCP28673 (T)13 y c f 6 /psbM intergenic 4 140–145 ATCP30287 (A)13 trnD(GUC)/trnY(GUA) intergenic 3 100–102 ATCP46615 (A)14 trnT(UGU)/trnL(UAA) intergenic 3 111–113 ATCP66701 (T)16 trnP(UGG)/psaJ intergenic 5 145–150 ATCP70189 (A)13 rpS12/clpP intergenic ATCCGCCCCTACGCTACTAT AGGTGGAATTTGCTACCTTTTT CGAACCCTCGGTACGATTAA TGGAGAAGGTTCTTTTTCAAGC GCGTTCCTTTCATTTAAGACG TGCACTCTTCATTCTCGTTCC CCCTATACCCTGAAATTTGACC CAGCTCGGCCCAATAATTAG AATTTTTTTCCATTGCACATTG TCAGAAATAGTCGAACGGTCG TCCACATCCTCCTTCTTTTTT CATTTGAAAACGTAAAGGCC CGGGTTGATGGATCATTACC GCAATGCACAAAAAAAGCCT 6 124–132 Brassica species Locus B. oleracea B. rapa B. napus B. nigra B. carinata B. juncea ATCP112 ATCP7905 ATCP28673 ATCP30287 ATCP46615 ATCP66701 ATCP70189 ✘ ✔✔ ✔✔ ✔ ✔✔ ✔ ✔✔ ✘ ✔✔ ✔✔ ✔ ✔✔ ✔ ✔✔ ✘ ✔✔ ✔✔ ✔ ✔✔ ✔ ✔✔ ✘ ✔✔ ✔✔ † ✔✔ ✔ ✔✔ ✘ ✔✔ ✔✔ † ✔✔ ✔ ✔✔ ✘ ✔✔ ✔✔ ✔ ✔✔ ✔ ✔✔ Range (bp) Table 2 Cross-species amplification in Brassica species using Arabidopsis chloroplast microsatellite primers ✔✔ — Strong amplification; ✔ — Weak amplification; ✘ — Poor or no amplification. †Primer ATCP30287 amplified two bands in both B. nigra and B. carinata. was the main drawback to the analysis of cytoplasmic variation, particularly below the species level. The discovery of polymorphic mononucleotide repeats in the chloroplast genomes of plants analogous to nuclear microsatellites, or simple sequence repeats, has provided a new approach to detecting cytoplasmic variation that had previously gone undetected using traditional restriction fragment length polymorphism (RFLP) studies. These chloroplast microsatellites have been used for the highresolution analysis of cytoplasmic diversity in both crop species and natural plant populations (Provan et al. 1999b, 2000). This report describes the development of chloroplast microsatellite markers in the weedy crucifer Arabidopsis thaliana (Brassicaceae). Despite Arabidopsis being the model organism for studies into the physiology, genetics and development of higher plants, very little work has been carried out on the analysis of natural populations of the species. Indeed, to date there have been no published studies investigating levels of cytoplasmic variation in Arabidopsis and only a limited number assessing levels of nuclear diversity in natural populations (Vander Zwan et al. 2000 and references therein). The complete chloroplast sequence of A. thaliana (EMBL accession number AP000423) was searched for mononucleotide repeats of n = 8 or greater using the findpatterns program (Genetics Computer Group). A total of 231 repeats were found and primers were designed to amplify seven mononucleotide repeat loci in noncoding regions using the primer program (Genetics Computer Group; Table 1). Primers were tested on 22 A. thaliana accessions from 11 populations in Europe and the USA, as well as on six Brassica species (see Table 2). Polymerase chain reaction (PCR) was carried out in a total volume of 10 µL containing 50 ng genomic DNA, 10 pmol 32P endlabelled forward primer, 10 pmol reverse primer, 1× PCR reaction buffer [20 mm Tris-HCl (pH 8.4), 50 mm KCl], 15 mm MgCl2, 0.05 U Taq polymerase (Gibco BRL). Reactions were carried out on a Techne GENIUS thermal cycler using the following parameters: initial denaturation at 94 °C for 3 min; 30 cycles of denaturation at 94 °C for 30 s, annealing at 55 °C for 30 s, extension at 72 °C for 30 s; final extension at 72 °C for 5 min. After addition of 10 µL loading buffer (95% formamide), products were resolved on 6% denaturing polyacrylamide gels containing 1× TBE buffer and 8 m urea at 80 W constant power for 2 h. Gels were transferred onto 3 mm blotting paper (Whatman) and exposed to X-ray film overnight at –70 °C. All seven loci were polymorphic in the sample studied, with between two and six alleles detected per locus (Table 1). © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2185 Combining the alleles from the seven linked loci gave 11 haplotypes in the 22 individuals. Intrapopulation variation was detected in several populations, highlighting the resolving power of the chloroplast microsatellite technique even in highly inbreeding species such as Arabidopsis, which is believed to have a selfing rate of ≈99%. Although no previous cytoplasmic studies have been carried out in Arabidopsis, it is unlikely that such levels of variation would be detected with RFLPs. This has been observed in chloroplast microsatellite studies of other inbreeding species, e.g. barley (Provan et al. 1999a), where diversity levels were far in excess of those revealed by chloroplast RFLPs. Due to high levels of conservation of both sequence and gene organization in plant chloroplast genomes, primers designed to amplify chloroplast microsatellites in one species have been shown to amplify polymorphic products in related species, even at the intergeneric level (Provan et al. 2000). Consequently, the primers developed for Arabidopsis were tested on several Brassica species (Table 2). Although primer pair ATCP112 did not amplify in the Brassica species and primers ATCP30287 and ATCP66701 gave poor and/or nonspecific amplification, the other four primer pairs amplified a single, strong product in all six species tested. This suggests that these primers may have considerable value in studying cytoplasmic variation within the Brassicaceae. In summary, these chloroplast microsatellite primers offer new opportunities to study levels and patterns of cytoplasmic variation within and between Arabidopsis natural populations and ecotypes. A comparison of chloroplast and nuclear microsatellite markers will provide new insights into the relative roles of seed and pollen movement in shaping the genetic structure of natural populations. Furthermore, their utility across the Brassicaceae means that this ability to discriminate seed and pollen movement will be of value in assessing modes of potential transgene escape in genetically modified oilseed rape (Brassica napus) and in possible hybrids between B. napus and its wild relatives. Acknowledgements The author would like to thank Joy Bergelson for providing the Arabidopsis DNA and Steve Millam for providing seeds of the Brassica species. References Ennos RA, Sinclair WT, Hu X-S, Langdon A (1999) Using organelle markers to elucidate the history, ecology and evolution of plant populations. In: Molecular Systematics and Plant Evolution (eds Hollingsworth PM, Bateman RM, Gornall RJ), pp. 1–19, Taylor & Francis, London. Provan J, Powell W, Hollingsworth PM (2000) Chloroplast microsatellites: new tools for studies in plant ecology and evolution. Trends in Ecology and Evolution, in press. Provan J, Russell JR, Booth A, Powell W (1999a) Polymorphic simple sequence repeat primers for systematic and population studies in the genus Hordeum. Molecular Ecology, 8, 505 – 511. Provan J, Soranzo N, Wilson NJ et al. (1999b) The use of uniparent-ally inherited simple sequence repeat markers in plant population studies and systematics. In: Molecular Systematics © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 and Plant Evolution (eds Hollingsworth PM, Bateman RM, Gornall RJ), pp. 35 –50, Taylor & Francis, London. Soltis DE, Soltis PS, Milligan BG (1992) Intraspecific chloroplast DNA variation: systematic and phylogenetic implications. In: Molecular Systematics of Plants (eds Soltis DE, Soltis PS, Doyle JJ), pp. 117–150, Chapman & Hall, New York. Vander Zwan C, Brodie S, Campanella JJ (2000) The intraspecific phylogenetics of Arabidopsis thaliana in worldwide populations. Systematic Botany, 25, 47–59. 2000 Graphicraft 1139 109PRIMER 02 NOTEs Limited, Hong Kong Characterization of 14 tetranucleotide microsatellite loci derived from sockeye salmon JEFFREY B. OLSEN ,* SHERI L. WILSO N,* ERIC J. K RETSC HMER,* K ENNETH C. J O NES† and JAMES E. SEEB* *Alaska Department of Fish and Game, Gene Conservation Laboratory, 333 Raspberry Road, Anchorage, Alaska 99518 –1599, USA, †Genetic Identification Services, 9552 Topanga Canyon Blvd., Chatsworth, California 91311, USA Keywords: conservation genetics, microsatellites, salmon, sockeye salmon Received 21 July 2000; revision accepted 15 August 2000 Correspondence: Jeffrey B. Olsen. Fax: +907 – 267–2442; E-mail: jeff_olsen@fishgame.state.ak.us The use of microsatellites for research and conservation of Pacific salmon (Oncorhynchus spp.) is increasing. Examples of applications include gene mapping (e.g. Lindner et al. 2000), pedigree analysis (e.g. Estoup et al. 1998), and population assignment (e.g. Olsen et al. 2000). Through these studies researchers have become more discriminating in their choice of microsatellites to meet specific project objectives and to improve genotyping efficiency. Examples of selection criteria include locus polymorphism, presence of null alleles, allele size range, polymerase chain reaction (PCR) annealing temperature, and PCR amplification quality. New microsatellites are needed to improve the selection of loci for each species of Pacific salmon. In particular, there is need for tetranucleotide microsatellites. This class of microsatellites does not generally exhibit the complicated shadow banding (‘stutter ’) observed in many dinucleotide microsatellites, and they provide sufficient range in polymorphism for various applications ( Jarne & Lagoda 1996). Here we report the development of primers for 14 novel tetranucleotide microsatellites in sockeye salmon (O. nerka). Sequences for 27 novel DNA fragments (~350 – 550 bp) containing TAGA tetranucleotide microsatellites were identified by Genetic Identification Services (GIS, Chatsworth CA) using an enrichment protocol similar to Edwards et al. (1996). Genomic DNA from a single sockeye salmon was partially restricted with a cocktail of seven blunt-end cutting enzymes (RsaI, HaeIII, BsrB 1, PvuII, StuI, ScaI, EcoR V). Fragments in the size range of 300 – 750 bp were adapted and subjected to magnetic bead capture (CPG Inc., Lincoln Park, NJ), using (TAGA)8 biotinylated capture molecules (Integrated DNA Technology, Coralville, IA). Captured molecules were amplified and 2186 P R I M E R N O T E S Table 1 Estimates of polymorphism for 14 novel sockeye salmon microsatellites. n, A, HO, HE refer to sample size, allele number, observed and expected heterozygosity, respectively. An asterisk denotes a significant difference (P < 0.05) between HO and HE. The forward primer is labelled for each locus and the annealing temperature is 56 °C for all primer pairs Locus One100 One101 One102 One103 One104 One105 One106 Repeat sequence of cloned allele Primer sequence (5′– 3′) (F, forward, R, reverse) (TAGA)18N14 (TAGA)18 (ATCT)26N12 (ATCT)13 (ATCT)10 F: CAATGCACTGTGATAGGAGG R: AGGGGAAGAAGAAGTTTTGG F: AAATGACTGAAATGTTGAGAGC R: TGGATGGATTGATGAATGG F: CATGGAGAAAAGACCAATCA R: TCACTGCCCTACAACAGAAG F: AATGTTGAGAGCTATTTCAATCC R: GATTGATGAATGGGTGGG F: ATCTTTATGGTGGCAAGTCC R: ATCTGGTACTTCCCTGATGC F: TCTTTAAGAATATGAGCCCTGG R: GCTCAAATAAACTTAAACCTGTCC F: TACCCTGCAAGACAGTGAGA R: GCTGTTTAGGAAGGAGGGTT F: TGCAGAGCCATACTAAACCA R: AAGAATTGAGAGATGCAGGG F: AGGGAGAGAAGAGAGGGAGA R: CCTCAGAAGTAGCATCAGCTC F: CCTCCATTTCAATCTCATCC R: ACAGAGAACAGTGAGGGAGC F: ATGACCAAGGAGCTTCTGC R: TATCCAGGTACTCCACTGGC F: GTGACCCAGACTCAGAGGAC R: CACAACCCATCACATGAAAC F: TCATTAATCTAGGCTTGTCAGC R: TGCAGGTAAGACAAGGTATCC F: CGCTATACATTTTCCATTTTCC R: TTTTTAAGTGGGAGAACTTGC (ATCT)27N16 (ATCT)11 (ATCT)15N4 (ATCT)10 (TAGA)9 One108 (ATCT)9N4 (ATCT)10N8(GTCT)10 (ATCT)21 One109 (TAGA)9 One110 (TAGA)21 One111 (TAGA)21 One112 (ATCT)28 One114 (TAGA)12N4 (TAGA)12 (ATCT)24 One115 restricted with HindIII to remove the adapters. The resulting fragments were ligated into the HindIII site of pUC19. Recombinant molecules were electroporated into Escherichia coli DH5alpha. Recombinant clones were selected at random for sequencing. Sequencing was preformed in an MJ Research PTC-200 thermocycler using ABI Prism Taq DyeDeoxy™ terminator chemistry. The sequences were visualized on an ABI 373 DNA sequencer. Primer pairs for 27 sequences were designed using the program Primer 3 (Rozen & Skaletsky 1996). Unlabelled primers, purchased from Operon Inc. (Alameda, CA), were tested for amplification effectiveness in four sockeye salmon using agarose gel electrophoresis. PCR was carried out in 10 µL volumes 10 mm Tris-HCl (pH 8.3), 50 mm KCl, 3.0 mm MgCl2, 0.2 mm each dNTP, 0.5 units AmpliTaq DNA polymerase (Perkin-Elmer Corp, Foster City, CA), 0.15 µm each primer, and 100 ng DNA template] using an MJ Research PTC-225 thermocycler. DNA amplifications involved the following profile: 92 °C (5 min); 25 cycles of 92 °C (30 s) + 56 °C (30 s) + 72 °C (30 s); 72 °C (30 min). The PCR product was electrophoresed for 2 h at 100 V in a 2% agarose gel, stained with ethidium bromide, and photographed under ultraviolet n Size range (bp) A HE HO GenBank no. 89 246–378 22 0.92 0.80* AF274516 89 182–344 28 0.94 0.92 AF274517 89 207–275 15 0.86 0.85 AF274518 89 167–447 29 0.93 0.94 AF274519 89 167–239 19 0.91 0.92 AF274520 89 127–151 6 0.44 0.42 AF274521 89 111–259 30 0.90 0.82 AF274522 89 184–244 16 0.90 0.91 AF274523 89 127–175 13 0.88 0.90 AF274525 89 235–287 13 0.89 0.87 AF274526 89 194–322 30 0.88 0.86 AF274527 88 127–241 27 0.90 0.87 AF274528 89 211– 295 22 0.93 0.89 AF274530 86 173–237 16 0.92 0.89 AF274531 light (312 nm). Four primer pairs failed to yield distinct PCR products. Fluorescein-labelled forward primers were purchased for the remaining 23 sequences from Perkin-Elmer Corp. (Foster City, CA). Fourteen of these 23 primer pairs yielded high-quality amplification product as determined using an ABI 377 – 96 DNA sequencer in GeneScan mode (ABI 1996a) to detect the labelled primers in a 4.5% denaturing polyacrylamide gel (Table 1). Estimates of polymorphism were obtained for the 14 loci by genotyping 89 sockeye salmon using the ABI 377 – 96. Allele scoring was performed with Genotyper software, version 2.0 (ABI 1996b). The number of alleles per locus ranged from 6 to 30 and averaged 20 (Table 1). The expected heterozygosity ranged from 0.44 to 0.94 and averaged 0.87. The observed and expected heterozygosity differed significantly (P < 0.05) at One100. The allelic size averaged 103 bp and ranged from 24 bp (One105) to 280 bp (One103). At least some of the microsatellites amplified in six related species of Oncorhynchus and only one locus (One100) failed to amplify in all species (Table 2). In most instances (27) the amplified fragment length was within the size range identified © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2187 Table 2 Results of cross-species testing of sockeye salmon microsatellite primers. Two individuals were tested for each species Locus (One) Species 100 101 102 103 104 105 106 108 109 110 111 112 114 115 Oncorhynchus gorbuscha O. keta O. tshawytscha O. kisutch O. mykiss O. clarki – – – – – – + +↓ +↓ + +↓ +↓ + + + – + + + +↓ +↓ +↓ +↓ +↓ – – – +↓ +↓ – – + – – – – – + + + – – – + – – + +↓ – – +↓ +↓ +↓ +↓ + + +↓ +↓ +↓ +↓ + + + +↓ + + + + – +↑ + + – + + – +↓ – +↓ – – – – – +, amplified at designed annealing temperature; –, did not amplify; ↓, amplified fragment smaller than smallest allele in sockeye; ↑, amplified fragment larger than largest allele in sockeye. in sockeye salmon. In 23 instances the amplified fragment was smaller than the smallest allele in sockeye salmon and in one instance the amplified fragment was larger than the largest allele in sockeye salmon (One112). These microsatellites should prove useful for a number of conservation genetic applications in sockeye salmon and, to a lesser degree, the other species examined here. Microsatellite characterization in central stoneroller Campostoma anomalum (Pisces: Cyprinidae) P E R O D I M S O S K I , G R E G O RY P. T O T H and M A R K J . B A G L E Y Acknowledgements National Exposure Research Laboratory, United States Environmental Protection Agency, 26 West Martin Luther King Drive, Cincinnati, OH 45268, USA Funding was provided by the State of Alaska and National Marine Fisheries Service through the Western Alaska research project. This is Alaska Department of Fish and Game professional paper 198. Keywords: Campostoma anomalum, central stoneroller, Cyprinidae, microsatellites Received 10 July 2000; revision accepted 15 August 2000 References ABI (Applied Biosystems Inc.) (1996a) GeneScan 672 Users Manual Rev. A. Perkin-Elmer Corp., Foster City, CA. ABI (Applied Biosystems Inc.) (1996b) Genotyper 2.0 Users Manual. Perkin-Elmer Corp., Foster City, CA. Edwards KJ, Barker JHA, Daly A, Jones C, Karp A (1996) Microsatellite libraries enriched for several microsatellite sequences in plants. Biotechniques, 20, 758–760. Estoup A, Gharbi K, SanCristobal M, Chavelet C, Haffray P, Guyomard R (1998) Parentage assignment using microsatellites in turbot (Scopthalamus maximus) and rainbow trout (Oncorhynchus mykiss) hatchery populations. Canadian Journal of Fisheries and Aquatic Sciences, 55, 715–723. Jarne P, Lagoda JL (1996) Microsatellites, from molecules to populations and back. Trends in Ecology and Evolution, 11 (10), 424– 429. Lindner KR, Seeb JE, Habicht C et al. (2000) Gene-centromere mapping of 312 loci in pink salmon by half-tetrad analysis. Genome, 43, 538– 549. Olsen JB, Bentzen P, Banks MA, Shaklee JB, Young S (2000) Microsatellites reveal population identity of individual pink salmon to allow supportive breeding of a population at risk of extinction. Transactions of the American Fisheries Society, 129, 232– 242. Rozen S, Skaletsky H (1996) Primer 3 design program. Code available at http://www-genome.wi.mit.edu/genome_software/ other/primer3.html. PRIMER 1140 2000 Graphicraft 10902 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Correspondence: Pero Dimsoski. Fax: (650) 638-6333; E-mail: dimsoski@usa.net The central stoneroller (Campostoma anomalum) is a small cyprinid fish that is native to streams and rivers of central and eastern North America. It can be found in a range of anthropogenically modified habitats, ranging from nearly pristine to highly polluted waters (Zimmerman et al. 1980), and has intermediate sensitivity to habitat degradation relative to other fishes in the region (Zimmerman et al. 1980; Gillespie & Guttman 1989). The species is the focus of intensive study by the United States Environmental Protection Agency due to its biological and distributional characteristics. An important aspect of this research is to understand the fine-scale genetic structure of the species across its native range, and to determine how this ‘genetic landscape’ relates to underlying environmental processes. To date, genetic analyses have focused on multi-locus fingerprints generated by the random amplified polymorphic DNA (RAPD) method to delineate levels of similarity among and within populations (Silbiger et al. 1998). Because allelic counts are highly sensitive to recent changes in population size, highly polymorphic microsatellite DNA markers should provide genetic information that is highly complementary to the RAPD data and may reveal finer levels of population structuring. Here, we report a suite of highly polymorphic microsatellite markers developed for the central stoneroller. 2188 P R I M E R N O T E S Table 1 Locus name, primer sequences, annealing temperature (TA), repeat motif of cloned allele, product size based on sequenced allele (bp), number of individual fish tested (n), number of alleles (NA), observed heterozygosity (HO), expected heterozygosity (HE) and GenBank accession number for the cloned sequences for 17 microsatellite primers developed for Campostoma anomalum Locus Primer sequence (5′ → 3′) TA (°C) Repeat motif Size (bp) n NA HO HE Accession number Ca1 AAGACGATGCTGGATGTTTAC CTATAGCTTATCCCGGCAGTA ACCTTTCCTTTCGTGTCGAGA GGACCCAGCGAGCACCT GGACAGTGAGGGACGCAGAC TCTAGCCCCCAAATTTTACGG CGGTATCGGTGCATCCCTAAA AACAGCGCGAGCGTCATTC TTGAGTGGATGGTGCTTGTA GCATTGCCAAAAGTTACCTAA CAGGTCTTGCCCACGTCTGAG CACCTGTGGAACCGGCTTGA ACACGGGCTCAGAGCTAGTC CAAATGTCAGGAGTTCTCCGA ACGCAGACATATTTTAGATG AATAATACAACTCGCTCTCA ATCAAGCCTGCCATGCAC ATCACTGTAGACTGCGACCAG CTGCACGGGTTTTAATATCTT AATGATGTCATCGCCATGTA TCCCTCACTGTGCCCTACA GGCGTAGCAATCATTATACCT GTGAAGCATGGCATAGCACA CAGGAAAGTGCCAGCATACAC GATCATTGATCCGCATGTCTC CTCCCTGACAGCAGCGACC GCGGAATAGCAGTCAATA GTTAAACTGTTCCTGTTACGGT TGATTTTATATCTTCGAGGAA AAACCCAACCGTTAGTCTAAT CGCGACCAGTTGTGAC GACGAGCGTATTCAGATTACA GTTTGAAGTGGGATTAACT GTTGTGTATACCTGGTTAAAG 58 (CA)24 112 10 6 0.60 0.78 AF277573 64 (CA)19 100 11 10 0.80 0.90 AF277574 55 (TAGA)14 243 13 10 0.70 0.80 AF277575 55 (CA)12 157 13 5 0.92 0.84 AF277576 51 (TAGA)15 149 13 8 0.67 0.83 AF277577 59 (CA)14CG(GA)6 204 13 5 0.42 0.72 AF277578 59 (CA)15 103 13 7 0.54 0.78 AF277579 53 (TAGA)20 183 13 12 0.61 0.89 AF277580 57 (CA)15 118 13 7 0.50 0.67 AF277581 57 (TAGA)16 243 13 7 0.42 0.78 AF277582 57 (TAGA)7 203 9 6 0.66 0.81 AF277583 57 (TAGA)10(CAGA)4(TAGA)2 238 13 6 0.38 0.70 AF277584 57 (CA)16 163 11 4 0.36 0.61 AF277585 54 (CA)13 90 13 9 0.90 0.87 AF277586 57 (CA)23 213 10 8 0.81 0.85 AF277587 54 (TAGA)7 213 13 3 0.25 0.55 AF277588 51 (TAGA)8 131 9 3 0.11 0.30 AF277589 Ca2 Ca3 Ca4 Ca5 Ca6 Ca7 Ca8 Ca9 Ca10 Ca11 Ca12 Ca13 Ca14 Ca15 Ca16 Ca17 A partial genomic library was constructed using the strategy described by Kandpal et al. (1994). After digestion of C. anomalum genomic DNA with Sau3AI restriction enzyme, DNA fragments ranging from 400 to 1500 bp were ligated to Sau3AI linkers. After removal of excess linkers by electrophoretic fractionation, fragments were amplified by polymerase chain reaction (PCR) using a primer complimentary to the Sau3AI linker. The whole genomic library was enriched for sequences containing CA repeats by hybridization to a biotinylated CA probe. Hybridized molecules were captured using VECTREX avidin D matrix (Vector Laboratories, Burlingame, California, USA). The microsatellite-enriched library was amplified by PCR and ligated into the pCR2.1 vector (Invitrogen, San Diego, California, USA). The transformed colonies were screened with a (CA)21 oligonucleotide probe conjugated to alkaline phosphatase (Lifecodes Corp. Stamford, California, USA). Additional microsatellite loci containing TAGA and CA motifs were identified from microsatellite-enriched libraries produced by Genetic Identification Services Inc. (Chatsworth, California, USA) by using a microsatellite enrichment procedure similar to the one described above. A total of 130 clones were sequenced using BigDye terminator sequencing chemistry (PE Biosystems, Foster City, California, USA) on an ABI 310 Genetic Analyser (PE Biosystems). Diand tetra-nucleotide repeat motifs were identified in 67 of the sequences. PCR primer pairs flanking repetitive regions were designed for 27 microsatellite loci using Oligo 6.21 (Molecular Biology Insights Inc., Cascade, Colorado, USA). Total genomic DNA was extracted (DNeasy, Qiagen Inc., Valencia, California, USA) from muscle tissue of 13 samples collected from throughout the C. anomalum species range. Between 9 and 13 C. anomalum samples and between 1 and 3 samples from five other species of cyprinids were genotyped for each microsatellite marker to confirm amplification and estimate the level of polymorphism. Each 15 µL PCR raction included 25 ng of template, 250 µm dNTP, 3 pmol of each primer, 2.5 mm MgCl2, 0.3 units Taq DNA polymerase (Perkin Elmer), 10 mm Tris–HCl, pH 8.3, and 50 mm KCl. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2189 Table 2 Cross-specific amplification of microsatellite loci isolated from Campostoma anomalum Locus Bluntnose minnows Pimephales notatus (n = 3) Fathead minnows Pimephales promelas (n = 3) Blacknose dace Rhinichthys atratulus (n = 3) Creek chubs Semotilus atromaculatus (n = 3) Zebra fish Danio rerio (n = 1) Ca1 Ca3 Ca6 Ca7 Ca9 Ca11 Ca12 Ca13 Ca14 Ca16 Ca17 4 1 4 — 4 — 3 — — 4 3 4 — 3 — — — 3 — — 5 — 4 3 — 3 — 2 6 — 2 — — 3 — — 2 — 4 5 — — — — 1 1 — — — — 1 1 — — — Fragments were resolved on 5% polyacrylamide gels. The data indicate the number of alleles counted from n genotyped individuals. ‘—’, no or unreadable amplification. Cycling was performed with a PE Biosystems Geneamp 9600 thermal cycler under the following conditions: 30 s at 94 °C; 27 cycles of 1 min at 92 °C, 1 min at 51–64 °C, depending on the specific primer set (Table 1), and 1.5 min at 72 °C; followed by 7 min at 72 °C. PCR products were separated on denaturing 5% polyacrylamide gels and visualized with Vistra Green (Amersham Life Science, Amersham, Buckingamshire, UK) fluorescent dye using a FluorImager 595 fluorescent scanner (Molecular Dynamics, Sunnyvale, California, USA). A total of 17 microsatellite markers were identified as highly polymorphic for C. anomalum (Table 1). The other 10 primer sets either failed to produce a reliable PCR product or were not polymorphic for the samples assayed. Two of the 17 primer sets that were informative for C. anomalum produced a PCR product of similar size for all of the other five cyprinid species tested (Table 2). In addition, nine primer sets produced a PCR product of similar size in at least one of the other cyprinid species. The extensive polymorphisms identified for these markers within C. anomalum, and their apparent applicability to other species, indicate that they will have utility for future population studies. Acknowledgements This research was supported in part by the appointment of PD to the Postgraduate Research Program at the National Exposure Research Laboratory administered by the Oak Ridge Institute for Science and Education through an inter-agency agreement between the US Department of Energy and the US Environmental Protection Agency. References Gillespie RB, Guttman SI (1989) Effects of contaminants on the frequencies of allozymes in populations of the central stoneroller. Environmental Toxicology, 8, 309–317. Kandpal RP, Kandpal G, Weissman SM (1994) Construction of libraries enriched for sequence repeats and jumping clones, and © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 hybridization selection for region-specific markers. Proceedings of the National Academy of Sciences USA, 91, 88 – 92. Silbiger RN, Christ SA, Leonard AC, et al. (1998) Preliminary studies on the population genetics of the central stoneroller (Campostoma anomalum) from the Great Miami River basin, Ohio. Environmental Monitoring and Assessment, 51, 481–495. Zimmerman EG, Merrit RL, Woten MC (1980) Genetic variation and ecology of stoneroller minnows. Biochemical Systematics and Ecology, 8, 447–453. 2000 1143 109PRIMER Graphicraft 2 00 NOTEs Limited, Hong Kong Dinucleotide microsatellite loci for Andrena vaga and other andrenid bees from non-enriched and CT-enriched libraries C. MOHRA, M. FELLENDORF, G. S E G E L B A C H E R and R . J . PA X T O N Zoological Institute, University of Tübingen, Auf der Morgenstelle 28, D-72076 Tübingen, Germany Keywords: Andrenidae, Apoidea, Hymenoptera, SSR Received 14 June 2000; revision accepted 15 August 2000 Correspondence: Dr R.J. Paxton. Fax: +49 7071 295634; E-mail: robert.paxton@uni-tuebingen.de Andrena is a large genus (>1000 species) of bees with a primarily Holarctic distribution (Michener 1979). Analysis of Andrena population genetics has been hampered by their limited allozyme variability (Ayasse et al. 1990). Microsatellites potentially make up this shortfall, although there are few loci described for this group of bees (Paxton et al. 1996). We developed microsatellites for the andrenid bee Andrena vaga Panzer 1799 using both non-enriched and enriched partial genomic libraries. DNA for cloning was isolated from the thorax of one male bee using phenol/chloroform, digested to completion with Sau3AI, resolved on an agarose gel, and fragments between 200 and 800 bp were isolated from the gel 2190 P R I M E R N O T E S Table 1 Description of 19 microsatellite loci for Andrena vaga and heterozygosities for eight females Locus Sequence (5′ → 3′) Repeat sequence Fragment size (and range) (bp) Annealing temp. (°C) HE HO Number of alleles GenBank accession no. vaga01 F: GTGCCAAGTCAGTTAGTGTGC R: GAAACACGTAGCGAACACG F: CTTCTCCAAGCCGAATCTTCC R: GATCGGCCTGGGAAATTCC F: GATTCGGGAACGACACTCG R: CGTTTATAGCGATGATGTCCG F: TTCTACGTTAGTCCGCAGG R: CTTAGTCCGTTAAGGAGCAAC F: GGAAGGTTGAGTGGAAATTG R: TGTCCGAAGTGAAGAGAACG F: GCTTTGGTTCCTCGTGTCG R: CCACTGAAACTCATCTAGGTACACG F: GATCCGAAAAGTTGAAGGTG R: CTACGTGACTTTCCTGTCCTC F: CCGTTGTAATCGAATGAACC R: GATGGAGGAAAGGGGAGA F: GGAATTCGTCGACGAAAGG R: CGATGGGTGTAGGTGGGAT F: CTTAGTCCGTTAAGGAGCAAC R: GGAACGAAAGTCTTCTCTTCTC F: CTGCCACCTCTGTACATGG R: CGTGTGAGCTAGAGTTCCATC F: CGACTTTGCTACAGCGATTC R: CGACTTGGATAGGCAGGG F: GGGTAACGAGAGAAGGGG R: GAGGAGTCGTGTTACGTGC F: GATCTTCTTACCTCCCCCC R: CTTCCTTTTGCTCCCTCTTG F: CCTTGTTACGCGTGCATAG R: TCGGAAACTGTACGTCGTC F: CTGTGTGGAAAGGTGATAACG R: GAAGGGAACAGTAATGGACAAG F: GTCGCTACACACTCGTTATCTTG R: CATGGATTCCAACGAATTCTC F: CGAGGGCAATCGACAGTG R: GCCGTTGAATTCACGTAGG F: GACGGACTCGGATACACCC R: CGAGTTGCCGCTAACTTTC (CT)20 186 (184–194) 65 0.76 0.63 4 G64722 (CT)27 232 (210–240) 65 0.91 0.88 8 G64906 (CT)17 107 (105–123) 65 0.82 0.63 7 G64907 (CT)17 228 (226–238) 60 0.88 0.75 7 G64908 (CT)16 318 (316–326) 56 0.88 0.75 6 G64909 (CT)15 194 (192–218) 60 0.24 0 2 G64910 (CT)29 171 (165–187) 56 0.88 0.75 7 G64911 (CT)21 117 (105–131) 56 0.91 0.63 8 G64912 (CT)21 178 (164–194) 63 0.86 0.88 7 G64913 (CT)18 153 (145–163) 56 0.89 1.00 8 G64914 (CT)16 218 (216–224) 65 0.65 0.50 4 G64915 (CT)20 135 (123–155) 60 0.68 0.25 6 G64916 (CT)23 160 (146–162) 56 0.82 0.38 6 G64923 (CT)17 183 (179–191) 60 0.79 0.63 7 G64917 (CT)19 149 (149–161) 60 0.80 0.38 5 G64918 (CT)33 219 (203–233) 56 0.87 0.38 7 G64919 (CT)18 126 (126–128) 56 0.23 0 2 G64920 (CT)20 166 (156–170) 60 0.77 0.50 5 G64921 (CT)14 100 (98–106) 65 0.69 0.63 4 G64922 vaga02 vaga03 vaga04 vaga05 vaga06 vaga08 vaga09 vaga12 vaga13 vaga14 vaga18 vaga19 vaga20 vaga21 vaga23 vaga25 vaga26 vaga27 using the QIAquick Gel Extraction Kit (Qiagen) following the manufacturer’s protocol. For the non-enriched library, the 200 – 800 bp fragments were ligated into plasmid vector pUC18/BamHI (Amersham/ Pharmacia). Highly competent E. coli (INVαF′ One Shot, Invitrogen) were transformed with plasmids, and resultant colonies were simultaneously screened for microsatellites using digoxigenin (DIG)-end labelled (GA)10 and (CA)10 exactly as described by Estoup & Turgeon (1996). We used filter hybridization (Armour et al. 1994) to generate a CT-enriched library, following methods described by Segelbacher et al. (2000) (see also Piertney et al. 1998). A 1 µg aliquot of the A. vaga 200 – 800 bp fragments was ligated to a SauL linker molecule, denatured and hybridized to a 1 cm2 piece of Hybond N+ membrane (Amersham/Pharmacia) to which synthetic (GA)n polymers had previously been bound (Schlötterer 1998, pp. 241– 244). After overnight hybridization at 65 °C in 2 × SSC and 0.1% SDS, nylon membranes were given three washes of 2 × SSC and 0.1% SDS, and then the enriched fragments were stripped from the membrane by heating to 95 °C for 5 min in water. The enriched fraction was precipitated and complementary strands were reformed in a polymerase chain reaction (PCR) (30 cycles consisting of 1 min at 94 °C, 1 min at 55 °C and 1 min at 72 °C) using the SauL-A oligonucleotide as a primer (Schlötterer 1998). Linkers were removed from the fragments by digestion with Sau3AI, and the fragments, now enriched for CT/GA sequences, were subsequently ligated into a plasmid, cloned and screened exactly as described above for the non-enriched library. From the non-enriched library, four of 732 screened colonies were positive (0.5%), and, from the enriched library, 154 of 435 screened colonies were positive (35.4%), demonstrating the utility of the enrichment protocol. Plasmid DNA was extracted from positive colonies, inserts were cycle-sequenced using Big Dye Terminator chemistry (Perkin Elmer), and fragments were resolved on an ABI Prism 377 automated sequencer. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2191 Table 2 Cross-species amplification of 19 pairs of Andrena vaga microsatellite primers with two individuals each of six other bees. Where a single PCR product was obtained, the number of alleles resolved is provided Locus (vaga) Species 01 02 03 04 05 06 08 09 12 13 14 18 19 20 21 23 25 26 27 Andrenidae Andrena agilissima Andrena scotica Andrena ferox — 2 1 — — — — 1 1 1 2 1 2 1 1 1 1 1 — 2 3 2 1 1 — 3 3 1 2 1 1 — — — — — 1 2 1 1 2 1 — 1 1 1 2 2 — 1 1 1 2 1 3 1 1 Halictidae Lasioglossum malachurum — — — 1 1 — 1 2 — — — — 1 — — — — — — Apidae Scaptotrigona postica — — 1 — 1 — 1 1 — — — — 1 — — — — — — Anthophoridae Nomada lathburiana — — — — — — — — — — — — 1 — — — — — — —, a multiple band, a smear, or no product was detected. The four colony plasmids from the non-enriched library each contained a unique (CT) n repeat, whilst 45 of 49 colony plasmids from the enriched library each contained a unique (CT)n repeat. PCR primers were designed on sequences flanking 22 perfect dinucleotide repeats using the software package Amplify, version 1.2 (www.wisc.edu/genetics/CATG/Amplify). DNA for PCR was extracted from thoracic tissue using a high-salt protocol (Paxton et al. 1996). PCR amplifications were performed in 10 µL reaction volumes using an MJ Research PTC-100 thermal cycler. Individual mixes consisted of 10 ng template DNA, 4 pmol of each primer, 75 µm of each dCTP, dGTP and dTTP, 6 µm dATP, 0.125 µCi [α33P]dATP, 1.5 mm MgCl2, 10 mm Tris–HCl, pH 8.8, 50 mm KCl, 0.1% Triton X-100, 200 µm spermidine and 0.4 units of thermostable DNA polymerase (Finnzymes). Samples were processed through one denaturing step of 3 min at 94 °C followed by 25 cycles consisting of 45 s at 94 °C, 30 s at the annealing temperature specified in Table 1, and 45 s at 72 °C, with a final elongation step of 10 min at 72 °C. Nineteen of 22 primer pairs gave an amplification product using A. vaga as template DNA, many with numerous alleles per locus (Table 1). Primers were also successful in amplifying DNA extracts of other andrenid bees, although less successful in amplifying DNA from phylogenetically distant taxa, namely anthophorid, apid and halictid bees (Table 2). These loci should prove useful in the analysis of the population genetic structure of many andrenid bees. Acknowledgements This research was funded by the German Research Council (Pa 632/2). We thank Manuela Giovanetti, Elizabeth Engels and Remko Leys for collection of bees. References Armour JAL, Neumann R, Gobert S, Jeffreys AJ (1994) Isolation of human simple repeat loci by hybridization selection. Human Molecular Genetics, 3, 599–605. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Ayasse M, Leys R, Pamilo P, Tengö J (1990) Kinship in communal nesting Andrena (Hymenoptera: andrenidae) bees is indicated by composition of Dufour’s gland secretions. Biochemical Systematics and Ecology, 18, 453–460. Estoup A, Turgeon J (1996) Microsatellite marker isolation with non-radioactive probes and amplification, version 12/1996, available at http://www.inapg.inra.fr/dsa/microsat/microsat.htm Michener CD (1979) Biogeography of the bees. Annals of the Missouri Botanical Garden, 66, 277–347. Paxton RJ, Thorén PA, Tengö J, Estoup A, Pamilo P (1996) Mating structure and nestmate relatedness in a communal bee, Andrena jacobi (Hymenoptera: andrenidae), using microsatellites. Molecular Ecology, 5, 511– 519. Piertney SB, MacColl ADC, Bacon PJ, Dallas JF (1998) Local genetic structure in red grouse (Lagopus lagopus scoticus): evidence from microsatellite DNA markers. Molecular Ecology, 7, 1645–1654. Schlötterer C (1998) Microsatellites. In: Molecular Genetic Analysis of Populations (ed. Hoelzel AR), pp. 237 – 261. Oxford University Press, Oxford. Segelbacher G, Paxton RJ, Steinbrück G, Trontelj P, Storch I (2000) Characterization of microsatellites in capercaillie Tetrao urogallus (AVES). Molecular Ecology, 9, 1934–1935. 2000 1142 109PRIMER Graphicraft 2 00 NOTEs Limited, Hong Kong Isolation and characterization of microsatellite loci in the dice snake (Natrix tessellata) B. GAUTSCHI,*§ A. WIDMER†§ and J . K O E L L A ‡ § *Institut für Umweltwissenschaften, Universität Zürich-Irchel, Winterthurerstraße 190, CH-8057 Zürich, Switzerland, †Geobotanisches Institut, ETH Zürich, Zollikerstraße 107, CH-8008 Zürich, Switzerland, ‡Laboratoire d’Ecologie, CC237, Université Pierre & Marie Curie, CNRS UMR 7625, 7 Quai St Bernard, 75252 Paris, France Keywords: conservation, dice snake, microsatellites, Natrix tessellata, PCR, primers Received 5 July 2000; revision accepted 15 August 2000 2192 P R I M E R N O T E S Table 1 Natrix tessellata microsatellite primer sequences, annealing temperatures (TA), population-specific allelic diversity (A), total number of alleles detected (Atotal) and observed (HO) and expected (HE) heterozygosities in samples from populations from Lake Lugano (Switzerland) (n = 10) and Lake Garda (Italy) (n = 19). Repeat motifs are derived from the sequenced clones (GenBank accession numbers AF269184 –AF269191) Locus Primer sequence (5′ → 3′) TA (°C) Size range (bp) Repeat motif Population A µNt1 60 129–137 (CA)15 60 172–226 (CA)21 63 142–156 (AC)16 58 116–126 (CA)2GA(CA)3GA(CA)4GA(CA)15 62 187–209 (CT)6CA(CT)14(GT)13TT(GT)4 58 171–185 (AC)17 58 152–166 (AC)15 64† 291–301 (GA)27 Lugano Garda Lugano Garda Lugano Garda Lugano Garda Lugano Garda Lugano Garda Lugano Garda Lugano Garda GGAGTAGCCATTATTGCCAAAG GCTCCGACCACACTTTAAGC* µNt2 TGGCACCATTTCAGTTTCTG GGGACCTCATCGAAACATTG* µNt3 GGCAGGCTATTGGAGAAATG GGCAAAACTCCAGGTGCTAC* µNt5 TGCTTTTCGGATTTGACATTC CTGCATTTGAAGCGTGGTAG* µNt6 TGCTGGCATGTGAAATCAAG GGGGCTGTTTTCTGTCAATC* µNt7 TTTGAAAGGAGAATGAATCGTG CGCGAGGAATCAGAATGAAC* µNt8 GGGGTATCGTCCTTCCAGAC* GCCAAGTGTTTCTTCAAGTGG µNt10 AATTACAGTAGGTAGGTAGTTAGGGAGG CTGTGCCAGCAGAAACACC* Atotal HO 2 4 3 7 10 8 5 6 4 5 6 5 6 11 10 6 6 5 5 5 3 3 6 5 0.000 0.158 0.600 0.369 0.000 0.053 0.700 0.632 0.600 0.474 0.900 0.632 0.700 0.632 0.400 0.368 HE 0.340 0.301 0.735 0.712 0.680 0.652 0.595 0.474 0.780 0.819 0.800 0.632 0.545 0.571 0.515 0.501 *Fluorescent-labelled primer. †For locus µNt10, a hot-start protocol was used. The PCR conditions are as described before but with HotStarTaq™ DNA polymerase and buffer (Tris – Cl (NH4)2SO4, 1.5 mm MgCl2; pH 8.7 (20 °C), Qiagen) and an initial denaturing step of 95 °C for 15 min. Correspondence: B. Gautschi. Fax: +41 1635 57 11; E-mail: babagaut@uwinst.unizh.ch §Former address: Experimentelle Ökologie, ETH Zürich, ETH Zentrum-NW, CH-8092 Zürich, Switzerland The dice snake, Natrix tessellata (Laurenti) 1768, has a large geographical distribution, ranging from Italy in the west to China in the east (Hecht 1930). While population sizes are often large in suitable habitats, they are small in many rangemarginal populations, such as along the rivers Mosel, Lahn and Nahe in Germany (Gruschwitz 1985), where populations are in danger of becoming extinct as a consequence of either stochastic catastrophic events or genetic erosion. On the other hand, allochthonous populations in Switzerland that result from introductions of a small number of founding individuals may be very large (Mebert 1993). To assess the levels of genetic variation within large natural populations, declining range-marginal populations and allochthonous populations, suitable molecular markers are necessary. Allozymes or mitochondrial DNA are not suitable for this purpose because levels of genetic variation in small populations are typically very low, or because relatively large amounts of fresh blood or tissue are necessary. Microsatellites, on the other hand, are often variable even in small and endangered populations and can be easily amplified from minute amounts of DNA recovered from blood samples, shed skin or faeces. We constructed a partial genomic library enriched for CA and GA repeats using a slight modification of the procedures described by Tenzer et al. (1999) and Gautschi et al. (2000). Briefly, total genomic DNA was isolated from blood samples using a standard phenol– chloroform extraction protocol (Sambrook et al. 1989). DNA was digested with Tsp509I (New England Biolabs), 200 – 700 bp fragments were isolated and ligated to TSPADSHORT/TSPADLONG linker sequences (Tenzer et al. 1999). DNA linker molecules were amplified according to Gautschi et al. (2000) using TSPADSHORT as the polymerase chain reaction (PCR) primer, and PCR products were hybridized to biotinylated (CA)13 and (GA)13 probes attached to streptavidincoated magnetic beads (Dynabeads M-280 streptavidin, Dynal, France) (see Tenzer et al. 1999 for details). Enriched fragments were again amplified and products were cloned using the Original TA Cloning® Kit (Invitrogen BV) following the manufacturer’s instructions. After dot-blotting of recombinant colonies onto Nylon membranes (Hybond N+, Amersham Pharmacia), oligonucleotide probes labelled using the ECL3′oligolabelling and detection system (Amersham Pharmacia) were used to screen for inserts containing CA and GA repeats. The hybridization was carried out in accordance with the manufacturer’s instructions. Plasmids from positive clones were sequenced as described in Gautschi et al. (2000) and the sequences submitted to GenBank (Table 1). Primer design was carried out using primer 3 software (Rozen & Skaletsky 1998), oligonucleotides were synthesized by Microsynth GmbH (Switzerland), and one primer for each pair was labelled with fluorescent dye (see Table 1). PCR amplification for polymorphism assessment was performed in a 10 µL reaction volume containing 10 ng of genomic DNA, 50 mm KCl, 1.5 mm MgCl2, 10 mm Tris –HCl (pH 9.0), 150 µm of each dNTP (Amersham Pharmacia), 0.5 µm each of forward and reverse primer and 0.5 units of Taq DNA polymerase (Amersham Pharmacia). We used the following © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2193 thermotreatment on a PTC-100™ Programmable Thermal Controller (MJ Research Inc.): 25 – 30 cycles at 95 °C for 30 s, the locus-specific annealing temperature (Table 1) for 30 s, and 72 °C for 30 s. Before the first cycle, a prolonged denaturation step (95 °C for 5 min) was included, and the last cycle was followed by an extra 8 min extension. The amplified products were diluted with double-distilled water containing GENESCAN-350 (TAMRA) Size Standard (PE Biosystems) and genotyped on an ABI Prism 310 Genetic Analyser using GeneScanAnalysis® Software version 2.1 and Genotyper® version 2.1 software (PE Biosystems). Observed and expected heterozygosities for each locus were calculated using Popgene version 1.32 (Yeh & Boyle 1997). All eight microsatellite loci reported here were variable in Natrix tessellata and detected between four and 11 alleles in the two populations studied. Likelihood ratio tests indicated significant deviations from Hardy–Weinberg equilibrium (HWE) at loci µNt1 and µNt3, suggesting that null alleles may be present at these loci. Genotype frequencies at all other loci conformed to HWE. These microsatellites will therefore provide a valuable tool for the analysis of genetic variation in natural and allochthonous populations of the dice snake and help to devise appropriate conservation management strategies for small and endangered populations. Acknowledgements The work was supported by the DGHT (Deutsche Gesellschaft für Herpetologie und Terrarienkunde), by the Barth-Fonds from the ETH Zürich (011/1994 - 28) and by a Swiss National Science Foundation Grant 31-49477.96 to Dr J.-P. Müller, B.G. and Professor B. Schmid. References Gautschi B, Tenzer I, Müller JP, Schmid B (2000) Isolation and characterization of microsatellite loci in the bearded vulture (Gypaetus barbatus) and cross-amplification in three Old World vulture species. Molecular Ecology, 9, 2193–2195. Gruschwitz M (1985) Status und Schutzproblematik der Würfelnatter (Natrix tessellata LAURENTI, 1768) in der Bundesrepublik Deutschland. Natur und Landschaft, 60, 353–356. Hecht G (1930) Systematik, Ausbreitungsgeschichte und Ökologie der europäischen Arten der Gattung Tropidonotus (Kuhl) H. Boie. Mitteilungen aus dem Zoologischen Museum in Berlin, 16, 244–393. Mebert K (1993) Untersuchung zur Morphologie und Taxonomie der Würfelnatter Natrix tessellata (LAURENTI) 1768 in der Schweiz und im südlichen Alpenraum. Diploma Thesis, University of Zurich. Rozen S, Skaletsky HJ (1998) Primer 3. Code available at http:// www-genome.wi.mit.edu/genome_software/other/primer3.html Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Tenzer I, degli Ivanissevich S, Morgante M, Gessler C (1999) Identification of microsatellite markers and their application to population genetics of Venturia inaequalis. Phytopathology, 89, 748– 753. Yeh FC, Boyle TJB (1997) Population genetic analysis of co-dominant and dominant markers and quantitative traits. Belgian Journal of Botany, 129, 157. PRIMER 1141 2000 Graphicraft 10902 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Isolation and characterization of microsatellite loci in the bearded vulture (Gypaetus barbatus) and cross-amplification in three Old World vulture species B . G A U T S C H I , * I . T E N Z E R , † J . P. M Ü L L E R , ‡ and B . S C H M I D * *Institut für Umweltwissenschaften, Universität Zürich-Irchel, Winterthurerstrasse 190, CH-8057 Zürich, Switzerland, †Institute of Plant Sciences, Pathology Group, Universitätstrasse 2, ETH-Zentrum, CH-8092 Zürich, Switzerland, ‡Bündner Natur-Museum, Masanserstrasse 31, CH-7000 Chur, Switzerland Keywords: Aegypius monachus, conservation genetics, cross-species amplification, Gypaetus barbatus, Gyps fulvus, Neophron percnopterus Received 25 June 2000; revision accepted 16 August 2000 Correspondence: B. Gautschi. Fax: + 41 1635 57 11; E-mail: babagaut@uwinst.unizh.ch During the last one hundred years the bearded vulture, Gypaetus barbatus, has suffered extreme population declines in Europe primarily because of hunting, but changes also in agriculture, and especially in grazing practices, have resulted in poor food conditions for carrion feeders. Small populations have survived in the Pyrenees, on Corsica and Crete. The population in the Alps was completely extinct by the beginning of the 20th century. To re-establish a self-sustaining population in the Alps, over 90 juvenile bearded vultures have been released since 1986, all originating from a captive population. The amount of genetic variability in the captive and released populations, genealogical relationships between individuals, and the degree of gene flow among wild populations in the past (represented by over 200 Museum specimens) and at present are important criteria in the development of a genetic management strategy. We describe the development of 14 microsatellite primers for conservation genetic analyses of the bearded vulture. We designed the primers with special emphasis on their later use for ancient DNA (aDNA) and tested their suitability for use in other Old World vulture species. We constructed a genomic library enriched for CA and GA repeats using a modification of the method described in Tenzer et al. (1999). Digestion of total genomic DNA with Tsp509I (New England Biolabs), isolation of 200 – 700 bp fragments and ligation to TSPADSHORT/TSPADLONG linker sequences were carried out according to Tenzer et al. (1999). The ligation produces blunt-ended molecules that were amplified in 40 25 µL reactions containing 10 mm Tris-HCl, pH 9.0; 50 mm KCl, 1.5 mm MgCl2; 0.1% TritonX 100; 0.2 mg/mL BSA, 150 µm of each dNTP, 1 µm of TSPADSHORT (primer), and 1.25 U of Taq DNA polymerase (Appligene oncor). The thermal profile on a PTC100™ Programmable Thermal Controller (MJ Research) was 30 cycles of 93 °C for 1 min, 55 °C for 1 min and 72 °C for 1 min. An initial 5 min extension step at 72 °C allowed the DNA polymerase to synthesize the nick between genomic DNA and linker sequences. Polymerase chain reaction (PCR) products were hybridized to biotinylated (CA)13 and (GA)13 probes that were immobilized onto Dynabeads M-280 Streptavidin (DYNAL, France). Hybridization was carried out 2194 P R I M E R N O T E S Table 1 Genetic characteristics of 14 bearded vulture microsatellite loci. Data on numbers of alleles and heterozygosities are based on genotypes of 30 bearded vulture individuals. Ta, locus-specific annealing temperature; HO, observed heterozygosity (direct count); HE, unbiased expected heterozygosity (Nei 1978). No significant departures from Hardy–Weinberg equilibrium were detected using likelihood ratio tests. The characteristics of repeat motifs and sizes are based on the sequenced clones (GenBank Accession nos: AF270729 –AF270742) Locus Primer Sequences (5′–3′) Repeat motif Ta (°C) BV 1 ATACTTTGGCTGCATGAAGTGC† GGTCTCACTCCTTGTGTCCC CAGCATGTTATTTTGGCTGC† TTGCTAAACCGGTTAGAAGTTG GTTCTGAGGGTAGAGGGACTG† GCTGAGCAGCTTCAGAAAGTC AATCTGCATCCCAGTTCTGC† CCGGAGACTCTCAGAACTTAAC TGAACTCCTGGAGACTTCCC† CTCCTTGTAGCGTTGCCTTC TGGCATGCTGCTATGAGAAC† GTGCTTTGCATGCTTTTACTC ATCTAGGGACATCGAGGAGC† ACAGGGATGCAGGTAAGCC TGTTTGCAAGCTGGAGACC† AAAAGCCTTGGGGTAAGCAC TCAGGTTTTGACGACCTTCC† GTGGTAACGGAGGAACAAGC AAAACAGAGTTTTCACATTTTCATAAG TTCAGGAAACAGAAGCATGAAC† GGCAGTGTGGAGCCTACATC† CTCCAGGGTCCTTGTTTGC CCCCTCACCTCACAGTCAC† GGAGTGATTTTCATTGTCTTGC TGATGTGCAGATGCGTGAC† GGACTCTGATGAAGCCAAGC GAACAGCACTGAACGTGAGC† GTTTCTCCTGACAGTGAAATAACTC (CA)14 58 (CA)11 BV 2 BV 5 BV 6 BV 7 BV 8 BV 9 BV 11 BV 12 BV 13 BV 14 BV 16 BV 17 BV 20 Size (bp) HO HE 3 101 0.400 0.371 58 4 136 0.633 0.705 (CA)17 62 6 197 0.733 0.708 (CA)11 60 7 115 0.633 0.586 (CA)15 55* 5 247 0.467 0.502 (CA)11 60 2 113 0.067 0.066 (TA)6 (CA)11 60 2 219 0.567 0.463 (CA)22 62 11 181 0.867 0.884 (CA)15 62 11 245 0.867 0.857 (CA)16 50 10 184 0.900 0.900 (CA)16 60 6 179 0.733 0.730 (GA)3(CA)3A13(GA)13 AACC(GA)8 (CA)11 62 13 221 0.867 0.825 58 2 199 0.267 0.325 (CA)13 58 3 141 0.267 0.320 No. of alleles *For this PCR, a hotstart protocol is needed. PCR conditions are as described before but with HotStarTaq™ DNA Polymerase and buffer (Tris-Cl (NH4)2SO4, 1.5 mm MgCl2; pH 8.7, Qiagen) and an initial denaturing step of 95 °C for 15 min. †Fluorescent labelled primer. as described in Tenzer et al. (1999). Retained fragments were amplified in a second PCR as above without the initial extension step. PCR products were digested with EcoRI (Amersham Pharmacia) in preparation for ligation with dephosphorylated pUC18 (precut EcoRI/BAP, Amersham Pharmacia). Following transformation of JM109 High Efficiency competent cells (Promega), plating onto selective agar media and dot-blotting colonies onto Nylon-Membrane (Hybond™-N +, Amersham Pharmacia), the library was screened for inserts containing CA and GA repeats using oligonucleotide probes labelled with ECL3′-oligolabelling and detection system (Amersham Pharmacia). Hybridization was carried out in accordance with the manufacturer’s instructions. Plasmids from positive clones were sequenced using M13 forward and reverse primers and the ABI PRISM® BigDye™ Terminator Cycle Sequencing Ready Reaction Kit (PE Biosystems). Sequences were analysed on an ABI Prism 310 Genetic Analyser and edited with Sequence Navigator Software (PE Biosystems). Primers were designed using Primer 3 software (Rozen & Skaletsky 1998). Where possible, we considered only primers that did not bind to a template thymine or cytosine residue at the 3′ end because these nucleotides are most likely to be degraded in aDNA (Pääbo 1989). One primer for each pair was labelled with fluorescent dye (Table 1). To assay variation among individuals, amplifications were performed in 10 µL volumes containing 10 ng of genomic DNA, 50 mm KCl, 1.5 mm MgCl2, 10 mm Tris-HCl (pH 9.0), 150 µm per dNTP, 0.5 µm of each primer, 0.5 U of Taq DNA Polymerase (Amersham Pharmacia), and the following thermotreatment: 25 – 30 cycles of 30 s at 95 °C, 30 s at locus specific annealing temperature (Table 1) and 30 s at 72 °C. An initial denaturation step (95 °C for 5 min) was included and the last cycle was followed by an 8-min extension. Amplified products were diluted and mixed with formamide containing GENESCAN-350(ROX) Size Standard (PE Biosystems) and the genotype was determined on an ABI Prism 310 Genetic Analyser using GeneScan Analysis® software version 2.1 and Genotyper ® software version 2.1 (PE Biosystems). We screened 30 bearded vulture individuals from the captive population (Table 1), 15 Egyptian vultures, Neophron percnopterus, 15 black vultures, Aegypius monachus, from Spain, and 10 griffon vultures, Gyps fulvus, from France (Table 2). © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2195 Table 2 Results of the cross-species amplification. Summarized are locus specific annealing temperature (Ta), PCR product size range and number of alleles for each of the species tested. When amplification did not result in a clear allelic pattern, received fragment sizes are listed below Egyptian Vulture (n = 15) Product size (bp) Locus Ta (°C) BV 1 BV 2 BV 5 BV 6 BV 7 BV 8 BV 9 BV 11 BV 12 BV 13 BV 14 BV 16 BV 17 BV 20 50 100 55 123 58* 190 58 95–117 No amplification 58 106 58* 224–228 58 148–150 58 235–239 50 170–176 55 161–163 58 179–185 55* 185 55 133–135 Griffon Vulture (n = 10) Number of alleles 1 1 1 5† 1 3 2 3 4 2 4 1 2 Ta (°C) Product size (bp) 50 91 55* 119 58* 177–183 58 118 –120 No amplification 58 106 58 207 58 152–162 58* 243–279 50 172–178 55 162–164 58 184–188 55* 185–187 55 132–138 Black Vulture (n = 15) Number of alleles 1 1 4 2 1 1 4 7§ 3 2 3 2 4 Ta (°C) Product size (bp) 58 91 55 118 58 189–201 55* 127–161 No amplification 58 103 58 205 58 164–180 58 227 50 174–176 55 158 55 160–170 55 185–187 55 136–140 Number of alleles 1 1 4 4‡ 1 1 7 1 2 1 4 2 3 *For this PCR, a hotstart protocol is needed. PCR conditions are as described before but with HotStarTaq™ DNA Polymerase and buffer (Tris-Cl (NH4)2SO4, 1.5 mm MgCl2; pH 8.7, Qiagen) and an initial denaturing step of 95 °C for 15 min. †Amplification products are (in bp): 95, 97, 105, 115, 117. ‡Amplification products are (in bp): 127, 133, 139, 161. §Amplification products are (in bp): 243, 245, 259, 263, 265, 267, 279. We received an individual genetic fingerprint for all analysed captive birds [probability of identity for sibs (PIsibs) = 7.8 × 10 –5; see Taberlet & Luikart 1999) ], showing that apart from the conservation genetic analysis mentioned above, these microsatellites will provide an important tool in the long term monitoring of the released population in the Alps. Acknowledgements We would like to thank Hans Frey (Veterinärmedizinisches Institut, Universität Wien, Austria), Juan Negro (Estación Biológica Doñana, Seville, Spain) and François Sarrazin (Université Pierre et Marie Curie, Paris, France) for providing blood samples. This work was supported by the Swiss National Science Foundation Grant 31–49477.96 to JPM, BG and BS. References Nei M (1978) Estimation of average heterozygosity and genetic distance from a small number of individuals. Genetics, 89, 583–590. Pääbo S (1989) Ancient DNA: Extraction, characterization, molecular cloning, and enzymatic amplification. Proceedings of the National Academy of Sciences of the USA, 86, 1939–1943. Rozen S, Skaletsky HJ (1998) Primer 3. Code available at http://www-genome.wi.mit.edu/genome_software/other/ primer3.html. Taberlet P, Luikart G (1999) Non-invasive genetic sampling and individual identification. Biological Journal of the Linnean Society, 68, 41–55. Tenzer I, degli Ivanissevich S, Morgante M, Gessler C (1999) Identification of microsatellite markers and their application to population genetics of Venturia inaequalis. Phytopathology, 89, 748– 753. PRIMER 1138 2000 Graphicraft 10900 2 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Microsatellites for Barren Brome (Anisantha sterilis) J . M . G R E E N , K . J . E D WA R D S , S . L . U S H E R , J . H . A . B A R K E R , E . J . P. M A R S H A L L , R . J . F R O U D - W I L L I A M S * and A . K A R P IACR-Long Ashton Research Station, Department of Agricultural Sciences, University of Bristol, Long Ashton, Bristol, BS41 9AF, UK, *Department of Agricultural Botany, School of Plant Sciences, The University of Reading, 2 Earley Gate, Reading, Berkshire RG6 6AU, UK Keywords: Anisantha sterilis, Bromus, microsatellites, weed Received 24 July 2000; revision accepted 16 August 2000 Correspondence: Dr Angela Karp. Fax: 01275 394007; E-mail: angela.karp@bbsrc.ac.uk Barren Brome (Anisantha sterilis: synonym Bromus sterilis) is a diploid grass weed of cereals which naturally occurs in field margins and waste ground. It is an inbreeding annual which can invade and compete with cereal crops especially when cereals are grown repeatedly with minimum cultivation (Cussans et al. 1994). Investigations into the genetic diversity within A. sterilis may provide indications of its ability to respond to future control measures. Here, we describe the identification of polymorphic microsatellites in A. sterilis for population genetic studies. A small-insert genomic library, enriched for microsatellites, was developed using a modified procedure of Edwards et al. (1996). Genomic DNA (2 µg) was digested with RsaI and SspI. An MluI adapter [21-mer: (5′-CTCTTGCTTACGCGTGGACTA-3′ ) and phosphorylated 25-mer: (5′-pTAGTCCACGCGTAAGCAAGAGCACA-3′ ) ] was ligated to the ends of the DNA 2196 P R I M E R N O T E S fragments. Five identical reactions (25 µL each) were prepared: 2 µL ligated DNA, 1× polymerase chain reaction (PCR) buffer (10 mm Tris-HCl, pH 8.5, 1.5 mm MgCl2, 50 mm KCl, 0.001% gelatine), 200 µm each dNTP, 300 ng 21-mer and 1 U Taq polymerase (GibcoBRL). Amplification proceeded for 20 cycles (94 °C for 20 s, 60 °C for 1 min and 72 °C for 3 min) using a Perkin Elmer 9600 Thermal Cycler. Reactions were pooled, purified by phenol– chloroform extraction, concentrated by ethanol precipitation and resuspended in 25 µL sterile distilled water (SDW). Oligonucleotides [ (CA)15, (CT)15] were cross-linked by UV irradiation to separate 0.7 cm2 nylon membranes (Hybond N+, Amersham), then used to hybridize the amplified DNA in one tube, at 45 °C overnight. Filters were washed four times in 2× SSC (20× SSC: 3 m NaCl, 0.3 m Na Citrate, pH 7) at 65 °C for 5 min and three times in 1× SSC at 65 °C for 5 min. Eluted DNA was ethanol precipitated and resuspended in 25 µL SDW. Five identical reactions (25 µL) were prepared: 2 µL DNA, 1× PCR buffer, 400 µm each dNTP, 200 ng 21-mer and 2 U Taq polymerase. Amplification proceeded for 25 cycles (94 °C for 30 s, 60 °C for 1 min and 72 °C for 3 min). Reactions were pooled, purified, concentrated and resuspended as before. DNA was digested with MluI and fragments were selected using a Size Sep™ 400 Spun Column (Pharmacia). Fragments were cloned into pJV1 vector and transformed into Epicurian coli® Competent Cells (XL1-Blue MRF′ Kan Supercompetent Cells, Stratagene). Plasmid DNA was extracted using Wizard™ Plus Minipreps DNA Purification System (Promega) and sequenced using the ABI Prism™ Dye Terminator Cycle Sequencing Ready Reaction Kit. Sequences were separated on the ABI Prism 377 DNA Sequencer. Primers were designed using primer version 0.5 (Whitehead Institute for Biomedical Research, Massachusetts) and synthesized by Genosys Inc. Table 1 Microsatellite library characterization Clones sequenced 97 Microsatellites >18 base pairs Of these: CA/GT repeats CT/GA repeats CT/GA & CA/GT repeats Duplicated sequences 56 41 6 6 3 Primers designed 17 Monomorphic Unscorable Multilocus No product Informative 2 3 2 1 9 Leaf samples were collected from 20 A. sterilis plants at an Oxfordshire farm and six A. diandra plants from farms across England. Genomic DNA was extracted using the Nucleon® Phytopure Extraction Kit (Amersham). DNA was also extracted from one seed of A. rigida (Australia). For genotyping, the forward primer was end-labelled with [γ 33P]-ATP (Amersham). Microsatellite amplification was performed in 12.5 µL reactions: 5 ng DNA, 1× PCR buffer, 200 µm each dNTP, 25 ng forward primer, 25 ng reverse primer, 0.5 U Taq polymerase. Amplification proceeded for 35 cycles (94 °C for 1 min, 54 °C for 1 min and 72 °C for 1 min) and one cycle of 72 °C for 10 min. PCR products were separated on 6% polyacrylamide denaturing gels using M13 control sequence as a size marker and exposed to Kodak Biomax MR-1 film overnight. Table 2 Primer sequences and characteristics of Anisantha sterilis microsatellites Locus Primer sequences (5′– 3′) Repeat motif Size range (bp) No. of alleles Gene diversity AS115 *† AS124 * AS133 † AS139 F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: (GT)19 117–131 3 0.585 (CA)5AG(CA)6AGCG(CA)20 182–184 2 0.320 (TG)15C(GT)2(TG)3A(GT)4 204–210 3 0.580 (TG)23 174–188 3 0.485 (CA)28AA(CA)10GT(CA)11 194–210 5 0.780 (GT)3(TG)2C(TG)2C(TG)8C(GT)14 178–187 4 0.685 (AC)3(AT)7AA(GT)16(A)5 144–170 4 0.700 (CA)12 126–140 3 0.545 (TG)15 146–150 3 0.535 AS147 * AS152 * AS184 * AS211 *† AS219 *† GTTGCTGCTGCCAGGCTGA TTAACAAAACAGGCAACACA GAATGTAGATAAAAACTGGTGT GCACTCACTTCATAAATTCAA ATGGACAACCATGGCGTGAGA TGATAGAAGTAATACGAGGCG AAACACCAAAAATAATTAAGG GCCCATCCAACATGTGCCAG ATTTTAGCTGATGTGCTTTTG ACTGTGGTGATCGTACCCGTG AAGGTTCAAAGTGTAAGGACG AGGAGAAGAAGAACGAGAGAA CGGAATGTTGTCAGAATAGTT ACGAACCGTGGAACTTGTTAC TTCTATGTAATCATGGCTTGC TCCAAGGACCGACCGATCTC CAGGAATTTGTCAGGTTAAG AGCTATAAAAGTAACCACATCA *Polymorphic in A. diandra; †cross-amplification in A. rigida. GenBank accession nos: AF285620 –AF285628. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2197 The results of sequencing and characterizing 97 library clones are shown in Table 1. Nine primer pairs amplified polymorphic markers producing 2–5 alleles per locus and gene diversity (D = 1−Σpi2) (Nei 1973) values from 0.32 to 0.78 (Table 2). No heterozygotes were observed. Crossamplification was investigated by testing the primers on the related species A. diandra and A. rigida. All nine loci were amplified from A. diandra and seven were polymorphic with between two and seven alleles per locus. For A. rigida, four loci were amplified (Table 2). Acknowledgements We acknowledge a Lawes Trust-University of Reading studentship award to Miss J. Green. We thank Chloe Aldam for technical help. IACR receives grant-aided support from the Biotechnology and Biological Sciences Research Council of England. References Cussans GW, Cooper FB, Davis DHK, Thomas MR (1994) A survey of the incidence of the Bromus species as weeds of winter cereals in England, Wales and parts of Scotland. Weed Research, 34, 361– 368. Edwards KJ, Barker JHA, Daly A, Jones C, Karp A (1996) Microsatellite libraries enriched for several microsatellite sequences in plants. Biotechniques, 20, 758–760. Nei M (1973) Analysis of gene diversity in subdivided populations. Proceedings of the National Academy of Sciences of the USA, 70, 3321– 3323. PRIMER 1159 2000 Graphicraft 1932 NOTEs Limited, Hong Kong Development and characterization of microsatellite loci from lynx (Lynx canadensis), and their use in other felids L . E . C A R M I C H A E L , W. C L A R K and C . STROBECK Department of Biological Sciences, University of Alberta, Edmonton, AB, Canada, T6G 2E9 Keywords: cross-species amplification, felids, lynx, microsatellites, primers Received 29 July 2000; revision accepted 18 August 2000 Correspondence: LE Carmichael. c/o Curtis Strobeck, Department of Biological, Sciences University of Alberta, Edmonton, Alberta, Canada, T6G 2E9. Fax: +780 492 9234; E-mail: lindsey_carmichael@hotmail.com On 24 March 2000, the United States Fish and Wildlife Service declared the Canadian lynx (Lynx canadensis) to be threatened throughout the contiguous United States (United States Fish and Wildlife Service Website: http://www.fws.gov/). Lynx conservation programmes have been attempted in Colorado (Kloor 1999) and are currently in development throughout the contiguous United States. Because an understanding of the genetics of wildlife populations may assist in their conservation, we set out to identify microsatellite markers that might facilitate population genetic studies of Canadian lynx. Muscle tissue chips from a single lynx were frozen in liquid nitrogen and ground to powder. High molecular weight genomic © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 DNA was then isolated by phenol extraction (Sambrook et al. 1989) and digested to completion with Sau3A. Fragments of 200 – 800 bp (size-selected as in Davis & Strobeck 1998) were cloned into BamHI-linearized M13 mp18 RF, and transformed into Escherichia coli strain DH5ãF′IQ (Gibco BRL) made competent using the SEM (18 °C) method of Inoue et al. (1990). The library was plated in 0.7% top agarose (containing X-gal and IPTG) at a density of 1000–2000 plaques per 150 mm Petri plate. Approximately 2600 recombinant clones were screened with a biotinylated (GT)12 oligonucleotide probe, and clones containing putative microsatellites identified using a nonradioactive detection kit (BluGene®, Gibco BRL). Forty-one insert-containing clones screened positive in the primary platings. These plaques were picked, regrown and replated at low density in a secondary hybridization/detection screen. Inserts from 24 confirmed positive clones were polymerase chain reaction (PCR) amplified using universal M13 forward and reverse primers. PCR products were then electrophoresed in 1.0% agarose (TAE) and gel-purified using the glass powder binding method of Vogelstein & Gillespie (1979). These purified products were cycle-sequenced using a dRhodamine Terminator sequencing kit (with Amplitaq DNA polymerase FS, ABI Prism, PE Applied Biosystems) and an ABI Prism 377 DNA sequencer. Primer pairs were designed for 10 microsatellite loci using oligo 4.0 (National Biosciences Inc.) and tested on lynx genomic DNA extracted from muscle samples (Alberta Fish and Wildlife). Six of these loci (Table 1) gave strong, clean PCR products. Furthermore, multiplexing allows the amplification of these six loci in three 15 µL reactions: Lc 106, Lc 110 and Lc 118 = 0.16 µm each primer, 160 µm dNTPs, 2 mm MgCl2, 0.36 U Taq DNA Polymerase (prepared as in Engelke et al. 1990) and approximately 50 ng genomic DNA [extracted using QIAamp™ spin columns (QIAGEN)] in PCR buffer (50 mm KC1, 10 mm Tris-HC1, pH8.8, 0.1% Triton-X 100, 0.16 mg/mL BSA); Lc 111 and Lc 120 = 0.16 µm each primer, 160 µm dNTPs, 2mm MgC12, 1.44 U Taq DNA Polymerase and 50 ng template DNA in PCR buffer; Lc 109 = 0.16 µm each primer, 120 µm dNTPs, 2 mm MgC12, 0.3 U Taq DNA Polymerase and 50 ng template DNA in PCR buffer. All cycling reactions were performed as in Davis & Strobeck (1998) and their products analysed on an ABI 377 Sequencer with Genescan and Genotyper software (Applied Biosystems). Twenty-nine lynx tissue samples were genotyped to estimate the variability of each locus (these samples do not represent a population as they were collected from a variety of sites in Alberta over approximately 15 years). Complete genotypes were obtained with a single exception: for one individual, locus Lc 109 could not be amplified. Size ranges and variability are given in Table 1. Mean number of alleles was 6.17, and there was no significant difference between observed and expected heterozygosity. Six additional felid species were also tested: cougar (Felis concolour); bobcat (Felis rufus); African lion (Panthera leo); Siberian tiger (Panthera tigris); Asian leopard cat (Felis bengalensis); and domestic cat (Felis catus, breed unknown). Table 2 summarizes the size range and number of alleles observed in each species. Variability in cougars does not exceed 2198 P R I M E R N O T E S Table 1 Size range of PCR products, variability, repeat motif and primer sequences for each locus. Expected heterozygosity (HE) was calculated using the formula (1−ΣPi2) Locus Size range (bp) No. of alleles HO HE Repeat motif Primer sequences (5′−3′) Lc 106 96 –108 7 0.793 0.786 (T)3(GT)17 Lc 109 172–182 6 0.893 0.801 (GT)18 Lc 110 91–103 7 0.828 0.815 (T)3(GT)14 Lc 111 140–154 6 0.586 0.619 (GT)17 Lc 118 133–145 7 0.759 0.766 (T)4(GT)22(T)2 Lc 120 196– 204 4 0.577 0.551 (T)3(GT)11(GA)13 F: TCTCCACAATAAGGTTAGC R: FAM– GGGATCTTAAATGTTCTCA F: AAGTGGCAAGATTACATTC R: TET– AACATCCTTTTATTCATTG F: CCTTTGTCACTCACCA R: TET– CGGGGATCTTCTGCTC F: GAGGATCATTGTGCAT R: FAM– ATCCACTCACCCTCTA F: TGGGGTGGGAACTCTC R: TET– AGTGCCCCAGATTTTT F: TGAGCCTGAGCATACATT R: HEX– GTTTGTGAGTTGGAGCC Accession no. AF288054 AF288055 AF288056 AF288057 AF288058 AF288059 FAM, TET and HEX are fluorescent dye labels (Gibco BRL). Table 2 Survey of amplification potential in six felid species. Size ranges are given in base pairs. Number of unique alleles observed/ number of alleles scored is provided in brackets. ‘–’ indicates no PCR product, while ‘ + ’ represents a multiple banding pattern Locus Cougar Felis concolour Bobcat Felis rufus African Lion Panthera leo Siberian Tiger Panthera tigris Asian Leopard Cat Felis bengalensis Domestic Cat Felis catus Lc 106 Lc 109 Lc 110 Lc 111 Lc 118 Lc 120 87–97 (2/16) 191–199 (2/18) 102–104 (2/12) − 112 –113 (2/16) 209 (1/20) 87–89 (2/6) 163–169 (3/6) 80 (1/6) 136–142* (3/6) 129–137 (3/6) 212–220 (3/6) 89 (1/2) 171 (1/2) 122–134 (2/2) − − 205–207 (2/2) 88 –98 (2/2) 163 (1/2) 124 (1/2) − − 202–208 (2/2) 93 –99 (2/2) 169 (1/2) 88 –90 (2/2) 146 (1/2) 111 (1/2) 201– 203 (2/2) + 167–177 (2/2) 96 (1/2) 140 (1/2) 114–116 (2/2) + *Lc 111 may include an additional Bobcat allele at 194 bp. that observed using domestic cat loci (data not shown), and patchy results for this species strongly suggest the existence of null alleles. However, the level of variation observed in lynx, and the ability of these primers to amplify microsatellites across a range of felid species, suggests they may be useful in a variety of population genetic studies. Acknowledgements We would like to thank Lisa Ostafichuk for her significant contribution to this work. The laboratory of Curtis Strobeck receives operating grants from Parks Canada and the Natural Sciences and Engineering Research Council of Canada. Kloor K (1999) Lynx and biologists try to recover after disastrous start. Science, 285, 320–321. Sambrook J, Fritch EF, Maniatus T (1989) Molecular Cloning: a Laboratory Manual. 2nd edn. Cold Spring Harbor Laboratory Press, New York. Vogelstein B, Gillespie D (1979) Preparative and analytical purification of DNA from agarose. Proceedings of the National Academy of Science of the USA, 76, 615–619. 2000 1147 109PRIMER Graphicraft 2 00 NOTEs Limited, Hong Kong Variation at tri- and tetranucleotide repeat microsatellite loci in the fruit bat genus Cynopterus (Chiroptera: Pteropodidae) References J . F. S T O R Z * Davis CS, Strobeck S (1998) Isolation, variability and crossspecies amplification of polymorphic microsatellite loci in the family Mustelidae. Molecular Ecology, 7, 1776–1777. Engelke DR, Krikos A, Bruck ME, Ginsburg D (1990) Purification of Thermus aquaticus DNA polymerase expressed in Escherichia coli. Analytical Biochemistry, 191, 396–400. Inoue H, Nojima H, Okayama H (1990) High efficiency transformation of Escherichia coli with plasmids. Gene, 96, 23 –28. Department of Biology, Boston University, 5 Cummington Street, Boston, MA, 02215, USA Keywords: Chiroptera, Cynopterus, microsatellite DNA, Pteropodidae Received 15 August 2000; revision accepted 22 August 2000 Correspondence: Jay F. Storz. *Present address: Department of Biology, Duke University, Box 90338, Durham, NC, 27708, USA. Fax: + 1 919 660 7293; E-mail: storz@duke.edu © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2199 Table 1 Primer sequences and characteristics of nine tri- and tetranucleotide repeat microsatellite loci used in the genetic analysis of Cynopterus sphinx from Pune, India (18°32′ N, 73°51′ E). Repeat numbers refer to cloned alleles and plus signs denote sequence interruptions between tracts of ≥ 2 repeat units. Ta, annealing temperature; n, number of bats genotyped per locus; NA, number of alleles per locus; HO, observed heterozygosity; HE, expected heterozygosity. Loci CSP-4, CSP-6, and CSP-7 segregated subsets of alleles that differed by 2 bp rather than 4 bp. It is not known whether this was due to interruptions within the array of tetranucleotide repeats or insertions/ deletions in flanking sequences Locus Primer sequences (5′– 3′) Repeat motif Ta (°C) Allele size range n CSP-1 F: GGGGAAACAAAGGAAAAGT R: AGAAAAGTGAGACCTGACAGAG F: CCCGATGATGGATTTCTAC R: CTGGGCTGTAATAAGTGCTC F: AACACCACCACCACCACTA R: TGTGGCAACAACTCAGACA F: GAGAGGACTCCGTTCTTTTAGA R: ATGGATGGGTGACAGATGA F: CATTTGTGGTAACTTGTGATG R: ACAGCAGTGAAACTTCCTCT F: TGAGGAGTGTTCCCGAGTA R: AAAAATCCCAACGCACAG F: CCACAAGAAACCCAATACTAAC R: CTTCCTAGCCCCACAATC F: CCAGGTGTTATGGGTTGA R: TGAGGTGTTGGGAGTTTG F: GGTCCCTCTGCTCTTCAG R: AGCATGGGGAATATAGTCAAG (ATC)2+4+5+3 55 191–218 (ATC)3+13+2 57 (ATC)8 CSP-2 CSP-3 CSP-4 CSP-5 CSP-6 CSP-7 CSP-8 CSP-9 NA HO HE GenBank accession no. 431 9 0.73 0.71 AF289705 113–134 431 7 0.78 0.74 AF289706 57 95 –107 431 5 0.37 0.38 AF289707 (CATC)12 57 139–163 431 10 0.79 0.78 AF289708 (ATGG)8(ACGG)4 55 110–170 431 12 0.76 0.73 AF289709 (CATC)10 55 127–219 431 14 0.81 0.85 AF289710 (TATC)3+8 57 231–265 431 17 0.82 0.82 AF289711 (TAGA)3+3+5+11 57 150–202 420 14 0.75 0.74 AF289712 (TAGA)3+7 57 278–298 431 5 0.49 0.47 AF289713 Species in the fruit bat genus Cynopterus (Chiroptera: Pteropodidae) are widely distributed across the Indomalayan region (Corbet & Hill 1992). The two most geographically widespread members of the genus are the short-nosed fruit bat (Cynopterus sphinx) and the lesser dog-faced fruit bat (C. brachyotis). There is considerable uncertainty surrounding the taxonomic relationship between C. sphinx and C. brachyotis, and the status of the many named forms within C. sphinx (Storz & Kunz 1999). The availability of polymorphic microsatellite markers for cynopterine fruit bats would greatly aid efforts to elucidate species boundaries and genetic correlates of morphological variation within species. The primary motivation for developing microsatellite markers for C. sphinx was to investigate the influence of polygynous mating and harem social organization on population genetic structure (Storz et al. 2000a,b). Efforts are also underway to investigate comparative levels of geographical differentiation in body size and microsatellites in Indian populations of C. sphinx (see Storz et al. 2000c). Genomic DNA was isolated from wing-membrane biopsy samples of C. sphinx using QIAamp extraction columns (Qiagen). Microsatellite loci were isolated from three genomic libraries enriched for tri- and tetranucleotide repeat motifs following the methods of Jones et al. (2000). Following partial digestion with a combination of seven blunt-end restriction endonucleases, size-selected genomic fragments (350 – 650 bp) were ligated to 20 bp oligonucleotide adapters that contained a HindIII restriction site. Genomic fragments were subjected to magnetic bead capture using the following 5′-biotinylated oligonucleotides: ATG8, CATC8, and TAGA8 (Integrated DNA Technologies). Captured fragments were Polymerase chain © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 reaction (PCR)-amplified using primers complementary to the adapter sequences. The resultant products were ligated into the HindIII restriction site of the plasmid pUC19. Recombinant plasmids were transfected into Escherichia coli strain DH5α by electroporation. Colonies were screened according to the protocol of Jones et al. (2000). Following PCR amplification, a total of 27 clones in the size range 350 – 650 bp were sequenced using Prism Cycle Sequencing kits and labelled dNTP’s (Applied Biosystems). Sequences were resolved on an ABI 373 automated sequencer (Applied Biosystems). All clone sequences contained at least one microsatellite locus. Primers were designed for a total of 21 microsatellite loci using the program Designer PCR version 1.03 (Research Genetics). Primer pairs were tested by amplifying DNA from eight individual C. sphinx sampled from various localities in peninsular India. Sixteen primer pairs amplified variable PCR products, as revealed by electrophoresis in 3.5% agarose gels followed by ethidium-bromide staining. Nine primer pairs that yielded the most consistent results were selected for further testing, and the forward primer of each pair was fluorescently labelled with 6-FAM, TET, or HEX (Applied Biosystems). PCR was performed using 20 µm of each primer, 5 mm dNTP’s, 25 mm MgCL2, 0.012 U of AmpliTaq DNA polymerase (Applied Biosystems), 10× PCR buffer (100 mm Tris-HCl buffer, pH 8.3, 500 mm KCl), ddH2O, and 10 ng of template DNA in a total reaction volume of 15 µL. Thermal cycling was performed in a GeneAmp PCR System 9700 (Applied Biosystems) under the following conditions: initial denaturation at 94 °C for 2 min followed by 35 cycles of denaturation at 94 °C for 30 s, annealing at 55– 57 °C (Table 1) for 45 s, and extension at 72 °C for 2200 P R I M E R N O T E S Table 2 Summary statistics for five tri- and tetranucleotide repeat microsatellite markers used in the genetic analysis of Cynopterus sphinx (from localities <18° N latitude) and C. brachyotis in peninsular India. n, number of bats genotyped per locus; NA, number of alleles per locus; HO, observed heterozygosity; HE, expected heterozygosity. In both species, locus CSP-7 segregated multiple alleles with lengths that differed by 2 bp, even though the cloned allele was a (TATC)n repeat Cynopterus sphinx (southern localities) Cynopterus brachyotis Locus Allele size range n NA* HO HE Allele size range n NA HO HE CSP-1 CSP-2 CSP-5 CSP-7 CSP-9 191– 224 119 – 134 130– 190 227– 285 286– 302 189 189 189 189 189 12 6(7) 11(16) 21 5(6) 0.79 0.74 0.82 0.78 0.55 0.86 0.71 0.81 0.84 0.60 176–227 101 110–166 229–263 270–282 111 20 111 111 111 12 1 11 17 4 0.61 0 0.29 0.72 0.45 0.69 0 0.34 0.86 0.49 *Numbers in parentheses refer to numbers of alleles observed in the complete sample of C. sphinx, from Pune and the southern localities (n = 620 bats). 50 s (with a final extension at 72° for 2 min 30 s). Allele sizes were quantified using an ABI Prism 377 automated sequencer and analysed using genescan software (PE Applied Biosystems). To assess levels of variation in C. sphinx and C. brachyotis, microsatellite genotypes were obtained for a total of 731 bats (620 C. sphinx and 111 C. brachyotis). A total of 431 adults and juveniles of C. sphinx from a single population in Pune, India (Storz et al. 2000b) were genotyped at all nine loci (Table 1). A total of 185 known mother–offspring pairs were examined, and no genotypic mismatches were observed at any locus. Using a subset of five microsatellite loci, an additional 189 C. sphinx that were sampled from localities in south-western India (see Storz et al. 2000c), and 111 C. brachyotis that were sampled from high-elevation wet forest sites in the Western Ghats were genotyped. In the total sample of C. sphinx (n = 620), mean number of alleles per locus was 12.4 (range = 6 – 21; Table 2). Although preliminary screening of 20 individuals indicated that CSP-2 was monomorphic in C. brachyotis, the remaining four loci segregated 4 –17 alleles. Relative to C. sphinx, homologous loci in C. brachyotis segregated alleles that were generally shorter in length (Table 2). These markers should open up many new opportunities for studying the population biology and phylogeography of Old World fruit bats. Acknowledgements Jones KC, Levine KF, Banks JD (2000) DNA-based genetic markers in black-tailed and mule deer for forensic applications. California Fish and Game, 86, in press. Storz JF, Balasingh J, Bhat HR et al. (2000c) Clinal variation in bodysize and sexual dimorphism in an Indian fruit bat, Cynopterus sphinx (Chiroptera: Pteropodidae). Biological Journal of the Linnean Society, 71, in press. Storz JF, Balasingh J, Nathan PT, Emmanuel K, Kunz TH (2000a) Dispersion and site-fidelity in a tent-roosting population of the short-nosed fruit bat (Cynopterus sphinx) in southern India. Journal of Tropical Ecology, 16, 117–131. Storz JF, Bhat HR, Kunz TH (2000b) Social structure of a polygynous tent-making bat, Cynopterus sphinx (Megachiroptera). Journal of Zoology (London), 251, 151–165. Storz JF, Kunz TH (1999) Cynopterus sphinx. Mammalian Species, 613, 1– 8. Graphicraft 2000 1149 00 91PRIMER 2 NOTEs Limited, Hong Kong Isolation and characterization of microsatellite DNA loci in Japanese flounder Paralichthys olivaceus (Pleuronectiformes, Pleuronectoidei, Paralichthyidae) Funding was provided by the National Geographic Society, the Lubee Foundation, and the National Science Foundation (DEB 97 – 01057). I thank H. R. Bhat, J. Balasingh, G. Marimuthu, P. T. Nathan, A. A. Prakash, and D. P. Swami Doss for assistance with field collections. *Tohoku National Fisheries Research Institute, Shinhama, Shiogama, Miyagi, 985 – 0001, Japan, †National Research Institute of Aquaculture, Nansei, Watarai, Mie, 516 – 0193, Japan References Keywords: DNA, Japanese flounder, microsatellites, Paralichthyidae, Paralichthys olivaceus Corbet GB, Hill JE (1992) The mammals of the Indomalayan Region: a systematic review. British Museum Publications, Oxford University Press, Oxford. M . S E K I N O * and M . H A R A † Received 15 August 2000; revision accepted 22 August 2000 Correspondence: M. Sekino. Fax: + 81 22-367-1250; E-mail: sekino@myg.affrc.go.jp © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2201 Japanese flounder Paralichthys olivaceus is an important species consisting coastal fisheries resources in Japan, and is of high commercial value. Interest has been directed toward resource enhancement, and accordingly, millions of P. olivaceus are released into Japanese coastal fisheries grounds every year (Furusawa 1997), yet little is known about reproductive success of the stocked fish. To promote effective stocking management, it is necessary to monitor the fate of stocked fish and their relatedness apart from naturally reproduced fish. Microsatellite DNA loci are expected to provide an invaluable tool for this purpose because of the power and ability of microsatellite markers in regard to resolution for genetic relatedness among individuals (Blouin et al. 1996) and parentage determination (O’Reilly et al. 1998). Here, we describe the characterization of microsatellites isolated from P. olivaceus that will be useful to address the stocking effects. The method described by Sekino et al. (2000) was used for cloning P. olivaceus microsatellites. In brief, genomic DNA was fragmented by sonication. Sonicated fragments were blunted by mung bean nuclease (Takara, Shiga, Japan), and the fragments ranging from 300 – 500 bp were recovered. The fragments were ligated into SrfI site of pCR-Script Amp SK(+) vector (Stratagene, La Jolla, CA, USA), and recombinant plasmid vector was transformed into XL2-Blue MRF′ ultracompetent cells (Stratagene). Single-stranded DNA was prepared, and selective second-strand DNA synthesis was employed using (CA)12 oligonucleotide and cloned pfu DNA polymerase (Stratagene). The resultant double-strand DNA was transformed into XL2-Blue MRF′ cells again and these transformants were referred to a (CA)nenriched library. From the library, 80 clones were randomly chosen, and plasmid DNAs were purified using GFX Micro Plasmid prep kit (Amersham Pharmacia Biotech, Uppsala, Sweden). The DNA sequences were determined in both directions using Thermo Sequenase™ cycle sequencing kit (Amersham Pharmacia Biotech) in combination with KS and T3 primers and subjected to an ALFexpress automated DNA sequencer (Amersham Pharmacia Biotech). Of the 80 clones, 59 contained one or more repeat sequences. We designed 27 polymerase chain reaction (PCR) primer pairs using a Premier software package (Premier Biosoft International, Palo Alto, CA, USA). To examine microsatellite polymorphisms, PCR was employed. PCR amplification was carried out in a 20 µL reaction volume, which included 20 pmols of each primer set (one primer in each pair was 5′ end-labelled with Cy5), 100 µm of each dNTP, 10 mm Tris-HCl (pH 8.3), 50 mm KCl, 1.5 mm MgCl2, 0.001% gelatin, 0.5 U of Ampli Taq GoldTM (Perkin Elmer, Foster City, CA, USA), and approximately 50 ng of template DNA using PC-960G gradient thermal cycler (Corbett Research, Mortlake, NSW, Australia). PCR amplification cycles were as follows: 12 min at 95 °C, 35 – 40 cycles of 30 s at 94 °C, 1 min at a primer-specific temperature, 1 min at 72 °C, and final elongation for 5 min at 72 °C. Analyses of PCR products were performed using ALFexpress sequencer in combination with an Allelelinks software package (Amersham Pharmacia Biotech). All 27 microsatellite loci were successfully amplified, © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 out of which we finally chose 16 primer sets (the remaining 11 having been rejected because their polymorphisms were low, and/or they produced unexpected PCR products in an initial sample of P. olivaceus) and assessed further microsatellite polymorphisms in a natural P. olivaceus population collected from the Japanese coast of the Japan Sea. As shown in Table 1, the number of alleles ranged from 4 – 40, and the observed and expected heterozygosity ranged from 0.43 – 0.99 and 0.43 – 0.97, respectively. All but one of the 16 loci conformed to Hardy–Weinberg’s (HW) equilib rium in the Markov-chain method (parameters used; 100 000 Markov-chain steps; 10 000 dememorization steps), using an Arlequin verion 1.1 software package (Schneider et al. 1997). At the Po31 locus, the observed genotype frequencies showed significant departure from HW expectations (P < 0.05) with a large discrepancy between the observed and expected heterozygosity (0.34 and 0.91, respectively). This may be explained by sampling errors due to limited sample size or substructuring of the samples, however, this seems unlikely because the observed genotype frequencies in all other 15 loci were consistent with the expectations. We believe that the presence of null alleles (Pemberton et al. 1995) may be a valid explanation causing these results. Further investigation of this topic is necessary. Microsatellite DNA loci described in the present study possess hypervariability, suggesting that these loci will be useful for genetic monitoring of stocked P. olivaceus in furthering our understanding of stocking effects. Acknowledgements We express gratitude to Dr H. Takahashi, National Research Institute of Agro-biological Resources, for the technical advises and contributions. References Blouin MS, Parsons M, Lacaille V, Lotz S (1996) Use of microsatellite loci to classfy individuals by relatedness. Molecular Ecology, 3, 393–401. Furusawa T (1997) Key problems of sea-farming associated with its perspective. In: Biology and Stock Enhancement of Japanese Flounder (eds Minami T, Tanaka M), pp. 117–126. Koseishakoseikaku, Tokyo. (in Japanese). O’Reilly PT, Herbinger C, Wright JM (1998) Analysis of parentage determination in Atlantic salmon (Salmo salar) using microsatellites. Animal Genetics, 29, 363–370. Pemberton JM, Slate J, Bancroft DR, Barrett A (1995) Nonamplifying alleles at microsatellite loci: a caution for parentage and population studies. Molecular Ecology, 4, 249–252. Schneider S, Kueffer JM, Roessli D, Excoffier L (1997) Arlequin version 1.1: A software for population genetic data analysis. Genetics and Biometry Laboratory, University of Geneva, Switzerland. Sekino M, Takagi N, Hara M, Takahashi H (2000) Microsatellites in rockfish Sebastes thompsoni (Scorpaenidae). Molecular Ecology, 9, 634–636. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Locus Core repeat sequence (5′−3′) Primer sequence (5′−3′) Anneal. (°C) Sample size No. of alleles Size range† (bp) HO HE P‡ GenBank accession no.§ Po1 (TG)3T2(TG)8 55 67 20 160– 216 0.68 0.71 0.84 AB046745 Po13 (TG)3GA(CA)13 58 69 23 206– 276 0.78 0.92 0.18 AB046746 Po20 58 69 40 239– 379 0.99 0.97 1.00 AB046748 Po25A (CACG)4(CA)4CG(CA)18 C(GT)3 (GATG)2A2CA(GATG)10 55 68 12 201– 253 0.76 0.76 0.26 AB046749 Po26 (CA)6CGCACGGA(CA)7 55 67 5 141–159 0.73 0.65 0.72 AB046750 Po31 (CA)4(GA)2(CA)11 57 69 25 129 –193 0.43 0.91 0.00* AB046751 Po33 (TG)5T2(TG)10 55 69 10 257– 290 0.74 0.68 0.82 AB046752 Po35 (CA)7 54 69 15 283– 333 0.81 0.78 1.00 AB046753 Po42 (CA)5(TA)13(CA)3 55 69 23 164– 224 0.88 0.91 0.67 AB046754 Po48 (CACG)4(CA)5 55 64 6 126 –142 0.44 0.43 0.69 AB046755 Po52 (CA)2CG(CA)6 GA(CA)5 58 64 4 155 –163 0.46 0.50 0.62 AB046756 Po56 (AC)20 55 69 26 139– 205 0.94 0.94 0.62 AB046757 Po58 (CA)11(GA)2GC(GA)9 52 69 27 101–159 0.84 0.90 0.52 AB046758 Po83 57 68 32 227– 313 0.91 0.93 0.18 AB046759 Po89 (CA)5AG(CG)2(TG)3 (CG)2(CA)15 TA3(CA)7 60 69 20 252– 327 0.86 0.90 0.44 AB046760 Po91 (CA)18 F-GCCTTTTGTCAGCCATTAACAGAGC R-CTGAGGCCAGACATGACATTACCTT F-CGGCCTAAACCTGGACATCCTCTCTA R-CGGGACAACGGAGGTTTGACTGAC F-TGCTCCTTCACCTGCACGGCCTCAAA R-TGCACCCTGACCTGTCACTGGGGATT F-TGAGGAGTCAGGTTTCAGGCCACT R-TCGCAGGAACACCCAGAGTACAGA F-ACACTGGGCCCTCTGTTAAACAC R-AGAGGAGAAAGGGCACCGAGATA F-AGGGTTAATTATAGAGGACGCAG R-CTGAAACAACAACTCAGAAGACG F-GTTGGTTTAACTGATTCATCTGCAG R-TTACATATCCCACAATGCTTCACTC F-TGGTTCTAGTGTTTGTCTGGTGA R-CCTACAGCACAGATATGACCTTT F-CGAGCGCTGTTTCAACTACGGTCATT R-ATGATGATCTAACCGTCCGGCTCCAT F-GCCTCCAGAAACATTTATGGGG R-TGTCTTGCCTCTGGTCCTTCTT F-TCAGACAGAGGAGCGGGGTTGTTGC R-GCTGTACCCAGGGTTCCGCTGAAGA F-TCGAGCGTAAACAAACCAGCTAACA R-GCTGAAAATCGCTTTAGCTTCCCAT F-GCCCCTCACTGAGACTGTGACA R-CAAGGTATGTGCATGAGCAGGC F-TGCGGTCATCATGTCTTTAAAATA R-AGCAAATGTTTGCTTTTGGATACA F-ATCAGAAGTCATCCATGCACTGGCAC R-AGCTACTTATCCACAGGTGTCGACGG F-AGGTTTCAAGGTGTTCATTGCGAGTC R-TAAAGGAAGTGCCTCACTGTGGAGAA 55 69 34 146– 246 0.96 0.94 0.97 AB046761 mean 20.1 — 0.76 0.80 — †Size is indicated as number of the base pairs of PCR products. ‡P is the exact P-value estimated by a test anologous to Fisher’s exact test described by Schneider et al. (1997). Significant departure of the observed genotype frequencies from H-W expectations was determined by adding *P < 0.05. §The nucleotide sequence data will appear in the DDJB/EMBL/GenBank nucleotide databases with the accession numbers. 2202 P R I M E R N O T E S Table 1 Core repeat and primer sequences, PCR amplification conditions, and results of variability of the 16 microsatellite loci in a Paralichthys olivaceus population. HO is observed and HE is expected heterozygosity P R I M E R N O T E S 2203 Polymorphic microsatellite loci for primitively eusocial Stenogastrine wasps Graphicraft 00 PRIMER 1156 2000 912 NOTEs Limited, Hong Kong YONG ZHU,* MONICA LANDI,* D AV I D C . Q U E L L E R , * S T E FA N O T U R I L L A Z Z I † and J O A N E . S T R A S S M A N N * *Department of Ecology and Evolutionary Biology, Rice University, PO Box 1892, Houston, TX 77251–1892, USA, †Department of Biologia Animale e Genetica, University of Firenze, Italy Keywords: cooperation, Eustenogaster, kin selection, microsatellite, sociality, wasp Received 6 July 2000; revision accepted 22 August 2000 Correspondence: Joan E. Strassmann. Fax: (713) 285 5232; E-mail: strassm@rice.edu Social wasps of the subfamily Stenogastrinae live in South-east Asia and comprise about 50 described species in six genera (Turillazzi 1996). Most species in this tropical subfamily of Vespidae have a colony with a small number of individuals and a simple temporal division of labour which makes them a suitable group for studying the origin of sociality in wasps (Turillazzi 1991; Strassmann et al. 1994). Microsatellite loci are useful tools for studying Stenogastrine population structure, relatedness within colonies and brood, and for determining males (Queller & Strassmann 1993). All of these are crucial factors for understanding Stenogastrine societies. In this paper, we describe 33 microsatellite loci isolated from Eustenogaster fraterna that are likely to be useful in this and many other species of the subfamily Stenogastrinae. We made a partical genomic library of E. fraterna following published protocols (Strassmann et al. 1996), but used a positive-selection plasmid (pZErO-2.1, Invitrogen) which eliminated the need for plasmid dephosphorylation. We cut genomic DNA with Sau3AI then ligated the fragments ranging from 300 – 900 bp into pZErO-2.1 plasmids. We transformed TOP10F′ cells to obtain a 15 000 clone library which was plated into nylon. Probing library replicates with oligoncleotides of all 10 trinucleotide motifs yielded 361 positives. Southern blots of plasmid DNA confirmed 121, 70 of which were sequenced. We designed polymerase chain reaction (PCR) primers for 31 clones containing five or more trinucleotide repeats, and two clones consisting of long dinucleotide repeats (Table 1). We evaluated these primers for heterozygosity on 24 individuals from six species following standard protocols (Strassmann et al. 1996). We extracted genomic DNA first and then performed PCR in a 10-µL volume with final concentrations of 50–200 ng genomic DNA, 250 nm of each primer, 100 µm of each dNTP, 50 mm KCl, 10 mm Tris-HCl pH 9.0, 0.1% Triton X-100, 1.55 mm MgCl2, 1.875 µCi 35S-dATP and 0.25 U Taq DNA polymerase under an oil overlay. We ran 40 cycles of 30 s at 92 °C denaturing, 30 s primer annealing (temperature varied depending on each primer and species) and 45 s extension at 72 °C, followed by 5 extra minutes at 72 °C to allow for the complete extension of all PCR fragments using a PTC-100 thermocycler (MJ Research). PCR products were run on 6% denaturing acrylamide gels, and an M13 sequence was used as a size standard. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Of the 33 primer pairs we designed, 29 yielded scorable microsatellite alleles. The number of alleles and expected heterozygosity of each variable locus in different species are detailed in Table 2. Out of 33 microsatellite loci, 27 were polymorphic in E. fraterna though heterozygosity varied from 0.20 – 0.81. Eighteen and 15 loci were polymorphic in E. calytodoma and Eustenogaster sp., respectively. Six loci were polymorphic in both Parischnogaster jacobsoni and P. alternata. Ten loci were polymorphic in Liostenogaster flavolineata. These results were congruent with the finding that polymorphisms of microsatellite loci decrease with increasing phylogenetic distance cross species (Ezenwa et al. 1998; Zhu et al. 2000). We analysed the relationship between repeat length and heterozygosity. Repeat length was represented by two measures: (i) the number of longest perfect, uninterrupted repeats; and (ii) total number of repeats including ones with base pair imperfections. We found significantly positive correlations between heterozygosity and repeat length (both perfect P = 0.038 and total P = 0.011, Spearman correlation). We had more than two microsatellite loci in E. fraterna for three repeat motifs, AAG, AAT and GAC. If we split the analysis for different motif types, we found no significant relationships between repeat length (either perfect or total) and heterozygosity for these three repeat motifs. Both correlations for the GAC motif were insignificant. Small sample size leads to a lack of power and might obscure any relationship between numbers of repeats and heterozygosity for these motifs. Acknowledgements We thank Rosli Hashim for permission to collect samples in Malaysia. We also thank Michael Henshaw for laboratory advice. This research was supported by NSF grants, IBN9975351 and IBN9808809. References Ezenwa VO, Peters JM, Zhu Y et al. (1998) Ancient conservation of trinucleotide microsatellite loci in Polistine wasps. Molecular Phylogenetics and Evolution, 10, 168–177. Queller DC, Strassmann JE (1993) Microsatellites and kinship. Trends in Ecology and Evolution, 8, 285–288. Strassmann JE, Hughes CR, Turillazzi S, Solis CR, Queller DC (1994) Genetic relatedness and incipient eusociality in stenogastrine wasps. Animal Behavior, 48, 813–821. Strassmann JE, Solis CR, Peters JM, Queller DC (1996) Strategies for finding and using highly polymorphic DNA microsatellite loci for studies of genetic relatedness and pedigrees. In: Molecular Methods in Zoology and Evolution (eds Ferraris J, Palumbi S), pp. 163 –178, 528 –549. Wiley, New York. Turillazzi S (1991) The Stenogastrinae. In: Social Biology of Wasps (eds Ross KG, Matthews RW), pp. 74 –98. Cornell University Press, Ithaca. Turillazzi S (1996) Polistes in perspective: comparative social biology and evolution in Belonogastrinae. In: Natural History and Evolution of Paper-wasps (eds By Turillazzi S, West-Eberhard), pp. 235–247, Oxford University Press, Oxford. Zhu Y, Queller DC, Strassmann JE (2000) A phylogenetic perspective on sequence evolution in microsatellite loci. Journal of Molecular Evolution, 50, 324–338. 2204 P R I M E R N O T E S Table 1 Characteristics of 28 microsatellite loci in social wasp Eustenogaster fraterna Locus Size (bp) Ta (°C) Repeat motif PO Primers sequences (5′−3′) Accession no. EF79TCT 176 51 (AAG)13 173 50 (AAG)15 EF91CTT 161 53 (AAG)7 EF92CTT 173 54 (AAG)9 EF97AAG 183 49 EF98AAG 246 57 (AAG)8AAT(AAG)2 AAT(AAG)2 (AAG)9 EF99AAG 153 51 (AAG)6 EF103AAG 258 53 (AAG)7 EF104AAG 125 52 (AAG)14 EF107AGA 217 55 (AAG)9 EF109AAG 178 53 (AAG)7AGA(AAG)5 EF131CAT 121 51 (CAT)4CGT(CAT)2 EF183AAG 124 49 (AAG)8GAG(AAG)4 EF184AAC 117 51 (AAC)12AAA(AAC)2 EF189TAA 206 48 (AAT)14 EF197TTC 141 50 (AAG)8 EF201TCT 175 50 (AAG)8 EF204TTC 187 54 (AAG)6ACG(AAG)2 EF211CTT 214 53 (AAG)7 EF213AAT 238 50 (AAT)10 EF217GA 237 52 (AG)4AA(AG)19 EF229AAG 175 52 (AAG)14TAG(AAG)3 EF238AAT 212 48 (AAT)12 EF280GCA 252 55 (CAG)3CAA(CAG)5 EF290CCT 121 54 (GAG)8AG(GAG)2 EF293CAG 197 55 (GAC)2AAC(GAC)6 EF299TGC 184 55 (GAC)9 EF318CAG 144 58 (GAC)15 TCGCTGTTCGACCATCG AATTCTTACCGCCGAATGG TGTCTTTCGCCTAACCG CCTCCTGCCTGTTTCTTG GACCGTTCCAACTGGCA CGACGTGTGAAATAAAGCAGGAG TCTACCGCCAACAGTCCCA CGAACGAGAAAAGTCCAAGCA GGGTTCCTTTATTAGTCCAAC TTCCTGGAGCATCCGTAAGC ATTGAGATGCAGAGAGCGTCGG AACAGGAGCACGGAGAAGAGGAAG CTGTCGTTCGTTTCGTTCTTCC AGTAGCGAGCAGATGATGATGATG TCCCTTCTCCTTCTCTTCGC CCTCCTTACTCCTTCTGGAC CGACCAGTGGCGTTTCA CCCTTACCGTTGAGACCCTG AAGCAAGGACGCACAGG ATCGACCGATGCACCGA CGCCTACAGAGTTCCTTG CGTCCTCGTTCATGGATTG TCATCTTCGTTGTCCTCG AAGCGGTTCTCTCGATG GCTCTTTGGGAATTTCTCG CGTTTCTCTTCGTCTTCG GCTCACATTTTTTCCCAGTCCC AATCTGCGTGCGTTGTTCTTG CGGATCTCGTAACGACTGATA GGAGCAAGTTGAAGGTACAA ACTCGGAAGCAACCTCG TGGAAAAGGCGGTAGAG GCGTGCCTCGAACATTA TGGAAAAGGCGGTAGAG GCGTTGTCCAGTCGTTTAACA TCGGCACGAAGACGATG AGGCTCTTCAGACGCTG TGGTGTAATCCGTGAGTGAG GCGATTTGAAGAAGCATTTAGTCG CAGGAAGTATATTAAGTGAAGCGTG GAAACTTTGCTCGCACACTG TCTATTCAGGGGAGGAAAAGC TGTAGGAAGAACGAAGGGTG GAGTGATTGATGGTCCGAGA GGATCACCGTGTAAAGACG CGATTTTTCTCGTTCGACGAAG CGCAGTCATCGCTTTTCA CCTAACCTACCCCAATCG TCTTGCGTTCGTTCGGA GCAGAGCGGAAAAAAGGG CGCAGTCATCGCTTTTCA GCATCGGCAACAGGAAA AGCTATCTCGGCTGTCG CTCCATCCATCCATCCA GTTTATCGCTCGTTGCTATCGG CTCCTATCCATCGCCCTTTCTC AF225508 EF80CTT F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R F R AF225495 AF225510 AF225511 AF225487 AF225488 AF225489 AF225479 AF225502 AF225497 AF225490 AF225480 AF225498 AF225481 AF225503 AF225504 AF225499 AF225505 AF225492 AF225483 AF225484 AF225506 AF225500 AF225493 AF225501 AF225494 AF225507 AF225486 Ta (°C), Annealing temperature; PO, primer orientation; F, forward primer; R, reverse primer. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2205 Table 2 Expected heterozygosities and number of alleles in different wasp species Expected heterozygosity (number of alleles) Locus/Species Eustenogaster fraterna E. calyptodoma E. sp. Parischnogaster jacobsoni P. alternata Liostenogaster flavolineata EF79TCT EF80CTT EF91CTT EF92CTT EF97AAG EF98AAG EF99AAG EF103AAG EF104AAG EF107AGA EF109AAG EF131CAT EF183AAG EF184AAC EF189TAA EF197TTC EF201TCT EF204TTC EF211CTT EF213AAT EF217GA EF229AAG EF238AAT EF280GCA EF290CCT EF293CAG EF299TGC EF318CAG Number of polymorphic loci 0.620 (3) 0.809 (7) 0.521 (3) 0.198 (2) 0.620 (3) 0.711 (5) 0.579 (3) 0.710 (5) 0.791 (6) 0.716 (5) 0.661 (3) 0.000 (1) 0.615 (3) 0.684 (5) 0.806 (6) 0.444 (2) 0.463 (2) 0.639 (2) 0.444 (2) 0.645 (3) 0.805 (7) 0.667 (4) 0.531 (3) 0.716 (4) 0.791 (6) 0.755 (6) 0.678 (4) 0.597 (4) 27 0.610 (3) 0.000 (1) 0.000 (1) 0.720 (4) 0.716 (4) 0.375 (2) 0.000 (1) 0.444 (2) 0.722 (4) 0.500 (2) 0.444 (2) 0.000 (1) + 0.480 (2) 0.625 (3) 0.000 (1) 0.375 (2) 0.375 (2) 0.000 (1) 0.000 (1) 0.833 (6) 0.444 (2) + 0.667 (4) + 0.722 (3) 0.500 (2) 0.667 (3) 18 0.560 (3) 0.000 (1) 0.000 (1) 0.480 (2) 0.625 (3) 0.480 (2) 0.000 (1) 0.444 (2) 0.560 (3) 0.560 (3) 0.444 (2) 0.000 (1) + 0.720 (4) 0.625 (3) 0.000 (1) 0.000 (1) 0.000 (1) + 0.000 (1) 0.500 (2) 0.000 (1) + 0.500 (2) + 0.500 (2) 0.500 (2) 0.625 (2) 15 + + 0.500 (2) 0.625 (3) + 0.000 (1) 0.630 (2) 0.000 (1) + + + 0.000 (1) + + + 0.000 (1) 0.445 (2) + + + + 0.000 (1) + 0.480 (2) + 0.000 (1) 0.480 (2) + 6 0.000 (1) + 0.500 (2) 0.000 (1) + 0.500 (2) 0.480 (2) 0.000 (1) + + 0. 000 (1) + + + + 0.000 (1) 0.000 (1) + + + + + + 0.500 (2) + 0.444 (2) 0.500 (2) + 6 0.000 (1) + 0.500 (2) 0.720 (4) 0.000 (1) + + + + 0.500 (2) 0.610 (3) 0.500 (2) + 0.611 (3) + 0.375 (2) 0.000 (1) 0.500 (2) + + + 0.000 (1) + 0.444 (2) + 0.000 (1) 0.500 (2) + 10 +, no PCR product. Microsatellites from the compact genome of the green spotted pufferfish (Tetraodon nigroviridis) Graphicraft 00 PRIMER 1158 2000 912 NOTEs Limited, Hong Kong G . H . Y U E , Y. L I , J . A . H I L L and L . O R B A N Laboratory of Fish Biotechnology, Institute of Molecular Agrobiology, 1 Research Link, NUS Campus, National University of Singapore, 117604 Singapore Keywords: fugu, genome programme, genomics, polymorphism, repeat Received 10 July 2000; revision accepted 26 August 2000 Correspondence: Laszlo Orban. Fax: +65 872 7007; E-mail: orban@ima.org.sg The green spotted pufferfish (Tetraodon nigroviridis) is an euryhaline species native to rivers and estuaries of South-East Asia (Kottelat et al. 1993). Beside the Japanese pufferfish (Fugu rubripes) (Brenner et al. 1993), T. nigroviridis is also becoming a model for cytogenetic and genomic studies (Grutzner et al. 1999; Roest Crollius et al. 2000a), because of its small genome (350 Mb). Green spotted pufferfish has not been bred in captivity and little is known about its biology, especially © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 reproduction. Most researchers rely on individuals collected from the wild, but very little is known about natural populations of the species. Microsatellites have been successfully applied to assess genetic diversity, population structure and individual relatedness in animals and plants. To our knowledge, no microsatellite has been characterized in T. nigroviridis. This paper describes the isolation and characterization of seven microsatellites from the genome of this pufferfish species. Thirty-two adult green spotted pufferfish were obtained from a local fish dealer in Singapore. Genomic DNA was extracted from muscle using a standard phenol–chloroform extraction protocol. A (CA)n-enriched plasmid library was constructed using DNA from one fish as described previously (Yue et al. 2000). More than 7500 clones were obtained, most of them (approximately 99%) were white on plates containing X-gal. The insert length of clones was checked by colony polymerase chain reaction (PCR) using M13-20 and M13-reverse primers. Out of 192 clones tested, 58 contained inserts in the size range of 250 –1000 bp. Colony PCR products of these clones were purified and sequenced as described (Yue et al. 2000). 2206 P R I M E R N O T E S Table 1 Characterization of six microsatellites on 32 individuals of the green spotted pufferfish ( Tetraodon nigroviridis) Locus Repeat motif Primer (5′− 3′) PF12 (CA)11 PF29 (GCA)13 PF39 (GT)8 PF41 (GA)2(CA)6GACTGAAG(CA)3 PF203 (GT)6(GA)7(GT)6(GA)8(GT)10 PF204 (GACA)10 F HEX-CAGGCCTGGACAAACAAAAC R ATCTTCAAAGTGGCGCTATCATT F HEX-TGAGCCGATCAAGTAGTGAG R GAATGATAGTGCTGCTGGGG F CTTGGATGTGACAGCGAAACAAAC R GCGCGTACGCACAGGCGGG F ACAAACACGGTCAACAAGCACTAC R ACAGGTGTTCTTTGGCGTGACA F TGGTGACCATTAGGGTAAGG R GGGGGTGAAACGACCTC F CTCGCCATGCAAAGAAAA R AAACGTTAAAGGTAGTGATGTGG Ta (°C) MgCl2 (mm) No. of alleles Size range (bp) HO HE GenBank Accession no. 55 1.5 22 176–240 0.95 0.91 AF283467 55 1.5 11 120–159 0.84 0.84 AF283468 60 1.5 5 158–180 0.59** 0.75 AF283470 55 1.5 17 162–198 0.59** 0.90 AF283471 45 3.0 12 220–258 0.22** 0.87 AF283472 50 1.5 9 130–154 0.72 0.78 AF283473 Ta, annealing temperature; HO, observed heterozygosity; HE, expected heterozygosity; **, Loci showing significant (P < 0.01) deviations from Hardy–Weinberg equilibrium by using chi-square analysis. HEX: Primer labelled with the fluorescent dye HEX. Out of the 58 clones sequenced, only 7 (~12%) contained microsatellite repeats, which is about 6 times lower than the percentage obtained from Asian arowana (Yue et al. 2000) by using the same enrichment procedure. This result might indicate low abundance of CA-repeats in the genome of T. nigroviridis. However, data from large scale sequencing performed on the T. nigroviridis genome do not support this hypothesis (Roest Crollius et al. 2000b). Alternative interpretation of the result might be that the CA-repeats in this compact genome are relatively short and are difficult to enrich by the method used. Among the seven microsatellites isolated, only five contained CA/GT repeats, the other two comprised GCA-repeats and GACA-repeats, respectively (Table 1). Primer pairs were designed to the flanking sequences of repeats using software PrimerSelect (DNASTAR). Thirty-two T. nigroviridis individuals were genotyped for the seven microsatellites as described previously (Yue et al. 2000), except using different annealing temperatures (in the range of 45 – 60 °C) and MgCl2 concentrations (1.5 or 3.0 mm; see Table 1). Six out of seven microsatellites showed specific products and polymorphism (Table 1), while locus PF33 was not polymorphic. The average number of alleles at the polymorphic loci was 12.7 (range: 5 – 22), whereas the average observed heterozygosity ranged from 0.22 – 0.95 with an average of 0.63 (Table 1). Three loci (PF12, PF29, PF204) conformed to Hardy–Weinberg expectations when tested using chi-square analysis, while the other three did not. A significant heterozygosity deficit was displayed at locus PF203, suggesting the appearance of null alleles. One duplex-PCR was established for the PF12 and PF29 loci according to Yue et al. (1999). The 25 µL reaction contained 10 mm Tris-HCl (pH 8.8), 150 mm KCl, 1.5 mm MgCl2, 100 µm of each dNTP, 0.2 µm each of PF12 primers, 0.4 µm each of PF29 primers, 30 ng genomic DNA and 1.0 U DyNAzyme II DNA-polymerase (Finnzymes). PCR cycling conditions were: an initial denaturation at 94 °C for 2 min, followed by 30 cycles of 94 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s, with a final extension at 72 °C for 5 min. The detection of PCR products and the sizing of alleles were performed on the ABI 377 sequencer as described previously (Yue et al. 2000). In order to become a good model not only for genomics, but also for genetics and developmental biology, T. nigroviridis must be bred routinely in captivity. The polymorphic microsatellite markers described here will assist the analysis of natural populations and breeding experiments. Acknowledgements We thank H. Roest Croellius for providing materials prior to publication. Funding was provided by the National Science and Technology Board of Singapore. References Brenner S, Elgar G, Sandford R, Macrae A, Venkatesh B, Aparicio S (1993) Characterization of the pufferfish (Fugu) genome as a compact model vertebrate genome. Nature, 366, 265–268. Grutzner F, Lutjens G, Rovira C, Barnes DW, Ropers HH, Haaf T (1999) Classical and molecular cytogenetics of the pufferfish Tetraodon nigroviridis. Chromosome Research, 7, 655–662. Kottelat M, Whitten AJ, Kartikasari SN, Wirjoatmodjo S (1993) Freshwater Fishes of Western Indonesia and Sulawesi. Periplus Editions (HK) Ltd, Indonesia, p. 242. Roest Crollius H, Jaillon O, Bernot A et al. (2000a) Estimate of human gene number provided by genome-wide analysis using Tetraodon nigroviridis DNA sequence. Nature Genetics, 25, 235 – 238. Roest Crollius H, Jaillon O, Dasilva C et al. (2000b) Characterisation and repeat analysis of the compact genome of the freshwater pufferfish Tetraodon nigroviridis. Genome Research, 10, 950–958. Yue GH, Beeckmann P, Bartenschlager H, Moser G, Geldermann H (1999) Rapid and precise genotyping of porcine microsatellites. Electrophoresis, 20, 3358–3363. Yue GH, Chen F, Orban L (2000) Rapid isolation and characterisation of microsatellites from the genome of Asian arowana (Scleropages formosus, Osteoglossidae, Pisces). Molecular Ecology, 9, 1007–1009. 2000 Graphicraft 1150 19PRIMER 32 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2207 Polymorphic DNA microsatellites identified in the yellow dung fly (Scathophaga stercoraria) T. W. J . G A R N E R ,* H . B R I N K M A N N ,† G . G E R L A C H ,† A . M E Y E R ,† P. I . WA R D ,‡ M . S P Ö R R I * and D . J . H O S K E N †‡ *Zoologisches Institut and ‡Zoological Museum, Universität Zürich-Irchel, Winterthurerstrasse 190, Zürich, Switzerland, †Faculty of Biology, Box 5560, University of Konstanz, D-78434 Konstanz, Germany Keywords: microsatellites, Scathophaga stercoraria, sperm competition, yellow dung fly Received 15 August 2000; revision accepted 2 September 2000 Correspondence: T. W. J. Garner. Fax: + 41 635 68 21; E-mail: twjg@zool.unizh.ch Sperm competition in yellow dung flies (Scathophaga stercoraria) has been extensively investigated since Parker’s (1970a) seminal work (e.g. Parker & Simmons 1991; Ward 1993; Hosken & Ward 2000; reviewed in Hosken 1999). These flies serve as a model system for understanding the mechanisms and outcomes of sperm competition in internal fertilizers. Invariably however, these investigations have been laboratory based, and typically involved competition between only two males. How the results of such studies relates to freeliving flies is unknown, but it is unlikely that the experimental conditions employed exist in nature, and therefore outcomes may not reflect true female sperm utilization patterns (Eady & Tubman 1996). This is exemplified by a study of sperm competition in pseudoscorpions, which showed that secondmale mating advantage breaks down when females mate with more than two males (Zeh & Zeh 1994). In addition, Ward (2000) has shown that females are able to subtly alter paternity patterns under conditions that are likely to be common in the field. With this in mind, our aim was to develop appropriate genetic markers to allow paternity to be accurately assigned in clutches laid by free-living female yellow dung flies. A subgenomic library enriched for CA repeat microsatellites was constructed following standard protocols outlined in Tenzer et al. (1999), with slight modifications. Genomic DNA isolated from a single S. stercoraria male using standard phenol– chloroform extraction and ethanol precipitation (Sambrook et al. 1989) was digested using Tsp509I (New England Biolabs). A 500 –1000 bp size fraction was isolated from a LM-MP agarose (Boehringer Mannheim) gel by first excising the appropriate size range from the gel. The gel fragment was melted in a 65 °C water bath and volume was increased to 500 µL using double distilled water. An equal volume of equilibrated phenol (pH 8.0) was added, the solution vortexed briefly and then put at – 80 °C for 30 min. The sample was then thawed and extraction was completed following standard phenol – chloroform extraction methods (Sambrook et al. 1989). This isolate was used for ligation with TSPADSHORT/ TSPADLONG linkers (Tenzer et al. 1999) and then amplified via the polymerase chain reaction (PCR), using TSPADSHORT as a primer. PCR was performed using the following conditions: Total reaction volume was 25 µL included 100 ng DNA, © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 1 U Taq DNA polymerase (Quantum-Appligene), 10 mm Tris-HCl, pH 9.0, 50 mm KCl, 1.5 mm MgCl2, 0.01% TritonX100, 0.2 mg BSA (Quantum-Appligene), 100 µm of each dNTP (Promega), and 1 µm of TSPADSHORT. PCR was performed on a Techne Genius thermocycler (Techne Ltd) using the following thermotreatment: 2 min at 72 °C, followed by 25 cycles of 1 min at 94 °C, 1 min at 55 °C, and 1 min at 72 °C. A total of 32 PCRs were carried out, pooled, cleaned and concentrated to minimize the likelihood of redundant products being detected during screening for positive clones. PCR products were hybridized to biotinylated (CA)20 probes bonded to streptavidin-coated magnetic beads (Dynabeads M-280 Streptavadin, DYNAL, France) and amplified again. These final PCR products were cloned following the Original TA Cloning® Kit (Invitrogen) protocol. White colonies were dot-blotted onto nylon membranes (Hybond™-N +, Amersham Pharmacia) and screened for CA repeats using the ECL 3′-oligolabelling and detection system (Amersham Pharmacia) and a 40mer CA oligonucleotide. All positive clones were sequenced following the ABI Prism® BigDye™ Terminator Cycle Sequencing Ready Reaction Kit protocol, version 2.0 (PE Biosystems) using M13 forward and reverse primers, and using the ABI 377 automated sequencing system (PE Biosystems). Primers were designed using Primer3 software (Rozen & Skaletsky 1998) and all oligonucleotides were synthesized by Microsynth GmbH (Switzerland). Initial tests for amplification and polymorphism were carried out at 55 °C and electrophoresed on 8%, nondenaturing, 14.5 cm × 17 cm acrylamide gels at 80 V overnight. Those primers that amplified polymorphic products using five test templates were used for all following analyses. Only field-caught male S. stercoraria were used for PCR analysis, as almost every field-caught female is already mated (Parker 1970b), and extraction from fertilized females could therefore result in contamination by sperm DNA. Each sample male was extracted using the QIAamp® DNA mini kit (Qiagen). Twenty males were used to characterize suitable primers, and PCR was carried out using approximately 100 ng of template DNA and the following cycle treatment; initial step of 3 min at 94 °C, followed by 27 cycles of 30 s at 94 °C, 30 s at 58– 61 °C (see Table 1), and 30 s at 72 °C, with a final extension step of 2 min at 72 °C. Total reaction volume was 25 µL and contained 10 mm Tris-HCl, pH 9.0, 50 mm KCl, 1.5 mm MgCl2, 0.01% Triton × 100, 0.2 mg BSA (QuantumAppligene), 100 µm of each dNTP (Promega), 0.5 µm of both forward and reverse primer, and 0.5 U Taq DNA polymerase (Quantum-Appligene). All products were electrophoresed on Spreadex™ EL-300 S-100 gels (Elchrom Scientific AG, Switzerland), using the SEA 2000™ advanced submerged gel electrophoresis apparatus (Elchrom Scientific AG, Switzerland). Gels were run at 100 V for 80–90 min, depending on allele sizes, then scored against the M3 Marker ladder (Elchrom Scientific AG, Switzerland). Expected and observed counts for homozygotes/heterozygotes were determined using genepop version 3x (Raymond & Rousset 1995) and homozygote excess was tested for using Chi-square analysis (null hypothesis rejected at P < 0.05). A minimum of five alleles were detected at each of the loci listed in Table 1. Tests for homozygote excess were only 2208 P R I M E R N O T E S Table 1 Primer sequence and related information for eight microsatellite loci developed for Scathophaga stercoraria. Both repeat motif and size of amplification product are based on that detected in the original sequenced clone (GenBank Accession nos: AF292121– 8). n, number of individuals tested; Ta, annealing temperature; HO, observed number of homozygotes; HE, unbiased average heterozygosity estimate (Nei 1978) Locus Primer Sequences (5′−3′) Repeat motif Ta (°C) n No. alleles Size (bp) HO HE SsCA3 CCTCAACCCCCTCACTCAC CATCATCATTTAAGTCAACATTAGAAA GACTTTGGTCCGTTGTAGTCC TTGGCGTCACCATACTCAAC AATAAAAACTCAACCAACCATACAC CCTTACTCGATAAGTTGGTATTTGTG TGTTTGCTGGTGCTACCG TGATCGTTGTTGTTTCATACG CACACACTCGCAGCTACACC AAACTTTAACTTCGATTTTTGCTG TGCCACTTTTGGTGCTTTC CAGCAAAAACCGGCAAAC GTTTGAAACCCTTAAGATAAAAACTC CCATCTTTCACGGGATTTTG AAAGAATTTTACGAATTGTGTCTGG CAACAAATGCAACAAATGACC (AC)1(A)2(AC)11 (A)3(C)2(A)3 (C)3AT(AC)11AT (AC)2(C)3 (TA)2GA(CA)4CG (CA)5 (CA)10 60 20 11 120 0.35 0.795 60 20 7 101 0.10 0.806 60 18 6 108 0.40 0.695 60 18 5 120 0.55 0.600 (C)4AT(AC)9 60 20 8 120 0.30 0.821 (CA)11(T)2(CA)2CG(CA)4CG (CA)4(T)2(CA)2(T)2(GTT)2 (CT)2(CA)5AACG (CA)10 (CA)6(A)2(CA)8 61 20 8 110 0.25 0.845 58 20 13 127 0.35 0.890 58 18 8 129 0.20 0.869 SsCA16 SsCA17 SsCA20 SsCA24 SsCA26 SsCA28 Ss63T7 significant at one locus, SsCa28, which may suggest one or more null alleles operating at this locus. Acknowledgements We would like to thank Tony Wilson and Jens Seckinger for their help. This work was funded in part by an Alexander Von Humboldt Stiftung awarded to DJH, and grants from the Swiss National Foundation (SNF 31–56902.99 to DJH, SNF 31– 46861.96 to PIW, SNF 31–40688.94 to H-U Reyer). AM would also like to thank the Deutsche Forschungsgemeinschaft and AM and GG acknowledge the contribution of the Verband der Chemischen Industrie. References Eady P, Tubman S (1996) Last-male sperm precedence does not break down when females mate with three males. Ecological Entomology, 21, 303–304. Hosken DJ (1999) Sperm displacement in yellow dung flies: a role for females. Trends in Ecology and Evolution, 14, 251– 252. Hosken DJ, Ward PI (2000) Copula in yellow dung flies (Scathophaga stercoraria): investigating sperm competition models by histological observation. Journal of Insect Physiology, 46, 1355 –1363. Nei M (1978) Estimation of average heterozygosity and genetic distance from a small number of individuals. Genetics, 89, 583– 590. Parker GA (1970a) Sperm competition and its evolutionary consequences in insects. Biological Reviews, 45, 525–567. Parker GA (1970b) Sperm competition and its evolutionary effect on copula duration in the fly, Scatophaga stercoraria. Journal of Insect Physiology, 16, 1301–1328. Parker GA, Simmons LW (1991) A model of constant random sperm displacement during mating: evidence from Scatophaga. Proceedings of the Royal Society of London, Series B, 246, 157–166. Raymond M, Rousset F (1995) genepop (Version 1.2): Population genetics software for exact tests and ecumenism. Journal of Heredity, 86, 248–249. Rozen S, Skaletsky HJ (1998) Primer3. Code available at http://www-genome.wi.mit.edu/genome_software/other/ primer3.html. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, New York. Tenzer I, degli Ivanissevich S, Morgante M, Gessler C (1999) Identification of microsatellite markers and their application to population genetics of Venturia inaequalis. Phytopathology, 89, 748–753. Ward PI (1993) Females influence sperm storage and use in the yellow dung fly Scathophaga stercoraria (L.). Behavioral Ecology and Sociobiology, 32, 313–319. Ward PI (2000) Cryptic female choice in the yellow dung fly Scathophaga stercoraria (L.). Evolution, in press. Zeh JA, Zeh DW (1994) Last-male sperm precedence breaks down when females mate with three males. Proceedings of the Royal Society of London, Series B, 257, 287–292. Graphicraft 2000 1151 00 91PRIMER 2 NOTEs Limited, Hong Kong Polymorphic microsatellite loci in vespertilionid bats isolated from the noctule bat Nyctalus noctula F. MAYER,* C. SCHLÖTTERER† and D. TAUTZ‡ *Institut für Zoologie II, Universität Erlangen-Nürnberg, Staudtstraße 5, D-91058 Erlangen, Germany, †Institut für Tierzucht und Genetik, Veterinärmedizinische Universität Wien, Josef-Baumann-Gasse 1, A-1210 Wien, Austria, ‡Institut für Genetik, Universität zu Köln, Weyertal 121, D-50931 Köln, Germany © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2209 Keywords: bat, Chiroptera, cross-species amplification, microsatellite, Nyctalus noctula Received 15 August 2000; revision accepted 2 September 2000 Correspondence: F. Mayer. Fax: + 49 91318528060; E-mail: fmayer@biologie.uni-erlangen.de Prolonged sperm storage increases the possibility of sperm competition because several males could contribute sperm during the sperm-storing period prior to fertilization. The longest sperm storing capacity among mammals is documented in the noctule bat. Females can store sperm in the uterus after copulation in autumn for up to six months until fertilization in spring (Racey 1973). Therefore, noctule bats are likely candidates to show high levels of multiple paternity that could be analysed most efficiently with highly polymorphic nuclear markers, such as microsatellite loci. Microsatellite loci of the noctule bat were isolated from a size-selected partial genomic library (Rassmann et al. 1991). Total genomic DNA was isolated from muscle tissue of a female bat and was digested with three restriction enzymes (AluI, HaeIII and RsaI). Fragments ranging from 300 – 600 bp in length were ligated in SmaI digested M13mp18 and M13mp19 cloning vectors (Yanisch-Perron et al. 1985). Ligation products were transformed into competent XL-1 Blue cells (Stratagene) which were plated onto LB pates containing X-gal and IPTG. A total of 5400 clones were screened with different probes specific for microsatellites. Radioactive 32P-labelled probes for di- and trinucleotide micosatellite loci were generated by slippage synthesis (Schlötterer & Tautz 1992) using the following pairs of oligonucleotides: (AG) 7/(TC)4, (GT)7/(CA)4, (TCC)5T/(GGA)3, (CCA)5/(GGT)3, (TGC)5/ (GCA)3 and (TCG)5T/(ACG)3. One hundred and forty-five ‘positive’ clones were detected with dinucleotide polymers, 54 with trinucleotide polymers, 11 with the 32P-end-labelled oligonucleotide (ATCC)3 and 14 with the 32P-end-labelled oligonucleotide (CTAT)5 representing 2.7, 1.0, 0.2 and 0.3%, respectively, of the total number of clones which were screened. Thirty-one of the 224 ‘positive’ clones were sequenced and polymerase chain reaction (PCR) primers were designed for 14 loci. A circular wing clip of 4 mm diameter was obtained from individual bats. Approximately 0.5 µg DNA was isolated after a 3-h incubation with 0.1 mg proteinase K in 500 µL digestion buffer (100 mm Tris-HCl, 100 mm NaCl, 2 mm EDTA, 42 mm dithiotreitol, 2% sodium dodecyl sulphate) following the protocol of Müllenbach et al. (1989). PCR amplifications were carried out in a 10-µL volume containing approximately 10 ng DNA, 1.5 mm MgCl2, 0.5 µm each primer, 0.025 mm each dNTP, 0.25 unit Goldstar Polymerase (Eurogentec) together with the reaction buffer provided by the supplier [final concentration: 75 mm Tris-HCl, pH 9.0, 20 mm (NH4)2SO4, 0.01% (w/v) Tween 20]. PCRs were performed in a Perkin Elmer DNA Thermal Cycler TC1 and consisted of 30 cycles of 94 °C for 30 s, the annealing temperature (Table 1) for 20 s and 72 °C for 30 s. For each microsatellite locus one primer was labelled with a fluorescent dye. The PCR products were separated on 6% Sequagel®XR gels (National Diagnostics) in a LI-COR DNA sequencer (model 4000 L), and genotypes were determined using RFLPscan™ (Scanalytics). Highly specific amplification products were obtained at 13 of the 14 loci. For each animal only one or two major amplification products were detected which were close to the length of the cloned allele. Allele sizes varied in multiples of the repeat size only and characteristic slippage bands could be detected at all loci. At two loci the proportion of homozygotes was much higher than expected under Hardy–Weinberg equilibrium. At locus P22a only one allele could be amplified in all males. In females no deviation from Hardy–Weinberg expectation was detected. This strongly suggests that locus P22a is located on the X chromosome. The deficit of heterozygotes at the other locus was not limited to males. Therefore, the presence of null alleles is a likely explanation. This locus was not evaluated further. Cross-species amplification was tested in 11 European bat species of the family Vespertilionidae. Nine loci could be amplified in other species and eight in other genera than the source species. Allelic variation was usually high and only in five cases the loci seemed to be monomorphic in a particular species (Table 2). The applicability of primer pairs is high among species within the family Vespertilionidae but seems to be low in other families (Burland et al. 1998). Length of amplification products can vary substantially among species. For example the amplification product of locus P217 was about 400 bp longer in the three species of Eptesicus and Plecotus than in all other species. Acknowledgements This work was supported by the Deutsche Forschungsgemeinschaft (DFG). References Burland TM, Barratt EM, Racey PA (1998) Isolation and characterization of microsatellite loci in the brown long-eared bat, Plecotus auritus, and cross-species amplification within the family Vespertilionidae. Molecular Ecology, 7, 136–138. Helversen Ov, Mayer F, Kock D (2000) Comments on the proposed designation of single neotypes for Vespertilio pipistrellus Schreber, 1774 (Mammalia, Chiroptera) and for Vespertilio pygmaeus Leach, 1825. Bulletin of Zoological Nomenclature, 57, 113–115. Müllenbach R, Lagoda PJL, Welter C (1989) An efficient saltchlorophorm extraction of DNA from blood and tissues. Trends in Genetics, 5, 391. Petri B, Pääbo S, Haeseler Av et al. (1997) Paternity assessment and population subdivision in a natural population of the larger mouse-eared bat Myotis myotis. Molecular Ecology, 6, 235–242. Racey PA (1973) The viability of spermatozoa after prolonged storage by male and female European bats. Periodicum Biologorum, 75, 201–205. Rassmann K, Schlötterer C, Tautz D (1991) Isolation of simplesequence loci for use in polymerase chain reaction-based DNA fingerprinting. Electrophoresis, 12, 113–118. Schlötterer C, Tautz D (1992) Slippage synthesis of simple sequence DNA. Nucleic Acids Research, 20, 211– 215. Yanisch-Perron C, Viera J, Messing J (1985) Improved M13 phage cloning vectors and host strains: nucleotide sequence of M13 mp18 and pUC19 vectors. Gene, 33, 103–119. Graphicraft 2000 1153 00 91PRIMER 2 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Annealing temperature (°C) Amplification of the cloned allele length (bp) Locus Primer Sequence N. noctula other species P2 Mü418 Mü419* ER22* ER23 Mü360* Mü361 Mü397* Mü398 ER28* ER29 ER47* Mü435 ER37* ER25 ER49* ER6 5′-ATATACTTAAGGATCAGAGC-3 5′-TATTGTTCTGTTCATTCAGT-3′ 5′-AAAACCAAAGTTATTTATTC-3′ 5′-CTTTCCTCAGAAATTATATC-3′ 5′-CCTGATAAAACCTGT-3′ 5′-CTGAATCGGTGTTTC-3′ 5′-TGGTGATTTGTTATG-3′ 5′-CACTTATCATTTTCA-3′ 5′-TCTAATCTCTTTCTGCACCC-3′ 5′-GGGGCATGGAAATTGAACAG-3′ 5′-CTTATCTAATCAATATACTTAAAA-3′ 5′-AAAATGCATCAATATATGAG-3′ 5′-CTTCTCCCTTCCCATAAATC-3′ 5′-TCTTATTTTGGGGGAAACTG-3′ 5′-TCCTAAGATTCTGTTCCTCC-3′ 5′-GGGCTGTATCATATGATTTT-3′ 48 48 98 52 — 52 ER36* ER5 ER1* ER2 5′-CAATTTAACTTTTCAACAAC-3′ 5′-TCTTCATTTCCTCTCCTCTC-3′ 5′-TCCATTTTTTCCCCTTCCCT-3′ 5′-GGTCTCCTTTTCTTCACTTTG-3′ P11 P13 P14 P19 © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 P20 P22a P217 P219 P223 microsatellite sequence GenBank accession no. 132 (AT)2(GT)2(AT)(GT)2 (AT)(GT)3(AT)2(GT)12 (GT)15 AF141645 AF273675 52 140 (TG)4C(GT)19 AF141646 40 — 112 (TG)17 AF273676 53 — 114 (AC)19 AF273677 45 40 176 (TA)21(TG)17TAT(TA)6 AF141647 48 40 113 (AT)5(AC)4AT(AC)10AT(AC)13 AF273678 48 48 251 AF141648 48 48 157 (CTAT)2CAT(CTAT)11 (CATCTAT)4(CTAT)2 (CATCTAT)2TAT(CTAT)3 (CAT)2CTAT (CTAT)7(CCAT)6 48 48 110 (CT)7CCCTC(CTAT)9 AF141650 AF141649 2210 P R I M E R N O T E S Table 1 Primer, annealing temperature and sequence of 12 microsatellite loci isolated from the noctule bat (Nyctalus noctula). Primers labelled at the 5′-end with the fluorescent dye IRD-41 or IRD-800 are marked with an asterisk © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Table 2 Polymorphism of microsatellite loci in different vespertilionid bat species. The source species was Nyctalus noctula. The loci NN8 and NN18 were previously published by Petri et al. (1997). No amplification is marked by a dash. Locus P22a is located on the X-chromosome. Therefore, the observed heterozygosity corresponds only to females at this locus. Crossspecies amplification of the locus P14 was not tested, due to weak signals in N. noctula. Heterozygosities are given if more than 10 individuals were scored within a species. Pipistrellus mediterraneus corresponds to the 55 kHz phonic type of the pipistrelle bat (Helversen et al. 2000) Species Parameter Nyctalus noctula Nyctalus leisleri Pipistrellus pipistrellus Pip. mediter. Pip. kuhli P2 number of individuals alleles detected observed heterozygosity expected heterozygosity 43 10 0.74 0.83 10 9 1.00 0.83 37 11 0.84 0.89 16 13 0.75 0.86 5 7 NN8 number of individuals alleles detected observed heterozygosity expected heterozygosity 43 12 0.72 0.79 10 12 0.70 0.87 55 30 0.76 0.93 P11 number of individuals alleles detected observed heterozygosity expected heterozygosity 41 11 0.90 0.84 — — P13 number of individuals alleles detected observed heterozygosity expected heterozygosity 42 15 0.95 0.91 5 10 P14 number of individuals alleles detected observed heterozygosity expected heterozygosity 40 9 0.92 0.86 NN18 number of individuals alleles detected observed heterozygosity expected heterozygosity P19 number of individuals alleles detected observed heterozygosity expected heterozygosity Pip. nathusii 4 4 Eptesicus nilssoni Vespertilio murinus 6 3 5 6 Plecotus austriacus Plecotus auritus Myotis myotis Myotis bechsteini — — — — — — 16 14 0.88 0.88 57 10 0.89 0.85 5 5 3 4 5 6 3 4 1 2 5 4 5 6 5 9 9 7 — — — — — — — — — — — — — — — — — — — — 25 14 0.92 0.83 20 11 0.95 0.85 3 4 2 3 5 1 5 4 10 11 0.80 0.87 16 13 0.69 0.89 — — — — — — ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? 30 8 0.83 0.78 10 9 0.90 0.82 53 12 0.83 0.85 21 9 0.67 0.79 5 1 37 16 0.86 0.90 — — — — — — — — ? ? ? ? ? ? ? ? ? ? ? ? 6 6 8 7 6 8 4 6 — — — — — — — — 14 10 0.93 0.81 — — 5 1 — — — — — — P R I M E R N O T E S 2211 Myotis 2212 P R I M E R N O T E S Table 2 Continued Species Myotis Parameter Nyctalus noctula P20 number of individuals alleles detected observed heterozygosity expected heterozygosity 36 14 0.83 0.92 P22a number females + males alleles detected observed heterozygosity expected heterozygosity P217 Nyctalus leisleri © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Pipistrellus pipistrellus Pip. mediter. Pip. kuhli 5 8 32 3 0.53 0.50 23 3 0.04 0.04 2 2 22 + 17 14 0.67 0.86 1+4 3 — — — — number of individuals alleles detected observed heterozygosity expected heterozygosity 34 18 0.88 0.90 5 8 45 12 0.89 0.81 P219 number of individuals alleles detected observed heterozygosity expected heterozygosity 36 7 0.72 0.75 — — P223 number of individuals alleles detected observed heterozygosity expected heterozygosity 38 13 0.76 0.77 5 5 Pip. nathusii Eptesicus nilssoni Vespertilio murinus Plecotus austriacus Plecotus auritus Myotis myotis Myotis bechsteini 3 6 5 1 5 5 3 4 15 4 0.67 0.56 32 18 0.94 0.92 9 7 — — — — — — — — — — — — — — — — 15 10 0.93 0.77 5 8 5 10 2 4 5 6 5 8 13 11 0.85 0.84 — — — — 42 14 0.76 0.81 14 8 0.79 0.80 4 3 — — 6 4 4 6 5 1 — — — — — — — — — — — — — — — — — — — — — — — — — — P R I M E R N O T E S 2213 Isolation and characterization of microsatellites in the seabird ectoparasite Ixodes uriae to amplify DNA using primers developed for I. ricinus, a common tick species of vertebrates in Europe (Delaye et al. 1998); no successful amplifications were achieved. In this note, we characterize nine microsatellite markers developed for I. uriae. DNA was extracted from 42 unfed larval ticks originating from Atlantic puffin (Fratercula arctica) hosts on Hornøya, Norway (70°22′N, 31°10′ W). A genomic library was constructed following Estoup et al. (1993). Eleven µg of larval DNA was restricted with the enzyme Sau3A. Resulting fragments were separated on a 1.5% low-melting-point agarose gel and fragments between 400 – 800 bp were isolated, purified using a Q1Aquick Gel Extraction Kit (Qiagen) and ligated into a pBluescript vector II Sk + plasmid (Stratagene). Ligation products were then transformed into XL1-Blue MRF′ Supercompetent cells (Stratagene) and the resulting colonies were blotted on Hybond-N + membranes which were hybridized with a mixture of two probes (CT)10 and (GT)10. Two thousand clones from the library gave 65 positively hybridized clones from which 48 were sequenced. Primers were designed for 10 loci using Primer 0.5 (Lincoln & Daly 1991). Nine loci were polymorphic and gave clear polymerase chain reaction (PCR) results of expected size (Table 1). Genomic DNA was prepared using a high-salt extraction method. PCR amplifications were performed in a 10-µL mixture containing 1 µL of genomic DNA (approximately 50 ng), 75 µm of each of dCTP, dTTP, dGTP, 7.5 µm of dATP, 0.4 µm of each primer, 1 µL of 10× Taq buffer (Tris-Cl, KCl (NH4)2SO4, 15 mm MgCl2, pH 8.7), 0.25 U Taq DNA polymerase (Qiagen) and 0.025 µCi [α33P]-dATP (Amersham). Amplifications were performed in a PTC100 thermocycler (MJ Research) as follows: initial denaturation of 3 min at 94 °C followed by 30 cycles (30 s at 94 °C, 30 s at annealing temperature specified in Table 1 and K A R E N D . M C C O Y and C L A I R E T I R A R D Laboratoire d’Ecologie, Université Paris VI — CNRS UMR 7625, 7 quai St. Bernard, 75005 Paris France Keywords: ectoparasite, Ixodidae, microsatellite, tick Received 10 August 2000; revision accepted 2 September 2000 Correspondence: Karen D. McCoy, Fax: + 33 1 44 27 35 16; E-mail: kmccoy@snv.jussieu.fr Ixodes uriae is a common parasite of many seabird species in the polar regions. Interest in this tick has risen in the past decade as more information is gathered on its potential impacts on host ecology and evolution. In particular, I. uriae is thought to affect the reproductive success and habitat choice of its seabird hosts (Boulinier et al. 2001), and to vector several avian arbo-viruses and disease agents (Chastel 1988), including the Lyme disease agent, Borrelia burgdorferi (Olsen et al. 1993). Thus, knowledge of dispersal ability of this ectoparasite within and among host populations is of vital importance if we are to understand its role in this hostparasite–disease interaction. The independent migratory abilities of hard ticks are considered to be weak, and direct examination of dispersal of these parasites at large spatial scales is not possible (McCoy et al. 1999). Thus, to examine patterns of dispersion, indirect approaches, such as using genetic markers, are more plausible. To address the question of population structure and gene flow of I. uriae within and among its seabird hosts, we first attempted Table 1 Characteristics of nine polymorphic loci developed for Ixodes uriae. The number of observed alleles (NA), observed (HO ) and expected (HE) heterozygosities were calculated using ticks (n refers to number of ticks) sampled from two Atlantic puffin (Fratercula arctica) colonies. Hardy–Weinberg equilibrium was tested separately for each colony (Raymond & Rousset 1995); no significant deviations were found after correcting for multiple tests (Rice 1989) Locus Repeat array n Size (bp) NA HO HE Ta * (°C) Accession no.† Primer sequence (5′− 3′) T1 63 158–164 7 0.63 0.70 57 AF293324 T3 (GA)3TA(GA)2-(GA)2(GA)4-(GA)2CA(GA)3 (CA)4AA(CA)3-(CA)7 64 112–114 2 0.05 0.08 57 AF293325 T5 (GA)15 63 180–244 12 0.73 0.86 55 AF293326 T22 (GA)7-(GA)13 64 157–187 11 0.83 0.83 57 AF293327 T35 (CT)12 60 142–162 10 0.80 0.79 57 AF293328 T38 (TC)13 64 155–167 6 0.44 0.55 57 AF293329 T39 (CA)31 63 188–255 26 0.89 0.94 55 AF293330 T44 (GT)4AG(GT)7AG(GT)7 63 153–185 6 0.32 0.33 57 AF293331 T47 (GT)5CT(GT)7 62 152–158 3 0.37 0.48 57 AF293332 F: CTTCAATCACGTGGGATGC R: GACTTGTGCCTCTCCCAAAG F: GCATTAGCGTCATAACATGAAC R: CTCTGTTTACCCTCTTCTTTGC F: AATTGGAAAGTAGCCATTCG R: ACTCTAATGCAACGGCGTATG F: CAGACGCCGACAAATTATCC R: GACGTTTGTTTGGTGCTGTG F: CTCCTTTCACTCGCTTGTCC R: TCCTTCAAGCGTGTATCCAG F: GCATAACCAGATTCCTCCTTTC R: CAAGTGAAAGAAAACGGTGAC F: AACCGCAATATTAGGTCAGC R: GTTTTGGTTTCGCTTGTTTAG F: CATAACCCGACTGTCTCACTG R: GAACCACACCCAGACAACG F: GAAACGCAATGACGTACAGG R: TAATAACGCCGCACAAGGAG *Ta, annealing temperature. †Accession no. of the sequences available from GenBank. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 2214 P R I M E R N O T E S 1 min at 72 °C) and a final elongation step of 10 min at 72 °C. PCR products were denatured and separated on 6% polyacrylamide and 8 m urea sequencing gels using a M13 sequence as a size marker. To characterize each locus, we genotyped ticks originating from two Atlantic Puffin colonies: Baccalieu Island, Newfoundland, Canada (48°08′ N, 52°48′ W) and Hornøya, Norway (Table 1). Expected heterozygosities were variable, ranging from 0.08 – 0.94 with an average of 0.61 (± 0.09). Neither tick population showed any significant deviation from Hardy–Weinberg equilibrium after correcting for multiple tests (Rice 1989). Cross-species amplification of primers was tested on I. ricinus. PCR amplifications were attempted for all nine loci using 10 ticks originating from Bern, Switzerland; PCR protocols were identical except that the annealing temperature used for all primers was 52 °C. No successful amplifications were achieved for any locus. In conclusion, based on their high polymorphism, the microsatellite markers developed for I. uriae should enable the examination of a diverse range of questions related to parasite dispersal among hosts over a range of spatial scales, from within colonies to between hemispheres. Likewise, patterns of parasite gene flow may provide insight into the large-scale movement of their seabird hosts. Such data will prove valuable for examining questions related to the evolution of local adaptation in this host-parasite system and for examining the epidemiology of tick-borne disease. Acknowledgements Many thanks to T. Boulinier, Y. Michalakis, E. Danchin, T. De Meeûs and F. Renaud for advice and support, and to T. De Meeûs for providing samples of Ixodes ricinus. This work was supported by the CNRS, Programme Environnement, Vie et Sociétés and the Institute Français Pour la Recherche et la Technologie Polaires (France), and by a Natural Sciences and Engineering Research Council of Canada Postgraduate Scholarship (Canada) to KM. References Boulinier T, McCoy KD, Sorci G (2001) Parasites and dispersal. In: Dispersal: Individual, Population and Community (eds Clobert J, Danchin E, Dhondt A, Nichols J), Oxford University Press, Oxford (in press). Chastel C (1988) Tick-borne virus infections of marine birds. In: Advances in Disease Vector Research (ed. Harris HK), pp. 25 –60. Springer-Verlag, New York. Delaye C, Aeschlimann A, Renaud F, Rosenthal B, De Meeûs T (1998) Isolation and characterization of microsatellite markers in the Ixodes ricinus complex (Acari: ixodidae). Molecular Ecology, 7, 360– 361. Estoup A, Solignac M, Harry M, Cornuet JM (1993) Characterisation of (GT)n and (CT)n microsatellites in two insect species: Apis mellifera and Bombus terrestris. Nucleic Acids Research, 21, 1427–1431. Lincoln S, Daly M (1991) Primer, Version 0.5. Whitehead Institute for Biomedical Research, Cambridge, MA. McCoy KD, Boulinier T, Chardine JW, Danchin E, Michalakis Y (1999) Dispersal and distribution of the tick Ixodes uriae within and among seabird host populations: the need for a population genetic approach. Journal of Parasitology, 85, 196–202. Olsen B, Jaenson TGT, Noppa N, Bunikis J, Bergstrom S (1993) A Lyme borreliosis cycle in seabirds and Ixodes uriae ticks. Nature, 362, 340–342. Raymond M, Rousset F (1995) genepop (v.1.2): population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. Rice WR (1989) Analyzing tables of statistical tests. Evolution, 43, 223–225. Graphicraft 2000 1154 00 91PRIMER 2 NOTEs Limited, Hong Kong Characterization of microsatellite loci in King George Whiting Sillaginodes punctata Cuvier and Valenciennes (Percoidei: Sillaginidae) L . H A I G H * and S . C . D O N N E L L A N † *South Australian Research and Development Institute, 2 Hamra Avenue, West Beach 5024, Australia †Evolutionary Biology Unit, South Australian Museum, Adelaide, 5000, Australia Keywords: fish, microsatellites, Sillaginidae, Sillaginodes, teleost Received 10 August 2000; revision accepted 2 September 2000 Correspondence: S. C. Donnellan. Fax: +61-8-82077222; E-mail: Donnellan.Steve@saugov.sa.gov.au Commonly known as ‘whiting’, the 25 species of the fish family Sillaginidae inhabit the western Pacific and Indian Oceans. The King George whiting, Sillaginodes punctata, is the most widespread whiting species in southern Australia where it forms significant fisheries, which have become subject to management controls due to catch declines (Kailola et al. 1993). If multiple stocks are present, then their identification can allow management to be more efficiently focused. In an allozyme study of stocks of Australian whiting, Dixon et al. (1987) identified too few usefully polymorphic loci in King George whiting. Microsatellite DNA markers can provide sufficient numbers of polymorphic markers in species that have low proportions of polymorphic allozyme loci. We describe the isolation and characterization of microsatellite loci from the King George whiting. We also evaluate the cross-species amplification of some of these loci on two other species of whiting of the genus Sillago, from southern Australian waters that also form significant fisheries. Microsatellites were isolated, using (AAAG)6 probes, with two methods, a polymerase chain reaction (PCR)-based procedure (Cooper et al. 1997) and magnetic bead enrichment (Gardner et al. 1999). A total of 32 clones were isolated by the first method and sequenced with the Sp6 vector primer using PE Applied Biosystems PRISM™ Dye Terminator Cycle Sequencing Kit with the products run on an ABI 373 instrument. Sequencing showed that inserts of 24 clones contained AAAG repeats. Of the 18 clones isolated with the second procedure, nine showed tandem repeats following sequencing. Of the 33 primer pairs designed, 19 produced amplifiable microsatellite loci. GenBank accession numbers for the sequenced clones are AF291469 – 80. Each of the 19 loci tested for variability were amplified by PCR using 50–100 ng DNA, 10 pmol each primer, 0.2 mm each © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2215 Table 1 Primer sequences and variability measures for nine microsatellite loci genotyped in King George whiting. HO and HE are the observed and expected heterozygosity, respectively Locus Repeat sequence of clone allele Sp2 (ATAG)8(AAAG)5 Sp7 (AAAG)7 Sp19 (AAAG)4 Sp22 (AAAG)7 Sp32 (AAAG)4(ACAG)(AAG)3 Sp35 (AAAG)4 Sp36 (AAAG)6 Sp38 (CCT)8 Sp39 (GTATC)11 Primer sequences (5′− 3′) F: ATGCGTGAAGATGGTGTCA R: CTGTTCTCAGCAGTGCTTCA F: AAGCTCATTTTCATCAGCGT R: CGGATCGGAATTTGAAGACA F: CGTGTAACCCAGAAACCTACT R: CATCGAAGCATTGCCTGTAA F: CTACTTCACTGCTGCACTCACA R: GGACCAACACAAGACACACAA F: ACACAGATCGCGCACTTGTA R: CACTGTCCTCGCTGTGGTGA F: TCCTAGCTACGATGATGGATG R: TCTGGTCAGATTCGTCGATGG F: CCTCAGTAAGCGCCAGTAATAGAC R: CCTACAGCGATTGGTACAGCAC F: CCGTGACCGGTTCCATTGAG R: TCCTCAACTGCGTCTGTGTTCA F: TTGCTGACCATGTCAAGTTGA R: CACCAGGACAAGGCTGATATG Fluoro-label type No. of alleles Size range (bp) Mean HO Mean HE HEX 29 215–391 0.671 0.768 0.0027 HEX 9 119 –147 0.466 0.530 0.0093 FAM 3 197–205 0.361 0.331 0.0304 HEX 23 119–223 0.774 0.853 0.0016 TET 3 142–158 0.363 0.411 –0.0024 FAM 6 124–144 0.438 0.404 0.0096 TET 4 106–118 0.239 0.263 –0.0025 FAM 4 274–283 0.019 0.538 * HEX 17 206–286 0.434 0.850 † FST *†See text for explanation of locus departures from Hardy–Weinberg equilibrium. of dNTP, 4 mm MgCl2, 1× Promega Taq Gold dilution buffer (10 mm Tris-HCl pH 8.3, 50 mm KCl) and 0.5 U Promega Taq Gold DNA polymerase in a 25-µL reaction volume. PCR cycling conditions for reactions involving King George whiting were: 94 °C for 3 min, 58 °C for 45 s, 72 °C for 1 min for one cycle; 94 °C for 45 s, 58 °C for 45 s, 72 °C for 1 min for 34 cycles; and 72 °C for 6 min, 26 °C for 10 s for one cycle. Genotyping of 10 individuals from across the species geographical range, showed that nine loci were variable (Table 1). A single primer from each variable pair was re-synthesized and labelled with ABI fluorescent dyes (Table 1) for genotyping on an ABI 377 instrument using the Genescan application (PE Applied Biosystems). Co-amplification was achieved for the following four groups of loci: Sp2–22, Sp7–19, Sp35 –39 and Sp32– 36 –38. Loci were combined for electrophoresis as follows: Sp2–22, Sp7–19 and Sp32 – 35 – 36 – 38 – 39. The nine microsatellite loci were genotyped for 288 individuals in 10 populations from across the species’ geographical range. Inspection of the genotype arrays showed, for locus Sp38, a small number of individuals typed as homozygous for rare putative alleles in four populations that otherwise contained only the common allele. In view of these potentially anomalous typings and the high frequency of the common allele, either fixed or P > 0.99, the locus was omitted from further consideration. Tests for conformity to Hardy–Weinberg proportions in the remaining loci, after sequential Bonferroni adjustment ( Hochberg 1988), produced significant results for locus Sp39 in all populations. Locus Sp39 was omitted from further analysis because of the possible presence of null alleles. Table 1 shows the values of FST, for the remaining individual loci estimated with genepop (Raymond & Rousset 1995). Individual locus FST values are low, the typical picture for subpopulation differentiation seen in many marine fishes (Ward et al. 1994). Cross-species amplifications, without extra optimization and at an annealing temperature of 50 °C, were successful for Sillago bassensis for Sp2, Sp11, Sp19, but were unsuccessful for Sp7, Sp22 and Sp32. For S. schomburgkii, Sp2, Sp7, Sp19, and Sp32 were successfully amplified. In all cases clean products were detected in the single individual of each species tested. Acknowledgements This work was supported by a Fisheries Research and Development Corporation grant number 95/008. References Cooper S, Bull CM, Gardner MG (1997) Characterisation of microsatellite loci from the socially monogamous lizard Tiliqua rugosa using a PCR-based isolation technique. Molecular Ecology, 6, 793–795. Dixon PI, Crozier RH, Black M, Church A (1987) Stock identification and discrimination of commercially important whitings in Australian waters using genetic criteria. FIRTA Project 83/16, final report 69pp. Centre for Marine Science, University of New South Wales, NSW, Australia. Gardner MG, Cooper S, Bull CM, Grant WN (1999) Isolation of microsatellite loci from a social lizard, Egernia stokesii, using a modified enrichment procedure. Journal of Heredity, 90, 301– 304. Hochberg Y (1988) A sharper Bonferroni procedure for multiple tests. Biometrika, 75, 800–802. Kailola PJ, Williams MJ, Stewart PC, Reichelt RE, McNee A, Grieve C (1993) Australian Fisheries Resources. Bureau of Resource Sciences and the Fisheries Research and Development Corporation, Canberra, Australia. Raymond M, Rousset F (1995) genepop (Version 3.1b): population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. Ward RD, Woodwark M, Skibinski DOF (1994) A comparison of genetic diversity levels in marine, freshwater, and anadromous fishes. Journal of Fish Biology, 44, 213–232. 2000 Graphicraft 1161 19PRIMER 32 NOTEs Limited, Hong Kong © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 2216 P R I M E R N O T E S Microsatellite primers from the Eurasian badger, Meles meles R . B I J L S M A ,* M . VA N D E V L I E T ,* C . P E RT O L D I ,† R . C . VA N A P E L D O O R N ‡ and L . VA N D E Z A N D E * *Department of Genetics, University of Groningen, Kerklaan 30, NL-9751 NN Haren, The Netherlands, †Department of Ecology and Genetics, University of Aarhus, Building 540, Ny Munkegade, DK-8000 Aarhus C, Denmark, ‡Alterra, PO Box 23, 6700 AA Wageningen, The Netherlands Keywords: badger, microsatellites, mustilidae, pine marten, primers Received 29 July 2000; revision accepted 2 September 2000 Correspondence: R. Bijlsma. Fax: + 31 50 3632348; E-mail: r.bijlsma@biol.rug.nl In man-dominated landscapes populations of once common species have become decreased both in range and density and been restricted to small habitat patches with reduced dispersal possibilities. Such fragmented populations become increasingly affected by stochastic population dynamics due to demographic, environmental and genetic risks, eventually leading to increased extinction probabilities (Bijlsma et al. 2000). The Eurasian badger (Meles meles L.) is a species that is threatened in many parts of Western Europe because of fragmentation and suffers greatly from agricultural activities (Moore et al. 1999). Moreover, increasing density of roads and traffic does not only considerably limit dispersal, but also highly increases mortality due to road-kills (Aaris Sørensen 1995). From a conservation perspective insights into the (meta)population structure of the badger is clearly needed. As allozyme variation was found to be low in badgers (Pertoldi et al. 2000), we have developed highly variable microsatellite markers that make noninvasive sampling and use of dead animals possible. To isolate microsatellite markers, muscle tissue was obtained from 23 badgers, killed by traffic accidents, by sampling a piece of the ear or the tail. DNA extraction and small insert libraries were constructed using standard procedures (Ausubel et al. 1987). Overnight incubation in lysis buffer (100 mm NaCl, 10 mm Tris-HCl pH 8.0, 25 mm EDTA, 0.5% SDS, 0.1 mg/mL proteinase K) at 55 °C was followed by two phenol extractions and one phenol:chloroform (24:1 v/v) extraction. Ethanol precipitated DNA was dried and dissolved in TE (10 mm Tris-HCL pH. 7.6, 1 mm EDTA). Total genomic DNA was digested to completion with MboI, size fractionated on a 1% agarose gel and the 200 –1000 bp fraction was recovered by electroelution. These fragments were ligated into BamHI digested pBluescript and used to transform competent XL1-Blue Escherichia coli cells to establish a small-insert library. This library was screened with synthetic (CA)7 and (GA)7 probes, end-labelled with [γ 32P]-ATP. Positive clones were sequenced using the T7-sequencing kit and [α35S]-dATP. Out of an initial 2400 clones, 43 positive recombinants were identified, eventually yielding seven usable microsatellite loci. Polymerase chain reactions (PCRs) were carried out in 10 µL volumes, containing 100 ng template DNA, 0.5 µm each primer, 0.2 mm dATP, dGTP and dTTP, 0.02 mm dCTP, 0.4 U Taq DNA polymerase (Pharmacia) and 0.14 µCi [α32P]-dCTP (3000 Ci/mmol) in buffer (50 mm KCl, 1.5 mm MgCl2 and 10 mm Tris-HCl, pH 9.0). After an initial 3 min at 94 °C, 30 cycles were performed with the following profile: 1 min at 94 °C, 2 min at Ta (optimal annealing temperature) and 1.5 min at 72 °C, followed by 10 min at 72 °C. Labelled PCR products were separated on a 5% denaturing polyacrylamide gel (Biozym, Sequagel XR) and exposed to medical X-ray film (Fuji) for 5 –16 h at –70 °C, using intensifying screens. A sequence ladder of pBluescript was used as size reference. The level of polymorphism was determined for a total of 105 badger samples collected from different localities in The Netherlands and Denmark. The characteristics of the seven microsatellite loci are shown in Table 1. All loci were found to be polymorphic and the mean number of alleles was 4.3 (range: 2 – 6) and mean expected heterozygosity (HE) was 0.45 (range: 0.15 – 0.65). Although this is within the range observed for other mustelid species (O’Connell et al. 1996; Dallas & Piertney 1998; Fleming et al. 1999), mean expected heterozygosity in the badger was lower than in these other species (means ranging from 0.55 to 0.84). Except for two subsamples of locus Mel 2, mainly due to very low expected numbers for some genotypes, no significant differences in expected and observed levels of heterozygosity were observed for all loci, indicating the absence of null-alleles. The badger primer sets were also evaluated for use in two other mustelid species. In the pine marten (Martes martes, n = 88), two primer sets failed to produce an amplification product, two were found to be polymorphic and the other three monomorphic (Table 1). In the otter (Lutra lutra, n = 5), all primer sets yielded an amplification product (data not shown). However, the sample size was too small to reliably estimate number of alleles and expected heterozygosity. Presently, the primer sets are used to assess the current genetic population structure of badger populations and of the pine marten. Acknowledgements We thank the ‘Vereniging Das en Boom’ (Beek-Ubbergen, NL) and the National Environmental Research Institute (Rønde, DK) for supplying the badger road-kills, and Alterra (Wageningen, NL) for the pine marten samples. References Aaris Sørensen J (1995) Road-kills of badgers (Meles meles) in Denmark. Annales Zoologici Fennici, 32, 31–36. Ausubel FM, Brent R, Kingston RE et al. (1987) Current Protocols in Molecular Biology. Wiley, New York. Bijlsma R, Bundgaard J, Boerema AC (2000) Does inbreeding affect the extinction risk of amall populations?: predictions from Drosophila. Journal of Evolutionary Biology, 13, 502–514. Dallas JF, Piertney SB (1998) Microsatellite primers for the Eurasian otter. Molecular Ecology, 7, 1248–1251. Fleming MA, Ostrander EA, Cook JA (1999) Microsatellite markers for American mink (Mustela vison) and ermine (Mustela erminea). Molecular Ecology, 8, 1352–1354. Moore N, Witherow A, Kelly P, Garthwaite D, Bishop J, Langton S, Cheeseman C (1999) Survey of badger Meles meles damage to © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2217 Table 1 Attributes of seven microsatellite loci derived from the badger (Meles meles) in 105 badgers and 88 pine martens (Martes martes) samples. Ta, annealing temperature; SR, size range observed in bp; A, number of alleles; HE, expected heterozygosity; HO, observed heterozygosity; NA, no amplification product. GenBank accession nos of these loci are AF300707–AF300714 Meles meles Martes martes Locus Repeat structure Primer sequences (5′−3′) Ta (°C) SR A HE HO SR A HE HO Mel 1 (GT)20 60 262–274 5 0.54 0.45 261–267 4 0.69 0.68 Mel 2 (GT)12 55 126–128 2 0.15 0.08 NA — — — Mel 3 (GT)13 60 128–134 4 0.69 0.67 NA — — — Mel 4 (GT)16 60 141–147 4 0.21 0.21 144 1 0 0 Mel 5 (GT)23 60 105–119 6 0.65 0.53 188 1 0 0 Mel 6 (GT)13AC(GA)4 60 149–155 4 0.33 0.29 137–139 2 0.29 0.31 Mel 7 (GT)21 CTGGGGAAAATGGCTAAACC AATGCAGGCTTTGCAATTCC TTGTGCGTATGCATGTGTGC TGCCCACGTTATAAACACTCC CTAAAACCACCACCACAATGC GTGTATAGCCTGCGAACAAGG TGAGTTTCCATCCTTGGTCC ATCTTTTTCCTGCTGAGACCC AATGTAAGGTACCCAGCATAGTCC GACACCATGTTAACCATATAAAGGG AAGTCCTCCTTGCAGTTTGG AGCAAGCTCTTGGTTCTTGG ATTCTTCCTTTTAGCTTTGGCC TCTCACAGTGTCAGCAGAAAGG 60 134–144 5 0.58 0.50 124 1 0 0 agriculture in England and Wales. Journal of Applied Ecology, 36, 974–988. O’Connell M, Wright JM, Farid A (1996) Development of PCR primers for nine polymorphic American mink Mustela vison microsatellite loci. Molecular Ecology, 5, 311–312. Pertoldi C, Loeschcke V, Madsen AB, Randi E (2000) Allozyme variation in the Eurasian badger Meles meles in Denmark. Journal of Zoology, 252, in press. Graphicraft 00 primer PRIMER 1162 2000 912 Notes NOTEs Limited, Hong Kong Characterization of microsatellite loci in the eastern oyster, Crassostrea virginica BONNIE L. BROWN,* DEAN E. FRANKLIN,* PAT R I C K M . G A F F N E Y , † M I N H O N G , ‡ D A N D E N D A N T O § and I RV K O R N F I E L D ¶ *Ecological Genetics Laboratory, Virginia Commonwealth University, Richmond, Virginia, USA, †College of Marine Studies, University of Delaware, Lewes, Delaware, USA, ‡Basic College of Medicine, Norman Bethune University of Medical Sciences, Chang Chun, Jilin Province, PR China, §Department of Biological Sciences, University of Maine, Orono, Maine, USA, ¶School of Marine Sciences, University of Maine, Orono, Maine, USA Keywords: Crassostrea virginica, genetics, microsatellite, oyster Received 10 August 2000; accepted 2 September 2000 Correspondence: B. L. Brown. Fax: + 804 8280503; BLBROWN@vcu.edu Oysters of the genus Crassostrea are of great ecological and economic value worldwide. Eastern oysters, C. virginica, were once a keystone species of western Atlantic estuaries but now are depleted in many areas due to the combined effects of overharvesting, habitat alteration, and diseases caused by introduced parasites (Brown & Paynter 1991). Hopes of restoring oysters to these regions are tied to development of © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 disease tolerant strains of the native C. virginica, and efforts to cultivate this and related species involve the use of markerassisted selection. The preferred character for such studies currently is the microsatellite (Hare & Avise 1997; Magoulas et al. 1998; Huvet et al. 2000). In each published instance to date, dinucleotide repeats were investigated, exhibiting extremely high heterozygosities and therefore reduced power for multilocus heterozygosity surveys. We surveyed a C. virginica library for the presence of tri- and tetranucleotide repeats. Genomic DNA was purified from C. virginica somatic tissue with an STE (10 mm Tris-HCl, pH 8.0, 1 mm EDTA, 100 mm NaCl, 2% SDS, 0.5 mg/mL proteinase K) extraction (Hillis et al. 1996), which included treating the first aqueous phase following organic extraction with RNase A (final concentration 0.03 µg/µL) for 30 min. Detection of microsatellite sequences in a size-selected (400 and 900 bp) partial genomic library (pBluescript II SK+) was performed as described by Rassmann et al. (1991). Transformant colonies were screened using a cocktail of two digoxygeninlabelled oligonucleotide probes [(ATG)7 and (AAAC)5] and DNA inserts of 13 positive colonies were sequenced. Primers were designed with target annealing temperatures of 50 – 55 °C and expected amplicon lengths between 80 and 220 bp. Polymerase chain reactions (PCRs) were performed in reactions containing 100 ng genomic DNA, 0.5 µm unlabelled forward primer, 0.25 µm unlabelled reverse primer, 0.25 µm labelled reverse primer, 0.2 mm each dNTP, 0.5 U of Taq polymerase (Display Systems Biotech), 10 mm Tris-HCl, pH 8.3, 2.5 mm MgCl2, 50 mm KCl, 0.01% Triton X-100, 0.0005% gelatin, and sufficient diH2O for a total volume of 15 µL. Amplification was conducted in PTC-100 thermal cyclers (MJ Research) using an initial denaturation at 94 °C for 2 min, followed by 30 cycles of 94° for 30 s, 50 – 55° for 30 s, and 72° for 15 s. Amplification products were resolved by ultrathin gel electrophoresis of fluorescent-labelled PCR products (using filter set 2218 P R I M E R N O T E S Table 1 Repeat structure, primer sequences, amplification characteristics, and polymorphism data for microsatellite loci examined in Crassostrea virginica. Observed numbers of alleles, heterozygosity values (observed and expected), and the P-values for exact tests of fit to Hardy–Weinberg equilibrium (HWE; for each P-value compared to Bonferroni-corrected alpha value, significant departure is shown by an asterisk ‘*’) were determined for native populations in Virginia (n = 40) and Connecticut (n = 44; for Cvi6, 8, and 11 only), USA, the latter shown in parentheses Locus Repeat © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Cvi6 (GAT)17 Cvi7 (CAAA)6 Cvi8 [(CAAA)2(CAA)]2 Cvi9 (CAT)14 Cvi11 (CAAA)5 Cvi12 (CAAA)6CAGAAAAA(CAAA)4 Cvi13 (CAAA)10 GenBank Accession no. Primer Sequences (5′−3′) (F = forward, R = reverse) Anneal (°C) Expected size (bp) Observed no. alleles HO HE HWE AF276247 F: AATATTACCACGTGACCTGTGATGAATCCTTGTAGC R: GTAAATATTGTATGTTCACTGTCCGGTCGTTGTGTTA F: TCGAAACCGAACCCTTCACCAG R: TAGTGTATATCAGTTCAGACAGGTCTTTTAATGG F: CTGAGCTTAGACTACAGCCCTACACCAG R: GATATCCTAAACCTACTCCTCTTTTGCATTTTTG F: TCCAGAATTTATAAGATACTAACGATAATATACTTTATAATCCGT R: ACGAAACCGACCACAACGACGACT F: CATCGGCCAGTGACTACCTTGTAAAAG R: GCGATAACACTAAATACTTTGTTTCGGCCC F: GAGTGAGAATTTCTCGGGTGGGGC R: ACTTTTTGTCACATTGACCATCCCATTTCA F: ACCGGAGATGGTGGTATTTCC R: GTGTTGCAAGACTTACAGAAGAAAC 50 198 50 196 13 (15) 9 0.54 (0.55) 0.58 0.87 (0.90) 0.77 <0.0001* (<0.0001*) <0.0001* 55 205 55 124 14 (6) 14 0.48 (0.18) 0.83 0.68 (0.47) 0.90 0.0001* (<0.0001*) 0.1207 55 153 55 117 4 (4) 10 0.51 (0.48) 0.70 0.64 (0.53) 0.81 0.0465 (0.2882) 0.0399 50 156 26 0.71 0.94 0.0027* AF276248 AF276249 AF276250 AF276252 AF276253 AF276254 P R I M E R N O T E S 2219 C and tamara size standard). Mendelian inheritance of alleles was determined by examining the amplified products in two or more full sib families per locus (both parents and 15 – 20 offspring in each family). To determine allele range and population-level variability, two wild groups of C. virginica were examined: one from Virginia Beach, Virginia (latitude 36°54′N, longitude 076°05′W; n = 40) and one derived from wild spat fall in Long Island Sound, Connecticut (latitude 41°06′N, longitude 73°25′W; n = 44). Primers also were tested with C. gigas (n = 5 each from two populations), C. angulata (n = 5), Saccostrea glomerata (formerly S. commercialis; n = 5) and Tiostrea chilensis (n = 5). Statistical analyses were performed using genepop version 3.1 (Raymond & Rousset 1995). Of the 10 primer sets, all amplified from C. virginica products of the size expected from insert sequences. Three yielded homologous products and seven loci (Cvi-6, Cvi-7, Cvi-8, Cvi-9, Cvi-11, Cvi-12, and Cvi-13) were polymorphic (Table 1). All seven polymorphic loci exhibited Mendelian segregation. The Virginia C. virginica population was surveyed for variation at all seven loci and the Connecticut population was surveyed for Cvi-6, Cvi-8, and Cvi-11. Only three of the seven loci conformed approximately to Hardy–Weinberg equilibrium (Cvi-9, Cvi-11, and Cvi-12). For all loci, observed heterozygosity was lower than expected, suggesting the common occurrence of segregating null alleles. No evidence for linkage was observed among these seven loci. Allelic distribution was significantly different between the two wild C. virginica populations (Fisher exact test P < 0.0001). When tested with C. gigas, C. angulata, and S. glomerata, four of the 10 primer sets (Cvi6, Cvi9, Cvi12, Cvi13) yielded various homologous products differing substantially in size from the allele sizes observed for C. virginica. No amplification was observed for T. chilensis. Acknowledgements We thank the following for providing tissue or DNA samples of oysters: Trafford Hill, Tallmadge Bros. Inc., Geoff Allan, Mike Heasman, Diarmaid Ó Foighil, John Scarpa, and Jon Waters. We acknowledge the assistance and advice of Tracy Hamm, Bill Eggleston and Jon Waters. This research was supported in part by funding from Chesapeake Scientific Investigations Foundation, Inc. and by a grant from the Maine Aquaculture Innovation Center (98 – 23). References Brown BL, Paynter KT (1991) Mitochondrial DNA analysis of native and selectively inbred Chesapeake Bay oysters, Crassostrea virginica. Marine Biology, 110, 343–352. Hare MP, Avise JC (1997) Population structure in the American oyster as inferred by nuclear gene genealogies. Molecular Biology and Evolution, 15, 119 –128. Hillis DM, Mable BK, Larson A, Davis SK, Zimmer EA (1996) Nucleic Acids IV: sequencing and cloning, In: Molecular Systematics 2nd edn. (eds Hillis DM, Moritz C, Mable B), pp. 342– 343. Sinauer Associates, Inc., Sunderland, Massachusetts, USA. Huvet A, Boudry P, Ohresser M, Delsert C, Bonhomme F (2000) Variable microsatellites in the Pacific oyster Crassostrea gigas and other cupped oyster species. Animal Genetics, 31, 71–72. Magoulas AN, Gjetvaj B, Terzoglou V, Zouros E (1998) Three © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 polymorphic microsatellites in Japanese oyster Crassostrea gigas (Thunberg). Animal Genetics, 29, 69 –70. Rassmann KC, Schlötterer C, Tautz D (1991) Isolation of simplesequence loci for use in polymerase chain reaction-based DNA fingerprinting. Electrophoresis, 12, 113–118. Raymond M, Rousset F (1995) genepop (vers 1.2): population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248–249. 2000 1148 109PRIMER Graphicraft 2 00 NOTEs Limited, Hong Kong Microsatellite markers for Rhytidoponera metallica and other ponerine ants M . C H A P U I S AT , * and J . N . PA I N T E R , † and R. H. CROZIER,‡ Department of Genetics, La Trobe University, Bundoora, Victoria 3083, Australia Keywords: microsatellite, ponerine ants, Rhytidoponera metallica, social insects Received 17 August 2000; revision accepted 4 September 2000 Correspondence: Michel Chapuisat. *Present address: Institute of Ecology, University of Lausanne, 1015 Lausanne, Switzerland. Fax: + 41 21 692 41 65; E-mail: Michel.Chapuisat@ie-zea.unil.ch Present addresses: †Department of Ecology and Systematics, University of Helsinki, PL 17, FIN-00014 Helsinki, Finland. ‡School of Tropical Biology, James Cook University, Townsville, Queensland 4811, Australia. The ant genus Rhytidoponera (subfamily Ponerinae) contains 104 described species (Bolton 1995) which are remarkably diverse in their social organization and mating system (Crozier & Pamilo 1996). The greenhead ant Rhytidoponera metallica is among the most common ants in Australia, and it possesses an unusual social structure, as the reproductive role is almost invariably taken by multiple mated workers in lieu of queens (Haskins & Whelden 1965). This secondary loss of queens and partitioning of reproduction between morphologically undifferentiated workers offers a good opportunity to study how altruism is maintained in societies with low relatedness (Hamilton 1972). Such studies require detailed genetic data on the social organization and mating system. Recently, microsatellite markers have been described in three species of ponerine ants (Doums 1999; Giraud et al. 1999; Tay & Crozier 2000), but only two microsatellites from the most related species proved useful in R. metallica. Therefore, we characterized eight new microsatellite markers for R. metallica, and tested for cross-species amplification in 10 other species of ponerine ants. A partial genomic library was constructed from 100 R. metallica workers, with gasters removed. DNA was extracted with a CTAB protocol (Hillis et al. 1990), digested to completion with Sau3A I and RsaI, size-selected for fragments between 300 and 900 bp (Crozier et al. 1999), and ligated into a pUC19 vector. The library was screened with an (AG)10 oligonucleotide probe end-labelled with 33P, and 62 positive recombinant clones were isolated. Thirty positive clones were sequenced, and primers were designed for 14 of them. These primers were assayed on a sample of workers collected from the You Yangs Regional Park in Victoria. DNA 2220 P R I M E R N O T E S Table 1 Characteristics of nine microsatellite loci for Rhytidoponera metallica. n, number of individuals analysed; N, number of nests analysed; HO, observed heterozygosity; HE, expected heterozygosity. Deviations from Hardy–Weinberg equilibrium are not significant (exact tests). GenBank accession nos: AF282988 –AF282998, AF292086 Locus Primer sequence (5′− 3′) Rmet3 F: TCTCGGAAAAGAAATAGAGACAG R: CATGTCTACCTGACCGAGAAC F: CATACTATCGCTTATCTCAGC R: GAACTAACCTCATCGTCCACT F: AGACTTCAATCACGAGAAGCG R: ATTGGCACTTGGTCGATAGG F: AAAACACGAGATACCGTCCTC R: CTGTTGACCCGCCTCCTG F: GTCATGGACGGAAATCGC R: TACCCCCATTCTATCTCGCA F: GGAGTTTCTACTCGCCTCTCG R: CTCATTCGTATCACGCAAGC F: CATTCGACCGCATTTTCC R: CGAGAGAGGGTGCGACAT F: TTTAGGGACAAGAGACATGGC R: ATTGATAGGTCGCGGTCTTG F: GACATACCGGGAGCGACC R: CGCCTTCTGACACCTTTGG Rmet4 Rmet7 Rmet8 Rmet10 Rmet12 Rmet15 Rmet16 Rh12 –13525 Core repeat in cloned allele n/N No. alleles Size range HO HE (GA)40 23/13 10 226–248 0.74 0.84 (CT)26 14/14 11 152–178 1.00 0.87 (AG)30 216/ 27 21 223–269 0.86 0.86 (CT)50 27/13 15 108–144 0.96 0.88 (CT)37 216/ 27 23 246–296 0.91 0.89 (GA)20 216/ 27 15 275–315 0.85 0.87 (AG)28 216/ 27 9 154–202 0.44 0.42 (CT)40 18/14 17 117–203 1.00 0.92 (CT)11 216/ 27 7 178–192 0.72 0.70 from individual workers was extracted by incubating three crushed legs in 250 µL of 5% Chelex at 95 °C for 20 min (Crozier et al. 1999). Amplification was carried out in 10 µL final volume with 10 mm Tris-HCl, 50 mm KCl, 0.1% Triton X-100, 1.5 mm MgCl2, 1.7 µm each dNTP, 0.03 – 0.05 µm forward primer end-labelled with 33P, 0.4 µm reverse primer, 5 µg of BSA, 0.4 U of Taq DNA polymerase (Promega) and 2 µL of template DNA. The polymerase chain reaction (PCR) profile consisted of a 3-min initial denaturation step at 94 °C, followed by 30 cycles of 30 s at 92 °C, 30 s at 50 °C and 30 s at 72 °C. PCR products were separated by electrophoresis through 6% denaturing polyacrylamide gels. Eight primer pairs yielded suitable amplification products. All eight markers were highly polymorphic, with between nine and 23 alleles detected in the study population (Table 1). Alleles were somewhat difficult to score for Rmet8 and Rmet16, because of stutter bands. Additionally, the previously unpublished marker Rh12 –13525, which was developed by W. Tek Tay for Rhytidoponera sp. 12, (Tay & Crozier 2000) had seven alleles in R. metallica (Table 1). The success of cross-species amplification in other genera was low (Table 2). Scorable amplification products were obtained in only 12 out of the 45 tests (27% of the five species assayed for nine markers). Polymorphism among three individuals was detected at a single marker in four species, i.e. in 9% of the 45 tests. In contrast, the success of cross-species amplification within the genus Rhytidoponera was very high (Table 2). Priming sites were well conserved among the Rhytidoponera species tested, resulting in strong amplification products in 40 out of the 45 tests (89%). Overall, scorable polymorphism among three individuals was detected in 23 out of the 45 tests (51%). In each species of Rhytidoponera, between three and eight markers were polymorphic, and this figure should increase when more individuals are analysed. Hence, this panel of microsatellites will permit detailed studies of kin structure, breeding system, gene flow and population structure across species of Rhytidoponera with variable social structures. Additionally, these markers might help to distinguish between the species yet to be described that are currently lumped into the metallica species-complex. Acknowledgements We thank Bruno Gobin, Juergen Liebig, Christian Peeters and Hanna Reichel for specimens of various ponerine species, Wee Tek Tay for the primers of the locus Rh12–13525, and Parks Victoria for permitting research in the You Yangs Regional Park. This work was supported by grants from the Swiss National Science Foundation and the Société Académique Vaudoise. References Bolton B (1995) A New General Catalogue of the Ants of the World. Harvard University Press, Cambridge, MA. Crozier RH, Kaufmann B, Carew ME, Crozier YC (1999) Mutability of microsatellites developed for the ant Camponotus consobrinus. Molecular Ecology, 8, 271–276. Crozier RH, Pamilo P (1996) Evolution of Social Insect Colonies: Sex Allocation and Kin Selection. Oxford University Press, Oxford. Doums C (1999) Characterization of microsatellite loci in the queenless Ponerine ant Diacamma cyaneiventre. Molecular Ecology, 8, 1957–1959. Giraud T, Blatrix R, Solignac M, Jaisson P (1999) Polymorphic microsatellite DNA markers in the ant Gnamptogenys striatula. Molecular Ecology, 8, 2143–2145. Hamilton WD (1972) Altruism and related phenomena, mainly © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2221 Table 2 Results of cross-species amplification in 10 other ant species from the subfamily Ponerinae Rmet3 Rmet4 Rmet7 Rmet8 Rmet10 Rmet12 Rmet15 Rmet16 Rh12–13525 Tribe ECTATOMMINI Rhytidoponera tasmaniensis R. victoriae R. purpurea R. impressa R. confusa Gnamptogenys menadensis +5 +2 +1 +1 +1 +1 +3 +1 +1 +1 +1 — — +1 +2 +5 +3 — +5 +1 — — — — +2 +2 +1 +1 +2 — +3 +2 +3 +2 +1 +1 +2 +1 +2 s s +2 +2 — +1 +1 +2 — +4 +2 +4 +4 +3 — Tribe PONERINI Diacamma cyaneiventre D. ceylonense Harpegnathos saltator Streblognathus aethiopicus +3 +2 +3 — — — — — — — — — — — — — — — +1 — — — — +1 +1 +1 +1 +1 — — — — — — — — + n, scorable amplification product with n alleles detected in three individuals. —, no scorable amplification product. s, present of supernumerary amplification products. in social insects. Annual Review of Ecology and Systematics, 3, 193– 232. Haskins CP, Whelden RM (1965) ‘Queenlessness’, worker sibship, and colony versus population structure in the Formicid genus Rhytidoponera. Psyche, 72, 8 7 – 112. Hillis DM, Larson A, Davis SK, Kimmer EA (1990) Nucleic acids III: Sequencing. In: Molecular Systematics (eds Hillis DM, Moritz C), pp. 318 – 370. Sinauer, Sunderland, MA. Tay WT, Crozier RH (2000) Microsatellite analysis of gamergate relatedness of the queenless ponerine ant Rhytidoponera sp. 12. Insectes Sociaux, 47, 188–192. Graphicraft 00 PRIMER 1152 2000 912 NOTEs Limited, Hong Kong Rapid and efficient identification of microsatellite loci from the sea urchin, Evechinus chloroticus C . P E R R I N and M . S . R O Y Department of Zoology, University of Otago, PO Box 56, Dunedin, New Zealand Keywords: biotin, Evechinus chloroticus, microsatellites, nonradioactive Received 10 August 2000; revision accepted 4 September 2000 Correspondence: M. S. Roy. Fax: + 64 3479 7584; E-mail: michael.roy@stonebow.otago.ac.nz The New Zealand Fiords are characterized by a seawardly flowing surface low salinity layer (LSL), produced by prodigious rainfall. Maintenance of salt balance occurs by a weak oceanic inflow below this LSL. Because the flow of sea water is inwards, planktonic larvae of the fiords are thought to be retained within natal fiords, which could have important consequences on gene flow. This hypothesis was supported by allozyme analyses of Evechinus chloroticus, a sea urchin endemic to New Zealand (Mladenov et al. 1997). Despite high levels of gene flow found amongst all coastal populations of New Zealand, a population sampled within one fiord © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 was found to be genetically differentiated. Our intention is to address the effects of oceanographic and hydrographic features of all 14 fiords on recruitment and population structuring of E. chloroticus. In order to do this we are using highly polymorphic microsatellite markers. Traditional colony hybridization methods used for microsatellite cloning are time-consuming and relatively inefficient. Several enrichment techniques have previously been published (Gardner et al. 1999; Inoue et al. 1999). However, these either use radioactivity or include a number of lengthy steps. We report here on an alternative easy, fast, efficient and nonradioactive method of cloning microsatellite markers from the sea urchin E. chloroticus. Size selected fragments (250 – 800 bp) of NdeII-digest genomic DNA from five individuals of E. chloroticus were ligated into pUC18 vector (Pharmacia). Inserts were amplified using universal primers (M13) and purified with High Pure PCR product purification Kit (Roche). In order to hybridize DNA to probes, 100 – 500 ng of size selected amplified DNA (250 – 800 bp) was mixed, in separate tubes, with 2 pmol of GA12 and GT12 5′-biotinylated repeat probes in 20 µL of extension solution containing: 0.2 mm of each dNTP, 2 mm MgCl2, 10 mm Tris-HCl (pH 8.3), 50 mm KCl, and 0.5 U Taq DNA polymerase (Roche). This mixture was subjected to one round of polymerase chain reaction (PCR) (5 min at 94 °C, 1 min at 55 °C, 10 min at 72 °C) using a PTC-100 thermal cycler (MJ Research). Purified products were added to Streptavidin MagneSphere Paramagnetic Particles (Promega) and incubated for 15 min at room temperature with 120 µL of 6× SSC/0.1% SDS, mixed continuously. After a series of washes in 150 µL of 6× SSC/0.1% SDS for 15 min: once at 60 °C, 65 °C, 70 °C, 75 °C and twice in 150 µL of 6× SSC at 80 °C, DNA was eluted with 100 µL of 0.1 m NaOH at 80 °C for 10 min. The solution was neutralized with 100 µL TE pH 7.5, purified and amplified as above. A further round of enrichment (hybridization, elution, PCR) was then undertaken. Size selected fragments (250 – 800 bp) of NdeII-digest from enriched inserts were ligated into pUC18 vector. Ligation 2222 P R I M E R N O T E S Table 1 Characteristics of height Evechinus chloroticus microsatellite loci. HO and HE are observed and expected heterozygosities, respectively, calculated with genetix 4.1 (Belkhir et al. 1996). PCR programmes are: (1) 31 cycles of 15 s at 94 °C, 10 s at annealing temperature, 10 s at 72 °C; and (2) 4 min at 94 °C, 31 cycles of 1 min at 94 °C, 1 min at annealing temperature, 1 min at 72 °C and finished by 10 min at 72 °C Locus Primer sequences (5′−3′) Repeat array PCR programme Annealing temp. (°C) No. of alleles Size range (bp) HO HE Accession no. C1 F: CTGCCCGGAAGTATTGTTATTG R: CATTTCGGCCACGGTCACT F: GAATAAACATTTACAAATCTGTC R: ATAAAAAGGGAAACGAAACAAGAA F: GATCGGTATGATAAACTT R: ATGCATGGGTAGGTGTG F: ACGGTTCGATTGAGAGAG R: TGACGGGGCAGGAAATGTG F: GATCATTGAGATGGCGATG R: GCACCCACACGTACGCGC F: CTGTGTTCTATTAAAATTGTCCTC R: TTGAAATTTGCTCTACCCCTATT F: CGACAAGTCCACCGTTCAACTCCA R: ATCTACTGTTGTTGCCTGTGTCAC F: ATCCCCTTCAAATGTTGCCTGATT R: GGCGTAACGGTAATGACCCTGTC (AG)23 1 53 24 110 –166 0.92 0.92 AF299134 (AG)11 2 55 6 77 – 89 0.77 0.77 AF299135 (CT)3CC(CT)3AT(CT)10 1 45 8 92 –102 0.74 0.73 AF299136 (AG)19 1 51 13 109 –133 0.63 0.86 AF299137 (GT)6 1 51 4 58 – 64 0.31 0.36 AF299138 (GA)4TAT(GA)7 2 58 7 85 – 97 0.14 0.51 AF299132 (CA)2CT(CA)2CT(CA)4TG(AC)2 1 51 6 91–109 0.07 0.28 AF299139 (AG)2(GA)5AA(AG)4 1 51 3 99 –111 0.07 0.07 AF299133 C29 G29 A34 © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 B14 A12 D1 A13 P R I M E R N O T E S 2223 reactions were transformed into Escherichia coli XL1-Blue competent cells. Recombinant clones were screened using two separate 10 µL PCR reactions, incorporating the repeat probe and either one of the M13 universal primers. Products were visualized on 2% agarose gel using ethidium bromide. Approximately 70% (19) and 45% (7) of the clones were positive for GA12 and GT12 probes, respectively. Screening of the same clones was attempted by using both M13 primers and the complementary probe in the same PCR reaction (as in Gardner et al. 1999). However, only 10% of clones for each probe were positive, indicating that this screening method is unreliable. Eighteen and six positive clones were amplified with M13 primers and sequenced using Big-Dye cycle sequencing kit (Applied Biosystems), separated on ABI 377 automated sequencer. Eighteen and five sequences contained microsatellites, respectively, and 11 and four were unique resulting in a final enrichment efficiency of approximately 45% for GA12 and 30% for GT12. Primer pairs were designed from sequences flanking repeats. PCR were performed in 10 µL reaction mixture: 20– 200 ng DNA, 0.2 mm of each dNTP, 1.5 mm MgCl2, 10 mm Tris-HCl (pH 8.3), 50 mm KCl, 0.5 µm of [γ 33]-ATP-labelled forward primer, 0.5 µm of reverse primer and 0.25 U Taq DNA polymerase. Different PCR regimes were used (Table 1). Alleles were separated on a 5% denaturing polyacrylamide gel (Long Ranger, FMC) and visualized by autoradiography. Eight polymorphic loci were identified and scored for 100 individuals from 14 sites along the fiords (Table 1). No linkage disequilibrium was detected between each pair of locus using genetix 4.1 (1000 permutations, P < 0.05) (Belkhir et al. 1996). To assess Wahlund effects, 26 individuals of the same site were analysed for deficit of heterozygotes. Loci A12 and D1 showed significant deviation from Hardy–Weinberg equilibrium (1000 permutations, P < 0.05) suggesting the possibility of null alleles. We also tested the utility of these primers for two individuals from each of Coscinasterias muricata (Asteroidea), Ophiactis savignyi and Amphipholis squamata (Ophiuroidea). Only the less polymorphic locus (A13) seems to amplify clearly in such a large range of echinoderms. Acknowledgements We thank Philip Mladenov and Steve Wing for help with sample collections, and Renate Sponer for technical advice. This work was funded by the Royal Society of New Zealand’s MARSDEN FUND, and approved by the Environmental Risk Management Authority of New Zealand (No. GMO00/UO021). References Belkhir K, Borsa P, Goudet J, Chikhi L, Bonhomme F (1996) GENETIX (Version 4.01), logiciel sous WINDOWS TM pour la génétique des populations. Laboratoire Génome, Populations, Interactions, Université de Montpellier II, Montpellier. Gardner MG, Cooper SJB, Bule CM, Grant WN (1999) Isolation of microsatellite loci from social lizard, Egernia stoksii, using a modified enrichment procedure. Journal of Heredity, 90 (2), 301–304. Inoue S, Takahashi K, Ohta M (1999) Sequence analysis of genomic © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 regions containing trinucleotide repeats isolated by a novel cloning method. Genomics, 57, 169–172. Mladenov PV, Allibone RM, Wallis GP (1997) Genetic differentiation in the New Zealand sea urchin Evechinus chloroticus (Echinodermata: Echinoidea). New Zealand Journal of Marine and Freshwater Research, 31, 261–269. 2000 Graphicraft 1155 19PRIMER 32 NOTEs Limited, Hong Kong Polymorphic microsatellite DNA markers in the African elephant (Loxondonta africana) and their use in the Asian elephant (Elephas maximus) L . S . E G G E RT ,* U . R A M A K R I S H N A N ,* N . I . M U N D Y † and D . S . W O O D R U F F * *Ecology, Behavior and Evolution, University of California San Diego, 9500 Gilman Dr., La Jolla, CA 92093 – 0116, USA, †Institute of Biological Anthropology, Oxford, OX2 6QS UK Keywords: elephants, genetic censusing, microsatellites Received 14 June 2000; revision received 10 August 2000; accepted 4 September 2000 Correspondence: Lori Eggert. Fax: (858) 534 – 7108; E-mail: leggert@biomail.ucsd.edu Poaching and rapid human population growth have put intense pressure on elephant populations, especially in the forests of west and central Africa. Conversion of rainforest to agriculture has resulted in the fragmentation and isolation of forest elephant populations in small reserves. Effective management of these populations will require information about census size, sex ratio, and the amount and distribution of genetic diversity. Although we can count savannah elephants from the ground or air, forest elephants are difficult to see in the dense vegetation and censusing them requires using indirect methods (Barnes & Jensen 1987). For our genetic characterization of African forest elephant populations, we developed a panel of microsatellite loci. Genomic DNA was extracted from tissue samples of four unrelated African zoo elephants using the QIAamp Blood and Tissue Kit (Qiagen), then pooled in equal concentrations. We digested 10 µg with MboI and ligated fragments of 200 – 500 bp into M13mp18 (Rassmann et al. 1991). Transformation of competent DH5αF′ Escherichia coli (GibcoBRL) was performed by electroporation. Cells were plated on YT media and plaques were replicated on nylon filters (MSI). The probes (CA)15 and (GA)15 were labelled with [γ32P]-dATP and hybridized with the plaque lifts. We selected 40 (1.6%) colonies that were strongly positive and isolated the DNA using the QiaPrep Spin Miniprep Kit (Qiagen). We sequenced these using the Sequenase 2.0 kit (Amersham Life Science), and determined that 32 contained microsatellites, 12 of which were uninterrupted and had sufficient flanking regions for primer design. Primer pairs were designed using primer 0.5 (Whitehead Institute, Cambridge, USA). We tested our primers on 10 African savannah elephants from the Frozen Zoo® of the Zoological Society of San Diego. Three primer sets revealed monomorphic loci and three were 2224 P R I M E R N O T E S Table 1 Characteristics of African elephant (Loxondonta africana) microsatellite loci and their use in the Asian elephant (Elephas maximus). Repeat motifs, primer sequences, allele numbers and sizes for elephants from the Frozen Zoo® and the forest elephants of Kakum National Park, expected (HE) and observed (HO) heterozygosity values for the Kakum elephants, and annealing temperatures for the loci developed in this study. Annealing temperatures (Ta) shown are for African elephants, these were lowered by 2 °C when amplifying DNA from Asian elephants. GenBank accession nos for the sequences of clones are AF 311670 –75 Allele sizes No. of alleles Kakum N. P. L. africana E. maximus HE HO Ta (°C) Locus Repeat Motif Primer sequences L. africana E. maximus LA1 (CA)10(TA)5 139–149 —* 6 — —* — 53 LA2 ((CA)6(CGTA))2(CA)6 227–241 226–234 3 4 0† 0 58 LA3 (CA)10 165–171 166–172 3 3 0.521 0.527 55 LA4 (CA)12(CGTA)4(CA)7 117–137 111–117 11 4 0.760 0.747 54 LA5 (CA)13 130–154 142–144 7 2 0.575 0.377‡ 52 LA6 (CA)13 F: TGGGTTGTTCCACCCTCTAC R: GTAACCGGGCAAGTGTGTG F: CTTGGTGGGAGTCATGACCT R: GGAGAAATGACTGCCCGATA F: TACTCTGCTCCTCTGCCTATCC R: GCAGAATTTTGGTCTTGGAGG F: GCTACAGAGGACATTACCCAGC R: TTTCCTCAGGGATTGGGAG F: GGGCAGCCTCCTTGTTTT R: CTGCTTCTTTCATGCCAATG F: AAAATTGACCCAACGGCTC R: TCACGTAACCACTGCGCTAC 158–214 155–159 7 3 0.542 0.563 57 *Locus does not amplify. †Locus monomorphic in this population. ‡Significant deviation from Hardy Weinburg expectation (P = 0.002, tested in genepop 3.2a, Raymond & Rousset 1995). unusable. To test our primers on Asian elephants (Elephas maximus), we used 12 samples from the Frozen Zoo®. Finally, our primers were screened on dung samples from 86 African forest elephants at Kakum National Park, Ghana. DNA from these samples was extracted using the protocol of Boom et al. (1990). To minimize the potential for allelic dropout or spurious alleles, genotypes were obtained from two different extractions of each sample in a ‘multiple tubes’ approach (Taberlet et al. 1996). Amplifications were performed in 10 µL volumes containing 20 –50 ng of template DNA, 1 µL reaction buffer (Promega), 0.2 µm radioactively labelled forward primer, 0.2 µm reverse primer, 0.2 µm dNTP mix, 1.5 mm MgCl2 and 0.5 U Taq DNA polymerase (Promega). Using a Hybaid thermocycler, the profile consisted of a denaturation step at 94 °C for 3 min, followed by 35 – 40 cycles of 94 °C denaturation for 30 s, 1 min of primer annealing at the temperatures shown in Table 1, and 1 min of primer extension at 72 °C. Alleles were separated in a 6% polyacrylamide gel, visualized by autoradiography, and scored by comparison with an M13 length standard. All six loci were highly polymorphic in African elephants with between three and 11 alleles (Table 1). The smaller number of alleles found in our Asian elephant samples is not surprising, as it is generally assumed that microsatellite loci will be more polymorphic in the species from which they are cloned than in related species (Ellegren et al. 1995). As the Frozen Zoo® samples do not represent natural populations, only expected and observed heterozygosity values for the Kakum elephants are shown. Previous work has shown that African forest elephants are genetically divergent from the savannah subspecies (Barriel et al. 1999), which may explain why locus LA1 could not be amplified in the Kakum samples. The significant deviation from the expected frequency of heterozygotes for locus LA5 may indicate the presence of one or more null alleles. However, we have no family groups with which to test for these. Although African and Asian elephants diverged from a common ancestor approximately 5 mya (Maglio 1973), five of the six primer pairs amplify in Asian elephants. While some of the loci have less alleles in Asian than in African elephants, we believe that these loci will be useful for population studies in both species. Acknowledgements We thank the Zoological Society of San Diego for archived samples of zoo elephants. Funding for this project was provided by a grant from the Academic Senate of the University of California San Diego. References Barnes RFW, Jensen KL (1987) How to count elephants in forests. Technical Bulletin of the African Elephant and Rhino Specialty Group, 1, 1– 6. Barriel V, Thuet E, Tassy P (1999) Molecular phylogeny of Elephantidae, extreme divergence of the extant forest African elephant. Comptes Rendus de l’Academie des Sciences Series III Sciences de la Vie, 322, 447–454. Boom RC, Sol JA, Salimans MMM, Jansen CL, van Dillen Werthein PME, van der Noordaa J (1990) Rapid and simple method for purification of nucleic acids. Journal of Clinical Microbiology, 23, 495–503. Ellegren H, Primmer CR, Sheldon BC (1995) Microsatellite ‘evolution’: directionality or bias? Nature Genetics, 11, 360–392. Maglio VJ (1973) Origin and evolution of the Elephantidae. Transactions of the American Philosophical Society, 63, 1–148. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 P R I M E R N O T E S 2225 Rassmann K, Schlötterer C, Tautz D (1991) Isolation of simplesequence loci for use in polymerase chain reaction-based DNA fingerprinting. Electrophoresis, 12, 113–118. Raymond M, Rousset F (1995) genepop (Version 3.2a): a population genetics software for exact tests and ecumenicism. Journal of Heredity, 86, 248– 249. Taberlet P, Griffin S, Goossens B et al. (1996) Reliable genotyping of samples with very low DNA quantities using PCR. Nucleic Acids Research, 24, 3189–3194. PRIMER 1157 2000 Graphicraft 1932 NOTEs Limited, Hong Kong The estuarine teleost, Acanthopagrus butcheri (Sparidae), shows low levels of polymorphism at five microsatellite loci E . S . YA P ,* P. B . S . S P E N C E R ,† J . A . C H A P L I N * and I . C . P O T T E R * *School of Biological Sciences and Biotechnology, Murdoch University, Perth, 6150, Western Australia †Perth Zoo, South Perth, 6951, Western Australia Keywords: Acanthopagrus butcheri, estuaries, microsatellite polymorphism, Sparidae Received 6 July 2000; revision received 2 September 2000; accepted 4 September 2000 Correspondence: E. S. Yap. Fax: + 61– 8 -9360– 6303; E-mail: esyap@central.murdoch.edu.au The black bream, Acanthopagrus butcheri, is a member of the family Sparidae that is found throughout southern Australia (Kailola et al. 1993). Information on the population genetic structure of this species is of value for two reasons. First, black bream is one of a relatively small number of teleosts that typically spends its entire life-cycle within estuaries. Thus, studies of this species can be used to test hypotheses about the role that estuaries play in promoting genetic differentiation in those teleosts that breed within these systems (e.g. Chaplin et al. 1998). Second, such information has important implications for the management of this species, which supports significant commercial and recreational fisheries in three Australian states (Kailola et al. 1993) and is a target of a developing inland aquaculture industry in south-western Australia. Microsatellite markers are particularly useful for elucidating the details of the population genetic structure of species that show low levels of polymorphism in other types of markers, such as allozymes and mitochondrial DNA (e.g. Shaw et al. 1999). The black bream is one such species (Chaplin et al. 1998; E. Yap et al. unpublished data). Here, we describe the isolation and characterization of microsatellite loci from black bream and then assess the levels of polymorphism at five loci. Genomic DNA was extracted from the muscle tissue of black bream using CTAB buffer and a phenol– chloroform extraction protocol. The DNA was digested to completion with Sau3A and size fractionated in an agarose gel. Fragments of 200– 600 bp were excised from the gel, purified and ligated into the BamHI site of the vector pGEM 3Zf(+) (Promega). The ligation products were transformed into ElectroMAX-DH10B cells (Life Technologies), which were then plated onto agar © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 containing ampicillin (100 mg/mL), IPTG and x-gal. The recombinant colonies (n = 446) were picked into 96-well microtitre plates, grown at 37 °C, vacuum blotted onto Hybond-N nylon membranes (Amersham), and screened with (CA)12, (AG)12, (TCC)5, (GACA)4, (GATA)4 and (GAA)5 oligo probes end-labelled with [α32P]-dATP. Plasmid DNA was isolated from 12 positive clones and then subjected to dye-terminator cycle sequencing. The sequencing products were electrophoresed and the sequences of the plasmids and inserts were determined using an ABI 373 Sequencer (Perkin Elmer). All inserts contained microsatellite loci. Primers, for use in polymerase chain reaction (PCR), were designed for six of these loci on the basis that they contained 15 or more repeat units and that the sequencing of their flanking regions was sufficient to permit primers to be generated. Five of the primer pairs amplified scorable alleles at the microsatellite loci (Table 1). The ‘optimised’ conditions for PCR amplification of these loci were: (i) 15 µL reaction mixture containing 50–100 ng DNA template, 1.5 mm MgCl2, 0.20 mm of each dNTPs, 20 – 40 nm of each primer, with 25% of the forward primer end-labelled with [γ33P]-ATP, 0.05 U Taq DNA polymerase, and 10 mm Tris-HCl with 50 mm KCl; and (ii) PCR profiles with an initial 5 min denaturation at 94 °C, followed by 26 cycles of 30 s denaturation at 94 °C, 30 s at annealing temperature (Table 1) and 90 s extension at 72 °C, and a final 7 min extension at 72 °C. Amplified alleles were resolved on a 6% denaturing polyacrylamide gel and their sizes estimated using pUC18 DNA sequencing standards. The levels of polymorphism at the five microsatellite loci were assessed using at least 40 black bream from nine water bodies in Western Australia and 10 individuals from Gippsland Lake in south-eastern Australia. One locus (pAb4D5) was monomorphic in all samples, while another locus (pAb2D11) was polymorphic only within the samples from south-eastern Australia (Table 1). Only one (pAb2B7) of the remaining three loci, which were polymorphic in all 10 populations, was represented by a total of more than seven alleles and had an expected heterozygosity of greater than 0.56 (Table 1). Thus, the black bream appears to contain relatively low amounts of microsatellite polymorphism, especially in Western Australia, and particularly in comparison with, for example, two species of marine sparid (see Takagi et al. 1997; Batargias et al. 1999). Nevertheless, the four polymorphic loci have revealed greater amounts of variation in black bream than allozyme genes (see Chaplin et al. 1998). In addition, the genotype frequencies at each of the pAb1H1, pAb2B7 and pAb2A5 loci, in each of samples of 38 or more black bream from nine water bodies in Western Australia, did not show any statistically significant departures from those expected under Hardy–Weinberg equilibrium conditions. The four polymorphic microsatellite loci should, therefore, be useful for addressing population-level questions about the black bream. Acknowledgements This work was made possible by the provision of a postgraduate scholarship to ESY from the Australian Agency for International Development (AusAID), and a Special Research Grant to JAC from Murdoch University. 2226 P R I M E R N O T E S Table 1 Characteristics of five microsatellite loci in samples of black bream (Acanthopagrus butcheri) from nine water bodies in Western Australia and from the Gippsland Lakes in Victoria, south-eastern Australia. The Western Australian samples are the same as those used by Chaplin et al. (1998) Locus GenBank accession no. pAb1H1 AF284351 pAb2B7 AF284352 pAb4D5 AF284353 pAb2A5 AF284354 pAb2D11 AF284355 Primer sequence (5′−3′) F: GGCTTTCATTTCCCCATTTGTG R: CACCTTTCTCCACGCCATAAA F: GGTGCGTGCATTGTTAATGTGT R: GATCTGCTTTCCTTTGACTCAGC F: ACCTCTTCATCTGCGTGACATCT R: GACAACACCCTCACTCAGCTGA F: AGTTACTTTCTCCAGAGTGGCGC R: GGCAACAGATAAGCACTGAGCATA F: CGGTCCAGTTTCACTCTGATGTT R: AACTGCTGTCATCGCCCTGTT Repeat unit* Ta (°C) Size range (bp) (TG)15 63 132–148 (TG)24 65 98 –128 (TG)60 54 (TG)19 (TG)15 No. of alleles n HE HO 5 268 0.37 0.44 14 274 0.70 0.72 199 1 50 0 0 63 105–119 7 273 0.56 0.62 65 106–110 4† 50 0.11 0.08 *determined from the sequenced insert; †polymorphic only within samples from the Gippsland Lakes. n, is the total number of individuals assayed per locus; Ta, is the optimal annealing temperature of each primer pair; HE, is the expected heterozygosity, calculated as 1 − Σ(fi2), where fi is the frequency of the ith allele; and HO , is the observed heterozygosity. References Batargias C, Dermitzakis E, Magoulas A, Zouros E (1999) Characterization of six polymorphic microsatellite markers in the gilthead seabream, Sparus aurata (Linneaus 1758). Molecular Ecology, 8, 897– 898. Chaplin JC, Baundais GA, Hill HS, McCulloch R, Potter IC (1998) Are assemblages of black bream (Acanthopagrus butcheri) in different estuaries distinct? International Journal of Salt Lake Research, 6, 303– 321. Kailola PJ, Williams MJ, Steward PC et al. (1993) Australian Fisheries Resources. Bureau of Resource Sciences and Fisheries Research and Development Corporation, Canberra. Shaw PW, Pierce GJ, Boyle PR (1999) Subtle population structuring within a highly vagile marine invertebrate, the veined squid Loligo forbesi, demonstrated with microsatellite markers. Molecular Ecology, 8, 407–417. Takagi M, Taniguchi N, Cook D, Doyle RW (1997) Isolation and characterisation of microsatellite loci from red sea bream Pagrus major and detection in closely related species. Fisheries Science, 63, 199–204. Graphicraft 00 PRIMER 2000 1160 912 NOTEs Limited, Hong Kong Fifty Seychelles warbler (Acrocephalus sechellensis) microsatellite loci polymorphic in Sylviidae species and their cross-species amplification in other passerine birds D . S . R I C H A R D S O N , F. L . J U RY , D . A . D AW S O N , P. S A L G U E I R O , J . K O M D E U R * and T. B U R K E Department of Animal and Plant Sciences, University of Sheffield, Sheffield, S10 2TN, UK, *Zoological Laboratory, University of Groningen, PO Box 14, 9750 AA Haren, The Netherlands Keywords: Acrocephalus, microsatellite, PCR, Seychelles warbler, Sylviidae Received 21 July 2000; revision received 2 September 2000; accepted 4 September 2000 Correspondence: T. Burke. Fax: + 44 (0) 114 222 0002; E-mail: T.A.Burke@Sheffield.ac.uk The cooperatively breeding Seychelles warbler, Acrocephalus sechellensis, is a rare endemic of the Seychelles islands. By 1959, anthropogenic disturbance had pushed this species to the verge of extinction and only 26 individuals remained, confined to the island of Cousin. The population has since recovered and has been the focus of intense study since 1985 (e.g. Komdeur 1992; Komdeur et al. 1997). We required a set of microsatellite markers to enable studies of mate choice, reproductive success and fitness. Genetic variability is relatively low within this species, possibly due to the recent population bottleneck. Consequently, many microsatellites had to be isolated and screened to provide sufficient polymorphic loci to enable parentage assignment and pedigree construction. We isolated 63 microsatellite loci from the Seychelles warbler and tested for their polymorphism in this and five other species of Sylviidae. We also examined the utility of a subset of these loci in 16 other passerine birds. DNA was extracted following Bruford et al. (1998). A genomic library enriched for (CA)n, (GA)n and (TTTC)n was prepared as described by Armour et al. (1994) using modifications suggested by Gibbs et al. (1997). DNA reactions were performed in a 10-µL volume containing 10 – 50 ng DNA, 1.0 µm of each primer, 0.2 mm of each dNTP, 0.05 units Taq DNA polymerase (Thermoprime Plus, Advanced Biotechnologies) and 1.0 – 2.0 mm MgCl2 (Table 1) in 20 mm (NH4)2SO4, 75 mm Tris-HCl pH 9.0, 0.01% (w/v) Tween. Polymerase chain reaction (PCR) amplification was performed in a Hybaid Touchdown thermal cycler. Initially, a touchdown cycle was performed with a reaction profile of 95 °C for 3 min, then 94 °C for 30 s, © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Table 1 Characterization of 50* polymorphic microsatellite loci from the Seychelles warbler (Acrocephalus sechellensis), and their polymorphism in five other members of the Sylviidae family Repeat motif Primer sequence (5′−3′) Ase2 AJ287385 [ (GAAA)2GCAA]3 Ase3 AJ287386 (CA)14CCA Ase4 AJ287387 (CA)11 Ase5 AJ287388 AAA(CA)12AAA Ase6 AJ287389 (CA)3G(CA)17 Ase7 AJ287390 (CT)13 Ase8 AJ287391 (GT)4TTT(GT)7 Ase 9 AJ287392 (CA)15 Ase10 AJ287393 (CCTTCCCT)7 Ase11 AJ287394 (AC)14 Ase12 AJ287395 (CA)11 Ase13 AJ287396 (GT)11 Ase16 AJ276374 (TCTCC)13 Ase18 AJ276375 (GT)12 Ase19 AJ276376 (CA)4GA(CA)5 Ase20 AJ276377 (CTTC/CTTT)10 Ase21 AJ276378 (CTTTT)2CTC(TTTC)8 Ase22 AJ276379 (GT)13 Ase25 AJ276382 (GAAA)31 F: TTGACAGAGTGTTATTCAATGTG R: GAGCAGATAATAGACCTTGCT F: ACAGGTATGGCGCTCAAGTC R: CTGAATCTTACACAGGAGACCGT F: TCTCCATCATCACCACAAAGC R: TTCCCATTGCCCTAGTTATTCCA F: TGAAACAAAATGGGATGGTCC R: CCTTTCTCGGAACTGATTGCTT F: TAAAAGCCAGCAGTGGAGCC R: CGAGCTTGCAGGGTTTCCT F: AATCAACTTCAAATGCTCACAG R: ACTACATGACTCCAGGCTCAG F: TACCTCTCCTTGGCTGAGCA R: CCAGCCCTAGCTGTTTCACC F: GACTGAAGTCCTTTCTGGCTTC R: CACCAGGAATACAAGTCCATTG F: CATTGGGGTACTATGGAAAGACC R: TCCTGAGTGGAAGGAACATAGG F: TCCCCAAATCTCTCAATTCC R: AGTTCTAAGCCTGCCTGTGC F: TCAAGGAAACACAACTACAGCC R: TTTCCTCACAGCCTTGACTG F: TGTGCTCCTCTGCTTTCC R: CAGATGGCCAGTGTTAGTCC F: TCAGTTCCTGAGTAAATGTCTC R: TGAATTACCCCTAAATACCTG F: ATCCAGTCTTCGCAAAAGCC R: TGCCCCAGAGGGAAGAAG F: TAGGGTCCCAGGGAGGAAG R: TCTGCCCATTAGGGAAAAGTC F: TCTAAAGCTGCCTGCCAGAA R: GCGGTTGCAGTGGACTTG F: TTAGAACCATTTGATAGTTGCCAC R: ATGGGTTTCTTGGGGAAGAG F: TGAACCATTGTCACCAACAC R: GCTTTAGTTCAGATGCCCAG F: GATGGCTATATGCTTCAAATGC R: TTGAAAGCCTTAAAGTGGGA Product size† (bp) Number of alleles/ number of individuals Ta (°C) MgCl2 conc. (mm) 60 1.5 97 2/7 60 1.5 101 60 1.0 61 HE CRW AW GRW EMW WW 0.71 0.50 1/6 0 1/3 3/2 1/2 3/7 0.86 0.60 1/6 4/4 1/3 1/2 1/2 103 2/25 0.40 0.37 0 3/4 1/3 0 0 1.0 110 1/7 0.00 0.00 1/6 1/4 1/3 1/2 2/2 60 1.5 119 4/25 0.76 0.70 1/6 2/4 1/3 0 0 60 1.5 123 2/7 0.83 0.53 2/6 1/4 2/3 1/2 0 TD 1.5 125 1/7 0.00 0.00 3/6 2/4 1/3 2/2 1/2 60 1.5 125 3/25 0.40 0.44 3/6 5/4 5/3 2/2 1/2 TD 1.5 127 3/25 0.64 0.56 9/5 0 1/3 1/2 0 60 1.5 128 2/5 0.40 0.53 7/6 4/4 5/3 3/2 0 60 1.5 128 1/7 0.00 0.00 4/6 4/4 1/3 3/2 2/2 62 1.5 132 3/25 0.52 0.54 5/5 7/4 1/3 2/2 1/2 58 1.5 155 4/7 0.70 5/6 0 6/3 1/2 0 60 1.5 176 3/25 0.56 0.50 1/6 5/4 3/3 1/2 3/2 60 2.0 177 4/8 0.88 0.64 3/6 3/4 1/3 4/2 3/2 TD 1.5 178 1/7 0.00 0.00 1/6 7/4 1/3 1/2 0 58 2.0 180 1/7 0.00 0.00 9/6 5/4 1/3 1/2 2/2 58 1.5 181 2/6 0.50 0.53 1/6 0 0 1/2 0 58 1.5 187 5/25 0.76 0.74 1/6 6/4 0 2/2 0 SW HO Number of alleles/number of individuals tested in 100.0 P R I M E R N O T E S 2227 Locus EMBL accession number © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Locus EMBL accession number Repeat motif Primer sequence (5′−3′) Ase26 AJ276383 (CTC)3(TC)12 Ase27 AJ276384 (TTTC)16 Ase29 AJ276386 (AC)7TTTG(AC)6 Ase32 AJ276635 (GT)13(TCAC)2(GT)9 Ase33 AJ289865 (AT)10 Ase34 AJ276636 (CT)11 Ase35 AJ276637 (GT)10 Ase36 AJ276638 (TGTGG)7 Ase37 AJ276639 (AC)9 Ase38 AJ276640 (CA)15 Ase40 AJ276642 (GT)10 Ase42 AJ276644 (GT)4(AT)6(GT)8(AT)2 Ase43 AJ276645 (TA)3(CA)8(TA)5 Ase44 AJ276646 (GT)18 Ase46 AJ276775 (TG)13 Ase47 AJ276776 (CA)10 … (CA)4 Ase48 AJ276777 (CCTTCT)6 Ase49 AJ276778 (AC)10 Ase50 AJ276779 (CA)12 F: GCTGGCCTTGCAAAAACTTC R: AACACCTCCCTGTCCCTGC F: TTAACATTGCATGCTCCTGC R: AGTCAAGGTACAGGCTAGATAGCC F: GATCAGTTTGGAGACGTTTTCT R: ACAGAGCCATAAGGAATGTGC F: AATGAGCAATACCATGACAGC R: GATCTTTCAGTCAGGAACAAGC F: CTTTGGAATGCCAGGCTGCT R: TGCTGGAACCACAGGACTT F: GTTAATTCTTTTGGCCCTCAGC R: GGAGACACCACACCAATGC F: GTCCTTGGTCCTTAGCATCTGT R: GCTCCTGTTGTTCTGGGAATAG F: AAGTTCCATGGGGTGAATGC R: GAGCGTGTTCCTCCAATTCC F: TAATTCATGGAGAAGCCCAG R: TCAAAACAACAGTTTTCACAGC F: ATCCGAGAACCCAATCACTT R: GCAGCATTACAGTCTCAAAGAAC F: CACTGCTCCAGGCACTCTG R: TCCAAGGCACACAAAGGTG F: CATGGGTAGGTTGGGATGTC R: AGGTGAGGGTATGCAAACATG F: ATTGTGTGGGATTTGCAT R: TTGCTGTGCAGTTTGCTTTT F: TTCCCGTAATTATGACCTCTCTTG R: ACCAGAACTTGTTGTCTGGGAG F: CTGGCTGTATCTTGGTGTGC R: CAGTGTTTTAGGTCTCCTGCTG F: GATCACATTTGGCATTTACTGAT R: ACTCTTTAGGGCAAGGCACT F: TTTATTTCCTGGACTGGAACAATC R: GAACATTGGGCTACTGGGC F: CCCCTGAAGTGTCCAACG R: ACTTTCCCAGCACATCTTGC F: CTGTGGAATGCTGTCTGGC R: ATGGACTCCCGTCTAACTTGC Ta (°C) MgCl2 conc. (mm) Product size† (bp) 60 1.5 60 Number of alleles/ number of individuals Number of alleles/number of individuals tested in SW HO HE CRW AW GRW EMW WW 203 1/7 0.00 0.00 1/6 5/4 1/3 2/2 2/2 1.0 204 4/25 0.64 0.60 1/6 1/4 3/3 2/2 1/2 62 1.5 207 2/7 0.14 0.14 1/6 1/4 1/3 1/2 2/2 58 1.5 218 1/7 0.00 0.00 1/6 5/4 0 0 0 TD 1.5 220 1/7 0.00 0.00 1/6 4/4 2/3 1/2 1/2 60 1.5 220 1/7 0.00 0.00 3/5 1/4 3/3 4/2 3/2 58 1.5 224 3/25 0.44 0.62 1/6 1/4 0 2/2 0 60 1.5 225 2/5 0.20 0.20 1/6 1/4 1/3 1/2 0 58 1.5 226 3/25 0.32 0.37 2/6 1/4 0 4/2 0 58 2.0 226 2/4 0.50 0.43 0 3/4 1/3 3/2 1/2 58 1.5 230 1/7 0.00 0.00 3/6 3/4 1/3 1/2 1/2 62 1.5 243 2/25 0.32 0.27 1/6 1/4 4/3 1/2 2/2 TD 1.5 250 1/7 0.00 0.00 2/6 3/4 1/3 1/2 2/2 TD 1.5 250 1/7 0.00 0.00 1/6 4/4 3/3 1/2 2/2 62 1.5 265 3/25 0.24 0.48 1/6 2/4 1/3 1/2 1/2 TD 1.5 267 1/7 0.00 0.00 4/6 0 1/3 1/2 2/2 58 1.0 270 4/25 0.56 0.53 7/5 7/4 5/3 3/2 0 58 1.5 272 2/7 0.00 0.26 1/6 2/4 1/3 1/2 2/2 60 1.5 272 1/7 0.00 0.00 1/6 6/3 2/3 2/2 2/2 2228 P R I M E R N O T E S Table 1 Continued © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Table 1 Continued Locus EMBL accession number Repeat motif Primer sequence (5′−3′) Ase51 AJ276780 (CA)12 Ase52 AJ276781 (CA)9(CA)5 Ase53 AJ276782 (CTT)22 (CTCCTT)10 Ase55 AJ276784 (GT)9 Ase56 AJ276785 (GT)18 Ase57 AJ276786 (AC)14 Ase58 AJ276787 (CTTTTT)27 Ase60 AJ276789 (GT)9GG(GT)8 Ase61 AJ276790 (GAAAAA)13 Ase62 AJ276791 (CT)2(GT)8 Ase63 AJ276792 (GAGAAA)8(GA)7 Ase64 AJ276793 (AGGG)9 (ATGG)12 F: AATTCCCCTAGACAGGCAGC R: TCACTGGAGAGCCAAATTCC F: TCTTAGCCTGCACTCATTTCA R: CAGTCACCGTAAGTTCATAGGC F: ATGGAGAATTCTGGGTGCTG R: CCCAATAATGAGGTAACACCAA F: GTGTGGACTCTGGTGGCTC R: TCCCAAAGCACTCAAACTAGG F: TTCACTGAGAAGTGAGAATGTG R: GTCCTTGATTGATTACAGGCT F: GCAAGTGCAGATGTTTCCCT R: CCAAAGCAGGACAATGCTG F: ATTCCAGGGATTGGGCAG R: CTCAAAGCGAAATTGAGCAGT F: CATGAAAAGGAACTCTCCAGC R: TTCCATCTCTGTTCTACTGCG F: AGGATTTTTAATGGGATATACACATCTG R: AGCCACATTTTAGCCCACAG F: TCGCCAGGTCGTGTGTAGTC R: CAAAACCGTGTCGGGGAG F: TTTGGGGTTTAGGAATAGCAGA R: GGCTTCAGCCTGAGAAAGTC F: CCACCTTTCATACTGGGGAG R: TTCAGCCAGTCAGTGTAGCC Ta (°C) MgCl2 conc. (mm) Product size† (bp) 60 1.5 60 Number of alleles/ number of individuals Number of alleles/number of individuals tested in SW HO HE CRW AW GRW EMW WW 277 1/7 0.00 0.00 1/6 7/4 2/3 2/2 1/2 1.5 278 1/7 0.00 0.00 1/6 2/4 1/3 1/2 1/2 60 1.5 285 2/7 0.43 0.54 1/6 8/4 0 0 0 62 1.5 292 1/7 0.00 0.00 1/6 6/4 2/3 2/2 2/2 60 1.5 298 3/25 0.44 0.40 5/6 5/4 2/3 3/2 0 TD 1.5 299 1/7 0.00 0.00 6/6 3/4 4/3 1/2 0 60 1.0 311 5/25 0.76 0.76 1/6 7/4 5/3 4/2 1/2 62 1.5 353 1/7 0.00 0.00 0 5/4 4/3 1/2 3/2 54 2.0 369 2/5 0.40 0.36 0 0 3/3 0 0 58 1.5 372 1/7 0.00 0.00 1/6 1/4 1/3 2/2 0 60 1.0 400 2/7 0.29 0.26 2/6 8/4 2/3 4/2 1/2 TD 1.5 412 2/8 0.50 0.40 7/6 1/4 3/3 1/2 1/2 P R I M E R N O T E S 2229 *An additional 13 loci were monomorphic in all species tested (EMBL accession numbers: AJ287384, AJ287397, AJ287398 AJ276380, AJ276381, AJ276385, AJ276387, AJ276634, AJ276641, AJ276643, AJ276647, AJ276783, AJ276788). †Size in cloned allele. SW, Seychelles warbler, Acrocephalus sechellensis; CRW, clamorous reed warbler, Acrocephalus stentoreus australis (M. Berg, personal communication); AW, aquatic warbler, Acrocephalus paludicola (P. Hedrich, personal communication); GRW, great reed warbler, Acrocephalus arundinaceus (B. Hansson, personal communication); EMW, European marsh warbler, Acrocephalus palustris (B. Hansson, personal communication); WW, willow warbler, Phylloscopus trochilus (B. Hansson, personal communication). Ta, annealing temperature; TD, Touchdown cycle; HO, observed heterozygosity; HE, expected heterozygosity; 0, no product detected. Number of alleles/Number of individuals tested (n = 4 unless stated) © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 Family* Species Ase8 Ase9 Ase13 Ase18 Ase19 Ase29 Ase34 Ase37 Ase40 Ase42 Ase43 Ase46 Ase48 Ase55 Ase56 Maluridae Pomatostomidae Laniidae Corvidae Cinclidae Sturnidae Certhiidae Paridae Paridae Hirundinidae Pycnonotidae Zosteropidae Sylviidae Sylviidae Sylviidae Sylviidae Sylviidae Sylviidae Sylviidae Nectariniidae Passeridae Fringillidae Superb fairy-wren, Malurus cyaneus White-browed babbler, Pomatostomus superciliosus Loggerhead shrike, Lanius ludovicianus Azure-winged magpie, Cyanopica cyana White-throated dipper, Cinclus cinclus European starling, Sturnus vulgaris Winter wren, Troglodytes troglodytes Blue tit, Parus caeruleus Long-tailed tit, Aegithalos caudatus Sand martin, Riparia riparia White-spectacled bulbul, Pycnonotus xanthopygos Seychelles grey white-eye, Zosterops modestus Aquatic warbler, Acrocephalus paludicola Sedge warbler, Acrocephalus schoenobaenus European marsh warbler, Acrocephalus palustris Great reed warbler, Acrocephalus arundinaceus Clamourous reed warbler, Acrocephalus stentoreus australis Seychelles warbler, Acrocephalus sechellensis Willow warbler, Phylloscopus trochilus Seychelles sunbird, Nectarinia dussumieri Seychelles fody, Fodia sechallarum European greenfinch, Carduelis chloris 0 1 0 0 0 1 7/6 0 — 0 0 1 2/4 — 2/2 1/4 3/6 1/7 1/2 0 0 0 1 1 0 1 1 1 2/6 1 — 1 1 1 5/4 1/8 2/2 5/4 3/6 3/25 1/2 1 1 1 1 1 1 1 1 1 2/6 1 — 1 1 1 7/4 — 2/2 1/4 5/5 3/25 1/2 1 1 1 0 1 1 1 1 1 5/6 1 1 0 1 1 5/4 16/40 1/2 3/4 1/6 4/25 3/2 1 1 1 0 1 0 0 0 1 3/6 0 — 1 1 0 3/4 — 4/2 1/4 3/6 4/8 3/2 0 0 1 1 1 0 0 1 1 5/6 1 1 1 1 1 1/4 — 1/2 1/4 1/6 2/7 2/2 1 1 1 1 1 1 1 1 1 2/6 1 1 1 1 1 1/4 — 4/2 3/4 3/5 1/7 3/2 1 1 1 0 1 0 0 0 0 3/6 0 16/680 0 0 1 1/4 — 4/2 0 2/6 3/25 0 0 0 0 1 1 0 0 1 1 2/5 0 1 1 1 1 3/4 — 1/2 1/4 3/6 1/7 1/2 1 1 1 0 1 1 1 1 1 — 1 1 1 0 1 1/4 5/8 1/2 4/4 1/6 2/25 2/2 1 1 1 1 1 1 1 1 1 3/6 1 0 1 1 1 3/4 1/8 1/2 1/4 2/6 1/7 2/2 1 1 1 1 1 0 1 1 1 3/6 0 1 1 1 1 2/4 — 1/2 1/4 1/6 3/25 1/2 0 0 1 0 1 0 0 0 1 — 0 0 1 0 1 7/4 — 3/2 5/4 7/5 4/25 0 0 1 1 0 1 0 1 1 1 5/6 1 — 1 1 1 6/4 — 2/2 2/4 1/6 1/7 2/2 1 1 1 0 0 0 1 0 1 6/6 0 — 0 0 0 5/4 — 3/2 2/4 5/6 3/25 0 1 0 1 Number of species tested for amplification % of species in which a product was amplified Number of species tested for variability % of species (tested for variability) with ≥3 alleles 20 50 7 29 21 95 8 50 20 100 7 43 21 90 8 75 20 55 7 86 21 86 7 14 21 100 7 57 21 38 6 67 21 86 7 29 21 90 7 29 22 95 8 25 21 80 7 14 20 52 5 100 20 90 7 29 20 50 6 83 *Following Sibley & Monroe (1990), except Seychelles warbler which follows Komdeur (1992). —, sample not tested; 0, no reliable product; 1, product visualized on agarose gel (not tested for variability). 2230 P R I M E R N O T E S Table 2 Cross-species utility of 15 Seychelles warbler (Acrocephalus sechellensis) microsatellite loci in 21 other passerine birds P R I M E R N O T E S 2231 annealing temperature X for 45 s, 72 °C for 45 s for two cycles each at X = 60 °C, 57 °C, 54 °C, 51 °C then 25 cycles at X = 48 °C, followed by 72 °C for 5 min. To optimize the PCR amplification of the loci found to be polymorphic, further PCRs consisted of one cycle at 95 °C for 3 min then 35 cycles at 94 °C for 1 min, annealing temperature (Table 1) for 30 s, 72 °C for 45 s, followed by 72 °C for 5 min. For the cross-species amplifications, a touchdown cycle was performed as above. PCR products were visualized on a 0.8% agarose gel stained with ethidium bromide. When testing for polymorphism, PCR products were run on 6% polyacrylamide gels and visualized by staining with silver (Promega) or by autoradiography (after PCR with one of the primers end-labelled with [γ 33P]dATP; Sambrook et al. 1989). We developed primers for 63 microsatellites, of which 50 were polymorphic in at least one of the tested species of Sylviidae (Table 1). Thirty loci were polymorphic, displaying up to five alleles, in a test panel of up to 25 unrelated Seychelles warblers. There was no significant difference at any locus between the observed and expected heterozygosity, though these comparisons were of limited power. All 50 loci found to be polymorphic in the Sylviidae were tested for polymorphism in six unrelated individuals of the winter wren, Troglodytes troglodytes (M. Berg, personal communication). Fifteen of the loci that were also found to be polymorphic in the winter wren were selected and tested for utility in 16 other species, representing 15 passerine families (Table 2; following Sibley & Monroe 1990). The high proportion of loci found to be polymorphic in the other Sylviidae will reduce or eliminate the need to develop new primers for future studies of these species. The crossspecies amplification suggests that, after further testing, many of the primers presented here may also be useful for detecting polymorphic loci in other passerine families (Table 2). Acknowledgements We thank M. Berg, N. Chaline, B. Hansson, P. Heidrich, R.C. Marshall and D.J. Ross for contributing data on the crossutility of primers. D. Bryant, M.C. Double, B.J. Hatchwell, J.G. Martinez, N. Mundy, J. Wetton, J. Wright, S. Yezerinac and R. Zilberman kindly supplied blood or DNA samples. This work was supported by the Natural Environment Research Council. References Armour JAL, Neumann R, Gobert S, Jeffreys AJ (1994) Isolation of human simple repeat loci by hybridization selection. Human Molecular Genetics, 3, 599 – 605. Bruford MW, Hanotte O, Brookfield JFY, Burke T (1998) Multilocus and single-locus DNA fingerprinting. In: Molecular Genetic Analysis of Populations: a Practical Approach, 2nd edn (ed. Hoelzel AR), pp. 287 – 336. IRL Press, Oxford. Gibbs M, Dawson DA, McCamley C, Wardle AF, Armour JAL, Burke T (1997) Chicken microsatellite markers isolated from libraries enriched for simple tandem repeats. Animal Genetics, 28, 401– 417. Komdeur J (1992) Importance of habitat saturation and territory quality for evolution of cooperative breeding in the Seychelles warbler. Nature, 358, 493 – 495. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 Komdeur J, Daan S, Tinbergen J, Mateman C (1997) Extreme adaptive modification in the sex ratio of Seychelles warbler’s eggs. Nature, 385, 522–525. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning: a Laboratory Manual. 2nd edn. Cold Spring Harbor Laboratory Press, New York. Sibley CG, Monroe BL (1990) Distribution and Taxonomy of Birds of the World. Yale University Press, New Haven. Graphicraft 2000 1164 00 91PRIMER primer 2 Notes NOTEs Limited, Hong Kong Variable microsatellite loci in red swamp crayfish, Procambarus clarkii, and their characterization in other crayfish taxa N ATA L I A M . B E L F I O R E and B E R N I E M A Y Department of Animal Science, University of California, Davis, 95616, USA Keywords: crayfish, heterologous, microsatellites, primers, Procambarus clarkii Received 10 August 2000; revision accepted 7 September 2000 Correspondence: Natalia M. Belfiore. Fax: + 530 752 0175; E-mail: nmbelfiore@ucdavis.edu The red swamp crayfish, Procambarus clarkii, is a temperate freshwater crayfish native to the south-eastern United States. It is heavily exploited as a fishery product and is used widely in aquaculture. Its economic importance led to widespread introductions on four continents. The species has been used extensively in laboratory studies, but studies of its population biology in the wild have been rare (Huner 1988). Previous population work using allozymes found low levels of genetic variation in two Procambarus species, including P. clarkii (Busack 1988). We developed two microsatellite libraries for P. clarkii (f. Cambaridae) from which 23 variable microsatellite loci were optimized. The 18 clearest markers were tested in representative taxa of the other two crayfish families (Parastacidae and Astacidae), as well as two cambarid species in Orconectes and one congeneric species; characterization is reported here. Genomic DNA was extracted from frozen (– 80 °C) tail muscle of a red swamp crayfish (Putah Creek, Yolo County California) using the Tris sodium chloride EDTA sodium dodecyl sulphate (SDS) (TNES)-urea buffer extraction protocol (Asahida et al. 1996) with the following modifications. Approximately 200 mg tissue were added to 700 µL extraction buffer, containing 4 m urea and 0.5% SDS, and 0.035 mg Proteinase K. After overnight incubation (37 °C), samples were extracted twice with phenol:chloroform:isoamyl alcohol (25:24:1) and once with chloroform:isoamyl alcohol (24:1). DNA was precipitated with 0.3 m sodium acetate pH 5.3 in a final ethanol concentration of 67%. The pellet was washed in 70% ethanol, air or vacuum dried, and resuspended in Tris low EDTA (TLE) buffer (10 mm tris + 0.1 mm EDTA, pH 8.0). Two subgenomic libraries were created by Genetic Identification Services (Chatsworth, CA) by partially digesting whole genomic DNA with a mixture of the following restriction enzymes: BsrBR1, Locus ID Primer sequences (5′− 3′) PclG-02 PclG-03 PclG-04 PclG-07 PclG-08 PclG-09 PclG-10 PclG-13 PclG-15 PclG-16 PclG-17 © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 – 2234 PclG-24 PclG-26 PclG-27 PclG-28 PclG-29 PclG-32 PclG-33 PclG-34 PclG-35 PclG-37 PclG-45 PclG-48 F: CTC R: TGG F: CTC R: AAG F: TAT R: TCA F: CCT R: GTG F: ACG R: CCG F: TAT R: TGT F: TGC R: CAA F: CTC R: TGA F: GGC R: GGC F: CTC R: TCA F: GTC R: AAG F: CAA R: CCG F: ATA R: TCG F: AAT R: TTT F: CTC R: AGA F: GAA R: TTT F: CCC R: TGT F: TTC R: CAA F: CAG R: CTC F: TCC R: TGC F: TAA R: TAA F: ATA R: CTT F: CTG R: AGA CCC CGA TCC CTT ATC GTA CCC GGT ATA GGT GCA TGG TCA TGG TCC AGA GTG TGG GGA TTA GGG AGC GGC CGC TAG TGT CTT AAG GGC AGA AGT TTG CCA GCT GAG GGA TCC AGG TCA CTT ATA CTA TAA TGA TTG TTC ATG ATT ACC ACA AGT AGT ACC GTG AAT CTG CCT TGT CGC TCC TGG GGC ACG CCA ATG TGG AAC GAA ATT CAC CCT TCA AAG GAA GAG AAG CAT GGC CTC TGC GCG AGC ATG TGG CGT TTC AGT AGC ACC CTT GTG AAC CAC TTG AGT ATA CAA AGA AGG GCG GGA TCT TTA GGT AAA TTG CGC AGA CCA CTT TCC ATT CTA GAA GAG AGA CGC CAT ATC CGT TTT GGA GGG TAT GTC GGG TTG GTA TGA AAC TTC GAT GGC CAG GGT CAC ATT GCT TCT CCT CAT AAT TCT TTG GTT CTC TAG GTC CCT CAT CTT ATT TGT GTG ACG TGT ACC TTG TTT AGA GGG ATT CCT CAG ATG ATA ACT TAT TGT GTG TCT AGT CTG TAG TCA ACT TTT CTC GTG GGT GTC CTT TCC GTG GGC GTT TTC ATA GTC ATA ATC TTG ATG TGT GAA CA GTA TGG TAT AGG TGT TAG TGA TCA ACA GAT GTG ACG TTT CAG AAA AGA GAA AAG AGG ACG GTG GAG ATT CCG ATA CAT CCC CAC TAA GGT GGT TTC GTC TTC TCT TCT TT GAT CAG GAA TAT TT GAT CA T GT GTC TTC TGT TTA ATT CTT CCT GA ATC GTG AAA AGA AGT ACC GAG AAG AAA AT GTA TGT AG TAT C GTA GAG CTG AAA ATC CTT GAC CTC GTA TCT AAT CTG AGT TCT GC CTC No. of alleles HO HE Repeat (cloned allele) Product size range (bp) n (GATA)3GAGAA(GATA)5 216– 224 25 3 0.56 0.61 1.5 0.5 (TCTA)20 216– 420 26 12 0.73 0.89 2.0 0.5 MgCl2 (mM) Primer (µM) AGA C (TCTA)3 … (TCTA)2 … (TCTA)29 … (TCTA)2 170– 290 26 15 0.77 0.89 2.0 0.5 GG TCA (TCTA)8 100–160 19 11 0.84 0.85 1.5 0.5 GAA (GATA)16 148– 220 18 11 0.56 0.82 1.0 0.3 (TCTA)14 80 –160 20 8 0.35 0.85 1.5 0.5 (TAGA)2TA(TAGA)16 90 – 176 10 6 0.40 0.65 1.5 0.5 (TCTA)12 130– 150 17 3 0.53 0.54 1.5 0.5 (TATC)2TGTC(TATC)17TATT(TATC)3 150–185 18 12 0.78 0.85 1.5 0.5 80 –160 19 11 0.95 0.86 1.5 0.5 GAG (TCTA)18TCTC(TATC)3 TAT TAT GAT T C A GTA GAC (TCTA)14 156–190 19 8 0.84 0.78 1.5 0.5 (GATA)3AATA(GATA)24 … (AC)8T(CA)31 280– 290 3 3 1.00 0.61 1.5 0.5 (CT)5(CA)41 210– 300 16 9 0.75 0.85 1.5 0.5 8 11 0.63 0.84 1.5 0.5 (TATC)4CATC(TATC)8 (GATA)22(GA)5 AAG AAC (TATC)9 G (CT)7 … (TC)37 … (CA)15 … (CA)5 AGT TCT ACC CAA ATT C GAG CAG G TAT TTT ATC (GT)21 (CA)4CG(CA)22TA(CA)15 (GT)6AA(GT)8AA(GT)11AA(GT)5 80 –150 210– 270 20 8 0.65 0.82 1.5 0.5 95 –165 19 7 0.58 0.82 1.5 0.5 150– 250 19 14 0.74 0.91 1.5 0.5 120–180 19 11 0.63 0.85 1.5 0.5 80 –160 4 6 0.75 0.75 1.5 0.5 152–190 18 6 0.56 0.68 1.5 0.5 (CA)4CG(CA)15CG(CA)13 80 – 180 20 12 0.85 0.90 1.5 0.5 (CA)3 … (GA)6 96 – 98 16 2 0.25 0.43 1.5 0.5 146– 190 17 8 0.59 0.84 1.5 0.5 (CA)12 2232 P R I M E R N O T E S Table 1 Summary of locus data for 23 microsatellite loci developed for Procambarus clarkii. GenBank Accession nos are AF290219-AF290941. n is the number of individuals screened; individuals were drawn from two or three (where n ≤ 10) to four (where n > 10) populations. HO and HE are the observed and expected heterozygosities, respectively, calculated across all populations due to small sample sizes (Genes in Populations version 2, May et al. 1995); sample sizes precluded reasonable inference of the presence of null alleles © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234 1 1 1 0 1 0 1(2) 0 0 0 1 – (2) 0 0 1 0 0 — 1(3) 1(1) 1(3) 1(2) 1 1(3) 0 0 0 0 1 1(1) 1 2(3) 1(3) 1 0 1(2) 3 1 2 1 0 1 0 0 0 0 0 1 1 1 1 — 0 2 1 1 2 1 0 1 0 0 0 0 0 1 1 1 0 — 0 3 5 3 — 1 0 1 0 0 2(2) 1 1 3 — 2 — 2(3) 1 7 Procambarus zonangulus (n = 4) Orconectes virilis (n = 2) Orconectes rusticus (n = 2) Pasifasticus leniusculus (n = 4) Cherax quadricarinatus (n = 4) This work was supported by a University of California Toxic Substances Research and Teaching fellowship and an National PclG-02 PclG-03 PclG-04 PclG-07 PclG-08 PclG-09 PclG-13 PclG-15 PclG-16 PclG-17 PclG-27 PclG-28 PclG-29 PclG-32 PclG-37 PclG-45 PclG-47 PclG-48 Acknowledgements Species EcoRV, HaeIII, PvuII, ScaI, and StuI. An oligonucleotide linker containing a HindIII site was ligated to fragments in the range of 300–700 bp. Magnetic beads were used to capture fragments containing (CA)n or (TAGA)n. These were ligated into the HindIII site of pUC19; the products were used to transform competent Escherichia coli DH5α. Of the positive clones initially screened, 82% (n = 11) (CA)n, and 58% (n = 12) (TAGA)n contained microsatellites. We plated additional clones and amplified approximately 300 recombinant clones by colony polymerase chain reaction (PCR) using the following protocol. We added a toothpick stab of each colony to 10 µL of 24 mm Tris-HCl (pH 8.4), 60 mm KCl, 0.075 mm each dNTP, 7.5 mm MgCl2, and 0.6 mm pUC19 forward and reverse sequencing primers. We incubated the mixture at 100 °C for 10 min then placed the tubes on ice. Five µL Taq solution (12 mm Tris-HCl, pH 8.4, 30 mm KCl, 0.5 U Taq DNA polymerase, recombinant, GIBCO) were added to each tube. Fifteen µL reactions (final conditions: 20 mm Tris-HCl, pH 8.4, 50 mm KCl, 0.05 mm each dNTP, 5 mm MgCl2, 0.4 mm each primer, 0.5 U Taq DNA polymerase) were placed in a preheated thermal cycler (MJ Research PTC 100) set to cycle as follows: 94 °C for 4.5 min, 25 cycles of 94 °C for 30 s, 57 °C for 30 s, 72 °C for 30 s, then 72 °C for 2 min. Approximately 1 µL product was run on a 3% TAE agarose gel made with 0.03× GelStar nucleic acid stain (BioWhittaker Molecular Products) to identify inserts of 300 – 800 bp. Colonies containing these inserts were grown overnight in Luria broth from which plasmids were purified using the QIAprep Spin Miniprep Kit (Qiagen). More than 150 clones were sequenced using the Big DyeTM Terminator cycle sequencing protocol and visualized on an ABI 377 DNA sequencer (Applied Biosystems) by Davis Sequencing (Davis, CA). Fifty-four primer pairs were designed from approximately 100 unique sequences using ‘PrimerSelect’ (DNAStar, Inc.). Ten to 20 ng DNA from up to four crayfish populations sampled within the Sacramento Valley, California, were combined with 20 mm Tris-HCl (pH 8.4), 50 mm KCl, 0.2 mm each dNTP and 0.5 U Taq DNA Polymerase in a 10 µL reaction volume; MgCl2 and primer concentrations are indicated in Table 1. Cycling conditions were 95 °C for 2 min, 30 cycles of 95 °C for 30 s, 56 °C for 30 s, 72 °C for 1 min, then 72 °C for 5 min. Amplification products were mixed 1:1 with 98% formamide loading dye, denatured for 3 min at 95 °C, placed on ice, then run on 5% denaturing acrylamide gels and stained by agarose overlay containing 0.5 µL SYBR GreenI nucleic acid stain (BioWhittaker Molecular Application). Staining otherwise followed Rodzen et al. (1998). Products were visualized on a Molecular Dymamics FluorImager 595. Locus details are reported in Table 1. Eighteen primer pairs were also tested on P. zonangulus, Orconectes virilis, O. rusticus, Pacifasticus leniusculus, and Cherax quadricarinatus. Amplification success is reported in Table 2. These results indicate the utility of these microsatellite loci for genetic studies involving P. clarkii, and their potential utility in related species. Table 2 Cross-species amplification with 18 of the primers listed in Table 1. n indicates number of individuals tested unless otherwise indicated in parentheses in each cell. Numbers in cells indicate the number of observed (presumed) alleles; ‘–’ indicates amplification but unclear; ‘0’ indicates no amplification or smear only P R I M E R N O T E S 2233 2234 P R I M E R N O T E S Institute of Environmental Health Sciences Superfund Research Fellowship to NM Belfiore. Many thanks to Drs W Perry and J Huner for samples. References Asahida T, Kobayashi T, Saitoh K, Nakayama I (1996) Tissue preservation and total DNA extraction from fish stored at ambient temperature using buffers containing high concentration of urea. Fisheries Science, 62 (5), 727–730. Busack CA (1988) Electrophoretic variation in the red swamp (Procambarus clarkii) and the White River crayfish (P. acutus) (Decapoda: Cambaridae). Aquaculture, 69, 211– 226. Huner JV (1988) Procambarus in North America and elsewhere. In: Freshwater Crayfish: Biology, Management and Exploitation (eds Holdich DM, Lowery RS), pp. 239–261. Timber Press, Portland. May B, Krueger CC, Eng W, Paul E (1995) Genes in Populations Version 2. http://animalscience.ucdavis.edu/extension/gene.htm. Rodzen JA, Agresti JJ, Tranah G, May B (1998) Agarose overlays allow simplified staining of polyacrylamide gels. BioTechniques, 25 (4), 584. © 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234