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
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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
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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.
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© 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
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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
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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
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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.
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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.
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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
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primer
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© 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
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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.
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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
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1139
109PRIMER
02
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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
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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
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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
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109PRIMER
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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.
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© 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
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© 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.
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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
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109PRIMER
Graphicraft
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NOTEs
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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.
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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
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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
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© 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.
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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.
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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.
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Notes
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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.
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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.
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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.
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Komdeur J, Daan S, Tinbergen J, Mateman C (1997) Extreme
adaptive modification in the sex ratio of Seychelles warbler’s
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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.
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© 2000 Blackwell Science Ltd, Molecular Ecology, 9, 2155 –2234