2019. Journal of Arachnology 47:190–201
Cost effective microsatellite isolation and genotyping by high throughput sequencing
,Sven K ¨
James B. Henderson
,Warren Brian Simison
and Gabriele Uhl
of Biogeography, University of Trier, Trier, Germany; E-mail: email@example.com;
Environmental Science, Policy and Management, University of California, Berkeley, USA;
Center for Comparative
Genomics, California Academy of Sciences, San Francisco, California, USA;
General and Systematic Zoology,
Zoological Institute and Museum, University of Greifswald, Germany;
Zoologisches Forschungsmuseum Alexander
Koenig, Bonn, Germany;
Institute of Zoology, Behavioural Biology, Universit¨
at Hamburg, Germany;
Bioscience, Aarhus University, Denmark;
Max Planck Institute for Evolutionary Biology, Pl ¨
Abstract. High throughput sequencing (HTS) has emerged as a valuable tool for the rapid isolation of genetic markers
for population genetics and pedigree analysis. HTS-based SNP (single nucleotide polymorphism) genotyping protocols like
RAD (Restriction-site associated DNA) sequencing or hybrid capture allow for the isolation of thousands of markers from
any non-model organism. However, these protocols are relatively laborious and expensive and the resulting high marker
density is not always necessary. Since HTS technology has also greatly simpliﬁed the process of microsatellite marker
isolation and genotyping, we develop microsatellite markers as a cost-efﬁcient and simple alternative to SNP genotyping.
We present low coverage genome sequencing data from seven distantly related spider species (Argiope bruennichi (Scopoli,
1772), Larinia jeskovi Marusik, 1987, Oedothorax retusus (Westring, 1851), Pisaura mirabilis (Clerck, 1757),
Australomisidia ergandros (Evans, 1995), Cheiracanthium punctorium (Villers, 1789), Theridion grallator Simon,1900)
and show the utility of HTS for microsatellite isolation. We also present a simple Illumina amplicon sequencing protocol to
genotype microsatellites from multiplex PCR amplicons in the Hawaiian happy face spider T. grallator. We discuss
advantages and drawbacks of the use of microsatellites for a range of research questions, and highlight an unexpectedly
fast decay and gain of repeat loci for T. grallator.
Keywords: Genome, paternity assessment, population genetics, amplicon sequencing
High throughput sequencing is currently revolutionizing
molecular ecology and systematics. Thousands of markers can
be isolated from any non-model organism with protocols like
RAD (Restriction-site associated DNA) sequencing (Peterson
et al. 2012) or sequence capture (Smith et al. 2013; Mayer et al.
2016) and whole genome and transcriptome sequencing are
now feasible (Ekblom & Galindo 2011; Ellegren 2014). The
resulting marker density has greatly contributed to an in-depth
understanding of evolutionary and ecological processes,
including in the ﬁeld of arachnology (Brewer et al. 2014).
Several spider genomes have recently been sequenced (Sang-
gaard et al. 2014; Babb et al. 2017; Schwager et al. 2017), the
spider tree of life has been tackled using high-density markers
(Bond et al. 2014; Ferna
´ndez et al. 2014, 2018) and genomic
and transcriptomic analyses have provided insights into
evolutionary divergence in spiders (Croucher et al. 2013;
Bechsgaard et al. 2015; Krehenwinkel et al. 2015; Settepani et
al. 2017). The recent isolation of Ultra Conserved Elements
(Starrett et al. 2017) and application of ddRAD (double
digest) sequencing protocols (Burns et al. 2017; Settepani et al.
2017) have additionally contributed a wealth of genetic
markers for spider research. However, RAD sequencing
(Burns et al. 2017) or sequence capture (Smith et al. 2013;
Cotoras et al. 2018) protocols are relatively laborious and
expensive and the analysis of such high throughput genotyping
data requires considerable computational resources. Depend-
ing on the question of research, a density of thousands of
SNPs may not always be necessary, and thus a simpler and
more cost-efﬁcient approach is desirable for some types of
studies. In this regard, microsatellites are noteworthy: They
are repeated short sequences of DNA that evolve rapidly and
hence are powerful markers for analyzing mating rates and
paternity success, for population and conservation genetics, as
well as pedigree analyses and recent evolutionary divergence
afer et al. 2008; Tuni et al. 2012; Krehenwinkel &
Tautz 2013; Zimmer et al. 2014; Krehenwinkel et al. 2016b).
While their isolation used to be laborious, microsatellites are
now routinely isolated from any species by shotgun sequenc-
ing of genomic DNA (Castoe et al. 2010; Malausa et al. 2011).
Thus, while little used for arachnids in the past (Brewer et al.
2014), microsatellites are becoming available for an increasing
number of spider species (R ¨
utten et al. 2001; Bilde et al. 2009;
da Silveira & Bonatto 2009; Hataway et al. 2011; Esquivel-
Bobadilla et al. 2013; Parmakelis et al. 2013; Planas et al.
2014). High throughput sequencing has not only simpliﬁed the
isolation of microsatellite markers but is also well-suited for
genotyping of microsatellites (Cao et al. 2014; Darby et al.
2016). In particular, Illumina amplicon sequencing provides a
rapid, accurate and very cost-efﬁcient alternative to the
laborious and expensive capillary electrophoresis protocols,
which were commonly used for microsatellite genotyping (Cao
et al. 2014; Darby et al. 2016). Large-scale amplicon
sequencing of multiplex PCRs can be routinely performed
for hundreds of samples in parallel (Fadrosh et al. 2014) and
microsatellite alleles can be directly called from amplicon
sequencing data by analyzing the read length distribution per
specimen. Several software solutions for microsatellite geno-
typing from Illumina amplicon sequencing data have been
published or are under development (Suez et al. 2016; Zhan et
* current address: Department of Environmental Systems Science,
ETH Zurich, Switzerland
al. 2017; Henderson, Russack, Krehenwinkel & Simison
unpublished data). Against this background, it is worthwhile
to consider the utility of microsatellite markers as a simple and
cost-efﬁcient alternative to high throughput SNP genotyping.
Here, we provide an assessment of the isolation and
genotyping processes of microsatellite markers. Our aims are
twofold. We ﬁrst analyze the feasibility of high throughput
sequencing for microsatellite marker isolation. For this, we
present low coverage genome sequencing data for seven
distantly related spider species from six families. These species
are important models in different ﬁelds of arachnology, from
behavioral ecology, to phylogeography and population
genetics. We identify tandem repeat contents in the analyzed
spider genomes, isolate markers from the genomic data and
present structural similarities of the repeat content between
genomes. Based on these results, we discuss the necessary
sequencing depth for microsatellite marker discovery in
spiders. We then provide a set of established markers for the
studied species. Secondly, we present an overview of the
workﬂow for microsatellite genotyping using Illumina ampli-
con sequencing, from PCR set up, to library preparation and
sequencing, and genotyping from raw read data, using the
Hawaiian happy face spider Theridion grallator Simon, 1900.
The phylogeography of the happy face spider has been studied
using mitochondrial sequence information and allozyme data
(Croucher et al. 2012); thus, by applying the approach to T.
grallator, we can compare directly the effectiveness of
microsatellite markers for recovering genetic differentiation
among populations. The well-understood genetic history of
the species also offers the potential to investigate the evolution
of repeat loci over time, e.g., decay or gain of microsatellites
between different populations. Based on our results, we
discuss promises and drawbacks of Illumina based microsat-
ellite analyses and highlight unique features of tandem repeat
evolution in spiders. Our results demonstrate the feasibility of
high throughput sequencing based microsatellite isolation and
genotyping, and may help in establishing microsatellites as a
more commonly used marker type for genetic studies on
Target species.—We targeted seven distantly related spider
species from six families to assess the utility of high
throughput sequencing for microsatellite marker isolation.
Argiope bruennichi (Scopoli, 1772) (Araneidae) serves as a
model species in research on the evolution of sexual
cannibalism and its consequences for mating systems (Fromh-
age et al. 2003; Schneider & Andrade 2011; Schneider 2014;
Schneider et al. 2015; Uhl et al. 2015) and has recently gained
importance as a model for evolutionary divergence during
contemporary range expansions (Krehenwinkel & Tautz 2013;
Krehenwinkel et al. 2015, 2016a). Larinia jeskovi Marusik,
1987 (Araneidae) is remarkable due to its mating behavior,
since males mutilate the female genitalia to prevent further
insemination by rival males (Mouginot et al. 2015, 2017).
Oedothorax retusus (Westring, 1851) (Linyphiidae) has re-
ceived considerable attention due to its mating strategy, in
which males perform gustatory courtship and plug the
female’s genital opening with secretion (Kunz et al. 2012,
2014). Pisaura mirabilis (Clerck, 1757) (Pisauridae) is known
as the nuptial gift spider, in which males provide females with
a prey item prior to mating that results in sexual conﬂict (Bilde
et al. 2007; Albo et al. 2013; Ghislandi et al. 2018).
Cheiracanthium punctorium (Villers, 1789) (Cheiracanthiidae)
is known for being the only medically important spider in
Central Europe and is currently rapidly expanding its range
(Muster et al. 2008; Krehenwinkel et al. 2016b). The thomisid
species Australomisidia ergandros (Evans, 1995) shows com-
munal hunting behavior (Ruch et al. 2014, 2015; Dumke et al.
2016) and brood care by which the female provides herself to
her offspring as food (Evans 1998). The happy face spider,
Theridion grallator Simon, 1900 (Theridiidae), is widely-
distributed across the rainforests of the Hawaiian Islands of
Oahu, Molokai, Maui, and Big Island, and is well known for
its conspicuous and eponymous color morphs that occur as a
balanced polymorphism in every population (Gillespie &
Oxford 1998). The diverse research questions ranging from
paternity and relatedness to population structure require fast
evolving genetic markers.
Microsatellite isolation by high throughput sequencing.—
Genomic DNA from six spider species was extracted from leg
muscle tissue using the Qiagen DNeasy blood and tissue kit
according to the manufacturer’s protocol (Qiagen, Hilden,
Germany). The taxonomy and origin of samples are shown in
Table 1. The untreated genomic DNA of ﬁve species was
sequenced, each on1/8th ﬂow cell of a 454 GS FLXþﬂow cell
in 2010 and 2011. Library preparation and sequencing were
performed according to the manufacturer’s protocols (Roche,
Basel, Switzerland; see Krehenwinkel & Tautz (2013) for more
details on the 454 sequencing and analyses). An additional
data set of the yellow sac spider Cheiracanthium punctorium
was generated by sequencing pooled genomic DNA on two
full ﬂow cells of an Illumina Miseq in 2014 (Illumina, San
Diego, CA, USA). Library preparation and sequencing were
performed using the V3 chemistry according to the manufac-
turer’s protocols and sequencing 300 bp paired end reads. The
454 and Illumina sequencing runs were both performed at the
Max Planck Institute for Evolutionary Biology in Pl¨
Germany. The paired end Illumina reads were quality trimmed
using PoPoolation (Koﬂer et al. 2011) with a minimum quality
threshold of 20 and adapters removed using Trimmomatic
(Bolger et al. 2014). A de novo assembly was generated
including both Illumina libraries and using CLC genomic
workbench at a minimum contig length of 1000 and including
a scaffolding step (CLC Bio, Boston, USA). For the happy
face spider Theridion grallator, we used a subset of ~20 Mb of
contigs from a preliminary genome assembly, based on
Illumina paired end sequencing data (Croucher, unpublished
data). We estimated the GC content of reads and assemblies
We identiﬁed tandem repeats in the Illumina assemblies of
C. punctorium and T. grallator and the raw 454 reads of the
remaining spiders using MSat Commander (Faircloth 2008)
with a minimum length of 10 repeats for mononucleotide
repeats, 10 repeats for di-, 6 repeats for tri-, 5 repeats for
tetra-, 4 repeats for penta- and hexanucleotide repeats. We
counted the number of repeat loci for each repeat class and
calculated the genome wide content of different repeat motifs
per megabase of sequence data (Table 1, 2). In order to
identify the tandem repeat content, we designed primers for
KREHENWINKEL ET AL.—MICROSATELLITES AND GENOTYPING FOR SPIDERS 191
all possible di, tri and tetranucleotide repeat loci using the
Primer3 plugin (Rozen & Skaletsky 1999) of MSat Com-
mander. A subset of the microsatellite loci was tested for
variability using ﬂuorescently labeled primers and following
genotyping as described in Krehenwinkel & Tautz (2013).
Amplicon sequencing for microsatellite genotyping.—We
used several Hawaiian populations of the happy face spider
Theridion grallator to explore the utility of Illumina amplicon
sequencing for microsatellite genotyping. Specimens of T.
grallator were collected in March 2015 on the Hawaiian
Islands of Oahu, Maui, Molokai and Hawai’i by beating
vegetation and hand collecting from the underside of leaves.
Two separate subpopulations per island were sampled (see
Table 3). All specimens were stored in 99%ethanol and
brought back to the University of California Berkeley for
further analyses. DNA extractions were performed on the
whole prosoma of each specimen using the Qiagen Puregene
Tissue kit according to the manufacturer’s protocol (Qiagen,
Valencia, USA). The DNA from 96 specimens was quantiﬁed
using a Qubit Fluorometer (Thermo Scientiﬁc, Waltham,
USA), diluted to approximately 20 ng/ll and distributed
among wells of a 96-well plate. Fifty primer pairs were tested
in a subset of three specimens from each island for
determining PCR ampliﬁcation efﬁciency. The targeted loci
contained di-, tri- or tetranucleotide repeats and the targeted
amplicons were less than 400 bp long, to achieve a good
overlap during read merging. Each primer pair was tested in
an annealing temperature gradient from 50–60 8C in incre-
ments of 2.5 8C. Gradient PCR were run for each primer pair
using the Qiagen Multiplex PCR kit according to the
manufacturer’s protocol and with 30 cycles. No Q-solution
was added. PCR products were screened on a 1.5 %agarose
gel. The 25 primer pairs which most consistently ampliﬁed
specimens from all four islands were chosen for further
analyses (see Supplementary Table 1, online at http://dx.doi.
org/10.1636/JoA-S-16-017.s1 for details on primer pairs and
repeat loci). This selection was done to avoid priming bias and
drop out of loci by sequence divergence between populations.
PCR reactions with primers of similar optimal annealing
temperature were then combined to multiplex reactions. We
added a 50tail to each primer and ampliﬁed the 25 primer
pairs in six multiplex PCRs of 30 cycles for all of the 96
specimens and otherwise as described above. PCRs were run in
10 ll volumes, with 0.5 ll of each 10 lM primer and 1 ll of the
20 ng/ll template. The added 50tails served as a priming site
for a second PCR round of 5 cycles, in which dual indices
(short, unique sequences for individual identiﬁcation) and
Illumina Truseq adapters were introduced. The concept of
PCR-based library preparation followed that described in
Lange et al. (2014). Before the second PCR, all multiplexes
were pooled into a single 96-well plate according to specimen
and placed for an approximate quantiﬁcation on a 1.5 %
agarose gel. A second PCR round was run with 8 forward
times 12 reverse combinations of indexed primers. This
indexing PCR was run as described above, but with only 5
cycles at 55 8C annealing temperature, 0.25 ll of each 10 lM
Table 1.—The upper table shows the taxonomy and sampling origin of the sequenced spider species, sequencing platform used, number of
sequenced bases, number of sequenced reads and estimates for the genome-wide GC-content for the six species. The lower table presents the
assembly statistics for the two assemblies used for marker isolation.
Family Genus Species Origin Platform Sequenced bases No. reads GC-content (%)
Araneidae Argiope bruennichi Germany 454 30.29*10
Araneidae Larinia jeskovi Poland 454 30.39*10
Linyphiidae Oedothorax retusus Germany 454 24.79*10
Pisauridae Pisaura mirabilis Germany 454 23.12*10
Theridiidae Theridion grallator Hawaii -- - 27
Thomisidae Australomisidia ergandros Australia 454 24.18*10
Cheiracanthiidae Cheiracanthium punctorium Baltic States MiSeq 18.78*10
Cheiracanthiidae Cheiracanthium punctorium Mediterranean MiSeq 12.37*10
Family Genus Species Origin Platform Assembly size
contig size (bp) GC-content (%)
Cheiracanthiidae Cheiracanthium punctorium Mediterranean MiSeq 148.60*10
Theridiidae Theridion grallator Hawaii HiSeq 22.00*10
Table 2.—Coverage of different tandem repeat motifs (TR) per megabase (Mb) of sequenced DNA for all spider species studied. The last two
columns show the number of primer pairs which could be designed on the recovered tandem repeats per Mb of DNA and the %of recovered
repeat motifs on which primers could be designed.
Species TR motifs/Mb Mono/Mb Di/Mb Tri/Mb Tetra/Mb Penta/Mb Hexa/Mb Primers/Mb %motifs with primers
A. bruennichi 340.84 293.89 35.55 7.13 2.48 1.39 0.45 3.93 8.36
L. jeskovi 103.30 78.79 17.67 3.16 1.48 2.01 0.07 1.88 7.71
O. retusus 118.20 102.3 3.15 4.36 5.28 2.22 0.89 2.74 17.23
P. mirabilis 225.90 113.74 94.27 5.02 9.09 3.37 0.43 9.13 8.14
A. ergandros 248.30 200.18 25.93 6.86 13.44 1.61 0.33 8.15 16.92
C. punctorium 464.78 434.22 10.91 9.65 8.08 1.83 0.11 19.18 62.72
T. grallator 218.19 203.22 13.02 0.6 0.75 0.55 0.05 2.5 16.70
192 JOURNAL OF ARACHNOLOGY
primer and 0.5 ll of the PCR template from the ﬁrst PCR
round. After each PCR, the products were puriﬁed from
remaining primer using AMpure XP beads (Beckman Coulter,
Brea, USA). The puriﬁed and dual indexed PCR products
were quantiﬁed using a Qubit, and then 10 ng of each sample
were pooled. The sample was sequenced on approximately 1/4
of a MiSeq ﬂow cell using the Illumina V3 chemistry, with 300
bp paired reads and according to the manufacturer’s protocol
(Illumina, San Diego, USA). The remainder of the ﬂow cell
contained microbial 16S and arthropod mitochondrial COI
samples. A ‘‘spike-in’’ of 15 %PhiX was added to the run.
Adapter sequences were trimmed from the raw reads using
Trimmomatic (Bolger et al. 2014). Paired reads were then
merged using PEAR (Zhang et al. 2014) with a minimum
overlap of 75 bp and a minimum quality of 30. Only those
assembled reads with at least 90 %of bases with Q30 or higher
were transformed into fasta ﬁles using the fastx toolkit
(Gordon & Hannon 2010). All sequences, starting with the
forward primer and ending with the reverse primer of each
specimen and each of the 25 microsatellite loci were ﬁltered
and saved to new ﬁles. This step was performed using UNIX
and served to demultiplex all separate loci. For allele-calling,
we measured the length of each amplicon and counted the
abundance of each length per specimen and per locus using the
programming language awk. By the previous ﬁltering of
sequences, which start and end exactly at the PCR primers, all
fragments can be measured in exact relation to each other. We
plotted the distribution of different fragment lengths for each
specimen and locus using a custom-made software (Hender-
son, Russack, Krehenwinkel & Simison unpublished data).
The approach is very comparable to that implemented in
classic fragment length analysis software for dye-labelled
allele-calls, e.g., Genemapper (Thermo Scientiﬁc, Waltham,
USA). However, here we plotted the abundance of reads for
different fragment lengths instead of the intensity of dye
ﬂuorescence (Fig. 1 A, B). The ﬁnal version of our simple and
user-friendly software solution for allele-calling will perform
all tasks from sequence adapter trimming and assembly, to
demultiplexing, allele-calling and exporting the called alleles in
a user-friendly graphical interface (Henderson, Russack,
Krehenwinkel & Simison unpublished data). Alleles were
called for each specimen and locus and the fragment size
distributions for each locus were manually inspected to edit
allele calls. Due to locus and population speciﬁc stutter
patterns and the possibility of ﬂanking indels contributing to
repeat length, this manual curation step proved essential. We
used a minimum coverage of 40 reads per locus and specimen
to call alleles. We additionally inspected a random subset of
unassembled reads for their microsatellite pattern and
compared the result with that identiﬁed based on assembled
reads pairs. This step served to exclude assembly-based
artifacts. MEGA (Tamura et al. 2013) was used to visualize
repeat containing sequences and to align subsamples of
sequences. We visually inspected the sequences of every
specimen for the presence of repeat motifs, indels outside of
repeat motifs and ﬂanking SNPs.
The genetic structure of T. grallator across the Hawaiian
Islands was evaluated by a STRUCTURE (Pritchard et al.
2000; Falush et al. 2003) analysis. STRUCTURE was run with
an admixture model, k-values from 1–10, 10 replicates per k-
value, 150,000 MCMC generations of which 50,000 were
removed as burnin. The optimal number of clusters was
identiﬁed using STRUCTURE HARVESTER (Earl et al.
2012) with the method described in Evanno et al. (2005).
values between populations were calculated in
Genepop (Raymond & Rousset 1995).
Microsatellite isolation by high throughput sequencing.—The
results of sequencing and assembly can be found in Table 1.
The 454 runs yielded about 100,000 reads and 30,000,000 bp
per species, the MiSeq runs between 15–20 million reads. The
assembly resulted in 148,602 contigs over 1 kb long, with a
median size of 1,024 and a total size of ~232 million bases
(Table 1). All spider genomes were characterized by a
relatively low GC content between 27–34 %. Depending on
the species, we recovered between 103–465 microsatellites per
megabase (Mb) of sequenced DNA (Table 2). The repeat
content in spider genomes was highly biased towards
mononucleotides and was variable between species. As
mononucleotide repeats are hard to genotype, we did not
design primers for this repeat type. Excluding mononucleo-
tides, only between 15–112 microsatellites could be recovered
per Mb. Due to the high AT content of spider genomes
(Sanggaard et al. 2014; Krehenwinkel et al. 2015), AT rich
repeats dominate in the isolated loci (Supplementary Table 1).
A relatively small subset of the identiﬁed repeat loci
contained sufﬁcient ﬂanking sequences to design primers.
Between 2–19 microsatellite markers could be recovered per
sequenced Mb of DNA. This corresponds to between 8–62 %
of the total number of recovered repeat loci. Moreover, a
considerable proportion of designed primer pairs had to be
dropped after initial analyses. On average, we could establish
Table 3.—Collection sites and sample numbers per population for the eight sampled populations of Theridion grallator on the Hawaiian
Island Population Locality N H
Big Island HiBiK15 Saddle Road, Kipuka15 12 0.277 0.351
Big Island HiBiM21 Saddle Road, Milemarker 21 8 0.265 0.388
Maui HiMaWA Waikamaoi, outside preserve 13 0.485 0.676
Maui HiMaWB Waikamoi, upper preserve 7 0.424 0.678
Molokai HiMoKA Kamakou, TNC cabin 17 0.324 0.508
Molokai HiMoKB Kamakou, boardwalk 14 0.318 0.463
Oahu HiOhPA Pahole 10 0.369 0.485
Oahu HiOhPB Pahole 10 0.332 0.474
KREHENWINKEL ET AL.—MICROSATELLITES AND GENOTYPING FOR SPIDERS 193
11 highly variable microsatellite markers out of 50 primer
pairs, which we designed per species and tested in PCR assays.
This low yield was due to two problems. First, some markers
showed highly population speciﬁc ampliﬁcation biases, leading
to dropouts if genetically distant populations were analyzed.
This problem was most evident in the analyses of divergent
Eastern and Western Palearctic populations of A. bruennichi
(Krehenwinkel et al. 2016a). Second, many repeat loci lacked
size variation in some spider species, particularly O. retusus
and A. ergandros. We provide a list of established primer
sequences, examples for their application and ampliﬁcation
bias or repeat variation issues in Supplementary Table 1.
Amplicon sequencing for microsatellite genotyping.—After
trimming and assembly, we recovered 26,425 high-quality
sequences per T. grallator specimen on average (68,787
standard deviation). Four specimens had to be removed from
the analysis due to low sequence coverage (12–16 sequences
only). We found 1,001 reads on average (61,488 standard
deviation) per microsatellite locus and specimen. One locus
was removed from the analysis because coverage was too low
(16 sequences per specimen on average). Our ﬁnal dataset
consisted of 92 specimens and 24 loci. Many recovered
microsatellite loci showed variable repeat motifs within and
among populations (Fig. 1A, B).
A STRUCTURE analysis of the microsatellite dataset
supported k ¼4 populations (Fig. 2), each of them
corresponding to one of the Hawaiian Islands and in line
with recent research by Croucher et al. (2012). An F
corroborated these results (Table 4). While populations within
islands showed little to no differentiation, we found consid-
erable divergence between islands.
Alignments of the microsatellite amplicons allowed more
detailed insights into the distribution of repeat patterns,
ﬂanking indels and SNPs in different populations of T.
grallator, and in comparison to the population from Maui, for
which the loci were designed (Fig. 3). Repeat sequences were
often imperfect, e.g., containing additional bases which break
the repeat pattern (70 %of loci; Fig. 3). One of the ampliﬁed
loci did not contain any tandem repeat, despite being
identiﬁed as a dinucleotide repeat in Msatcommander. Many
microsatellites appeared to be population-speciﬁc and were
absent in other populations. Specimens from Maui, which
were also used as template to design microsatellites, carried
repeat motifs for 23 out of 24 analyzed loci. In most other
populations, we found an absence of many repeat loci, with
about 1/3 of loci not showing the repeat in comparison to
Maui (Fig. 4, Table 5). Overall, 13 out of 24 loci showed a
missing repeat in at least one population. The decay of repeats
was often part of the standing variation, with some specimens
in a population carrying repeats and others not. Most of the
loci, for which repeats were absent in other populations,
carried the loss of the repeat pattern in the standing variation
Figure 1.—(A) Subsection of genotyping plots for the allelic distribution of a CT-dinucleotide repeat locus (TG_MS41) for two happy face
spider specimens from Maui. The plots show the abundance (Y-axis) of reads of different fragment length (X-axis). Each bar indicates the
abundance of one fragment length in a mixture of sequences, with the red dot indicating the called allele length. The upper specimen is
heterozygous and the lower is homozygous.
194 JOURNAL OF ARACHNOLOGY
on Maui. Across the phylogeny of the happy face spider
(Croucher et al. 2012), our data suggest considerable gains and
losses of repeat loci within about 1 million years of inter-island
colonization of the species (Table 5, Fig. 4).
Apart from losses of repeats, additional factors contributed
to size differences in PCR amplicons (Fig. 3, Table 5). About
25 %of the analyzed loci carried a second tandem repeat
motif, often right next to the targeted one. Moreover, indels
outside of the targeted tandem repeat pattern had a
substantial contribution to amplicon size differences. Depend-
ing on the population, up to 90%of the analyzed loci carried
ﬂanking indels in the amplicon. Even after a complete loss of
the repeat motif, these indels contributed to variable fragment
sizes. This was particularly important in populations outside
of Maui, where a signiﬁcant proportion of the repeat motifs
Microsatellite isolation by high throughput sequencing.—Our
results show that microsatellite markers can be routinely
isolated by low coverage sequencing from any spider genome.
Simple high throughput sequencing of untreated genomic
DNA will yield sufﬁcient markers for certain types of
population studies in spiders. It is important to aim for long
reads to provide sufﬁcient ﬂanking regions for primer design.
The best combination of read length and high output is
currently offered by the Illumina MiSeq system. With its V3
Figure 1.—(B) Subsection of genotyping plots for the allelic distribution of an AT-dinucleotide repeat (TG_MS1) for two happy face
specimens from Hawaii Island. The plots show the abundance (Y-axis) of reads of different fragment length (X-axis). Each bar indicates the
abundance of one fragment length in a mixture of sequences, with the red dot indicating the called allele length. The upper specimen is
heterozygous and the lower is homozygous.
Table 4.—Pairwise F
between Hawaiian populations of the happy face spider Theridion grallator for the microsatellite fragment length
dataset generated by Illumina amplicon sequencing. The population names correspond to those in Table 3.
HiBiK15 HiBiMM21 HiMaWaA HiMaWaB HiMoKaA HiMoKaC HiOhPeA
HiMaWaA 0.302 0.277
HiMaWaB 0.348 0.315 0.000
HiMoKaA 0.372 0.341 0.250 0.255
HiMoKaC 0.403 0.375 0.273 0.275 0.000
HiOhPeA 0.497 0.463 0.337 0.354 0.437 0.460
HiOhPeB 0.517 0.483 0.346 0.365 0.446 0.467 0.012
KREHENWINKEL ET AL.—MICROSATELLITES AND GENOTYPING FOR SPIDERS 195
Figure 2.—Results (right) of a STRUCTURE analysis of the microsatellite dataset assuming k ¼4 populations. Colors correspond to that of
islands in the sampling map (left).
Figure 3.—Alignment of sequences of happy face spiders (Theridion grallator) from different Hawaiian Islands for a CT-dinucleotide repeat
(MS41). Each island is represented with 8 sequences. The repeat motif is only found in populations on Maui and absent on all other islands. The
repeat motif is not perfect, with a Thymine inserted instead of Cytosine in some specimens. Flanking SNPs support the signal of the repeat
pattern. A ﬂanking deletion is present in all specimens from Big Island and a subset of those from Oahu.
196 JOURNAL OF ARACHNOLOGY
chemistry, it is possible to obtain up to 20,000,000 paired reads
of 2 3300 bp per run. About 50 microsatellite loci should
provide sufﬁcient resolution for some population genetic
applications depending on the question in focus, as well as
for paternity analysis. Considering the recovery of 2–19 primer
pairs per sequenced Mb of DNA, this translates to a minimum
of 25 Mb of DNA that needs to be sequenced for
microsatellite recovery. Our analyses in T. grallator suggest a
very fast turnover of repeat loci in spider genomes. Due to the
fast evolution of the repeat content, it is probably not possible
to predict the expected repeat content for a target species from
other related taxa. The repeat content has to be determined de
novo for each species. We found a high dropout rate due to the
lack of variation in repeat length for some taxa, e.g., many loci
could not be used for further analysis in these taxa.
Consequently, we recommend a higher sequencing coverage
of about 250 Mb per specimen for the development of
microsatellite markers. Considering the throughput of an
Illumina MiSeq, this still translates to 80 separate spider
species, for which microsatellites could be isolated in a single
sequencing run. At a cost of about $1,600 USD per run,
microsatellites for any spider species can be isolated for about
$50 USD, including the cost for library preparation. Using
enrichment protocols, the output of isolated loci could be
increased substantially (Malausa et al. 2011).
Microsatellite genotyping using Illumina amplicon sequenc-
ing.—Microsatellite amplicons can be rapidly sequenced and
genotyped using paired-end Illumina sequencing of multiplex
PCRs. High throughput sequencing approaches come with
many advantages over traditional capillary electrophoresis of
dye-labelled amplicons. At the same time, the results of high
throughput sequencing approaches are highly congruent with
those of capillary electrophoresis (Vartia et al. 2016; Zhan et
al. 2017). The high coverage of Illumina sequencing avoids the
need to balance ampliﬁcation of every single marker in a
multiplex reaction. While the ﬂuorescence of a single,
overrepresented marker can outshine all other loci in a
multiplex, the read count information for each locus is
independent from that of the others. Even without previous
optimization of multiplexes, we recover a very high number of
loci (24 out of 25) and specimens (92 out of 96) in our analysis.
There is also no need to select non-overlapping groups of
marker sizes for multiplex reactions, as is typical for dye-
labelled reactions. The only limit of an Illumina based
approach is the current maximum size of paired read lengths
of 2 3300 bp. Assuming that each sequence is supposed to
read through the tandem repeat, a large overlap is desired for
read merging. Thus, amplicons should not exceed 400bp. Our
approach of plotting read length abundance proﬁles per locus
(Henderson, Russack, Krehenwinkel & Simison unpublished
data) is comparable to classic allele-calling software. Recent
work suggests automated calling options (Suez et al. 2016) or a
Figure 4.—Losses and gains of tandem repeat loci over the microsatellite-based phylogeny of the four happy face spider populations analyzed.
The root of the tree and the divergence ages for the major clades are in accordance with Croucher et al. (2012). The bar plot shows the number of
loci with a repeat motif present in all specimens (black), present in only a subset of the population, e.g., part of the standing variation of the
population (grey) and absent (white).
Table 5.—Proportion of loci, out of 24 sequenced loci, for
Theridion grallator populations from four islands, which (1) contain
the tandem repeat motif targeted by primer design, (2) contain an
additional tandem repeat motif, (3) contain indels outside of the
tandem repeat motif, (4) carry variable SNPs.
Big Island Maui Molokai Oahu
Repeat motif present 0.71 0.96 0.71 0.63
Second repeat motif present 0.25 0.29 0.25 0.25
InDels present 0.67 0.92 0.83 0.75
SNPs present 0.96 0.96 1.00 1.00
KREHENWINKEL ET AL.—MICROSATELLITES AND GENOTYPING FOR SPIDERS 197
mixture of automated and manual curation of such datasets
(Zhan et al. 2016).
For the genotyping of small to moderate numbers of
markers, sequencing of microsatellite amplicons will be
cheaper and require less laboratory experience than current
SNP genotyping protocols, e.g., RAD sequencing. To generate
the 24-locus dataset for each specimen, we required six
multiplex PCRs, two clean up reactions and one indexing
PCR, adding to $4 USD. Sequencing required another $4
USD per specimen. However, after further optimization of our
multiplex PCRs, the number of PCRs and necessary
sequencing coverage and processing cost could be consider-
ably reduced. Recovering a higher genetic diversity and having
a better population assignment power at low marker densities
(Yang et al. 2011), microsatellites might even be preferable
over SNPs for small-scale population studies. A combination
of microsatellite isolation and genotyping by high throughput
sequencing could allow the rapid and cost-efﬁcient analysis of
paternity or population structure.
While microsatellites can be valuable markers, microsatellite
repeats can show highly complex and unpredictable patterns
of evolution (Ellegren 2000) making modelling difﬁcult. This
can hamper the interpretation of microsatellite data and
requires careful evaluation, e.g., by manual curation of the
data before downstream analysis. Furthermore, it should be
investigated whether more traditional enrichment marker
design protocols as described in Nolte et al. (2005) and Leese
et al. (2008) could be combined with HTS techniques, in order
to further increase the effectiveness of the procedure.
Implications of a rapid turnover of microsatellite repeat loci in
spiders.—The decay and possible gain of tandem repeat loci is
a prominent pattern in our data, with almost half of the
investigated loci not showing the repeat in at least one
population of T. grallator. Only populations from Maui, on
which the microsatellite loci used in this study were designed,
consistently show all repeat loci. Recent molecular analyses
suggest a stepping stone mode of colonization of the spiders
from Oahu to Maui Nui (Maui and Molokai) and on to the
Big Island within the past million years (Croucher et al. 2012).
With only a few thousand years of coalescence time, the happy
face spider is a young taxon. As an annual species, this
corresponds to only a few thousand generations. A turnover of
almost half of all repeat loci in this timeframe suggests a rapid
decay and gain of repeat loci. The fast turnover may also be
the reason for the lack of variation we found in many
microsatellite sequences for some spider species, particularly
O. retusus and A. ergandros. The lack of variation may also be
the result of ascertainment bias (Ellegren et al. 1995). By
choosing relatively long repeat motifs in our primer design, the
maximum repeat motif length for some loci may be reached
already. It is then likely by chance that evolution will decrease
the size of repeat motifs in distant groups.
A study on interspeciﬁc decay rates of repeat motifs in
Diptera and Hymenoptera, suggest losses of motifs in 30–40 %
of loci after 2 million generations (Stolle et al. 2013). Spiders
might thus show a faster rate of repeat motif evolution than
other arthropod taxa. In contrast to Diptera and Hymenop-
tera, all spiders have large genome sizes (Gregory 2001). This
might result in more genomic regions with little functional
constraint, allowing for a faster random evolution and decay
of repetitive sequences. Moreover, spider genomes generally
show a low GC content (Sanggaard et al. 2014; Krehenwinkel
et al. 2015). A low GC content in tandem repeats and their
ﬂanking sequence has been suggested to possibly contribute to
an increased mutation rate (Schl¨
otterer 2000). Microsatellites
are often associated with transposable elements (Ellegren
ecz et al. 2007). Their emergence by repeat
mobilization could explain the rapid appearance and losses
of repeats in T. grallator. However, little is currently known
about microsatellite evolution in invertebrates (Chapuis et al.
2015), and future studies will have to explore this topic in more
Practically, the rapid decay and gain of repeat loci means
that called allele sizes for microsatellites in distant populations
are often only based on ﬂanking indels and not the evolving
repeat. This urges care in cross-species ampliﬁcation, still a
popular approach (Moodley et al. 2015). This problem is
probably not speciﬁc for spiders. In particular, distances based
on models of evolution for the actual repeat motif might be
biased by repeat loss. Another practical issue with the lack of
size variation in repeat motifs is the need to explore large
numbers of microsatellite loci to identify sets of variable
markers for some spider species. However, due to the
simplicity of multiplex PCRs and the high throughput of
current amplicon sequencing protocols, the scoring of large
numbers of microsatellite markers is still a worthwhile
approach for studying population genetics in spiders.
Current high throughput sequencing technology allows the
rapid and cost-efﬁcient isolation of large numbers of
microsatellite markers from spiders as we have shown in
seven distantly related spider species. Moreover, Illumina
amplicon sequencing is well suited for genotyping of
microsatellite markers. As we highlight in an exemplary
species, amplicon sequencing based microsatellite genotyping
offers a greatly simpliﬁed workﬂow over currently used
capillary electrophoresis-based protocols. Provided that mi-
crosatellite data are carefully analyzed, our results demon-
strate that microsatellite markers can be a useful alternative to
SNP genotyping for population genetics and pedigree
We cordially thank Thoomke Br¨
uning, Sarah Frehse, Nadja
Gogrefe and Nicole Thomsen for assistance during lab work.
We thank Andrew Rominger, Ellie Armstrong and Nate Yuen
for help collecting happy face spider specimens. Anna Sellas
and the California Academy of Sciences’ Center for Compar-
ative Genomics, are acknowledged for support during
sequencing. Jun Ying Lim kindly provided a map of the
Hawaiian Archipelago. We acknowledge the Nature Conser-
vancy and the Hawaiian authorities for providing the
necessary permits to collect happy face spiders. Natalie
Graham provided helpful comments on the manuscript.
Diethard Tautz provided helpful discussion and access to
high throughput sequencing machines. HK was funded by a
PhD fellowship of the Studienstiftung des Deutschen Volkes
and by a postdoctoral fellowship of the German Research
Foundation (DFG). The research was supported by NSF
grant DEB 1241253 to RG and by the DFG to GU (Uh/87-7,
198 JOURNAL OF ARACHNOLOGY
RTG 2010). TB was supported by The Danish Council for
Independent Research grant number 4002-00328B.
Albo, M.J., T. Bilde & G. Uhl. 2013. Sperm storage mediated by
cryptic female choice for nuptial gifts. Proceedings of the Royal
Society of London B 280:20131735, doi: 10.1098/rspb.2013.1735.
Babb, P.L., N.F. Lahens, S.M. Correa-Garhwal, D.N. Nicholson,
E.J. Kim, J.B. Hogenesch et al. 2017. The Nephila clavipes genome
highlights the diversity of spider silk genes and their complex
expression. Nature Genetics 49:895.
Bechsgaard, J., B. Vanthournout, P. Funch, S. Vestbo, R.A. Gibbs, S.
Richards et al. 2015. Comparative genomic study of arachnid
immune systems indicates loss of beta-1, 3-glucanase-related
proteins and the immune deﬁciency pathway. Journal of Evolu-
tionary Biology 29:277–291.
Bilde, T., C. Tuni, A. Cariani, A. Santini, C. Tabarroni, F. Garoia et
al. 2009. Characterization of microsatellite loci in the subsocial
spider Stegodyphus lineatus (Araneae: Eresidae). Molecular Ecol-
ogy Resources 9:128–130.
Bilde, T., C. Tuni, R. Elsayed, S. Peka
´r & S. Toft. 2007. Nuptial gifts
of male spiders: sensory exploitation of the female’s maternal care
instinct or foraging motivation? Animal Behaviour 73:267–273.
Bolger, A.M., M. Lohse & B. Usadel. 2014. Trimmomatic: a ﬂexible
trimmer for Illumina sequence data. Bioinformatics 30:2114–2120.
Bond, J.E., N.L. Garrison, C.A. Hamilton, R. Godwin, M. Hedin &
I. Agnarsson. 2014. Phylogenomics resolves a spider backbone
phylogeny and rejects a prevailing paradigm for orb web evolution.
Current Biology 24:1765–1771.
Brewer, M.S., D.D. Cotoras, P.J. Croucher & R.G. Gillespie. 2014.
New sequencing technologies, the development of genomics tools,
and their applications in evolutionary arachnology. Journal of
Burns, M., J. Starrett, S. Derkarabetian, C.H. Richart, A. Cabrero &
M. Hedin. 2017. Comparative performance of double-digest RAD
sequencing across divergent arachnid lineages. Molecular Ecology
Cao, M. D., E. Tasker, K. Willadsen, M. Imelfort, S. Vishwanathan,
S. Sureshkumar et al. 2014. Inferring short tandem repeat variation
from paired-end short reads. Nucleic Acids Research 42:e16.
Castoe, T.A., A.W. Poole, W. Gu, A.P. Jason de Koning, J.M. Daza,
E.N. Smith et al. 2010. Rapid identiﬁcation of thousands of
copperhead snake (Agkistrodon contortrix) microsatellite loci from
modest amounts of 454 shotgun genome sequence. Molecular
Ecology Resources 10:341–347.
Chapuis, M.P., C. Plantamp, R. Streiff, L. Blondin & C. Piou 2015.
Microsatellite evolutionary rate and pattern in Schistocerca
gregaria inferred from direct observation of germline mutations.
Molecular Ecology 24:6107–6119.
Cotoras, D.O., K. Bi, M.S. Brewer, D.R. Lindberg, S. Prost & R.G.
Gillespie. 2018. Co-occurrence of ecologically similar species of
Hawaiian spiders reveals critical early phase of adaptive radiation.
BMC Evolutionary Biology 18:100.
Croucher, P.J., G.S. Oxford, A. Lam, N. Mody & R.G. Gillespie.
2012. Colonization history and population genetics of the color-
polymorphic Hawaiian happy-face spider Theridion grallator
(Araneae, Theridiidae). Evolution 66:2815–2833.
Croucher, P.J., M.S. Brewer, C.J. Winchell, G.S. Oxford & R.G.
Gillespie. 2013. De novo characterization of the gene-rich tran-
scriptomes of two color-polymorphic spiders, Theridion grallator
and T. californicum (Araneae: Theridiidae), with special reference
to pigment genes. BMC Genomics 14:862.
da Silveira, L.C.T. & S.L. Bonatto. 2009. Isolation and characteriza-
tion of 12 dinucletiotide microsatellite loci in Paratrechalea
galianoae (Araneae, Trechaleidae), a nuptial gift-spider. Molecular
Ecology Resources 9:539–541.
Darby, B.J., S.F. Erickson, S.D. Hervey & S.N. Ellis-Felege. 2016.
Digital fragment analysis of short tandem repeats by high-
throughput amplicon sequencing. Ecology and Evolution 6:4502–
Dumke, M., M.E. Herberstein & J.M. Schneider. 2016. Scrounging or
producing: individual feeding tactics change with group size in a
communally foraging spider. Proceedings of the Royal Society of
London B 283:20160114.
Earl, D.A. 2012. STRUCTURE HARVESTER: a website and
program for visualizing STRUCTURE output and implementing
the Evanno method. Conservation Genetics Resources 4:359–361.
Ekblom, R. & J. Galindo. 2011. Applications of next generation
sequencing in molecular ecology of non-model organisms. Hered-
Ellegren, H. 2000. Microsatellite mutations in the germline: implica-
tions for evolutionary inference. Trends in Genetics 16:551–558.
Ellegren, H. 2004. Microsatellites: simple sequences with complex
evolution. Nature Reviews Genetics 5:435–445.
Ellegren, H. 2014. Genome sequencing and population genomics in
non-model organisms. Trends in Ecology & Evolution 29:51–63.
Ellegren, H., C.R. Primmer & B.C. Sheldon. 1995. Microsatellite
‘evolution’: directionality or bias? Nature Genetics 11:360.
Esquivel-Bobadilla, S., O.A. Lozano-Garza, F.J. Garc´
I.D.L.A. Barriga-Sosa & M.L. Jim´
enez. 2013. Development and
characterization of 14 microsatellite loci in the beach wolf spider
(Arctosa littoralis), using next-generation sequencing. Conserva-
tion Genetics Resources 5:261–263.
Evanno, G., S. Regnaut & J. Goudet. 2005. Detecting the number of
clusters of individuals using the software STRUCTURE: a
simulation study. Molecular Ecology 14:2611–2620.
Evans, T.A. 1998. Offspring recognition by mother crab spiders with
extreme maternal care. Proceedings of the Royal Society of
London B 265:129–134.
Fadrosh, D.W., B. Ma, P. Gajer, N. Sengamalay, S. Ott, R.M.
Brotman & J. Ravel 2014. An improved dual-indexing approach
for multiplexed 16S rRNA gene sequencing on the Illumina MiSeq
platform. Microbiome 2:1.
Faircloth, B.C. 2008. Msatcommander: detection of microsatellite
repeat arrays and automated, locus-speciﬁc primer design.
Molecular Ecology Resources 8:92–94.
Falush D., M. Stephens & J.K. Pritchard. 2003. Inference of
population structure using multilocus genotype data linked loci
and correlated allele frequencies. Genetics 164:1567–1587.
´ndez, R., G. Hormiga & G. Giribet. 2014. Phylogenomic
analysis of spiders reveals nonmonophyly of orb weavers. Current
´ndez, R., R.J. Kallal, D. Dimitrov, J.A. Ballesteros, M.A.
Arnedo, G. Giribet & G. Hormiga. 2018. Phylogenomics,
diversiﬁcation dynamics, and comparative transcriptomics across
the Spider Tree of Life. Current Biology 28:1489–1497.
Fromhage, L., G. Uhl & J.M. Schneider. 2003. Fitness consequences
of sexual cannibalism in female Argiope bruennichi. Behavioral
Ecology and Sociobiology 55:60–64.
Ghislandi, P.G., S. Peka
´r, M. Matzke, S. Schulte-Doinghaus, T. Bilde
& C. Tuni. 2018. Resource availability, mating opportunity, and
sexual selection intensity inﬂuence the expression of male
alternative reproductive tactics. Journal of Evolutionary Biology
Gillespie, R.G. & G.S. Oxford. 1998. Selection on the color
polymorphism in Hawaiian happy-face spiders: evidence from
genetic structure and temporal ﬂuctuations. Evolution 52:775–783.
Gordon, A. & G.J. Hannon. 2010. Fastx-toolkit. FASTQ/A short-
reads preprocessing tools (unpublished) Online at http://
Gregory, T.R. 2001. Animal genome size database. 2001. Online at
Hataway, R.A., D.H. Reed & B.P. Noonan. 2011. Development of 10
KREHENWINKEL ET AL.—MICROSATELLITES AND GENOTYPING FOR SPIDERS 199
microsatellite loci in the wolf spider Arctosa sancterosae (Araneae:
Lycosidae). Conservation Genetics Resources 3:271–273.
Koﬂer, R., P. Orozco-terWengel, N. De Maio, R.V. Pandey, V.
Nolte, A. Futschik et al. 2011. PoPoolation: a toolbox for
population genetic analysis of next generation sequencing data
from pooled individuals. PloS ONE 6:e15925.
Krehenwinkel, H. & D. Tautz. 2013. Northern range expansion of
European populations of the wasp spider Argiope bruennichi is
associated with global warming–correlated genetic admixture and
population-speciﬁc temperature adaptations. Molecular Ecology
Krehenwinkel, H., M. Graze, D. R¨
odder, K. Tanaka, Y.G. Baba, C.
Muster et al. 2016a. A phylogeographical survey of a highly
dispersive spider reveals eastern Asia as a major glacial refugium
for Palaearctic fauna. Journal of Biogeography 43:1583–1594.
Krehenwinkel, H., D. R¨
odder, M. Na
c& M. Kuntner.
2016b. Rapid genetic and ecological differentiation during the
northern range expansion of the venomous yellow sac spider
Cheiracanthium punctorium in Europe. Evolutionary Applications
Krehenwinkel, H., D. R¨
odder & D. Tautz. 2015. Eco-Genomic
analysis of the poleward range expansion of the wasp spider
Argiope bruennichi shows rapid adaptation and genomic admix-
ture. Global Change Biology 21:4320–4332.
Kunz, K., S. Garbe & G. Uhl. 2012. The function of the secretory
cephalic hump in males of the dwarf spider Oedothorax retusus
(Linyphiidae: Erigoninae). Animal Behaviour 83:511–517.
Kunz, K., M. Witthuhn & G. Uhl. 2014. Do the size and age of
mating plugs alter their efﬁcacy in protecting paternity? Behav-
ioural Ecology and Sociobiology 68:1321–1328.
Lange, V., I. B¨
ohme, J. Hofmann, K. Lang, J. Sauter, B. Sch ¨
one et al.
2014. Cost-efﬁcient high-throughput HLA typing by MiSeq
amplicon sequencing. BMC Genomics 15:1.
Leese, F., C. Mayer & C. Held. 2008. Isolation of microsatellites from
unknown genomes using known genomes as enrichment templates.
Limnology & Oceanography: Methods 6:412–426.
Malausa, T., A. Gilles, E. Meglecz, H. Blanquart, S. Duthoy, C.
Costedoat et al. 2011. High-throughput microsatellite isolation
through 454 GS-FLX Titanium pyrosequencing of enriched DNA
libraries. Molecular Ecology Resources 11:638–644.
Mayer, C., M. Sann, A. Donath, M. Meixner, L. Podsiadlowski, R.S.
Peters et al. 2016. BaitFisher: A software package for multispecies
target DNA enrichment probe design. Molecular Biology and
Evolution 33:1875–1886. Online at https://doi.org/10.1093/
ecz, E., S.J. Anderson, D. Bourguet, R. Butcher, A. Caldas, A.
Cassel-Lundhagen et al. 2007. Microsatellite ﬂanking region
similarities among different loci within insect species. Insect
Molecular Biology 16:175–185.
Moodley, Y., J.F. Masello, T.L. Cole, L. Calderon, G.K. Muniman-
da, M.R. Thali et al. 2015. Evolutionary factors affecting the cross-
species utility of newly developed microsatellite markers in
seabirds. Molecular Ecology Resources 15:1046–1058.
Mouginot, P., J. Pr¨
ugel, U. Thom, P.O.M. Steinhoff, J. Kupryjano-
wicz & G. Uhl. 2015. Securing paternity by mutilating female
genitalia in spiders. Current Biology 25:1–5.
Mouginot, P., G. Uhl & L. Fromhage. 2017. Evolution of external
female genital mutilation: why do males harm their mates? Royal
Society Open Science 4:171195.
Muster, C., A. Herrmann, S. Otto & D. Bernhard. 2008. Zur
Ausbreitung humanmedizinisch bedeutsamer Dornﬁnger-Arten
Cheiracanthium mildei und C. punctorium in Sachsen und
Brandenburg (Araneae: Miturgidae). Arachnologische Mitteilun-
Nolte, A.W., K.C. Stemshorn & D. Tautz. 2005. Direct cloning of
microsatellite loci from Cottus gobio through a simpliﬁed
enrichment procedure. Molecular Ecology Notes 5:628–636.
Parmakelis, A., K. Balanika, S. Terzopoulou, F. Rigal, R.R. Beasley,
K.L. Jones et al. 2013. Development of 28 polymorphic
microsatellite markers for the endemic Azorean spider Sancus
acoreensis (Araneae, Tetragnathidae). Conservation Genetics
Peterson, B.K., J.N. Weber, E.H. Kay, H.S. Fisher & H.E. Hoekstra
2012. Double digest RADseq: an inexpensive method for de novo
SNP discovery and genotyping in model and non-model species.
PLoS ONE 7:e37135.
Planas, E., L. Bernaus & C. Ribera. 2014. Development of novel
microsatellite markers for the spider genus Loxosceles (Sicariidae)
using next-generation sequencing. Journal of Arachnology 42:315–
Pritchard, J.K., M. Stephens & P. Donnelly. 2000. Inference of
population structure using multilocus genotype data. Genetics
R Core Team 2016. R: A language and environment for statistical
computing. R Foundation for Statistical Computing, Vienna,
Austria. Online at https://www.R-project.org/
Raymond, M. & F. Rousset. 1995. GENEPOP on the Web (Version
3.4). Online at http://wbiomed.curtin.edu.au/genepop/ Updated
from Raymond & Rousset.
Rozen, S.H. & H. Skaletsky. 1999. Primer3 on the WWW for general
users and for biologist programmers. Pp. 365–386. In Bioinfor-
matics Methods and Protocols. Vol. 132: Methods in Molecular
Biology. (S. Misener & S.A. Krawetz eds.). Humana Press, New
Ruch, J., M. Dumke & J.M. Schneider. 2015. Social network
structure in group-feeding spiders. Behavioral Ecology & Sociobi-
Ruch, J., M.E. Herberstein & J.M. Schneider. 2014. Offspring
dynamics affect food provisioning, growth and mortality in a
brood-caring spider. Proceedings of the Royal Society of London
utten, K.B., I. Schulz, K. Olek & G. Uhl. 2001. Polymorphic
microsatellite markers in the spider Pholcus phalangioides isolated
from a library enriched for CA repeats. Molecular Ecology Notes
Sanggaard, K.W., J.S. Bechsgaard, X. Fang, J. Duan, T.F. Dyrlund,
V. Gupta et al. 2014. Spider genomes provide insight into
composition and evolution of venom and silk. Nature Communi-
afer, M.A., B. Misof & G. Uhl. 2008. Effects of body size of both
sexes and female mating history on male behaviour and paternity
success in a spider. Animal Behaviour 76:75–86.
otterer, C. 2000. Evolutionary dynamics of microsatellite DNA.
Schneider, J.M. 2014. Sexual cannibalism as a manifestation of sexual
conﬂict. In Sexual Conﬂict. (B. Rice & S. Gavrilets eds.). Cold
Spring Harbor Perspectives in Biology. doi: 10.1101/cshperspect.
Schneider, J.M. & M.C.D. Andrade. 2011. Mating behaviour and
sexual selection. Pp. 215–275. In Spider Behaviour: Variability and
Versatility. (M.E. Herberstein, ed.). Cambridge University Press,
Schneider, J., G. Uhl & M.E. Herberstein. 2015. Cryptic female
choice within the genus Argiope: A comparative approach. Pp. 55–
77. In Cryptic Female Choice in Arthropods: Patterns, Mecha-
nisms and Prospects. (A.V. Peretti & A. Aisenberg, eds.). Springer,
Schwager E.E., P.P. Sharma, T. Clarke, D.J. Leite, T. Wierschin, M.
Pechmann et al. 2017. The house spider genome reveals an ancient
whole-genome duplication during arachnid evolution. BMC
Settepani, V., M.F. Schou, M. Greve, L. Grinsted, J. Bechsgaard & T.
Bilde. 2017. Changes in mating system and life history traits with
the evolution of sociality leads to depleted genomic diversity at
200 JOURNAL OF ARACHNOLOGY
both population and species level. Molecular Ecology 26:4197–
Smith, B.T., M.G. Harvey, B.C. Faircloth, T.C. Glenn & R.T.
Brumﬁeld. 2013. Target capture and massively parallel sequencing
of ultraconserved elements (UCEs) for comparative studies at
shallow evolutionary time scales. Systematic Biology 63:83–95.
Starrett, J., S. Derkarabetian, M. Hedin, R.W. Bryson Jr., J.E.
McCormack & B.C. Faircloth. 2017. High phylogenetic utility of
an ultraconserved element probe set designed for Arachnida.
Molecular Ecology Resources 17:812–823.
Stolle, E., J.H. Kidner & R.F. Moritz. 2013. Patterns of evolutionary
conservation of microsatellites (SSRs) suggest a faster rate of
genome evolution in Hymenoptera than in Diptera. Genome
Biology and Evolution 5:151–162.
Suez, M., A. Behdenna, S. Brouillet, P. Gra¸ca, D. Higuet & G. Achaz.
2016. MicNeSs: genotyping microsatellite loci from a collection of
(NGS) reads. Molecular Ecology Resources 16:524–533.
Tamura, K., G. Stecher, D. Peterson, A. Filipski. & S. Kumar. 2013.
MEGA6: molecular evolutionary genetics analysis version 6.0.
Molecular Biology and Evolution 30:2725–2729.
Tuni, C., S. Goodacre, J. Bechsgaard & T. Bilde. 2012. Moderate
multiple parentage and low genetic variation reduces the potential
for genetic incompatibility avoidance despite high risk of
inbreeding. PLOSOne, 7(1): e29636. doi:10.1371/journal.pone.
Uhl, G., S.M. Zimmer, D. Renner & J.M. Schneider. 2015. Exploiting
a moment of weakness: male spiders escape sexual cannibalism by
copulating with molting females. Scientiﬁc Reports 5:16928, DOI:
Vartia, S., J.L. Villanueva-Ca˜
nas, J. Finarelli, E.D. Farrell, P.C.
Collins, G.M. Hughes et al. 2016. A novel method of microsatellite
genotyping-by-sequencing using individual combinatorial barcod-
ing. Royal Society Open Science 3:150565.
Yang, X., Y. Xu, T. Shah, H. Li, Z. Han, J. Li & J. Yan. 2011.
Comparison of SSRs and SNPs in assessment of genetic
relatedness in maize. Genetica 139:1045–1054.
Zhan, L., I.G. Paterson, B.A. Fraser, B. Watson, I.R. Bradbury, P.
Nadukkalam Ravindran et al. 2017. MEGASAT: automated
inference of microsatellite genotypes from sequence data. Molec-
ular Ecology Resources 17:247–256.
Zhang, J., K. Kobert, T. Flouri & A. Stamatakis. 2014. PEAR: a fast
and accurate Illumina Paired-End reAd mergeR. Bioinformatics
Zimmer, S.M., H. Krehenwinkel & J.M. Schneider. 2014. Rapid
range expansion is not restricted by inbreeding in a sexually
cannibalistic spider. PLoS ONE 7:e95963.
Manuscript received 6 March 2016, revised 7 January 2019.
The following data sets are available in the Dryad Digital repository
(Online at doi:10.5061/dryad.c4d7cc0)
1. 454 reads and Assemblies for all studied species
2. Isolated primer sequences for all studied spider species
3. Primer sequences, which have already been tested for
variability and PCR ampliﬁcation success.
4. Illumina reads for microsatellite genotyping of Hawaiian
happy face spider populations.
KREHENWINKEL ET AL.—MICROSATELLITES AND GENOTYPING FOR SPIDERS 201