ArticlePDF Available

Abstract and Figures

Whole genome sequencing is instrumental for the study of genome variation in natural populations, delivering important knowledge on genomic modifications and potential targets of natural selection at the population level. Large dormant eggbanks of aquatic invertebrates such as the keystone herbivore Daphnia, a microcrustacean widespread in freshwater ecosystems, provide detailed sedimentary archives to study genomic processes over centuries. To overcome the problem of limited DNA amounts in single Daphnia dormant eggs, we developed an optimized workflow for whole genome amplification (WGA), yielding sufficient amounts of DNA for downstream whole genome sequencing of individual historical eggs, including polyploid lineages. We compare two WGA kits, applied to recently produced Daphnia magna dormant eggs from laboratory cultures, and to historical dormant eggs of Daphnia pulicaria collected from Arctic lake sediment between 10 and 300 years old. Resulting genome coverage breadth in most samples was ~70%, including those from >100-year-old isolates. Sequence read distribution was highly correlated among samples amplified with the same kit, but less correlated between kits. Despite this, a high percentage of genomic positions with single nucleotide polymorphisms in one or more samples (maximum of 74% between kits, and 97% within kits) were recovered at a depth required for genotyping. As a by-product of sequencing we obtained 100% coverage of the mitochondrial genomes even from the oldest isolates (~300 years). The mitochondrial DNA provides an additional source for evolutionary studies of these populations. We provide an optimized workflow for WGA followed by whole genome sequencing including steps to minimize exogenous DNA.
Content may be subject to copyright.
Mol Ecol Resour. 2021;00:1–16.
|
1wileyonlinelibrary.com/journal/men
Received: 17 April 2021 
|
Revised: 3 September 2021 
|
Accepted: 7 September 2021
DOI: 10.1111/1755-0998.13524
RESOURCE ARTICLE
Refining the evolutionary time machine: An assessment of
whole genome amplification using single historical Daphnia
eggs
Christopher James O’Grady1,2,3| Vignesh Dhandapani3|
John K. Colbourne3| Dagmar Frisch3,4
This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium,
provided the original work is properly cited.
© 2021 The Authors. Molecular Ecology Resources published by John Wiley & Sons Ltd.
1School of Life Sciences, University of
Warwick, Coventry, UK
2Cell and Gene Therapy Catapult, London,
UK
3School of Biosciences, University of
Birmingham, Birmingham, UK
4Leibniz Institute of Freshwater Ecology
and Inland Fisheries (IGB), Berlin, Germany
Correspondence
Dagmar Frisch, Leibniz Institute of
Freshwater Ecology and Inland Fisheries
(IGB), Berlin, Germany.
Email: dagmar.frisch@igb-berlin.de
Funding information
Midlands Integrative Biosciences Training
Partnership; H2020 Marie Skłodowska-
Curie Actions, Grant/Award Number:
658714; NERC Biomolecular Analysis
Facility, Grant/Award Number: NBAF998
Abstract
Whole genome sequencing is instrumental for the study of genome variation in natu-
ral populations, delivering important knowledge on genomic modifications and po-
tential targets of natural selection at the population level. Large dormant eggbanks
of aquatic invertebrates such as the keystone herbivore Daphnia, a microcrustacean
widespread in freshwater ecosystems, provide detailed sedimentary archives to study
genomic processes over centuries. To overcome the problem of limited DNA amounts
in single Daphnia dormant eggs, we developed an optimized workflow for whole
genome amplification (WGA), yielding sufficient amounts of DNA for downstream
whole genome sequencing of individual historical eggs, including polyploid lineages.
We compare two WGA kits, applied to recently produced Daphnia magna dormant
eggs from laboratory cultures, and to historical dormant eggs of Daphnia pulicaria
collected from Arctic lake sediment between 10 and 300 years old. Resulting genome
coverage breadth in most samples was ~70%, including those from >100- year- old
isolates. Sequence read distribution was highly correlated among samples amplified
with the same kit, but less correlated between kits. Despite this, a high percentage
of genomic positions with single nucleotide polymorphisms in one or more samples
(maximum of 74% between kits, and 97% within kits) were recovered at a depth re-
quired for genotyping. As a by- product of sequencing we obtained 100% coverage of
the mitochondrial genomes even from the oldest isolates (~300 years). The mitochon-
drial DNA provides an additional source for evolutionary studies of these populations.
We provide an optimized workflow for WGA followed by whole genome sequencing
including steps to minimize exogenous DNA.
KEYWORDS
ancient DNA, Daphnia, population genomics, SNP analysis, whole genome sequencing
2 
|
    O ’GR ADY et al.
1 | INTRODUCTION
Ancient or historical genomic data from natural populations is a key
resource for an in- depth understanding of how organisms adapt to
their environment. This is one of the most compelling and challeng-
ing tasks in evolutionary ecology, especially regarding the current
unprecedented environmental change. A unique approach gaining
momentum is the study of propagules of various plant or animal taxa
preserved in layered aquatic sediments to reconstruct biological and
environmental history (Ellegaard et al., 2020; Orsini et al., 2013).
These propagules contain dormant embryos in early development
(inside, e.g., eggs, seeds, cysts), providing DNA that is degraded to
varying degrees, as well as intact DNA , and allowing the direct ob-
servation of evolutionary change across centuries or even millen-
nia (Brede et al., 2009; Cordellier et al., 2021; Frisch et al., 2014;
Härnström et al., 2011; Mergeay et al., 2006; Pollard et al., 2003;
Weider et al., 1997). The exploitation of such resources together
with modern molecular tools, targeting many key members of the
aquatic food web, is instrumental in the study of evolutionary pro-
cesses over thousands of generations in relation to environmental
change, and can potentially be performed at genomic resolution of
individual isolates.
With recent declines in sequencing costs due to development
of high- throughput technologies, whole genome sequencing (WGS)
has emerged as an important molecular tool in evolutionary biology
and has been applied to a plethora of different biological systems
(Dettman et al., 2012; Ellegren, 2014; Hohenlohe et al., 2018; Stiller
& Zhang, 2019). WGS allows the analysis of genetic variation at
thousands of genomic loci to test relationships between phenotypic
and genotypic adaptations in genome- wide association studies (De
La Torre et al., 2019; Rajpurohit et al., 2018; Sella & Barton, 2019).
At the population level, and in particular if long- term time series data
including from ancient DNA are available, WGS can provide invalu-
able genomic detail, shedding light on evolutionary patterns and
processes (Leonardi et al., 2017; Parks et al., 2015).
As one of the notable examples, the population genetics of
the ecological and genomic model Daphnia (Crustacea, Cladocera)
has been studied over historical time frames and associated with
changes in the lake environment, genotyping either individual eggs
(Brede et al., 2009; Frisch et al., 2014, 2016; Limburg & Weider,
2003; Orsini et al., 2012) or by WGS of pooled egg DNA (Cordellier
et al., 2021). WGS of individual eggs from sedimentar y archives
would allow long- term population genomic studies not only of
Daphnia, but also of other key members of the aquatic foodweb with
diapause stages that are preserved in aquatic sediments (e.g., vari-
ous cladocerans, copepods, rotifers, large branchiopods, as well as
algae or fungi) at high resolution.
A major obstacle for WGS of individual dormant eggs of key zoo-
plankton taxa is their limited cell number, and thus minute amount
of DNA. The dormant eggs of several planktonic crustaceans (co-
pepods, cladocerans) contain an embr yo in the late blastula or early
gastrula stage with between 500 and 3000 cells depending on taxon
(von Baldass, 1941; Chen et al., 2018; Reed et al., 2021), and even
fewer in rotifers (18– 160 cells, Boschetti et al., 2011). For example,
given a haploid genome size of ~200 Mb (Colbourne et al., 2011)
and ~1000 cells in a dormant embryo of Daphnia pulex (von Baldass,
1941), the DNA content of a triploid embryo can be estimated at
~60 0 pg. Low- input libr ary method s for WG S of individual zooplank-
ton specimens are available but currently require an input of at least
0.35 ng DNA. While this is a small fraction of the DNA that can be
extracted from adult zooplankton, averaging 16 ng per individual
(Beninde et al., 2020), it exceeds the total amount of DNA available
from dormant material. DNA loss during extraction and purification
from historical material can be as high as 49%– 90%, depending on
DNA fragmentation (Barta et al., 2014). Extracting enough DNA
from dormant Daphnia embryos thus poses technical limitations
that are difficult to overcome, further reducing the likelihood that
enough DNA can be extracted from dormant material. The situation
is exacerbated for historical dormant Daphnia eggs or those of other
taxa, due to DNA degradation, posing additional problems for DNA
sequencing (Rizzi et al., 2012).
To overcome the problem of DNA limitation in individual eggs,
it is possible to combine eggs from individual sediment strata for a
pooled sequencing approach. Such a strategy has several main dis-
advantages (Schlötterer et al., 2014): it is not a suitable method to
infer haplotypes and linkage disequilibrium; low- frequency variants
can be difficult to distinguish from sequencing errors; and technical
difficulties (pipetting, DNA quantification) can result in unequally
represented individuals in the DNA pool, especially at small sample
sizes. It can also lead to information loss on individual genotypes and
accuracy of population genomic parameters such as FST estimates
(Dorant et al., 2019).
An alternative approach to gain sufficient amounts of genetic
starting material is by performing whole genome amplification
(WGA). This method uses cell material without prior DNA extraction,
thus minimizing potential loss of DNA during the extraction process,
and amplifies genomic DNA from extremely low starting concen-
trations in the picogram range. However, a previous study that ap-
plied WGA to individual, dormant Daphnia eggs had limited success
with only one of three eggs producing amplified Daphnia DNA (Lack
et al., 2018). Multiple displacement amplification (MDA), a widely
used PCR- free WGA method, utilizes a high- fidelity φ29 DNA poly-
merase which extends from hexamer primers that randomly bind
to targets across the genomic template (Dean et al., 2001, 2002).
This results in the generation of large DNA products, with an av-
erage length of ~10 kb (capable of reaching over 100 kb), that can
have a strong coverage of the target genome (Blanco et al., 1989;
Handyside et al., 2004; Lasken & Egholm, 2003; Paez, 2004). MDA is
often favoured over PCR- WGA techniques, for example degenerate
oligonucleotide- primed PCR, as PCR- based methods can result in
the production of small DNA fragments (>1 kb; Telenius et al., 1992;
Wells et al., 1999; Zhang et al., 1992) that contain several nonspe-
cific amplification artefacts (Cheung & Nelson, 1996). Additionally,
PCR- WGA methods can show a significant amplification bias to-
wards specific loci, and consequently products may not give a com-
plete coverage of loci (Dean et al., 2002). MDA is highly sensitive and
   
|
 3
O’GRA DY et al .
is particularly vulnerable to DNA contamination, which can compete
or co- amplify with the desired DNA template during WGA and cause
issues during downstream analyses (Blainey & Quake, 2011; Woyke
et al., 2011). Great care must therefore be taken to eliminate sources
of contamination during the amplification step.
In contrast to Beninde et al. (2020), who provided a detailed low-
input library preparation protocol for adult zooplankton individuals
from both fresh and maximally 29- year- old ethanol- preserved mate-
rial, our goal was to develop an optimized WGA- WGS workflow for
several- centuries- old, dormant egg isolates including improved de-
contamination steps. Extending a study of individual Daphnia where
WGA of dormant eggs had limited success (Lack et al., 2018), we
use recently produced Daphnia magna dormant eggs from laboratory
cultures (days old) and Daphnia pulicaria dormant eggs isolated from
lake sediment (between 10 and 300 years old), and compared two
commercially available single- cell WGA kits based on MDA tech-
nology. We test the success of reducing exogenous DNA through
the application of different concentrations and durations of bleach,
or several washes with PBS (phosphate- buffered saline) to samples
prior to WGA .
In a sequencing experiment, we analyse mapping efficiency and
genome- wide read distribution and compare these between species
and eggs of various age for a total number of 16 dormant eggs aged
up to 300 years old. We compare read distribution patterns, cover-
age breadth and uniformity, and identify contaminants. Finally, we
test the utility of these kits for detecting genomic variants in both
nuclear and mitochondrial genomes.
2 | MATERIALS AND METHODS
2.1  |  Egg collection
All eggs used for whole genome amplification were isolated from
ephippia of two species: Daphnia magna Straus, 1820, and Daphnia
pulicaria Forbes, 1893. For D. magna, we used eggs from recently
pr oduce d ephi ppia (sexu al eg gs) that ar e routi nely re move d from lab-
oratory cultures maintained in the Daphnia facility of the University
of Birmingham, UK (DM1- DM7, unknown origin). Ephippia of
Arctic, triploid populations of Daphnia pulicaria (asexually produced
eggs) were collected in 2015 from sediment of two lakes in West
Greenland (Kangerlussuaq area). Details on the lakes and sediment
dating can be found in Dane et al. (2020). Briefly, we sampled ephip-
pia from sediment corresponding to several historical time periods
in two lakes: Lake SS4 (Braya Sø): c. 2010, c. 1880, c. 1720, and Lake
SS381: c. 2010, c. 1840 (Table 1).
2.2  |  Pre- WGA preparation and cleaning of
Daphnia eggs
Eggs were removed from ephippia (decapsulated) and transferred
to sterile 1× PBS shortly before use. Decapsulated eggs were in-
spected under a stereomicroscope to ensure that eggs were in
good condition (judged by colour and appearance). Visually un-
damaged eggs were washed in a 5% or 10% wash solution made
TAB LE 1  Species identity and specifics of dormant eggs used as template in whole genome amplification
Sample Species Ploidy Age Pretreatment kit
DNA
(µg)
DM1 D. magna 2nLaboratory 10% , < 2 s REPLI- g 22.27
DM2 D. magna 2nLaboratory 10%, 20 s REPLI- g 28.66
DM3 D. magna 2nLaboratory 10%, < 2 s REPLI- g 3 7.93
DM4 D. magna 2nLaboratory 5%, < 2 s REPLI- g 33.58
DM5 D. magna 2nLaboratory 5%, 20 s REPLI- g 22.80
DM6 D. magna 2nLaboratory 10%, < 2 s Trueprime 7.06
DM7 D. magna 2nLaboratory 10%, < 2 s Trueprime 7.48
DP1 D. pulicaria 3n~10 years (SS4) 10%, < 2 s REPLI- g 39.4 8
DP2 D. pulicaria 3n~140 years (SS4) 5%, < 2 s Trueprime 9.21
DP3 D. pulicaria 3n~140 years (SS4) 10%, < 2 s Trueprime 9.18
DP4 D. pulicaria 3n~300 years (SS4) 10%, < 2 s REPLI- g 42.85
DP5 D. pulicaria 3n~300 years (SS4) 10%, < 2 s REPLI- g 26.50
DP6 D. pulicaria 3n~10 years (SS1381) 1× PBS Trueprime 8.30
DP7 D. pulicaria 3n~10 years (SS1381) 1× PBS Trueprime 6.00
DP8 D. pulicaria 3n~180 years (SS1381) 1× PBS Trueprime 6.64
DP9 D. pulicaria 3n~180 years (SS1381) 1× PBS Trueprime 5.14
Note: Additional information is given on pretreatment (% of the bleach solution and exposure time, or 1× PBS buffer, e.g., "10% < 2 s" indicates
exposure for less than 2 s to a 10% bleach solution made from 12% industrial bleach), the applied WGA kit and the total product of WGA- DNA
obtained. SS4 and SS1831 are two lakes in West Greenland near Kangerlussuaq (for details see Section 2) from which sediment cores with ephippia
were extracted.
4 
|
    O ’GR ADY et al.
from industrial strength (12%) bleach. Exposure to the wash so-
lution was either instantly (<2 s) or for 20 s (Table 1), followed
by five separate rinses in sterile 1× PBS to remove any remaining
bleach. Alternatively, eggs were washed by five to eight rinses in
1× PBS , by pla cing a row of PBS drop let s on a gl ass slid e, an d wash-
ing each egg individually by carefully and repeatedly drawing them
up with a pipette (sterile tip) in each of the droplet s. Rinse controls
contained the PBS solution used for the last rinse, while negative
controls contained sterile PBS. Bleaching (including the final PBS
rinse) was performed in a SCANLAF Mars Safety Class 2 laminar
flowhood in sterile conditions. The alternative procedure of PBS
rinsing was performed in a clean, dedicated room with thorough
bleaching of all surfaces prior to processing eggs. Bleached and
rins ed egg s wer e kept on ice for brief per iod s in sterile 1× PBS unti l
further processing.
2.3  |  Whole genome amplification
WGA was performed using two PCR- free kits: Expedeon TruePrime
Single Cell WGA kit (hereafter: TruePrime), and Qiagen REPLI- g
Single Cell Kit (hereafter: REPLI- g). Positive controls (extracted
Daphnia DNA) were included in WGA. Rinse controls and negative
controls were included to monitor possible amplification of contam-
inating DNA. Prior to WGA, egg membranes were pierced with a
sterile 10- μl pipette tip to allow exposure of embryonic cells, and
kept in the respective amount of 1× PBS required for the first step
of the reaction in each test kit. DNA concentration in WGA products
was quantified with a microplate Reader (Tecan infinite F200 pro),
or a Qubit 2.0 Fluorometer (Invitrogen) and the Qubit dsDNA HS
Assay kit (Invitrogen). The size distribution of WGA products was
determined by agarose gel electrophoresis to analyse the impact of
pretreatment steps (bleaching or washing in PBS) on the DNA frag-
ments produced by WGA.
2.4  |  Whole genome library
preparation and sequencing
WGA samples DM1DM7 (D. magna eggs) an d DP1– DP5 (D. pulicaria
eggs) were used to prepare single- end (SE) libraries with an insert
size of 300 bp using a PCR- free workflow with the KAPA HyperPrep
Kit (Roche) following the manufacturer's instructions. Sequencing
of 100- bp SE libraries was performed on the Illumina HiSeq2500
platform at the Environmental Omics sequencing facility, University
of Birmingham, UK. WGA samples DP6– DP9 (D. pulicaria eggs)
were used to prepare paired- end (PE) libraries with an insert size of
350 bp with the TruSeq DNA PCR- free gel- free library preparation
kit (Illumina) according to the manufacturer's instructions. PE library
preparation and sequencing (150 - bp PE libraries) was performed
at Edinburgh Genomics, The University of Edinburgh, UK. The raw
sequence files are available in the open- access repository Zenodo
(O'Grady et al., 2021).
2.5  |  Bioinformatic and statistical analysis
All analyses involving R packages were completed with R version
3.6.2 (R CoreTeam, 2019). The nature of the study system with lim-
ited access to historical eggs, led to some imbalance of the study
design. We therefore refrained from a formal factorial statistical
analysis for the comparison of different pretreatments and sequenc-
ing strategies.
2.5.1  |  Quality control and mapping
We used fastqc (Andrews, 2015) to check read quality, followed by
adapter trimming and removal of leading and trailing low- quality
bases with trimmomatic (Bolger et al., 2014). Following quality control,
reads were mapped using bwa- mem with default settings (Li & Durbi n,
2009) to the respective reference genome assembly (D. magna ge-
nome assembly daphmag2.4, GenBank accession GCA_001632505.1;
Daphnia pulex genome assembly [http://genome.jgi.doe.gov/Dappu
1/Dappu1.downl oad.html] [Colbourne et al., 2011]; D. pulex mito-
chondrial genome, GenBank Accession NC _00 084 4 [Crease, 1999]).
Mapping statistics were computed with qualimap (García- Alcalde
et al., 2012) prior to variant calling. Duplicate reads were removed
from mapped reads using markduplicates from the Picard Toolkit
(Broad Institute, 2019).
2.5.2  |  Nuclear DNA variant calling and analysis
Nuclear single nucleotide polymorphisms (SNPs) were called in
D. magna using the available SE libraries. SNPs in the triploid Arctic
D. pulicaria egg s were called onl y from PE samples because se quenc-
ing depth of SE samples was insufficient for calling variants in a trip-
loid organism (Maruki & Lynch, 2017). Nuclear and mirochondrial
variants were called with freebayes version 1.3.2 (Garrison & Marth,
2012), excluding reads with a mapping quality <40, base quality <24
and a minimum alternate allele fraction of 0.01, ploidy = 2 (D. magna
nuclear DNA [ncDNA], D. pulic aria mitochondrial DNA [mtDNA]) and
ploidy = 3 in D. pulicaria ncDNA. After variant calling, nuclear SNPs
were hard- filtered with vcffilter (Garrison, 2016) applying all of the
following settings: “QUAL > 1to ensure the exclusion of variants
of very low quality, “QUAL/AO > 10” to include only variants where
each observation contributes at least 10 log units (~Q10 per read),
“SAF > 0 & SAR > 0” to avoid strand bias, and “RPR > 1 & RPL > 1” to
require at least two reads on each side of the variant.
Genomic positions with high- confidence SNPs present in one or
more samples were compared between selected samples to assess
the percentage of loci that could be called in all selected samples
(i.e., that were amplified and sequenced at the depth required for
genotyping). This comparison was used primarily to estimate the
repeatability of WGA and subsequent WGS, and thus for the result-
ing capacity to call variants at the multisample level. For D. magna,
we compared the four samples with the highest number of SNPs
   
|
 5
O’GRA DY et al .
(two REPLI- g amplified samples: DM2, DM3, two TruePrime ampli-
fied samples: DM6, DM7). For D. pulicaria, we compared all four PE
samples (only TruePrime amplified). Results were visualized with the
R packages eulerr version 6.1.0 (Larsson, 2020) and ggvenndiagram
version 0.3 (Gao & Yi, 2019).
Transition- to- transversion ratios for SNPs (Ti:Tv) were calculated
after applying a minor allele frequency (MAF) threshold of 0.05, and
not allowing missing data (R packages seqarray 1.26.2, Zheng et al.,
2017; and seqvartools 1.24.1, Gogarten et al., 2021).
2.5.3  |  mtDNA variant calling and analysis
To analyse mtDNA, we used all available D. pulicaria samples
(DP1– DP9). For mitochondrial SNPs, the same filters as for nuclear
SNPs were applied except "QUAL/AO > 10" to avoid filtering calls
of the alternate allele from samples with PE sequencing due to their
consistently higher depth compared to the SE samples. Identity- by-
state (IBS) was calculated by applying an MAF threshold of 0.05, not
allowing missing data (R package seqarray 1.26.2, Zheng et al., 2017).
SNPs in D. pulicaria mtDNA were visualized with the packages cir-
clize version 0.4.11 (Gu et al. 2014), and snprelate (Zheng et al., 2021).
2.5.4  |  Read distribution
This analysis focused on reads mapped to the N50 scaffolds of the
D. magna and D. pulex genomes. Coverage depth was normalized be-
tween samples of binned reads (bin size 10 or 100 kb, normalized
reads = number of reads per bin/average number of reads across
bins). Normalized read coverage was visualized with the packages cir-
clize version 0.4.11 (Gu, 2014), and ggplot2 version 3.3.2 (Wickham,
2016). Read distribution was compared within species between
samples by correlation analysis (Pearson's correlation coefficient)
of normalized binned reads. For this purpose, we removed a single
outlier present in all D. pulicaria samples (position 420,001430,000,
scaffold 38). We tested uniformity of read distribution according to
the standard model for random sequencing by fitting the distribu-
tion of normalized read coverage to a Poisson distribution (Lander &
Waterman, 1988). Uniformity of read distribution was quantified by
the evenness score (Oexle 2016). In short, this metric is calculated as
the coefficient of variation for non- normalized data.
2.5.5  |  Outlier identification and exogenous DNA
The REPLI- g amplified D. pulicaria samples DP1, DP4 and DP5 con-
tained obvious outliers with preferential amplification. They were
defined as regions with a mapping rate 10 times higher than the
mean normalized count (100- kb bins). Sequences belonging to these
outlier regions were extracted and searched against the nucleotide
database with blastn. Exogenous DNA was identified in two sam-
ples with low mapping efficiency (DP4, DP5). For this, unmapped
reads were called from bam files with samtools version 1.4 (Li &
Durbin, 2009) and converted to PE fastq files using bedtools version
2 (Quinlan & Hall, 2010). Following this step, soapdenovo- 127mer ver-
sion 2 (Luo et al., 2012) was used to de novo- assemble the unmapped
reads with kmer size of 23 bp and default parameters. The resulting
contigs were searched against the NCBI nucleotide database with
blastn (using the command line "blastn - task megablast - db NCBI_nt_
db - query infile - evalue 1e- 10 0 - out outfile - max_target_seqs 1 - num_
threads 10 - outfmt "6 qseqid sseqid sciname qlen slen qstart qend sstart
send length evalue pident nident mismatch gaps"). For graphical repre-
sentation we used krona (Ondov et al., 2011).
3 | RESULTS
3.1  |  Whole genome amplification
WGA products were separated by agarose gel electrophoresis to
determine the impact of pretreatment steps (bleaching or washing
in PBS) on WGA product fragment sizes. Regardless of the pretreat-
ment process, strong bands for DNA fragments >10 kb were de-
tected in all samples, suggesting a minimal impact of pretreatment
on WGA product size (examples in Figure S1). WGA products ob-
tained from one of the rinse controls (1× PBS, DNA amplified from
the last PBS wash of an unbleached sample) also produced high-
intensity bands (from 500 bp to >10 kb). This DNA probably resulted
from amplified contaminant DNA carried over from one of the pre-
vious serial washes, suggesting that PBS washes do remove exog-
enous DNA from egg surfaces, but that a higher number of washes
may be advisable for more complete removal. WGA of rinse controls
obtained from the PBS after bleach- washing did not yield any prod-
uct, suggesting that the application of bleach completely removes
external, exogenous DNA. No amplified DNA was present in any of
the negative controls (sterile PBS).
Mean WGA- DNA concentration was lower in the samples am-
plified by TruePrime (7.38 µg) in comparison with REPLI- g (31.76 µg;
Table 1). These values were within the ranges suggested by the re-
spective WGA kit manufacturers (~40 µg WGA- DNA for REPLI- g,
and 3– 4 µg when starting from a single cell for TruePrime).
3.2  |  Read mapping to nuclear and mitochondrial
reference genomes
Both SE (DP1– DP5) and PE (DP6– DP9) sequencing was performed
for D. pulicaria, the latter with about 10- fold higher read numbers
and related higher coverage depth (Table 2). For Daphnia magna,
only SE libraries were sequenced (DM1– DM7). With one excep-
tion (DM4), all D. magna libraries mapped to the reference genome
with high efficiency >98%, indicating no clear pattern of mapping
efficiency to the nuclear genome related to the applied pretreat-
ment in the several- days- old dormant eggs produced in cultures
(DM1– DM7). Lower mapping efficiency and/or a smaller fraction of
6 
|
    O ’GR ADY et al.
TAB LE 2  Mapping details and number of variants called in (a) nuclear DNA of Daphnia magna (DM), and Daphnia pulicaria (DP), and (b) mitochondrial DNA of D. pulicaria
Nuclear genome D. magna and D. pulicaria
Sample Age Pretreatment Kit Seq Number of reads Mapped reads (%)
Coverage
breadth (%) Coverage depth Evenness score SNP loci
DM1 Laboratory 10%, <2 s R - g SE 8,488,215 98.72 74.95 60.76 301,779
DM2 Laboratory 10%, 20 s R - g SE 17, 74 2 , 8 8 2 98 .76 77. 4 3 13 0 .76 414,782
DM3 Laboratory 10% , <2 s R - g SE 24, 0 69, 767 98.74 78.63 18 0.75 4 30,909
DM4 Laboratory 5%, <2 s R - g SE 38 ,765 55.73 1.47 0.02 0. 55 NA
DM5 Laboratory 5%, 20 s R - g SE 10,167,137 98.64 76 .36 70.7 7 33 9, 023
DM6 Laboratory 10% , <2 s TP SE 6,392,740 99.18 66.75 50.68 99,9 27
DM7 Laboratory 10%, <2 s TP SE 14,8 00, 247 99.04 78.05 11 0.71 357, 8 4 7
DP1 ~10 years 10%, <2 s R - g SE 12,990,625 48.36 41.45 30.38 NAa
DP2 ~140 years 5%, <2 s TP SE 12,088,977 87.36 50.11 50.62 NAa
DP3 ~140 years 10% , <2 s TP SE 16,643,733 84.62 51.8 2 70.66 NAa
DP4 ~300 years 10%, <2 s R - g SE 8,579,357 0.36 0.33 0.01 0.23 NAa
DP5 ~300 years 10%, <2 s R - g SE 7,950, 28 3 2 .55 0.93 0.07 0.16 NAa
DP6 ~10 years 1× PBS TP PE 85,442,043 90.53 69.21 51 0.74 1,732,886
DP7 ~10 years 1× PBS TP PE 134,031,117 90.95 69. 5 8 78 0.74 1,734,987
DP8 ~180 years 1× PBS TP PE 95,665,657 91.65 6 9.50 58 0.73 1,742,4 63
DP9 ~180 years 1× PBS TP PE 93,262,683 91.85 69.5 5 57 0.73 1,741,613
(b) Mitochondrial genome D. pulicaria
Sample Age (years) Seq Mapped reads (%) Coverage breadth (%) Coverage depth SNP loci
DP1 ~10 SE 73.87 99. 97 61,542 20 0
DP2 ~140 SE 0.68 99.94 498 200
DP3 ~140 SE 0.25 99.93 252 200
DP4 ~300 SE 0.13 9 9.93 71 200
DP5 ~300 SE 4.55 99. 95 2315 20 0
DP6 ~10 PE 3.32 99.97 25,740 200
DP7 ~10 PE 0.86 99.9 5 10,240 200
DP8 ~180 PE 0.72 99. 97 6155 200
DP9 ~180 PE 0.62 99.95 5197 200
Note: Age = see Table 1; kit = WGA kit (R- g = REPLI- g, TP = TruePrime); Seq = Sequencing strategy (SE = single end, PE = paired end); Number of reads = total number of reads used for mapping; Mapped
reads = fraction of reads mapped to the respective reference genome; Coverage breadth = fraction of reference genome with at least 1× coverage; Average coverage depth = mean number of reads per
genomic position; Evenness score = evenness of coverage which quantifies uniformity of read distribution; SNP loci = number of single nucleotide variants compared to the respective reference genome.
aNo variants called in these samples due to insufficient sequencing depth for triploid variants.
   
|
 7
O’GRA DY et al .
the genome covered was observed in the SE sequenced sedimen-
tary eggs DP1– DP5 (Table 2). These eggs were pretreated with vari-
ous bleach concentrations, suggesting a potential negative effect of
bleaching for sedimentary eggs regardless of their age, compared
to the several- days- old laboratory- produced eggs. Due to the ex-
perimental setup in which all bleached sedimentary samples were
SE sequenced, and all nonbleached Daphnia pulicara eggs were PE
sequenced, it is not possible to separate effects of pretreatment
and sequencing effort on the mapping efficiency within this species.
Nevertheless, contamination was low in the nonbleached sedimen-
tary eggs DP6– DP9, with ~90% mapping rate, and ~70% coverage
breadth (Table 2a), indicating efficient removal of exogenous DNA
from egg surfaces with PBS and maintenance of DNA integrity be-
fore WGA.
High- throughput sequencing of libraries prepared from WGA-
DNA from 13 of the 16 tested dormant Daphnia eggs successfully
mapped with between 48% and 99% of reads (mean 88%, median
92%) in both SE and PE libraries to the respective Daphnia nuclear
genomes, suggesting that WGA largely resulted in amplification of
the target DNA (Table 2a). This was achieved for ephippia produced
in laboratory cultures as well as those collected from lake sediment
of different age with up to ~180- year- old eggs. Maximum cover-
age breadth (i.e., the fraction of the nuclear genome covered) was
similar in both Daphnia species (between 70% and 80%, Table 2a).
Moderate mapping efficiency was recorded for two relatively young
eggs at 48% (DP1, from ~10- year- old sediment) and 56% (DM4,
from a laboratory culture) and resulted in coverage of a lower frac-
tion of the nuclear reference genomes (lower coverage breadth).
Mapping to the nuclear genome failed almost entirely in the two
oldest eggs where WGA was attempted (DP4 and DP5, ~300 years
old, Table 2a), although the WGA- DNA yield was similar to that of
other eggs (Table 1), suggesting amplification of contaminant DNA.
However, egg age did not have a consistent effect on mapping effi-
ciency; for example, SE libraries for the ~10- year- old D. pulicaria egg
mapped with an efficiency of 48% (DP1), while the SE libraries from
two ~140- year- old eggs mapped with an efficiency of 84%– 87%
(DP2, DP3). Likewise, PE libraries of D. pulicaria eggs were mapped
with high efficiency (~90%) regardless of age (Table 2).
In contrast to the nuclear genome, we found that reads obtained
from all historical eggs of D. pulicaria including the oldest samples
(~300 years old) could be mapped to the mitochondrial genomes of
the target species, resulting in high average coverage depth between
71× (DP4) and 60,000× (DP1) and a near to 100% coverage breadth
of the mitochondrial genome (Table 2b). The extremely high cover-
age observed for DP1 indicates preferential amplification of mtDNA
encoded in the mitochondrial and/or nuclear genome (see also next
section).
3.3  |  Read distribution
Patterns of genome- wide normalized read distribution differed be-
tween species, WGA kits and sequencing strategy (Figure 1). Results
for the evenness score that quantifies uniformity of read distribu-
tion did not suggest differences for libraries obtained with either
WGA kit from the laboratory- produced eggs of D. magna (Figures 1a
and S2). In contrast, the read distribution pattern differed markedly
in D. pulicaria (Figure 1c e). Samples from this species were repre-
sented by historical, sedimentary eggs and therefore the DNA may
have been compromised to different degrees, providing inferior am-
plification substrate. The evenness scores obtained from sequenc-
ing sedimentary eggs (only D. pulicaria, Figure 1c,d) suggest lower
uniformity of read distribution (Table 2a; Figure S2) in the REPLI- g
amplified samples (DP1, DP4, DP5), while historical eggs amplified
with TruePr ime (DP2, DP3, DP6DP9) provide d a more unifor m cov-
erage with less pronounced outliers even in samples >100 years old
(Figure 1c– e).
Read distribution patterns in D. pulicaria (Figure 1c– e) revealed
several regions of the genome that were preferentially amplified.
These outliers included genomic regions with sequences, for ex-
ample, of the Pokey transposon or of several introns, but also for
segments of mtDNA encoded in the nuclear genome (Table S1). The
most extreme outlier identified in TruePrime- amplified DNA (a seg-
ment of nuclear mtDNA on scaffold 38, Table S1) was observed in
both SE and PE libraries of the historical eggs.
To test the repeatability of read distribution patterns between
samples within each species, we computed pairwise correlation
coefficients (excluding the outlier on scaffold 38) for read counts
within 10- kb bins (Figures 2, S3 and S4). For both species, samples
amplified by the same WGA kit were strongly correlated (mean r:
D. magna REPLI- g = .731; D. magna TruePrime = .767; D. pulicaria
REPLI- g [SE] = .643; D. pulicaria TruePrime [SE] = .470, D. pulicaria
TruePrime [PE] = .938). Weak correlations were observed between
samples amplified using different kits (mean r: D. magna = .149;
D. pulicaria = .085).
Owing to over- represented regions and preferentially amplified
regions detailed above, the read distributions of none of the ampli-
fied samples had a significant fit to a Poisson distribution expected
under the standard model for random sequencing (Table S2).
3.4  |  Exogenous DNA
In samples where WGA failed almost entirely to produce target DNA
(DP4 and DP5, Table 2a), but yielded a similar amount of DNA as for
other eggs, we used a blast search to identify the taxon origin of the
unmapped reads. This DNA was identified mostly as contaminant
DNA from various bacterial and invertebrate taxa, as well as human
DNA, but diversity and percentages of the contaminant taxa dif-
fered between the two eggs (Figure 3).
3.5  |  Variant calling
For D. magna we called single nucleotide variants (SNPs) in the six
samples with >6 million reads (DM1– DM3, DM5– DM7). For the
8 
|
    O ’GR ADY et al.
triploid D. pulicaria eggs we could only use the four PE samples for
confident SNP calling because deeper sequencing with higher cov-
erage depth is needed for higher ploidy genomes.
The number of identified SNPs per sample in D. magna was be-
tween 99,927 and 430,909 in comparison with the D. magna reference
genome (Table 2). In the D. magna samples with the highest number of
cal led SN P s (REP L I- g : DM2, DM 3 , Tru ePrime: DM7, Tabl e 2, Fi g u re 4a) ,
a tot al of 442,976 unique and shared SNP loci were recorded. All three
samples could be genotyped at the required depth at 327,802 of these
positions (74% of the potential genomic positions of SNPs in these
three samples). SNP loci recorded in the lower- depth D. magna sample
DM6 were nested almost entirely in those of the higher- depth sample
DM7 (both amplified with TruePrime, Figure 4b). In D. pulicaria, the
total number of SNP loci identified in the four PE samples in compari-
son to the D. pulex reference genome was 1,758,439. All four samples
could be genotyped with the required depth at 1,712,889 (or 97.4%)
of these genomic positions (Figure 4c).
We estimated the Ti:Tv ratio (Figure 4d) to gauge the variability
of this metric between the samples studied and to assess whether
their values were within the ranges reported for Daphnia in general.
We found that the ratios differed between species but that variation
among samples of the same species was small (D. pulicaria: 1.30 in
all four samples, D. magna: 1.43– 1.47). Differences of ti:tv ratios be-
tween amplification kits (only D. magna) were small (REPLI- g: 1.43
1.45, TruePrime: 1.461.47).
SNP calling in the mitochondrial genomes (Figure 5) was per-
formed with all available D. pulicaria samples. The analysis revealed
the presence of 200 biallelic SNPs that differed between the two
lake populations sampled. However, within- population samples
were identical with the exception of DP1 (Lake SS4) and DP9 (Lake
1381) that differed from other samples of their respective popula-
tion by a single SNP.
4 | DISCUSSION
WGA and subsequent WGS are staples of modern single- cell ge-
nome studies, and are widely applied to the study of human diseases
FIGURE 1 Visualization of the genome- wide normalized coverage pattern resulting from WGA- DNA of dormant eggs of Daphnia magna
and D. pulicaria, with comparison of two commercial MDA kits. Bars in circular graphs (N50 scaffolds) represent normalized coverage in
100- kb bins, and 10- kb bins in linear graphs. (a) D. magna dormant eggs from cultures (SE libraries), amplified with REPLI- g (DM1DM5, light
blue) and TruePrime (DM6, DM7, orange). (b) Detail of (a) for the largest D. magna scaffold (scaffold 512). (c) D. pulicaria sedimentary eggs of
different age (SE libraries). Amplified with REPLI- g (three outer rings: DP1, DP4, DP5) and with TruePrime (two inner rings: DP2, DP3). Red
arrows point to outliers with preferred amplification in DP1, DP4 and DP5 (details in Table S1). (d) D. pulicaria sedimentary eggs of different
age amplified with TruePrime (PE libraries). From outer to inner rings: DP6DP9. (e) Detail of (c) for the largest D. pulex scaffold (scaffold 1).
Sample order of circular graphs identical to linear graphs
5
24
84
115
139
190
1581
1663
2121
2190
2227
2244
2372
2385
2452
2486
2569
2581
2850
2861
2865
2994
3025
3124
3258
3276
3326
3376
7
243
248
311
337
389
446
512
547
568
626
642
687
725
781
868
872
915
930
944
966
996
1005
1036
1253
1274
1348
1361
1409
1574
1579
1764
1867
1937
2066
2076
2101
REPLI-g, fresh dormant egg, culture
TruePrime,fresh dormant egg,culture
(a) (b)
(c)
(d) (e)
DP6 DP1DP4DP2 DP5DP3DP7DP8DP9
0100 200300 400
Scaffold 1
DM6
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
0
200
400
DM1DM2DM4DM3DM5
0100 200300
Scaffold 512
DM7
D. pulicaria
normalised
coverage
(SE)
REPLI-g, ~10yold egg
REPLI-g, ~300yold egg
TruePrime,~140y old egg
Outlier
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
39
41
43
44
45
47
49
50
38
40
42
46
48
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
39
41
43
44
45
47
49
50
38
40
42
46
48
1
TruePrime,~10 yold egg
TruePrime,~180 yold egg
D. pulicaria
normalised
coverage
(PE)
D. pulicaria
normalised coverage (10k bins)
D. magna
normalised
coverage
(SE)
D. magnanormalisedcoverage (10k bins)
   
|
 9
O’GRA DY et al .
(Huang et al., 2015). Other promising but less common applications
include phylogenomics (Ahrendt et al., 2018; Zhang et al., 2019) and
metagenomics of microbial communities (Xu & Zhao, 2018). Suitable
application for population genomics and evolutionary studies using
Daphnia dormant eggs has been suggested (Lack et al., 2018), but
to date a comprehensive study involving the comparison between
multiple sedimentary eggs of different historical age and species, ap-
plying different pretreatments and amplification kits is not available.
MDA has superior qualities when the goal is the discovery of single
nucleotide variants, due to high fidelity of the φ29 polymerase and
associated low error rates, while PCR- based WGA such as MALBAC
may perform better for detecting copy number variation (de Bourcy
et al., 2014; Chen et al., 2014). We therefore tested two MDA- WGA
kits to provide a detailed workflow (Figure 6) for successful and re-
peatable amplification and sequencing of DNA for variant calls from
dormant eggs of Daphnia.
4.1  |  Pretreatment to minimize exogenous DNA
Sources of DNA contamination may originate from the WGA rea-
gents, or are introduced when handling samples (Rinke et al., 2014).
Contamination may also derive from nontarget exogenous DNA on
the biological isolate (often of microbial origin), which is especially
common in historical samples containing highly degraded ancient
DNA (Pilli et al., 2013). Decontamination procedures of equipment,
which include the application of bleach and UV light, can be per-
formed prior to WGA to prevent the amplification of exogenous
DNA (Woyke et al., 2011). It is also recommended to use a thor-
oughly decontaminated laminar flowhood during all stages of WGA ,
preferably situated in a dedicated clean room.
Overall, our data suggest that amplification of exogenous DNA
can be kept to a minimum when all careful steps are followed to avoid
contamination. However, in historical samples where DNA is already
damaged, exogenous DNA may be present in higher amounts than
the target DNA , and thus be preferentially amplified, and even over-
whelm the amplification process. This was particularly obvious in the
oldest samples tested here (~300 years old). Apart from egg age,
DNA integrity may also be highly dependent on the preservation
conditions of the lake sediment: DNA preservation can vary strongly
between different lakes and may be related to a number of variables,
including temperature, salt concentration and pH (Ellegaard et al.,
2020; Giguet- Covex et al., 2019).
Eggs produced in the laboratory that were only several days old
did not show any signs of DNA degradation even after bleaching
with a higher bleach concentration, and/or for a longer exposure
time to bleach. In comparison to these samples, the DNA integrity
and the resulting quality of WGA- DNA sedimentary eggs pretreated
with diluted bleach appeared to be impaired. A possible explanation
is the probable presence of microfissures in the egg membranes of
historical, sedimentary eggs that are prone to increase with dormant
egg age. To test this idea, microscopic studies comparing eggs of dif-
ferent age are needed.
Washing with PBS did not appear to affect the DNA in dormant
eggs of D. pulicaria of different historical age (~10 and ~180 years
old), which produced high- quality WGA- DNA. However, the ob-
served higher quality of DNA libraries prepared from the PBS-
treated eggs may also be largely due to a higher sequencing effort
for these samples, as the PBS treatment was only performed prior
to PE sequencing. It is also possible that these eggs had a generally
higher quality than those used for bleaching, which were sampled
from sediment of a neighbouring lake. Based on our findings, we
cannot conclude with certainty that a pretreatment with PBS is su-
perior to bleach, and further tests should be performed. However,
serial washes with PBS appeared to effectively remove contamina-
tion in the tested samples, and thus may be a more cautious alterna-
tive to bleaching to remove possible contaminants on sedimentary
egg surfaces that could overwhelm DNA amplification, particularly
if the external structure of eggs and their DNA is more strongly im-
paired with increasing age.
4.2  |  Mapping success and patterns of read
distribution
Mapping of WGA- DNA produced with either of the two kits tested
here was highly successful in most isolates, with a median of 92% of
FIGURE 2 Correlation matrix (Pearson's r) of pairwise
comparisons between normalized read counts of samples in 10-
kb bins. (a) Pairwise comparisons of Daphnia magna samples. (b)
Pairwise comparisons of Daphnia pulicaria samples. All pairwise
comparisons had an associated p- value < .01
DP7
DP9
DP8
DP6
DP3
DP2
DP5
DP4
DP1
PE
PESE
SE
DP4DP5 DP2DP3 DP6DP7 DP8
SE
SE
REPLI-g
TruePrime
DM7
DM6
DM5
DM4
DM3
DM2
DM6
DM5
DM4DM3DM2DM1
(a)
(b)
Pearson'sr
0.25
0.50
0.75
0.90
Pearson'sr
0.25
0.50
0.75
0.90
10 
|
    O ’GR ADY et al.
reads mapped to their respective reference genomes, even of eggs
as old as 180 years. Coverage breadth in most samples was between
70% and 80%. These values are similar to that previously reported
for a dormant Daphnia egg, but below those obtained from Daphnia
bulk sequencing (Lack et al., 2018), and higher than of MALBAC
amplified individual sperm cells of Daphnia (53%, Xu et al., 2015).
Indeed, incomplete genome coverage is commonly observed in
WGA data in various taxa (e.g., de Bourcy et al., 2014; Huang et al.,
2015; Picher et al., 2016).
Deviation of read distributions from a Poisson distribution ex-
pected under a random sequencing model (Lander & Waterman,
1988) was observed in all samples. However, this is not unexpected
as it has been described previously that this model is inadequate for
single- cell sequencing due to the possibility of locus dropout (Daley
& Smith, 2014). Despite this, we found the regions of the genome
that are amplified to be remarkably repeatable among samples of
the same species, with highly significant correlation coefficients
within amplification kits (average r of ~.7). The below- average r val-
ues were generally associated with failed target amplification or low
read depth overall, specifically in the SE samples; but above- average
values were found in the PE samples with high read depth. However,
correlations bet ween WGA kits were not stron g, so for good compa-
rability between samples, it is recommended to apply only one kit.
4.3  |  Variant calling in nuclear and
mitochondrial genomes
The highly reproducible patterns of read distribution within
kits are largely responsible for the success of SNP calling across
samples, demonstrating the suitability of this method for popu-
lation genomic applications, particularly for dormant eggs, and
thus for its utility for studying genome evolution using sediment
archives. In D. magna, >70% of the SNP positions could be called
in all three samples with >14 million reads. Perhaps not surpris-
ingly these values were even higher in the PE samples of D. puli-
caria with >80 million reads, suggesting in general that sequencing
strategy and depth strongly influence fidelity of the variant call
also when applied to WGA- DNA.
Transition to transversion ratios have been suggested as a qual-
ity indicator for human SNP discovery (Wang et al., 2015). However,
because these ratios vary between species, for example averaging
1.54 in several strains of D. magna (Ho et al., 2020), or 0.45 in C. ele-
gans (Denver et al., 2009), comparisons should be made within spe-
cies. Our range between 1.43 to 1.47 for nuclear DNA of D. magna
was well within the ratio measured by Ho et al. (2020), and for D. pu-
licaria (Ti:Tv 1.30) was similar to that reported for the closely related
D. pulex (1.58, Keith et al., 2016). In this study, we can also apply the
Ti:Tv ratio as an indicator for the reliability of the WGA procedure
within and across kits, with highly similar values for D. magna eggs,
or almost identical values for D. pulicaria.
Other studies have identified possible limitations of the reliabil-
ity of WGA , such as coverage uniformity, reproducibility and allelic
dropout rate (de Bourcy et al., 2014; Huang et al., 2015). The sub-
strates that thes e studie s use d wer e cell li nea ges from whi ch in div id-
ual cells were subjected to single cell WGA (scWGA) and compared
to bulk sequencing of a multicellular sample from the same lineage.
For historical isolates of dormant eggs from the sediment, such a
strategy cannot be applied. However, an effective substitute to test
the reliability of SNPs from these historical samples was the use of
FIGURE 3 Taxon identity of nontarget DNA amplified from two Daphnia pulicaria dormant eggs. (a) DP5, (b) DP4. Labels only shown for
those taxa that constitute >1% of the total exogenous sequences identified
(a)
(b)
   
|
  11
O’GRA DY et al .
asexually produced dormant eggs such as those of the Arctic D. pu-
licaria population from which our samples originated. WGA of these
samples allowed 97% of all SNP loci detected in the PE sequences to
be genotyped in all triploid isolates. Preliminary analysis of variation
between these genotyped eggs showed a maximum difference of
3% of the roughly 1.7 million SNPs between individuals (unpublished
data). Encouraging results were also reported for MDA- amplified
DNA from individual adults of a D. pulicaria clone compared to DNA
from pooled individuals of the same clone (Lack et al., 2018). These
authors found only a slight loss of heterozygosity in the amplified
DNA, but concordance of structural variation (insertions, deletions,
duplications and translocation, but not of inversions) between both
sample types (Lack et al., 2018).
An added benefit of WGA with both MDA kits tested here was
the possibility of obtaining high coverage of the full mitochon-
drial genome for both species. This is of particular interest for the
historical samples from which full mitochondrial genomes and
high- quality SNP calls could be retrieved even of the oldest samples
(~300- year- old eggs). This opens a promising avenue for gathering
information of genome- wide mutation rates and spectra of the mi-
tochondrial genome stored in sedimentary archives across extended
time periods and thousands of generations, probably surpassing the
time range tested here.
5 | CONCLUSION
Although the method described here was tested on dormant eggs of
Daphnia, it could be widely applied to sedimentary dormant stages
of a variety of taxa, potentially providing access to the evolutionary
history not only of single taxa but also to that of entire aquatic com-
munities. Such taxa could include key members of the plankton in the
freshwater and marine to hypersaline ecosystems that produce dor-
mant propagules at an early embryonic stage with a limited number
FIGURE 4 Results of SNP analysis in Daphnia magna and D. pulicaria. Venn diagrams (a– c) represent the number of genomic positions
where SNPs were located in one or more samples and that were sequenced in all or a subset of samples (shared and unique genomic positions).
This comparison was used to estimate the repeatability of whole genome amplification and thus for the resulting capacity to call variants at
multisample level. (a) Comparison between three D. magna samples with the highest number of SNPs identified, amplified by REPLI- g (DM2,
DM3) and TruePrime (DM7). (b) Comparison between the two D. magna samples amplified by TruePrime (DM6, DM7). (c) Comparison between
four D. pulicaria samples, amplified with TruePrime. (d) Transition:transversion ratio analysed for SNPs of both Daphnia species
DM2∩DM3 ∩DM7:
268,071 DM3
repli-G TruePrime
DM7DM6
DM2
DM3:
5,188
DM2:
2,027
DM7:
3,801
DM2∩
DM3:
78,314
DM3∩DM7:
79,336
DM3∩DM7:
3,706
Daphnia magna
Daphnia pulicaria Transition:Transversion ratio
(a)
(b)
(c) (d)
DM6:
3,561
DM7:
261,481
DM6∩D7:
96,366
DP6
DP7 DP8
DP9
1965
(0.11%)
1823
(0.1%)
7130
(0.41%)
1712889
(97.41%)
3536
(0.2%)
1735
(0.1%) 2814
(0.16%)
994
(0.06%)
3766
(0.21%)
3500
(0.2%)
7765
(0.44%)
2054
(0.12%)
3536
(0.2%)
2244
(0.13%)
2688
(0.15%)
0.0
0.5
1.0
1.5
TiTv
species
magna
pulicaria
DM1
DM2
DM3
DM5
DM6
DM7
DP6
DP7
DP8
DP9
R
R
e
e
p
p
l
l
i
i
-
-
G
G
T
T
r
r
u
u
e
e
P
P
r
r
i
i
m
m
e
e
12 
|
    O ’GR ADY et al.
of cells, such as the dormant eggs of calanoid copepods, cysts of the
brine shrimp Artemia, algal dormant stages and plant seeds. Other
methods that have successfully been applied to individual small
planktonic crustaceans, such as low- DNA- input sequencing libraries
(B enin de et al., 2020) , cou l d be te s ted on dorma nt st ages isola ted fr om
historical sediment layers. However, their library preparation proto-
col, which was optimized for adult specimens, requires an amount of
purified template DNA that may exceed the DNA content available
from dormant embryos. In such cases, a technology which does not
require a DNA extraction step such as the MDA methodology tested
here could be a more practical approach. Since the protocol presented
here does not require DNA extraction but uses the entire egg mate-
rial as template, the risk of losing precious material can be minimized.
Our data reveal that both of the tested amplification kits pro-
vided high- quality DNA for most Daphnia egg isolates, and that the
amplified DNA could efficiently be applied to WGS and subsequent
genome- wide studies at the population level. However, differences
were observed with respect to the age of dormant eggs, where our
results suggest a superior performance of TruePrime (compared with
REPLI- g) for application to eggs of sedimentar y origin and thus to
prospectively degraded DNA. A possible explanation could be that
the primase TthPrimPol used in the TruePrime kit shows translesion
activity, allowing re- initiation of the replication fork when encoun-
tering damaged DNA, and thus continued amplification of damaged
substrate (Picher & Blanco, 2014). Due to the similarity of TthPrimPol
to human PrimPol, another mechanism could explain our results:
human PrimPol can reprime DNA synthesis following a lesion, allow-
ing the φ29 DNA polymerase to continue the amplification process
close to the region in which the lesion was found (Mourón et al.,
2013), possibly improving evenness during the process.
FIGURE 5 SNP positions and pairwise clustering of individuals by IBS of nine Daphnia pulicaria mitochondrial genomes recovered
from historical sedimentary dormant eggs from two lakes in West Greenland (Lake SS1381, green label, and SS4, blue label). Samples
with yellow labels were amplified with TruePrime, and the remaining three with REPLI- g. (a) Circular plot of detected SNP loci. The
four outer rings represent samples from Lake SS1381, the five inner rings from SS4. SNP colours are labelled according to their state
(turquoise = homozygous [variant allele], grey = homozygous [reference allele], blue = heteroplasmic [both reference and variant
allele present]). (b) Hierarchical clustering tree resulting from the fraction of identical genomic positions (identity- by- state) in the nine
mitochondrial genomes compared
   
|
  13
O’GRA DY et al .
Ultimately, our data indicate that for optimal results, preliminary
trials are recommended using both kits tested here (and if possible
others) on the dormant egg population in question.
ACKNOWLEDGEMENTS
D.F. received funding from the European Union’s Horizon 2020
research and innovation programme under the Marie Skłodowska-
Curie grant agreement No. 658714 and NERC Biomolecular
Analysis Facility Pilot Project Grant NBAF998. C.O.G. received
funding from the Midlands Integrative Biosciences Training
Partnership (MIBTP). We are grateful to Stephen Kissane for prep-
aration and sequencing of SE libraries, and to Caroline Sewell for
supplying D. magna ephippia from the Daphnia facility, University
of Birmingham. PE library preparation and sequencing were carried
out by Edinburgh Genomics, the University of Edinburgh, which is
partly supported with core funding from NERC (UKSBS PR18037).
FIGURE 6 Recommended workflow
for whole genome amplification
and whole genome sequencing with
downstream variant call and variant
filtering. Recommendation for sequencing
depth of higher ploidy genomes see
Maruki and Lynch (2017). Steps that need
further testing are marked by a red line.
For further details see text
fresh egg,
impeccable
quality
REPLI-g
scWGA
TruePrime
scWGA
TruePrime
scWGA
serialPBS
wash (5-10x)
sedimentary
egg, variable
quality
transferegg to PBS volumerequired by WGA kit
individual
dormant egg
pierce egg with sterile pipettetip (e.g. 10μl tip)to
breakmembrane and expose contents
PCR-free paired end librarypreparation and HTP
sequencingtodesired depth(min. 5-10x for
diploids,>20xfor triploids)
10' bleach wash
final PBS rinse(s)
QualityControl
Raw reads:
FastQC,Trimming
(Adapter, low
quality bases)
Mapping:
BWA-mem
BIOIFORMATICS
(example workflow)
PRE-TREATMENT
,AGW
,SEIRARBIL
WGS
QualityControl Mapping:
qualimap (mapping stats)
laminarflow hood
VariantCall: freebayes
(diploids & polypoids)
hardfilterwith vcffilter for
diploids & polyploids
SNPanalysis: various
toolsfor population
genomic analysis, e.g.
SeqArray forIBS(diploids
& polyploids)
14 
|
    O ’GR ADY et al.
CONFLICT OF INTEREST
The authors declare no conflicts of interest.
AUTHOR CONTRIBUTIONS
D.F. conceived the idea and obtained funding for the study. D.F.,
C.O.G. and J.K.C. designed the experiments. C.O.G. performed the
laboratory work. D.F. and C.O.G. analysed the data and wrote the
paper with input from J.K.C. and V.D.
DATA AVAILAB ILITY STATE MEN T
The sequence data supporting the findings reported here are availa-
ble in the open access repository Zenodo at https://doi.org/10.5281/
zenodo.5256276.
ORCID
Vignesh Dhandapani https://orcid.org/0000-0002-5745-2409
John K. Colbourne https://orcid.org/0000-0002-6966-2972
Dagmar Frisch https://orcid.org/0000-0001-9310-2230
REFERENCES
Ahrendt, S. R., Quandt, C. A., Ciobanu, D., Clum, A., Salamov, A.,
Andreopoulos, B., Cheng, J.- F., Woyke, T., Pelin, A., Henrissat, B.,
Reynolds, N. K., Benny, G. L., Smith, M. E., James, T. Y., & Grigoriev,
I. V. (2018). Leveraging single- cell genomics to expand the fungal
tree of life. Nature Microbiology, 3(12), 1417– 1428. htt ps://doi .
o r g / 1 0 . 1 0 3 8 / s 4 1 5 6 4 - 0 1 8 - 0 2 6 1 - 0
Andrews, S. (2015). FASTQC a quality control tool for high throughput
sequence data. Babraham Institute. https://www.Bioin forma tics.
B a b r a h a m . A c . U k / P r o j e c t s / F a s t q c /
Barta, J. L., Monroe, C., Teisberg, J. E., Winters, M., Flanigan, K., & Kemp, B.
M. (2014). One of the key characteristics of ancient DNA, low copy
number, may be a product of its extraction. Journal of Archaeological
Science, 46, 281– 289. https://doi.org/10.1016/j.jas.2014.03.030
Beninde, J., Möst, M., & Meyer, A. (2020). Optimized and affordable
high- throughput sequencing workflow for preserved and nonpre-
served small zooplankton specimens. Molecular Ecology Resources,
20(6), 1632– 1646. https://doi.org/10.1111/1755- 0998.13228
Blainey, P. C., & Quake, S. R. (2011). Digital MDA for enumeration of
total nucleic acid contamination. Nucleic Acids Research, 39(4), e19.
https://doi.org/10.1093/nar/gkq1074
Blanco, L., Bernad, A., Lázaro, J. M., Martín, G., Garmendia, C., & Salas,
M. (1989). Highly efficient DNA synthesis by the phage ϕ 29 DNA
polymerase. Journal of Biological Chemistry, 264(15), 8935– 8940.
h t t p s : // d o i . o r g / 1 0 . 1 0 1 6 / s 0 0 2 1 - 9 2 5 8 ( 1 8 ) 8 1 8 8 3 - x
Bolger, A. M., Lohse, M., & Usadel, B. (2014). Trimmomatic: A flexible
trimmer for Illumina sequence data. Bioinformatics, 30(15), 2114–
2120. https://doi.org/10.1093/bioin forma tics/btu170
Boschetti, C., Leasi, F., & Ricci, C. (2011). Developmental stages in diapausing
eggs: An investigation across monogonont rotifer species. Hydrobiologia,
662, 149– 155. https://doi.org/10.1007/s1075 0- 010- 0490- 6
Brede, N., Sandrock, C., Straile, D., Spaak, P., Jankowski, T., Streit, B., &
Schwenk, K. (2009). The impact of human- made ecological changes
on the genetic architecture of Daphnia species. Proceedings of the
National A cademy of Science s of the United States of Ame rica, 106(12),
4758– 4763. https://doi.org/10.1073/pnas.08071 87106
Broad Institute. (2019). Picard toolkit. Broad Institute, GitHub Repository.
https://Github.Com/Broad insti tute/Picar d/Relea ses/Tag/2.25.0
Chen, L., Barnett, R. E., Horstmann, M., Bamberger, V., Heberle, L., Krebs,
N., Colbourne, J. K., Gómez, R., & Weiss, L. C. (2018). Mitotic activ-
ity patterns and cytoskeletal changes throughout the progression
of diapause developmental program in Daphnia. BMC Cell Biology,
19(1), 30. https://doi.org/10.1186/s1286 0- 018- 0181- 0
Chen, M., Song, P., Zou, D., Hu, X., Zhao, S., Gao, S., & Ling, F. (2014).
Comparison of multiple displacement amplification (MDA) and mul-
tiple annealing and looping- based amplification cycles (MALBAC)
in single- cell sequencing. PLoS One, 9(12), e114520. https://doi.
org/10.1371/journ al.pone.0114520
Cheung, V. G., & Nelson, S. F. (1996). Whole genome amplification using
a degenerate oligonucleotide primer allows hundreds of genotypes
to be performed on less than one nanogram of genomic DNA.
Proceedings of the National Academy of Sciences, 93(25), 14676–
14679. https://doi.org/10.1073/pnas.93.25.14676
Colbourne, J. K., Pfrender, M. E., Gilber t, D., Thomas, W. K., Tucker,
A., Oakley, T. H., & Boore, J. L. (2011). The ecoresponsive ge-
nome of Daphnia pulex. Science, 331(6017), 555– 561. https://doi.
org/10.1126/scien ce.1197761
Cordellier, M., Wojewodzic, M. W., Wessels, M., Kuster, C., & von Elert,
E. (2021). Next- generation sequencing of DNA from resting eggs:
Signatures of eutrophication in a lake’s sediment. Zoology, 145,
125895. https://doi.org/10.1016/j.zool.2021.125895
Crease, T. J. (1999). The complete sequence of the mitochondrial genome
of Daphnia pulex (Cladocera: Crustacea). Gene, 233( 12 ) , 8 9 – 9 9 .
h t t p s : // d o i . o r g / 1 0 . 1 0 1 6 / S 0 3 7 8 - 1 1 1 9 ( 9 9 ) 0 0 1 5 1 - 1
Daley, T., & Smith, A. D. (2014). Modeling genome coverage in single-
cell sequencing. Bioinformatics, 30(22), 3159– 3165. https://doi.
org/10.1093/bioin forma tics/btu540
Dane, M., Anderson, N. J., Osburn, C. L., Colbourne, J. K., & Frisch, D.
(2020). Centennial clonal stability of asexual Daphnia in Greenland
lakes despite climate variability. Ecology and Evolution, 10(24),
14178– 14188. https://doi.org/10.1002/ece3.7012
de Bourcy, C. F. A., De Vlaminck, I., Kanbar, J. N., Wang, J., Gawad, C., &
Quake, S. R. (2014). A quantitative comparison of single- cell whole
genome amplification methods. PLoS One, 9(8), e105585. https://
doi.org/10.1371/journ al.pone.0105585
De La Torre, A. R., Wilhite, B., & Neale, D. B. (2019). Environmental
genome- wide association reveals climate adaptation is shaped by
subtle to moderate allele frequency shifts in loblolly pine. Genome
Biology and Evolution, 11(10), 2976– 2989. https://doi.org/10.1093/
gbe/evz2 20
Dean, F. B., Hosono, S., Fang, L., Wu, X., Faruqi, A. F., Bray- Ward, P.,
Sun, Z., Zong, Q., Du, Y., Du, J., Driscoll, M., Song, W., Kingsmore,
S. F., Egholm, M., & Lasken, R. S. (2002). Comprehensive human
genome amplification using multiple displacement amplification.
Proceedings of the National Academy of Sciences of the United States
of America, 99(8), 5261– 5266. https://doi.org/10.1073/pnas.08208
9499
Dean, F. B., Nelson, J. R ., Giesler, T. L., & Lasken, R. S. (2001). Rapid am-
plification of plasmid and phage DNA using Phi29 DNA polymerase
and multiply- primed rolling circle amplification. Genome Research,
11(6), 1095– 1099. https://doi.org/10.1101/gr.180501
Denver, D. R., Dolan, P. C., Wilhelm, L. J., Sung, W., Lucas- Lledo, J. I.,
Howe, D. K., Lewis, S. C., Okamoto, K., Thomas, W. K., Lynch, M.,
& Baer, C. F. (2009). A genome- wide view of Caenorhabditis el-
egans base- substitution mutation processes. Proceedings of the
National Academy of Sciences, 106 (38), 16310– 16314. https://doi.
org/10.1073/pnas.0904 8 95106
Dettman, J. R., Rodrigue, N., Melnyk, A. H., Wong, A., Bailey, S. F., & Kassen,
R. (2012). Evolutionary insight from whole- genome sequencing of
experimentally evolved microbes. Molecular Ecology, 21(9), 2058–
2077. https://doi.org/10.1111/j.1365- 294X.2012.05484.x
Dorant, Y., Benestan, L., Rougemont, Q., Normandeau, E., Boyle, B.,
Rochette, R., & Bernatchez, L. (2019). Comparing Pool- seq, Rapture,
and GBS genotyping for inferring weak population structure: The
American lobster (Homarus americanus) as a case study. Ecology and
Evolution, 9(11), 6606– 6623. https://doi.org/10.1002/ece3.5240
   
|
  15
O’GRA DY et al .
Ellegaard, M., Clokie, M. R. J., Czypionka, T., Frisch, D., Godhe, A., Kremp,
A., Letarov, A., McGenity, T. J., Ribeiro, S., & John Anderson, N.
(2020). Dead or alive: Sediment DNA archives as tools for track-
ing aquatic evolution and adaptation. Communications Biology, 3(1),
1 6 9 . h t t p s : / /d o i . o r g / 1 0 . 1 0 3 8 / s 4 2 0 0 3 - 0 2 0 - 0 8 9 9 - z
Ellegren, H. (2014). Genome sequencing and population genomics in
nonmodel organisms. Trends in Ecology and Evolution, 29(1), 51– 63.
https://doi.org/10.1016/j.tree.2013.09.008
Frisch, D., Morton, P. K., Culver, B. W., Edlund, M. B., Jeyasingh, P. D., &
Weider, L. J. (2016). Paleogenetic records of Daphnia pulicaria in
North American lakes reveal the impact of cultural eutrophication.
Global Change Biology, 23(2), 708– 718. https://doi.org/10.1111/
gcb.13 445
Frisch, D., Morton, P. K., Roy Chowdhury, P., Culver, B. W., Colbourne, J.
K., Weider, L. J., & Jeyasingh, P. D. (2014). A millennial- scale chroni-
cle of evolutionary responses to cultural eutrophication in Daphnia.
Ecology Letters, 17(3), 360– 368. https://doi.org/10.1111/ele.12237
Gao, C.- H., & Yi, L. (2019). ggVennDiagram: A “ggplot2” im-
plement of Venn Diagram. https://github.com/gaosp ecial/ ggVen
nDiagram
García- Alcalde, F., Okonechnikov, K., Carbonell, J., Cruz, L. M., Götz,
S., Tarazona, S., Dopazo, J., Meyer, T. F., & Conesa, A. (2012).
Qualimap: Evaluating next- generation sequencing alignment data.
Bioinformatics, 28(20), 2678– 2679. https://doi.org/10.1093/bioin
forma tics/bts503
Garrison, E. (2016). Vcflib, a simp le C++ library for parsing and manipulating
VCF files. Retrieved from https://github.com/vcfli b/vcflib
Garrison, E. & Marth, G. (2012). Haplotype- based variant detection from
short- read sequencing. ArXiv Preprint ArXiv:1207.3907.
Giguet- Covex, C., Ficetola, G. F., Walsh, K., Poulenard, J., Bajard, M.,
Fouinat, L., Sabatier, P., Gielly, L., Messager, E., Develle, A. L.,
David, F., Taberlet, P., Brisset, E., Guiter, F., Sinet, R., & Arnaud, F.
(2019). New insights on lake sediment DNA from the catchment:
Importance of taphonomic and analytical issues on the record qual-
ity. Scientific Reports, 9(1), 14676. https://doi.org/10.103 8/s4159
8 - 0 1 9 - 5 0 3 3 9 - 1
Gogar ten, S. M., Zheng, X., & Stilp, A . (2021). SeqVarTools: Tools for vari-
ant data. R package version 1.30.0. https://github.com/smgog arten/
Se qVa rTo ols
Gu, Z., Gu, L., Eils, R., Schlesner, M., & Brors, B. (2014). Circlize implements
and enhances circular visualization in R. Bioinformatics, 30(19),
28112812. https://doi.org/10.1093/bioin forma tics/btu393
Handyside, A. H., Robinson, M. D., Simpson, R. J., Omar, M. B., Shaw, M.
A., Grudzinskas, J. G., & Rutherford, A. (2004). Isothermal whole
genome amplification from single and small numbers of cells: A
new era for preimplantation genetic diagnosis of inherited dis-
ease. Molecular Human Reproduction, 10 (10), 767772. https://doi.
org/10.1093/moleh r/gah101
Härnström, K., Ellegaard, M., Andersen, T. J., & Godhe, A. (2011). Hundred
years of genetic structure in a sediment revived diatom popula-
tion. Proceedings of the National Academy of Sciences of the United
States of America, 10 8(10), 4252– 4257. https://doi.org/10.1073/
pnas.10135 28108
Ho, E. K. H., Macrae, F., Latta, L. C., McIlroy, P., Eber t, D., Fields, P. D.,
& Schaack, S. (2020). High and highly variable spontaneous muta-
tion rates in Daphnia. Molecular Biology and Evolution, 37(11), 3258–
3266 . https://doi.org/10.1093/molbe v/msaa142
Hohenlohe, P. A., Hand, B. K., Andrews, K. R., & Luikart, G. (2018).
Population genomics provides key insights in ecology and evolu-
tion. In O. Rajora (Ed.), Population genomics. Springer. https://doi.
org/10.1007/13836_2018_20
Huang, L., Ma, F., Chapman, A., Lu, S., & Xie, X. S. (2015). Single- cell whole-
genome amplification and sequencing: Methodology and applications.
Annual Review of Genomics and Human Genetics, 16(1), 79– 102. https://
doi.org/10.1146/annur ev- genom - 09041 3- 025352
Keith, N., Tucker, A. E., Jackson, C. E., Sung, W., Lucas Lledó, J. I., Schrider,
D. R., & Lynch, M. (2016). High mu tatio nal rates of la rge- sca le dupli-
cation and deletion in Daphnia pulex. Genome Research, 26(1), 60–
69. https://doi.org/10.1101/gr.191338.115
Lack, J. B., Weider, L. J., & Jeyasingh, P. D. (2018). Whole ge-
nome amplification and sequencing of a Daphnia resting
egg. Molecular Ecology Resources, 18(1), 118– 127. https://doi.
org /10.1111/1755- 0998 .12720
Lander, E. S., & Waterman, M. S. (1988). Genomic mapping by finger-
printing random clones: A mathematical analysis. Genomics, 2(3),
231– 239. https://doi.org/10.1016/0888- 7543(88)900 07 - 9
Larsson, J. (2020). eulerr: Area- proportional euler and venn diagrams
with ellipses. R package version 6.1.0. Retrieved from https://
c r a n . r - p r o j e c t . o r g / p a c k a g e =euler r.
Lasken, R. S., & Egholm, M. (2003). Whole genome amplification:
Abundant supplies of DNA from precious samples or clinical
specimens. Trends in Biotechnology, 21(12), 531– 535. https://doi.
org/10.1016/j.tibte ch.2003.09.010
Leonardi, M., Librado, P., Der Sarkissian, C., Schubert, M., Alfarhan, A.
H., Alquraishi, S. A., Al- Rasheid, K. A. S., Gamba, C., Willerslev, E.,
& Orlando, L. (2017). Evolutionary patterns and processes: Lessons
from ancient DNA. Systematic Biology, 66(1), e1e29. https://doi.
org/10.1093/sysbi o/syw059
Li, H., & Durbin, R. (2009). Fast and accurate short read alignment with
Burrows- Wheeler transform. Bioinformatics, 25(14), 17541760.
https://doi.org/10.1093/bioin forma tics/btp324
Limburg, P. A., & Weider, L. J. (2002). “Ancient” DNA in the resting egg
bank of a microcrustacean can serve as a palaeolimnological data-
base. Proceedings of the Royal Society of London Series B- Biological
Sciences, 269(1488), 281– 287.
Lu o, R., Li u, B., Xi e, Y., Li , Z., Hua ng, W., Yu an, J., He , G., Che n, Y., Pan ,
Q. I., Liu, Y., Tang, J., Wu, G., Zhang, H., Shi, Y., Liu, Y., Yu, C.,
Wang, B. O., Lu, Y., Han, C., … Wang, J. (2012). SOAPdenovo2:
An empirically improved memory- efficient short- read de novo
assembler. GigaScience, 1(1), 18. https://doi.org/10.1186/
2 0 4 7 - 2 1 7 X - 1 - 1 8
Maruki, T., & Lynch, M. (2017). Genotype calling from population-
genomic sequencing data. G3: Genes|genomes|genetics, 7(5), 1393–
1404. https://doi.org/10.1534/g3.117.039008
Mergeay, J., Ver sc huren, D., & De Mee ster, L. (20 06). Invasion of an asex-
ual Ame ric an wat er flea clon e thr oug hou t Afr ica and rapid disp lac e-
ment of a native sibling species. Proceedings of the Royal Societ y B-
Biological Sciences, 273(1603), 2839– 2844.
Mourón, S., Rodriguez- Acebes, S., Martínez- Jiménez, M. I., García-
Gómez, S., Chocrón, S., Blanco, L., & Méndez, J. (2013). Repriming
of DNA synthesis at stalled replication forks by human PrimPol.
Nature Structural & Molecular Biology, 20 (12), 1383– 1389. https://
doi.org/10.1038/nsmb. 2719
O'Gra dy, C. , Dhand apa ni, V., Colb ourne, J.K ., & Fris ch, D. (2021). Data fr om:
Refining the evolutionary time machine: an assessment of whole ge-
nome amplification using single historical Daphnia eggs. Molecular
Ecology Resources. https://doi.org/10.5281/zenodo.5256276
Ondov, B. D., Bergman, N. H., & Phillippy, A. M. (2011). Interactive
metagenomic visualization in a Web browser. BMC Bioinformatics,
12(1), 3 85. https://doi.org/10.1186/1471- 2105- 12- 385
Orsini, L., Schwenk, K., De Meester, L., Colbourne, J. K., Pfrender, M. E.,
& Weider, L. J. (2013). The evolutionary time machine: Forecasting
how populations can adapt to changing environments using dor-
mant propagules. Trends in Ecology & Evolution, 28, 274– 282.
https://doi.org/10.1016/j.tree.2013.01.009
Orsini, L., Spanier, K. I., & De Meester, L. (2012). Genomic signature of
natural and anthropogenic stress in wild populations of the wa-
terflea Daphnia magna: Validation in space, time and experimen-
tal evolution. Molecular Ecology, 21(9), 2160– 2175. https://doi.
org /10.1111/j.1365- 294X. 2011.05429.x
16 
|
    O ’GR ADY et al.
Paez, J. G. (2004). Genome coverage and sequence fidelity of 29
polymerase- based multiple strand displacement whole genome
amplification. Nucleic Acids Research, 32(9), e71. https://doi.
org/10.1093/nar/gnh069
Parks, M., Subramanian, S., Baroni, C., Salvatore, M. C., Zhang, G., Millar, C. D.,
& Lambert, D. M. (2015). Ancient population genomics and the study
of evolution. Philosophical Transactions of the Royal Society B: Biological
Sciences, 370, 20130381. https://doi.org/10.1098/rstb.2013.0381
Picher, Á. J., & Blanco, L. (2014). Patent No. International Publication
Nu mbe r W O2 01414 03 09A1. https://paten ts.google.com/paten t/
W O 2 0 1 4 1 4 0 3 0 9 A 1 / e n
Picher, Á. J., Budeus, B., Wafzig, O., Krüger, C., García- Gómez, S., Martínez-
Jiménez, M. I., Díaz- Talavera, A., Weber, D., Blanco, L., & Schneider, A.
(2016). TruePrime is a novel method for whole- genome amplification
from single cells based on TthPrimPol. Nature Communications, 7(1),
13296. https://doi.org/10.1038/ncomm s13296
Pilli, E., Modi, A., Serpico, C., Achilli, A., Lancioni, H., Lippi, B., Bertoldi,
F., Gelichi, S., Lari, M., & Caramelli, D. (2013). Monitoring DNA con-
tamination in handled vs. directly excavated ancient human skele-
tal remains. PLoS O ne, 8(1), e52524. https://doi.org/10.1371/journ
al.pone.0 052524
Pollard, H. G., Colbourne, J. K., & Keller, W. (2003). Reconstruction of
centuries- old Daphnia communities in a lake recovering from acid-
ification and metal contamination. Ambio, 32(3), 214– 218. https://
d o i . o r g / 1 0 . 1 5 7 9 / 0 0 4 4 - 7 4 4 7 - 3 2 . 3 . 2 1 4
Quinlan, A. R., & Hall, I. M. (2010). BEDTools: A flexible suite of utilities
for comparing genomic features. Bioinformatics, 26(6), 841842.
https://doi.org/10.1093/bioin forma tics/btq033
R CoreTeam. (2019). R: A language and environment for statistical computing.
R Foundation for Statistical Computing. http://www.R- proje ct.org
Rajpurohit, S., Gefen, E., Bergland, A. O., Petrov, D. A., Gibbs, A. G., &
Schmidt, P. S. (2018). Spatiotemporal dynamics and genome- wide
association genome- wide association analysis of desiccation toler-
ance in Drosophila melanogaster. Molecular Ecology, 27(17), 3525–
354 0. https://doi.org/10.1111/mec.14814
Reed, K. A., Lee, S. G., Lee, J. H., Park, H., & Covi, J. A. (2021). The ul-
trastructure of resurrection: Post- diapause development in an
Antarctic freshwater copepod. Journal of Structural Biology, 213(1),
107705. https://doi.org/10.1016/j.jsb.2021.107705
Rinke, C., Lee, J., Nath, N., Goudeau, D., Thompson, B., Poulton, N.,
Dmitrieff, E., Malmstrom, R., Stepanauskas, R., & Woyke, T. (2014).
Obtaining genomes from uncultivated environmental microorgan-
isms using FACS- based single- cell genomics. Nature Protocols, 9(5),
1038– 1048. https://doi.org/10.1038/nprot .2014.067
Rizzi, E., Lari, M., Gigli, E., De Bellis, G., & Caramelli, D. (2012). Ancient
DNA studies: New perspectives on old samples. Genetics Selection
Evolution, 44(1), 21. https://doi.org/10.1186/1297- 9686- 44- 21
Schlötterer, C., Tobler, R., Kofler, R., & Nolte, V. (2014). Sequencing pools
of individuals— Mining genome- wide polymorphism data without
big funding. Nature Reviews Genetics, 15(11) , 749– 763. ht tps://doi.
org/10.1038/nrg3803
Sella, G., & Barton, N. H. (2019). Thinking about the evolution of com-
plex traits in the era of genome- wide association studies. Annual
Review of Genomics and Human G enetics, 20, 461493. https://doi.
o r g / 1 0 . 1 1 4 6 / a n n u r e v - g e n o m - 0 8 3 1 1 5 - 0 2 2 3 1 6
Stiller, J., & Zhang, G. (2019). Comparative phylogenomics, a stepping
stone for bird biodiversity studies. Diversity, 11(7), 115. https://doi.
org /10.3390/d1107 0115
Telenius, H., Carter, N. P., Bebb, C. E., Nordenskjöld, M., Ponder, B.
A. J., & Tunnacliffe, A. (1992). Degenerate oligonucleotide-
primed PCR: General amplification of target DNA by a sin-
gle degenerate primer. Genomics, 13(3), 718– 725. https://doi.
o r g / 1 0 . 1 0 1 6 / 0 8 8 8 - 7 5 4 3 ( 9 2 ) 9 0 1 4 7 - K
von Baldass, F. (1941). Entwicklung von Daphnia pulex. Zoologische
Jahrbücher. Abteilung Für Anatomie Und Ontogenie Der Tiere, 67,
1– 6 0 .
Wang, J., Raskin, L., Samuels, D. C., Shyr, Y., & Guo, Y. (2015). Genome
measures used for quality control are dependent on gene func-
tion and ancestry. Bioinformatics, 31(3), 318– 323. https://doi.
org/10.1093/bioin forma tics/btu668
Weider, L. J., Lampert, W., Wessels, M., Colbourne, J. K., & Limburg, P.
(1997). Long- term genetic shifts in a microcrustacean egg bank as-
sociated with anthropogenic changes in the Lake Constance eco-
system. Proceedings of the Royal Society of London Series B- Biological
Sciences, 264(13 88), 16131618.
Wells, D., Sherlock, J. K., Handyside, A. H., & Delhanty, J. D. (1999).
Detailed chromosomal and molecular genetic analysis of single
cells by whole genome amplification and comparative genomic hy-
bridisation. Nucleic Acids Research, 27(4), 1214– 1218. https://doi.
org/10.1093/nar/27.4.1214
Wickham, H. (2016). ggplot2 Elegant Graphics for Data Analysis (Use R!).
S p r i n g e r . h t t p s : / / d o i . o r g / 1 0 . 1 0 0 7 / 9 7 8 - 0 - 3 8 7 - 9 8 1 4 1 - 3
Woyke, T., Sczyrba, A ., Lee, J., Rinke, C., Tighe, D., Clingenpeel,
S., Malmstrom, R., Stepanauskas, R., & Cheng, J.- F. (2011).
Decontamination of MDA reagents for single cell whole genome
amplification. PLoS One, 6(10), e26161. https://doi.org/10.1371/
journ al.pone.0026161
Xu, S., Ackerman, M. S., Long, H., Bright, L., Spitze, K., Ramsdell, J. S., &
Lynch, M. (2015). A male- specific genetic map of the microcrus-
tacean Daphnia pulex based on single- sperm whole- genome se-
quencing. Genetics, 201(1), 31– 38. https://doi.org/10.1534/genet
ics.115.179028
Xu, Y., & Zhao, F. (2018). Single- cell metagenomics: Challenges and ap-
plications. Protein & Cell, 9(5), 501– 510. https://doi.org/10.1007/
s 1 3 2 3 8 - 0 1 8 - 0 5 4 4 - 5
Zhang, F., Ding, Y., Zhu, C. D., Zhou, X., Orr, M. C., Scheu, S., & Luan,
Y. X. (2019). Phylogenomics from low- coverage whole- genome se-
quencing. Methods in Ecology and Evolution, 10, 507– 517. https://
doi.org/10.1111/20 41- 210X.13145
Zhang, L., Cui, X., Schmitt, K., Hubert, R., Navidi, W., & Arnheim, N.
(1992). Whole genome amplification from a single cell: Implications
for genetic analysis. Proceedings of the National Academy of Sciences
of the United States of America, 89(13), 5847– 5851. ht tps://doi.
org /10.1073/pnas.89.13.5847
Zheng, X., Gogarten, S. M., Lawrence, M., Stilp, A., Conomos, M. P.,
Weir, B. S., Laurie, C., & Levine, D. (2017). SeqArray- a storage-
efficient high- performance data format for WGS variant calls.
Bioinformatics, 33(15), 22512257. https://doi.org/10.1093/bioin
f o r m a t i c s / b t x 1 4 5
Zheng, X., Levine, D., Shen, J., Gogarten, S. M., Laurie, C., & Weir, B. S.
(2012). A high- performance computing toolset for relatedness and
principal component analysis of SNP data. Bioinformatics, 28(24),
3326– 3328. https://doi.org/10.1093/bioin forma tics/bts606
SUPPORTING INFORMATION
Additional supporting information may be found in the online ver-
sion of the article at the publisher’s website.
How to cite this article: O’Grady, C. J., Dhandapani, V.,
Colbourne, J. K., & Frisch, D. (2021). Refining the
evolutionary time machine: An assessment of whole genome
amplification using single historical Daphnia eggs. Molecular
Ecology Resources, 00, 1– 16. h t tps://d o i .
org /10.1111/1755- 0998.13524
ResearchGate has not been able to resolve any citations for this publication.
Article
Full-text available
The copepod, Boeckella poppei, is broadly distributed in Antarctic and subantarctic maritime lakes threatened by climate change and anthropogenic chemicals. Unfortunately, comparatively little is known about freshwater zooplankton in lakes influenced by the Southern Ocean. In order to predict the impact of climate change and chemicals on freshwater species like B. poppei, it is necessary to understand the nature of their most resilient life stages. Embryos of B. poppei survive up to two centuries in a resilient dormant state, but no published studies evaluate the encapsulating wall that protects theses embryos or their development after dormancy. This study fills that knowledge gap by using microscopy to examine development and the encapsulating wall in B. poppei embryos from Antarctica. The encapsulating wall of B. poppei is comprised of three layers that appear to be conserved among crustacean zooplankton, but emergence and hatching are uniquely delayed until the nauplius is fully formed in this species. Diapause embryos in Antarctic sediments appear to be in a partially syncytial mid-gastrula stage. The number of nuclei quadruples between the end of diapause and hatching. Approximately 75% of yolk platelets are completely consumed during the same time period. However, some yolk platelets are left completely intact at the time of hatching. Preservation of complete yolk platelets suggests an all-or-none biochemical process for activating yolk consumption that is inactivated during dormancy to preserve yolk for post-dormancy development. The implications of these and additional ultrastructural features are discussed in the context of anthropogenic influence and the natural environment.
Article
Full-text available
The results presented in this paper indicate that the ϕ29 DNA polymerase is the only enzyme required for efficient synthesis of full length ϕ29 DNA with the ϕ29 terminal protein, the initiation primer, as the only additional protein requirement. Analysis of ϕ 29 DNA polymerase activity in various in vitro DNA replication systems indicates that two main reasons are responsible for the efficiency of this minimal system: 1) the ϕ29 DNA polymerase is highly processive in the absence of any accessory protein; 2) the polymerase itself is able to produce strand displacement coupled to the polymerization process. Using primed M13 DNA as template, the ϕ29 DNA polymerase is able to synthesize DNA chains greater than 70 kilobase pairs. Furthermore, conditions that increase the stability of secondary structure in the template do not affect the processivity and strand displacement ability of the enzyme. Thus, the catalytic properties of the ϕ29 DNA polymerase are appropriate for a ϕ29 DNA replication mechanism involving two replication origins, strand displacement and continuous synthesis of both strands. The enzymology of ϕ 29 DNA replication would support a symmetrical model of DNA replication.
Article
Full-text available
Climate and environmental condition drive biodiversity at many levels of biological organization, from populations to ecosystems. Combined with paleoecological reconstructions, palaeogenetic information on resident populations provides novel insights into evolutionary trajectories and genetic diversity driven by environmental variability. While temporal observations of changing genetic structure are often made of sexual populations, little is known about how environmental change affects the long-term fate of asexual lineages. Here, we provide information on obligately asexual, triploid Daphnia populations from three Arctic lakes in West Greenland through the past 200-300 years to test the impact of environmental change on the temporal and spatial population genetic structure. The contrasting ecological state of the lakes, specifically regarding salinity and habitat structure may explain the observed lake-specific clonal composition over time. Palaeolimnological reconstructions show considerable regional environmental fluctuations since 1,700 (the end of the Little Ice Age), but the population genetic structure in two lakes was almost unchanged with at most two clones per time period. Their local populations were strongly dominated by a single clone that has persisted for 250-300 years. We discuss possible explanations for the apparent population genetic stability: (a) persistent clones are general-purpose genotypes that thrive under broad environmental conditions, (b) clonal lineages evolved subtle genotypic differences unresolved by microsatellite markers, or (c) epigenetic modifications allow for clonal adaptation to changing environmental conditions. Our results motivate research into the mechanisms of adaptation in these populations, as well as their evolutionary fate in the light of accelerating climate change in the polar regions.
Article
Full-text available
Genomic analysis of hundreds of individuals is increasingly becoming standard in evolutionary and ecological research. Individual‐based sequencing generates large amounts of valuable data from experimental and field studies, while using preserved samples is an invaluable resource for studying biodiversity in remote areas or across time. Yet, small‐bodied individuals or specimens from collections are often of limited use for genomic analyses due to a lack of suitable extraction and library preparation protocols for preserved or small amounts of tissues. Currently, high‐throughput sequencing in zooplankton is mostly restricted to clonal species, that can be maintained in live cultures to obtain sufficient amounts of tissue, or relies on a whole‐genome amplification step that comes with several biases and high costs. Here, we present a workflow for high‐throughput sequencing of single small individuals omitting the need for prior whole‐genome amplification or live cultures. We establish and demonstrate this method using 27 species of the genus Daphnia, aquatic keystone organisms, and validate it with small‐bodied ostracods. Our workflow is applicable to both live and preserved samples at low costs per sample. We first show that a silica‐column based DNA extraction method resulted in the highest DNA yields for non‐preserved samples while a precipitation‐based technique gave the highest yield for ethanol‐preserved samples and provided the longest DNA fragments. We then successfully performed short‐read whole genome sequencing from single Daphnia specimens and ostracods. Moreover, we assembled a draft reference genome from a single Daphnia individual (> 50× coverage) highlighting the value of the workflow for non‐model organisms.
Article
Full-text available
The rate and spectrum of spontaneous mutations are critical parameters in basic and applied biology because they dictate the pace and character of genetic variation introduced into populations which is a prerequisite for evolution. We use a mutation-accumulation (MA) approach to estimate mutation parameters from whole genome sequence data from multiple genotypes from multiple populations of Daphnia magna, an ecological and evolutionary model system. We report extremely high base substitution mutation rates (µ-n,bs = 8.96 x 10-9/bp/generation [95% CI: 6.66-11.97 x 10-9/bp/generation] in the nuclear genome and µ-m,bs= 8.7x10-7/bp/generation [95% CI: 4.40-15.12 x 10-7/bp/generation] in the mtDNA), the highest of any eukaryote examined using this approach. Levels of intraspecific variation based on the range of estimates from the 9 genotypes collected from three populations (Finland, Germany, and Israel) span 1 and 3 orders of magnitude, respectively, resulting in up to a ∼300-fold difference in rates among genomic partitions within the same lineage. In contrast, mutation spectra exhibit very consistent patterns across genotypes and populations, suggesting the mechanisms underlying the mutational process may be similar, even when the rates at which they occur differ. We discuss the implications of high levels of intraspecific variation in rates, the importance of estimating gene conversion rates using an MA approach, and the interacting factors influencing the evolution of mutation parameters. Our findings deepen our knowledge about mutation and provide both challenges to and support for current theories aimed at explaining the evolution of the mutation rate, as a trait, across taxa.
Article
Full-text available
DNA can be preserved in marine and freshwater sediments both in bulk sediment and in intact, viable resting stages. Here, we assess the potential for combined use of ancient, environmental, DNA and timeseries of resurrected long-term dormant organisms, to reconstruct trophic interactions and evolutionary adaptation to changing environments. These new methods, coupled with independent evidence of biotic and abiotic forcing factors, can provide a holistic view of past ecosystems beyond that offered by standard palaeoecology, help us assess implications of ecological and molecular change for contemporary ecosystem functioning and services, and improve our ability to predict adaptation to environmental stress. Ellegaard et al. discuss the potential for using ancient environmental DNA (eDNA), combined with resurrection ecology, to analyse trophic interactions and evolutionary adaptation to changing environments. Their Review suggests that these techniques will improve our ability to predict genetic and phenotypic adaptation to environmental stress.
Article
Full-text available
Over the last decade, an increasing number of studies have used lake sediment DNA to trace past landscape changes, agricultural activities or human presence. However, the processes responsible for lake sediment formation and sediment properties might affect DNA records via taphonomic and analytical processes. It is crucial to understand these processes to ensure reliable interpretations for “palaeo” studies. Here, we combined plant and mammal DNA metabarcoding analyses with sedimentological and geochemical analyses from three lake-catchment systems that are characterised by different erosion dynamics. The new insights derived from this approach elucidate and assess issues relating to DNA sources and transfer processes. The sources of eroded materials strongly affect the “catchment-DNA” concentration in the sediments. For instance, erosion of upper organic and organo-mineral soil horizons provides a higher amount of plant DNA in lake sediments than deep horizons, bare soils or glacial flours. Moreover, high erosion rates, along with a well-developed hydrographic network, are proposed as factors positively affecting the representation of the catchment flora. The development of open and agricultural landscapes, which favour the erosion, could thus bias the reconstructed landscape trajectory but help the record of these human activities. Regarding domestic animals, pastoral practices and animal behaviour might affect their DNA record because they control the type of source of DNA (“point” vs. “diffuse”).
Article
Full-text available
Understanding the genomic basis of local adaptation is crucial to determine the potential of long-lived woody species to withstand changes in their natural environment. In the past, efforts to dissect the genomic architecture in gymnosperms species have been limited due to the absence of reference genomes. Recently, the genomes of some commercially important conifers, such as loblolly pine, have become available, allowing whole-genome studies of these species. In this study, we test for associations between 87k SNPs, obtained from whole-genome re-sequencing of loblolly pine individuals, and 270 environmental variables and combinations of them. We determine the geographic location of significant loci and identify their genomic location using our newly constructed ultra-dense 26k SNP linkage map. We found that water availability is the main climatic variable shaping local adaptation of the species, and found 821 SNPs showing significant associations with climatic variables or combinations of them based on the consistent results of three different GEA methods. Our results suggest that adaptation to climate in the species might have occurred by many changes in the frequency of alleles with moderate to small effect sizes, and by the smaller contribution of large effect alleles in genes related to moisture deficit, temperature and precipitation. Genomic regions of low recombination and high population differentiation harbored SNPs associated with groups of environmental variables, suggesting climate adaptation might have evolved as a result of different selection pressures acting on groups of genes associated with an aspect of climate rather than on individual environmental variables.
Article
Full-text available
Birds are a group with immense availability of genomic resources, and hundreds of forthcoming genomes at the doorstep. We review recent developments in whole genome sequencing, phylogenomics, and comparative genomics of birds. Short read based genome assemblies are common, largely due to efforts of the Bird 10K genome project (B10K). Chromosome-level assemblies are expected to increase due to improved long-read sequencing. The available genomic data has enabled the reconstruction of the bird tree of life with increasing confidence and resolution, but challenges remain in the early splits of Neoaves due to their explosive diversification after the Cretaceous-Paleogene (K-Pg) event. Continued genomic sampling of the bird tree of life will not just better reflect their evolutionary history but also shine new light onto the organization of phylogenetic signal and conflict across the genome. The comparatively simple architecture of avian genomes makes them a powerful system to study the molecular foundation of bird specific traits. Birds are on the verge of becoming an extremely resourceful system to study biodiversity from the nucleotide up.
Article
Hatching resting stages of ecologically important organisms such as Daphnia from lake sediments, referred to as resurrection ecology, is a powerful approach to assess changes in alleles and traits over time. However, the utility of the approach is constrained by a few obstacles, including low and/or biased hatching among genotypes. Here, we eliminated such bottlenecks by investigating DNA sequences isolated directly (i.e. without hatching) from resting eggs found in the sediments of Lake Constance spanning pre-, peri-, and post-eutrophication. While we expected genome-wide changes, we specifically expected changes in alleles related to pathways involved in mitigating effects of cyanobacterial toxins. We used pairwise FST-analyses to identify transcripts that showed strongest divergence among the four different populations and a clustering analysis to identify correlations between allele frequency shifts and changes in abiotic and biotic lake parameters. In a cluster that correlated with the increased abundance of cyanobacteria in Lake Constance we find genes that have been reported earlier to be differentially expressed in response to the cyanobacterial toxin microcystin and to microcystin-free cyanobacteria. We further reveal the enrichment of gene ontology terms that have been shown to be involved in microcystin-related responses in other organisms but not yet in Daphnia and as such are candidate loci for adaptation of natural Daphnia populations to increased cyanobacterial abundances. In conclusion this approach of investigating DNA extracted from Daphnia resting stages allowed to determine frequency changes of loci in a natural population over time.