Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
Contents lists available at ScienceDirect
Journal of Experimental Marine Biology and Ecology
journal homepage: www.elsevier.com/locate/jembe
An effective procedure for DNA isolation and voucher recovery from
millimeter-scale copepods and new primers for the 18S rRNA and
cytb genes
Erin E. Easton ⁎, David Thistle
Department of Earth, Ocean and Atmospheric Science, Florida State University, Tallahassee, FL 32306-4320, USA
a r t i c l e
i n f o
Article history:
Received 10 February 2014
Received in revised form 29 May 2014
Accepted 30 June 2014
Available online xxxx
Keywords:
DNA extraction
Crustacea
Nuclear
Mitochondrial
Copepoda
Method
a b s t r a c t
Many investigators need to determine whether individuals belong to the same species. DNA-sequence data have
helped with this task, but current procedures of DNA isolation from millimeter-scale crustaceans, such as
harpacticoid copepods, leave little to no voucher material for morphological analysis, and many procedures
yield only enough DNA for a single amplification reaction. We therefore developed a DNA-isolation procedure
that yielded essentially intact exoskeletons and sufficient DNA for multiple polymerase chain reactions.
DNA-amplification success of our DNA-isolation procedure was relatively insensitive to (1) the length of preservation time from sample collection to DNA isolations and (2) the length of time the DNA was stored at −20 °C
after isolation. An additional benefit of our procedure is therefore that the DNA isolated is relatively stable.
Primers available for the nuclear 18S rRNA gene and the mitochondrial cytochrome oxidase b (cytb) gene are
known not to work for many harpacticoids. We therefore designed primers that would amplify and sequence
an ~750-base-pair fragment of the 18S rRNA gene and others that would amplify and sequence an ~450-basepair fragment of the cytb gene. Both primer sets worked for at least 12 harpacticoid families.
© 2014 Elsevier B.V. All rights reserved.
1. Introduction
Investigators studying ecology, biogeography, or biodiversity must
often decide whether individuals are conspecific. Traditionally, these
decisions were based on morphological characters, but unfortunately,
speciation often occurs without changes in taxonomically important
characters (see, e.g., Knowlton, 1993; Wake et al., 1983), causing
individuals of different species to be morphologically indistinguishable
(i.e., to be cryptic species). Problems also arise if morphological variability within a species is greater than differences ordinarily found between
closely related species. In such cases, an investigator using morphological procedures may exclude some individuals that belong to the species
in question. Further, the taxonomic importance of some morphological
characters may not yet be recognized (see, e.g., Knowlton, 1993; Norris,
2000), so investigators could assign individuals of more than one
species to a single species.
As analysis of DNA sequences has become more common,
investigators have been able to use them to recognize conspecific individuals (see, e.g., Hajibabaei et al., 2007; Knowlton, 2000; Vogler and
Abbreviations: 18S, nuclear small-subunit ribosomal RNA; bp, base pair; cytb,
mitochondrially encoded cytochrome b; FSU, Florida State University; PCR, polymerase
chain reaction; rRNA, ribosomal RNA.
⁎ Corresponding author.
E-mail address: [email protected] (E.E. Easton).
http://dx.doi.org/10.1016/j.jembe.2014.06.016
0022-0981/© 2014 Elsevier B.V. All rights reserved.
Monaghan, 2007). When both morphological and DNA procedures are
used on the same individuals and the results agree, an investigator can
have much greater confidence in the assignment of individuals to
species.
Recent work has demonstrated the need for a combined molecular
and morphological approach to the study of copepods (Bron et al.,
2011; Garlitska et al., 2012), which are millimeter-scale crustaceans
that are abundant and speciose in marine and freshwater environments
(Humes, 1994). For example, potential cryptic species have been
discovered (Rocha-Olivares et al., 2001; Schizas et al., 1999), and
overlooked morphological differences of taxonomic importance have
been documented (Easton et al., 2010; Rocha-Olivares et al., 2001;
Schizas et al., 1999, 2002; Staton et al., 2005). These issues hinder our
ability to understand the biodiversity and the species-specific ecological
roles of copepods (Bron et al., 2011; Garlitska et al., 2012) because
individuals cannot be assigned to species with confidence. With
combined approaches to assigning individuals to species, ecologists
can determine their species-level contributions to such issues as carbon
cycling and responses to environmental perturbations.
Although combined approaches have been used, current procedures
need improvement. First, many of the DNA-isolation procedures in
current use destroy the individual (see, e.g., Braga et al., 1999; Burton,
1998; Caudill and Bucklin, 2004; Edmands, 2001; Lindeque et al.,
1999; Papadopoulos et al., 2005; Street et al., 1998; Thum, 2004;
Vestheim et al., 2005) or leave very little of it (see, e.g., Bucklin et al.,
136
E.E. Easton, D. Thistle / Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
2003; Easton et al., 2010; Vestheim et al., 2005) for morphological examination. Because genetic data may reveal the need for additional
morphological analysis, as much as possible of each individual should
be retained after processing for DNA.
Second, many current DNA-isolation procedures allow only a single
DNA-amplification reaction (e.g., those of Bucklin et al., 2003; Burton,
1998; Caudill and Bucklin, 2004; Lindeque et al., 1999; Street et al.,
1998), so if in the course of a study, DNA amplification must be repeated
or additional target regions amplified, no DNA remains for these
reactions. DNA-isolation procedures for millimeter-scale individuals
should therefore yield sufficient DNA template for multiple polymerase
chain reactions (PCR; Saiki et al., 1988).
These first two problems can be solved with DNA-release procedures. Unfortunately, these methods are not generally known to yield
DNA stable enough to permit DNA to be isolated (or PCR amplified)
after more than a few weeks of storage (see, e.g., Giraffa et al., 2000;
Hajibabaei et al., 2005; Kim et al., 2012). Circumstances can arise
under which DNA isolation (or PCR amplification) can only be done
months after collection (or DNA isolation), for example, when new
primers must be developed.
Fourth, although universal primers are available for the genes (e.g.,
COI, cytb, 18S) commonly used for species-level biogeography studies,
they tend to be unsatisfactory for use on millimeter-scale copepods
(E.E.E., personal observation). In particular, they often fail because
complementary sequences do not exist in the target genes for many
species of copepods, especially those of the order Harpacticoida. Without primers that will amplify and sequence species belonging to many
families, some fundamental studies cannot be done.
The goals of our study were to develop a DNA-isolation procedure for
single, millimeter-scale copepods that (1) yields an essentially complete
voucher for morphological analysis, (2) yields sufficient DNA template
for multiple PCR amplifications, and (3) yields DNA that is stable for
more than one year. We also developed (4) amplification and sequencing primers for a target region in the nuclear 18S ribosomal RNA (rRNA)
gene and one in the mitochondrial cytochrome oxidase b (cytb) gene
that work for individuals from many harpacticoid families.
2. Material and methods
2.1. Sample collection and preparation
2.1.1. Cultured individuals
We used cultures of a shallow-water harpacticoid species,
Amphiascoides atopus Lotufo and Fleeger 1995 (Miraciidae), to develop
and test our procedure. We maintained them in 1-L flasks with 550 ml
of artificial seawater (salinity = 30) made with Instant Ocean® Aquarium Sea Salt (Spectrum Brands, Madison, WI) and deionized water. We
fed the cultures ~ 0.01 g of crushed TetraMin® Tropical Crisps (Tetra
Holding, Blacksburg, VA) every two to four weeks. (Cultures of this
species are also maintained by Adelaide Rhodes, Harte Institute for
Gulf of Mexico Studies, Corpus Christi, TX.) Living individuals were
placed in 100% ethanol at −20 °C for at least 24 h before DNA isolation.
2.1.2. Deep-sea individuals
We tested our procedure on benthic deep-sea copepods, mostly
members of the Harpacticoida. We collected sediment samples with a
version of the Barnett et al. (1984) multiple corer (Ocean Instruments
MC 800 Multi Core, San Diego, CA) from stations on the continental
rise off the west coast of the United States of America (Table 1). The
overlying water and the top centimeter of sediment from a given core
were collected, combined, preserved with cold 95% ethanol, and stored
at −20 °C.
In the laboratory, we used sieves to separate the 300-μm fraction
from the 30-μm fraction for each sample. Organisms in the 300-μm fraction were stained overnight in a solution of 200 ml of 100% ethanol and
0.25 g of rose bengal. For the 30-μm fraction, we used Ludox® HS-40
Table 1
Average depth, latitude, and longitude of stations from which we collected copepods.
Station
Depth
(m)
Latitude
(decimal degree)
Longitude
(decimal degree)
1
2
3
4
5
6
7
8
3247
3593
3673
2733
3682
2717
3854
2699
44.00
42.56
39.99
40.00
36.80
36.68
32.87
32.80
−130.39
−131.92
−125.88
−125.45
−123.70
−122.82
−120.62
−120.37
(E.I. du Pont de Nemours, Wilmington, DE) to separate most organisms
from most of the sediment (see Burgess, 2001). Organisms in the 30-μm
fraction were stained overnight in a solution of 200 ml of 100% ethanol
and 0.3 g of Congo red. We used a stainless-steel loop to remove copepods from both size fractions under a dissecting microscope. Copepods
were stored in 100% ethanol at −20 °C.
2.2. DNA isolation
2.2.1. InstaGene™ Matrix
To isolate DNA, we used the DNA-releasing solution InstaGene™ Matrix (Bio-Rad Laboratories, Hercules, CA). It contained a Chelex®-based
ionic resin thought to bind to PCR-inhibiting and DNA-degrading compounds (see Walsh et al., 1991, and references therein).
2.2.2. DNA isolation from individuals
2.2.2.1. Pre-DNA-isolation imaging. In studies of single individuals, we
used a stainless-steel loop to transfer each individual from ethanol to
100 μl of nuclease-free water (Integrated DNA Technologies, Coralville,
IA) in a well of a custom-made, acrylic, depression-well plate (~0.5-ml
volume wells with a diameter of ~ 15 mm). After the individual
rehydrated for at least 10 min, we transferred it to a drop of glycerin
on a single-concavity slide (Fisher Scientific, Pittsburgh, PA), placed
the individual under a #1.5 cover slip, and made lateral and dorsal
images of it (at 5 × to 20 × magnification, as appropriate) with a
Moticam 2500 camera (Motic, Richmond, British Columbia, Canada)
mounted on an Axioskop compound microscope (Carl Zeiss Microscopy,
Oberkochen, Germany). After imaging, we returned the individual to
the original well for at least 10 min to remove glycerin and ethanol
and then transferred it to a new well that contained 100 μl of
nuclease-free water.
2.2.2.2. DNA isolation. After ~10 min, we isolated DNA with InstaGeneTM
Matrix following a modified version of the manufacturer's protocol. The
InstaGene™ Matrix was mixed at room temperature for a minimum of
60 s at a moderate speed (set at 4.5 on a 6-point scale) on a Thermix®
Stirring Hot Plate 310 T (Fisher Scientific). We transferred 100 μl of
the material to a sterile, 0.5-ml, Axygen® MAXYMum Recovery™
microcentrifuge tube (Corning, Tewksbury, MA) using a 1000-μl
pipette. (Note that a large-bore pipet tip is required for maintenance
of the original concentration of 6% InstaGene™ Matrix.)
We transferred the ethanol-free (see Section 2.2.2.1.) individual to
this tube of InstaGene™ Matrix with a stainless-steel loop that had
been sterilized in 5% hydrogen peroxide. After we confirmed visually
that the individual was submerged, we incubated the tube in a water
bath at 56 °C for 30 min, vortexed it on a Maxi Mix II (Thermolyne,
Dubuque, IA) at maximum speed for 10 s, shook the solution to the bottom of the tube by hand, confirmed that the copepod was submerged,
and placed the tube back in the water bath overnight. We then vortexed
the tube for 10 s at maximum speed, shook the solution to the bottom of
the tube by hand, and confirmed that the copepod was submerged. The
E.E. Easton, D. Thistle / Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
tube was then placed in a heating block for 8 min at 100 °C and
subsequently vortexed for 10 s at maximum speed. We then shook
the solution to the bottom of the tube by hand. After confirming that
the copepod (now essentially an exoskeleton) was submerged, we
centrifuged the tube at 11,200 relative centrifugal force for 2 min to
pull the exoskeleton and Chelex®-based resin to the bottom of the tube.
2.2.2.3. Supernatant transfer and exoskeleton recovery. We immediately
transferred the supernatant to a sterile, 0.5-ml, Axygen® MAXYMum
Recovery™ microcentrifuge tube, using a small (200-μl) pipet to minimize the amount of Chelex®-based resin transferred to the new tube.
During this step, we monitored the exoskeleton to confirm that it
remained in the original tube and to ensure we did not crush it with
the pipet tip. Deionized water was then added to the original tube to
prevent the exoskeleton from drying out. If we lost track of the specimen or it became stuck to the inside of the pipet tip, we filled the
pipet with the deionized water that had been added to the original
tube and left the filled pipet tip in the tube so that the exoskeleton
would remain submerged. In all cases, we retained the pipet tip to transfer the exoskeleton to a 50-mm-diameter petri dish. To recover an
exoskeleton stuck to the inner wall of a pipet tip, we cut off and
discarded the filtered end of the tip and then washed the exoskeleton
from the remaining portion into the petri dish with deionized water.
With a stainless-steel loop, we recovered the exoskeleton under a
dissecting microscope and transferred it to glycerin on a depression
slide for long-term preservation. We then imaged it as above.
2.2.3. DNA isolation from bulk Amphiascoides atopus
To obtain sufficient DNA to use as positive controls and to test our
primers, we isolated DNA from bulk A. atopus as above but with these
differences. (1) We did not image the individuals. (2) We made four
bulk DNA isolations by adding 50 or 100 individuals to 100 μl of
InstaGeneTM Matrix and 50 or 100 individuals to 200 μl of InstaGeneTM
Matrix. (3) We did not attempt to confirm that all individuals remained
submerged during processing. (4) We did not transfer the supernatant
to a new tube.
2.3. PCR amplification
2.3.1. Standard procedures
After DNA isolation, we stored the supernatant at − 20 °C in the
microcentrifuge tube. Before each use of the supernatant, we thawed
it, vortexed it at maximum speed for 10 s to homogenize it, and centrifuged it for 2 min at 11,200 relative centrifugal force; this last step
helped minimize the amount of the Chelex®-based resin transferred
to the PCR mixture, where it could inhibit amplification (Montero-Pau
et al., 2008).
For PCR amplification, we used a Mastercyler® pro S thermocycler
(Eppendorf AG, Hamburg, Germany). We added 1, 2, or 10 μl of the
supernatant (or a dilution of the supernatant) to the following PCR mixture to yield a total volume of 50 μl: (1) 1:5 dilution of 5 × iProof™ HF
Buffer (20 mM Tris–HCl pH 7.4 at 25 °C, 0.1 mM EDTA, 1 mM DTT,
100 mM KCl, 0.5% Tween 20, 0.5% Nonidet P 40, 200 μg ml−1 BSA, 50%
glycerol; Bio-Rad Laboratories), (2) 200 μM of each deoxyribonucleotide triphosphate (USB Corporation, Cleveland, OH), (3) 1.0 μM of
each primer (see Table 2), and (4) 1 unit of iProof™ High-Fidelity DNA
polymerase (Bio-Rad Laboratories). See Sections 2.3.3 and 2.3.4 for
details and for differences between PCR amplification of the two genes.
PCR products and the 100 bp Plus DNA Ladder (Bioneer, Alameda,
CA) were subjected to electrophoresis at 100 V. We used a gel made
from 1.2% Certified™ Molecular Biology agarose (Bio-Rad Laboratories)
and 1× sodium-borate acid (Brody and Kern, 2004). Initially, we stained
the gels with ethidium bromide but switched to Gelstar® nucleic-acidgel stain (Lonza Rockland, Rockland, ME) because it is more sensitive
(allowing us to use 5 μl rather than 10 μl of PCR product). For the
ethidium-bromide-stained gels, we used a BioDoc-It® 220 (UVP,
137
Table 2
Primers designed for amplification and/or sequencing of copepods, reported in the
orientation they were most commonly used. Primer names begin with an abbreviation
for the gene name, followed by the approximate base-pair position in the given gene,
and end with an H for harpacticoid. For 18S primers, an r indicates that the primer amplifies in the reverse direction from primers ending in H. For consistency with the naming
convention of Burger et al. (2007), the amplification directions of cytb primers are not indicated in the primer name. We used the International Union of Pure and Applied Chemistry single-letter codes for nucleotides.
Gene
Primer
Sequence (5′ to 3′)
18S
18S
18S
18S
18S
18S
18S
cytb
cytb
cytb
cytb
cytb
cytb
cytb
cytb
18s150H
18s583Hr
18s648H
18s1075H
18s1343H
18s1538Hr
18s1871Hr
cb415H
cb424Ha
cb436H
cb867Hb
cb876Ha,b
cb897Hb
cb900Hb
cb922Hb
CTG CGG TAA TTC TGG AGC TAA TAC ATG C
GG CTG CTG GCA CCA GAC TTG CCC TCC
TCC GTT AAA AAG YTC GTA GTT KGA
CGA AGG CGM TCA GAT ACC GCC CTA G
CTC GAT TCR GTG GGT GGT GGT GCA TG
CAT CTA AGG GCA TCA CAG ACC
CAC CTA CGG AAA CCT TGT TAC GAC
GCY TTY TTA GGY TAT GTN YTN CCY TGR GG
GGY TAT GTN YTN CCY TGR GGD CAR AT
CCY TGR GGD CAR ATR TCH TTY TGR GG
AAR TAY CAY TCH GGY TGA ATR TG
GCR TAN GCR AAT ARR AAR TAY CAY TCH GG
GGD AYD GMH CGY AAA ATD GCR TAN GC
TTA TTW GGD AYD GMH CGY AAA ATD GCR TA
GCH AYN ACN CCY CCY AAY TTA TTW GG
a
b
Modified from Boore and Brown (2000).
Designed to amplify in the reverse direction from the other cytb primers in this table.
Upland, CA) to visualize and to make images. For the Gelstar®-stained
gels, we used a Dark Reader® 46B Transilluminator (Clare Chemical
Research, Dolores, CO) to visualize them and a Nikon Coolpix 7600
camera (Nikon, Melville, NY) to make images.
2.3.2. Choice of genetic markers
We chose to work with the 18S rRNA gene because (1) it has been
used in metagenomic studies of sediments (e.g., Bik et al., 2012; Tang
et al., 2012), (2) it has been used in previous studies of Harpacticoida
(e.g., by Burton and Lee, 1994; Burton et al., 2005; Easton et al., 2010),
and (3) its rate of nucleotide substitution is sufficiently low that we
could design primers that could be used for many Harpacticoida families. We chose the cytb gene because its rate of nucleotide substitution
is high enough to make it useful for species-level studies of copepods
(Easton et al., 2010; Schizas et al., 1999, 2002; Staton et al., 2005).
2.3.3. PCR amplification of the nuclear 18S rRNA gene
2.3.3.1. Design of primers. We found that primers typically used to amplify regions of the 18S rRNA gene (Blaxter et al., 1998; Spears et al., 1992;
Zarlenga et al., 1994) did not work for many species of deep-sea
copepods. We therefore designed primers (Table 2) from an alignment
of copepod 18S rDNA sequences obtained by Holly Bik (University of
California, Davis) from a metagenomic study of material from our
stations. We then confirmed the presence of our primer sequences in
all the complete harpacticoid 18S rDNA gene sequences available on
GenBank®.
2.3.3.2. Amplification of individual DNA templates. After DNA isolation, we
used the primer pair 18S1075H and 18S1871Hr (Table 2) to amplify an
~795-bp fragment of the 18S gene. In general, we used 1, 2, or 10 μl of
DNA isolate. DNA was denatured at 95 °C for 5 min. It was then subjected to 40 cycles of denaturing at 95 °C for 40 s, annealing at 68 °C for
1 min, and extension at 72 °C for 3 min. The amplification ended with
a final extension at 72 °C for 15 min. We stored PCR products at 4 °C
until they were visualized by electrophoresis and subsequently
prepared for sequencing.
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E.E. Easton, D. Thistle / Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
2.3.3.3. Amplification of bulk DNA templates. Bulk DNA templates were
amplified as above, with the following exceptions. (1) We added 1 or
2 μl of a 1:9 dilution of the supernatant (1 μl of the supernatant to 9 μl
of nuclease-free water) to the PCR reaction mix. (2) When testing the
different primers (Table 2) with bulk DNA template, we used annealing
temperature gradients of 50–65, 50–68, and 55–70 °C.
2.3.4. PCR amplification of the mitochondrial cytb gene
2.3.4.1. Design of primers. Primers typically used to amplify regions of the
cytb gene (Boore and Brown, 2000; Burger et al., 2007) did not work for
many deep-sea individuals in our study. We therefore designed primers
(Table 2) from an alignment of cytb sequences obtained from our studies of the harpacticoid mitochondrial genome (Easton et al., 2014). We
then searched the harpacticoid cytb gene sequences in GenBank to
confirm the presence of our primer sequences.
2.3.4.2. Amplification of individual DNA templates. After testing the
primers (see Section 2.3.4.3), we determined that a nested PCR
approach increased amplification success and minimized amplification
of nontarget products. For the first round of PCR, we used the primer
pair cb415h and cb922h (Table 2) to amplify an ~ 500-bp fragment.
We added 5 or 10 μl of the supernatant to the PCR-reaction mix
described above. DNA was initially denatured at 98 °C for 3 min, then
subjected to 12 cycles of denaturing at 98 °C for 35 s, annealing at
47 °C for 35 s, and extension at 72 °C for 1 min. These cycles were
followed by an additional 36 cycles of denaturing at 98 °C for 35 s,
annealing at 53 °C for 35 s, and extension at 72 °C for 1 min before
the amplification ended with a final extension at 72 °C for 7 min. We
stored PCR products at 4 °C until they were used for the second round
of PCR.
For the second round of PCR, we used the primer pair cb424h and
cb897h (Table 2) to amplify an ~475-bp fragment. We added 1 or 2 μl
of the first-round product to the PCR reaction mix described above.
The thermocycler parameters from the first round were used with the
following differences: 10 cycles of denaturing at 98 °C, annealing at
53 °C, and extension at 72 °C followed by 30 cycles of denaturing at
98 °C, annealing at 58 °C, and extension at 72 °C. We stored PCR
products at 4 °C until they were visualized by electrophoresis and
subsequently prepared for sequencing.
2.3.4.3. Amplification of bulk DNA templates. Bulk DNA templates were
amplified as above, except that we used (1) 1 μl of the supernatant in
the first round of the nested-PCR amplification and (2) different
thermocycler parameters when testing the different primers (Table 2).
For these tests, we used the thermocycler parameters from the second
round with the following differences: the first 10 cycles of annealing
were at a gradient of 47–53 °C and the latter 30 cycles at a gradient of
53–68 °C.
2.3.5. Amplification success as proxy for DNA-isolation success
We considered DNA isolation to be successful if we visualized a
target-sized band after gel electrophoresis of 18S rDNA PCR products.
We chose not to quantify the DNA because of the presence of dye that
had leached from the specimen into the supernatant, inconsistent
evaporation of the supernatant during extractions (E.E.E., personal
observation), and differences among individuals in the condition and
amount of tissue (e.g., individuals differed in size). We used the 18S
target region rather than the cytb as a proxy for DNA-isolation success
because the latter has a higher rate of nucleotide substitution, and
therefore sequences complementary to the primers are less likely to
be present in all the deep-sea-copepod taxa in our study.
2.4. PCR-product purification and sequence assembly
PCR products were sequenced at the Florida State University
Sequencing Facility (Tallahassee, FL) or at the High Throughput Genomics Center (Seattle, WA). Before sequencing, we purified PCR products
with the CONCERTTM Rapid Purification System (Life Technologies,
Carlsbad, CA) per the manufacturer's protocol. At the sequencing facility, purified PCR products were cycle sequenced with a PRISM® Big Dye
™ Terminator Ready Reaction Kit (Applied Biosystems, Foster City, CA).
Electrophoresis of reaction mixtures was done on an Applied
Biosystems PRISM® 3100 Genetic Analyzer. PCR products sequenced
at the High Throughput Genomics Center (http://www.htseq.org)
were purified at their facility with Exo-Sap™ before labeling with Applied Biosystems reagents and subsequent electrophoresis on their Applied Biosystems 3730xl DNA Analyzer. At both facilities, the
sequencing primers were 18S1343H and 18S1538Hr for the 18S rRNA
gene and cb436H and cb897H for the cytb gene (Table 2). (We did not
attempt to sequence PCR products if the target bands were absent or
barely visible in images of the gels because we knew from previous experience that sequencing would have failed.).
We used Geneious™ Pro v.5.6.6 (Biomatters, 2012) to assemble and
edit a consensus sequence from sequenced PCR products. De novo
assemblies were run with the default settings for highest sensitivity,
and these assemblies were edited manually, if necessary. The authenticity of the sequences was assessed by a BLAST search of the National
Center for Biotechnology Information (2013) database.
2.5. Morphological vouchers and dissections
Morphological identification of a copepod individual to species
required that the individual be dissected so that appendages could be
examined under a compound microscope. We used needles made
from 0.25-mm-diameter tungsten wire (see Tindall, 1960) to dissect individuals in the glycerin drop in which the exoskeleton had been stored
after DNA extraction. We mounted the antennules, antennae, maxillipeds, and pereiopods 1 to 5 routinely and other parts as needed on a microscope slide. Each part was placed in a separate drop of the following
version of Hoyer's mounting medium: 10.0 g of distilled water, 8.0 g of
gum Arabic (a.k.a. acacia), 30.0 g of chloral hydrate, 2.0 g of glycerin,
0.07 g of potassium iodine, and 0.10 g of iodine crystals. We placed an
8-mm-diameter, #1.5 cover slip on each drop. Twenty-four hours or
more later, the cover slip was sealed to the slide with clear fingernail
polish (which prevented evaporation), and the parts were imaged as
described in Section 2.2.2.1.
3. Results
3.1. Goal 1, recover voucher material
We did preliminary tests of our procedure with A. atopus and recovered the exoskeleton for each of the 17 individuals. Of the 915 individual
deep-sea copepods from which we isolated DNA, we recovered the exoskeletons of 891 (97.4%); 838 (91.6%) were suitable for imaging and
morphological analyses. After DNA isolation, little or no tissue remained
in most individuals. The exoskeleton, appendages, and most setae
remained intact (Fig. 1), although caudal setae were often lost during
processing.
3.2. Goal 2, procure sufficient DNA template for multiple PCR amplifications
We had positive PCR amplification of the 18S target region for the
DNA isolated from all 17 individual A. atopus. For DNA isolated from
the 915 individual deep-sea copepods, 18S amplification was successful
for 884 (96.6%) of them. After DNA isolation, ~ 80 μl of supernatant was
available for PCR amplifications, so we had sufficient template for at
least eight reactions. We were able to amplify DNA from 97.5% (n =
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139
0.470). In particular, only three of the 214 isolates that we amplified
after 12 mo of storage failed (Fig. 4). All three of these isolates had amplified for at least 12 mo after DNA isolation.
3.4. Goal 4, develop new primers for the 18S rRNA gene and the cytb gene
For the 18S rRNA gene, the primers we designed amplified 96.6%
of our deep-sea individuals and sequenced the complete target
region for 733 (82.9%) of the 884 amplified. For the cytb target region, we attempted PCR amplification with our primers for 400 of
the 884 individuals for which we successfully amplified the 18S target region. Of these individuals, we successfully amplified 385
(96.3%) and assembled consensus sequences for 342 (88.8% of the
385 amplified). Among the deep-sea individuals for which we
amplified both genes were members of 12 families of harpacticoid
copepods: Aegisthidae, Ameiridae, Argestidae, Canthocamptidae,
Cletodidae, Dactylopusiidae, Ectinosomatidae, Idyanthidae, Miraciidae,
Nannopodidae, Pseudotachidiidae, and Zosimeidae.
4. Discussion
4.1. Goal 1, recover voucher material
Fig. 1. Pre- (A) and post- (B) DNA-isolation images of a male deep-sea copepod (lateral
view), showing that the exoskeleton survived the procedure with little or no damage.
The specimen was stained with Congo red. Image (A) shows a specimen in good condition
with internal tissues present; (B) shows the exoskeleton in good condition with the testis
still visible as well as some sediment in the gut.
122) of the individuals stained with rose Bengal and 96.5% (n = 793) of
those stained with Congo red.
3.3. Goal 3, isolate stable DNA
We isolated DNA from deep-sea individuals that had been preserved
in 95% ethanol for between 24 and 52 mo. Grouping individuals by the
number of months they had been preserved (Fig. 2) revealed that
preservation time had essentially no effect on the success of PCR
amplification (R2 = 0.052, p = 0.346).
After DNA was isolated, it was stored at − 20 °C for up to 32 mo
(Fig. 3). The success of PCR amplification of this stored DNA was
essentially independent of storage time (Fig. 4, R2 = 0.031, p =
DNA-isolation procedures currently used for copepod studies (e.g.,
by Braga et al., 1999; Bucklin et al., 2003; Burton, 1998; Caudill and
Bucklin, 2004; Easton et al., 2010; Edmands, 2001; Lindeque et al.,
1999; Papadopoulos et al., 2005; Street et al., 1998; Thum, 2004;
Vestheim et al., 2005) recover little or no morphological voucher material. In contrast, our procedure allowed us to recover the exoskeleton
after DNA isolation in a condition suitable for morphological analysis
(Fig. 1) in most cases (91.5%), so morphological reanalysis was possible
if DNA-sequence data and morphological data disagreed. Few of these
current DNA-isolation procedures use Chelex®-based methods or
other DNA-release methods (e.g., Edmands, 2001; Montero-Pau et al.,
2008) known to isolate DNA without physically damaging specimens
(see Hajibabaei et al., 2005). We believe that other DNA-release
methods could be modified by addition of a specimen-recovery step,
but additional tests would be necessary to determine their relative successes for combined morphological and molecular studies of copepods.
Our isolation technique is relatively quick (b60 min for one individual and b80 min for a batch of 10), excluding imaging and the overnight
incubation, which can be omitted if time is limited. During preliminary
tests (data not shown), we successfully amplified all 22 A. atopus we
Fig. 2. Graph showing that PCR success did not decline with months preserved in ethanol before DNA isolation. The number above each bar indicates the number of specimens preserved
for that duration.
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E.E. Easton, D. Thistle / Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
Fig. 3. Gel images of the PCR product of the ~798-base-pair (bp) 18S target region, showing the stability of the DNA after isolation. The DNA was isolated on 18 November 2010.
The image on the left is of the results of the third amplification, which was done five
days after DNA isolation; we used 2 μl of template and loaded 10 μl of PCR product and
1 μl of the 100 bp Plus DNA Ladder (Bioneer, Alameda, CA) on the gel. The image on the
right is of the results of the twenty-third amplification, on 15 January 2013 (nearly
26 mo after isolation); we used 2 μl of template for PCR amplification and loaded 5 μl of
PCR product and 0.5 μl of the ladder on the gel. Both gels were visualized after electrophoresis at 100 V for 30 min, but the gel on the left was stained with ethidium bromide and the
gel on the right with Gelstar® nucleic-acid-gel stain. The right-pointing arrow indicates
the 1000-bp band, and the left-pointing arrow the 500-bp band of the 100-bp Plus DNA
Ladder.
Exoskeletons that were not recovered typically disappeared (1) during
transfer of the supernatant to the new tube or (2) during the recovery of
the specimen from the original tube. The losses occurred, at least in part, because leaching out of most of the stain made the exoskeletons difficult to
see with the naked eye. Holding the microcentrifuge tube against a white
background made them easier to see. We also retained the pipet tip used
to transfer the supernatant so that we could recover exoskeletons that
had stuck to them (see Section 2.2.2.3). Future investigators may wish
(1) to use magnifying glasses to improve visibility of the exoskeletons
throughout the procedure and (2) to experiment with stains, because our
impression was that rose Bengal resisted leaching better than Congo red.
Another source of loss or damage was drying caused by inadvertent
failure to keep exoskeletons submerged throughout the procedure (see
Section 2.2). Dried-out individuals became brittle, collapsed on themselves, and tended to stick to the pipet tip and microcentrifuge tube.
Once they were stuck, washing them into a petri dish with deionized
water often failed to free them. In these cases, we had to free them
with a stainless-steel loop, which often caused breakage and lost appendages. In general, collapsed and damaged exoskeletons were not
suitable for morphological analysis, a frustrating development because
we had often obtained good sequence data from these individuals.
4.2. Goal 2, procure sufficient DNA template for multiple PCR amplifications
tested without the overnight incubation and did not detect a difference
in PCR success from those incubated overnight. Our approach saved
time because we did the morphological analysis only on individuals
from which we had already obtained gene-sequence data. Further,
most exoskeletons were virtually free of tissue and stain (Fig. 1), so
characters were easier to see and dissections were easier than those
on the pre-DNA-isolation individuals.
We lost or damaged 8.5% of the exoskeletons. This value overestimates the loss expected in future studies because it includes the individuals we processed before modifications (discussed below) were made
to reduce loss and damage.
Although ~20 μl of the initial 100-μl volume was lost to evaporation
and during supernatant-transfer and specimen-recovery steps, enough
DNA isolate remained for at least 8 attempts at PCR amplification. In
our study, we did as many as 33 amplifications from a single DNA
isolate. Our ability to do multiple amplifications on DNA isolated from
individuals from at least 12 harpacticoid families shows that our
procedure is not taxon specific (see Section 3.4.).
Although generally little to no internal tissue (not quantified)
remained in our exoskeletons after DNA isolation, we observed some
in which substantial amounts of tissue remained. We were able to
amplify DNA isolated from many of these individuals and conclude
that not all the tissue must be digested for successful PCR amplification.
Fig. 4. Graph showing that PCR success did not decline with months (over 12) stored at −20 °C after DNA isolation. The number above each bar indicates the number of specimens stored
for that duration before the final PCR amplification.
E.E. Easton, D. Thistle / Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
We were unable to amplify the DNA isolated from 3.4% of 915
individuals. We do not know why they failed. One possibility is that
we isolated insufficient DNA. Given the small size (b1 mm) and therefore low biomass of an individual copepod, some failures due to low
DNA yield are to be expected. Second, some individuals were probably
dead when we collected them; if so, their DNA had begun to decay. A
third possibility is poor binding of primers due to poorly conserved
complementary sequences in some taxa (see Section 4.4 for evidence
from sequence data).
4.3. Goal 3, isolate stable DNA
Concerns have been raised about the ability of Chelex®-based
procedures, such as ours, to isolate stable DNA. First, Hajibabaei et al.
(2005) observed that such procedures have low or no PCRamplification success for individuals preserved for longer than 24 mo
before DNA isolation. In contrast, we could PCR-amplify DNA isolated
from individuals preserved for as long as 52 mo (Fig. 2; see also Söller
et al., 2000). Because our study differs from Hajibabaei et al.'s (2005)
study in many ways (e.g., taxon, preservation methods, DNA-isolation
procedure), we cannot be sure which steps of our procedure increased
our success relative to theirs.
A second criticism involves the storage time between DNA isolation
and PCR amplification. Some authors (e.g., Giraffa et al., 2000;
Hajibabaei et al., 2005; Kim et al., 2012) report that Chelex®-based procedures, including InstaGene™ Matrix, produce DNA that can be stored
at –20 °C for as little as one month, unless buffered (as in Söller et al.,
2000), before PCR-amplification success declines (Greenspoon et al.,
1998; Hajibabaei et al., 2005). With our samples, we observed no meaningful decrease in PCR amplification success with time at − 20 °C
(Figs. 3 and 4). Again, the reason for our success after long-term storage
is unclear. Future investigators may want to determine whether success
differs with taxon, Chelex®-based reagents, the amount of Chelex®resin remaining in the supernatant, the number of freeze–thaw cycles
(see Greenspoon et al., 1998), or the ratio of Chelex®-based resin to biomass, which should be maintained at a high enough ratio to bind to PCR
inhibitors (see Söller et al., 2000).
In sum, our procedure can isolate DNA from millimeter-scale
individuals preserved for up to 52 mo, and that isolated DNA was stable
for up to 32 mo of storage at − 20 °C. This stability has many
advantages, e.g., when logistics constrain the rate of sample processing
or when sequencing difficulties are encountered that delay processing.
4.4. Goal 4, develop new primers for the 18S rRNA gene and the cytb gene
Our new cytb primers amplified the target region more than 96% of
the time, a substantial improvement over the ~ 50% success found by
E.E.E. when she used the universal cytb primers during a pilot study (unpublished) that included species from the Canthocamptidae,
Harpacticidae, Miraciidae, and Nannopodidae. Subsequent work, including Easton et al. (2014), revealed that the primer-binding regions
differed at several loci, a problem that could have interfered with the
binding of the universal primers.
For those deep-sea individuals that had positive PCR results,
sequencing success was 82.9% for the 18S gene and 88.8% for the cytb
gene. The higher sequencing-success rate for the cytb target region
may have arisen (1) because our primers for this gene are degenerate
and therefore suitable for a broader range of species or (2) because we
only amplified this gene region for individuals for which we had obtained contaminant-free 18S-sequence data.
More than half of the failures arose because one sequencing primer
did not provide good sequence data, even after multiple attempts. These
failures, as well as failures of both primers, could be due to (1) insufficient
DNA concentration of the target region, (2) interference from nontargetlength products or other contaminants, or (3) poor sequencing-primer
binding. In future research, sequencing primers such as those in Table 2
141
or species-specific primers may solve this problem. We successfully amplified individuals from at least 12 families, so our primers are not limited
to a small number of closely related species.
4.5. Imaging
Although not strictly part of our procedure of obtaining gene sequences, our routine of imaging each individual before and after DNA
isolation has advantages. The images, which take a few minutes to
make, provide a reasonable approximation to the usual habitus drawings, which take hours. Also, a person with very little training can
make the images, whereas only an expert can make useful habitus
drawings.
We suspect that other types of imaging could be done after the
individual has passed through the DNA-isolation procedure and before
it is dissected. For example, Terue C. Kihara (personal communication)
recovered copepod exoskeletons after DNA isolation, stained them in
Congo red, and produced 3D images (Michels, 2007; Michels and
Buntzow, 2010). Further, if the exoskeleton will not be dissected, scanning electron microscopy might be possible.
4.6. Potential future applications of this procedure
Our procedure may be useful in other types of studies where the investigator needs both the identity of the individual and enough DNA for
multiple amplifications. For example, ecological studies of copepod
health, as indicated by the presence of epibionts (e.g., alveolates, bacteria, fungi) and studies of copepod decomposition and gut contents as indicated by associated biota (e.g., bacteria, diatoms) could be done on
single individuals. Our procedure isolates sufficient DNA to amplify
and sequence a region of the 18S rRNA gene for marine alveolates associated with our individuals (E.E.E., personal observation). In addition,
similar Chelex®-based procedures, including InstaGene™ Matrix, are
sensitive enough to isolate DNA from algae (Richlen and Barber, 2005;
Simonelli et al., 2009), bacteria (Drake et al., 1996; Giraffa et al., 2000;
Kim et al., 2012), trematodes (Enk et al., 2010), and the gut contents
of copepods (Simonelli et al., 2009). Because our procedure and other
Chelex®-based procedures have successfully amplified and sequenced
target regions for the associated fauna, investigators should be able to
use our procedure to isolate DNA for such species-level ecological studies. For example, DNA isolated from individuals could be used to amplify
copepod DNA as well as bacterial DNA for determination of specieslevel differences in diet and in symbionts.
Currently, copepods sequenced from environmental DNA studies
often cannot be identified because of the lack of available data
(F. Sinniger, personal communication) and are therefore only identified
to phylum (see, e.g., Bik et al., 2012), so evaluating biodiversity and ecology of copepods in these studies is difficult. Our approach could expand
understanding of copepod ecology if paired with such studies. For
example, next-generation sequencing of DNA isolated by our methods
would produce sequences for thousands of individuals that would
provide a database to which environmental DNA sequences could be
compared. The morphological component of our study could then be
focused on those individuals that are most abundant.
5. Summary
We developed a procedure that isolates DNA from individual,
millimeter-scale copepods. Our procedure has several advantages for
future copepod studies. (A) DNA can be isolated from whole individuals
rather than just pieces, so more DNA can be isolated. (B) Individuals can
be recovered after DNA isolation in condition suitable for morphological
analysis and imaging. Recovery of individuals allows them to be
retained as voucher specimens and to be reanalyzed for new specieslevel characters when DNA-sequence data show that individuals previously thought to be conspecific belong to different species. Because
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E.E. Easton, D. Thistle / Journal of Experimental Marine Biology and Ecology 460 (2014) 135–143
specimens are recovered intact, individuals can be identified after rather
than before DNA isolation, so no time is wasted on morphological analysis of specimens for which DNA-sequence data could not be obtained.
(C) Sufficient DNA is isolated from a single individual to allow multiple
PCR amplifications and therefore to allow multiple attempts at successful sequencing and the amplification of several target genes. (D) Stable
DNA can be isolated from specimens preserved in ethanol at −20 °C for
up to 52 mo, so archived specimens can be used when fresh specimens
cannot be collected. (E) The isolated DNA is stable and is therefore suitable for PCR amplification after at least 32 mo at − 20 °C, so among
other things, individuals are less likely to become impossible to analyze
if processing is delayed for some reason. Finally, (F) we designed new
primers for the 18S rRNA and the cytb genes, which work for at least
12 families of harpacticoids.
Acknowledgements
D. Oliff at the FSU Oceanography Instrument Shop made the
depression-well plates. M. Schram made the stainless-steel loops.
S. Miller at the FSU Sequencing Facility performed sequencing reactions.
J. W. Fleeger (then at Louisiana State University) gave us the cultures.
The manuscript was improved by comments from L. E. Gillies,
M. Huettel, T. (Spears) Terebelski, A. B. Thistle, and two anonymous
reviewers. Assistance in the laboratory or at sea was provided by
C. Armstrong, S. Bode, S. Bourgoin, M. Bublitz, E. Carroll, R. Carvalho,
K. Christiano, R. Coker, V. Cruz, E. Darrow, S. Dorado, K. Easton, J. Fields,
L. E. Gillies, M. Rohal, L. Rose, R. Rowland, A. S. McInnes, C. Sackmann,
F. Stephenson, M. Volkenandt, J. Voutour, G. D. F. Wilson, and the crew
of the R.V. Point Sur. This material is based on work supported by the
National Science Foundation under Grant No. 0727243 to DT. We are
grateful for this support. [RH]
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