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Aquatic Invasions (2017) Volume 12, Issue 2: 177–189
DOI:
https://doi.org/10.3391/ai.2017.12.2.06
© 2017 The Author(s). Journal compilation © 2017 REABIC
Open Access
177
Research Article
Detection of a new non-native freshwater species by DNA metabarcoding
of environmental samples – first record of Gammarus fossarum in the UK
Rosetta C. Blackman
1,
*
, Drew Constable
2
, Christoph Hahn
1,3
, Andrew M. Sheard
4
, Jessica Durkota
5
,
Bernd Hänfling
1
and Lori Lawson Handley
1
1
Evolutionary and Environmental Genomics Group, School of Environmental Sciences, University of Hull, Cottingham Road,
Hull, HU6 7RX, UK
2
Environment Agency, Bromholme Lane, Brampton, Huntingdon, Cambs, PE28 4NE, UK
3
Institute of Zoology, University of Graz, Universitaetsplatz 2, A-8010 Graz, Austria
4
Independent author, Beverley, UK
5
Department of Geography, University College London, Gower Street, London, WC1E 6BT, UK
Author e-mails: rosiecblackman@gmail.com (RCB), drew.constable@environment-agency.gov.uk (DC), C.Hahn@hull.ac.uk (CH),
andysheard@me.com (AMS), j.durkota@ucl.ac.uk (JD), B.Haenfling@hull.ac.uk (BH), L.Lawson-Handley@hull.ac.uk (LLH)
*
Corresponding author
Received: 23 September 2016 / Accepted: 20 March 2017 / Published online: 24 April 2017
Handling editor: Michal Grabowski
Abstract
We report the discovery of a non-native gammarid, Gammarus fossarum (Koch, 1836) (Crustacea, Amphipoda), in UK rivers.
Gammarus fossarum is a common freshwater gammarid in many parts of mainland Europe, but was previously considered absent
from the UK. Gammarus fossarum was detected in a number of UK rivers following DNA metabarcoding of a mini-barcode
region of the COI gene in macroinvertebrate kick samples, and environmental DNA (eDNA) from water and sediment samples.
Subsequent morphological analysis and standard DNA barcoding showed that the species could be reliably identified and
separated from Gammarus pulex (Linnaeus, 1758), the most dominant and widespread native freshwater gammarid in the UK. Our
data demonstrate extensive geographical coverage of G. fossarum in the UK, spanning distant river catchments. At present there is
no data to confirm the likely origin of G. fossarum’s introduction. Subsequent re-examination of historic archive material shows
the species to have been present in the UK since at least 1964. This study is among the first to demonstrate the potential of eDNA
metabarcoding for detection of new non-native species.
Key words: environmental DNA, metabarcoding, passive detection, early warning, cryptic species, Gammaridae, non-native
Introduction
Amphipods are successful invaders in freshwater
ecosystems, with many invasive non-native species
(INNS) having been observed to adversely impact
indigenous species within Europe over the last century
(Bij de Vaate et al. 2002; Grabowski et al. 2007).
The introduction of non-native amphipods may not
only lead to displacement of native congeners (e.g.
Dick and Platvoet 2000; MacNeil and Platvoet 2005;
Kinzler et al. 2009), but may also impact on ecosystem
structure and functioning (MacNeil et al. 2011; Piscart
et al. 2011; Constable and Birkby 2016) and introduce
novel pathogens to newly colonised areas (Bacela-
Spychalska et al. 2012).
Once non-native species are widely established,
efforts to reduce their impacts are often problematic,
hence management strategies are strongly focused
on preventing introductions or spread (e.g. the “check,
clean, dry” campaign in the UK). Early detection is
key to such strategies, either to improve the success
of eradication programs or to prevent further estab-
lishment and dispersal (Roy et al. 2014; Dejean et al.
2012). For freshwater macroinvertebrates, INNS detection
R.C. Blackman et al.
178
methods typically rely on sampling programmes and
morphological identification. However, the standard
UK monitoring method for macroinvertebrates, a
three minute kick sample, will typically recover 62%
of families and 50% of species at a site (Furse et al.
1981). This can present considerable challenges when
dealing with rare or elusive species. Morphological
identification can also prove difficult when identifying
taxonomically similar or cryptic species, or juvenile
life stages, and is highly dependent on the taxonomical
expertise of the investigator. Emerging molecular
detection methods may provide significant benefit
for detecting non-native species in aquatic environments
(Darling and Mahon 2011; Lawson Handley 2015).
One new and rapidly developing method is the
use of environmental DNA (eDNA) (Taberlet et al.
2012a, b; Rees et al. 2014; Lawson Handley 2015),
which refers to cellular or extracellular DNA that
can be extracted directly from environmental samples
without prior separation of taxa (Taberlet et al.
2012a). Environmental DNA has been successfully
used in numerous studies to detect specific taxa
using a targeted approach based on standard or
quantitative PCR (Dejean et al. 2012; Dougherty et
al. 2016). In an alternative approach, called “meta-
barcoding”, entire species assemblages are analysed
by PCR with broadly conserved primers, followed
by Next Generation Sequencing (NGS: see Lawson
Handley 2015; Hänfling et al. 2016; Port et al. 2016;
Valentini et al. 2016 for further detail). Environ-
mental DNA metabarcoding has been successfully
used in a small number of studies, for example, to
describe entire communities of vertebrates (e.g.
Lawson Handley 2015; Hänfling et al. 2016; Port et
al. 2016; Valentini et al. 2016) and invertebrates
(Deiner et al. 2016) from marine, lake and river
samples. Metabarcoding has excellent potential as an
early warning tool for detection of non-native species
from samples collected from invasion pathways or
natural/semi-natural habitats (Mahon and Jerde 2016;
Lawson Handley 2015). For example, the technique
was recently used as an early detection method for
screening ship ballast, and detected non-indigenous
zooplankton in Canadian ports (Brown et al. 2016).
Environmental DNA metabarcoding has also identified
non-native fish species present in samples from the
live bait trade (white perch, Morone americana
(Gmelin, 1789) Mahon et al. 2014) and in river
samples (northern snakehead, Channa argus (Cantor,
1842) Simmons et al. 2015). However the number of
applications of metabarcoding for detection of non-
native species has so far been limited.
In this paper we describe the detection of
Gammarus fossarum (Koch, 1836), a newly recognised
freshwater amphipod to the UK, using macroinverte-
brate community and eDNA metabarcoding. The
species was found in several UK rivers following a
preliminary non-targeted sampling programme for
macroinvertebrate communities based on metabarcoding
of a 313 bp mini-barcode region of the cytochrome c
oxidase subunit I (COI) gene, and was subsequently
confirmed using a combination of morphological
analysis and standard full-length COI DNA barcoding
(via Sanger sequencing). This study demonstrates
the power of eDNA metabarcoding for detection of
non-native species in natural habitats.
Methods
Metabarcoding surveys
Sampling
Field surveys were carried out in March 2015 within
8 UK river catchments (Figure 1, Maps A–H, excluding
E). At each site (n = 65) environmental variables
including water depth, width, substrate type and
surrounding habitat were recorded. Three sample
types were collected at each site: a three minute
macroinvertebrate kick sample (Murray-Bligh 1999)
for identification by microscopy analysis and high
molecular weight DNA extraction from pools of
individuals; and water and sediment samples were
collected for eDNA extraction. Two litres of water
was sampled from the surface by collecting 4 × 500 ml
from points across the river width using a sterile
bottle. Sediment samples were collected from points
across the river width using a trowel, and the
material was placed in a 42 fluid oz. sterile Whirl-pak®
bag (Cole-Palmer, Hanwell, London). All sampling
equipment was sterilized in 10% commercial bleach
solution for 10 minutes then rinsed with 10% MicroSol
detergent (Anachem, UK) and purified water between
samples. Sample bottles filled with ddH2O were
taken into the field and later filtered as sample blanks.
Macroinvertebrate community sample processing
All macroinvertebrates from each kick sample were
sorted and identified to the lowest taxonomic level
possible, before being stored in sterile 50 ml falcon
tubes filled with 100% ethanol. For DNA extraction,
samples were dried to remove the ethanol and the
entire macroinvertebrate community was lysed in a
Qiagen Tissue Lyser® with Digisol (50mM Tris, 20M
EDTA, 120 mM NaCl and 1% SDS) (3 × 30 sec).
Samples were then incubated overnight at 55 °C with
SDS and Proteinase K. DNA from a 200 μl subsample
of the lysed tissue was extracted using the DNeasy
Blood & Tissue Kit® (Qiagen, Hilden, Germany)
according to the manufacturer’s protocol.
Detection of a new non-native freshwater species by DNA metabarcoding
179
Figure 1. Distribution of Gammaridae species detected during this study. – Gammarus fossarum, – Gammarus pulex and – both
species present. A – River Hull, B – River Bain, C – River Cam, D – River Colne, E – Nailbourne, F – River Frome, G – Rivers Taff and Ely
and H – River Ribble. See supplementary information Table S1 for further site information (Pebesma et al. 2005; Wickham. 2009; Bivand et
al. 2013; Bivand et al. 2016; Gallic. 2016) Contains OS data © Crown copyright and database right (2016).
R.C. Blackman et al.
180
Environmental DNA sample processing
Water samples were filtered within 24 hours through
sterile 47 mm diameter 0.45 μm cellulose nitrate
membrane filters and pads (Whatman, GE Healthcare,
UK), using Nalgene filtration units attached to a
vacuum pump. Sediment samples were stored at
−20 °C within 12 hours of sampling. The sample was
defrosted, mixed and 200 ml of sediment placed in a
sterile measuring cylinder with 500 ml of molecular
grade water, then inverted 10 times and left to stand
for 30 s, the supernatant was then poured off into a
sterile container. This procedure was repeated twice.
Two hundred and fifty millilitres of the supernatant
was then prefiltered through sterile 20 μm filter paper
(Whatman, GE Healthcare, UK), and the filtrate
subsequently filtered through 0.45 μm cellulose
nitrate filters, as for the water samples. Filter papers
were stored in sterile petri dishes at −20 °C until
extraction. Filtration blanks (2 L purified water) were
run before the samples for each filtration run to test for
contamination at the filtration stage (n = 5). Filtration
equipment was sterilized in 10% commercial bleach
solution for 10 minutes then rinsed with 10% MicroSol
detergent and purified water after each filtration.
Environmental DNA from both water and sediment
samples was extracted using PowerWater® DNA
Isolation Kit (MoBio Laboratories, Inc. Carlsbad,
USA) following the manufacturer’s instructions.
PCR, library prep and sequencing
We chose to use COI for metabarcoding because this
region has the broadest taxonomic coverage for
macroinvertebrates in public sequence databases and
is the most widely used DNA barcode for taxonomic
discrimination in this group. A 313 bp fragment
(“mini-barcode”) was targeted using the primers
described in Leray et al. (2013). For library prepa-
ration we used a nested tagging protocol, modified from
the Illumina 16S two-step metabarcoding protocol
(Illumina 2011) as outlined in Kitson et al. (2015).
In the first step, PCRs were performed with
modified versions of the primers jgHCO2198 TAIA
CYTCIGGRTGICCRAARAAYCA and mICOIintF
GGWACWGGWTGAACWGTWTAYCCYCC (Leray
et al. 2013). In addition to the standard primer
sequence, primers included one of eight unique forward
or 12 unique reverse 8-nucleotide Molecular Identi-
fication Tags (MID), plus a bridge site, which acts as
a binding site for PCR 2 (see Kitson et al. 2015 for
full details). PCRs were carried out in 25 μl volumes
with MyFi High-Fidelity Taq (Bioline, UK) containing:
10 μM of each primer, and 2 μl of undiluted DNA
template. PCRs were performed on an Applied Bio-
systems Veriti Thermal Cycler with the following
profile: initial denaturation at 95 °C for 1 min,
followed by 45 cycles of denaturation at 98 °C for
15 s, annealing at 51 °C for 15 s and extension at 72 °C
for 30 s, with a final extension time of 10 min at 72 °C.
This included PCR and filtering blanks (n = 3 and
n = 5, respectively) and single species positives: Triops
cancriformis (Bosc, 1801) (n = 2) and Harmonia
axyridis (Pallas, 1773) (n = 2). PCR products were
confirmed by gel electrophoresis on a 2% agarose
gel stained with ethidium bromide. PCRs were
carried out three times and then pooled. Pooled PCR
products were then purified using the E.Z.N.A Cycle
Pure Kit® (VWR International, Leicestershire).
In the second PCR step, Illumina adapters and
additional forward and reverse MID tags were added
in a second PCR with 10 μM of each tagging primer
and 2 μl of purified PCR product. PCR settings were:
initial denaturation at 95 °C for 3 min, followed by
12 cycles of denaturation at 98 °C for 20 s, annealing
at 72 °C for 1 min and extension at 72 °C for 5 mins,
with a final extension time of 10 mins at 4 °C (Kitson
et al. 2015).
Samples were then classified into five categories
based on the strength of band produced on ethidium
bromide-stained agarose gels. Negative controls (inclu-
ding filtration blanks) produced no bands on the
agarose gel so were categorised with samples with
the lowest band strengths when being added to the
library. All positive control (i.e. extracted tissue)
samples were categorised as high band strength.
Volumes of the samples were then pooled according
5 band strength categories: 10 μl for the lowest band
strength, then decreasing volumes of 8 μl, 6 μl, 4 μl,
and 2 μl for increasing band strength. The library was
then pooled and cleaned using AMPure XP beads
following the recommended manufacturer’s protocol
(Agencourt AMPure XP, Beckman Coulter Inc. US).
The library was run at a 12 pM concentration on an
Illumina MiSeq, at the in-house facility at the Uni-
versity of Hull, using the 2 × 300 bp V3 chemistry.
Specimen confirmation – microscopy and standard
DNA barcode sequencing:
Verification of the results from DNA metabarcoding
was carried out using a combination of morphological
identification and standard DNA barcoding (by
Sanger sequencing).
Gammarus fossarum is a well-studied diverse species
complex, which has three well established cryptic species
(types A, B and C) with a further 36–53 different cryptic
lineages being identified through phylogenetic studies
(Weiss et al. 2014; Copilaş-Ciocianu and Petrusek
2015). Species within this complex are known to differ
Detection of a new non-native freshwater species by DNA metabarcoding
181
Figure 2. Picture of Gammarus fossarum
found in the River Taff, UK, 7/6/2016,
A) male adult specimen, B) male uropod III
and C) male plumose hairs on inside of exopod
of uropod III (↗); and picture of male
Gammarus pulex features for comparison
D) uropod III and E) plumose hairs on inner
and outer edge of exopod of uropod III (↗)
(Photographs by D. Constable).
in their ecology both in terms of their environmental
requirements and geographic distribu-tions (Copilaş-
Ciocianu and Petrusek 2015; Eisenring et al. 2016).
The G. fossarum complex belongs to the G. pulex-
group, which means it has small oval or kidney
shaped eyes (less than twice as long as wide) and the
pereopods 5–7 are armed with spines and few setae
(Pinkster 1972). Within the UK, these features alone
would help to separate it from G. duebeni, G. tigrinus
and G. zaddachi. It can be distinguished from all
R.C. Blackman et al.
182
five known UK freshwater Gammarus residents by
examining uropod III. In G. fossarum the ratio
length of the endopod versus the exopod is about
0.5, whilst in the other five it is >0.5, typically 0.75
(see Figure 2B and 2D respectively). Another
feature of G. fossarum is that only the inside margin
of the exopod has plumose setae, whilst the other
five have plumose setae on both inner and outer
margins (see Figure 2C and 2E respectively). The
latter feature should however be used with caution,
as plumose setae on the outer margin of the exopod
can show up in very old males of G. fossarum
(Meijering 1972).
A post hoc morphological examination of UK
Gammarus specimens was carried out to confirm the
presence of G. fossarum. Since the entire macro-
invertebrate samples from the original sampling
program had been lysed for metabarcoding, new
specimens were collected by hand net from two
catchments where G. fossarum was detected by
metabarcoding in close proximity to previously
sampled sites; River Taff, Wales (n = 38) on 7/6/2016
and River Frome, England (n = 39) on 27/6/2016.
Additional, archived specimens obtained from the
Nailbourne (Little Stour catchment), England (n = 2)
on 20/4/2013, were also analysed; (see Table 1 and
Figure 1, Maps: E, F and G). Collected individuals
were then subject to morphological examination and
identified using Karaman and Pinkster (1977), Eggers
and Martens (2001) and Piscart and Bollache (2012).
Microscopic identification was carried out on all
specimens collected for morphological confirmation.
Both G. fossarum (n = 37) and G. pulex (n = 1) were
identified from individuals collected from the River
Taff and only G. fossarum (n = 39) was found in a
sample from the River Frome. Standard DNA
barcoding was performed on some of the individuals
identified morphologically as G. fossarum (n = 3) and
G. pulex (n = 1) from the River Taff, and G. fossarum
from the Nailbourne (Little Stour catchment) (n = 2).
DNA was extracted using the DNeasy Blood &
Tissue Kit® (Qiagen, Hilden, Germany) according to
the manufacturer’s protocol. The full length COI
DNA barcoding fragment was amplified (Folmer et
al. 1994) using the following protocol: PCRs were
performed in 25 μl volumes with MyTaq (Bioline,
UK), 10 μM of each primer and 2 μl of DNA template.
The PCR profile consisted of: initial denaturation at
95 °C for 1 min, followed by 35 cycles of denaturation
at 95 °C for 15 s, annealing at 50 °C for 15 s and
extension at 72 °C for 10 s, with a final extension
time of 10 min at 72 °C. PCR products were checked
on agarose gels and commercially sequenced using
HCO2198 (Macrogen Europe, Amsterdam, Netherlands).
Bioinformatics
Processing of Illumina read data and taxonomic
assignment were performed using a custom bioin-
formatics pipeline (metaBEAT, v.0.97.7-global; see
Github reference 1) as described previously (Hänfling
et al. 2016), with minor modifications. For each
sample, raw Illumina sequences were filtered to
retain only read pairs containing the expected
forward/reverse in-line barcode combination (perfect
matches only) using the program process_shortreads
from the Stacks v1.20 program suite (Catchen et al.
2013) and subsequently quality trimmed using the
program Trimmomatic v0.32 (Bolger et al. 2014).
Specifically, read quality was assessed across 5 bp
sliding windows starting from the 3’-end, and reads
were clipped until the per window average read
quality reached a minimum of phred 30. Any reads
shorter than 100 bp after the quality clipping were
discarded. To remove PCR primers and spacer
sequences the first 30 bp of the reads was clipped off.
Remaining sequence pairs were merged into single
high quality reads using the program FLASH
v1.2.11 (Magoč and Salzberg 2011). For any read
pairs not merged successfully, only the forward read
was retained for downstream analyses. Sequences
were clustered at 97% identity using vsearch v1.1
(see Github reference 2). Any clusters represented
by less than three sequences were excluded from
further analyses, as these likely represent sequencing
error. Each of the remaining distinct sequence clusters
was collapsed to a single representative sequence (aka
centroid). Only centroid sequences of the expected
length as determined by the primers (313 bp ± 5%)
were retained for downstream analyses. To obtain a
final set of non-redundant (nr) queries for taxonomic
assignment, centroid sequences across all samples
were clustered globally at 97% identity using
vsearch v1.1. The global set of nr queries was subjected
to a BLAST (Zhang et al. 2000) search (blastn)
against a custom reference database consisting of
gammarid sequences from Weiss et al. (2014) and
two CO1 sequences from T. cancriformis (GenBank
accession numbers EF189678.1 and JX110644.1)
and H. axyridis (accession numbers KU188381.1 and
KU188380.1), respectively. Taxonomic assignment
was performed using a lowest common ancestor
(LCA) approach. In brief, after the BLAST search
the algorithm identifies the most significant matches
to the reference database (top 10% bit-scores) for
each of the query sequences. If only a single taxon is
present in this list of matches then the query is
assigned directly to this taxon. If more than one taxon
is present, the query is assigned to the lowest
taxonomic level that is shared by all taxa in the list.
Detection of a new non-native freshwater species by DNA metabarcoding
183
Queries yielding best BLAST matches below a bit-
score of 80 or with less than 85% identity were
binned as “unassigned”. To assure full reproducibility
of our analyses we have deposited the entire workflow
in an additional dedicated Github repository (see
Github reference 3). To reduce the possibility of
false positives based on our single species positive
samples and in order to obtain a conservative estimate
of the distribution of G. fossarum in the UK, we
only report G. fossarum as present at a given site if it
was supported by at least 1% of the total quality
trimmed reads per sample.
Phylogeny
Phylogenetic analysis was performed to further
confirm the identity of the putative Gammarus sp.
sequences obtained as part of the current study. We
downloaded a previously published CO1 dataset
(Weiss et al. 2014; Copilaş-Ciocianu and Petrusek
2015) from Genbank, comprising 89 sequences of G.
fossarum, six G. pulex (Linnaeus, 1758) sequences
and a single sequence each from four further outgroup
species (G. balcanicus (Schaferna, 1922), G. glabratus
(Hou and Li, 2003), G. roeselii (Gervais, 1835) and
G. tigrinus (Sexton, 1939) (Radulovici et al. 2009;
Hou et al. 2011; Feckler et al. 2012; Weiss et al.
2014). This set of previously published sequences
was extended by the sequences obtained via standard
full-length DNA barcoding and mini-barcode meta-
barcoding. Prior to phylogenetic analysis we extracted
the most abundant sequence, i.e. haplotype, from
each sample from the initially obtained 97% sequence
clusters assigned to G. fossarum and G. pulex,
respectively. Nucleotide sequences of G. fossarum
and G. pulex used in the phylogenetic analysis were
deposited in Genbank (GenBank accession KY464959–
KY464977). Phylogenetic analysis was performed in
the Reprophylo environment (Szitenberg et al. 2015).
In brief, sequences were aligned using the program
MAFFT v7.123b (Katoh and Standley 2013) and the
alignment was trimmed using the program trimAl
v1.2rev59 (Capella-Gutiérrez et al. 2009). Maximum-
likelihood tree inference was performed using RAxML
v8.0.12 (Stamatakis 2014). The full, detailed analysis
is provided as Jupyter notebook in the dedicated Github
repository (Github reference 3), which also contains
the alignment underlying the phylogenetic tree and
further supplementary information.
Comparison of data from eDNA/DNA and
microscopy analysis
A correlation was performed to compare the
Gammaridae abundance data generated from the kick
sample microscopy analysis and the DNA/eDNA
metabarcoding. Specifically, the relationship between
DNA/eDNA data (read count) and data from micro-
scopy analysis (biomass calculated from average
Gammaridae specimen weight) was investigated by
calculating Pearson’s Correlation Coefficient in R
v3.1.3 (R Core team 2013). Note that G. fossarum
and G. pulex sequencing data have been combined
here as the species were not distinguished during the
initial morphological determination.
Results
Metabarcoding survey
The total sequence read count passing quality
control, before removal of chimeric sequences, was
4,290,271. We quantified the level of possible conta-
mination using sequence information from single
species positive samples, which enabled us to choose
a suitable threshold level (1% of total sample reads)
for filtering and removal of low level contamination.
This conservative threshold is comparable to recent,
similar studies (e.g. Hänfling et al. 2016; Port et al.
2016). After applying this threshold, over the 195
samples the total read count was 933,457.
Gammarus fossarum was detected in 28 sites in
total: 25 via metabarcoding, 1 site by morphological
identification, 1 site by standard DNA barcoding and
1 site by morphological identification and DNA bar-
coding (See Table 1 and Supplementary material
Table S1). Of the 25 metabarcoding samples, G.
fossarum was found in: 25 DNA macroinvertebrate
samples, 8 water eDNA samples and 9 sediment eDNA
samples. G. pulex was detected in 27 of the sites in
the metabarcoding DNA macroinvertebrate samples
only and a single site using Sanger sequencing.
A full breakdown of gammarid sequences per
sample and proportion of gammarid biomass per
sample are included in Supplementary material
Table S1. A further 36 freshwater macroinvertebrate
families were detected by metabarcoding: data from
these non-gammarid species form part of a wider
macroinvertebrate data set which is being analysed
separately and will be published elsewhere.
The average read count of the samples with
gammarid species present was 3512. At those sites
the proportion of G. fossarum reads per sample ranged
from 1.68 – 100% in the macroinvertebrate DNA,
1.67 – 55.35% in the water eDNA and 1.59 – 18.05%
in sediment eDNA samples (Table S1). Similarly, G.
pulex reads ranged from 1.65 – 97.41% in the DNA
macroinvertebrate samples. There was a significant
positive correlation between the percentage of
Gammarus biomass in the sample, and the percentage
R.C. Blackman et al.
184
Table 1. Specimen identification and identification method for morphologically identified and DNA barcoded specimens. (*Specimens
collected from the River Frome were subject to morphological identification only. **Specimens collected from Nailbourne were DNA
sequenced only due to damaged specimens).
Coordinates G. fossarum G. pulex
Unique ID Catchment Site Name Lat Long Micro-
scopy
DNA
sequen-
cing
Micro-
scopy
DNA
sequen-
cing
DC003 Taff Forest Farm Country Park 51.516 −3.242 ✓ ✓
DC004 Taff Forest Farm Country Park 51.516 −3.242 ✓ ✓
DC005 Taff Forest Farm Country Park 51.516 −3.242 ✓ ✓
DC006 Taff Forest Farm Country Park 51.516 −3.242 ✓ ✓
DC007-045 Frome East Stoke 50.681 −2.185 ✓*
JD001 Nailbourne Adj Saint Ethelburga well 51.126 1.087 ✓**
JD002 Nailbourne Adj Saint Ethelburga well 51.126 1.087 ✓**
of Gammarus sequence reads (Pearson’s r = 0.747,
df = 46, P = 1.098 × 10-9, Supplementary material
Figure S1). Importantly, Gammarus sequences were
detected when gammarids constituted as little as
2.6% of the total biomass (Table S1).
Verification of Gammarus fossarum by microscopy
Gammarus fossarum was not identified morpho-
logically in any samples surveyed in March 2015
prior to metabarcoding. Of the 38 gammarid specimens
recovered from the River Taff on 7/6/2016, 37 G.
fossarum morphological identifications were made.
Adult males ranged between 8–12 mm (n = 21) and
adult females 7–10 mm (n = 15). Four females were
ovigerous. The other gammarid specimen encountered
was a male G. pulex (13 mm). Of the 39 gammarid
specimens collected from the River Frome on
27/6/2016, all were identified as G. fossarum
morphologically. Adult males of this population
ranged from 8–11.5 mm (n = 24) and adult females
7–9 mm (n = 15). Again, four ovigerous females were
recorded. The relative abundance of size distribution
in the two sampled populations can be seen in the
Supplementary information (Figure S2). The two
individuals collected from the Nailbourne on
20/4/2013 were not verified using microscopy as the
specimens were too heavily damaged for morpho-
logical identification.
The size ranges encountered for G. fossarum fall
within the expected range for the species, with
Goedmakers (1972), Pinkster (1972), Karaman and
Pinkster (1977) and Piscart and Bollache (2012) repor-
ting that the largest males typically reach 14–15 mm.
Verification of Gammarus fossarum by DNA
barcoding
Morphological identifications were confirmed by
DNA sequencing for specimens collected from the
River Taff (n = 4): 3 specimens of G. fossarum and a
single G. pulex. The individuals collected from the
Nailbourne (n = 2) were also both identified as G.
fossarum using subsequent DNA barcoding (see
Table 1).
Phylogeny
The phylogeny (Figure 3) is congruent with the
findings of the morphological identification. The G.
cf. fossarum and G. cf. pulex sequences cluster with
their respective lineages (identified in Weiss et al.
2014; Copilaş-Ciocianu and Petrusek 2015). Gammarus
fossarum sequences obtained by both metabarcoding
and standard DNA barcoding show little divergence
and cluster together in the phylogeny, indicating
closely related sequences. The G. fossarum sequen-
ces obtained in the current study group with high
statistical support within Clade 11, as defined using
the distance based Automatic Barcode Gap Discovery
(ABGD) approach in Weiss et al. (2014). Sequences
further group in a subclade with samples from south-
western Germany, Southern Black Forest and
Eastern Sauerland in Germany, i.e. clade 14, as
delineated using the tree-based GMYC in Weiss et al.
(2014). Aligning the UK G. fossarum specimens within
Clade 11 confirms previous studies which show this
clade to be the most widely distributed across Europe
within the species complex (Copilaş-Ciocianu and
Petrusek 2015; Weiss and Leese 2016).
Discussion
Non-targeted detection by direct and environmental
DNA metabarcoding has the potential to revolutionise
early warning systems for non-native species, but
this utility of the new technology has so far been
demonstrated only a limited number of times
(Mahon et al. 2014; Brown et al. 2016). In this study,
Detection of a new non-native freshwater species by DNA metabarcoding
185
Figure 3. Maximum likelihood phylogenetic tree
for the COI gene – based on sequences obtained
from previously published and newly obtained
Gammaridae sequences. The mini-barcode
(metabarcoding) and standard COI barcode
sequences from this study are represented in blue
and red, respectively. (See supplementary material
Table S2, for accession numbers and origin of
individual sequences). GMYC – General Mixed
Yule Coalescent, ABGD – Automatic Barcode Gap
Discovery (Puillandre et al. 2011) indicate the
approaches used by Weiss et al (2014) to detect the
different clades in their study.
R.C. Blackman et al.
186
G. fossarum, a newly recognised non-native species
for the UK, was detected during the course of a
wider metabarcoding survey of macroinvertebrate
communities. The identification of G. fossarum was
subsequently confirmed by microscopy and standard
DNA barcoding. The sequences generated from this
study indicate that the UK populations of G.
fossarum sampled here fall within the previously
identified Clade 11, sensu Weiss et al. (2014), of this
highly diverse species complex (Figure 3). Importantly
this is the most widely distributed clade within the
G. fossarum complex (Weiss et al. 2014; Copilaş-
Ciocianu and Petrusek 2015; Weiss and Leese 2016).
Gammarus fossarum was found in seven distant
river catchments within the UK, indicating a wide-
spread distribution (Figure 1). Initial detection of G.
fossarum was made using non-targeted meta-
barcoding of macroinvertebrate DNA, water eDNA
and sediment eDNA samples. Of the sites where G.
fossarum was detected using this method (n = 25),
G. fossarum was detected in all 25 DNA macro-
invertebrate samples (100%), in 8 of water (32%)
and 9 sediment (36%) samples. The lower detection
of G. fossarum in eDNA samples compared to
macroinvertebrate samples is not surprising due to
the dilution of eDNA and effects of flow on DNA
availability in lotic systems.
At 23 of the 28 sites (including post hoc samples)
where G. fossarum was present it was the only
Gammaridae species detected. This suggests it is not
only widespread in the UK but could also be the
dominant gammarid in some locations, possibly even
having displaced the native G. pulex locally. With
the new species discovery, recent re-examination of
historical archived gammarid samples was undertaken
from available Environment Agency and Natural
History Museum (NHM), London, collections. Material
from the Environment Agency had overlooked
records of G. fossarum dating back to 2005 from the
River Len, Maidstone, Kent (51.2619ºN; 0.56451ºE)
whilst re-examination of material from the NHM
revealed the earliest record to date, 1964 from the
River Darent, Kent. This shows that G. fossarum has
remained undetected and overlooked by conventional
means for a substantial length of time.
Gammarus fossarum is indigenous and wide-
spread in mainland Europe, and typically inhabits
springs and upper reaches of mountainous streams,
with G. pulex being more dominant in lower river
sections (Nijssen 1963; Goedmakers 1972; Karaman
and Pinkster 1977; Chen et al. 2012). This distribution
pattern is linked to G. fossarum’s comparative
preference for shallower streams and higher current
velocities, and its reduced tolerance of low dissolved
oxygen conditions (Meijering 1971; Peeters and
Gardeniers 1998). It may also be found in middle
sections of rivers and is able to coexist with G. pulex
(Janetzky 1994; Piscart and Bollache 2012; Copilaş-
Ciocianu et al. 2014). In such areas of coexistence,
G. fossarum will often occupy faster flowing areas
where vegetation is sparse or absent, and G. pulex
will be found near marginal shore zones, with reduced
currents and rich vegetation growth (Karaman and
Pinkster 1977). The distributions of G. fossarum in
this study covered a range of habitats, mainly
lowland rivers (altitude <90 m) with the exception of
the Nailbourne spring, adjacent to Saint Ethelburga
Well and Maiden Newton on the Upper Frome, with
altitudes of 106 m and 109 m, respectively (see
Supplementary information). The river depths at G.
fossarum locations were shallow, seldom reaching
more than 20 cm. It is important that further explora-
tion of UK upland systems is undertaken as the sites
surveyed for this study were mostly lowland, and at
this stage are an indication of habitat suitability rather
than preference for G. fossarum in the UK. Of our
five study sites where G. fossarum and G. pulex co-
existed, all had a mean depth >20 cm and featured
both fast and slow currents as well as vegetative
marginal areas, however there appears to be no other
pattern in the distribution of sites where both species
were found to co-exist. Four of the five sites were
from the metabarcoding samples, the percentage
read count for both species varied substantially, hence
no species dominance can be inferred from this data
(see Supplementary material Table S1).
Gammarus fossarum is the third non-native
freshwater gammarid to be found in the UK within
the last six years, following the discoveries of
Dikerogammarus villosus in 2010 (MacNeil et al.
2010) and Dikerogammarus haemobaphes in 2012
(Aldridge 2013). The record is rather unforeseen and
the species has not been included on the UK’s non-
native species watch list with more focus being
placed on Ponto-Caspian species that have invaded
western Europe (Gallardo and Aldridge 2015). A
detailed risk assessment of the threat that G.
fossarum poses to native Gammaridae within the UK
does not currently exist; further research into how G.
pulex and G. fossarum co-exist within UK habitats
should be carried out to decide if this action is
warranted. However, the importance of this discovery
as a new non-native species to the UK should not be
overlooked as it has important implications for
future ecological assessments.
In conclusion, we detected a newly recognised
non-native species to UK fauna using non-targeted
DNA metabarcoding, and confirmed its presence
using microscopy and standard DNA barcoding. It is
well known that the effectiveness of INNS control or
Detection of a new non-native freshwater species by DNA metabarcoding
187
management relies heavily upon early detection
(Lodge et al. 2006; Vander Zanden et al. 2010). In
future, for other species, non-targeted monitoring of
high risk invasion pathways using eDNA may ensure
that early eradication or containment are possible
management options (Davis 2009; Hulme 2009; Jerde
et al. 2011; Thomsen et al. 2012; Lawson Handley
2015). It is important that future research should
now focus on establishing the true distribution,
ecology and potential implications of G. fossarum
within the UK, as well as exploring how the non-
targeted eDNA metabarcoding approach can be used
to detect non-native species.
Acknowledgements
This work was funded by the UK Environment Agency. We are
particularly grateful to Dr Kerry Walsh and Dr Alice Hiley for
initiating the metabarcoding study and for support throughout. We
gratefully acknowledge Amanda Arnold, Frances Attwood, Charlotte
Davey, Carole Fitzpatrick, Tim Jones, Mel Lacan, Ellen Moss and
Paul Nichols for providing invaluable help during field work, Julia
Day, Robert Donnelly, Helen Kimbell, James Kitson and Graham
Sellers for contributing to laboratory work and Miranda Lowe and
Ben Price from the Natural History Museum, London and Susan
Skipp from the Environment Agency for allowing access to archive
material. We would like to thank F. Leese and anonymous reviewers
for their constructive criticism on the initial submission which greatly
helped strengthen the manuscript. The views expressed in this paper
are our own, and not necessarily of the institutions we represent.
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Github reference 1: https://github.com/HullUni-bioinformatics/metaBEAT; v.0.97.7-global
Github reference 2: https://github.com/torognes/vsearch
Github reference 3: https://github.com/HullUni-bioinformatics/Blackman_et_al_Gfossarum_UK. DOI: https://doi.org/10.5281/zenodo.495075
Supplementary material
The following supplementary material is available for this article:
Table S1. Specimen identification, identification method and site information for metabarcoding samples.
Table S2. Information on specimens from own and published studies that were used in the phylogenetic tree.
This material is available as part of online article from:
http://www.aquaticinvasions.net/2017 /Supplements/AI_2017_Blackman_etal_SupplementaryTables.xls
Figure S1. Correlation of the % Gammarus biomass in the sample, and the percentage Gammarus sequence reads.
Figure S2. Frequency distribution of body length of male and female Gammarus fossarum individuals collected from the River
Taff and the River Frome.
This material is available as part of online article from:
http://www.aquaticinvasions.net/2017 /Supplements/AI_2017_Blackman_etal_SupplementaryFigures.pdf
