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ORIGINAL PAPER
The role of invasive tunicates as reservoirs of molluscan
pathogens
Katie E. Costello .Sharon A. Lynch .Rob McAllen .Ruth M. O’Riordan .
Sarah C. Culloty
Received: 4 March 2020 / Accepted: 15 October 2020
ÓSpringer Nature Switzerland AG 2020, corrected publication 2020
Abstract Ascidian tunicates frequently display
rapid expansion when introduced beyond their native
range and are considered successful invaders. This
invasive potential may be exacerbated by a warming
climate, allowing for the occupation of environmental
niches previously held by native species. Research
into tunicate invasion ecology is prevalent, but less is
known about their role in pathogen maintenance. This
study investigated the impact of invasive tunicates on
the maintenance of pathogens that affect commercial
bivalves, including the cultured species Ostrea edulis
(European flat oyster) and Crassostrea gigas (Pacific
oyster), and the fished species Cerastoderma edule
(Common cockle). Focal pathogens included ostreid
herpesvirus OsHV-1 lVar, Vibrio aestuarianus,Bon-
amia ostreae and Minchinia spp. The range of
pathogens in their molluscan hosts was determined
and the tunicates Botrylloides violaceus,Didemnum
vexillum and Styela clava were then screened for these
same pathogens, using both field samples from oyster
culture sites and marinas and a series of laboratory
cohabitation trials. Sample sites reflected areas close
to and further away from known pathogen sources.
PCR, Sanger sequencing and histology confirmed the
presence of B. ostreae and Minchinia mercenariae-
like in S. clava, and V. aestuarianus was confirmed by
qPCR in B. violaceus and D. vexillum. Furthermore,
histology confirmed Minchinia mercenariae-like
sporonts in S. clava suggesting that the tunicate can
facilitate replication of this species. S. clava also
maintained B. ostreae in tanks with no oysters present.
The results indicate that tunicates can act as reservoirs
of infection in areas where disease occurs and
potentially transport diseases to uninfected sites.
Keywords Invasive Tunicates Oysters
Aquaculture Pathogens
Introduction
The Subphylum Tunicata is a diverse group of
invertebrates belonging to the Phylum Chordata, and
members of this group are globally recognised as
significant contributors to biofouling communities,
particularly species within the Class Ascidiacea; the
sea squirts (Rosa et al. 2013; Comeau et al. 2015; Zhan
K. E. Costello S. A. Lynch R. McAllen
R. M. O’Riordan S. C. Culloty
School of Biological, Earth and Environmental Sciences,
University College Cork, Cork, Ireland
K. E. Costello (&)S. A. Lynch
R. M. O’Riordan S. C. Culloty
Aquaculture and Fisheries Development Centre,
Environmental Research Institute, University College
Cork, Cork, Ireland
e-mail: katiecostello@ucc.ie
S. C. Culloty
MaREI Centre, Environmental Research Institute,
University College Cork, Cork, Ireland
123
Biol Invasions
https://doi.org/10.1007/s10530-020-02392-5(0123456789().,-volV)(0123456789().,-volV)
et al. 2015). A number of tunicate species, both
colonial and solitary, have become high profile
invasive species worldwide due to their significant
impacts on the aquaculture sector, including costs
associated with their control and removal when
fouling occurs, and their ability to compete with
commercial shellfish for food resources (Hillock and
Costello 2013; Casso et al. 2018).
Botrylloides violaceus (orange sheath tunicate) is a
colonial tunicate native to the northwest Pacific but
now established in temperate regions around the globe
(Simkanin et al. 2013) and known for its invasive
tendencies. This species can become competitively
dominant over commercial species, thus impeding
settlement (Cordell et al. 2012) and fouling cultured
stock (Paetzold et al. 2012). Didemnum vexillum
(carpet sea squirt) is another high-profile colonial
tunicate that has steadily increased its global range
from Japan (thought to be its native range) to New
Zealand, North America and Europe. Similar to B.
violaceus,D. vexillum can impact commercial aqua-
culture practices, for example heavy growth results in
shellfish valve openings or siphons becoming
obstructed (Ferguson et al. 2017).
Styela clava (leathery sea squirt/club tunicate) is a
solitary ascidian native to the Northwest Pacific and its
high tolerance of fluctuating environmental conditions
has seen it become a successful invader, expanding its
range from the Pacific coasts of Asia and Russia to
extend throughout the northern and southern hemi-
spheres (Goldstien et al. 2010). This tunicate attaches
to hard substrates such as rocks and bivalve shells, and
when natural substrates are unavailable it acts as a
biofouling organism on artificial structures and aqua-
culture gear. Mature adults are hermaphroditic and
oviparous, and densities can reach 500–1000 individ-
uals/m
2
(Wong et al. 2011), as such the cleaning of
culture gear has been known to make shellfish
production more labour intensive (Bourque et al.
2007).
Interactions between invasive species, pathogens
and native communities are complex, as invaders that
have not lost their associated parasites along the
invasion pathway may transmit novel parasites to
native hosts (Dunn and Hatcher 2015) or become
infected by native pathogens in the invaded range.
Furthermore, invasive species may also inhibit or
promote the transmission of pathogens by native
species (Goedknegt et al. 2016).
While many studies have focused on tunicates as
model organisms for understanding invasion success,
less is known about their role in the maintenance of
parasites and pathogens and their subsequent interac-
tions with commercial bivalves. Franchi and Ballarin
(2017) noted that tunicate physiology results in natural
defences against parasites and pathogens, as the tunic
acts as a physical barrier and the vascularised oral
siphon and pharynx have circulating haemocytes that
trigger immune responses that can lead to inflamma-
tion and phagocytosis of foreign material. Despite
these defences, tunicates have been shown to be
susceptible to specific pathogens, for example Styela
clava has been established as a potential carrier of the
flagellated protozoan Azumiobodo hoyamushi—the
causative agent of Soft Tunic Syndrome (Kumagai
et al. 2014). This disease widely affects cultured
tunicates such as the edible ascidian Halocynthia
roretzi and has caused economic losses in the aqua-
culture sector in Korea and Japan.
Rosa et al. (2013) tested the ability of biofouling
ascidians including Styela clava and Botrylloides
violaceus to distribute potentially harmful algal cells
to new geographic ranges by exposing them to
cultured strains of algae capable of forming harmful
algal blooms. Algal cells were found to pass through
the ascidian digestive system and remain capable of
re-establishing bloom populations. Rueckert et al.
(2015) also detected a number of apicomplexan
parasites within the genus Lankesteria infecting
Pacific ascidians and Lynch et al. (2016) found four
parasite groups including ciliates and trematodes in
the European sea squirt Ascidiella aspersa.
Determining the ability of bacteria, protists and
viruses to maintain themselves in the environment can
inform how they contribute to disease cycling, partic-
ularly in commercially important bivalves. However,
this can prove difficult due to free-living life stages,
the potential ability to persist short-term outside a host
and the use of alternative hosts. Haplosporidia
belonging to the genus Minchinia have traditionally
been detected in a range of bivalves including clams,
cockles and mussels (Ramilo et al. 2018). Other
haplosporidian species, including Bonamia ostreae
and Haplosporidium nelsoni, are well studied within
their hosts but knowledge of their life-history, diver-
sity and mechanisms of transmission is limited (Ward
et al. 2019). B. ostreae is an intra-haemocytic parasite
of the European flat oyster Ostrea edulis that induces
123
K. E. Costello et al.
physiological disorder and mortality. All oyster life-
stages are susceptible, although individuals over two
years are more adversely affected, and the pathogen is
present throughout the year but infections appear to
increase from Autumn and peak in late winter/early
Spring. Laboratory experiments suggest that an inter-
mediate host is not required, and the pathogen is
capable of surviving in seawater for up to one week
(OIE 2019). It is also known that B. ostreae is capable
of utilising alternative hosts, for example in a labora-
tory setting the brittle star Ophiothrix fragilis caused
transmission of B. ostreae to naive O. edulis (Lynch
et al. 2007).
The bacterium Vibrio aestuarianus is associated
with mortality events in populations of the Pacific
cupped oyster Crassostrea gigas in Europe, with adult
oysters of marketable size primarily affected and
mortalities mainly occurring in Summer (Lupo et al.
2019). The ostreid herpesvirus OsHV-1 lVar is
another pathogen associated with C. gigas mortality
events, with larval, spat and juvenile life-stages the
most susceptible. Transmission is primarily direct
although the virus is also hypothesised to travel via
vector particles in the water column. Mortalities occur
more frequently in summer water temperatures
although temperature effects differ between geo-
graphic locations, for example Europe and Australia
(OIE 2019). The virus has also been detected in other
phyla, for example the shore crab Carcinus maenas
can transmit OsHV-1 lVar from crabs exposed to the
virus in the wild to naive crabs in a laboratory setting
(Bookelaar et al. 2018).
Comprehensive information on the entire host
range for pathogens remains difficult to achieve, and
this can have implications for monitoring pro-
grammes, as these tend to focus on known susceptible
hosts and may therefore miss other potential hosts.
Accordingly, applied research of potential alternative
host species could provide more robust conclusions
about the host range of pathogens and also potential
carriers, reservoirs and vectors (Carnegie et al. 2016).
Climate change is predicted to increase introduc-
tions of problematic species, and affect the range,
abundance and impacts of invasive species (Beaury
et al. 2019). This study selected tunicates as a model
taxon that is heavily represented by species with
invasive tendencies, which may expand their range
under shifting climate conditions. Three species of
tunicate that are considered invasive outside of their
native range, Botrylloides violaceus,Didemnum vex-
illum and Styela clava, were screened for parasites and
pathogens that are known to affect commercial bivalve
species. These pathogens included the herpesvirus
OsHV-1 lVar, a virus that can cause mass mortalities
in C. gigas (Prado-Alvarez et al. 2016), and Vibrio
aestuarianus, a bacterium also linked to C. gigas
mortality events (McCleary and Henshilwood 2015).
Screening was also conducted for the Phylum
Haplosporidia, in particular Bonamia ostreae, the
intra-haemocytic parasite that affects O. edulis and is
the causative agent of bonamiosis (Lynch et al. 2010,
Prado-Alvarez et al. 2015), and Minchinia spp. as the
haplosporidia Minchinia tapetis and Minchinia merce-
nariae-like have also been detected in common
cockles (Cerastoderma edule) in Cork Harbour and
are known to cause losses for the cockle sector
(Albuixech-Martı
´et al. 2020). Screening was also
conducted for a broader range of haplosporidian
species, as previous research conducted in similar
sample sites to this study has indicated the presence of
Haplosporidium nelsoni in C. gigas and O. edulis, and
a further Haplosporidium sp., most likely
Haplosporidium armoricanum,inO. edulis (Lynch
et al. 2013a).
The aim of this study was to focus on the potential
impact of invasive tunicates on the maintenance of
pathogens known to be problematic within the aqua-
culture and shellfishery sectors. This was investigated
by determining the current status of pathogens known
to cause significant mortalities in Ostrea edulis and
Crassostrea gigas sampled from aquaculture sites in
Ireland, and also by reviewing the pathogens currently
affecting the fished species Cerastoderma edule; then
establishing if these same pathogens are present in
invasive tunicates cohabiting with the oysters at the
aquaculture sites. The study also aimed to determine
whether pathogens can be detected in invasive tuni-
cates sampled from an area with no aquaculture but
potentially other sources of introduction such as heavy
recreational shipping traffic. The final aim was to
establish the potential role of tunicates as reservoirs of
infection using laboratory cohabitation trials.
123
The role of invasive tunicates as reservoirs of molluscan pathogens
Materials and methods
Study sites
Sampling for tunicates and oysters was conducted
under licence from Ireland’s National Parks and
Wildlife Service and the Marine Institute. Sampling
in 2018 was conducted in Summer (July) and late
Autumn/Winter (October-November) to ensure that
pathogen screening encompassed both cold winter and
elevated summer water temperatures. Sampling was
conducted in two locations within Cork Harbour and
sea temperatures were sourced from Copernicus
(Fig. 1and Table 1). The first sample site was an
oyster culture site in Rossmore, Co. Cork where both
the oysters Ostrea edulis (n = 60) and Crassostrea
gigas (n = 60) and tunicate Styela clava (n = 90) were
collected.
Sampling of the colonial tunicate Botrylloides
violaceus in 2018 was conducted in the second
location in Co. Cork, a marina in Crosshaven, to
ascertain whether growth in an area with heavy
recreational shipping would result in the presence of
pathogens due to incidental transport.
Summary of pathogen screening
The oysters, as natural hosts of the pathogens of
interest, were screened to determine presence/absence
of the pathogens for which they are the primary host
and prevalence if present, before screening the tuni-
cates to determine if proximity to the oysters resulted
in a positive detection. In cases where the number of
tissue samples is higher than the number of individuals
(Table 1) it is because multiple tissue types were
collected per individual (see Sample Processing—
2018 and 2019 Samples for a detailed description).
Ostrea edulis was screened using specific primers
for Bonamia ostreae and Haplosporidium nelsoni
(MSX). General haplosporidian primers were then
also used to screen O. edulis and ensure that any
further haplosporidian species (e.g. Haplosporidium
spp., Minchinia spp.) were detected if present. Cras-
sostrea gigas was screened for Vibrio aestuarianus
and OsHV-1 lVar, as these are both pathogenic to C.
gigas but not O. edulis.C. gigas were also screened
with general primers for haplosporidian spp., to ensure
that any pathogens within this phylum were detected.
Styela clava was then screened for B. ostreae, OsHV-1
lVar, general haplosporidian spp. and V. aestuari-
anus.Botrylloides violaceus was screened for V.
aestuarianus, OsHV-1 lVar and haplosporidian spp.
using general primers (Table 2) (see Molecular
Screening for a detailed description).
Laboratory trials: 2019
Samples for the subsequent laboratory cohabitation
trials in 2019 were obtained from both Co. Cork
(Rossmore & Rostellan), Dungarvan, Co. Waterford
and Carlingford, Co. Louth (Fig. 1and Table 3). All
sampling locations were oyster aquaculture sites and
tunicate species were collected from the trestles.
Trial 1: Cohabitation trial between Bonamia
ostreae infected Ostrea edulis and Styela clava
to determine if S. clava is a reservoir or carrier
of infection
150 Ostrea edulis were randomly sampled from
Rossmore, Cork Harbour and of these an initial
sample (n = 30) was screened to assess the prevalence
of Bonamia ostreae. The remaining oysters were
divided into three experimental 50 L tanks, each
Fig. 1 Map of sample sites for 2018 and 2019 samples
123
K. E. Costello et al.
Table 1 Summary of oyster and tunicate samples collected from Cork Harbour in 2018
Species Site Date collected Avg. monthly sea temperature (°C) Individuals n tissue samples
a
Cork Harbour
Crassostrea gigas Rossmore 11-07-2018 16.47 30 30
Crassostrea gigas Rossmore 2-11-2018 11.93 30 30
Ostrea edulis Rossmore 11-07-2018 16.47 30 30
Ostrea edulis Rossmore 23-11-2018 11.93 30 60
Styela clava (intertidal) Rossmore 4-07-2018 16.47 30 76
Styela clava (subtidal) Rossmore 11-07-2018 16.47 30 144
Styela clava (subtidal) Rossmore 2-11-2018 11.93 30 162
Crosshaven Marina
Botrylloides violaceus Crosshaven 16-07-2018 16.57 30
b
90
Botrylloides violaceus Crosshaven 25-10-2018 14.38 10
b
30
a
See ‘Sample Processing—2018 and 2019 Samples’ for detailed explanation of column ‘n tissue samples’
b
Botrylloides violaceus is a colonial organism, therefore ‘individuals’ refers to the different lobes of three large colonies that
extended along the underside of marina pontoons
Table 2 Summary of the pathogens screened in each oyster and tunicate species in 2018
Species Crassostrea gigas
(n = 60)
Ostrea edulis
(n = 60)
Botrylloides violaceus
(n = 40
a
)
Styela clava
(n = 90)
Bonamia ostreae X4X4
Haplosporidium
nelsoni
X4XX
Haplosporidian spp. 444 4
OsHV-1 lVar 4X44
Vibrio aestuarianus 4X44
a
Botrylloides violaceus is a colonial organism, therefore ‘individuals’ refers to the different lobes of three large colonies that extended
along the underside of marina pontoons
Table 3 Summary of samples for laboratory trials 2019
Species Site Date collected Avg. monthly sea temperature (°C) n individuals n tissue samples
a
Ostrea edulis Rossmore 5-04-2019 10.08 150 300
Styela clava (intertidal) Rostellan 18-06-2019 12.07 32 248
Crassostrea gigas Dungarvan 4-07-2019 15.37 150 150
Didemnum vexillum Carlingford 6-08-2019 14.72 28
b
140
a
See ‘Sample Processing—2018 and 2019 Samples’ for detailed explanation of column ‘n tissue samples’
b
Didemnum vexillum is a colonial organism with an amorphous shape, therefore ‘individuals’ refers to different (and often
heterogeneous) colonies that extended along the underside of oyster trestles
123
The role of invasive tunicates as reservoirs of molluscan pathogens
containing 40 oysters, and held in a constant temper-
ature room at 14 °C, a salinity of 33–35 and a 12:12 h
light/dark regime.
Ostrea edulis were held for 75 days to allow the
intensity and prevalence of infection with Bonamia
ostreae to increase. After 75 days, 32 Styela clava of
all sizes were randomly sampled from Rostellan, Cork
Harbour. Tunicates growing on algae rather than
directly on the oyster trestles were collected, and a
piece of the algal substrate was taken too, as this
ensured the tunicates were not damaged in the
collection process and could survive the transfer to
the tanks. An initial sample of the tunicates (n = 8)
was screened for B. ostreae and the remaining 24
placed in three 50 L tanks, with eight tunicates per
tank; this resulted in one control tank with eight S.
clava only, and two experimental tanks each contain-
ing the B. ostreae-infected oysters and tunicates (8 S.
clava and 35 O. edulis 92 tanks). The final control
tank contained 35 O. edulis only. The trial ran for 60
days.
As both Minchinia tapetis and Minchinia merce-
nariae-like have been recorded in Cerastoderma edule
from Cork Harbour (Albuixech-Martı
´et al. 2020), a
subsample of eight Styela clava was also screened for
both Minchinia tapetis and Minchinia mercenariae-
like at the end of the trial, using PCR with primers
specific to each species, histology as per Ford et al.
(2009) and Sanger sequencing. This subsample
encompassed two tunicates from the initial baseline
screening, three from the oyster cohabitation tanks and
three from the control tank with tunicates only.
Trial 2: Cohabitation trial between Vibrio
aestuarianus infected Crassostrea gigas
and Didemnum vexillum
150 Crassostrea gigas were randomly sampled from
Dungarvan, Waterford (selected because OsHV-1
lVar and Vibrio aestuarianus are both present at this
location (Bookelaar 2018)) and of these an initial
sample (n = 30) was screened to assess the prevalence
of V. aestuarianus. The remaining oysters were
divided into three experimental 50 L tanks, each
containing 40 oysters, and held in a constant temper-
ature room at 16 °C and a salinity of 33–35. After 34
days there were 42 oyster mortalities and a further 10
were removed and screened for V. aestuarianus. The
remaining oysters (n = 68) were subsequently divided
into two tanks, 34 per tank.
28 colonies of Didemnum vexillum/colonial spp.
were collected from Carlingford, Co. Louth. The
target species was D. vexillum as this is a prolific
global invader, and ID guides were used to identify
this species in the field, however given the morpho-
logical similarities between colonial tunicate species,
and the heterogeneous nature of colonies fouling the
oyster trestles, other species were also collected and
included in the trials. A tissue sample from all 28
colonies were sequenced at the end of the trial using
the primer pair F16–R497 (Price et al. 2005)to
confirm the community composition. An initial sam-
ple (n = 7 colonies) was removed to assess the
baseline prevalence of V. aestuarianus. The remaining
21 colonies were placed in three 50 L tanks with seven
colonies per tank; this resulted in one control tank with
seven D. vexillum/colonial tunicate spp. colonies only,
and two experimental tanks each containing the V.
aestuarianus-infected oysters and tunicates (7 D.
vexillum/colonial tunicate spp. colonies and 34 C.
gigas 92 tanks). The trial ran for 11 days.
In all laboratory experiments tanks were checked
twice daily and any moribund or dead oysters and
tunicates removed and processed. Tanks were con-
stantly aerated using airlines with airstones and
oysters and tunicates were fed standard marine
invertebrate food 3–4 times per week.
Sample processing—2018 and 2019 samples
Oyster species were processed by carefully opening
the shell without disrupting the tissue within and
draining the excess cavity fluid and seawater onto
clean tissue. In Ostrea edulis, heart and gill smears
were prepared as per Bonamia ostreae screening
(Flannery et al. 2014). Smears were screened for B.
ostreae using a Leica DM500 microscope at the 409
lens. Each slide was observed for five minutes and the
results categorised into class 0: no cells visible after
five minutes, class 1: 1–10 cells, class 2: 11–100 cells
and class 3: heavy infection throughout the slide. Gill
smears were used as a second reference on occasions
where cells were difficult to distinguish within the
heart smears.
The Ostrea edulis heart and piece of gill tissue were
then used for DNA extraction (after smearing) using
10% Chelex Ò100 resin and subsequent PCR
123
K. E. Costello et al.
analysis. In Crassostrea gigas a piece of gill tissue
only was removed for DNA extraction. Oyster sam-
pling in 2018 consisted of 60 O. edulis individuals
with 90 tissue samples screened (gill only in July 2018,
heart and gill for all future screening), and 60 C. gigas
individuals with 60 tissue samples screened (Table 1).
Oyster sampling in 2019 trials consisted of 150 O.
edulis individuals with 300 tissue samples screened,
and 150 C. gigas individuals with 150 tissue samples
screened (Table 3). In both oyster species a transverse
section consisting of gill, gonad, digestive tract and
mantle was then preserved in Davidson’s fixative as
per the OIE (2019) and transferred to 70% ethanol for
long-term storage.
Solitary tunicate samples (Styela clava) were
prepared for molecular work by measuring the length,
wet weight and drained weight then removing the
tunic and dissecting the soft tissue. Tunicates ranged
from 2.7 cm to 11.8 cm in length, therefore the
number of tissue samples was scaled to the size of
the individual (minimum 2 tissue samples and max-
imum 12) and taken from base to oral siphon to ensure
all tissue types were preserved. DNA extraction was
performed in the same manner as the oyster tissue. S.
clava sampling in 2018 consisted of 90 individuals
with 382 tissue samples screened (Table 1). S. clava
sampling in 2019 trials consisted of 32 individuals
with 248 tissue samples screened (Table 3). Samples
also had a histological section consisting of one piece
from base to oral siphon fixed in Davidson’s solution
and prepared as per Flannery et al. (2014), and samples
from 2019 also had a sample (again base to oral
siphon) prepared for in situ hybridisation (ISH) as per
Lynch et al. (2008). Images were captured on a Leica
DM500 microscope using LAS 4.12.0 software.
Colonial samples were also preserved for molecular
work and long-term storage for histology. In the case
of Botrylloides violaceus each small lobe had 3 tissue
sections removed for DNA extraction (120 tissue
samples in total) and 1 section per lobe fixed in
Davidson’s solution (40 in total). In the case of
Didemnum vexillum each colony had 5 tissue sections
removed for DNA extraction (140 tissue samples in
total) and 5 sections fixed in Davidson’s solution (140
in total).
Molecular screening
Prior to molecular screening the quality and concen-
tration of DNA was tested using a NanoDrop
TM
1000
Spectrophotometer. Conventional PCR assays were
initially conducted on oysters, for pathogens of which
they are the natural hosts (see Table 2). If positive
results were obtained from the oysters, the tunicates
were then screened for those pathogens. Screening for
Bonamia ostreae consisted of the BO-BOAS primer
pair (Cochennec et al. 2000) and amplifications were
carried out in a total volume of 50 ll, including 1 ll
DNA, 25.75 ll ddH
2
0, 10 ll59Green GoTaqÒ
Reaction Buffer (Promega), 10 ll dNTPs (1.25 mM
stock solution), 2 ll MgCl
2
(25 mM), 0.25 ll
GoTaqÒDNA Polymerase (Promega) and 0.5 ll each
of forward and reverse primers. Reactions were
carried out on a thermocycler at 94 °C for five minutes
(one cycle), then one minute each at 94 °C, 55 °C and
72 °C for 35 cycles and finally 72 °C for ten minutes.
Haplosporidian primer pairs consisted of MSXA-
MSXB for Haplosporidium nelsoni (causative agent
of multinucleate sphere X disease (MSX)), and the
HapF1-HapR3 pair for generic haplosporidian species.
Haplosporidian amplifications were carried out in a
total volume of 20 ll, including 1 ll DNA, 10 ll
ddH
2
0, 4 ll59Green GoTaqÒReaction Buffer
(Promega), 4 ll dNTPs (1.25 mM stock solution),
0.2 ll GoTaqÒDNA Polymerase (Promega) and
0.4 ll of forward and reverse primers. Reactions were
carried out on a thermocycler at 95 °C for three
minutes, then 95 °C (30 s), 59 °C (MSXA-MSXB)/
48 °C (HapF1-HapR3) (30 s) and 72 °C (30 s HapF1-
HapR3/one minute MSXA-MSXB) for 35 cycles and
finally 72 °C for five minutes.
The primer pair OHVA-OHVB was used to detect
the presence of OsHV-1 lVar as per Lynch et al.
(2013b), with the addition of 1.5 ll dMSO to the mix.
The primer pair DnaJf420-DnaJr456 along with the
probe DnaJp441 were used to detect Vibrio aestuar-
ianus using qPCR with a total volume of 25 ll,
including 5 ll DNA, 5.62 ll ddH
2
0, 12.5 ll Taq-
ManÒ, 0.38 ll probe and 0.75 ll each of forward and
reverse primers. The qPCR thermal profile was as per
McCleary and Henshilwood (2015).
The primer pairs TAP-For/Rev and MER-For/Rev
were used for Minchinia tapetis and Minchinia
mercenariae-like respectively (on a subset of Styela
clava from 2019) as per Albuixech-Martı
´et al. (2020).
123
The role of invasive tunicates as reservoirs of molluscan pathogens
PCR negative controls consisted of double distilled
H
2
O (ddH
2
O) and two positive controls for each
screen. Positive controls for V. aestuarianus were
obtained from the Marine Institute in the form of a
stock solution and dilutions had Ct (cycle threshold)
values ranging from 27 to 34, a mean Ct below 37 was
considered positive. In the case of B. ostreae, positive
controls were not used in the initial PCR on O. edulis
samples but DNA that was isolated in those samples
was used as a positive control in all subsequent
screenings. Results were visualised using gel elec-
trophoresis on a 2% agarose gel. All primer pairs are
listed in Table 4and on occasions where Sanger
sequencing was utilised, samples were sent to Source
BioSciences, Co. Waterford.
Results
2018 field samples (Cork Harbour)
Table 5outlines the screening for pathogens in the two
oyster species over the summer and winter samples. In
Cork Harbour Crassostrea gigas samples were nega-
tive for OsHV-1 lVar in both July 2018 and Novem-
ber 2018, but positive for Vibrio aestuarianus, with a
higher prevalence in July (10%) than November
(3.3%). Haplosporidian spp. were detected in C. gigas
samples using general haplosporidian primers, with
3.3% and 10% prevalence of infection for July and
November respectively. Subsequently, one sample
from July and three samples from November were
sequenced but were unsuccessful in identifying the
haplosporidian species present in C. gigas. Pacific
oysters were not screened for Bonamia ostreae as they
are not the primary host for the pathogen.
26.6% of Ostrea edulis from the November samples
were positive for haplosporidian spp. using general
primers (HapF1-HapR3), and Sanger sequencing of
one sample (after purification using a Qiagen QIA-
quickÒGel Extraction Kit) identified it as Bonamia
ostreae (100% BLASTn). Bonamia ostreae-specific
primers then detected this pathogen in O. edulis
samples from both months, with a higher prevalence in
November (46%) compared to July (3.3%). O. edulis
samples were all negative for Haplosporidium nelsoni
(MSX). Flat oysters were not screened for OsHV-1
lVar or Vibrio aestuarianus.
Microscopic examination of O. edulis heart smears
determined that 56.6% of oysters had no visible
Bonamia ostreae cells, with 70% infections class 0 in
July and 43.3% class 0 in November. For oysters with
a visible infection, class 1 infections were the most
prevalent, again in both months. A single class 3
infection was detected in November (Fig. 2).
Styela clava were screened from Cork Harbour and
were negative for both haplosporidian spp. and OsHV-
1lVar (Table 6). Samples were not screened for
Haplosporidium nelsoni (MSX) as it had not been
detected in the oyster samples. Vibrio aestuarianus
Table 4 List of forward
and reverse primers used for
PCR/qPCR in this study
Primer Sequence 50-30Amplicon
BO-BOAS
(Cochennec et al. 2000)
CATTTAATTGGTCGGGCCGC
CTGATCGTCTTCGATCCCCC
300 bp
HapF1-HapR3
(Renault et al. 2000)
GTTCTTTCWTGATTCTATGMA
AKRHRTTCCTWGTTCAAGAYGA
190 bp
OHVA-OHVB
(Lynch et al. 2013b)
TGCTGGCTGATGTGATGGCTTTGG
GGATATGGAGCTGCGGCGCT
385 bp
MSXA-MSXB
(Renault et al. 2000)
CGACTTTGGCATTAGGTTTCAGACC
ATGTGTTGGTGACGCTAACCG
573 bp
TAP-For/Rev
(Albuixech-Martı
´et al. 2020)
ATCTAACTAGCTGTCGCTAACTCGT
CTTTCAAGATTACCCGGCTCTGC
165 bp
MER-For/Rev
(Albuixech-Martı
´et al. 2020)
ATCTAACTAGCTGTCACTATGGAAAA
ACGCACATTAAAGATTGCCCAGCTCTTT
170 bp
DnaJp441 (probe) FAM-AGG GCA CGT CGG C-MGB N/A
DnaJf420-DnaJr456
(McCleary and Henshilwood 2015)
GTGAAGGGACGGGTGCTAAG
CCATGACAAGTGCCACAAGTCT
123
K. E. Costello et al.
was detected in S. clava in the July samples (6.6%,
lowest Ct 35.5). Bonamia ostreae was also detected in
S. clava in July (5%) and November (10%), and four
samples were sequenced with one confirmed as B.
ostreae (90.6% match, BLASTn). This study also
calculated the wet weight and drained weight of S.
clava to ascertain the potential for tunicates to
transport water containing B. ostreae cells, with 27%
of tunicate biomass from 62 samples (from field trials
and subsequent laboratory cohabitation trials) com-
prised of water contained within the tunic.
Botrylloides violaceus sampled from the marina in
Crosshaven were negative for haplosporidian spp. and
OsHV-1 lVar. However, Vibrio aestuarianus was
detected in B. violaceus samples from July (33.3%,
lowest Ct 35.5). B. violaceus samples were not
screened for H. nelsoni (MSX).
2019 laboratory cohabitation trials
Trial 1: Cohabitation trial between Bonamia ostreae
infected Ostrea edulis and Styela clava
Of the 150 Ostrea edulis collected on 5th April 2019,
30 were instantly removed and screened for Bonamia
ostreae using PCR, with the prevalence of infection
recorded at 13.3%. Oysters were then maintained in
the laboratory for 75 days to allow a higher prevalence
and intensity of infection to develop before cohabita-
tion trials began. 15 live and moribund animals were
screened before the addition of tunicates. B. ostreae
prevalence was 100%, and 46.6% of these were
animals with class 3 infections as evidenced with the
heart smears. This 100% infection was considered the
baseline oyster infection when adding S. clava. The
prevalence of B. ostreae in O. edulis decreased from
the start to the end of the trial, from 100% to 84.3%,
and the prevalence of infection in the control O. edulis
tank (oysters only) was 60% (Fig. 3).
Styela clava screened at the start of the study
showed a 25% prevalence of Bonamia ostreae and this
was considered the baseline tunicate infection. At the
end of the trial, after cohabiting with infected oysters
for two months, this prevalence was 18.75%. The
prevalence of infection in the S. clava in the control
tank, i.e. tunicates only, was 37.5%.
Table 5 PCR (Haplosporidian spp., Haplosporidium nelsoni,
Bonamia ostreae and OsHV-1 lVar), qPCR (Vibrio aestuar-
ianus) and B. ostreae/Ostrea edulis heart smear results for the
detection of pathogens in Crassostrea gigas and O. edulis (NS
= Not Screened, NA = Not Applicable)
Species Date Generic Haplosporidian
spp. PCR (%)
B. ostreae
PCR (%)
Heart
smears (%)
H. nelsoni (MSX)
PCR (%)
OsHV-1 lVar
PCR (%)
V. aestuarianus
qPCR (%)
C.
gigas
11-07-
2018
3.3 NS NA NS 0 10.0
C.
gigas
2-11-
2018
10.0 NS NA NS 0 3.3
O.
edulis
11-07-
2018
0 3.3 30.0 0 NS NS
O.
edulis
23-11-
2018
26.6 46.6 56.6 0 NS NS
Fig. 2 Different classes of Bonamia ostreae infection in Ostrea
edulis heart smears with a sample size of 30 in each month (see
text for each class category)
123
The role of invasive tunicates as reservoirs of molluscan pathogens
All S. clava screened for Minchinia spp. (samples
that were positive for Bonamia ostreae, n = 8) had
100% prevalence of infection of M. mercenariae-like
using PCR, with six samples sequenced for confirma-
tion (100% BLASTn: Minchinia sp. ex Cerastoderma
edule clone 3 small subunit ribosomal RNA gene,
partial sequence; 96.34% BLASTn: Minchinia merce-
nariae small subunit ribosomal RNA gene, partial
sequence). These positive samples encompassed indi-
vidual tunicates from the trial baseline sample,
experimental tanks and control tank, and these same
samples were all negative for M. tapetis. Histology
also revealed sporonts and spore-like structures
belonging to Minchinia mercenariae-like (Fig. 4).
Haplosporidian cells were visualised in a subsam-
ple of Styela clava (n = 5) from the trials using in situ
hybridisation and histology. Bonamia ostreae-specific
in situ hybridisation revealed cells that fit the profile of
B. ostreae, but there was also the potential for them to
be considered aspecific ‘loose’ content due to deteri-
oration of the tunicate tissue during the hybridisation
process. However histological examination of the
coinfected tunicate samples revealed uninucleate
haplosporidian cells consistent with either B. ostreae
or Minchinia mercenariae-like in addition to the
sporonts and spore-like structures belonging to the M.
mercenariae-like species (Fig. 4).
Trial 2: Cohabitation trial between Vibrio
aestuarianus infected Crassostrea gigas
and Didemnum vexillum
Of the 150 Crassostrea gigas collected on 4th July
2019, 30 were instantly removed and screened for
Vibrio aestuarianus using qPCR, with the prevalence
of infection recorded at 93.3%. In the 32 days prior to
the addition of the tunicates there were 42 oyster
mortalities, all of which were screened and had 100%
V. aestuarianus infection. Tunicates were added to
tanks on 6th August and prior to addition of tunicates a
further ten oysters were screened, with 50% oysters
infected with V. aestuarianus. This 50% was consid-
ered the baseline infection of oysters at the start of the
cohabitation trial. Due to the limited number of oysters
as a result of the early high mortality rate there was no
Table 6 Molecular results for the detection of pathogens in Botrylloides violaceus and Styela clava (NS = Not Screened)
Species Date Haplosporidian spp.
PCR (%)
Bonamia ostreae
PCR (%)
H. nelsoni (MSX)
PCR (%)
OsHV-1 lVar
PCR (%)
Vibrio aestuarianus
qPCR (%)
B.
violaceus
16-07-
2018
0 NS NS 0 33.3
B.
violaceus
25-10-
2018
0NSNS00
S. clava 4-07-
2018
0 6.6 NS 0 0
S. clava 11-07-
2018
0 3.3 NS 0 6.6
S. clava 2-11-
2018
010NS00
Fig. 3 % prevalence of Bonamia ostreae in Ostrea edulis and
Styela clava using PCR (control screened at end of cohabitation
trial)
123
K. E. Costello et al.
tank of oysters only as a control. There were no further
oyster mortalities while the cohabitation trial was
running and at the end of the trial all remaining oysters
were screened (n = 68), with prevalence of intensity
just 0.03%.
The total prevalence of infection for the baseline
tunicate sample (7 heterogeneous colonies) was 71.4%
and the total prevalence of infection in the experi-
mental sample (14 heterogeneous colonies) was 50%.
No Vibrio aestuarianus was detected in the control
tunicates (7 heterogeneous colonies) that were held
alone in a tank without any oysters being present.
Given the heterogeneous nature of the tunicate
colonies in Carlingford, and the morphological sim-
ilarities between species, a sample from each of the 28
colonies was sequenced at the end of the trial to
determine the full species composition of the 28
colonies. Sanger sequencing confirmed 11 Didemnum
vexillum (invasive) colonies and 9 Aplidium glabrum
(native) colonies. There was also one sample each of
the tunicate genera Botryllus/Botrylloides sp., Styela
sp. and Ascidiella sp. The remaining five sequences
failed but were visually identified (by comparing
morphology to other sequenced individuals) as D.
vexillum (94) and A. glabrum (91). Infection was
higher in D. vexillum than A. glabrum/other at both the
baseline sample (100% in D. vexillum, 33.3% in A.
glabrum/other species) and at the end of the cohab-
itation trial with mortalities included (55.5% in D.
vexillum, 40% in A. glabrum/other). In the control tank
with no oysters present there was no infection detected
in any tunicate species either from mortalities or at
completion of the trial (Fig. 5).
Fig. 4 (a) and (b) uninucleate haplosporidian ‘fried egg’ cells
consistent with Bonamia ostreae or Minchinia mercenariae-like
in the connective tissue of Styela clava (c) vacuoles in Styela
clava epithelial tissue arising from Minchinia mercenariae-like
spore-like stage with polar nuclei visible (d)Minchinia merce-
nariae-like sporonts in Styela clava connective tissue
123
The role of invasive tunicates as reservoirs of molluscan pathogens
Discussion
This study used OIE-recommended diagnostic tech-
niques and current literature to demonstrate that of a
number of molluscan pathogens recorded in Cork
Harbour, Bonamia ostreae,Minchinia mercenariae-
like and Vibrio aestuarianus are currently present in
commercial bivalves. Furthermore, field surveys and
laboratory trials indicated that these same pathogens
are present in invasive tunicates cohabiting with
oysters at the aquaculture sites. Field surveys also
demonstrated that V. aestuarianus can occur in
invasive tunicates present in marinas removed from
aquaculture sites, raising the possibility that transport
of pathogens may occur from aquaculture sites and
remain in reservoirs outside the zone of infection, for
example via currents or recreational shipping. Addi-
tionally, the haplosporidian M. mercenariae-like was
capable of replicating in Styela clava, suggesting that
this tunicate is not only a carrier of the pathogen but
potentially a viable host. S. clava were also capable of
maintaining a second haplosporidian pathogen, B.
ostreae, without the presence of the primary host
Ostrea edulis. The presence of both B. ostreae and M.
mercenariae-like in S. clava individuals also demon-
strates that the tunicates are susceptible to coinfection.
For both Styela clava/Bonamia ostreae and Didem-
num vexillum/Vibrio aestuarianus the prevalence of
intensity of the respective pathogens was higher at the
start of the trial than at the end of the trial. The filter
feeding mechanism of the tunicates may explain how
they take in the disease and it is possible that in a larger
aquaculture area the tunicates are filtering more so that
although the geographic spread is significantly larger
than in an enclosed tank the increased filtering could
result in a greater intake of pathogenic cells and a
higher infection.
In the case of Vibrio aestuarianus it is possible that
Didemnum vexillum and other colonial spp. may need
to be in contact with oysters to maintain the disease, as
both the baseline sample and the experimental treat-
ments proved positive despite a low prevalence of V.
aestuarianus in the cohabitating Crassostrea gigas at
the end of the trial which suggested that the remaining
oysters were ‘survivors’ that had not suffered Vibrio-
induced mortality. Conversely the control treatment of
tunicates only was negative for V. aestuarianus.
In the case of Bonamia ostreae,Styela clava in
control tanks maintained the disease for two months
without the presence of oysters. Furthermore, the
percentage prevalence of infection was higher in the
control than the baseline and experimental samples,
which indicates that replication in the system may be
possible and the pathogen may have been transmitting.
This would again suggest that S. clava is potentially a
host, not just a carrier or reservoir. The fact that B.
ostreae prevalence was higher in control tunicates
than tunicates cohabiting with oysters may also be
because the oysters were filtering more quickly and
therefore picking up B. ostreae cells so the infection
was reduced in tunicates.
Another key point is that the percentage prevalence
of B. ostreae was higher in O. edulis cohabiting with S.
clava, rather than in the O. edulis control tank,
meaning a cumulative effect of S. clava and O. edulis
on B. ostreae percentage prevalence cannot be ruled
out. O. edulis in this study came from a site where a
selective breeding programme commenced in 1988 for
over 30 years (Lynch et al. 2014) and the slight
reduction in B. ostreae in the oysters from the baseline
to the end of the trial may support that they have been
bred to be resistant to B. ostreae and can maintain it at
a sub-lethal level.
This study focused on Didemnum vexillum as the
target species for laboratory trials, however this
species was growing in heterogeneous colonies and
it is therefore possible that the interwoven nature of
colonial tunicates means they may circle pathogens
Fig. 5 % prevalence of Vibrio aestuarianus in Crassostrea
gigas,Didemnum vexillum and Aplidium glabrum/other species
123
K. E. Costello et al.
both from zooid to zooid within the colony, but also
from one species to another thus heightening the
potential for disease transmission. Furthermore, given
the detection of Vibrio aestuarianus it is highly likely
that other Vibrio spp. that are potentially pathogenic to
commercial bivalves could also be carried by
tunicates.
Although the impact of these pathogens on tunicate
health was not investigated in this study, the combined
results across the solitary and colonial forms indicate
that invasive tunicates can potentially act as reservoirs
of pathogens. Other studies have indicated that in
many cases such reservoirs can transmit the infection
back to susceptible bivalves, for example when
Crassostrea gigas infected with Bonamia ostreae
were held with naive Ostrea edulis, the O. edulis
cavity fluid then tested positive for the pathogen
(Lynch et al. 2010). B. ostreae has also been detected
in a number of benthic macroinvertebrates, including
the native European sea squirt Ascidiella aspersa and
other phyla including phylum Crustacea (Carcinus
maenas) and phylum Cnidaria (Actinia equina)
(Lynch et al. 2007). This circling of the disease
through diverse invertebrate groups could suggest that
there are multiple avenues for reservoirs to come in
contact with the disease.
Small and Pagenkopp (2011), noted that the
environment itself can act as a reservoir and this is
demonstrated by Bonamia ostreae, as the OIE states
that it can live for up to one week in the water column.
This also means that Styela clava could potentially act
as a mechanical vector and carry water with the protist
present to different sites if transferred on improperly
cleaned aquaculture gear or non-biosecure stock
shipments.
Tunicates were used as a model taxon in this study,
however it is important to note that invasive species
across different phyla may be also able to move
significant pathogens. Davies et al. (2019) investigated
the globally invasive shore crab (Carcinus maenas)in
Wales and found it infected with Hematodinium sp.
and Hematodinium perezi. The disease was also
present in seawater eDNA samples, possibly due to
the release of infectious stage dinospores from mori-
bund individuals. Shore crabs in the study site were
cohabiting with the commercially valuable edible
brown crab, and this demonstrates that ability of
disease reservoirs to enable pathogens to persist in
habitats utilised by commercial shellfish.
The presence of disease in invasive tunicates is of
significant interest to the aquaculture sector, particu-
larly in the context of climate change, as species
potentially expand their invaded range (Hellmann
et al. 2008). When coupling the mechanical impacts of
the tunicates themselves, for example competition for
space, with their potential to interact with pathogens
affecting commercial bivalve species, they emerge as
a threat that warrants serious consideration and
enhanced biosecurity. The Food and Agriculture
Organization of the United Nations recognised the
risk of invasive species in the 2030 Agenda for
Sustainable Development, and noted the need to
introduce measures to eradicate or control such
species (FAO 2017). A number of aspects need to be
addressed to lessen invasive impacts; reproductive
strategies need to be identified, transport via shipping
or other pathways such as aquaculture needs to be
minimised and mechanisms identified to remove and
inactivate fouling organisms. Examples of these
mechanisms include controls such as those described
in Hillock and Costello (2013) with a study on
desiccation methods for Styela clava, or Turrell et al.
(2018) with a study of different bath treatments that
could be applied to commercial bivalve consignments
to induce full mortality in Didemnum vexillum.
It is necessary to develop a global approach to
ensure aquaculture transfer takes management of both
invasive species and disease into account, thereby
developing legislation and codes of practice. In 2016
shelled molluscs constituted 58.8% of the combined
production of marine and coastal aquaculture (FAO
2018) and growth projections for the future are
positive, meaning that molluscan aquaculture will
play an essential role in global food resources
(Rodgers et al. 2015; Arzul et al. 2017). However,
this sector is intrinsically linked to the movement of
invasive species and disease cycling is a further
constraint on growth. Guidelines and legislation are
only as good as the knowledge that informs them, and
it is not feasible to protect against species on which
there is little knowledge about health impacts or
potential roles in disease cycling. Accordingly, it is
necessary to broaden the understanding of the how
pathogens utilise invasive species, and the subsequent
impact on the commercial sector.
This study raises questions relating to the viability
of tunicates as sources of infection and suggests that
the taxon should be taken into account in risk
123
The role of invasive tunicates as reservoirs of molluscan pathogens
assessments and disease management, particularly as
pathogens can endure in species that are not their true
host (Lynch et al. 2010). Screening of the tunicates
focused specifically on pathogens that had previously
been detected in the natural hosts, however further
work could expand on the diversity and seasonality of
pathogen species, as if tunicates are reservoirs then a
range of pathogens might be present in them at times
of the year not found in oysters. It also demonstrates
that it is important to have good monitoring networks
and collaborative efforts so that information on
invasive species with potential impacts is widely
shared and available within the aquaculture sector
(Brenner et al. 2014).
Acknowledgements This work was supported by the Bluefish
Project (Grant Agreement No. 80991), part-funded by the
European Regional Development Fund (ERDF) through the
Ireland Wales Co-operation Programme. The authors would like
to thank Gavin Deane of the Royal Cork Yacht Club, the Hugh-
Jones of Atlantic Shellfish Ltd. and all at Carlingford Oyster
Company and Dungarvan Shellfish Ltd. for their assistance
during fieldwork. We would also like to thank Sara Albuixech-
Martı
´, Kate Mahony, Gary Kett, Ashley Bennison and Alex
McGreer for their support during the laboratory trials. Lastly, we
would like to thank Dr Eileen Dillane, Dr Elizabeth Cotter, Luke
Harman and Allen Whitaker for their technical assistance.
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The role of invasive tunicates as reservoirs of molluscan pathogens
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