Canadian Journal of Zoology
The life history and feeding ecology of velvet shell, Velutina velutina (Gastropoda: Velutinidae), a specialist predator of ascidians
Journal: Canadian Journal of Zoology
Manuscript ID cjz-2018-0327.R1
Manuscript Type: Article
Date Submitted by the 03-Jun-2019 Author:
Complete List of Authors: Sargent, Philip; Northwest Atlantic Fisheries Centre, Fisheries and Oceans Canada Hamel, Jean-Francois; Society for the Exploration and Valuing of the EnvironmentDraft Mercier, Annie; Memorial University of Newfoundland, Ocean Sciences
Is your manuscript invited for consideration in a Special Not applicable (regular submission) Issue?:
Velutina velutina, velvet shell, velutinid, gastropod, invasive species, Keyword: specialist predator, ascidian
https://mc06.manuscriptcentral.com/cjz-pubs Page 1 of 42 Canadian Journal of Zoology
1
The life history and feeding ecology of velvet shell, Velutina velutina (Gastropoda: Velutinidae), a specialist predator of ascidians
P. S. Sargent*, J-F. Hamel, and A. Mercier
P. S. Sargent1 Department of Ocean Sciences, Memorial University, St. John’s (Newfoundland and Labrador) Canada A1C 5S7 Email: [email protected]
J-F Hamel Society for the Exploration and ValuingDraft of the Environment (SEVE), Portugal Cove-St. Philips (Newfoundland and Labrador) Canada A1M 2B7 Email: [email protected]
A. Mercier Department of Ocean Sciences, Memorial University, St. John’s (Newfoundland and Labrador) Canada A1C 5S7 Email: [email protected]
* Corresponding Author: Philip S. Sargent Department of Fisheries and Oceans Canada, Northwest Atlantic Fisheries Centre,
80 East White Hills Road, St. John’s, Newfoundland and Labrador, Canada, A1C 4N1
Email: [email protected] Phone: 1 (709) 772-4278 Fax: 1 (709) 772-5315
1 Current Contact Information for P. S. Sargent: Department of Fisheries and Oceans Canada, Northwest Atlantic Fisheries Centre, 80 East White Hills Road, St. John’s, Newfoundland and Labrador, Canada, A1C 4N1 Email: [email protected]
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 2 of 42
2
The life history and feeding ecology of velvet shell, Velutina velutina
(Gastropoda: Velutinidae), a specialist predator of ascidians
P. S. Sargent*, J-F. Hamel, and A. Mercier
Abstract
Velvet shell, Velutina velutina (O. F. Müller, 1776), is a specialist predator of ascidians, like other members of the gastropod family Velutinidae. Globally, invasive ascidians have become problematic, ecologically and economically, yet ecological knowledge of velutinids remains limited. This study outlines the life history and feeding ecology of V. velutina in eastern CanadaDraft based on laboratory work complemented by field observations. The life history of V. velutina is closely linked with ascidians, which serve as prey and protection for their egg capsules. Egg capsules were embedded within tunics of Aplidium glabrum (Verrill, 1871) and Ascidia callosa Stimpson, 1852, but the latter was preferred. Seasonal behavioural shifts were consistent annually and corresponded with seawater temperature cycles. Feeding dominated during the coldest months (January – May), growth occurred as water temperature increased to the annual maximum (June and July), transitioned to mating during the warmest period
(July/August), and egg capsule deposition dominated as water temperature declined
(November – January). Larvae hatched between January and July after 2 – 4 months of development. Velvet shell preyed on all ascidian species presented during this study, including golden star tunicate, Botryllus schlosseri (Pallas, 1766), and vase tunicate,
https://mc06.manuscriptcentral.com/cjz-pubs Page 3 of 42 Canadian Journal of Zoology
3
Ciona intestinalis (Linnaeus, 1767), two non-indigenous species, although solitary
species were preferred.
Keywords Velutina velutina, velvet shell, velutinid, gastropod, invasive species, specialist predator,
ascidian
Introduction
Members of the gastropod family Velutinidae (formerly Lamellariidae) are
specialized predators of ascidians that closely associate with their prey (Fretter and
Graham 1962; Ghiselin 1964; Behrens 1984; Dias and Delboni 2008) and deposit their egg capsules within the tunic of ascidiansDraft (Diehl 1956; Strathmann 1992; Page 2002; Dias and Delboni 2008). The close association of velutinids and ascidians has even been
described as a symbiotic (sensu lato) relationship (Queiroz and Sales 2016).
Available literature on velutinids consists primarily of anatomical and
taxonomical descriptions (e.g. Gulbin and Golikov 1997; 1999), observational references
to biogeography, depth distributions, and habitat associations (e.g. Fretter and Graham
1981; Gulbin 2005). Scarcity of ecological and biological information on velutinids may
be due in part to their cryptic nature and the fact that few specimens have ever been
collected (Lambert 1980). The mantle of most velutinid species partially or completely
covers a thin and weakly calcified shell (Gulbin and Golikov 1997) and often mimics the
colour and texture of the tunic of ascidians (Ghiselin 1964; Lambert 1980; Behrens
1984). This renders velutinids almost indistinguishable from their prey (Ghiselin 1964;
Behrens 1980; Lambert 1980). Furthermore, velutinids are commonly misidentified as
dorid nudibranchs (Jeffreys 1867; Behrens 1984; Dias and Delboni 2008) and some
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 4 of 42
4
species are nocturnal, remaining concealed during the day (Sarma and Pattanaik 1986). In
general, our understanding of the ecological role of velutinids in subtidal communities is
limited.
The velvet shell, Velutina velutina (O. F. Müller, 1776), is one of the most
common and widely distributed velutinid species in northern seas (Derjugin 1950). It is a
sublittoral species with a boreal-arctic distribution (Gulbin and Golikov 1999; Gulbin
2005) found to depths of 1000 m (Fretter and Graham 1981; Gulbin 2005). In the
northwest Atlantic, this species extends from the Arctic to as far south as Cape Cod
(Massachusetts, USA) but becomes increasingly rare and occurs deeper towards its
southern range (Fretter and Graham 1981). Members of the genus Velutina have an
external shell and the mantle may onlyDraft cover a portion of the shell’s surface along the
edge of the aperture (Fretter and Graham 1981). To our knowledge, the only ecological
studies of V. velutina were conducted in northern Europe and examined predation on and
egg deposition in the ascidian Styela coriacea (Alder and Hancock, 1848) (Diehl 1956),
copulation (Diehl 1956), and planktonic larval development (Lebour 1935; Thorson
1946; Mileikovsky 1960; Fretter and Pilkington 1970; Thiriot-Quiévreux 1974).
A better understanding of the ecology of velutinids would not only increase our
knowledge of this poorly understood gastropod family but may also be relevant to the
control of some aquatic invasive species. Non-indigenous ascidians have proven to be
very successful invasive species globally (Lambert 2007). Invasive ascidians may have
significant impacts both ecologically, by altering community structure (Lambert and
Lambert 2003; Valentine et al. 2007), and economically, by fouling man-made structures, especially shellfish aquaculture infrastructure (Carver et al. 2003; LeBlanc et al. 2007;
https://mc06.manuscriptcentral.com/cjz-pubs Page 5 of 42 Canadian Journal of Zoology
5
Rocha et al. 2009). Yet, knowledge of biotic resistance of benthic marine communities to
such invasions is lacking (Lambert 2007; Epelbaum et al. 2009). Determining the
response of benthic communities to invasive species (e.g. presence and role of potential
predators such as velutinids) is crucial to understand and mitigate species invasions.
The purpose of the present study was to investigate the life history and feeding
ecology of V. velutina and provide insight into the role of velutinids within the subtidal
communities they inhabit. It combined laboratory studies complemented by field
observations to determine diet and feeding habits (preferences, rates, and periodicity) as
well as seasonal behavioural patterns. Mating, egg laying, larval development and
hatching were characterized and the role of environmental factors in the control of these
behaviours were also examined. As Drafta specialist predator of ascidians, V. velutina may
represent a natural means of mitigating the impacts of some non-indigenous ascidians.
Materials and methods
Collections and field sites
Most field observations and collections of V. velutina were made
opportunistically (77.8% of dives) while conducting dives for other purposes. Field
observations of habitat associations and behaviours of V. velutina were recorded2 and
used to complement observations recorded in the laboratory. Over the course of 41 dives
(mean = 36 min dive time) conducted between November 2005 and August 2011 around
insular Newfoundland, 105 velutinids were observed and 75 individuals were collected
and transferred to Memorial University’s Oceans Science Centre3. Five additional
2 as per Table S1 and S2, respectively 3 Tables S1 and S3
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 6 of 42
6 individuals were collected over four dives between March and May 2017 (mean = 44 min dive time) and transferred to the Northwest Atlantic Fisheries Centre3. In 2006 and 2007, samples of callused tunicate, Ascidia callosa Stimpson, 1852, were collected from two sites4 and examined for the presence of developing or previously hatched egg capsules and settled larvae of V. velutina.
Holding conditions and maintenance
Unless otherwise stated for specific experiments, velutinids were maintained in laboratory microcosms consisting of tanks (23 and 40 L) supplied with flow-through unfiltered seawater (1.5 – 3.0 L min-1) at ambient temperature under either natural or fluorescent lighting (25 – 70 lux) set on timers to follow the natural daylight cycle.
Seawater temperature in tanks was recordedDraft on a daily to weekly basis. An inline chiller was used between July and October to maintain temperature below 14 °C, as observed in their natural habitat.
Velutinids (juveniles and adults) were provided various ascidian species ad libitum. Based on availability, velutinids were maintained primarily on Halocynthia pyriformis (Rathke, 1806) with opportunistic additions of other indigenous species, including Aplidium glabrum (Verrill, 1871), Didemnum albidum (Verrill, 1871), Ascidia callosa Stimpson, 1852, Boltenia echinata (Linnaeus, 1767), Molgula citrina Alder and
Hancock, 1848, and Molgula griffithsii (MacLeay, 1825). Pieces of encrusting algae, including Lithothamnion glaciale Kjellman, 1883 and Clathromorphum compactum
(Kjellman) Foslie, 1898, and pieces of shale rock, encrusted with L. glaciale, were placed in tanks for shelter.
4 Table S3
https://mc06.manuscriptcentral.com/cjz-pubs Page 7 of 42 Canadian Journal of Zoology
7
Feeding
General feeding behaviour
Feeding behaviour was recorded weekly from April 2006 to June 2011.
Observations of velutinids were classified5 and feeding positions on ascidians were
confirmed by the presence of feeding holes. Approximate length of proboscis relative to
shell length of feeding velutinids was noted.
To examine fine-scale behavioural shifts, as well as length of time spent feeding
on ascidians, complementary daily observations (from September 2008 to September
2009) were made on five isolated adult velutinids fed the following ad libitum: Aplidium
glabrum, Ascidia callosa, Boltenia echinata, Halocynthia pyriformis, and Molgula
citrina. Intervals between feeding bouts,Draft and number of ascidians consumed over time
were recorded. Some ascidian prey where dissected to assess the proportion of remaining
tissues/organs (n = 16) while others were retained to assess mortality rates from predation
events (n = 8).
Prey preferences: feeding experiments
A series of feeding experiments were conducted in the laboratory to test the null
hypothesis that V. velutina exhibits no preferences among available ascidian prey species
(see Table 1). For experiments with non-indigenous ascidian species, every 1 – 3 d up to
30% of water and all wastes from tanks were removed and bleached for 24 h before being
discarded, to avoid propagation. Before each experiment, ascidians were acclimated for 2
– 3 d and velutinids were introduced after being starved for 4 d to standardize hunger
levels. When predation was suspected, velutinids were removed from ascidians for
5 Table S2
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 8 of 42
8 confirmation and thereafter replaced haphazardly in their tank. Confirmation of a predation event was recorded when a hole was drilled through the tunic.
Growth
Shell length (SL) of 42 velutinids (mean initial SL = 15.7 mm, range = 6.7 – 24.2 mm) was measured monthly from October 2007 to January 2010 as the maximal shell diameter to the nearest 0.1 mm to examine seasonal growth patterns.
Mating and egg laying
Interactions between individual V. velutina6 were recorded weekly in the laboratory. Grouped or paired individuals were examined for copulation, and whether they were acting as males, females, or both simultaneously. Copulation was described based on 81 observed copulation eventsDraft from July 2006 to December 2008. Egg laying and associated behaviours were recorded weekly in the laboratory. Numbers of intact and hatched egg capsules were also recorded in the laboratory to estimate the total number of capsules deposited annually per velutinid. Complementary data were gathered from 5 isolated adult velutinids from September 2008 to September 2009 to examine behavioural responses and preferences for egg laying in different ascidian species (Ascidia callosa,
Molgula citrina, Boltenia echinata, and Aplidium glabrum). Relative position and behaviour of individual velutinids were recorded every 1 – 4 d, while number of egg capsules laid by each velutinid, and in which ascidian species, were recorded weekly.
Larval development, hatching, and settlement
Egg capsules of V. velutina laid in the laboratory and collected from the field were sampled periodically throughout their development (from initial deposition to
6 as defined in Table S2
https://mc06.manuscriptcentral.com/cjz-pubs Page 9 of 42 Canadian Journal of Zoology
9
hatching). To examine developing larvae, the cap was removed from each capsule and
larvae were extracted with a pipette, counted, and photographed using a stereomicroscope
(Nikon SMZ1500). Egg capsules deposited in the laboratory were counted and examined
weekly for signs of hatching. Dates of when capsules first started hatching and when the
last egg capsule hatched were recorded. Observations of veliger larvae naturally hatching
from capsules were recorded on two occasions and information on larval behaviour was
noted. Observations of settled larvae from ascidians collected in the field were also
reported.
Seasonal behaviour
Seasonal behavioural patterns were determined by recording weekly relative
position in the tank and behaviour ofDraft individual V. velutina7 between October 28, 2006
and May 3, 2009. During that period the mean number individuals in the tank was 18 but
ranged from a high of 30 individuals in 2007 to a low of 5 individuals at the end in 2009.
Velutinids were numbered with India ink on their dorsal shell surface to track them
individually. As ink labels faded over time, they were retraced as necessary.
Statistical analysis
Prey preference results were scored as presence/absence of a predation event on
each species. Choices among native species and between B. schlosseri and native species
were analysed using generalized linear models (GLM) with binomial distributions and
logit links. Results for the pairwise feeding experiment with C. intestinalis were analyzed
using a GLM with a binomial distribution. Monthly shell growth was compared against
monthly change in water temperature using a GLM. GLMs and ANOVAs on them were
7 as per Table S2
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 10 of 42
10
run in R (R Core Team 2018). Descriptive summary data are provided as mean ±
standard deviations.
Results
Field observations and habitat characteristics
Velutinids were observed (n = 110) and collected (n = 80; mean = 12.6 mm SL;
range 0.6 – 26.9 mm SL) primarily on vertical bedrock substrates or on undersides of
rocks at depths between 5.0 and 16.5 m8. Individuals were collected mostly in autumn
(October, n = 14; November, n = 16) in association with Aplidium glabrum and Ascidia callosa for deposition of egg capsules, or in spring (April, n = 13; May, n = 14) when feeding on various ascidians9.
Feeding Draft
Velutina velutina (O. F. Müller, 1776) preyed on all indigenous solitary ascidian species collected sympatrically (Ascidia callosa, Boltenia echinata, Halocynthia pyriformis, Molgula citrina) and non-sympatrically (Molgula griffithsii) but preyed less frequently on indigenous colonial species (Aplidium glabrum and Didemnum albidum).
In feeding trials with non-indigenous species, V. velutina showed limited feeding on the
colonial Botryllus schlosseri but consumed most internal organs (≥ 90%) when preying
on the solitary Ciona intestinalis. Velutina velutina (O. F. Müller, 1776) preyed on a wide
size range of ascidians, from those that were considerably smaller than the individual
velutinid to many times its size.
8 see Table S3 9 See Table S1
https://mc06.manuscriptcentral.com/cjz-pubs Page 11 of 42 Canadian Journal of Zoology
11
During the multi-choice feeding experiment with native ascidian species,
predation was observed on B. echinata (n = 6) and H. pyriformis (n = 5), but was not
detected on A. callosa, A. glabrum or D. albidum. Results of the ANOVA performed on
the GLM confirmed that prey species was significant in predicting the number of
predation events (Chi-square, χ² = 66.375, df = 2, P < 0.001). In the complementary
multi-prey choice experiment including the non-indigenous species Botryllus schlosseri,
V. velutina preyed on all species examined but preyed more on the solitary species (H.
pyriformis, n = 35; M. citrina, n = 10; B. schlosseri, n = 2; A. glabrum n = 1), and A.
glabrum was only preyed on after all solitary species had been consumed. Results of the
analyses indicated that prey species was a significant predictor of number of predation
events (Chi-square, χ² = 41.627, df =Draft 4, P < 0.001), and that all species were preyed upon
significantly less often than H. pyriformis (P < 0.003). In the pair-wise feeding
experiment with the non-indigenous Ciona intestinalis, predation was detected on all
species but A. callosa (C. intestinalis, n = 2; B. echinata, n = 2; H. pyriformis, n = 1; M.
citrina, n = 1). Prey species was again significant in predicting the number of predation
events (Chi-square, χ² = 230.01, df = 5, P < 0.001) but due to the low number of
predation events it was not possible to determine specific species preferences. Results of
these feeding experiments reject the null hypothesis that V. velutina exhibits no
preferences among available ascidian prey species and demonstrates that solitary ascidian
species are preferred.
Feeding behaviour in V. velutina (Fig. 1A) included: cleaning the prey ascidian’s
tunic surface with the radula; drilling a hole through the tunic; inserting the proboscis
through the hole (Fig. 1B – 1D); and extending the proboscis up to 1.5 times its shell
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 12 of 42
12 length inside the prey to consume internal organs (Fig. 1E, 1F). Predation was not observed on dead ascidians but velutinids occasionally fed through recently abandoned holes drilled by congeners.
Velutina velutina (O. F. Müller, 1776) exhibited feeding behaviour for 84.6 ± 25.9 d and preyed on 5.2 ± 2.4 ascidians annually. Average time between feeding events was
45.5 ± 65.6 d but one individual went 245 d between feeding events. Velutina velutina
(O. F. Müller, 1776) demonstrated preferred feeding positions for each ascidian species
(Fig. 2). Field observations were similar, as 71.4% of velutinids that drilled or fed on H. pyriformis did so near the base, and the single observation of predation on M. citrina was through the side. In contrast, feeding on A. callosa in the field was observed twice through the side and once on top, nearDraft a siphon, as the base of the ascidian was inaccessible. Ascidians dissected post-feeding showed up to ~95% of internal tissues consumed (Fig. 1E, 1F) and all ascidians that were preyed upon eventually died.
Growth
Average growth of V. velutina was negligible throughout most of the year but increased significantly in June and July, which corresponded with the peak in water temperature (Fig. 3). Results of the ANOVA performed on the GLM indicated that change in water temperature is significant in predicting shell growth (Chi-square, χ² =
57.452, df = 1, P < 0.001). However, some smaller individuals (< 10 mm initial SL) exhibited growth throughout the year. Of the 42 individuals measured monthly, 21 survived to provide annual growth rates. Annual growth rates declined with increasing initial SL but beyond 16.9 mm initial SL growth stabilized at 0.5 ± 0.5 mm yr-1.
https://mc06.manuscriptcentral.com/cjz-pubs Page 13 of 42 Canadian Journal of Zoology
13
Maximum growth rate recorded for an individual was 3.8 mm mo-1 (July 2009, initial SL
= 12.3 mm) and 7.9 mm yr-1 (initial SL = 9.0 mm).
Mating
Mating behaviours of V. velutina included aggregation, pairing, and copulation,
all of which showed some degree of overlap. Most notable was the overlap between
aggregating and pairing behaviours which typically co-occurred between late June and
early December, while copulation (Fig. 4A) persisted until late January. In the laboratory,
aggregating (n = 97 individuals) and pairing (n = 103 individuals) velutinids were mainly
concealed (80.4% and 65.0% of observations, respectively). Aggregations of up to 9
individuals were observed in clusters or in a line beneath rocks, coralline algae, or
ascidians. Likewise, in the field, aggregationsDraft of up to 7 individuals were observed under
rocks, in crevices of vertical rock walls, or near the bases of Ascidia callosa. All field
observations of pairings (n = 4 pairs) were on the undersides of rocks. In the laboratory,
copulation (n = 153 observations) occurred more frequently concealed beneath rocks and
coralline algae (41.2%) than on A. callosa (24.8%), whereas in the field, copulation (n =
6 individuals) occurred more frequently on A. callosa (66.7%) than in rock crevices
(33.3%).
Velutina velutina (O. F. Müller, 1776) copulated face to face but somewhat off-
centred to align reproductive organs located on their right sides (Fig. 4A). Velutinids
were observed undergoing sex role alternation during copulation as one pair, observed for
1.75 h, initially acted in one sex role but upon separation at the end of this period were
found to have switched sex roles. Laboratory observations of copulation were
predominantly unilateral (91.3%) but a few pairs (3.8%) exhibited simultaneous
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 14 of 42
14
reciprocal copulation. During one reproductive season (2008/2009), individual velutinids
copulated on average 4.1 ± 1.3 times. Individuals acted as either male or female in
roughly equal proportions (43.6% as males, 38.5% as females) and less frequently as
male and female simultaneously (10.3%). The smallest individual V. velutina observed
copulating was 10.7 mm SL, yet one smaller individual (9.0 mm SL) was observed laying
an egg capsule, indicating this species may be mature by 9.0 mm SL.
Egg laying
Velutina velutina (O. F. Müller, 1776) can store spermatozoa for extended periods
or else is capable of self-fertilizing. Three of five individuals isolated to examine fine-
scale behavioural shifts and feeding rates were observed mating prior to isolation. These
velutinids were initially deprived of Draftsuitable ascidian species for egg capsule deposition.
Consequently, some attempted to lay capsules on other substrates including the surface of
Boltenia echinata, a clam shell, and even the tank surface, but these capsules did not develop. When later provided suitable ascidian species, all isolated velutinids began laying egg capsules, between 148 and 178 d after isolation. However, larvae from only three of those isolated velutinids developed and hatched.
Velutina velutina (O. F. Müller, 1776) only deposited egg capsules in Aplidium glabrum and Ascidia callosa but preferred the latter. One isolated velutinid was provided
A. glabrum and A. callosa simultaneously, but deposited capsules only in A. callosa.
Another isolated velutinid, initially provided A. glabrum, began to deposit capsules within it, but when A. callosa was later introduced, it deposited its remaining capsules in
A. callosa. Velutinids associated with A. glabrum and A. callosa in summer (late July –
early August), 3 – 4 months prior to initiation of egg laying in autumn. During this time,
https://mc06.manuscriptcentral.com/cjz-pubs Page 15 of 42 Canadian Journal of Zoology
15
maturing gonads with bright orange oocytes were often observed through the posterior
portion of the shell.
While embedding egg capsules, V. velutina firmly attached to an ascidian’s tunic
(Fig. 4B). If forcibly removed, velutinids were often observed holding the capsule in their
ventral pedal gland, a circular indentation near the center of the foot. After interrupting
capsule deposition, capsules exposed on the tunic surface did not develop viable larvae,
and often burst open. However, egg capsules that were partially embedded when the
velutinid was removed would embed themselves in the ascidian tunic with no further
assistance from the parent, and the cap formed a bubble on the tunic surface. If
uninterrupted, velutinids embedded egg capsules in the ascidian tunic, leaving a striated
cap flush on the tunic surface (Fig. 4C).Draft As larvae began to develop, egg capsules
expanded in size with larvae taking up the lower portion of the capsule (Fig. 4D). Veliger
larvae of V. velutina were observed hatching from egg capsules in Ascidia callosa on two
occasions in April 2006. Most larvae emerged in groups of 2 – 3, hovered briefly, and
then swam to the surface where they remained. During the first hatching observation, the
host A. callosa broadcasted gametes soon after velutinid larvae began to emerge (Fig.
4E). When all larvae emerged, an empty cavity was left in the ascidian’s tunic (Fig. 4F).
Over time the empty capsule filled in with growth of new tunic tissue and a dark circle
remained as a scar where the capsule opening was located.
Based on data from 2006/2007 and 2007/2008, individual velutinids laid up to 18
egg capsules annually (mean = 8.2). In the laboratory and in the field, multiple velutinids
often converged on an individual A. callosa to deposit egg capsules (Fig. 4B). Up to 9
velutinids in the laboratory and 2 velutinids in the field, exhibited egg laying behaviour
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 16 of 42
16 on an individual A. callosa. In the laboratory, as many as 53 intact capsules were recorded in an individual A. callosa. In the field, one large A. callosa (L: 90.0 × W: 63.3
× H: 45.5 mm) contained more than 30 intact capsules with evidence of at least 30 recently hatched capsules. In contrast, even a very small A. callosa (L: 9.9 × W: 8.3 × H:
4.8 mm), smaller than most adult V. velutina, was found containing 2 capsules. Mean number of larvae per capsule was 1392 and ranged from 408 – 1944 in the field and 538
– 3356 in the laboratory.
Larval development, hatching, and settlement
Egg capsules of V. velutina were opened periodically and larvae were photographed throughout their development (Fig. 5). Eggs (Fig. 5A) developed into 4- cell embryos after about 3 – 4 d (Fig.Draft 5B). They reached early veliger stage after ~2 weeks of development (Fig. 5D, E) and became fully developed echinospira veligers
(Fig. 5F, G, H) ~3 weeks of age. There was no evidence that V. velutina used nurse eggs to feed developing offspring. Larvae hatched (Fig. 4E) from early January to mid-July
(Table 2) and swam in the water column as planktonic echinospira veligers (Fig 5F, G,
H). In the communal tank, egg capsules laid at the start of each egg laying period (late
October/early November), took on average 71 d to hatch (range 60 – 81 d). Water temperature during those three incubation periods averaged 4.9 ºC (1.1 – 9.0 ºC).
However, from three isolated velutinids in 2009, egg capsule deposition started near the end of the egg laying season (mid-February/mid-March) and took on average 111 d to hatch (range 109 – 113 d), as water temperature averaged 2.1ºC (-1.9 – 10.5 ºC).
Settled larvae of V. velutina were observed on 8 individual A. callosa (17.4%) collected in April 2006 (Fig. 5I) but none were observed on A. callosa collected in March
https://mc06.manuscriptcentral.com/cjz-pubs Page 17 of 42 Canadian Journal of Zoology
17
or November 2007. All settled larvae under laboratory conditions died unexpectedly soon
after. Settled velutinid larvae were not detected on any other ascidian species or substrate.
Seasonal behaviour
Velutina velutina (O. F. Müller, 1776) exhibited consistent annual succession of
behaviours (Fig. 6, Table 2). The proportion of feeding V. velutina increased as water
temperature decreased in winter and rapidly declined as water temperature increased in
summer (Fig. 6). Rapid transitions between feeding and mating modes occurred between
June and July (Fig. 6). Mating was the dominant behaviour observed during the warmest
time of year, late summer to late autumn (4.2 – 9.9 °C). Egg laying then became the
dominant behaviour as water temperature started to decline in late November (from 7.2
°C in 2007) and in early December (fromDraft 5.7 °C in 2008), peaking in December/early
January and progressively declined as water temperature reached its lowest level of the
year in late March/early April (Fig. 6). Velutinids were generally concealed under
coralline algae and rocks and fed near the base of ascidians from April to October. From
November to March, which coincided with the egg laying period, velutinids were more
exposed, mobile, or observed on the sides or tops of ascidians. Behavioural observations
from the field (n = 69) were consistent with the succession of behaviours recorded in the
laboratory (Fig. 6). Feeding field observations (n = 15) occurred mainly (73.3%) from
April to June. Mating in the field occurred (n = 12) primarily (91.7%) from October to
December. Egg laying behaviour in the field (n = 20) was mostly (80.0%) between
November and April. Further, evidence of developing or recently hatched egg capsules of
V. velutina in A. callosa collected from the field were found primarily in the spring.
Overall, 39.1% of A. callosa (n = 46) showed evidence of egg capsules in April 2006,
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 18 of 42
18
13.2% (n = 38) in March 2007, and only 3.8% (n = 52) in November 2007. Occurrences of egg capsules may have been underestimated as biofouling on A. callosa made identification of capsules difficult.
Discussion
Velutina velutina (O. F. Müller, 1776) was consistently found on hard substrates, such as vertical rock walls and undersides of rocks, near ascidians, upon which they fed and deposit egg capsules, as reported in northern Europe (Jeffreys 1867; Diehl 1956;
Fretter and Graham 1981). Unlike some previous reports, V. velutina was not found associated with mud, sand (North and Baltic Seas; Ankel 1936), gravel/pebble mixtures, or macroalgae beds (Gulbin and Golikov 1999). In general, velutinids live on variety of substrates provided ascidians are nearbyDraft (Gulbin 2005).
Velutina velutina (O. F. Müller, 1776) preyed upon all ascidian species tested in the present study but preferred solitary species. This preference was also reported in the
North and Baltic Seas (Ankel 1936; Diehl 1956), and in San Juan Islands (northeast
Pacific; Young 1985). Preferred feeding locations on each ascidian species were detected, which may be related to mechanical, physical, and chemical anti-predator defences exhibited by different ascidians (see Lambert 2005), and how velutinids overcome those defences. During this study, predation on Halocynthia pyriformis occurred predominantly near the attachment base, as was likewise reported by Diehl (1956) for Styela coriacea.
This feeding location may reduce risk of exposure of V. velutina to potential predators and, as suggested by Young (1985), this area of the tunic may be softer, compared to the anterior end. In contrast, Boltenia echinata exhibits large, flexible spines over a thick leathery tunic, which in this study restricted predation by V. velutina to the bare base or
https://mc06.manuscriptcentral.com/cjz-pubs Page 19 of 42 Canadian Journal of Zoology
19
siphons. Moreover, V. velutina preyed less frequently on A. callosa than other solitary
ascidians, likely due to its thick tunic and because it bioaccumulates vanadium (Stacey
2009), a heavy metal that may inhibit predation (Stoecker 1980; Young 1986). The low
predation observed on A. callosa may also be explained by the fact that V. velutina used
them to lay their egg capsules.
Here, V. velutina commonly formed spawning aggregations on or near A. callosa,
within which they deposit egg capsules. Many gastropods aggregate for reproduction on
their feeding grounds at sites suitable for egg deposition (Fretter 1984) to optimize use of
suitable sites for larval development where predation on capsules is limited (Martel et al.
1986). Velutina velutina (O. F. Müller, 1776) began aggregating in summer and persisted
until egg laying concluded in the spring.Draft In Atlantic Canada, gastropods such as
Buccinum undatum Linnaeus, 1758 and Nucella lapillus (Linnaeus, 1758), likewise
aggregate for mating and spawning, until egg capsules are laid (Feare 1970; Martel et al.
1986). Spawning aggregations of N. lapillus may consist of thirty or more individuals
that do not feed between bouts of copulation and egg laying (Fretter and Graham 1962).
Velutina velutina (O. F. Müller, 1776) also copulated between egg laying bouts but, in
contrast, fed throughout the spawning period.
Members of the genus Velutina are simultaneous hermaphrodites (Ankel 1936;
Fretter 1984). Here, copulation was primarily unilateral with individuals behaving as
male or female in roughly equal proportions but occasionally individuals behaved as male
and female simultaneously. Diehl (1956) also observed mutual copulations between
individual V. velutina in the laboratory. For slow moving or sparsely distributed species it
is advantageous to be hermaphroditic such that any encounter with a mature conspecific
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 20 of 42
20
may lead to fertilization (Tomlinson 1966; Ghiselin 1969). This may also be important
for species that occur in low abundance, like most velutinid species. Velutina velutina (O.
F. Müller, 1776) also exhibited polygamy, as reported for other gastropods (see Murray
1964; Martel et al. 1986; Johannesson et al. 2016).
The present study suggests that V. velutina can store viable spermatozoa for up to
6 months until a suitable ascidian host for egg laying becomes available. Eupleura
caudata (Say, 1822) and Urosalpinx cinerea (Say, 1822) may store spermatozoa for 6 to
9 months (Hargis and MacKenzie 1961), and Littorina saxatilis (Olivi, 1792) for at least a year (Johannesson et al. 2016). Sperm storage may allow selection of favourable time and location to deposit egg capsules (Martel et al. 1986). Polygamy and long-term sperm storage likely evolved in species withDraft limited mobility, like V. velutina, to prevent
impoverishment of their gene pools (Murray 1964).
The egg laying period for V. velutina occurred over five months (early November
to late March), starting as water temperature declined and ending as the lowest annual
temperature was reached. In Norway, V. velutina was observed to deposit egg capsules
between January and March (E. Svensen, personal communication, 2018), whereas
Mileikovsky (1960) suggested that reproduction of V. velutina in the Barents, Norwegian,
and White Seas occurred over one month or less, soon after the lowest annual water
temperatures. However, Mileikovsky’s (1960) report was based on accounts of early
larvae of V. velutina in the plankton, rather than direct observation, as in this study. In the
northeast Pacific, Marsenina rhombica (Dall, 1871) and V. plicatilis (O. F. Müller, 1776)
also spawns in winter, from January to March (McCloskey 1973; Strathmann 1992),
https://mc06.manuscriptcentral.com/cjz-pubs Page 21 of 42 Canadian Journal of Zoology
21
while in northern Scotland Lamellaria perspicua (Linnaeus, 1758) spawns from February
to May (Jeffreys 1867).
Velutina velutina (O. F. Müller, 1776) preferred to deposit egg capsules in the
solitary ascidian Ascidia callosa, consistent with previous reports that members of the
genus Velutina spawn primarily in solitary ascidians. Reports from Europe found V.
velutina deposited egg capsules in Styela coriacea in the Baltic Sea (Diehl 1956) and in
Ascidia virginea Müller, 1776 in the Norwegian Sea (Moen and Svensen 2004). In the
northeast Pacific, Strathmann (1992) reported V. plicatilis deposited egg capsules in
Ascidia paratropa (Huntsman, 1912) and Chelyosoma productum Stimpson, 1864.
Locally, V. velutina may prefer to deposit egg capsules in A. callosa because its thick
tunic and bioaccumulation of vanadiumDraft (Stacey 2009) may deter predation (Stoecker
1980; Young 1986). Velutina velutina (O. F. Müller, 1776) laid up to 18 egg capsules
annually and spawning aggregations led to deposition in an individual A. callosa of 50 or
more egg capsules in the laboratory and more than 30 capsules in the field. Similarly,
Dias and Delboni (2008) reported that Lamellaria mopsicolor Ev. Marcus, 1958 may
deposit up to 20 egg capsules annually, and up to 53 egg capsules per ascidian host were
collected in the field.
Members of the superfamily Cypraeoidea are reported to prepare the initial point
of egg capsule deposition by creating a hollow in the tunic of an ascidian with their
radula (Ankel 1936; Diehl 1956). Here however, V. velutina used the ventral pedal gland
to forcibly embed egg capsules within the tunic with no evidence that the radula was
used. The ventral pedal gland is very muscular with a protrusible central region that can
be used as a ramrod to push the egg capsule into position (Fretter and Graham 1994). The
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 22 of 42
22 relatively soft tunics of A. callosa and A. glabrum may allow V. velutina to use its pedal gland alone to deposit egg capsules. Diehl (1956) reported that after V. velutina partially imbedded a capsule in the tunic of Styela coriacea, the tunic grew around it, leaving only the cap exposed on the surface, as was observed here. Diehl (1956) suggested this reaction was either caused by the capsules themselves, as seemed to be the case here, or due to a teasing motion of the velutinid’s radula, which was not observed. However, during this study, egg capsules were typically fully embedded by the parent V. velutina.
Feeding and spawning periods of V. velutina overlapped during this study, and feeding peaked as spawning activity declined. In contrast, Diehl (1956) suggested that V. velutina did not feed during its spawning period, to avoid consuming ascidians in which it deposits egg capsules. However, V.Draft velutina preyed very little on A. callosa in this study, thus avoiding destruction of its developing larvae or future spawning grounds.
Egg capsules of V. velutina contained between 400 – 3400 larvae in this study.
This is similar to Lamellaria perspicua, which reportedly contain at least 1000, but usually well over 3000 larvae per capsule (Ankel 1935). In contrast, egg capsules of L. mopsicolor contained only 13 – 164 larvae (Dias and Delboni 2008). There was no evidence in this study that V. velutina use nurse eggs to nourish developing larvae. This concurs with Diehl (1956), who reported that larvae of V. velutina are nourished by a yolk reservoir in the hepatopancreas.
Only planktonic larval development of V. velutina has been previously described
(Lebour 1935; Thorson 1946; Diehl 1956; Fretter and Pilkington 1970; Thiriot-Quiévreux
1974). This is the first study to document the complete development of V. velutina from egg capsule deposition to larval hatching. Egg capsules of V. velutina took 2 – 4 months
https://mc06.manuscriptcentral.com/cjz-pubs Page 23 of 42 Canadian Journal of Zoology
23
to hatch, depending on when they were laid. Capsules deposited at the start of the
spawning period were subject to warmer water temperatures (average 4.9 ºC) and their
larvae developed more rapidly than those in capsules deposited later in the period when
water temperatures were colder (average 2.1 ºC) and near the annual minimum (-1.9 ºC).
In the northeast Pacific, Strathmann (1992) found that larvae of V. plicatilis hatched in 36
d at 10 – 13 ºC, while in Brazil, Dias and Delboni (2008) found that larvae of Lamellaria
mopsicolor began hatching after 6 d at 23 – 26 ºC. Water temperature is a well-known
driver of larval development times (Scheltema 1967; Hoegh-Guldberg and Pearse 1995).
Egg capsules hatched over roughly seven months during the coldest months of the
year (January – April) until water temperatures reached the annual maximum (around
mid-July). Northern European studiesDraft have not reported hatching times for V. velutina but
several recorded the occurrence of their larvae in the plankton. Near Plymouth, UK,
Fretter and Shale (1973) collected larvae of V. velutina as early as April, while Lebour
(1935) reported them from May through the summer. Mileikovsky (1960) found Stage I
larvae of V. velutina as early as mid-March in the Norwegian Sea, and suggested larvae
entered the plankton in April in the Barents Sea, and in late May in the White Sea, where
they persisted until early July.
Adult V. velutina exhibited seasonal growth that peaked in summer, as water
temperature increased to its maximum, and stalled the rest of the year, similar to other
gastropod species (Largen 1967; Ota and Tokeshi 2000). Smaller individuals that were
likely immature exhibited growth throughout the year, as has also been reported for other
gastropods (Ota and Tokeshi 2000; Ishida 2004).
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 24 of 42
24
The consistent succession of behaviours exhibited by V. velutina during this study primarily followed annual changes in seawater temperature. Feeding dominated during the coldest temperatures of the year and declined as temperature increased to its maximum in summer. At the end of the feeding period, velutinids exhibited growth, as water temperature reached its maximum, before transitioning to mating. Ishda (2004) reported that the gastropod Drupella margariticola (Broderip, 1833) annually reallocate energy from growth to reproduction. Here, behaviour alternated between feeding and mating before eventually transitioning to mating in June and July (2007 and 2008). This alternation seemed partly related to food supply in the laboratory. As prey became depleted, velutinids switched to mating but when prey supply was replenished feeding resumed. In August, despite availableDraft prey, velutinids entered mating season throughout warmer months. Egg laying started as water temperature decreased in late autumn, and ended when temperature reached the lowest annual level in the spring. Although water temperature may influence the timing of spawning, multiple factors (e.g. photoperiod, physiological changes, prey availability, water temperature) likely work in combination or succession to fine tune this behaviour in V. velutina, as in other marine taxa (Giese
1959).
In this study, V. velutina was more active and exposed during the egg laying period but tended to be concealed and inactive the remainder of the year, possibly driven by predator avoidance. From November to March, potential predators may be less active
[e.g. cunner, Tautogolabrus adspersus (Walbaum, 1792); Green and Farwell 1971], absent from shallow waters (<15 m) [e.g. juvenile Atlantic cod, Gadus morhua Linnaeus,
1758; Hanson 1996; Cote et al. 2004], or engaged in reproductive activities [e.g. polar sea
https://mc06.manuscriptcentral.com/cjz-pubs Page 25 of 42 Canadian Journal of Zoology
25
star, Leptasterias (Hexasterias) polaris (Müller and Troschel, 1842); Himmelman et al.
1982; Hamel and Mercier 1995].
To date, little attention has focused on the role of indigenous predators in limiting
the success of non-indigenous species (Harley et al. 2013) and studies that examined
potential predators of non-indigenous ascidians have focused on generalist rather than
specialist predators (e.g. Osman and Whitlatch 2004; Simoncini and Miller 2007;
Epelbaum et al. 2009). The dynamics of specialist predators are tightly linked to one or a
few prey species and may play a role in structuring communities by controlling their prey
species densities (Snyder and Ives 2001). Members of the gastropod superfamily
Cypraeoidea have been previously reported to prey on invasive ascidian species.
Gitternberger (2007) reported LamellariaDraft sp. (likely L. perspicua) and Trivia arctica
(Pulteney, 1799) preyed on the colonial species, Didemnum sp. (later identified in
Lambert 2009 as D. vexillum Kott, 2002) in the Netherlands. Here, V. velutina preyed on
the non-indigenous C. intestinalis, but showed limited predation on the colonial species,
B. schlosseri. However, due to the relatively low abundance and low feeding rates of V.
velutina in this study and the rapid colonization rates and high abundance reported for C.
intestinalis (Carver et al. 2003; Ramsay et al. 2008), it may take time for V. velutina to
exhibit biotic resistance against populations of C. intestinalis. Therefore, predator-prey
interactions between V. velutina and C. intestinalis should continue to be monitored.
Acknowledgements
For invaluable assistance in the field we thank S. Bettles, R. Boland, W. Coffey,
J. Flight, C. Lewis, K. Matheson, D. Mouland, R. Murphy, A. Noseworthy, A. Storch, R.
O’Donnell, and B. Wringe. We thank S. Baillon, G. Doyle, K. Gale, N. Laite, J. So, and
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 26 of 42
26
Z. Sun for assistance in laboratory maintenance and monitoring. We thank J. Stacey for allowing us to examine Ascidia callosa specimens for velutinid egg capsules and settled larvae; E. Svensen for providing observations of Velutina velutina feeding and spawning in Norway; E. Geissinger and E. Pedersen for providing advice on statistical analyses; and V. Ramírez for her suggestions and support. We also appreciated the constructive comments and suggestions of two anonymous reviewers.
References
Ankel, W.E. 1935. Das Gelege von Lamellaria perspicua L. Z. Morphol. Öekol. Tiere, 30: 635-647.
Ankel, W.E. 1936. Prosobranchia. In Die Tierwelt der Nord-und Ostsee. IX b. Edited by E. Grimpe and E.
Wägler Akademische Verlagsgesellschaft, Leipzig. pp. 1-240. Behrens, D.W. 1980. The Lamellariidae of Draftthe North Eastern Pacific. Veliger, 22(4): 323-339. Behrens, D.W. 1984. Lamellariids: masters of disguise. Opisthobranch, 16(4): 42-44.
Callahan, A.G., Deibel, D., McKenzie, C.H., Hall, J.R., and Rise, M.L. 2010. Survey of harbours in
Newfoundland for indigenous and non-indigenous ascidians and an analysis of their cytochrome c
oxidase I gene sequences. Aquat. Invasions, 5(1): 31-39. doi: 10.3391/ai.2010.5.1.5.
Carver, C., Chisholm, A., and Mallet, A. 2003. Strategies to mitigate the impact of Ciona intestinalis (L.)
biofouling on shellfish production. J. Shellfish Res. 22(3): 621-632.
Cote, D., Moulton, S., Frampton, P.C.B., Scruton, D.A., and McKinley, R.S. 2004. Habitat use and early
winter movements by juvenile Atlantic cod in a coastal area of Newfoundland. J. Fish Biol. 64(3):
665-679. doi:10.1111/j.1095-8649.2004.00331.x.
Derjugin, K. 1950. New data on systematic, morphology and biogeography of the genus Velutina Flem.
(Mollusca, Gastropoda, Lamellariidae). Explor. Far East Seas USSR, 2: 7-27.
Dias, G.M., and Delboni, C.G.M. 2008. Colour polymorphism and oviposition habits of Lamellaria
mopsicolor. Mar. Biodivers. Rec. 1: 1-4. doi:10.1017/S1755267206005550.
Diehl, V.M. 1956. Die Raubschnecke Velutina velutina als Feind und Bruteinmieter der Ascidie Styela
coriacea. Kieler Meeresforsch, 12: 180-185.
https://mc06.manuscriptcentral.com/cjz-pubs Page 27 of 42 Canadian Journal of Zoology
27
Epelbaum, A., Pearce, C., Barker, D., Paulson, A., and Therriault, T. 2009. Susceptibility of non-
indigenous ascidian species in British Columbia (Canada) to invertebrate predation. Mar. Biol.
156(6): 1311-1320. doi:10.1007/s00227-009-1172-7.
Feare, C.J. 1970. The reproductive cycle of the dog whelk (Nucella lapillus). Proc. Malacol. Soc. Lond.
39(2-3): 125-137. doi:10.1093/oxfordjournals.mollus.a065087.
Fretter, V. 1984. Prosobranchs. In The mollusca, Vol. 7 Reproduction. Edited by A.S. Tompa, N.H.
Verdonk, and J.A.M. van den Biggelaar. Academic Press, New York. pp 1-45.
Fretter, V., and Graham, A. 1962. British prosobranch molluscs. Their functional anatomy and ecology.
Ray Society, London.
Fretter, V., and Graham, A. 1981. The Prosobranch Molluscs of Britain and Denmark. Part 6–Cerithiacea,
Strombacea, Hipponiacea, Calyptraeacea, Lamellariacea, Cypraeacea, Naticacea, Tonnacea,
Heteropoda. J. Mollus. Stud. (Suppl.) 9: 285-362. Fretter, V., and Graham, A. 1994. British prosobranchDraft molluscs, their functional anatomy and ecology. The Ray Society, London.
Fretter, V., and Pilkington, M.C. 1970. Prosobranchia. Veliger larvae of Taenioglossa and Stenoglossa.
Conseil International pour L’Exploration de la Mer Zooplankton Sheets, 129-132: 1-26.
Fretter, V., and Shale, D. 1973. Seasonal changes in population density and vertical distribution of
prosobranch veligers in offshore plankton at Plymouth. J. Mar. Biol. Assoc. UK, 53(3): 471-492.
doi:10.1017/S0025315400058719.
Ghiselin, M.T. 1964. Morphological and behavioral concealing adaptations of Lamellaria stearnsii, a
marine prosobranch gastropod. Veliger, 6(3): 123-124.
Ghiselin, M.T. 1969. The evolution of hermaphroditism among animals. Q. Rev. Biol. 44(2): 189-208.
doi:10.1086/406066.
Giese, A.C. 1959. Comparative physiology: annual reproductive cycles of marine invertebrates. Annu. Rev.
Physiol. 21: 547-576. doi:10.1146/annurev.ph.21.030159.002555.
Green, J.M., and Farwell, M. 1971. Winter habits of the cunner, Tautogolabrus adspersus (Walbaum
1792), in Newfoundland. Can. J. Zool. 49(12): 1497-1499. doi:10.1139/z71-218.
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 28 of 42
28
Gulbin, V. 2005. Prosobranch family Velutinidae (Gastropoda) in cold and temperate waters of the
northern hemisphere: history, biogeography, evolution and chorology. Ocean Sci. J. 40(1): 45-54.
Gulbin, V., and Golikov, A. 1997. A review of the prosobranch family Velutinidae in cold and temperate
waters of the Northern Hemisphere. I. Capulacmaeinae. Ophelia, 47(1): 43-54.
Gulbin, V., and Golikov, A. 1999. A review of the prosobranch family Velutinidae in cold and temperate
waters of the Northern Hemisphere. III. Velutininae. Genera Ciliatovelutina and Velutina.
Ophelia, 51(3): 223-238.
Hamel, J-F., and Mercier, A. 1995. Prespawning behavior, spawning, and development of the brooding
starfish Leptasterias polaris. Biol. Bull. 188(1): 32-45.
Hanson, J.M. 1996. Seasonal distribution of juvenile Atlantic cod in the southern Gulf of St. Lawrence. J.
Fish Biol. 49(6): 1138-1152. doi:10.1111/j.1095-8649.1996.tb01784.x.
Hargis, W.J., and MacKenzie, C.I. 1961. Sexual behavior of the oyster drills: Eupleura caudata and Urosalpinx cinerea. Nautilus, 75(1):Draft 7-16. Harley, C.D., Anderson, K.M., Lebreton, C.A-M., MacKay, A., Ayala-Díaz, M., Chong, S.L., et al. 2013.
The introduction of Littorina littorea to British Columbia, Canada: potential impacts and the
importance of biotic resistance by native predators. Mar. Biol. 160(7): 1529-1541.
doi:10.1007/s00227-013-2206-8.
Himmelman, J.H., Lavergne, Y., Cardinal, A., Martel, G., and Jalbert, P. 1982. Brooding behaviour of the
northern sea star Leptasterias polaris. Mar. Biol. 68(3): 235-240. doi:10.1007/BF00409590.
Hoegh-Guldberg, O., and Pearse, J.S. 1995. Temperature, food availability, and the development of marine
invertebrate larvae. Am. Zool. 35(4): 415-425. doi:10.1093/icb/35.4.415.
Hooper, R. 1975. Bonne Bay marine resources: An ecological and biological assessment. Parks Canada
Atlantic Region Office.
Ishida, S. 2004. Life history of the muricid gastropod, Cronia margariticola (Broderip, 1833): growth
mode transition with season and sexual maturity. Benthos Res. 59(1): 35-44.
doi:10.5179/benthos1996.59.1_35.
https://mc06.manuscriptcentral.com/cjz-pubs Page 29 of 42 Canadian Journal of Zoology
29
Jeffreys, J.G. 1867. British conchology, or an account of the Mollusca which now inhabit the British Isles
and the surrounding seas. Volume IV. Marine Shells, in continuation of the Gastropoda as far as
the Bulla family. J. Van Voorst, London.
Johannesson, K., Saltin, S.H., Charrier, G., Ring, A-K., Kvarnemo, C., André, C., and Panova, M. 2016.
Non-random paternity of offspring in a highly promiscuous marine snail suggests postcopulatory
sexual selection. Behav. Ecol. Sociobiol. 70(8): 1357-1366. doi:10.1007/s00265-016-2143-x.
Lambert, C.C., and Lambert, G. 2003. Persistence and differential distribution of nonindigenous ascidians
in harbors of the Southern California Bight. Mar. Ecol. Prog. Ser. 259: 145-161.
doi:10.3354/meps259145.
Lambert, G. 1980. Predation by the prosobranch mollusk Lamellaria diegoensis on Cystodytes lobatus, a
colonial ascidian. Veliger, 22(4): 340-344.
Lambert, G. 2005. Ecology and natural history of the protochordates. Can. J. Zool. 83(1): 34-50. doi:10.1139/z04-156. Draft Lambert, G. 2007. Invasive sea squirts: a growing global problem. J. Exp. Mar. Biol. Ecol. 342(1): 3-4.
doi:10.1016/j.jembe.2006.10.009.
Lambert, G. 2009. Adventures of a sea squirt sleuth: unraveling the identity of Didemnum vexillum, a
global ascidian invader. Aquat. Invasions, 4(1): 5-28. doi:10.3391/ai.2009.4.1.2.
Largen, M.J. 1967. The influence of water temperature upon the life of the dog-whelk Thais lapillus
(Gastropoda: Prosobranchia). J. Anim. Ecol. 36(1): 207-214. doi:10.2307/3022.
LeBlanc, N., Davidson, J., Tremblay, R., McNiven, M., and Landry, T. 2007. The effect of anti-fouling
treatments for the clubbed tunicate on the blue mussel, Mytilus edulis. Aquaculture, 264(1-4): 205-
213. doi:10.1016/j.aquaculture.2006.12.027.
Lebour, M.V. 1935. The echinospira larvae (Mollusca) of Plymouth. Proc. Zool. Soc. London, 35: 163-174.
Ma, K.C.K., Deibel, D., Law, K.K.M., Aoki, M., McKenzie, C.H., and Palomares, M.L.D. 2017. Richness
and zoogeography of ascidians (Tunicata: Ascidiacea) in eastern Canada. Can. J. Zool. 95: 51-59.
doi:10.1139/cjz-2016-0087.
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 30 of 42
30
Martel, A., Larrivée, D.H., and Himmelman, J.H. 1986. Behaviour and timing of copulation and egg-laying
in the neogastropod Buccinum undatum L. J. Exp. Mar. Biol. Ecol. 96(1): 27-42.
doi:10.1016/0022-0981(86)90011-0.
McCloskey, L. 1973. Development and ecological aspects of the echinospira shell of Lamellaria rhombica
Dall (Prosobranchia; Mesogastropoda). Ophelia, 10(1-2): 155-168.
Mileikovsky, S. 1960. Reproduction and larval development in Velutina velutina (Gastropoda
Prosobranchia, Lamellariidae) in the White, Barents, and Norwegian Seas. Proc. Murmansk Mar.
Biol. Institute AN. SSSR. 2: 162-171.
Moen, F.E., and Svensen, E. 2004. Marine fish and invertebrates of northern Europe. AquaPress, Essex,
UK.
Murray, J. 1964. Multiple matings and effective population size in Cepea nemoralis. Evolution, 18(2): 283-
291. doi:10.1111/j.1558-5646.1964.tb01601.x. Ota, N., and Tokeshi, M. 2000. A comparitiveDraft study of feeding and growth in two coexisting species of carnivorous gastropods. Mar. Biol. 136(1): 101-114. doi:10.1007/s002270050013.
Osman, R.W., and Whitlatch, R.B. 2004. The control of the development of a marine benthic community
by predation on recruits. J. Exp. Mar. Biol. Ecol. 311(1): 117-145.
doi:10.1016/j.jembe.2004.05.001.
Page, L. 2002. Larval and metamorphic development of the foregut and proboscis in the caenogastropod
Marsenina (Lamellaria) stearnsii. J. Morphol. 252(2): 202-217. doi:10.1002/jmor.1099.
Queiroz, V., and Sales, L. 2016. A new color pattern for the ascidian-symbiontic Lamellaria mopsicolor
(Mollusca: Caenogastropoda) in northeastern Brazil, with a discussion of its symbiotic lifestyle.
Pan-Am. J. Aquat. Sci. 11(2): 123-129.
R Core Team. 2018. R: a language and environment for statistical computing. R Foundation for Statistical
Computing, Vienna, Austria. Available from https://www.R-project.org/ [accessed 1 May 2019].
Ramsay, A., Davidson, J., Landry, T., and Arsenault, G. 2008. Process of invasiveness among exotic
tunicates in Prince Edward Island, Canada. Biol. Invasions, 10(8): 1311-1316.
doi:10.1007/s10530-007-9205-y.
https://mc06.manuscriptcentral.com/cjz-pubs Page 31 of 42 Canadian Journal of Zoology
31
Rocha, R.M., Kremer, L.P., Baptista, M.S., and Metri, R. 2009. Bivalve cultures provide habitat for exotic
tunicates in southern Brazil. Aquat. Invasions, 4(1): 195-205. doi:10.3391/ai.2009.4.1.20.
Sargent, P.S., Wells, T., Matheson, K., McKenzie, C.H., and Deibel, D. 2013. First record of vase tunicate,
Ciona intestinalis (Linnaeus, 1767) in coastal Newfoundland waters. Bioinvasions Rec. 2(2): 89-
98. doi:10.3391/bir.2013.2.2.01.
Sarma, A., and Pattanaik, P. 1986. Lamellaridae (Gastropoda) off Visakhapatnam, east coast of India.
Indian J. Mar. Sci. 15(2): 123-124.
Scheltema, R.S. 1967. The relationship of temperature to the larval development of Nassarius obsoletus
(Gastropoda). Biol. Bull. 132(2): 253-265.
Simoncini, M., and Miller, R.J. 2007. Feeding preference of Strongylocentrotus droebachiensis
(Echinoidea) for a dominant native ascidian, Aplidium glabrum, relative to the invasive ascidian
Botrylloides violaceus. J. Exp. Mar. Biol. Ecol. 342(1): 93-98. doi:10.1016/j.jembe.2006.10.019. Snyder, W.E., and Ives, A.R. 2001. GeneralistDraft predators disrupt biological control by a specialist parasitoid. Ecology, 82(3): 705-716. doi:10.1890/0012-9658(2001)082[0705:GPDBCB]2.0.CO;2.
Stacey, J.E. 2009. The ecophysiology of iron and vanadium accumulation by North Atlantic ascidians. PhD
thesis, Department of Biology, Memorial University, St. John's, Newfoundland and Labrador.
Stoecker, D. 1980. Chemical defenses of ascidians against predators. Ecology, 61(6): 1327-1334.
doi:10.2307/1939041.
Strathmann, M.F. 1992. Reproduction and development of marine invertebrates of the northern Pacific
coast: data and methods for the study of eggs, embryos, and larvae. University of Washington
Press, Seattle.
Thiriot-Quiévreux, C. 1974. Anatomie interne de véligères planctoniques de Prosobranches
Mésogastropodes au stade proche de la métamorphose. Thalassia Jugoslavica, 10(1-2): 379-399.
Thorson, G. 1946. Reproduction and larval development of Danish marine bottom invertebrates, with
special reference to the planktonic larvae in the Sound (Øresund). Med. Kom. Dan. Fisk.
Havunders Plankton, 4: 1−523.
Tomlinson, J. 1966. The advantages of hermaphroditism and parthenogenesis. J. Theor. Biol. 11(1): 54-58.
doi:10.1016/0022-5193(66)90038-5.
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 32 of 42
32
United States Navy. 1951. Report on marine borers and fouling organisms in 56 important harbors and
tabular summaries of marine borer data from 160 widespread locations. Bureau of Yards and
Docks, Dept. Navy, Washington, DC.
Valentine, P.C., Collie, J.S., Reid, R.N., Asch, R.G., Guida, V.G., Blackwood, D.S., Whitlatch, R., and
Bullard, S. 2007. The occurrence of the colonial ascidian Didemnum sp. on Georges Bank gravel
habitat-ecological observations and potential effects on groundfish and scallop fisheries. J. Exp.
Mar. Biol. Ecol. 342(1): 179-181. doi:10.1016/j.jembe.2006.10.038.
Young, C. 1985. Abundance patterns of subtidal solitary ascidians in the San Juan Islands, Washington, as
influenced by food preferences of the predatory snail Fusitriton oregonensis. Mar. Biol. 84(3):
309-321. doi:10.1007/BF00392501.
Young, C. 1986. Defenses and refuges: alternative mechanisms of coexistence between a predatory
gastropod and its ascidian prey. Mar. Biol. 91(4): 513-522. doi:10.1007/BF00392603. Yund, P.O., Collins, C., and Johnson, S.L. 2015.Draft Evidence of a native northwest Atlantic COI haplotype clade in the cryptogenic colonial ascidian Botryllus schlosseri. Biol. Bull. 228(3): 201-216. doi:
10.1086/BBLv228n3p201.
https://mc06.manuscriptcentral.com/cjz-pubs Page 33 of 42 Canadian Journal of Zoology
33
Fig. 1. Velvet shell (Velutina velutina) feeding on Halocynthia pyriformis. A – multiple
V. velutina attached near the base of H. pyriformis; B – V. velutina with proboscis
inserted through tunic of H. pyriformis; C – close-up of V. velutina proboscis inserted
through tunic of H. pyriformis; D – feeding hole of V. velutina near the base of H.
pyriformis; E - dissected intact H. pyriformis; F – dissected H. pyriformis after preyed
upon by V. velutina. Scale bars: A = 17 mm; B = 9.5 mm; C = 1.5 mm; D = 15 mm; E =
15 mm; F = 14 mm.
Fig. 2. Relative feeding positions of velvet shell (Velutina velutina) on common
indigenous ascidian species of Newfoundland, Halocynthia pyriformis, Boltenia
echinata, Ascidia callosa, and MolgulaDraft citrina. Shaded areas indicate observed feeding
and unshaded areas indicate drilling. BP = base papillae (on H. pyriformis only); B =
base; NB = near base; S = side; NT = near top; T = top; NS = near siphons; TS = through
a siphon.
Fig. 3. Average monthly growth rates (mm mo-1) ± Standard Error of velvet shell
(Velutina velutina) and daily tank water temperature data for 2008 (upper panel) and
2009 (lower panel).
Fig. 4. Velvet shell (Velutina velutina) copulation, embedding egg capsules, and hatching
larvae. A – copulation; B – egg capsules being embedded in Ascidia callosa; C – egg
capsules recently embedded in the tunic of an A. callosa; D – egg capsule with larvae
settled to bottom; E – larvae hatching as A. callosa simultaneously releases spawn; F –
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 34 of 42
34 empty egg capsule in A. callosa tunic. Scale bars: A = 16 mm; B = 14 mm; C = 3 mm; D
= 3 mm; E = 23 mm; F = 3 mm.
Fig. 5. Velvet shell (Velutina velutina) larval development. A – Newly deposited eggs; B
– 4-cell stage embryos; C – transition stage between late embryos and early larvae; D – early veliger larvae; E – early veliger with echinospira clearly visible; F – retracted veliger showing its well-developed echinospira; G – frontal view of well-developed veligers swimming within an egg capsule; H – lateral view of a well-developed echinospira veliger; I – just settled larvae (arrows) on tunic surface of Ascidia callosa, insert showing one larva (length 920 µm). Scale bars: A = 360 µm; B = 380 µm; C = 200 µm; D = 295 µm; E = 150 µm; F = 230Draft µm; G = 700 µm; H = 800 µm.
Fig. 6. Laboratory and field behavioural activity patterns of velvet shell (Velutina velutina) and corresponding laboratory water temperatures in 2007 (upper panel) and
2008 (lower panel). Note: Numbers adjacent to symbols for field observations indicate the number of individuals observed exhibiting the behaviour at that time.
https://mc06.manuscriptcentral.com/cjz-pubs Page 35 of 42 Canadian Journal of Zoology
1
Table 1. Summary of feeding experiments conducted on velvet shell (Velutina velutina) in the laboratory.
Total Number Experiment Sample Frequency of Prey Species Experimental Set-Up Duration of Outcome Type Sizes Observations Observations Each ascidian species had roughly equal surface area (2800 – 3200 Solitary species: mm2) tank-1; Ascidians placed Ascidia callosa, 3 tanks, Multi-species haphazardly on 21 × 28 cm Twice daily Number of Boltenia echinata, 3 V. velutina prey choice with perforated plastic canvas sheets 7.5 d velutinid-1 predation Halocynthia pyriformis; tank-1 135 indigenous and attached with monofilament (May 2006) (0800-1000, events on Colonial species: (9 velutinids ascidian species line (1 canvas sheet tank-1); 2000-2100) each species Aplidium glabrum, total) Flow through ambient seawater; Didemnum albidum Ascidians preyed on were not replaced during the experiment B. schlosseri (4 colonies); H. pyriformis (2 smallDraft individuals); Single Non-indigenous species: M. citrina (2 individuals); and non-indigenousa Botryllus schlosseri; A. glabrum (5 colonies) placed Number of 98 d Once every prey species Indigenous species: simultaneously in 1 tank; Static 1 tank, predation (February to 1-7 d 194 versus multiple Halocynthia pyriformis, conditions maintained near 5 V. velutina events on June 2007) velutinid-1 indigenous prey Molgula citrina, ambient water temperature; each species species Aplidium glabrum Deceased ascidians were removed but not replaced during the experiment Non-indigenous species: C. intestinalis (1 individual) tank-1 Pairwise Ciona intestinalis; paired with an indigenous ascidian 5 tanks, Number of non-indigenousa Indigenous species: (1 individual); Static conditions 1 V. velutina 44 d Once every predation versus Ascidia callosa, maintained near ambient water tank-1 (May to 1-4 d 105 events on indigenous prey Boltenia echinata, temperature; When predation (5 velutinids June 2017) velutinid-1 each species species Halocynthia pyriformis, confirmed, both ascidians replaced total) Molgula citrina to start a new feeding trial a – Both golden star tunicate (Botryllus schlosseri Pallas, 1766) and vase tunicate (Ciona intestinalis Linnaeus, 1767) (formerly C. intestinalis type B) are considered cryptogenic in northeastern North America (Yund et al. 2015; Ma et al. 2017); however, they are likely non-indigenous to Newfoundland waters given limited reports of these species in the area (United States Navy 1951; Hooper 1975) until recently (Callahan et al. 2010; Sargent et al. 2013; Ma et al. 2017). Therefore, the interaction of V. velutina with these two non-indigenous ascidian species in Newfoundland was examined.
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 36 of 42
2
Table 2. Combined laboratory and field observations of copulation, egg capsule laying and hatching for velvet shell (Velutina velutina) from 2006 to 2009.
First Egg Final Egg First Egg Final Egg Years First Copulation Final Copulation Capsule Laid Capsule Laid Capsule Hatch Capsule Hatch
2006 – 2007 Jun. 24, 2006 Jan. 14, 2007 Nov. 4, 2006 Mar. 25, 2007 Jan. 23, 2007 N/Aa
2007 – 2008 May 26, 2007b Jan. 21, 2008 Oct. 28, 2007 Apr. 5, 2008 Jan. 6, 2008 Jun. 29, 2008
2008 – 2009 Jun. 23, 2008 Dec. 14, 2008 Nov. 6, 2008 Mar. 22, 2009 Jan. 4, 2009 Jul. 13, 2009
a – All Ascidia callosa died before egg capsules could all hatch. Some capsules were still intact on May 21, 2007. b – Lack of food in tank at this time may have induced early copulation.
Draft
https://mc06.manuscriptcentral.com/cjz-pubs Page 37 of 42 Canadian Journal of Zoology
Draft
Fig. 1. Velvet shell (Velutina velutina) feeding on Halocynthia pyriformis. A – multiple V. velutina attached near the base of H. pyriformis; B – V. velutina with proboscis inserted through tunic of H. pyriformis; C – close-up of V. velutina proboscis inserted through tunic of H. pyriformis; D – feeding hole of V. velutina near the base of H. pyriformis; E - dissected intact H. pyriformis; F – dissected H. pyriformis after preyed upon by V. velutina. Scale bars: A = 17 mm; B = 9.5 mm; C = 1.5 mm; D = 15 mm; E = 15 mm; F = 14 mm.
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 38 of 42
70 Halocynthia pyriformis Boltenia echinata 60 n = 575 n = 39
50
40
30
20
10 Draft
0 BP B NB S NT T NS TS B NB S NT T NS TS 70 Ascidia callosa Molgula citrina
60 n = 28 n = 19
Percentage of Observations of Percentage 50
40
30
20
10
0 B NB S NT T NS TS B NB S NT T NS TS
Relative Feeding Position
https://mc06.manuscriptcentral.com/cjz-pubs Page 39 of 42 Canadian Journal of Zoology
Draft
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 40 of 42
Draft
Fig. 4. Velvet shell (Velutina velutina) copulation, embedding egg capsules, and hatching larvae. A – copulation; B – egg capsules being embedded in Ascidia callosa; C – egg capsules recently embedded in the tunic of an A. callosa; D – egg capsule with larvae settled to bottom; E – larvae hatching as A. callosa simultaneously releases spawn; F – empty egg capsule in A. callosa tunic. Scale bars: A = 16 mm; B = 14 mm; C = 3 mm; D = 3 mm; E = 23 mm; F = 3 mm.
https://mc06.manuscriptcentral.com/cjz-pubs Page 41 of 42 Canadian Journal of Zoology
Draft
Fig. 5. Velvet shell (Velutina velutina) larval development. A – Newly deposited eggs; B – 4-cell stage embryos; C – transition stage between late embryos and early larvae; D – early veliger larvae; E – early veliger with echinospira clearly visible; F – retracted veliger showing its well-developed echinospira; G – frontal view of well-developed veligers swimming within an egg capsule; H – lateral view of a well-developed echinospira veliger; I – just settled larvae (arrows) on tunic surface of Ascidia callosa, insert showing one larva (length 920 µm). Scale bars: A = 360 µm; B = 380 µm; C = 200 µm; D = 295 µm; E = 150 µm; F = 230 µm; G = 700 µm; H = 800 µm.
https://mc06.manuscriptcentral.com/cjz-pubs Canadian Journal of Zoology Page 42 of 42
12 10 8 6 4 2 0 -2 2MM2 Temperature (°C) 1E 2007 3 E 100
80
60
40
20
0
Proportion of Individuals (%) Jan Feb Mar Apr May Draft Jun Jul Aug Sep Oct Nov Dec Jan 12 10 8 6 4 2008/2009 2 0
-2 Temperature (°C) F Field Feeding 2F 1F M Field Mating 2008 6MMM 4 2 2M E Field Egg Laying 2 E 2 E 2 E 100 Feeding Mating 80 Egg Laying
60
40
20
0
Proportion of Individuals (%) Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec Jan Date
https://mc06.manuscriptcentral.com/cjz-pubs