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Larvae from deep-sea methane seeps
disperse in surface waters
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Shawn M. Arellano1,†, Ahna L. Van Gaest1, Shannon B. Johnson2,
Robert C. Vrijenhoek2 and Craig M. Young1
1
2
Research
Cite this article: Arellano SM, Van Gaest AL,
Johnson SB, Vrijenhoek RC, Young CM. 2014
Larvae from deep-sea methane seeps disperse
in surface waters. Proc. R. Soc. B 281:
20133276.
http://dx.doi.org/10.1098/rspb.2013.3276
Received: 16 December 2013
Accepted: 9 April 2014
Subject Areas:
ecology
Keywords:
“Bathymodiolus” childressi, Bathynerita
naticoidea, vertical migration, cold seep,
dispersal
Author for correspondence:
Shawn M. Arellano
e-mail: [email protected]
†
Present address: Shannon Point Marine Center,
Western Washington University, 1900 Shannon
Point Road, Anacortes, WA 98221, USA.
Oregon Institute of Marine Biology, University of Oregon, PO Box 5389, Charleston, OR 97420, USA
Monterey Bay Aquarium Research Institute, 7700 Sandholdt Road, Moss Landing, CA 95039, USA
Many species endemic to deep-sea methane seeps have broad geographical
distributions, suggesting that they produce larvae with at least episodic longdistance dispersal. Cold-seep communities on both sides of the Atlantic share
species or species complexes, yet larval dispersal across the Atlantic is expected
to take prohibitively long at adult depths. Here, we provide direct evidence that
the long-lived larvae of two cold-seep molluscs migrate hundreds of metres
above the ocean floor, allowing them to take advantage of faster surface currents
that may facilitate long-distance dispersal. We collected larvae of the ubiquitous
seep mussel “Bathymodiolus” childressi and an associated gastropod, Bathynerita
naticoidea, using remote-control plankton nets towed in the euphotic zone of
the Gulf of Mexico. The timing of collections suggested that the larvae might disperse in the water column for more than a year, where they feed and grow to
more than triple their original sizes. Ontogenetic vertical migration during a
long larval life suggests teleplanic dispersal, a plausible explanation for the
amphi-Atlantic distribution of “B.” mauritanicus and the broad western Atlantic
distribution of B. naticoidea. These are the first empirical data to demonstrate a
biological mechanism that might explain the genetic similarities between eastern
and western Atlantic seep fauna.
1. Introduction
Species endemic to deep-sea methane seeps can be broadly distributed, despite
their reliance on chemosynthetic primary productivity. Cold-seep sites on the
east side of the Atlantic share nearly 12% of their megafauna with seeps on
the west sides of the Atlantic [1]. These include mussels in the subfamily Bathymodiolinae, which always bear chemoautotrophic symbionts and, therefore,
are found exclusively at cold seeps, hydrothermal vents, sunken wood or
whale falls. Members of the “Bathymodiolus” childressi species complex (genus
uncertain) form extensive beds at cold seeps that range in depth from 500 to
more than 2000 m throughout the Gulf of Mexico [2]. Recent morphological
and genetic analyses reveal that one member of this complex, “B.” mauritanicus
occurs at cold seeps distributed along the Atlantic Equatorial Belt from the
Barbados accretionary prism across to the Nigerian margin seeps [1,3–5].
This broad amphi-Atlantic distribution suggests at least episodic connectivity
via dispersive larvae, yet dispersal in slow deep-sea currents is expected to take
too long for larvae to cross this ocean [6]. Connections among the Atlantic Equatorial Belt seeps would be facilitated by ontogenetic vertical migration because
dispersal distances are expected to increase in the faster currents of surface
waters compared with the deep-sea [6,7]. However, initial hypotheses suggested
that the larvae of bathymodiolin mussels probably do not migrate to surface
waters, because migration would increase advection of larvae away from the
required chemosynthetic habitats [8–10]. Moreover, in situ collections of hydrothermal vent larvae have suggested that transport takes place mostly at depth
[11,12]. While indirect evidence from morphology, isotopic analysis and lipid
profiles suggests that some vent and seep larvae might undergo vertical
migrations (reviewed in [13,14]), direct evidence for ontogenetic vertical
migration by organisms that rely on chemoautotrophy is scarce [15].
& 2014 The Author(s) Published by the Royal Society. All rights reserved.
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(a) Plankton sampling
We sampled plankton throughout the water column above the
Brine Pool NR1 cold seep (approx. 650 m depth), located approximately 180 km south of New Orleans, LA, in the Gulf of Mexico
(278430 2400 N, 918160 3000 W). We sampled nine times, in March
2002, December 2002, February 2003 and November 2003, using
a Multiple Opening and Closing Net Environmental Sampling
System (MOCNESS) towed through 50–100 m intervals. Each
150-mm net was towed at the maximum depth of its sampling
interval for 10 min then pulled obliquely through 50- or 100-m
at about 20–25 m min21 before it was closed and the next net
was opened. Larval samples were either preserved in 95% ethanol
or fixed overnight in 10% seawater-buffered formalin then stored
in 70% ethanol. Each larva was photographed under 10–20
magnification and tentatively identified based on morphology
(shell length, shape and colour) [20]. Larvae were then processed
for identification to species based on either gene sequencing or
scanning electron micrograph examination of shell morphology.
(b) Molecular identifications
Individual formalin-fixed larvae were extracted [21], amplified and
sequenced for approximately 300 bp of the cytochrome-c-oxidase
subunit I (COI) locus with the methods of Hoos et al. [22]. These
methods included two rounds of PCR; the primers COIG/H
were used in the first round [5]. The second round of PCR included
1 ml of product from the first round of PCR and the primers InternalF: 50 -AGA GTT CAT CCA GTC CCA-30 and InternalR: 50 -TGC
TAT GCC AGT TTT AGC TGC-30 designed by P. Hoos. Individual
ethanol-preserved larvae were extracted, amplified and sequenced
for COI as in Johnson et al. [23]. Adults of “B.” childressi collected
from the Brine Pool cold seep were sequenced for COI for comparison. Sequences were then compared against the blast database
(http://blast.ncbi.nlm.nih.gov/) to confirm the identities. For “B.”
childressi, Kimura-2-parameter distances were also calculated with
MEGA (v. 6.01) [24] among other known bathymodiolin mussel
species from the Gulf of Mexico and the Atlantic Ocean. For one
(c) Scanning electron micrograph identifications
Scanning electron micrographs (SEMs) of larval shells were taken
on a JEOL 6400F field emission scanning electron microscope.
Shells were cleaned in 5% sodium hypochlorite solution, rinsed
with distilled water, air-dried and mounted on adhesive carbon
discs for SEM [27,28]. Procedures to accurately document the
shapes and dimensions of the larval bivalve shells were modified
from those in Fuller et al. [29]. We also used larval tube traps
placed at the Brine Pool cold seep to collect late-stage larvae of
both species for comparison with plankton samples (figure 1).
Briefly, larval tube traps were 30-cm tall PVC pipes (5-cm diameter opening, aspect ratio ¼ 6 : 1) that were mounted on 2 kg
iron discs, filled with 10% buffered formalin [30] and placed at
the Brine Pool approximately 250 days [31]. For larval bivalves,
we measured height and length of the prodissoconch II, shell
length, straight hinge length of the prodissoconch I (if possible),
provinculum length and number of teeth and compared them to
those of “B.” childressi in Arellano & Young [13]. Length is the
greatest dimension approximately parallel to the provinculum
and height is the greatest dimension starting from and perpendicular to the hinge line. For neritid larval shells, we compared shell
shape, shell length and aperture shape of MOCNESS-collected
larvae with those collected at the Brine Pool.
(d) Laboratory cultures
Both species were cultured in the laboratory. Adults of both
species and egg capsules of B. naticoidea were collected using
the Johnson-Sea-Link I and II submersibles (Harbor Branch
Oceanographic Institution) from the Brine Pool cold seep.
Adults of both species were maintained at approximately 78C
in the laboratory at the Oregon Institute of Marine Biology.
Complete culturing procedures and results for “B.” childressi
are detailed in [13,32]. B. naticoidea began to lay egg capsules on
“B.” childressi mussel shells in the laboratory from winter 2003
to 2005. Egg capsules were separated from each other immediately
after deposition and placed in 2-ml wells filled with cold (78C)
0.45 mm-filtered seawater (FSW) until hatching. The water was
changed once a week until hatching from May to July. Once
hatched, veligers from each capsule were placed into either 175-ml
glass dishes with FSW or combined with many capsules that
hatched on the same day into 2-l glass jars, with 10 mg l21 chloramphenicol. Veligers were fed a mixture of Thalassiosira
pseudonana and Isochrysis galbana at concentrations of 5000 –
10 000 cells ml21, and water was changed every other day [33].
(e) Temperature tolerances of larvae
Thermal tolerances of B. naticoidea larvae were tested by exposing
them to a range of temperatures found throughout the water
column using an aluminium thermal gradient block [34].
B. naticoidea veligers (20 days post-hatching) in three replicate
20-ml scintillation vials (1 larva ml21) of cold (7 – 88C) FSW
were place into five temperature treatments (15, 25, 29, 32 and
358C). Per cent survival was scored after 72 h. Because all treatments except one resulted in either 100% survival or 0%
survival, data were not analysed statistically. Thermal tolerances
for trochophore larvae of “B.” childressi have been previously
published in Arellano & Young [32]. Those data were checked
2
Proc. R. Soc. B 281: 20133276
2. Material and methods
bivalve larval morphotype, we were unable to successfully
sequence the COI amplicon; instead, we successfully sequenced
approximately 550 bp of the 18S ribosomal RNA amplicon with
the DNA extraction methods of Johnson & Geller [25] and the primers from Giribet et al. [26] and compared it to the blast database
for identification at the family level. Sequences were deposited in
GenBank under accession numbers: KF739294–KF739297,
KJ576847, KJ576848 and KJ585667.
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Within the Gulf of Mexico, there is minimal genetic differentiation among populations of “B.” childressi [16]. Similarly,
the amphi-Atlantic species, “B.” mauritanicus, which is distinct
from “B.” childressi (approx. 5% sequence divergence for mitochondrial COI), exhibits minimal (less than or equal to 0.42%)
genetic distances between eastern and western Atlantic populations [5]. These genetic findings support a hypothesis of
infrequent or historical connectivity of “B.” mauritanicus via
widespread larval dispersal [1,4]. The neritid gastropod
Bathynerita naticoidea is usually found in association with bathymodiolin mussels in the Gulf of Mexico and is also known
from the southern Barbados Prism at depths from 400 to
1700 m, where it is the most abundant snail in mussel beds
[17]. Bathynerita-like neritids are found in Miocene cold-seep
deposits from Italy [18] and a Middle Eocene deposit in western Washington, USA [19], suggesting that widespread
dispersal has been a life-history feature of this clade for
millions to tens-of-millions of years.
To determine whether cold-seep larvae migrate from the
deep to the sea surface, we towed a remote-control plankton
net sampling system through discrete depth horizons above
cold seeps in the Gulf of Mexico in multiple years. We collected larvae of two molluscs, the mussel “B.” childressi and
the snail B. naticoidea, which are both endemic to cold seeps
and widespread throughout the Gulf of Mexico.
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(b)
(c)
(d )
3
(e)
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(a)
Proc. R. Soc. B 281: 20133276
(f)
(g)
(i)
(k)
(h)
(l)
( j)
Figure 1. Field-collected larvae and juveniles of Bathynerita naticoidea (a –h) and “Bathymodiolus” childressi (i – l ). (a) Newly hatched veliger larva of B. naticoidea
from laboratory culture. (b) Veliger larva collected 10 November 2003, 650– 700 m depth. (c) Aperture (ventral) view of veliger larva collected 11 February 2003,
0–100 m depth. (d) Dorsal view of the larva depicted in (c). (e) Recently settled juvenile collected from the sea floor at the Brine Pool cold seep. Arrow marks the
transition between the larval shell (protoconch) and the juvenile shell (scale bar for (a–e): 500 mm). ( f ) SEM, ventral view, of a larva collected from the plankton 11
February 2011, 0–100 m depth, showing the larval operculum occluding the large aperture (scale bar, 200 mm). (g) SEM of the shell apex of a larva collected in
February from the upper 200 m of the water column (scale bar, 100 mm). (h) SEM of the shell apex of a larva collected in a larval tube trap on the sea floor at the Brine
Pool cold seep (scale bar, 100 mm). (i) Early D-shell larval stage of “B.” childressi cultured in the laboratory. ( j) Bathymodiolin veliger larva collected from the plankton
in November 2003, 300–350 m depth. (k) “B.” childressi veliger collected 11 February 2003, 300–400 m depth (scale bar for (i–k): 200 mm). (l ) Newly settled
juveniles of “B.” childressi captured on settlement plates on the bottom at the Brine Pool cold seep [31]. Darker portions of the shell are the prodissoconchs
(larval shells) and lighter portions are the dissoconchs, representing juvenile growth after settlement (scale bar, 1 mm).
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depth
collection date
ID method
number
length (mm)
accession no.
Bathynerita naticoidea
0 – 100
11 Feb. 2003
SEM
11
389.6– 667.8
—
300– 400
500– 550
11 Feb. 2003
15 Nov. 2003
light microscopy
light microscopy
1
1
418
402.6
—
—
650– 700
10 Nov. 2003
COI, 16S
1
676.5
KF739294, KJ576848
0 – 100
200– 300
11 Feb. 2003
11 Feb. 2003
SEM
COI
3
2
417.8– 437.8
430, 400
—
KF739295, KF739297
300– 400
500– 550
11 Feb. 2003
8 Mar. 2002
COI, light microscopy
light microscopy
2
1
449.0, 459.0
500
KJ576847
—
0 – 100
11 Feb. 2003
light microscopy
1
350
—
200– 250
300– 350
15 Dec. 2002
15 Nov. 2003
light microscopy
18S
1
1
324.3
274.8
—
KJ585667
“Bathymodiolus” childressi
Bathymodiolinae veligers
Table 2. Larval shell dimensions measured by SEM for “Bathymodiolus” childressi veligers found in plankton tows taken from 0 to 100 m depth on 11 February
2003 (table 1). PI and PII are the prodissoconchs I and II. Hinge is the length of the hinge line. Means and standard deviations (in italics; n ¼ 5) of shell
dimensions given for “B.” childressi are from recent settlers collected from the Brine Pool cold seep [13].
PI
PII
provinculum
hinge
length
length
height
length
no. of teeth
“Bathymodiolus” childressi
89.41
1.94
113.35
2.02
442.56
8.84
391.92
7.39
210.15
10.94
29 – 31
0
veliger 1
88.82
115.48
437.76
382.63
191.85
31
veliger 2
veliger 3
—
70.77
—
87.82
425.81
417.8
378.17
344.06
177.26
163.39
31
29
for normality and heteroscedasticity, arcsine transformed, then
analysed with a one-way analysis of variance followed by a
two-sided Dunnett’s test against the control (78C) [32].
3. Results and discussion
Eleven B. naticoidea and three “B.” childressi veligers were collected in the top 100 m of the Gulf of Mexico above the Brine
Pool cold seep in February 2003 (table 1 and figure 1). Additional
veligers of each species were collected at greater depths in
November and February. Larval shells of mytilid mussels and
neritid snails were easily identified due to their distinctive
shell shapes (figure 1). We further narrowed our search for
“B.” childressi and B. naticoidea based on colour and size: “B.”
childressi and other bathymodiolin larvae are a distinctive pink
colour [13,31], and B. naticoidea veligers are smaller than the
coastal and estuarine neritid larvae that may be present in the
Gulf of Mexico. Identifications based on morphology viewed
under light microscopy were further corroborated with either
analysis of the larval shell using SEM (table 2 for “B.” childressi)
or sequencing (table 3 for “B.” childressi). COI and 16S mtRNA
sequences of our B. naticoidea larva (table 1) were 99% similar
to B. naticoidea isolate GM.1 (EU732361, EU732198).
A similar multistep approach to larval identification has
been used to identify species of mytilid mussel post-settlers
[20] and hydrothermal vent larvae [35]. Table 1 includes individuals identified based on morphology as viewed under light
microscopy only if we could confirm the identity of individuals
with similar morphotypes via SEM or sequencing. Thus, we consider our estimates of the numbers of larvae of these two species
that we captured in the plankton to be conservative; many of the
individuals we tentatively identified as “B.” childressi or B. naticoidea based on morphology were not included in table 1
because methodological limitations prevented some identities
from being confirmed. For one bivalve morphotype (figure 1
and table 1), we were unable to successfully sequence the COI
amplicon; however, sequence comparison of the 18S rRNA
amplicon showed 100% similarity to other members of the Bathymodiolinae in the GenBank database. In addition to “B.”
childressi, at least four species of bathymodiolin mussels are
known from the Gulf of Mexico seeps. While Idas macdonaldi
and Tamu fisheri can be found on the upper Louisiana slope,
Bathymodiolus heckerae and B. brooksi are generally found at the
deeper Gulf of Mexico seep sites (e.g. Alaminos Canyon,
Atwater Valley, Florida Escarpment) from approximately
2000–3000 m [3]. Besides some estimated larval size ranges
[13], there are virtually no data available on the larval development or reproductive timing of these other bathymodiolin
species. Nevertheless, while we cannot confirm that those bathymodiolin larvae that we identified with 18S rRNA sequencing
are “B.” childressi, the timing of their collection from November
Proc. R. Soc. B 281: 20133276
species
4
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Table 1. Cold-seep mollusc larvae collected in MOCNESS plankton tows. All larvae were preliminarily identified by examining morphological characters under
light microscopy. Some identities were confirmed with SEM or gene sequencing as noted. Lengths are of the prodissoconch II or protoconch II; ‘n.d.’ indicates
no data collected due to specimen damage.
3.67
“B.” aff. childressi
9.13
13.73
14.51
B. tangaroa (AY608439)
B. brooksi (HF545110)
Gigantidas horikoshii
16.00
16.05
16.38
B. heckerae (DQ513441)
B. boomerang (DQ513449)
Idas macdonaldi
17.68
20.88
Idas sp. (FJ937190)
Tamu fisheri (HF545104)
(AY649804)
16.00
B. azoricus (FJ766924)
(HF545113)
5.59
B. platifrons (AB250695)
(AY649801, EU288164)
“B.” mauritanicus
3.67
0.00
“B.” childressi (EU288173)
(DQ513438)
0.00
“B.” childressi Brine Pool
larvae (table 1)
20.88
17.68
16.38
16.05
16.00
16.00
14.51
13.73
9.13
5.59
3.67
3.67
0.00
Brine Pool
20.88
17.68
16.38
16.05
16.00
16.00
14.51
13.73
9.13
5.59
3.67
3.67
B. childressi
18.44
17.04
15.79
13.24
14.32
13.21
13.42
12.11
6.09
1.80
0.00
childressi
B. aff.
18.44
17.04
15.79
13.24
14.32
13.21
13.42
12.11
6.09
1.80
B. mauritanicus
16.59
17.01
13.61
13.21
14.29
13.18
13.37
11.56
6.11
B. platifrons
17.23
14.76
16.40
13.80
14.89
11.58
12.29
11.04
B. tangaroa
14.98
16.40
15.32
10.12
11.16
8.58
14.57
B. brooksi
17.90
18.18
18.78
17.96
19.13
15.04
horikoshii
Gigantidas
16.64
16.98
16.40
5.61
7.09
B. azoricus
20.81
18.74
19.87
1.35
B. heckerae
Proc. R. Soc. B 281: 20133276
B. childressi
18.57
17.13
19.00
macdonaldi
Idas
17.59
19.30
B. boomerang
18.67
Idas sp.
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larvae
Table 3. Kimura-2-Parameter % distance matrix of “Bathymodiolus” childressi larvae collected in MOCNESS tows (from table 1) and representative mussel taxa known from Gulf of Mexico (in bold), Atlantic Ocean, Gulf of Cadiz and West
African hydrocarbon seeps and hydrothermal vent sites.
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5
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temperature (°C)
6
Nov 2003
Dec 2004
Feb 2003
Mar 2002
100
Proc. R. Soc. B 281: 20133276
depth (m)
200
300
400
500
600
Figure 2. Temperature-depth profiles of the water column above the Brine Pool NR1 (278430 2400 N, 918160 3000 W) cold seep in November 2003, December 2004,
February 2003 and March 2002. Water column profiles were taken concurrently with MOCNESS tows; December 2004 is given as a representation of the temperaturedepth profile during the December 2002 tow.
100
80
percent survival
to February (table 1) is consistent with the reproductive season
of “B.” childressi [36].
Our larval cultures of both species provided indirect evidence for vertical migration and long larval durations: neither
species could be reared long enough to reach metamorphosis
and the larvae of both species were tolerant of temperatures
found above the permanent thermocline. Egg capsules of
B. naticoidea held in the laboratory at 88C released swimming
veligers (length: x̄ + s.d. ¼ 170.6 mm + 4.9; n ¼ 28) after
approximately four months of encapsulated development.
The larvae were reared in the laboratory for up to 90 days
after release and consumed microalgae but did not undergo
metamorphosis. “B.” childressi embryos develop more than
twice as slow as those of their shallow-water mytilid relatives,
reaching D-shell veligers after 8 days [13]. As with B. naticoidea,
we were unable to rear the larvae of “B.” childressi to settlement, but by comparing settlement times to known
spawning seasons, Arellano & Young [13] predicted that “B.”
childressi larvae remain in the plankton anywhere from 2–3
months to more than 1 year before settling.
We tested the ability of our cultured larvae to survive at a
range of temperatures (7–358C) they might encounter if
migrating vertically through the water column. The ambient
temperature at the Brine Pool cold seep is 7–88C year round,
while the sea surface above the Pool ranges from 20–258C in
the winter (figure 2) to upwards of 308C in the summer [32].
High percentages of larvae survived at all temperatures,
including at those temperatures representative of the sea surface (figure 3) [32]. Tolerances of early mussel larvae to
temperature are not as broad as those of B. naticoidea larvae
(figure 3) [32], suggesting that “B.” childressi larvae must
either widen their tolerances as they grow or acclimate to the
gradual increase in water column temperature (figure 2)
during migration to the shallow depths where we found them.
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5 10 15 20 25 30 35 40 5 10 15 20 25 30 35 40 5 10 15 20 25 30 35 40 5 10 15 20 25 30 35 40
0
60
40
*
20
0
7
15
20
25
temperature (°C)
29 32 35
Figure 3. Thermal tolerances of cold-seep molluscan larvae. Open circles are
mean percent survival (+1 s.d.) for 20-day-old veligers of Bathynerita
naticoidea after 72 h exposed to 15, 25, 29, 32 and 358C (n ¼ 3). Survival
was 100% from 15 to 298C, but there were no survivors at 358C. Blackened circles
are mean per cent survival (+1 s.d.) of trochophore larvae of “Bathymodiolus”
childressi after 24 h of exposure to 7, 15, 20 and 258C (n ¼ 4) from Arellano
& Young [32]. Survival was significantly lower than the control at only 258C (indicated by *; Dunnett’s t: p ¼ 0.002) [32]. Sea surface temperatures above the Brine
Pool in the Gulf of Mexico typically reach 20–308C throughout the year (figure 2).
The size range and timing of occurrence of larvae collected
in MOCNESS plankton samples are consistent with a prediction of long larval durations for both species. Previous
histological work has shown that gametogenic cycles are
strongly periodic for both species, with extended spawning
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Acknowledgements. We thank the captains and crews of the R. V. Seward
Johnsons I & II. Technical assistance for SEM work was given by the
staff of the Center for Advanced Materials Characterization in
Oregon (CAMCOR) at the University of Oregon. Two anonymous
reviewers greatly improved this manuscript. S.M.A. collected data
on “B.” childressi and drafted the manuscript. A.V.G. collected data
on B. naticoidea. S.B.J. and S.M.A. completed molecular identifications. C.M.Y. conceived the broader project on dispersal of deepsea larvae, supervised the project and led cruises. R.C.V. provided
oversight for the molecular identifications. All authors approved
the final manuscript.
Data accessibility. Sequences for “B.” childressi and B. naticoidea veligers
were deposited in GenBank under accession numbers: KF739294–
KF739297, KJ576847, KJ576848 and KJ585667.
Funding statement. This work was supported by National Science Foundation grant nos. OCE-118733, OCE-0527139 and OCE-1030453 to
C.M.Y. S.M.A. was supported by an NSF graduate research fellowship, a Ford Foundation pre-doctoral fellowship and the MBARI
summer internship (David and Lucile Packard Foundation).
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Proc. R. Soc. B 281: 20133276
would explain the lack of genetic differentiation among
populations of “B.” childressi throughout the Gulf of Mexico
[16] and the equally widespread distribution of B. naticoidea
throughout the Gulf of Mexico and at the Barbados Accretionary Prism. Using physical oceanographic data, we have
modelled dispersal of “B.” childressi and B. naticoidea in the
upper water column and near the bottom; these models
show that maximum dispersal distance is increased by ontogenetic migration [6]. Molluscan larvae with a 1-year larval
period originating in the Gulf of Mexico have the potential
of dispersing up the entire eastern seaboard of the US in a
single generation [6], a result that explains the recent discovery of extensive beds of B. childressi mussels (identified by
Katharine Coykendall, United States Geological Survey) in
Northwest Atlantic canyons [40]. Similarly, our results with
“B.” childressi suggest a mechanism by which the larvae of
its sister-species, “B.” mauritanicus, might drift in equatorial
surface currents across the tropical Atlantic, connecting disjunct metapopulations off Barbados and West Africa and in
the Gulf of Cadiz [1,5]. Teleplanic (far-wandering) molluscan
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[41]. Our data now extend the concept of teleplanic larval
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chemosynthetic environments.
rspb.royalsocietypublishing.org
periods from October to March annually [13,36,37]. In this
study, adult B. naticoidea deposited egg capsules in the
laboratory between October 2002 and March 2003, with peak
oviposition between late December and February. Similar
cycles of oviposition were observed in the field, where capsules
were common in November and February 2003 but none were
found in July 2004 [37]. Assuming all populations of B. naticoidea in the Gulf of Mexico have a reproductive season from
October to March, and considering the lengths of larvae
obtained from MOCNESS tows in February (389–676 mm;
table 1), the larvae of B. naticoidea would be planktonic for at
least 7 to 12 months, tripling in size before settling at cold
seeps. Settlement sizes were confirmed by examining
the protoconch lengths of settlers collected in tubes traps
(x + s.d. ¼ 667.6 + 44.02 mm; n ¼ 12) and of post-settlement
juveniles collected at the Brine Pool (x + s.d. ¼ 622.4 +
10.7 mm; n ¼ 2; figure 1).
We collected “B.” childressi veligers that were nearly
settlement-size (x + s.d. ¼ 427.1 + 10.0 mm; n ¼ 3) near
the surface in February 2003 (table 1). If these collected
“B.” childressi veligers developed from eggs spawned at the
beginning of the spawning season (October 2003), they
would be 4.5 months old with calculated growth rates of
3.2 mm d21. This growth rate is comparable to those of veligers
of the related intertidal mussel Mytilus edulis developing at
68C
with high food rations (approx. 3.4 mm d21 with
10–40 cells ml21) [38]. On one hand, this may suggest that
“B.” childressi veligers migrate upwards slowly, finding
enough food below the photic zone to grow very quickly.
However, our laboratory cultures show that early larvae of
“B.” childressi grow two to four times slower than does the
intertidal mussel M. trossulus at the same temperature and
salinity [32]. Alternatively, the settlement-sized “B.” childressi
we collected between the surface and 100 m depth in February
might have been spawned during the previous season and,
thus, could have been up to 16.5 months old.
Although feeding larvae of some deep-sea taxa have been
found in shallow plankton tows [39], direct evidence of ontogenetic vertical migration has never been shown for animals
endemic to highly specialized and isolated chemosynthesisbased habitats like cold seeps, hydrothermal vents or
wood- or whale-falls. Transport in the upper water column
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