Aquaculture 318 (2011) 199–206
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Aquaculture
j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / a q u a - o n l i n e
Methods for generating triploid green sea urchin embryos: An initial step in
producing triploid adults for land-based and near-shore aquaculture
S. Anne Böttger a,c, Celeste C. Eno b,c, Charles W. Walker c,⁎
a
b
c
Department of Biology, West Chester University, 750 South Church Street, West Chester, PA 19383, United States
Genetics, 425-G Henry Mall, University of Wisconsin, Madison, WI 53705, United States
Molecular, Cellular and Biomedical Sciences, University of New Hampshire, 46 College Road, Durham, NH 03824, United States
a r t i c l e
i n f o
Article history:
Received 12 August 2010
Received in revised form 7 April 2011
Accepted 11 April 2011
Available online 21 April 2011
Keywords:
Triploidy
Strongylocentrotus droebachiensis
Sea urchin
Near-shore aquaculture
Nutritive phagocytes
Gametogenesis
a b s t r a c t
Here we provide methods for generating triploid green sea urchin embryos (Strongylocentrotus
droebachiensis). These are the first triploid sea urchins of any species that have been produced at any life
stage and represent an initial step in a larger effort to yield triploid adult green sea urchins for large-scale
land-based and near-shore commercial aquaculture ventures. Following fertilization, triploid mollusks and
fish result by preventing release of the second polar body. Adult triploids in all of these organisms are sterile
and may be larger than diploids based on accelerated growth of somatic tissues. Similar methods are
impossible for sea urchins since haploid (1n) ova are released from the ovary after completion of both meiotic
divisions. We have produced triploid green sea urchin embryos (3n = 63) by fusing two 1n ova (each n = 21)
and fertilizing the diploid products of these fusions with highly diluted spermatozoa (n = 21). Mechanical,
proteolytic and acidic treatments were employed to remove the jelly coat and vitelline membrane prior to
fusion. Results indicate that acidic removal of the extracellular structures, subsequent fusion with poly(Arg) in
1.5 ml microcentrifuge tubes and fertilization with highly diluted sperm yield 60.64 ± 1.39% triploid embryos
that reach prism pluteus stage (with spicules). Triploidy was verified at all developmental stages by counting
chromosomes. We discuss scaling up our laboratory methods for embryos to the hatchery environment and
also issues unique to the reproductive biology of sea urchins that will be vital for commercial aquaculture
ventures to consider if triploid adult green sea urchins can ultimately be produced.
© 2011 Elsevier B.V. All rights reserved.
1. Introduction
In the last fifteen years, a fishery for the green sea urchin,
Strongylocentrotus droebachiensis, has become the seventh largest
marine fishery overall in the Northeastern United States and the third
largest in Maine behind lobsters and aquacultured Atlantic salmon
(Andrew et al., 2002). Early unregulated harvesting and continued
over-fishing have drastically depleted once abundant field populations throughout the Gulf of Maine. Developing sustainable alternatives to direct harvest of the green sea urchin are vital to the future of
this commercial enterprise. It is of significant interest to commercial
aquaculture to produce adult triploid sea urchins for aquaculture in
Abbreviations: DAPI, 4', 6-diamidino-2-phenylindole; DMSO, dimethyl sulfoxide;
FSW, filtered seawater; NP, nutritive phagocyte.
⁎ Corresponding author at: Molecular, Cellular and Biomedical Sciences, Center for
Marine Biology and Marine Biomedical Research Group, University of New Hampshire,
46 College Road, Durham, NH 03824, United States. Tel.: + 1 603 862 2111; fax: + 1 603
862 3784.
E-mail address: [email protected] (C.W. Walker).
0044-8486/$ – see front matter © 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.aquaculture.2011.04.027
land-based facilities or near-shore lease sites. In land-based facilities,
the taste of urchin gonads would not be negatively influenced by the
initiation of gametogenesis (Unuma and Walker, 2009, 2010). In lease
sites, urchins would feed on naturally available food sources and
growth of somatic tissues (nutritive phagocytes, NPs) in the germinal
epithelia of gonads in such animals would not rely on the use of
expensive formulated feeds and would naturally develop taste
preferred by consumers.
Triploid bivalve mollusks and fish may have increased somatic
growth and provide year-round marketable products because they are
sterile (Allen and Downing, 1986; Barber and Mann, 1991; Cox et al.,
1996; Eversole et al., 1996; Hand et al., 1998; Ruiz-Verdugo et al., 2000;
Piferrer et al., 2009). Sterility presumably depends on the failure of
three sets of chromosomes to successfully form synaptonemal complexes prior to meiosis I (Liu et al., 2004). Another benefit of sterility in
triploids is the reduced risk of genetic cross-contamination when
triploids are released into natural populations of diploid individuals
(Purdom and Lincoln, 1973; Purdom, 1973; Gjedrem, 1976; Allen and
Stanley, 1979; Thorgaard and Gall, 1979). Triploidy can be induced
in salmon (Salmo salar) and rainbow trout (Salmo gardneri) by heat
shock (Thorgaard et al., 1981) or by treatment of newly fertilized “eggs”
One wash in FSW
–Sperm dilution: 15 drops undiluted
sperm to 10 ml FSW
–Fertilization medium: 10 ml jelly
water and 40 ml FSW with a single
layer of ova
–Fertilization: 1 ml diluted sperm into
50 ml of fertilization medium
containing eggs
–Fertilization cut-off after 10 min
Two washes in FSW
100 ml Ca-free SW containing
0.2 g cysteine and 120 mg
pronase for 30 min
10 ml eggs in 90 ml Ca-free SW,
passed 10 times through
100 ml FSW pH 5 for 27 min
pH readjusted to 7 by
addition of 1N NaOH,
wash in FSW
–Three times 100 ml (300 ml total,
100 ml/treatment) of 75 ml ddH2O
and 225 ml Ca-free SW containing
1.101 g CaCl2 and 198 μl poly(Arg)
(10 mg/ml).
–Eggs (10 ml) from each treatment
were resuspended in 90 ml of above
fusion mixture and 1.5 ml
eggs/solution distributed into 30
microcentrifuge tubes (for each
denuding treatment)
–55 min incubation
100 ml FSW containing
6.006 g urea and 1.1 mg
CaCl2 for 10 min
Wash in 100 ml FSW
Fertilization
Wash
Acidic
Eggs were denuded (by removal of both the jelly coat and vitelline
membrane) using three different procedures, mechanical, proteolytic
and acidic (Table 1). (1) For mechanical removal of extracellular
structures, ova were resuspended in filtered seawater (FSW) at
10% vol/vol. The suspension was gently passed into a glass beaker
through a nylon mesh screen (210 μm) pre-wetted with FSW (Foltz
et al., 2004). The mesh was rinsed with FSW and this process was
repeated 9 times. After ten gentle passages through the nylon mesh,
ova were allowed to settle by gravity, collected by pipetting and
Mechanical
2.3. Removal of the jelly coat and vitelline membrane
Two washes with
Ca-free SW
Spawning was induced using 2 ml of 0.5 M KCl per animal
injected through the peristomial membrane into the coelom of the
mature adults (Carpizo-Ituarte et al., 2002; Foltz et al., 2004), and
gametes were collected (dry for sperm and in calcium-free seawater
for eggs) following established protocols described by Strathmann
(1987).
Proteolytic
2.2. Spawning and gamete collection
Fusion
One hundred adult northeastern green sea urchins (Strongylocentrotus
droebachiensis) between 50 and 75 mm in diameter were collected
by SCUBA in March, 2010 from a lease site in the Piscataqua River
(Portsmouth, New Hampshire). They were transported in ambient
seawater to the Coastal Marine Laboratory of the University of New
Hampshire, maintained in a two tiered A-frame tank system at the
ambient water temperature (5–10 °C) and salinity (33 ppt) of collection
within the facilities flowing seawater system and fed ad libitum with a
commercially available formulated feed (Wenger International, Sabetha,
KS) prior to the induction of spawning. Spawning in the laboratory on
the campus of the University of New Hampshire occurred within 2 days
of collection.
Wash
2.1. Animal collection and maintenance
Denuding
2. Materials and methods
Wash
(= secondary oocytes) with cytocalasin B (Barber and Mann, 1991;
Nell et al., 1996) or with 6-dimethylaminopurine (6-DMAP) (Desrosiers
et al., 1993; Gerard et al., 1994; Nell et al., 1996; Zhang et al., 1998; Norris
and Preston, 2003; Liu et al., 2004). Following fertilization, methods
described for these organisms block second meiotic division and
result in retention of the second polar body yielding triploid embryos
that develop into sterile triploid adults.
Since methods that block release of the second polar body are not
possible for sea urchins because both meiotic divisions occur within
the ovary and prior to ovulation and fertilization, triploids have never
been produced for any species of sea urchin (Walker et al., 2005,
2006). Retrieval of primary oocytes from dissected green sea urchin
ovaries is not practical for commercial purposes because most oocytes
obtained in this way are immature and meiotically incompetent
(Wessel et al., 2004; Walker et al., 2005) and the numbers of embryos
that might result from suppressing polar body release would be
extremely low.
In this study we describe methodology for producing triploid
green sea urchin embryos (n = 63) and maintaining them to the prism
pluteus stage with spicules (Stephens, 1972). Our methods have
yielded the only viable triploid embryos ever generated from any
species of sea urchin and are a successful first step in an effort to
generate adult triploid green sea urchins. We discuss scaling up our
laboratory methods for embryos to the hatchery environment and
also address issues unique to the reproductive biology of sea urchins
that will be vital for commercial land-based and near-shore
aquaculture ventures to consider if triploid adult green sea urchins
can ultimately be produced.
Wash
S.A. Böttger et al. / Aquaculture 318 (2011) 199–206
Table 1
Methods for denuding (proteolytic, mechanical and acidic), fusion and fertilization for producing triploid Strongylocentrotus droebachiensis embryos, including all washes conducted and times of treatments. One liter Ca-free SW was produced
by addition of 27.8 g NaCl, 0.754 g KCl, 7.3 g MgCl2 6H2O, 4.31 g MgSO4 7 H2O and 1.24 g H3BO3 to 1000 ml ddH2O.
200
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S.A. Böttger et al. / Aquaculture 318 (2011) 199–206
resuspended in calcium-free seawater. (2) For proteolytic removal,
ova were treated with calcium-free seawater containing 0.2% cysteine
and 1.2 mg ml−1 pronase for 30 min (Bennett and Mazia, 1981a,b)
followed immediately by exposure to 1 M urea containing 0.1 mM
CaCl2, after which the denuded ova were resuspended in calcium-free
seawater. (3) Acidic removal was accomplished by exposing ova for
20 min to acidified FSW at pH 5. Following exposure, ova were
washed three times and resuspended in calcium-free seawater
(Bennett and Mazia, 1981a,b). In all of these treatments, the resulting
jelly water (possibly containing elements of the jelly coat and
essential to inducing the acrosomal reaction in spermatozoa) was
collected, maintained on ice and used later during fertilization (Foltz
et al., 2004).
To confirm that the jelly coat had been removed, subsamples of
denuded ova from all three treatments described above were stained
with Azure B (which stains the jelly coat) and were viewed with a
Zeiss Axioplan II MOT equipped with an AxioCam MR camera and
AxioVision 4.3 software (Carl Zeiss, Inc., Thornwood, NJ). Following
fertilization, success in removal of vitelline membrane was determined by the absence of a fertilization membrane that is derived from
an existing vitelline membrane by the addition of structural proteins
from the cortical granules of the fertilized ovum.
control diploid and triploid embryos at the gastrula stage (when
chromosome numbers were again counted) were transferred to a
local hatchery and placed in 300 l vats filled with seawater at ambient
local temperatures.
2.7. Chromosome counts
Embryos resulting from control fertilizations and after fusion and
subsequent fertilization were collected before and after gastulation
and treated according to Eno et al. (2009). To prepare chromosomes at
gastrula and pluteus developmental stages, both fused and unfused
embryos were treated for 2 h with colchicine (Sigma) at a final
concentration of 1 mg/ml to suspend cell division at metaphase,
collected by centrifugation (1000 rpm for 5 min), suspended in 1 M
urea and dissociated into their component cells by pipetting. The
dissociated cells were collected by centrifugation, treated with 8%
sodium citrate for 10 min and fixed in methanol: acetic acid (3:1). The
fixed cells were stained with DAPI to stain their chromosomes,
squashed manually on microscope slides and viewed with a Zeiss
Axioplan II MOT, an AxioCam MR camera and AxioVision 4.3 software
(Carl Zeiss, Inc., Thornwood, NJ).
2.4. Fusion of denuded ova
2.8. Statistical analysis
2.5. Fertilization of fused ova
All ova (fused and un-fused) were fertilized in new tripour beakers
(4.5 cm internal diameter) by adding 15 drops of undiluted sperm to
10 ml of sterile FSW and adding 1 ml of diluted sperm and 10 ml of
jelly-water (collected as described above during mechanical removal
of the jelly coat and vitelline membrane) to fused ova and 40 ml FSW
(Table 1). Ova were distributed in a single layer at the bottom of the
beaker after addition of spermatozoa; mixing was accomplished using
a plastic pipet. Each time fused ova were fertilized, batches of
untreated and un-fused ova were simultaneously fertilized to serve as
controls. When N95% of the control ova had produced a fertilization
membrane, resulting zygotes in all treatments were washed three
times and returned to labeled containers with FSW.
Data were expressed as mean ± SEM. Differences among values
for: (1) removal of the jelly coat and vitelline membrane, (2) fusion by
the different methods used and (3) fertilization and development of
resulting embryos were evaluated using descriptive statistics and a
One-Way ANOVA on Ranks followed by a Tukey pairwise comparison
(SigmaStat). All statistical analyses were preceded by assessments of
the assumption of normality (Kolmogorov–Smirnov Test) and homoscedacity (Spearman Rank Correlation).
(A)
% Jelly Coat and Vitelline
Membrane Removal
Two green sea urchin ova (1n) denuded as above were fused to
generate the “diploid” entities used for fertilizations. Calcium-free
seawater diluted to 75% of its original volume with distilled H2O and
CaCl2 was added to reach a working concentration of 25 mM. 6.75 μl of
poly(Arg) (120,000 MW, Sigma in 1 ml DMSO) was added to 10 ml of
the diluted seawater and denuded ova were suspended in this
medium (Table 1).
Containers tested for fusion efficiency were: (1) 1.5 ml microcentrifuge tubes, (2) 96 well microtiter plates, (3) 6 ml round-bottom
tubes and (4) 15 ml Falcon tubes. Following treatment ova were
washed with 1.5 mg/ml arginase in calcium-free seawater and
returned to sterilized FSW. After 55 min, diameters of ova containing
two pronuclei were measured with an ocular micrometer. This
procedure was repeated eight times.
successful
100
unsuccessful
damaged
50
0
Mechanical
(B)
Proteolytic
Acidic
(C)
2.6. Zygotes and development of embryos
Resulting zygotes were allowed to develop at 8 °C for 24 h until the
control embryos had reached the blastula stage when chromosome
numbers were first counted. A sub-sample of 30 blastulae was
withdrawn from all treatments and embryos were scored into four
groups depending on their developmental stage (blastula, undeveloped, delayed or abnormal development). The same embryos were
also sub-divided based on the presence or absence of the fertilization
membrane and their diameters were measured (μm) to account for
unsuccessful removal of the vitelline membrane. Subsequently,
Fig. 1. Removal of the jelly coat and vitelline membrane: A) percent success ± SEM for
removal of the jelly coat and vitelline membrane using the following three
treatments: mechanical (passing through a 210 μm mesh screen), proteolytic
(pronase and cysteine) and acidic (treatment in seawater at pH 5); B) untreated
ova with fertilization membranes; scale bar — 100 μm and C) unfertilized ova with cell
membranes touching following treatments to remove the jelly coat and vitelline
membrane. Scale bar — 100 μm.
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S.A. Böttger et al. / Aquaculture 318 (2011) 199–206
3. Results
14.24% ± 7.81 respectively), with no significant difference between
containers (p= 0.67).
3.1. Removal of the jelly coat and vitelline membrane
Mechanical removal of the vitelline membrane did not yield significant success (p = 0.255), but did produce large numbers of ova
with incomplete or no removal (42.01% ± 28.87) or damage (13.32% ±
9.99). A significant number (p b 0.001) of denuded ova (= successfully
removal of both the jelly coat and the vitelline membrane) resulted
when ova were subjected to proteolytic (97.92% ± 1.94 all results ±
SEM) or acidic treatments (73.54% ± 6.15) (Fig. 1A–C). Since we
required denuded ova to initiate fusion these results indicate that
mechanical denuding was neither as successful nor as consistent
as that resulting from proteolytic and acidic treatments. Though
denuding was most successful following proteolytic treatment,
resulting ova were not fertilizable. Acidic removal was used in this
study to prepare for fusion of ova and subsequent fertilization.
3.3. Fertilization
Ova that had been successfully denuded either by proteolytic or
acidic treatment did not develop a fertilization membrane (100% ± 0
and 78.26% ± 1.9 respectively, all results ± SEM). For example, only
21.74% ± 5.4 of ova successfully developed a fertilization membrane
following acidic treatments (Fig. 3) while the majority of fertilized
eggs did not develop a fertilization membrane. Following mechanical
treatment, only 24.14% ± 4.3 of ova did not produce a fertilization
membrane, while 75.86% ± 10.4 of ova had a fertilization membrane,
a result that was not significantly different (p = 0.058) from untreated ova where fertilization membranes were evident in 93.1% ±
7.3 of ova.
3.4. Laboratory rearing, development and size of embryos
3.2. Fusion
As determined by the presence of two pro-nuclei and by size
measurements, the highest numbers of fusing and fused ova occurred in
1.5 ml microcentrifuge tubes (41.44% ± 7.45 of ova achieved fusion and
15.81% ± 4.6 were completing fusion, all results± SEM) (Fig. 2A–C).
When compared with results from the other containers used, the
numbers of viable fused ova were significantly higher in microcentrifuge tubes (p= 0.023). For example, fusion of ova in Falcon tubes
(15 ml) resulted in 25.01% ± 6.2, fused ova, however, since these tubes
were generally filled with more ova, a large number were crushed (up
to 30.57% ± 7.65). Also, both 96-well plates and 6 ml round bottom
tubes, yielded low numbers of completed fusions (6.59% ± 5.4 and
Percentage (%)
60
(A)
fusing
3.5. Transferring laboratory methodology to a hatchery
fused
On two occasions, subsamples of control diploid and of triploid
gastrulae were transferred from the laboratory to two 300 l round
Plexiglas vats at a local sea urchin hatchery and maintained at
ambient temperature and salinity in seawater with aeration via an air
stone. In these vats, triploid gastrulae developed to the prism pluteus
stage more slowly than diploid gastrulae with a delay of approximately 30 h. In these trials, both control diploid and triploid embryos
40
20
0
In 10 separate trials subsequent to fertilization, proteolytically
denuded ova (100% ± 0, all results ± SE) did not undergo cleavage to
2-cell stage (Fig. 4A). Mechanically denuded ova resulted in a large
number of zygotes that did not cleave (34.84% ± 9.56), although a few
blastulae did result (10.32% + 1.45). Acidically denuded ova produced
a significantly higher numbers of normal blastulae lacking both the
jelly coat and vitelline membrane (p b 0.001) than any other treatment (60.64% + 1.39).
Removal of the jelly coat and vitelline membrane by mechanical
and acidic treatments yielded significantly larger (p b 0.001) blastulae
(317 μm + 12.13 and 321.2 μm + 10.54 respectively) than untreated
ova or ova that were not denuded and then treated mechanically or
with acidified seawater (199.2 μm + 1.01, 201.3 μm + 2.3 and
200.9 μm + 2.4 respectively) (Fig. 4B).
1.5 ml
microcentrifuge
tube
96 well plate
6 ml
round bottom
tube
15 ml Falcon
tube
120
% fertilization membrane
% absence
Experimental Containers
(B)
(C)
80
40
0
Fig. 2. Fusion of ova: A) fusion (% ± SEM) of ova occurred in all four containers tested:
(1.5 ml microcentrifuge tubes, 96 flat-bottom well plates, 6 ml round bottom tubes and
15 ml Falcon tubes); the highest percentage of fusion was obtained in 1.5 ml
microcentrifuge tubes. Fusion occurs as: B) 2 ova attach; scale bar — 200 μm followed
by C) partial fusion (two figure eight examples) and complete fusion (four examples,
showing larger size following fusion). Scale bar — 200 μm.
untreated
mechanical
proteolytic
acidic
Fig. 3. Fertilization of ova with two haploid nuclei: fertilization was observed in all
treatments except the proteolytic treatment. However following fertilization, untreated
ova resulted in zygotes surrounded by the fertilization membrane and acidic denuded
ova lacked a fertilization membrane.
S.A. Böttger et al. / Aquaculture 318 (2011) 199–206
120
(A)
% Development
untreated
proteolytic
mechanical
acidic
80
40
0
membrane
no
no
no
membrane
membrane
membrane
membrane
membrane
membrane
Blastula
Undeveloped
Delayed
Abnormal
(B)
Diameter in (µm)
300
200
100
0
normal blastula
unsuccessful
mechanical
unsuccessful
acidic
mechanical
acidic
Treatments
Fig. 4. Development of triploid embryos: A) for all treatments, development was
assessed at the blastula stage and results were classified as blastulae, undeveloped ova,
delayed stages (anything below blastula) and abnormal development (irregularities
and malformations during development) for embryos displaying and lacking the
fertilization membrane. Acidically treated ova resulted the highest numbers of blastulae
that lacked a fertilization membrane. B) Size (μm) was measured for untreated and for
mechanical and acidically treated blastulae. The treated embryos were divided into
those with successful and unsuccessful removal of the jelly coat and vitelline
membrane. Embryos that resulted in successful removal of the jelly coat and vitelline
membrane had significantly larger blastomeres and blastulae. All results ± SEM.
perished in these small vats soon after attaining the prism pluteus
stage.
3.6. Chromosome counts
Chromosome counts were made according to methods developed
specifically for strongylocentrotid sea urchins and described in Eno
et al. (2009). Normal green sea urchins have 42 chromosomes in their
diploid genomes (n = 21) and include two large, eight medium and
ten small pairs plus one putative sex pair (Fig. 5A). Resulting triploid
blastulae and gastrulae contained 63 chromosomes (a diploid set and
an extra maternal set of chromosomes) with representatives from all
of the documented size classes (Fig. 5B) In subsamples of 50 gastrulae,
77.42 ± 1.57% were triploid while 21.58 ± 1.16% were diploid.
4. Discussion
We discuss: Scaling up our successful laboratory methodology for
the production of triploid green sea urchin embryos to a hatchery
environment (Section 4.1) and Issues unique to the reproductive
biology of sea urchins that will be vital for commercial near-shore
aquaculture ventures to consider if triploid adult green sea urchins
can ultimately be produced (Section 4.2).
203
4.1. Scaling up our current successful laboratory methodology for the
production of triploid green sea urchin embryos in a hatchery environment
Generation of triploids in animals that are fertilized as primary
oocytes (molluscs) or as secondary oocytes arrested at metaphase of
second meiotic division (fish) is a relative straightforward process
involving the treatment of oocytes with chemical or physical shock to
prevent release of the second polar body. Generating triploids in sea
urchins cannot be accomplished using this methodology since haploid
ova are released from the gonopores and fertilized in the water
column. Previous studies have indicated that fusion of the haploid ova
of sea urchins can be accomplished by a variety of means (Wilson,
1953; Bennett and Mazia, 1981a,b; Richter et al., 1981; Sekirina et al.,
1983; Vassetzky and Sekirina, 1985; Vassetzky et al., 1986). Results of
these early studies did not yield triploids, nor have they been adapted
for use in aquaculture for any sea urchin and they are probably not
useful since we have shown that proteolytic or chemical removal of
extracellular structures results in non-fertilizable ova.
Scaling up our methods for the generation of triploid green sea
urchin embryos from the research laboratory to a commercial
hatchery environment is underway. Each step in this process is
discussed below to assist those interested in commercial application
of our methods.
4.1.1. Spawning, gamete collection and removal of the vitelline
membrane
This suite of procedures prepares 1n ova (that exist in the ovary
before ovulation, Walker et al., 2005) for fusion and subsequent
fertilization. The procedure for spawning sea urchins with 0.5 M KCl is
outlined in Foltz et al. (2004). In nature, sea urchin ova are fertilized
ecto-somatically in the water surrounding feeding adults. As a result,
both sexes produce prodigious numbers of gametes (millions) that
are more than sufficient to yield hundreds of thousands of embryos
using our methods. Removal of the jelly coat and the vitelline
membrane is necessary before fusion of ova can take place. The jelly
coat is removed mechanically during filtration. We describe three
methods for removal of the vitelline membrane. Acidic removal was
the most successful and was used in this study to prepare ova for
fusion and subsequent fertilization. It should also be possible to
remove the vitelline membrane using the Cleland reagent (Epel et al.,
1970) although we did not attempt this method in our study.
4.1.2. Fusion, fertilization and embryo culture
We determined that fusion of denuded ova occurred optimally in
1.5 ml microcentrifuge tubes (41.44% ± 7.45 of ova achieved fusion
and 15.81% ± 4.6 were completing fusion). Each of these tubes
contains approximately 1000 ova. To scale up this laboratory
procedure, 500 tubes would yield 250,000 triploid embryos. In our
hands, embryos reached the gastrula stage after 3 days in laboratory
culture (8 °C) and early prism pluteus after 6 days in the hatchery post
fertilization (fert 3/7/2010; gast 3/11/2010; prism pluteus 3/13/
2010). Diploid green sea urchin plutei begin feeding after the mouth
has formed and appropriate phytoplankton (Isochrysis galbana,
Dunaliella tertiolecta and Rhodomonas lens) must be cultured in
adequate quantities to supply feeding embryos as soon as the mouth
is formed. It is important to point out that the triploid embryos we
generated have gastrulated successfully. In sea urchins, most new
expression of zygotic genes begins at gastrulation. Development to
the gastrula stage depends upon the expression of genes or use of
proteins placed in the oocytes during oogenesis and before meiosis
(Davidson, 1987). Expression of novel, lethal combinations of genes
united during fertilization are potentially fatal at gastrulation. The fact
that triploid embryos did gastrulate successfully confirms that
“maternal stored RNA” remains and is accessible during early
development and the observation that triploid embryos transferred
to a commercial hatchery developed successfully to prism stage
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S.A. Böttger et al. / Aquaculture 318 (2011) 199–206
(A)
(B)
Fig. 5. Chromosome counts for diploid and triploid blastulae: A) 42 randomly numbered chromosomes from a normal diploid blastula at 24 h after fertilization; (B) 63 randomly
numbered chromosomes from a triploid blastula at 24 h after fertilization. The latter were produced by fusing two green sea urchin ova (each 1n) to generate the “diploid” entities
we fertilized using 1n normal sperm, scale bars — 5 μm.
suggests that the equivalence of zygotic RNA transcribed from the
triploid cell nucleus was available and translated successfully during
development.
4.1.3. Chromosome counts
This process easily differentiates triploids from embryos with an
alternative ploidy and provides a means for monitoring the success of
the fusion process prior to transfer of embryos to a hatchery
environment. The protocol is described in detail by Eno et al. (2009)
and employs a simple compound microscope at 60–100× magnification.
4.1.4. Transfer to hatchery, feeding embryos and maintenance
to metamorphosis
This portion of the scaling up procedure depends on the availability
of a hatchery. The hatchery used in this study is maintained by Dr. Larry
Harris and located in Portsmouth, New Hampshire. In our case, parallel
trials for control diploid gastrulae and for triploid gastrulae were carried
out in 300-liter vats containing filtered and UV-sterilized seawater.
Control diploid and triploid embryos were both cultured at 10 °C with
aeration provided by an air stone to create current within the vat.
Stocking density was 4 embryos/ml of medium with an ultimate final
density expected to be 300,000 to 600,000 settled juveniles per vat.
Larvae were fed a combination of Isochrysis galbana, Dunaliella tertiolecta
and Rhodomonas lens daily and overfeeding was avoided to yield achieve
minimum mortality (Kelly, 2002). Still in this initial trial, mortality
resulted for both diploid and triploid embryos following the prism
pluteus stage. We suspected that the mortality that we observed for
both diploid and triploid embryos resulted from over-aeration in these
small containers that prevented normal filter feeding by the plutei, but
we do not have evidence to back up this idea. Normally in this hatchery,
successful development of diploid embryos to metamorphosis and
settlement occurs at 21 days in containers three to five times larger than
the ones we used in these trials. Resulting diploid juveniles are then
maintained for 5 to 7 days before extraction and transfer to juvenile
culture systems at the hatchery. In future efforts to generate triploids,
the use of larger tanks, with less vigorous aerations should yield triploid
embryos that will also develop beyond the prism pluteus stage of
development.
4.1.5. Maintenance of triploid juveniles in land-based aquaculture
ventures and out-planting and maintenance of juveniles in near-shore
lease sites
Because their sterility can reduce the energy required for reproduction, triploid adult sea urchins should prove useful for both land-based
and near shore aquaculture ventures. It should be recognized that there
is anecdotal evidence that some individuals view the use of triploid
organisms negatively.
Diploid juvenile green sea urchins are typically maintained in
recirculating or flow-through culture systems at hatcheries until they
are approximately 10–20 mm in diameter (unpublished data from
Dr. Larry Harris, UNH). Juvenile urchins can then be maintained in
land-based aquaculture systems or be transferred to near-shore lease
sites until they reach harvest size at 52 mm diameter. Previous and
on-going studies show that transfer to lease sites during winter
months when predators are dormant results in higher survival and
under appropriate culture conditions, juveniles reach transfer size
within 6 months. Sea urchins remain within lease sites if adequate
food is available (Dumont et al., 2006; Lauzon-Guay et al., 2006).
Since reversion of presumably triploid organisms to reproductively
active individuals has occurred in some cases, especially for male
triploid fish and shellfish (Nell et al., 1996; Piferrer et al., 2009), follow
up studies to monitor this potential outcome for triploid sea urchins are
essential. While the behavioral issues recognized in fish that result from
matings between triploid males and diploid females would not be an
important issue for sea urchins, broadcasting gametes from triploid sea
urchins might be even more important (Kirk et al., 2004). These
precautionary studies should be carried out for both land- and near
shore-based aquaculture ventures using triploid sea urchins, since such
reversion may have genetic consequences for wild sea urchin populations (Kirk et al., 2004).
4.2. Issues unique to the reproductive biology of sea urchins that will be vital
for commercial near-shore aquaculture ventures to consider if triploid adult
green sea urchins can ultimately be produced
Gametogenesis and intra-gonadal nutrient storage and utilization
are linked processes in sea urchin reproduction (Walker et al., 2005,
2006). Initial increase in size of sea urchin gonads of both sexes results
as somatic cells within the germinal epithelium called nutritive
phagocytes (NPs), store extensive nutrient reserves prior to gametogenesis (principally the major yolk protein, MYP, Unuma et al., 2003).
Urchin gonads of both sexes that contain predominantly NPs are
preferred as a commercial product (Walker et al., 2005; Unuma and
Walker, 2009, 2010). In the green sea urchin, shorter day-lengths
characteristic of fall photoperiod are closely correlated with initiation
of gametogenesis and commercial quality of these gonads declines as
gametes of both sexes are produced (Walker and Lesser, 1998; Unuma
and Walker, 2009, 2010). Several studies of a variety of sea urchins
have addressed the possibility that photoperiod can be manipulated
at various times of the year to suspend gametogenesis in order to
emphasize the desirable size and sensory qualities present in green
S.A. Böttger et al. / Aquaculture 318 (2011) 199–206
sea urchin gonads harvested from the wild in the fall when NPs are
maximally developed and result in the best tasting commercial
product (Walker and Lesser, 1998; Walker et al., 2005, 2006; Dumont
et al., 2006; Böttger et al., 2006; Unuma and Walker, 2009, 2010).
While it is possible to delay gametogenesis and to generate large
commercially valuable gonads for the green sea urchin that contain
predominantly NPs for use in land-based aquaculture (Walker et al.,
2005; Böttger et al., 2006), methods for achieving this same result for
the near-shore aquaculture of the green sea urchins have not yet been
developed. Generation of sterile adult triploids is advantageous for
culture of green sea urchins in their natural environment where
photoperiod is ambient and where diploid green sea urchins initiate
gametogenesis in the fall.
By generating adult triploid green sea urchins, we would begin to
experimentally unlink gametogenesis and intragonadal nutrient
storage in NPs and over-emphasize the nutrient storage phase of
the green sea urchin annual gametogenic cycle. Gonads in triploid sea
urchin of both sexes should be sterile and would not produce gametes
in response to the autumn photoperiod cue (Walker and Lesser,
1998). We do not know if autumn photoperiod will or will not lead to
mobilization of nutrients from NPs in adult triploid sea urchins
maintained in the wild, resulting in reduction in the quality of the
commercial product. Experiments designed to test this idea can only
be accomplished when adult triploid green sea urchins are available.
However, it is reassuring that in nature, adult green and other sea
urchins continue to incorporate copious nutrients in NPs following the
autumn photoperiod cue and the initiation of gametogenesis (Walker
et al., 2005; Unuma et al., 2010). An additional benefit of near-shore
aquaculture of triploid adult green sea urchins is that they will
consume food in their natural environment and as a result, their
gonads should develop taste that is expected by the industry.
Acknowledgments
We thank Dr. Larry G. Harris and Chris Hill for their help with
animal acquisition and with the culture of triploid sea urchin embryos
in their hatchery in Portsmouth, New Hampshire and additionally to
Dr. Harris for comments in the discussion on hatchery culture and
transfer to lease sites. Funding for this study was provided by UNH Sea
Grant (Development Funds) and the Northeast Regional Aquaculture
Center (04-15) to CWW.
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C.W. Walker is a Professor of Molecular, Cellular and
Biomedical Sciences at the University of New Hampshire,
Durham, NH. He received a BS from Miami University,
Oxford, Ohio and an MS and PhD from Cornell University,
Ithaca, New York. His interests include molecular aspects
of gametogenesis and also aquaculture of the green sea
urchin and molecular biology of the p53 pathway in
hemocyte cancer of the soft shell clam, Mya arenaria. His
laboratory was the first to clone c-myc and p53 genes
from any invertebrate and opsin from the green sea
urchin.
S. Anne Böttger graduated in 2001 with a PhD in
Biology from the University of Alabama at Birmingham,
studying effects of anthropogenic pollutants on physiology, immunology and development of sea urchins.
She was a postdoctoral fellow at the University of New
Hampshire (2002–2008) investigating effects, etiology
and natural occurrence of cancer in softshell clams and
culture techniques of sea urchins. She is currently an
Assistant Professor at West Chester University in
Pennsylvania.
Celeste Eno graduated in 2006 with a BS in Marine and
Freshwater Biology followed by MS in Zoology in 2009
from the University of New Hampshire. Masters work
included the study of the chromosomes of two sea urchins,
Strongylocentrotus droebachiensis and S. purpuratus. She is
currently pursuing a Ph. D. in Genetics at University of
Wisconsin-Madison.