Culture of triploid greenlip abalone (Haliotis laevigata Donovan) to market size: Commercial implications

Aquaculture 271 (2007) 130 – 141 www.elsevier.com/locate/aqua-online

Culture of triploid greenlip abalone (Haliotis laevigata Donovan) to market size: Commercial implications Graeme A. Dunstan a,b,⁎, Nick G. Elliott a,b , Sharon A. Appleyard a,b , Bronwyn H. Holmes a , Natalie Conod a,1 , Mark A. Grubert c,2 , Mark A. Cozens d a

c

CSIRO Marine and Atmospheric Research, Tasmania 7001, Australia b CSIRO Food Futures National Research Flagship, Australia Tasmanian Aquaculture and Fisheries Institute, University of Tasmania, Tasmania, Australia d School of Medicine, University of Tasmania, Tasmania, Australia Received 5 February 2007; received in revised form 13 June 2007; accepted 21 June 2007

Abstract Triploid greenlip abalone (Haliotis laevigata) were produced via a 15 min exposure to 100 μM 6-dimethylaminopurine at 20 min post-fertilization and grown under commercial conditions for up to 48 months. Differential mortality of the ploidy classes resulted in a decline in the proportion of triploid abalone from 83% in trochophores (2 h post-hatch), 55% in post-larvae (29 days post-hatch) and 50% in adults. A random sample of 540 juvenile triploid and diploid abalone from the same cohort were tagged, biopsied (for ploidy determination), weighed and measured at 13 months. Measurements of the tagged abalone for growth were recorded at 17, 25, 32, 36, 39, 42, and 48 months, and sampling for meat yield performed at 36, 42 and 48 months. There were significant differences in the length and weight of diploid and triploid abalone by 13 months, with the latter being impaired by the equivalent of 23 to 25 days growth by this time. Comparisons of growth from 13 to 48 months showed that diploid abalone were consistently 11–15% longer than their triploid siblings. However, larger triploid abalone yielded up to 30% higher meat weight than diploid abalone of comparable length during the spring–summer maturation periods (at 36 and 48 months), making harvesting of triploid abalone for canned product at this time cost effective. Diploid abalone exhibited compensatory growth and had meat weights comparable to those of triploids between the maturation periods (at 42 months). The fatty acid composition of the meat from 48 month old greenlip abalone was not affected by ploidy, maturity or sex. Half of the triploid abalone were male and most of the remainder did not develop gonad. Attempts to spawn triploid abalone were unsuccessful as females did not mature and those males which did mature, failed to respond to artificial spawning stimuli. Crown Copyright © 2007 Published by Elsevier B.V. All rights reserved. Keywords: Abalone; Diploid; Fatty acids; Growth; Sterility; Triploid

⁎ Corresponding author. CSIRO Marine and Atmospheric Research, GPO Box 1538, Hobart, Tasmania 7001, Australia. Tel.: +61 3 62325274; fax: +61 3 62325000. E-mail address: [email protected] (G.A. Dunstan). 1 Present address: Tasmanian Institute of Agricultural Research, University of Tasmania, Tasmania, Australia. 2 Present address: Department of Primary Industry, Fisheries and Mines, Northern Territory, Australia.

1. Introduction Triploid animals are produced by inhibiting the release of the first or second polar body through the application of a chemical or environmental stress soon after fertilization. The retention of an extra set of chromosomes within the

0044-8486/$ - see front matter. Crown Copyright © 2007 Published by Elsevier B.V. All rights reserved. doi:10.1016/j.aquaculture.2007.06.023

G.A. Dunstan et al. / Aquaculture 271 (2007) 130–141

developing embryo results in an animal containing three sets of chromosomes per cell (triploid) instead of the usual two sets (diploid). The production of triploid animals in aquaculture is undertaken for potential growth advantages and/or to produce infertile animals (Liu et al., 2004a). Accelerated growth of triploids is most obvious during sexual maturation when diploids reduce body growth rate to divert energy and nutrients into gametogenesis, while triploids continue to invest in somatic growth (Ihssen et al., 1990). Improved growth rates in triploid molluscs have been reported in blue mussel (Mytilus edulis) (Brake et al., 2004), catarina scallop (Argopecten ventricosus) (Ruiz-Verdugo et al., 2000, 2001), Indian oyster (Crassostrea madrasensis) (Mallia et al., 2006) and other edible oyster species (Nell, 2002). Guo and Allen (1994a) suggested that triploid dwarf surf clams (Mulinia lateralis) grew larger than diploid clams not because of energy diversion from gametogenesis but through polyploidy gigantism, whereby the organism produces larger cells to accommodate the extra DNA. A similar conclusion was drawn for enlarged larvae of Haliotis discus hannai by Okumura and Yamamori (2002). However, the growth advantage of triploids over diploids animals has not always been shown to be the case. Examples of this include the Pacific oyster (Crassostrea gigas; Desrosiers et al., 1993), lion-paw scallop (Nodipecten subnodosus; Maldonado-Amparo et al., 2004), and a variety of other molluscan (Beaumont and Fairbrother, 1991) as well as finfish species (Ihssen et al., 1990). While reduced gametogenesis (Ruiz-Verdugo et al., 2000; Brake et al., 2004), an inability to spawn (Brake et al., 2004) and low fertilization rates (Nell, 2002) have been associated with triploidy, triploids may not always be 100% sterile (Maclean, 2003). Given the variable effects of triploidy, it is unwise to make broad generalizations regarding the benefits (or otherwise) of this procedure (Ihssen et al., 1990). Induction of triploidy in abalone (Haliotis discus hannai, H. diversicolor, H. diversicolor supertexta, H. midae, H. rufescens, H. asinina, and H. rubra) has been reported with varying degrees of success under laboratory or small scale conditions (e.g. Arai et al., 1986; Stepto and Cook, 1998; Yang et al., 1998a,b; Zhang et al., 1998; Maldonado et al., 2001; Norris and Preston, 2003; Liu et al., 2004a,b,c). Few studies have examined triploid abalone grown to maturity. This study investigates the effect of triploid induction on the growth, meat yield, meat fatty acid composition and reproductive ability of greenlip abalone (Haliotis laevigata) reared in a commercial system.

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2. Materials and methods 2.1. Induction, culture and ploidy sampling Spawning of greenlip abalone (H. laevigata) broodstock was induced by exposure to flowing UV-irradiated seawater. Eggs from 17 females (induced in three tanks) were mixed in a 20-l bucket with sperm from six males (induced in one tank), thereby creating the potential for 102 crosses. After 15 min contact time, the eggs were rinsed of excess sperm and prepared for the triploid induction procedure. Previous trials with H. laevigata had shown polar body I to have begun to emerge at ∼5 min post-fertilization (50% emergence by 12–15 min), and polar body II had begun to emerge at ∼20 min (50% emergence by 25–28 min) (at 19 °C ± 1 °C). The meiosis II triploids were produced by exposing a batch of the fertilized eggs to a 100 μM solution of 6-DMAP (6-dimethylaminopurine), 20 min after fertilization (Desrosiers et al., 1993; Norris and Preston, 2003; Liu et al., 2004b). Following 15 min exposure to 6-DMAP, the eggs were rinsed and placed in a hatching tray. The diploid abalone of the current study were refractory rather than naïve (i.e. untreated) diploids as they were from the same batch of fertilized eggs subjected to the 6-DMAP induction as the triploids but ploidy was not affected by the treatment. Bulk samples of trochophore larvae (2 h post-hatch) and veliger larvae (85 h post-hatch) were harvested in replicate for ploidy determination by flow cytometry. The remaining larvae were settled onto settlement plates at six days post-hatch and a bulk sample of postsettlement juveniles harvested for flow cytometry at 29 days post-hatch. The remaining juveniles were transferred to a commercial rectangular, 10,000 l concrete growout tank on an abalone farm at 6 months. The animals in the tank were maintained according to the standard growout practices on the farm. The tank was supplied with flow-through seawater at ambient temperature (10–20 °C) with aeration. The abalone were fed a commercial formulated feed and the tank cleaned three times per week. At 13 months a random sample of 540 individuals (from an estimated 20,000 abalone in total) were tagged, weighed, measured and a cephalic tentacle removed for flow cytometry. The tagged abalone were maintained with their untagged siblings in a single tank for the duration of the trial, and selected tagged abalone were re-sampled at later dates. Five tagged animals of known ploidy (three triploid and two diploid) were sacrificed at 32 months and samples of the cephalic tentacle, foot meat, digestive gland and gonad removed

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for flow cytometry. After annual removal of untagged animals from the tank to maintain total tank biomass, the final population in the tank at 48 months was estimated to be 3000 abalone.

(t1), and (t2 − t1) is the number of days between the two measurements.

2.2. Flow cytometry

With the onset of sexual maturity, tagged animals were systematically sacrificed at three sampling times (36 months n = 20, 42 months n = 77 and 48 months n = 68) to examine the effect of triploidy on meat and shell production. Length, total wet weight, wet meat and shell weights were recorded and the visual gonad index (VGI) assessed according to the following criteria: 0 — no gonad development, sex indeterminable; 1 — gonad flat, sex determinable; 2 — gonad partially distended; 3 — gonad fully distended, including conical tip (Kikuchi and Uki, 1974a; Ebert and Houk, 1984).

Samples were added to 800 μl Citrate/DMSO buffer and stored at −20 °C. The samples were macerated and rinsed through a cotton wool plugged 1 ml pipette tip into a flow cytometer tube immediately prior to analysis. The DNA was stained by adding 100–200 μl of the macerated sample to 20 μg/ml Propidium Iodide (Sigma) from a stock solution (with 50 ml Milli-Q water and 1% Triton X (Sigma) added) and left to stand for 5 min. The samples were then run through a flow cytometer (Coulter Elite ESP Cell sorter, Miami USA) and 3000–10000 events collected using a 488 nm air cooled argon laser (Uniphase San Jose CA USA) and 575 nm band pass filter (BP). Data was collected as a histogram using the red integrated linear fluorescence channel (575BP). Ploidy results were compared against a known diploid control, which was set at the 200 channel marker, with triploids at 300 and tetraploids at 400. Percentage of triploid and diploid larvae and postsettlement juveniles were estimated by comparing the relative proportion of triploid and diploid cells from the flow-cytogram of each bulk sample (Stepto and Cook, 1998; Maldonado et al., 2001; Norris and Preston, 2003; Liu et al., 2004a,b,c). While this approach is widely used and relatively simple, it only provides an estimate of the percentage ploidy, as the animals of different ploidy types may have different numbers of cells per animal (e.g. due differences in the rate of development). Percentages of older triploid and diploid abalone were calculated from the results of the individually tagged and biopsied animals. 2.3. Cohort growth Measurements of maximum shell length (digital calipers) and wet weight (electronic balance) were taken from tagged animals at 17, 25, 32, 36, 39, 42 and 48 months. The specific growth rate (SGR — which provides a growth estimate that is independent of initial size and considers exponential growth) was calculated according to the equation: SGR ¼ ðLnðm2 Þ  Lnðm1 ÞÞ=ðt2  t1 Þ where, Ln is log normal, m2 is the measurement (length or weight) at time 2 (t2), m1 is the measurement at time 1

2.4. Meat yield and shell production

2.5. Meat fatty acid composition A longitudinal section of lyophilized meat (foot muscle) was taken from 48 month old triploid (n = 12) and diploid (n = 14) abalone of both sexes and across the range of available VGI scores. A 30 mg sub sample of each homogenized meat section was transferred to a 10 ml reaction tube and 5 ml of a 10:1:1 solution of nanograde methanol: nanograde chloroform: ultrapure hydrochloric acid added. After purging with high purity nitrogen, the tubes were tightly capped (Teflon lined caps) and heated to 80 °C for 2 h, during which time they were removed and vigorously shaken every 15 min. After cooling to room temperature, 2.5 ml of Milli-Q water and 1.8 ml of 4:1 nanograde hexane:nanograde chloroform were added, the tubes shaken well and centrifuged at 2500 rpm for 5 min. The upper hexane: chloroform layer containing most of the fatty acid methyl esters (FAME) was transferred to a 1.8 ml vial and dried under high purity nitrogen. To ensure complete extraction, a further two extractions of the lower methanol layer in the reaction tube with 1.8 ml of 4:1 hexane:chloroform followed by shaking, centrifugation, transfer and drying were performed. Following quantitative addition of tricosanoic acid methyl ester (23:0) internal standard in nanograde dichloromethane, the vials were flushed with high purity nitrogen and stored at − 20 °C until analysis. The FAME were quantified with an Agilent Technologies 6890N gas chromatograph fitted with a HP-5 cross-linked (5% Phenyl)methyl-polysiloxane fused silica capillary column (50 m long, 0.32 mm i.d., 0.17 μm film thickness), a flame ionization detector (at 310 °C), a split/splitless injector (at 290 °C) and an Agilent Technologies 7683A auto injector. Helium was the carrier gas. Samples were

G.A. Dunstan et al. / Aquaculture 271 (2007) 130–141

injected in splitless mode at an oven temperature of 50 °C. After 1 min, the oven temperature was raised to 150 °C at 30 °C min− 1, then to 250 °C at 2 °C min− 1, then to 300 °C at 5 °C min− 1. This temperature was maintained for a further 15 min. Peaks were quantified with GC Chemstation Rev. A. 10.01 and identified with authentic standards. 2.6. Broodstock conditioning A selection of 50 previously untagged 36 month old male and female abalone from the mixed triploid/diploid cohort was transferred to an experimental abalone conditioning facility, tagged and biopsied for ploidy identification. The sexes were kept in separate 150 l conditioning tanks supplied with flow-through sea water at a rate of 1.5 l min− 1 and a temperature of 16 °C. The tanks were aerated and the abalone fed a commercially available broodstock conditioning diet (Adam and Amos Pty. Ltd., South Australia). Maturation was assessed (using the VGI) after 30, 42 and 222 days of conditioning, and attempts were made to spawn the animals in separate spawning trays and exposed to UV irradiated seawater that was heated from 16 °C to 21 °C over 5 h then allowed to cool to 17.5 °C (Kikuchi and Uki, 1974b; Grubert and Ritar, 2005). Two further attempts to condition and spawn the triploid greenlip abalone were made, however these were also unsuccessful (data not presented). 2.7. Statistical analyses The mean shell length and total wet weight of triploid and diploid abalone were compared using t-tests (assuming equal variances). The power regressions for wet meat weight against shell length of triploid and diploid abalone were log10-transformed and analysis of covariance (ANCOVA) performed using Statistica 7.

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Fifty five percent of the 540 abalone tagged at 13 months were triploid (triploid n = 279, diploid n = 226, unknown [aneuploid or mosaic] n = 35), and at 32 months 51% of the tagged sub-population were triploid. After this time, destructive sampling for meat yield meant that the proportion of surviving triploids could not be calculated. Re-sampling of a cephalic tentacle of triploid animals at various intervals during the trial established that triploid animals had not reverted to the diploid condition. In three triploid individuals, analyses of the tentacle, foot meat, digestive gland and gonad, confirmed that these tissues were triploid. The relationship of abalone total wet weight to shell length for all measures up to 42 months was not influenced by ploidy, as neither triploid nor diploid abalone were either relatively heavier (per length) or longer (per weight) (Fig. 1). Total wet weight increased in proportion to approximately the cube of the shell length for both ploidy types. Unfortunately, the diet was changed on the abalone farm prior to the 48 month's measurements, which resulted in reduced weights per length equally for both triploid and diploid abalone at the last measurement (data not shown in Fig. 1). However, it was clear that there were many more large diploid abalone than large triploid abalone (Fig. 1), and at every measurement the diploid abalone were on average longer than triploid abalone by 11 to 15% (Table 1). The first measurement of the tagged triploid and diploid abalone revealed that there was a significant difference (P = 0.001) in average length and weight by 13 months (Table 1). The diploid abalone were significantly larger than the triploid abalone and this was consistent for both length and weight with time (Table 1).

3. Results 3.1. Cohort growth and survival In this single mass spawning, chemical induction resulted in an estimated proportion of triploidy of 83% ± 1 (mean ± sd) in trochophore larvae (∼2 h post-hatch), 76% ± 5 in veliger larvae (∼85 h post-hatch) and 55% ± 4 in post-settlement juveniles (29 days post-hatch). Survival at settlement of the cohort was between 20 and 25%. Using this figure and ploidy percentages, the mortality rate from trochophore to settlement of triploid abalone was estimated to be 85% and of diploids to be 40%.

Fig. 1. Total wet weight (g) versus shell length (mm) of triploid (open circles) and diploid (closed circles) greenlip abalone between 13 months and 42 months (trend lines overlap and are indistinguishable).

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Table 1 Average biometric data of the tagged and dissected triploid and diploid abalone (asterisk denotes significant difference or effect) Age (months)

13

Ploidy

Diploid

Triploid

Tagged abalone (n) Shell length (mm) t-test significance Total wet weight (g) t-test significance Dissected abalone (n) Shell length (mm) Wet meat wt. (g) Wet shell wt. (g) Wet meat wt./shell wt. ratio VGI score (0–3) log10-transformed equation for wet meat wt. vs shell length r2 for wet meat wt. vs shell length ANCOVA test of significance (p value) Length effect Ploidy effect Length × ploidy interaction

227 24.7 ± 3.48 b0.001⁎ 2.0 ± 0.81 b0.001⁎ – – – – – – –

279 109 21.8 ± 3.18 69.6 ± 5.96 b0.001⁎ 1.3 ± 0.58 45.3 ± 12.2 b0.001⁎ – 10 – 68.7 ± 5.04 – 9.7 ± 2.52 – 14.4 ± 3.70 – 0.68 ± 0.06 – 2.1 ± 0.42 – y = 0.0194x− 0.3558 – 0.8058

–

36 Diploid

0.0000⁎ 0.0558 0.0288⁎

During the spring–summer maturation periods (32– 36 months and 42–48 months) the average triploid growth rate (31 and 11 μm/day respectively) exceeded that of the diploid abalone (28 and 8 μm/day respectively) but this was not significant (Fig. 2). During most other time periods, triploid growth rate equalled or was less than diploid growth rate (Fig. 2). The SGR (length) of triploid abalone typically equalled or exceeded that of diploid abalone, especially during the spring maturation periods (Fig. 3). The absolute growth rate and SGR of both triploid and diploid abalone varied with season and time to a similar degree, with the highest growth rates at 13– 17 months, and the lowest during the maturation periods (Figs. 2 and 3).

Fig. 2. Average absolute growth rates (shell length in μm/day) of triploid and diploid greenlip abalone over specific time periods in months (mo) (mean ± 1.96 SE).

42 Triploid

Diploid

107 60.2 ± 5.09

96 75.6 ± 6.37 b0.001⁎ 28.4 ± 7.42 54.4 ± 14.8 b0.001⁎ 10 42 62.3 ± 4.94 76.1 ± 6.57 8.5 ± 2.68 17.9 ± 6.03 10.2 ± 2.50 18.9 ± 5.03 0.82 ± 0.07 0.94 ± 0.11 0.6 ± 0.56 1.1 ± 0.13 y = 0.0291x− y = 0.0211x− 0.9071 0.3718 0.9539 0.9261

0.0000⁎ 0.7152 0.5350

48 Triploid

Diploid

86 66.1 ± 5.03

40 75.9 ± 5.49 b0.001⁎ 35.4 ± 8.29 46.2 ± 9.42 b0.001⁎ 35 40 67.6 ± 5.71 75.9 ± 5.49 12.4 ± 3.71 12.8 ± 3.22 12.3 ± 3.06 19.4 ± 4.19 1.00 ± 0.10 0.66 ± 0.09 0.7 ± 0.26 2.2 ± 0.32 y = 0.0219x− y = 0.0165x− 0.4079 0.1594 0.9424 0.7190

Triploid 28 68.3 ± 4.97 33.5 ± 8.07 28 68.3 ± 4.97 10.3 ± 2.87 13.6 ± 3.10 0.76 ± 0.09 0.9 ± 0.39 y = 0.0220x− 0.5037 0.8599

0.0000⁎ 0.0634 0.0355⁎

3.2. Meat yield and shell production Triploid abalone yielded more meat per animal than diploid abalone of a comparable length during both maturation periods (Fig. 4a and c). This difference was not apparent during winter (Fig. 4b). A measure of this is the power exponent of the fitted regression curve. Total wet weight increased to the cube of the length in both triploid and diploid greenlip abalone (Fig. 1) and if all three fractions of wet meat, shell and viscera were accumulating at a constant rate they too should be increasing as a function of the cube of the length. The wet meat weight of triploid abalone increased by a power of 4.0, 3.4 and 3.4 of the length at 36, 42 and 48 months, respectively (Fig. 4). However, wet meat

Fig. 3. Specific growth rates (length) of triploid and diploid greenlip abalone over specific time periods in months (mo) (mean ± 1.96 SE).

G.A. Dunstan et al. / Aquaculture 271 (2007) 130–141

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10% decreasing to 3% higher wet meat weights than diploid abalone (for 63 to 80 mm abalone). Expressing these data a different way shows that during the maturation periods a diploid abalone would need to be between 11 and 14% (at 36 months) or 6 and 11% (at 48 months) longer than a triploid abalone to yield the same wet meat weight. Diploid abalone generally had a slightly (but not significantly) higher average wet shell weight than triploid abalone at comparable shell lengths, and shell weight to shell length was not affected by season (data not shown). The higher wet meat to shell weight ratio for a given length for triploid compared to diploid abalone reflected the increased meat and to a lesser extent, lower shell weight in triploid abalone (Fig. 5). There were no significant differences in wet meat or shell weights between sexes within a ploidy type (data not shown). Diploid H. laevigata had higher average VGI scores than their triploid siblings (Table 1), with very few of the latter developing significant gonad. The average VGI score for triploid H. laevigata at first maturation period, the following winter and the second maturation period was 0.6, 0.7 and 0.9, respectively. Corresponding figures for diploid H. laevigata were 2.1, 1.1 and 2.2, respectively (Table 1). 3.3. Meat fatty acid composition

Fig. 4. Wet meat weight by shell length of triploid (open circles) and diploid (closed circles) greenlip abalone at the three sampling periods. a) Maturation period at 36 mo—enhanced meat production in triploid abalone. b) Winter at 42 mo—no difference in wet meat weight. c) Maturation period 48 mo—enhanced meat production in triploid abalone.

weight of diploid abalone increased by only a power of 3.1 and 2.8 of the length, during the two maturation periods but accelerated to a power of 3.7 of the length during the winter (Fig. 4). These seasonal trends were particularly evident when expressed as percentage differences between the regression formulas for 36, 42 and 48 months (for the length ranges for which there was data for both triploid and diploid abalone). By regression differences, triploid abalone at 36 months had 16% increasing to 30% higher wet meat weights than diploid abalone (for 62 to 70 mm abalone). Similarly, triploid abalone at 48 months had 1% increasing to 21% higher wet meat weights than diploid abalone (for 61 to 82 mm abalone). However, during the winter (at 42 months) triploid abalone had

There were no significant differences in meat fatty acid composition or concentration within or between a ploidy type within sex (Table 2). Similarly, there were no significant differences in fatty acid compositions and concentrations with varying gonad stage or shell length (data not shown). The fatty acid compositions of the abalone meat samples were typical of other abalone, with the most abundant fatty acids being 16:0, 18:1n-9, 20:5n-3, 22:2(7,13) and 22:5n-3.

Fig. 5. Average wet meat weight to wet shell weight ratio verses length for all dissection data from triploid and diploid greenlip abalone.

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Table 2 Major fatty acids (N1% of total fatty acids) in the foot muscle of triploid and diploid greenlip abalone (percentage of total fatty acids and concentration in mg g− 1 dry weight of foot muscle) and biometric parameters of the abalone analysed (mean ± SD) Ploidy

Diploid abalone

Sex

Immature

Female

Male

Immature

Female

Male

n analysed Shell length (mm) Total wet weight (g) Wet meat weight (g) Wet shell weight (g)

2 71.5 ± 12.4 37.5 ± 20.1 10.4 ± 4.8 16.8 ± 10.2

6 77.3 ± 6.6 47.9 ± 10.2 12.3 ± 3.0 20.8 ± 5.5

6 74.6 ± 5.4 43.9 ± 9.6 12.4 ± 3.1 18.5 ± 4.0

5 68.9 ± 5.5 35.9 ± 13.9 11.6 ± 5.4 14.1 ± 4.6

1 68.3 30.9 9.0 12.8

6 68.2 ± 6.7 33.5 ± 11.1 10.1 ± 2.9 13.3 ± 6.7

Percentage of total fatty acids a 14:0 16:0 18:0 16:1n-7 18:1n-9 b 18:1n-7 18:2n-6 20:4n-6 20:5n-3 22:2-7,13 c 22:5n-3 22:6n-3 sum SFA d sum brFA d sum MUFA d sum PUFA d n-3/n-6 ratio total n-3 total n-6

1.1 21.9 7.0 1.4 8.3 6.2 5.9 4.4 8.6 6.3 10.6 2.3 32.8 1.5 22.5 43.2 1.9 22.7 12.0

1.1 22.3 7.2 1.2 8.3 6.1 5.2 4.2 8.7 6.7 10.3 2.1 33.4 1.5 22.6 42.5 2.0 22.2 11.2

1.1 22.5 7.2 1.2 8.4 6.2 5.5 3.9 8.9 6.7 9.9 2.1 33.4 1.3 23.0 42.3 2.0 22.0 11.1

0.9 23.0 6.8 1.2 8.2 6.1 5.8 4.1 8.6 6.4 10.0 2.3 33.4 1.4 23.0 42.3 1.9 21.9 11.6

1.1 22.0 7.1 1.1 8.2 6.3 5.7 3.5 9.3 6.6 10.9 2.0 33.0 1.1 22.7 43.3 2.1 23.3 11.0

1.2 22.5 7.1 1.2 8.2 6.7 5.3 3.9 9.0 6.1 9.8 2.1 33.5 1.5 23.5 41.5 2.0 22.0 11.0

muscle) 0.8 1.6 1.3 1.9 0.4 1.9

0.7 1.7 1.2 1.9 0.4 1.9

0.7 1.6 1.1 1.8 0.4 1.8

0.7 1.9 1.3 2.2 0.4 2.0

0.7 1.7 1.1 1.8 0.4 1.9

Concentration of main PUFA d (mg g-1 dry weight of foot 20:4n-6 0.7 20:5n-3 1.5 22:2-7,13 c 1.1 22:5n-3 1.8 22:6n-3 0.4 Total FAME d (% dry matter) 1.7

Triploid abalone

a Minor fatty acids (less than 0.5% of total fatty acids) not listed but included in the calculations included; 15:0, 17:0, 19:0, 20:0, 22:0, 24:0, i15:0, a15:0, i16:0, i17:0, a17:0, i18:0, i19:0, a19:0, 15:1, 16:1n-9, 16:1n-5, 17:1n-8, 17:1n-4, 18:1n-5, 19:1n-10, 20:1n-9 (&20:3n-3), 20:1n-7, 20:1n-7t, 22:1n-11 (&22:2n-6), 22:1n-9 (&22:3n-3), 24:1n-11, 24:1n-9, 18:2n-9, 18:3n-6, 18:2n-3, 20:2n-9, 20:2-5,11 NMID, 20:2-5,13 NMID (&20:3n-6), 20:2n-6, 21:5n-3, 22:5n-6, 22:4n-6, 22:3n-6, 22:2–7,15 NMID. b Contains small amounts of 18:3n-3. c Non-methylene interrupted Diene. d SFA saturated fatty acids, brFA branched chain fatty acids, MUFA monounsaturated fatty acids, PUFA polyunsaturated fatty acids, FAME fatty acid methyl esters.

3.4. Broodstock conditioning Of the three attempts to condition adult abalone from the cohort, only the data from the first attempt is presented. Diploid male abalone generally achieved a higher average VGI score than triploid males after 30 days of conditioning (Table 3), with both groups losing condition by 42 days. No male triploid abalone exceeded VGI stage 2 by 42 days, but six male triploid

abalone reached VGI stage 3 by 222 days; suggesting that triploid males could be conditioned up to the level required for spawning. However attempts to induce spawning failed. Diploid females also achieved a higher average VGI score than triploid females after 30 days of conditioning (Table 3). The average VGI score of diploid females then dropped at successive inspections (i.e. 42 and 222 days). This loss of condition was most probably due

G.A. Dunstan et al. / Aquaculture 271 (2007) 130–141 Table 3 Frequency of occurrence of Visual Gonad Index (VGI) scores relative to sex, ploidy and conditioning time Sex

Male

Ploidy (n)

Diploid (12)

Triploid (27)

Female

Diploid (14)

Triploid (22)

Conditioning time (days) 30 42 222 30 42 222 30 42 222 30 42 222

VGI score 0

1

2

3

– 17 – 33 69 15 – 21 27 91 86 95

8 42 50 52 27 19 21 29 73 9 14 5

25 33 50 15 4 44 21 36 – – – –

67 8 – – – 22 57 14 – – – –

to an unknown number of females spontaneously spawning in the (monosex) tank at 42 days. The resulting eggs were assayed by flow cytometry and shown to be from diploid abalone. No triploid female exceeded VGI stage 1, with most remaining barely identifiable as female. 4. Discussion 4.1. Cohort growth and survival — implications for production Meiosis II triploids were induced for the present study because induction of meiosis I triploids relative to meiosis II triploids in abalone has been reported to result in low induction rates and high mortalities (Zhang et al., 1998; Liu et al., 2004c). Triploid induction using 6-DMAP in abalone has been shown to be up to 100% in H. rubra larvae (Liu et al., 2004a) and up to 56% in H. discus hannai (Zhang et al., 1998) and can vary significantly between different species of abalone (Zhang et al., 1998; Norris and Preston, 2003; Liu et al., 2004b). In the current study, chemical shock at polar body II resulted in approximately 83% triploidy in H. laevigata larvae. This number had decreased to 55% by settlement due to the higher mortality of triploid abalone (85%) relative to diploid abalone (40%) prior to settlement. Similarly, mortality in 6-DMAP induced triploid H. rubra was higher (70%) than for diploid abalone (35%) for day 7 larvae (Liu et al., 2004a). These authors suggested that most mortality had occurred in larvae by day 2, but do note that high mortalities are often experienced during the second week after larval settlement. Similarly, larval or juvenile mortality is often higher in triploid than diploid animals (e.g. Ihssen et al., 1990; Desrosiers et al., 1993; Stepto and Cook, 1998; Zhang et al., 1998; Ruiz-Verdugo

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et al., 2000; Maldonado et al., 2001; Liu et al., 2004a,b,c; Maldonado-Amparo et al., 2004; Mallia et al., 2006), possibly due to the triploid induction method used (Mallia et al., 2006). There is very little data on growth rates of triploid compared to diploid abalone, and that which is available has only been measured for juvenile abalone less than an average 5 mm in length. Four month old triploid H. discus hannai induced with 6-DMAP were larger than the naïve control diploid abalone (Zhang et al., 1998). In contrast, 5 month old triploid H. rubra induced with Cytochalasin B were smaller than refractory diploid abalone (Liu et al., 2004a). The first measurement of the tagged triploid and refractory diploid H. laevigata cohort revealed that there was a significant difference in length and weight by 13 months. It is not known if the initial differential growth was due to survival of only smaller triploid abalone, or that triploidy induction impaired early growth. Using the formula for SGR (length), we estimated that triploid induction had impaired growth within the first 13 months of life by 23 to 25 days compared to the diploid abalone. This lag is an issue for commercial production, especially when selling whole live, frozen and/or by weight, as after 4 years the diploid abalone were on average between 11 and 15% larger than their triploid siblings. This occurred even though the specific growth rate of the triploid abalone typically exceeded that of the diploid abalone after the 13 month measurement. While growth rates of these triploid compared to diploid gastropod molluscs differed between studies, triploid bivalve molluscs generally grow faster than diploids (e.g. Nell, 2002). Research on the European flat oyster (Ostrea edulis) has shown that blocking meiosis I resulted in significantly faster growth rates than triploid meiosis II oysters or control diploids and this was attributed to higher heterozygosity and reduced energy expenditure (Hawkins et al., 1994). Similarly, Mallia et al. (2006) have shown that meiosis I triploid C. madrasensis had a growth advantage over meiosis II triploids which in turn had a growth advantage over diploids. Meiosis II C. gigas triploids grown at different sites grew faster than sibling diploids (Garnier-Géré et al., 2002). 4.2. Meat yield and shell production — implications for timing of harvest We hypothesised that with the development of gonad in H. laevigata, the average growth rate of diploid abalone should be lower relative to that of triploids, as the former expends more energy and nutrients for gonad

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development at the expense of somatic growth. This was the case on an absolute (micrometers/day) basis, however interestingly during both maturation periods, growth rates of both triploid and diploid abalone were depressed relative to other times of the year. During both maturation periods the larger diploid abalone had higher VGI scores, while very few of the triploid abalone showed signs of significant gonad development. Triploid abalone continued growing and adding to the meat during these maturation periods, which supports the hypothesis that energy and nutrients were directed to meat production and not gonad development. Triploid lion-paw scallop (N. subnodosus) also had lower growth rates and slightly (but not significantly), higher meat weights (17%) than refractory diploids (naïve control diploids had intermediate values) (Maldonado-Amparo et al., 2004). In contrast, 14 month old triploid scallops (Chlamys nobilis), 12 month old triploid catarina scallops (A. ventricosus), and 12 month old triploid oysters (C. madrasensis) had higher growth rates and meat yields than the diploid controls (Komaru and Wada, 1989; Ruiz-Verdugo et al., 2000; Mallia et al., 2006). Abalone total wet weights were proportional to approximately the cube of their length; 3.06 and 3.07 for H. laevigata (current study), 2.95 for H. laevigata (Reaburn and Edwards, 2003) and 2.82 for H. rubra (Dunstan, unpublished data), 2.99 for H. midae (Britz et al., 1997). During the maturation periods of the current study the wet meat weight of diploid H. laevigata also increased at approximately the cube of the length (3.1 and 2.8), in line with the total wet weight and shell weight. However, during the winter, wet meat weight of diploid abalone increased at a power of 3.7 of the abalones' length. This also suggests that meat production by diploid abalone was affected by a shifting of metabolic resources to gonad production during the maturation period. This was not evident in triploid abalone; meat production was continuous and high relative to total body growth at these three times (power of 4.0, 3.4, 3.4). Once spawned (or having produced gonad which is then mostly reabsorbed), the diploid H. laevigata recovered meat levels in line with triploid abalone (on a per length basis), resulting in no net benefit to off-season triploid meat production. Similar to H. laevigata, triploid C. gigas grew throughout the year, while diploid oysters exhibited distinct decreases in soft tissue weight and glycogen stores with onset of the spawning season (Allen and Downing, 1986). As diploid abalone generally had a slightly heavier shell and lower wet meat weights (and therefore a lower meat to shell weight ratio) than triploid abalone for a given shell length, processing would result in higher meat yield per kilo of whole live triploid abalone than

diploid abalone (after grading by length). This was particularly evident during the breeding season when the diploid abalone had further reduced meat yields for a given length, relative to triploid abalone. During the maturation periods (36 and 48 months) the percentage differences between the meat weight/shell length regressions increased with abalone size indicating that the wet meat gain in triploid relative to diploid abalone was proportionally greater for larger abalone. Also during the maturation periods, a diploid H. laevigata would need to be between 11 and 14% (at 36 months) or 6 and 11% (at 48 months) longer to yield the same wet meat weight as a triploid abalone. The diploid abalone in this study were on average 15% (at 36 months) and 11% (at 48 months) longer than triploid abalone. We propose that the effect of being triploid compromised the growth of other (non-meat) parts of the body such as shell and viscera (including gonad), leading to an overall reduced body size and growth, relative to diploid abalone of the same age. Therefore, for commercial abalone stocks graded by length and sold by meat weight (e.g. to canneries), harvesting of triploid abalone during the maturation period and either triploid or diploid abalone at other times could confer a cost benefit by yielding more meat while having to transport less weight to the processing plant. In contrast, for commercial abalone stocks sold by total weight (and independent of meat content), harvesting diploid abalone at any time could confer a cost benefit to the abalone farmer, but it may not be cost effective for the consumer. 4.3. Meat fatty acid composition If differences between the fatty acid compositions of the meat of triploid and diploid, or male and female abalone were to be detected, this would be evident at the height of the breeding season when significant differences in meat yield were detected, and when lipids such as fatty acids may be translocated to the gonad for gametogenesis. In finfish muscle, seasonal accumulation of lipids with feeding and depletion with breeding has been outlined in the literature (e.g. Kołakowska et al., 2003). However, very few significant differences were detected between the relative fatty acid compositions of triploid and diploid Chinese catfish (Clarias fuscus) or amago (Oncorhynchus masou ishikawae), although the triploids did have higher total amounts of fatty acids or lipids in the flesh compared to diploids late in the breeding season (Qin et al., 1998; Saito et al., 1997). These differences in the amount of lipid in triploid finfish may be due to the accumulation of lipid in the form of fatty acids through the diet, but unlike

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diploids, they do not utilize it during the breeding season for gamete production. In contrast, molluscs do not store lipids in the muscle — their main form of energy store in the muscle is carbohydrate (as glycogen) which is stored and therefore elevated in the muscle of triploids relative to diploids especially during the spawning season when the latter deplete their stores for energy used in reproduction (Ruiz-Verdugo et al., 2001; Garnier-Géré et al., 2002). In the current study however, there was no difference in the total muscle fatty acid content or composition of H. laevigata meat from either sex (as reported previously; Grubert et al., 2004), or from either ploidy type. Similarly, lipid levels did not vary between ploidy types in C. gigas (Garnier-Géré et al., 2002). Ploidy induction and having an extra set of chromosomes per cell did not affect H. laevigata lipid metabolism with respect to fatty acid composition of the abalone foot. The elevated 18:2n-6 and reduced 20:4n-6 (relative to previous studies of wild or seaweed fed abalone) in all H. laevigata foot samples was indicative of the diet being a formulated feed — and the elevated 22:5n-3 relative to 22:6n-3 is atypical of most marine animals but is typical of abalone (Dunstan et al., 1996; Grubert et al., 2004). 4.4. Broodstock conditioning — implications for sterility Sterility induction in aquaculture species can be an attractive feature for farmers with enhanced stock. For example, animals such as abalone can be sold live into the market place, where they could be intercepted by other breeders and used as broodstock. When such animals come from an expensive selective breeding program this can put the farmers' financial advantage at risk. Therefore, fast growing sterile animals are highly sought after from a production standpoint. Sterility also ensures that cultured animals that escape into the environment do not interbreed with, or their offspring compete with the native fauna (Ihssen et al., 1990; Beaumont and Fairbrother, 1991). Many triploid H. laevigata of adult size displayed little gonad material, and very few female triploid abalone were identified in the main commercial population. Overall, this study demonstrated that female H. laevigata were more adversely affected by triploid induction than male H. laevigata, and were possibly sterile, as has been found for other species (Ihssen et al., 1990). Assuming a 1:1 sex ratio as exhibited in the diploid H. laevigata of the current study, and other species of abalone (Litaay and De Silva, 2003 and references therein), and that half the triploid abalone were male and almost half being immature suggests that these immature triploid abalone were genetically

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female. Higher male to female ratios were identified in M. lateralis — with high levels of immature animals presumed to be female (Guo and Allen, 1994a). In the Sydney rock oyster (Saccostrea commercialis) the opposite was found, namely a lower male to female ratio with high levels of immature animals shown to be male (Cox et al., 1996). High male to female ratios were also shown in triploid scallops (C. nobilis, Komaru and Wada, 1989), while no female triploid mussels (M. galloprovincialis, Kiyomoto et al., 1996 and M. edulis, Brake et al., 2004) and no male triploid soft-shell clams (Mya arenaria, Allen et al., 1986) were identified. Further to this, the male gonad of triploids of the normally hermaphroditic catarina scallop (A. ventricosus) ceased development, and resulted in the unusual situation of all female gonads (Ruiz-Verdugo et al., 2000). Such variability in the molluscs has been attributed to the different sex determining models, including whether the dominant sex determining gene is paternally or maternally derived so as to explain the resultant ratios. In the case of H. laevigata, as the male to female ratio was approximately 1:1 and as triploid males predominated, it is possible that the sex determining mechanism is of an X:Y type, with XX females, XY males and paternally derived dominant “maleness” genes similar to M. lateralis, mammals and many other vertebrates (Guo and Allen, 1994a). Triploid female H. laevigata did not develop gonad past stage 1 after 222 days at an optimum temperature and diet for conditioning (Grubert and Ritar, 2005). In comparison, triploid male H. laevigata could be conditioned to stage 3, however three separate attempts to induce these males to spawn, failed. In other studies, female triploid dwarf surf clam were less fecund than triploid males (Guo and Allen, 1994a), and triploid scallops (C. nobilis), Sydney rock oysters (S. commercialis) and mussels (M. edulis), developed gonad but did not spawn (Komaru and Wada, 1989; Cox et al., 1996; Brake et al., 2004). Furthermore, while gametes were shed from triploid C. gigas, the amount of gonad in female triploids was reduced relative to that in male triploids, which in turn was reduced relative to both sexes of diploid C. gigas (Allen and Downing, 1986). Similarly, triploid M. galloprovincialis have been reported to produce a small number of spermatozoa, however their viability was not tested (Kiyomoto et al., 1996). Unfortunately, because triploidy induction is typically incomplete and the triploids induced may not be 100% sterile, the production of a sterile population of a particular species is not guaranteed. For example, only 95–99% of triploid N. subnodosus were sterile (Maldonado-Amparo et al., 2004). Triploid C. gigas eggs

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and sperm were found to be viable when crossed together and when crossed with diploid gametes (Guo and Allen, 1994b), and female triploid Japanese pearl oyster (Pinctada fucata martensii) crossed with a diploid male produced progeny (Komaru and Wada, 1994). Even though in both cases survival to spat (3– 4 months) was very low compared to pure diploid crosses, the triploids were clearly not sterile. 4.5. Conclusions As stated previously, for commercial abalone stocks graded by length and sold by meat weight, harvesting of triploid abalone during the maturation period could be cost effective for the farmer. However, for commercial abalone stocks sold by total weight, harvesting diploids at any time may be beneficial for the farmer, but not the consumer. Diploid female H. laevigata were on average slightly (but not significantly) larger and heavier than diploid males. Similarly, wild-caught female H. laevigata have been reported to grow faster than males (Shepherd and Hearn, 1983). This difference was not evident in the triploid H. laevigata. While triploid female H. laevigata were producing less gonad than triploid males, there was no growth or wet meat weight advantage in producing only triploid females. Nonetheless, there could be an economic advantage to producing all female stocks of H. laevigata if sterility in triploid females can be guaranteed. Acknowledgements We gratefully acknowledge the assistance of farm managers Miles Cropp (Abalone Farms Australia) and Mike Wing (Cold Gold) for the use of their facilities, and Drs. Belinda Norris and Peter Rothlisberg (CSIRO) for assistance with generating the triploid abalone. Also Drs. Arthur Ritar and Craig Mundy for the use of their abalone conditioning system (Tasmanian Aquaculture and Fisheries Institute), Dean Broughan and Jason Bartlett for technical assistance during the project and Dr. Belinda J. Norris and three anonymous reviewers for their valuable comments on the manuscript. References Allen Jr., S.K., Downing, S.L., 1986. Performance of triploid Pacific oysters, Crassostrea gigas (Thunberg). I. Survival, growth, glycogen content, and sexual maturation in yearlings. J. Exp. Mar. Biol. Ecol. 102, 197–208. Allen, S.K., Hidu, H., Stanley, J.G., 1986. Abnormal gametogenesis and sex ratio in triploid soft-shell clams (Mya arenaria). Biol. Bull. 170, 198–210.

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