trial: reducing costs and increasing survival .... Staining and tagging trial . .... One such example is Chesapeake Bay in the USA, where the annual income ...
Recreational Fishing Initiatives Fund: Project 2016/03
Blue Swimmer Crab (Portunus armatus) stocking trial: reducing costs and increasing survival Greg I. Jenkins1, Robert Michael1, James R. Tweedley2,3, David Oberstein3, Neil R. Loneragan2,3 & Danielle J. Johnston4 1
Australian Centre for Applied Aquaculture Research, South Metropolitan TAFE, Fremantle, Western Australia Centre for Sustainable Aquatic Ecosystems, Harry Butler Institute, Murdoch University, Perth, Western Australia 3 School of Veterinary and Life Science, Murdoch University, Perth, Western Australia 4 Department of Primary Industries and Regional Development, Hillarys, Western Australia 2
July 2018 Recipient: Australian Centre for Applied Aquaculture Research Grantor: The Government of Western Australia Department of Primary Industries and Regional Development (as a delegate of the Minister of Fisheries)
Copyright Recreational Fishing Initiative Fund, South Metropolitan TAFE and Murdoch University This work is copyright. Except as permitted under the Copyright Act 1968 (Cth), no part of this publication may be reproduced by any process, electronic or otherwise, without the specific written permission of the copyright owners. Information may not be stored electronically in any form whatsoever without such permission.
Disclaimer The authors do not warrant that the information in this document is free from errors or omissions. The authors do not accept any form of liability, be it contractual, tortious, or otherwise, for the contents of this document or for any consequences arising from its use or any reliance placed upon it. The information, opinions and advice contained in this document may not relate, or be relevant, to a reader’s particular circumstances. Opinions expressed by the authors are the individual opinions expressed by those persons and are not necessarily those of the publisher, research provider or those associated with the Recreational Fishing Initiative Fund.
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Table of Contents Acknowledgments ............................................................................................................ 4 Non-technical summary of research findings ..................................................................... 5 Section 1: Background information and project aims ......................................................... 7 Section 2: Culture of larval Portunus armatus, without the use of live microalgae ........... 11 Specific aim ........................................................................................................................... 11 Methods and results ............................................................................................................. 11 Broodstock ........................................................................................................................ 11 Larviculture ....................................................................................................................... 12 Megalopae ........................................................................................................................ 15 Discussion ............................................................................................................................. 18 Section 3: Staining/tagging trial and hatchery and field survival experiments .................. 22 Specific aim ........................................................................................................................... 22 Materials and methods ........................................................................................................ 22 Aquaculture and staining of Portunus armatus ................................................................ 22 Survival of Portunus armatus crablets in various types of habitat enrichment ............... 23 Estimates of the survival of megalopae and crablets in a controlled external environment ........................................................................................................................................... 27 Results................................................................................................................................... 30 Staining and tagging trial .................................................................................................. 30 Survival of Portunus armatus crablets in various types of habitat enrichment ............... 34 Estimates of the survival of megalopae and crablets in a controlled external environment ........................................................................................................................................... 35 Discussion ............................................................................................................................. 39 Staining and tagging trial .................................................................................................. 39 Survival of Portunus armatus crablets in various types of habitat enrichment ............... 42 Estimates of the survival of megalopae and crablets in a controlled external environment ........................................................................................................................................... 44 References...................................................................................................................... 46
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Acknowledgments This project was made possible by the Recreational Fishing Initiatives Fund and supported by Recfishwest and the WA Department of Primary Industries and Regional Development. Additional financial and logistical support was provided by the Australian Centre for Applied Aquaculture Research (ACAAR) and Murdoch University. Gratitude is expressed to the crab team at the Department of Primary Industries and Regional Development for catching broodstock crabs and to Charlie Maus, Jason Crisp, Clara Obregón Lafuente and Sarah Poulton for their help with the field and laboratory experiments. David Coaker and the team at Leigh Grange Fisheries are also thanked for their help and support with the grow-out of the hatchery-reared crablets.
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Non-technical summary of research findings This project has two main components; (i) aquaculture, i.e. to culture and release larval/juvenile crabs, without the use of live microalgae and (ii) experimental, i.e. to conduct staining/tagging trials and hatchery survival experiments. During this milestone period 36,000 late stage megalopae were cultured from the 1,420,000 zoea produced by four late stage berried females collected in the wild and induced to release their eggs in a single hatchery run undertaken at Australian Centre for Applied Aquaculture Research (ACAAR) in November 2017. Larvae were reared exclusively on Reed Shellfish diet, rotifers and Artemia. Survival remained relatively high (37%) right up until the last day of megalopae, when a large mortality event occurred prior to moulting into crablets. Surviving megalopae were harvested and counted and survival was determined to be around 6%. A total of 23,000 of these megalopae were released into the cooling water intake pond at the disused South Fremantle Power Station, 5,000 were kept for crablet tank trials at Murdoch University and 8,000 were kept at ACAAR for on growing and were later released into an inland aquaculture facility to assess the feasibility for their survival in a nursery/growout environment. The experimental component of this project set out to: (i) test the utility of a chemical stain (Alizarin Complexone) and physical tag (Visible Implant Elastomer) to denote hatchery-reared individuals, so that any crablets released into the wild could be identified; (ii) determine the survival of hatchery-reared crablets in a range of different habitats; and (iii) compare the survival of hatchery-reared megalopae and crablets in the wild to determine which size-atrelease provides the best survival rates. Staining crabs with Alizarin Complexone proved unsuccessful as the stain was not detected following mounting in resin and sectioning or dissection. However, Visible Implant Elastomer tags implanted in large crablets and adults were clearly seen under both normal and UV light. A 25 day tank trial in five different habtats types including four types of habitat enrichment (i.e. sand, pebbles, artificial turf and BIOBLOK) and a control (bare tank) showed that increasing habitat complexity may help reduce cannibalism, but that substantial cannibalism will still occur.
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The third experiment involved release of megalopae and crablets into a cooling water intake pond for a disused power station south of Fremantle. Following the successful acquisition of a number of approvals, a water exchange occurred to increase salinity and 23,000 megalopae were released and weekly monitoring was carried out after stocking. No crabs were caught during this sampling, indicating that those megalopae died, possibly due to the input of low salinity ground water. Rather than expose remaining 623 crablets to this unfavourable environment, they were all transported from the ACAAR hatchery in Fremantle to an inland saline aquaculture facility ~400 km north-east of Perth in Morawa. These crablets grew rapidly, i.e. from between 20-30 mm carapace width on January 12 to between 40-70 mm carapace width six weeks later, and some were still alive as of July 2018. Unfortunately, as the transfer resulted in the crabs being located at a commercial aquaculture facility six hours away and thus regular monitoring was not able to be maintained. Due to the lack of survival of the megalopae and thus the transfer of the crablets to a different environment no comparison of the trade-off of releasing larger numbers of megalopae vs smaller numbers of crablets were able to be undertaken. Given the results of the current study and the hatchery capacity in Perth it seems likely that any substantial culture effort would either (i) release at the megalopae size (and thus at the point when cannibalism starts), which, in aquaculture-based enhancement projects for other crab species has not proven to be successful or (ii) utilise an outdoor nursery/grow out environment, such as earthen ponds used in inland saline aquaculture, enriched with appropriate vertical habitat, for a period of time sufficient to allow growth to a large size before release, without becoming maladapted to the wild.
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Section 1: Background information and project aims Global exploitation of portunid crabs increased seven fold between the 1970s and 2000s with, for example, landings of the Blue Swimmer Crab (Portunus pelagicus complex, which includes Portunus armatus) and the Gazami Crab (Portunus tribuberculatus), increasing from ~525,000 tonnes in 2002 to ~820,000 tonnes in 2014 (FAO, 2016). These species constitute nearly 50% of global landings of portunids. A combination of anthropogenic influences, however, such as habitat loss, overfishing, eutrophication and reduced recruitment, have led to a global decrease in catch rate, as well as the collapse of several portunid fisheries (Miller et al., 2005). One such example is Chesapeake Bay in the USA, where the annual income generated from the Blue Crab (Callinectes sapidus) fishery has been reduced by 70% from US$ 900,000 to $ 300,000 in only 15 years, leading to the initiation of a large multi-year restocking project (Zohar et al., 2008). The Blue Swimmer Crab Portunus armatus is a highly sort after commercial and recreational species that is distributed throughout the tropics, with highest abundances occurring in the Indo-West Pacific region. In Australia, this species is distributed throughout coastal and estuarine waters and is the focus of commercial and recreational fisheries (Johnston et al., 2016). In Western Australia, P. armatus is by far the most popular species targeted by recreational fishers, with an estimated ~900,000 crabs being caught annually by boat-based fishers alone (Ryan et al., 2015) with significant additional recreational catches being taken by fishers wading in the shallows. Moreover, 70-80% of this fishing pressure occurs in the Peel-Harvey Estuary, which is located in a metropolitan area, ~70 km south of the capital city of Perth (Ryan et al., 2015). In 2013/14, boat-based recreational fishers caught an estimated 38-56 tonnes of P. armatus from the Peel-Harvey Estuary, which comprised ~75-80% of the total crab catch by fishers in the West Coast Bioregion and is roughly equivalent to the commercial catch of 57 tonnes in this estuary in 2015/16 (Johnston et al., 2016). With population of the City of Mandurah and surrounding areas growing rapidly and the increasing popularity of recreational fishing in Western Australia, fishing pressure from the recreational sector is likely to intensify. Moreover, fishing pressure for P. armatus in the Peel-Harvey and Swan-Canning estuaries has increased following the collapse and closure of Cockburn Sound 7
to commercial and recreational crab fishing (Johnston et al., 2011). These authors suggested that aquaculture-based enhancement or restocking P. armatus in Cockburn Sound, i.e. the culture of and production of crabs in aquaculture and their release into the marine environment, may be a potential method of rebuilding this fishery. Aquaculture-Based Enhancement (ABE) is a management approach involving the supplementation of stock via the release of hatchery-reared individuals in an attempt to restore or enhance the productivity of a fishery and includes restocking, stock enhancement and sea ranching (Bell et al., 2008; Taylor et al., 2017). Over time, ABE projects aim to increase the population by increasing spawning stock and optimise harvest by overcoming recruitment limitations. One of the most well-documented portunid ABE projects was completed in Chesapeake Bay, where low recruitment along with environmental degradation and high fishing pressure resulted in substantial declines in catches of C. sapidus (Miller et al., 2005). Using a multidisciplinary approach, crablets were successfully produced in aquaculture and released into the estuary, where they contributed to breeding stock and, in turn, helped restore the population of this portunid (Zmora et al., 2005; Zohar et al., 2008). Note that despite the success of the trial, only experimental scale releases were undertaken and a fullscale commercial restocking program was never initiated (Zohar et al., 2008). Although aquaculture techniques to produce hatchery-reared brachyurans have been developed and trials have shown these can be effective, albeit at experiment scales, a number of challenges need to be addressed, which could explain why, among portunid crabs, only the culture of Portunus trituberculatus has been scaled up to a commercial operation (Tweedley et al., 2017a). Principally, these challenges involve developing methods to (i) culture larvae to the crablet stage without live microalgae to reduce the costs of culture and (ii) reduce the high mortality rate in the hatchery due to cannibalism. For example, live microalgae was required for the culture and production of the Western School Prawn Metapenaeus dalli (Crisp et al., 2017a), but was one of the major costs of producing cutured prawns for release due to the need to employ a dedicated algal technician (Jenkins et al., 2017b; Tweedley et al., 2017b). Cannibalism is most influential when the larvae metamorphose to megalopae, and
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their asynchronous development results in the faster developing larvae becoming megalopae before others (Marshall et al., 2005).
The claws (chelipeds) develop during the megalopal stage and this allows the megalopae to exhibit raptorial capabilities on the less well-developed zoea (Romano and Zeng, 2017). This occurred in an earlier trial to grow P. armatus in Western Australia, where a 94% reduction in the number of individuals was observed in 48 h due to cannibalism and negative interactions during the metamorphosis between megalopae and crablets (Jenkins et al., 2017a). Therefore, as cannibalism has the potential to dramatically reduce the number of hatcheryreared crabs produced in a given aquaculture run, the future viability of any aquaculturebased enhancement program, including that for P. armatus, relies on reducing mortality in the hatchery by lowering instances of aggressive behaviour and cannibalism. Lower stocking densities and the provision of greater substrate complexity have reduced juvenile cannibalism in a range of brachyuran species (e.g. Romano and Zeng, 2017). Moreover, several crab species actively select more complex habitats, such as seagrass, macroalgae, pebbles and crushed shells over finer sand substrates (Van Montfrans et al., 2003) and artificial habitats have also been successful in increasing survival in the hatchery (Johnston et al., 2006; Rodriguez et al., 2007). Blankenship and Leber (1995) stated that “one of the most critical components of any enhancement effort is the ability to quantify success or failure”. This, however, requires a mechanism to distinguish between hatchery-reared and wild stock to enable post-release monitoring to be undertaken (Hodson, 2016). This has been done for fish by staining otoliths (Partridge et al., 2009) and applying external tags (e.g. t-bar tags). However, external tags poses a mortality risk for crustaceans during the moulting of their exoskeleton and, due to a lack of ‘bony structures’ for chemical stains to bond to, there is a need to develop successful methods of marking hatchery-reared crabs. The aims of this study were thus to: 1. culture and release larval/juvenile crabs without the use of live microalgae (Section 2)
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2. test the utility of a chemical stain (Alizarin Complexone) and physical tag (Visible Implant Elastomer) to denote hatchery-reared individuals, so that any crablets released into the wild can be identified (Section 3) 3. determine the survival of hatchery-reared crablets in a range of different habitats (Section 3); and 4. compare the survival of hatchery-reared megalopae and crablets in the wild to determine which size-at-release (megalopae or crablets) exhibits the best survival rates (Section 3).
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Section 2: Culture of larval Portunus armatus, without the use of live microalgae Specific aim The aim of this component of the project was to modify the culture protocols for Blue Swimmer Crabs (Portunus armatus) used in the precursor project (Jenkins et al., 2017a), which was based on the novel techniques developed by Jenkins et al. (2017b) as part of the Western School Prawn restocking project (Tweedley et al., 2017b). However, in an effort to reduce the cost of larval rearing, and thus the potential expenditure of any future aquaculture-based enhancement, the culture was to be attempted without the use of live microalgae.
Methods and results The text below describes the methods used to culture Blue Swimmer Crabs without the use of live microalgae and the number of megalopae and crablets produced from that aquaculture run. Broodstock Four late stage berried females (Fig. 1.1.) were collected from the Swan River on Friday 27 th October by DPRID research staff. These crabs were caught at 19 °C before being transferred into 300 L spawning tanks where they were slowly brought up to 23 °C. It only took two nights for spawning to occur with each tank containing thousands of hatched zoea the following morning (Table 1.1).
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Fig. 1.1. (left) Late stage female being transferred to a spawning tank and (right) 300 L spawning tanks and flow through zoea collectors.
Table 1.1. Relationship between carapace length and Zoea hatched. Crab Carapace length (mm) Number of zoea produced and their fate 1 145 1,240,000 (Euthanised) 2 155 1,420,000 (Stocked) 3 N/A N/A 4 N/A N/A
Larviculture Zoea were harvested from the spawning tanks using 250 µm collectors (Fig. 1.2.), counted and then stocked into both 3,000L larval tanks and the 5,000 L parabolic tank at a maximum stocking density of 100/L. See Appendices for detailed larval rearing protocol. Because of the relatively high survival observed across all tanks during the zoea stages (Table 1.2), individuals in the 5 tonne parabolic tank was harvested and euthanised to make room available to transfer later stage megalopae from the two 3 tonne larval tanks. This was done on Day 8 with 155,000 zoea 3 being counted from the 5 tonne parabolic tank (37% survival to this stage).
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Fig. 1.3. Submerged 250µm zoea collector.
Table. 1.2. Number of larvae stocked in various tanks. Number of larvae stocked Tank 426,000 5 t parabolic tank 300,000 3 t larval tank 300,000 3 t larval tank
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The photographs below outline the larval development through the main life stages.
Day 2: Zoea 1. Eyes sessile, abdomen 5 Day 4: Zoea 2. Eyes stalked, abdomen 5 segments. segments.
Day 7: Zoea 3. Eyes stalked, abdomen 6 segments, pleopod buds forming along lower ventral side of abdomen.
Day 10: Zoea 4. Eyes stalked, abdomen 6 Day 11: Zoea 4. Halfway through process of segments, pleopod buds well developed. moulting into megalopa.
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Day 12: Megalopa. Possesses biting claws and Day 16: Late stage megalopa. Abdomen pointed joints at ends of legs. Able to swim freely beginning to curve beneath cephlathorax. using pleopods.
Day 19: Crablet 1. Abdomen completely curved Day 22: Crablet 1. Terminal segments of the fifth beneath cepthlathorax. pair or periopods are fully developed and now function as swimmers.
Megalopae On day 15, during late stage Megalopae (Fig. 1.4), a large, unexplained mortality event occurred. The following day, both larval tanks were drained, harvested and counted. A total of 36,000 live, day 16 megalopae were collected. Of these, 23,000 were taken to the trial pond
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(Fig. 1.5), 5,000 were held in 1,000 L crablet tank for later use by Murdoch (Fig. 1.6) and 8,000 held for on growing in 5,000 L parabolic tank (Fig. 1.7). On 24 November approximately 300 crablets (day 26) were harvested from the 1,000 L tank and taken to Murdoch for tank trials (see later). On 22 December, 647 crablets (Day 54) were harvested and delivered chilled (15 °C) to Morawa for on-growing pond trials (Fig. 1.8). There was a large variation in the weight of these 54 day old crablets, which ranging from 0.057 – 1.435 g.
Fig. 1.4. Late stage megalopae prior to the mass mortality event.
Fig. 1.5. Adding seawater to the cooling water intake pond at South Fremantle Power Station.
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Fig. 1.6. A 1,000 L crablet tank.
Fig. 1.7. The 5,000 L parabolic tank enriched with habitat prior to stocking.
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Fig. 1.8. Crablets being transferred into pond at Morowa.
Discussion The aim of this component of the project was to test whether blue swimmer crabs could be successfully cultured using commercially available concentrated algal pastes rather than live algae which is typically used in crustacean larval culture. Although a large, unexplained mortality event occurred at day 15, this was several days after feeding with microalgal paste had ceased and was suspected to be the result of a bacterial outbreak. The regular addition of concentrated algal paste and low water exchange may have contributed to the increased bacterial loads within the culture tanks, however this could be mitigated in the future by modifying culture strategies, e.g. increasing water exchanges. However, overall, this component was deemed successful as survival remained relatively high during the nine day microalgae feeding stage with all crab larvae successfully metamorphosing through each zoeal stage, which shows that crab larvae were able to feed and grow on commercial algal pastes.
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Appendix 1. Production protocol used in the culture run.
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Appendix 2. Crab feeding protocol used in the culture run.
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Section 3: Staining/tagging trial and hatchery and field survival experiments Specific aim The aims of this component of the project were to (1) test the utility of a chemical stain (Alizarin Complexone) and physical tag (Visible Implant Elastomer) to denote hatchery-reared individuals, so that any crablets released into the wild can be identified; (2) determine the survival of hatchery-reared crablets in a range of different habitats; and (3) compare the survival of hatchery-reared megalopae and crablets in the wild to determine which size-atrelease (megalopae or crablets) exhibits the best survival rates.
Materials and methods Aquaculture and staining of Portunus armatus Female Portunus armatus carrying eggs, i.e. berried crabs, were collected from the PeelHarvey Estuary by staff from the Department of Primary Industries and Regional Development (DPIRD) in October 2016. These crabs were transferred to the Australian Centre for Applied Aquaculture Research (ACAAR) hatchery in Fremantle. As the berried females were collected from waters with a temperature of 19 °C, they were placed in to holding tanks and the water temperature raised to 25 °C over several hours to induce faster egg development and the release of the eggs. After being held for around seven days the eggs on the carapace began to change colour from yellow to black-grey, at which point each female crab was transferred into a 300 L spawning tank. Spawning, i.e. egg release, occurred overnight with each female producing between 300,000 and 800,000 zoea, which were stocked in culture tanks at a stocking density of 120 L-1. Zoea were fed a diet of algae and Artemia (following Zmora et al., 2005) and moulted through the megalopae stage, and eventually to crablets. At an age of 22 Days Post Hatch (DPH), a portion of the crablets were transferred to a solution of 10 ppm Alizarin Complexone for one hour. Following this, they were stocked in 1,000 L tanks with substrate to help reduce negative interactions between crablets. The crablets remained in this tank for a further 53 days and fed daily with a mixture of commercial prawn
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feed (Microelite or FrippaK), minced fresh fish and fish pellets. After 71 DPH, the tank was drained and the 89 crabs that had survived were euthanized in an ice slurry and frozen.
In the laboratory, the crabs were defrosted, their carapace width and weight measured to the nearest 1 mm and 0.01 g, respectively. A proportion of the crablets were dried at 60 °C for 48 hours embedded in resin, cut into ~1 mm thick sections and viewed under a high-power microscope to determine if the purple colour of the Alizarin Complexone stain had been retained in any of the hard parts of the crabs (e.g. the gastric mill and eyestalks). The above procedure was repeated using freshly defrosted (and not dried) crablets. Photographs of the resultant sections were taken under reflected light using a Leica DFC425 digital camera attached to a Leica MZ75 dissecting microscope. A third group of crablets was dissected and examined under a dissecting microscope to see if any stain was retained in any part of the body . Crablets from the above experiment and larger crabs (i.e. > 100 mm carapace width) were tagged using a 10:1 Visible Implant Elastomer (VIE) tag (Northwest Marine Technology Inc., Atlanta, USA). The elastomer components (i.e. the coloured component and the curing agent) were dispensed from separate syringes into a small plastic mixing cup in a 10:1 ratio. The components were mixed together for one minute using a thin wooden spatula and the resultant elastomer drawn in to a 0.3 cc injecting syringe. This procedure was conducted twice using two different colours, i.e. red and green. An injecting syringe containing a coloured elastomer was loaded into the Manual Elastomer Injector and the contents injected through the carapace into the body cavity or muscle. Photographs of the crabs were taken under normal light and a deep violet light of 405 nm, under which the VIE tags fluoresce. These light conditions were created using the VI light supplied by Northwest Marine Technology with the tags.
Survival of Portunus armatus crablets in various types of habitat enrichment Berried crabs were collected from the Swan-Canning Estuary by staff from DPIRD and the resultant eggs cultured using the same methodology described above, except that zoea were
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fed algal paste, not live microalgae (Appendix 1.1). At 30 DPH, a proportion of the crablets were transferred from the hatchery at ACAAR by road to a controlled temperature room at Murdoch University (Fig. 2.1). The room contained twenty 54 L glass aquaria (W 30 x L 60 x H
30 cm) filled with 40 L of high quality 1 µm filtered seawater at a salinity of ~35 ‰ drawn from a saline aquifer, aerated constantly and maintained at a temperature of ~24 °C. Each aquaria was randomly allocated one of four types of habitat enrichment or was a control (i.e. bare tank with no natural or artificial habitat; Fig. 2.2). The four types of enhancement were: (i) builders sand (grain size = 125–250 µm), (ii) pebbles (grain size =8-12 mm), (iii) artificial turf (pile height = 35 mm) and (iv) BIO-BLOK®, which is a series of polyethylene net tubes welded together (Fig. 2.2). As a total of 254 crablets were available for this experiment, 50 crabs were transferred to a single aquaria, each allocated to one of the four treatments and one control tank. Water quality was measured daily using a Yellow Spring Instrument 556 Water Quality Meter to ensure the 100W aquarium heaters were maintaining the correct water temperature and that any losses of water through evaporation, and thus increases in salinity, were accounted for by the addition of correct quantities of dechlorinated freshwater. Moreover, 50% (20 L) water changes, using 1 μm filtered seawater transported from the ACAAR hatchery were completed weekly. This was deemed sufficient due to the very low stocking density (three crabs and < 0.1 g wet weight per 100 cm2). Crabs were fed daily with 0.2 g of hydrated fish feed (Vitalis algae pellets 1.5 mm in diameter and comprising 38.5% protein and 8.8% fat) and visual observations indicated that this level of food was sufficient as the vast majority, but not all pellets, were consumed each day. The provision of habitat and the propensity of the crabs to burry or hide, meant that visual estimates were unlikely to provide a reliable estimate of actual crab abundance. Moreover, moving habitats in the aquaria to facilitate more accurate counts could incite aggressive behaviour in the crabs and/or directly result in their mortality (e.g. by damaging them). Thus, instead, a baited ‘plate trap’ was developed to sample the crabs during this experiment (Fig. 2.3). A 10 cm diameter lid from a screw top jar with a lip of 2 cm was modified by drilling
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four ~2 mm holes in the sides of the lid (90° apart) to allow the insertion of fishing line to create a lifting mechanism (Fig. 2.3). Around 15 holes of the same size were drilled into the bottom of the lid to allow water to drain during the lifting phase, while any remaining food pellets and crabs were retained. Note that these holes were substantially smaller than the carapace width of the crabs and thus no crabs were lost during the lifting process.
Fig. 2.1. Conceptual figure detailing the methodology followed during this project.
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Fig. 2.2. Photographs showing (a) the arrangement of the aquaria in the controlled temperature room at Murdoch University and close-ups of (b) the aquaria without habitat enhancement (bare/control treatment) and those with (c) sand, (d) pebbles, (e) artificial turf and (f) BIO-BLOK.
Fig. 2.3. Photographs showing (a) a baited plate trap deployed in an aquaria containing artificial turf, (b) aerial view of a trap containing bait and a crablet (black circle), (c) close-up of the crablet in (b) and (d) a crablet feeding on a pellet.
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Crablets were counted using the plate trap each day for the first week and every two days for the next two weeks, after which the experiment was terminated (i.e. 15 counts over the 24 day duration of the experiment). Each count was conducted by adding ~0.2 g feed to the plate trap together with a large metal sinker (to counteract against the buoyancy of the plastic lid). The trap was lowered to the bottom of each tank and left for one hour before raising it and counting any crabs before releasing them into the tank, together with any remaining food. At the end of the experiment, the tanks were drained and any surviving crabs were counted, measured and frozen for comparison with Alizarin stained crabs (see earlier). The total number of crablets in all aquaria (i.e. treatment and control combined) on each sampling occasion was correlated against the number of days post-release using a one-way Pearson’s correlation in Minitab 18. The null hypothesis of no significant relationship between the number of crabs and the number of days since the experiment started was rejected if the significance value (P) associated with the test statistic (r) was < 0.05.
Estimates of the survival of megalopae and crablets in a controlled external environment To assess the survival of the different life stages of hatchery-reared P. armatus in the wild (megalopae vs crablets), a suitable location that could be stocked was required. An initial idea of building small fine mesh cages and deploying them in the shallow waters of the SwanCanning or Peel-Harvey estuaries was rejected due to: (i) the very small size of the crablets (~2 mm total length) and (ii) the risk of the cages being tampered with by members of the public, thus reducing the reliability of the results. Instead, permission was sought to release crablets in a confined “natural” environment, i.e. the cooling water intake pond of the disused South Fremantle Power Station, which was 502 m2 in area and up to 2 m deep (Fig. 2.4). This site has undergone remediation work since the power station ceased operating in 1985 and is listed by the Department of Health and Department of Water and Environmental Regulation as decontaminated. The selection of this site over an ‘open’ estuarine environment, was based on the fact that it is a confined space where the restocked crablets cannot escape, therefore enabling effective monitoring of their survival. Permission to use the land was granted from Cockburn City Council, Department of Lands, State Electricity Commission of Western Australia (Western Power) and Synergy. 27
Fig. 2.4. (a) Photograph showing the cooling water intake pond adjacent to the disused South Fremantle Power Station and (b) a satellite image of the study site, showing the intake pond where stocking occurred denoted by the red box. Circles show sites at which water quality measurement were collected. Red = intake pond; blue = large intake pond and yellow = ocean. Baseline benthic macroinvertebrate and fish data were also collected from the four sites in the intake pond (red circles).
As outlined conceptually in Figure 2.1, the aim of this novel component of the study was to compare the survival of megalopae and crablets in the hatchery and wild. The megalopae produced from the culture run were split into three batches, with 5% transported to Murdoch for the habitat enrichment experiment (see above), 45% transferred to a large 50,000 L parabolic prawn tank at ACAAR filled with BIO-BLOK and nets to provide three-dimensional structure and kept in total darkness to reduce cannibalism, and the remaining individuals were released into the intake pond (Fig. 2.4). The aim was to compare survival in the hatchery and wild (intake pond) after one month and to determine whether releasing crabs at a large size resulted in greater survival. This design (Fig. 2.1) allowed the cost and benefits of (i) releasing megalopae vs crablets in the wild and (ii) holding megalopae in the hatchery vs releasing them into the wild to be determined.
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Prior to releasing the megalopae into the cooling water intake pond, water quality was measured. As this showed that the initial salinity was 3 ‰, a total of 936,000 L of water were pumped out (~53 % of the 1,762,000 L volume of the pond) and replaced with seawater (salinity = 35 ‰). This raised the salinity to 20 ‰. One of the benefits of releasing the hatchery-reared megalopae/crablets into the cooling water intake pond was that the absence of cages allowed the individuals to be predated on by other inhabitants of the pond, thus providing a more representative assessment of the likely survival in the wild. To determine the types and abundance of predators and potential prey present, a baseline survey of the benthic macroinvertebrate and fish fauna were conducted at each of the four sites shown in Figure 2.4b. A sample of sediment was collected from the deeper (> 1 m) waters of each site using an Ekman grab (Wildco, Yulee, FL, USA) that collected substratum from an area of 225 cm2 and to a depth of 15 cm (Tweedley et al., 2016a). Each sediment sample was preserved in 5 % formalin buffered in ‘estuary’ water and subsequently wet sieved though a 500 µm mesh and stored in 70 % ethanol. Using a dissecting microscope, any benthic invertebrates were removed from any sediment retained on the mesh and identified to the lowest possible taxonomic level (usually species). The benthic macroinvertebrate samples contained very few individuals and of a limited number of species. In total, they contained three chironomid larvae; two individuals of the amphipod Austrochiltonia subtenuis and a Ceratopogonid larva. The composition of the fish fauna of the intake pond was determined by deploying a 21.5 m seine net at each site. The net, which comprised two 10 m long wings (6 m of 9 mm mesh and 4 of 3 mm mesh) and a 1.5 m bunt (3 mm mesh), fished to a maximum depth of 1.5 m and swept an area of 116 m2 (Potter et al., 2016). All fish collected were identified quickly and returned alive to the water. The following five fish species were recorded in the pond, with their abundances expressed as a density 100 m-2; Flathead Goby (Callogobius depressus) 7.32, Black Bream (Acanthopagrus butcheri) 5.17, Sculptured Goby (Callogobius mucosus) 4.31, Bridled Goby (Arenigobius bifrenatus), 1.72 and Pink Snapper (Chrysophrys auratus) 0.64. Note that the individuals belonging of both sparid species were juveniles, i.e. < 10 cm in total length. 29
A total of 23,000 megalopae (22 DPH and thus about to metamorphose into crablets) was transferred by road from the ACAAR hatchery to the intake pond. Water quality (temperature, salinity and dissolved oxygen concentration) was monitored at each of the four sites using a Yellow Spring Instrument 556 Water Quality Meter in the intake pond at time of stocking and weekly for 28 days. Additional sampling of the larger intake pond and the ocean (Fig. 2.4b) was conducted in the final three weeks to provide comparison data. Post-release monitoring of the crablets was conducted 24 hours post-release using an Ekman grab. Three replicate cores were obtained from the shallow areas near the bank, intermediate and deeper waters of each of the four sites. Given the failure to catch crablets using this method, on each of the following four sampling occasions, a baited plate net was employed as this was larger in area (2,500 vs 225 cm2) and the addition of the bait would likely increase the catch rate. Note that this ‘net’ is similar to those used on commercial prawn farms to determine abundance of prawn stocks. Each net consisted of a 50 cm x 50 cm metal frame, overlaid with 0.2 mm mesh. Rope, 50 cm long and 0.5 cm thick, was tied from each corner to a central 3 cm diameter ring above the centre of the net to provide a stable attachment point for 2 m of lifting rope and a marker float and to prevent tilting of the net (and thus the spilling of contents during each set and retrieval). Approximately 10 g of Vitalis algae pellets were hydrated in ‘estuary’ water to ensure they would be negatively buoyant and placed on the net together with a lead weight to facilitate fast sinking. On each sampling occasion, a single plate net was baited, deployed and left in the water for one hour before being retrieved and any crablets counted and returned alive to the pond.
Results Staining and tagging trial When the sections of dried crabs were viewed under the compound microscope, no evidence of the purple Alizarin Complexone stain was detected (Fig. 2.5). Moreover, the drying process led to the internal parts of the crabs shrinking inside the carapace, becoming brittle and occasionally, if the sections were cut too thin (< 1 mm thick), falling out or disintegrating. The
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sectioning of ‘fresh’ crablets was slightly more successful in that more of the crablet was visible in the section, however, once again, no evidence of the stain was observed in the tissue or exoskeleton (Fig. 2.6). Visual analysis of the gastric mill, eyestalks and other internal structure of the crablets under a dissecting microscope also failed to detected any Alizarin Complexone. Crablets of ~10 mm carapace width were tagged successfully into the body cavity using the red VIE tags. The colour of the tag could be seen through the darker dorsal side of the carapace under both visual and VI light, although the tags were far clearer when fluoresced under the VI light (Fig. 2.7a, b). When injected into the perieopods, both the green and red VIE tags were visible under normal light despite the patterning on the carapace, with the red colour slightly more noticeable to the naked eye (Fig. 2.7c). Under florescence, both tags were clearly visible on the 2nd pereopod and stand out against the colouring of the carapace (Fig. 2.7c, d). The green VIE tag was clearly visible when implanted on the underside of the merus of the 5th perieopod (i.e. the swimming leg) under both light conditions (Fig. 2.7e, f). Finally, both colours of tag were employed in ‘dots’ and ‘dashes’ on the underside of the thoracic sternite (below the gills) and were clearly distinct from the white carapace. This demonstrates that the tags could be arranged in, for example, a Morse code arrangement to distinguish between marked crabs (i.e. #4 vs #95).
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Fig. 2.5. Photographs taken through a compound microscope of sections of dried crablet to detect evidence of uptake of the stain Alizarin Complexone by the crabs.
Fig. 2.6. Photographs taken through a compound microscope of sections of ‘fresh’ crablet to detect evidence of uptake of the stain Alizarin Complexone by the crabs.
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Fig. 2.7. Photographs of crablets and crablet body parts injected with red and/or green Visible Implant Elastomer tags taken under visible (a, c, e, g) and (b, d, f, h) VI light.
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Survival of Portunus armatus crablets in various types of habitat enrichment On November 24 2017, 50 crablets were stocked into each tank. After 24 days, a single crablet was found in each of the bare (control), pebble, artificial turf aquaria, with two individuals in the BIO-BLOK and none in the sand treatments (i.e. survival ranged from 0 to 4%). The average size of the five crablets at the end of the experiment was 9 mm in carapace width (5 mm carapace length) and the average weight was 0.09 g wet-weight. However, the size of the crablets varied markedly ranging from 6 to 12 mm carapace width (4 to 7 mm carapace length) and 0.01 to 0.11 g in weight. The largest and smallest crabs were both found in the BIO-BLOK treatment at the end of the experiment (Fig. 2.8).
Fig. 2.8. The two crablets, of markedly different sizes, found in the BIO-BLOK treatment at the end of the habitat enrichment experiment, 24 days post-release.
Crablets were caught using plate traps in at least one of the five treatments on eight of the 15 sampling occasions (Fig. 2.9a). A greater total number of crablets (≥ 3) were caught on the first three sampling occasions with ≤ 1 being recorded in the remaining twelve events. No crablets were caught on six of these occasions. Pearson’s correlation detected a significant relationship between the total number of crablets and the number of days since the crablets were released into the aquaria (r = -0.671; p = 0.006; n = 15; Fig. 2.9b). The frequency of occurrence of crablets was greatest in the sand treatments (four occasions), followed by the BIO-BLOK, artificial turf and sand (all three occasions) and lastly the pebbles (one occasion). Visual analyses at the time of sampling indicated that some food pellets were found on the bottom of the aquaria in each treatment and that, particularly, in the BIO-BLOK treatment, crablets were observed in the tank but were not collected in the plate traps. 34
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Days post-release Fig. 2.9. Number of (a) crablets found in each of the treatment aquaria on each sampling occasion and (b) the relationship between the total number of crablets in all treatments on each sampling occasion. Dashed line shows the predicted line using a least squares regression.
Estimates of the survival of megalopae and crablets in a controlled external environment Despite collecting three replicate Eckman grabs samples at each of the four sites in the intake pond in the first week post-release no crablets were recorded. It was thus decided to switch methodology to the baited plate net, which may attract any remaining crablets to the net and thus be more effective than the unbaited grab samples. Monitoring using the baited plate nets set for one hour also failed to catch a crablet at any of the sites in any of the four weekly
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sampling trips. No crablets were caught following the release of 23,000 megalopae into the small intake pond (502 m2). Salinity in the intake pond declined markedly over the course of the study ranging from a maximum of ~20 ‰ immediately post release (week zero) to 11 ‰ after a week and only six in the final two weeks of the experiment (weeks three and four; Fig. 2.10a). In contrast, salinity in the larger intake pond remained lower and relatively consistent; it was ~3 ‰ in week two and 2 ‰ in the following two weeks. The salinity of the ocean in those three weeks was constant at ~35 ‰ (Fig. 2.10a). In the third week post-release, a halocline was present with salinities at each of the sites on the surface of the water columns being relatively ‘fresher’ than those on the bottom (i.e. 4.7 vs 6.7 ‰; data not shown). Water temperature also declined over the duration of the experiment from a peak of ~32 °C at the start of the experiment (i.e. immediately post-release) to a minima of ~20 °C in week four (Fig. 2.10b). Typically temperatures in the intake pond were slightly higher than those in the larger intake pond and the ocean by between 2 and 4 °C in weeks two and three, respectively, although they were all similar after 4 weeks (~20 °C). Daytime dissolved oxygen concentrations recorded at ~10:00 am were always normoxic (i.e. ≥ 2 mgL-1) ranging between ~6 and 12 mg L-1, and were broadly similar to that in the larger intake pool and the ocean (Fig. 2.10c).
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Due to the relatively large decline in salinity in the intake pond (i.e. 20 to 6 ‰) and the absence of crablets in either of the Eckman grab or baited plate net samples (indicating that the megalopae released into the intake pond had undergone large mortality), it was decided that conditions in the intake pond were not, at that time, conducive to the growth of P. armatus crablets. Thus, the remaining 623 crablets retained in the ACAAR hatchery were transferred by road to an inland saline aquaculture facility (Leigh Grange Fisheries) in Morawa (~ 400 km north-east of Perth) on December 22 2017 (Fig. 2.11). They were stocked in to a 15 m x 15 m and 2 m deep polyethene lined earthen pond (covering 225 m2; or 450 m3; stocking density = 2.8 crabs m-2), supplemented with BIO-BLOK to enrich the habitat on December 21 2017. The crablets were fed daily with 1 kg of pelletised fish food. By January 12, the crablets were between 20 and 30 mm carapace width and by February 26 they had continued to grow rapidly to reach between 40 and 70 mm carapace width (Fig. 2.12). Hatchery technicians lifted up the BIO-BLOK and, on one occasion, found 14 crablets sheltering in the three-dimensional structure this medium provides. Baited plate net samples collected during the morning typically yielded between one and three crablets, with crabs also spotted sheltering along the edges of the pond too (Dave Croaker, Leigh Grange Fisheries; personal observation). The aquaculturists are confident they can continue to hold the crablets until they reach market size. However, as these tanks are not heated it is not known when this might occur. It has not as of July 2018.
(a)
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Fig. 2.11. Crablets grown in the Australian Centre for Applied Aquaculture Research in Fremantle being released into the outdoor lined earthen ponds of Leigh Grange Fisheries in Morawa.
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Fig. 2.12. Crablets sampled from the earthen ponds of Leigh Grange Fisheries in Morawa in late February 2018 (approximately four months post hatch), again showing the large difference in size among individual crablets/crabs (i.e. 40 – 70 mm carapace width).
Discussion This component of this project set out to: (i) test the utility of a chemical stain (Alizarin Complexone) and physical tag (Visible Implant Elastomer) to denote hatchery-reared Portunus armatus, so that any crablets released into the wild can be identified. (ii) Determine the survival of hatchery-reared crablets in a range of different habitats. (iii) Compare the survival of hatchery-reared megalopae and crablets in the wild to determine which size-atrelease provides the best survival rates. The focus of this research was to help improve the feasibility of any future aquaculture-based enhancement project for P. armatus, by increasing survival in the hatchery and wild by reducing mortality and determining the best size-atrelease and testing various methods of distinguishing between wild and hatchery-reared individuals to allow and monitoring and evaluation regime to be established. Staining and tagging trial The staining of crablets (22 DPH) with 10 PPM Alizarin Complexone was not successful as the stain was not detected in individuals, either following mounting in resin and sectioning or
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after dissection. This is in direct contrast to the results obtained during the staining of numerous fish species (Beckman and Schulz, 1996; Thorrold et al., 2002; Partridge et al., 2009). Typically this stain, and similar compounds, have been used to mark otoliths and, in some species, have been shown to remain visible for at least 15 years (Cottingham et al., 2015). Moreover, at higher concentrations (i.e. > 300 ppm and thus markedly greater than the 10 ppm used in the current study), such stains are able to mark scales and fin rays under fluorescent wavelengths of light (Liu et al., 2009). The lack of staining success in the current study may reflect the fact that brachyurans contain fewer internal structures composed of calcium salts with which the stain reacts with (Puchtler et al., 1969), e.g. they do not have an otolith, but also that these crustaceans undergo regular moulting, which is more frequent in the early life stages. For example, Alizarin Complexone stained the carapace of juvenile Western School Prawns (Metapenaeus dalli), however, after several moults, the newly formed carapace was free of the stain and only the exuviae retained the stain. This explains why the exoskeleton (including the eyestalks, chelipeds and walking legs) does not retain the stain. Work by Brösing (2014) has shown that parts of the foregut and hindgut are also subjected to moulting. Although some calcium (and potentially some stain) is resorbed from the exoskeleton in aquatic crustaceans most is lost to the external environment (Greenaway, 1985; Brösing, 2014). While it is possible that some of the Alizarin Complexone was resorbed and incorporated in to the new exoskeleton and fore and hind guts, it is relevant that the quantity of stain employed in the current study (i.e. 10 ppm), was at the lower end of the range employed by a suite of studies, which have used concentrations of up to 400 ppm (Beckman and Schulz, 1996; Lagardère et al., 2000; Liu et al., 2009). Alternative tagging methods, such as elemental tagging using strontium and rare earth elements has been used for a variety of fishes, producing long-lasting signatures in the otoliths, vertebrae, opercula and spines (Thorrold et al., 2002). However, Levin et al. (1993) found that most elements were either not absorbed or not retained for a sufficient time periods to function as a tag even if that were incorporated in to the carapace. Similar results were obtain by Anastasia et al. (2003) for cobalt, silver, europium and gadolinium for brachyuran larvae, although selenium was retained above background levels for some weeks
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and could be worthy of further investigation. Another approach is the use of genetic markers as an effective method of identifying hatchery-reared individuals on the basis of work done on several penaeids (Bravington and Ward, 2004; see also Crisp et al., 2018). Bravington and Ward (2004) noted, however, that a greater suite of markers would need to be developed for aquaculture-based enhancement programs, such as the current one, where wild berried broodstock were collected, as the genotype of the father(s) are unknown. While the Alizarin Complexone was not successful, the Visible Implant Elastomer tags were able to be implanted in to both large crablets and adults and were clearly visible under both normal and UV light. Such tags, although mainly used on fish (Catalano et al., 2011; FitzGerald et al., 2011), have been used on a range of invertebrates including cephalopods (Zeeh and Wood, 2009; Barry et al., 2011), penaeids (Dinh et al., 2011) and brachyurans (Davis et al., 2004; Morgan et al., 2006). Although the several studies have shown that the tags remain visible for extended periods of time, for example, six months to two years, (Woods and James, 2003; FitzGerald et al., 2011), the results are likely species specific and thus further studies would need to be done of P. armatus crablets to further evaluate their potential performance. It is encouraging that Liu et al. (2011) found that large tagged Mud Crabs (Scylla paramamosain; 157-250 mm carapace width) did not suffer enhanced mortality over an eight-week period, despite undergoing three ecdyses. In contrast, T-bar tagging had a 100% failure rate in P. armatus during moulting (McPherson, 2002), albeit Meynecke et al. (2015) had more success using this method in Giant Mud Crabs (Scylla serrata). In the current study we tagged individuals that had been euthanised, but Liu et al. (2011) provide a methodology for live animals. These authors selected six locations on the ventral surface of the juvenile crab, adjacent to the second, third and fourth pair of pereiopods (Fig. 2.13). Prior to tagging, they immersed the crabs in iced water for 6–8 seconds to induce temporary shock, with the individual immersed again for 3–4 seconds if another injection was made. On each occasion, elastomer was transversely injected into the tissue between the cortex and muscle, forming a linear shape about 2 mm long. As the elastomer comes in a range of ten colours (Fig. 2.13), there is the potential to use the six injection points and suite
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of colours to enable a coding system to be develop that could distinguish between cohorts or culture runs of crabs.
F
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Fig. 2.13. The marks of juvenile Mud Crab (Scylla paramamosain). (A) Six injection sites and injected dot photographed under natural light. (B) Under deep purple light from VI light. (C) The marking retention after the (D) first, (E) and third (F) moult. Taken from Liu et al. (2011). (F and G) the ten colours of Visual Implant Elastomer tags. Photo from https://www.nmt.us/visible-implantelastomer/.
Survival of Portunus armatus crablets in various types of habitat enrichment Cannibalism, which typically starts following metamorphosis to megalopae, is a major cause of mortality within the hatchery environment and has the ability to severely limit production. For example, this prevented the larger-scale production of the Alaskan Red King Crab Paralithodes camtschaticus (Zmora et al., 2005; Romano and Zeng, 2017; Tweedley et al., 2017a). Moreover, even at lower levels, cannibalism increases the costs of producing larger crabs and thus reduces the cost-effectiveness of any aquaculture-based enhancement. In the current experiment, cannibalism was prolific in most treatments, despite the fact that the
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rearing protocol involved providing more benthic food than was required as recommended by several authors (Baylon et al., 2004; Romano and Zeng, 2017). This may reflect the stocking density in the experimental tanks, which at 50 individuals per 54 L glass aquaria with a surface area of 1,800 cm2 equated to a stocking density of 270 crablets m-2. This is markedly greater than the 7, 10 and 25 individuals m-2 used by Smallridge (2002), that after 90 days had a survival of 100%. It is worth noting, however, that at higher stocking densities of 50 and 100 individuals m-2 the same authors achieved survival rates of 57 and 23%, respectively. In the current study, a 25 day tank trail employing a range of types of habitat enrichment (control [no habitat], sand, pebbles, artificial turf and BIO-BLOK) showed that increasing habitat complexity may help reduce cannibalism, but that substantial cannibalism will still occur. This was not surprising as the provision of habitats with structural complexity including seagrass, macroalgae, paving stones, pebbles and crushed shells have been show to increase survival compared to bare habitats (Van Montfrans et al., 2003; Marshall et al., 2005; Romano and Zeng, 2017). That being said, given the extent of cannibalism it is unlikely that any culture effort would be based on bare or unenriched tanks. Although, due to low number of crablets, we were not able to statistically determine the best of the enriched treatments, at the end of the experiment the BIO-BLOK treatment contained greatest number of crablets. From visual observations made during the experiment, this was the only enrichment treatment that provided vertical relief and thus smaller crablets could climb up the structure to escape larger individuals. Similarly, artificial net shelters have been used successfully by Rodriguez et al. (2007) and Zmora et al. (2005). Therefore, given the swimming ability of P. armatus the provision of vertical habitat may help reduce cannibalism more than benthic habitats/shelters alone. It is also noteworthy that, in the ACAAR hatchery, the crablets were held in the dark, i.e. a 0:24 photoperiod, which was considered to help reduce cannibalism (R. Michael, personal obs.), although this may reduce the growth rate of the crablets as has been shown in crab larvae (Azra and Ikhwanuddin, 2015). While the stocking densities in the experiments undertaken in the current project were deliberately high, i.e. to potentially allow a repeated measures experiment to determine the best experimental treatments, it does raise the question that if even the crablets in the best
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treatment (i.e. overfed and provided with vertical habitat enhancement) were subjected to cannibalism, would this prevent the aquaculture-based enhancement project being undertaken in the future? Given the results of this and the field experiment and the hatchery capacity in Perth, it seems likely that any substantial culture effort would (i) have to release at the megalopae size (and thus at the point when cannibalism starts), which, in other species of crabs has not proven successful (Tweedley et al., 2017a) or (ii) utilise an outdoor nursery/grow out environment, such as earthen ponds used in inland saline aquaculture for a period of time sufficient to allow growth to a large size, without becoming maladapted to the wild. Although future studies are needed to determine the length of that time. It is also worth noting that Lebata et al. (2009) found that conditioning juveniles in earthern ponds for 4-6 weeks prior to release enhanced survival and that the ponds used by Turano (2012) for Callinectes sapidus did include predators. Moreover that for the congener Portunus pelagicus survival at a stocking density of 3 crabs m-2 was 47% but could be increased to 58% through the provision of shelters, in this case bent plastic plates (Oniam et al., 2011).
Estimates of the survival of megalopae and crablets in a controlled external environment The third experiment involved release of crablets and megalopae into a cooling water intake pond of a disused power station. However, despite 23,000 megalopae being released, subsequent monitoring, done weekly for one month, failed to catch a single crab, indicating that those megalopae likely died in the early stages of the experiment. Before the experiment started, 936,000 L of cooling pool water (salinity of 3‰) were exchanged, using a pump, with a similar amount of full-strength seawater (salinity = 35‰), raising the salinity in the pond to 20. While this was lower than the ~35‰ the megalopae were raised in at the aquaculture facility a study by Fujaya et al. (2015) found that, of the species in the Blue Swimmer Crab complex, P. armatus and P. pelagicus were the most suitable for rearing in a ‘brackish’ water pond. However, although later stage juveniles of most crustaceans the are less sensitive to changes in salinity (Kinne, 1963, 1964; Crisp et al., 2017b), the subsequent decline in the cooling intake pond from 20 to 10 within the first week of the study may have been too substantial. With the broodstock having been collected from microtidal south-western
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Australian estuaries, where many of the local fish and penaeid species have adapted to spawn around summer, when rainfall is low and thus the physico-chemical conditions in the estuary are stable (Potter et al., 2015; Tweedley et al., 2016b; Broadley et al., 2017), these crabs may not be able to tolerate ‘rapid’ changes in salinity. Given the lack of survival of the megalopae, the release site of crablets was moved from the cooling water intake pond to a dedicated grow-out pond at an inland saline aquaculture facility were they grew rapidly and are still alive. The cooling water intake pond was chosen over a ‘natural’ site in an estuary due to the potential for interference in the natural site from inquisitive people. Although this experiment failed, likely due to the groundwater transport of low-salinity water from the larger cooling water intake pond that goes under the power station itself, which are a salinity of ~2‰, manmade ponds located on private property are likely to prove very useful in future experiments if the aquaculture-based enhancement of P. armatus was to be trialled again. For example, earthern ponds with and without artificial grass habitats and predators have been used successfully in trials to produce C. sapidus in the USA (Turano, 2012). The stocking density employed in the current study (2.8 m-2) was similar to that used by Oniam et al. (2011) for P. pelagicus. While the crabs in the current study grew rapidly in the grow-out ponds, doubling in size from 20-30 mm to 40-70 CW in only six weeks, we are unsure of the survival rate and this warrant further investigation if an optimal method for culturing and potentially restocking P. armatus is to be developed. It is noteworthy that Oniam et al. (2011) achieved a survival of 58% after 120 days using plastic plates as artificial habitat. Given the highly cannibalistic nature of this species and the fact that releases of megalopae have generally been unsuccessful (Tweedley et al., 2017a), the use of saline inland aquaculture ponds, like that investigated for a number of finfish species in Western Australia including for Barramundi (Lates calcarifer) and Black Bream (Acanthopagrus butcheri) (Partridge and Jenkins, 2002; Doupé et al., 2005; Partridge et al., 2008).
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