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NSW Department of Primary Industries – Fisheries Research Report Series:

20

Manual for hatchery production of Sydney rock oysters (Saccostrea glomerata)

by Wayne O’Connor, Michael Dove, Ben Finn and Stephan O’Connor

October 2008

ISSN 1449-9959

NSW Department of Primary Industries – Fisheries Research Report Series

This series presents scientific and technical information on general fisheries research and the documents in the series are intended to be progress reports on ongoing investigations. Titles in this series may be cited as publications, with the correct citation on the front cover.

Fisheries Research in New South Wales Fisheries research activities in the NSW Department of Primary Industries are based at various centres throughout the state. The studies conducted cover commercial and recreational fisheries and aquaculture, and conservation issues in coastal and riverine areas. The major role of the research is to provide information upon which relevant fisheries management policies and strategies are developed, monitored and assessed in terms of the Department’s obligations under the NSW Fisheries Management Act, 1994. Title: Manual for hatchery production of Sydney rock oysters (Saccostrea glomerata) Authors: Wayne O’Connor, Michael Dove, Ben Finn and Stephan O’Connor Published By: NSW Department of Primary Industries (now incorporating NSW Fisheries) Postal Address: PO Box 21, Cronulla, NSW, 2230 Internet: www.dpi.nsw.gov.au

© NSW Department of Primary Industries, the Fisheries Research and Development Corporation and Seafood CRC This work is copyright. Except as permitted under the Copyright Act, no part of this reproduction may be reproduced by any process, electronic or otherwise, without the specific written permission of the copyright owners. Neither may information be stored electronically in any form whatsoever without such permission. DISCLAIMER The publishers do not warrant that the information in this report is free from errors or omissions. The publishers do not accept any form of liability, be it contractual, tortuous or otherwise, for the contents of this report for any consequences arising from its use or any reliance placed on it. The information, opinions and advice contained in this report may not relate to, or be relevant to, a reader’s particular circumstance. ISSN 1449-9959 [Note: Prior to July 2004, this report series was published as the ‘NSW Fisheries Resource Assessment Report Series’ with ISSN number 1440-057X]

Hatchery Manual for Sydney Rock Oysters, Wayne O’Connor et al.

TABLE OF CONTENTS

Table of contents ................................................................................................................................................1 List of figures ................................................................................................................................................2 List of tables ................................................................................................................................................2 Acknowledgements ............................................................................................................................................3 1. INTRODUCTION ........................................................................................................................................4 2. SEAWATER...............................................................................................................................................5

2.1. Water storage...................................................................................................................... 5 2.2. Seawater delivery................................................................................................................ 7

3. BROODSTOCK ..........................................................................................................................................8 3.1. Introduction ........................................................................................................................ 8 3.2. Conditioning systems ........................................................................................................ 10 3.3. Feeding broodstock .......................................................................................................... 11

4. SPAWNING .............................................................................................................................................13 4.1. Thermal and salinity induction spawning......................................................................... 13 4.2. Strip spawning gametes .................................................................................................... 15

5. EMBRYOS...............................................................................................................................................17 6. LARVAE .................................................................................................................................................18

6.1. Larval equipment .............................................................................................................. 18 6.2. Larval sampling and measuring techniques ..................................................................... 22 6.3. Larval feeds ...................................................................................................................... 24 6.4. Algal cell counts ............................................................................................................... 25 6.5. Calculating larval feed rates ............................................................................................ 26 6.6. Hatchery time-line for SRO production............................................................................ 28

7. SETTLEMENT..........................................................................................................................................34 7.1. Introduction ...................................................................................................................... 34 7.2. Epinephrine....................................................................................................................... 35 7.3. Procedures for the use of epinephrine.............................................................................. 37 7.4 Grading............................................................................................................................. 39

8. NURSERY ...............................................................................................................................................40 8.1. Downweller systems.......................................................................................................... 40 8.2. Fluidised spat bottle system.............................................................................................. 40 8.3. Daily protocols for the spat bottle system......................................................................... 43 8.4. Diets and food rations for juvenile SRO spat ................................................................... 44

9. TRANSPORT............................................................................................................................................45 9.1. Transportation of spat ...................................................................................................... 45 9.2. Transportation of broodstock ........................................................................................... 46 9.3. Counting spat for delivery ................................................................................................ 46

10. HEALTH .................................................................................................................................................48 10.1. Introduction ...................................................................................................................... 48 10.2. Methods for sample collection.......................................................................................... 48 10.3. Sample investigation......................................................................................................... 48 10.4. Formaldehyde/formalin and ethanol ................................................................................ 48

11. REFERENCES ..........................................................................................................................................50 12. APPENDICES...........................................................................................................................................51

12.1. Daily larval record sheet .................................................................................................. 51 12.2. Larval feed calculation ..................................................................................................... 52

NSW DPI – Fisheries Research Report Series: No. 20 1


LIST OF FIGURES

Figure 1. Map of Port Stephens showing Halifax Point and Shoal Bay in relation to Port Stephens Fisheries Centre (PSFC), Taylors Beach. .................................................................................... 5

Figure 2. Oyster hatchery, PSFC, Taylors Beach, NSW. ........................................................................... 6 Figure 3. Water storage tanks at PSFC. ...................................................................................................... 6 Figure 4. Seawater delivery filtration system at PSFC. .............................................................................. 7 Figure 5. Example of an oyster in ripe and poor reproductive condition. .................................................. 9 Figure 6. Broodstock conditioning system. .............................................................................................. 10 Figure 7. Sydney rock oysters on spawning tables. .................................................................................. 14 Figure 8. Timeline of one cycle for spawning Sydney rock oysters. ........................................................ 14 Figure 9. Freshly released eggs illustrating the “water hardening” process. ............................................ 15 Figure 10. ‘Stripping’ or removing of gametes from a ‘ripe’ Sydney rock oyster using a pasteur

pipette. ....................................................................................................................................... 16 Figure 11. Developmental stages of bivalve larvae. . ................................................................................. 17 Figure 12. Larval retention screens and apparatus used during water exchanges in 20,000L tanks and

1000L tanks. .............................................................................................................................. 19 Figure 13. Oyster larvae retained on screens. ............................................................................................. 20 Figure 14. Stages of development throughout the bivalve larval cycle.. .................................................... 21 Figure 15. Equipment used to sample, observe and measure larvae........................................................... 22 Figure 16. Larval sampling from a 20L bucket. ......................................................................................... 23 Figure 17. Sedgewick-Rafter slide.............................................................................................................. 23 Figure 18. Sydney rock oyster larval feeding curve. .................................................................................. 24 Figure 19. Algae cultured in 10 L carboys and 500 L bags. ....................................................................... 25 Figure 20. Schematic of the counting grid from an Improved Neubauer Haemocytometer. ...................... 26 Figure 21. The mean larval growth of Sydney rock oyster production runs conducted since 2003. .......... 30 Figure 22. Unbonate SRO larvae. ............................................................................................................... 31 Figure 23. Sydney rock oyster pediveligers. The eyespot is clearly visible and the foot is shown

protruding from the mantle cavity. ............................................................................................ 32 Figure 24. Recently settled Sydney rock oyster spat. Note, the newly formed gill arch and gill buds....... 34 Figure 25. Pictures and components of a downwelling unit used to rear SRO spat. .................................. 35 Figure 26. Fluidised spat bottle system. Inset shows close-up of spat in the base of the bottle. ................ 41 Figure 27. Spat being placed on mesh for transport. .................................................................................. 46

LIST OF TABLES

Table 1. Larval screen sizes used at the PSFC and their diagonal measurements. .................................. 21 Table 2. Estimation of spat numbers based on the height of spat within the spat bottle for the PSFC

system........................................................................................................................................ 42 Table 3. Flow rates used for SRO spat at various sizes and densities in the spat bottle system at

PSFC.......................................................................................................................................... 42 Table 4. SRO spat/ml for different size grades settled using epinephrine bitartrate. .............................. 47

2 NSW DPI – Fisheries Research Report Series: No. 20


ACKNOWLEDGEMENTS

The development of techniques for the production of SRO has benefited from the considerable efforts of a number of researchers over many years. Among those to make significant contributions have been Baughn Wisely, John Holliday, John Nell, Lindsay Goard, Ken Frankish John Diemar and Mike Heasman. We also thank those researchers and hatchery operators who have assisted and provided advice in the process of improving SRO production protocols and establishing the mechanisms for the provision of SRO seed to industry, notably Martin John, Rod Grove-Jones and Nik Duyst. The assistance of Patrick Hone and Peter Horvat (FRDC) in the preparation of this manual is gratefully acknowledged.



1. INTRODUCTION

Known in New South Wales as the Sydney rock oyster (SRO), Saccostrea glomerata (Gould 1850) is found along much of the eastern Australian coastline (Lamprell and Healy 1998) as well as across the Tasman Sea in New Zealand (Anderson and Adlard 1994). Although it was once farmed in both countries, the culture of S. glomerata is now largely confined to estuaries in southern Queensland and New South Wales, where it forms the basis of one of the oldest and largest aquaculture industries in the eastern states. In New South Wales (NSW), the industry currently generates approximately $36 million in sales annually. Overall production of S. glomerata has been decreasing since the mid 1970s. The causes of the decline are varied and include competition from other faster growing oyster species, disease and declining water quality (White 2002). To assist industry, NSW DPI Fisheries commenced breeding programs for S. glomerata in 1990 with the intention of producing faster growing oysters and later incorporating disease resistance. Despite the success of this program, industry adoption of the benefits of genetic research have however been greatly constrained by the chronic failure of attempts to produce commercial quantities of S. glomerata in hatcheries in NSW. Hatchery production of S. glomerata commenced in the early 1980s but has been plagued by recurrent mass mortality syndromes in both larvae and spat. In a review of hatchery production attempts for S. glomerata undertaken at the NSW DPI Fisheries, Port Stephens Fisheries Centre (PSFC) between 1985 and 1999, Heasman et al. (2000) found 57% of larval runs failed within eight days of fertilisation. For these batches where larvae survived to settlement, approximately half of the batches of spat suffered sudden mortality episodes in which 60 – 90% of the spat were lost. These losses were also characteristic of the attempts to produce SRO at a number of private hatcheries during that time. It is worth noting, that some bivalve species are more amenable to culture than others. The hatchery at the PSFC has successfully produced an extensive range of bivalve species and has been responsible for the development of large scale rearing techniques for a number of species. In this context, the production of SRO has been by far the most problematic. To attempt to overcome production constraints, NSW DPI Fisheries, with strong support from the Fisheries Research and Development Corporation (FRDC), began to implement simple but significant modifications to existing bivalve hatchery facilities, rearing equipment and operating protocols at PSFC. These changes followed advice received at the Sydney Rock Oyster Hatchery and Nursery Health Workshop, a commissioned hatchery audit and HACCP plan, and the outcomes of a recent hatchery research grant from FRDC (FRDC 2003/209). The outcome of the amendments to hatchery facilities and protocols, and ongoing research has been that significant improvements in the reliability of SRO production at the PSFC have been made. Indeed between 2003 and December 2005, no failures in larval batches occurred and over 30 million selectively bred oysters were distributed to industry. This manual documents the new equipment and procedures that were in use at the PSFC over that time.



2. SEAWATER

2.1. Water storage

A fundamental requirement for successful bivalve production is an adequate source of good quality seawater and the ability to maintain the quality of that seawater. All seawater used within the PSFC mollusc hatchery is collected from sites within Port Stephens (Figure 1) and then transported to the hatchery by truck. Seawater collected can be of variable quality dependent upon antecedent seasonal and climatic factors, chiefly fresh water influx to Port Stephens and offshore conditions. The salinity of the trucked water generally ranges between 33‰ and 35‰. Water temperature is seasonal and varies from a high of approximately 24 – 25°C in the summer months (December to February) to 14 – 16°C during winter months (June to August).

Figure 1. Map of Port Stephens showing Halifax Point and Shoal Bay in relation to Port Stephens Fisheries Centre (PSFC), Taylors Beach.



Figure 2. Oyster hatchery, PSFC, Taylors Beach, NSW. On arrival at the PSFC hatchery, seawater is filtered through two parallel 25 µm filter bags before it is stored in a clean 45,000 L storage tank (Figure 3). There are four 45,000L tanks at the PSFC hatchery with a further two 55,000L tanks used during periods of high water demand. The seawater is held for a minimum settlement period of three days in these tanks. This settlement process allows any suspended material from the water column to settle and ensures that there is a shift in the bacterial microflora away from potentially pathogenic species (e.g., Vibrio spp.) towards more benign flora (Frankish et al. 1991).

Figure 3. Water storage tanks at PSFC.



Outlets on the storage tanks are placed so that only approximately 40,000L can be pumped from each tank to the hatchery. The residual seawater on the bottom of the tank contains material that has settled from the water column and is discarded. Tanks are then cleaned using freshwater, scrubbed and allowed to dry for at least one day. Before major larval production runs, the storage tanks are cleaned using Virkon S (Antec International), scrubbed and allowed to dry for at least one week.

2.2. Seawater delivery

Seawater is pumped from the storage tanks via a 75mm flexible hose into the hatchery where it is passed through two 1µm (nominal) filter bags, configured in series (Figure 4). The delivery lines within the hatchery are all constructed from 50mm PVC pipe. The lines and valves can all be dismantled for cleaning at any time and have been designed so that almost all points within the pipe work can be visually inspected and all pipes can be drained completely.

Figure 4. Seawater delivery filtration system at PSFC. Before commencing a production run, seawater delivery lines are dismantled, physically cleaned by either brushes or by passing a cleaning ‘pig’ through the line before they are dried for at least five days. All valves in the hatchery are ball valves that can be fully dismantled, cleaned and dried before being reassembled for use. After the delivery lines are reconstructed they are filled with chlorinated water for 24 – 48 hours. The lines are then flushed with fresh seawater and allowed to dry for a further 24 hours. At the end of each day, used filter bags are removed from the filter housing, washed and placed into chlorinated water. The concentration of chlorinated water used for cleaning throughout the hatchery is 50mL of chlorine (13% hypochlorite w/v) dispersed in 200L of freshwater (or 0.25mL/L). After 24 hours the filter bags are removed from the chlorinated water, washed again and allowed to dry for a further 24 hours. Three sets of filter bags are required for both the hatchery and the storage tanks to allow adherence to these protocols.



3. BROODSTOCK

3.1. Introduction

There are more than 30 oyster producing estuaries spread along the coastline of NSW (28° to 37° S) which enables collection of sexually mature SROs for most of the year. However, when oysters in good reproductive condition are unobtainable (commonly between June to September), or broodstock of particular genetic lines are needed, conditioning within the hatchery is necessary for successful spawning. Broodstock conditioning usually entails oysters of reasonable reproductive condition being brought into the hatchery and allowed to reproductively mature or ripen over time under culture conditions. By obtaining broodstock from a variety of coastal locations the hatchery is vulnerable to the translocation of diseases and external pathogens into the hatchery. Therefore, very strict quarantine protocols are followed before broodstock enter the hatchery (refer to Section 3.2 for quarantine protocols). It is common practice at the PSFC hatchery that broodstock used for production runs are brought into the hatchery and kept for a period of 2 – 8 weeks before spawning induction. The period of conditioning depends on the reproductive condition of the oyster when they enter the hatchery. Oysters with little or no reproductive development are avoided because of excessive time periods and food demands associated with conditioning stock from a “resting” or “spent” reproductive state. The reproductive condition of an oyster is usually determined by a visual inspection of the gonad. When assessing reproductive condition, the size of the gonad, its turgor, its colour and the degree to which the gonad is filled with gametes are all used. Gonads are ranked on a scale of 1 to 5, with 1 representing gonads that have recently spawned or have no gametogenic development; through to 5 indicating ripe, ready to spawn oysters. For example, an oyster that is ready to spawn will usually have a gonad that: occupies approximately 70% of the shell cavity (or left valve); is firm, has a white creamy colour; and, has visible “veining” within the gonadal tissue (Figure 5).



Figure 5. Example of an oyster in A) ripe and B) poor reproductive condition. Unlike some other bivalves such as scallops, visual determination of reproductive condition involves sacrificing the oyster. Therefore, small sub-samples of approximately six oysters are often observed at weekly intervals to assess the reproductive development of the broodstock.



3.2. Conditioning systems

The PSFC hatchery uses closed recirculating systems to condition oysters. Each system is comprised of a 1000 – 1400L reservoir tank and a 150L holding tank in which the oysters are held (Figure 6). A pump submersed in the reservoir tank transfers water to the holding tank to cause water to circulate inward. A raised drainpipe is located in the centre of the holding tank. This allows faeces to accumulate in the centre of the header tank where they are manually siphoned from the system daily.

Figure 6. Broodstock conditioning system. Conditioning systems undergo a complete water exchange every second day and care is taken to avoid any stimuli that could encourage oysters to spawn. Variations in water temperature are the most likely stimuli to cause spawning and so water is heated to a matching temperature before the water exchange takes place. This process becomes more critical as the broodstock move closer to sexual maturity (ripe). Oyster emersion and handling or fluctuations in salinity can also cause premature spawning. The temperature range for conditioning SROs is usually between 22ºC and 26ºC and can be manipulated depending on the rate at which the oysters need to be conditioned. Higher temperatures are thought to promote more rapid conditioning.



The presence of gametes in the water used to hold broodstock can also stimulate spawning. To safeguard against this, batches of broodstock are usually divided between two conditioning systems. This control measure ensures that if one conditioning system spawns prematurely, another batch of conditioned broodstock is readily available. In addition, water exchanges for concurrently held batches of broodstock are made on alternate days so that the likelihood of inadvertent spawning in all broodstock batches is further reduced. The movement of oysters between estuaries in NSW is strictly governed by a series of legislative requirements designed to prevent the spread of diseases and certain fouling organisms. Therefore, before broodstock are permitted to enter the hatchery, strict protocols are followed to ensure that all fouling organisms are removed. It is preferable that this is done at the point of origin, although all oysters are inspected again on arrival. Broodstock oysters are bathed in an iodine solution (10mL of Providone-Iodine Antiseptic Solution is added to 10L of freshwater) and scrubbed clean with a stiff brush. As a rule, broodstock are also held in a separate and isolated area of the hatchery, well away from larval or spat rearing areas. The health of broodstock is visually assessed on collection from the estuary and daily inspections of broodstock are done when they are held in the conditioning tanks.

3.3. Feeding broodstock

The diet for conditioning adult oysters always comprises a minimum of at least three or more microalgae. Those commonly used at the PSFC are Chaetoceros muelleri, Tetraselmis chui, Pavlova lutheri and Tahitian Isochrysis aff. galbana. Other favoured species include Skeletonema costatum and Thalassiosira pseudonana, all of which are obtained from the CSIRO laboratories, Hobart. Broodstock are fed to satiation with their diet generally comprising approximately equal proportions of each species, with the exception of Chaetoceros muelleri, which is often used to make up to 50% of the diet. This diet can be fed in batches, at least twice daily, although continuous drip-feeding is considered to be the most effective means of food delivery.

3.3.1. Feed rates

The algal consumption of SROs varies according to a number of factors, most notably their size and the water temperature and salinity. As a rule of thumb, we allow the dry weight equivalent of 6 billion T. Isochrysis cells/oyster/day for planning purposes and feeding usually starts at this rate. However, feed rates are very quickly modified in accordance with the rate at which the oysters are removing the food from the water column. Ideally, we are looking for most of the algae to be removed between feeds and thus observing the colour of the culture water provides a good indicator that the correct feeding regime is being used. In addition careful observation will also show whether or not faeces or pseudofaeces are being produced. Excessive feed rates can lead to high pseudofaecal production which is inefficient and of less benefit to the oyster.

Number of oysters X Maximum Algal Consumption of Adult Oyster

Per Day = Total algal Feed

Per Day



Example: Total daily feed for 100 broodstock Total Algal Feed / Day = 100 x (6.0x10 9) algal cells = 6.0 x 10 11 algal cells Feed ratio = 1/3 of each algal species (33.3% of total diet for each species). * Dry weight factors vary between algal species due to the differing cell sizes between algal species.

Dry weight factors (From Nell and O’Connor, 1991) (Where T. Iso = 1, Pav = 0.8, C. muelleri = 0.67) Total algal feed = 6.0 x 1011 or (600 x 109) algal cells Total P. lutheri = 0.8 x (600 x 109) = L 3 x count (cells L-1) Total T. Isochrysis = 1 x (600 x 109) = L 3 x count (cells L-1) Total C. muelleri = 0.67 x (600 x 109) = L 3 x count (cells L-1)



4. SPAWNING

4.1. Thermal and salinity induction spawning

Thermal or heat induced spawning techniques are the mainstay for spawning SROs. Broodstock are removed from the conditioning system and cleaned using Providone-Iodine antiseptic solution and then left out of water in a cool dry area (16 – 20oC) for 24 hours before spawning induction. The broodstock are placed on the spawning table with the right (or flat) valve of the oyster facing upright. This orientation makes it easier to observe the initial release of gametes. At PSFC we have a specially constructed table for spawning (Figure 7). The table is painted black so that when eggs and sperm, that are cream/white in colour, are released they are more easily seen. The table is also divided into four bays so that several groups of oysters can be spawned separately. The number of broodstock used depends on the task at hand and the size and condition of the oysters to be spawned. While it is possible to have all the oysters spawn, it is wise to allow a generous margin for error. For commercial scale production runs we may use as many as 400 million eggs, so we would commonly use approximately 100 mature broodstock. This number also allows for at least 20 males and 20 females to be used in each run to ensure a measure of genetic variability within each batch. Sexually mature oysters are placed on the spawning table in seawater that is the same temperature as that from which they were being held for conditioning, commonly 24°C, 35‰ salinity and filtered to 1µm. The oysters are allowed to acclimate for 15 minutes and should be observed to ensure they are open and filtering seawater. The temperature is then steadily increased by 4 – 5ºC (up to a maximum of 30ºC) over 30 minutes using an immersion heater equipped with a pump to circulate the water. The oysters are held at 30ºC for 15 minutes before freshwater is added to reduce the salinity from 35‰ to 22‰. This reduction in salinity is measured using a hydrometer or a calibrated salinity meter, but can be achieved by simply adding an additional 50% of the current table volume in freshwater (i.e., if the table is currently holding 100L of seawater, add 50L of freshwater). The oysters are held at 22‰ for 15 minutes. If spawning has not commenced, the spawning table is drained and the induction procedure is repeated. This thermal stimulation and salinity reduction process is known as a “cycle” (Figure 8). It is not uncommon for oysters to progress through a number of cycles before starting to spawn. Generally, reproductively mature oysters will spawn within 2 – 3 cycles. Any protracted delays in spawning (for example more than four cycles) are viewed as sign of poor reproductive condition which may impact larval performance. When individual oysters commence spawning, they are removed from the spawning table and placed in individual 500mL plastic food containers filled with seawater (26oC; 33 – 35‰), where they continue to spawn.



Figure 7. Sydney rock oysters on spawning tables.

Figure 8. Timeline of one cycle for spawning Sydney rock oysters. Fecundity (the number of eggs released) in SROs is high, with the female capable of producing in excess of 20 million eggs per spawning, while a male produces many times this number of sperm. A spawning male can be identified as having a “cigar” like trail or a continuous milky stream of sperm liberated from the exhalent siphon with a “pearly” lustre being observed at the top of the individual container in which the male is held. Females tend to “puff” the eggs out with a contraction of the adductor muscle and closure of the valves. When the container in which the females spawn into is held up to the light, individual eggs can be observed in the water column. The size of an egg ranges from approximately 50 – 60µm. Once spawning is complete, the eggs from selected females are pooled and gently washed through a 63µm screen and retained on a 20µm with 26ºC seawater. The eggs are then suspended in seawater in a 20L bucket of ambient seawater. This screening process assists to remove any debris produced during the spawning period (e.g., shell fragments, faeces and pseudofaeces from the broodstock). If spawning is protracted, i.e., some oysters spawn on one cycle and some on the next, then a number of fertilisations may be done so that eggs and sperm are not left for long periods of time (>60 minutes) before fertilisation occurs. Sperm can be placed in a refrigerator and chilled to 4°C or placed on a bed of ice to stop it from deteriorating over time if the spawning period becomes protracted.



While the eggs from females that spawn poorly or that are clumped on release are discarded, the quality of the remaining eggs is checked before fertilisation occurs. A 0.5mL sample is then taken from the bucket containing the pooled eggs to examine egg quality and to determine the number of eggs present in the bucket using light microscopy (magnification 100X). Sperm from each of the selected males is pooled and a sample taken to check it’s motility using light microscopy (magnification 200X). Fertilisation is then conducted as soon as practically possible (generally within 60 minutes of spawning). A general rule for fertilisation has been to supply enough sperm so that at least one sperm is seen at the periphery of each egg. After a 5 minutes interval a second sample is taken to examine the egg/sperm ratio, percentage fertilisation and to ensure that the eggs are water hardening (taking on a rounded shape, see Figure 9). Once the majority of eggs are fertilised they are stocked into an incubation tank.

Figure 9. Freshly released eggs illustrating the “water hardening” process. Before spawning, an incubation tank is filled with seawater and heated using immersion heaters to 26ºC. The tank is gently aerated and any immersion heaters are removed before the zygotes are stocked.

4.2. Strip spawning gametes

Many oyster hatcheries around the world obtain gametes by physically removing them from the gonad. Known as “strip spawning”, this is a fast efficient method of obtaining gametes. The procedure varies from scarifying the surface of the gonad and washing the gametes free to inserting a Pasteur pipette beneath the gonad epithelium to gently suck the gametes out (Figure 10) and resuspending them in seawater. The efficacy of strip spawning varies with species and works comparatively poorly with SROs. Generally the technique is avoided because the percentage development of larvae (<10%) is far inferior to that achieved with thermal stimulation. However, for breeding programs where absolute control of parentage is required, the gametes from individual oysters can be kept separate and specific males can be fertilized with predetermined females. Its other application is for eliciting spawning. The introduction of small quantities of sperm to the water during thermally induced spawning acts as an additional stimulant for spawning. Stripped sperm can be treated in a microwave oven to reduce the risk of uncontrolled fertilisation occurring on the spawning table.



Figure 10. ‘Stripping’ or removing of gametes from a ‘ripe’ Sydney rock oyster using a pasteur pipette.



5. EMBRYOS

Upon release, eggs have a characteristic pear shape due to crowding in the gonad, however after a short time in seawater these eggs take on a rounder shape, referred to as “water hardening” (Figure 9). Under the microscope the nucleus can be seen as a large transparent area surrounded by densely packed yolk granules. Many sperm may attach to the egg but usually only one penetrates to complete fertilisation. Upon fertilisation the egg contracts; assuming a spherical shape, and the cytoplasm becomes so dense that the nucleus is no longer visible. The first visual sign of development is polar body release, which depends on the water temperature, salinity and the quality of the eggs. Usually polar bodies should start to appear well within 20 minutes and the second polar body, which is harder to see, appears within the first 40 minutes post-fertilisation. At 26°C, development continues to the stage of a free-swimming trochophore in approximately 6 hours and the first hinged D-veligers appear 16 hours post-fertilisation (Figure 11).

Figure 11. Developmental stages of bivalve larvae. (Source: Helm et al. 2004, page 101).

Development rates for Sydney rock oysters are generally high (>90%) and there appears to be a small window of opportunity between 18–20 hours where the early developing D-veligers can be screened off using a 35µm screen and segregated from the slower developing larvae. This window for segregation is used in production runs to try to collect the best quality larvae. NB: This assumes that the gametes were collected and fertilised at the same time, which is not always the case in SRO spawnings.



6. LARVAE

Research into the rearing of Sydney rock oysters at PSFC began in the mid 1970’s and in April 1981 larvae were first successfully reared beyond metamorphosis to yield spat (Frankish et al. 1991). From this time until 1987 poor larval retention rates of approx 35% to pediveliger stage were observed, which resulted in changes being made to stocking densities and algal consumption (Frankish et al. 1991). In 1987, batch after batch failed, with food consumption, growth and development halting prior to early umbo stage. Since that time until the late 1990’s successful batches were scarce and successful runs were largely confined to the period between September and January. This yield period coincides with the natural conditioning and sexual maturation of the broodstock used for the spawning. At PSFC between 1985 and 2003, 56% of all larval batches failed with 57% of these failures being within the first eight days (Heasman et al. 2000). Larval batches over this time conducted between December and June period failed 70% of the time and, of those runs that did yield spat, approximately half suffered 60 – 90% spat mortality (Heasman et al. 2000).

6.1. Larval equipment

At PSFC the larval rearing room consists of four 20,000L fibreglass larval rearing tanks. At any one time only two of these tanks are used allowing a spare tank for each batch to be maintained that can be thoroughly cleaned and dried awaiting the next water exchange. Each tank has a 50mm flanged outlet approximately 30mm from the base of the tank and attached to the flange is a 50mm ball valve. Both the flanged outlet and 50mm valve can be fully dismantled, cleaned and chlorinated after each water exchange to ensure that there is no microbial build up in the system. The tanks and screens used to collect larvae during water exchanges are illustrated in Figures 12 and 13. The larvae are collected on a series of immersed screens, with the larvae being graded over a primary screen before the effluent is passed over a finer backup screen where any larvae passing through the primary screen are retained. From the backup screen the effluent is then directed into the hatcheries drainage system. Following use, this equipment is dismantled and cleaned using Virkon S solution, scrubbed and then rinsed with freshwater after each water exchange.



A.

B.

Figure 12. Larval retention screens and apparatus used during water exchanges in A) 20,000L tanks and B) 1000L tanks.



Figure 13. Oyster larvae retained on screens. Duplicate sets of larval screens, ranging from 20µm (used for washing eggs) – 265µm (used for grading freshly metamorphosed spat from larvae) are maintained. Two sets allow tanks to be drained concurrently and helps prevent cross contamination between the larval batches. Common mesh screen sizes and their diagonal measurement are listed in Table 1. The four main developmental stages of bivalve larvae, and the approximate size at which each of these stages occur, is displayed in Figure 14. Screen maintenance is important as these screens are the fundamental mechanism for retaining larvae. Before every water exchange, screens are checked for any visible holes in the fabric or any degradation of the silicon seals around the screen. Any small holes (1 – 2mm) can be repaired using an eye-dropper and PVC glue. Any hole larger than this size requires replacement of the screen material. Spare mesh of each screen size is kept on hand so that repairs can be made quickly. Embryo and larval tanks are heated using titanium immersion heaters. The heaters however are removed before embryos and larvae are added and the room is heated to 26.5°C so that the tank temperature remains stable at about 25.5 – 26°C.



Table 1. Larval screen sizes used at the PSFC and their diagonal measurements.

Nominal Size

Diagonal (µm) Nominal Size

Diagonal (µm)

35 40.00 142 200.82 45 63.64 150 212.13 55 74.95 170 240.42 63 89.00 180 254.56 85 120.21 190 268.70 90 123.00 200 283.00

100 141.00 212 299.81 108 152.00 236 333.75 118 166.88 250 354.00 124 175.36 265 374.77 132 179.00 292 412.95

Figure 14. Stages of development throughout the bivalve larval cycle. (Source: Helm et al.

2004 page 101).



6.2. Larval sampling and measuring techniques

Commonly used equipment to sample, observe and measure larvae is displayed in Figure 15. At each water exchange, larvae are rinsed into a 20L bucket and gently suspended uniformly into the water column using a perforated plunger (Figure 15). Two replicate 1mL samples are taken using an automatic pipette and each is placed on a separate Sedgewick-Rafter Chamber for counting slide (Figure 17). Using a binocular microscope (Figure 15) fitted with an ocular micrometer (40X magnification) the number of larvae within each sample is determined and the antero-posterior shell lengths of 30 or more larvae are measured to gauge the growth and survival of the larval batch.

Figure 15. Equipment used to sample, observe and measure larvae.



Figure 16. Larval sampling from a 20L bucket.

Figure 17. Sedgewick-Rafter slide. A sample of larvae should be examined live to assess their general health then fixed with a drop of 10% formalin and seawater (Section 10.4) to allow rapid accurate counting and sizing. These counts and sizes are also used to determine feed rates and to adjust larval densities when required.



6.3. Larval feeds

Larval feeding commences when the first D-veligers are observed, usually after 16 – 18 hours has elapsed. Total number of D-veliger larvae and their average shell size enables the algal feed rate to be determined from the equation and line displayed on Figure. 18. These data are derived from a succession of SRO larval runs over many years. The feed curve is used as a guide only and is altered in accordance with daily observations of the larval cultures. If excess food in the culture water is observed the feeding rate may be reduced, conversely if larvae have a light gut colouration the feeding rate may be increased. Inexperience can sometimes introduce a tendency to over-feed but this should be avoided as the consequences can be equally as damaging to larval performance as under-feeding. The daily larval ration is always divided over a number of feeds with at least a minimum of a morning and afternoon feed.

Figure 18. Sydney rock oyster larval feeding curve. Four species of algae are fed to larvae during a larval run. These algal species are T. Isochrysis, P. lutheri, C. calcitrans and C. meulleri. Both T. Isochrysis and P. lutheri are fed continuously throughout the larval run and both individually represent 25% of the total daily feed ration. At PSFC C. calcitrans is fed to the larvae (in conjunction with T. Isochrysis and P. lutheri) for the first seven days at a rate of 50% of the total daily feed ration. After Day 7, C. calcitrans is phased out of the diet and C. meulleri is introduced (approx 10% per day) until C. meulleri forms 50% of the total daily feed ration. Initially, the larvae are fed from 10L carboys (Figure 19) containing concentrated algal cell cultures for the first 14 days. At about this time food consumption increases to the point at which it becomes necessary to move to larger 500L bag cultures (Figure 19). By the time settlement occurs the larvae are fed solely from bag cultures.



A. B.

Figure 19. Algae cultured in A) 10 L carboys and B) 500 L bags.

6.4. Algal cell counts

Algal cell counts form an important part of the feeding regime. Before calculating the daily larval feeds, the cell densities of the algal cultures are determined. Algal counts for each species of algae are done on a daily basis. Preparing an algal sample

1. Collect sample from algal culture. 2. Add a drop of formalin and gently agitate sample. 3. Clean haemocytometer slide and cover slip gently using a tissue. 4. Place cover slip on haemocytometer. 5. Using a 1mL pipette, draw some algae and then load the chambers of the haemocytometer by

gently releasing the algal sample from the pipette so that it is drawn in between the cover slip and the haemocytometer surface.

6. Place the haemocytometer onto microscope platform. Cell counts

1. Focus the microscope on the haemocytometer grid. 2. Determine the number of algal cells visible within 1mm x 1mm square (Figure 20). There are

nine larger pronounced squares on a haemocytometer that are 1 mm2, each square is again subdivided into a number of smaller grids. The pronounced centre square on the haemocytometer (circled on Figure 20) is divided into 25 separate sub-squares. Count all the algal cells in this square and multiply this number by 10,000 (volume conversion factor for 1 mm2) to determine the number of algal cells in 1mL. The pronounced squares surrounding the centre square contain 25 separate sub-squares – again you need to count the cells within the 25 sub-squares to ascertain how many cells there are in 1mm2 and then multiply by the volume conversion factor (10,000).



Figure 20. Schematic of the counting grid from an Improved Neubauer Haemocytometer. Example: Algal cell count If 764 cells of algae are counted in the 25 sub-squares in the centre of the Improved Neubauer Haemocytometer, then the algal count for the culture sampled will be 764 x 10,000 = 7,640,000 or 7.64 x 106 cells/mL. If 84 cells of algae are counted, then the number of algal cells per mL of algal culture is 84 x 10,000 = 840,000 or 8.4 x 105 cells/mL. Note: The more 1mm2 sections of the Improved Neubauer Haemocytometer that are counted and averaged the more accurate the algal density estimate will be.

6.5. Calculating larval feed rates

Larval feed rates are determined according to the feeding equation displayed in Figure 18, which is based on the number of cells of T. Isochryisis to be fed to each larva each day. To determine the total daily feed requirement for a batch of larvae it is necessary to know: the number of larvae; the average size of larvae; the algal diet required; and, the density in cells mL-1 of each algal species to be fed. Because algal species differ in size, we allow a correction factor to account for smaller or larger species (see below). These correction factors have been determined on the basis of the individual dry weights of each alga. Dry weight correction factors: T. Iso = 1, C. muelleri = 1, P. lutheri = 0.8, C. calc = 1.2. Total algal feed = Total No. of larvae x cells/larvae/day (from feed curve) Algal species = Dry weight x Total algal feed No. of diets (*) x Algal cell count



Example: Larval feed calculation Scenario: Calculate the amount of algae required to feed 7.80 x 106 larvae that are 162µm in size. The larval diet needs to be comprised of three algal species (P. lutheri, T. Isochrysis and C. calcitrans), all of equal quantities (33% of total daily feed for each algal species). Algal cell counts:

P. lutheri = 7.65 x 106 cells mL-1

T. Iso = 8.15 x 106 cells mL-1

C. calc= 5.45 x 106 cells mL-1

Dry weight correction factors:

P. lutheri = 0.8, T. Iso = 1, C. calc = 1.2 Algal feed ration from growth chart (Fig. 6.7) = 8200 algal cells/larva/day Total algal feed = Total No. of larvae x cells/larvae/day = 7.8 x 106 x 8200 = 64 x 109 cells of algae Algal species = Dry weight x Total algal feed No. of diets (*) x Algal cell count Total Pav = 0.8 x ( 64 x 109 ) / 3 x ( 7.65 x 106 ) = 5.1 x 1010 / 2.3 x 107

= 2226 mL Total T. Iso = 1 x ( 64 x 109 ) / 3 x ( 8.15 x 106 ) = 6.4 x 1010 / 2.4 x 107

= 2667 mL Total C. calc = 1.2 x ( 64 x 109 ) / 3 x ( 5.45 x 106 ) = 7.7 x 1010 / 1.6 x 107

= 4813 mL * Divide each of the total feeds by two to determine the morning and evening feeds. Appendix 2 has another worked example for calculating the daily larval feed ration.



6.6. Hatchery time-line for SRO production

8 – 10 weeks before spawning

The first step in production is the collection of mature broodstock. At the very least this collection occurs at least one week prior to spawning and the stock are placed in a conditioning system (regardless of the time of year) for quarantine purposes and assessment of their reproductive condition. If the larval run is planned for winter or spring, broodstock may require a conditioning period of up to 8 – 10 weeks before the spawning date. 1 – 2 weeks before spawning

Preparation within the hatchery starts 1 – 2 weeks before the designated spawning date. Hatchery dry-out procedures are implemented ensuring complete dry-out of the hatchery for a period of at least one week. During this time, all seawater delivery lines and valves are dismantled and cleaned, water storage tanks are cleaned, larval rearing tanks and valves are dismantled and cleaned, and hatchery floors are disinfected with chlorine solution. Preparation starts in the algal culture room. The inoculation of the cultures to be used for larval food usually commences about one week before the spawning date, but will vary with algal species and the growth rates being experienced. The temperature controlled rooms at PSFC are set to 26.5°C two days before spawning. This reduces the need for constantly leaving immersion heaters in the larval rearing tanks once the larval run starts as the room temperature maintains the water temperature at a constant 25°C. At PSFC the water storage tanks are filled and settled for one week prior to use. Before spawning

The day before the spawning the seawater delivery system lines are reconnected and fresh seawater is pumped through the lines to flush the system. The old set of 1µm filter bags are replaced with a new set and the incubation tanks are filled with 1µm filtered seawater from storage tanks. Immersion heaters are placed into the incubation tanks and the water is heated to 26°C. The tank is gently aerated and the airlines and ceramic weights (used to hold the airlines down in the water) are autoclaved before use. Broodstock are taken out of the conditioning system, cleaned using a Providone-Iodine antiseptic solution, chipped, scrubbed and left to dry overnight. Spawning

On the day of spawning the broodstock are arranged on the spawning table and thermal spawning induction techniques are applied. This thermal process is repeated until spawning occurs. Spawners are immediately placed into individual containers of seawater and allowed to continue spawning. Selected eggs are washed through a 63µm screen with 1µm filtered seawater, pooled into a 20L bucket, homogenised and counted using replicated 1ml samples. Selected sperm is pooled and assessed for motility. The sperm are added to the eggs at a ratio of one sperm visible per egg and fertilisation is conducted. Replicated 1ml samples are taken 15 minutes later to ensure fertilisation has commenced after which incubation tanks are stocked. At PSFC larvae (regardless of the volume of culture water) are initially stocked at between 10 – 15 embryos/mL with gentle aeration. A half daily feed ration (calculated from the daily feed curve) of mixed algal species (Pav, T. Iso, C. cal) is usually added after 16 hours so that larvae can commence feeding as soon as they are



capable. Filter bags from the seawater delivery line are removed at the end of each day and placed into a chlorine bath for a 24 hour period (See Section 2.2). It is recommended that the hatchery have at least two sets of filter bags so that one set can be used while the other is chlorinated. Day 1

Larval rearing tanks are filled using 1µm filtered seawater as close to the time of water exchange as possible. In summer this can be done immediately prior to the first water exchange on the newly developed larvae. However in winter it can take hours for the heaters to warm the tank and thus tanks are some times filled and heated the evening prior to water exchange. Heaters do not remain in the tanks with the embryos or larvae and are removed as soon as the desired temperature is reached. The fresh tank of seawater is gently aerated. As a general rule a 10% feed ration is fed immediately upon arrival on the following morning. This is also done on subsequent water exchange days; to ensure that algae remain readily available to larvae as the water exchange takes place. Before exchanging water, a sample of the larvae at 18 – 20 hours is collected. Microscopic examination is used to determine the extent of larval development (“D” stage in 16 hours is normal), size, gut content, motility and general health. If greater than 80% D-veliger stage larvae are present, the incubation tank is gently dropped onto a 35µm screen. The larvae retained on the 35µm screen are then washed into a known volume of 26ºC seawater (usually a 20L bucket) and 2 x 1ml samples are taken. The larval samples are used to determine the total number of larvae present as well as larval size and percentage development. (Refer to Section 6.2 for Larval Sampling and Measuring Techniques). NB. Draining large tanks (20,000L) can take some time, therefore, batches of larvae are collected from the screens and counted periodically (about every 30 minutes) and then placed into the new tank of seawater. This is done to prevent the larvae remaining on the screens for protracted periods of time. Two samples of approximately 100 larvae are collected each water change. One is fixed in 10% formalin and the other in 70% ethanol. These samples are stored in case of problems so that an archive of larvae is available for health testing and pathology. The SRO feed curve (Figure 18) is used to calculate the total daily feed ration required. Divide the total daily feed ration into morning and evening feeds. The morning feed ration is fed to the new larval rearing tank before the tank is stocked with larvae. At this point and throughout the larval run, the mean larval size of the batch can be compared to the mean of all SRO run’s that have been conducted at the PSFC since 2003 (Figure 21). This will give an indication of the relative performance of the larval batch.



Figure 21. The mean larval growth of Sydney rock oyster production runs conducted since

2003. Once the density of the larvae from the incubation tank is obtained, the new larval rearing tanks are stocked at a density of 8 larvae/mL. The empty incubation tank is then washed out with freshwater and cleaned using Virkon S solution. The tank is then scrubbed and rinsed using freshwater before being left to dry for at least 24 hours. Air lines and air stones from the incubation tank are placed into a chlorine bath (0.25mL of chlorine/L of freshwater) for 24 hours, washed with freshwater and hung up to dry for a further 24 hours. Before leaving, the evening feed ration is added to the tank, the water temperature is checked and a larval sample from the top of the water column is taken to ensure that the larvae look healthy. Day 2

Larval water exchanges are made every second day. On arrival in the morning the larvae are fed immediately using the quantity of food supplied in the previous evenings PM ration. If the larvae are growing normally this will be slightly less than required, but this can be adjusted once larval samples have been taken and growth has been determined. Collect larval samples and inspect live larvae before they are fixed for measurement. Calculate the required quantity of algae and make any adjustments to the morning feed that are required. Do not be concerned if you have overfed slightly, simply account for this in the evening feed. Feed the balance of the daily feed ration as late in the afternoon as possible and take a small sample of larvae to inspect their health. In practice feeding usually occurs at 7:00 – 8:00 am and 5:00 – 6:00 pm.



Day 3

A new larval tank is filled, heated and aerated (as Day 1). Add 10% of the previous evenings feed ration to the larval tank before starting the water exchange. Commence batch water change using a 55μm primary screen and a 35μm back-up screen. The larvae are washed from the screen into a 20L bucket and sampled to establish total numbers, gut content, development, motility and general health. The known number of larvae is stocked into the new tank to which 50% of their daily feed ration has been added. The balance of the daily feed ration is added in the evening in accordance with the feed curve. The used larval rearing tank is washed out and all equipment is cleaned using the protocols described in Day 1. Days 4 – 6

Days 4 – 6 are as per the protocols described above in Days 2 – 3. The water change for Day 5 is normally done using a 65µm screen with a 55μm backup. These screen sizes are only a guide. Larval growth rates differ; therefore the screen sizes used on each water change should be chosen in accordance with the size of the larvae and following consultation of the screen size chart (Table 1). It is important to note the diagonal measurement for each screen mesh as this determines the size of larvae that can pass through the screen. Stocking densities at this time are usually still between 5 – 7 larvae/mL, although by Day 7 stocking densities are reduced to approximately 4 larvae/mL by restocking the largest larvae (retained on the primary screen). Days 7 – 18

The above pattern regarding water changes, feeds, daily larval samples and sanitation is continued throughout the larval cycle until settlement. From Day 7 (or when larvae exceed 100 µm in size) to Day 10 the proportion of C. calcitrans in the diet is incrementally reduced (10% per day) and replaced with C. muelleri. Usually at Day 7, when the larvae are approximately 120 – 130 µm in size, the first umbonate larvae are present (Figure 22).

Figure 22. Unbonate SRO larvae.



The development of an eyespot is another indicative characteristic seen at approximately Day 13 of the larval run as the larvae exceed 220 µm in size. The eyespot (Figure 23) is an important milestone in the larval development as it indicates to the technician that the larvae are approaching the pediveliger stage. Screen sizes are increased at each water change and are selected on the basis of the larval growth. Larval management becomes increasingly important as larvae approach settlement. Better settlement rates are obtained in larval batches that are very uniform in size. Day 18 – 19

Routine procedures as per Day 17 are conducted. It is common that at this time the numbers of crawling pediveligers are increasing (Figure 23). The extension of the foot can be observed whilst the larvae are swimming or stationary. It is very difficult to identify pediveligers using a sample that has been fixed using 10% formalin and seawater. Therefore it is suggested that a fixed sample and a live sample of the larvae are collected to clearly identify such behavioural characteristics.

Figure 23. Sydney rock oyster pediveligers. The eyespot is clearly visible and the foot is shown protruding from the mantle cavity.



Days 20 – 22

Water exchanges are made using 212µm screen 180µm backup. Larvae retained on the 212µm screen are washed into a 20L bucket and sampled to establish total numbers, development and general health. It is essential at these later stages that close attention is given to larval development. Strong indicators that the larvae are ready to settle are when more than 35% of larvae are crawling pediveligers and when there are between 3 – 5 gill buds (Chapter 7) present on 50% of the pediveligers. Larvae retained on the 212µm screen are then put to set using epinephrine bitartrate (refer to Chapter 7). If a significant number of larvae are retained on 180µm screen, they are restocked into a new larval rearing tank until they are large enough to be retained on a 212µm screen and the pediveliger larvae show signs that they are ready to metamorphose and settle as spat.



7. SETTLEMENT

7.1. Introduction

The term settlement refers to the metamorphosis of oyster larvae to spat. A number of physical characteristics are evident in SRO larvae indicating when settlement is imminent. The larvae have eyespots and have usually reached approximately 300μm in size. The foot has developed and a percentage of the larvae can be observed both swimming with their foot protruding and attempting to crawl on fixed surfaces or substrate. At this stage it is also important to monitor the development of gill buds in the larvae (Figure 24). At least three (preferably 4 – 5) rudimentary gill buds on the gill arch should be observed under the microscope at 100X magnification before settlement protocols are commenced. Gill buds are quite difficult to observe in a fixed sample, therefore it is recommended to observe larvae in live samples.

Figure 24. Recently settled Sydney rock oyster spat. Note, the newly formed gill arch and gill buds.

Two distinctly different methods have been used to settle SROs. Originally larvae were placed on screens with ground scallop shell and allowed to settle and metamorphose (Frankish et al. 1991). More recently this method has been abandoned in favour of epinephrine induced set techniques.



7.2. Epinephrine

Upon removal from the larval rearing tank, larvae are rinsed into a known volume of seawater and total numbers determined. For large quantities of larvae it is best if the first epinephrine treatment is conducted in a suitable tank. We use a 90L cylindroconical tank. Smaller quantities of larvae can be treated in a 20L bucket. The treatment of the larvae is carried out as follows and immediately following the treatment larvae are stocked into a downweller system at a density of 500,000 larvae per screen (Figure 25).

Figure 25. Pictures and components of a downwelling unit used to rear SRO spat.



The procedures used are a modification of those used for Pacific oysters (Crassostrea gigas) and have been designed to expose larvae at the same epinephrine concentration on each occasion. These procedures routinely provide settlement percentages of between 40 – 70%, depending on larval quality and the number of treatments applied. Because larvae within batches develop at different rates and are never uniformly ready to set at the one time, multiple treatments of epinephrine are necessary to obtain maximum spat yields. Some batches of larvae require up to five treatments with epinephrine. The frequency of the epinephrine treatments varies from run-to-run in accordance with the response of the larvae to the treatment. For this reason it is important to sample regularly between treatments to determine percentage set and to observe the progression of the remaining larvae. Before beginning epinephrine treatments check that the epinephrine is within its “use by date” and has been stored according to the manufacturer’s instructions. Epinephrine is purchased before each run; so that it is fresh when used. Old batches of epinephrine that are discoloured (brownish hue) are less effective in inducing settlement. Epinephrine is made up freshly as required. We do not make or store stock solutions. NB. Epinephrine is not freely available to hatcheries in Australia and guidance is required for its use. Hatcheries wishing to use epinephrine are required to join the National Aquaculture Council (NAC) and meet a series of guidelines and record keeping protocols for use. An advantage that epinephrine settled spat holds over shell settled spat is the ability for the spat to be segregated from the larvae earlier. The larvae/spat are graded over a 265µm screen within 1 – 2 days of the first epinephrine treatment, depending on percentage settlement. Spat retained on 265µm screens are placed onto a separate screen where they spend another two days in the downweller before being transferred into the spat bubbler system. 265µm retained spat are held in the downweller system for a further two days after being separated from larvae because the spat are still quite sticky and tend to clump together. Therefore, it is hard to prevent individual spat contained in the spat bubbler system from being ejected, due to the high flow rate which keeps spat fluidised. Larvae or spat that pass through the 265µm mesh are either re-treated with epinephrine or placed back on to a screen. Separating the spat from larvae improves hatchery efficiency, as spat can be placed into fluidised spat bottle systems earlier, where they grow faster than in downweller systems and, consequently, can be transferred to field nurseries sooner. An additionally benefit that arises from having epinephrine settled spat is that 180µm screens can be stocked at double the density of shell set spat. This significantly increases the number of spat a hatchery can carry in downweller tanks.



7.3. Procedures for the use of epinephrine

Chemical: Epinephrine Bitartrate Salt Sigma E-4375

C9H13NO3-C4H6O6

FW=333.3 General use of epinephrine: 1.2g of epinephrine per 50g of larvae in 20L of seawater in a conical tank for 60 minutes treatment. Method: Culture larvae until the percentage of crawling pediveligers is 15% or more and at least three gill buds (preferably 4 or 5) can be observed. Note: When retained on a 212µm screen, 1 million larvae are assumed to weigh 18g. First Epinephrine Treatment

1. Determine number of oyster larvae and then estimate their combined weight (1 million larvae = 18g).

2. Weigh out epinephrine based on 1.2g per 50g larvae and dissolve in small volume of fresh water.

3. In a cylindroconical tank, add 20L of seawater per 50g larvae. The seawater used should be at the same temperature as the larvae were being held (26ºC).

4. Turn on air, making sure that the bubbling effect of the water is not vigorous but turning the water column over.

5. Add larvae and epinephrine solution to conical tank. 6. Cover tank with black plastic. Epinephrine denatures in light, therefore the tank needs to be

covered with black plastic so that no light can penetrate onto the larvae (If possible carry out this procedure in a dark room).

7. Drain tank after 60 minutes onto a suitable screen (a 180µm screen is easier to work with than a 212µm screen).

8. Gently brush off any adherent larvae from the walls of the tank and add to the 180µm screen. 9. After treatment is completed give the larvae/spat a thorough wash with fresh seawater. 10. Add treated larvae to a 180µm settlement screen in the set system (250,000 – 500,000 larvae

per screen). 11. 2 hours after being placed onto screens the larvae/spat should once again be given a wash with

fresh seawater. A soft tipped brush together with a gentle fanned flow of seawater should be used to gently brush off any recently settled spat. This process stops spat from attaching to the screen material.

Repeated epinephrine treatments

We have found it necessary to treat Sydney rock oyster larvae with epinephrine several times to achieve the maximum percentage settlement. We have used two approaches based on the number of larvae to be treated. First, to reduce the number of oysters to be treated, the larvae are gently sieved on a 265µm screen to remove already settled spat, the numbers of remaining larvae are determined and they are treated as for Day 1. This sieving is optional on Day 2, as very few spat will have reached a size at which they will be retained.



Small numbers

1. For small numbers (a few set screens) we find it more convenient to treat the larvae on the screens.

2. Remove screens from set system and place them into a shallow tub (Preferably one of a similar size to the screen that has a flat bottom so that large volumes of epinephrine solution are not required to cover the screen).

3. Weigh out the quantity of epinephrine required using the same calculations as for Day 1. For example, to treat a single set screen with 500,000 larvae, 0.22g of epinephrine is used in 3.6L of seawater.

4. Add epinephrine solution to each screen and treat for 60 minutes (we have not found it necessary to aerate the shallow tub).

5. Cover the tub with black plastic (We also carry out this procedure in a dark room). 6. After the treatment is completed, rinse the screen gently and return the screen to the set system. 7. Two hours later, larvae/spat should once again be given a wash with fresh seawater as

described previously (refer to ‘11’ above). If there are large quantities of larvae to be treated on Day 2 or any subsequent day, it becomes more convenient to pool the larvae together and treat as for Day 1 using the conical tank. If it is necessary to treat larvae again on Day 4 or 5, use the Day 3 protocol of grading spat from larvae in conjunction with either of the Day 2 methods deemed to be necessary. It may be of some advantage at this stage of the treatment process to increase the exposure time of the larvae to the epinephrine solution although the total exposure time should not exceed 90 minutes. Example: Scenario: 2 million spat are being put to set using epinephrine bitartrate. Calculate the weight (g) of epinephrine that is required and the volume of water that the spat are to be treated in. Weight of larvae: If 1.0 x 106 larvae = 18g Then 2.0 x 106 larvae = 36g Epinephrine required: 1.2 g of epinephrine = 50g of larvae = (36 ÷ 50) x 1.2 = 0.72 x 1.2 = 0.864 g of epinephrine Volume of seawater for Epinephrine treatment: 20L = 50g of larvae = (36 ÷ 50) x 20 = 0.72 x 20 = 14.4L of seawater



7.4 Grading

At PSFC grading is conducted using a wet screen method. Larvae/spat are placed onto a desired screen size submerged in seawater and sieved gently. The larger size grade will be retained on the screen, whilst the smaller size grade will pass through. For SROs the first post settlement grading commences between one and two days after the initial epinephrine treatment. The initial grading for epinephrine treated spat is conducted using a 265µm screen. This screen size is useful in separating the metamorphosed spat from larvae. From this time grading should be carried out at regular intervals and spat should be placed on screens or in bottles specific to their size grade.



8. NURSERY

8.1. Downweller systems

SRO larvae are settled in a downwelling settlement system (Figure 25). This system has been designed to grow spat to a size of approximately 1000µm (retained on a 500µm screen) before they are transported to field nurseries. The downweller system consists of a fibreglass 1400L tank that holds ten 45cm diameter screens (Figure 25). Initially, ready to set larvae that have been treated with epinephrine are stocked on a 180µm screen at a density of 250,000 to 500,000 larvae. A spray bar provides flow to the larvae by way of a submersible pump located in the reservoir tank. This tank is aerated and a titanium 4KW immersion heater maintains the water temperature at 24 – 25ºC. Every second day the screens are transferred into a new system with fresh pumps, hoses, airlines and spray bars. The used tank and fittings are cleaned using Virkon S solution, before being rinsed with freshwater and allowed to dry. The feeding rates for larvae/spat held in downwelling systems start at approximately 40,000 – 50,000 cells/larva/day and are adjusted according to the rate of algal consumption (Section 8.4). After approximately two weeks the spat are large enough to be retained on a 500µm mesh. At this stage spat are removed from the hatchery to field nursery upwelling systems. It is worth emphasizing that in our experience at the PSFC, if water temperatures in the field are appropriate and the nursery site is well chosen, the move from the hatchery to the field promotes significantly faster spat growth than that of spat that are maintained in the hatchery.

8.2. Fluidised spat bottle system

The spat bottle is an alternative method to the conventional downwelling early nursery system. While it does not eliminate the use of downwelling systems completely it does offer an economical, low cost alternative method that enhances both spat rearing and production (Figure 26).



Figure 26. Fluidised spat bottle system. Inset shows close-up of spat in the base of the bottle. The spat bottle system was originally developed by John Bayes and was supplied to the PSFC by Rodney Grove-Jones (Grofish, South Australia). It involves one or more conical based bottles connected to a reservoir tank. Water from the reservoir is delivered via silicon tubing into a glass tube that extends to the base of each bottle. This water flow continually suspends the spat in the water column in a fluidised bed. The effluent from each bottle escapes through an outlet near the top of the bottle. Effluent water is directed into a common gutter before passing through a 200µm screen (to trap pediveliger larvae) and returns to the reservoir tank. There is an adjustable clamp placed on the silicon tubing at the top of the glass rod to control the flow rate to each bottle. Correct flow is critical. Insufficient flow can lead to reduced algal availability and mortality through overcrowding. A flow rate that is excessive can physically eject the spat from the bottles. The side of the bottle can be marked to show the total volume of spat which can be used to monitor spat growth. By marking or recording the volume of the spat on the side of the bottle, daily increments in growth can be readily observed. For more accurate growth measurements, turn the water flow off and allow the spat to settle before recording the measurement. Table 2 lists data from PSFC that allows an estimate of the number of spat to be made based on the height of spat within a spat bottle. \



Table 2. Estimation of spat numbers based on the height of spat within the spat bottle for the PSFC system.

Measurement Volume No. of spat (500 µm retained)

5cm 70ml 333200 6cm 105ml 499800 7cm 150ml 714000 8cm 210ml 999600 9cm 245ml 1166200

10cm 300ml 1428000 12cm 420ml 1999200 13cm 495ml 2356200 15cm 620ml 2951200

The algal cell density in the reservoir tanks is maintained at 100 – 150 x 10³ cells/mL so that the spat within the bottles are offered a constant supply of algal feed. The flow rate is adjusted to expel spat feeding and faecal wastes from the bottle. The bottle system promotes faster growth rates of SRO spat than downwellers and reduces the time taken to grow spat to a size suitable for field nurseries. In comparative trials, the average growth rate of spat (measured as shell height) reared in the bottle system was 7.4% per day compared with that of spat reared in downweller system screens at 5.0% increase in shell height per day. The spat bottle system is relatively inexpensive to construct and requires minimal floor space, although the system uses more seawater and food than the equivalent downweller system. For this reason, we have adapted the spat bottles to run on partially recirculated water to increase the efficiency of algal and seawater use which makes this type of rearing system more suited for use at the PSFC hatchery. Bottle systems are easy to clean and maintain. Because the spat are continuously suspended throughout the fluidised bed, fouling of the spat is minimised and they are markedly cleaner than those spat held in a downweller systems. This however does not reduce the need for regular (daily) thorough cleaning of the system. Spat can be transferred to the bottle system as soon as they can be separated from the pediveligers, usually when they exceed 350µm in size. It is difficult to remove all pediveligers at this time, even with stringent size grading. Those pediveligers that are inadvertently stocked into spat bottles are usually rapidly lost from the bottles in the effluent water stream and can be recaptured on the 200µm back-up screens. Stocking density for each bottle depends on the size of the spat and the water flow rate required to keep the bed of spat fluidised. Generally we have initially stocked about two million spat per bottle and reduce the density as they grow. While it is not possible to be prescriptive about flow rates for the bottles Table 3 provides an example of the relationship between flow rate, density and size for each spat bottle at PSFC. Table 3. Flow rates used for SRO spat at various sizes and densities in the spat bottle system at

PSFC. Screen mesh size on which spat

are retained Flow rate for 500,000 spat Flow rate for 1 million spat

300µm 1.27 L / min 2.25 L / min 500µm 2.0 L / min 2.45 L / min 670µm 3.1 L / min N/A



8.3. Daily protocols for the spat bottle system

Operational protocol

1. Adjust hose clamp to halt the seawater flow into the spat bottle. 2. Take the glass tube gently out of the spat bottle. 3. Take empty bottle out of clamp on the bottle rack. 4. Place bottle into bottle holder. 5. Place a plastic funnel over the top of the bottle. 6. Stock spat into the spat bottle. We usually stock approximately two million 500µm retained

spat into a single bottle (maximum: 4 million spat per bottle). 7. Remove bottle from bottle holder and mount back into position on the bottle rack. 8. Release the pressure on the hose clamp to allow water flow through the glass tube. 9. Place glass tubing into the spat bottle. Note: there should NOT be any back pressure on the

glass tube as it is placed into the bottom of the bottle. If the glass tube does not pass through the spat easily adjust the clamp to increase the water flow.

10. As the bottle fills the movement of the spat looks like a “rolling boil”. This movement should propel the spat upwards into the water column, before they gently sink and slide back down the shoulder of the bottle. The spat are then propelled upwards again and this process is repeated over and over.

11. The flow rate will need to be adjusted on a daily basis according to the size of the spat. As spat size increases, the flow rate is increased. If the flow rate is adjusted correctly faeces should be visible in the water column. This is also a good indication that the spat are feeding well.

12. It is suggested that on a daily basis the water flow be turned off and the spat allowed to settle to the bottom of the bottle. Using a whiteboard marker, mark the volume of spat. Doing this daily will give an indication of spat growth. As the spat grow it is not uncommon to observe a 2 cm increase growth over a 24 hour period.

13. AM: Check for growth overnight and adjust the water flow. 14. PM: Mark the height of spat on bottle as well as the date. 15. Spat are cleaned daily. The spat are removed from the bottles, placed on a screen (180µm)

then rinsed with freshwater. Clean the bottle thoroughly with freshwater and a soft tipped bottle cleaner. It is important not to scratch the plastic sides of the bottles as this reduces the visibility into the bottles.

16. A sub-sample of spat should be removed from the bottle and examined and sized using the microscope every day.



8.4. Diets and food rations for juvenile SRO spat

8.4.1. Species composition of the diet and calculating food ration

While undergoing metamorphosis in the downwelling system the larvae/spat are undergoing a great deal of physiological change. It is not uncommon for feed consumption to temporarily stabilise or reduce slightly. At this stage, continue to calculate feed rations according to the larval feed equation (usually 40,000 to 50,000 cells/larva/day), but more importantly, visually inspect tanks to assess whether the feed rate is excessive or needs to be increased. With experience it is possible to determine the need for increased or decreased feed rations by the colour and clarity of the culture water. In the first few days after settlement the spat diet is the same as that used for the latter stages of the larval cycle. After this time period, it is recommended that the percentage of diatoms (e.g., C. muelleri) in the diet is slightly increased and that other species, such as T. chui, are introduced to comprise 10% of the total daily food ration. Spat are fed in the morning and evening of each day. The period after spat settlement is the stage of the run at which the most algae is consumed, thus it is important to ensure that algal production is ready to cope with consumption when large numbers of spat are held in the hatchery. Progressively, as the larvae metamorphose and the spat grow, the feed rate is adjusted so that the algal cell density in the culture water is between 100,000 to 150,000 cells/mL when added to the tanks in the morning and afternoon, dependant on food consumption rates.



9. TRANSPORT

9.1. Transportation of spat

Hatchery production invariably involves transportation of spat from hatcheries to field nurseries and requires considerable care. Prior to this research program, sudden mortalities of spat in field nurseries were common and severe. During the three years of this research and the transport of tens of millions of spat to field nurseries, the only two mortality events that occurred happened immediately after transport. The size of SRO spat transferred from hatcheries is determined by grading using mesh screens and as a general rule only oysters retained on 500µm mesh and above are sent to field nurseries. This does not necessarily reflect any weakness on the part of the spat. Rather it reflects the difficulties of maintaining fine mesh screens in a field based nursery. For transport, spat are removed from water, rinsed and then retained on pieces of fine mesh (voile or other suitable material can be purchased cheaply from haberdasheries for this purpose) as shown in Figure 27. The mesh is then bundled up and the top fixed with an elastic band before being placed on a layer of moist (not wet) towelling in an insulated foam box. Transportation times vary with the proximity of the nursery, however we endeavour to keep emersion times to a minimum and always less than 24 hours. During transport care is taken to hold the temperatures within the box between 16 and 22°C. Temperature data recorders are used from time-to-time to monitor transport temperature. However, an inexpensive and useful alternative is to insert a thermometer through the lid of the box such that the bulb is reading internal temperature adjacent to the spat, while the temperature scale can be read from outside of the box. Air-conditioned vehicles are used for transport so that the chance of variation beyond the desired temperature range is reduced. The foam boxes are kept closed while in the car for a prolonged period as air-conditioners reduce the humidity and a moist humid atmosphere is preferred during transport. Spat are also frequently transported by air although again care is needed. If the spat are being freighted it is important to ensure the cargo hold is pressurised. Given the recent reduction in internal airfares it is sometimes viable to send someone to pick up the spat and carry them as hand luggage, ensuring their care.



Figure 27. Spat being placed on mesh for transport.

9.2. Transportation of broodstock

Although SRO broodstock are very robust and tolerant, the transportation of ripe adult oysters should be undertaken as quickly as possible to minimise stress on the oyster and to reduce the chances of the transport acting as a trigger for spawning. Adult oysters are transported in a similar fashion to spat (Section 9.1). All broodstock sent from the Oyster Hatchery at PSFC are chipped and cleaned of any fouling and scrubbed in a Providone-Iodine solution. Broodstock are then packed (usually) in a polystyrene foam box that has either wet paper towel or newspaper lining the bottom of the box. The oysters are then placed in the container and wet paper towel or newspaper is used to cover the oysters. If needed, an ice-brick can be taped to the lid of the box to keep the oysters cool, although if this is to be done, a layer of cardboard is placed over the oysters so that they do not come into direct contact with the brick. The preferred transport temperature for broodstock is generally between 16 – 22°C and transport times are kept to a minimum, usually less than 24 hours. It is a good practice to speak with the courier company about any special requirements they have for packaging and transporting live oysters to avoid valuable broodstock becoming stranded in transit.

9.3. Counting spat for delivery

It is not physically possible to count the number of spat produced from a larval run. However, the number of spat can be estimated by passing spat through a series of mesh screens, counting the number of spat that occupy a known volume unit, then measuring the total volume of spat in that size grade. The “voluming” of spat is an essential husbandry tool in determining the number of spat on a screen, in a nursery system or the volume required to fulfil a particular order. Generally, counting



is left until the spat are large enough to be retained on a 500µm screen. Prior to this, estimates of the numbers of spat on particular screens are made based on the number of larvae stocked and the percentage that have metamorphosed. Because spat numbers are determined by volume and are extrapolated from a table developed over years of SRO production (Table 4), it is important that the spat are graded appropriately. The first step in determining numbers is to grade spat into tight size ranges. These ranges are determined by the grading screens available to us at the PSFC and are listed below. The spat from each size range are placed in a measuring cylinder in water and the cylinder is gently tapped until the spat pack down and any further tapping does not reduce their volume. The volume of spat is recorded and then their number is determined from Table 4. In 2007, The Select Oyster Company Pty Ltd (SOCo) in conjunction with NSW DPI and FRDC produced a DVD entitled “Handling Fast Growing and Disease Resistant Oyster Spat”. This DVD shows in detail the methods used to accurately grade and estimate the number of hatchery and nursery sized spat. Further information about this DVD is listed in the reference section of this manual. Table 4. SRO spat/ml for different size grades settled using epinephrine bitartrate.

Screen (µm) Epinephrine Set (No. of spat/ml) 500 4760 670 2122

1000 1262 1250 672 1400 478 1800 240 2000 145 2240 68.4 3000 34.4 4000 19 7000 5.37

10,000 3.84 For example, the approximate number of spat in a 500ml sample that has passed through a 670µm screen and has been retained on a 500µm screen is 2.38 million. Notes:

• These numbers are conservative estimates. • These numbers assume that the spat have been settled using epinephrine. • Spat grow rapidly, so they should be volumed immediately after grading, not at some later

date.



10. HEALTH

10.1. Introduction

At PSFC larval and spat sampling for disease is an ongoing process, undertaken routinely with each run. Disease sampling commences prior to fertilisation with egg samples being taken, and continues through until the spat are transported into the field. For each larval run a complete catalogue of samples are kept so that in the event of reduced growth or mortality, the samples may be analysed to investigate possible causes. Most commonly, samples are taken at two-day intervals usually coinciding with water change days. However, if there is any suspicion of ill-health or any sign of mortality, sampling is increased to a daily basis or as needed. At PSFC two methods of sampling for disease are commonly used, and reflect differing analytical procedures used to identify problems. Samples are collected for histopathology (the sectioning, staining and diagnosis of diseased tissue using microscopy) and Polymerase chain reaction (PCR) testing.

10.2. Methods for sample collection

Fixation and storage of eggs/larvae for future examination using Polymerase Chain Reaction (PCR):

Collect 100 – 400mg (or 0.1 – 0.4mL) wet weight of eggs/larvae on sampling sieve. Rinse thoroughly with ammonium formate solution (63g of ammonium formate in 2L of distilled water). Store samples in absolute ethanol in labelled 5mL screw cap vials.

Storage and fixation of eggs/larvae samples for future sectioning for histopathology examination:

Collect 1 – 2000 larvae on sampling sieve.

Fix and store in 5mL labelled vial in 10% formalin and seawater (Section 10.4)

10.3. Sample investigation

If you have larvae or spat health concerns samples need to be submitted to a Regional Veterinary Laboratory and, if in NSW, notify the Animal and Plant Biosecurity Branch, NSW DPI (PH: 02 4982 1232). Veterinarians may not always be able to provide definitive answers as to the exact cause or causes of the problem; however, they may be able to rule out particular infectious diseases. The cost of testing ten samples is around $200.

10.4. Formaldehyde/formalin and ethanol

Chemicals used to preserve spat are toxic and hazardous at varying degrees. Use of these chemicals requires adequate training and full compliance with occupational health and safety requirements. For information on the specific risks, safe handling and safety equipment for these chemicals please read the appropriate Material Safety Data Sheet available from chemical suppliers or the internet.



Contact with eyes and skin should be prevented. Do not breathe fumes, use only in a well-ventilated area and always wear suitable protective clothing. Chemicals for fixation can be obtained from aquacultural supply companies, chemical suppliers, chemists or veterinarians. The recipe for 10% formalin and seawater solution is one volume of concentrated formalin (usually supplied as 37% formaldehyde solution) to nine volumes of clean, filtered seawater.




11. REFERENCES

Anderson, T.J. & R.D. Adlard. 1994. Nucleotide sequence of a rDNA internal transcribed spacer synonymy of Saccostrea commercialis and Saccostrea glomerata. Journal of Molluscan Studies 60:196–197.

Frankish, K.R., L.J. Goard & W.A. O’Connor. 1991. The development of hatchery rearing techniques of the Sydney rock oyster (Saccostrea commercialis) at the Brackish Water Fish Culture Research Station, Salamander Bay, NSW, 2301. 26 pp.

Heasman, M.P. 2004. Sydney rock oyster hatchery health workshop 8 – 9 August 2002, Port Stephens, NSW. NSW Fisheries Final Report Series: 61. Nelson Bay. 115 pp.

Helm, M.M., N. Bourne & A. Lovatelli. 2004. Hatchery Culture of Bivalves: A Practical Manual. FAO Fisheries Technical Paper No. 471. FAO, Rome 177 pp.

Lamprell, K. & J. Healy. 1998. Bivalves of Australia, Volume 2. Backhuys Publishers, Leiden, The Netherlands.

Nell, J.A. & W.A. O’Connor. 1991. The evaluation of fresh algae and stored algal concentrates as a food source for Sydney rock oyster, Saccostrea commercialis (Iredale and Roughley), larvae. Aquaculture 99:277–284.

Select Oyster Company. 2007. Handling Fast Growing and Disease Resistant Oyster Spat DVD. www.soco.com.au.

White, I. 2002. ‘Safeguarding Environmental Conditions for Oyster Cultivation in New South Wales.’ Report (Number 010801) for the NSW Healthy Rivers Commission.

http://www.soco.com.au/
















12. APPENDICES

12.1. Daily larval record sheet

Date………….. Batch No…………. Day……… Temp……… Larval Size…………. Predicted Size…………….. Observations……………………………………………………………………………………………………………………No. Larvae………………. Consumption (cells/larva/day)………………….. Total Consumption (cells/mL/day)…………………….. 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33

34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51

52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69

algae V = pcvf/C species size factor (f) date count(C) Proportion Vol (am) cells/mL Vol (pm) cells/mL Screen…… Backup…..

total


12.2. Larval feed calculation

Example: Scenario: Calculate the daily feed for 7.8 x 106 larvae that are 162µm’s in size. Then calculate the amount (L) of algae for each species using a ratio of 25% Pavlova lutheri, 25% Tahitian Isochrysis and 50% Chaetoceros muelleri. Algal cell counts: P. lutheri = 6.95 x 109 cells L-1 (N.B. counts are expressed in cells per litre in this example) T. Iso = 8.15 x 109 cells L-1

C. muelleri = 5.45 x 109 cells L-1

Dry weight correction factors: T. Iso = 1, C. muelleri = 1, P. lutheri = 0.8 Total algal feed = Total no. of larvae x Cells / larvae / Day Total algal feed 7.8 x 106 x 8200 = 64 x 109 or (6.396 x 1010) Algal species = Dry weight x Total algal feed No. of diets (*) x Algal cell count Total 25% Pav = 0.8 x ( 64 x 109 ) / 4 x ( 6.95 x 109 ) (N.B. units are L) = 51.20 / 27.80 = 1.84 L Total 25% T. Iso = 1 x ( 64 x 109 ) / 4 x ( 8.15 x 109 ) (N.B. units are L) = 64.00 / 32.6 = 1.96 L Total 50 % C. muel = 2 x ((0.67 x ( 64 x 109 ) / 4 x ( 5.45 x 109 )) (N.B. units are L) = 2 x ( 42.88 / 21.80 ) = 2 x ( 1.97 ) = 3.94 L * Note: When dealing with algal diets of differing ratios or percentages it is important to find a common denominator. See the following example: Example: 10% Pav, 40% T. Iso and 50% Muel. Then the common denominator would be 10 and the equations would be as follows. Pav = 1 x ( 0.8 x Total Algal Feed / 10 x Algal cell count) T. Iso = 4 x ( 1 x Total Algal Feed / 10 x Algal cell count) Muel = 5 x ( 0.67 x Total Algal Feed / 10 x Algal cell count)



Other titles in this series:

ISSN 1442-0147 (NSW Fisheries Research Report Series) No. 1 Otway, N.M. and Parker, P.C., 1999. A review of the biology and ecology of the grey nurse shark

(Carcharias taurus) Rafinesque 1810. 36pp. No. 2 Graham, K.J., 1999. Trawl fish length-weight relationships from data collected during FRV

Kapala surveys. 105pp. No. 3 Steffe, A.S., Chapman, D.J. and Murphy, J.J., 1999. A description of the charter fishing boat

industry operating in the coastal and estuarine waters of New South Wales during 1997-98. 33pp. No. 4 Reid, D.D. and Smith, I.R., 1998. The 1998 Pacific oyster survey. 14pp. No. 5 Walford, T. and Pease, B., 2000. Strategies and techniques for sampling adult anguillid eels.

Proceedings of a workshop held at FRI, Cronulla, Australia, August 1999. 176pp. No. 6 Heasman, M. and Lyall, I., 2000. Proceedings of the workshop held on 3 March 2000 at the

Sydney Fish Markets: Problems of producing and marketing the flat oyster Ostrea angasi in NSW. 57pp.

No. 7 Heasman, M., 2003. Proceedings of the Sydney Rock Oyster Hatchery Workshop held on 8 and 9 August 2002 at Port Stephens, NSW. 164pp.

No. 8 Allan, G.A., 2003. Proceedings of the Aquafin CRC Snapper Workshop held on 26 September 2002 at the Convention Centre, Melbourne (Aquafin CRC 2001/208). 107pp.

No. 9 Faragher, R.A., 2004. Hooking mortality of trout: a summary of scientific services. 9pp. No. 10 Daly, T., 2004. Summary of Proceedings from the Perkinsus Workshop held at the Cronulla

Fisheries Centre on 3 September 2003. 32pp. ISSN 1449-9959 (NSW Department of Primary Industries – Fisheries Research Report Series) No. 11 Baumgartner, L., 2005. Fish in Irrigation Supply Offtakes: A literature review. 22pp. No. 12 Ganassin, C. and Gibbs, P., 2005. Descriptions of the wildlife species that commonly occur in the

marine and estuarine waters of NSW. 88pp. No. 13 Nell, J., 2006. Manual for mass selection of Sydney rock oysters for fast growth and disease

resistance. 57pp + 110pp attachments. No. 14 Gilligan, D. and Rayner, T., 2007. The distribution, spread, ecological impacts and potential

control of carp in the upper Murray River. 25pp. No. 15 Baumgartner, L., 2007. Fish communities of the Nepean River in the vicinity of Pheasants Nest

Weir. 18pp. No. 16 Gilligan, D., 2007. Annual progress report towards achievement of the Lower Murray Darling

Catchment Action Plan 2004 – 2015: Fish Community Monitoring 2005/06. 42pp. No. 17 Gale, R., Silberschneider, V. and Stewart, J., 2007. A biological and economic assessment of the

2001 change in the Minimum Legal Length (MLL) of snapper in NSW. Report to the NSW Ocean Trap & Line Management Advisory Committee, December 2007. 43 pp.

No. 18 Rowling, K., 2008. Review of ‘Bobbin Gear’ in the NSW Ocean Trawl Fishery. 15 pp. No. 19 Baumgartner, L., Cameron, L., Faragher, B. and Pogonoski, J., 2008. An assessment of the trout

fishery in Oberon Dam and the Fish River. 23pp. No. 20 O’Connor, W., Dove, M., Finn, B. and O’Connor, S., 2008. Manual for hatchery production of

Sydney rock oysters (Saccostrea glomerata). 53pp.


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