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Batch-tagging blacklip abalone (Haliotis rubra) for identification of hatchery-reared individuals on natural coastal reefs in New South Wales, Australia.

ABSTRACT The identification of hatchery-reared larvae and juveniles is fundamental to assessing the success of their release when restocking. Hatchery-reared Haliotis rubra larvae and juveniles were successfully batch-tagged with distinct and persistent marks, enabling unambiguous differentiation from wild conspecifics when recaptured. Larvae were batch-tagged with the epifluorescent dye calcein. Experiments demonstrated that the batch-tagged larval shell was clearly visible in the spire of juvenile shells after 260 days. The recapture of batch-tagged and released larvae from natural reefs after 533 days at liberty also confirmed the persistence of this tag. A reliable and cost-effective method for batch-tagging juveniles was achieved with the use of an artificial diet that resulted in a distinctive blue-green coloration of the shell. This coloration differentiated released juveniles from wild conspecifics, was easily observed with the naked eye, and persisted on the spire of individuals for 777 days at liberty. These batch-tagging protocols allow large numbers of H. rubra larvae and juveniles to be distinctly tagged for long periods of time, enabling reliable estimation of survival after release and individual growth. It is likely that these techniques could also be applied to other abalone species.

KEY WORDS: tag-recapture, batch-tagging, restocking, abalone, calcein, Haliotis rubra


Measuring the success of releasing animals from hatcheries in wild populations is difficult, because reared individuals must be differentiated from wild stocks (Blankenship & Leber 1995, Hilborn 1998, Caddy & Defeo 2003, Bell et al. 2005). The success or otherwise of a number of stock enhancement or restocking programs has been difficult to quantify as a result of the inability to identify released individuals or their progeny. The inability to measure the success of releasing individuals into wild populations has raised concern over the continued investment of resources into such programs (Hilborn 1998, Hilborn 2004, Bell et al. 2005). However, many enhancement programs undoubtedly contribute individuals to the fishable stock and support extensive fishing industries (Chiu Liao 1999, Kitada 1999, Simizu & Uchino 2004, Werthemier et al. 2004, Bell et al. 2005).

Reliable identification of released and recaptured individuals with unambiguous differentiation from wild conspecifics requires effective tagging. Recapturing tagged individuals enable estimates of postrelease survival, and if the size at release is known, individual growth estimates can be determined. Tagging methods can be defined within 4 broad categories, including physical, chemical, electronic, and biological tags (Bell et al. 2005). Although alternative categories have been described (Rothlisberg & Preston 1992, Thorrold et al. 2002), the specific methods used for tagging individuals remain the same (Rothlisberg & Preston 1992, Thorrold et al. 2002, Bell et al. 2005). Physical tags can include identifying labels that are fixed to or inserted through external structures. Chemical tagging can range from stable isotope analysis to immersion in or injection of fluorescent compounds (usually antibiotics). Electronic tags can be as simple as implanting a passive integrated transponder tag, to the use of more expensive pop-up satellite archival tags. Biological tags can range from inheritable genetic markers and geographically distinct parasites to external morphological markings, either natural or imposed from diet, branding, or removal of some physical structure from an individual (e.g., tail or fin clipping).

Tags require a range of attributes to identify individuals and to determine the success of a restocking program (Rothlisberg & Preston 1992). Depending upon the objectives of the program, the tag must be able (1) to mark early life history stages; (2) to be retained and detectable in subsequent life history stages; (3) to be unique to the local population; (4) to be harmless to the tagged animal and any consumer, and be acceptable to the public; (5) to be inexpensive to mark and detect; (6) to allow rapid marking and detection; (7) to allow for multiple marking; and (8) to be transmitted to subsequent generations. Although all these attributes are highly desirable for any tag, only a subset of these attributes may be needed for some restocking objectives (see Rothlisberg & Preston 1992, Bell et al. 2005). For example, evaluating the contribution of released individuals to subsequent generations requires a heritable tag, but this is unnecessary when the objective of the study is to determine the survival of released individuals.

Many tagging methods focus on the unique identification (individually numbered or coded tags) of recaptured individuals and their survival, and several methods have been successfully applied to invertebrates (see Nielson 1992, Bell et al. 2005). However, tagging to identify individuals uniquely can often be problematic, because of extensive handling, and there are logistical difficulties if large numbers of individuals are required to be tagged. These issues are particularly relevant as they can affect the survival and/or growth of individuals released to investigate the efficacy of restocking local populations (McCormick et al. 1994, Shepherd et al. 2000).

Batch-tagging is an efficient method for tagging large numbers of individuals with a common tag that is added simultaneously to all individuals within a group. In addition to estimates of survival, this approach can also facilitate measurement of an individual's growth if the tag (1) can be incorporated into an actively growing structure (2) does not affect growth or affects it in a measurable way, and (3) persists in the individual through to its recapture.

Methods for batch-tagging marine invertebrate larvae or very young juveniles have been developed (see reviews by Levin 1990, Thorrold et al. 2002). In particular, immersion or injection with fluorochrome dye, such as calcein, has been effectively used to batch-tag a range of invertebrate larvae (Rowley & Mackinnon 1995), including molluscs (Moran 2000, Moran & Marko 2005). Fluorochrome dyes can be incorporated into calcified tissues, such as shell or bone, as they are deposited, providing a mark that is persistent and visible in the structure with the aid of an appropriate filtered light source (Day et al. 1995, Rowley & Mackinnon 1995, Collin & Voltzow 1998, Moran & Marko 2005). However, a number of studies have identified, or inferred, reduced survivorship and/or growth associated with tagging using fluorochrome dyes, including calcein (Brooks et al. 1994, Bumguardner & King 1996, Thebault et al. 2006).

Calcein has been used to mark the shell of larval Haliotis kamtschatkana in vitro (Collin & Voltzow 1998). Although Collin and Voltzow (1998) used the deposition of calcein in calcium structures to describe larval development, they did demonstrate that the mark is retained within the shell during settlement and metamorphosis, and that it remains embedded in the growing shell at the top of the spire. That calcein is deposited in the spire of the larval shell, and can be observed, provides a means of positively identifying recaptured individuals that were tagged and released as larvae. However, there are no data describing the effect of tagging larvae on their survival or growth, the persistence of such a tag in the natural environment, or the success of tagging larval or early juvenile Haliotis rubra.

Groups of juvenile or adult abalone have been batch-tagged either through a diet of varying alga (as observed by Leighton (1961), Olsen (1968), Hahn (1989), and Schiel (1993)), artificial feed (Gallardo et al. 2003), or immersion in or injection of fluorochrome dyes (Pirker & Schiel 1993, Day et al. 1995). Abalone fed artificial diets produce a distinct blue-green coloration of the shell (Gallardo et al. 2003), providing a distinct and persistent tag, that remains in the spire of the shell as the individual grows. Newly deposited shell deposition returns to a red-brown color, in common with wild conspecifics, after a natural algal diet is consumed (Gallardo et al. 2003), such as upon release to a natural reef.

The objective of this study was to develop and test methods that enable batches of larval and juvenile H. rubra to be reliably and consistently tagged, and subsequently identified on natural reefs over periods of time ranging from 1 day to more than 2 y. The ability to readily distinguish released juveniles enables effective assessment of restocking success, including postrelease survival and estimates of individual growth rate.


Batch- Tagging Larvae

Calcein Concentration and Immersion Time

A laboratory experiment was done to determine (1) the combination of concentration and immersion time that would pro vide a distinct and persistent chemical tag in the larval shell of individuals that could be detected after 6 mo and (2) whether calcein could be used to batch-tag H. rubra larvae without substantially affecting their survival and growth.

The survival of settled larvae and subsequent juveniles was observed, and the shell length of juveniles from treatments with the highest concentration and longest immersion time was measured. The persistence and level of epifluorescence was measured from the larval shell embedded in the spire of each juvenile. It was predicted that the levels of epifluorescence would be greater at higher concentrations and for longer immersion times. Furthermore, it was predicted that there would be no substantial effect of batch-tagging larvae with calcein on their growth as juveniles.

Developed larvae (6 days after fertilization) were added at a concentration of about 100 larvae/mL to 1 of 2 larval rearing tanks (180 L) containing calcein (product no. C8075; Sigma-Aldrich, Sydney, Australia) at a concentration of either 0.05 g/L or 0.1 g/L in sodium bicarbonate (0.1 g/[L.sup.1]) buffered seawater. The temperature of the seawater in the tanks was maintained at 19 [+ or -] 0.5[degrees]C. Larvae were held in each tank for 24 h and 48 h, with aeration and without water flow.

After 24 h and 48 h, approximately 3,000 larvae from each treatment tank were removed. These 4 groups (2 concentrations and 2 immersion times) of larvae were placed into separate, 14-L settlement tanks. Each settlement tank contained 12 polyvinyl carbonate sheets (150 X 200 mm), stacked on their edge and separated by about 30 mm. These sheets had been conditioned in aerated seawater for approximately 4 wk, with each covered by an adventitious algal biofilm able to support abalone larval settlement (Ebert & Houk 1984, Hahn 1989). Each tank had a constant flow of seawater at ambient temperature (~19[degrees]C). Settled larvae were grown on the plates until food was limiting (~75 days). They were then removed from the plates and returned to the 14-L tanks and fed a commercially available artificial abalone diet (Adam and Amos Abalone Foods Pty Ltd., South Australia; hereafter referred to as an artificial diet) for the term of the experiment (260 days). Twenty individuals from each treatment were haphazardly sampled at 15, 23, 30, 37, 56, 75, 97, 120, and 260 days after settlement. Each sample was preserved in 70% alcohol and stored in the dark. The spire of each shell in all samples was observed under a compound microscope with illumination via a 50-W reflected, ultraviolet light source through a blue-light (I2) filter block. This allowed observation of the calcein epifluorescence bound in the larval shell.

The level of epifluorescence was scored on a graduated, subjective visual index from 0-4. A score of 0 indicated no discernible epifluorescence and no positive identification of a batch-tagged larval shell in the spire of the juvenile shell, whereas a score of 4 indicated a bright and obvious mark with clear definition of the larval shell in the juvenile shell. Scores of 2 and more indicated unambiguous positive identification of calcein in the larval shell.

The effects of the factors calcein concentration (0.05 g/L and 0.1 g/L), immersion time (24 h and 48 h), and time (15, 23, 30, 37, 56, 75, 97, 120, and 260 days postsettlement) were analyzed using the data of the subjective scores of epifluorescence. Cochran's C test was used to determine the homogeneity of variances (Underwood 1997). Analysis of variance was used to test for differences among treatments.

Effect of Larval Batch-Tagging on Growth

The maximum shell length of each individual, from all samples taken from the treatment with the highest concentration and longest soak time, was measured. These data were compared with predicted lengths of juveniles of the same age, calculated from minimum and maximum daily growth rates reported for batches of abalone reared in the same hatchery facility and treated similarly, without having been exposed to a batch-tagging treatment as larvae (after Heasman et al. 2004).

Persistence of the Larval Tag in the Field

In addition to the laboratory experiment described here, juveniles were collected from 3 locations along the New South Wales (NSW) coast where batch-tagged larvae (calcein at 0.05 g/L for 48 h) had been released 553 days previously (Chick et al. in prep.). Any fouling on the spire of each shell was carefully removed using dissecting tools while viewing the shell under a dissecting microscope. The spire of the shell was examined for the presence of a calcein-labeled larval shell, as described for samples observed in the laboratory experiment and using image analysis software (public domain NIH Image program, U.S. National Institutes of Health, nih-image/).

Batch-Tagging Juveniles

Hatchery Growth of Juveniles

Juveniles used in field experiments were settled as larvae on plates and grown until reaching a length of about 1.5 mm. Once removed from plates, early juveniles were held in flow-through seawater tanks and fed an artificial diet. This diet resulted in the blue-green coloration of the juvenile shell. This shell color is distinctly different from that of the natural red-brown shell of wild conspecifics (pers. obs.; pers. comm., Scoresby Shepherd, South Australian Research and Development Institute, 2009). The relationship between the maximum length of abalone in the hatchery and the size of the batch-tagged shell at the time juveniles were released was described for batches of juveniles reared in the hatchery over the size range (2-15 mm)--i.e., the length of juveniles commonly used in field experiments investigating the restocking of H. rubra in NSW (Chick et al. in prep.).

Persistence of the Juvenile Tag in the Field

Juveniles reared in the hatchery and fed an artificial diet were released to 10 locations along the NSW coast. Abalone were recaptured from these locations after more than 750 days (Chick et al. in prep.). The presence and persistence of the blue-green shell in spire of these recaptured individuals was assessed to support the use of this method to batch-tag juveniles. It was predicted that the coloration of the hatcheryreared juvenile shell would persist in the spire of recaptured individuals. Furthermore, it was predicted that the coloration of newly deposited shell would change to a natural red-brown, enabling the individual size at release of recaptured abalone to be determined (length of blue-green shell), in addition to growth (red-brown shell) subsequent to release. Observation of the blue-green shell and the delineation between this and the subsequent red-brown shell in some cases required the removal of natural fouling.



Batch-Tagging Larvae

Calcein Concentration and Immersion Time

The calcein tagged larval shell was visible in the spire of more than 95% of abalone shells from all treatments investigated (Fig. 1). Higher and more consistent levels of epifluorescence were observed in the batch-tagged larval shell of juveniles when larvae were immersed for 48 h, irrespective of the concentration of calcein (Fig. 1). However, differences in epifluorescence among concentrations of calcein varied within immersion times (Table 1). There was no significant difference in the level of epifluorescence associated with the time (e.g., loss of epifluorescence) that abalone were sampled after settlement (Table 1). The larval shell could be easily distinguished in [greater than or equal to] 85% of the abalone treated at the lowest concentration of calcein (0.05 g/L) for the shortest immersion time (24 h). However, within these treatments at any one sampling time, up to 15 % of samples were faint, providing an ambiguous identification of the calcein tagged larval shell and attaining a score of 1.


Effect of Larval Batch-Tagging on Growth

The mean length of juveniles batch-tagged as larvae in a calcein concentration of 0.1 g/L for 48 h (i.e., the highest concentration and longest immersion time) was within the bounds reported for batches of juveniles reared in the hatchery of the same age and not subject to batch-tagging as larvae (24-50 [micro]m/day on plates and 50-100 [micro]m/day for the following year on an artificial diet [after Heasman et al. 2004]; Fig. 2). The mean length of juveniles sampled 75, 97, and 120 days after tagging indicates the growth rate during this time was close to the lower range of that predicted from juveniles not batch-tagged as larvae. However, by 250 days after tagging, the mean length of early juveniles batch-tagged as larvae was more medially positioned within the minimum and maximum predicted lengths for juveniles not batch-tagged as larvae.

Persistence of the Larval Tag in the Field

After 553 days, 9 juvenile H. rubra were recovered from 3 locations where batch-tagged larvae were released for investigation of the calcein-labeled larval shell. Five of the 9 individuals (56%) were positively identified, with scores of epifluorescence ranging between 3 and 4 (Fig. 3).

Batch- Tagging Juveniles

Hatchery Growth of Juveniles

As expected, there was a strong linear relationship between the length of the juvenile shell and time since settlement in the hatchery (Fig. 4). The length of the juvenile shell and hence the size of the tag at the time of release was a linear function of the time held in the hatchery. Juveniles held in the hatchery and fed an artificial diet attained a length of ~7 mm after 180 days and ~20 mm after 360 days.


Persistence of the Juvenile Tag in the Field

Hatchery-reared juveniles recaptured after more than 750 days were unambiguously identifiable from wild conspecifics, following the removal of any fouling of the shell (Fig. 5), as a result of the distinctive blue-green shell attained prior to release from the artificial diet (Fig. 6). Within 30 days of release, new increments of red-brown shell were clearly visible on hatchery-reared juveniles observed in situ at multiple release locations (pers. obs.).


A distinct and persistent tag enables individuals to be identified at a subsequent time, and this can be fundamental for any investigation in which individuals are released and recaptured for demographic studies, including an enhancement or restocking program for which survival and growth are used to measure its success. The objective of this research was to develop and test methods that enable batches of larval and juvenile abalone to be reliably and consistently tagged and subsequently identified when recaptured from natural reefs after an extended period. Such a tagging program objective would enable a comprehensive assessment of an ongoing enhancement or restocking program (Rothlisberg & Preston 1992, Bell et al. 2005).

Simple and reliable methods of batch-tagging H. rubra larvae were developed and tested. Batch-tagging of larvae and juveniles enabled released individuals to be positively identified and differentiated from wild conspecifics when recaptured. The methods described in this article to batch-tag larvae and juveniles addressed a substantial proportion of those attributes desirable for a tag to identify individuals for a stock enhancement or restocking program (Rothlisberg & Preston 1992). The results from this research demonstrated that the use of either method to batch-tag larvae or juveniles (1) enabled the earliest life history stages of H. rubra to be tagged, (2) provided a tag that is detectable in subsequent life history stages, (3) allowed released individuals to be differentiated from conspecifics, (4) was harmless to the tagged individual, (5) was relatively inexpensive, and (6) was relatively quick for tagging larvae (48 h). Detection of both larval and juvenile tags in particular was relatively efficient with very little sample preparation time required, because they could be detected in whole, unprocessed shells. An additional advantage of both these methods is that the noninvasive preparation of samples could be used to identify live animals and allow rerelease, despite the difficulties of requiring a compound microscope and light source to identify the calcein-tagged larval shell.


Persistence of the calcein-tagged larval shell in the spire of juveniles was demonstrated by unambiguous identification of the calcein tag in more than 95% of abalone treated in the laboratory experiment and was retained with a similar level of epifluorescence over time, for up to 260 days after settlement. Nevertheless, the higher levels of epifluorescence observed in the treatment where larvae were immersed for 48 h combined with the negligible difference between the level of epifluorescence among the different calcein concentrations within this treatment, together with a reduced cost, supports batch-tagging of larvae at a concentration of 0.05 g/L for 48 h as a standard protocol for larvae released to natural reefs to investigate their use for stock enhancement or restocking.


More important, the treatment of larvae with calcein did not appear to affect larval survival or settlement, and results indicate that there was no substantial difference in the growth of juveniles after having been batch-tagged at 0.1 g/[L.sup.-1] for 48 h. Batch-tagged larvae reared in the hatchery appeared to grow at a similar rate to individuals from other batches and not tagged as larvae, although the rates of growth prior to removal from plates was toward the lower range predicted from untagged individuals. Indeed, the growth rates for batch-tagged individuals were similar to those described for many other haliotids over similar times (see the review by Kawamura et al. 1998), and toward the higher range of those described for H. rubra by Daume et al. (2000).

It is likely that a proportion of batch-tagged larvae recaptured as juveniles was not positively identified. For example, if the larval shell had been physically worn from the spire of juveniles during the course of their life history, or fouling of the shell or removal of fouling during sampling damaged the spire, then the level of detection would be reduced. There is also some evidence that the intensity of a calcein tag can degrade through time (Leips et al. 2001) and when exposed to sunlight (Bashey 2004). Results from the work described in this article do not indicate any substantial reduction in the intensity of the calcein tag after 260 days postsettlement, although the hatchery-reared juveniles were mostly observed under shelters in the hatchery tanks, and hence out of direct sunlight. Furthermore, the positive identification of juveniles recaptured from beneath boulders on natural reefs after being released as batch-tagged larvae 553 days previously provides additional support to the value of this batch-tagging protocol and its application. These results add substantial support to the growing body of evidence regarding the efficacy of chemically tagging larvae for identification in stock enhancement or restocking programs, and ecological investigations regarding their distribution and abundance (see reviews by Thorrold et al. 2002, Moran & Marko 2005).

The artificial diet fed to hatchery-reared juveniles, removed from plates at ~1.5 mm, provided the opportunity to use the persistent and blue-green shell coloring generated by the diet as a batch-tag. Other significant advantages of this method included the ability to positively identify individuals in the field at the surface and underwater, and it enabled large numbers of very small juveniles (<5 mm) to be batch-tagged without excessive handling and subsequent high rates of associated mortality.


There was strong anecdotal evidence from short-term recaptures (<1 mo) of batch-tagged individuals at multiple locations (Chick et al. in prep.) that changes in shell coloration of released juveniles from blue-green to a natural red-brown occurred rapidly. There was scant evidence of the persisting blue-green shell eroding or becoming faint to preclude positive identification of hatchery released and recaptured abalone through time. However, although there is strong evidence for the success of this method of tagging over the timescales reported, further investigation is needed into the longer term persistence of this tag in the wild and the relationship between persistence and size of the tag. This is particularly important if small juveniles are the basis of any future restocking program. The ability to measure the size at release upon recapture (by measuring the maximum length of the blue-green portion of shell) provides a simple means to measure individual growth rates of released juveniles both at the surface and in situ.

It is possible that the conspicuous blue-green tag could increase susceptibility to visual predators (Catalano et al. 2001), which might compromise the use of this method for restocking. However, it is likely that the survival of tagged juveniles could be increased if they are released onto areas of reef where their exposure to visual predators is limited, possibly by the use of shelters, until they are able to disperse to more cryptic habitats (Tegner & Butler 1985, McCormick et al. 1994, Shepherd et al. 2000). It is also possible that such conspicuous coloration could be minimized by supplementing artificial diets with natural food sources or with specific nutritional additions to the diet so that a conspicuous single band of shell color or less conspicuous coloration of the whole shell could be used to tag batches. Furthermore, the manipulation of diet in the hatchery could provide a means of uniquely tagging different batches of juveniles prior to release.

A limitation of batch-tagging is that it does not allow for the identification of individuals that have been recaptured multiple times and does not, therefore, enable the determination of any sampling efficiency estimates (Dixon et al. 2006). For example, multiple recapture data from batch-tagging cannot demonstrate the survival of individuals that were not observed during initial sampling times, as is possible with tagging methods that identify each specific released individual. Batch-tagging is therefore unable to provide a scaling parameter to support the reassessment of initial estimates of survival, when some individuals may not have been observed, and therefore were not included in initial estimates of survival.

The tagging methods described in this paper provide the first documentation for successfully batch-tagging hatchery-reared larval and juvenile H. rubra to distinguish from wild stocks after an extended period. Although these protocols cannot provide a means to assess intergenerational effects of releasing individuals among wild populations (Rothlisberg & Preston 1992), they demonstrate that large numbers of larvae and juveniles can be reliably and consistently batch-tagged at relatively low cost within a short amount of time. Furthermore, application of these approaches can enable reliable estimation of survival and individual growth for abalone released onto natural coastal reefs.


Much of this work was supported through the Fisheries Research and Development Corporation (project no. 1998/ 219), the NSW Department of Primary Industries (Fisheries and Aquaculture), and contributions from the abalone industry in NSW, in particular Southern Ocean Seafoods (NSW) Pty Ltd and Twofold Bay Quality Bait Supplies Pty Ltd. The South Australian Research and Development Institute (Aquatic Sciences) provided valuable support for completion of this article. Peter Gibson, Craig Brand, Judy Upston, Duncan Worthington, and Craig Blount provided invaluable laboratory and field support. Mike Heasman, Nick Savva, Craig Brand, and Peter Gibson supported the rearing of all abalone. Valuable advice and comments on this research and improvements on the manuscript were provided by Michael J. Kingsford, Duncan Worthington, Cameron Dixon, Stephen Mayfield, Craig Blount, and an anonymous reviewer.


Bashey, F. 2004. A comparison of the suitability of alizarin red S and calcein for inducing a nonlethally detectable mark in juvenile guppies. Trans. Am. Fish. Soc. 133:1516-1523.

Bell, J. D., P. C. Rothlisberg, J. L. Munro, N. R. Loneragan, W. J. Nash, R. D. Ward & N. L. Andrew, editors. 2005. Restocking and stock enhancement of marine invertebrate fisheries. Adv. Mar. Biol. 49:243-286.

Blankenship, H. L. & K. M. Leber. 1995. A responsible approach to marine stock enhancement. Am. Fish. Soc. Symp. 15:167-175.

Brooks, R. C., R. C. Heidinger & C. C. Kohler. 1994. Mass-marking otoliths of larval and juvenile walleyes by immersion in oxytetracycline, calcein, or calcein blue. N. Am. J. Fish. Manage. 14:143-150.

Bumguardner, B. W. & T. L. King. 1996. Toxicity of oxytetracycline and calcein to juvenile striped bass. Trans. Am. Fish. Soc. 125:143-145.

Caddy, J. F. & O. Defeo. 2003. Enhancing or restoring the productivity of natural populations of shellfish and other marine invertebrate resources. FAO Fisheries technical paper no. 448. Rome, Italy: Food and Agriculture Organization of the United Nations. 159 pp.

Catalano, M. J., S. R. Chipps, M. A. Bouchard & D. H. Wahl. 2001. Evaluation of injectable fluorescent tags for marking centrarchid fishes: retention rate and effects on vulnerability to predation. N. Am. J. Fish. Manage. 21:911-917.

Chiu Liao, I. 1999. How can stock enhancement and sea ranching help sustain and increase coastal fisheries? In: B. R. Howell, E. Moksness & T. Svasand, editors. Stock enhancement and sea ranching. Oxford: Blackwell Science. pp. 132-149.

Collin, R. & J. Voltzow. 1998. Initiation, calcification, and form of larval "archaeogastropod" shells. J. Morphol. 235:77-89.

Daume, S., A. Krsinich, S. Farrel & M. Gervis. 2000. Settlement and early growth and survival of Haliotis rubra in response to different algal species. J. Appl. Phycol. 12:479-488.

Day, R. W., M. C. Williams & G. P. Hawkes. 1995. A comparison of fluorochromes for marking abalone shells. Mar. Freshw. Res. 46: 599-605.

Dixon, C. D., R. W. Day, S. M. H. Huchette & S. A. Shepherd. 2006. Successful seeding of hatchery-produced juvenile greenlip abalone to restore wild stocks. Fish. Res. 78:179-185.

Ebert, E. E. & J. L. Houk. 1984. Elements and innovations in the cultivation of red abalone Haliotis rufescens. Aquaculture 39:375-392.

Gallardo, W. G., M. N. Bautista-Teruel, A. C. Fermin & C. L. Marte. 2003. Shell marking by artificial feeding of the tropical abalone Haliotis asinina Linne juveniles for sea ranching and stock enhancement. Aquacult. Res. 34:839-842.

Hahn, K. O. 1989. Nutrition and growth in abalone. In: K. O. Hahn, editor. Handbook of culture of abalone and other marine gastropods. Boca Raton, FL: CRC Press. pp. 135-156.

Heasman, M., R. Chick, N. Savva, D. Worthington, C. Brand, P. Gibson & J. Diemar. 2004. Enhancement of populations of abalone in NSW using hatchery-produced seed. FRDC no. 1998/219. NSW final report series no. 62. Sydney, Australia: New South Wales Fisheries. 269 pp.

Hilborn, R. 1998. The economic performance of marine stock enhancement projects. Bull. Mar. Sci. 62(2):661-674.

Hilborn, R. 2004. Population management in stock enhancement and sea ranching. In: K. M. Leber, S. Kitada, H. L. Blankenship & T. Svasand, editors. Stock enhancement and sea ranching: developments, pitfalls and opportunities. Oxford: Blackwell Science. pp. 201-210.

Kawamura, T., R. D. Roberts & H. Takami. 1998. A review of the feeding and growth of post-larval abalone. J. Shellfish Res. 17:615-625.

Kitada, S. 1999. Effectiveness of Japan's stock enhancement programmes: current perspectives. In: B. R. Howell, E. Moksness & T. Svasand, editors. Stock enhancement and sea ranching. Oxford: Blackwell Science. pp. 103-131.

Leighton, D. L. 1961. Observations on the effects of diet on shell coloration in the red abalone Haliotis rufescens Swainson. Veliger 4:29-32.

Leips, J., C. T. Baril, F. H. Rodd, D. N. Reznick, F. Bashey, G. J. Visser & J. Travis. 2001. The suitability of calcein to mark poeciliid fish and a new method of detection. Trans. Am. Fish. Soc. 130:501-507.

Levin, L. A. 1990. A review of methods for labeling and tracking marine invertebrate larvae. Ophelia 32:115-144.

McCormick, T. B., K. Herbinson, T. S. Mill & J. Altick. 1994. A review of abalone seeding, possible significance and a new seeding device. Bull. Mar. Sci. 55:680-693.

Moran, A. L. 2000. Calcein as a marker in experimental studies newly-hatched gastropods. Mar. Biol. 137:893-898.

Moran, A. L. & P. B. Marko. 2005. A simple technique for physical marking of larvae of marine bivalves. J. Shellfish Res. 24:567-571.

Nielson, L. A. 1992. Methods of marking fish and shellfish. Am. Fisheries Soc. Spec. Publ. 23:208.

Olsen, D. A. 1968. Banding patterns in Haliotis. 2: some behavioural considerations and the effect of diet on shell coloration for Haliotis rufescens, Haliotis corrugata, Haliotis sorenseni and Haliotis assimilis. Veliger 11:135-139.

Pirker, J. G. & D. R. Schiel. 1993. Tetracycline as a fluorescent shellmarker in the abalone Haliotis iris. Mar. Biol. 116:81-86.

Rothlisberg, P. C. & N. P. Preston. 1992. Technical aspects of stocking: batch marking and stock assessment. In: D. A. Hancock, editor. Recruitment processes. Australian Society for Fish Biology Workshop, August 21, 1991. Canberra: Bureau of Rural Resources Proceedings No. 16. Australian Government Printing Office. pp. 187-191.

Rowley, R. J. & D. I. Mackinnon. 1995. Use of the fluorescent marker calcein in biomineralisation studies of brachiopods and other marine organisms. Bull. Inst. Oceanogr. Numero Spec. (Monaco) 14:111-120.

Schiel, D. R. 1993. Experimental evaluation of commercial-scale enhancement of abalone Haliotis iris populations in New Zealand. Mar. Ecol. Prog. Ser. 97:167-181.

Shepherd, S. A., P. A. Preece & R. W. G. White. 2000. Tired nature's sweet restorer? Ecology of abalone (Haliotis spp.) stock enhancement in Australia. In: A. Campbell, editor. Workshop on rebuilding abalone stocks in British Columbia: Canadian Special Publication of Fisheries and Aquatic Sciences. Nanaimo, British Columbia, Canada. National Research Council of Canada, pp. 84-97.

Simizu, T. & K. Uchino. 2004. Effects of extensive seeding on abalone, Haliotis discus discus, abundance on the pacific coast of Boso Peninsula, Japan. J. Shellfish Res. 23:1209-1211.

Tegner, M. J. & R. A. Butler. 1985. The survival and mortality of seeded and native red abalones, Haliotis rufescens, on the Palos Verdes peninsula. Calif. Fish Game 71:150-163.

Thebault, J., L. Chauvaud, J. Clavier, R. Fichez & E. Morize. 2006. Evidence of a 2-day periodicity of striae formation in the tropical scallop Comptopallium radula using calcein marking. Mar. Biol. 149: 257-267.

Thorrold, S. R., G. P. Jones, M. E. Hellberg, R. S. Burton, S. E. Swearer, J. E. Neigel, S. G. Morgan & R. R. Warner. 2002. Quantifying larval retention and connectivity in marine populations with artificial and natural markers. Bull. Mar. Sci. 70:291-308.

Underwood, A. J. 1997. Experiments in ecology: their logical design and interpretation using analysis of variance. Melbourne: Cambridge University Press.

Werthemier, A. C., W. R. Heard & W. W. Smoker. 2004. Effects of hatchery releases and environmental variation on wild-stock productivity: consequences for sea ranching of pink salmon in Prince William Sound, Alaska. In: K. M. Leber, S. Kitada, H. L. Blankenship & T. Svasand, editors. Stock enhancement and sea ranching: developments, pitfalls and opportunities. Oxford: Blackwell Science. pp. 307-326.


New South Wales Department of Primary Industries (Fisheries and Aquaculture), Cronulla Fisheries Research Centre of Excellence, PO Box 120, Cronulla, New South Wales 2230, Australia; School of Marine and Tropical Biology, James Cook University, Townsville, Queensland 4811, Australia

* Current address: South Australian Research and Development Institute, PO Box 120, Henley Beach, South Australia 5022, Australia. E-mail:
Summary of analysis of variance in the level of epifluorescence
of the calcein-tagged larval shell in the spire of abalone 15,
23, 30, 37, 56, 75, 97, 120, and 260 days after being treated
in 2 concentrations of calcein (0.05 g/L and 0.1 g/L) for 2
immersion times (24 h and 48 h).

Source                 df    Mean Square      F Ratio

Concentration (C)       1       21.01         89.35 **
Immersion time (I)      1      421.67      1,793.10 **
C x I x T               1       10.03         42.67 **
Time (T)                8        0.20          0.86 (ns)
C X T                   8        0.14          0.60 (ns)
I X T                   8        0.22          0.94 (ns)
C x I x T               8        0.34          1.46 (ns)
Error                 684        0.24

* P < 0.05, ** P <0.01 (Cochran's C = 0.0531, ns; variances,
36; df, 19);  ns, not significant.
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Author:Chick, Rowan C.
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:8AUST
Date:Apr 1, 2010
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