Capture Identification Culture Techniques

TRAINING COURSE REPORT

COMPONENT 2A - Project 2A1

February 2009

PCC development

Capture, identifi cation and culture Capture, identifi cation and culture techniquestechniques

of coral reef fi sh larvaeof coral reef fi sh larvae

Author: Viliame Pita WaqalevuAuthor: Viliame Pita Waqalevu

(French Polynesia)(French Polynesia)

Phot

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edit:

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LUA

Photos credit: Eric Clua (if no specifi c mention). Printed at SPC • Copyright : CRISP

CRISP is funded by the following partners :

CRISP Coordination Unit (CCU)Programme Manager: Eric CLUA

SPC - PO Box D5 - 98848 Noumea Cedex New Caledonia

Tel./Fax: (687) 26 54 71E-mail: [email protected]

T he Initiative for the Protection and Management of Coral Reefs in the Pacifi c (CRISP), sponsored

by France and prepared by the French Development Agency (AFD) as part of an inter-ministerial project from 2002 onwards, aims to develop a vision for the future of these unique ecosystems and the communi-ties that depend on them and to introduce strategies and projects to conserve their biodiversity, while de-veloping the economic and environmental services that they provide both locally and globally. Also, it is designed as a factor for integration between deve-loped countries (Australia, New Zealand, Japan and USA), French overseas territories and Pacifi c Island de-veloping countries.

The initiative follows a specifi c approach designed to:- associate network activities and fi eldwork projects;- bring together research, management and develop-ment endeavours;- combine the contributions of a range of scientifi c disciplines, including biology, ecology, economics, law and social sciences;- address the various land and marine factors aff ecting coral reefs (including watershed rehabilitation and management);- avoid setting up any new body but supply fi nancial resources to already operational partners wishing to develop their activities in a spirit of regional coo-peration. This is why the initiative was prepared on the basis of a call for proposals to all institutions and networks.

The CRISP Programme comprises three major compo-nents, which are:

Component 1A: Integrated Coastal Management and Watershed Management- 1A1: Marine biodiversity conservation planning - 1A2: Marine Protected Areas- 1A3: Institutional strengthening and networking- 1A4: Integrated coastal reef zone and watershed managementComponent 2: Development of Coral Ecosystems- 2A: Knowledge, benefi cial use and management of coral ecosytems- 2B: Reef rehabilitation- 2C: Development of active marine substances- 2D: Development of regional data base (ReefBase Pacifi c)Component 3: Programme Coordination and Deve-lopment- 3A: Capitalisation, value-adding and dissemina- tion of CRISP results- 3B: Coordination, promotion and development of CRISP activities- 3C: Support to alternative livelihoods- 3D: Vulnerability of ecosystems and species- 3E: Economic task force

The CRISP programme is implemented as part of the policy developed by the Secretariat of the Pacifi c Regional Environment Programme for a contribution to conservation and sustainable development of coral reefs in the Pacifi c

The CRISP Coordinating Unit (CCU) was integrated into the Secretariat of the Pacifi c Community in April 2008 to insure maximum coordination and synergy in work relating to coral reef management in the region.

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Ambassade de France à Fidji

UNIVERSITY OF THE SOUTH PACIFIC School of Marine Studies, Laucala Campus, Fiji

Capture, Identification and Culture techniques of coral reef fish larvae

(French Polynesia)

By Viliame Pita Waqalevu

Training course conducted from the 10th January to 15th February 2009

Supervisors: David LECCHINI (IRD - UR 128 CoReUs)

Christophe BRIE (Tropical Fish Tahiti)

Pascal DAYEZ-BURGEON (French Embassy at Fiji Islands)

Institut de recherchepour le développement

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REMERCIEMENTS

Au terme de ces deux mois de recherche, je tiens à remercier toutes les personnes qui, de près ou

de loin, scientifiquement, financièrement ou moralement, ont contribué à l'aboutissement de ce rapport.

Je désire remercier David Lecchini, Chargé de recherche à l'Institut de Recherche pour le

Développement (UR 128 Coreus) et Christophe Brié (Tropical Fish Tahiti) qui m'ont permis de réaliser

ce stage. Je leur sais gré de m'avoir fait confiance tout au long de ce travail.

Je désire aussi remercier Mr. Pascal Dayez-Burgeon, Conseiller de coopération et d’action

culturelle à l’Ambassade de France à Fidji. Sans son aide, ce stage n’aura jamais vu le jour. Vinaka…

Ce travail a été réalisé au Centre de Recherche Insulaire et Observatoire de l’Environnement à

Moorea. Je tiens à remercier très chaleureusement Serge Planes, Yannick Chancerelle et René Galzin

d'avoir entrepris de nombreuses démarches pour le bon déroulement du stage et m'avoir permis de loger

au Centre de Recherche. Un immense merci aussi à Pascal, Benoit et Franck pour leur aide de tous les

jours.

Je tiens enfin à remercier mes collègues de Moorea : Eric, Loic, Cécile, Jennifer, Elisabeth, …

pour avoir mis la bonne ambiance au centre de recherche.

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FINANCEMENT DE L'ETUDE

* Financement par l'ANR (Agence Nationale de la Recherche) : L'étude a été financée en partie par un

financement ANR Jeunes Chercheurs (ANR-06-JCJC-0012-01; D. Lecchini; Coral Reefs; novembre 2006

/ octobre 2009).

* Financement par l’Ambassade de France à Fidji : Mr. Pascal Dayez-Burgeon a financé le voyage de

l’étudiant Viliame Pita Waqalevu de Fidji à Tahiti.

* Financement par le programme CRISP (Coral Reef Initiative in the South Pacific) : L'étude a été

financée aussi en partie par un financement CRISP (Composante C2A, R. Galzin & D. Lecchini; janvier

2007 / décembre 2009). L’initiative pour la protection et la gestion des récifs coralliens dans le Pacifique,

engagée par la France et ouverte à toutes les contributions, a pour but de développer pour l’avenir une

vision de ces milieux uniques et des peuples qui en dépendent ; elle se propose de mettre en place des

stratégies et des projets visant à préserver leur biodiversité et à développer les services économiques et

environnementaux qu’ils rendent, tant au niveau local que global. Elle est conçue en outre comme un

vecteur d’intégration régionale entre états développés et pays en voie de développement du Pacifique. Le

CRISP est un programme mis en œuvre dans le cadre de la politique développée par le Programme

Régional Océanien pour l’Environnement afin de contribuer à la protection et la gestion durable des récifs

coralliens des pays du Pacifique.

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ABSTRACT

Fish larvae were sampled daily over 4 weeks using the crest net method on the Island of Moorea in the

Society Archipelago, French Polynesia. A total of 71 species were identified with a total abundance of

1204 larvae. Handpicked larvae were then cultured to juvenile stage in an aquarium and 67 fish larvae

photographed during larval stage and 16 larvae during juvenile stages, in order to observe the

metamorphosis between these two stages in the larvae’s life cycle. Larvae were fed three times a day with

artemia and artificial pellet feed during the duration of culture and mortality was minimized to <5%.

Light traps were also trialed on three different sites on two nights around the new moon on the North

Coast of Moorea in order to compare methods of larval capture and to observe which method yielded the

higher species richness and abundance. Upon completion of this study all larvae and juveniles were

released back into the lagoon.

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1) INTRODUCTION

In coral reef ecosystems, the life cycle of most fish species includes a planktonic larval phase (in

the open ocean), which usually lasts from three to six weeks, followed by a sedentary reef phase (in the

lagoon) for the juveniles and adults (for review, see Werner 1988 – Fig. 1). During the oceanic phase, the

larvae may move far from their native island due to currents and/or their swimming abilities. Then larvae

return to the reef (natal or not) to continue their development into juveniles, then to adults. Generally

larvae enter the lagoon across the reef crest by night (colonisation phase, Dufour and Galzin 1993). In the

hours following this colonisation, larvae undergo metamorphosis and choose suitable habitats (settlement

phase) based mainly on the characteristics of coral habitat and the presence or absence of conspecifics

(individuals of same species) as well as other species (for review, see Doherty 2002).

Figure 1: Description of life cycle of coral reef fish

Lagoon

Ocean

Life cycle of coral reef fish

Oceanicdispersion

Larvae

Genital products& eggs

Reproduction

Recruitment

Juveniles

Adults

Settlement

Colonisation

Lagoon

Ocean

Life cycle of coral reef fish

Oceanicdispersion

Larvae

Genital products& eggs

Reproduction

Recruitment

Juveniles

Adults

Settlement

Colonisation

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Thus, most coral reef fishes have a larval oceanic pelagic phase that ultimately ends with

settlement onto suitable benthic habitats on the reef. The success of the transition between the two

different environments determines the fish’s settlement levels (Dufour and Galzin 1993). There is a great

need to understand the complexity with which coral reef fish larvae choose an appropriate habitat. Studies

of similar scope need to be undertaken with greater enthusiasm as its ramifications greatly affect Pacific

Islanders and the natural stock of reef fish. As shown by the study undertaken by the University of the

South Pacific and University of Perpignan (PhD student, Julien Grignon), fish larval capture, culture and

release can be used as a tool for reef restocking, ornamental and aquarium trade or for food fish, local or

live fish exportation.

Colonization of fish involves the settlement of post-flexion larvae upon completion of pelagic

phase and the transition into juvenile involves some nature of habitat association within the lagoon. The

colonization process was observed to usually occur at night and in higher numbers during new moon

periods than in full moon periods (Lecchini et al. 2004). This process concerns mainly post-flexion larval

stage and juvenile stage (Dufour and Galzin 1993) as these are the larvae that are able to drift easily and

passively over the reef crest (Sale 1991). Therefore, the most suitable method that can be utilized to

capture these larvae is the crest net which actively filter larvae that are swimming in the water flowing

from the ocean into the lagoon. Crest nets have a number of advantages over other methods (light trap, net

either towed or dropped in the water column): (1) fish larvae are caught just before settlement, which

would give a suitable measure of larval flux and supply; (2) the high energy and turbulence of the reef

crest minimizes net avoidance by larvae; (3) since the net is put up for the night, the larvae cannot see the

net thus it is a passive gear for easy larval capture (Dufour et al. 1996). Moreover, in marine organisms

that have a relatively sedentary stage and dispersive propagules, the size of adult stocks is set by a

dynamic balance between input of propagules (larval colonisation) and their subsequent loss via death.15

In coral reef fish, the mortality of larvae having colonised may eliminate up to 90% of the total population

within three first post-colonisation months. The larval supply at colonisation represents a real natural

ichthyologic production of a fish adult stock, in number of individuals. As fish larvae stock is numerically

more important than adult stock and as catches of aquarium fish are based upon a number of individuals

(and less on biomass or size), it is preferable to encourage fishing pressure on larvae stock and rear them

with aquaculture methods to increase their survival. Effectively, in the wild, 90% of larvae disappear

before adult age. The adult breeding stock would be thus preserved and the colonisation rate would be the

exploitable theoretical limit not to be exceeded (over-fishing).

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This study aims to answer 4 questions: [1] How to capture larval fish using the crest net method.

[2] How to identify post-flexion fish larvae to the lowest possible taxonomic classification. [3] How to

rear larvae in aquarium using artificial feed whilst attempting to minimize mortality. [4] Comparing the

effectiveness of crest net to a light trap in larval capture.

The training course is in line with the Research Program of David Lecchini: “Effects of alternate

coral reef states on the attraction, settlement and subsequent survival of crustacean & fish larvae (French

Polynesia, 2006- 2010)”. The biodiversity is declining, and habitat destruction and degradation are now

commonplace. Examples of degradation can be found throughout marine ecosystems, including estuaries,

saltmarshes, soft-bottoms, hard-bottoms, and coral reefs. The degradation in coral ecosystems is usually

characterized by coral mortality from natural and anthropogenic stressors (e.g., disease, hurricane

damage, pollution, temperature-induced bleaching). This decrease of coral cover opens space on most

reefs and causes substantial increases in cover and biomass of rapidly growing fleshy and filamentous

macroalgae which, in turn, limits the recovery of coral populations and then modify fish and invertebrates

communities. Thus, areas experiencing perturbance often exhibit declines in adult populations, leading to

a higher rate of extirpation than in pristine habitat, and the persistence of species in the area becomes

reliant on the "rescue" effect of recruitment. The potential of the areas population to be supplemented by

recruits, however, depends on whether pelagic larvae detect an appropriate habitat in that area and then

settle and persist in that habitat. The effect of habitat decline on the potential of the area to attract and

sustain the faunal assemblage (i.e., recruitment potential) remains unclear. We do not know if the decline

of marine organisms is due to increased mortality of juveniles and adults of reef organisms or due to a

decrease in the degraded reef’s recruitment potential, which could decline if 1) its properties have

changed sufficiently to decrease its inherent attractiveness to planktonic larvae; 2) larval ability to locate

preferred microhabitats has declined; or 3) newly settled individuals’ ability to survive to recruitment has

decreased. We will test the hypothesis of effects of alternate reef states on the attraction, settlement, and

subsequent survival of crustacean and fish larvae. If the recruitment potential of some coral islands has

decreased, these meta-populations of reef organisms will continue their rapid decline, as recruitment will

not be able to replace and to sustain the adult populations on the reefs. The research program “Effects of

alternate coral reef states on the attraction, settlement and subsequent survival of crustacean & fish larvae

(French Polynesia, 2006- 2010)” is funded by ANR (French Government). Three axes make up the

program: Recruitment success of marine larvae according to the alternate coral reef states (axis 1) ; How

the alternate reef states influence the settlement rates and microhabitat selection of larval fishes and

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crustacean (axis 2) ; How alternate reef states influence the survival of larval fishes and crustacean (axis

3). My training course is a part of axis 1.

2) MATERIALS AND METHODS

a) Study area

Moorea Island is a high volcanic island, located in the Society Archipelago approximately 16 km

from the larger Island of Tahiti. There is a barrier reef that encircles the island which encloses a lagoon

800 to 1300m wide. The reef is intersected by several passes and two deep bays on the North coast (Cook

Bay & Opunohu Bay). The circulation of water is such that breaking waves bring oceanic water into the

lagoon and water within the barrier reef exists via the pass. The average annual tidal range at Moorea is

approximately 0.3 m.

Site suitability is an important aspect of choosing an area to place the crest net. There was only

one sampling site and this study site was located on the West coast of Moorea (17°31’7.38”S,

Figure 2: Moorea Island, French Polynesia (source: Google Earth)

149°55’20.89”W) between two Marine Protected

Area’s of Tetaiuo and Taotaha. This site was

chosen as the study site because wave energy is

low in this area and there is a deep region

immediately behind the reef crest for the nets

collector to be placed. The prevailing wind on

this side of the island was North Westerly. The

nearest pass is Taota pass located south from the

sampling area.

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b) Methodology

Fish larvae were sampled using a technique similar to Dufour and Galzin (1993). The crest net is

a technique that is most suitable to capture larvae upon entering the lagoon and just before settlement. The

net is 1.5 m wide x 0.75 m high x 5 m long and the mesh was of 1 mm size. Four hinged panels of 0.7

mm enlarged the mouth area of the net to 6 m (Fig. 3). The net was divided into three chambers: 1) the

mouth where larvae enter, and the codends which is made of 2 parts; 2) the separator is designed in such a

way that fish larvae are allowed to stay in sufficient water even if the water level is extremely low, it

decreases mortality due to dessication and low oxygen level greatly; 3) and the collector where the larvae

are captured. The whole structure is fastened

secured by steel cables which are bolted

firmly onto the reef-rock; this is to prevent

the net from being swept away during times

of strong current. The codend was attached

during the evening as larval settlement

usually occurs at night (Dufour and Galzin

1993).

Figure 3: Schematic view and photo of crest net used at Moorea

Upon collection of the larvae, the net is put in “off” mode, where the mouth of the net and its

wings are laid down on the reef. The codend is detached from its base attachment and is released from the

first chamber. It is important to ensure that no larvae are trapped within the first chamber before releasing

the codend. All the contents of the codend are then emptied into a large bucket/cooler where any excess

algae or rocks are removed. Oxygen is added at minimal amount in order to minimize stress and

mortality. The nets were left on the crest without the codend and with the wing and net placed in “off”

mode during the day.

Sorting of fish larvae was undertaken at the laboratory where fish were grouped into similar

Genera. Larval identification was undertaken using the meristics and morphology characteristics of the

fish with the CRISP larvae identification guide handbook (Galzin et al. 2006) as an aid. Pictures were

taken of all the fish larvae caught as this would become an important future aid in identifying larva and

juvenile (Annex 1 & 2). Alcohol was not used to kill the fish before taking pictures as it was observed

that alcohol removed some colors from the fish, thus, freshwater was used as an alternative method. Some

of the larvae were kept alive and moved to a larger more permanent glass tank for the rearing process into

juvenile stage.

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Larvae that were to be cultured into juveniles were fed artificial fish-food bought from Ridley’s

Aquafeed Limited and this consisted of pellets which were crumbled into pieces or sieved into really

small particles (Annex 2). This was supplemented with live feed in the form of artemia. The larvae were

fed three times a day, first with the pellets and then followed with artemia forty five minutes later, as the

fish will not eat the pellets if they are fed first with artemia. It was observed that the fish, at first, did not

eat the pellet however, after three weeks they started eating more of the artificial feed.

3) RESULTS

Fish larval capture with crest nets - A total of 1208 larvae belonging to 71 species were collected over

four sampling weeks. The number is relatively small due to the fact that big waves and rough weather

conditions were encountered often and this prevented the net from being put up and the current strength

often broke and separated the codend from the net. So, after 24 days of sampling, the average was 50.3

larval fish per day. The most abundant families were Acanthuridae (11 species) Pomacentridae (8 species)

and Chaetodontidae (6 species – see Annex 1). Figure 4 shows the colonization species richness of the

different larvae that were caught.

Figure 4: Larval species richness at colonization

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The most abundant species were Acanthurus triostegus (total abundance = 325), Stegastes

fasciolatus (204), Chromis viridis (71) and Chrysiptera leucopoma (65). The period that when there was

the highest species diversity during the new moon, suggesting that species richness was dependent on

lunar cycle. It was observed that fish larval colonization was also linked with the lunar cycle (Fig. 5).

` Figure 5: Larval colonization and lunar cycle

We observed a colonization peak just before and after the new moon, and another just prior to the

first quarter. This general trend of fish larval colonization varied at species level (Fig. 6a,b). For example,

Acanthurus triostegus and Stegaste fasciolatus were observed to colonize the reef throughout the 4

different lunar periods and thus had a constant colonization pattern. In contrast, species like Canthigaster

solandri, Synodus binotatus, Mulloidichthys flavolineatus, and Parupeneus spp colonized at only certain

times during the lunar period (Fig 6a,b). This shows that colonization was done by peak whereby large

numbers of larvae enter the lagoon via the reef crest.

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Figure 6a: Most abundant species (ranked 1-5)

Figure 6b: Most abundant species (ranked 6-10)

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Fish larval capture with light traps - Three light traps were used for sampling during two nights around

the new moon. Two were placed on the outer reef and one on the reef crest on the north coast of Moorea.

We collected 60 larvae belonging to the Families Acanthuridae, Apogonidae, Chaetodontidae,

Holocentridae, and Pomacentridae. A total of 13 species were collected (Annex 2). The most abundant

species were Neoniphon argenteus and Stegastes fasciolatus.

Fish larval identification - Among 71 species captured at larval stage with crest nets, we were able to

identify 67 to the lowest possible taxonomic level (for more details, see Annex 3 & 4).

Fish larval culture - Certain larvae were chosen for rearing in aquaria and these were fed with artemia

and artificial food. The mortality rate was low (<5%), showing that the conditions in the aquarium was

suitable for the fish to grow into juveniles.

4) DISCUSSION

Coral reef fish larvae have been sampled with the use of crest nets since 1988 (Dufour and Galzin 1993)

and has been proved a successful method of larval capture at the end of the larva’s pelagic life stage.

Colonization pattern of larvae are dependent on the lunar cycle, as was observed in the results. This

pattern also varies at species level, where different species prefer to colonize at different times; this could

be due to certain ecological processes (e.g. presence of predator, food, strength of waves, etc). There are a

few setbacks to the crest net method of larval capture as was noticed during the study, these include:

• It is highly dependent on weather conditions, unlike the light trap, this method of capture does not

have a high capability to work in rough weather conditions;

• Site selection of where the crest net is to be placed is very important. Factors such as wave and

current direction and strength, tidal range, etc, have to be considered to make this method operate

effectively. Therefore, site surveys have to be undertaken impeccably to prevent failures in the

future. Crest nets require low tidal range (e.g. 40 cm annually, as in Moorea, Tahiti) to prevent

larvae from escaping at extremely high tides and having high mortality at really low tides;

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• Since the net is placed on the crest, where energy is the highest on the reef, its design is essential

as abrasion on corals is liable to damage the net. The net design also has to be practical in order to

make operation (insertion and removal) easy to undertake.

Light traps were put up in order to observe the diversity of larvae caught and to find out which larval

capture technique is best for this type of survey. For capture of pre-flexion fish it is best to use light trap,

however, there is very little diversity in the fish caught in this method than that which was observed to be

caught by the crest net. There were more technical difficulties (e.g. light not working, or fish not getting

attracted to the light) that prevented the success of capture with the light trap method. The light trap

method is biased to fish that an attracted by light (active gear technique) therefore only fish that are

attracted to light will get caught in the net. Larvae that are caught by the crest net are involuntarily carried

into the codend by currents flowing over the crest into the lagoon thus making it a passive gear technique.

Fish larvae are brought from the study site to the aquarium in a large icebox with constant oxygen

supply to minimize stress for the larvae. Upon arrival at the aquarium, fish were sorted according to the

same species or family. Small fish like Pomacentrids and Chaetodons were put in containers whilst larger

fish (Mulliods and Priacanthids) were left to swim freely in the tank (Annex 5). It was important to

minimize stress throughout this process. Some fish may become stressed and change color. For example,

Chenochaetus striatus orange horizontal bands become more pronounced when they are stressed thus

making it look more like Acanthurus lineatus.

The larvae that were to be cultured into juveniles were moved into the aquarium for culture. There are

several factors that are important in the successful culture of larvae, these include; water quality, water

exchange system, practicality and placement of essential components of the aquaria and feed. The

aquarium had a “flow-through system” where water is constantly circulated within the tank. This system

is preferred over the “closed-system” as clean water is always entering the tank, and ‘difficult’ species

like Chaetodontidae require high water quality. Good filtration is lacking, as at times of high rainfall the

water quality decreases greatly. It is important, in order to reduce mortality, for a sand filter to placed

between the collection tank line and the seawater pump so that water entering the aquaria has reduced silt

and sedimentation thus reducing extra effort put into bottom siphoning the tanks. Certain components in

the aquaria like, placement of oxygen pipeline and seawater pipeline needs to be rearranged so that it is

easier and more practical to work in the aquaria. Feed for the larvae consisted of a combination of

artificial pellet food and live feed in the form of artemia (Annex 5). Tank color is also important in fish

identification as some fish, in order to camouflage themselves, change color according to the color of the

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aquaria. For example Rhinecanthus aculeatus is a pale yellow larvae with lateral black striations (Annex

4) but when placed in white aquaria, they become white and the black striations are almost

incomprehensible. Overall, mortality was low therefore, despite the various technical problems, it can be

said that the culture process was a success.

Pictures were taken of the fish larvae and after rearing to juvenile stage, pictures were taken again in

order to identify the various changes in meristic characteristics (Annex 3,4). This was undertaken in the

Optic laboratory at the CRIOBE station which was highly equipped and suitable for the requirements of

this study (Annex 5). Fish that did not require being cultured or having a picture taken were released back

into the lagoon on the evening after collection.

To conclude, the crest net method is a good approach in capture of fish larvae just before

colonization, which is subsequent to the pelagic stage of the life cycle of reef fish. It is an effective

method that requires minimum maintenance and effort in obtaining fish larvae that enter the lagoon via

reef crest. There is however, more need to study the morphological and meristical changes that the occur

between the larval and juvenile transition.

5) REFERENCE

Allen, G. (2002) A Field Guide for Anglers and Divers: Marine Fishes of the Great Barrier Reef and

Sout-East Asia. Western Australian Museum, Francis Street, Perth, Western Australia.

Allen, G., Steene, R., Humann, P., Deloach. (2005) Reef Fish Identification: Tropical Pacific. New World

Publications, Inc. Jacksonville, Florida, USA. Odyssey Publishing, Inc. El Cajon, California

USA.

Bond, CE. (1996) Biology of Fishes: Second Edition. Saunders College Publishing.

Doherty PJ (2002) Variable replenishment and the dynamics of reef fish populations. In: Sale PF (ed)

Coral reef fishes: dynamics and diversity in a complex ecosystem. Academic Press, San Diego,

pp 327-358

Dufour, V. (1994) Comparison of the Colonization of Fish Larvae in Coral Reefs of Two Islands of

French Polynesia, the Atoll of Rangiroa (Tuamotu Archipelago) and the High Island of Moorea

(Society Archipelago). Atoll research Bulletin No. 399. 12 pp.

Dufour, V., Galzin, R. (1993) Colonization Patterns of Reef Fish Larvae to the Lagoon at Moorea Island,

French Polynesia. Marine Ecology Progress Series 102, 143-52

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Dufour, V., Riclet, E., Lo-Yat, A. (1996) Colonization of Reef Fishes at Moorea Island, French

Polynesia: Temporal and Spatial Variation of the Larval Flux. Marine & Freshwater Resources,

47, 413-22

Google Earth Pro 4.0 Beta (2006). Google Earth Software. Google ©.

Grignon, J. (2007) Culturing Fish Larvae : A New Tool for Restocking Coral Reefs in Fiji. Melanesian

Geo, May-September. Pp 34 – 35.

Helfman, GS., Collete, BB., Facey, DE. (1997) The Diversity of Fishes. Blackwell Science, Inc. USA.\

Lecchini D., Dufour V., Carleton J., Strand S. & R. Galzin, 2004. Study of the fish larval flux at Moorea

Island: is the spatial scale significant? Journal of Fish Biology, 65: 1142-1146

Maamaatuaiahutapu, M., Remoissenet, G., Galzin, R. (2006) Guide d’identification des larves de poisons

récifaux de Polynésie française. Coral Reef Initiative for the South Pacific. Éditions Téthys 104p.

Myers, RF. (1999) Micronesian Reef Fishes: A Field guide for Divers and Aquarists. Coral Graphics,

Territory of Guam.

Randall, JE. (2005) Reef and Shore Fishes of the South Pacific: New Caledonia to Tahiti and the Pitcairn

Islands. University of Hawai’i Press, Honolulu.

Randall, JE., Allen, GR., Steene, RC. (1990) The Complete Divers’ & Fishermen’s Guide to Fishes of the

Great Barrier Reef and Coral Sea. University of Hawai’I Press, Honolulu.

Sale, PF. (1991) The Ecology of Reef Fishes on Coral Reefs. Academic Press Limited, London.

Sale, PF. (2002) Coral Reef Fishes: Dynamics and Diversity in a complex Ecosystem. Academic Press

Limited, London. Elsevier Science USA.

Werner, E.E. (1988). Size, scaling and the evolution of complex life cycles. In: Ebenman, B., Perssons, L.

(Eds.) Size-structured populations. Springer-Verlag, Berlin, pp 61-81.

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ANNEX 1:

CAPTURE OF FISH LARVAE WITH CREST NET

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Species List ABDAcanthurus triostegus 325Stegastes fasciolatus 204Chromis viridis 71Chrysiptera leucopoma 65Apogonidae 51Canthigaster solandri 38Synodus binotatus 35Ptereleotris microlepis 30Mulloidichthys flavolineatus 28Chrysiptera glouca 25Parupeneus multifasciatus 25Neoniphon argenteus 23Pomacentrus pavo 22Ctenochaetus striatus 21Siganus argenteus 19Naso brevirostris 15Stegastes nigricans 15Albula glossodonta 13Sargocentron microstoma 13Chaetodon citrinellus 12Apogon exostigma 11Myripristis kuntee 10Zebrasoma scopas 10Bothus mancus 9Chaetodon auriga 7Gymmapogon 7Priacanthus hamrur 7Acanthurus lineatus 6Acanthurus olivaceus 6Naso unicornis 6Ostohinchus augustatus 6Parupeneus barberinus 6Acanthurus xanthopterus 5Chaetodon ephippium 5Naso lituratus 5Apogon fraenatus 3Myripristis violacea 3Naso annulatus 3Naso hexacanthus 3Pseudobalistes flavimarginatus 3

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Rhinecanthus aculeatus 3Valenciennea strigata 3Aulostomus chinensis 2Apogonichthys ocellatus 2Canthigaster bennetti 2Canthigaster valentini 2Clupeidae 2Dactyloptena orientalis 2Lutjanus kasmira 2Myripristis bernndti 2Abudefduf sexfasciatus 1Arothron hispidus 1Aulostomus chinensis 1Balistoides viridescens 1Carapus honei 1Centropyge flavissimus 1Chaetodon pelewensis 1Chaetodon ulietensis 1Cheilodipterus quinquelineatus 1Dascyllus aruanus 1Fistularia commersonii 1Heteropriacanthus cruentatus 1Lutjanus fulvus 1Lutjanus monostigma 1Monotaxis grandoculis 1Myripristis adusta 1Neoniphon sammara 1Parupeneus cyclostomus 1Pseudocheilinus octotaenia 1Sargocentron spiniferum 1Thalassoma amblycephalum 1

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ANNEX 2:

CAPTURE OF FISH LARVAE WITH LIGHT TRAP

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SPECIES LIST ABDAcanthurus triostegus 1Acanthurus xanthopterus 4Apogon fraenatus 2Apogonidae 1Chaetodon auriga 2Chaetodon ephppium 1Dascyllus reticulatus 5Lutjanus monostigma 2Myripristis augusta 1Myripristis berndt 1Neoniphon argenteus 30Sargocentron microstoma 1Stegastes fasciolatus 9TOTAL 60

ANNEX 3:

IDENTIFICATION GUIDE OF CORAL REEF FISH LARVAE

FAMILY – ACANTHURIDAE

(Surgeonfishes)

Surgeonfishes are named for the sharp spine or spines that they possess on the caudal peduncle region. This family is divided into 3 sub‐families:

1. Acanthurinae : Single caudal spine which folds into a horizontal groove on the side of the peduncle (genera: Acanthurus, Zebrasoma, Paracanthurus and Ctenochaetus),

2. Nasinae : One or two fixed keel‐like spines on each side of the peduncle (genus: Naso), 3. Prionurinae: Three to six spines extending from peduncle region (genus: Prionurus).

Most members of Acanthuridae family are herbivorous and graze or browse on algae, however, a few species (Acanthurus and many Naso) feed on zooplankton. The genus Ctenochaetus are mainly detrivores. Acanthurid fish are diurnal and are often observed to sleep in small caves or crevices in the reef (Randle et al, 1990).

Surgeonfishes, in general, have a long period of development in the pelagic realm. Thus as a result, many species are widely distributed: 9 species from Indo‐Pacific region have extended their region to the tropical eastern pacific. The post‐larval stage of surgeonfish is orbicular, scaleless with narrow vertical ridges on the body, transparent and possesses a silvery color over the abdomen (Randall 2005).

Sub‐Family ACANTHURINAE

Acanthurus triostegus (Linnaeus, 1758)

Convict Surgeonfish

LARVAL STAGE JUVENILE STAGE ADULT STAGE

STANDARD LENGTH: 25 ± 5 mm

SEASON OF CAPTURE: January – February

SIMILAR/DIFFERENT CHARACTERISTICS

GUIDE TO IDENTIFYING Acanthurus Triostegus AT LARVAL STAGE:

* General body shape of larvae is similar to Acanthurus genus

* 4 vertical stripes on trunk (plus 1 stripe over the eye and caudal peduncle‐not so pronounced)

* Abdomen extending to head is silver in color, whereas trunk is transparent.

LARVAL CHARACTERISTICS:

Larva is translucent beginning from pelvic fin to caudal peduncle, whilst abdominal region and head is silver in color (to conceal the heart and main blood arteries, to prevent easy visual detection form predator in open water). Black vertical stripes extending from base of dorsal fin to base of the pelvic region.

SIMILARITIES/DIFFERENCES BETWEEN LARVAE‐JUVENILE AND ADULT

SIMILARITIES ‐ General body shape remains the same. Black vertical stripes are retained (4 stripes including 1 on caudal peduncle and 1 over the eye) and are more pronounced in adult.

DIFFERENCES ‐ Adult color is light olivaceous gray color and larval transparency is lost. Adult possesses a sharp, forward‐pointing, erectile spine on each side of caudal peduncle which folds down into a groove.

JUVENILE CHARACTERISTICS:

Larvae Loses transparency after metamorphosis into juvenile stage. Changes to white and stripes become more pronounced upon transition from juvenile stage. Juvenile is 40 ± 5 mm in length.

MAXIMUM TOTAL LENGTH: 260 mm

B

A

Acanthurus olivaceus (Forster, 1801)

Orangespot surgeonfish





GUIDE TO IDENTIFYING Acanthurus olivaceus AT LARVAL STAGE:


* Transparent posteriorly from abdominal region to caudal peduncle. Abdominal region is silver whilst pectoral, anal and caudal fins are yellow.

Transparent posteriorly from abdominal region to caudal peduncle with yellow caudal fin and yellow soft ray fins (pectoral & anal) – A. Head and abdominal region is silver in color. Upon metamorphosis into juvenile, it is yellow all over. B shows transition into juvenile stage.


Juvenile completely yellow. Juvenile is usually 46 ± 5 mm in length. Develops an orange band with light blue rim extending posteriorly from upper end of gill opening‐occurs before change from yellow to brown. Tip of anal fin is blue in color. Tail develops a lunate shape and caudal spine develops.



SIMILARITIES ‐ General body shape remains the same.

DIFFERENCES ‐ Larvae are transparent with silver head and abdominal region, pectoral, anal and caudal fins are yellow. Juvenile is yellow and develops orange band with blue rim upon transition into adult stage, caudal spine is seen in this stage. Adult is dark brownish gray with posterior to upper end of gill opening possessing a bright orange horizontal band, with a deep blue border, the head and anterior half of body usually abruptly paler than the posterior half, tail is lunate.

Acanthurus lineatus (Linnaeus, 1758)

Lined Surgeonfish



SEASON OF CAPTURE: January‐February

GUIDE TO IDENTIFYING Acanthurus lineatus AT LARVAL STAGE:


*Alternate bands of yellow and dark blue. Often when under stress, blue band becomes more prominent

*Anal, dorsal and caudal fins have small tinge of orange or yellow.

LARVAL CHARACTERISITCS:

Translucent brown extending posteriorly from abdomen to caudal region. Abdominal region and head silver in color. Alternate orange and dark blue striations longitudinally posteriorly from operculum. Tail is lunate.



Transition from larvae to juvenile involves the body becoming whiter in color and loosing larval transparency and translucency. Horizontal striations remain the same in color (alternate yellow and blue) as in larval stage however, striation pattern becomes more pronounced on the head of juvenile


SIMILARITIES ‐ General body shape remains the same.

DIFFERENCES ‐ Upper three quarter portion of adult head and body alternately horizontally banded yellow and black‐edged blue. Lower quarter of adult body is lavender to pale blue. Caudal spine is very long, this is absent in larvae.

Acanthurus xanthopterus (Valenciennes, 1835)

Yellowfin Surgeonfish




GUIDE TO IDENTIFYING Acanthurus xanthoptera AT LARVAL STAGE:


*Transparent posteriorly from abdomen to caudal peduncle

*Abdomen and head is striking silver in color. On forehead, in between eyes is a dark region

*To prevent confusion with Acanthurus nigricauda at larval stage, the size of larvae should be within 21 – 30 mm, if it is larger than this range, it is not A. xanthoptera.


Transparent posteriorly from abdomen to caudal peduncle. Abdomen and head is silver. Dark region just above the eyes on forehead.


Larvae can be easily mistaken for Acanthurus nigricauda, however, upon metamorphosis to juvenile there is a stark difference. Pectoral fin is yellow at the tip and brownish grey at the base. Dorsal and anal fins contain several light blue bands alternating with dull yellow. Distinct whitish bad on caudal peduncle.


SIMILARITIES ‐ General body shape remains the same and bands observed in juvenile stage are also present in adult. Pale yellow band around eye is also present but more distinct in adult stage.

DIFFERENCES ‐ Adult color pattern can change quickly from uniform purplish gray to one with alternating, very irregular, lengthwise bands of dark yellowish grey and light blue grey. There is a prominent broad yellow band extending anteriorly to eye and one irregularly posteriorly to eye and extending to upper portion of operculum. Pale blue area around caudal peduncle near caudal spine.

Zebrasoma scopas (Cuvier, 1829)

Brushtail Tang




GUIDE TO IDENTIFYING Zebrasoma scopas AT LARVAL STAGE:

*General body shape of larvae is similar to Zebrasoma genus

*Transparent posteriorly from abdomen with alternate vertical black striations extending into dorsal and anal fins

*Abdomen is silver, dark area above eye on forehead and mouth is transparent with irregular black striations.


General body shape of larvae is similar to Zebrasoma genus. Transparent posteriorly from abdomen. Vertical pale black striations on body extending to dorsal and anal fins. Abdomen and head is silver. Transparent extending anteriorly from the eye up to mouth with irregular pale black striations following the contours of the mouth. Dark area above the eye on forehead.


Forebody pale brown with gold/yellow spots on head. Rear body is dark brown to purple. Thin alternate paired vertical yellow and bluish grey bars on lower three quarter of body. Abdomen region has white tinge. White tail spine.



SIMILARITIES ‐ General body shape remains the same. Tail spine white as in juvenile.

DIFFERENCES ‐ Larval transparency is lost. Alternate vertical striations during larval and juvenile stage become dark brush‐like patch of bristles in front of caudal spine and tiny horizontal pale blue dots or lines on head and body during adult stage.

Ctenochaetus striatus (Qouy & Gaimard, 1825)

Lined Bristletooth




GUIDE TO IDENTIFYING Ctenochaetus striatus AT LARVAL STAGE:

*Alternate orange and black horizontal striations on the body.

*Silver abdomen and head

* In order to prevent confusion between C. striatus and Acanthurus lineatus, look for black spot located on posterior base of dorsal and anal fin and C. striatus is significantly smaller than A. lineatus.


Abdomen and head is silver. Body is translucent with alternate horizontal orange and black banded striations. Dorsal, anal and caudal fins are translucent grey. Black spot located above and below caudal peduncle on posterior base of dorsal and anal fin.


Horizontal striations become pronounced and posterior most tips of caudal fin is yellow. Two yellow stripes located across the eye. Head region pale grayish brown in color. Posterior tips of caudal fin orange.



SIMILARITIES ‐ General body shape remains the same. Striation pattern similar throughout life‐cycle (i.e.) horizontal pattern on the trunk extending from the operculum to caudal region. Dark spot located on the base of dorsal fin is retained in adult.

DIFFERENCES ‐ Adult can be dark brown to black with narrow orange stripes and black dorsal, anal, and caudal fin. Pale spots also found on head and around the eye. Pale alternate striations can be found on the body and extending to fins.

Sub‐Family NASINAE

Naso annulatus (Qouy & Gaimard, 1825)

Whitemargin Unicornfish

LARVAL STAGE ADULT STAGE



GUIDE TO IDENTIFYING Naso annulatus AT LARVAL STAGE:

*General body shape is similar to Naso genus

*Abdomen and head silver with body near transparent

*First dorsal and anal spine is long and decreases posteriorly

*Two sharp spines at the base of abdomen

*Usually with a length of 30 – 34 mm range, if bigger, it is most likely N. brevirostris.


Abdomen and head is silver in color. Translucent posteriorly from abdomen to caudal region. Long first dorsal and anal spine and decreasing in height posteriorly. Two sharp spines at the base of the abdomen.



White ring between caudal spines around caudal peduncle. Shiny rear flank develops after 6 months into juvenile stage.


SIMILARITIES ‐ General body shape similar between larvae and juvenile stage.

DIFFERENCES ‐ Adult is olivaceous brown with no dark markings but can rapidly change color to pale bluish gray. White lips, caudal fin with blackish sub‐marginal band with white posterior border. Long tapering horn on forehead. Adult has less body‐depth to length ratio compared to larvae.

PELVIC SPINE

Naso brevirostris (Valenciennes, 1835)

Spotted Unicornfish




GUIDE TO IDENTIFYING Naso annulatus AT LARVAL STAGE:

*General body shape is similar to Naso genus

*Abdomen and head silver with body near transparent

*First dorsal and anal spine is long and decreases posteriorly

*Two sharp spines at the base of abdomen

*Usually with a length of 37 – 43 mm range, if smaller, it is most likely N. anulatus.



Abdomen and head is silver. Dark sports (pigmentation) on body and around the head. First dorsal and anal spine is long. Two sharp spines at the base of the abdomen.


Greyish brown, darker on upper half of body and lighter close to base of abdomen. Juveniles of about 10cm develop horn which appears as a bump on forehead. Caudal fin slightly rounded and white except on the base.


SIMILARITIES ‐ First dorsal and anal spines are long in larval and juvenile stages.

DIFFERENCES ‐ Adult is olivaceous brown to grey with small dark spots on head and vertical rows of spots and lines on flank. Caudal fin is white with small dusky spot at the base and is truncate to slightly rounded. Two peduncle plates on each side of caudal region. A broad‐based tapering horn on forehead‐more than half a head length in front of mouth.

Naso hexacanthus (Bleeker, 1855)

Sleek Unicornfish




GUIDE TO IDENTIFYING Naso hexacanthus AT LARVAL STAGE:

* General body shape is similar to Naso genus

*Abdomen and head silver with body translucent grey

*First dorsal and anal spine is long and decreases posteriorly (two sharp spines at the base of abdomen)

*Dark circular dots (pigmentation) on the upper portion of the body and dark line over the eye

*Usually large in size 40+ mm.


Abdomen and head is silver. Body is translucent grey. Dark circular sports (pigmentation) on upper portion of body. Dark line over the eye. First dorsal and anal spine is long. Caudal fin is emarginate. Two sharp spines at the base of the abdomen.


Caudal fin is emarginate. General body shape is maintained but with lesser body‐depth to length ration.


SIMILARITIES ‐ First dorsal and anal spines are long and caudal fin emarginate in larval and juvenile stages.

DIFFERENCES ‐ Adults are brown to bluish grey dorsally and shading ventrally to yellowish. Is capable of changing color rapidly to pale blue. Margin of operculum and preopercle often dark brown. Pale white line extending horizontally across the forehead and ending at the eye. Two peduncle plates with large knife like spines. Caudal fin truncate.


Naso lituratus (Forster, 1801)

Orangespine Unicornfish




GUIDE TO IDENTIFYING Naso lituratus AT LARVAL STAGE:


* Abdomen and head silver with body dark translucent grey (flank is shiny)

*Pale yellow blotch above the eye, circular black dots located in the upper portion of body closer to head and white dots randomly cover flank and extending to dorsal fin.



Abdomen is silver with body translucent grey. Pale yellow blotch above eye and on mouth. Flank is shiny. Circular black dots located in the upper portion of body closer to head. White dots randomly cover flank and extending to dorsal fin. Long first dorsal and anal fin. 2 spines at the base of the abdomen. Caudal fin with pale yellow posterior border. Base of anal fin pale yellow with dark posterior border.


Yellow‐edged black area extending from forehead and bordering mouth. Black dorsal fin with white posterior border. Yellow anal fin. Peduncle plates are yellow and juvenile develop caudal spines at 5‐6 months.


SIMILARITIES ‐ First dorsal and anal spines are long in larval and juvenile stages. Narrow yellow edged black area extending from the eye to the mouth present in both juvenile and (more pronounced) in adult.

DIFFERENCES ‐ Adults are brownish gray with a yellowish nape. Peduncular plates bright orange. Lips orange‐yellow. Dorsal fin black basally, white posteriorly, with a blue margin and narrow blue band at base. Caudal fin with a sub‐marginal yellow band posteriorly.

Naso unicornis (Forsskål, 1775)

Bluespine Unicornfish




GUIDE TO IDENTIFYING Naso unicornis AT LARVAL STAGE:


* Abdomen and head dull silver with upper portion of body yellowish brown and lower half transparent (flank is shiny)

* Circular black dots located in the upper mid‐dorsal portion of body

* Two black dots on caudal peduncle.



Abdomen is silver. Mid‐upper portion of body is yellowish brown and lower half of the body is translucent. Flank is shiny. Circular black dots located in the upper mid‐dorsal portion of body. Long first dorsal and anal fin. Caudal fin transparent with two black dots on caudal peduncle.


Juvenile is near uniformly olivaceous brown. Two peduncle plates are blue with caudal spines developing at around 4 months. Caudal fin emarginate.


SIMILARITIES ‐ First dorsal and anal spines are long in larval and juvenile stages. Peduncular plates are blue as in adults.

DIFFERENCES ‐ Adults are olivaceous gray. Relatively short horn which does not project past the mouth. Dorsal and anal fins are yellowish with narrow blue stripes. Well developed forward pointing caudal spines. Caudal fin truncate with filamentous lobes.

FAMILY – CHAETODONTIDAAE

(Butterflyfishes)

Butterflyfish are best known for the intricate patterns and striking colors that make them a popular choice for aquarists and snorkelers on reef ecosystems. This family contains 116 species all of which occur mainly in tropical seas around coral reefs (Randall et al, 1990).

Butterflyfish lack the spine at the corner of the preopercle which is what makes them different from Angelfishes. They have scaly axillary process at the base of the pelvic fins which is absent in Angelfishes (Randall, 2005). They have deep and highly compressed bodies covered with moderately small ctenoid scales extending onto median fins. They possess small protractile mouths that have a band or rows of tiny brushlike teeth in the jaws. Butterflyfish have a single continuous dorsal fin with the anterior‐most interspinous membranes deeply incised and rounded to a slightly emarginated tail (Myers, 1999). Chaetodons possess a unique post‐larval stage called the tholichthys larva, which has large bony plates on the head and on the anterior of the body (Randall et al, 1990).

Butterfly fish are highly diurnal and many species feed on live coral polyps whilst others consume small invertebrates, tubeworms, algae or zooplankton (Helfman et al, 1997). Most butterflyfish are solitary or maybe found in monogamous pairs that are most often a lifelong partnership. Although most species inhabit coral‐rich reefs, some Chaetodons are found to be associated with silty coastal areas, while some others gather in huge shoals high above the reef to feed on drifting plankton (Allen, et al, 2003).

Chaetodon auriga (Forsskål, 1775)

Threadfin Butterflyfish




GUIDE TO IDENTIFYING Chaetodon auriga AT LARVAL STAGE:

*Black spot on posterior end of dorsal fin

*Silver head and fading to yellow posteriorly

*Black band over eye.



Larva is silver and fading to yellow at caudal region. One black vertical striation over the eye. Pelvic, and caudal fins transparent. Dorsal fin transparent at first quarter and becoming yellow posteriorly. Dark spot on posterior end of dorsal fin.


Body is white and fading to yellow at caudal region. Dark band over the eye. “Chevron” pattern on side. Black spot on posterior end of dorsal fin. Caudal fin yellow.


SIMILARITIES ‐ Black bar on eye. Black spot on posterior end of dorsal fin.

DIFFERENCES ‐ Adult is white overall with bright yellow on posterior part of the body and caudal fin. Chevron pattern on flank. Mouth of adults is elongated compared to larval and juvenile.

Chaetodon citrinellus (Cuvier, 1831)

Speckled Butterflyfish




GUIDE TO IDENTIFYING Chaetodon citrinellus AT LARVAL STAGE:

*Oblique to horizontal dark spots on flank

*Dark band over the eye. Silver and fading to yellow posteriorly

*Caudal fin transparent. First dorsal spine is long.



Larva is silver and fading to yellow at caudal region. One black vertical band over the eye. Small oblique to horizontal dark spots on flank. Caudal fin transparent. First dorsal spine is black and dorsal fin is yellow posteriorly. Anal fin transparent at the base with black margin.


Juvenile is pale yellow, head still silver. Prominent black band over the eye. Oblique to horizontal black spots on flank. Anal fin pale yellow at the base with black margin.


SIMILARITIES ‐ Black bar on eye. Oblique to horizontal dark spots on flank. Pale yellow in color.

DIFFERENCES ‐ Adult has greater body depth to length ratio. Anal fin pale yellow at the base followed by white with black edging.

Chaetodon ephippium (Cuvier, 1831)

Saddled Butterflyfish




GUIDE TO IDENTIFYING Chaetodon ephippium AT LARVAL STAGE:

*Larva is silver and body is bordered yellow

*One vertical black band over the eye.

*Dark area near posterior end of the body.



Larva is silver and body is bordered by yellow. One black vertical band over the eye. Anal fin transparent. Dark area posteriorly on body.


Juvenile is white with mouth, dorsal fin, pectoral fin and anal fin yellow in color. Prominent black band over the eye. Black area posteriorly on back and adjacent dorsal fin. Black band with white edging on caudal peduncle. Caudal fin transparent (at 2 months).


SIMILARITIES ‐ Pelvic fin is yellow. Short narrow black bar over the eye in adult.

DIFFERENCES ‐ Adult is bluish‐grey with wavy blue lines on lower body. Large white bordered black patch on upper rear body. Orange area from snout to ventral fins. Filament extending posteriorly from upper part of soft portion of dorsal fin.

Chaetodon pelewensis (Kner, 1868)

Dot‐Dash Butterflyfish




GUIDE TO IDENTIFYING Chaetodon pelewensis AT LARVAL STAGE:

*Larva is silver with black vertical line over the eye

*Tip of dorsal fin spines yellow

*First four dorsal spines are long

*Pale oblique dark striations on upper posterior end of body.


Larva is silver. One black vertical band over the eye. First four dorsal spines extremely long. Tips of dorsal spines are yellow. Oblique dark striations on upper posterior end of body. Caudal fin transparent.


Juvenile is pale tan with oblique rows of dark spots. Caudal region orange in color. Black stripe over the eye.



SIMILARITIES ‐ Oblique dark spots on upper end of posterior. Caudal region is orange.

DIFFERENCES ‐ Adult is pale tan. Dark spots becoming solid bands on upper half of body. Dark orange band over the eye.

Chaetodon trifascialis (Quoy & Gaimard, 1825)

Chevron Butterflyfish




GUIDE TO IDENTIFYING Chaetodon pelewensis AT LARVAL STAGE:

*Larva is white with black vertical line over the eye and posterior end of body

*Mouth, lower abdomen and dorsal fin yellow

*Yellow band over caudal peduncle.

*Large eyes.


Larva is white with mouth and caudal region yellow. One black vertical band over the eye and one vertical black striation over posterior of body. Yellow band over caudal peduncle. Mid‐section of body is white and caudal fin is transparent.


Juvenile is white with yellow mouth and forehead, pectoral fin and caudal fin. Black band over the eye and oon posterior end of body. Pale chevron pattern on mid‐section of body.


SIMILARITIES ‐ Vertical black band over the eye. Chevron pattern on body.

DIFFERENCES ‐ Body is white with dorsal and anal fin yellow. Caudal fin is black with yellow margin.

Chaetodon ulientensis (Cuvier, 1831)

Pacific Double‐Saddle/Double‐Barred Butterflyfish




GUIDE TO IDENTIFYING Chaetodon ulietensis AT LARVAL STAGE:

*Larvae is small and silver overall in color over‐laid with one dark and two pale black stripes.

*Three vertical black bars on the body. Darker bar located over the eye.


Larva is silver anteriorly and fading posteriorly to gold. One black vertical striation over the eye and two pale vertical striations across the body (i.e. one thin band after the operculm and another thicker band before the caudal peduncle). Pelvic, and dorsal fins are transparent.



Body is white and fading to yellow at caudal region with alternate thin black striations (more pronounced than in larvae‐3 stripes, 1 dark band over the eye, 1 after operculum and 1 before caudal peduncle). Second and third band do not reach the base of the body. There is a black spot on the caudal peduncle. Dorsal and anal fin tips yellow.


SIMILARITIES ‐ Three black bars on body. Black spot on caudal peduncle present in juvenile and adult stages.

DIFFERENCES ‐ Adult is white overall with bright yellow on posterior part of the body and caudal fin. Series of blackish vertical lines on its sides overlaid by two broad, blackish bars. Mouth of adults is elongated compared to larval and juvenile. Body depth to length ratio is greater in adult.

FAMILY – POMACENTRIDAE

(Damselfishes)

Damselfish are among the hardiest of aquarium fish, but their aggressive territorial nature of many species does not make them a popular choice for small scale aquariums. This family is divided into 4 sub‐families:

1. Amphiprioninae: These include the Anemonefishes which live in close association with large sea anemones (genus: Amphiprion, Premnas).

2. Chrominae: Primarily planktivores that are closely tied to a particular patch of coral (genera: Chromis, Dascyllus).

3. Lepidozyginae: This small elongate Damsel occurs in aggregations, often with anthiases, on the upper edges of current swept seaward reefs. (genus: Lepidozygus)

4. Pomacentrinae: Contains the bulk of this family.(genera: Abudefduf, Amblyglyphidodon, Cheiloprion, Chrysiptera, Dischistodus, Hemiglyphidodon, Neoglyphidodon, Neopomacentrus, Plectroglyphidodon, Plectroglyphidodon, Pomacentrus,Pomachromis, Stegastes)

There are approximately 28 genera and 321 species of Damselfishes that occur worldwide, and this makes them one of the most abundant groups of coral reef fish (Randall, 2003). Damsels are elongate to orbicular with compressed bodies and a single continuous dorsal fin. The majority of the species dwell in shallow coral reef areas or rocky substrates of the tropics, some though are found in temperate waters.

Some Damselfish feed primarily on coral polyps, most other Damsels are omnivorous, feeding on algae and many different small benthic invertebrates. They are highly diurnal and find shelter in the crevices of reefs.

Sub‐Family CHROMINAE

Chromis viridis (Cuvier, 1830)

Blue‐green Chromis




GUIDE TO IDENTIFYING Chromis viridis AT LARVAL STAGE:

* Larvae is small (5‐7 mm)

* Upper portion of head is pale blue in color

* To prevent confusion with Pomacentrus pavo, C. viridis has a shorter abdomen to tail length compared to P. pavo

*Body proceeding posteriorly from abdomen is transparent.


Larva is transparent beginning from pelvic fin to caudal peduncle, whilst abdominal region and head is silver in color. Head is pale blue.


Larva loses transparency after metamorphosis. Changes to iridescent light blue to green color on upper half of body and fading to white ventrally. Juvenile has large eyes.



SIMILARITIES ‐ General body shape remains the same. Iridescent light blue to green color is found throughout all life stages.

DIFFERENCES – Adult has a faint dusky spot at the upper base of its pectoral fin.

Dascyllus aruanus (Linnaeus, 1758)

Humbug Dascyllus




GUIDE TO IDENTIFYING Dascyllus aruanus AT LARVAL STAGE:


* 2 thick pale black striations over body

* Blue dots over abdomen and black dots between the eyes

*Body is transparent


Larva is transparent. Two pale black vertical bands over abdominal region and on posterior end of body. Circular and shiny blue dots over abdominal region. Black Dots on forehead, between the eyes.


Juvenile is white overlayed with 3 prominent black vertical striations extending to dorsal and anal fins (1 over the eye, 1 over midsection and 1 over caudal region). Caudal peduncle white and caudal fin transparent.


SIMILARITIES – General body shape is maintained. White overlaid with black vertical striations.

DIFFERENCES – Adult has a large white spot between the eyes.


Sub‐Family POMACENTRINAE

Abudefduf sexfasciatus (Lacepède, 1801)

Scissortail Sergeant






* Bluish black color

* Abdomen shiny (pearl‐like), *Mouth, dorsal, anal and caudal fins are transparent.


Larva is blue‐black in color. Abdomen is shiny. Mouth is translucent. Anal, dorsal and caudal fins are transparent. Pale and non‐distinct vertical striations on flank.


Juvenile is pale blue‐green in color. Dark vertical striations on body, first band beginning behind operculum and last band on caudal peduncle. 5 bands in total. Anal, dorsal and caudal fins transparent.


SIMILARITIES – Body is pale blue‐green color overlaid with 5 black bars.

DIFFERENCES – In adult, 3 black bars extend onto dorsal fin and 1 extends onto anal fin. Caudal fin grey with a black band in each lobe, upper band connects dorsally with caudle peduncle bar.


Chrysiptera leucopoma (Cuvier)

Synonyms ‐ C. amabilis (De Vis), C. brownriggii (Bennett, 1828)

Surge Damselfish





* Yellow body with bright horizontal blue line extending from forehead to posterior end of dorsal fin/brown body with white vertical band in midsection. Sometimes there is a combination of both types as seen in the above picture

* Dorsal fin is bright orange

*Tiny black dot at the base of the dorsal fin.


2 types of larvae: 1) yellow body with bright horizontal blue bar extending from forehead to posterior end of dorsal fin. Dorsal fin is bright orange. 2) brown body with white vertical band in midsection of body, blue horizontal bar not pronounced.


Juvenile is either: 1) yellow with bright horizontal bar extending from forehead to posterior end of dorsal fin with two black spots at the base of the dorsal fin, or, 2) brown with two vertical white bands, 1 on midsection and second on caudal peduncle. Two dark spots on the base of dorsal fin.


SIMILARITIES – General bod shape is maintained. Similar pattern observed throughout life stages (regardless of whether yellow or brown), habitat shift may cause coloration to change. Black dot on the base of caudal peduncle.

DIFFERENCES – In adult, the horizontal blue band (in the yellow C. leucopoma) is thicker and there is counter shading from light blue to yellow ventrally.


Pomacentrus pavo (Bloch, 1787)

Peacock Damselfish




GUIDE TO IDENTIFYING Pomacentrus pavo AT LARVAL STAGE:

* Transparent body with yellow caudal fin

* Abdomen and head is silver with a tinge of blue

*To prevent confusion with chromis viridis, this species has a body which is more elongated compared to the Blue‐green damsel.


Larvae are transparent. Abdomen and head are silver with a slight tinge of pale blue. Caudal region and fins are pale yellow.


Juvenile is blue with a slightly silver abdomen. Dark shiny blue dots on head and over body. Posterior end of dorsal and anal fin are yellow. Caudal region and caudal fin is yellow.


SIMILARITIES – Blue body with yellow caudal fin.

DIFFERENCES – In adult, there is a line or row of pale green dots on each scale. Irregular blue‐to‐bluish green lines on head and operculum


ANNEX 4:

PHOTOGRAPH OF CORAL REEF FISH LARVAE

Abudeduf sexfasciatus‐9 mm Acanthurus lineatus‐43 mm Acanthurus olivaceaus‐30 mm

Acanthurus triostegus‐27 mm Acanthurus xanthopterus‐29 mm Antennarius commersonii‐20 mm

Apogon exostigma‐30 mm Apogonichthys ocellatus‐17 mm Apogonidae‐9 mm

Arothron hispidus‐10 mm Aulostomous chinensis‐155 mm Bothus mancus‐38 mm

Canthigaster bennetti‐28 mm Canthigaster solandri‐16 mm Carapus honei‐110 mm

Cephalopholis argus‐21 mm Chaetodon auriga‐19 mm Chaetodon citrinellus‐27 mm

Chaetodon ephippium‐16 mm Chaetodon pelewensis‐30 mm

Chaetodon trifascialis‐10 mm

Chaetodon ulientensis‐18 mm Chromis viridis‐9 mm

Chrysiptera glauca‐14 mm

Chrysiptera leucopoma‐15 mm

Chrysiptera leucopoma‐16 mm

Ctenochaetus striatus‐34 mm

Dactyloptena orientalis‐42 mm

Dactyloptena orientalis‐42 mm

Fistularia commersonii‐155 mm

Gymmapogon‐8 mm Gymmapogon‐11 mm Gymmapogon‐21 mm

Lutjanus fulvus‐27 mm Lutjanus kasmira‐21 mm Mulloidichthys flavolineatus‐78 mm

Myripristis berndti‐60 mm Myripristis kuntee‐63 mm Naso annulatus‐32 mm

Naso brevirostris‐40 mm

Naso hexacanthus‐44 mm Naso lituratus‐65 mm

Naso unicornis‐65 mm Neoniphon argentus‐33 mm Ostorhinchus augustatus‐16 mm

Parupeneus barberinus‐44 mm Parupeneus cylostomous‐67 mm Parupeneus mutifasciatus‐57 mm

Pomacentrus pavo‐17 mm

Priacanthus hamrur‐84 mm

Pseudobalistes flavimarginatus‐28 mm

Ptereleotris microlepis‐23 mm Rhinecanthus aculeatus‐25 mm

Sargocentron microstoma‐60 mm

Scorpaenodes guamensis‐9 mm Siganus argenteus‐40 mm

Stegastes fasciolatus‐13 mm

Stegastes nigricans‐10 mm Synodus binotatus‐43 mm Valencienna strigata‐25 mm

Zebrasoma scopas‐22 mm

ANNEX 5:

PHOTOGRAPHS OF THE TRAINING COURSE

Figure 1: Setting up the crest net for the night Figure 2: Attaching the codends securely

Figure 3: Sorting of fish larvae Figure 4: Fish put into tanks Figure 5: Artificial habitat

Sorting process involves sorting of larvae, putting similar species in the same tanks and creating artificial habitats for larvae

Figure 6: CRIOBE aquarium Figure 7: Larvae sorting shelf Figure 8: Aquarium maintenance

After collection of larvae from the study site, the larvae are brought to the aquarium for sorting and culture.

Figure 9: Photography apparatus Figure 10: Microscope photography apparatus

Upon identification, certain species are photographed and standard length recorded. Some larvae were donated for the BioCode project.

Figure 11: Dust pellet feed Figure 12: Crumble pellet feed Figure 13: Artemia feed

Fish larval feed consisting of artificial pellet food, sieved into dust for small fish and fish that have small mouths and crumble for the larger larvae. Live food in the form of artemia are fed to the fish, 45 minutes after they are fed with pellets.Larvae/juvenile are fed three times a day.

Figure 14: Releasing the juveniles

The juveniles were released back into the lagoon at Papetoai, Moorea, upon completion of the training workshop.

Documents

Capture Identification Culture Techniques