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Development of Captive Breeding Techniques for
Marine Ornamental Fish: A Review
Jonathan A. Moorhead a & Chaoshu Zeng a
a Aquaculture Discipline, School of Marine and Tropical Biology, James Cook University,
Townsville, Australia
Available online: 06 Oct 2010
To cite this article: Jonathan A. Moorhead & Chaoshu Zeng (2010): Development of Captive Breeding Techniques for Marine
Ornamental Fish: A Review, Reviews in Fisheries Science, 18:4, 315-343
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Reviews in Fisheries Science
, 18(4):315–343, 2010
Copyright C
Taylor and Francis Group, LLC
ISSN: 1064-1262 print
DOI: 10.1080/10641262.2010.516035
Development of Captive Breeding
Techniques for Marine Ornamental
Fish: A Review
JONATHAN A. MOORHEAD and CHAOSHU ZENG
Aquaculture Discipline, School of Marine and Tropical Biology, James Cook University, Townsville, Australia
The increasingly popular aquarium hobby is fueling the rapid growth of the aquatic ornamental industry, particularly
the trade of marine ornamental species. However, currently there is a heavy reliance on wild caught marine ornamentals
to satisfy consumer demand. As public awareness of the plight of marine ecosystems grows, the often destructive and
unmanaged exploitation of coral reefs for the marine ornamental trade has raised concerns. Therefore, there is consensus
that urgent actions need to be taken to limit destructive exploitation of coral reefs, and to ensure the sustainability of the
marine ornamental industry. An obvious and vital action of such efforts would be the development of captive breeding
techniques for marine ornamental species, in particular, coral reef fish. Research efforts towards captive breeding of marine
ornamental species, also known as marine ornamental aquaculture, stand to supplement or replace the supply of wild
caught specimens for the marine ornamental trade, and potentially help boost reef recovery efforts through restocking.
However, unfortunately, the marine ornamental aquaculture sector is still in its infancy, receiving limited research attention,
and, in turn, has experienced very slow development compared to the technical and industrial advances made in foodfish
aquaculture. While it is true that at present, multiple bottlenecks have severely limited the progress of marine ornamental
aquaculture, through careful appraisal and adaptation of culture techniques developed for foodfish, and by addressing the
specific needs of marine ornamental aquaculture, significant progress could be made for the marine ornamental aquaculture
industry. With this objective in mind, this review attempts to summarize the major bottlenecks facing the marine ornamental
aquaculture industry, and to highlight weaknesses in the current state of research. Major areas in need of increased research
efforts include broodstock management, such as the development of specific broodstock diets and broodstock husbandry,
spawning induction via hormone technologies that are tailored to the size and sensitivity of small broodstock ornamentals,
and comprehensive, species-specific larval rearing techniques, including system design and larval culture conditions as well
as larval feeds and nutrition.
Keywords marine ornamental fish, captive breeding, broodstock management, larval rearing, culture techniques
1. INTRODUCTION
The trade of aquatic organisms for home and public aquar-
iums and water gardens, along with associated equipment
and accessories, has become a multi-billion dollar industry
known as the aquatic-ornamental industry (AOI) (Larkin, 2003;
Wabnitz et al., 2003; Pelicice and Agostinho, 2005; Prang,
2007). On the whole, the most recent estimated value of the
AOI was in the vicinity of 15 billion US dollars (Prang, 2007;
Wittington and Chong, 2007). Within the AOI, among a myriad
Address correspondence to Chaoshu Zeng, School of Marine and Trop-
ical Biology, James Cook University, Townsville 4814, Australia. E-mail:
chaoshu.zeng@jcu.edu.au
of aquatic organisms traded, freshwater and marine fish species
are the most popular and dominant groups (Lecchini et al., 2006;
Moreau and Coomes, 2006). And on a unit weight basis, orna-
mental fish form the most valuable fisheries commodity in the
world (Hardy, 2003).
Over the past decade, technical advances in responsible cap-
tive care and life support system technologies have made aquar-
ium care easier and more accessible to common households,
especially with respect to maintaining marine organisms. This
has resulted in a rapid increase in demand for marine ornamental
fish, particularly coral-reef species, which has also contributed
to recent rapid growth of the AOI (Chan and Sadovy, 1998;
Rinkevich and Shafir, 1998; Sales and Janssens, 2003; Shuman
et al., 2004; Livengood and Chapman, 2007; Calado, 2006).
315
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316 J. A. MOORHEAD AND C. ZENG
Of the estimated 1,500 to 1,600 ornamental fish species com-
monly traded, near half are marine (Olivier, 2003; Livengood
and Chapman, 2007; Wittington and Chong, 2007). Marine fish
differ significantly from freshwater fish when it comes to the vol-
ume and value of specimens traded on the ornamental market.
In terms of volume, world trade of marine ornamental species
makes up less than 10% of total ornamental fish traded; however,
owing to their strong unit value, the proportion is higher when
estimated as a percentage of world trade by value, with strong
trading growth in recent years (Olivier, 2001, 2003; Livengood
and Chapman, 2007; Wittington and Chong, 2007).
Another striking difference between marine and freshwater
ornamental fish is the origins from which they are sourced.
Unlike their fresh-water counterparts that are mostly captive
bred (approximately 90%), it is estimated that between 90 to
99% of marine fish species traded on the ornamental market
are collected directly from the wild (Chapman et al., 1997;
Ostrowski and Laidley, 2001; Wilson et al., 2001; Tlusty, 2002;
Moe, 2003; Olivier, 2003; Tissot and Hallacher, 2003; Calado,
2006). This is of particular concern considering that a large
portion of coral-reef fish species are collected from poorly man-
aged fisheries in the Southeastern Asian and Caribbean region.
In countries of these regions, such as the Philippines and Indone-
sia, destructive capture methods, including explosives, poisons
(such as sodium cyanide), muro-ami and kayakas fishing tech-
niques (involving smashing of corals and rocks to scare fish into
nets), are widely used (Rubec, 1988; Rubec et al., 2001a; Wood,
2001a; Ziemann, 2001; Calado, 2006). At the point of collec-
tion, the use of such destructive methods inflicts severe and
often permanent collateral damage to coral reef ecosystems; de-
stroying reef structure, and killing coral heads and other fish of
no collection interest, along with target species (Rubec et al.,
2001b; Wood, 2001a). Furthermore, post-collection mortality
due to delayed and chronic chemical toxicity, poor handling,
and transport stress compounds the deleterious effects (Rubec
et al., 2001b; Wood, 2001a). Continued exploitation of reefs
in a destructive, poorly managed manner, has been reported to
reduce the yields of both ornamental and food fish from reef
fisheries, ultimately affecting the capacity of exploited reefs to
recover (Rubec et al., 2001a; Ziemann, 2001). In some instances,
collection of specimens for the marine ornamental market itself
has been directly linked to localized declines in several popu-
lar ornamental species (Lubbock and Polunin, 1975; Ziemann,
2001; Tissot and Hallacher, 2003; Shuman et al., 2005).
Aside from direct exploitation, the world’s coral reefs also
face the severe effects of global climate change and increased
impacts from anthropogenic activity, highlighting the need for
multi-faceted management and recovery programs to curb reef
declines and encourage recovery (Rubec, 1988; Wood, 2001a;
Bellwood et al., 2004; Shuman et al., 2004). As a part of effec-
tive coral reef management plans, there exists great potential for
a marine ornamental reef fish captive breeding program (Wood,
2001a; Zieman, 2001; Tlusty, 2002; Bellwood et al., 2004). Such
a program could complement coral reef management through
controlled releases of hatchery produced fingerlings of heavily
exploited species, and provide a significant contribution to scien-
tific knowledge of reef fish biology (Rinkevich and Shafir, 1998;
Ostrowski and Laidley, 2001; Wood, 2001a; Zieman, 2001).
In its present state, marine ornamental aquaculture (MOA)
is still in its infancy, with a limited number of species being
produced at an economically viable scale (Wood, 2001b; Holt,
2003). To date, captive larval rearing successes have been largely
limited to small, experimental, or hobbyist scales. Additionally,
very few scientific publications exist documenting aspects per-
tinent to captive culture of ornamental reef fish species (Shei
et al., 2010). However, there is great potential for significant
growth in this sector of the ornamental industry, owing to high
product value and its significance to coral reef conservation.
There are growing interests in marine ornamental aquacul-
ture from the commercial aquarium trading industry, marine
conservation, and research community (Watson and Hill, 2006).
However, for MOA to grow and reach commercial success, tech-
niques need to be developed to yield high quality and quantity
of eggs, and to culture a large quantity of larvae successfully
through to a marketable end point (Brooks et al., 1997; Coward
et al., 2002; Tlusty, 2002; Holt, 2003). This requires robust sci-
entific experimentation, with a view to make such information
freely available to the public. The aim of this review is to high-
light major bottlenecks/barriers to the development of MOA,
with focus on the key issues of broodstock management and
larval rearing. Aquaculture techniques and methods developed
for marine foodfish that have potential to be adapted for use in
the MOA will also be reviewed. Ultimately, this review aims to
elucidate the needs and appropriate avenues by which research
progress can be made, in order to substantially advance captive
breeding techniques for marine ornamental fish species.
2. BROODSTOCK MANAGEMENT
In general, broodstock management encompasses all the ap-
propriate measures to enable captive fish to produce a high
quantity of fertilized eggs of high quality. It requires competent
knowledge in gonad maturation and spawn induction techniques
of broodstock fish, and successful egg fertilization procedures
(Mylonas et al., 2009). Marine fish, in particular coral reef fish,
occupy diverse habitat niches and display variable and complex
reproductive modes and strategies (Brock and Bullis, 2001; Holt
and Riley, 2001; Coward et al., 2002; Wittenrich, 2007). One of
the major challenges MOA faces is the need to accommodate
these reproductive strategies in order to produce a diverse range
of ornamental fish species that satisfy market demands (Watson
and Shireman, 1996; Ostrowski and Laidley, 2001).
Reproduction in broodstock fish encompasses three key de-
velopmental stages, i.e., gonad and gamete development (go-
nadogenesis and gametogenesis), final oocyte maturation, and
gamete release (ovulation/oviposition in females and ejacula-
tion of milt in males) (Schreck et al., 2001; Zohar and Mylonas,
2001; Coward et al., 2002). In reality, this process is highly
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 317
complex, involving multiple, and sometimes very different de-
velopmental stages. The quality of resulting gametes comes
under the influence of a range of variables, including aspects
of the behavioral, physical, and chemical environment (Munro,
1990; Sumpter, 1990; Coward et al., 2002; Wittenrich, 2007).
The ability to condition broodstock to maximize gamete qual-
ity and output is a major prerequisite to a successful captive
breeding program, but at the same time represents a signifi-
cant bottleneck for both foodfish and marine ornamental fish
aquaculture (Ostrowski and Laidley, 2001; Zohar and Mylonas,
2001; Coward et al., 2002; Mylonas et al., 2009). The major
critical control points in this process include careful broodstock
selection, and the combination of appropriate broodstock hus-
bandry, tank design, and manipulative techniques that induce
reproductive activity and spawning of the target fish.
2.1. Broodstock Selection
As a critical first step in obtaining a high quantity of good
quality gametes, appropriate broodfish should be chosen (Brock
and Bullis, 2001; Schreck et al., 2001; Olivotto et al., 2003). At
a basic level, broodstock can be selected based on size, age, and
external attributes; however, a well-informed choice will also
take into account social interaction and behavioral influences
on sex and sex change, morphological and biochemical char-
acteristics of the gametes produced by the broodfish, the risk
of disease transmission to other broodstock and larvae, and the
genetic background of the broodfish. All these factors are im-
portant in any foodfish hatchery and, thus, must be considered
when culturing marine ornamental fish to maximize hatchery
production and efficiency.
2.1.1. Broodstock Age and Size
In some foodfish species, adult age and spawning history has
been reported to have a bearing on egg size and chemical con-
tent of eggs produced by females, and sperm quality of males
(Vuthiphandchai and Zohar, 1999; Kamler, 2005). Gamete qual-
ity generally follows a trend in which very young and old age
females produce smaller eggs of lower nutritional content as
compared to those of average age (Kamler, 2005). Likewise,
very young and old males produce lower quality sperm com-
pared to middle aged males, particularly with respect to sperm
concentration and spermatocrit, as well as storage potential and
longevity (Vuthiphandchai and Zohar, 1999). At present, un-
fortunately, it seems no research exists regarding this in ma-
rine ornamental fish. Considering that broodstock supply for
marine ornamental fish currently relies almost exclusively on
wild caught specimens (Sales and Janssens, 2003; Pavlov and
Emel’yanova, 2006), the absence of age and spawning history
information concerning wild caught broodfish may limit the
possibility of selecting broodfish based on their age. However,
creating an information base that documents the change in ga-
mete quality with age for marine ornamental species is likely
to be useful in the future. With such information, marine or-
namental aquaculturists may be able to make more informed
choices on broodfish selection and determine when an individ-
ual is reaching the ‘retirement’ age.
The size of a marine ornamental broodfish may also be an im-
portant selection consideration, as it has been shown to influence
fecundity and egg size in many foodfish species (Bromage et al.,
1992; Kolm, 2002; Kamler, 2005). This influence is particularly
pronounced in species that display large size variation between
reproductively mature females (Kamler, 2005). The positive re-
lationship between egg size/number and broodfish size is con-
sidered largely universal among foodfish (Kamler, 2005), which
has also been observed in several marine ornamental fish. For
example, large females of the banggai cardinalfish, Pterapogon
kauderni, and the blueband goby, Valenciennea strigata,were
found to produce more and larger eggs than smaller conspecifics
(Reavis, 1997; Kolm, 2002).
2.1.2. Social Interaction and Behavior-Mediated Sex Change
When selecting broodstock to establish breeding pairs or
groups, size and age must also be considered relative to their
potential influences on social behavior (e.g., competition be-
tween males for females, agonistic, and territorial behaviors),
the social environment and sex change, particularly in social
and haremic fish (Warner, 1988; Sakai, 1997). Social and be-
havioral factors influence intra-specific interactions and the es-
tablishment of social hierarchies, affect the stability of the so-
cial environment, stress levels, sex change of individuals, and
ultimately the reproductive capacity and likelihood of sponta-
neous spawning (Warner, 1988; Fox et al., 1997; White et al.,
2002; Forrester et al., 2006; Mylonas et al., 2009). The pathway
and degree to which these interactions may influence reproduc-
tion in teleosts depends on the sexual morphology (gonocho-
rism or hermaphroditism), reproductive strategy (monogamy or
polygamy), spawning mechanism (such as oviparity and ovo-
viviparity; Coward et al., 2002), and mode of reproduction (de-
mersal or pelagic spawners) of the species concerned. Such
behavioral interactions may be accentuated in a closed system
where an individual is unable to avoid or escape, or find an al-
ternate mating partner or group. For example, sex change is a
common phenomenon among coral reef fish; a change in social
environment brought about by a sex change event, may expose
an individual or group to agonistic behaviors, causing stress, and
leading to low reproductive output, poor gamete quality, ‘repro-
ductive failure,’ or even mortality (Kjesbu, 1989; McCormick,
1998; Nordeide, 2007).
The collection and/or captive history of an individual is also
a point of consideration, particularly with respect to selecting
hermaphroditic fish. In the wild, reproductive development and
sex change that comes under the control of the social envi-
ronment and dominant conspecifics, may limit the reproductive
opportunity of an individual, or suppress maturation all together
(Fox et al., 1997). However, once collected, the individual may
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318 J. A. MOORHEAD AND C. ZENG
be liberated from the suppressive effects of its social group, and
thus, have an opportunity to progress to a reproductively ex-
pressive or receptive state. The implications of this are that the
individuals’ behavior changes from that of a submissive sex or
subordinate, to an active and potentially aggressive competitor
for reproductive opportunity. This may mean, for example, that
larger females of protogynous species will more likely change
sex, becoming a ‘bachelor male’ (Aldenhoven, 1986), and chal-
lenge or compete with male conspecifics in an artificial breeding
group, particularly if its history included long periods of seclu-
sion from dominant conspecifics. Such a case may necessitate
the choice of smaller females to facilitate successful pairing,
in favor of a more common situation of selecting larger fish
for greater fecundity and higher quality gametes. For example,
attempts to pair similar sized animals of the protogynous se-
quential hermaphrodite; bicolor angelfish, Centropyge bicolor,
often resulted in intense aggression, physical damage, or death
of one of the animals (personal observation). It appeared that
closely sized animals paid more attention to each other and
more aggressively attempted to establish dominance, compared
to when the size difference of conspecifics was large (personal
observation). The reverse scenario may be true when selecting
males of species whom are protandrous hermaphrodites.
Even in less aggressive species, the interaction between size
and behavior of male and female fish during reproduction could
influence gamete output. For example, male size in the pater-
nal mouthbrooding banggai cardinalfish, Pterapogon kauderni,
influenced the number of eggs provided by the female for in-
cubation. Females were observed to adjust their egg output, or
cannibalize excess eggs based on the size of their male partner
(Kolm, 2002). Furthermore, male size in wild blueband gob-
ies, Valenciennea strigata, was suggested to have a bearing on
fecundity and egg survivorship, leading to an active choice of
females to leave their partner, if a larger single male was avail-
able (Reavis, 1997). Similarly, it has been observed in wild blue
devil damselfish, Chrysiptera cyanea, that larger sized males
experienced greater spawning success, as male size influenced
the decision of females to spawn (Gronell, 1989).
In summary, a thorough knowledge of the way in which
broodstock size influences social interaction and reproduc-
tive behavior, and vice versa, could be critical, and a factor
to consider when selecting broodstock of marine ornamental
fish.
2.1.3. Gamete Quality
Hatchery productivity can be substantially enhanced if the
appropriate steps are taken to obtain high quality gametes
(Gim´
enez et al., 2006). On occasion, when a large number of
potential broodstock are in breeding condition, selection may be
made based on gamete quality. Assessing gametes also serves
to identify readiness for spawning and avoids selecting brood-
fish with under- or over-ripe gametes (Kjørsvik et al., 1990;
Bromage et al., 1994).
The difficulty in obtaining sufficient high quality eggs and lar-
vae is often considered a major limiting factor in foodfish aqua-
culture, particularly of marine species (Kjørsvik et al., 1990;
Brooks et al., 1997; Aristizabal et al., 2009). With respect to as-
sessing fish eggs for quality, high fertilization rate and survivor-
ship of developing eggs, and a good number of healthy larvae at
hatching and first feeding are the major criteria (Bromage et al.,
1992; Brooks et al., 1997; Lahnsteiner and Patarnello, 2005).
Unfortunately, many factors that affect egg quality in fish are
not well understood (Brooks et al., 1997). Despite the lack of
knowledge on this issue, several morphological and biochemi-
cal criteria have been used as indicators of egg quality, however,
their effectiveness appears to vary with species. Common mor-
phological and physical parameters used include size of egg,
egg yolk and lipid-vesicle size, egg shape, and cell morphology
(e.g., cell symmetry at early cleavage stages), as well as egg
buoyancy, dry weight, and larval morphology at first feeding.
The biochemical parameters include carbohydrate metabolism
and lytic enzyme activity, as well as lipid, amino acid, and/or
vitamin content of eggs (McEvoy, 1984; Carrillo et al., 1989;
Kjørsvik et al., 1990, 2003; Bromage et al., 1994; Kjørsvik,
1994; Shields et al., 1997; Lahnsteiner and Patarnello, 2003,
2004a, 2004b, 2005). Depending on species, one or a combina-
tion of these measurements may be employed to select female
broodstock before larval culture efforts commence.
Although receiving less attention, milt quality, including
sperm density, motility, and fertilizing capacity of spermato-
zoa, has also been shown as an important factor influencing
fertilization and hatching success of fish (Kamler, 2005; Pavlov
and Emel’yanova, 2006). Pavlov (2006) described a method for
assessing sperm motility in fish, which has potential for use in
screening male broodstock candidates. Pavlov and Emel’yanova
(2006) further implemented this technique in assessing three reef
fish commonly traded on the ornamental market; i.e., the brown
tang, Zebrasoma scopas, the scissortail sergeant, Abudefduf sex-
fasciatus, and the domino damselfish, Dascyllus trimaculatus,
but found little difference in sperm motility between species.
All three species tested possessed spermatozoa of compara-
tively low initial velocity, retaining activity for at least 5 min,
and showing viability after being stored for several hours below
4.5◦C. The technique may be useful for monitoring male perfor-
mance and assessing sperm quality of other marine ornamental
fish, particularly in relation to factors, such as diet, stress, size,
and age.
Since gamete quality comes under the influence of multi-
ple factors, including environmental variation, the physiolog-
ical and endocrinological status of the broodfish, their diet,
and genetic factors (Brooks et al., 1997; Coward et al., 2002;
Aristizabal et al., 2009), measurements of gamete quality can
also serve as an indicator of inappropriate captive conditions or
broodfish health. Therefore, attention should be paid to the ap-
plication of gamete quality monitoring and control techniques
in marine ornamental aquaculture, as an important step towards
quality control and the optimization of conditions conducive to
reproduction.
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 319
2.1.4. Disease and Biosecurity
Prevention, rapid detection, and control of disease outbreaks
are critical issues in all hatcheries with broodstock being one
of the primary control points for such management (Brock and
Bullis, 2001; Adams and Thompson, 2006). Pathogens that can
be vertically transmitted to gametes from broodstock include
bacteria, fungi, parasites, and, more prominently, viruses (Brock
and Bullis, 2001). Common pathogens reported to affect marine
ornamental fish, potentially posing a threat to MOA, include
bacteria such as Mycobacterium spp., Vibr i o spp., Aeromonas
spp., Pseudomonas spp., and myxobacteria, as well as parasites
such as Cryptocaryon, Amyloodinium, Uronema, and mono-
genetic trematodes (Lipton, 1993; Francis-Floyd and Klinger,
2003; Tendencia, 2004; Zanoni et al., 2008). Some of these
pathogens are identifiable and treatable (Francis-Floyd and
Klinger, 2003), however, avoiding contaminated broodfish is the
first line of defense, with gametes being sourced ideally from
pathogen-free broodstock (Brock and Bullis, 2001). Although
research continues in the development of specific pathogen-free
broodstock in many foodfish and shrimp, this is not partic-
ularly practical for marine ornamental fish at this stage of
development. Characteristically, broodstock sourced from wild
populations, such as most marine ornamental fish, have been ex-
posed to many endemic pathogens, and are then further exposed
to pathogens commonly found in aquarium and aquaculture
systems (Brock and Bullis, 2001; Francis-Floyd and Klinger,
2003). Therefore, considering the lack of certified pathogen-free
marine ornamental fish, there is a need for vigilance in identi-
fying new pathogens, and diligence to develop techniques and
maintain control over outbreaks of diseases and their vertical
transmission (Brock and Bullis, 2001; Adams and Thompson,
2006).
Apart from inspection for external signs of disease while
selecting broodstock, there are several technologies that allow
more accurate screening and identification of pathogens. These
include both traditional methods of bacteriology, virology, par-
asitology, and mycology, which are often supported by his-
tology, for the detection of easily cultured pathogens (Adams
and Thompson, 2006), and more advanced cell cultures, and
immunological and molecular methods for more complicated
cases (Villena, 2003; Adams and Thompson, 2006). For ex-
ample, investigation into screening methods for barfin flounder
(Verasper moseri) broodstock suggested that both a PCR test
and ELISA antibody titers could help prevent vertical trans-
mission of viral-nervous necrosis from broodstock (Watanabe
et al. 2000). As breakthroughs are largely made in foodfish dis-
ease management and biosecurity, the MOA industry stands to
benefit. However, considering the potential of increasing eco-
nomic importance of MOA, there is also a need for more re-
search into pathologies that affect ornamental fish specifically.
Furthermore, efforts to improve biosecurity, diagnostic support,
quarantine, and prophylactic and probiotic treatments, should
also be made as the MOA sector grows (Francis-Floyd and
Klinger, 2003; Zanoni et al., 2008).
2.1.5. External Attributes and Genetic Selection
As future progress is made in marine ornamental aquacul-
ture, broodstock selection may inherit an added dimension, that
is the quest to both track and maintain genetic pools for conser-
vation purposes (i.e., potential for restocking), and add value to
the commercialization of aquacultured marine ornamental fish.
The commercial arm of the AOI is highly selective in the species
that are collected, cultured, and traded (Lecchini et al., 2006).
Species that display vibrant coloration, unique and interesting
behaviors and habits, or some degree of novelty, in addition to
their suitability for aquaria, are key criteria dictating the value
and interests for the species traded and the level of their ex-
ploitation (Lecchini et al., 2006; Sinha and Asimi, 2007; Willis,
2007). Those species that find popularity among consumers
are often judged for quality based on appearance and physi-
cal attributes, such as coloration and finnage (Watson and Hill,
2006).
Clearly, the MOA has not evolved to the level seen in fresh-
water ornamental aquaculture; i.e., producing a large range of
hybrids and ‘fancy’ strains of fish. In its present state, MOA is
more likely to concentrate on the elucidation and manipulation
of non-genetic factors, such as diet and environmental condi-
tions, as these often precede consideration of genetic effects in
broodstock selection and management for early stages of de-
velopment of an aquaculture sector (Butts and Litvak, 2007).
However, in the long term and with the future development of
the industry, genetic manipulations will inevitably become a
very important and powerful tool.
Traditionally, efforts directed toward genetic improvement of
aquaculture broodstock have involved selective breeding tech-
niques, such as selection of favorable phenotypes, crossbreed-
ing, and hybridization (Liu and Cordes, 2004). The first concern
for commercial breeders of marine ornamental fish is likely to
obtain broodstock with attractive appearance and physical at-
tributes, in the hopes that these attributes will be expressed in
their offspring, and, thus, ensure good return for culture ef-
forts. However, the application of modern genetic techniques
for broodstock selection could be a valuable tool for the future.
Particularly as it relates to identifying and mapping monogenic,
and more so, polygenic (quantitative) gene loci associated with
desirable phenotypes (Khoo et al., 2003). Both for foodfish
aquaculture and MOA, knowledge of linkages between easily
amplified polymorphic DNA markers and polymorphic quanti-
tative trait loci (QTLs), has application in selective breeding for
improving traits of cultured species, such as growth, color, pat-
terning, finnage, and disease resistance, and is termed marker-
assisted selection (MAS) (Poompuang and Hallerman, 1997;
Liu and Cordes, 2004). Additionally, as many marine ornamen-
tal fish are sexually dimorphic and fetch a different price based
on their sex, genetic techniques may be used in the selection of
broodstock that display some degree of autosomal sex determi-
nation like that of Tilapia, or to produce monosex populations
in dioecious species (Piferrer and Lim, 1997; Poompuang and
Hallerman, 1997).
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320 J. A. MOORHEAD AND C. ZENG
On the whole, DNA marker technology and gene manip-
ulations have not impacted aquaculture in a significant way
until recently (Liu and Cordes, 2004; De-Santis and Jerry,
2007). There are few publications focused on ornamental fish
species in this area of biotechnology, and few if any QTLs
have been mapped for marine ornamental fish. However, there
is confidence that with continued progress in genomic re-
search, and in particular QTL mapping, MAS will realize sig-
nificant use for efficient and precise broodstock selection for
aquaculture, including marine ornamental fish culture, both
for commercial and conservation purposes (Fernando et al.,
1997; Taniguchi, 2003; Liu and Cordes, 2004; Chistiakov et al.,
2006).
2.2. Broodstock Husbandry and Reproduction Manipulation
Once broodstock have been obtained, appropriate condi-
tions, including diet and physical conditions, must be identified
and maintained to provide an environment that is conducive to
sexual maturation and, ideally, natural reproduction (Mylonas
et al., 2010). However, not all fish will spawn naturally un-
der captive conditions. Therefore, as an integral part of an ef-
fective broodstock management plan, invasive techniques may
become necessary for obtaining viable gametes, particularly
when a species exhibits reduced reproductive activity or spawn-
ing asynchrony under captive conditions (Coward et al., 2002;
Mylonas et al., 2009). While invasive techniques may be re-
quired, proper husbandry practices are still a major contributor
to the resulting quality of gametes (Kjørsvik et al., 1990; Brooks
et al., 1997; Coward et al., 2002). Therefore, appropriate hus-
bandry for broodstock and the need for manipulation or strip-
ping to obtain mature gametes, are not mutually exclusive with
respect to obtaining quality gametes.
Considerable attention must be paid to provide appropriate
conditions associated with reproduction in marine ornamental
fish, particularly in light of the high diversity of living and
breeding modes, and strategies displayed among species. The
main areas that often encompass broodstock husbandry and
reproductive manipulation are the control of environment, diets
and feeding, the direct administration of reproductive hormones,
and in some cases, invasive stripping techniques (Coward et al.,
2002; Melamed et al., 2002).
2.2.1. The Physical and Chemical Characteristics of the
Culture Environment
A fish’s response to and utilization of a particular habitat is
influenced by its interactions with both biotic and abiotic fac-
tors in the environment, and its ability to exploit the habitat
for the acquisition of food, refuge, or territory (Bellwood and
Wainwright, 2001; Fulton et al., 2001; Gill and Andrews, 2001;
Wilson et al., 2001). In the wild, a fish may typically choose
an appropriate niche habitat within a broader environment that
suits its needs, or be forced to move on or adapt to survive.
However, in a captive setting, a fish has very limited freedom to
choose its environment and often a more limited choice of niche
habitat. Therefore, with respect to basic husbandry of fish, the
onus is on the caregivers to provide an environment in which
the animal can live and grow well. This can be achieved through
sound knowledge of the biology of the fish and a careful assess-
ment of its tolerances and preference for certain environmental
conditions.
Similarly, reproduction in many fish also comes under the
influence of a series of environmental factors, some of which
play a specific role during reproduction periods. Therefore, fur-
ther to their basic husbandry needs, the environmental factors
that influence reproduction need to be identified and replicated
in a captive setting, in order to stimulate reproductive responses
(Coward et al., 2002). In captivity, many fish species have
successfully spawned when the correct physical and chemical
environmental conditions are provided (Kodric-Brown, 1988;
Clifton, 1995; Peter and Yu, 1997; Gordon and Bok, 2001;
Holt and Riley, 2001; Zohar and Mylonas, 2001; Olivotto et al.,
2006a). Among many factors, tank design, substrate, and water
conditions are some of the most important considerations.
2.2.1.1. Tank Design. On a broad scale, the aquaculture in-
dustry relies heavily on what has been classified as undomesti-
cated stocks. Therefore, system engineering and successful cap-
tive culture often hinges on the animal’s ability to cope and adapt
to artificial surroundings (Koolhaas et al., 1999; Rasmussen
et al., 2005; Watson and Hill, 2006). With this in mind, greater
success may be achieved in encouraging target species to spawn
under controlled conditions if the engineered environment more
closely replicates natural conditions in which the animal is com-
monly found.
At present, both industry and researchers of marine orna-
mental fish aquaculture show distinct favor towards culture
of demersal spawners, particularly members of the families
Pomacentridae, Gobiidae, Sygnathidae, and Pseudochromidae
(Fig. 1). These species tend to form strong pair bonds, pro-
duce regular clutches of large eggs, and display some degree of
parental egg care (Brown et al., 2003; Watson and Hill, 2006).
One reason for such a trend appears to be the ready acceptance of
many demersal fish for relatively simple, space efficient tank de-
signs, and artificial spawning substrate. Clownfish, or anemone-
fish (Amphiprion spp.), are a classic example, which have been
shown to spawn in tanks as small as 37 L, attaching their ben-
thic eggs to terracotta pots or tiles, as substitutes for natural rock
surfaces (Hoff, 1996; Wittenrich, 2007). However, overall, tank
design remains a stumbling block for product diversification of
MOA.
In general, tank design has a significant effect on the physical
characteristics of the holding environment, and the behavior of
the organisms being held (Ross et al., 1995; Rasmussen et al.,
2005). The influence of tank design on growth performance is a
recognized concept in grow-out aquaculture (Ross et al., 1995;
Rasmussen et al., 2005). It has also been recognized that tanks
proven adequate for growth may be inadequate for broodstock
conditioning (Ostrowski and Laidley, 2001).
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 321
Figure 1 (A) Percentage of demersal and pelagic spawning marine ornamental fish species that have been cultured commercially or as a part of research; and
(B) proportion of various families within demersal spawning species that have been cultured commercially or as a part of research. (Data drawn from 49 sources;
principal, Arvedlund et al., 2000; Wabnitz et al., 2003.)
Tank size, water volume, and depth, as well as stocking
density, can all influence the reproductive success of the fish
being held (Mylonas et al., 2010). For example, Planas et al.
(2008) reported that for the Spiny seahorse, Hippocampus gut-
tulatus, deep tanks and good plumbing design eliminated the
occurrence of gas bubble disease; a commonly occurring ail-
ment in seahorses kept in shallow tanks with fine water bub-
bles, and facilitated breeding activity. In assessing the specific
needs of broodstock fish, it may be useful to examine and
collate information recorded from field studies. For example,
Lecchini et al.(2003) pointed out that there are definite pat-
terns in the spatial distributions and abundance of reef fishes,
which in many instances are not only explained by biological
factors, but also physical factors, such as water depth and hy-
drodynamic conditions, including water movement and wave
exposure. An understanding of a fish’s natural distribution and
abundance gradients may give insight into its specific physi-
cal requirements. Important clues as to the physical require-
ments of certain reef fish may also be found through under-
standing their physiological capabilities. For example, studies
on the swimming capabilities of wrasses (Labridae) showed
that swimming performance, as a function of pectoral fin as-
pect ratio (pectoral fin aspect ratio =(length of pectoral fin
leading edge)2/total fin area; Fulton et al., 2001), influences
their distribution (Bellwood and Wainright, 2001; Fulton et
al., 2001). When applying this knowledge to the engineering
of a captive environment, inferences may be made as to the
suitable current regimes that should be created for broodstock
of these fish. It is unclear whether strong swimming fish may
prefer environments of high wave and current energy at every
stage of their life history and especially during spawning. How-
ever, it may be postulated with some confidence that fish with
weak swimming performance (possessing a low fin aspect ra-
tio), may fail to progress through certain stages of reproductive
maturity if faced with a stressful, high turbulence environment
(Watson and Hill, 2006). It should also be noted that an envi-
ronment in which a fish is commonly found might be different
to where it spawns. Once gonad maturity is reached, some fish
may require a change in wave and current regimes, tank dimen-
sions, or even water quality, to initiate gamete release (Mylonas
et al., 2009). In summary, replication of the physical environ-
ment that typifies a species’ preferred niche in the wild may re-
duce stress and encourage broodstock to progress through early
gonad development, maturation, and finally gamete release in
captivity.
It has been postulated that as focus shifts from benthic spawn-
ing species to more popular and higher value pelagic-spawning
marine ornamentals, such as pygmy angelfish (Centropyge spp.),
surgeonfishes (Acanthuridae) and wrasses (Labridae), a move
from shallow tank designs, towards larger and/or deeper designs
has to occur (Ostrowski and Laidley, 2001). However, there are
conflicting reports in this regard. For example, Job et al. (1997)
recommend a tank size of 1,000 L for the bicolor angelfish,
Centropyge bicolor, while Olivotto et al. (2006a) noted that a
300 L tank was adequate to elicit natural spawning in the closely
related lemonpeel angelfish, Centropyge flavissimus, despite the
absence of several courtship behaviors seen in the wild (Bauer
and Bauer, 1981).
Broadly, despite ongoing development towards larger tank
designs to accommodate pelagic spawning species, hatcheries
built for marine ornamentals are still much smaller than that used
for foodfish, but often require more technologically advanced
equipment (Watson and Hill, 2006). Although at present, the
culture of some marine ornamental fishes may demand expen-
sive system designs and techniques to elicit spawning, Mylonas
et al. (2009) suggested that the domestication process is likely
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322 J. A. MOORHEAD AND C. ZENG
to facilitate spontaneous spawning in captivity, which may help
to simplify system design in the future.
2.2.1.2. Substrate. Vast majorities of marine ornamental fish
depend upon reefs for food, shelter and for larval settlement.
Provisions of suitable substrate that simulate natural reef struc-
ture are therefore very important in marine ornamental aqua-
culture. More importantly, they are crucial in eliciting a natural
spawning response in some fish species, particularly benthic
spawning species (Friedlander, 2001; Ostrowski and Laidley,
2001). However, different views exist on substrate type and ef-
fectiveness. Natural live substrate has been used successfully
to facilitate spawning of marine ornamental fishes in captivity.
For example, Olivotto et al.(2006a, 2003) successfully spawned
lemonpeel angelfish, Centropyge flavissimus, and the yellow-
tailed damselfish, Chrysiptera parasema using live rocks. And
a study on the banggai cardinalfish, Pterapogon kauderni,even
utilized live sea urchins (Diadema sp.), an animal the banggai
cardinalfish exploits for protection in the wild (Allen, 2000),
during breeding experiments. Despite successes with a natu-
ral substrate, such as live rock, their use may not be ideal for
commercial production of marine ornamental fish. This is be-
cause natural live substrate is likely to carry a range of nuisance
and pathogenic organisms that could severely affect broodstock
health, and directly or indirectly affect gamete viability, fertiliza-
tion, and hatching rate (Brock and Bullis, 2001; Ostrowski and
Laidley, 2001). In their experiments leading to captive spawning
of the flame angelfish Centropyge loriculus, and the potter’s an-
gelfish, Centropyge potteri, Ostrowski and Laidley (2001) made
use of artificial structures, such as PVC scaffolding, and noted
the ease in which they could be removed, cleaned, and sterilized
to reduce the pathogen loading in broodstock holding systems.
Similarly, successful conditioning and spawning of the bicolor
angelfish, Centropyge bicolor, the blueband goby, Valencien-
nea strigata, and the blue devil damselfish, Chrysiptera cyanea,
has been achieved with the use of artificial substrate including
stacked PVC pipe and terracotta roof tiles (personal observa-
tion). Since mass production of marine ornamental fish is the
aim of commercial ventures and also for conservation initia-
tives, it seems obvious that research should weigh toward the
development of artificial analogues to natural reef structures for
captive breeding.
2.2.1.3. Water Conditions. Control and manipulation of the
physical and chemical characteristics of the culture water is criti-
cal for the husbandry and conditioning of broodstock fish (Brock
and Bullis, 2001; Schreck, 2001; Sales and Janssens, 2003). Un-
like many foodfish species, most marine ornamentals come from
ecosystems that display chemical stability, or ‘oligotrophic’ con-
ditions (Watson and Hill, 2006). Broadly speaking, water quality
parameters may be roughly drawn from literature that investi-
gates the tolerances of marine fishes. For example, a review
by Camargo et al. (2005) suggests that NO3-N be maintained
at less than 20 mgL−1for most marine fishes. Unfortunately,
very few scientific documents detail the specific tolerances of
marine ornamental fishes to water quality. It is useful, therefore,
to draw general guidelines for marine ornamental broodstock
based on maintenance conditions described in literature, partic-
ularly where successful spawning has occurred (Table 1).
When attempting to trigger gonad maturation and/or illicit
spawning, manipulations of temperature and photoperiod are
often effective for many teleost fish, including marine ornamen-
tals, and these factors can also influence fecundity and gamete
quality (Carrillo et al., 1989; Pankhurst et al., 1996; Peter and
Yu, 1997; Koger et al., 1999; Holt and Riley, 2001; Coward
et al., 2002; Holt, 2003; Kamler, 2005; Mylonas et al., 2009).
It has been reported that changes, particularly increases in tem-
perature and photoperiod, that simulate the diurnal and seasonal
characteristics of the breeding season, often trigger reproductive
activities in fish, particularly for those dwelling in more tem-
perate regions (Hoff, 1996; Richardson et al., 1997; Boef and
Le Bail, 1999; Gordon and Bok, 2000; Holt and Riley, 2001).
For example, Olivotto et al. (2006a) reported that manipula-
tion of photoperiod and temperature in captivity resulted in a
spawning response from the lemonpeel angelfish, Centropyge
flavissimus. Research has also shown that spawning periodicity
and frequency of various anemonefish (Amphiprion spp.) ap-
pears dependant on temperature and photoperiod (Hoff, 1996;
Richardson et al., 1997; Gordon and Bok, 2000).
However, in addition to photoperiod, lighting conditions also
include light intensity and spectral quality, both of which have
received less attention compared to photoperiod. According to
Boef and Le Bail (1999), light intensity and spectral quality
can have an affect on fish growth. These characteristics of light
vary with water depth, and the receptiveness and reaction to
change in light intensity and spectral quality varies with fish
species (Boef and Le Bail, 1999). There is no known literature
that investigates the affect of light intensity or spectral quality
on reproductive performance of broodstock of marine ornamen-
tals. However, this concept may be an important factor in a
captive setting as incorrect intensity and spectral quality may
stress fish. Conversely, light intensity and spectra that simulate
certain water depths where natural spawning occurs may be a
key factor stimulating gonad maturation and gamete release of
some species maintained in often shallow broodstock tanks.
For the basic husbandry of marine ornamental fish, a simplis-
tic but effective approach to water quality is to maintain values
within the ranges that the animals experience in the wild. How-
ever, considering that water quality ideals and tolerances vary
with species, age, and associated interactions among parameters
(Poxton and Allouse, 1982; Cuenco et al., 1985; Woods, 2003),
future research taking these factors into account is clearly ben-
eficial.
Although the water quality requirements of marine orna-
mental fish are generally uncompromising compared to that of
many foodfish, the unique economics, and the often significantly
smaller systems used in ornamental aquaculture allow the incor-
poration of expensive and precise system components to main-
tain high water quality, while remaining profitable (Watson and
Hill, 2006). However, research efforts are required to determine
which and how water quality parameters may affect aspects of
reproductive performance of a particular marine ornamental fish
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Tab l e 1 List of species-specific water quality conditions of marine ornamental fish where captive reproduction has occurred
Salinity, Temperature, Ammonia Nitrite Nitrate Dissolved Photoperiod
Common Name Species =pH ◦C (TAN), mgL−1(NO−
2), mgL−1(NO−
3), mgL−1Oxygen, mgL−1(Light:Dark) References
Clown anemonefish Amphiprion percula 28–34 — 24.0–28.0 — — — — — ¨
Onal et al., 2008; Gordon and
Hecht, 2002
False clown anemonefish Amphiprion ocellaris 26–30 8.0–8.5 25.5–30.0 <0.02 <0.02 — — 15h:9h Furuta et al., 2005; Yasir and Qin,
2007; Avella et al., 2007
Black anemonefish Amphiprion melanopus 33–35 8.0–8.2 27.0–30.0 — — — — 14h:10h Arvedlund et al., 2000; Green and
McCormick, 1999
Yellowtail anemonefish Amphiprion clarkii 30 8.0–8.2 27.7–28.5 <0.03 <0.03 — — 14h:10h Olivotto et al., 2008a, 2008b
Sebae anemonefish Amphiprion sebae 33–35 — 28.0–32.0 — — — — — Ignatius et al., 2001
Skunk anemonefish Amphiprion akallopisos 27–33 8.0–8.1 25.0–31.0 0 0 <6 — 12–14h:12–10h Gordon and Bok, 2001
Twoband anemonefish Amphiprion bicinctus — — 25.0–26.0 — — — — 12h:12h Maroz and Fishelson, 1997
Black damsel Neopomacentrus cyanomus 22–28 7.5–8.5 24.0-32.0 <0.1 — — 4.5–6.5 — Rajasekar et al., 2009
Yellow tailed damsel Neopomacentruc nemurus 22–28 7.5–8.5 24.0–32.0 <0.1 — — 4.5–6.5 — Rajasekar et al., 2009
Blue damsel Pomacentrus caeruleus 22–28 7.5–8.5 24.0—32.0 <0.1 — — 4.5–6.5 — Rajasekar et al., 2009
Domino damsel Dascyllus trimaculeatus 22–28 7.5–8.5 24.0–32.0 <0.1 — — 4.5–6.5 — Rajasekar et al., 2009
Yellow-tailed damselfish Chrysiptera parasema 28–30 8.2 27.0–28.0 — — — — 13h:11h Olivotto et al., 2003
Blue Devil damselfish Chrysiptera cyanea 29–37 8.0–8.2 27.5–29.5 <0.02 <0.02 <6 — Natural Unpublished data from author
Semicircle angelfish Pomacanthus semicirculatus 30–33 — 22.8–31.7 — — — — 12h:12h Leu et al., 2009
Cherubfish Centropyge argi 32–36 8.2 20.0–24.0 <0.03 <0.03 — — 10–13h:14–11h Bauer and Bauer, 1981 Holt and
Riley, 2001
Orangeback angelfish Centropyge acanthops 27–30 — 20.0–24.0 — — — — — Bauer and Bauer, 1981
Twospined angelfish Centropyge bispinosus 27–30 — 20.0–24.0 — — — — — Bauer and Bauer, 1981
Orange angelfish Centropyge fisheri 27–30 — 20.0–24.0 — — — — — Bauer and Bauer, 1981
Lemonpeel angelfish Centropyge flavissimus 27–36 8.2 20.0–30.0 <0.03 <0.03 — — 13h:11h Bauer and Bauer, 1981; Olivotto et
al., 2006a
Flame angelfish Centropyge loriculus 27–30 — 20.0–24.0 — — — — — Bauer and Bauer, 1981
Blue Mauritius angelfish Centropyge debelius 32 8.2 22.0–24.0 0 0 <10 — 11–14h:13–10h Baensch and Tamaru, 2009
Spiny seahorse Hippocampus guttulatus 34–39 7.8–8.3 15.0–25.0 <0.1 <0.03 <10 — 14–15.5h:10–8.5h Faleiro et al., 2008; Planas et al.,
2008
Long-snouted seahorse Hippocampus reidi 27–30 8.0–8.4 22.0–28.5 <0.03 <0.03 — — 12–14h:12–10h Hora and Joyeux, 2009; Olivotto et
al., 2008c
Lined seahorse Hippocampus erectus 34–36 7.5–8.1 27.5–28.5 — — — 6–7 14h:10h Lin et al., 2008
Coral seahorse Hippocampus barbouri 33–36 7.9–8.2 25.1–26.8 <0.05 <0.1 — — — Payne, 2003
Big belly seahorse Hippocampus abdominalis 33–35 7.9–8.2 10.6–19.5 — — — 7.73–8.97 15h:9h Woods, 2000b, 2000a
Three spotted seahorse Hippocampus trimaculatus 31–34 7.3–7.9 30.2–30.3 0 <0.07 — 5.95–6.7 16h:8h Murugan et al., 2009
Common seahorse Hippocampus kuda 32–33 8.3–8.7 26.0–29.3 <0.05 <0.05 <0.05 7.4–7.8 — Lin et al., 2006, 2007
White’s seahorse Hippocampus whitei — — 20.0 — — — — 12h:12h Wong and Benzie, 2003
West Australian seahorse Hippocampus subelongatus 35 8.1–8.3 22.5–23.5 <0.1 <0.1 — ∼7.7–8.5 12h:12h Payne and Rippingale, 2000
Knysna seahorse Hippocampus capensis 25 — 22.0 — — — — 16h:8h Lockyear et al., 1997
Sunrise dottyback Pseudochromis flavivertex 30 8.0–8.2 26.5–27.5 <0.03 <0.03 — — 14h:10h Olivotto et al., 2006b
Cleaner goby Gobiosoma evelynae 30 8.2 25.0 <0.03 <0.03 — — 13h:11h Olivotto et al., 2005
Blueband goby Valenciennea strigata 29–37 8.0–8.2 27.5–29.5 <0.02 <0.02 <6 — Natural Unpublished data from author
Blackline fangblenny Meiacanthus nigrolineatus — 8.0 23.0–25.0 — — — — — Fishelson, 1975
Striped blenny Meiacanthus grammistes 30 8.2 28.0 <0.03 <0.03 — — 13h:11h Olivotto et al., 2010
Forktail blenny Meiacanthus atrodorsalis 29–37 8.0–8.2 27.5–29.5 <0.02 <0.02 <6 — Natural Unpublished data from author
White Tiger goby Priolepis nocturna 32 8.2 30.0 <0.02 <0.02 <20 — 14h:10h Wittenrich et al., 2007
Barber goby Elacatinus figaro 34 8.2 26.0 <0.02 <2.0 — — 12h:12h Shei et al., 2010
Banggai cardinalfish Pterapogon kauderni 34–36 7.8–8.0 24.5–25.5 0 0 — — 12h:12h Vagelli, 1999
Spotfin hogfish Bodianus puchellus 32–36 8.2 22–25.5 <0.03 <0.03 — — 11–13h:13–11h Holt and Riley 2001
Bluehead wrasse Thalassoma bifasciatum 32–36 8.2 22.0–29.0 <0.03 <0.03 — — 10–13h:14–11h Holt and Riley 2001
Clown wrasse Halichoeres maculipinna 32–36 8.2 25.0–27.5 <0.03 <0.03 — — 11–13h:13–11h Holt and Riley 2001
323
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324 J. A. MOORHEAD AND C. ZENG
in a captive setting. This should also include chemical inputs that
are for therapeutic or prophylactic treatments. Such treatmeants
are likely to be applied more regularly in marine ornamental
operations as compared to more strictly regulated foodfish op-
erations. For instance, the heavy metal copper, an active ingredi-
ent in copper sulphate solutions for treating pathogens (Lipton,
1993), has been found to accumulate in tissues, impact phys-
iological and osmoregulatory mechanisms of fish, and at high
concentrations, cause death (Dethloff et al., 1999; Burridge and
Zitko, 2002; Grosell et al., 2004; Oliva et al., 2007). Although
trace levels of copper have been shown to benefit larval culture
of false clownfish, Amphiprion ocellaris, overdose is likely to
have lasting detrimental effects (Furuta et al., 2005). This high-
lights the need for caution and the necessity to develop alterna-
tive treatments, such as vaccination, biocontrols, and probiotics,
which are showing promise in foodfish aquaculture (Gatesoupe,
1999; Vine et al., 2004; Bondad-Reantaso et al., 2005; Adams
and Thompson, 2006).
2.2.2. Broodstock Diet
Compared to foodfish, research related to the dietary nu-
tritional needs of ornamental fish, and the production of di-
ets specifically designed for broodstock are lagging far behind
(James and Sampath, 2004a, 2004b). Despite the obvious signif-
icance of such research, information concerning the nutritional
requirements of both freshwater and marine ornamental fish is,
at best, patchy (Blom and Dabrowski, 2000; Sales and Janssens,
2003; Vijayagopal et al., 2008). The foods used to feed ma-
rine ornamentals are often not specifically designed for them,
but rather for foodfish (Tamaru et al., 2001). Formulated di-
ets that are commercially available and claim to be designed
for ornamental fish are generally packed in expensive small
packages targeting the hobbyist market, making them econom-
ically unsuitable for large-scale operations (Chong et al., 2003;
Mosig, 2007). Moreover, the creditability of such claims is also
doubtable. Clearly, for marine ornamental culture, broodstock
dietary requirements, including food type, ration, feeding fre-
quency, and the nutritional contents of diets, should receive
closer attention as it is known to influence the general health,
condition, fecundity, and gamete and larval quality of marine
fish (Brooks et al., 1997; Izquierdo et al., 2001; Kamler, 2005;
Donelson et al., 2008).
2.2.2.1. Food Type. On the whole, in order to achieve con-
sistent performance from broodstock, there is an industry-wide
push in aquaculture to develop fully formulated feeds with
known nutritional compositions. However, at present even for
the foodfish industry, few formulated diets completely satisfy
the dietary needs (including palatability and nutritional content)
of broodstock fish. As a result, formulated feeds are often used as
a supplement to fresh or frozen raw natural foods, such as ‘trash
fish,’ when feeding broodstock fish in the hatchery (Izquierdo
et al., 2001).
Likewise, based on their feeding habits, marine ornamental
broodstock fish are generally fed one or a combination of raw
or processed natural foods, such as squid, fish, mussels, prawns,
Artemia, aquatic or terrestrial plant matter, supplemented with
formulated diets or cultured live prey (e.g., live Artemia or
mysis) for conditioning (Job et al., 1997; Ignatius et al., 2001;
Ostrowski and Laidley, 2001; Olivotto et al., 2003, 2005, 2006a,
2006b; Hopkins et al., 2005; Wittenrich et al., 2007). The need
to feed broodstock fish with raw or live prey is not only based
on a concern for nutritional requirements, but also in response
to higher acceptability and palatability of raw or live foods
observed over formulated diets. This phenomenon is proba-
bly also linked to the wild origins of most marine ornamental
broodstock, particularly when their natural diet is made up of
live organisms. Members of the family Syngnathidae, including
seahorses and pipefishes, are a prime example of marine orna-
mentals that display strict and almost obligate preference for
live or frozen feeds that resemble their natural prey, particularly
when the broodstock are sourced from the wild. In freshwater
ornamental aquaculture, even for species that are considered
fully domesticated, live foods are still fed either exclusively or
commonly with formulated diets (Lim et al., 2003; James and
Sampath, 2004b).
This reliance on raw/live feeds for marine ornamental brood-
stock poses several potential areas for investigation. First, it
may be that this reliance dissipates with successive generations
of captive breeding, therefore, investigation into how the do-
mestication process may affect preference and acceptability of
diet types may prove worthwhile. Second, processed foods that
emulate the physical and chemical features of live natural foods
may need to be developed. This could include the addition of
chemical attractants and the incorporation of visual stimuli to
encourage feeding behavior, i.e., arousal, search, and consum-
mation (Davis et al., 2006). Finally, there is a clear disadvantage
in the use of live/raw foods, owing to the risk of disease trans-
mission and often variable reproductive performance as a result
of inconsistent nutritional values of such diets (Izquierdo et al.,
2001; Sales and Janssens, 2003). Therefore, the development
of formulated diets that are not only acceptable but also meet
the specific nutritional needs of broodstock fish should be the
ultimate goal in marine ornamental aquaculture.
2.2.2.2. Ration and Feeding Frequency. Feeding ration and
frequency are also important considerations when conditioning
broodstock fish (Izquierdo et al., 2001). They can ultimately
affect the reproductive performance of broodstock and the qual-
ity of gametes and larvae produced (Izquierdo et al., 2001;
Donelson et al., 2008). Food restriction can negatively affect re-
production, resulting in delayed gonadal maturation and spawn-
ing (Izquierdo et al., 2001), increased instances of filial cannibal-
ism of eggs under parental care (Okuda and Yanagisawa, 1996;
Okuda et al. 2004), and the production of smaller larvae that dis-
play poor survival (Donelson et al., 2008). On the other hand,
overfeeding can also lead to food wastage, resulting in water
quality problems, which could in turn lead to poor reproductive
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 325
performance (Chang et al., 2005). Although a safe default is to
feed fish regularly to satiation, significant savings in feed and
labor costs can be achieved if feeding ration and frequency are
optimized, possibly in conjunction with the use of automatic
feeders.
2.2.2.3. Broodstock Nutrition. The nutrition of broodstock
fish food can significantly affect reproductive performance as
well as the quality of gametes and larvae they produce (Izquierdo
et al., 2001; Tocher, 2003; Watanabe and Vassallo-Agius, 2003;
Chong et al., 2004; Khan et al., 2005; Jaya-Ram et al., 2008).
Unfortunately, this is not a particularly well-studied area in
finfish aquaculture (Izquierdo et al., 2001; Khan et al., 2005),
with a majority of studies focusing on nutritional requirements
for somatic growth (Izquierdo et al., 2001). The lack of studies
on foodfish broodstock nutrition is probably related to the high
cost, long experimental durations, and large facilities required
to undertake such studies (Brooks et al., 1997; Izquierdo et al.,
2001).
Research on foodfish to date has shown that fecundity and
gamete quality in finfish can be optimized by maintaining
proper protein and lipid levels in their diets (Pustowka et al.,
2000; Emata and Borlongan, 2003; Tocher, 2003; Watanabe
and Vassallo-Agius, 2003; Khan et al., 2005). It has been
further shown that reproductive performance and gamete and
larval quality in fish are influenced by levels of certain es-
sential fatty acids, such as n-3 and n-6 highly unsaturated
fatty acids (HUFA), vitamin E, vitamin C, and carotenoids
(Sandnes, 1984; Ciereszko and Dabrowski, 1995, 2000; Bell et
al., 1996; Dabrowski and Ciereszko, 1996, 2001; Asturiano et
al., 2001; Izquierdo et al., 2001; Coward et al., 2002; Watanabe
and Vassallo-Agius, 2003; Dom´
ınguez et al., 2005; Jaya-Ram,
et al., 2008; Furuita et al., 2009). For example, females of the
freshwater ornamental swordtail, Xiphophorus helleri, require
a dietary protein level of at least 30% to support reproductive
processes (Chong et al., 2004). A recent study of another fresh-
water ornamental, the zebrafish, Danio rerio, has also shown
that a direct supply of dietary HUFA, such as docosahexaenoic
acid (DHA), eicosapentaenoic acid (EPA), and arachidonic acid
(ARA), to female diets, benefits reproduction (Jaya-Ram et al.,
2008). In contrast to foodfish and freshwater ornamental fish,
little research appears to have been done to elucidate the specific
nutritional needs of marine ornamental broodfish.
Out of consideration for cost, it is common practice in fresh-
water ornamental aquaculture that inert diets formulated for
foodfish are used, instead of commercially available ornamental
diets, and often the same diet is used for both growout and condi-
tioning broodfish (Garci´
a-Ulloa and G´
omez-Romero, 2005). If
a specified diet does exist for an ornamental fish, they are often
formulated based on extrapolation of formulations for foodfish,
and are often too expensive to use at a production level (Sales
and Janssens, 2003). For marine ornamentals, a commonly prac-
ticed ‘rule of thumb’ approach is to combine various diets in
an attempt to satisfy the nutritional requirements of broodfish.
Considering that the nutritional requirements of fish can vary
substantially between species and at different life stages, there
will be obvious benefits if specific diets could be developed
for broodstock of marine ornamental fish based on vigorous
research.
At present, there is a lack of high quality formulated foods
for ornamental fish (Chong et al., 2003; James and Sampath,
2004b); therefore, it comes as no surprise that there are numer-
ous problems with the sole use of inert feeds, and that raw and
live feeds are considered superior to inert feeds for broodstock
conditioning (Degani, 1993; Chong et al., 2003; Kaiser et al.,
2003; Velu and Munuswamy, 2003; James and Sampath, 2004b;
Garci´
a-Ulloa and G´
omez-Romero, 2005). Hence, there is an ur-
gent need for research on nutritional requirements of marine
ornamental fish, which in turn will allow the formulation of
appropriate diets. It is true that at the present, research in this
area for marine ornamentals lags behind that of foodfish. How-
ever, their smaller size, relative ease of maintenance, ability to
reach sexual maturity in a shorter time period, and often short
spawning intervals, means that studies on broodstock nutrition
in marine ornamental fish can be relatively easy, and, hence,
could quickly catch up and serve as ideal models for closely
related foodfish species.
2.2.3. Spawning Induction and Stripping
In basic terms, final oocyte maturation and ovulation in
teleosts requires a surge in the secretion of gonadotropins
(GtHs), particularly luteinizing hormone (LH), by the pituitary
gland (Brooks et al., 1997; Ng et al., 1997; Mylonas et al., 2009).
In some species, environmental manipulation to stimulate final
oocyte maturation and ovulation may fail. First, this may be due
to the fact that creating the correct set of conditions to elicit
the release of gonadotropin by broodstock fish may be unviable
or beyond current knowledge (Zohar and Mylonas, 2001; Hill
et al., 2009; Mylonas et al., 2009). And, second, fish in captivity
may display severe physiological responses to stressors present
in captive environments, antagonizing processes leading to ovu-
lation (Schreck et al., 2001; Zohar and Mylonas, 2001). In such
cases, hormone treatments and/or manual stripping may be used
to initiate final phases of oocyte maturation and to obtain ga-
metes if they are not released naturally (Schreck et al., 2001;
Zohar and Mylonas, 2001; Coward et al., 2002; Pavlov, 2006).
Even for species that can spawn naturally in captivity, hormonal
treatments and stripping can still be used as management tools
to enhance hatchery efficiency (Mylonas et al., 2009).
Fish species that do require hormone and/or invasive treat-
ments to obtain viable gametes generally experience one of
following three ‘reproductive problems’: (1) the failure to com-
plete vitellogenesis or spermatogenesis; (2) the absence of final
oocyte maturation; and/or (3) the absence of spawning (Zohar
and Mylonas, 2001). In foodfish aquaculture, many species, in-
cluding the European eel, Anguilla anguilla, the grey mullet,
Mugilis mugilis, the rainbow trout, Oncorhynchus mykiss, and
the striped bass, Morone saxatilis, encounter at least one of
these problems and thus require hormone induction (Bromage
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326 J. A. MOORHEAD AND C. ZENG
et al., 1992; Zohar and Mylonas, 2001). Freshwater ornamental
species, such as the red-tailed black shark, Labeo bicolor, and
rainbow shark, Epalzeorhynchos frenatus, are also examples of
species that display the inability to reach final oocyte maturation
and/or ovulation under captive conditions, and thus require hor-
mone treatments (Watson and Shireman, 1996; Hill and Yanong,
2002; Brown et al., 2003; Hill et al., 2005). Hormone treatments
have also helped achieve year-round production for seasonally
spawning species in freshwater ornamental aquaculture (Burten
et al., 1998; Brown et al., 2003).
With respect to marine ornamental fish, hormone induction
has not been used extensively and only a handful of reports exist
on the subject (Ostrowski and Laidley, 2001). However, to date,
results appear promising. For example, in combination with strip
spawning, Moe (1997) utilized human chorionic gonadotropin
(hCG) to successfully obtain viable gametes from wild-caught
French and grey angelfish (Pomacanthus spp.) and Emel’yanova
et al. (2006) induced maturation and ovulation in the scopas
tang, Zebrasoma scopas, via double injection of the hormone,
surfagon.
It must be noted, however, that many marine ornamental
fish are smaller compared to either Pomacanthus or Zebrasoma
species. This will not only add difficulty in injecting, but also
cause a higher degree of stress in small fish from handling
and delivery of hormones via injection, and thus may hinder
or antagonize the hormone induction process (Clearwater and
Pankhurst, 1997; Hill et al., 2005). To negate these problems,
methods of less- or non-invasive hormone administration that
have been used in foodfish and freshwater ornamental aquacul-
ture, may be adopted as viable alternatives for marine ornamen-
tal fish (Thomas and Boyd, 1989; Burton et al., 1998; Roelants
et al., 2000). For instance, Burton et al. (1998) demonstrated
that Artemia-mediated delivery of gonadotropin-releasing hor-
mone analogue (GnRHa) to broodstock of the freshwater car-
dinal tetra, Paracheirodon axelrodi, could successfully induce
ovulation, if the Artemia were exposed to a 33.3% GnRHa so-
lution for 30–60 min prior to feeding them to the fish. Similarly,
hormones could be delivered via formulated diets fed to brood-
stock fish, and may yield similar results as Artemia-mediated
delivery with an additional advantage of precise dosage control.
Furthermore, investigations by Hill et al. (2005) into the topi-
cal gill application of ovaprim, dissolved in a dimethyl sulfox-
ide solvent, to the red-fin shark, Epalzeorhynchos erythrurus,
broodstock also showed that this less invasive method could
be effective in inducing ovulation. Such techniques may prove
effective if adapted for marine ornamental broodstock condi-
tioning, and thus stand as a potential avenue for further research
in marine ornamental aquaculture. Furthermore, for ornamental
species that are priced based on the sex of the animal, hormones
may also be used for the production of mono-sex populations
(Piferrer and Lim, 1997).
Despite their many advantages, hormone induction tech-
niques could however have detrimental effects on egg fertiliza-
tion and hatching rates. Such negative effects are often linked to
inappropriate adoption of methods; incorrect dosage and timing
of hormone administration/stripping, stresses incurred by the
administration and failure to fertilize eggs soon enough after
ovulation (Mylonas et al., 1992; Ohta et al., 1996; Coward et al.,
2002; Hill et al., 2005; Kamler, 2005; Mylonas et al., 2009).
Therefore, hormone induction techniques and stripping should
be used as a last resort only after efforts to induce natural spawn-
ing in marine ornamental broodstock fishes prove unsuccessful.
3. GAMETE PHYSIOLOGY, EMBRYOLOGY, AND
LARVAL MORPHOLOGY AT HATCH
Development of captive breeding techniques for targeted
aquaculture species requires a sound knowledge of their gamete
physiology, embryology, and early larval ontogeny (Coward
et al., 2002). Such baseline studies could help in the estab-
lishment and refinement of broodstock management and larval
rearing techniques. For instance, developing indices and criteria
or benchmarks unique to target species for gauging the quality
of gametes, the development rates and characteristics of em-
bryos, or newly hatched larvae, could provide information to
help monitor broodstock reproductive health and assess larval
quality prior to a rearing attempt (Coward et al., 2002; Kamler,
2005; Yasir and Qin 2007; ¨
Onal et al., 2008). Such knowledge
may include gamete size, embryonic development stages and
durations, larval size and stage of development, endogenous re-
serves or condition indices at hatch, and functional morphology,
such as mouth gape at first feeding and ontogeny of sensory or-
gans, as well as swimming ability of larvae (Brooks et al., 1997;
Kamler, 2005).
Embryology studies may also be extrapolated to refine man-
agement techniques. For example, observations of the clownfish
Amphiprion ocellaris by Yasir and Qin (2007), suggest that the
rate of ‘body turnover’ in late stage eggs dictates hatching suc-
cess. It is also accepted that the biotic and abiotic environment
in which eggs are incubated has a bearing on embryonic de-
velopment, and eventually the hatching rate and viability of
the newly hatched larvae (Brooks et al., 1997; Skjermo and
Vadstein, 1999; Olafsen, 2001). Therefore, baseline knowledge
of embryology and early larval ontogeny will help identify how
the environment may affect developing embryos and resulting
larvae, helping develop techniques to improve hatching rate and
survival of newly hatched larvae (Kamler, 2005). To date, basic
information is available for some marine ornamental species
with respect to gamete physiology, embryo development and
ontogeny of newly hatched larvae (Fishelson, 1976; Green and
McCormick, 2001; Gordon and Hecht, 2002; Olivotto et al.,
2003, 2005, 2006a, 2006b; Wittenrich et al., 2007; Yasir and
Qin, 2007; ¨
Onal et al., 2008). Further research to elucidate
species-specific developmental characteristics, environmental,
physiological and immunological capabilities and tolerances of
embryos and newly hatched larvae should benefit the establish-
ment and refinement of rearing techniques.
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 327
4. LARVAL REARING
Larval rearing has presented itself as a major challenge and
a persistent bottleneck for contemporary aquaculture develop-
ment (Dhert et al., 1998; Planas and Cunha, 1999; Holt, 2003;
Papandroulakis et al., 2004; Battaglene and Cobcroft, 2007).
The need to develop techniques for rearing fish past critical lar-
val stages is essential to success in commercial production of
any aquaculture species, and marine ornamentals are no excep-
tion (Papandroulakis et al., 2004). Since research and techniques
in larval rearing of marine ornamental fish lag far behind that of
foodfish and freshwater ornamentals, these areas should be the
main focus of future studies (Holt, 2003). Key issues include
system design, physical parameters of the culture environment,
and larval feeds and nutrition (Planas and Cunha, 1999; Ignatius
et al., 2001; Olivotto et al., 2003; Papandroulakis et al., 2004).
4.1. Water Quality and System Design
Like that of broodstock, a general lack of knowledge in the
area of water quality requirements for larval culture of marine
ornamental fish, means that there is a reliance on drawing broad
guidelines from literature where larval culture has been success-
ful (Table 2). As water quality ideals and tolerances will vary
with species and developmental stages, another common and
effective approach is to imitate the environmental conditions
observed in the wild. For instance, compared to many foodfish
and freshwater ornamental species, oligotrophic conditions of-
ten characterize the habitats of marine ornamentals, and, hence,
many marine ornamentals are thought to have a low tolerance to
ammonia, nitrite, and nitrate (Watson and Hill, 2006). However
this is no substitute for properly designed research on a species-
by-species basis. For example, Frakes and Hoff (1982) recorded
a survival of 37% after 21 days’ larval culture of the clown-
fish, Amphiprion ocellaris, when subjecting them to a NO3-N
concentration of approximately 16 mgL−1. However, rate of lar-
val metamorphosis, and subsequent growth and survival, were
reduced when larvae were exposed to a high NO3-N level of
approximately 100 mgL−1(Frakes and Hoff, 1982). Limited
information in this area calls for more research in this field as
quantifying larval water quality tolerances and ideals is impor-
tant for both the establishment, and the refinement of culturing
protocols for marine ornamental fish.
System design for larval rearing is another important area that
is dependent on many other factors, including rearing density
and culture methods (e.g., ‘clear’ or ‘green water’), the level of
contact with the external environment (closed/indoors, semi-
closed/protected or open/exposed), and the water exchange
regimes and rate (static, recirculating or flow through) (Planas
and Cunha, 1999; Papandroulakis et al., 2004). Although there
are a large number of areas to cover with respect to system de-
sign, a collation of studies in which larvae of marine ornamentals
were reared shows the adoption of a key culture system, i.e., the
almost exclusive use of a closed ‘mesocosm’ system (Table 3).
The mesocosm system is seen as an integration of the pos-
itive aspects of live algae addition to the culture water, such
as system stability, better prey quality and light dissipation,
and the positives of an intensive system, i.e., maximum con-
trol over system water parameters and larval stocking den-
sity (Job et al., 1997; Planas and Cunha, 1997; Papandroulakis
et al., 2004). Marine fish larvae have very limited immunologi-
cal capabilities, relying mostly on nonspecific mechanisms and
phagocytosis, to respond to challenges posed by obligate and
opportunistic pathogens (Skjermo and Vadstein, 1999; Olafsen,
2001). Therefore, it is important to note that studies have indi-
cated that live algae greenwater (and/or the microbial ‘cocktail’
it introduces), as a part of mesocosm systems, may antagonize
some bacteria types (such as opportunistic pathogenic bacteria)
through the production of antibiotics, and possibly promote bac-
teria that benefit larvae (Gatesoupe, 1999; Hargreaves, 2006;
Vine et al., 2006). Such beneficial bacteria, known as probi-
otics, may act directly on larvae as an ingested probiont that
enhances intestinal micro-flora; or indirectly as either a pas-
sive bio-control of pathogenic and toxin-releasing bacteria or as
a bio-filtration enhancer (Gatesoupe, 1999; Hargreaves, 2006;
Vine et al., 2006). Unfortunately, a complete understanding of
the effects of microalgae on larviculture has not yet been attained
(van der Meeran et al., 2007), marine ornamental research in this
sense therefore should contribute to knowledge concerning the
benefits of greenwater and the harnessing of these benefits for
the development of efficient rearing systems.
The eyes of first feeding marine fish larvae are an impor-
tant sensory organ for prey identification and capture, as most
marine fish larvae are visual feeders (Naas et al., 1996). There-
fore, lighting conditions (including intensity, spectral quality,
and photoperiod) are an important environmental parameter
that needs to be considered when rearing marine fish larvae
(Naas et al., 1996; Boeuf and Le Bail, 1999). Unfortunately,
few studies record light intensity or manipulation of light spec-
trum used in larval culture of marine ornamental fish. However,
there is evidence to suggest that the effect of these parame-
ters of light may be significant. For instance, it has been re-
ported that on mass-scale rearing of the three-spotted seahorse,
Hippocampus trimaculatus, survival of pelagic stage juveniles
subjected to three different light intensities was the highest
at a light intensity of 2,000 lux (Murugan et al., 2009). Pe˜
na
et al. (2004) concluded that larval feeding incidence of the spot-
ted sand bass, Paralabrax maculatofasciatus, increased with an
increasing light intensity from 0 to 700 lux. Job and Shand
(2001) found that larvae of ochre striped cardinalfish, Apogon
compressus, the ambon damselfish, Pomacentrus amboinensis,
and the spinecheek anemonefish, Premnas biaculeatus, appear
well adapted to light wavelengths (493–507 nm) experienced in
shallow coral reef waters. Also, an extrapolation on the findings
of Job and Bellwood (2007), concerning ultraviolet sensitivity
of coral reef fishes, is that prey perception by larvae may be
improved by manipulating light wavelengths.
With respect to photoperiod, it appears that the light durations
used by most authors in rearing marine ornamental fish larvae are
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Tab l e 2 List of species-specific water quality conditions for marine ornamental fish in which larval rearing has been attempted
Temperature, Ammonia (TAN), Nitrite Nitrate Dissolved
Common Name Species Salinity pH ◦CmgL
−1(NO−
2), mgL−1(NO−
3), mgL−1Oxygen, mgL−1References
Clown anemonefish Amphiprion percula 30–35 8.1–8.3 24.0–27.0 0–0.02 0–0.01 <25 ∼6.5–8.0 Gordon et al., 1998; ¨
Onal et al., 2008
False clown anemonefish Amphiprion ocellaris — — 26.0–27.0 — — <440 — Frakes and Hoff, 1982; Furuta et al., 2005
Black anemonefish Amphiprion melanopus — — 27.0–29.0 — — — — Green and McCormick, 1999
Yellowtail anemonefish Amphiprion clarkii 30 8.0–8.2 27.7–28.5 <0.03 <0.03 — — Olivotto et al., 2008a, 2008b
Sebae anemonefish Amphiprion sebae 28–35 8.1–8.5 26.5–33.4 0 — — 3.6–5.6 Ignatius et al., 2001
Yellow-tailed damselfish Chrysiptera parasema 30 — 28.0 — — — — Olivotto et al., 2003
Hawaiian dascyllus Dascyllus albisella 35 — 25.5–27.5 — — — — Danilowicz and Brown, 1992
Whitetail dascyllus Dascyllus aruanus 35 — 24.0–28.5 — — — — Danilowicz and Brown, 1992
Semicircle angelfish Pomacanthus semicirulatus 33–35 7.9–8.3 26.0–28.2 — — — 5.62–8.09 Leu et al., 2009
Lemonpeel angelfish Centropyge flavissimus 32–36 — 26.0–28.0 — — — — Olivotto et al., 2006a
Blue Mauritius angelfish Centropyge debelius — — 25.0–26.0 <0.25 — — — Baensch and Tamaru, 2009
Forktail blenny Meiacanthus atrodorsalis 34–38 8.0–8.2 26.0–29.0 <0.02 <0.02 <0.5 — Unpublished data from author
Lyretail blenny Meiacanthus reticulatus 34–38 8.0–8.2 26.0–29.0 <0.02 <0.02 <0.5 — Unpublished data from author
Striped blenny Meiacanthus grammistes 30 8.2 28.0 <0.03 <0.03 — — Olivotto et al., 2010
White Tiger goby Priolepis nocturna 32 8.2 30.0 <0.02 <0.02 <20 — Wittenrich et al., 2007
Barber goby Elacatinus figaro 34 8.2 26.0 <0.02 <2.0 — — Shei et al., 2010
328
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 329
Tab l e 3 Summary of culture systems, culture methods, tank background, water flow and photoperiod used for rearing larvae of various marine ornamental fish
(Culture method: GW—greenwater; CW—clearwater. Culture system: C—closed; SC—semi-closed; O—open. Tank background: Bw—black walls, Bb—black
bottom; Gw—grey walls; Gb—grey bottom; Ww—white walls; Wb—White bottom. Water flow: S—static; R—recirculating; FT—flow through)
Culture Culture Tank Water Photoperiod
Common Name Species Method System Background Flow (Light:Dark) References
Clown anemonefish Amphiprion
percula
GW C BwBb S 14h:10h Job et al., 1997
GW C Bw S 16h:8h ¨
Onal et al., 2008
GW C BwBb S 12h:12h Gordon and Hecht, 2002
False clown anemonefish Amphiprion
ocellaris
GWaC BwWb S 14–16h:10–8h Hoff, 1996
GW C Bw R/FT∗∗∗ 24h:0h Avella et al., 2007
Black anemonefish Amphiprion
melanopus
GW C BwBb S 14h:10h Job et al., 1997
GW C BwBb S 14h:10h Green and McCormick, 2001
Sebae anemonefish Amphiprion sebae GW C — S — Ignatius et al., 2001
Yellowtail anemonefish Amphiprion
clarkii
GW C Bw R/FT∗∗∗ 14h:10h Olivotto et al., 2008a
Spinecheek anemonefish Premnas
biaculeatus
GW C BwBb S 14h:10h Job et al., 1997
GW C BwBb S 14h:10h Job and Shand, 2001
GW C BwBb S 14h:10h Job and Bellwood, 2007
Yellow-tailed damselfish Chrysiptera
parasema
GW C Covered (color
unknown)
S 24h:0h Olivotto et al., 2003
Whitetail dascyllus Dascyllus aruanus GW C — FT∗24h:0h∗∗ Danilowicz and Brown, 1992
Hawaiian dascyllus Dascyllus
albisella
GW C — FT∗24h:0h∗∗ Danilowicz and Brown, 1992
Ambon damselfish Pomacentrus
amboinensis
GW C BwBb S 14h:10h Job et al., 1997
GW C BwBb S 14h:10h Job and Shand, 2001
GW C BwBb S 14h:10h Job and Bellwood, 2007
Chinese demoiselle Neopomacentrus
bankieri
GW C BwBb S 14h:10h Job et al., 1997
Semicircle angelfish Pomacanthus
semicirculatus
GW C — S — Leu et al., 2009
Lemonpeel angelfish Centropyge
flavissimus
GW C GwGb S 24h:0h Olivotto et al., 2006a
Blue Mauritius angelfish Centropyge
debelius
GW C BwWb S 16h:8h Baensch and Tamaru, 2009
Yellow-striped
cardinalfish
Apogon
cyanosoma
GW C BwBb S 14h:10h Job et al., 1997
Blue-eyed cardinalfish Apogon
compressus
GW C BwBb S 14h:10h Job et al., 1997
GW C BwBb S 14h:10h Job and Shand, 2001
GW C BwBb S 14h:10h Job and Bellwood, 2007
Sunrise Dottyback Pseudochromis
flavivertex
GW C Bw R or FT∗∗∗ — Olivotto et al., 2006b
Cleaner Goby Gobiosoma
evelynae
GW C Bw R or FT∗∗∗ 24h:0h Olivotto et al., 2005
Five-lined cardinalfish Cheilodipterus
quinquelineatus
GW C Bw S 14h:10h Job et al., 1997
Forktail blenny Meiacanthus
atrodorsalis
GW C BwWb S 24h:0 Unpublished data from
author
Lyretail blenny Meiacanthus
reticulatus
GW C BwWb S 24h:0 Unpublished data from
author
Striped blenny Meiacanthus
grammistes
GW C Bw R or FT∗∗∗ 13h:11h Olivotto et al., 2010
White Tiger goby Priolepis nocturna CW C BwBb S/R (100 mL
min−1)b
14h:10h Wittenrich et al., 2007
Barber goby Elacatinus figaro GW C BwWb S 24h:0h Shei et al., 2010
Manderinfish Synchiropus
splendidus
— C — S 24h:0h Sadovy et al., 2001
∗Water exchange of between 12% and 29% total tank volume per day directly from the ocean.
∗∗Consisted of a 14 h ‘High-light’ period and a 10 h ‘Low-light’ period.
∗∗∗Unclear whether water was recirculated or flow through.
aSpirulina used to ‘green-up’ water.
bStatic replicates placed inside a larger water bath with recirculated, filtered water.
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330 J. A. MOORHEAD AND C. ZENG
either emulating natural summer conditions or are extended up
to 24 h (Table 3). Several studies have investigated the effect of
varied photoperiod on larval fish culture (Barahona-fernandes,
1979; Tandler and Helps, 1985; Duray and Kohno, 1988; Bar-
low et al., 1995; Martin-Robichaud and Peterson, 1998; Down-
ing and Litvak, 1999; Arvedlund et al., 2000). Many concluded
that extended photoperiods longer than 14 h light phase resulted
in increased survival and growth (Barahona-fernandes, 1979;
Tandler and Helps, 1985; Duray and Kohno, 1988; Barlow et al.,
1995; Arvedlund et al., 2000). However, Martin-Robichaud and
Peterson (1998) reported that for striped bass, Morone saxatilis,
a short photoperiod of 8 h light:16 h dark appeared to facilitate
better swim-bladder inflation in early larvae, leading to greater
survival as compared to a 16 h light:8 h dark photoperiod. Mean-
while, results in support of extended photoperiods did appear
to vary between species and sometimes between different de-
velopmental stages of the same species. There also appears to
be some conjecture as to the effectiveness of unnatural constant
light phases (24 h light:0 h dark) used for culture. The find-
ings of Arvedlund et al. (2000) led to the argument that the
anemonefish, A. melanopus, larvae benefited from an extended
photoperiod, but may require a dark period to optimize growth.
Meanwhile, in foodfish, Barlow et al. (1995) concluded that an
extended light period from 16 to 24 h results in better growth of
barramundi, Lates calcarifer, but also noted that these fish still
displayed a circadian feeding rhythm with periods of feeding in-
activity. Better survival and growth was found with a 24 h light
period compared to natural photoperiods for larval rabbitfish,
Siganus guttatus, and gilthead seabream, Sparus aurata (Tan-
dler and Helps, 1985; Duray and Kohno, 1988). Further, to these
findings for foodfish, more recent research on the ornamental
yellow-tailed damselfish, Chrysiptera parasema, noted that a 24
h light period was indispensable for successful rearing of larvae
(Olivotto et al., 2003). Consequently, the constant light period
appears to have been adopted by these authors as a standard pho-
toperiod in recent studies on larvae culture of various marine
ornamental reef fish, including the cleaner goby, Gobiosoma
evelynae, the sunrise dottyback, Pseudochromis flavivertex, and
the lemonpeel angelfish; Centropyge flavissimus (Olivotto et al.,
2003, 2005, 2006a, 2006b). However, caution ought to be exer-
cised with attempts to compare among studies or to formulate a
‘best practice’ of photoperiod for different species, as responses
to photoperiod seem species specific and linked to associated
variables, including light intensity and spectral quality (Boef
and Le Bail, 1999; Downing and Litvak, 1999; Trotter et al.,
2003).
Tank color has also been shown to have an influence on lar-
val survival and growth, and coloration of juvenile stages (Yasir
and Qin, 2009). It is suggested that appropriate backgrounds
may provide the right contrast for better prey visualization and,
hence, improved capture efficiency (Job et al., 1997; Martin-
Robichaud and Peterson, 1998; Green and McCormick, 2001).
For example, Pe˜
na et al. (2005) reported that although feeding
incidence was not affected by tank color in Paralabrax macu-
latofasciatus, feeding success (number of prey in the digestive
tract) was improved by a darker tank. Furthermore, many fish
larvae display phototaxis, which dictates their response to light,
including reflected light off tank walls (Naas et al., 1996). Photo-
taxis responses have been reported to be linked to mortality and
jaw malformation of larvae of striped trumpeter, Latris lineata,
as a result of ‘walling’ behavior (Cobcroft et al., 2001; Cobcroft
and Battaglene, 2009). For culture of marine ornamental larvae,
standard practice appears to be that of using black-walled tanks
(Table 3). This appears to follow the conclusion that black-
walled tanks best emulate natural conditions experienced by
pelagic larvae in the wild, and tend to prevent or reduce the
negative affects of ‘walling’ (Naas et al., 1996; Cobcroft et al.,
2001; Green and McCormick, 2001; Cobcroft and Battaglene,
2009). However, the effect of tank color appears to vary depend-
ing on the species being investigated and the light intensity used.
For example, Downing and Litvak (1999) concluded that black-
walled tanks impaired growth of larval haddock, Melanogram-
mus aeglefinus, at low light intensity, as compared to a white tank
background. Similarly, Woods (2000b) reported better attack
rate and prey capture success of the planktonic stage juveniles
of pot-bellied seahorses, Hippocampus abdominalis, in clear
jars compared to black and white background jars. However, in
an attempt to build on the findings of Woods (2000b), Martinez-
Cardenas and Purser (2007) used a different photoperiod, light
intensity, and tank volume for their experiments, and found no
significant difference in growth or survival of the planktonic
stage, Hippocampus abdominalis, when culturing them in clear,
white, yellow, red, blue, and black background tanks. This sug-
gests that caution again needs to be exercised when attempting
to formulate a ‘best practice’ for tank color, as this is likely to
be species specific and stage specific, and involve consideration
of light intensity, spectrum, photoperiod, tank shape and size,
and the interactions between these parameters.
Clearly, the interactions between light characteristics, tank
color and design, and water turbidity (such as the use of green-
water) are all likely to influence the performance of larvae and
make it a very complex issue. However, as a ‘rule of thumb’,
lighting conditions that maximize contrast between prey and the
surrounding environment, promoting regular feeding incidence
and feeding success, as well as limiting the occurrence of light
trapping and walling phenomena, are the underlying goals of
light manipulation in larval rearing of marine fish (Downing
and Litvak, 1999).
4.2. Larval Diet
Understandably, larval diets are critical to the success of lar-
val culture of fish species and there are many factors in play
that determine whether a diet is suitable or not for a particular
stage of larvae of a target species. These factors include both
physical characteristics, such as size, shape, buoyancy, mobility,
and coloration of diets as well as chemical features, including
inclusion levels of various nutrients, enzymes, and attractants.
During larval ontogeny, it is well recognized that high mortality
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occurring at the onset of exogenous feeding of larvae represents
a stumbling block for the development of both marine orna-
mental and foodfish aquaculture (Watanabe and Kiron, 1994;
Rønnestad et al., 1999; Holt, 2003; Y´
ufera and Darias, 2007).
Multiple papers and literature reviews have addressed this issue,
with feed type, size, density, and provision of correct nutrition
being the major focuses (Y´
ufera and Darias, 2007), of which
will be discussed in the following section.
4.2.1. Feed Types, Sizes, and Density
Gut analyses of wild caught marine fish larvae have revealed
a wide range of ingested prey, including various life stages
of copepods, protozoans, dinofagellates, and larvae of vari-
ous marine organisms (Watanabe and Kiron, 1994; Holt 2003;
McKinnon et al., 2003). In contrast, captive larval rearing of
marine fish has relied mainly on two traditional live prey; the
rotifer, Brachionus spp. and the nauplii of brine shrimp, Artemia
spp. (Watanabe and Kiron, 1994; Battaglene and Fielder, 1997;
Sargent et al., 1997; Wullur et al., 2009). The major advantage
of rotifers and Artemia is the relative ease of mass produc-
tion; however, as they are not natural prey for many marine
fish larvae, the larval culture of some species, particularly reef
fish, has been hindered by the inadequacy of these two live foods
(Rodriguez and Hirayama, 1997; Schipp et al., 1999; Holt, 2003;
McKinnon et al., 2003; Olivotto et al., 2008a, 2008b). Difficulty
in raising tropical reef fish larvae, of both foodfish and ornamen-
tal species, commonly arises due to their typically small size,
small mouth gape, and simple digestive system at the time of
first feeding (Rønnestad et al., 1999; Holt, 2003; McKinnon et
al., 2003). For example, in larvae of the tropical groupers and
snappers, prized foodfish species, small mouth gape dictates
the size requirement of first foods to be less than 100 µm. De-
spite the development of super small strain rotifers (ss-rotifers)
that can be ingested by the larvae, survival is still poor if these
rotifers are used alone (Schipp et al., 1999; McKinnon et al.,
2003).
With respect to marine ornamentals, similar problems exist in
larval rearing (Ostrowski and Laidley, 2001). To date, among the
few marine ornamentals that have been successfully cultured, a
majority of them are characterized either by the possession of a
large yolk sac, relatively large larvae and mouth gape sizes, or
advanced development at hatching or first feeding (Leis, 1991;
Fisher et al., 2000). However, many marine ornamental fish
larvae are very small at hatching and first feeding (<3 mm), and,
therefore, demand foods that are smaller than even the smallest
strain of Brachionus rotifers (Holt, 2003). Furthermore, the low
ingestion rates of rotifers, the traditional live prey used for first
feeding of fish larvae, is also evident in some marine ornamental
larvae, with cases of refused ingestion despite its acceptable size
being reported (Young, 1994; Ostrowski and Laidley, 2001).
There are several examples of successful use of alternatives
to Artemia and rotifers, such as various copepod species, wild
zooplankton, ciliates, and dinoflagelates, in research scale rear-
ing of early larvae. However, at present, in one way or another,
potential replacements or supplements to rotifers for early lar-
vae have repeatedly fallen short of prerequisites that make them
feasible for mass production and industrial use, and a similar
situation exists for the development of inert microdiets to to-
tally replace Artemia nauplii for older larvae (Rodriguez and
Hirayama, 1997; Stoettrup and Norsker, 1997; Holt, 2003).
This unavoidably affects the development of marine ornamental
aquaculture. Among various options, copepods appear to have
better potential to supplement or replace rotifers and Artemia
for marine ornamental larval culture, given the fact that ma-
rine ornamentals generally produce far fewer larvae compared
to most foodfish species, and given recent development and
refinement of mass culture techniques, particularly for tropical
copepod species (Schipp et al., 1999; Marcus and Murray, 2001;
McKinnon et al., 2003; Milione and Zeng, 2007, 2008; Camus
and Zeng, 2008, 2009; Camus et al., 2009). The use of copepods
has been shown to be effective in addressing problems of size,
digestibility, attractiveness, and nutritional value in the culture
of various foodfish (van der Meeren, 1991; McEvoy et al., 1998;
Schipp et al., 1999; Rajkumar and Kumaraguru vasagam, 2006)
as well as some marine ornamental species (Ignatius et al., 2001;
Olivotto et al., 2006a, 2008a, 2008b, 2010; Baensch and Tamaru,
2009). Additionally, marine ciliates, even smaller in size, have
been suggested as a potential live prey that may bridge the gap
between first feeding and the acceptance of rotifers or copepod
nauplii (Nagano et al., 2000a, 2000b; Olivotto, 2005). Labo-
ratory trials on very small marine ornamental larvae, such as
the palette surgeonfish, Paracanthurus hepatus, and the cleaner
goby, Gobiosoma evelynae, as well as the seven-band grouper,
Epinephelus septemfasciatus, have shown some promise in this
respect (Nagano et al., 2000a, 2000b; Olivotto, 2005). However,
the low nutritional value of ciliates in some of these trials was
apparent (Nagano et al., 2000a, 2000b), and research on their
culture techniques to ensure a consistent supply is also lack-
ing. Therefore, more work on mass production techniques of
ciliates, and investigation into the potential to manipulate their
nutritional value to satisfy the needs of first feeding larvae are
required before they become a viable option.
Another interesting development is recent investigation into
alternative rotifer species to the genus Brachionus. Chigbu and
Suchar (2006) reported the successful isolation and mass cul-
ture of the marine rotifer Colurella dicentra, which displayed
an average lorica length of 93 µm and a width of 49 µm, while
Wullur et al. (2009) documented the culture of the rotifer Pro-
ales similis, which displayed an average body length and width
of 83 µm and 40 µm, respectively. These rotifers are substan-
tially smaller than Brachionus rotundiformis, the super small
Brachionus strain commonly used for feeding tropical fish lar-
vae, which have an average lorica length and width of 134 µm
and 102 µm, respectively (Wuller et al., 2009). With the use of
similar culture methods, these rotifers can be cultured to achieve
comparable densities to that of Brachionus rotifers. However,
although Wuller et al. (2009) found a higher feeding incidence
of seven-band grouper, Epinephelus septemfasciatus,larvaeon
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332 J. A. MOORHEAD AND C. ZENG
Paoles similus as compared to Brachionus rotundiformis, there
was no clear evidence that larval survival was improved. Hence,
there are doubts as to whether the use of these alternative rotifer
species would increase the likelihood of successfully culturing
small-mouthed fish larvae. Further investigation is obviously re-
quired to verify the potential of these alternative rotifer species
for the culture of small-mouthed fish larvae.
In the past decades, inert formulated microdiets have at-
tracted substantial interest because of their advantages in easy
and precise manipulation of nutrient components, low produc-
tion cost, and off-the-shelf convenience. While they represent
the ultimate goal in larval culture (Y´
ufera et al., 1999; Langdon
et al., 2007), unfortunately, the current state of development
sees them largely unstable in water, displaying poor buoyancy
and residence time in the water column, and often unattractive to
fish larvae, as well as unable to contribute to or promote enzyme
activity in larval guts (Gordon et al., 1998; Chong et al., 2002;
Kanazawa, 2003; Langdon et al., 2007). As a consequence, over-
all, formulated diets have shown limited success in completely
replacing live feeds in the culture of early larvae of marine fish
species, including ornamentals (Gordon et al., 1998; Y´
ufera
et al., 1999; Cahu and Zambonino Infante, 2001; Kanazawa,
2003). For example, Gordon et al. (1998), and Gordon and
Hecht (2002) concluded that even for larvae of the anemone-
fish, A. percula, a relatively hardy species for larvae culture,
the earliest time for weaning onto a formulated diet with-
out significantly affecting survival was 7–9 days after hatch.
Nevertheless, with the continuous development and improve-
ment of microdiets, they may increasingly become a more
amenable alternative to live feeds, and stand to incur signifi-
cant cost savings to hatcheries by phasing out costly and labor-
intensive live feed production in the future (L ´
opez-Alvarado
et al., 1994; Cahu and Zambonino Infante, 2001; Langdon et al.,
2007).
Although not necessarily a bottleneck to the larval culture of
marine ornamental fish, prey density could be a consideration
in larval rearing of certain species, and can have a significant
bearing on feeding success, water quality, and ultimately, lar-
val survival and growth (Houde, 1975, 1978; Duray et al., 1996;
Puvanendran and Brown, 1999). Prey density optima and thresh-
olds appear to be both species and stage specific (MacKenzie
and Kiørboe, 1995; Puvanendran and Brown, 1999; Laurel et al.,
2001) and depend on factors, such as larval density (Houde,
1975, 1977) and mobility, foraging strategies employed by lar-
vae (MacKenzie and Kiørboe, 1995; Laurel et al., 2001), light
and turbidity levels (Grecay and Targett, 1996; Pe˜
na et al., 2005),
and water movement in the rearing system (Sundby and Fossum,
1990; MacKenzie and Kiørboe, 1995). In marine ornamental fish
larvae culture, while prey densities applied are often mentioned,
they appear to vary depending on the author, and variables that
may interact to affect their efficacy are rarely taken into account.
Hence, caution should be taken in committing to a certain prey
density and assuming it is a ‘cover-all’ for all species, stages,
and systems.
4.2.2. Larval Nutrition
The early life stages of marine fish are often characterized
by a priority for rapid growth, physiological development, and
high rates of food intake and metabolism; however, often for
early larvae, all of these have to be achieved with reliance
on a set of rudimentary digestive organs with low capacities
to process and assimilate nutritional resources (Segner et al.,
1993; Watanabe and Kiron, 1994; Izquierdo et al., 2000; Cahu
and Zambonino Infante, 2001; Kim et al., 2001; Y´
ufera et al.,
2003; Rønnestad et al., 2003; Morais et al., 2004; Arag˜
ao et al.,
2007; Rønnestad et al., 2007). Therefore, upon diets being in-
gested by fish larvae, whether a diet can be digested and provide
sufficient nutrition for sustaining larval development arise as
a critical issue (Kanazawa, 2003; Kolkovski et al., 2004). A
thorough understanding of the digestive function and process-
ing capacity of larvae, from the onset of exogenous feeding,
will form the basis of successful diet formulations, particu-
larly with respect to reducing reliance on live foods (Rønnestad
et al., 2007). Many marine larvae have a rudimentary, relatively
undifferentiated gut, and possess a limited ability to efficiently
process complex food materials in both natural and formulated
diets (Kim et al., 2001; Rønnestad et al., 2007). It appears that
patterns of enzyme production and activity vary during fish on-
togeny, are species-specific, and have been reported to be under
pre-programmed genetic control, and influenced by diet quality
and quantity (P´
eres et al., 1998; Cahu and Zambonino Infante,
2001; Kim et al., 2001; Arag˜
ao et al., 2007). Larval enzymatic
capacity is thus an important factor that dictates the nutritional
suitability of a particular diet for marine larvae (Izquierdo et al.,
2000; Cahu and Zambonino Infante, 2001; Kim et al., 2001) and
requires species-specific and stage-specific research. In addition
to larval enzymatic capacity, among a host of topics associated
with the nutritional make-up of larval diets, there are several ma-
jor nutrient components that have attracted particular attention
and are likely to be important for marine ornamental fish, in-
cluding lipids (especially essential fatty acids), protein, vitamins
and minerals, as well as carotenoids.
4.4.2.1. Lipids. Lipids form an important part of marine fish
eggs, are the sources of the essential fatty acids (EFA), and to-
gether with free amino acids (FAA), form the most important
energy sources for developing embryos and larvae (Rainuzzo et
al., 1997; Izquierdo et al., 2000; Cahu and Zambonino Infante,
2001; Kanazawa, 2003; Tocher, 2003). It has been suggested
that the lipid composition found in marine fish eggs, should ap-
proximately indicate the lipid requirement of marine fish larvae
(Sargent et al., 1999). Of various lipids in the diets of marine
larvae, phospholipids (PL) and highly unsaturated fatty acids
(HUFA), are two lipid components that appear to have received
the highest attention in past research (Cahu and Zambonino
Infante, 2001; Kanazawa, 2003; Tocher, 2003; Glencross,
2009).
The essentiality of PL in the diet of marine fish larvae is well
established and they have been suggested as a superior source
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DEVELOPMENT OF CAPTIVE BREEDING TECHNIQUES FOR MARINE ORNAMENTAL FISH: A REVIEW 333
of HUFA as compared to neutral lipids (Izquierdo et al., 2000;
Cahu and Zambonino Infante, 2001; Kanazawa, 2003; Tocher,
2003; Tocher et al., 2008; Cahu et al., 2009). Although the
roles that various PL fractions play is not yet clear, dietary PL
broadly improves digestive functions and skeletal development
of fish larvae, enhancing their development, growth and survival
(Cahu and Zambonino Infante, 2001; Tocher et al., 2008; Cahu
et al., 2009). Both the dietary source and classes of PL, and
the criteria by which their effects are measured, play a role in
determining the optimal PL level in marine fish larvae (Cahu
et al., 2009). Clearly, much work is still to be done to elucidate
the specific functions of various fractions and classes of PL,
and the optimal dietary requirement for the larvae of various
fish species, particularly in regards to marine ornamental fish
species.
Aside from PL, a large body of research has focused on
the nutritional values of essential fatty acids. This research
has shown clear evidence of the importance of HUFAs for
larval marine fish culture (Sargent et al., 1997; Planas and
Cunha, 1999; Sargent et al., 1999; Kanazawa, 2003; McKin-
non et al., 2003; Tocher, 2003; Glencross, 2009). Among HU-
FAs, of particular value to fish larvae are docosahexaenoic acid
(DHA; 22:6n-3) and eicosapentaenoic acid (EPA; 20:5n-3) of n-
3 HUFA, and arachidonic acid (ARA; 20:4n-6) of the n-6 HUFA
(Rainuzzo et al., 1997; Planas and Cunha, 1999; Sargent et al.,
1999; McKinnon et al., 2003). For the culture of most larval
fish, effective delivery of HUFA is achieved via enrichment
of rotifers or Artemia using emulsified fish oils or HUFA rich
microalgae (Watanabe, 1993; Ozkizilcik and Chu, 1994; South-
gate and Lou, 1995; Barclay and Zeller, 1996; Southgate, 1996;
Brown et al., 1997; Harel et al., 2002). Like in many foodfish
species, the importance of essential fatty acids to marine orna-
mental larvae has been clearly demonstrated, achieving better
survival and growth (Olivotto et al., 2003, 2005, 2006a, 2006b,
2008a; Avella et al., 2007), as well as decreasing anomalies in
the development of the central nervous system (Avella et al.,
2007). Furthermore, of additional interest to the marine orna-
mental industry are findings that suggest levels of DHA, EPA,
and ARA in larval diets could effect their pigmentation, which is
of critical importance in determining the sale value of many or-
namental species (Rainuzzo et al., 1997; Copeman et al., 2002;
Bell et al., 2003). For example, Avella et al. (2007) highlighted
a clear correlation between fatty acid enrichment and reduced
incidence of miss-bands in the cultured clownfish, Amphiprion
ocellaris, although the underlying mechanisms for such a cor-
relation need further investigation.
4.4.2.2. Protein. As the source of amino acids for tissue
growth and energy, protein is another critical nutrient to the
larvae of marine fish (Watanabe and Kiron, 1994). However,
unlike in juveniles and adult fish, it has been suggested that in
the absence of HCl- and pepsin-secretion in first feeding larvae,
no preparatory acid denaturation of ingested proteins and low
proteolytic activity are common for early larvae (Rønnestad et
al., 1999; Helland et al., 2003; Rønnestad et al., 2003; Arag˜
ao
et al., 2007). This renders alkaline enzymatic attack of complex
proteins difficult or inefficient for the larvae, limiting their access
to the total amino acid pool provided in their diet (Rønnestad
et al., 2003). Therefore, the supply of hydrolyzed protein and/or
essential amino acids, particularly free amino acids (FAA), to
diets of marine larvae appears to have greater importance than
complex proteins (Rønnestad et al., 1999; Cahu and Zambonino
Infante, 2001; Arag˜
ao et al., 2007).
Many amino acids (AA) are important as metabolic fuel, form
the basis of body protein synthesis and are chemo-attractants to
feeding fish larvae (Kolkovski et al., 1997; Rønnestad et al.,
1999, 2003; Cahu and Zambonino Infante, 2001; Brown et al.,
2005; Arag˜
ao et al., 2007). Although many marine fish larvae
lack the ability for acid hydrolysis of complex proteins, and
appear to have a slow rate of nutrient absorption, it has been
reported that they have a high capacity to digest protein hy-
drolyzate, and a high retention efficiency of amino acids, of
which FAA can be readily absorbed without digestion (Cahu
and Zambonino Infante, 2001; Morais et al., 2004; Rønnestad
et al., 2007). This may explain why marine larvae tend to have
a high preference for live preys, which are known to be high
in FAA, and display better survival when fed them, as opposed
to formulated diets (Helland et al., 2003; Morais et al., 2004;
Rønnestad et al., 2007). Considering that AA are the building
blocks for proteins and an important energy source, they need
to be provided in the appropriate levels and ratios to maximize
protein synthesis while also providing adequate metabolic fuel
(Arag˜
ao et al., 2007). Therefore, there is a need to develop
methods to manipulate the AA profile in both live and formu-
lated larval diets to satisfy the daily requirements of marine fish
larvae.
4.4.2.2. Vitamins, Minerals, and Carotenoids. The effects of
vitamins on fish larvae, and the way they are metabolized and
stored, can relate to their solubility characteristics, i.e., water-
soluble or fat-soluble, and their concentration in the diet and
culture media (Furuita et al., 2009). The functions of vitamins
are very diverse and some of them must be provided within op-
timum ranges to avoid pathologies related to deficiency or hy-
pervitaminosis (Dedi et al., 1995; Furuita et al., 2009). Despite
the importance of maintaining vitamins within narrow optima
for fish, the vitamin requirements of most larval fish are largely
unknown (Kanazawa, 2003; Furuita et al., 2009). Vitamin re-
quirements that have been investigated for fish larvae include Vi-
tamin C, Vitamin A, Vitamin E, and riboflavin (Dedi et al., 1995;
Merchie et al., 1996, 1997; Gapasin et al., 1998; Cahu et al.,
2003; Kanazawa, 2003; Glencross, 2006; Souto et al., 2008;
Furuita et al., 2009). Among various vitamins, ascorbic acid
(Vitamin C) and retinoic acid (related to Vitamin A) have been
shown to affect larval development, survival, and pigmentation
in larvae of marine fish species, such as the Japanese flounder,
Paralichtys olivaceus, the milkfish, Chanos chanos, the barra-
mundi, Lates calcarifer, the Japanese eel, Anguilla japonica, and
the turbot, Scophthalmus maximus (Dedi et al., 1995; Merchie
et al., 1996, 1997; Gapasin et al., 1998; Cahu et al., 2003;
Kanazawa, 2003; Glencross, 2006; Furuita et al., 2009). Sim-
ilarly, riboflavin was reported to improve growth and survival
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334 J. A. MOORHEAD AND C. ZENG
of white sea bream, Diplodus sargus (Souto et al., 2008), while
Vitamin E (largely in the form of α-tocopherol) is also consid-
ered important as it can act as an antioxidant, protecting HUFA
from oxidative damage (St´
ephan et al., 1995; Furuita et al.,
2009).
As inorganic compounds, various minerals are required by
all animals, and like vitamins, often need to be maintained
within strict ranges for normal cellular metabolic activities
(Watanabe et al., 1997). Minerals serve many biological and
physiological functions, including contributions to skeletal for-
mation, maintenance of colloidal systems, regulation of acid-
base equilibrium, and form important components of hormones
and enzymes (Watanabe et al., 1997; Bury et al., 2005; Ngyuyen
et al., 2008). Depending on the duration and degree of depri-
vation, mineral deficiencies can cause biochemical, structural,
and functional abnormalities (Watanabe et al., 1997; Ngyuyen
et al., 2008; Matsumoto et al., 2009). The specific requirements
of fish for minerals have been sparsely researched, particularly
as most minerals are either minor or trace nutrients in terms of
dietary requirements (Watanabe et al., 1997; Bury et al., 2005).
Multiple factors, including diet characteristics, mineral chemi-
cal state, and the waterborne mineral concentration, could affect
the bioavailability and level of absorption of a particular min-
eral to fish larvae (Watanabe et al., 1997; Bury et al., 2005). For
marine fish larvae, dietary minerals that are considered highly
important include zinc, manganese, iron, cobalt, selenium, and
iodine (Watanabe et al. 1997). For example, both zinc and man-
ganese are considered essential nutrients for normal growth and
skeletal development in larvae of the red seabream, Pagrus ma-
jor, and have been found to further contribute to stress tolerance
in larvae (Ngyuyen et al., 2008). Meanwhile, Olivotto et al.
(2003, 2006a) added potassium iodide (KI) to the culture water
of larvae of Centropyge flavissimus and Chrysiptera parasema
to promote metamorphosis and settlement.
Finally, carotenoids have recently received attention in the
growout of freshwater ornamental fish, as they are recognized
as important for skin coloration of fish (Guoveia et al., 2003;
Garci´
a-Ulloa and G´
omez-Romero, 2005; Wallat et al., 2005;
Sinha and Asimi, 2007). They are likely to be significant for
growout stages of marine ornamental fish as coloration is clearly
an important value criteria for marine ornamentals. Carotenoids
can also play important roles as pro-vitamin A, antioxidants,
immunoregulators, and are suggested to have a function in dis-
ease resistance (Dom`
ınguez et al., 2005). Carotenoids have been
reported to be a beneficial nutrient to marine fish larvae. For ex-
ample, dietary supplementations of carotenoids, such as astax-
anthin and β-carotene, have been reported to benefit larvae of
salmonids, the Japanese and the spotted parrotfish, Oplegnathus
fasciatus and Oplegnathus punctatus, by promoting better sur-
vival, growth, and increased resistance to bacterial and fun-
gal diseases (Johnson and An, 1991; Christiansen et al., 1995;
Tachibana et al., 1997; Dom`
ınguez et al., 2005). Furthermore,
astanxanthin has also been incorporated into formulated diets to
improve visibility of the food particles to the fish larvae (Cahu
and Zambonino Infante, 2001).
Though generally only small quantities are needed, vitamins,
minerals, and carotenoids form important dietary ingredients for
marine fish larvae. However, further research is clearly needed
to determine the dietary requirements of these nutrients as well
as to elucidate the underlining mechanisms of their beneficial
effects for the larvae of marine ornamental fish.
5. CONCLUSION
The marine ornamental aquaculture industry is in its infancy
with limited specific research and multiple restricting bottle-
necks. Tapping into the growing knowledge base that exists for
foodfish has allowed the rudimentary development of the indus-
try. However, there is urgent need to addresskey bottlenecks and
challenges that cripple its progress. This review has surveyed
the available knowledge base, and attempted to identify areas in
which future research can be directed to refine captive culture
techniques for marine ornamental fish. Areas identified to be of
particular importance are:
•The optimization of broodstock diet and in particular nutrition
in order to maximize reproductive performance
•Investigation into non-invasive hormone delivery techniques
that cater for the smaller size of ‘challenging’ marine orna-
mental broodstock species that fail to spawn naturally under
captive conditions
•Optimization of system design and physical parameters for
larvae, with a view to maximize feeding response, health,
survival, and growth of the larvae, and decrease production
costs
•Exploitation of alternative, nutritive live prey and develop-
ing their mass culture methods for feeding larvae with small
mouth gapes at first feeding
•A better understanding of larval nutritional requirements and
on that basis, development of nutritionally balanced formu-
lated feeds that are attractive to larvae to optimize growth,
survival, and produce healthy pigmentation for marine orna-
mental fish
As interest in marine ornamental aquaculture grows, research
findings in the above areas stand to contribute significantly to
the development of an economically viable marine ornamental
aquaculture industry, and to the overall knowledge base of fish
aquaculture, general biology as well as coral reef conservation.
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