Methods of farming sexually propagated corals andoutplanting for coral reef rehabilitation; with listof references for coral reef rehabilitation
through active restoration measure

Makoto Omori and Kenji Iwao*
Akajima Marine Science Laboratory* Correspondence: M. Omori makomori@amsl.or.jp

K. Iwao iwao@amsl.or.jp
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Table of Contents
Ⅰ. English edition
1. Introduction
2. Sampling of gametes (egg and sperm)
2–1. Prediction of spawning
2–2. Sampling of bundles immediately after spawning
2–2a. Ex situ bundle sampling
2–2b. In situ bundle sampling
3. Maintenance of fertilized eggs
3–1. Fertilization
3–2. Determination of the fertilization rate
4. Sampling of slicks after synchronous spawning
Column 1. Spawning induction
5. Rearing of coral larvae
5–1. Rearing larvae in aquaria on land
Column 2. Rearing embryos and larvae using a flow-through trapping tank
5–2. Collecting brooding coral planulae
5–3. Mass production of larvae using a floating pond
6. Transportation of larvae
7. Induction of settlement and metamorphosis
Column 3. Change in settlement capability of larvae
Column 4. Larval settlement and metamorphosis
8. Substrates
8–1. Quality and structure of an artificial substrate
Column 5. Structure of the artificial substrate
8–2. Larval settlement on the substrate
Column 6. Introduction of larvae to substrate in the sea
Column 7. Acquisition of zooxanthellae
9. Culture of juvenile corals
9–1. Commencement of rearing juvenile corals
9–2. Maintenance during rearing
10. Outplanting
Column 8. Transplantation, outplanting, and translocation
Column 9. Protecting outplanted juvinile corals from predation and nibbling by fish
11. Growth and spawning of outplanted coral colonies
Column 10. Cross fertilization rate and larval settlement rate of the outplanted corals
12. Cost estimate

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Acknowledgments

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References

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Ⅱ.

Japanese edition

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List of references for coral reef rehabilitation through active restoration measure

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Afterward

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Ⅰ．English edition

1. Introduction
Restoration of coral reefs through methods
such as culture and transplanting of
reef-building corals (hereafter they refer corals) are still in the developmental stage. In
contrast to land-based silviculture practices,
the lack of state-of-the-art restoration protocols for reef rehabilitation impedes the development of technical and ecological disciplines.
The lessons learned from our efforts to refine
the coral farming technique at Akajima Marine Science Laboratory (AMSL) have been
described in the study by Hatta et al. (2004)
and Guest et al. (2010). However, these manuals do not include culturing juvenile corals
and outplanting techniques (see Column 8).
More than 100 Acropora colonies reared fromeggs and outplanted at AMSL grew to ＞15–
30 cm in diameter after 4–5 years and they are
spawning every year, indicating that this culturing technique could be used to assist incoral reef rehabilitation. The present overview
illustrates the entire practice of reef rehabilita-tion using sexually propagated corals based on
our studies (Fig. 1).
Corals have two different reproductive strategies that are important to their cultivationsuch as broadcast spawning and brooding.
Corals may also be either hermaphroditic or
gonochoristic. Broadcasting species releasegametes (eggs and sperm) into the water column for external fertilization followed by larval development, whereas brooding species
fertilize within the polyp and fully formed larvae (called planulae) are released during
spawning. Broadcast spawning can be highly
synchronized within species. Broadcasters
usually spawn only once each year, whereas
many brooders can release planulae duringconsecutive months. The majority of coral
studied for cultivation to date are hermaphroditic broadcast spawners including Acroporidae and Faviidae species, which occupy approximately 63% of all coral species. The
remaining species are gonochoric broadcasters such as Poritidae and Fungiidae (approxi-

Fig. 1. Flowchart of the techniques for reef restoration with outplanting of sexually propagated corals.
Numbers on the shoulder of box mean article in the text; shadow areas indicate ex situ operations

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mately 22%), and hermachroditic or gonochoric brooders such as Pocilloporiidae(approximately 15% in total) (Guest et al.
2010).
Corals of the genus Acropora are highly diverse and abundant in the western Pacific
Ocean, including Okinawa. They are broadcast spawners, and as their growth rate ishigher than other scleractinian taxa, they are
considered to be the most important genus forreef rehabilitation. Many species of Acropora
participate in synchronous mass spawningand release millions of eggs. However, the initial loss of embryos and larvae is very highand many die before and after settlement. Synchronous spawning allows for the collection of
gametes in the sea or in a container on land,fertilized, and the larvae used for coral farming. Alternatively, the embryos and larvae may
be collected while they are floating on the sea
surface. The larvae settle on substrate andmetamorphose to juvenile corals in due course.
The methods for farming and outplanting in
the present study mainly consider acroporid
species. When the method concerns otherspecies, each of the species names are mentioned individually.
Three approaches are currently available tofarm corals for active reef rehabilitation. The
first approach is to enhance coral recruitment
on artificial substrates by stocking (reseeding)planula larvae and hoping that the corals willgrow well in the sea. Bundle1) collection→fertilization (or sampling of slicks2))→rearing ofplanula larvae→stocking (reseeding) the larvae on substrates in the sea using mesh enclosure→growth of corals.
The second is that natural settlement of coral larvae from the sea is simply encouraged
using artificial substrates on coral reefs.
The third approach is to enhance mass culture of juvenile corals in a nursery followed byoutplanting the juveniles on degraded reefs.
Bundle collection→fertilization in aquaria onland (or sampling of slicks)→rearing of planula larvae→induction of larval settlement onto
1) Bundle: Eggs and sperm of hermaphroditic species areusually bundied together as buoyant packages called
bundles at the time of release from the polyp.
2) Slick: Aggregates of eggs and embryos floating on the sea
surface.
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artificial substrate→farming juvenile colony→outplanting
The former two approaches are easy and do
not require a long-term effort to maintain andfarm the corals. However, loss of settled larvae
is considerable, as the majority of them usually die because of natural causes in the sea. At
present, we consider that these approaches arestill not favored as a reef rehabilitation technique because of the lack of evidence for apositive long-term effect, although various artificial substrata with specially designed structures are being devised to reduce mortality.
2. Sampling of gametes
2–1. Prediction of spawning
Simultaneous mass spawning of coral, mainly Acropora, can be seen around the time of
the full moon in May near the Yaeyama Islands of the Ryukyu Archipelago and in Juneat the Kerama Islands, including Akajima Island, and Okinawa Island (Hayashibara et al.
1993, Heyward et al. 1987) (Fig. 2). Spawning
may be delayed by 1 month or split into 2
months depending on the lunar cycle, watertemperature, and sexual maturity.
Reference to past spawning records at eachlocation is recommended to predict the exact
timing of spawning. Observation of the colonies during the spawning period from 3 days
before to 7 days after the full moon is recommended. The eggs in coral polyp change fromwhite or cream to a pastel color, often pink ororange, as the spawning date approaches (Fig.
3). Thus, examining gonad maturity by break-Fig. 2. Pattern of coral spawning at Akajima Island (Numberof coral species spawned during each spring, middle,
and neap tide periods) (after Hayashibara et al. 1993)

Fig. 3. Coral gonads immediately before spawningthey are released (Fig. 4). Spawning of somecorals such as Faviidae may be difficult to predict because appearance of the bundle is notdefinite. Fish around the coral may swim excitedly around spawning time and this behavior may be an additional indication of spawning. Spawning times vary depending on thehours of sunlight. Some species including
Acropora tenuis spawn immediately after sunset (19:30–20:30 h), and many other species
spawn 2:30–3:30 h after sunset (peak from
21 : 30–22 : 30 h) at Akajima (Fig. 5, see Fukami
et al. 2003). Although density of their distribution is not large, a few species may spawn
between these time zones, while the others
may spawn in August or around the occurrence of the new moon.
2–2. Sampling of bundles immediately after
spawning
Each acroporid polyp releases bundles. The
bundles rise to the sea surface because of thebuoyancy of the eggs. They subsequently dissociate and meet the gametes released fromother colonies for fertilization.

Fig. 4. Bundles appearing at the mouth of the coral

Fig. 5. Coral spawning

ing a branch from a colony is a useful indicator of the upcoming spawn. As the density ofcoral colonies has decreased recently, spawning may be delayed or split into 2 days depending on the location and even at an island.
On the night of spawning gamete bundles appear in the mouth of the polyps 1–1.5 h before

2–2a. Ex situ sampling of bundles
The bundles may be collected ex situ after
obtaining fragments (15–20 cm in length andwidth) from donor colonies using a hammer
and chisel. These fragments are maintained inshallow seawater near a land-based aquaria fora few days before spawning. The fragments of
Acropora are brought to the aquaria 1−2 h before the spawning is predicted, and each isolated fragment is maintained in a container(20 L). For corymbose corals such as Acropora
tenuis, 3 colonies of approximately 30 cm indiameter, allow the collection of approximately 1×106 eggs. The fragments used for sampling the bundles should be transplanted backto the home sites within 2 weeks after spawning to minimize the stress exerted on the coral.
2–2b. In situ bundle sampling
Bundles can be easily collected by setting up
a funnel-shaped device (bundle collector) underwater over the coral colony (Fig. 6). A rigid
cone with stainless steel wire is used to construct the bundle collector by wrapping the5

Fig. 6. Bundle collector (Photo courtesy of Dr. M. Hatta)

conical form with fine nylon mesh (≤300 μm).
A transparent plastic screw cap bottle (500 mL
to 1 L) is attached at the tip of the cone foreasy attachment and detachment and a polystyrene foam float is used to keep the plastic
bottle buoyant.
A few donor colonies are selected beforehand and are examined daily to predictspawning. When the bundles emerge, thebundle collector is placed over the colony andsecured using string and nails. The bundle collector is allowed to set for a few hours before
the expected spawning time. It is advisable toprepare whole procedure beforehand so thatthe work on the spawning night is minimized.
If spawning does not occur, the bundle collector is removed shortly to avoid damage to thecoral and prevent loss of the device.
Spawning may be completed within approximately 30 min. Because the bundles are concentrated in the cap bottle, it is required to detach the bottle immediately after spawning,cover the cap in water, and bring it back to theaquaria for fertilization. The eggs may die
from the lack of oxygen if they are left in thebottle at a high density for a long time.
3. Maintenance of fertilized eggs
3–1. Fertilization
Because corals such as Acropora have physiological barriers to prevent the crossing of
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gametes from the same parent, multiple colonies are required for successful fertilization.
Cross-fertilization with many colonies enhances the genetic and genotypic diversity ofthe population.
The collection of bundles from at least 3 colonies, but preferably 6 or more colonies is recommended for successful fertilization. Fertilization of gametes from 2 colonies may resultin a low fertilization rate depending on the
combination and status of the donor colonies.
In addition, the bundles can be collected from
cryptic species that are not easily distinguished based on morphology.
When the spawning is ceased, gather thebundles at the surface of container and stir thewater with a spatula or hand to separate theeggs and sperm. If 6 donor colonies are available, we recommend that the colonies be divided into 2 separate containers for cross-fertilization. If bad eggs or eggs of cryptic species
are contaminated, unfertilized eggs and abnormal embryos occur, and they can deteriorate rearing water and lead to mass decay ofthe entire egg supply during rearing.
For experimental purpose of fertilization orin case if the origin of gametes must be clarified, use the gametes from 1 fragment in a
separate container by stirring bundles at thesurface gently with a spatula. Collect the eggs
with a pipette or plastic cup and transfer themto another container filled with fresh, filteredseawater. The eggs should be rinsed twice withfresh, filtered seawater by repeating this treatment. Sperm are collected from the opaque residual solution after collecting the eggs. Then,the rinsed eggs are mixed with the spermfrom another fragment(s) from different container(s). The spatula, pipette, and cup are prepared for each container to prevent the contamination of sperm from other colonies.
Egg density should be such that only a thinlayer of eggs cover the surface at the time of
insemination. Sufficient sperm should be used
to turn the seawater opaque (105–107/mL). Insemination should be completed within 4 h after spawning (Fig. 7). Allow fertilization to occur for 20–30 min, stirring occasionally andthen skim the fertilized eggs from the surface
using a glass or plastic cup. Transfer the fertilized eggs twice from 1 container (5–10 L) toFig. 7. Fertilization and development of embryo

another to wash away excess sperm that may
cause the water quality to deteriorate. The
rinsed eggs are maintained in a large rearing
tank (100–500 L). Fertilized eggs should bemaintained scattering thinly over 10–20% of
the surface. Avoid stirring the water for 12 hafter fertilization, because early developmentof the embryos is delicate.

Fig. 8. Slicks drifting ashore inside a harbor

3–2. Determination of the fertilization rate
The embryos develop into morulas and blastulas 4–8 h after fertilization. The fertilization
ratio is measured by counting unfertilized(round in shape with no cell division) and fertilized eggs using a dissecting microscope.
Under normal conditions the fertilization ratio should be ＞90%–95%.
4. Sampling of slicks after synchronous
spawning
In many cases, a long narrow slick will formon the sea surface from night-time until themorning after synchronous spawning. These
slicks often remain in harbors or drift onto theshore at the Ryukyu Archipelago where fringing reefs develop. When the slicks appear in a
harbor or shore, they should be collected soonafter spawning, as the blastomeres from the2-cell to the morula stage can easily dissociateand die from wave impact against the pier orshore (Fig. 8).
An offshore slick can be collected by boat in
the morning after spawning (Fig. 9). Slick collection is recommended, as it minimizes landbased contaminants. However, slicks may not
be found because of ocean conditions.
Skim the slicks off the sea surface using a

Fig. 9. Offshore slicks

dipper and place them into a bucket with freshseawater. The eggs and embryos that are contained in the slicks are mostly Acropora species, but Montipora and other species can also
be found. Eggs and embryos of Montipora are
distinguished from those of Acropora because
they are smaller in size and possess zooxanthellae. The status of the embryos should be
examined using a dissecting microscope before
rearing. The embryos may not survive if significant dissociation of the blastomere and/or unfertilized eggs is observed. If a large quantity offoreign matter has contaminated the water, agood result could not be expected.
7Column 1. Spawning induction
There are a very few successful cases of artificially induced coral spawning. However,it can be accomplished with some Acropora
and Montipora species using hydrogen peroxide (Fig. 10) (Hayashibara et al. 2004).
Add hydrogen peroxide (2 mM for 3 h exposure time or 5 mM for 2 h) after transferring the fragments of a mature colony fromthe sea to a container on land. After that,rinse the fragments to remove the hydrogen
peroxide and transfer them into clean, continuous flowing seawater for spawning.
Spawning by artificial induction occurs from
20 : 00 to 23 : 00 h under natural lighting,similar to natural spawning in the sea. A period of at least 9 h is needed between induction and spawning (initiation of spawningwith higher probability requires 16 h). If thetime is ＜9 h until the estimated spawninghour, induction should be postponed until
the following day to secure enough time between induction and spawning. Iwao (2000)
indicated that if the fragments are maintained in the light for ＞9 h and then in thedark by covering the container with a dark
curtain, spawning occurs within 2–3.5 h.
Although fertilization of gametes and subsequent larval development has been confirmed by this method, this treatment may
considerably damage the coral colonies including secretion of mucus or emission of
zooxanthellae, which may lead to death. Because the degree of damage caused by hydrogen peroxide varies according to species
and condition of the colonies, observing thestatus of the colonies during processing is
recommended. The concentration of hydrogen peroxide and the length of exposure
should be adjusted with the condition of thefragments for successful artificial spawninginduction.
5. Maintenance and rearing of coral larvae
5–1. Rearing larvae in aquaria on land
Fresh seawater is very important when rearing embryos and planula larvae (Fig. 11).
Twelve hours after fertilization, the seawaterin the rearing tank should be changed once
every 8–12 h until the larvae become compe8

Fig. 10. Spawning induction using hydrogen peroxide(Experiment for comparison)tent to settle onto a substrate. During the first
2 days, as embryos are drifting to the surface,seawater in the rearing tanks can be changed
easily. However, because the larvae distributewidely in the tank, changing all of the water
becomes difficult. Seawater can only bechanged partially. In addition, in the earlystages, the larvae tend to gather at the surface
edge of the rearing tanks and this causes massmortality because of oxygen deficiency. Thus,
the seawater should be mixed gently to avoid
dense concentration of the larvae. Iwao devised an apparatus (trapping tank) that enables the rearing water to be replaced continuously with fresh, filtered seawater without
losing the embryos and larvae from the rearing tank (Column 2).
The recommended density of larvae perrearing tank (300–500 L) is ＜2000 larvae/L ofseawater. Divide the larvae into several rearing tanks to maintain suitable conditions andat low density without changing the water. Using eggs from bundles, Shimomura et al.
(2002) successfully raised 50×104 larvae/tonof seawater (Fig. 12).

Fig. 11. Rearing larvae and maintenanceto obtain an accurate number because the larvae are unevenly distributed on the surface
during the early developmental period. In addition, larval distribution is affected by wind if
a large outdoor floating pond is used. Sampling and counting are needed at least 3–5
times after gentle stirring the water to obtainreliable values.

Fig. 12. Rearing larvae in a large water tank

When embryos and larvae collected fromslicks are reared in tanks, the seawater quality
may rapidly deteriorate because of the presence of foreign matter and decaying unfertilized eggs; thus, the density should be lowered.
Water temperature should be maintained at＜28°
C. Light intensity at the surface should
be lowered to 70%–50% using a black shadescreen when rearing outdoors.
The duration of larval settlement capability
varies depending on the species and temperature. The usual period needed for the acroporid planulae to begin settling is 5.5 days at a
C or 5 days at 27°
C.
water temperature of 26°
The density and total number of larvae
raised can be estimated by counting the number of larvae in 0.1 L of seawater sampled fromthe rearing tank. However, it is often difficult

Column 2. Rearing embryos and larvae
using a flow-through trappingtank
Larval rearing in land-based aquaria istroublesome with changing water. This is particularly true in the case of large-scale rearing. We have devised a set of drainage pipes
covered a large area with nylon filter mesh
(60–100 μm) (Fig. 13). Fresh seawater runs
from large seawater holding tank to a roundtrapping tank through 2 thin vinyl tubes andproduces a gentle circular flow in the trapping tank to prevent dense aggregation ofembryos and larvae at the surface. It is possible to rear 50–70×103 acroporid larvae using
a 100 L trapping tank and save labor.
This flow-through system is also useful torear larvae from gonochoric broadcasters
and brooders that spawn eggs without bundles or release planulae over an extended period of several hours or those in which there9

Fig. 14. Collection of larvae in a running water aquarium

Fig. 13. Devices for rearing coral larvae in a trapping tank.
Seawater is sent from a seawater holding tank to trapping tanks through thin vinyl tubes.

leasing time is unknown. When we maintained fragments of the gonochoric broadcast spawners Fungia sctaria and Ctenactis
echinata in the holding tank, eggs and sperm
were released and they were subsequently
fertilized in the holding tank. The fertilized
eggs were sent to the trapping tank andreared there until the larvae were capable ofsettling. In addition, we have successfully
used this system for collecting planulae fromblooding corals such as Pocillopora damicornis and Seriatopora hystrip.
5–2. Collecting brooding coral planulae
Brooding coral such as Pocillopora damicornis may release planulae several times a year,
and larval rearing time is generally shorter
than that for broadcast spawners because thelarvae are already fully developed and ready tosettle. These observations seem to be advantageous for artificial production of juvenile coral. However, because it is difficult to ascertain
whether the coral is ready to release planulae,
selection of adult colonies to collect larvae isdifficult. A number of collectors have to be set
over colonies. Large size bundle collectors
similar to that shown in Fig. 6 or plankton net(mesh opening 100–200 μm) with a screw capbottle and float are used to collect the larvae.
The collector must be set over a colony tightly
so that the larvae do not escape from opening.
It is necessary to set an upturned funnel in the10mouth of screw cap bottle so that larvae in thebottle will not escape back down into the collector (Guest et al. 2010). It is also advisable toopen windows with net on the screw cap bottle to prevent stagnation inside the bottle if thecollector is left overnight.
Ex situ collection of planula larvae is easierthan in situ collection. As shown in Fig. 14,
donor colonies or coral fragments can betransported to a land-based flow through tankbefore the predicted day of planulation. The
seawater in the holding tank allows the larvae
to flow through an overflow pipe and be transferred to a separate collecting tank (trappingtank) that contains a mesh filter (see Column
2). The trapping tank should be periodicallymonitored each morning for the presence of
planula larvae. Any larvae detected are collected at once and transferred to a clean container.
5–3. Mass production of larvae using a floating pond
A large number of larvae can be reared in a
floating pond (Omori et al. 2004). The floating
pond (2.0 m×2.0 m, 1.0 m deep) is made of vinyl sheets used for eel aquaculture (Fig. 15). A
submersible pump is used to deliver the seawater through a hose with small holes that are
attached around the edge of the pond. Freshseawater is gently sprayed against the walls toprevent the larvae from attaching to the wallsof the vinyl sheet. Four screen windows arepresent on the sides of the pond and 1 at thebottom to allow the outflow of seawater. The
screen windows should be cleaned if they become clogged with microalgae and other matter to ensure continuous flow of seawater.
Omori et al. (2007) succeeded in rearing＞420×103 coral larvae (134 larvae/L) in eachpond.

Fig. 15. Large-scale rearing of coral larvae in a floating pond (a). Sketch (b) and dimensions (c)6. Transportation of larvae
Embryos can be transported when they attain the shape of a ball. However, the recommended time for transport is 3–4 days after
fertilization when the larvae are well ciliatedand move actively. At this stage, the larvae arestronger and can tolerate various stressors. A
container with 2000 larvae/L can be transported to various places in Japan using a generalparcel delivery service at normal temperature
with survivorship ＞90%. Sending live larvae
by air from AMSL to several public aquariums
in Europe using 50-mL vials is succeeded (Petersen et al. 2005).
To reduce the stress from vibration duringhanding and transportation involving a longdelivery time, larval density to be low in asealed container without air space is advised.
A collapsible polyethylene container (10 L or
20 L) is suitable when transporting a large
number of larvae into an underwater net enclosure in order to enhance larval settlementon substrate. This type of container can besqueezed to release the larvae underwaterwith a SCUBA diver.

7. Induction of settlement and metamorphosis
The mechanisms of settlement and metamorphosis of hermatypic coral larvae have notbeen fully revealed. However, for acroporid
larvae in Okinawa these processes can be induced with extracts from Hydrolithon reinboldii, a species of crustose coralline algae (Morseet al. 1996). The algal tissue or algal chips areadded to a vessel containing the larvae in filtered seawater to induce settlement. The chips
are collected by scraping the surface of the algae using a knife. They can be preserved bysoaking in a rifampicin water solution(2 mg/L) followed by washing and dehydrating
before freezing. If the chips remain frozen,
their effect can be maintained for at least 1
year. The larvae will settle in 6–24 h after adding the thawed chips once they become competent to settle onto a substrate. In many cases, the larvae settle at the bottom of thecontainer near the chips or sometimes on thechips. A thalloid red alga, Peyssonnelia sp., hasa similar effect on acroporid coral (Iwao
1997). Settlement rate is 60%–90%.
Inducing settlement with coralline algal
chips is easy, but collecting the algae may bedifficult and preparing the chips is tedious. On
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an experimental scale, the use of neuropeptide
Hym-248 promotes higher, more stable induction of settlement and metamorphosis in acroporid larvae (Iwao et al. 2002).
In farming corals, we use a “conditioned”
substrate that is maintained in the sea for 4–8
weeks with the surface covered in coralline algae and biofilm to enhance settlement of corallarvae (Column 4). The conditioned substrate
may maintain its effect even after it is desiccated for certain period.
Column 3. Change in settlement capabilityof larvae
The fertilized eggs repeat cleavage and develop into planula larvae capable of metamorphosis and settlement. The duration toreach this stage varies depending on the species. Acroporid larvae take 5–5.5 days at
C–27°
C, whereas faviid species take 2–3
26°days. The time taken by the larvae to maintain competency to settle also varies by species. For example, Acropora tenuis can settle
for ＞20 days, but the competence becomes
highest for only 1–2 days and is lowered
during subsequent 2 weeks in A. nasuta,
suggesting species-specific differences in thewindows of opportunity for successful settlement and metamorphosis on the substrates (Fig. 16). The reseeding process infarming corals should be taken when larval
competence peaked.

such as Seriatopora hystrix and Heliopora
coerulea may settle on any material, whereas
other species choose a location in responseto an environmental signal from the seafloor. Settlement and metamorphosis occur
consecutively after stimulation by multiple
signals but they are separate phenomena. If
no signals are provided, larvae remain in thewater column for a considerably long period
without settlement or metamorphosis evenwhen they are competent to do so. On theother hand, if some stimulation is given, thelarvae may metamorphose while suspended
in the water column without settling.
Besides crustose coralline algae (Morse etal. 1988, 1996), bacteria on the coralline algae
and a microbial biofilm lacking coralline algae play a role in triggering settlement andmetamorphosis (Negri et al. 2002, Webster et
al. 2004). A significant relationship is observed between the number of settled larvae
and coralline algae covering the substrate in anatural setting (Fig. 17). Biofilms comprisecomplex distinct bacterial and microalgal
communities, varying by time and depth.
Thus, the effect could be different with species. Webster et al. (2004) found that 2-weekold biofilms induce metamorphosis of Acropora microphthalma in ＜ 10% of larvae,
whereas the metamorphosis rate increased
significantly to 41% on an 8-week-old biofilm. The settlement rate of Favites halicora
larvae was 25% on a 2-week-old immersed
substrate in the sea, whereas it increased to
68% on a 2-month-old substrate (Guest et al.
2010). In contrast, some Porites and Acropora
species settle preferentially on unconditioned

Fig. 16. Change in settlement capability of coral larvae

Column 4. Larval settlement and metamorphosis
Larval settlement and early post-settlement survival of coral explain much of thevariance in the adult population. Some coral
12

Fig. 17. Relationship between percentage cover of coralline
algae and number of coral polyps settled onto the substrate (Yamaki 2010)

red substrata (Mason et al. 2007). The preferred red substrata have their spectra dominated by wavelengths 580–590 nm that issimilar to the fluorescent emission peaks ofcoralline algae, indicating that the larvae determine settlement using light signals. (Thisphenomenon was not however confirmed byour experiment with Acropora tenuis.)
In nature, some species select a settlement
location in relation to chemicals originatingfrom coralline algae, whereas others attach toand metamorphose on biofilms. Larvae utilize
a variety of environmental cues such as biological constituents and color of the substrate.
This behavior appears to be a strategy of eachspecies to optimize survivorship and growth.
8. Substrates
8–1. Quality and construction of an artificial
substrate
Shells and various substrates made of concrete, slate, unglazed ceramic tile, polycarbonate, and fiberglass-reinforced plastic have beenused. However, it is difficult to determine
which one is best. Nondurable materials suchas wood or bamboo are not recommended. A
concrete substrate should be placed in the sea
for some time to decrease its alkaline content.
Although it is unclear whether this is becauseof the quality or surface structure of the substrate, it has been reported that larvae exhibit
greater settlement affinity for substrate coatedwith calcium carbonate with electrochemical
method than for unglazed ceramic tiles (Kihara et al. 2013).
Because more larvae settle inside irregularareas than on smooth surfaces, the substrates
with several grids or ridges and numerous
hollows are considered to be more effective forsettlement. However, for induction of larval
settlement biological conditioning of the surface of substrate is more important than raw
material or surface structure of substrate. Before larval settlement induction, other biota(e.g., macroalgae, colonial tunicates, sponges,
and sea anemones) than crustose coralline
algae and biofilm should be removed by gently
brushing the substrate surface.
We used a square concrete or unglazed ceramic tile (10 cm×10 cm) as a substrate. How-

Fig. 18. Coral pegs. A number of polyps settle on head of thepeg (a, b). Juvenile coral grown there (c).

ever, the square tiles are not suitable for transportation and outplanting once juvenile corals
grow on them. In addition, a considerable
amount of epoxy is required to attach them tothe reef. The square tiles that had nicks like achocolate bar were devised so that they could
be split into several pieces in the sea. However,juvenile corals often grew over the nicks.
A “coral peg” is currently being utilizedalong with the square tile, as it allows for easytransport and outplanting than the tile. The
total height of the coral peg is 60 mm; thehead is made of concrete cement mixed withquartz sand and is 18 mm in diameter and
10 mm in height. The leg (plastic) is 10 mm indiameter and 50 mm in height (Omori and
Iwao 2009) (Fig. 18). The juvenile coral grewon the head of the coral peg. They can betransported and fixed onto a crevasse of thereef using epoxy.
Column 5. Structure of the artificial substrate
The microscale turbulence produced by
13

the complex shape of the substrate surface
serves to retain the approaching larvae on
the settlement surface. It is known empirically that coral larvae tend to settle more on
the edge or a groove with approximately
1 mm wide and 1 mm deep of a substrate
than its flat surface. However, the relationship between the physical movement of water and behavior of coral larvae on a complex substrate structure has not been studied
in detail. On the other hand, surface irregularities reduce the danger of predation byfish and scraping by sea urchins after settlement. According to Nozawa (2008), while all
juvenile corals of 3 species including Echinophylla aspera that settled on a plain tile surface died within the first 4 months at sea,others that settled in the small crevices of
3–4 mm deep on the substrate survived.
Nozawa (2012) monitored juvenile E. asperaand Favites abdita that settled on artificial
substrates with projections at 5-, 15-, and25-mm intervals for 2 years in the sea. He
found that coral survivorship on the 5-mm
refugia was significantly higher than that inthe 2 larger intervals.
It must remind that crevice on substrate isalso excellent space for survival of algae thatare competing with coral spats.
8–2. Larval settlement on the substrate
Reseeding (larval settlement on a substrate)
must be accomplished at the peak period ofcompetency. The competency should be tested
twice per day by placing 10–20 larvae and coralline algae chips in a culture dish for 12 h (seetext 7). Reseeding work should be prepared inadvance considering that in many cases larval
settlement competency is highest at half or 1
day after its capability is indicated (Column
3). Larval settlement will be completed in 1–5
days (Fig. 19).
Reseeding can be performed in aquaria onland using a tank (100–500 L). Coral larvae arereleased into a tank containing many substrates (Fig. 20). Settlement rate varies by timing, but usually ＞60% of acroporid larvae settle onto tiles within 1 day, if larval density is100–130 larvae/L. If reseeding is accomplishedoutdoors, the aquaria should be shaded to re14

Fig. 19. Juvenile corals on a conditioned substrate (right, enlarged photo)Fig. 20. Larval settlement on ceramic tiles

duce sunlight by 50%. If the number of spat
on a substrate is lower than the expected number after 1 day, more larvae should be addedon the second day. More larvae may settle
onto one side of the substrate when the title isused. If larval settlement on both sides is desired, the tile should be turned over on thesecond day. Water in the tank is gently agitated throughout the settlement by bubbling air
through the water column. The substrates
should not to be hit directly by air bubbles.
The water should be carefully changed during
the reseeding process using a filtering apparatus to avoid losing any larvae.
We recommend that the initial density
should be 0.5–1.5 spats (polyps)/cm2 on a substrate. On the one hand, if the density is very
high as such that an individual spat attach toneighboring spat, it may cause significantly
high mortality (＞90%) because of infection
during the early post-settlement period. On theother hand, coral spat will tend to form large
chimeras at very high densities and this maylead to higher survival and increased growthrate if fused colonies reach an escape size more
rapidly than single spat (Puill-Stephan et al.
2012, Raymondo and Maypa 2004).

Column 6. Introduction of larvae to a substrate in the sea
It is possible to enhance settlement of larvae onto a substrate, underwater, by releasing the larvae into an enclosure using a net.
AMSL and the then Tetra Co. Ltd. have constructed a nursery using a conditioned concrete block (1.3 m wide) surrounded by aseat (5.5 m long, 5.5 m wide, and 6.0 m high)at Naha Harbor, Okinawa. The concrete
block was covered with fine mesh net, andapproximately 1.64 million larvae cultured
at Akajima were released into the enclosure
by SCUBA divers (Aota et al. 2003).
Heyward et al. (2002) developed anothertechnique using a movable floating poolcontaining coral larvae cultured from a slick
collection that was placed over the area to bereseeded. Fresh seawater was pumped inside

Fig. 21. An example of a method that delivers larvae to the enclosure on the seabed from a floating pool (Heyward
et al. 2002)

the floating pool to raise the water level sothat the larvae were sent by hydraulic pressure to the settlement substrates in an enclosure of camping mosquito net on the seafloor through a tube connected to theunderside of the pool (Fig. 21).
The results of these in situ reseeding experiments demonstrate that early recruitment can be significantly enhanced. However, most settled juvenile corals die because ofnatural causes in the sea within the first few
months (e.g., Sato 1985). Thus, this method
is not favored as a reef rehabilitation technique until positive evidence of a long-term
effect has been demonstrated. Hayashibara(2009) devised a special substrate for settlement of the branched coral larvae and indicate possible result for this method (see also
Suzuki et al. (2012)).
Column 7. Acquisition of zooxanthellae
Offsprings of many coral (approximately
80% of broadcast spawners) acquire symbiotic zooxanthellae (Symbiodinium spp.) horizontally from the natural environment(Baird et al. 2009). Although planktonic
planulae may acquire zooxanthellae, generally the coral spats begin to acquire the symbionts within 5–7 days after settlement. It isessential for them to acquire zooxanthella
during this period. A failure may lead todeath. Establishing symbiosis may be enhanced by introducing giant clam or coralfragments from outside to the rearing tankat the time of reseeding or immediately after
settlement. This process is easy, although theeffectiveness of the introduction of zooxanthella in the rearing tank for the spatʼs survivorship has not been distinct as yet. The coral will turn yellowish when the number ofzooxanthellae in the larvae increases. The
genetic type (clade) of Symbiodinium spp. ismostly type C in adult coral, but it is mostly
type A and/or type D when coral spats are
acquired (Abrego et al. 2009, Yamashita etal. 2013). Introduction of the cultured stockof planktonic type A and/or D into the tankmay usher more definite results (see Suzuki
et al. 2013).

15

9. Culturing juvenile coral
9–1. Commencement of rearing
In the sea, nibbling fish and/or coral eating
gastropods, sea urchins, and amphipods reduce the number of newly settled juvenile corals. Corals are also vulnerable to sedimentation and overgrowth of algae, colonialtunicates, and bryozoans. It is recommended
to rear corals in an environment where thesefactors that lead to mortality are nonexistent.
However, there are currently no methods toeliminate these factors, unless the juvenile
corals are raised on land using large flowthrough seawater aquaria.
At AMSL we rear the juvenile corals on 15
pairs of ceramic tiles (30 faces, 10 cm×10 cmeach) or coral pegs that are maintained in anursery cage (80 cm×55 cm×30 cm) suspended at a depth of 1.5–3.0 m from the sea surface(Fig. 22). One hundred individuals ofalgae-grazing juvenile top shell snails Trochus
niloticus (5–10 mm in basal shell diameter) are
reared in each cage (i.e. ca. 70 inds./m2) (Fig.
23). Grazing trails of the juvenile top shellsnails are nearly harmless to coral spats, unlessdensity and/or size of the grazers exceed
threshold (Tamura 2008). Other algae-grazing
snails (We used Clypeomorus batillariaeformis
in Palau.) may be used to algal control, but it
is necessary to check if they do not scrape or
dislodge the corals while creeping. Althoughmany sea urchins are also grazers, Diadema
should not be used, as they clash and eat thejuvenile corals (Sato 2010). Sedimentation
may cause newly settled coral to die. Thus, thesubstrate tiles are arranged vertically orobliquely in the cage.
9–2. Maintenance during rearing
The condition of juvenile corals in the nursery cages should be checked periodically andalgae (top shell snails appear to avoid foliose
algae), sea anemones, sponges, and colonialtunicates should be removed from the cages
and substrates as much as possible using a
brush and scraper. If juvenile coral are rearedon land using large flow-through nursery
aquaria, in addition to algae, sea anemones
can be successfully removed by releasing juveniles of algae-grazing rabbitfish Siganus spp.
juvenile, and/or butterfly fish Chaetodon
16

Fig. 22. Nursery cage and ceramic tiles

Fig. 23. Co-culture of juvenile corals (arrows) and top shellsnails in a nursery cage

Fig. 24. Culture of juvenile corals in nursery cages (Photo
courtesy of Mr. H. Kobayashi, Asahi Shimbun)

Fig. 25. One and half year old Acropora tenuis in a nursery cage

kleinii. Water movement is inevitably stagnated in the nursery cages causing a negative impact on coral growth. Coral species such as
Acropora digitifera and A. hyacinsus requireconsiderable water flow to sustain growth(Boch and Morse 2012). When the juvenile
corals grow 1–2 cm in diameter, the mesh sizeof the nuresry cages may be changed or the
top cover of the cage may be removed so thatwater flow is enhanced and algae in the nursery cages are removed by fish.
When the juvenile corals grow to a size of＞5 cm in diameter, they could be outplanted(Figs. 24 and 25). This is important, as the unit
for both propagation and eventual reef rehabilitation, that each substrate contains 1–3 living corals, not the total number of living corals.
10. Outplanting
The juvenile corals are outplanted with theirartificial substrates onto the outcrops near
their native reef (Fig. 26). If the outplantingsite is far from the native site, we choose a location with physical and chemical conditions
as well as a coral species composition similar
to the native site. If the corals die at a prospective rehabilitation site because of an episodic
disturbance, dead coral heads or coral rubble
should be used to identify the site.
Smaller colonies (＜2 cm) may be perishedbecause of various factors including nibblingby fish or predation by corallivorous muricid
snails. Larger colonies may reduce their size
following outplanting when their branch tips
are removed by fish but they survive and re-Fig. 26. Underwater outplanting operation (Photo courtesy of
Mr. H. Kobayashi, Asahi Shimbun)

sume growing.
Outplanting immediately before the stormy
season and/or during the year when sea temperatures are highest should be avoided tominimize risk of death. Bleaching may occur
and diseases may be prevalent during thewarmest months of the year. Thus, operating
during cooler months and in sheltered reef environments is recommended where the outplanted coral will be protected from wave action.
The juvenile corals can be fixed to convex
surfaces of the coral limestone outcrops using
stainless steel wire or adhesives (refer to Okubo and Omori 2001). Marine adhesives (e.g.,
Konishi marine epoxy bond, Epoxyclay
AquaTM, S-dine Joiner, etc.) are sufficient to secure corals on artificial substrates to the reef.
Before fixing, the point of attachment should
be brushed lightly with a wire brush to remove
algae and sessile invertebrates on the surface.
For sites such as the side of a steep outcrop or
areas of high wave action, the substrate should
be secured with a concrete nail or stainlesssteel rod in addition to the stainless steel wireand epoxy. As the movement of sand andgravel easily damages coral on a flat seabed,
outplanting to such site should be avoided.
The coral colonies and substrates should befirmly fixed on the side or top surface of an
outcrop, at least 50 cm above the sea bed.
There should also be a distance of at least
30 cm between the outplanted colony and a
nearby coral to provide ample space for future
growth.
With no regard to how they were fixed, the17

success of the outplanting is totally dependenton whether the coral and substrates are firmly
fixed on the reefs. Therefore, it is very important to check all outplanted coral within 1 day
after the operation is finished. After outplanting, the living coral gradually extends over thewire and adhesives and reaches toward thenatural substrates. If outplanted colony is
moved by waves, the coral tissue will not reachthe natural substrates. The outplanting is considered a success when living tissue reached
the outcrop. The growth rate of the living tissue varies under similar environments depending on the species. If living tissue is distant from the natural substrate, it may retard
attachment of the coral tissue to the outcrop
and be easier to die.
The epoxy is composed of a resin and ahardener. The chemical reaction between the 2parts does not occur until they are mixed. The
2 parts are mixed immediately before underwater application. Because the time taken by
the epoxy to remain workable and harden may
differ by manufacture and water temperature,detailed and advanced planning of the underwater operation is necessary.
The effect of epoxy on marine invertebrates
including the outplanted corals has not beenstudied in detail, because the chemical compound of each product are seldom shown.
Thus, care should be taken to minimize directcontact between live coral tissue and epoxy.
Although we have not encountered a negative
impact of underwater adhesives on coral, theeffect should be investigated further for cases
in which large amounts of epoxy will be used.
The outplanting operation is the most expensive process of reef rehabilitation, because
considerable time is spent to secure the coral
to the reef along with the costs accompanying
the use of boats and SCUBA divers. Analysisof several studies suggests that only 10–15
substrates (with corals) can be fixed per person per hour. Coral colonies grown on sticklike substrates, such as coral pegs, can be slotted into predrilled holes or natural holes of thereef with a small amount of adhesive. Thus, anoperation using the coral pegs can be performed faster and cheaper than attaching flattiles.

18

Column 8. Transplantation, outplanting,and translocation
The term “transplantation” is commonlyused for removal of an entire coral colony ora fragment from one place to resettle in another. The operation can be divided into 4
parts as follows: 1. Direct transplantation ofcoral fragments that were clipped/fragmented from donor colonies and “coral of opportunity” to the reef. 2. Transplantation ofnursery-grown asexually propagated fragments. 3. Transplantation of nursery-grownsexually propagated colonies. 4. Transplantation of entire coral colonies to rescue theones that would otherwise be destroyed orseverely damaged by coastal and/or underwater development. Survaival rate of nursery-reared fragments that were grown to an
adequate size on substrates is higher thanfragments that were clipped and/or collectedin the sea and directly transplanted onto thereef (Shafir et al. 2010). We propose that theterm “transplantation” be used for the direct
removal of fragments onto a reef (1). The removal of nursery-reared corals (2 and 3)
should be expressed by the term “outplanting”. Removal of entire coral colonies or
those together with a hard substrate for mitigation purposes (4) should be termed as“translocation.”
Column 9. Protecting outplanted juvenile
corals from nibbling by fish
We have outplanted cultured, juvenile colonies of Acropora tenuis (1-year-old; approximately 3 cm in diameter) on the Reef BallsTM
deployed at Lukes Reef, Palau in March
2007. However, 6 months after outplanting
nearly all of the coral were broken, as well assome parts of the tile substrate. The culprit
was suspected to be parrotfish, although further investigation is needed. Similar damage
was observed at Akajima Island with the A.
tenuis juvenile colonies (3–4 cm in diameter)a few days after outplanting. Based on ourobservation, nibbling of the barred filefish
Cantherhines dumerilii and others were
thought to be the cause. Predation exclusion
experiments were conducted by placing protective cages over the corals (Fig. 27). Pre-Fig. 27. Outplanting of 1-year-old juvenile colonies in protective cages

liminary results showed that nibbling/predation by fish interfered with the cages, butalgae had grown on some cages. Therefore,
the cages must be removed at an appropriate
time (＜1 year), by so doing, grazing fish
may remove the algae.
11. Growth and spawning of outplanted coral colonies
When juvenile Acropora tenuis in mid-water
nursery cages, which were of 1.5 years postfertilization, grew to approximately 6 cm in diameter, approximately 2000 colonies were outplanted with the tile substrate onto bommies
at Akajima Island in December 2006. The coral grew well (89% survival 6 months after outplanting) until they were seriously damagedby waves driven by the typhoon Man-yi in
July 2007. An approximate of 60 surviving colonies grew to 20–25 cm in diameter and morethan seventeen colonies had spawned at
4-year-old initially (Iwao et al. 2010, Omori etal. 2008).
Up to 2013, we have outplanted ＞6000 juvenile corals cultured from eggs on limestone
outcrops at Majanohama, Akajima Island.
Most of the corals matured 4 years later.
Spawning of the colonies has continued every
year since 2009 (Fig. 28). This indicates thatthe cultured coral had grown into “a mature
population” capable of producing the next
generation. At present, many kinds of damselfish and butterfly fish swim in the outplanting
area and gobies and crustaceans inhabit thecorals, indicating that the present restoration

Fig. 28. Spawning of outplanted Acropora tenuis (5-year-old)at Akajima Island

technique has certainly enhanced the rich biodiversity of the coral reef (Fig. 29)
Column 10. Cross fertilization and larval settlement rates of the outplanted corals
The merit of reef rehabilitation techniquesthat utilize outplanted corals cultured fromsexual propagation is greater compared tothat of asexually derived coral fragments,
which can reduce genetic diversity of a population. Gametes from the same colony areunable to cross-fertilize in many coral species. Low genetic diversity is a concern forreef restoration when seed fragments areraised asexually from only a few donor colonies. This can lead to lower fertilization rates
among seed fragments, and reducing thelong-term benefits of reef restoration in particular areas. Additionally, low genetic diversity can compound the effects of increased
ocean temperature and other environmental
stressors, further jeopardizing the health of areef.
We tested fertilization rates between gametes collected from 6 cultured colonies ofAcropora tenuis (5-year-old) that were cultured from eggs and outplanted and 3 wildcolonies of the same species. The tests
showed that all 4 crosses, i.e., between outplanted colonies; between natural colonies;
and between outplanted and natural colonies, resulted in fertilization rates ＞98.5%.
19

Fig. 29. Outplanted corals that were cultured from eggs in June 2005. The photograph was taken at Akajima Island in February 2010.

No difference in settlement rate (＞95%) of
the larvae (3- and 6-day-old) was observed
between the wild and cultured colonies aswell as in the crosses between the wild andcultured colonies (Iwao 2013). It is expectedthat the cultured colonies contribute to the
expansion of the population into neighboring areas.

boat transportation, but including ropes,
cages, rearing containers, and epoxy bond)was US$8,639. The total cost was US$28,806(Table 1). Assuming that 10 colonies remainedon each tile, 3,200 juvenile colonies could be
produced after 1 year of culture; excluding thecost for outplanting 320 substrates (US$4,795),
the cost of producing 1 colony would be
US$7.50 (750 JPY).

12. Cost estimate
We estimated the cost of cultivating sexually
propagated coral using our experimental
study in Palau in 2007. During the 1 year experiment our expenditures were as follows:
Personnel cost (operational costs) for production of the juvenile colonies using 320 tile substrates and 10 nursery cages was US$20,167.
The cost of equipment and consumables (setup costs excluding SCUBA tank rental and

Aknowledgements
Nippon Foundation provided financial support for the research at AMSL. A number ofvisiting researchers such as M. Hatta and A.
Watanuki and the successive staff members of
the AMSL helped our study. We would like toexpress our appreciation for their cooperation
and assistance. The authors would like tothank Enago (www.enago.jp) for the English
language review.

20

Table 1. Set-up and operational costs for producing juvenile coral colonies using 320 tiles in 10 nursery cages and outplanting inPalau.

Staff
involved(number)

Personhours

Equipment andconsurmablecost (US$)

Collection of donorcolonies

2

4

15

2 dives

Prediction of
spawning night

2

20

134

Snokeling

Collection of eggs
and insemination

3

15

2,173

Rearing of larvae

375

Larval settling onto
tiles

2

60

Preconditioning oftiles in the sea

2

8

Culturing Juvenile
corals in nursery

Setting the cages

330

Maintenance ofJuvenile corals innursery

Clearing andmonitoring

2

Outplanting

Transportation andoutplanting

5

Activities

DetailSampling

Rearing coral larvae

Was SCUBA
needed?

Was a boat
needed?

1 transport

Total operationalor running costs
(US$)
333

1,000

250
1,250

1,113

1,000

Snokeling

2 transports

667

4,436

15 dives

5 transports

2,500

108

140

54 dives

27 transports

9,000

50

628

25 dives

5 transports

4,167

subtotal

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