Sexual production of corals for reef restoration in the Anthropocene

Abstract

Coral-reef ecosystems are experiencing frequent and severe disturbance events that are reducing global coral abundance and potentially overwhelming the natural capacity for reefs to recover. While mitigation strategies for climate warming and other anthropogenic disturbances are implemented, coral restoration programmes are being established worldwide as an additional conservation measure to minimise coral loss and enhance coral recovery. Current restoration efforts predominantly rely on asexually produced coral fragments—a process with inherent practical constraints on the genetic diversity conserved and the spatial scale achieved. Because the resilience of coral communities has hitherto relied on regular renewal with natural recruits, the scaling-up of restoration programmes would benefit from greater use of sexually produced corals, which is an approach that is gaining momentum. Here we review the present state of knowledge of scleractinian coral sexual reproduction in the context of reef restoration, with a focus on broadcast-spawning corals. We identify key knowledge gaps and bottlenecks that currently constrain the sexual production of corals and consider the feasibility of using sexually produced corals for scaling-up restoration to the reef- and reef-system scales.

Full text

MARINE ECOLOGY PROGRESS SERIES
Mar Ecol Prog Ser
Vol. 635: 203–232, 2020
https://doi.org/10.3354/meps13206 Published February 6
1. INTRODUCTION
Up to half of the world’s tropical corals have been
lost in the last 50 yr (Pandolfi et al. 2003, Wilkinson
2008, Burke et al. 2011, De’ath et al. 2012, Hughes et
al. 2017, 2018), and over one-third of scleractinian co -
ral species are now at increased risk of extinction
(Car penter et al. 2008). Three pan-tropical mass-
bleaching events occurred in the last 4 decades —
includ ing unprecedented back-to-back bleaching
and mortality events on the Great Barrier Reef (GBR)
in 2016 and 2017 — collectively signalling an acceler-
ation in global reef decline (Sweet & Brown 2016,
Hughes et al. 2018). The increasing frequency, inten-
sity, and spatial scale of these thermal-stress events
no longer allows sufficient time between disturbances
© The authors and the Australian Institute of Marine Science
2020. Open Access under Creative Commons by Attribution
Licence. Use, distribution and reproduction are un restricted.
Authors and original publication must be credited.
Publisher: Inter-Research · www.int-res.com
*Corresponding author: c.randall@aims.gov.au
REVIEW
Sexual production of corals for reef restoration in
the Anthropocene
Carly J. Randall1,*, Andrew P. Negri1, Kate M. Quigley1, Taryn Foster2,
Gerard F. Ricardo1, Nicole S. Webster1,3, Line K. Bay1, Peter L. Harrison4, Russ C.
Babcock5, Andrew J. Heyward2
1Australian Institute of Marine Science, PMB 3, Townsville, QLD 4810, Australia
2Australian Institute of Marine Science, Indian Ocean Marine Research Centre, University of Western Australia, Crawley,
WA 6009, Australia
3Australian Centre for Ecogenomics, University of Queensland, Brisbane, QLD 4072, Australia
4Marine Ecology Research Centre, Southern Cross University, Lismore, NSW 2480, Australia
5Oceans and Atmosphere, Commonwealth Scientific and Industrial Research Organisation, St Lucia, QLD 4072, Australia
ABSTRACT: Coral-reef ecosystems are experiencing frequent and severe disturbance events that
are reducing global coral abundance and potentially overwhelming the natural capacity for reefs
to recover. While mitigation strategies for climate warming and other anthropogenic disturbances
are implemented, coral restoration programmes are being established worldwide as an additional
conservation measure to minimise coral loss and enhance coral recovery. Current restoration
efforts predominantly rely on asexually produced coral fragments —a process with inherent prac-
tical constraints on the genetic diversity conserved and the spatial scale achieved. Because the
resilience of coral communities has hitherto relied on regular renewal with natural recruits, the
scaling-up of restoration programmes would benefit from greater use of sexually produced corals,
which is an approach that is gaining momentum. Here we review the present state of knowledge
of scleractinian coral sexual reproduction in the context of reef restoration, with a focus on broad-
cast-spawning corals. We identify key knowledge gaps and bottlenecks that currently constrain
the sexual production of corals and consider the feasibility of using sexually produced corals for
scaling-up restoration to the reef- and reef-system scales.
KEY WORDS: Coral reproduction · Sexual reproduction · Coral restoration · Gametogenesis ·
Embryogenesis · Spawning · Settlement · Climate change
O
PEN
PEN
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Mar Ecol Prog Ser 635: 203–232, 2020
for coral communities to recover to their pre-distur-
bance cover and composition (Osborne et al. 2017,
Hughes et al. 2018, Lough et al. 2018, Ortiz et al.
2018). Furthermore, coral-bleaching events are no
longer constrained to years with extreme El Niño−
Southern Oscillation (ENSO) conditions (Hughes et
al. 2018), and are affecting reefs regardless of conser-
vation status and management regime (Selig et al.
2012), leading to calls for additional action, including
direct restoration interventions (see Anthony et al.
2017, van Oppen et al. 2017). At present, active coral
restoration is spatially limited (e.g. Hein et al. 2017),
and coral restoration programmes targeting entire
reefs and reef systems will require new approaches to
re-establish ecosystem functions at ecologically rele-
vant scales.
Most coral restoration programmes to date have
leveraged the clonal structure of coral colonies and
utilised ‘coral gardening’ techniques, whereby corals
are fragmented or microfragmented (asexually) and
outplanted onto degraded reefs or artificial reef struc-
tures (Bowden-Kerby 2003, Rinkevich 2005, 2008,
Forsman et al. 2006, 2015, Shaish et al. 2010, Johnson
et al. 2011, Young et al. 2012, Lirman & Schop meyer
2016). An alternative, and increasingly advocated ap-
proach, is to generate and deploy sexually produced
corals (e.g. Heyward et al. 2002, Edwards & Gomez
2007, Harrison 2011, Nakamura et al. 2011, Guest et
al. 2014, Omori & Iwao 2014, Chamberland et al.
2015, 2017, Harrison et al. 2016, dela Cruz & Harrison
2017, Pollock et al. 2017, Omori 2019, see also www.
secore.org).
Future restoration approaches involving the mass
production of corals are likely to benefit from combin-
ing traditional asexual propagation methods with ad-
vances in the sexual production of corals. Firstly, col-
lecting sexual propagules while leaving adult colonies
on the reef is a more ecologically sustainable ap-
proach. Secondly, the use of sexual propagules from
spawning events is arguably the most cost-effective
and feasible way to produce the large numbers of
corals required for restoration (Edwards 2010,
Doropoulos et al. 2019). Finally, and perhaps most im-
portantly, approaches using sexual reproduction pro-
mote genetic diversity, which is central to species con-
servation (Baums 2008, van Oppen et al. 2015, 2017).
The sexual generation of corals also enables selective
breeding of individuals with potentially advantageous
traits, such as temperature tolerance (van Oppen et al.
2017), which may already exist in bleaching survivors
and populations from naturally extreme or marginal
environments (e.g. Barshis et al. 2013, Howells et al.
2016a). Given the rapid and accelerating rates of en-
vironmental change associated with increasing green-
house-gas emissions, it is essential to maintain and
potentially enhance the acclimatisation and adapta-
tion potential supported by high genetic diversity
within existing coral populations. Consequently, res-
toration programmes that produce corals sexually
have started to gain momentum over the last decade.
In this review, we summarise the current state of
knowledge of sexual reproduction in scleractinian
corals within the context of reef restoration, with a
focus on broadcast spawning, which is both the dom-
inant mode of sexual reproduction and currently is
more amenable to large-scale larval culture than the
collection of planulae from brooding species. Addi-
tional information from studies of brooding corals is
included where relevant. We highlight key knowl-
edge gaps and bottlenecks in the coral life cycle
where human intervention may optimise or acceler-
ate processes for application in reef restoration. We
also summarise key knowledge needs to guide
research in the broader coral restoration community,
and consider the feasibility of recovering the struc-
ture and function of degraded coral reefs through
interventions with sexually produced corals.
2. THE CORAL LIFE CYCLE
Scleractinian corals have a biphasic life cycle com-
posed of a dominant sessile, benthic polyp phase and
a motile planula larval phase (our Fig. 1; Harrison &
Wallace 1990, Richmond & Hunter 1990, Harrison
2011). Polyps form the basic unit of a coral colony,
and the polyp phase is dominated by somatic growth
and asexual budding that creates new and geneti-
cally identical polyps. Once a coral colony reaches an
adequate size and age, a cycle of sexual reproduction
commences (Fig. 1).
The modes of sexual reproduction have been identi-
fied for approximately half of the estimated 900 ex tant
species of hermatypic scleractinians (Veron 2000),
and 4 general patterns have emerged: (1) hermaphro-
ditic broadcast spawning; (2) hermaphroditic brood-
ing; (3) gonochoric broadcast spawning; and (4) gono-
choric brooding (Fadlallah 1983, Harrison 1985, 2011,
Richmond & Hunter 1990, Baird et al. 2009, Madin et
al. 2016). While most corals with known reproductive
patterns follow one of these strategies, some species
do not conform to the typical dicho to mies (i.e. brood-
ing vs. spawning and hermaphroditic vs. gonochoric;
Harrison 2011, Guest et al. 2012), and it is likely that
the diversity of recorded reproductive patterns will
continue to increase as more taxa are studied.
204

Randall et al.: Coral sexual production for reef restoration
Gonochoric species have separate sexes, while her-
maphroditic corals develop both eggs and sperm
within polyps or colonies. Most hermaphroditic
corals are simultaneous hermaphrodites, while some
are sequential (Harrison 2011), and a few exhibit
bidirectional sex change (i.e. Loya et al. 2009, Eyal-
Shaham et al. 2019). Broadcast-spawning corals
release gametes into the water column for external
fertilisation and larval development. Brooders typi-
cally undergo internal fertilisation and release well-
developed planula larvae, although some species
produce asexually brooded larvae (Harrison 2011).
Over 80% of species whose reproductive modes
are known spawn gametes for external fertilisation
(Baird et al. 2009, Harrison 2011).
3. GAMETOGENESIS, SPAWNING, AND REPRO-
DUCTIVE SYNCHRONY
Coral reproductive cycles are aligned with envi-
ronmental conditions that improve survival and fit-
ness. Environmental conditions can regulate repro-
ductive cycles in 2 ways: as ultimate factors, which
exert evolutionary selective pressures to maximise
reproductive success (Babcock et al. 1986, Harrison
205
Fig. 1. Stages of broadcast-spawning coral life cycle (after Jones et al. 2015), with stages of gametogenesis in the centre (after
Vargas-Ángel et al. 2006). Text sections of the present paper are indicated by the numbers in the filled grey circles, and key
restoration research priorities for each developmental stage are indicated with icons as defined in Table 2. Gametes develop
within or attached to mesenteries that contain oocytes and/or spermatocytes and progress through 4 stages, over several
months or more (I−IV; e.g. Szmant-Froelich et al. 1980, Szmant-Froelich 1985, Harrison & Wallace 1990, Glynn et al. 1991, Ver-
meij et al. 2004, Madsen et al. 2014). Oocytes (top row of inner cycle) begin as enlarging interstitial cells adjacent to, or within,
the mesenterial mesoglea (Stage I; Madsen et al. 2014), and slowly accumulate cytoplasm around the nucleus (Stage II). Over
time, oocytes increase in size, and yolk forms around the nucleus (vitellogenesis, Stage III), with final maturation (Stage IV) oc-
curring as the cortical layer and vitelline membrane are complete, as the yolk granulates and the egg becomes visibly pig-
mented in many species, and finally as the nucleus moves toward one side of the egg (Stage IV). Sperm development (bottom
row of inner cycle) begins with the clustering of spermatogonia adjacent to, or within, the mesoglea (Stage I), which is followed
by meiosis and the development of distinct spermatocytes with large nuclei in Stage II. The number of spermatocytes and
spermatids within the spermary increases during Stage III as meiosis continues, and they become peripherally arranged with
prominent central lacunae. In Stage IV, spermatids have little cytoplasm, develop a flagellum, and arrange themselves in
bouquet arrays within the spermaries

Mar Ecol Prog Ser 635: 203–232, 2020
& Wallace 1990, Oliver et al. 1998), and as proximate
factors, which cue and synchronise cycles. Ultimate
factors include seasonal temperature cycles that may
optimise physiological performance (Babcock et al.
1986, Keith et al. 2016), wind speeds (van Woesik
2010, Heyward & Negri 2012) and tidal phases (Bab-
cock et al. 1986, 1994) that may maximise fertilisation
success and dispersal, and diurnal cycles that may
allow for predator avoidance (Harrison et al. 1984). In
the context of coral restoration, we are unlikely to be
able to manipulate ultimate factors, and doing so is
risky and may decrease fitness. Therefore, we focus
here on proximate factors that act mechanistically,
which are more amenable to experimental manipula-
tion, and that are most relevant to restoration.
Proximate factors that synchronise gametogenic
and spawning cycles (Fig. 1) are thought to be regu-
lated by a hierarchy of exogenous environmental
cues (e.g. annual, seasonal, lunar, and daily, as de -
tailed in the following sections) that interact with
endo genous biorhythms at successively finer time -
scales to optimise reproductive success (Harrison &
Wallace 1990, Hoad ley et al. 2016). There is a rapidly
growing understanding of the endogenous and
molecular mechanisms that underpin reproductive
cycles and synchronisation (e.g. Sorek et al. 2014,
Kaniewska et al. 2015, Hoadley et al. 2016), which
shows great promise for our ability to manipulate
gametogenesis and spawning for restoration. How-
ever, it is critical that such manipulations retain the
reproductive characteristics of wild populations to
allow for interbreeding with existing populations
after deployment.
3.1. Annual and seasonal controls of
gametogenesis
For most broadcast spawners, gametogenesis fol-
lows an annual cycle; oogenesis takes place slowly,
over multiple months (4−12 mo), while spermatogen-
esis occurs more quickly (1−8 mo), with sperm matu-
ration coinciding with the final stages of oogenesis,
usually a few days or weeks prior to an annual-
spawning event (our Fig. 1; Wallace 1985, Szmant
1986, Harrison & Wallace 1990, Vermeij et al. 2004,
Vargas-Ángel et al. 2006, Chin et al. 2014). However,
multiple gametogenic cycles per year have been
described in some coral populations (e.g. Stobart et
al. 1992, Penland et al. 2004, Mangubhai & Harrison
2008a), and many brooding species have several
overlapping gametogenic cycles each year (Kojis
1986, Szmant 1986, Harrison & Wallace 1990, Foster
& Gilmour 2018), with some releasing larvae on a
daily basis (Nietzer et al. 2018).
Temperature has long been considered an impor-
tant factor governing reproductive seasonality and
often is correlated with gametogenic cycles in broad-
cast spawners and some brooders (Shlesinger & Loya
1985, Babcock et al. 1986, Kojis 1986, Hayashibara
et al. 1993, Vargas-Ángel et al. 2006, Nozawa 2012,
Keith et al. 2016). Oogenesis usually commences
asynchronously during cooler periods in autumn or
winter months (Wallace 1985, Szmant 1986, Harrison
& Wallace 1990, Vargas-Ángel et al. 2006); then, as
seawater temperatures rise rapidly in mid-spring,
oocyte diameter increases and oocytes gradually de -
velop from Stage I through Stage IV, culminating in
synchronised maturation (Stage IV) during periods of
warm seawater temperatures (our Fig. 1, inner cycle;
Wallace 1985, Willis et al. 1985, Szmant 1986, Var-
gas-Ángel et al. 2006, Madsen et al. 2014, Keith et al.
2016). While temperature clearly affects the rates of
physiological processes in corals, it likely plays a sec-
ondary role in determining the timing of spawning,
either by affecting growth and final maturation di -
rectly (Nozawa 2012) or through interactions be -
tween endogenous clock mechanisms (e.g. Kani ew -
ska et al. 2015) and temperature.
Solar insolation is another important cue that syn-
chronises gametogenesis, particularly for equatorial
corals that experience a small annual range in sea-
water temperatures (Penland et al. 2004, van Woesik
et al. 2006, Brady et al. 2009). Light is thought to
entrain oscillations of a molecular clock (~24 h perio-
dicity) through 2 interacting transcription/translation
feedbacks that form an endogenous rhythm, the
‘speed’ of which is in synchrony with seasonal
changes in day length (Hoadley et al. 2016). Light
acts as a proximate cue for reproduction, but is also a
fundamental energy source driving reproductive
effort. Reduction in solar insolation with increasing
latitude or water depth may constrain reproductive
effort, resulting in protracted gamete development
(i.e. ‘slower speeds’) and lower overall fecundity
(Kojis & Quinn 1984, Harii et al. 2001, Feld man et al.
2018, Shlesinger et al. 2018).
While spawning usually occurs once per year in a
synchronous-spawning event, it can also be protracted,
occurring as sequential events over several months
and seasons (Shlesinger & Loya 1985, Heyward 1987,
Hayashibara et al. 1993, Penland et al. 2004, Man gub -
hai & Harrison 2008b, Fogarty et al. 2012, Bouw -
meester et al. 2015). Biannual spawning within coral
populations has been observed in Western Australia
(Gilmour et al. 2009, 2016), and split-spawning — a
206

Randall et al.: Coral sexual production for reef restoration
form of asynchrony that occurs somewhat predictably
every 2−3 yr — happens either when the same colony
spawns over 2 or 3 consecutive months, or when differ-
ent colonies from the same population spawn on suc-
cessive lunar cycles (Willis et al. 1985, Mangubhai &
Harrison 2008b, Baird et al. 2009, 2012, Foster et al.
2015, 2018, Gilmour et al. 2016).
3.2. Lunar controls of spawning
Remarkably, gamete release within species can be
synchronised to within minutes, and often is re -
stricted to a single night of the year. Within seasons,
spawning is correlated with the lunar cycle and is
likely cued directly by moonlight detected through
photoreceptors (possibly cryptochromes or opsins,
see Hoadley et al. 2016) that sense moonlight inten-
sity (Levy et al. 2007, Kaniewska et al. 2015, but see
Linden et al. 2018). Some brooding species may use
similar processes to coordinate planulae release,
whereas other brooders do not exhibit lunar periodic-
ity in planulation (Jokiel et al. 1985, Vermeij et al.
2003, Zakai et al. 2006, Linden et al. 2018, Nietzer et
al. 2018). No data are available on photosensitivity of
cnidarian cryptochromes; however, they may form
part of the circadian clock rather than serve as photo-
sensors (Hoadley et al. 2016). Manipulating the day
of spawning or planulation based on offset artificial
moonlight cycles could be used to distribute coral
production in aquaculture facilities throughout the
month, reducing bottlenecks in culture processes
that could occur when relying on single monthly
spawnings. Manipulating lunar and seasonal tem-
perature and light cycles to split spawning across
months also could potentially increase coral produc-
tion in controlled aquaculture systems.
3.3. Diel controls of spawning
While several species have been observed to spawn
during daylight (e.g. Mangubhai et al. 2007, Eyal-
Shaham et al. 2019), the final and shortest temporal
cue for the majority of spawning species is the time af-
ter sunset on the night of spawning; conspecifics con-
sistently spawn a certain number of minutes to hours
after sunset (Willis et al. 1985, Babcock et al. 1986,
Vize 2006), and similar diel cycles of planulae release
are evident among some brooding species (Jokiel et
al. 1985, Fan et al. 2006). This timing can persist even
following artificial induction of gamete re lease
(Hayashibara et al. 2004) and when diel cycles are off-
set to manipulate the timing of spawning (Babcock
1984). This behaviour is directly controlled by local
photoperiod and does not appear to be entrained by a
biological clock (Brady et al. 2009).
3.4. Predicting and controlling gametogenesis and
spawning for restoration
Identifying the underlying drivers of synchronous
spawning among more taxa will enable the efficient
and reliable production of a greater diversity of coral
species for restoration. For example, more reliable
information about spawning cycles for daytime or
dawn spawners (e.g. Bronstein & Loya 2011, Eyal-
Shaham et al. 2019) would allow production effort to
be spread across species throughout the day and
night, and identification of multiple spawning events
within a population (e.g. Stobart et al. 1992) could
provide additional opportunities throughout the year
for spawn collection. Furthermore, reliably obtaining
spawned gametes from other important coral taxa
(beyond reliance on acroporid and merulinid species)
is important to retain functional diversity of restored
ecosystems.
Manipulation of endogenous rhythms and opti-
mised aquaculture conditions may underpin future
hatchery-style production of corals. Practitioners
could (1) maintain multiple populations on off-set
annual cycles to produce gametes several times per
year, (2) accelerate gametogenesis so that 1 popula-
tion can be spawned several times per year, and (3)
control the hour of spawning to spread effort
throughout the day. Research has already begun to
test the feasibility of some of these options. Firstly, by
artificially mimicking seasonal and daily insolation
cycles, lunar-irradiance cycles, and seasonal temper-
ature fluctuations, Craggs et al. (2017) replicated nat-
ural gametogenic cycles in GBR corals that were
transported and maintained for >1 yr in an aquarium
in the UK. This is a first step in maintaining synchro-
nously spawning populations in artificially controlled
environments. Secondly, maintaining optimal light
intensity to support photosynthesis and providing
additional food sources for corals ex situ may partly
overcome nutrient limitations for gametogenesis,
allowing for multiple gametogenic cycles per year
(e.g. Szmant-Froelich et al. 1980).
There may be potential trade-offs, however, be -
tween increased gamete production and other vital
processes such as somatic-tissue growth, calcification,
and immune responses (Harrison & Wallace 1990).
Furthermore, the quality and viability of gametes re-
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Mar Ecol Prog Ser 635: 203–232, 2020
sulting from nutrient enrichment may be inferior to
those produced under natural conditions, and thus
further investigation is needed prior to implementa-
tion in any restoration programme (e.g. Ward & Harri-
son 2000). Lastly, the long-term effects of out-of-sea-
son-spawned corals on future spawning synchrony
and fitness are not known and must be investigated.
3.5. Capturing spawn slicks for restoration
Synchronously spawning corals, with colony fecun-
dities reaching 106m−2 (e.g. Álvarez-Noriega et al.
2016, Howells et al. 2016b), may release billions of
eggs and sperm per hectare of reef. Under calm con-
ditions, these positively buoyant eggs can form
highly concentrated and species-rich surface slicks
(our Fig. 2a; Butler 1980, Oliver & Willis 1987), and if
the gametes are healthy, could provide a natural
source of sexually produced propagules for restora-
tion (Kawaguti 1940, Heyward et al. 1999, Omori &
Fujiwara 2004, Harrison et al. 2016).
Spawn slicks are generally pink or white in colour
and can be observed as discrete patches or narrow
strips on the sea surface extending several kilometres
from a reef (Butler 1980, Oliver & Willis 1987). The size
and location, the species composition,
the viability, and the longevity of a slick
are all closely tied both to oceanographic
and environmental conditions and to
coral composition and abundance. When
and where slicks will form, and how long
they persist, is controlled by surface
oceanographic features, such as eddies
or fronts between water bodies (Wolanski
& Hamner 1988, Jones et al. 2006), and is
de pendent on wind speed. Wind speeds
between 2 and 4 m s−1 are associated
with the highest probability of slick for-
mation (Romano & Marquet 1991, van
Woesik 2010), while slicks are unlikely to
form at wind speeds above 6−7 m s–1 (Ro-
mano 1996). Although coral spawning
often occurs during calm periods (van
Woesik 2010), weather monitoring over a
decade at selected reefs on the GBR
showed that wind speed exceeded 5 m
s−1 for ~50% of major spawning events
(Heyward & Negri 2012). Similarly, in the
Philippines, slight differences in local
turbulence from year to year have influ-
enced the extent of slick formation and
their speed of dispersal (Jamo diong et al.
2018). Therefore, local meteorological and oce ano -
graphic conditions during coral spawning have impor-
tant roles in predicting slick formation at a given reef.
The species composition and viability of the slick
are closely tied to the abundance of adult corals on
the reef from the most fecund spawning families,
such as Acroporidae and Merulinidae. The biological
and physical attributes of propagules, including
oocyte diameter, buoyancy, and developmental rate,
also vary by species and consequently affect the
propensity of each species to end up in slicks. A slick
sample from the GBR, for example, contained early-
stage larvae, in a size range mostly between 200 and
800 µm, with modal peaks around 400 and 550 µm
and a mean diameter of 484 µm (Fig. 2b,c), indicating
a diversity of species. Extrapolation from these sizes
to a calculation of packed cell volume suggests that a
dense surface slick just 1 mm thick could contain sev-
eral million eggs, embryos, or larvae per square
metre, although viability within slicks can be highly
variable (Oliver & Willis 1987). Poor viability in some
slicks may be caused by rainfall creating low-salinity
stress (Harrison et al. 1984, Richmond 1993) or anoxic
conditions in the surface layer from microbial decom-
position of spawn, caused by either low fertilisation
success or sequential days of spawning (Simpson et
208
n = 821
0
2
4
6
8
10
12
14
16
18
20
0 200 400 600 800 1000
Frequency (%)
Mean larval diameter (µm)
ab
c
Fig. 2. (a) Multi-species coral surface slick at Lizard Island, at 07:00 h, the
morning after spawning. (b) Photomicrograph of a sample from the surface
slick in (a), showing various coral embryos and larvae and other flotsam. (c)
Coral larval size−frequency distribution measured from the slick shown in
(a,b). 'n' in (c) indicates the number of larvae measured to estimate the size–
frequency distribution. Photos: A. Heyward

Randall et al.: Coral sexual production for reef restoration
al. 1993, Guillemette et al. 2018). Be cause the forma-
tion, size, and viability of slicks are influenced by the
abundance of adult colonies, the accessibility of coral
slicks for restoration may diminish in parallel with a
decline in coral abundance.
Spawn slicks could be utilised in restoration by: (1)
holding a slick above a target reef to mimic natural
retention and increase recruitment (e.g. Golbuu et al.
2012, Harrison et al. 2016); (2) collecting and trans-
porting larvae from a slick to a recruitment-limited
reef location for release (Heyward et al. 1999, 2002,
dela Cruz & Harrison 2017, Doropoulos et al. 2019);
or (3) collecting and settling larvae from a slick onto
artificial substrates for grow-out and deployment
(e.g. Chamberland et al. 2017). Efforts to collect and
deliver captive larvae back to the reef and manipu-
late larval settlement have so far utilised floating
ponds and hoses (Heyward et al. 2002, Omori et al.
2004), mesh enclosures (dela Cruz & Harrison 2017),
tents (Edwards et al. 2015), plastic bags (Suzuki et al.
2012), foam ring-seeder devices (Cooper et al. 2014),
and Perspex boxes (Quigley et al. 2018b). However,
coral restoration programmes targeting entire reefs
and reef systems would need to implement such
methods at much larger scales than have previously
been attempted (Doropoulos et al. 2019).
In early trials, harvested wild slicks produced com-
petent larvae in Western Australia (Heyward et al.
2002) and Okinawa (Omori et al. 2004). Those stud-
ies, and subsequent restoration efforts using captive-
spawned larvae in Palau (Edwards et al. 2015) and
the Philippines (dela Cruz & Harrison 2017), demon-
strated enhanced settlement rates through increased
larval supply. Nonetheless, increased settlement
rates on artificial surfaces did not elevate the num-
bers of surviving 1 yr old corals above that observed
in adjacent areas with natural recruitment in Palau
(Edwards et al. 2015). In contrast, larval enhance-
ment on 4 replicate 24 m2degraded reef plots with
low natural recruitment in the Philippines increased
recruitment rates of Acropora tenuis compared with
reference plots, and re-established a breeding popu-
lation within 3 yr (dela Cruz & Harrison 2017). These
examples demonstrate that a positive outcome from
larval restoration is possible in some situations, but
the cost-effectiveness and scalability of this approach
remain challenging.
There is a limited period for the capture of a surface
slick with concentrated and viable embryos and lar-
vae. Coral embryos are structurally fragile during the
first hours of development (Heyward & Negri 2012)
but become more robust after gastrulation. Conse-
quently, careful isolation and containment of a slick
may be feasible in the hour or 2 immediately after
spawning, prior to cleavage (which may be protracted
for multi-species slicks), but otherwise is best delayed
until the cohort is at least 24 h old (Omori & Fujiwara
2004). Subsequently, a period of days (but generally
not weeks) is available for culture and transport. Ex-
tended transit times risk larval settlement during
transport, particularly for cultures of brooded larvae,
which are usually released at an advanced larval
stage and are competent to settle quickly after re -
lease. Therefore, slick harvesting should consider lar-
val viability and concentration at or near the surface,
bearing in mind early embryo fragility and changes to
larval distribution in the water column during devel-
opment (Section 4.1). These trade-offs provide several
technical challenges for the future large-scale use of
slicks for coral restoration.
4. EMBRYOGENESIS AND
LARVAL DEVELOPMENT
Embryogenesis and larval development are among
the most sensitive processes in the coral life cycle
(Figs. 1 & 3). Understanding where and when these
processes take place, the risks and sensitivities asso-
ciated with these stages, and their environmental
drivers will allow culture conditions to be optimised
for the ex situ production of corals.
4.1. Environmental controls on embryogenesis
and larval development
Early embryogenesis (our Fig. 3f−h; first cleavage
through blastula) usually occurs overnight at the sea
surface (Babcock & Heyward 1986, Hayashibara et al.
1997, Ball et al. 2002, Okubo & Motokawa 2007,
Okubo et al. 2013, 2017). The lipid-rich embryos of
most species (Arai et al. 1993) are positively buoyant
and passively transported, while cilia develop on the
outer surface, leading to ‘rotary’ swimming. The tim-
ing of the onset of rotary swimming varies across spe-
cies but can occur as early as 12 h (e.g. Pavona decus-
sata) or as late as 3 d (e.g. Pseudosiderastrea spp.)
after fertilisation (Oku bo et al. 2013). Directional
swimming of the larvae is achieved hours to days after
rotary swimming, when they begin exhibiting photo-
tactic responses and become capable of adjusting
their positions in the water column (our Fig. 3j,k;
Lewis 1974, Harrison & Wallace 1990). The timing of
larval competency for settlement also varies across
species (reviewed in Jones et al. 2015) and can occur
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Mar Ecol Prog Ser 635: 203–232, 2020
as early as 24 h after spawning (e.g. Fungia fungites)
or can require a week or more of larval development
(e.g. Acropora austera).
Environmental factors that affect fertilisation suc-
cess and can cause embryonic abnormalities include
elevated inorganic nutrient concentrations (reviewed
in Fabricius et al. 2005, Humphrey et al. 2008, Lam et
al. 2015, Richmond et al. 2018), pCO2(Albright 2011,
Albright & Mason 2013), sediments (reviewed in
Jones et al. 2015), and pollutants (reviewed in Hud-
spith et al. 2017). Rates of embryogenesis are also
temperature-dependent; warm temperatures have
been shown to accelerate development in several
species, including acroporids and mussids (Bassim et
al. 2002, Bassim & Sammarco 2003, Negri et al. 2007,
Randall & Szmant 2009a, Heyward & Negri 2010,
Chua et al. 2013, Graham et al. 2017). Warmer ocean
temperatures can also reduce fertilisation success
(Negri et al. 2007, Albright & Mason 2013) and
increase the likelihood of embryonic abnormalities,
leading to increased mortality (Bassim et al. 2002,
Bassim & Sammarco 2003, Negri et al. 2007, Randall
& Szmant 2009a,b). In a study of Acropora palmata, a
4°C increase in temperature decreased larval sur-
vival by ~60%, with most losses occurring during the
process of gastrulation (Randall & Szmant 2009a).
Coral embryos lack the protective external mem-
brane of some other metazoans; hence, early embry-
onic stages are fragile and readily fragment into
smaller groups of cells (Heyward & Negri 2012). Dur-
ing windy and turbulent conditions, for example, early
morulae (2−16-cell stages; Fig. 3m) can fragment and
then continue cleaving, eventually developing into
proportionally smaller larval clones, sometimes one-
eighth the normal size (Heyward & Negri 2012). The
deliberate fragmentation of early embryos to generate
smaller larvae in culture may increase larval numbers
and compress the larval competency period, as smaller
larvae generally reach competency sooner (Figueiredo
et al. 2013). However, larger fragments (blastomeres)
are more likely to survive, and the resulting small re-
cruits may be disadvantaged, as the time required to
grow to size-escape thresholds may be longer (Ray-
mundo & Maypa 2004, Doropoulos et al. 2012b). Fur-
thermore, only fragments with animal hemispheres
can develop into primary polyps (Okubo et al. 2017).
4.2. Optimising embryogenesis and larval
development for restoration
In culture, direct handling of embryos should be
minimised, and mass cultures should initially have no
aeration and slow water movement for the first 18−
24 h, unless deliberately generating small embryos
(Edwards 2010, Omori & Iwao 2014, Pollock et al.
2017). Aeration and water flow can be increased
once embryos reach the gastrula stage (our Fig. 3i)
~12− 24 h post-fertilisation (Okubo et al. 2013).
Exposure to elevated-temperature conditions dur-
ing early-life histories is a potential tool for accelerat-
ing development (Nozawa & Harrison 2000, 2007,
Randall Szmant 2009a, Heyward & Negri 2010) and
enhancing the thermal tolerance of restored corals
(Putnam & Gates 2015, van Oppen et al. 2015, 2017).
Nonetheless, possible benefits of accelerated rates of
development and stress-hardening may be coun-
tered by lower fertilisation rates, higher rates of
developmental abnormalities, disease, mortality, and
downstream consequences on general culture
health. Thermal tolerance selection through stress-
hardening of coral larvae, and its role in long-term
acclimatisation, have not yet been quantified, and
thus constitute critical research priorities.
5. SETTLEMENT AND METAMORPHOSIS
The successful recruitment of corals includes both
settlement (Section 5) and post-settlement survival
(Section 6), and reef restoration would benefit from
the optimisation of both processes.
5.1. Larval sensory systems and the processes of
settlement and metamorphosis
The transition from a motile planula larva to a ses-
sile polyp (i.e. ‘settlement’) is a multi-stage process
that includes selection of a settlement substrate, fol-
lowed by attachment to that substrate, metamorpho-
sis from larva to polyp, and finally acquisition of pho-
tosymbionts (Section 7) for those species that acquire
them from the environment (Harrison & Wallace
1990). Although planktonic larvae are transported by
ocean currents, they can vertically modulate their
position in the water column (Tay et al. 2011), which
increases their likelihood of intercepting reef sub-
strate (Raimondi & Morse 2000, Szmant & Meadows
2006). Once at or near the seabed, larvae (whether
from spawning or brooding species) sense the sub-
stratum and actively search for a suitable attachment
site (reviewed in Gleason & Hofmann 2011, Jones et
al. 2015) using a range of sensory capabilities (our
Fig. 4; Young 1995). These sensory capabilities com-
prise the detection and discrimination of light fre-
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Randall et al.: Coral sexual production for reef restoration
quencies (Lewis 1974, Babcock & Mundy 1996,
Mundy & Babcock 1998, Strader et al. 2015), gravity,
hydrostatic pressure (Stake & Sammarco 2003), pos-
sibly sound (Vermeij et al. 2010), and biochemical
signals (Morse et al. 1996, Negri et al. 2001, Gleason
et al. 2009, Tebben et al. 2015) (our Fig. 4). While the
larvae of many coral species require a chemical cue
of biological origin (inducer or morphogen) to induce
settlement, this may not be a stringent requirement
for all corals, as larvae of some species appear to set-
tle spontaneously (Loya 1976, Harrison & Wallace
1990, Baird & Morse 2004). A broad range of cues
(Section 5.2) can initiate settlement. Receptors on the
ovoid larva detect the cues, and this triggers a cas-
211
a b c d
e f g
h i j k
l m n o
Fig. 3. Coral developmental stages. (a) Egg−sperm bundles setting inside mouths of polyps in Acropora loripes. (b) Intact,
packed egg−sperm bundles of Montipora digitata immediately after release. (c) Acropora longicyathus egg−sperm bundles
dissociating, releasing individual eggs and sperm from the bundle centres. (d) Unfertilised eggs of A. spathulata after 30 min
of rounding out. (e) Unfertilised eggs of M. digitata under fluorescence microscopy showing variable green-fluorescent pro-
tein signals. (f) Early cleavage of fertilised M. digitata eggs. Note the presence of endosymbiotic dinoflagellates in these verti-
cal transmitters. (g) A. tenuis 3 h after fertilisation in early cleavage. (h) Delicate ‘prawn-chip’ stage of Montipora capitata 15 h
after fertilisation. Note the presence of endosymbiotic dinoflagellates. (i) A. loripes 16 h after fertilisation, rounding out into the
gastrula stage. (j) Platygyra daedalea larvae beginning to elongate. (k) Fully developed and competent larvae of M. digitata
with dense endosymbionts. Note the fully differentiated epidermis that lacks symbionts. (l) Mycedium elephantotus chimeras
formed from the fusion of 4−8 individual embryos. (m) A. millepora recruits of various size classes resulting from fragmentation
of blastomeres of 2-, 4-, or 8-cell embryos during early cleavage. (n) A. spathulata spat (single polyps) 2 d post-settlement.
Note the absence of endosymbionts. (o) Montipora digitata coral spat settled in an aggregation, 2 d post-settlement. Note the
presence of endosymbionts. Scale bars = 1 mm. Photos in (a), (g), and (m): Andrew Negri; (c): Andrew Heyward; (b), (d−f),
(h−l), and (n,o): Carly Randall

Mar Ecol Prog Ser 635: 203–232, 2020
cade of internal biochemical and molecular signals
(Grasso et al. 2011) that result in attachment of the
aboral end, and initiation of metamorphosis into a
sessile polyp (our Fig. 3m−o; Harrison & Wallace
1990).
The time to settlement competency in larvae of
broadcast spawners is species-specific and may fol-
low a normal distribution within a cohort. Typically,
first settlement is observed around 3 d after spawn-
ing at water temperatures of 26−28°C (Connolly &
Baird 2010, Figueiredo et al. 2013, Jones et al. 2015),
with a majority competent within 4−6 d. By contrast,
brooded larvae often may settle within a few hours
after release from parent polyps, although some
planulae remain competent for months (Stephenson
1931, Harrigan 1972, Shlesinger & Loya 1985, Rich-
mond 1987, Harrison & Wallace 1990).
Metamorphosis of most species occurs within 24 h of
substrate attachment (but see Nozawa & Harrison
2000), when the larva has morphed into a disc-shaped
structure with 6 incipient mesenteries radiating out-
ward from a central mouth (Figs. 1 & 3m−o). Tentacles
usually become apparent within 48 h (our Fig. 3o;
Harrison & Wallace 1990, Heyward & Negri 1999).
While metamorphosis usually closely follows attach-
ment, the cues for attachment and metamorphosis can
be distinct and temporally uncoupled, with attach-
ment occurring days to weeks before metamorphosis
(e.g. Platygyra daedalea, Nozawa & Harrison 2000;
some Acropora species, Harrison 2006).
The hierarchy of cues, from subtle physical signals
to strong and specific chemical inducers, also differs
among species, enabling larvae to identify species-
specific settlement sites that may optimise post-
settlement survival and fitness (Morse et al. 1988, Rai-
mondi & Morse 2000, Baird et al. 2003, Baird & Morse
2004, Golbuu & Richmond 2007, Gleason & Hofmann
2011). Settlement on light-exposed upper surfaces, for
example, may favour energy acquisition via photo-
synthesis by symbionts; however, in this orientation,
the ~1 mm juvenile polyps may be more prone to pre-
dation, overgrowth by algae, and smothering by sedi-
ments (Vermeij 2006, Gleason & Hofmann 2011,
Jones et al. 2015). Attachment of larvae to dark under-
surfaces may reduce these hazards, but could cause
reduced growth from light limitation or competition
212
Fig. 4. Schematic diagram of the general physical, chemical, and biological factors (cues) that guide larvae to settlement loca-
tions and influence and trigger larval settlement on the reef, with example references for each factor. 1Babcock & Mundy
(1996), 2Mundy & Babcock (1998), 3Strader et al. (2015), 4Roberts (1997), 5Hata et al. (2017), 6Babcock & Davies (1991), 7Jones
et al. (2015), 8Vermeij et al. (2010), 9Edmunds et al. (2001), 10Randall & Szmant (2009a), 11Stake & Sammarco (2003), 12Goreau
et al. (1981), 13Doyle (1975), 14Fadlallah (1983), 15Mason et al. (2011), 16Foster & Gilmour (2016), 17Da-Anoy et al. (2017),
18Whalan et al. (2015), 19Tebben et al. (2015), 20Gleason et al. (2009), 21Morse et al. (1996), 22Negri et al. (2001)

Randall et al.: Coral sexual production for reef restoration
from encrusting taxa. A successful strategy for the
acroporids is to settle near underside edges somewhat
protected from predation, grazing, and physical dis-
lodgement, yet receiving enough light to promote
colony growth (Mizrahi et al. 2014, dela Cruz & Harri-
son 2017). Larvae of many species also favour settling
in microrefugia such as in corners, crevices, or
hollows of similar size to their diameter (Petersen et
al. 2005, Nozawa 2008, Okamoto et al. 2008, Doropou-
los et al. 2012b, 2016, Whalan et al. 2015). Older juve-
niles, however, tend to be found on exposed surfaces,
reflecting the strong influence of post-settlement mor-
tality on determining adult coral distributions (Bab-
cock & Mundy 1996, Mizrahi et al. 2014, Doropoulos
et al. 2016, dela Cruz & Harrison 2017).
5.2. Settlement cues and inhibitors
Larval settlement can be triggered by a broad
range of abiotic cues, including surface topography
(Whalan et al. 2015) and colour (Mason et al. 2011,
Foster & Gilmour 2016); however, biochemical induc-
ers produced by crustose coralline algae (CCA) (our
Fig. 3n; Morse et al. 1996, Harrington et al. 2004,
Tebben et al. 2015) and microbial biofilms (Harrigan
1972, Webster et al. 2004, Tran & Hadfeld 2011,
Sharp et al. 2015) are considered more influential in
triggering settlement in many species (Gleason &
Hofmann 2011, Jones et al. 2015). Choice ex -
periments have demonstrated clear coral−algal
specificity for some species (Morse et al. 1996, Har-
rington et al. 2004, Ritson-Williams et al. 2010, 2016,
Davies et al. 2014); e.g. Acropora spp. larvae settle
at higher rates on cryptic Titanoderma prototypum
(Harring ton et al. 2004). Less-preferred CCA species
may overgrow the juvenile coral or shed surface lay-
ers, resulting in the dislodgement of the coral (Har-
rington et al. 2004).
The specific components of CCA and biofilms that
induce settlement remain elusive. Reported chemical
inducers associated with CCA include a sulphated
glycosaminoglycan (Morse & Morse 1991), a macro-
diolide (Kitamura et al. 2009), glycoglycerolipids and
polysaccharides (Tebben et al. 2015), and mixtures of
a bromotyrosine derivative and carotenoids (Kita-
mura et al. 2007). Tetrabromopyrrole (TBP) isolated
from the biofilm bacterium Pseudoalteromonas sp.
induces rapid metamorphosis in some species, but is
not always preceded by attachment and is relatively
unstable (Tebben et al. 2011, Sneed et al. 2014). Sim-
ilarly, the neuroactive signalling peptide GLW-amide
induces metamorphosis in acroporid corals and can
be readily synthesised, but permanent attachment of
larvae is not always achieved (Iwao et al. 2002, Erwin
& Szmant 2010, Tebben et al. 2015).
A range of toxicants, including metals, pesticides,
petroleum products, and other industrial products,
have been shown to impact larval settlement (Negri
& Heyward 2000, Reichelt-Brushett & Harrison 2000,
Negri et al. 2005, Lam et al. 2015, Hudspith et al.
2017), and the physical blocking of cues can impede
settlement as well. A very thin layer of sediment, for
example, can prevent settlement on substrates with
strong cues, and when removed, a legacy impact on
the inductive capacity of the substrate may remain
(Ricardo et al. 2017). Elevated temperature and pCO2
exposure can also directly reduce settlement rates
and increase post-settlement mortality (Randall &
Szmant 2009a,b, Albright et al. 2010, Heyward &
Negri 2010, Albright & Langdon 2011, Doropoulos et
al. 2012a, but see Putnam et al. 2008), but can also
negatively impact the biota that induce coral settle-
ment, causing the broad-scale impairment of natural
recruitment (Kuffner et al. 2008, Doropoulos et al.
2012a, Webster et al. 2013, Fabricius et al. 2015,
2017).
5.3. Optimising coral settlement for restoration
Understanding the cues, and particularly the bio-
chemical inducers, that control larval settlement can
facilitate the attachment of mass-cultured larvae onto
natural or artificial substrata for restoration.
Successful settlement on any surface typically fol-
lows ‘biological conditioning’ for several weeks, for
the development of microbial biofilms and recruitment
of CCA (Harrigan 1972, Harrison & Wallace 1990,
Webster et al. 2004), or artificial induction (Guest et al.
2014, Omori & Iwao 2014, Chamberland et al. 2017).
Small (~5 mm) chips of live or dead CCA have reliably
been used to attract and trigger larval settlement onto
surfaces in experimental studies (Morse et al. 1996,
Heyward & Negri 1999) and this method could be
applied to settle cultured larvae onto most natural and
artificial surfaces for deployment in restoration.
However, scaling-up this process requires the har-
vesting or culturing of CCA, and the species used may
not actively induce settlement of all target species
(Baird & Morse 2004, Davies et al. 2014).
Applying natural biochemical inducers from CCA
and biofilms onto surfaces for reef rehabilitation has
been considered for over 2 decades (D. E. Morse et
al. 1994, A. Morse et al. 1996). This approach is
dependent on extracting the active chemicals pro-
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Mar Ecol Prog Ser 635: 203–232, 2020
duced by the CCA or biofilm and immobilising the
chemicals on a surface (D. E. Morse et al. 1994, A.
Morse et al. 1996). The application of these chemicals
to artificial surfaces for reef restoration would require
mass isolation or synthesis of complex biochemicals,
which can be unstable and costly, and none have so
far proven to have universal activity across species.
The application of natural and artificial inducers to
settle larvae en masse requires more fundamental
research, and currently, field or laboratory condition-
ing of surfaces to develop multi-species CCA and
bacterial biofilms represents a less controlled but
more feasible approach to settle a diversity of larvae
for restoration.
Site selection by competent larvae is an important
consideration for larval reseeding. Changes in settle-
ment behaviour commence in the water column
before a physical encounter between a larva and the
substrate occurs, where waterborne chemicals asso-
ciated with benthic organisms or pollutants may
attract or repel them. Because allelopathic effects can
extend well beyond the substrate boundary layer
(Birrell et al. 2005, 2008, Morrow et al. 2016), they
may drive larval selection or rejection of a settlement
site. The degree to which larvae are capable of
actively selecting among settlement sites through
vertical positioning, however, is a matter of debate
(Baird et al. 2014, Dixson et al. 2014, Hata et al.
2017), and may be limited to a small spatial scale, and
to the crawling phase after the larvae have contacted
the substrate.
Given the highly sensitive nature of larvae to phys-
ical, biological, and chemical agents, it is critical to
evaluate and mitigate potential settlement inhibitors,
particularly at a site where recruitment of larvae is a
core restoration strategy (Beyer et al. 2018). The risks
posed by settlement inhibitors at candidate restora-
tion sites can be quantified by comparing in situ
water quality measurements with thresholds derived
from laboratory assays. In addition, there is mounting
evidence that stressors and inhibitors can interact
cumulatively to reduce coral settlement (Humanes et
al. 2017) and thus should be considered in the selec-
tion and management of sites for restoration.
5.4. Artificial substrates for use in reef restoration
The artificial substrate most suitable for reef resto-
ration depends on the goal and scale of the project
(Spieler et al. 2001, Chamberland et al. 2015, 2017,
Barton et al. 2015), and practitioners should consider
the effectiveness, cost, environmental impact and the
feasibility of use in large-scale applications. Almost
any solid material can be used to settle coral larvae;
however, materials differ in their composition and
succession of colonising communities, resulting in dif-
ferences in their abilities to attract and induce meta-
morphosis, and to reduce competition with other ben-
thic organisms. For example, ceramic tiles are widely
used for assessing settlement in situ, in part because
they provide colonising CCA with a competitive
advantage over other encrusting and fouling organ-
isms (Harriott & Fisk 1987). As settlement cues are
often species-specific, the settlement surface and sub-
sequent successional biofouling community likely
bias the settling-coral community.
The shape and topography of artificial substrates
can be designed and manufactured to produce an
optimal settlement habitat. Larvae are drawn to small
crevices and areas free of sediments and grazers (our
Section 5.1; Babcock & Davies 1991, Doropoulos et al.
2016, Ricardo et al. 2017). Several organisations are
developing ‘seeding units’ designed to maximise re -
cruitment success by incorporating natural micro -
refugia (e.g. Chamberland et al. 2017). Past restora-
tion projects have improved recruit survival by using
grates, meshes, or poles that provide a range of
angles for settlement, while also allowing sediments
to pass through (Omori et al. 2006, Suzuki et al. 2011,
Higa & Omori 2014, Ng et al. 2016). Three-dimen-
sional printing technology is increasingly applied to
help replicate the complex shapes of naturally occur-
ring reefs (Mohammed 2016). Not all materials are
easily shaped, however, and malleability and ease of
manufacture of complex surfaces to improve restora-
tion success need to be carefully considered.
Ease of deployment and suitability for the receiv-
ing environment are also important considerations.
Substrates that need to be manually and individually
attached to the seafloor (e.g. small recruitment sub-
strates) are likely to be the most expensive and least
practical to up-scale. By contrast, substrates that can
be deployed from the surface with less effort, such as
tetrapods designed to maximise the chance of wedg-
ing into the reef structure and increase retention
(Chamberland et al. 2017), could be more economical
at scale, noting that ideal substrates may vary by
environment. Finally, it is imperative to consider the
potential long-term environmental and ecological
impacts of the selected material before introduction
into the environment, where it may act as, or gener-
ate, pollution or marine debris. Some plastics have
been associated with coral diseases (Lamb et al.
2018), and microplastics generated from the break-
down of plastics in the marine environment impact
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Randall et al.: Coral sexual production for reef restoration
many organisms, including corals (Hall et al. 2015,
Reichert et al. 2018).
6. RECRUITMENT AND POST-SETTLEMENT
SURVIVAL
Early post-settlement survival is a primary bottle-
neck and challenge for coral restoration. Coral spat
are particularly susceptible to predation, competi-
tion, and stochastic disturbances, and need to grow
quickly to escape this vulnerable phase. Most newly
settled corals are also aposymbiotic and must estab-
lish symbioses with a suite of partners to survive.
Reducing predation and competition for young re -
stored corals, both directly and indirectly, and pro-
viding recruits with targeted symbiotic partners, rep-
resent opportunities to overcome this bottleneck.
6.1. Density-dependent processes in recruitment
and post-settlement survival
Settlement success can increase with larval density
(Heyward et al. 2002, Suzuki et al. 2012, Doropoulos
et al. 2018), but dense cultures may be suboptimal for
larval health (Guest et al. 2010, Pollock et al. 2017) or
carry downstream risks. Higher larval densities can
increase gregarious settlement (our Fig. 3m, o) up to
a density-dependent parabolic threshold (Suzuki et
al. 2012, Doropoulos et al. 2017, 2018), which can
accelerate growth and improve survival to a point
(Suzuki et al. 2012). The availability of substrate with
appropriate settlement cues can also interact with
density-dependent biological processes; clustered
patterns in the benthic community can encourage the
clustering of spat around a settlement cue. On de -
graded reefs, a reduction in quality substrate may fur-
ther encourage clustered settlement around rare cues
and promote density-dependent recruitment bottle -
necks (Vermeij & Sandin 2008, Albright et al. 2010,
dela Cruz & Harrison 2017, Fabricius et al. 2017).
Mortality in the first year after settlement can be
extremely high (>30−99%) (Loya 1976, Babcock
1985, Babcock & Mundy 1996, Wilson & Harrison
2005, Davies et al. 2013, Suzuki et al. 2018). Stochas-
tic processes like accidental grazing and storms con-
tribute significantly to juvenile mortality (Mumby
1999, Davies et al. 2013, Trapon et al. 2013), and the
drivers of mortality shift with life stage and size. For
example, small juveniles 1−2 mo old are more sus-
ceptible to accidental grazing (Trapon et al. 2013)
compared with larger 10−14 mo old juveniles (Davies
et al. 2013). Generally, mortality pressures continue
to act on coral juveniles until they reach a size-
escape threshold, or size refuge, when mortality sig-
nificantly declines (Babcock & Mundy 1996, Doro -
poulos et al. 2012b, dela Cruz & Harrison 2017). This
size refuge may differ across species but can be
around 5 mm in diameter or 3−9 mo old (Babcock &
Mundy 1996, Doropoulos et al. 2012b). Furthermore,
juvenile mortality after settlement can vary in a
density-dependent manner. Dense settlement, for
example, may lead to higher predation through pred-
ator attraction (Gallagher & Doropoulos 2017), or
alternatively could lead to decreased predation by
reaching size-escape thresholds more quickly (Ray-
mundo & Maypa 2004, Doropoulos et al. 2012b).
6.2. Maximising post-settlement survival
in reef restoration
A variety of methods have been proposed to either
reduce predation on, or competition with, spat to
overcome post-settlement survival bottlenecks for
coral restoration. Firstly, substrates have been engi-
neered with multiple surface orientations and micro-
topography, such as corners, crevices, holes, and div-
ots, to offer a refuge from accidental grazing and to
limit sedimentation impacts (Nozawa 2008, Cham-
berland et al. 2017). Materials with embedded an-
tifouling compounds that target and prevent direct
competition from other benthic organisms are being
explored (Tebben et al. 2014), as are materials that
have unpalatable embedded compounds that aim to
reduce grazing pressure and deter corallivores.
Secondly, co-culturing techniques have been tri-
alled, whereby coral recruits or fragments are
reared alongside grazers, in a multi-trophic aqua-
culture system, and then deployed together. For
example, co-culturing of coral spat with herbivorous
gastropods such as Trochus spp. and Clypeomorus
spp. snails (Omori 2005, Villanueva et al. 2013,
Omori & Iwao 2014, Toh et al. 2016) and echino-
derms (Toh et al. 2016, Craggs et al. 2019) has been
found to increase spat survival. Thus, herbivorous
gastropods, echinoderms, and fishes, which already
are routinely used to control algae in large-scale
coral nurseries, offer opportunities for enhanced
production efficiency. Furthermore, Spadaro (2014)
reported preliminary success in deploying large
herbivorous brachyuran crabs with high site fidelity
at coral restoration locations in the Caribbean, indi-
cating that in situ co-culturing also may be benefi-
cial at restoration sites.
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Mar Ecol Prog Ser 635: 203–232, 2020
216
6.3. Size matters: improving restoration outcomes
with chimeras
Coral larvae tend to settle gregariously (our Fig.
3m,n) and occasionally form natural chimeras (Amar
et al. 2008, dela Cruz & Harrison 2017), which can
rapidly increase the size of the juvenile by an order
of magnitude, and accelerate growth to the size-
escape threshold (Raymundo & Maypa 2004,
Doropoulos et al. 2012b, 2018, Suzuki et al. 2012).
Chimeras de velop from the fusion of genetically dis-
tinct coral embryos (our Fig. 3l; Jiang et al. 2015),
juveniles (our Fig. 3m,n; e.g. Raymundo & Maypa
2004, Amar et al. 2008) or adults (Puill-Stephan et
al. 2009), and may confer several ecological advan-
tages such as faster growth, earlier sexual matura-
tion, and increased competitive ability (Rinkevich &
Weissman 1987, 1992, Puill-Stephan et al. 2009,
Rinkevich et al. 2016). The fusion of Pocillopora
damicornis larvae resulted in higher growth and
survival of juveniles (Raymundo & Maypa 2004),
and chimeras produced from brooded Stylophora
pistillata larvae exhibited in creased survival com-
pared with individual juveniles (Amar et al. 2008).
Yet, the ability of corals to discriminate self from
non-self (allorecognition) and to tolerate chimeric
colonies varies among species. Allorecognition in
corals is driven by mechanisms of genetic histocom-
patibility (Heyward & Collins 1985, Heyward &
Stoddart 1985, Stoddart et al. 1985), which develop
as juveniles mature (Hidaka 1985, Hidaka et al.
1997). Chimerical full-sibling juveniles tend to
exhibit the highest rates of stable post-settlement
survival compared with half and non-sibling juve-
niles (Nozawa & Loya 2005, Puill-Stephan et al.
2012b), and chimera formation at the larval or early
recruit stage, before the maturation of immune
recognition mechanisms, can in crease fusion suc-
cess (Hidaka 1985, Hidaka et al. 1997, Wilson
& Grosberg 2004, Puill-Stephan et al. 2012a,b,
Schweinsberg et al. 2015). Thus, forming chimeras
from related individuals early in ontogeny, during
the embryo and larval stages, is likely to have the
best outcome. Whether coral chimera formation can
be controlled to yield a net increase in surviving
recruits remains an area of current research (Cooper
et al. 2014, Barton et al. 2015, Forsman et al. 2015).
7. CORAL SYMBIOSES
Central to a healthy coral are relationships with
associated microorganisms, which have profound
implications for coral health, stress tolerance, and
acclimation response (Mieog et al. 2009). Mutually
beneficial symbioses with photosynthetic dinoflagel-
lates (Symbiodiniaceae, sensu LaJeunesse et al.
2018) and enduring partnerships with an array of
bacterial, archaeal, fungal, protistan, and viral asso-
ciates together form the coral holobiont (Bourne et al.
2016). Understanding when and how coral symbioses
are established, and the capacity for these relation-
ships to be manipulated and maintained is an active
area of research in coral restoration.
7.1. Establishment, maintenance, and specificity of
symbiotic partnerships
The most well studied coral symbionts are dinofla-
gellates of the family Symbiodiniaceae with densi-
ties >106cells cm−2 providing up to 90 % of the
coral’s nutritional requirements through transloca-
tion of photosynthates and other essential nutrients
(Muscatine & Porter 1977). The amount of carbon
and nitrogen translocated to the coral host, however,
can vary substantially among species and Symbio-
diniaceae clades (Yellowlees et al. 2008, Davy et al.
2012, Tremblay et al. 2014) and under different
environmental conditions (Little et al. 2004, Rey -
nolds et al. 2008, Cantin et al. 2009, Hume et al.
2015). Bacterial and archaeal symbionts also occur
at densities as high as 106cells cm−2 (Garren &
Azam 2012), with the diversity of these prokaryotic
communities often ex ceeding thousands of distinct
taxa (Sunagawa et al. 2010, Blackall et al. 2015).
While these microbes cycle essential nutrients (car-
bon, nitrogen, sulphur, and phosphate) and provide
trace metals, vitamins, and other cofactors (Bourne
et al. 2016), examples of specific prokaryotic sym-
bionts being unequivocally assigned functional roles
are rare.
Early molecular studies indicated that some coral
species form exclusive partnerships with a single
Symbiodiniaceae type, whereas other species are
known to associate with multiple types that vary in
their relative abundance over time and space (Little
et al. 2004, Abrego et al. 2009a,b, Fabina et al.
2012, Byler et al. 2013, Boulotte et al. 2016, Poland
& Coff roth 2017). These symbionts can be shared
via paren tal gametes (i.e. vertical transmission),
assimilated solely from the surrounding environ-
ment (i.e. horizontal transmission), or acquired
through a mixture of both strategies (Byler et al.
2013, Quigley et al. 2018a). Vertical transmission of
a locally adapted Symbiodiniaceae community may

Randall et al.: Coral sexual production for reef restoration
benefit species with localised dispersal (Underwood
et al. 2007, Sher man 2008, Noreen et al. 2009,
Warner et al. 2016), but could also be disadvanta-
geous if environmental conditions change or if lar-
vae are dispersed to novel environments. Alterna-
tively, a horizontally acquired community may offer
more flexibility under variable environmental con-
ditions (Abrego et al. 2012, Boulotte et al. 2016,
Quigley et al. 2017a, 2018a). Genetic analyses of
relatedness and preferential selection experiments,
however, have demonstrated that for both transmis-
sion modes, Symbiodiniaceae communities are, at
least in part, regulated by their host and are not
random (Yamashita et al. 2014, Quigley et al.
2017a, 2018a).
For most coral species, the exact point at which a
Symbiodiniaceae symbiosis is initiated, and the envi-
ronmental origins of horizontally transmitted sym-
bionts, remain unclear. Symbiodiniaceae, which can
also be free-living, have been recovered from sea -
water and substrates of coral reefs, but their diversity
and availability for host uptake is not well known
(Adams et al. 2009, Cumbo et al. 2013, Quigley et al.
2017b). Larvae and juveniles of some Acropora spe-
cies can establish symbiosis with Symbiodiniaceae
shortly after exposure to local sediments (Adams et
al. 2009, Cumbo et al. 2013). Yet, the diversity of
Symbiodiniaceae residing within sediments is much
greater than the diversity in corals, indicating that
juvenile corals selectively uptake Symbiodiniaceae
and/or certain types and sizes have higher infectivity
(Yamashita et al. 2013, 2014, 2018, Biquand et al.
2017, Quigley et al. 2017b).
How the Symbiodiniaceae community changes
during juvenile development is also poorly under-
stood. For example, the brooding coral Porites astreo -
ides showed substantial variation in the Symbiodini-
aceae community throughout ontogeny and under
different environmental conditions (Reich et al.
2017). Yet, parental effects (e.g. the influence of the
maternal environment during gamete development)
also explain a significant amount of the variation in
symbiont composition in the spawning species Acro-
pora tenuis (Quigley et al. 2016). Thus, it re mains
largely unknown how flexible or deterministic the
host−symbiont community is for the vast majority of
coral species.
The mode of transmission of prokaryotic symbi -
onts, including bacterial, archaeal, fungal, protistan,
and viral associates, constitutes a critical re search
priority. There is some indication that brooding spe-
cies transmit at least some of these sym bi onts verti-
cally, whereas species that rely on external fertilisa-
tion acquire many of these symbionts horizontally
(Apprill et al. 2009, Sharp et al. 2012). However,
spawning corals can vertically transmit some compo-
nents of the microbiome via mucus contact during
spawning (Leite et al. 2017), and coral-associated
bacteria can exhibit chemotaxis towards chemicals
released from the coral, which likely influences the
establishment and maintenance of species-specific
host−microbe interactions (Tout et al. 2015).
7.2. Environmental control over symbiotic
partnerships
Symbiotic partnerships between corals and micro-
organisms are sensitive to environmental perturba-
tions, and symbiotic dysbiosis (i.e. imbalance) can
represent a significant challenge to coral survival.
The ability of some species to establish flexible part-
nerships with tolerant symbionts (Symbiodiniaceae
or bacteria), however, could confer a plasticity that
underpins transgenerational acclimatisation (Web-
ster & Reusch 2017). For instance, juvenile Acropora
species can increase the relative abundance of toler-
ant Symbiodiniaceae under higher seawater temper-
atures (Abrego et al. 2012, Yorifuji et al. 2017), while
some adult corals can acquire increased thermal tol-
erance by changing their dominant Symbiodiniaceae
type (Berkelmans & van Oppen 2006). At higher sea-
water temperatures, the larvae of some species can
establish symbiosis with novel and thermally tolerant
symbionts, while reducing their association with
heat-sensitive Symbiodiniaceae (Cumbo et al. 2018).
Furthermore, corals containing thermally tolerant
algal symbionts are much more abundant on reefs
that have been severely affected by recent climate
change (Baker 2003, Baker et al. 2004), and Symbio-
diniaceae associations can vary by host depth and
light conditions (Iglesias-Prieto et al. 2004, Bongaerts
et al. 2013, Nitschke et al. 2018).
The environmental sensitivity of coral-associated
bacterial communities is also well established, with
a shift towards opportunistic microorganisms and
potential pathogens under ocean warming and ocean
acidification (reviewed in Bourne et al. 2016). This
sensitivity has been primarily documented with adult
corals, with little understanding of microbial re -
sponses in early life-history stages. Furthermore,
while the loss of symbionts has been correlated with
declining host health, there are few examples of
favourable symbiotic bacterial shifts that enhance
growth or confer a competitive advantage under
environmental change.
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Mar Ecol Prog Ser 635: 203–232, 2020
7.3. Controlling symbiont communities in
restored corals
Although there is some support for symbiont
switching in adult corals (Boulotte et al. 2016, Quigley
et al. 2019), early juveniles may be the opportune age
to initiate infection with both environmentally tolerant
Symbiodiniaceae communities, and beneficial-bacte-
rial partners. This opportunity window precedes the
development of an immune response during the juve-
nile stage and varies across coral taxa, but may take a
few months to years (Little et al. 2004, Abrego et al.
2009b, Puill-Stephan et al. 2012a). Administering a
‘probiotic’ treatment, which incorporates a community
of beneficial symbionts, could allow researchers to
control the early ‘infection’ of coral settlers with select
communities, thereby increasing resilience and opti-
mising growth and fitness. As a first step, Damjanovic
et al. (2017) demonstrated that a single microbial in-
oculation of adult corals drove a shift in the micro-
biome. Similarly, Rosado et al. (2018) showed that a
consortium of native and putatively beneficial micro-
organisms administered as a probiotic to adult corals
was able to partially mitigate coral bleaching in a lab-
oratory thermal-stress experiment. Defining the onto-
genetic variability in the microbiome and the species-
specific life-history window when corals have
considerable flexibility to associate with a diverse
range of environmentally acquired and potentially
stress-tolerant symbionts prior to community speciali-
sation should be a future research priority.
8. AGE AND SIZE AT SEXUAL MATURATION
It can take 3 to 8 yr or more for a coral to reach sex-
ual maturation (Kojis & Quinn 1981, Babcock 1991,
dela Cruz & Harrison 2017). Reducing this duration
to promote early maturation has many potential res-
toration applications. Yet, the factors that govern the
onset of sexual maturation in corals, and how pheno-
typically plastic that onset may be, are largely un -
known.
8.1. Is sexual maturation determined
by age or size?
For most colonial organisms, including scleractini-
ans, a critical-size threshold must be reached to
trigger the onset of sexual maturation, beyond which,
fecundity increases with size until a colony is fully re-
productive (Connell 1973, Loya 1976, Kojis & Quinn
1981, 1985, Harriott 1983, Szmant-Froelich 1985, Wal-
lace 1985, Babcock 1991, Soong & Lang 1992, Hall &
Hughes 1996). Only some corals in a population ma-
ture at the size threshold (Szmant-Froelich 1985, Bab-
cock 1991, dela Cruz & Harrison 2017), however, and
estimates of that size have only been described for a
handful of species, ranging widely from 2.3 cm2in the
brooder Favia fragum to 1600 cm2in the spawner
Acropora palmata (Soong & Lang 1992). Soong (1993)
identified a significant positive correlation between
size at first reproduction and maximum colony size for
7 brooding and spawning Caribbean species, sug-
gesting that larger species generally mature later and
at a larger size than smaller ones (MacArthur &
Wilson 1967, Szmant-Froelich 1985), although this
has yet to be investigated for most species. The onset
of sexual maturation is difficult to predict, as it is likely
to be influenced by spatio-temporal variation in bio-
logical condition (Harvell & Grosberg 1988) and by
variation in coral growth rates, which are in turn influ-
enced by many factors, including competition (Rinke-
vich & Loya 1985, Tanner 1995), light availability
(Huston 1985, Lough & Barnes 2000), nutrient profiles
and concentrations (Tomascik & Sander 1985, Fabri-
cius 2005), symbiont communities (Jones & Berkel-
mans 2010), and host genetics (Drury et al. 2017).
Consequently, variations in these factors all have the
potential to affect the timing of sexual maturation.
Fragmentation and partial-mortality events have
the potential to reduce a colony’s size below the criti-
cal threshold for reproduction. In these cases, the
colony becomes small, while the polyps remain ‘old’,
leading Connell (1973) to pose 2 questions: (1) Is sex-
ual maturation in corals determined by polyp age or
colony size?; and (2) Will a reduction in colony size re-
sult in a regression to a juvenile state? Kojis & Quinn
(1981, 1985) experimentally fragmented ma ture adult
Goniastrea colonies into various size classes and
found that sexual reproduction ceased when the total
number of polyps was reduced below 30, even after
2 yr of recovery. However, when they compared re-
productive colonies of the same size but different
ages, they found that older colonies were more fe-
cund, indicating that age also influences re productive
output. Similarly, Szmant-Froelich (1985) found that
fragmenting adult Orbicella annularis colo nies below
200 cm2in surface area reduced or prevented gonad
development. Subsequent fragmentation studies of
spawning and brooding species found that both
colony size and polyp age can influence fecundity
(Ward 1995, Smith & Hughes 1999, Za kai et al. 2000,
Okubo et al. 2007, Kai & Sakai 2008), and led Graham
& van Woesik (2013) to suggest that the reproductive
218

Randall et al.: Coral sexual production for reef restoration
response to fragmentation below the critical-size
threshold may be binary and species-specific, where
some species maintain gametogenesis while others
regress to an immature state. Research over 30 yr has
identified at least 7 considerations when predicting
whether fragmented corals will reproduce: (1) size of
the fragment; (2) age of the fragment; (3) shape of the
fragment and location of polyps; (4) mode of polyp
budding; (5) timing of fragmentation; (6) duration of
fragment isolation; and (7) species (Table 1).
219
Table 1. Key factors to consider when fragmenting corals for restoration, to maintain the greatest possible reproductive output
Key
consideration
Description Restoration optimisation References
Size of the
fragment
If a coral is fragmented below the mini-
mum-size threshold, the likelihood of
regression to a prepubescent state is high
and may result in resorption of oocytes
Maintain fragments well above the
minimum-size threshold (mea-
sured by excluding infertile
margins). If not known, select
colonies >25 cm in diameter, as
most studies to date indicate
colonies of this size should be
sexually mature
Kojis & Quinn (1985),
Szmant-Froelich (1985),
Lirman (2000),
Zakai et al. (2000),
Okubo et al. (2007)
Age of the
colony
There may be a minimum age at which a
colony can reach sexual maturation.
When colonies of the same size but
different ages are compared, the older
colony may be more fecund
If known, select colonies well
above the minimum age of sexual
maturation to maximise spawning
output. If not known, select
colonies >25 cm in diameter, as
most studies to date indicate
colonies of this size should be
sexually mature
Kojis & Quinn (1985)
Shape of the
fragment and
location of
polyps
Colonies exhibit considerable spatial
variability in polyp fecundity. Colony
margins and axial polyps are often
infertile, and downward-facing tissue on
branches and vertical surfaces on mound-
ing corals have lower fecundity
Optimise reproductive output by
collecting centrally located
fragments and fragments with
upper-facing surfaces. Limit the
creation of new colony margins
Wallace (1985),
Soong & Lang (1992),
Van Veghel & Bak
(1994),
Nozawa & Lin (2014)
Mode of polyp
budding
Interior polyps newly produced by
intratentacular budding were fully fecund
in Favia fragum, whereas polyps newly
produced by extratentacular budding in
Pseudodiploria spp. were infertile
Anticipate potentially higher
reproductive output from frag-
ments of species that bud intraten-
tacularly
Soong & Lang (1992)
Timing of
fragmentation
Timing within the gametogenic cycle can
influence fecundity post-fragmentation.
Fragmentation during early vitellogenesis
(i.e. yolk development) has been shown to
result in resorption of oocytes
Fragment late in the gametogenic
cycle but limit fragmentation in the
days immediately prior to spawn-
ing to avoid stress-spawning
Okubo et al. (2007)
Duration of
fragment
isolation
Some species may exhibit ‘reproduction
adaptation’, whereby polyps on fragments
that have been isolated for several years
(e.g. on individual branches) may become
fecund even if the fragment remains
below the critical-size threshold
If collecting fragments near the
minimum-size threshold, select
those that appear to have been
isolated for a long duration, with
healed and growing margins
Szmant-Froelich (1985)
Species Some species regress to an immature
state when fragmented below the mini-
mum-size threshold (i.e. Favia chinensis)
and others remain reproductively active
(e.g. Goniastrea aspera and Pseudodiplo-
ria strigosa)
If known, select colonies that
remain reproductively active post-
fragmentation. When fragmenting
species that regress, create
fragments above the critical-size
threshold
Kai & Sakai (2008),
Graham & van Woesik
(2013)

Mar Ecol Prog Ser 635: 203–232, 2020
8.2. Is it possible to artificially accelerate
sexual maturation?
Accelerating sexual maturation through the
growth and isogenic fusion of microfragments (sensu
Page et al. 2018) in a process called ‘re-skinning’
(Forsman et al. 2015) has the potential to reduce gen-
eration times and consequently increase coral-recov-
ery rates and restoration outputs. Preliminary results
from fused microfragments of Orbicella faveolata
(Forsman et al. 2015) indicate that sexual maturation
may be artificially accelerated by promoting the
fusion and growth of the colonies to the critical-size
threshold (Page et al. 2018). More research is
needed, however, to evaluate the suitability of this
technique for other species and growth morpholo-
gies. For example, in some massive corals, the fecun-
dity of individual polyps will be influenced not only
by colony size (area) but also by tissue depth, which
may be limited in re-skinned corals.
9. FEASIBILITY OF LARGE-SCALE
RESTORATION
Restoring degraded reefs with sexually propagated
corals is being considered or applied on reefs around
the world at various spatial scales and with numerous
approaches. Comprehensive desktop analyses com-
paring a multitude of approaches are currently
underway to examine the benefits, risks, and feasi-
bility of both influencing the long-term health and
survival of corals, and facilitating acclimatisation and
adaptation of coral populations at scale (The National
Academies of Sciences, Engineering, and Medicine
2019; www. gbrrestoration. org). In Australia, a range
of scalable methods are being considered that rely on
the sexual propagation of corals, either through the
capture and redirection of wild spawn, or from land-
and/or sea-based aquaculture production facilities.
Preliminary analyses indicate that there are common
bottlenecks to achieve cost and scale across all
approaches, relating to key biological and ecological
attributes of coral reproduction. For example, high in
situ mortality post-settlement means (1) nursery-
reared corals will need to be kept for weeks or
months to pass a size-escape threshold, (2) the pro-
duction numbers required to achieve scale will need
to be very large, and (3) the estimated cost per
deployed coral surviving to adulthood will be inflated
(Omori 2019). The extent to which early growth and
survival of corals can be enhanced will materially
affect the cost and scale at which sexually produced
corals can be feasibly used. Analyses thus far have
also clarified that, while extensive effort needs to be
applied to technological developments in coral sex-
ual propagation, a significant amount of attention
also needs to be directed toward investigating funda-
mental aspects of ecological benefits and risks, such
as quantifying trade-offs in growth and thermal toler-
ance, and identifying the down-stream effects of
spawning corals out of season. This research is
required to adequately assess the feasibility of the
proposed methods.
10. CONCLUSIONS
The success of coral restoration and adaptation
interventions will vary among species, sites, reefs,
and regions. Incorporating reef-building species that
occupy a variety of ecological niches, encompass a
range of reproductive modes, include a suite of mor-
phological or functional forms, and include a diver-
sity of species, will be required to facilitate and max-
imise recovery rates, with the end goal of restoring
and maintaining ecosystem functions.
Based on current knowledge, we suggest that a
combination of restoration approaches will deliver
the greatest benefit, and that restoration interven-
tions should (1) incorporate sexually reproduced
corals across a variety of life-history stages, (2) con-
sider the ecology of the target restoration site (Ladd
et al. 2018) and implement deployment methods con-
sistent with local ecological processes, (3) incorpo-
rate pre-adapted or stress-hardened coral stock, or
incorporate methods that support natural processes
of adaptation, (4) protect key ecological functions, (5)
conserve sociocultural values, such as those held by
Traditional Owners and a diverse range of stakehold-
ers, and (6) be logistically feasible and at a cost point
for deployment at large scale. Whether it is feasible
to achieve these targets through scaled-up coral res-
toration remains an active area of investigation
(Guest et al. 2014) that will require well-developed
and integrated research and development plans (our
Table 2).
Overcoming the research challenges outlined here
will require an unprecedented level of collaboration
across nations, research groups, and public and pri-
vate sectors. To address fundamental research ques-
tions and achieve long-term success, researchers and
practitioners working in reef restoration must also
draw on expertise from vastly different scientific dis-
ciplines such as microbiology, genetics, restoration
ecology, aquaculture, materials science, and engi-
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Randall et al.: Coral sexual production for reef restoration 221
• Lar ge exper iment al system s with fi ne-sc ale cont rol over ex situ
enviro nmenta l condi tions, w ith and wi thout of f-set se asona l and
diel cycles
• Histologic al samp ling to ide ntif y the timin g of game togene sis in
the fiel d
• Fiel d monitor ing of key fun ction al group s (e.g. mass ive Porites)
to more accura tely identif y in situ spawning patterns
• Development of an integrated on-line system for collecting and
synthesizing regional or global spawning observations
• Field monito ring for t he pres ence or a bsenc e of slick fo rmati on,
and coincident collection of environmental data
• Remotely s ensed d etecti on of spawn s licks
• Field samplin g for hist ologic al and ge netic id entif icatio n of
speci es comp ositio n and dive rsity w ithin sl icks, an d embr yo
viability
• Fi eld expe riment ation to c ontai n, colle ct and tr anspor t spawn
slicks
• Field an d lab expe rimen ts and mod ellin g to under stand a nd
quanti fy pro cesse s (inclu ding sp erm den sity an d gamete q ualit y)
affec ting cor al fer tiliza tion suc cess
• L aborat ory-b ased bi ochemi cal and l arva l-beha viour as says to
identi fy inducer s and inhibitor s for a wider rang e of coral taxa , and
define r elevant e ffec tive dose s
• Evalu ation of t he micro habit at and env ironme ntal fa ctors,
including influence of biofilm succession, and conditioning timing
and dur ation on s ettle ment and p ost-settlem ent sur vival in the lab
and fie ld, for a wid er rang e of spec ies
• Laborator y and fi eld expe rimen ts to build b asic kn owledg e
around h ow (hori zontal ly, vertic ally or d ual mode t ransm issio n)
and whe n symbio sis gets e stabl ished
• Genomic t echniq ues to dete rmine u ptake and o ntogen etic
variab ility i n the micr obiome f rom emb ryo to juve nile
• Laboratory inoculation experiments followed by out-planting
and time-series sampling to evaluate the feasibility of establishing
and maintaining select symbiotic partners
• Lab orator y exper iment s to evaluate t he impac t of stre ss
harde ning on em bryos a nd lar vae
• Development of ecologically safe and effective juvenile
deploy ment sub strate s and ef ficien t method s to deplo y
• Laboratory choice and no-choice settlement experiments with a
suite of sh apes an d substr ate typ es to iden tify op timal su bstrat es
for post-settlement survival
• Field exp erime nts to tra ck post- deploy ment sur vival a nd
compare substrate performance across reefs, reef zones and
speci es and un der dif ferent l evels of co mpetit ion and p redati on
• Fie ld exper iments t o investig ate dens ity- depend ent ef fects of
settlement on survival
• Laboratory exper iements to tes t the effect of nut rition and
probio tic trea tments o n growt h and sur vival, a nd evalua te a suite
of delive ry met hods
• Lab orator y and fi eld expe riment s with adu lt, fra gmente d,
microfragmented, and isogenically fused coral colonies to identify
the facto rs governing th e onset of sexual m aturati on for a diversit y
of species
• L abor atory ex perim ents to de termin e the facto rs that p romote
early m aturati on and id entif y what phy siolog ical/e cologi cal
trade -off s are ass ociate d with ear ly matur ation ( if any)
• Tempora l field s amplin g with his tologi cal ana lyses to i dentif y
minimum size-thresholds for sexual maturation for a diversity of
species
• Labor atory e xperim ents to de termin e the eff ects of fu sion an d
fissi on of colo nies on r eprod uctive o utput, i nclud ing the ef fects of
timing , fragme nt size, ag e, etc.
• Development of automated high-throughput aquaculture
production
• Time se ries ge nomics s ampli ng and an alysis o f batch cul tures
throughout embryogenesis and larval development (culture
genetics) to identify the most successful genotypes and assess
the risk o f ‘lab- adapted ’ coral s on poten tial per formance on re efs
• Ability to predict more accurately which species will spawn,
where and when
• Aquaculture systems that enable manipulation of seasonal and
lunar cycles to synchronously spawn corals ‘out-of-season’
• Ability to adjust diel timing of coral spawning to increase culture
efficiencies
• Better understanding of the molecular mechanisms and
environmental factors underpinning function of endogenous
reproductive rhythms in corals
• Knowledge to assess the risk of spawning asynchrony in
restored corals
• Predict ive mode ls to est imate wh en and wh ere spaw n slick s will
form
• Ability to target slicks to maximize the diversity of species
harve sted for r estora tion
• Abi lity to ef fecti vely har vest an d trans port s pawn sli cks for re ef
restoration
• Know ledge o f stockin g densi ties th at are ade quate to en sure
self-sustaining fertilization and larval production
• Identi ficat ion of cue s to induc e and impr ove sett lement o f a
divers ity of co ral ta xa
• Set of ‘b est-pr actic es’ for set tling a di versi ty of spe cies (e.g.
culture conditions, densities, durations, flow rates, substrate
conditioning protocols)
• Abili ty to characte rise and ident ify optimal ca ndidate restor ation
sites
• Ability to induce settlement at specifically targeted locations or
optimal ‘receiving sites’
• Ef fective u ptake an d transm issio n method s to maxi mize sym-
biosi s succes s and long-term m ainten ance
• Identific ation of sy mbioti c partn ers tha t may enha nce cor al
perfo rmanc e in a warmi ng clima te
• Stres s-hard ening m ethods to i mprove th ermal to leran ce of
embryos and larvae
• Sp ecies -spec ific ab ility to m aximi ze post-d eploym ent sur vival
and growth
• Ec ologic ally ef fectiv e and cost- effi cient s ubstra ta for out plan-
ting corals
• Nutriti on and pro biotic p rofile s, conc entra tions, a nd deliv ery
method s that opt imize grow th and post-set tleme nt sur vival
• More accurate models of population growth and response to
restor ation fo r a divers ity of sp ecies
• Ability to sh orten g enera tion tim es by indu cing ea rly matu ration
in selected strains of coral or young, isogenically fused corals
• Ability to o ptimis e fragm entati on and oth er asexu al prop agatio n
method s to maxi mise rep roduc tive outp ut
• Ability to p ropag ate cora ls in lar ge quant ities an d at low cos t
• Esti mates of th e risk of r earing ‘ labor atory- adapte d’ coral s that
may resu lt in mala dapted g enoty pes that h ave impai red
performance on reefs
Describing and controlling gametogenic
and spawning cycles
Characterization of spawn slicks
Identification of larval settlement cues
and inhibitors
Establishing symbioses
Overcoming the post-settlement
survival bottleneck
Identifying the factors that control
the onset of sexual maturity
Age and size at sexual maturation
Coral aquaculture research
and development
Areas of research that target
key knowledge gaps
Approaches to address knowledge gaps Expected outcomes
Table 2. High-priority knowledge gaps identified in the review and corresponding suggested approaches. Expected outcomes of the
research for each section of the review are presented based on current state of knowledge

Mar Ecol Prog Ser 635: 203–232, 2020
neering. Key knowledge gaps in the science of coral
sexual reproduction are reported in Table 2 and offer
a list of high-priority research questions to quickly
move the field toward potentially feasible and cost-
effective restoration. We emphasise that no approach
will be successful without swift and effective efforts
to mitigate greenhouse gas emissions. Mitigation
must go hand-in-hand with these restoration ap -
proaches, as intervention is intended only to temper
decline and accelerate natural recovery at more local
scales while global efforts take effect to slow the rate
of ocean warming and acidification.
Acknowledgements. We thank B. Schaffelke for providing
valuable comments on an earlier draft. This work was par-
tially funded by the Australian Government through the
Reef Restoration and Adaptation Program, a collaboration of
leading experts formed to create a suite of innovative meas-
ures to help preserve and restore the Great Barrier Reef.
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Editorial responsibility: Tim McClanahan,
Mombasa, Kenya
Submitted: April 8, 2019; Accepted: November 25, 2019
Proofs received from author(s): January 21, 2020