Reproductive phenology and sexual propagation of the pink sea fan Eunicella verrucosa Pallas, 1766 for coral restoration

Reproductive phenology and sexual propagation of the pink sea fan Eunicella verrucosa Pallas, 1766 for coral restoration | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Reproductive phenology and sexual propagation of the pink sea fan Eunicella verrucosa Pallas, 1766 for coral restoration Christina Egger, Catarina Melo, Bailey Marquardt, Aschwin H. Engelen, and 6 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-5741857/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 21 Jul, 2025 Read the published version in Coral Reefs → Version 1 posted 11 You are reading this latest preprint version Abstract The widespread decline of coral-dominated ecosystems due to human disturbances has highlighted the urgent need for active habitat restoration. Coral restoration using sexually produced individuals instead of clonal fragments is essential to reduce impacts on donor populations and promote genetic diversity, which is vital for adaptability to environmental changes. However, for most coral species, particularly those in temperate and deep-water (> 50 m), critical knowledge of reproduction and larval ecology for ex situ sexual propagation is lacking. To address this gap, in this study, we provide the first report of spawning of the octocoral Eunicella verrucosa in the North-East Atlantic and describe details on larval development and settlement. The annual reproductive timing in South-West Portugal was determined from samples collected as fisheries bycatch from a single population source and monitored for comparison across distinct durations and conditions. The species exhibited split-spawning over about one month (mid September – mid October), with 3 major events approximately every 2 weeks. Spawning patterns suggest lunar periodicity but shifted between colonies kept in distinct conditions. Oocytes were positively buoyant and developed into swimming larvae after 3 days. Settlement trials using substrates like natural rock, CCA, and gorgonian skeleton, showed larvae behaviour testing the substrates about two weeks post-spawning, and settlement activity continuing over three months. Fully developed recruits were observed after one month, with sclerite production starting before tentacle development. New settlement continued for up to three months, indicating a prolonged competency period. This study provides crucial data for coral restoration efforts using ex situ sexual propagation of this vulnerable species. broadcast spawning pelagic larval duration settlement competency octocorallia Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Figure 9 Figure 10 INTRODUCTION Coral ecosystems are experiencing a significant decline globally, primarily due to a combination of climate change and other anthropogenic pressures (Carpenter et al. 2008; Hughes et al. 2017b; Hughes et al. 2018). The present threats to coral reefs extend beyond rising temperatures. Ocean acidification adversely affects coral calcification and overall health, making them more susceptible to diseases and environmental stressors (Feng et al. 2016; Pendleton et al. 2016). Local human activities, such as pollution, overfishing, and coastal development, exacerbate these challenges, often leading to more immediate and severe impacts than climate change alone (Ferrario et al. 2014; Høegh-Guldberg et al. 2018). However, such effects are much less studied in temperate and deep corals than in tropical ones, in which widespread damage has been well documented. Coral reefs have lost approximately 50% of their living coral cover since the 1950s, with a corresponding 60% decline in fish populations associated with these habitats (Eddy et al. 2021; Hughes et al. 2017b). This loss not only threatens biodiversity but also undermines the ecosystem services that coral reefs provide and which are vital for the livelihoods of millions of people (Hughes et al. 2017a; Hughes et al. 2017b; Mercado-Molina & Suleimán-Ramos 2023). The altered trophic dynamics further destabilizes these ecosystems (Robinson et al. 2019). When coral reefs continue to degrade, the potential for recovery diminishes, creating a feedback loop that perpetuates the decline of these critical marine environments (Carpenter et al. 2008; Pandolfi et al. 2003; Pratchett et al. 2012). As conservation alone appears no longer enough for the long-term persistence of many ecosystems (Hughes et al. 2017a; O’connor et al. 2020; Orth et al. 2020; Smith et al. 2023), the United Nations and European Union have outlined the critical role of biological restoration to restore degraded ecosystems (UN Decade on Ecosystem Restoration 2021–2030 https://www.decadeonrestoration.org/ , the EU Biodiversity Strategy for 2030, the EU Nature Restoration Law), with the marine realm being given increasing attention (Marine Strategy Framework Directive (MSFD). Restoration efforts in shallow, warm marine ecosystems have notably advanced more quickly than those in deep, cold, and temperate ecosystems. This disparity is largely due to a combination of ecological, logistical, and financial challenges. Shallow tropical marine environments, including coral reefs, mangroves, and seagrass meadows, have received the most attention in restoration initiatives, primarily because they are more accessible and offer more perceived immediate socio-economic benefits (Bayraktarov et al. 2020; Williams et al. 2017). Colder and deeper marine habitats have received far less restoration a attention and efforts compared to their tropical counterparts (Ros et al. 2019). Damage to non-tropical corals has been accumulating over a long period, but it has often been overlooked due to the lack of visibility of these ecosystems. However, the ecological importance of these habitats cannot be overstated. They provide essential habitats for many marine species, including commercially valuable fish, yet they are as vulnerable as tropical corals (Bongiorni et al. 2010; Buhl-Mortensen et al. 2018; Buhl-Mortensen et al. 2017; Danovaro et al. 2010; Lange et al. 2023). This disparity is largely due to the technical challenges and high costs associated with deep-sea restoration, as well as a general lack of awareness and understanding of these habitats among policymakers and the public (Ounanian et al. 2018; Ros et al. 2019). This is evident in the absence of clear, consistent definitions for deep-sea, cold-water, and temperate habitats. Many coral species exist over broad depths and geographical ranges (Buhl-Mortensen et al. 2018), making it difficult to define their distributions and even harder to protect or restore them. Furthermore, the ecological complexity and slow recovery rates of deep-sea ecosystems add to the challenges of restoration. Many deep-sea species have long lifespans, and the intricate relationships within these ecosystems mean that restoration efforts take much longer to yield visible results. This delays investment and research in these areas (Prouty et al. 2016; Ros et al. 2019). As a result, while shallow marine ecosystems benefit from established restoration practices, deeper ecosystems remain in the early stages of restoration science, highlighting a critical gap in marine conservation (Ounanian et al. 2018; Ros et al. 2019). To counteract this, the essential scientific information for efficient restoration plans and techniques is needed, to allow to at least catch up with progress in restoration methods for shallow water counterparts. In coral restoration, there are several approaches, all of which gained noticeable increase over the last years (Boström-Einarsson et al. 2020; Suggett & Van Oppen 2022). The most common and widely applied method is asexual propagation, achieved by the fragmentation of donor colonies or from opportunistic coral sources like fishing-bycatch, broken pieces during storms (Boch & Morse 2012; Garrison & Greg 2008; Plucer-Rosario & Randall 1987). The corals can be grown in nurseries and subsequently out planted on the reef or directly transplanted (Forrester et al. 2019). Asexual propagation, however, carries the risk of limited genetic diversity or potentially dismissed impacts on the donor colonies and may not be self-sustaining (Baums et al. 2019; Henry et al. 2021; Van Oppen et al. 2015). A relatively new but steadily growing approach is restoration through sexual propagation, where sexually produced recruits can then be reintroduced on the reef (Henry et al. 2021). Coral spawn can be either collected in situ and then reared in the laboratory or field (Chamberland et al. 2015; Heyward et al. 2002; Suzuki et al. 2020) or the parent colonies can be kept and spawn in captivity (Dela Cruz & Harrison 2020; Henry et al. 2021; O’neil et al. 2021; Pollock et al. 2017). Ex situ aquaculture offers the opportunity to deliver a consistent supply of corals to support research and assist in complementary reef restoration (Lam et al. 2023). Sexual propagation offers the significant advantage of producing many genetically unique individuals thereby enhancing genetic diversity. This diversity enhances a population's ability to adapt to changing environments by providing a rich source of genetic variability for natural selection. Additionally, precise manipulations and targeted selection techniques, such as assisted evolution during the larval stage, can further increase the resilience of the new generation (Baums 2008; Doropoulos et al. 2019; Van Oppen et al. 2017; Van Oppen et al. 2015), though long-term trade-offs still need to be assessed. Despite the growing recognition for the need to actively restore coral habitats, the implementation of coral restoration at large scales in non-tropical corals has been very limited (but see Montseny et al. 2021b; Montseny et al. 2020) and it is so far restricted to asexual propagation (Boch et al. 2019; Roik et al. 2015; Ros et al. 2019). In contrast, in tropical coral reefs, methods have been developed and simplified, especially for upscaling restoration through non-scientific organisations that implement them in the field (SECORE; Bayraktarov et al. 2019; Boström-Einarsson et al. 2020). We are lacking such methodologies and protocols for deeper reefs, despite the growing interest of non-scientific conservationists and explorational divers in these underexplored ecosystems (Ankamah-Yeboah et al. 2020; Mengerink et al. 2014). When it comes to sexual propagation in deep-sea corals, the first major challenge is the limited understanding of their biology and reproductive patterns. Many of these species, particularly those found below diving depth, remain understudied due to logistical and financial difficulties (Montseny et al. 2021a; Randall et al. 2020). The second challenge lies in our ability to keep these corals healthy in captivity for extended periods, which is essential for manipulating and inducing spawning. Despite these hurdles, addressing them is crucial for developing effective restoration strategies. Deep-water and cold-temperate coral habitats differ significantly from tropical shallow-water reefs in terms of physiology, environmental conditions, and species composition (Bridge et al. 2013; Menza et al. 2007; Price et al. 2019). Coral gardens that are dominated by gorgonians can form loose to dense forest-like aggregations, often referred to as marine animal forests (Rossi et al. 2017). These habitats extend from shallow depths to several hundred meters below the surface. They typically consist of a few key gorgonian species that create a three-dimensional structure, providing essential habitat and support for a diverse array of marine organisms (Baillon et al. 2012; Buhl-Mortensen et al. 2017; Watling et al. 2011). The pink sea fan, Eunicella verrucosa , plays such an important, habitat-forming role in biotopes of the Eastern Atlantic and the Mediterranean Sea. It grows on hard substrate over a large latitudinal and depth range (2-200m) (Chimienti 2020; Coz et al. 2012; Pikesley et al. 2016; Sartoretto & Francour 2012). Its trans-equatorial distribution spans from western Ireland and SW British Isles to Angola in Western Africa and includes the Mediterranean Sea (Carpine 1963; Carpine & Grasshoff 1975; Chimienti 2020; Grasshoff 1992). Like other temperate and cold-water corals, these species exhibit long lifespans and slow growth rates (0.62-3.33cm/year in height: Sartoretto & Francour 2012), making their populations particularly susceptible to anthropogenic disturbances such as anchoring and fishing (Watling & Norse 1998). These human activities can exacerbate the already significant impacts of climate change, which include increased frequency of natural disturbances like storms and marine heatwaves (Hall-Spencer et al. 2007; Munro & Munro 2003; Sheehan et al. 2017; Sheehan et al. 2013). Eunicella verrucosa is listed as vulnerable in the IUCN Red List of Threatened Species since 1996 and is considered to have a high to medium-high extinction risk due to exploitation. Impacts resulting from bottom contact fishing gear are one of the major risk factors for the species in South Portugal, where E. verrucosa is one of coral species most numerous caught as fishing bycatch (Dias et al. 2020). Very little is known about the biology and life cycle of Eunicella verrucosa . Munro (2004) studied some populations in the UK, revealing that they are gonochoric, with the female reproductive cycle extending beyond 12 months. Broadcast spawning has been suggested as the mode of reproduction, with spawning likely occurring in August or September (Munro 2004). This is a limitation for restoration because reproductive traits such as planktonic larval duration (PLD), larval behaviour (buoyancy and swimming behaviour) or mode of reproduction of the corals (brooder or spawner) are crucial information for sexual propagation, as well as to help understand natural population dynamics that can directly influence the planning of restoration strategies (Fogarty & Botsford 2007; Jones et al. 2007; Marti-Puig et al. 2013; Waller et al. 2023). The aim of this study was to characterize the reproductive and larval biology of Eunicella verrucosa and test methods of sexual propagation in captivity, thus addressing the conservation needs and information gaps highlighted above. Specifically, we document for the first time the reproductive phenology of the species in populations from SW Portugal, comparing colonies maintained in captivity for short- and long-term (1 and 2 years). We also describe larval development and settlement preferences using experiments with the overall objective of obtaining sexually produced recruits for habitat restoration. METHODS Inference of Reproduction Window The timing of reproduction in E. verrucosa was inferred by examining the stages of gamete development in samples collected as fishing bycatch around Cape St. Vincent (southern Portugal) in 2020 and 2021 (see Dias et al. 2020 for details on fishing areas) at the Centro de Ciências do Mar (CCMAR). Sampling focused on the Summer and early Autumn periods following available information on reproductive patterns for the species in the UK (Munro 2004) and other Eunicella spp. in the Mediterranean (Coma et al. 1995b; Gori et al. 2012). The presence and maturation state of the gametes was assessed through polyp dissection and histological sectioning at the at CCMAR. All samples were fixed in 10% Formalin, washed three times with distilled water and gradually dehydrated to EtOH 70% until further analysis. Dissection of the polyps was performed under a stereomicroscope, first to distinguish female and male individuals by identifying their gonads (oocytes and spermaries) where possible. The number of gonads per polyp and branch order (sensu Brazeau & Lasker 1988) was then counted and the individuals with higher number of reproductive polyps were selected for histological sectioning. For the histological study, branchlets of approximately 2 cm were cut from the first and second order branches of four colonies. Selected samples for histology were hydrated and treated in 2.1 M EDTA (pH 8) for two days to decalcify the sclerites embedded in the coral tissue, following the method used for sea bream scales (Vieira et al. 2011). After decalcification, the samples were washed several times with deionized water to remove any residual EDTA and then dehydrated through a graded ethanol series. The samples were saturated in xylene, followed by impregnation and embedding in low melting point paraffin wax (Histosec, Merck) using an automatic embedding processor. Serial sections of 5 µm were cut from the paraffin block using a manual rotary microtome (Leica RM 2135, Germany) and mounted on glass slides coated with 3-aminopropyltriethoxysilane (APES; Sigma-Aldrich, Madrid, Spain). For each wax block containing a piece of coral branch, 2–3 sections were cut from the base of the polyps to sample the gonadal tissue. The sections were dried overnight at 37°C, then cooled to room temperature for storage or staining. The sections on the slides were dewaxed and rehydrated before being stained with hematoxylin and eosin (H&E) according to the method described by Najafpour et al. (2020). They were then mounted in Tissue-Tek Resin (Sakura Finetek) and covered with a glass coverslip. The stained slides were visualized under a light microscope to identify the reproductive structures of the polyps. The gametogenic maturation classification system defined by Waller (2005) was used to describe the ripening process of the gametes. Spawning Observations and Larval Rearing The observations on spawning and settlement were conducted on corals collected as fishing bycatch from approximately the same location as the corals used to determine the spawning window: Cape St. Vincent in southern Portugal. However, observations were made on corals collected during three consecutive years (2021, 2022 and 2023) and kept at two separate facilities: the marine field station Ramalhete of CCMAR (corals from 2023) and the Oceanário de Lisboa (ODL; corals from 2021 and 2022). Shorter-term monitoring during the reproductive season (CCMAR) In 2023, coral colonies were collected from early July to mid-August. A total of 70 colonies, varying in colour, size, and health, were collected in several batches. The colonies were dissected, sexed, and then transferred to tank systems at Ramalhete Marine Station in Faro for spawning observations. Approximately 50 colonies were maintained in a cooled, semi dark outdoor flow-through system, and 20 in a dark indoor, semi-closed, temperature-controlled system at 15–16ºC. Water flow was provided by wavemaker pumps and corals were fed once daily with frozen rotifers, copepods and red zooplankton, which were squeezed through a 150µm net to avoid oversized food particles. During feeding water in- and outflow was stopped for 1–3 hours to avoid the food being washed out. The tank bottoms were hoovered daily to provide good water quality. During the expected spawning window, perforated PVC cylinders with a 150µm mesh fitting were attached to the outflow of the tanks to collect released offspring. A venturi airlift was also installed at the water surface of the outdoor tanks. The air lift, outflow filters and colonies were monitored daily for gamete and/or larval release. Additionally, branches were regularly dissected to check for the presence of mature gametes. During spawning, oocytes and embryos were collected both directly from the water column using a transfer pipette and from the two airlifts (in case of the outdoor tanks) and the outflow filters, by gently washing them into plastic beakers. Eggs, embryos, and larvae were kept in plastic jars and boxes (1–10 L) sorted by collection date in a temperature-controlled room at 16 ºC. Gentle aeration was provided via air tubes producing a few bubbles per second, and approximately two thirds of the water was changed every other day using a 5mm hose with a small, perforated tube with a 150µm plankton net wrapping at the end to avoid larvae being sucked out. The intensity of spawning each day was categorized into low (10–250 propagules collected) and high (> 250 propagules collected) spawning. Releases below 10 eggs were not considered as an event. Longer-term monitoring (1–2 years, ODL) A total of 15 colonies collected in 2021 and 9 colonies from 2022 were kept together in a closed indoor system at the Oceanário de Lisboa. The life support system included an EcoDrift 4.2 pump from Aqua Medic Direct and a Hydor Seltz L 700 pump in each aquarium, a 50 µm filter bag at the inlet of a 1300 L sump containing bioballs, a 20 µm cartridge filter, two UV lights, a Frimar C1000 chiller and a HydroAir pump (model AV50-20N-S). The aquariums had indirect sunlight and moonlight from east turn windows and indirect light from the room ceiling (from 7h30 to 16h30). The moon cycle was simulated using a 54W actinic light set with a timer and an intensity adjuster, following the moon cycle at Sagres (from the website Timeanddate.Com, n.d.). Temperature modulation was based on seasonal variation patterns. The corals were fed live and frozen zooplankton and phytoplankton three times a day. The food concentration was adjusted based on field variations. From the 26th of July till the 16th of November, egg collectors were placed in the surface skimmer of each aquarium from 16h00 to 8h00. Water flow was reduced, and circulation pumps were turned off to prevent egg and embryo damage. In the morning, the collectors were rinsed with saltwater to flush all gametes into a plastic beaker, allowing their observation under a stereomicroscope. The collected eggs, spermic sacs and embryos were placed in 4 L circular boxes with 125 µm mesh on the bottom and with slow water flow, connected to the main system, where the breeding colonies were maintained. Embryogenesis and Larval Development (CCMAR) Embryonic and larval development stages were observed and imaged using a ZEISS Stemi 508 stereo microscope with a ZEISS Axiocam 208 colour camera system, at Ramalhete Marine Station (CCMAR). Qualitative observations were conducted on > 10 batches of 50–600 larvae each throughout the development period to ensure consistency. Samples of various developmental stages were collected for scanning electron microscopy (SEM). These samples were fixed overnight in 4% glutaraldehyde buffered in 0.1–0.5 M Sørensen’s phosphate buffer (pH 7.1), at 4 ºC. The following day, the samples were washed three times in pure Sørensen’s phosphate buffer and transferred into 30% EtOH where they were stored until further analysis. All SEM imaging was performed at the SNSB - Bavarian State Collection for Zoology in Munich (Germany). Samples were then dehydrated using a graded acetone series (30%, 50%, 75%, 95% and 100%). Specimens were soaked for 10 min at each level, then twice in 100% acetone. Subsequently they were critical point dried using a Polaron E3000, mounted on SEM stubs with self-adhesive carbon stickers, and gold-coated for 3 minutes in an argon atmosphere using a Polaron SC510. Three embryos or larvae of each stage of development were analysed with a LEO 1430 VP SEM at a voltage of 15 kV (method described in Melzer et al. 2021; Torres et al. 2021). Measurements were done on > 5 propagules per stadium from LM images, using the scale bar and Adobe Photoshop© 22.2.0 and compared with the SEM images for consistency. Larval Behaviour (CCMAR) Larvae, kept in plastic jars and boxes (1–10 L), were provided with a substrate of bare rock and CCA-covered rock when they were between 1 and 2 weeks old. Larval behaviour was documented qualitatively by close observation of the individual batches of each spawning event to ensure consistency. The larvae were visually observed directly in their rearing jars within the cold room. Microscopic observations including video records of the substrate probing behaviour were conducted under the Stemi stereo microscope. Larval Swimming Behaviour To document the swimming behaviour of 4–6 days old larvae, three replicates of 19, 15 and 16 larvae, respectively, were temporarily placed in a 10x10x2 cm jar with a black background and filmed using a Canon 550D camera with Canon EF 50mm F1.8 STM objective, at Ramalhete Marine Station (CCMAR). Larval Survivorship and Substrate Choice and Settlement (Experiment 1) A settlement experiment was conducted to document larval settlement and metamorphosis, as well as to test settlement preferences using natural substrate. Three types of substrate were offered for settlement choice: rocky substrate collected as bycatch along with the corals, which were encrusted with a variety of taxa, including natural biofilms and/or crustose coralline algae (CCA) that are presumably implicated in inducing coral settlement and metamorphosis (Zelli et al., 2020); chips of an unidentified species of CCA retrieved with the substrate aforementioned; and pieces of conspecific bare gorgonian skeleton, on which larvae of other octocorals have been observed to settle (Coelho & Lasker, 2014; Weinberg & Weinberg, 1979; authors personal observations). Larval settlement was quantified for two cohorts of larvae spawned on October 11 (Cohort 1) and September 30 (Cohort 2), which were 7- and 17-d old at the start of the experiment, respectively. For each cohort, we used 5 replicates of 50 larvae each that were challenged with the three types of substrates. All replicates were maintained in open plastic Tupperware containers of ca. 8 cm diameter and 4 cm tall with ~ 250 ml of seawater. 50% water changes were done after each counting every 2–3 days. The containers were kept in a separate room with same ambient temperature as the coral indoor tank system and pH was regularly checked to monitor for variations due to organic or inorganic substrates. On the 28th of October (day 11 of the experiment of Cohort 1 and day 12 of the experiment of Cohort 2) the experiments had to be transferred to a different room kept at 18 ºC due to a complete failure of the air conditioning in the original room. Larval settlement (i.e. attachment and metamorphosis) was counted as the attachment of larvae or the presence of recruits on a substrate in each of the 5 replicates. Monitoring of larval settlement was performed every second day at the beginning of the experiment and alternating for each cohort (experiments started on 16th and 17th of October for Cohorts 1 and 2, respectively). After 11 days, the monitoring intervals were increased to 3–4 days until 34 days post-start and to every 7 days thereafter. At day 62 (Cohort 2) and 63 (Cohort 1), more substrate was added to each replicate to test for a potential settlement bottleneck caused by the occupation of available space by settlers as settling rates seemed to decrease. To evaluate substrate choice, settlers on each substrate were counted. For the evaluation of general attachment and settlement, settlers were counted for each replicate over all three substrates and the total and average number of settlers across all replicates was calculated. To compare settlement between the cohorts, we used the age of the larvae. After settlement, primary polyps were transferred to the tanks containing the mother colonies and provided with the same food. Early Life Ecology Short-term monitoring (CCMAR) Larval settlement and post-settlement mortality was monitored over a period of 9 months. Single recruits were followed over the period of one year to document their development. All recruits were imaged using the previously described stereo microscope. Long-term monitoring (ODL) Larvae from the major spawning events observed in Oceanário de Lisboa in September and October were transferred from the circular boxes to four 4l rectangular boxes with 200 µm mesh on the side and water entrance from the main system 12 days after fertilization. Larvae were provided with natural rocks with CCA for settlement, which were collected from the wild 2 months earlier, as well as with basaltic rocks conditioned in the gorgonian system for over 7 months. The microalgae Chaetoceros calcitrans and Tisochrysis lutea were provided to the larvae since 5 days after (presumed) fertilisation. The same microalgae and living Acartia tonsa and Brachionus plicatilis and frozen Copepods (Ocean Aquaculture ®) were fed after the first primary polyps were observed. The polyps were kept in the same boxes until 197 days, when 2 of them were transferred to one of the aquaria containing the adult colonies. Gradually all polyps were transferred to the same aquarium. Monitoring of larval settlement and polyp survival was conducted every two weeks until month 3 and every month after that using a Nikon SMZ745T stereo microscope in Oceanário de Lisboa. Recruits Survival (Experiment 2) To estimate total settlement success and recruit survival we followed 4 cohorts of eggs (3 cohorts from short-term monitoring at CCMAR of 226, 200 and 100 eggs, respectively, and 1 cohort of 1621 eggs from long-term monitoring at ODL) and 4 cohorts of larvae (2 cohorts of 250 larvae from Experiment 1 and 2 more cohorts of 250 and 180 larvae, respectively, all from short-term monitoring at CCMAR). RESULTS Inference of Reproduction Window The gamete development data for samples of E. verrucosa collected in 2020 and 2021suggests that reproduction occurred in late summer or early autumn in both years. For 2020, polyp dissections of multiple female colonies revealed a significant decrease in the number of oocytes from June to September, indicating potential spawning over this period (see Fig. S1 in supplement 1). Histological sectioning of samples from 2021 pointed towards a reproductive window to be in a similar period to that of 2020, with female colonies still having late stage 4 vitellogenic oocytes on August 5th and 26th and in mid-September (Fig. 1 a-c). The spermatocytes showed a ripening process from small, growing (stage 2) spermatocytes in early August until stage 3 spermatocytes in mid-September with empty lumen. This shows that the accelerated ripening indicates the proximity of the spawning period. Polyp dissections of male and female colonies collected in August 2023 for spawning observations revealed vitellogenic or late vitellogenic (stage 3 or 4) oocytes and visible spermatocytes in most of the colonies (data not shown). In all the polyp dissections and histological sections made we never observed embryos or larvae inside the polyps which indicates that E. verrucosa is a gonochoric broadcast spawner. Spawning Observations Short-term monitoring (CCMAR) For the 2023 reproductive season, spawning at Ramalhete Marine Station was first observed on September 2nd, when minor release of positively buoyant eggs and developing embryos were floating in both the inside and outside tanks systems synchronously, and accumulated in the collection filters attached to the water outflow. Most of the spawning then occurred as split spawning in 3 major events: the first one over two consecutive days in September 12–13, the second one on the 26th and 30th of September, and the third one on the 11th of October (Fig. 2 ). Each large spawning event was preceded and followed by minor egg releases (Fig. 2 ). The spawning peaks appeared to correspond with the moon phase, with the largest events occurring between 5 days before and 3 days after the full or new moon, and the highest spawning output consistently happening 2–3 days before the full or new moon. The eggs released were ~ 300–400 µm in diameter, all positively buoyant and accumulated in the overflow collection filters after being released into the water column, thereby confirming E. ver rucosa to be a broadcast spawner. Long-term monitoring (ODL) For colonies of E. verrucosa kept in captivity at Oceanário de Lisboa for over two years, egg release started earlier with minor releases first observed on August 4, extending into the 14th of November despite of very low spawning after the 23rd of October (only 0–3 oocytes) (Fig. 2 ). The major spawning events occurred on the 6th and 7th of September (6 days before the large event documented at Ramalhete) and on the 5th, 6th and 15th of October (Fig. 2 ). The first four major events occurred 6–7 days after the full moon, with the timing of spawning for the first three starting between 5:15 hrs and 6:10 hrs after moon rising. At the major spawning event in September almost all eggs and embryos were positively buoyant, however, at those occurring in October only a minor fraction had positive buoyancy. Embryogenesis (CCMAR) Embryos collected at CCMAR developed into a gastrula within 24 h of egg collection from the tank system. Fertilisation rates of the eggs seemed to be very high, as all eggs observed started cleaving. Once eggs started cleaving, the embryos were observed to sink down in the rearing containers, indicating negative buoyancy. We observed partial cleavage (meroblastic) of the eggs until the 8-cell stage (Fig. 3 a, d and video in supplement 2). The same observations were made for embryos reared at ODL. The greater part of the yolk remained in the initial (egg) cell at the beginning. In some cases, eggs started segmenting into one big and four to five smaller compartments that did not cleave entirely, returning instead to the initial round shape only to start segmenting again and then proceed to cleave into 8 and finally 16 evenly sized cells (Fig. 3 e, f and video in supplement 2). The cleavage was spiral and above the 32 cleavages, the pattern continued in irregular-sized cells with six squares and an irregular-shaped embryo (Fig. 3 g, i). While the six squared cells became more equal in size with further cleavage, the embryo remained somewhat irregularly shaped and developed several infoldings and invaginations during gastrulation (Fig. 3 j, k, l). Larval Development and Behaviour (CCMAR) Seventy-two hours after fertilization, the first oval-shaped larvae (~ 350µm long) started moving and actively swimming upwards to the water surface in the rearing containers where they congregated (Fig. 4 a, b and videos in supplement 3). Five to six days after egg collection, larvae were already more elongated (Fig. 4 a, d) and approximately 3 times as long as wide (~ 750µm long, ~ 250 µm wide). The same observations were made for larvae collected at CCMAR and ODL. At the age between 3 to 9 days old, the larval ciliation was sparse, with short cilia covering the larvae body during this period (Fig. 4 c) and ciliation increasing until day 17 of age (Fig. 4 e, g). The increase in larval ciliation coincided with an increase in swimming activity, with larvae between 7- and 12-days following spawning observed to start to partially leave the water surface and standing upright in the water column. After 13 days, larvae were observed to frequently swim up and down and starting to probe substrate (see video 1 in supplement 4). During that phase, larvae developed an oral and aboral pole, denser ciliation, and had a clear propulsion direction (Fig. 4 e-k). With advancing age, the larvae became more transparent and shorter, however stayed motile until over 100 days of age (Fig. 4 j, k). Larval Settlement, Metamorphosis and Early Life Ecology Short-term monitoring (CCMAR) The first attachment of larvae to the substrate offered in the settlement experiments at Ramalhete Marine Station was observed 15 days after spawning, when the larvae temporarily attached with the aboral pole to the substrate provided (Fig. 5 a and video 2 in supplement 4). Onset of settlement started shortly after and continued over the next two months. After successful settlement, metamorphosis occurred over several days (Fig. 5 b), during which the settlers sequentially developed mesenteries, a mouth and tentacles. The primary polyps appeared healthy, apparently began feeding, and started developing sclerites within a few weeks (Fig. 5 b, c). These sclerites formed in eight vertical rows surrounding the mesenteries (Fig. 5 c) and later developed into acuminate leaflets enclosing the mesenteries similar to petals (Fig. 6 b). When stimulated or stressed, these leaflets would form a complete, enveloping spherical shell around the contracting polyp (Fig. 6 a). Growth in the aquarium was slow. Within the first 3 months, the settlers grew from 0.5 mm to a maximum length of 1–2 mm (Fig. 6 d). By 9 months, they had expanded from 0.5 mm to 1 mm in width and reached up to 5 mm in height (Fig. 6 ). The first deposition of an internal skeleton was observed at 9 months post-settlement, or 11–12 months post-spawning (Fig. 6 b, e). Shortly after, branching of the second polyp was noted (Fig. 6 c, e). The internal axis in the primary polyp began to develop, bending away from the mouth region (Fig. 6 b), while the second polyp branched in the opposite direction from the axis below the primary polyp (Fig. 6 c, e). By the time the third polyp developed, it emerged from the axis between the first and second polyps, while the fourth polyp developed below them (Fig. 6 f). As they grew, the recruits also started to deposit a holdfast on the substrate to which they were attached (Fig. 6 c, e). Longer-term monitoring (ODL) At Oceanário de Lisboa, the first recruits were observed at 20 and 21 days after the major spawning event (6th-7th September). Settlement continued over a period of 3–4 months, similar to the observations made at CCMAR. All larvae settled in CCA rocks, and no settlers were observed on the basaltic rocks. From the October major spawning events in which a majority of oocytes were negatively buoyant, none of the larvae settled. The first tentacles with pinnules were observed at 36 days after spawning and the first observation of a secondary polyp occurred 204 days or 6.5 months after spawning, shortly after the transfer to the aquarium with the adult colonies. Settlers with 3 or more polyps were observed almost one year after spawning. Larval Survivorship and Substrate Choice and Settlement (Experiment 1) In the settlement experiment conducted at CCMAR, we observed two small settlement peaks at 15 days after spawning in Cohort 1 (released on 11th October, 7 days old when experiment started) and at 19 days in Cohort 2 (released on 30th of September 17-days old when experiment started) (Fig. 7 ). Those peaks did not lead to successful settlement, as the number of settlers counted in the following monitoring day decreased to 0 again, either indicating a temporary detachment of the ‘settler’ or failure to complete metamorphosis (Fig. 7 ). The number of settlers then increased slowly until 78 days of larval age in Cohort 1 and 89 days of larval age in Cohort 2, followed by a steeper increase in settlement until 97 days of larval age to a maximum of 19 settlers in Cohort 1 (C1) and until 96 days of larval age to a maximum of 29 settlers in Cohort 2 (C2). The increase in settlement occurred 9 (C1) and 19 days (C2) after the addition of new substrate when larvae were 69 and 80 days old in C1 and C2, respectively. There was no obvious change in settling behaviour after the room change at 11 (C1) / 12 (C2) days of running experiment (18 and 29 days old larvae) (Fig. 7 ). Beyond a larval age of ~ 95 days no more additional settling was observed. Figure 8 shows the average counts of survivors (larvae + settlers) over the same range of time. The number of survivors shows a steady decrease until the same point in time as above (~ 95 days) and then turns to an almost constant level. This indicates that larvae are competent to settle during the entire pelagic larval duration, until all larvae have either settled or died. Substrate choice appeared to vary over time during the settlement period in both Cohorts 1 and 2, as well as between cohorts. Initially, settlers appeared to preferentially attach to bare gorgonian skeleton, but this preference decreased over time, with larvae at later stages of development settling more on the rocky substrates provided (Fig. 9 ). In cohort 1 (C1, orange colour in Fig. 9 ), the maximum number of settlers on rock was 9, on CCA (coralline crusts) was 4, and on gorgonin was 6, observed on days 97 and 100. During days 15 to 21, settlement was exclusively on CCA and gorgonin. From days 24 to 55, settlers were observed only on rock and CCA, with the number of settlers on rock slowly increasing from 1 to 9, and on CCA from 1 to 4. From day 85 onward, settlers appeared on all three substrates, with the number of settlers on gorgonin increasing from 4 to 6 by day 97. This number remained constant until the experiment's conclusion on day 100. In cohort 2 (C2, pink colour in Fig. 9 ), the highest number of settlers was recorded on days 73 and 80, with 19 settlers on rock, 2 on CCA, and 8 on gorgonin. During days 17 to 21, attachment was only observed on gorgonin. From day 24 onward, settlement was observed on both rock (settlers increased from 1 to 19 by day 73) and gorgonin (settlers increased from 1 to 8 by day 73). Settlement on CCA was minimal, with only 1 to 2 settlers recorded on days 49, 73, 80, and 89. On day 89, the number of settlers on rock decreased slightly to 17 while the number of settlers on gorgonin remained constant. Only 1 settler remained on CCA. Recruits Survival (Experiment 2) From the 526 eggs monitored at Ramalhete Marine Station in 3 settlement batches, 56 larvae (11%) settled. After 4 months, 33 of these settlers were still alive, yielding a survival rate of 59% of settlers. At more than 6 months post-settlement, 36% of initial settlers were still alive resulting in an overall survival rate from egg to 6–7 months post-settlement of 3.8% (see Fig. 10 a for data on individual cohorts). Out of the 930 larvae monitored across 4 batches, 61 (6.6%) successfully settled (see Fig. 10 b for data on individual cohorts). After 4 months post-settlement, 35 of these settlers were still alive, and after more than 6 months 17 settlers were still alive resulting in a survival rate of 28% of the settlers. This corresponds to an overall survival rate of 1.8% from the original 930 larvae. Out of 1621 eggs, 29 larvae settled at Oceanário de Lisboa within 4–5 months after fertilization (1.8%). 6–7 months after fertilization 25 settlers were still alive, corresponding to a survival rate of the settlers of 86% and a total survival rate after 6–7 month of 1.5%. At 9 months after fertilization, some mortality was detected, and the survival rate decreased to 59% of the settlers (1.0% total). 10 settlers were still alive 12 months after fertilization, corresponding to a survival rate of 34% within the settlers (0.61% total). DISCUSSION Reproductive Cycle and Spawning The histological investigations revealed that female oocytes were already close to mature (stage 4) at least 1.5 months before spawning. In contrast, the spermaries seemed to ripen more quickly from immature (stage 2) to nearly mature (stage 3) while becoming noticeably larger within the same period. These findings suggest a distinct difference in the maturation timelines of oocytes and spermaries. Similar patterns have been observed in the related species such as Eunicella singularis and Paramuricea clavata from the Mediterranean (Coma et al. 1995b; Ribes et al. 2007; Weinberg & Weinberg 1979) and in E. verrucosa populations in the UK (Munro 2004), as well as in other octocorals (reviewed in Kahng et al. 2011). Polyp dissection and histologic section data indicated a spawning window in late summer/early autumn, later confirmed by ex situ observations in aquaria. This contrasts with Mediterranean gorgonians, including E. singularis , and Eunicella cavolini , which spawn in spring or early summer (Gori et al. 2012; Ribes et al. 2007; von Koch 1887; Weinberg & Weinberg 1979). In the UK, E. verrucosa colonies have been inferred to spawn in August or September but no direct observations were ever made (Munro 2004). The difference between the water bodies of Atlantic Ocean and Mediterranean Sea in terms of temperature regimes and stratification, currents and tides likely explain the variation in timing. Regional variation in the timing of spawning of coral populations is a well described phenomenon in corals, also across the Mediterranean Sea, and is often related to temperature variations (Foster & Gilmour 2020; Kersting et al. 2013; Osman et al. 2024; Sakai et al. 2024).Several environmental cues and variables are known to influence spawning in tropical corals, including solar insolation, moonlight intensity, day length, and temperature (Hatta et al. 1999; Kaniewska et al. 2015; Paxton et al. 2016; Van Woesik et al. 2006). However, for cold- and deep-water species, where some of these cues may be absent or differ significantly, our understanding is limited, partially due to the challenges of studying these species at depth. In our study, coral colonies were maintained in both indoor and outdoor tank systems. The indoor system was kept at a stable temperature of 14–16°C, while the outdoor system, although cooled, experienced natural temperature fluctuations that exceeded 18°C during the summer. Despite these differences, spawning occurred simultaneously in both systems in at least one of the major spawning events (September 12–13), suggesting that temperature is not the primary driver of spawning synchrony, although it may still influence the gamete maturation. Interestingly, colonies collected in previous years and maintained for 1–2 reproductive cycles under different conditions at the ODL spawned during the same general period, though not on the same exact days. The natural seasonal temperature fluctuations and moon cycle (i.e. light intensity) were simulated within ODL’s closed tank system to mirror in situ conditions as best as possible based on available information, which supports the hypothesis that temperature plays an important role in regulating the annual gametogenic cycle of corals kept in captivity (Sakai et al. 2024). The incomplete synchronization of the time of spawning between colonies kept at CCMAR (two tank systems) and ODL likely result from fine-tune responses of corals to other environmental conditions specific to each captivity system, as well as to inter-colony variability (Gilmour et al. 2016; Monfared et al. 2023). The spawning period of Eunicella verrucosa lasted several weeks and appeared to be tentatively correlated with the lunar cycle, with peak spawning activity occurring around the full and new moons for colonies kept at CCMAR. Extended spawning periods and correlation with the lunar cycle have been reported for some temperate and cold-water corals (Coma et al. 1995a; Gori et al. 2012; reviewed in Waller et al. 2023; Weinberg & Weinberg 1979), and moonlight is widely recognized as a likely environmental cue triggering spawning in broadcast spawning corals (Kaniewska et al. 2015; Randall et al. 2020; Sorek & Levy 2014). While many cold-water corals inhabit depths beyond the reach of moonlight, E. verrucosa spans a wide depth range (~ 10–200 m depth) (Grasshoff 1992), including shallow-water habitats where moonlight can still be detected (Kaartvedt et al. 2019). This suggests that spawning of E. verrucosa may be influenced by the lunar patterns as observed in other temperate gorgonians like P. clavata (Coma et al. 1995a), but likely relies on a combination of additional cues to regulate the exact timing of spawning. At ODL, the spawning showed a less pronounced correlation with the lunar phase, with two of the three spawning peaks occurring over two consecutive days 6–7 days after the full moon (Fig. 2 ). This correlation, which differs slightly from the observations made at CCMAR, appear to suggest an effect of the artificial moonlight provided at ODL on the time of spawning: in 3 nights of major spawning recorded at the ODL’s closed system, egg release was observed to start between 5:15 hrs and 6:10 hrs after the simulated moon rise. On the other hand, at CCMAR only the outdoor tank system was exposed to direct moonlight while the indoor system was not, yet spawning still occurred during the full and new moon phases in both systems. This indicates that moonlight is not the sole trigger for spawning; gravitational forces during these lunar phases may also play a significant role, or the internal biological clock of the corals may remain entrained, even in the absence of external cues. The observed temporal shift between the spawning maxima of CCMAR and ODL may be related to the fact that the CCMAR samples were relatively fresh, whereas the ODL samples had been kept in tanks for 1–2 years. Our observations confirm that Eunicella verrucosa is a broadcast spawner, as previously suggested by (Munro 2004). This contrasts with other members of the genus like E. singularis and E. cavolini , which are internal brooders (Ribes et al. 2007; Theodor 1967; von Koch 1887; Weinberg & Weinberg 1979) further highlighting that reproductive mode is relatively plastic in corals (Kahng et al. 2011; Kerr et al. 2011). While broadcast spawning is the dominant reproductive strategy in shallow-water scleractinian corals, approximately half of the studied octocoral species are internal or surface brooders (Kahng et al. 2011). Fewer deep-water species, however, have been studied, making it difficult to identify broader patterns (Eckelbarger et al. 1998; Rakka et al. 2021; Rakka et al. 2017; Sun et al. 2010; Sun et al. 2009; Sun et al. 2011; Waller 2005; Waller et al. 2023). Embryonic development Embryonic development in E. verrucosa showed several notable differences in comparison with other members of the genus. First, while egg segmentation in closely related species like E. singularis is described as holoblastic (Weinberg & Weinberg 1979) in which the eggs cleave into two equal-sized blastomeres that then further divide into 4, 16, and so on until the gastrulation stage, we observed partial (meroblastic) cleavage in E. verrucosa . Most of the yolk remained in the original egg cell and the cleavage process was incomplete, with blastomeres not fully separated by membranes (Brun-Usan & Salazar-Ciudada 2020). Most studies on coral embryonic development report holoblastic cleavage patterns (Brun-Usan & Salazar-Ciudada 2020; Linares et al. 2008; Okubo et al. 2013; Rakka et al. 2021), similar to what has been observed in E. singularis . However, it is not unusual for cleavage patterns to vary across closely related species, particularly among nonbilaterian invertebrates like cnidarians (Brun-Usan & Salazar-Ciudada 2020). Meroblastic cleavage is common in yolk-rich eggs and occurs because the dense yolk interferes with the formation of the cytoskeleton during cell division (Adamska et al. 2011; Martin 1997). This suggests that the eggs of E. verrucosa exhibit this pattern due to high yolk content. The presumed high lipid content remains to be assessed but may help explain the long pelagic larval duration (PLD) for the species in this study (Viladrich et al. 2021). In other cnidarians, chaotic cleavages and temporal syncytial stages—where random blastomeres fuse—have been observed, resulting in different cleavage patterns even between individuals (Brun-Usan & Salazar-Ciudada 2020). Such cleavage patterns may help explain the observations made for E. verrucosa embryos, where partially cleaved embryos containing one large blastomere and four to five smaller ones, sometimes fused back into a single large cell before progressing to the final cleavage stage, which produced 8 blastomeres. Between the 8- and 32-cell stages, the embryos followed a more organized cleavage pattern. However, beyond the 32-cell stage, the embryos became asymmetrical again, with some blastomeres remaining very large while others were much smaller. These variations were seen among multiple individual embryos. As cell sizes became more uniform and the embryos developed regular 6-sided shapes, the embryos exhibited relatively deep and extreme infoldings reminiscent of a tangle more than the "raisin-like" formation, that has been described in other tropical gorgonians (Lasker, personal communication, Lasker & Kim 1996; Tonra et al. 2021). Larval development, behaviour and settlement The eggs (300–400 µm diameter) and larvae (500–750 µm length, 250–350 µm width) of E. verrucosa were larger than those of the surface-brooding species like P. clavata (eggs 250–350, larvae 500–800 µm in length but very thin, Linares et al. 2008), but smaller than those of the internal brooder E. singularis (larvae 2500 µm in length, 500 µm in width, Weinberg & Weinberg 1979) from the Mediterranean Sea. Although larval behavioural traits were not quantified, some preliminary qualitative findings are worth noting. Similar to many other broadcast spawning coral species, the eggs of E. verrucosa were mostly positively buoyant for a few hours. However, the embryos became negatively buoyant once they started cleaving and remained so for up to three days until the onset of larval swimming activity. A significant decrease in propagule buoyancy during embryogenesis has been previously reported (e.g., Coelho & Lasker 2016a) and is likely the result of the temporal decrease in lipid content that occurs in several species of broadcast spawning corals as larvae develop (Figueiredo et al. 2012; Harii et al. 2007). The larvae of E. verrucosa were consistently observed to swim upwards, especially in the first two weeks, suggesting active swimming behaviour that likely compensates for the decrease in buoyancy (although this has not been tested). SEM images of E. verrucosa larvae during this stage revealed that ciliation appeared incomplete. At a later stage, the larvae continued swimming actively, but with more up-and-downward movement including substrate probing behaviour. The further development of cilia and changes in swimming behaviour seemed crucial for larvae to become more agile and competent to probe the substrate and settle. Despite offering settlement substrate to larvae at two different points in time (Cohort 1, 7 day-old larvae vs. Cohort 2, 17 day-old larvae), both cohorts began substrate probing and settlement around a similar time at days 15 and 19, respectively (Fig. 7 ). While studies documenting settlement dynamics in temperate and cold-water coral species remain sparse, especially in species that broadcast spawn, such a prolonged delay in the onset of competency to settle has been previously documented in a deep-sea and temperate octocorals (Rakka et al. 2021; Zelli et al. 2020). Estimating the onset of settlement competency in suboptimal ex situ conditions compared to the wild remains challenging, and we cannot exclude that our estimate is potentially an overestimate of the duration of the pre-competency period in nature. However, our data seemed remarkably consistent across two different larval cohorts used in the settlement experiments conducted at CCMAR and an additional cohort followed in the entirely separate system at t ODL, which suggests delayed onset of settlement may be common in temperate and cold-water broadcast spawners presumably due to a longer larval development period as a result at low ambient seawater temperature. One factor that determines behaviour and settlement of the larvae is the potential survival of the larvae in the water under laboratory conditions (PLD), which can be quite long with some species. For example, Corallium rubrum larvae can survive for up to 42 days (Martínez-Quintana et al. 2015), P. clavata maximal 64 days and E. singularis > 78 days (Guizien et al. 2020). (von Koch 1887) also observed that E. cavolini larvae could remain in the water column for months under water flow. In contrast to such previous studies on temperate and cold-water octocorals, our observations for E. verrucosa showed that despite overall low settlement rates, larvae remained competent to settle for a relatively long period, with a maximum PLD of up to 132 days under laboratory conditions. For example, larvae of the Mediterranean internal brooder E. singularis can settle within 13 days (Theodor 1967; Weinberg & Weinberg 1979; Zelli et al. 2020), and P. clavata have been observed to metamorphose into polyps without attaching to the substratum between 8–25 days (Linares et al. 2008). For E. verrucosa , we only recorded the total number of settlers at any given monitoring time without tracking individual recruits, so there may have been an underestimation of settlement if some settlers died unnoticed. In addition, our observation that E. verrucosa has an extremely long PLD is concordant with the mode of reproduction. Broadcast spawners typically have longer PLDs than brooding species (Harrison & Wallace 1990). Furthermore, it is in accordance with the patterns of embryonic cleavage reported here, suggesting extremely yolk-rich propagules that may enable a long settlement competency period. Coral larvae settlement is known to be influenced by various complex environmental and biological cues. These cues are not yet fully understood, and knowledge is predominantly focused on tropical shallow-water coral species. These cues can be chemical or physical signals, with chemical compounds produced by crustose coralline algae (CCA) and/or the microbial communities and biofilms associated with the substrate having previously shown to induce settlement in multiple species of corals (Gómez-Lemos et al. 2018; Sneed et al. 2014; Tebben et al. 2015). We therefore selected substrates previously shown to encourage coral settlement in temperate octocorals: gorgonian bare skeleton for E. singularis (Weinberg & Weinberg 1979), CCA covered rocks for C. rubrum (Zelli et al. 2020) and rocks from the same area as the breeding stock used in our experiments. However, no clear preference for any substrate was observed, only trends that shifted with larval age (Fig. 9 ). The hierarchical induction of settlement through certain cues is subject to a number of recent studies (Jorissen et al. 2021; Petersen et al. 2023; Wahab et al. 2023). Some CCA species may even provide positive microbial cues that initiate the settlement process, but then also deter attachment and metamorphosis, leading larvae to choose alternative substrates (Jorissen et al. 2021). E. verrucosa does not appear to be limited to a particular substrate type, and despite intense probing behaviour, the larvae did not avoid any of the three substrate types provided. Initially, however, the larvae of both cohorts seemed to show a preference for the gorgonian skeleton. Later, the majority of settlers chose to settle on rock, with fewer settling on the gorgonian skeleton and CCA. The gorgonian skeleton may have provided strong cues at first, but these cues might have faded, or the larvae may have determined, after initial attachment and mechanical testing, that the material quality was insufficient for settlement. The lower number of settlers on the gorgonian skeleton and CCA could also be due to post-settlement detachment of recruits. For example, the gorgonian skeleton develops a slimy biofilm, and living CCA has defence mechanisms to avoid overgrowth, such as chemical defences and the sloughing off from epithelial cells (Gómez-Lemos & Díaz-Pulido 2017). The rock, on the other hand, was not bare but covered with a diverse array of microorganisms. Therefore, the settlement cue was likely a combination of abiotic factors (surface structure) and biotic cues (microfilm and bioactive molecules produced by organisms). Since we did not test the substrates individually, we cannot rule out cross-influences among them, and future experiments are necessary to further investigate settlement cues. Once larvae settled and started metamorphosis, survival rates were high, but growth was slow (2-3mm in length in the first 3 months, budding after 9 months post settlement) compared with the description of von Koch (1887) on E. cavolini who observed the formation of an internal skeleton already after several weeks and budding shortly after. Implications on Larval dispersal and connectivity Incorporating population dynamics and connectivity patterns into marine conservation and restoration strategies, especially for sessile invertebrates like corals, is critical for effective biodiversity preservation (Jones et al. 2007; Marti-Puig et al. 2013; Possingham et al. 2015). The primary goal of coral restoration efforts is to rehabilitate a degraded habitat by reintroducing a healthy, self-sustaining breeding population, based on a robust, scientifically grounded restoration framework. However, such frameworks are largely absent in most coral restoration projects (Boström-Einarsson et al. 2020; Mcdonald et al. 2016). For instance, dispersal potential and patterns of population connectivity are crucial considerations for conservation, yet key biological traits that influence larval dispersal such as planktonic larval duration (PLD), larval behaviour (e.g., buoyancy and swimming), and reproductive strategies (brooding vs. spawning), are often unknown or remain overlooked in restoration planning and implementation (Coelho & Lasker 2016b; Cowen et al. 2007; Cowen & Sponaugle 2009; Randall et al. 2020; Suggett & Van Oppen 2022). Our findings reveal that the PLD of E. verrucosa is exceptionally long as is the settlement competency period, which started late (earliest at day 15). This suggests that coral larvae can disperse over a large distance, which has important implications for population connectivity. However, genetic studies by Holland et al. (2017) detected significant genetic structure at regional scale suggesting that dispersal over 500-2000km is infrequent. And the more recent genome-wide analyses by (Macleod et al. 2024) further highlighted regional clustering and a pattern of isolation by distance. In a subsequent analysis, Macleod et al. (2024) modeled larval dispersal under two hypothetical PLDs (14 and 21 days) due to the absence of data for the species and to represent the central ranges of PLDs observed in other octocorals. These assumptions deviate considerably from our observations under laboratory conditions. While the effective PLD of E. verrucosa in the field may be lower than that observed in the laboratory due to larval mortality (e.g., predation, drift to unsuitable areas) (Sciascia et al. 2022), settling competency most likely only starts above 15 days of age and has to potential to extend up to 4 months. Thus, the estimated mean and maximum dispersal distances obtained by Macleod et al. (2024) likely underestimate dispersal potential and the frequency of ‘rare’ dispersal events, thus highlighting the shortcomings of generalizing biological traits in modelling studies of dispersal and population connectivity that may be used for informing conservation practitioners. First steps into restoration through sexual propagation While data on reproductive and early life ecological traits are crucial for a better understanding of the biology and population dynamics of E. verrucosa in particular, and corals in general, this information is also the base for our overall goal: developing methods and workflows for restoration through sexual propagation. Coral collection and rearing of parent colonies While numerous approaches have been developed to obtain coral offspring from tropical scleractinian corals (see Randall et al. 2020 for an overview), methods for octocorals and especially cold- or deep-water coral species remain limited (Fava et al. 2010; Montseny et al. 2021a). On-site collection is only feasible for species with predictable spawning times and those located at dive-accessible depths. Obtaining offspring from aquarium-kept corals caught as fisheries bycatch offers the advantage of accessing large biomass that would otherwise perish, resulting in a substantial spawning output and diverse genotypes, providing a good representation of the population (Montseny et al. 2021b). However, relying on destructive fishing-techniques is unsustainable and should generally not be encouraged as the impact of fishing practices should be mitigated and/or removed. The collection of parent corals from bycatch is opportunistic, dependent on the availability, timing, location, and depth of fishing activities, all of which are uncontrollable factors. Additionally, the condition of the colonies can vary, as they are often entangled and damaged in nets, experiencing high stress levels that may negatively affect reproductive output and gamete nutritional reserves. However, we view coral bycatch as a valuable opportunity to establish a large-scale coral nursery that could be maintained for many years, providing a consistent annual supply of offspring. Although the corals at CCMAR were kept under less controlled conditions of lighting and temperatures, and other factors such as feeding with frozen food, several colonies produced eggs the following year, indicating the persistence of their gametogenic cycle despite being kept in suboptimal conditions. In contrast, at the Oceanário de Lisboa, the breeding colonies were maintained over 1–2 years in a closed system with precise management of water quality, high-quality food, and regulation of key environmental parameters such as moonlight and temperature. This led not only to the completion of the reproductive cycle but also to spawning, settlement, and the development of healthy recruits. This demonstrates the potential for ex-situ sexual reproduction of E. verrucosa , eliminating the need to collect adult corals annually, which is crucial in the event of limited bycatch availability or depletion of wild populations due to anthropogenic or natural disasters. Although propagule release and/or capture efficiency was higher at Oceanário (~ 5500 eggs from 15 colonies) compared to the Ramalhete Marine Station (~ 3000 eggs from 70 colonies), settlement success was significantly greater at Ramalhete. This discrepancy may be attributed to less suitable substrates or less competent larvae at Oceanário, possibly due to suboptimal maintenance conditions and insufficient energy reserves in the cultured colonies, as opposed to the fresh, wild-collected colonies used for the experiemnts conducted at CCMAR. This suggests that keeping colonies in captivity in the long-term may not yet be the most-efficient restoration strategy. Future studies should therefore focus on improving reproductive output, enhancing settlement success, and increasing recruit survival. This could be achieved by boosting nutrient input for parent colonies, providing adequate space for individuals, optimizing temperature control, and adjusting lighting to better replicate their natural environment year-round—measures likely to improve both adult survival and offspring production. Meeting the nutritional needs of deep and cold-water corals remains a major challenge, given their reliance on external food sources. Although this has been the focus of several studies (Cocito et al. 2013; Gori et al. 2012; Rakka et al. 2021; Ribes et al. 1999), optimal feeding regimes have yet to be developed. Reproduction, settlement and survival of recruits in captivity The broadcast spawning of eggs and their release over an extended period in E. verrucosa offers significant advantages for restoration efforts. Broadcast spawning is a mode of reproduction that is far more feasible for egg collection and rearing, whereas brooding species are more challenging in this regard (Randall et al. 2020). We collected spawn using air-driven suction traps positioned in the tank, by filters in the outflow, and manually retrieved eggs with a pipette. Although all methods worked similarly well, pipetting proved to be highly time-consuming, whereas a portion of the eggs or embryos collected in the filters were observed to have been squeezed and damaged. Additionally, we were unable to quantify the number of eggs potentially lost due to contact with the walls or air bubbles. The trade-off between methods highly depends on the availability of human resources; however, filters and traps, particularly with enhancements, proved highly effective during spawning events when no human presence was available such as during night-time release. In many species, the limitation of spawning events to just one or a few nights per year represents a major bottleneck (Randall et al. 2020). However, with E. verrucosa , the prolonged release over more than a month allows for more flexible human and facility resources. Investigating the drivers behind E. verrucosa 's spawning synchronicity would further enhance the predictability and management of spawn collection. However, this may require several more years of observation and experimenting. During embryo and larval rearing, maintaining the larvae in large plastic containers (5–10 L) with gentle aeration seemed to result in good survival rates. We observed larval settlement of up to 16% (Fig. 10 ), which, while lower than the much higher settlement rates reported for tropical corals (Jorissen et al. 2021; Tebben et al. 2015), represents a successful settlement outcomes for the first trials with this species. Settlement rates in cold- and deep-water corals are generally much lower across the up-to-date studied species, and comparative analyses are challenging due to the limited reports on settlement success or data on settlement dynamics (Linares et al. 2008; Rakka et al. 2021; Weinberg & Weinberg 1979). An exception is the documented high settlement rate of up to 50 ± 7.8% in the temperate gorgonian E. singularis , although it is a shallow-water species, under laboratory conditions (Viladrich et al. 2022). Studies like Zelli et al. (2020), highlight the great potential that lies in widening our understanding on settlement cues also in temperate and cold-water corals. Settlement was the bottleneck in this study and enhancement could greatly benefit not only the reproductive success of E. verrucosa in captivity but also other threatened cold-water coral species. Once the larvae settled and metamorphosed, survival rates were high over a period of 6–7 months in comparison to the recruits of P. clavata in the field, which were close to zero (Linares et al. 2008). Growth, however, was slow as it took 6.5–12 months to develop the secondary polyps. For E. singularis studied under lab conditions, for example, growth has been reported to be much faster, with budding observed after just two weeks (Weinberg & Weinberg 1979). Following the settlement experiments conducted at CCMAR, we did not provide the recruits with any food beyond what was available to feed adult colonies (e.g., frozen rotifers, copepods). The feeding regime for recruits kept at ODL, which included living microalgae and zooplankton, seemed to positively affect growth, with the first budding observed only after 6.5 months (compared to that observed at CCMAR at 11–12 months after spawning). This underscores the importance of further research into optimal nourishment for recruits, as studies on tropical coral larvae have shown that improved feeding strategies can significantly boost early growth (Petersen et al. 2008; Toh et al. 2014). Substrate type also appeared to play a crucial role in recruit survival. Natural rock substrates generally proved to be suitable for settlement, in particular those with numerous crevices and holes. On the other hand, surface structures tended to accumulate debris, which likely negatively affected recruit survival. Nearly all surviving recruits were observed growing on smooth, exposed surfaces that remained free of debris and received adequate water flow for self-cleaning, emphasizing the importance of current-exposed, sediment-free surfaces for successful coral settlement and growth. CONCLUSIONS This study provides the first description of the spawning and early life ecology of the widely distributed, forest-forming, and vulnerable pink sea fan E. verrucosa . This information is vital for managing, protecting, and restoring habitats dominated by this valuable species, also providing key data for dispersal and connectivity models. Our findings revealed late onset of competency to settle and an extended period over which larvae remained competent to settle (up to 3–4 months). Our results on settlement and recruit survival also represent early steps toward breeding sexually derived corals for restoration. We demonstrate that larvae can be successfully settled onto natural substrate, recruits survive and grow under laboratory conditions, and adult colonies can be maintained in captivity through multiple reproductive cycles, completing gamete development and spawning thus allowing for the production of sexually-derived recruits ex situ . This study also marks the first steps toward cold-water coral breeding for this vulnerable species, laying the groundwork for establishing coral nurseries for restoration. Working with such sensitive organisms presents significant challenges in planning, so our approach prioritized flexibility, allowing us to adapt our strategy continuously. As a result, the data collected may not always be as complete or systematic as desired. Nonetheless, the findings of this study pave the way for broader research and restoration efforts. A key aspect of the study was the collaboration between a scientific research centre (CCMAR) and a major facility aimed at public visitors and education, the Oceanário de Lisboa, highlighting the potential of such institutional collaborations as platforms for large-scale restoration initiatives. Declarations ACKNOWLEDGEMENTS This work was funded by the 2020–2021 Biodiversa+ and Water JPI joint call for research projects, under the Bio-divRestore ERA-NET Cofund (GA N°101003777), with the EU and the Portuguese Foundation for Science and Technology (FCT) (DivRestore/0013/2020); and by project CORALFORESTS financed by the 2022 Fundação Belmiro de Azevedo Award in the field of Conservation, restoration and monitoring of biodiversity in Portugal. This study also received Portuguese national funds from FCT through projects UIDB/04326/2020 (DOI:10.54499/UIDB/04326/2020) and LA/P/0101/2020 (DOI:10.54499/LA/P/0101/2020). C. Egger was supported by FCT Doctoral Scholarship SFRH/BD/151455/2021. We thank Capucine Récasens and Laura Balsalobre for their help examining gamete development, particularly polyp dissections and histological sectioning, as well as Elsa Couto and Deborah Power for technical and laboratory support with the histological examinations. We also thank João Reis and André Lopes for their technical support at the Ramalhete Marine Station. We are especially grateful to master Casimiro and his fishing crew for the collaboration and help collecting samples for this study along the Sagres region in SW Iberia. SUPPLEMENTARY INFORMATION Supplement 1: Polyp dissection to infer spawning window: https://doi.org/10.6084/m9.figshare.28113371 Supplement 2 (video): Eunicella verrucosa time lapse video of embryonal development: https://doi.org/10.6084/m9.figshare.28070732 Supplement 3 (video): Eunicella verrucosa larvae swimming behaviour: https://doi.org/10.6084/m9.figshare.28070870 Supplement 4 (video): Eunicella verrucosa larvae testing substrate and attaching: https://doi.org/10.6084/m9.figshare.28072046 Conflict of interest On behalf of all authors, the corresponding author states that there is no conflict of interest. Author information Christina Egger, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal Catarina Melo, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal Bailey Marquardt, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal Aschwin H. Engelen, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal Roland R. Melzer, SNSB-Bavarian State Collection for Zoology, Munich, Germany Elsa Santos, Oceanário de Lisboa, Lisboa, Portugal Margarida Fernandes, Oceanário de Lisboa, Lisboa, Portugal Núria Baylina, Oceanário de Lisboa, Lisboa, Portugal Ester A. Serrao, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal Márcio A. Coelho, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal References Adamska, M., Degnan, B. M., Green, K., & Zwafink, C. (2011). What sponges can tell us about the evolution of developmental processes. Zoology, 114(1), 1–10. https://doi.org/10.1016/j.zool.2010.10.003. Ankamah-Yeboah, I., Xuan, B. B., Hynes, S., & Armstrong, C. W. (2020). Public perceptions of deep-sea environment: Evidence from Scotland and Norway. Frontiers in Marine Science, 7, 137. https://doi.org/10.3389/fmars.2020.00137. Baillon, S., Hamel, J.-F., Wareham, V. E., & Mercier, A. (2012). Deep cold-water corals as nurseries for fish larvae. Frontiers in Ecology and the Environment, 10(7), 351–356. Baums, I. B. (2008). A restoration genetics guide for coral reef conservation. Molecular Ecology, 17(12), 2796–2811. https://doi.org/10.1111/j.1365-294X.2008.03787.x. Baums, I. B., Baker, A. C., Davies, S. W., Grottoli, A. G., Kenkel, C. D., Kitchen, S. A., Kuffner, I. B., LaJeunesse, T. C., Matz, M. V., & Miller, M. W. (2019). Considerations for maximizing the adaptive potential of restored coral populations in the western Atlantic. Ecological Applications, 29(8), e01978. https://doi.org/10.1002/eap.1978. Bayraktarov, E., Brisbane, S., Hagger, V., Smith, C. S., Wilson, K. A., Lovelock, C. E., Gillies, C. L., Steven, A. D. L., & Saunders, M. I. (2020). Priorities and Motivations of Marine Coastal Restoration Research. Frontiers in Marine Science, 7. https://doi.org/10.3389/fmars.2020.00484. Bayraktarov, E., Stewart-Sinclair, P. J., Brisbane, S., Boström‐Einarsson, L., Saunders, M. I., Lovelock, C. E., Possingham, H. P., Mumby, P. J., & Wilson, K. A. (2019). Motivations, Success, and Cost of Coral Reef Restoration. Restoration Ecology, 27(5), 981–991. https://doi.org/10.1111/rec.12977. Boch, C. A., DeVogelaere, A., Burton, E., King, C., Lord, J. P., Lovera, C., Litvin, S. Y., Kuhnz, L. A., & Barry, J. (2019). Coral Translocation as a Method to Restore Impacted Deep-Sea Coral Communities. Frontiers in Marine Science, 6. https://doi.org/10.3389/fmars.2019.00540. Boch, C. A., & Morse, A. N. C. (2012). Testing the effectiveness of direct propagation techniques for coral restoration of Acropora spp. Ecological Engineering, 40, 11–17. https://www.sciencedirect.com/science/article/pii/S0925857411003934. Bongiorni, L., Mea, M., Gambi, C., Pusceddu, A., Taviani, M., & Danovaro, R. (2010). Deep-Water Scleractinian Corals Promote Higher Biodiversity in Deep-Sea Meiofaunal Assemblages Along Continental Margins. Biological conservation, 143(7), 1687–1700. https://doi.org/10.1016/j.biocon.2010.04.009. Boström-Einarsson, L., Babcock, R. C., Bayraktarov, E., Ceccarelli, D., Cook, N., Ferse, S. C. A., Hancock, B., Harrison, P., Hein, M., Shaver, E., Smith, A., Suggett, D., Stewart-Sinclair, P. J., Vardi, T., & McLeod, I. M. (2020). Coral restoration – A systematic review of current methods, successes, failures and future directions. PLoS One, 15(1). https://doi.org/10.1371/journal.pone.0226631. Brazeau, D. A., & Lasker, H. R. (1988). Inter-and intraspecific variation in gorgonian colony morphology: quantifying branching patterns in arborescent animals. Coral Reefs, 7, 139–143. https://doi.org/10.1007/BF00300973. Bridge, T. C. L., Hoey, A. S., Campbell, S. J., Muttaqin, E., Rudi, E., Fadli, N., & Baird, A. H. (2013). Depth-Dependent Mortality of Reef Corals Following a Severe Bleaching Event: Implications for Thermal Refuges and Population Recovery. F1000research, 2, 187. https://doi.org/10.12688/f1000research.2-187.v1. Brun-Usan, M., & Salazar-Ciudada, I. (2020). The Evolution of Cleavage in Metazoans. In L. Nuno de la Rosa & G. B. Müller (Eds.), Evolutionary Developmental Biology: Springer Nature Switzerland AG. Buhl-Mortensen, L., Buhl-Mortensen, P., Rungruangsak-Torrissen, K., Schwach, V., Hjort, J., Jakobsen, T., Ozhigin, V., Bergh, Ø., Hamre, J., & Torgersen, T. (2018). Cold temperate coral habitats. In C. D. Beltran & E. T. Camacho (Eds.), Corals in a changing world (Vol. 9). Rijeka, Croatia: InTech. Buhl-Mortensen, P., Buhl-Mortensen, L., & Purser, A. (2017). Trophic ecology and habitat provision in cold-water coral ecosystems. Marine Animal Forests. The Ecology of Benthic Biodiversity Hotspots. Springer, Cham. https://doi.org/10.1007/978-3-319-21012-4_20. Carpenter, K. E., Abrar, M., Aeby, G. S., Aronson, R. B., Banks, S., Bruckner, A. W., Chiriboga, A., Cortés, J., Delbeek, J. C., DeVantier, L., Edgar, G. J., Edwards, A. J., Fenner, D., Guzmán, H. M., Hoeksema, B. W., Hodgson, G., Johan, O., Licuanan, W. Y., Livingstone, S. R., Lovell, E. R., Moore, J. A., Obura, D., Ochavillo, D., Polidoro, B., Precht, W. F., Quibilan, M. C. C., Reboton, C. T., Richards, Z. T., Rogers, A. D., Sanciangco, J. C., Sheppard, A., Sheppard, C., Smith, J. E., Stuart, S. N., Turak, E., Veron, J., Wallace, C. C., Weil, E., & Wood, E. (2008). One-Third of Reef-Building Corals Face Elevated Extinction Risk From Climate Change and Local Impacts. Science, 321(58888), 560–563. https://doi.org/10.1126/science.1159196. Carpine, C. (1963). Contribution à la connaissance des Gorgones Holaxonia de la Mediterranean occidentale. Bull. Inst. océanogr. Monaco, 60(1270), 1–52. Carpine, C., & Grasshoff, M. (1975). Les gorgonaires de la Méditerranée. Bull. Inst. océanogr. Monaco, 71(1430), 1-140. Chamberland, V. F., Vermeij, M. J., Brittsan, M., Carl, M., Schick, M., Snowden, S., Schrier, A., & Petersen, D. (2015). Restoration of critically endangered elkhorn coral (Acropora palmata) populations using larvae reared from wild-caught gametes. Global Ecology and Conservation, 4, 526–537. https://doi.org/10.1016/j.gecco.2015.10.005. Chimienti, G. (2020). Vulnerable forests of the pink sea fan Eunicella verrucosa in the Mediterranean Sea. Diversity, 12(5), 176. https://doi.org/10.3390/d12050176. Cocito, S., Ferrier-Pagès, C., Cupido, R., Rottier, C., Meier-Augenstein, W., Kemp, H., Reynaud, S., & Peirano, A. (2013). Nutrient acquisition in four Mediterranean gorgonian species. Marine Ecology Progress Series, 473, 179–188. https://doi.org/10.3354/meps10037. Coelho, M. A., & Lasker, H. R. (2016a). Larval behavior and settlement dynamics of a ubiquitous Caribbean octocoral and its implications for dispersal. Marine Ecology Progress Series, 561, 109–121. https://doi.org/10.3354/meps11941. Coelho, M. A., & Lasker, H. R. (2016b). Larval dispersal and population connectivity in Anthozoans. In S. Goffredo & Z. Dubinsky (Eds.), The Cnidaria, Past, Present and Future: The world of Medusa and her sisters (pp. 291–315): Springer. Coma, R., Ribes, M., Zabala, M., & Gilil, J. (1995a). Reproduction and cycle of gonadal development in the Mediterranean gorgonian Paramuricea clavata. Mar. Ecol. Prog. Ser, 117(1–3), 173–183. https://doi.org/10.3354/meps117173. Coma, R., Zabala, M., & Gili, J. (1995b). Sexual reproductive effort in the Mediterranean gorgonian Paramuricea clavata. Marine Ecology Progress Series, 185–192. https://www.jstor.org/stable/44634830. Cowen, R. K., Gawarkiewicz, G., Pineda, J., Thorrold, S. R., & Werner, F. E. (2007). Population connectivity in marine systems an overview. Oceanography, 20(3), 14–21. https://doi.org/10.5670/oceanog.2007.26. Cowen, R. K., & Sponaugle, S. (2009). Larval dispersal and marine population connectivity. Annual review of marine science, 1(1), 443–466. https://doi.org/10.1146/annurev.marine.010908.163757. Coz, R., Ouisse, V., Artero, C., Carpentier, A., Crave, A., Feunteun, E., Olivier, J.-M., Perrin, B., & Ysnel, F. (2012). Development of a new standardised method for sustainable monitoring of the vulnerable pink sea fan Eunicella verrucosa. Marine Biology, 159, 1375–1388. https://doi.org/10.1007/s00227-012-1908-7. Danovaro, R., Corinaldesi, C., D’Onghia, G., Galil, B. S., Gambi, C., Gooday, A. J., Lampadariou, N., Luna, G. M., Morigi, C., Olu, K., Polymenakou, P. N., Ramirez-Llodra, E., Sabbatini, A., Sardà, F., Sibuet, M., & Τσελεπίδης, Α. (2010). Deep-Sea Biodiversity in the Mediterranean Sea: The Known, the Unknown, and the Unknowable. PLoS One, 5(8). https://doi.org/10.1371/journal.pone.0011832. Dela Cruz, D. W., & Harrison, P. L. (2020). Enhancing coral recruitment through assisted mass settlement of cultured coral larvae. PLoS One, 15(11), e0242847. https://doi.org/10.1371/journal.pone.0242847. Dias, V., Oliveira, F., Boavida, J., Serrão, E. A., Gonçalves, J. M., & Coelho, M. A. (2020). High coral bycatch in bottom-set gillnet coastal fisheries reveals rich coral habitats in southern Portugal. Frontiers in Marine Science, 7, 603438. https://doi.org/10.3389/fmars.2020.603438. Doropoulos, C., Vons, F., Elzinga, J., Ter Hofstede, R., Salee, K., Van Koningsveld, M., & Babcock, R. C. (2019). Testing industrial-scale coral restoration techniques: harvesting and culturing wild coral-spawn slicks. Frontiers in Marine Science, 6, 658. https://doi.org/10.3389/fmars.2019.00658. Eckelbarger, K., Tyler, P., & Langton, R. (1998). Gonadal morphology and gametogenesis in the sea pen Pennatula aculeata (Anthozoa: Pennatulacea) from the Gulf of Maine. Marine Biology, 132, 677–690. https://doi.org/10.1007/s002270050432. Eddy, T. D., Lam, V. W. Y., Reygondeau, G., Cisneros-Montemayor, A. M., Greer, K., Palomares, M. L. D., Bruno, J. F., Ota, Y., & Cheung, W. W. L. (2021). Global Decline in Capacity of Coral Reefs to Provide Ecosystem Services. One Earth, 4(9), 1278–1285. https://doi.org/10.1016/j.oneear.2021.08.016. Fava, F., Bavestrello, G., Valisano, L., & Cerrano, C. (2010). Survival, growth and regeneration in explants of four temperate gorgonian species in the Mediterranean Sea. Italian Journal of Zoology, 77, 44–52. https://doi.org/10.1080/11250000902769680. Feng, E. Y., Keller, D. P., Koeve, W., & Oschlies, A. (2016). Could Artificial Ocean Alkalinization Protect Tropical Coral Ecosystems From Ocean Acidification? Environmental Research Letters, 11(7). https://doi.org/10.1088/1748-9326/11/7/074008. Ferrario, F., Beck, M. W., Storlazzi, C. D., Micheli, F., Shepard, C. C., & Airoldi, L. (2014). The Effectiveness of Coral Reefs for Coastal Hazard Risk Reduction and Adaptation. Nature Communications, 5(1). https://doi.org/10.1038/ncomms4794. Figueiredo, J., Baird, A. H., Cohen, M. F., Flot, J.-F., Kamiki, T., Meziane, T., Tsuchiya, M., & Yamasaki, H. (2012). Ontogenetic change in the lipid and fatty acid composition of scleractinian coral larvae. Coral Reefs, 31, 613–619. https://doi.org/10.1007/s00338-012-0874-3. Fogarty, M. J., & Botsford, L. W. (2007). Population connectivity and spatial management of marine fisheries. Oceanography, 20(3), 112–123. http://dx.doi.org/10.5670/oceanog.2007.34. Forrester, G. E., Chan, M., Conetta, D., Dauksis, R., Nickles, K., & Siravo, A. (2019). Comparing the efficiency of nursery and direct transplanting methods for restoring endangered corals. Ecological Restoration, 37(2), 81–89. https://doi.org/10.3368/er.37.2.81. Foster, T., & Gilmour, J. (2020). Egg Size and Fecundity of Biannually Spawning Corals at Scott Reef. Scientific Reports, 10(1). https://doi.org/10.1038/s41598-020-68289-4. Garrison, V., & Greg, W. (2008). Storm-generated coral fragments–A viable source of transplants for reef rehabilitation. Biological conservation, 141(12), 3089–3100. https://doi.org/10.1016/j.biocon.2008.09.020. Gilmour, J. P., Underwood, J. N., Howells, E. J., Gates, E., & Heyward, A. J. (2016). Biannual spawning and temporal reproductive isolation in Acropora corals. PLoS One, 11(3), e0150916. Gómez-Lemos, L. A., & Díaz-Pulido, G. (2017). Crustose Coralline Algae and Associated Microbial Biofilms Deter Seaweed Settlement on Coral Reefs. Coral Reefs, 36(2), 453–462. https://doi.org/10.1007/s00338-017-1549-x. Gómez-Lemos, L. A., Doropoulos, C., Bayraktarov, E., & Diaz-Pulido, G. (2018). Coralline algal metabolites induce settlement and mediate the inductive effect of epiphytic microbes on coral larvae. Scientific Reports, 8(1), 17557. https://doi.org/10.1038/s41598-018-35206-9. Gori, A., Viladrich, N., Gili, J. M., Kotta, M., Cucio, C., Magni, L., Bramanti, L., & Rossi, S. (2012). Reproductive cycle and trophic ecology in deep versus shallow populations of the Mediterranean gorgonian Eunicella singularis (Cap de Creus, northwestern Mediterranean Sea). Coral Reefs, 31, 823–837. https://doi.org/10.1007/s00338-012-0904-1. Grasshoff, M. (1992). Die Flachwasser-Gorgonarien von Europa und Westafrika-(Cnidaria, Anthozoa) (Vol. 149). Frankfurt a. M.: Senckenbergischen Naturforschenden Gesellschaft Frankfurt a. M. Guizien, K., Viladrich, N., Martínez-Quintana, Á., & Bramanti, L. (2020). Survive or swim: different relationships between migration potential and larval size in three sympatric Mediterranean octocorals. Scientific Reports, 10(1), 18096. https://doi.org/10.1038/s41598-020-75099-1. Hall-Spencer, J. M., Pike, J., & Munn, C. B. (2007). Diseases affect cold-water corals too: Eunicella verrucosa (Cnidaria: Gorgonacea) necrosis in SW England. Diseases of aquatic organisms, 76(2), 87–97. https://doi.org/10.3354/dao076087. Harii, S., Nadaoka, K., Yamamoto, M., & Iwao, K. (2007). Temporal changes in settlement, lipid content and lipid composition of larvae of the spawning hermatypic coral Acropora tenuis. Marine Ecology Progress Series, 346, 89–96. https://doi.org/10.3354/meps07114. Harrison, P. L., & Wallace, C. C. (1990). Reproduction, dispersal and recruitment of scleractinian corals. In Coral Reefs (Vol. 25, pp. 133–207): Elsevier. Hatta, M., Fukami, H., Wang, W., Omori, M., Shimoike, K., Hayashibara, T., Ina, Y., & Sugiyama, T. (1999). Reproductive and genetic evidence for a reticulate evolutionary history of mass-spawning corals. Molecular Biology and Evolution, 16(11), 1607–1613. https://doi.org/10.1093/oxfordjournals.molbev.a026073. Henry, J. A., O’Neil, K. L., Pilnick, A. R., & Patterson, J. T. (2021). Strategies for integrating sexually propagated corals into Caribbean reef restoration: experimental results and considerations. Coral Reefs, 40(5), 1667–1677. https://doi.org/10.1007/s00338-021-02154-2. Heyward, A., Smith, L., Rees, M., & Field, S. (2002). Enhancement of coral recruitment by in situ mass culture of coral larvae. Marine Ecology Progress Series, 230, 113–118. https://doi.org/10.3354/MEPS230113. Høegh-Guldberg, O., Kennedy, E. V., Beyer, H. L., McClennen, C., & Possingham, H. P. (2018). Securing a Long-Term Future for Coral Reefs. Trends in Ecology & Evolution, 33(12), 936–944. https://doi.org/10.1016/j.tree.2018.09.006. Holland, L., Jenkins, T., & Stevens, J. (2017). Contrasting patterns of population structure and gene flow facilitate exploration of connectivity in two widely distributed temperate octocorals. Heredity, 119(1), 35–48. https://doi.org/10.1038/hdy.2017.14. Hughes, A. R., Grabowski, J. H., Leslie, H. M., Scyphers, S. B., & Williams, S. L. (2017a). Inclusion of Biodiversity in Habitat Restoration Policy to Facilitate Ecosystem Recovery. Conservation Letters, 11(3). https://doi.org/10.1111/conl.12419. Hughes, T. P., Barnes, M. L., Bellwood, D. R., Cinner, J. E., Cumming, G. S., Jackson, J. B. C., Kleypas, J., Leemput, I. A. v. d., Lough, J., Morrison, T. H., Palumbi, S. R., Nes, E. H. v., & Scheffer, M. (2017b). Coral Reefs in the Anthropocene. Nature, 546(7656), 82–90. https://doi.org/10.1038/nature22901. Hughes, T. P., Kerry, J. T., Baird, A. H., Connolly, S. R., Dietzel, A., Eakin, C. M., Heron, S. F., Hoey, A. S., Hoogenboom, M. O., & Liu, G. (2018). Global warming transforms coral reef assemblages. Nature, 556(7702), 492–496. https://doi.org/10.1038/s41586-018-0041-2. Jones, G. P., Srinivasan, M., & Almany, G. R. (2007). Population connectivity and conservation of marine biodiversity. Oceanography, 20(3), 100–111. https://doi.org/10.5670/oceanog.2007.33. Jorissen, H., Galand, P. E., Bonnard, I., Meiling, S. S., Raviglione, D., Meistertzheim, A.-L., Hédouin, L., Banaigs, B., Payri, C., & Nugues, M. M. (2021). Coral Larval Settlement Preferences Linked to Crustose Coralline Algae With Distinct Chemical and Microbial Signatures. Scientific Reports, 11(1). https://doi.org/10.1038/s41598-021-94096-6. Kaartvedt, S., Langbehn, T. J., & Aksnes, D. L. (2019). Enlightening the ocean’s twilight zone. ICES Journal of Marine Science, 76(4), 803–812. https://doi.org/10.1093/icesjms/fsz010. Kahng, S. E., Benayahu, Y., & Lasker, H. R. (2011). Sexual reproduction in octocorals. Marine Ecology Progress Series, 443, 265–283. https://doi.org/10.3354/meps09414. Kaniewska, P., Alon, S., Karako-Lampert, S., Hoegh-Guldberg, O., & Levy, O. (2015). Signaling cascades and the importance of moonlight in coral broadcast mass spawning. Elife, 4. https://doi.org/10.7554/eLife.09991. Kerr, A. M., Baird, A. H., & Hughes, T. P. (2011). Correlated evolution of sex and reproductive mode in corals (Anthozoa: Scleractinia). Proceedings of the Royal Society B: Biological Sciences, 278(1702), 75–81. Kersting, D. K., Casado, C., López-Legentil, S., & Linares, C. (2013). Unexpected Patterns in the Sexual Reproduction of the Mediterranean Scleractinian Coral Cladocora Caespitosa. Marine Ecology Progress Series, 486, 165–171. https://doi.org/10.3354/meps10356. Lam, K.-W., McRae, C. J., Zhang, X.-C., Ye, Z.-M., Qiu, Y.-T., Jiang, M.-Q., Cheng, T.-H., Chen, G. K., & Fan, T.-Y. (2023). Consistent monthly reproduction and completion of a brooding coral life cycle through ex situ culture. Diversity, 15(2), 218. https://doi.org/10.3390/d15020218. Lange, K., Maguer, J.-F., Reynaud, S., & Ferrier-Pagès, C. (2023). Nutritional Ecology of Temperate Octocorals in a Warming Ocean. Frontiers in Marine Science, 10. https://doi.org/10.3389/fmars.2023.1236164. Lasker, H. R., & Kim, K. (1996). Larval development and settlement behavior of the gorgonian coral Plexaura kuna (Lasker, Kim and Coffroth). Journal of Experimental Marine Biology and Ecology, 207(1), 161–175. https://doi.org/10.1016/S0022-0981(96)02625-1. Linares, C., Coma, R., Mariani, S., Díaz, D., Hereu, B., & Zabala, M. (2008). Early life history of the Mediterranean gorgonian Paramuricea clavata: implications for population dynamics. Invertebrate Biology, 127(1), 1–11. https://doi.org/10.HH/j.1744-7410.2007.00109.x. Macleod, K. L., Jenkins, T. L., Witt, M. J., & Stevens, J. R. (2024). Rare, long-distance dispersal underpins genetic connectivity in the pink sea fan, Eunicella verrucosa. Evolutionary Applications, 17(3). https://doi.org/10.1111/eva.13649. Marti-Puig, P., Costantini, F., Rugiu, L., Ponti, M., & Abbiati, M. (2013). Patterns of genetic connectivity in invertebrates of temperate MPA networks. Advances in Oceanography and Limnology, 4(2), 138–149. https://doi.org/10.4081/AIOL.2013.5341. Martin, V. (1997). Cnidarians, the jellyfish and hydras. In S. F. Gilbert & A. M. Raunio (Eds.), Embryology: construction the organism (pp. 57–86). Sunderland, MA: Sinauer Associates. Martínez-Quintana, A., Bramanti, L., Viladrich, N., Rossi, S., & Guizien, K. (2015). Quantification of larval traits driving connectivity: the case of Corallium rubrum (L. 1758). Marine Biology, 162, 309–318. https://doi.org/10.1007/s00227-014-2599-z. McDonald, T., Gann, G., Jonson, J., & Dixon, K. (2016). International standards for the practice of ecological restoration–including principles and key concepts.(Society for Ecological Restoration: Washington, DC, USA.). Soil-Tec, Inc.,© Marcel Huijser, Bethanie Walder. https://doi.org/10.1111/rec.13035. Melzer, R., Spitzner, F., Šargač, Z., Hörnig, M., Krieger, J., Haug, C., Haug, J., Kirchhoff, T., Meth, R., & Torres, G. (2021). Methods to study organogenesis in decapod crustacean larvae II: analysing cells and tissues. Helgoland Marine Research, 75(1), 2. https://doi.org/10.1186/s10152-021-00547-y. Mengerink, K. J., Van Dover, C. L., Ardron, J., Baker, M., Escobar-Briones, E., Gjerde, K., Koslow, J. A., Ramirez-Llodra, E., Lara-Lopez, A., & Squires, D. (2014). A call for deep-ocean stewardship. Science, 344(6185), 696–698. https://doi.org/10.1126/science.1251458. Menza, C. W., Kendall, M. S., & Hile, S. D. (2007). The Deeper We Go the Less We Know. Revista De Biología Tropical, 56(0). https://doi.org/10.15517/rbt.v56i0.5575. Mercado-Molina, A. E., & Suleimán-Ramos, S. E. (2023). Outplants of the Threatened Coral Acropora Cervicornis Promote Coral Recruitment in a Shallow-Water Coral Reef, Culebra, Puerto Rico. Sustainability, 15(24). https://doi.org/10.3390/su152416548. Monfared, M. A. A., Sheridan, K., Dixon, S. P., Gledhill, M., & Le Berre, T. (2023). Coral Spawning Patterns of Acropora Across Two Maldivian Reef Ecosystems. Peerj, 11, e16315. https://doi.org/10.7717/peerj.16315. Montseny, M., Linares, C., Carreiro-Silva, M., Henry, L.-A., Billett, D., Cordes, E., Smith, C., Papadopoulou, N., Bilan, M., Girard, F., Burdett, H., Larsson, A., Strömberg, S., Viladrich, N., Barry, J., Baena, P., Godinho, A., Grinyó, J., Santín, A., Morato, T., Sweetman, A., Gili, J., & Gori, A. (2021a). Active Ecological Restoration of Cold-Water Corals: Techniques, Challenges, Costs and Future Directions. Frontiers in Marine Science, 8. https://doi.org/10.3389/fmars.2021.621151. Montseny, M., Linares, C., Viladrich, N., Biel, M., Gracias, N., Baena, P., Quintanilla, E., Ambroso, S., Grinyó, J., Santín, A., Salazar, J., Carreras, M., Palomeras, N., Magí, L., Vallicrosa, G., Gili, J.-M., & Gori, A. (2021b). Involving fishers in scaling up the restoration of cold-water coral gardens on the Mediterranean continental shelf. Biological conservation, 262. https://www.sciencedirect.com/science/article/pii/S0006320721003530. Montseny, M., Linares, C., Viladrich, N., Capdevila, P., Ambroso, S., Díaz, D., Gili, J. M., & Gori, A. (2020). A new large-scale and cost‐effective restoration method for cold‐water coral gardens. Aquatic Conservation: Marine and Freshwater Ecosystems, 30(5), 977–987. https://doi.org/10.1002/aqc.3303. Munro, C., & Munro, L. (2003). Climate change impacts on seafan populations. Reef Research, RR Report 6/2003 RR 08. https://www.marine-bio-images.com/wp-content/uploads/2020/05/Report-RR-08-Jun-2003-Climate-pdf.pdf. Munro, L. (2004). Determining the reproductive cycle of Eunicella verrucosa. Reef Research, RR Report 3/2003 ETR 07. https://www.marine-bio-images.com/RR_Eunicella_PDFS/Report_RR12Jul2004reproductive%20cycle%20pdf.pdf. Najafpour, B., Dorafshan, S., Heyrati, F. P., Canário, A. V. M., & Power, D. M. (2020). Comparativeo ontogeny of the digestive tract of Oncorhynchus Mykiss Salmo Trutta caspius triploid hybrids to their Parental Species. Aquaculture Nutrition, 27(2), 427–438. https://doi.org/10.1111/anu.13196. O’Connor, E., Hynes, S., Chen, W., Papadopoulou, N., & Smith, C. J. (2020). Investigating Societal Attitudes Toward Marine Ecosystem Restoration. Restoration Ecology, 29(S2). https://doi.org/10.1111/rec.13239. O’Neil, K. L., Serafin, R. M., Patterson, J. T., & Craggs, J. R. (2021). Repeated ex situ spawning in two highly disease susceptible corals in the family Meandrinidae. Frontiers in Marine Science, 8, 669976. https://doi.org/10.3389/fmars.2021.669976. Okubo, N., Mezaki, T., Nozawa, Y., Nakano, Y., Lien, Y. T. K., Fukami, H., Hayward, D. C., & Ball, E. E. (2013). Comparative Embryology of Eleven Species of Stony Corals (Scleractinia). PLoS One, 8(12), e84115. Orth, R. J., Lefcheck, J. S., McGlathery, K., Aoki, L. R., Luckenbach, M. W., Moore, K. A., Oreska, M. P. J., Snyder, R. A., Wilcox, D. J., & Lusk, B. (2020). Restoration of Seagrass Habitat Leads to Rapid Recovery of Coastal Ecosystem Services. Science Advances, 6(41). https://doi.org/10.1126/sciadv.abc6434. Osman, E. O., Suggett, D. J., Attalla, T. M., Casartelli, M., Cook, N., El-Sadek, I., Gallab, A., Goergen, E. A., Garcias-Bonet, N., Glanz, J. S., Pereira, P. H., Ramirez-Sanchez, M., Santoro, E. P., Stead, A., Yoder, S., Benzoni, F., & Peixoto, R. S. (2024). Spatial Variation in Spawning Timing for Multi-Species Acropora Assemblages in the Red Sea. Frontiers in Marine Science, 11. https://doi.org/10.3389/fmars.2024.1333621. Ounanian, K., Carballo-Cárdenas, E. C., Tatenhove, J. P. M. v., Delaney, A., Papadopoulou, N., & Smith, C. J. (2018). Governing Marine Ecosystem Restoration: The Role of Discourses and Uncertainties. Marine Policy, 96, 136–144. https://doi.org/10.1016/j.marpol.2018.08.014. Pandolfi, J. M., Bradbury, R. H., Sala, E., Hughes, T. P., Bjorndal, K. A., Cooke, R. G., McArdle, D., McClenachan, L., Newman, M. J., & Paredes, G. (2003). Global trajectories of the long-term decline of coral reef ecosystems. Science, 301(5635), 955–958. https://doi.org/10.1126/science.1085706. Paxton, C. W., Baria, M. V. B., Weis, V. M., & Harii, S. (2016). Effect of elevated temperature on fecundity and reproductive timing in the coral Acropora digitifera. Zygote, 24(4), 511–516. https://doi.org/10.1017/S0967199415000477. Pendleton, L. H., Comte, A., Langdon, C., Ekstrom, J. A., Cooley, S. R., Suatoni, L., Beck, M. W., Brander, L., Burke, L., Cinner, J. E., Doherty, C., Edwards, P., Gledhill, D. K., Jiang, L. Q., Hooidonk, R. J. V., Teh, L., Waldbusser, G. G., & Ritter, J. (2016). Coral Reefs and People in a High-Co2 World: Where Can Science Make a Difference to People? PLoS One, 11(11), e0164699. https://doi.org/10.1371/journal.pone.0164699. Petersen, D., Wietheger, A., & Laterveer, M. (2008). Influence of different food sources on the initial development of sexual recruits of reefbuilding corals in aquaculture. Aquaculture, 277(3), 174–178. https://doi.org/10.1016/j.aquaculture.2008.02.034. Petersen, L.-E., Kellermann, M. Y., Fiegel, L. J., Nietzer, S., Bickmeyer, U., Abele, D., & Schupp, P. J. (2023). Photodegradation of a Bacterial Pigment and Resulting Hydrogen Peroxide Release Enable Coral Settlement. Scientific Reports, 13(1). https://doi.org/10.1038/s41598-023-30470-w. Pikesley, S. K., Godley, B. J., Latham, H., Richardson, P. B., Robson, L. M., Solandt, J.-L., Trundle, C., Wood, C., & Witt, M. J. (2016). Pink sea fans (Eunicella verrucosa) as indicators of the spatial efficacy of Marine Protected Areas in southwest UK coastal waters. Marine Policy, 64, 38–45. https://doi.org/10.1016/j.marpol.2015.10.010. Plucer-Rosario, G., & Randall, R. H. (1987). Preservation of rare coral species by transplantation and examination of their recruitment and growth. Bulletin of Marine Science, 41(2), 585–593. Pollock, F. J., Katz, S. M., Water, J. A. J. M. v. d., Davies, S. W., Hein, M. Y., Torda, G., Matz, M. V., Beltran, V. H., Buerger, P., Puill-Stephan, E., Abrego, D., Bourne, D. G., & Willis, B. L. (2017). Coral Larvae for Restoration and Research: A Large-Scale Method for Rearing Acropora Millepora Larvae, Inducing Settlement, and Establishing Symbiosis. Peerj. https://doi.org/10.7717/peerj.3732. Possingham, H. P., Bode, M., & Klein, C. J. (2015). Optimal conservation outcomes require both restoration and protection. PLoS biology, 13(1), e1002052. https://doi.org/10.1371/journal.pbio.1002052. Pratchett, M. S., Bay, L. K., Coker, D. J., Cole, A. J., & Lawton, R. (2012). Effects of Coral Bleaching on Coral Habitats and Associated Fishes. 59–67. https://doi.org/10.7882/fs.2012.012. Price, D. M., Robert, K., Callaway, A., lacono, C. L., Hall, R., & Huvenne, V. (2019). Using 3D Photogrammetry From ROV Video to Quantify Cold-Water Coral Reef Structural Complexity and Investigate Its Influence on Biodiversity and Community Assemblage. Coral Reefs, 38(5), 1007–1021. http://doi.org/10.1007/s00338-019-01827-3. Prouty, N. G., Fisher, C. R., Demopoulos, A. W., & Druffel, E. R. (2016). Growth rates and ages of deep-sea corals impacted by the Deepwater Horizon oil spill. Deep Sea Research Part II: Topical Studies in Oceanography, 129, 196–212. https://doi.org/10.1016/j.dsr2.2014.10.021. Rakka, M., Maier, S., Van Oevelen, D., Godinho, A., Bilan, M., Orejas, C., & Carreiro-Silva, M. (2021). Contrasting metabolic strategies of two co-occurring deep-sea octocorals. Scientific Reports, 11(1), 10633. https://doi.org/10.1038/s41598-021-90134-5. Rakka, M., Orejas, C., Sampaio, I., Monteiro, J., Parra, H., & Carreiro-Silva, M. (2017). Reproductive biology of the black coral Antipathella wollastoni (Cnidaria: Antipatharia) in the Azores (NE Atlantic). Deep Sea Research Part II: Topical Studies in Oceanography, 145, 131–141. https://doi.org/10.1016/j.dsr2.2016.05.011. Randall, C. J., Negri, A. P., Quigley, K. M., Foster, T., Ricardo, G. F., Webster, N. S., Bay, L. K., Harrison, P. L., Babcock, R. C., & Heyward, A. J. (2020). Sexual production of corals for reef restoration in the Anthropocene. Marine Ecology Progress Series, 635, 203–232. https://www.int-res.com/abstracts/meps/v635/p203-232. Ribes, M., Coma, R., & Gili, J.-M. (1999). Heterogeneous feeding in benthic suspension feeders: the natural diet and grazing rate of the temperate gorgonian Paramuricea clavata (Cnidaria: Octocorallia) over a year cycle. Marine Ecology Progress Series, 183, 125–137. https://doi.org/10.3354/meps183125. Ribes, M., Coma, R., Rossi, S., & Micheli, M. (2007). Cycle of gonadal development in Eunicella singularis (Cnidaria: Octocorallia): trends in sexual reproduction in gorgonians. Invertebrate Biology, 126(4), 307–317. https://doi.org/10.1111/j.1744-7410.2007.00101.x. Robinson, J. P. W., Wilson, S. K., Jennings, S., & Graham, N. A. J. (2019). Thermal Stress Induces Persistently Altered Coral Reef Fish Assemblages. Global change biology, 25(8), 2739–2750. https://doi.org/10.1111/gcb.14704. Roik, A. K., Röthig, T., Roder, C., Muller, P. J., & Voolstra, C. R. (2015). Captive Rearing of the Deep-Sea Coral Eguchipsammia fistula from the Red Sea Demonstrates Remarkable Physiological Plasticity. Peerj, 3(e734). https://doi.org/10.7717/peerj.734. Ros, Z. D., Dell’Anno, A., Morato, T., Sweetman, A. K., Carreiro-Silva, M., Smith, C. J., Papadopoulou, N., Corinaldesi, C., Bianchelli, S., Gambi, C., Cimino, R., Snelgrove, P. V. R., Dover, C. L. V., & Danovaro, R. (2019). The Deep Sea: The New Frontier for Ecological Restoration. Marine Policy, 108. https://doi.org/10.1016/j.marpol.2019.103642. Rossi, S., Bramanti, L., Gori, A., & Orejas, C. (2017). Marine animal forests: Springer Nature. Sakai, Y., Yamamoto, H. H., & Maruyama, S. (2024). Long-Term Aquarium Records Delineate the Synchronized Spawning Strategy Of Acropora Corals. Royal Society Open Science, 11(5). https://doi.org/10.1098/rsos.240183. Sartoretto, S., & Francour, P. (2012). Bathymetric distribution and growth rates of Eunicella verrucosa (Cnidaria: Gorgoniidae) populations along the Marseilles coast (France). Scientia Marina, 76(2), 349–355. https://doi.org/10.3989/scimar.03262.16B. Sciascia, R., Guizien, K., & Magaldi, M. G. (2022). Larval dispersal simulations and connectivity predictions for Mediterranean gorgonian species: sensitivity to flow representation and biological traits. ICES Journal of Marine Science, 79(7), 2043–2054. https://doi.org/10.1093/icesjms/fsac135. Sheehan, E., Rees, A., Bridger, D., Williams, T., & Hall-Spencer, J. (2017). Strandings of NE Atlantic gorgonians. Biological conservation, 209, 482–487. https://doi.org/10.1016/J.BIOCON.2017.03.020. Sheehan, E. V., Stevens, T. F., Gall, S. C., Cousens, S. L., & Attrill, M. J. (2013). Recovery of a temperate reef assemblage in a marine protected area following the exclusion of towed demersal fishing. PLoS One, 8(12), e83883. https://doi.org/10.1371/journal.pone.0083883. Smith, C. J., Verdura, J., Papadopoulou, N., Fraschetti, S., Cebrian, E., Fabbrizzi, E., Monserrat, M., Drake, M., Bianchelli, S., Danovaro, R., Malak, D. A., Ballesteros, E., Benjumea Tesouro, T., Boissery, P., D’Ambrosio, P., Galobart, C., Javel, F., Laurent, D., Orfanidis, S., & Mangialajo, L. (2023). A Decision-Support Framework for the Restoration of Cystoseira Sensu Lato Forests. Frontiers in Marine Science, 10. https://doi.org/10.3389/fmars.2023.1159262. Sneed, J. M., Sharp, K. H., Ritchie, K. B., & Paul, V. J. (2014). The chemical cue tetrabromopyrrole from a biofilm bacterium induces settlement of multiple Caribbean corals. Proc Biol Sci, 281(1786). https://doi.org/10.1098/rspb.2013.3086. Sorek, M., & Levy, O. (2014). Coral spawning behavior and timing. In Annual, lunar, and tidal clocks: Patterns and mechanisms of nature's enigmatic rhythms (pp. 81–97). Tokyo: Springer. Suggett, D. J., & van Oppen, M. J. H. (2022). Horizon scan of rapidly advancing coral restoration approaches for 21st century reef management. Emerging Topics in Life Sciences, 6(1), 125–136. https://doi.org/10.1042/ETLS20210240. Sun, Z., Hamel, J.-F., Edinger, E., & Mercier, A. (2010). Reproductive biology of the deep-sea octocoral Drifa glomerata in the Northwest Atlantic. Marine Biology, 157, 863–873. https://doi.org/10.1007/S00227-009-1369-9. Sun, Z., Hamel, J., & Mercier, A. (2009). Planulation of deep-sea octocorals in the NW Atlantic. Coral Reefs, 28(3), 781. https://doi.org/10.1007/s00338-009-0505-9. Sun, Z., Hamel, J. F., & Mercier, A. (2011). Planulation, larval biology, and early growth of the deep-sea soft corals Gersemia fruticosa and Duva florida (Octocorallia: Alcyonacea). Invertebrate Biology, 130(2), 91–99. https://doi.org/10.1111/j.1744-7410.2011.00229.x. Suzuki, G., Okada, W., Yasutake, Y., Yamamoto, H., Tanita, I., Yamashita, H., Hayashibara, T., Komatsu, T., Kanyama, T., & Inoue, M. (2020). Enhancing coral larval supply and seedling production using a special bundle collection system “coral larval cradle” for large-scale coral restoration. Restoration Ecology, 28(5), 1172–1182. https://doi.org/10.1111/rec.13178. Tebben, J., Motti, C. A., Siboni, N., Tapiolas, D. M., Negri, A. P., Schupp, P. J., Kitamura, M., Hatta, M., Steinberg, P. D., & Harder, T. (2015). Chemical mediation of coral larval settlement by crustose coralline algae. Scientific Reports, 5(1), 10803. https://doi.org/10.1038/srep10803. Theodor, J. (1967). Contribution a l'étude des Gorgones (VII): Ecologie et comportement de la planula. Vie et Milieu, 291–302. https://hal.sorbonne-universite.fr/hal-02951553. Toh, T. C., Ng, C. S. L., Peh, J. W. K., Toh, K. B., & Chou, L. M. (2014). Augmenting the post-transplantation growth and survivorship of juvenile scleractinian corals via nutritional enhancement. PLoS One, 9(6), e98529. https://doi.org/10.1371/journal.pone.0098529. Tonra, K. J., Wells, C. D., & Lasker, H. R. (2021). Spawning, embryogenesis, settlement, and post-settlement development of the gorgonian Plexaura homomalla. Invertebrate Biology, 140(2), e12319. https://doi.org/10.1111/ivb.12319. Torres, G., Melzer, R., Spitzner, F., Šargač, Z., Harzsch, S., & Gimenez, L. (2021). Methods to study organogenesis in decapod crustacean larvae. I. larval rearing, preparation, and fixation. Helgoland Marine Research, 75(1), 1–21. https://doi.org/10.1186/s10152-021-00548-x. Van Oppen, M. J., Gates, R. D., Blackall, L. L., Cantin, N., Chakravarti, L. J., Chan, W. Y., Cormick, C., Crean, A., Damjanovic, K., & Epstein, H. (2017). Shifting paradigms in restoration of the world's coral reefs. Global change biology, 23(9), 3437–3448. https://doi.org/10.1111/gcb.13647. Van Oppen, M. J., Oliver, J. K., Putnam, H. M., & Gates, R. D. (2015). Building coral reef resilience through assisted evolution. Proceedings of the National Academy of Sciences, 112(8), 2307–2313. https://doi.org/10.1073/pnas.1422301112. Van Woesik, R., Lacharmoise, F., & Köksal, S. (2006). Annual cycles of solar insolation predict spawning times of Caribbean corals. Ecology Letters, 9(4), 390–398. https://doi.org/10.1111/j.1461-0248.2006.00886.x. Vieira, F. A., Gregório, S. F., Ferraresso, S., Thorne, M. A., Costa, R., Milan, M., Bargelloni, L., Clark, M. S., Canario, A. V., & Power, D. M. (2011). Skin healing and scale regeneration in fed and unfed sea bream, Sparus auratus. BMC Genomics, 12, 1–19. https://doi.org/10.1186/1471-2164-12-490. Viladrich, N., Bramanti, L., Tsounis, G., Coppari, M., Dominguez-Carrió, C., Pruski, A., & Rossi, S. (2021). Estimations of free fatty acid (FFA) as a reliable proxy for larval performance in Mediterranean octocoral species. Mediterranean Marine Science. https://doi.org.10.12681/mms.27151. Viladrich, N., Linares, C., & Padilla-Gamiño, J. L. (2022). Lethal and Sublethal Effects of Thermal Stress on Octocorals Early Life‐history Stages. Global change biology, 28(23), 7049–7062. von Koch, G. (1887). Die Gorgoniden des Golfes von Neapel und der angrenzenden Meeresabschnitte: Erster Theil einer Monographie der Anthozoa Alcyonaria. In Fauna und Flora des Golfes von Neapel und der angrenzenden Meeres-Abschnitte / hrsg. von der Zoologischen Station zu Neapel (Vol. 9). Berlin: R. Friedlander & sohn. Wahab, M. A., Ferguson, S., Snekkevik, V. K., McCutchan, G., Jeong, S. Y., Severati, A., Randall, C. J., Negri, A. P., & Díaz-Pulido, G. (2023). Hierarchical Settlement Behaviours of Coral Larvae to Common Coralline Algae. Scientific Reports, 13(1). https://doi.org/10.1038/s41598-023-32676-4. Waller, R. G. (2005). Deep-water Scleractinia (Cnidaria: Anthozoa): current knowledge of reproductive processes. In A. Freiwald & J. M. Roberts (Eds.), Cold-water corals and ecosystems (Vol. Erlangen Earth Conference Series, pp. 691–700). Berlin, Heidelberg: Springer. Waller, R. G., Goode, S., Tracey, D., Johnstone, J., & Mercier, A. (2023). A review of current knowledge on reproductive and larval processes of deep-sea corals. Marine Biology, 170(5), 58. https://doi.org/10.1007/s00227-023-04182-8. Watling, L., France, S. C., Pante, E., & Simpson, A. (2011). Biology of deep-water octocorals. Advances in marine biology, 60, 41–122. https://doi.org/10.1016/B978-0-12-385529-9.00002-0. Watling, L., & Norse, E. A. (1998). Disturbance of the seabed by mobile fishing gear: a comparison to forest clearcutting. Conservation biology, 12(6), 1180–1197. https://doi.org/10.1046/j.1523-1739.1998.0120061180.x. Weinberg, S., & Weinberg, F. (1979). The life cycle of a gorgonian: Eunicella singularis (Esper, 1794). Bijdragen tot de Dierkunde, 48(2), 127–140. https://doi.org/10.1163/26660644-04802003. Williams, S. L., Ambo-Rappe, R., Sur, C., Abbott, J. M., & Limbong, S. R. (2017). Species Richness Accelerates Marine Ecosystem Restoration in the Coral Triangle. Proceedings of the National Academy of Sciences, 114(45), 11986–11991. https://doi.org/10.1073/pnas.1707962114. Zelli, E., Quéré, G., Lago, N., Di Franco, G., Costantini, F., Rossi, S., & Bramanti, L. (2020). Settlement dynamics and recruitment responses of Mediterranean gorgonians larvae to different crustose coralline algae species. Journal of Experimental Marine Biology and Ecology, 530, 151427. https://doi.org/10.1016/j.jembe.2020.151427. Additional Declarations No competing interests reported. Cite Share Download PDF Status: Published Journal Publication published 21 Jul, 2025 Read the published version in Coral Reefs → Version 1 posted Editorial decision: Revision requested 09 Apr, 2025 Reviews received at journal 27 Feb, 2025 Reviews received at journal 08 Feb, 2025 Reviews received at journal 08 Feb, 2025 Reviewers agreed at journal 22 Jan, 2025 Reviewers agreed at journal 19 Jan, 2025 Reviewers agreed at journal 16 Jan, 2025 Reviewers invited by journal 09 Jan, 2025 Editor assigned by journal 06 Jan, 2025 Submission checks completed at journal 03 Jan, 2025 First submitted to journal 31 Dec, 2024 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-5741857","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":405761883,"identity":"3c79892e-668e-4f1b-8bae-de66c7bcddb1","order_by":0,"name":"Christina Egger","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAA+klEQVRIiWNgGAWjYDACCQaGA2ASAmxggoS0JMDVpBGnhYEhAc49TFgL/+zehwd//rDI429vv/zi457zdmtnJDDe+IDPkjvHDQ7zJEgUS5w5U2Y549nt5G1nDjBbzsCjxUAiDeiYBInEhhs5acY8B24nmx1vYJPmIaDl4A+glvn336QZ/zlwLtnsMAOb9B8CWg4AHZa44Qb74ccMBw7YgW3B532JG0CH8aRJJG48k8PG2HMgOcHszMFmyx48WvhnpDF//GFTlzjv+PHHH34csLM3u5F88MYPfNYgAI8ZKDoSGxgYG4jTwMDA/hgUG/bEKh8Fo2AUjIKRAwB+aldewsNewAAAAABJRU5ErkJggg==","orcid":"","institution":"CCMAR (Centre of Marine Sciences)","correspondingAuthor":true,"prefix":"","firstName":"Christina","middleName":"","lastName":"Egger","suffix":""},{"id":405761884,"identity":"70556470-babe-41e3-bba2-c3aea200ef05","order_by":1,"name":"Catarina Melo","email":"","orcid":"","institution":"CCMAR (Centre of Marine Sciences)","correspondingAuthor":false,"prefix":"","firstName":"Catarina","middleName":"","lastName":"Melo","suffix":""},{"id":405761885,"identity":"1f4e9120-d9d1-4664-82c1-31451993e21b","order_by":2,"name":"Bailey Marquardt","email":"","orcid":"","institution":"CCMAR (Centre of Marine Sciences)","correspondingAuthor":false,"prefix":"","firstName":"Bailey","middleName":"","lastName":"Marquardt","suffix":""},{"id":405761886,"identity":"e16a4804-c3bb-4cc5-83a1-265124d572e1","order_by":3,"name":"Aschwin H. Engelen","email":"","orcid":"","institution":"CCMAR (Centre of Marine Sciences)","correspondingAuthor":false,"prefix":"","firstName":"Aschwin","middleName":"H.","lastName":"Engelen","suffix":""},{"id":405761887,"identity":"6706375c-0264-479c-aea5-22bc3c6ede66","order_by":4,"name":"Roland R. Melzer","email":"","orcid":"","institution":"SNSB-Bavarian State Collection for Zoology","correspondingAuthor":false,"prefix":"","firstName":"Roland","middleName":"R.","lastName":"Melzer","suffix":""},{"id":405761888,"identity":"a3edd465-0755-4d7c-ad7d-e46be8d72bf8","order_by":5,"name":"Elsa Santos","email":"","orcid":"","institution":"Oceanário de Lisboa","correspondingAuthor":false,"prefix":"","firstName":"Elsa","middleName":"","lastName":"Santos","suffix":""},{"id":405761889,"identity":"00380524-271c-4520-8e4b-4a0e1488a0ea","order_by":6,"name":"Margarida Fernandes","email":"","orcid":"","institution":"Oceanário de Lisboa","correspondingAuthor":false,"prefix":"","firstName":"Margarida","middleName":"","lastName":"Fernandes","suffix":""},{"id":405761890,"identity":"f0ee8a81-bb4b-42e0-b715-e257df26a287","order_by":7,"name":"Núria Baylina","email":"","orcid":"","institution":"Oceanário de Lisboa","correspondingAuthor":false,"prefix":"","firstName":"Núria","middleName":"","lastName":"Baylina","suffix":""},{"id":405761891,"identity":"f84961b9-77eb-4af8-8156-a3307780ae06","order_by":8,"name":"Ester A. Serrao","email":"","orcid":"","institution":"CCMAR (Centre of Marine Sciences)","correspondingAuthor":false,"prefix":"","firstName":"Ester","middleName":"A.","lastName":"Serrao","suffix":""},{"id":405761892,"identity":"7a518634-ec9b-444a-8374-4ed77aae4068","order_by":9,"name":"Márcio A. Coelho","email":"","orcid":"","institution":"CCMAR (Centre of Marine Sciences)","correspondingAuthor":false,"prefix":"","firstName":"Márcio","middleName":"A.","lastName":"Coelho","suffix":""}],"badges":[],"createdAt":"2024-12-31 12:08:07","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-5741857/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-5741857/v1","draftVersion":[],"editorialEvents":[{"content":"https://doi.org/10.1007/s00338-025-02705-x","type":"published","date":"2025-07-21T15:58:23+00:00"}],"editorialNote":"","failedWorkflow":false,"files":[{"id":74953499,"identity":"c5088eae-54d1-411e-9227-96401ff313df","added_by":"auto","created_at":"2025-01-28 16:45:01","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":6856495,"visible":true,"origin":"","legend":"\u003cp\u003eHistological sections of example polyps of Eunicella verrucosafrom three different dates: \u003cstrong\u003ea-c\u003c/strong\u003e Female polyps with visible oocytes (arrows); \u003cstrong\u003ea\u003c/strong\u003e Stage 4 late vitellogenic oocyte in early August (05/08/2021), \u003cstrong\u003eb\u003c/strong\u003e late August (26/08/2021) and \u003cstrong\u003ec\u003c/strong\u003e stage 4 late vitellogenic oocyte mid-September (16/09/2021); \u003cstrong\u003ed-f\u003c/strong\u003e Male polyps with visible spermatocytes (asterisk); \u003cstrong\u003ed\u003c/strong\u003e small, growing (stage 2) spermatocytes in early August (05/08/2021), \u003cstrong\u003ee\u003c/strong\u003e Developing (stage 2) spermatocytes late August (26/08/2021) and \u003cstrong\u003ef\u003c/strong\u003e stage 3 spermatocytes in mid-September with empty lumen (16/09/2021)\u003c/p\u003e","description":"","filename":"floatimage1.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/8cd6828aeae535d380a6aae2.png"},{"id":74953797,"identity":"25ee0854-d00a-4718-af54-f5222850d50a","added_by":"auto","created_at":"2025-01-28 16:53:02","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":2280362,"visible":true,"origin":"","legend":"\u003cp\u003eSpawning intensity of the pink sea fan Eunicella verrucosa kept in captivity across the 2023 lunar cycle. The qualitative spawning index is as follows: high (\u0026gt;250 propagules); low (10-250 propagules). Egg release below 10 propagules was not considered as an event.\u003c/p\u003e","description":"","filename":"floatimage2.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/cfc3603b792d525a044e4759.png"},{"id":74953798,"identity":"de1ddab8-e717-4d74-aff3-12ff83a3d955","added_by":"auto","created_at":"2025-01-28 16:53:02","extension":"jpg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":6482544,"visible":true,"origin":"","legend":"\u003cp\u003eLight and Scanning Electron Microscopy images of the embryonic development of the pink sea fan, Eunicella verrucosa, across 72 hours. \u003cstrong\u003ea\u003c/strong\u003e Light Microscopy images displaying the variations in embryonic development. \u003cstrong\u003eb\u003c/strong\u003e-\u003cstrong\u003el\u003c/strong\u003eSEM micrographs. \u003cstrong\u003eb \u003c/strong\u003eEgg starting to cleave. \u003cstrong\u003ec\u003c/strong\u003e Magnification\u003cstrong\u003e \u003c/strong\u003eof\u003cstrong\u003e \u003c/strong\u003ecleaving area\u003cstrong\u003e \u003c/strong\u003ein a 4-cell embryo.\u003cstrong\u003e d\u003c/strong\u003e Meroblastic cleavage between 4- and 8-cell stadia; \u003cstrong\u003ee \u003c/strong\u003eEarly 16-cell stadium embryo. \u003cstrong\u003ef \u003c/strong\u003e16-cell stadium embryo. \u003cstrong\u003eg \u003c/strong\u003e64-cell stadium embryo with irregular sized cells and shape. \u003cstrong\u003eh \u003c/strong\u003e12h old embryo above 64-cell stadium. \u003cstrong\u003ei \u003c/strong\u003eMagnification\u003cstrong\u003e \u003c/strong\u003eof 6 angled cells of a 12-24h old embryo. \u003cstrong\u003ej\u003c/strong\u003e 24h old embryo with several invaginations. \u003cstrong\u003ek\u003c/strong\u003e Magnification of a 48h old embryo with 6 angled cells and microvilli. \u003cstrong\u003el\u003c/strong\u003e 48 h old embryo with pronounced invaginations.\u003c/p\u003e","description":"","filename":"floatimage3.jpg","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/19ace0460c7d565c450fafb0.jpg"},{"id":74953503,"identity":"d95b005d-d53a-415a-8af7-dc8c714ec7a6","added_by":"auto","created_at":"2025-01-28 16:45:02","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":7761758,"visible":true,"origin":"","legend":"\u003cp\u003eLight and Scanning Electron Microscopy images of the larval development of the pink sea fan, Eunicella verrucosa, over 90 days. \u003cstrong\u003ea\u003c/strong\u003e Light Microscopy images displaying changes in shape over time. \u003cstrong\u003eb-l\u003c/strong\u003e SEM micrographs: \u003cstrong\u003eb\u003c/strong\u003e Young larva (5 days old) at the onset of swimming activity;\u003cstrong\u003e c\u003c/strong\u003e Magnification of the preliminary sparse ciliation of an early larva (9 days). \u003cstrong\u003ed\u003c/strong\u003e Elongated 9-day old larva. \u003cstrong\u003ee\u003c/strong\u003eCiliation at the anterior part of a 17-day old larva. \u003cstrong\u003ef \u003c/strong\u003e17-day old larva. \u003cstrong\u003eg\u003c/strong\u003e Ciliation at the posterior part of a 17-day old larva. \u003cstrong\u003eh\u003c/strong\u003e20-day old larvae with visible posterior pole. \u003cstrong\u003ei\u003c/strong\u003e Magnification the apical pole of a 39-day old larva. \u003cstrong\u003ej\u003c/strong\u003e 39-day old larva. \u003cstrong\u003ek\u003c/strong\u003eMagnification of the dense ciliation of a 39-day old larva. \u003cstrong\u003el\u003c/strong\u003e 39-day old, substrate probing larva.\u003c/p\u003e","description":"","filename":"floatimage4.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/1d00dfcc0dff6af809f16c49.png"},{"id":74953505,"identity":"0f62d622-e730-4e62-bdd5-c487c9bf0db3","added_by":"auto","created_at":"2025-01-28 16:45:02","extension":"jpg","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":3449494,"visible":true,"origin":"","legend":"\u003cp\u003eAttachment, settlement and metamorphosis of the pink sea fan, Eunicella verrucosa, larvae. \u003cstrong\u003ea\u003c/strong\u003e Loose attachment of larvae to rocky substrate with the posterior pole after 15-17 days. \u003cstrong\u003eb\u003c/strong\u003e Settlers metamorphosing and developing eight mesenteries and a mouth. \u003cstrong\u003ec\u003c/strong\u003e Development of tentacles, pinnules and sclerites. \u003cstrong\u003ed\u003c/strong\u003e Three-month-old recruits growing in length.\u003c/p\u003e","description":"","filename":"floatimage5.jpg","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/1893f70d327430aa986b3531.jpg"},{"id":74953511,"identity":"0fa81ae5-eef1-44f7-8479-346f92d288da","added_by":"auto","created_at":"2025-01-28 16:45:02","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":4390143,"visible":true,"origin":"","legend":"\u003cp\u003eDevelopment of two exemplary recruits of the pink sea fan Eunicella verrucosa over the period of one year. \u003cstrong\u003ea\u003c/strong\u003e Three-month-old recruit retracted. \u003cstrong\u003eb\u003c/strong\u003e The same recruit 6 months old, growing an internal skeleton. \u003cstrong\u003ec\u003c/strong\u003e The same recruit, ten months old, budding the second polyp and secreting a holdfast. \u003cstrong\u003ed\u003c/strong\u003e Six-month-old recruit. \u003cstrong\u003ee \u003c/strong\u003eThe same recruit nine months old with internal skeleton and second polyp depositing the holdfast. \u003cstrong\u003ef\u003c/strong\u003e The same recruit one month later with third and fourth polyp budding and increase in hights. Abbreviations: \u003cstrong\u003eh\u003c/strong\u003e, holdfast; \u003cstrong\u003eis\u003c/strong\u003e, internal skeleton. \u003cstrong\u003ep\u003c/strong\u003e, pinnules; \u003cstrong\u003em\u003c/strong\u003e, mouth; \u003cstrong\u003eme\u003c/strong\u003e, mesenteries; \u003cstrong\u003es\u003c/strong\u003e, sclerites; \u003cstrong\u003e1\u003c/strong\u003e, primary polyp; \u003cstrong\u003e2\u003c/strong\u003e, secondary polyp; \u003cstrong\u003e3\u003c/strong\u003e, third polyp; \u003cstrong\u003e4\u003c/strong\u003e, fourth polyp.\u003c/p\u003e","description":"","filename":"floatimage6.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/f9bf908a7d337497b963388a.png"},{"id":74953508,"identity":"0052d1cb-dad3-42fa-a7a5-73e8c1d78d78","added_by":"auto","created_at":"2025-01-28 16:45:02","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":682287,"visible":true,"origin":"","legend":"\u003cp\u003eTotal number and average over the 5 replicates of pink sea fan, Eunicella verrucosa, settlers over time for two cohorts: Cohort 1 - spawning of October 11\u003csup\u003eth\u003c/sup\u003e, challenged with substrate at 7 days of age; Cohort 2 – spawning of 30\u003csup\u003eth\u003c/sup\u003e of September, challenged with substrate at 17 days of age. The data presented represent the cumulative number of settlers irrespective of the type of substrate on which larvae settled (see Methods). The vertical lines indicate the points in time when rooms were changed and substrates were replaced, respectively.\u003c/p\u003e","description":"","filename":"floatimage7.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/3f5e974b463d877030c62ca9.png"},{"id":74953504,"identity":"67d0625b-0f25-4ff7-8a20-0792a0fc1102","added_by":"auto","created_at":"2025-01-28 16:45:02","extension":"png","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":570197,"visible":true,"origin":"","legend":"\u003cp\u003eLarval and recruit survivorship (average over the 5 replicates) of pink sea fan Eunicella verrucosa of the two cohorts: Cohort 1 - spawning of October 11th, challenged with substrate at 7 days of age; Cohort 2 – spawning of 30th of September, challenged with substrate at 17 days of age. The data presented represent the cumulative number of settlers irrespective of the type of substrate on which larvae settled (see Material and Methods). The vertical lines indicate the points in time when rooms were changed and substrates were replaced, respectively.\u003c/p\u003e","description":"","filename":"floatimage8.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/61753e563ccbb6de243c7741.png"},{"id":74953527,"identity":"16fea2a1-8ef4-4391-a019-233ee604304d","added_by":"auto","created_at":"2025-01-28 16:45:03","extension":"png","order_by":9,"title":"Figure 9","display":"","copyAsset":false,"role":"figure","size":1307544,"visible":true,"origin":"","legend":"\u003cp\u003eSettlement preferences of pink sea fan, Eunicella verrucosa, larvae when provided with rock (solid bar), crustose calcareous algae (small doted bar) and bare gorgonian skeleton (larger dotted bar). The data presented are for two cohorts of larvae: Cohort, challenged with substrate 7 days after spawning (orange); and Cohort 2, challenged with substrate after 17 days (pink).\u003c/p\u003e","description":"","filename":"floatimage9.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/b280a29c97c28cfd7f6e9a60.png"},{"id":74953510,"identity":"a30cb5ba-42b0-402e-8e8b-7f8c36e6c719","added_by":"auto","created_at":"2025-01-28 16:45:02","extension":"png","order_by":10,"title":"Figure 10","display":"","copyAsset":false,"role":"figure","size":488338,"visible":true,"origin":"","legend":"\u003cp\u003eTotal settlement success and survivorship of 3 cohorts of eggs from CCMAR, one cohort of eggs from Oceanário de Lisboa (\u003cstrong\u003ea\u003c/strong\u003e) and 4 cohorts of larvae from CCMAR (\u003cstrong\u003eb\u003c/strong\u003e) of the pink sea fan Eunicella verrucosa over the period of 7 months post settlement.\u003c/p\u003e","description":"","filename":"floatimage10.png","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/4ac381f836d4e66469213540.png"},{"id":87756791,"identity":"9e38bddd-4009-437b-b717-644d13cdfa9e","added_by":"auto","created_at":"2025-07-28 16:09:22","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":35551538,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-5741857/v1/34063b61-f3de-49e7-bfae-bef9ed2a4d38.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"Reproductive phenology and sexual propagation of the pink sea fan Eunicella verrucosa Pallas, 1766 for coral restoration","fulltext":[{"header":"INTRODUCTION","content":"\u003cp\u003eCoral ecosystems are experiencing a significant decline globally, primarily due to a combination of climate change and other anthropogenic pressures (Carpenter et al. 2008; Hughes et al. 2017b; Hughes et al. 2018). The present threats to coral reefs extend beyond rising temperatures. Ocean acidification adversely affects coral calcification and overall health, making them more susceptible to diseases and environmental stressors (Feng et al. 2016; Pendleton et al. 2016). Local human activities, such as pollution, overfishing, and coastal development, exacerbate these challenges, often leading to more immediate and severe impacts than climate change alone (Ferrario et al. 2014; H\u0026oslash;egh-Guldberg et al. 2018). However, such effects are much less studied in temperate and deep corals than in tropical ones, in which widespread damage has been well documented. Coral reefs have lost approximately 50% of their living coral cover since the 1950s, with a corresponding 60% decline in fish populations associated with these habitats (Eddy et al. 2021; Hughes et al. 2017b). This loss not only threatens biodiversity but also undermines the ecosystem services that coral reefs provide and which are vital for the livelihoods of millions of people (Hughes et al. 2017a; Hughes et al. 2017b; Mercado-Molina \u0026amp; Suleim\u0026aacute;n-Ramos 2023). The altered trophic dynamics further destabilizes these ecosystems (Robinson et al. 2019). When coral reefs continue to degrade, the potential for recovery diminishes, creating a feedback loop that perpetuates the decline of these critical marine environments (Carpenter et al. 2008; Pandolfi et al. 2003; Pratchett et al. 2012). As conservation alone appears no longer enough for the long-term persistence of many ecosystems (Hughes et al. 2017a; O\u0026rsquo;connor et al. 2020; Orth et al. 2020; Smith et al. 2023), the United Nations and European Union have outlined the critical role of biological restoration to restore degraded ecosystems (UN Decade on Ecosystem Restoration 2021\u0026ndash;2030 \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://www.decadeonrestoration.org/\u003c/span\u003e\u003cspan address=\"https://www.decadeonrestoration.org/\" targettype=\"URL\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e, the EU Biodiversity Strategy for 2030, the EU Nature Restoration Law), with the marine realm being given increasing attention (Marine Strategy Framework Directive (MSFD).\u003c/p\u003e \u003cp\u003eRestoration efforts in shallow, warm marine ecosystems have notably advanced more quickly than those in deep, cold, and temperate ecosystems. This disparity is largely due to a combination of ecological, logistical, and financial challenges. Shallow tropical marine environments, including coral reefs, mangroves, and seagrass meadows, have received the most attention in restoration initiatives, primarily because they are more accessible and offer more perceived immediate socio-economic benefits (Bayraktarov et al. 2020; Williams et al. 2017). Colder and deeper marine habitats have received far less restoration a attention and efforts compared to their tropical counterparts (Ros et al. 2019). Damage to non-tropical corals has been accumulating over a long period, but it has often been overlooked due to the lack of visibility of these ecosystems. However, the ecological importance of these habitats cannot be overstated. They provide essential habitats for many marine species, including commercially valuable fish, yet they are as vulnerable as tropical corals (Bongiorni et al. 2010; Buhl-Mortensen et al. 2018; Buhl-Mortensen et al. 2017; Danovaro et al. 2010; Lange et al. 2023). This disparity is largely due to the technical challenges and high costs associated with deep-sea restoration, as well as a general lack of awareness and understanding of these habitats among policymakers and the public (Ounanian et al. 2018; Ros et al. 2019). This is evident in the absence of clear, consistent definitions for deep-sea, cold-water, and temperate habitats. Many coral species exist over broad depths and geographical ranges (Buhl-Mortensen et al. 2018), making it difficult to define their distributions and even harder to protect or restore them. Furthermore, the ecological complexity and slow recovery rates of deep-sea ecosystems add to the challenges of restoration. Many deep-sea species have long lifespans, and the intricate relationships within these ecosystems mean that restoration efforts take much longer to yield visible results. This delays investment and research in these areas (Prouty et al. 2016; Ros et al. 2019). As a result, while shallow marine ecosystems benefit from established restoration practices, deeper ecosystems remain in the early stages of restoration science, highlighting a critical gap in marine conservation (Ounanian et al. 2018; Ros et al. 2019). To counteract this, the essential scientific information for efficient restoration plans and techniques is needed, to allow to at least catch up with progress in restoration methods for shallow water counterparts.\u003c/p\u003e \u003cp\u003eIn coral restoration, there are several approaches, all of which gained noticeable increase over the last years (Bostr\u0026ouml;m-Einarsson et al. 2020; Suggett \u0026amp; Van Oppen 2022). The most common and widely applied method is asexual propagation, achieved by the fragmentation of donor colonies or from opportunistic coral sources like fishing-bycatch, broken pieces during storms (Boch \u0026amp; Morse 2012; Garrison \u0026amp; Greg 2008; Plucer-Rosario \u0026amp; Randall 1987). The corals can be grown in nurseries and subsequently out planted on the reef or directly transplanted (Forrester et al. 2019). Asexual propagation, however, carries the risk of limited genetic diversity or potentially dismissed impacts on the donor colonies and may not be self-sustaining (Baums et al. 2019; Henry et al. 2021; Van Oppen et al. 2015). A relatively new but steadily growing approach is restoration through sexual propagation, where sexually produced recruits can then be reintroduced on the reef (Henry et al. 2021). Coral spawn can be either collected \u003cem\u003ein situ\u003c/em\u003e and then reared in the laboratory or field (Chamberland et al. 2015; Heyward et al. 2002; Suzuki et al. 2020) or the parent colonies can be kept and spawn in captivity (Dela Cruz \u0026amp; Harrison 2020; Henry et al. 2021; O\u0026rsquo;neil et al. 2021; Pollock et al. 2017). Ex situ aquaculture offers the opportunity to deliver a consistent supply of corals to support research and assist in complementary reef restoration (Lam et al. 2023). Sexual propagation offers the significant advantage of producing many genetically unique individuals thereby enhancing genetic diversity. This diversity enhances a population's ability to adapt to changing environments by providing a rich source of genetic variability for natural selection. Additionally, precise manipulations and targeted selection techniques, such as assisted evolution during the larval stage, can further increase the resilience of the new generation (Baums 2008; Doropoulos et al. 2019; Van Oppen et al. 2017; Van Oppen et al. 2015), though long-term trade-offs still need to be assessed.\u003c/p\u003e \u003cp\u003eDespite the growing recognition for the need to actively restore coral habitats, the implementation of coral restoration at large scales in non-tropical corals has been very limited (but see Montseny et al. 2021b; Montseny et al. 2020) and it is so far restricted to asexual propagation (Boch et al. 2019; Roik et al. 2015; Ros et al. 2019). In contrast, in tropical coral reefs, methods have been developed and simplified, especially for upscaling restoration through non-scientific organisations that implement them in the field (SECORE; Bayraktarov et al. 2019; Bostr\u0026ouml;m-Einarsson et al. 2020). We are lacking such methodologies and protocols for deeper reefs, despite the growing interest of non-scientific conservationists and explorational divers in these underexplored ecosystems (Ankamah-Yeboah et al. 2020; Mengerink et al. 2014). When it comes to sexual propagation in deep-sea corals, the first major challenge is the limited understanding of their biology and reproductive patterns. Many of these species, particularly those found below diving depth, remain understudied due to logistical and financial difficulties (Montseny et al. 2021a; Randall et al. 2020). The second challenge lies in our ability to keep these corals healthy in captivity for extended periods, which is essential for manipulating and inducing spawning. Despite these hurdles, addressing them is crucial for developing effective restoration strategies.\u003c/p\u003e \u003cp\u003eDeep-water and cold-temperate coral habitats differ significantly from tropical shallow-water reefs in terms of physiology, environmental conditions, and species composition (Bridge et al. 2013; Menza et al. 2007; Price et al. 2019). Coral gardens that are dominated by gorgonians can form loose to dense forest-like aggregations, often referred to as marine animal forests (Rossi et al. 2017). These habitats extend from shallow depths to several hundred meters below the surface. They typically consist of a few key gorgonian species that create a three-dimensional structure, providing essential habitat and support for a diverse array of marine organisms (Baillon et al. 2012; Buhl-Mortensen et al. 2017; Watling et al. 2011).\u003c/p\u003e \u003cp\u003eThe pink sea fan, \u003cem\u003eEunicella verrucosa\u003c/em\u003e, plays such an important, habitat-forming role in biotopes of the Eastern Atlantic and the Mediterranean Sea. It grows on hard substrate over a large latitudinal and depth range (2-200m) (Chimienti 2020; Coz et al. 2012; Pikesley et al. 2016; Sartoretto \u0026amp; Francour 2012). Its trans-equatorial distribution spans from western Ireland and SW British Isles to Angola in Western Africa and includes the Mediterranean Sea (Carpine 1963; Carpine \u0026amp; Grasshoff 1975; Chimienti 2020; Grasshoff 1992). Like other temperate and cold-water corals, these species exhibit long lifespans and slow growth rates (0.62-3.33cm/year in height: Sartoretto \u0026amp; Francour 2012), making their populations particularly susceptible to anthropogenic disturbances such as anchoring and fishing (Watling \u0026amp; Norse 1998). These human activities can exacerbate the already significant impacts of climate change, which include increased frequency of natural disturbances like storms and marine heatwaves (Hall-Spencer et al. 2007; Munro \u0026amp; Munro 2003; Sheehan et al. 2017; Sheehan et al. 2013).\u003c/p\u003e \u003cp\u003e \u003cem\u003eEunicella verrucosa\u003c/em\u003e is listed as vulnerable in the IUCN Red List of Threatened Species since 1996 and is considered to have a high to medium-high extinction risk due to exploitation. Impacts resulting from bottom contact fishing gear are one of the major risk factors for the species in South Portugal, where \u003cem\u003eE. verrucosa\u003c/em\u003e is one of coral species most numerous caught as fishing bycatch (Dias et al. 2020).\u003c/p\u003e \u003cp\u003eVery little is known about the biology and life cycle of \u003cem\u003eEunicella verrucosa\u003c/em\u003e. Munro (2004) studied some populations in the UK, revealing that they are gonochoric, with the female reproductive cycle extending beyond 12 months. Broadcast spawning has been suggested as the mode of reproduction, with spawning likely occurring in August or September (Munro 2004). This is a limitation for restoration because reproductive traits such as planktonic larval duration (PLD), larval behaviour (buoyancy and swimming behaviour) or mode of reproduction of the corals (brooder or spawner) are crucial information for sexual propagation, as well as to help understand natural population dynamics that can directly influence the planning of restoration strategies (Fogarty \u0026amp; Botsford 2007; Jones et al. 2007; Marti-Puig et al. 2013; Waller et al. 2023).\u003c/p\u003e \u003cp\u003eThe aim of this study was to characterize the reproductive and larval biology of \u003cem\u003eEunicella verrucosa\u003c/em\u003e and test methods of sexual propagation in captivity, thus addressing the conservation needs and information gaps highlighted above. Specifically, we document for the first time the reproductive phenology of the species in populations from SW Portugal, comparing colonies maintained in captivity for short- and long-term (1 and 2 years). We also describe larval development and settlement preferences using experiments with the overall objective of obtaining sexually produced recruits for habitat restoration.\u003c/p\u003e"},{"header":"METHODS","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eInference of Reproduction Window\u003c/h2\u003e \u003cp\u003eThe timing of reproduction in \u003cem\u003eE. verrucosa\u003c/em\u003e was inferred by examining the stages of gamete development in samples collected as fishing bycatch around Cape St. Vincent (southern Portugal) in 2020 and 2021 (see Dias et al. 2020 for details on fishing areas) at the Centro de Ci\u0026ecirc;ncias do Mar (CCMAR). Sampling focused on the Summer and early Autumn periods following available information on reproductive patterns for the species in the UK (Munro 2004) and other \u003cem\u003eEunicella\u003c/em\u003e spp. in the Mediterranean (Coma et al. 1995b; Gori et al. 2012). The presence and maturation state of the gametes was assessed through polyp dissection and histological sectioning at the at CCMAR. All samples were fixed in 10% Formalin, washed three times with distilled water and gradually dehydrated to EtOH 70% until further analysis. Dissection of the polyps was performed under a stereomicroscope, first to distinguish female and male individuals by identifying their gonads (oocytes and spermaries) where possible. The number of gonads per polyp and branch order (sensu Brazeau \u0026amp; Lasker 1988) was then counted and the individuals with higher number of reproductive polyps were selected for histological sectioning. For the histological study, branchlets of approximately 2 cm were cut from the first and second order branches of four colonies. Selected samples for histology were hydrated and treated in 2.1 M EDTA (pH 8) for two days to decalcify the sclerites embedded in the coral tissue, following the method used for sea bream scales (Vieira et al. 2011). After decalcification, the samples were washed several times with deionized water to remove any residual EDTA and then dehydrated through a graded ethanol series. The samples were saturated in xylene, followed by impregnation and embedding in low melting point paraffin wax (Histosec, Merck) using an automatic embedding processor. Serial sections of 5 \u0026micro;m were cut from the paraffin block using a manual rotary microtome (Leica RM 2135, Germany) and mounted on glass slides coated with 3-aminopropyltriethoxysilane (APES; Sigma-Aldrich, Madrid, Spain). For each wax block containing a piece of coral branch, 2\u0026ndash;3 sections were cut from the base of the polyps to sample the gonadal tissue. The sections were dried overnight at 37\u0026deg;C, then cooled to room temperature for storage or staining. The sections on the slides were dewaxed and rehydrated before being stained with hematoxylin and eosin (H\u0026amp;E) according to the method described by Najafpour et al. (2020). They were then mounted in Tissue-Tek Resin (Sakura Finetek) and covered with a glass coverslip. The stained slides were visualized under a light microscope to identify the reproductive structures of the polyps. The gametogenic maturation classification system defined by Waller (2005) was used to describe the ripening process of the gametes.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eSpawning Observations and Larval Rearing\u003c/h3\u003e\n\u003cp\u003eThe observations on spawning and settlement were conducted on corals collected as fishing bycatch from approximately the same location as the corals used to determine the spawning window: Cape St. Vincent in southern Portugal. However, observations were made on corals collected during three consecutive years (2021, 2022 and 2023) and kept at two separate facilities: the marine field station Ramalhete of CCMAR (corals from 2023) and the Ocean\u0026aacute;rio de Lisboa (ODL; corals from 2021 and 2022).\u003c/p\u003e\n\u003ch3\u003eShorter-term monitoring during the reproductive season (CCMAR)\u003c/h3\u003e\n\u003cp\u003eIn 2023, coral colonies were collected from early July to mid-August. A total of 70 colonies, varying in colour, size, and health, were collected in several batches. The colonies were dissected, sexed, and then transferred to tank systems at Ramalhete Marine Station in Faro for spawning observations. Approximately 50 colonies were maintained in a cooled, semi dark outdoor flow-through system, and 20 in a dark indoor, semi-closed, temperature-controlled system at 15\u0026ndash;16\u0026ordm;C. Water flow was provided by wavemaker pumps and corals were fed once daily with frozen rotifers, copepods and red zooplankton, which were squeezed through a 150\u0026micro;m net to avoid oversized food particles. During feeding water in- and outflow was stopped for 1\u0026ndash;3 hours to avoid the food being washed out. The tank bottoms were hoovered daily to provide good water quality. During the expected spawning window, perforated PVC cylinders with a 150\u0026micro;m mesh fitting were attached to the outflow of the tanks to collect released offspring. A venturi airlift was also installed at the water surface of the outdoor tanks. The air lift, outflow filters and colonies were monitored daily for gamete and/or larval release. Additionally, branches were regularly dissected to check for the presence of mature gametes.\u003c/p\u003e \u003cp\u003eDuring spawning, oocytes and embryos were collected both directly from the water column using a transfer pipette and from the two airlifts (in case of the outdoor tanks) and the outflow filters, by gently washing them into plastic beakers. Eggs, embryos, and larvae were kept in plastic jars and boxes (1\u0026ndash;10 L) sorted by collection date in a temperature-controlled room at 16 \u0026ordm;C. Gentle aeration was provided via air tubes producing a few bubbles per second, and approximately two thirds of the water was changed every other day using a 5mm hose with a small, perforated tube with a 150\u0026micro;m plankton net wrapping at the end to avoid larvae being sucked out. The intensity of spawning each day was categorized into low (10\u0026ndash;250 propagules collected) and high (\u0026gt;\u0026thinsp;250 propagules collected) spawning. Releases below 10 eggs were not considered as an event.\u003c/p\u003e\n\u003ch3\u003eLonger-term monitoring (1–2 years, ODL)\u003c/h3\u003e\n\u003cp\u003eA total of 15 colonies collected in 2021 and 9 colonies from 2022 were kept together in a closed indoor system at the Ocean\u0026aacute;rio de Lisboa. The life support system included an EcoDrift 4.2 pump from Aqua Medic Direct and a Hydor Seltz L 700 pump in each aquarium, a 50 \u0026micro;m filter bag at the inlet of a 1300 L sump containing bioballs, a 20 \u0026micro;m cartridge filter, two UV lights, a Frimar C1000 chiller and a HydroAir pump (model AV50-20N-S). The aquariums had indirect sunlight and moonlight from east turn windows and indirect light from the room ceiling (from 7h30 to 16h30). The moon cycle was simulated using a 54W actinic light set with a timer and an intensity adjuster, following the moon cycle at Sagres (from the website Timeanddate.Com, n.d.). Temperature modulation was based on seasonal variation patterns. The corals were fed live and frozen zooplankton and phytoplankton three times a day. The food concentration was adjusted based on field variations.\u003c/p\u003e \u003cp\u003eFrom the 26th of July till the 16th of November, egg collectors were placed in the surface skimmer of each aquarium from 16h00 to 8h00. Water flow was reduced, and circulation pumps were turned off to prevent egg and embryo damage. In the morning, the collectors were rinsed with saltwater to flush all gametes into a plastic beaker, allowing their observation under a stereomicroscope. The collected eggs, spermic sacs and embryos were placed in 4 L circular boxes with 125 \u0026micro;m mesh on the bottom and with slow water flow, connected to the main system, where the breeding colonies were maintained.\u003c/p\u003e\n\u003ch3\u003eEmbryogenesis and Larval Development (CCMAR)\u003c/h3\u003e\n\u003cp\u003eEmbryonic and larval development stages were observed and imaged using a ZEISS Stemi 508 stereo microscope with a ZEISS Axiocam 208 colour camera system, at Ramalhete Marine Station (CCMAR). Qualitative observations were conducted on \u0026gt;\u0026thinsp;10 batches of 50\u0026ndash;600 larvae each throughout the development period to ensure consistency. Samples of various developmental stages were collected for scanning electron microscopy (SEM). These samples were fixed overnight in 4% glutaraldehyde buffered in 0.1\u0026ndash;0.5 M S\u0026oslash;rensen\u0026rsquo;s phosphate buffer (pH 7.1), at 4 \u0026ordm;C. The following day, the samples were washed three times in pure S\u0026oslash;rensen\u0026rsquo;s phosphate buffer and transferred into 30% EtOH where they were stored until further analysis. All SEM imaging was performed at the SNSB - Bavarian State Collection for Zoology in Munich (Germany). Samples were then dehydrated using a graded acetone series (30%, 50%, 75%, 95% and 100%). Specimens were soaked for 10 min at each level, then twice in 100% acetone. Subsequently they were critical point dried using a Polaron E3000, mounted on SEM stubs with self-adhesive carbon stickers, and gold-coated for 3 minutes in an argon atmosphere using a Polaron SC510. Three embryos or larvae of each stage of development were analysed with a LEO 1430 VP SEM at a voltage of 15 kV (method described in Melzer et al. 2021; Torres et al. 2021). Measurements were done on \u0026gt;\u0026thinsp;5 propagules per stadium from LM images, using the scale bar and Adobe Photoshop\u0026copy; 22.2.0 and compared with the SEM images for consistency.\u003c/p\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eLarval Behaviour (CCMAR)\u003c/h2\u003e \u003cp\u003eLarvae, kept in plastic jars and boxes (1\u0026ndash;10 L), were provided with a substrate of bare rock and CCA-covered rock when they were between 1 and 2 weeks old. Larval behaviour was documented qualitatively by close observation of the individual batches of each spawning event to ensure consistency. The larvae were visually observed directly in their rearing jars within the cold room. Microscopic observations including video records of the substrate probing behaviour were conducted under the Stemi stereo microscope.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eLarval Swimming Behaviour\u003c/h3\u003e\n\u003cp\u003eTo document the swimming behaviour of 4\u0026ndash;6 days old larvae, three replicates of 19, 15 and 16 larvae, respectively, were temporarily placed in a 10x10x2 cm jar with a black background and filmed using a Canon 550D camera with Canon EF 50mm F1.8 STM objective, at Ramalhete Marine Station (CCMAR).\u003c/p\u003e\n\u003ch3\u003eLarval Survivorship and Substrate Choice and Settlement (Experiment 1)\u003c/h3\u003e\n\u003cp\u003eA settlement experiment was conducted to document larval settlement and metamorphosis, as well as to test settlement preferences using natural substrate. Three types of substrate were offered for settlement choice: rocky substrate collected as bycatch along with the corals, which were encrusted with a variety of taxa, including natural biofilms and/or crustose coralline algae (CCA) that are presumably implicated in inducing coral settlement and metamorphosis (Zelli et al., 2020); chips of an unidentified species of CCA retrieved with the substrate aforementioned; and pieces of conspecific bare gorgonian skeleton, on which larvae of other octocorals have been observed to settle (Coelho \u0026amp; Lasker, 2014; Weinberg \u0026amp; Weinberg, 1979; authors personal observations). Larval settlement was quantified for two cohorts of larvae spawned on October 11 (Cohort 1) and September 30 (Cohort 2), which were 7- and 17-d old at the start of the experiment, respectively. For each cohort, we used 5 replicates of 50 larvae each that were challenged with the three types of substrates. All replicates were maintained in open plastic Tupperware containers of ca. 8 cm diameter and 4 cm tall with ~\u0026thinsp;250 ml of seawater. 50% water changes were done after each counting every 2\u0026ndash;3 days. The containers were kept in a separate room with same ambient temperature as the coral indoor tank system and pH was regularly checked to monitor for variations due to organic or inorganic substrates. On the 28th of October (day 11 of the experiment of Cohort 1 and day 12 of the experiment of Cohort 2) the experiments had to be transferred to a different room kept at 18 \u0026ordm;C due to a complete failure of the air conditioning in the original room.\u003c/p\u003e \u003cp\u003eLarval settlement (i.e. attachment and metamorphosis) was counted as the attachment of larvae or the presence of recruits on a substrate in each of the 5 replicates. Monitoring of larval settlement was performed every second day at the beginning of the experiment and alternating for each cohort (experiments started on 16th and 17th of October for Cohorts 1 and 2, respectively). After 11 days, the monitoring intervals were increased to 3\u0026ndash;4 days until 34 days post-start and to every 7 days thereafter. At day 62 (Cohort 2) and 63 (Cohort 1), more substrate was added to each replicate to test for a potential settlement bottleneck caused by the occupation of available space by settlers as settling rates seemed to decrease.\u003c/p\u003e \u003cp\u003eTo evaluate substrate choice, settlers on each substrate were counted. For the evaluation of general attachment and settlement, settlers were counted for each replicate over all three substrates and the total and average number of settlers across all replicates was calculated. To compare settlement between the cohorts, we used the age of the larvae. After settlement, primary polyps were transferred to the tanks containing the mother colonies and provided with the same food.\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eEarly Life Ecology\u003c/h2\u003e \u003cdiv id=\"Sec12\" class=\"Section3\"\u003e \u003ch2\u003eShort-term monitoring (CCMAR)\u003c/h2\u003e \u003cp\u003eLarval settlement and post-settlement mortality was monitored over a period of 9 months. Single recruits were followed over the period of one year to document their development. All recruits were imaged using the previously described stereo microscope.\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eLong-term monitoring (ODL)\u003c/h2\u003e \u003cp\u003eLarvae from the major spawning events observed in Ocean\u0026aacute;rio de Lisboa in September and October were transferred from the circular boxes to four 4l rectangular boxes with 200 \u0026micro;m mesh on the side and water entrance from the main system 12 days after fertilization. Larvae were provided with natural rocks with CCA for settlement, which were collected from the wild 2 months earlier, as well as with basaltic rocks conditioned in the gorgonian system for over 7 months. The microalgae \u003cem\u003eChaetoceros calcitrans\u003c/em\u003e and \u003cem\u003eTisochrysis lutea\u003c/em\u003e were provided to the larvae since 5 days after (presumed) fertilisation. The same microalgae and living \u003cem\u003eAcartia tonsa\u003c/em\u003e and \u003cem\u003eBrachionus plicatilis\u003c/em\u003e and frozen Copepods (Ocean Aquaculture \u0026reg;) were fed after the first primary polyps were observed. The polyps were kept in the same boxes until 197 days, when 2 of them were transferred to one of the aquaria containing the adult colonies. Gradually all polyps were transferred to the same aquarium. Monitoring of larval settlement and polyp survival was conducted every two weeks until month 3 and every month after that using a Nikon SMZ745T stereo microscope in Ocean\u0026aacute;rio de Lisboa.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eRecruits Survival (Experiment 2)\u003c/h2\u003e \u003cp\u003eTo estimate total settlement success and recruit survival we followed 4 cohorts of eggs (3 cohorts from short-term monitoring at CCMAR of 226, 200 and 100 eggs, respectively, and 1 cohort of 1621 eggs from long-term monitoring at ODL) and 4 cohorts of larvae (2 cohorts of 250 larvae from Experiment 1 and 2 more cohorts of 250 and 180 larvae, respectively, all from short-term monitoring at CCMAR).\u003c/p\u003e \u003c/div\u003e"},{"header":"RESULTS","content":"\u003cdiv id=\"Sec16\" class=\"Section2\"\u003e \u003ch2\u003eInference of Reproduction Window\u003c/h2\u003e \u003cp\u003eThe gamete development data for samples of \u003cem\u003eE. verrucosa\u003c/em\u003e collected in 2020 and 2021suggests that reproduction occurred in late summer or early autumn in both years. For 2020, polyp dissections of multiple female colonies revealed a significant decrease in the number of oocytes from June to September, indicating potential spawning over this period (see \u003cb\u003eFig. S1\u003c/b\u003e in supplement 1). Histological sectioning of samples from 2021 pointed towards a reproductive window to be in a similar period to that of 2020, with female colonies still having late stage 4 vitellogenic oocytes on August 5th and 26th and in mid-September (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003ea-c). The spermatocytes showed a ripening process from small, growing (stage 2) spermatocytes in early August until stage 3 spermatocytes in mid-September with empty lumen. This shows that the accelerated ripening indicates the proximity of the spawning period. Polyp dissections of male and female colonies collected in August 2023 for spawning observations revealed vitellogenic or late vitellogenic (stage 3 or 4) oocytes and visible spermatocytes in most of the colonies (data not shown). In all the polyp dissections and histological sections made we never observed embryos or larvae inside the polyps which indicates that \u003cem\u003eE. verrucosa\u003c/em\u003e is a gonochoric broadcast spawner.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec17\" class=\"Section2\"\u003e \u003ch2\u003eSpawning Observations\u003c/h2\u003e \u003cdiv id=\"Sec18\" class=\"Section3\"\u003e \u003ch2\u003eShort-term monitoring (CCMAR)\u003c/h2\u003e \u003cp\u003eFor the 2023 reproductive season, spawning at Ramalhete Marine Station was first observed on September 2nd, when minor release of positively buoyant eggs and developing embryos were floating in both the inside and outside tanks systems synchronously, and accumulated in the collection filters attached to the water outflow. Most of the spawning then occurred as split spawning in 3 major events: the first one over two consecutive days in September 12\u0026ndash;13, the second one on the 26th and 30th of September, and the third one on the 11th of October (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). Each large spawning event was preceded and followed by minor egg releases (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). The spawning peaks appeared to correspond with the moon phase, with the largest events occurring between 5 days before and 3 days after the full or new moon, and the highest spawning output consistently happening 2\u0026ndash;3 days before the full or new moon. The eggs released were ~\u0026thinsp;300\u0026ndash;400 \u0026micro;m in diameter, all positively buoyant and accumulated in the overflow collection filters after being released into the water column, thereby confirming E. ver\u003cem\u003erucosa\u003c/em\u003e to be a broadcast spawner.\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e \u003cdiv id=\"Sec19\" class=\"Section2\"\u003e \u003ch2\u003eLong-term monitoring (ODL)\u003c/h2\u003e \u003cp\u003eFor colonies of \u003cem\u003eE. verrucosa\u003c/em\u003e kept in captivity at Ocean\u0026aacute;rio de Lisboa for over two years, egg release started earlier with minor releases first observed on August 4, extending into the 14th of November despite of very low spawning after the 23rd of October (only 0\u0026ndash;3 oocytes) (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). The major spawning events occurred on the 6th and 7th of September (6 days before the large event documented at Ramalhete) and on the 5th, 6th and 15th of October (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). The first four major events occurred 6\u0026ndash;7 days after the full moon, with the timing of spawning for the first three starting between 5:15 hrs and 6:10 hrs after moon rising. At the major spawning event in September almost all eggs and embryos were positively buoyant, however, at those occurring in October only a minor fraction had positive buoyancy.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec20\" class=\"Section2\"\u003e \u003ch2\u003eEmbryogenesis (CCMAR)\u003c/h2\u003e \u003cp\u003eEmbryos collected at CCMAR developed into a gastrula within 24 h of egg collection from the tank system. Fertilisation rates of the eggs seemed to be very high, as all eggs observed started cleaving. Once eggs started cleaving, the embryos were observed to sink down in the rearing containers, indicating negative buoyancy. We observed partial cleavage (meroblastic) of the eggs until the 8-cell stage (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003ea, d and video in supplement 2). The same observations were made for embryos reared at ODL. The greater part of the yolk remained in the initial (egg) cell at the beginning. In some cases, eggs started segmenting into one big and four to five smaller compartments that did not cleave entirely, returning instead to the initial round shape only to start segmenting again and then proceed to cleave into 8 and finally 16 evenly sized cells (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003ee, f and video in supplement 2). The cleavage was spiral and above the 32 cleavages, the pattern continued in irregular-sized cells with six squares and an irregular-shaped embryo (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eg, i). While the six squared cells became more equal in size with further cleavage, the embryo remained somewhat irregularly shaped and developed several infoldings and invaginations during gastrulation (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003ej, k, l).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec21\" class=\"Section2\"\u003e \u003ch2\u003eLarval Development and Behaviour (CCMAR)\u003c/h2\u003e \u003cp\u003eSeventy-two hours after fertilization, the first oval-shaped larvae (~\u0026thinsp;350\u0026micro;m long) started moving and actively swimming upwards to the water surface in the rearing containers where they congregated (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003ea, b and videos in supplement 3). Five to six days after egg collection, larvae were already more elongated (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003ea, d) and approximately 3 times as long as wide (~\u0026thinsp;750\u0026micro;m long, ~\u0026thinsp;250 \u0026micro;m wide). The same observations were made for larvae collected at CCMAR and ODL. At the age between 3 to 9 days old, the larval ciliation was sparse, with short cilia covering the larvae body during this period (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003ec) and ciliation increasing until day 17 of age (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003ee, g). The increase in larval ciliation coincided with an increase in swimming activity, with larvae between 7- and 12-days following spawning observed to start to partially leave the water surface and standing upright in the water column. After 13 days, larvae were observed to frequently swim up and down and starting to probe substrate (see video 1 in supplement 4). During that phase, larvae developed an oral and aboral pole, denser ciliation, and had a clear propulsion direction (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003ee-k). With advancing age, the larvae became more transparent and shorter, however stayed motile until over 100 days of age (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003ej, k).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec22\" class=\"Section2\"\u003e \u003ch2\u003eLarval Settlement, Metamorphosis and Early Life Ecology\u003c/h2\u003e \u003cdiv id=\"Sec23\" class=\"Section3\"\u003e \u003ch2\u003eShort-term monitoring (CCMAR)\u003c/h2\u003e \u003cp\u003eThe first attachment of larvae to the substrate offered in the settlement experiments at Ramalhete Marine Station was observed 15 days after spawning, when the larvae temporarily attached with the aboral pole to the substrate provided (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003ea and video 2 in supplement 4). Onset of settlement started shortly after and continued over the next two months.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eAfter successful settlement, metamorphosis occurred over several days (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eb), during which the settlers sequentially developed mesenteries, a mouth and tentacles. The primary polyps appeared healthy, apparently began feeding, and started developing sclerites within a few weeks (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eb, c). These sclerites formed in eight vertical rows surrounding the mesenteries (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003ec) and later developed into acuminate leaflets enclosing the mesenteries similar to petals (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eb). When stimulated or stressed, these leaflets would form a complete, enveloping spherical shell around the contracting polyp (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003ea). Growth in the aquarium was slow. Within the first 3 months, the settlers grew from 0.5 mm to a maximum length of 1\u0026ndash;2 mm (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003ed). By 9 months, they had expanded from 0.5 mm to 1 mm in width and reached up to 5 mm in height (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003e). The first deposition of an internal skeleton was observed at 9 months post-settlement, or 11\u0026ndash;12 months post-spawning (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eb, e). Shortly after, branching of the second polyp was noted (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003ec, e). The internal axis in the primary polyp began to develop, bending away from the mouth region (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eb), while the second polyp branched in the opposite direction from the axis below the primary polyp (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003ec, e). By the time the third polyp developed, it emerged from the axis between the first and second polyps, while the fourth polyp developed below them (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003ef). As they grew, the recruits also started to deposit a holdfast on the substrate to which they were attached (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003ec, e).\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e \u003cdiv id=\"Sec24\" class=\"Section2\"\u003e \u003ch2\u003eLonger-term monitoring (ODL)\u003c/h2\u003e \u003cp\u003eAt Ocean\u0026aacute;rio de Lisboa, the first recruits were observed at 20 and 21 days after the major spawning event (6th-7th September). Settlement continued over a period of 3\u0026ndash;4 months, similar to the observations made at CCMAR. All larvae settled in CCA rocks, and no settlers were observed on the basaltic rocks. From the October major spawning events in which a majority of oocytes were negatively buoyant, none of the larvae settled. The first tentacles with pinnules were observed at 36 days after spawning and the first observation of a secondary polyp occurred 204 days or 6.5 months after spawning, shortly after the transfer to the aquarium with the adult colonies. Settlers with 3 or more polyps were observed almost one year after spawning.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cdiv id=\"Sec25\" class=\"Section3\"\u003e \u003ch2\u003eLarval Survivorship and Substrate Choice and Settlement (Experiment 1)\u003c/h2\u003e \u003cp\u003eIn the settlement experiment conducted at CCMAR, we observed two small settlement peaks at 15 days after spawning in Cohort 1 (released on 11th October, 7 days old when experiment started) and at 19 days in Cohort 2 (released on 30th of September 17-days old when experiment started) (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). Those peaks did not lead to successful settlement, as the number of settlers counted in the following monitoring day decreased to 0 again, either indicating a temporary detachment of the \u0026lsquo;settler\u0026rsquo; or failure to complete metamorphosis (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). The number of settlers then increased slowly until 78 days of larval age in Cohort 1 and 89 days of larval age in Cohort 2, followed by a steeper increase in settlement until 97 days of larval age to a maximum of 19 settlers in Cohort 1 (C1) and until 96 days of larval age to a maximum of 29 settlers in Cohort 2 (C2). The increase in settlement occurred 9 (C1) and 19 days (C2) after the addition of new substrate when larvae were 69 and 80 days old in C1 and C2, respectively. There was no obvious change in settling behaviour after the room change at 11 (C1) / 12 (C2) days of running experiment (18 and 29 days old larvae) (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). Beyond a larval age of ~\u0026thinsp;95 days no more additional settling was observed. Figure\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003e shows the average counts of survivors (larvae\u0026thinsp;+\u0026thinsp;settlers) over the same range of time. The number of survivors shows a steady decrease until the same point in time as above (~\u0026thinsp;95 days) and then turns to an almost constant level. This indicates that larvae are competent to settle during the entire pelagic larval duration, until all larvae have either settled or died.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eSubstrate choice appeared to vary over time during the settlement period in both Cohorts 1 and 2, as well as between cohorts. Initially, settlers appeared to preferentially attach to bare gorgonian skeleton, but this preference decreased over time, with larvae at later stages of development settling more on the rocky substrates provided (Fig.\u0026nbsp;\u003cspan refid=\"Fig9\" class=\"InternalRef\"\u003e9\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eIn cohort 1 (C1, orange colour in Fig.\u0026nbsp;\u003cspan refid=\"Fig9\" class=\"InternalRef\"\u003e9\u003c/span\u003e), the maximum number of settlers on rock was 9, on CCA (coralline crusts) was 4, and on gorgonin was 6, observed on days 97 and 100. During days 15 to 21, settlement was exclusively on CCA and gorgonin. From days 24 to 55, settlers were observed only on rock and CCA, with the number of settlers on rock slowly increasing from 1 to 9, and on CCA from 1 to 4. From day 85 onward, settlers appeared on all three substrates, with the number of settlers on gorgonin increasing from 4 to 6 by day 97. This number remained constant until the experiment's conclusion on day 100.\u003c/p\u003e \u003cp\u003eIn cohort 2 (C2, pink colour in Fig.\u0026nbsp;\u003cspan refid=\"Fig9\" class=\"InternalRef\"\u003e9\u003c/span\u003e), the highest number of settlers was recorded on days 73 and 80, with 19 settlers on rock, 2 on CCA, and 8 on gorgonin. During days 17 to 21, attachment was only observed on gorgonin. From day 24 onward, settlement was observed on both rock (settlers increased from 1 to 19 by day 73) and gorgonin (settlers increased from 1 to 8 by day 73). Settlement on CCA was minimal, with only 1 to 2 settlers recorded on days 49, 73, 80, and 89. On day 89, the number of settlers on rock decreased slightly to 17 while the number of settlers on gorgonin remained constant. Only 1 settler remained on CCA.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec26\" class=\"Section3\"\u003e \u003ch2\u003eRecruits Survival (Experiment 2)\u003c/h2\u003e \u003cp\u003eFrom the 526 eggs monitored at Ramalhete Marine Station in 3 settlement batches, 56 larvae (11%) settled. After 4 months, 33 of these settlers were still alive, yielding a survival rate of 59% of settlers. At more than 6 months post-settlement, 36% of initial settlers were still alive resulting in an overall survival rate from egg to 6\u0026ndash;7 months post-settlement of 3.8% (see Fig.\u0026nbsp;\u003cspan refid=\"Fig10\" class=\"InternalRef\"\u003e10\u003c/span\u003ea for data on individual cohorts).\u003c/p\u003e \u003cp\u003eOut of the 930 larvae monitored across 4 batches, 61 (6.6%) successfully settled (see Fig.\u0026nbsp;\u003cspan refid=\"Fig10\" class=\"InternalRef\"\u003e10\u003c/span\u003eb for data on individual cohorts). After 4 months post-settlement, 35 of these settlers were still alive, and after more than 6 months 17 settlers were still alive resulting in a survival rate of 28% of the settlers. This corresponds to an overall survival rate of 1.8% from the original 930 larvae.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eOut of 1621 eggs, 29 larvae settled at Ocean\u0026aacute;rio de Lisboa within 4\u0026ndash;5 months after fertilization (1.8%). 6\u0026ndash;7 months after fertilization 25 settlers were still alive, corresponding to a survival rate of the settlers of 86% and a total survival rate after 6\u0026ndash;7 month of 1.5%. At 9 months after fertilization, some mortality was detected, and the survival rate decreased to 59% of the settlers (1.0% total). 10 settlers were still alive 12 months after fertilization, corresponding to a survival rate of 34% within the settlers (0.61% total).\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e"},{"header":"DISCUSSION","content":"\u003cdiv id=\"Sec28\" class=\"Section2\"\u003e \u003ch2\u003eReproductive Cycle and Spawning\u003c/h2\u003e \u003cp\u003eThe histological investigations revealed that female oocytes were already close to mature (stage 4) at least 1.5 months before spawning. In contrast, the spermaries seemed to ripen more quickly from immature (stage 2) to nearly mature (stage 3) while becoming noticeably larger within the same period. These findings suggest a distinct difference in the maturation timelines of oocytes and spermaries. Similar patterns have been observed in the related species such as \u003cem\u003eEunicella singularis\u003c/em\u003e and \u003cem\u003eParamuricea clavata\u003c/em\u003e from the Mediterranean (Coma et al. 1995b; Ribes et al. 2007; Weinberg \u0026amp; Weinberg 1979) and in \u003cem\u003eE. verrucosa\u003c/em\u003e populations in the UK (Munro 2004), as well as in other octocorals (reviewed in Kahng et al. 2011). Polyp dissection and histologic section data indicated a spawning window in late summer/early autumn, later confirmed by \u003cem\u003eex situ\u003c/em\u003e observations in aquaria. This contrasts with Mediterranean gorgonians, including \u003cem\u003eE. singularis\u003c/em\u003e, and \u003cem\u003eEunicella cavolini\u003c/em\u003e, which spawn in spring or early summer (Gori et al. 2012; Ribes et al. 2007; von Koch 1887; Weinberg \u0026amp; Weinberg 1979). In the UK, \u003cem\u003eE. verrucosa\u003c/em\u003e colonies have been inferred to spawn in August or September but no direct observations were ever made (Munro 2004). The difference between the water bodies of Atlantic Ocean and Mediterranean Sea in terms of temperature regimes and stratification, currents and tides likely explain the variation in timing. Regional variation in the timing of spawning of coral populations is a well described phenomenon in corals, also across the Mediterranean Sea, and is often related to temperature variations (Foster \u0026amp; Gilmour 2020; Kersting et al. 2013; Osman et al. 2024; Sakai et al. 2024).Several environmental cues and variables are known to influence spawning in tropical corals, including solar insolation, moonlight intensity, day length, and temperature (Hatta et al. 1999; Kaniewska et al. 2015; Paxton et al. 2016; Van Woesik et al. 2006). However, for cold- and deep-water species, where some of these cues may be absent or differ significantly, our understanding is limited, partially due to the challenges of studying these species at depth. In our study, coral colonies were maintained in both indoor and outdoor tank systems. The indoor system was kept at a stable temperature of 14\u0026ndash;16\u0026deg;C, while the outdoor system, although cooled, experienced natural temperature fluctuations that exceeded 18\u0026deg;C during the summer. Despite these differences, spawning occurred simultaneously in both systems in at least one of the major spawning events (September 12\u0026ndash;13), suggesting that temperature is not the primary driver of spawning synchrony, although it may still influence the gamete maturation. Interestingly, colonies collected in previous years and maintained for 1\u0026ndash;2 reproductive cycles under different conditions at the ODL spawned during the same general period, though not on the same exact days. The natural seasonal temperature fluctuations and moon cycle (i.e. light intensity) were simulated within ODL\u0026rsquo;s closed tank system to mirror \u003cem\u003ein situ\u003c/em\u003e conditions as best as possible based on available information, which supports the hypothesis that temperature plays an important role in regulating the annual gametogenic cycle of corals kept in captivity (Sakai et al. 2024). The incomplete synchronization of the time of spawning between colonies kept at CCMAR (two tank systems) and ODL likely result from fine-tune responses of corals to other environmental conditions specific to each captivity system, as well as to inter-colony variability (Gilmour et al. 2016; Monfared et al. 2023).\u003c/p\u003e \u003cp\u003eThe spawning period of \u003cem\u003eEunicella verrucosa\u003c/em\u003e lasted several weeks and appeared to be tentatively correlated with the lunar cycle, with peak spawning activity occurring around the full and new moons for colonies kept at CCMAR. Extended spawning periods and correlation with the lunar cycle have been reported for some temperate and cold-water corals (Coma et al. 1995a; Gori et al. 2012; reviewed in Waller et al. 2023; Weinberg \u0026amp; Weinberg 1979), and moonlight is widely recognized as a likely environmental cue triggering spawning in broadcast spawning corals (Kaniewska et al. 2015; Randall et al. 2020; Sorek \u0026amp; Levy 2014). While many cold-water corals inhabit depths beyond the reach of moonlight, \u003cem\u003eE. verrucosa\u003c/em\u003e spans a wide depth range (~\u0026thinsp;10\u0026ndash;200 m depth) (Grasshoff 1992), including shallow-water habitats where moonlight can still be detected (Kaartvedt et al. 2019). This suggests that spawning of \u003cem\u003eE. verrucosa\u003c/em\u003e may be influenced by the lunar patterns as observed in other temperate gorgonians like \u003cem\u003eP. clavata\u003c/em\u003e (Coma et al. 1995a), but likely relies on a combination of additional cues to regulate the exact timing of spawning. At ODL, the spawning showed a less pronounced correlation with the lunar phase, with two of the three spawning peaks occurring over two consecutive days 6\u0026ndash;7 days after the full moon (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). This correlation, which differs slightly from the observations made at CCMAR, appear to suggest an effect of the artificial moonlight provided at ODL on the time of spawning: in 3 nights of major spawning recorded at the ODL\u0026rsquo;s closed system, egg release was observed to start between 5:15 hrs and 6:10 hrs after the simulated moon rise. On the other hand, at CCMAR only the outdoor tank system was exposed to direct moonlight while the indoor system was not, yet spawning still occurred during the full and new moon phases in both systems. This indicates that moonlight is not the sole trigger for spawning; gravitational forces during these lunar phases may also play a significant role, or the internal biological clock of the corals may remain entrained, even in the absence of external cues. The observed temporal shift between the spawning maxima of CCMAR and ODL may be related to the fact that the CCMAR samples were relatively fresh, whereas the ODL samples had been kept in tanks for 1\u0026ndash;2 years.\u003c/p\u003e \u003cp\u003eOur observations confirm that \u003cem\u003eEunicella verrucosa\u003c/em\u003e is a broadcast spawner, as previously suggested by (Munro 2004). This contrasts with other members of the genus like \u003cem\u003eE. singularis\u003c/em\u003e and \u003cem\u003eE. cavolini\u003c/em\u003e, which are internal brooders (Ribes et al. 2007; Theodor 1967; von Koch 1887; Weinberg \u0026amp; Weinberg 1979) further highlighting that reproductive mode is relatively plastic in corals (Kahng et al. 2011; Kerr et al. 2011). While broadcast spawning is the dominant reproductive strategy in shallow-water scleractinian corals, approximately half of the studied octocoral species are internal or surface brooders (Kahng et al. 2011). Fewer deep-water species, however, have been studied, making it difficult to identify broader patterns (Eckelbarger et al. 1998; Rakka et al. 2021; Rakka et al. 2017; Sun et al. 2010; Sun et al. 2009; Sun et al. 2011; Waller 2005; Waller et al. 2023).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec29\" class=\"Section2\"\u003e \u003ch2\u003eEmbryonic development\u003c/h2\u003e \u003cp\u003eEmbryonic development in \u003cem\u003eE. verrucosa\u003c/em\u003e showed several notable differences in comparison with other members of the genus. First, while egg segmentation in closely related species like \u003cem\u003eE. singularis\u003c/em\u003e is described as holoblastic (Weinberg \u0026amp; Weinberg 1979) in which the eggs cleave into two equal-sized blastomeres that then further divide into 4, 16, and so on until the gastrulation stage, we observed partial (meroblastic) cleavage in \u003cem\u003eE. verrucosa\u003c/em\u003e. Most of the yolk remained in the original egg cell and the cleavage process was incomplete, with blastomeres not fully separated by membranes (Brun-Usan \u0026amp; Salazar-Ciudada 2020). Most studies on coral embryonic development report holoblastic cleavage patterns (Brun-Usan \u0026amp; Salazar-Ciudada 2020; Linares et al. 2008; Okubo et al. 2013; Rakka et al. 2021), similar to what has been observed in \u003cem\u003eE. singularis\u003c/em\u003e. However, it is not unusual for cleavage patterns to vary across closely related species, particularly among nonbilaterian invertebrates like cnidarians (Brun-Usan \u0026amp; Salazar-Ciudada 2020). Meroblastic cleavage is common in yolk-rich eggs and occurs because the dense yolk interferes with the formation of the cytoskeleton during cell division (Adamska et al. 2011; Martin 1997). This suggests that the eggs of \u003cem\u003eE. verrucosa\u003c/em\u003e exhibit this pattern due to high yolk content. The presumed high lipid content remains to be assessed but may help explain the long pelagic larval duration (PLD) for the species in this study (Viladrich et al. 2021).\u003c/p\u003e \u003cp\u003eIn other cnidarians, chaotic cleavages and temporal syncytial stages\u0026mdash;where random blastomeres fuse\u0026mdash;have been observed, resulting in different cleavage patterns even between individuals (Brun-Usan \u0026amp; Salazar-Ciudada 2020). Such cleavage patterns may help explain the observations made for \u003cem\u003eE. verrucosa\u003c/em\u003e embryos, where partially cleaved embryos containing one large blastomere and four to five smaller ones, sometimes fused back into a single large cell before progressing to the final cleavage stage, which produced 8 blastomeres. Between the 8- and 32-cell stages, the embryos followed a more organized cleavage pattern. However, beyond the 32-cell stage, the embryos became asymmetrical again, with some blastomeres remaining very large while others were much smaller. These variations were seen among multiple individual embryos. As cell sizes became more uniform and the embryos developed regular 6-sided shapes, the embryos exhibited relatively deep and extreme infoldings reminiscent of a tangle more than the \"raisin-like\" formation, that has been described in other tropical gorgonians (Lasker, personal communication, Lasker \u0026amp; Kim 1996; Tonra et al. 2021).\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eLarval development, behaviour and settlement\u003c/h3\u003e\n\u003cp\u003eThe eggs (300\u0026ndash;400 \u0026micro;m diameter) and larvae (500\u0026ndash;750 \u0026micro;m length, 250\u0026ndash;350 \u0026micro;m width) of \u003cem\u003eE. verrucosa\u003c/em\u003e were larger than those of the surface-brooding species like \u003cem\u003eP. clavata\u003c/em\u003e (eggs 250\u0026ndash;350, larvae 500\u0026ndash;800 \u0026micro;m in length but very thin, Linares et al. 2008), but smaller than those of the internal brooder \u003cem\u003eE. singularis\u003c/em\u003e (larvae 2500 \u0026micro;m in length, 500 \u0026micro;m in width, Weinberg \u0026amp; Weinberg 1979) from the Mediterranean Sea. Although larval behavioural traits were not quantified, some preliminary qualitative findings are worth noting. Similar to many other broadcast spawning coral species, the eggs of \u003cem\u003eE. verrucosa\u003c/em\u003e were mostly positively buoyant for a few hours. However, the embryos became negatively buoyant once they started cleaving and remained so for up to three days until the onset of larval swimming activity. A significant decrease in propagule buoyancy during embryogenesis has been previously reported (e.g., Coelho \u0026amp; Lasker 2016a) and is likely the result of the temporal decrease in lipid content that occurs in several species of broadcast spawning corals as larvae develop (Figueiredo et al. 2012; Harii et al. 2007). The larvae of \u003cem\u003eE. verrucosa\u003c/em\u003e were consistently observed to swim upwards, especially in the first two weeks, suggesting active swimming behaviour that likely compensates for the decrease in buoyancy (although this has not been tested). SEM images of \u003cem\u003eE. verrucosa\u003c/em\u003e larvae during this stage revealed that ciliation appeared incomplete. At a later stage, the larvae continued swimming actively, but with more up-and-downward movement including substrate probing behaviour. The further development of cilia and changes in swimming behaviour seemed crucial for larvae to become more agile and competent to probe the substrate and settle.\u003c/p\u003e \u003cp\u003eDespite offering settlement substrate to larvae at two different points in time (Cohort 1, 7 day-old larvae \u003cem\u003evs.\u003c/em\u003e Cohort 2, 17 day-old larvae), both cohorts began substrate probing and settlement around a similar time at days 15 and 19, respectively (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). While studies documenting settlement dynamics in temperate and cold-water coral species remain sparse, especially in species that broadcast spawn, such a prolonged delay in the onset of competency to settle has been previously documented in a deep-sea and temperate octocorals (Rakka et al. 2021; Zelli et al. 2020). Estimating the onset of settlement competency in suboptimal \u003cem\u003eex situ\u003c/em\u003e conditions compared to the wild remains challenging, and we cannot exclude that our estimate is potentially an overestimate of the duration of the pre-competency period in nature. However, our data seemed remarkably consistent across two different larval cohorts used in the settlement experiments conducted at CCMAR and an additional cohort followed in the entirely separate system at t ODL, which suggests delayed onset of settlement may be common in temperate and cold-water broadcast spawners presumably due to a longer larval development period as a result at low ambient seawater temperature.\u003c/p\u003e \u003cp\u003eOne factor that determines behaviour and settlement of the larvae is the potential survival of the larvae in the water under laboratory conditions (PLD), which can be quite long with some species. For example, \u003cem\u003eCorallium rubrum\u003c/em\u003e larvae can survive for up to 42 days (Mart\u0026iacute;nez-Quintana et al. 2015), \u003cem\u003eP. clavata\u003c/em\u003e maximal 64 days and \u003cem\u003eE. singularis\u003c/em\u003e\u0026thinsp;\u0026gt;\u0026thinsp;78 days (Guizien et al. 2020). (von Koch 1887) also observed that \u003cem\u003eE. cavolini\u003c/em\u003e larvae could remain in the water column for months under water flow. In contrast to such previous studies on temperate and cold-water octocorals, our observations for \u003cem\u003eE. verrucosa\u003c/em\u003e showed that despite overall low settlement rates, larvae remained competent to settle for a relatively long period, with a maximum PLD of up to 132 days under laboratory conditions. For example, larvae of the Mediterranean internal brooder \u003cem\u003eE. singularis\u003c/em\u003e can settle within 13 days (Theodor 1967; Weinberg \u0026amp; Weinberg 1979; Zelli et al. 2020), and \u003cem\u003eP. clavata\u003c/em\u003e have been observed to metamorphose into polyps without attaching to the substratum between 8\u0026ndash;25 days (Linares et al. 2008). For \u003cem\u003eE. verrucosa\u003c/em\u003e, we only recorded the total number of settlers at any given monitoring time without tracking individual recruits, so there may have been an underestimation of settlement if some settlers died unnoticed. In addition, our observation that \u003cem\u003eE. verrucosa\u003c/em\u003e has an extremely long PLD is concordant with the mode of reproduction. Broadcast spawners typically have longer PLDs than brooding species (Harrison \u0026amp; Wallace 1990). Furthermore, it is in accordance with the patterns of embryonic cleavage reported here, suggesting extremely yolk-rich propagules that may enable a long settlement competency period.\u003c/p\u003e \u003cp\u003eCoral larvae settlement is known to be influenced by various complex environmental and biological cues. These cues are not yet fully understood, and knowledge is predominantly focused on tropical shallow-water coral species. These cues can be chemical or physical signals, with chemical compounds produced by crustose coralline algae (CCA) and/or the microbial communities and biofilms associated with the substrate having previously shown to induce settlement in multiple species of corals (G\u0026oacute;mez-Lemos et al. 2018; Sneed et al. 2014; Tebben et al. 2015). We therefore selected substrates previously shown to encourage coral settlement in temperate octocorals: gorgonian bare skeleton for \u003cem\u003eE. singularis\u003c/em\u003e (Weinberg \u0026amp; Weinberg 1979), CCA covered rocks for \u003cem\u003eC. rubrum\u003c/em\u003e (Zelli et al. 2020) and rocks from the same area as the breeding stock used in our experiments. However, no clear preference for any substrate was observed, only trends that shifted with larval age (Fig.\u0026nbsp;\u003cspan refid=\"Fig9\" class=\"InternalRef\"\u003e9\u003c/span\u003e). The hierarchical induction of settlement through certain cues is subject to a number of recent studies (Jorissen et al. 2021; Petersen et al. 2023; Wahab et al. 2023). Some CCA species may even provide positive microbial cues that initiate the settlement process, but then also deter attachment and metamorphosis, leading larvae to choose alternative substrates (Jorissen et al. 2021).\u003c/p\u003e \u003cp\u003e \u003cem\u003eE. verrucosa\u003c/em\u003e does not appear to be limited to a particular substrate type, and despite intense probing behaviour, the larvae did not avoid any of the three substrate types provided. Initially, however, the larvae of both cohorts seemed to show a preference for the gorgonian skeleton. Later, the majority of settlers chose to settle on rock, with fewer settling on the gorgonian skeleton and CCA. The gorgonian skeleton may have provided strong cues at first, but these cues might have faded, or the larvae may have determined, after initial attachment and mechanical testing, that the material quality was insufficient for settlement. The lower number of settlers on the gorgonian skeleton and CCA could also be due to post-settlement detachment of recruits. For example, the gorgonian skeleton develops a slimy biofilm, and living CCA has defence mechanisms to avoid overgrowth, such as chemical defences and the sloughing off from epithelial cells (G\u0026oacute;mez-Lemos \u0026amp; D\u0026iacute;az-Pulido 2017). The rock, on the other hand, was not bare but covered with a diverse array of microorganisms. Therefore, the settlement cue was likely a combination of abiotic factors (surface structure) and biotic cues (microfilm and bioactive molecules produced by organisms). Since we did not test the substrates individually, we cannot rule out cross-influences among them, and future experiments are necessary to further investigate settlement cues.\u003c/p\u003e \u003cp\u003eOnce larvae settled and started metamorphosis, survival rates were high, but growth was slow (2-3mm in length in the first 3 months, budding after 9 months post settlement) compared with the description of von Koch (1887) on \u003cem\u003eE. cavolini\u003c/em\u003e who observed the formation of an internal skeleton already after several weeks and budding shortly after.\u003c/p\u003e \u003cdiv id=\"Sec31\" class=\"Section2\"\u003e \u003ch2\u003eImplications on Larval dispersal and connectivity\u003c/h2\u003e \u003cp\u003eIncorporating population dynamics and connectivity patterns into marine conservation and restoration strategies, especially for sessile invertebrates like corals, is critical for effective biodiversity preservation (Jones et al. 2007; Marti-Puig et al. 2013; Possingham et al. 2015). The primary goal of coral restoration efforts is to rehabilitate a degraded habitat by reintroducing a healthy, self-sustaining breeding population, based on a robust, scientifically grounded restoration framework. However, such frameworks are largely absent in most coral restoration projects (Bostr\u0026ouml;m-Einarsson et al. 2020; Mcdonald et al. 2016). For instance, dispersal potential and patterns of population connectivity are crucial considerations for conservation, yet key biological traits that influence larval dispersal such as planktonic larval duration (PLD), larval behaviour (e.g., buoyancy and swimming), and reproductive strategies (brooding vs. spawning), are often unknown or remain overlooked in restoration planning and implementation (Coelho \u0026amp; Lasker 2016b; Cowen et al. 2007; Cowen \u0026amp; Sponaugle 2009; Randall et al. 2020; Suggett \u0026amp; Van Oppen 2022). Our findings reveal that the PLD of \u003cem\u003eE. verrucosa\u003c/em\u003e is exceptionally long as is the settlement competency period, which started late (earliest at day 15). This suggests that coral larvae can disperse over a large distance, which has important implications for population connectivity.\u003c/p\u003e \u003cp\u003eHowever, genetic studies by Holland et al. (2017) detected significant genetic structure at regional scale suggesting that dispersal over 500-2000km is infrequent. And the more recent genome-wide analyses by (Macleod et al. 2024) further highlighted regional clustering and a pattern of isolation by distance. In a subsequent analysis, Macleod et al. (2024) modeled larval dispersal under two hypothetical PLDs (14 and 21 days) due to the absence of data for the species and to represent the central ranges of PLDs observed in other octocorals. These assumptions deviate considerably from our observations under laboratory conditions. While the effective PLD of \u003cem\u003eE. verrucosa\u003c/em\u003e in the field may be lower than that observed in the laboratory due to larval mortality (e.g., predation, drift to unsuitable areas) (Sciascia et al. 2022), settling competency most likely only starts above 15 days of age and has to potential to extend up to 4 months. Thus, the estimated mean and maximum dispersal distances obtained by Macleod et al. (2024) likely underestimate dispersal potential and the frequency of \u0026lsquo;rare\u0026rsquo; dispersal events, thus highlighting the shortcomings of generalizing biological traits in modelling studies of dispersal and population connectivity that may be used for informing conservation practitioners.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec32\" class=\"Section2\"\u003e \u003ch2\u003eFirst steps into restoration through sexual propagation\u003c/h2\u003e \u003cp\u003eWhile data on reproductive and early life ecological traits are crucial for a better understanding of the biology and population dynamics of \u003cem\u003eE. verrucosa\u003c/em\u003e in particular, and corals in general, this information is also the base for our overall goal: developing methods and workflows for restoration through sexual propagation.\u003c/p\u003e \u003cdiv id=\"Sec33\" class=\"Section3\"\u003e \u003ch2\u003eCoral collection and rearing of parent colonies\u003c/h2\u003e \u003cp\u003eWhile numerous approaches have been developed to obtain coral offspring from tropical scleractinian corals (see Randall et al. 2020 for an overview), methods for octocorals and especially cold- or deep-water coral species remain limited (Fava et al. 2010; Montseny et al. 2021a). On-site collection is only feasible for species with predictable spawning times and those located at dive-accessible depths. Obtaining offspring from aquarium-kept corals caught as fisheries bycatch offers the advantage of accessing large biomass that would otherwise perish, resulting in a substantial spawning output and diverse genotypes, providing a good representation of the population (Montseny et al. 2021b). However, relying on destructive fishing-techniques is unsustainable and should generally not be encouraged as the impact of fishing practices should be mitigated and/or removed.\u003c/p\u003e \u003cp\u003eThe collection of parent corals from bycatch is opportunistic, dependent on the availability, timing, location, and depth of fishing activities, all of which are uncontrollable factors. Additionally, the condition of the colonies can vary, as they are often entangled and damaged in nets, experiencing high stress levels that may negatively affect reproductive output and gamete nutritional reserves. However, we view coral bycatch as a valuable opportunity to establish a large-scale coral nursery that could be maintained for many years, providing a consistent annual supply of offspring. Although the corals at CCMAR were kept under less controlled conditions of lighting and temperatures, and other factors such as feeding with frozen food, several colonies produced eggs the following year, indicating the persistence of their gametogenic cycle despite being kept in suboptimal conditions. In contrast, at the Ocean\u0026aacute;rio de Lisboa, the breeding colonies were maintained over 1\u0026ndash;2 years in a closed system with precise management of water quality, high-quality food, and regulation of key environmental parameters such as moonlight and temperature. This led not only to the completion of the reproductive cycle but also to spawning, settlement, and the development of healthy recruits. This demonstrates the potential for \u003cem\u003eex-situ\u003c/em\u003e sexual reproduction of \u003cem\u003eE. verrucosa\u003c/em\u003e, eliminating the need to collect adult corals annually, which is crucial in the event of limited bycatch availability or depletion of wild populations due to anthropogenic or natural disasters.\u003c/p\u003e \u003cp\u003eAlthough propagule release and/or capture efficiency was higher at Ocean\u0026aacute;rio (~\u0026thinsp;5500 eggs from 15 colonies) compared to the Ramalhete Marine Station (~\u0026thinsp;3000 eggs from 70 colonies), settlement success was significantly greater at Ramalhete. This discrepancy may be attributed to less suitable substrates or less competent larvae at Ocean\u0026aacute;rio, possibly due to suboptimal maintenance conditions and insufficient energy reserves in the cultured colonies, as opposed to the fresh, wild-collected colonies used for the experiemnts conducted at CCMAR. This suggests that keeping colonies in captivity in the long-term may not yet be the most-efficient restoration strategy. Future studies should therefore focus on improving reproductive output, enhancing settlement success, and increasing recruit survival. This could be achieved by boosting nutrient input for parent colonies, providing adequate space for individuals, optimizing temperature control, and adjusting lighting to better replicate their natural environment year-round\u0026mdash;measures likely to improve both adult survival and offspring production. Meeting the nutritional needs of deep and cold-water corals remains a major challenge, given their reliance on external food sources. Although this has been the focus of several studies (Cocito et al. 2013; Gori et al. 2012; Rakka et al. 2021; Ribes et al. 1999), optimal feeding regimes have yet to be developed.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec34\" class=\"Section3\"\u003e \u003ch2\u003eReproduction, settlement and survival of recruits in captivity\u003c/h2\u003e \u003cp\u003eThe broadcast spawning of eggs and their release over an extended period in \u003cem\u003eE. verrucosa\u003c/em\u003e offers significant advantages for restoration efforts. Broadcast spawning is a mode of reproduction that is far more feasible for egg collection and rearing, whereas brooding species are more challenging in this regard (Randall et al. 2020). We collected spawn using air-driven suction traps positioned in the tank, by filters in the outflow, and manually retrieved eggs with a pipette. Although all methods worked similarly well, pipetting proved to be highly time-consuming, whereas a portion of the eggs or embryos collected in the filters were observed to have been squeezed and damaged. Additionally, we were unable to quantify the number of eggs potentially lost due to contact with the walls or air bubbles. The trade-off between methods highly depends on the availability of human resources; however, filters and traps, particularly with enhancements, proved highly effective during spawning events when no human presence was available such as during night-time release. In many species, the limitation of spawning events to just one or a few nights per year represents a major bottleneck (Randall et al. 2020). However, with \u003cem\u003eE. verrucosa\u003c/em\u003e, the prolonged release over more than a month allows for more flexible human and facility resources. Investigating the drivers behind \u003cem\u003eE. verrucosa\u003c/em\u003e's spawning synchronicity would further enhance the predictability and management of spawn collection. However, this may require several more years of observation and experimenting.\u003c/p\u003e \u003cp\u003eDuring embryo and larval rearing, maintaining the larvae in large plastic containers (5\u0026ndash;10 L) with gentle aeration seemed to result in good survival rates. We observed larval settlement of up to 16% (Fig.\u0026nbsp;\u003cspan refid=\"Fig10\" class=\"InternalRef\"\u003e10\u003c/span\u003e), which, while lower than the much higher settlement rates reported for tropical corals (Jorissen et al. 2021; Tebben et al. 2015), represents a successful settlement outcomes for the first trials with this species. Settlement rates in cold- and deep-water corals are generally much lower across the up-to-date studied species, and comparative analyses are challenging due to the limited reports on settlement success or data on settlement dynamics (Linares et al. 2008; Rakka et al. 2021; Weinberg \u0026amp; Weinberg 1979). An exception is the documented high settlement rate of up to 50\u0026thinsp;\u0026plusmn;\u0026thinsp;7.8% in the temperate gorgonian \u003cem\u003eE. singularis\u003c/em\u003e, although it is a shallow-water species, under laboratory conditions (Viladrich et al. 2022). Studies like Zelli et al. (2020), highlight the great potential that lies in widening our understanding on settlement cues also in temperate and cold-water corals. Settlement was the bottleneck in this study and enhancement could greatly benefit not only the reproductive success of \u003cem\u003eE. verrucosa\u003c/em\u003e in captivity but also other threatened cold-water coral species. Once the larvae settled and metamorphosed, survival rates were high over a period of 6\u0026ndash;7 months in comparison to the recruits of \u003cem\u003eP. clavata\u003c/em\u003e in the field, which were close to zero (Linares et al. 2008). Growth, however, was slow as it took 6.5\u0026ndash;12 months to develop the secondary polyps. For \u003cem\u003eE. singularis\u003c/em\u003e studied under lab conditions, for example, growth has been reported to be much faster, with budding observed after just two weeks (Weinberg \u0026amp; Weinberg 1979). Following the settlement experiments conducted at CCMAR, we did not provide the recruits with any food beyond what was available to feed adult colonies (e.g., frozen rotifers, copepods). The feeding regime for recruits kept at ODL, which included living microalgae and zooplankton, seemed to positively affect growth, with the first budding observed only after 6.5 months (compared to that observed at CCMAR at 11\u0026ndash;12 months after spawning). This underscores the importance of further research into optimal nourishment for recruits, as studies on tropical coral larvae have shown that improved feeding strategies can significantly boost early growth (Petersen et al. 2008; Toh et al. 2014). Substrate type also appeared to play a crucial role in recruit survival. Natural rock substrates generally proved to be suitable for settlement, in particular those with numerous crevices and holes. On the other hand, surface structures tended to accumulate debris, which likely negatively affected recruit survival. Nearly all surviving recruits were observed growing on smooth, exposed surfaces that remained free of debris and received adequate water flow for self-cleaning, emphasizing the importance of current-exposed, sediment-free surfaces for successful coral settlement and growth.\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e"},{"header":"CONCLUSIONS","content":"\u003cp\u003eThis study provides the first description of the spawning and early life ecology of the widely distributed, forest-forming, and vulnerable pink sea fan \u003cem\u003eE. verrucosa\u003c/em\u003e. This information is vital for managing, protecting, and restoring habitats dominated by this valuable species, also providing key data for dispersal and connectivity models. Our findings revealed late onset of competency to settle and an extended period over which larvae remained competent to settle (up to 3\u0026ndash;4 months). Our results on settlement and recruit survival also represent early steps toward breeding sexually derived corals for restoration. We demonstrate that larvae can be successfully settled onto natural substrate, recruits survive and grow under laboratory conditions, and adult colonies can be maintained in captivity through multiple reproductive cycles, completing gamete development and spawning thus allowing for the production of sexually-derived recruits \u003cem\u003eex situ\u003c/em\u003e.\u003c/p\u003e \u003cp\u003eThis study also marks the first steps toward cold-water coral breeding for this vulnerable species, laying the groundwork for establishing coral nurseries for restoration. Working with such sensitive organisms presents significant challenges in planning, so our approach prioritized flexibility, allowing us to adapt our strategy continuously. As a result, the data collected may not always be as complete or systematic as desired. Nonetheless, the findings of this study pave the way for broader research and restoration efforts. A key aspect of the study was the collaboration between a scientific research centre (CCMAR) and a major facility aimed at public visitors and education, the Ocean\u0026aacute;rio de Lisboa, highlighting the potential of such institutional collaborations as platforms for large-scale restoration initiatives.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eACKNOWLEDGEMENTS\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was funded by the 2020\u0026ndash;2021 Biodiversa+ and Water JPI joint call for research projects, under the Bio-divRestore ERA-NET Cofund (GA N\u0026deg;101003777), with the EU and the Portuguese Foundation for Science and Technology (FCT) (DivRestore/0013/2020); and by project CORALFORESTS financed by the 2022 Funda\u0026ccedil;\u0026atilde;o Belmiro de Azevedo Award in the field of Conservation, restoration and monitoring of biodiversity in Portugal. This study also received Portuguese national funds from FCT through projects UIDB/04326/2020 (DOI:10.54499/UIDB/04326/2020) and LA/P/0101/2020 (DOI:10.54499/LA/P/0101/2020). C. Egger was supported by FCT Doctoral Scholarship SFRH/BD/151455/2021. We thank Capucine R\u0026eacute;casens and Laura Balsalobre for their help examining gamete development, particularly polyp dissections and histological sectioning, as well as Elsa Couto and Deborah Power for technical and laboratory support with the histological examinations. We also thank Jo\u0026atilde;o Reis and Andr\u0026eacute; Lopes for their technical support at the\u0026nbsp;Ramalhete Marine Station. We\u0026nbsp;are especially grateful to master Casimiro and his fishing crew for the collaboration and help collecting samples for this study along the Sagres region in SW Iberia.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eSUPPLEMENTARY INFORMATION\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eSupplement 1:\u0026nbsp;Polyp dissection to infer spawning window:\u003cbr\u003e\u0026nbsp;https://doi.org/10.6084/m9.figshare.28113371\u003c/p\u003e\n\u003cp\u003eSupplement 2 (video): \u003cem\u003eEunicella verrucosa\u003c/em\u003e time lapse video of embryonal development: \u0026nbsp;https://doi.org/10.6084/m9.figshare.28070732\u003c/p\u003e\n\u003cp\u003eSupplement 3 (video): \u003cem\u003eEunicella verrucosa\u003c/em\u003e larvae swimming behaviour: https://doi.org/10.6084/m9.figshare.28070870\u003c/p\u003e\n\u003cp\u003eSupplement 4 (video): \u003cem\u003eEunicella verrucosa\u003c/em\u003e larvae testing substrate and attaching: \u0026nbsp;https://doi.org/10.6084/m9.figshare.28072046\u003c/p\u003e\n\u003ch3\u003e\u003cstrong\u003eConflict of interest\u003c/strong\u003e\u003c/h3\u003e\n\u003cp\u003eOn behalf of all authors, the corresponding author states that there is no conflict of interest.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthor information\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eChristina Egger, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal\u003c/p\u003e\n\u003cp\u003eCatarina Melo, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal\u003c/p\u003e\n\u003cp\u003eBailey Marquardt, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal\u003c/p\u003e\n\u003cp\u003eAschwin H. Engelen, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal\u003c/p\u003e\n\u003cp\u003eRoland R. Melzer, SNSB-Bavarian State Collection for Zoology, Munich, Germany\u003c/p\u003e\n\u003cp\u003eElsa Santos, Ocean\u0026aacute;rio de Lisboa, Lisboa, Portugal\u003c/p\u003e\n\u003cp\u003eMargarida Fernandes, Ocean\u0026aacute;rio de Lisboa, Lisboa, Portugal\u003c/p\u003e\n\u003cp\u003eN\u0026uacute;ria Baylina, Ocean\u0026aacute;rio de Lisboa, Lisboa, Portugal\u003c/p\u003e\n\u003cp\u003eEster A. Serrao, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal\u003c/p\u003e\n\u003cp\u003eM\u0026aacute;rcio A. Coelho, Centre of Marine Sciences (CCMAR/CIMAR LA), Campus de Gambelas, Universidade do Algarve, 8005-139 Faro, Portugal\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\u003cli\u003e\u003cspan\u003eAdamska, M., Degnan, B. M., Green, K., \u0026amp; Zwafink, C. (2011). What sponges can tell us about the evolution of developmental processes. Zoology, 114(1), 1\u0026ndash;10. https://doi.org/10.1016/j.zool.2010.10.003.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eAnkamah-Yeboah, I., Xuan, B. B., Hynes, S., \u0026amp; Armstrong, C. W. (2020). Public perceptions of deep-sea environment: Evidence from Scotland and Norway. Frontiers in Marine Science, 7, 137. https://doi.org/10.3389/fmars.2020.00137.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBaillon, S., Hamel, J.-F., Wareham, V. E., \u0026amp; Mercier, A. (2012). Deep cold-water corals as nurseries for fish larvae. Frontiers in Ecology and the Environment, 10(7), 351\u0026ndash;356.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBaums, I. B. (2008). A restoration genetics guide for coral reef conservation. Molecular Ecology, 17(12), 2796\u0026ndash;2811. https://doi.org/10.1111/j.1365-294X.2008.03787.x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBaums, I. B., Baker, A. C., Davies, S. W., Grottoli, A. G., Kenkel, C. D., Kitchen, S. A., Kuffner, I. B., LaJeunesse, T. C., Matz, M. V., \u0026amp; Miller, M. W. (2019). Considerations for maximizing the adaptive potential of restored coral populations in the western Atlantic. Ecological Applications, 29(8), e01978. https://doi.org/10.1002/eap.1978.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBayraktarov, E., Brisbane, S., Hagger, V., Smith, C. S., Wilson, K. A., Lovelock, C. E., Gillies, C. L., Steven, A. D. L., \u0026amp; Saunders, M. I. (2020). Priorities and Motivations of Marine Coastal Restoration Research. Frontiers in Marine Science, 7. https://doi.org/10.3389/fmars.2020.00484.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBayraktarov, E., Stewart-Sinclair, P. J., Brisbane, S., Bostr\u0026ouml;m‐Einarsson, L., Saunders, M. I., Lovelock, C. E., Possingham, H. P., Mumby, P. J., \u0026amp; Wilson, K. A. (2019). Motivations, Success, and Cost of Coral Reef Restoration. Restoration Ecology, 27(5), 981\u0026ndash;991. https://doi.org/10.1111/rec.12977.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBoch, C. A., DeVogelaere, A., Burton, E., King, C., Lord, J. P., Lovera, C., Litvin, S. Y., Kuhnz, L. A., \u0026amp; Barry, J. (2019). Coral Translocation as a Method to Restore Impacted Deep-Sea Coral Communities. Frontiers in Marine Science, 6. https://doi.org/10.3389/fmars.2019.00540.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBoch, C. A., \u0026amp; Morse, A. N. C. (2012). Testing the effectiveness of direct propagation techniques for coral restoration of Acropora spp. Ecological Engineering, 40, 11\u0026ndash;17. https://www.sciencedirect.com/science/article/pii/S0925857411003934.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBongiorni, L., Mea, M., Gambi, C., Pusceddu, A., Taviani, M., \u0026amp; Danovaro, R. (2010). Deep-Water Scleractinian Corals Promote Higher Biodiversity in Deep-Sea Meiofaunal Assemblages Along Continental Margins. Biological conservation, 143(7), 1687\u0026ndash;1700. https://doi.org/10.1016/j.biocon.2010.04.009.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBostr\u0026ouml;m-Einarsson, L., Babcock, R. C., Bayraktarov, E., Ceccarelli, D., Cook, N., Ferse, S. C. A., Hancock, B., Harrison, P., Hein, M., Shaver, E., Smith, A., Suggett, D., Stewart-Sinclair, P. J., Vardi, T., \u0026amp; McLeod, I. M. (2020). Coral restoration \u0026ndash; A systematic review of current methods, successes, failures and future directions. PLoS One, 15(1). https://doi.org/10.1371/journal.pone.0226631.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBrazeau, D. A., \u0026amp; Lasker, H. R. (1988). Inter-and intraspecific variation in gorgonian colony morphology: quantifying branching patterns in arborescent animals. Coral Reefs, 7, 139\u0026ndash;143. https://doi.org/10.1007/BF00300973.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBridge, T. C. L., Hoey, A. S., Campbell, S. J., Muttaqin, E., Rudi, E., Fadli, N., \u0026amp; Baird, A. H. (2013). Depth-Dependent Mortality of Reef Corals Following a Severe Bleaching Event: Implications for Thermal Refuges and Population Recovery. F1000research, 2, 187. https://doi.org/10.12688/f1000research.2-187.v1.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBrun-Usan, M., \u0026amp; Salazar-Ciudada, I. (2020). The Evolution of Cleavage in Metazoans. In L. Nuno de la Rosa \u0026amp; G. B. M\u0026uuml;ller (Eds.), Evolutionary Developmental Biology: Springer Nature Switzerland AG.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBuhl-Mortensen, L., Buhl-Mortensen, P., Rungruangsak-Torrissen, K., Schwach, V., Hjort, J., Jakobsen, T., Ozhigin, V., Bergh, \u0026Oslash;., Hamre, J., \u0026amp; Torgersen, T. (2018). Cold temperate coral habitats. In C. D. Beltran \u0026amp; E. T. Camacho (Eds.), Corals in a changing world (Vol. 9). Rijeka, Croatia: InTech.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBuhl-Mortensen, P., Buhl-Mortensen, L., \u0026amp; Purser, A. (2017). Trophic ecology and habitat provision in cold-water coral ecosystems. Marine Animal Forests. The Ecology of Benthic Biodiversity Hotspots. Springer, Cham. https://doi.org/10.1007/978-3-319-21012-4_20.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCarpenter, K. E., Abrar, M., Aeby, G. S., Aronson, R. B., Banks, S., Bruckner, A. W., Chiriboga, A., Cort\u0026eacute;s, J., Delbeek, J. C., DeVantier, L., Edgar, G. J., Edwards, A. J., Fenner, D., Guzm\u0026aacute;n, H. M., Hoeksema, B. W., Hodgson, G., Johan, O., Licuanan, W. Y., Livingstone, S. R., Lovell, E. R., Moore, J. A., Obura, D., Ochavillo, D., Polidoro, B., Precht, W. F., Quibilan, M. C. C., Reboton, C. T., Richards, Z. T., Rogers, A. D., Sanciangco, J. C., Sheppard, A., Sheppard, C., Smith, J. E., Stuart, S. N., Turak, E., Veron, J., Wallace, C. C., Weil, E., \u0026amp; Wood, E. (2008). One-Third of Reef-Building Corals Face Elevated Extinction Risk From Climate Change and Local Impacts. Science, 321(58888), 560\u0026ndash;563. https://doi.org/10.1126/science.1159196.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCarpine, C. (1963). Contribution \u0026agrave; la connaissance des Gorgones Holaxonia de la Mediterranean occidentale. Bull. Inst. oc\u0026eacute;anogr. Monaco, 60(1270), 1\u0026ndash;52.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCarpine, C., \u0026amp; Grasshoff, M. (1975). Les gorgonaires de la M\u0026eacute;diterran\u0026eacute;e. Bull. Inst. oc\u0026eacute;anogr. Monaco, 71(1430), 1-140.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eChamberland, V. F., Vermeij, M. J., Brittsan, M., Carl, M., Schick, M., Snowden, S., Schrier, A., \u0026amp; Petersen, D. (2015). Restoration of critically endangered elkhorn coral (Acropora palmata) populations using larvae reared from wild-caught gametes. Global Ecology and Conservation, 4, 526\u0026ndash;537. https://doi.org/10.1016/j.gecco.2015.10.005.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eChimienti, G. (2020). Vulnerable forests of the pink sea fan Eunicella verrucosa in the Mediterranean Sea. Diversity, 12(5), 176. https://doi.org/10.3390/d12050176.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCocito, S., Ferrier-Pag\u0026egrave;s, C., Cupido, R., Rottier, C., Meier-Augenstein, W., Kemp, H., Reynaud, S., \u0026amp; Peirano, A. (2013). Nutrient acquisition in four Mediterranean gorgonian species. Marine Ecology Progress Series, 473, 179\u0026ndash;188. https://doi.org/10.3354/meps10037.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCoelho, M. A., \u0026amp; Lasker, H. R. (2016a). Larval behavior and settlement dynamics of a ubiquitous Caribbean octocoral and its implications for dispersal. Marine Ecology Progress Series, 561, 109\u0026ndash;121. https://doi.org/10.3354/meps11941.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCoelho, M. A., \u0026amp; Lasker, H. R. (2016b). Larval dispersal and population connectivity in Anthozoans. In S. Goffredo \u0026amp; Z. Dubinsky (Eds.), The Cnidaria, Past, Present and Future: The world of Medusa and her sisters (pp. 291\u0026ndash;315): Springer.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eComa, R., Ribes, M., Zabala, M., \u0026amp; Gilil, J. (1995a). Reproduction and cycle of gonadal development in the Mediterranean gorgonian Paramuricea clavata. Mar. Ecol. Prog. Ser, 117(1\u0026ndash;3), 173\u0026ndash;183. https://doi.org/10.3354/meps117173.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eComa, R., Zabala, M., \u0026amp; Gili, J. (1995b). Sexual reproductive effort in the Mediterranean gorgonian Paramuricea clavata. Marine Ecology Progress Series, 185\u0026ndash;192. https://www.jstor.org/stable/44634830.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCowen, R. K., Gawarkiewicz, G., Pineda, J., Thorrold, S. R., \u0026amp; Werner, F. E. (2007). Population connectivity in marine systems an overview. Oceanography, 20(3), 14\u0026ndash;21. https://doi.org/10.5670/oceanog.2007.26.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCowen, R. K., \u0026amp; Sponaugle, S. (2009). Larval dispersal and marine population connectivity. Annual review of marine science, 1(1), 443\u0026ndash;466. https://doi.org/10.1146/annurev.marine.010908.163757.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCoz, R., Ouisse, V., Artero, C., Carpentier, A., Crave, A., Feunteun, E., Olivier, J.-M., Perrin, B., \u0026amp; Ysnel, F. (2012). Development of a new standardised method for sustainable monitoring of the vulnerable pink sea fan Eunicella verrucosa. Marine Biology, 159, 1375\u0026ndash;1388. https://doi.org/10.1007/s00227-012-1908-7.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDanovaro, R., Corinaldesi, C., D\u0026rsquo;Onghia, G., Galil, B. S., Gambi, C., Gooday, A. J., Lampadariou, N., Luna, G. M., Morigi, C., Olu, K., Polymenakou, P. N., Ramirez-Llodra, E., Sabbatini, A., Sard\u0026agrave;, F., Sibuet, M., \u0026amp; Τσελεπίδης, Α. (2010). Deep-Sea Biodiversity in the Mediterranean Sea: The Known, the Unknown, and the Unknowable. PLoS One, 5(8). https://doi.org/10.1371/journal.pone.0011832.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDela Cruz, D. W., \u0026amp; Harrison, P. L. (2020). Enhancing coral recruitment through assisted mass settlement of cultured coral larvae. PLoS One, 15(11), e0242847. https://doi.org/10.1371/journal.pone.0242847.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDias, V., Oliveira, F., Boavida, J., Serr\u0026atilde;o, E. A., Gon\u0026ccedil;alves, J. M., \u0026amp; Coelho, M. A. (2020). High coral bycatch in bottom-set gillnet coastal fisheries reveals rich coral habitats in southern Portugal. Frontiers in Marine Science, 7, 603438. https://doi.org/10.3389/fmars.2020.603438.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDoropoulos, C., Vons, F., Elzinga, J., Ter Hofstede, R., Salee, K., Van Koningsveld, M., \u0026amp; Babcock, R. C. (2019). Testing industrial-scale coral restoration techniques: harvesting and culturing wild coral-spawn slicks. Frontiers in Marine Science, 6, 658. https://doi.org/10.3389/fmars.2019.00658.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eEckelbarger, K., Tyler, P., \u0026amp; Langton, R. (1998). Gonadal morphology and gametogenesis in the sea pen Pennatula aculeata (Anthozoa: Pennatulacea) from the Gulf of Maine. Marine Biology, 132, 677\u0026ndash;690. https://doi.org/10.1007/s002270050432.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eEddy, T. D., Lam, V. W. Y., Reygondeau, G., Cisneros-Montemayor, A. M., Greer, K., Palomares, M. L. D., Bruno, J. F., Ota, Y., \u0026amp; Cheung, W. W. L. (2021). Global Decline in Capacity of Coral Reefs to Provide Ecosystem Services. One Earth, 4(9), 1278\u0026ndash;1285. https://doi.org/10.1016/j.oneear.2021.08.016.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFava, F., Bavestrello, G., Valisano, L., \u0026amp; Cerrano, C. (2010). Survival, growth and regeneration in explants of four temperate gorgonian species in the Mediterranean Sea. Italian Journal of Zoology, 77, 44\u0026ndash;52. https://doi.org/10.1080/11250000902769680.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFeng, E. Y., Keller, D. P., Koeve, W., \u0026amp; Oschlies, A. (2016). Could Artificial Ocean Alkalinization Protect Tropical Coral Ecosystems From Ocean Acidification? Environmental Research Letters, 11(7). https://doi.org/10.1088/1748-9326/11/7/074008.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFerrario, F., Beck, M. W., Storlazzi, C. D., Micheli, F., Shepard, C. C., \u0026amp; Airoldi, L. (2014). The Effectiveness of Coral Reefs for Coastal Hazard Risk Reduction and Adaptation. Nature Communications, 5(1). https://doi.org/10.1038/ncomms4794.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFigueiredo, J., Baird, A. H., Cohen, M. F., Flot, J.-F., Kamiki, T., Meziane, T., Tsuchiya, M., \u0026amp; Yamasaki, H. (2012). Ontogenetic change in the lipid and fatty acid composition of scleractinian coral larvae. Coral Reefs, 31, 613\u0026ndash;619. https://doi.org/10.1007/s00338-012-0874-3.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFogarty, M. J., \u0026amp; Botsford, L. W. (2007). Population connectivity and spatial management of marine fisheries. Oceanography, 20(3), 112\u0026ndash;123. http://dx.doi.org/10.5670/oceanog.2007.34.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eForrester, G. E., Chan, M., Conetta, D., Dauksis, R., Nickles, K., \u0026amp; Siravo, A. (2019). Comparing the efficiency of nursery and direct transplanting methods for restoring endangered corals. Ecological Restoration, 37(2), 81\u0026ndash;89. https://doi.org/10.3368/er.37.2.81.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFoster, T., \u0026amp; Gilmour, J. (2020). Egg Size and Fecundity of Biannually Spawning Corals at Scott Reef. Scientific Reports, 10(1). https://doi.org/10.1038/s41598-020-68289-4.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGarrison, V., \u0026amp; Greg, W. (2008). Storm-generated coral fragments\u0026ndash;A viable source of transplants for reef rehabilitation. Biological conservation, 141(12), 3089\u0026ndash;3100. https://doi.org/10.1016/j.biocon.2008.09.020.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGilmour, J. P., Underwood, J. N., Howells, E. J., Gates, E., \u0026amp; Heyward, A. J. (2016). Biannual spawning and temporal reproductive isolation in Acropora corals. PLoS One, 11(3), e0150916.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eG\u0026oacute;mez-Lemos, L. A., \u0026amp; D\u0026iacute;az-Pulido, G. (2017). Crustose Coralline Algae and Associated Microbial Biofilms Deter Seaweed Settlement on Coral Reefs. Coral Reefs, 36(2), 453\u0026ndash;462. https://doi.org/10.1007/s00338-017-1549-x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eG\u0026oacute;mez-Lemos, L. A., Doropoulos, C., Bayraktarov, E., \u0026amp; Diaz-Pulido, G. (2018). Coralline algal metabolites induce settlement and mediate the inductive effect of epiphytic microbes on coral larvae. Scientific Reports, 8(1), 17557. https://doi.org/10.1038/s41598-018-35206-9.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGori, A., Viladrich, N., Gili, J. M., Kotta, M., Cucio, C., Magni, L., Bramanti, L., \u0026amp; Rossi, S. (2012). Reproductive cycle and trophic ecology in deep versus shallow populations of the Mediterranean gorgonian Eunicella singularis (Cap de Creus, northwestern Mediterranean Sea). Coral Reefs, 31, 823\u0026ndash;837. https://doi.org/10.1007/s00338-012-0904-1.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGrasshoff, M. (1992). Die Flachwasser-Gorgonarien von Europa und Westafrika-(Cnidaria, Anthozoa) (Vol. 149). Frankfurt a. M.: Senckenbergischen Naturforschenden Gesellschaft Frankfurt a. M.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGuizien, K., Viladrich, N., Mart\u0026iacute;nez-Quintana, \u0026Aacute;., \u0026amp; Bramanti, L. (2020). Survive or swim: different relationships between migration potential and larval size in three sympatric Mediterranean octocorals. Scientific Reports, 10(1), 18096. https://doi.org/10.1038/s41598-020-75099-1.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHall-Spencer, J. M., Pike, J., \u0026amp; Munn, C. B. (2007). Diseases affect cold-water corals too: Eunicella verrucosa (Cnidaria: Gorgonacea) necrosis in SW England. Diseases of aquatic organisms, 76(2), 87\u0026ndash;97. https://doi.org/10.3354/dao076087.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHarii, S., Nadaoka, K., Yamamoto, M., \u0026amp; Iwao, K. (2007). Temporal changes in settlement, lipid content and lipid composition of larvae of the spawning hermatypic coral Acropora tenuis. Marine Ecology Progress Series, 346, 89\u0026ndash;96. https://doi.org/10.3354/meps07114.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHarrison, P. L., \u0026amp; Wallace, C. C. (1990). Reproduction, dispersal and recruitment of scleractinian corals. In Coral Reefs (Vol. 25, pp. 133\u0026ndash;207): Elsevier.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHatta, M., Fukami, H., Wang, W., Omori, M., Shimoike, K., Hayashibara, T., Ina, Y., \u0026amp; Sugiyama, T. (1999). Reproductive and genetic evidence for a reticulate evolutionary history of mass-spawning corals. Molecular Biology and Evolution, 16(11), 1607\u0026ndash;1613. https://doi.org/10.1093/oxfordjournals.molbev.a026073.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHenry, J. A., O\u0026rsquo;Neil, K. L., Pilnick, A. R., \u0026amp; Patterson, J. T. (2021). Strategies for integrating sexually propagated corals into Caribbean reef restoration: experimental results and considerations. Coral Reefs, 40(5), 1667\u0026ndash;1677. https://doi.org/10.1007/s00338-021-02154-2.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHeyward, A., Smith, L., Rees, M., \u0026amp; Field, S. (2002). Enhancement of coral recruitment by in situ mass culture of coral larvae. Marine Ecology Progress Series, 230, 113\u0026ndash;118. https://doi.org/10.3354/MEPS230113.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eH\u0026oslash;egh-Guldberg, O., Kennedy, E. V., Beyer, H. L., McClennen, C., \u0026amp; Possingham, H. P. (2018). Securing a Long-Term Future for Coral Reefs. Trends in Ecology \u0026amp; Evolution, 33(12), 936\u0026ndash;944. https://doi.org/10.1016/j.tree.2018.09.006.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHolland, L., Jenkins, T., \u0026amp; Stevens, J. (2017). Contrasting patterns of population structure and gene flow facilitate exploration of connectivity in two widely distributed temperate octocorals. Heredity, 119(1), 35\u0026ndash;48. https://doi.org/10.1038/hdy.2017.14.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHughes, A. R., Grabowski, J. H., Leslie, H. M., Scyphers, S. B., \u0026amp; Williams, S. L. (2017a). Inclusion of Biodiversity in Habitat Restoration Policy to Facilitate Ecosystem Recovery. Conservation Letters, 11(3). https://doi.org/10.1111/conl.12419.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHughes, T. P., Barnes, M. L., Bellwood, D. R., Cinner, J. E., Cumming, G. S., Jackson, J. B. C., Kleypas, J., Leemput, I. A. v. d., Lough, J., Morrison, T. H., Palumbi, S. R., Nes, E. H. v., \u0026amp; Scheffer, M. (2017b). Coral Reefs in the Anthropocene. Nature, 546(7656), 82\u0026ndash;90. https://doi.org/10.1038/nature22901.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHughes, T. P., Kerry, J. T., Baird, A. H., Connolly, S. R., Dietzel, A., Eakin, C. M., Heron, S. F., Hoey, A. S., Hoogenboom, M. O., \u0026amp; Liu, G. (2018). Global warming transforms coral reef assemblages. Nature, 556(7702), 492\u0026ndash;496. https://doi.org/10.1038/s41586-018-0041-2.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eJones, G. P., Srinivasan, M., \u0026amp; Almany, G. R. (2007). Population connectivity and conservation of marine biodiversity. Oceanography, 20(3), 100\u0026ndash;111. https://doi.org/10.5670/oceanog.2007.33.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eJorissen, H., Galand, P. E., Bonnard, I., Meiling, S. S., Raviglione, D., Meistertzheim, A.-L., H\u0026eacute;douin, L., Banaigs, B., Payri, C., \u0026amp; Nugues, M. M. (2021). Coral Larval Settlement Preferences Linked to Crustose Coralline Algae With Distinct Chemical and Microbial Signatures. Scientific Reports, 11(1). https://doi.org/10.1038/s41598-021-94096-6.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKaartvedt, S., Langbehn, T. J., \u0026amp; Aksnes, D. L. (2019). Enlightening the ocean\u0026rsquo;s twilight zone. ICES Journal of Marine Science, 76(4), 803\u0026ndash;812. https://doi.org/10.1093/icesjms/fsz010.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKahng, S. E., Benayahu, Y., \u0026amp; Lasker, H. R. (2011). Sexual reproduction in octocorals. Marine Ecology Progress Series, 443, 265\u0026ndash;283. https://doi.org/10.3354/meps09414.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKaniewska, P., Alon, S., Karako-Lampert, S., Hoegh-Guldberg, O., \u0026amp; Levy, O. (2015). Signaling cascades and the importance of moonlight in coral broadcast mass spawning. Elife, 4. https://doi.org/10.7554/eLife.09991.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKerr, A. M., Baird, A. H., \u0026amp; Hughes, T. P. (2011). Correlated evolution of sex and reproductive mode in corals (Anthozoa: Scleractinia). Proceedings of the Royal Society B: Biological Sciences, 278(1702), 75\u0026ndash;81.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKersting, D. K., Casado, C., L\u0026oacute;pez-Legentil, S., \u0026amp; Linares, C. (2013). Unexpected Patterns in the Sexual Reproduction of the Mediterranean Scleractinian Coral Cladocora Caespitosa. Marine Ecology Progress Series, 486, 165\u0026ndash;171. https://doi.org/10.3354/meps10356.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eLam, K.-W., McRae, C. J., Zhang, X.-C., Ye, Z.-M., Qiu, Y.-T., Jiang, M.-Q., Cheng, T.-H., Chen, G. K., \u0026amp; Fan, T.-Y. (2023). Consistent monthly reproduction and completion of a brooding coral life cycle through ex situ culture. Diversity, 15(2), 218. https://doi.org/10.3390/d15020218.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eLange, K., Maguer, J.-F., Reynaud, S., \u0026amp; Ferrier-Pag\u0026egrave;s, C. (2023). Nutritional Ecology of Temperate Octocorals in a Warming Ocean. Frontiers in Marine Science, 10. https://doi.org/10.3389/fmars.2023.1236164.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eLasker, H. R., \u0026amp; Kim, K. (1996). Larval development and settlement behavior of the gorgonian coral Plexaura kuna (Lasker, Kim and Coffroth). Journal of Experimental Marine Biology and Ecology, 207(1), 161\u0026ndash;175. https://doi.org/10.1016/S0022-0981(96)02625-1.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eLinares, C., Coma, R., Mariani, S., D\u0026iacute;az, D., Hereu, B., \u0026amp; Zabala, M. (2008). Early life history of the Mediterranean gorgonian Paramuricea clavata: implications for population dynamics. Invertebrate Biology, 127(1), 1\u0026ndash;11. https://doi.org/10.HH/j.1744-7410.2007.00109.x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMacleod, K. L., Jenkins, T. L., Witt, M. J., \u0026amp; Stevens, J. R. (2024). Rare, long-distance dispersal underpins genetic connectivity in the pink sea fan, Eunicella verrucosa. Evolutionary Applications, 17(3). https://doi.org/10.1111/eva.13649.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMarti-Puig, P., Costantini, F., Rugiu, L., Ponti, M., \u0026amp; Abbiati, M. (2013). Patterns of genetic connectivity in invertebrates of temperate MPA networks. Advances in Oceanography and Limnology, 4(2), 138\u0026ndash;149. https://doi.org/10.4081/AIOL.2013.5341.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMartin, V. (1997). Cnidarians, the jellyfish and hydras. In S. F. Gilbert \u0026amp; A. M. Raunio (Eds.), Embryology: construction the organism (pp. 57\u0026ndash;86). Sunderland, MA: Sinauer Associates.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMart\u0026iacute;nez-Quintana, A., Bramanti, L., Viladrich, N., Rossi, S., \u0026amp; Guizien, K. (2015). Quantification of larval traits driving connectivity: the case of Corallium rubrum (L. 1758). Marine Biology, 162, 309\u0026ndash;318. https://doi.org/10.1007/s00227-014-2599-z.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMcDonald, T., Gann, G., Jonson, J., \u0026amp; Dixon, K. (2016). International standards for the practice of ecological restoration\u0026ndash;including principles and key concepts.(Society for Ecological Restoration: Washington, DC, USA.). Soil-Tec, Inc.,\u0026copy; Marcel Huijser, Bethanie Walder. https://doi.org/10.1111/rec.13035.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMelzer, R., Spitzner, F., Šargač, Z., H\u0026ouml;rnig, M., Krieger, J., Haug, C., Haug, J., Kirchhoff, T., Meth, R., \u0026amp; Torres, G. (2021). Methods to study organogenesis in decapod crustacean larvae II: analysing cells and tissues. Helgoland Marine Research, 75(1), 2. https://doi.org/10.1186/s10152-021-00547-y.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMengerink, K. J., Van Dover, C. L., Ardron, J., Baker, M., Escobar-Briones, E., Gjerde, K., Koslow, J. A., Ramirez-Llodra, E., Lara-Lopez, A., \u0026amp; Squires, D. (2014). A call for deep-ocean stewardship. Science, 344(6185), 696\u0026ndash;698. https://doi.org/10.1126/science.1251458.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMenza, C. W., Kendall, M. S., \u0026amp; Hile, S. D. (2007). The Deeper We Go the Less We Know. Revista De Biolog\u0026iacute;a Tropical, 56(0). https://doi.org/10.15517/rbt.v56i0.5575.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMercado-Molina, A. E., \u0026amp; Suleim\u0026aacute;n-Ramos, S. E. (2023). Outplants of the Threatened Coral Acropora Cervicornis Promote Coral Recruitment in a Shallow-Water Coral Reef, Culebra, Puerto Rico. Sustainability, 15(24). https://doi.org/10.3390/su152416548.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMonfared, M. A. A., Sheridan, K., Dixon, S. P., Gledhill, M., \u0026amp; Le Berre, T. (2023). Coral Spawning Patterns of \u003cem\u003eAcropora\u003c/em\u003e Across Two Maldivian Reef Ecosystems. Peerj, 11, e16315. https://doi.org/10.7717/peerj.16315.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMontseny, M., Linares, C., Carreiro-Silva, M., Henry, L.-A., Billett, D., Cordes, E., Smith, C., Papadopoulou, N., Bilan, M., Girard, F., Burdett, H., Larsson, A., Str\u0026ouml;mberg, S., Viladrich, N., Barry, J., Baena, P., Godinho, A., Griny\u0026oacute;, J., Sant\u0026iacute;n, A., Morato, T., Sweetman, A., Gili, J., \u0026amp; Gori, A. (2021a). Active Ecological Restoration of Cold-Water Corals: Techniques, Challenges, Costs and Future Directions. Frontiers in Marine Science, 8. https://doi.org/10.3389/fmars.2021.621151.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMontseny, M., Linares, C., Viladrich, N., Biel, M., Gracias, N., Baena, P., Quintanilla, E., Ambroso, S., Griny\u0026oacute;, J., Sant\u0026iacute;n, A., Salazar, J., Carreras, M., Palomeras, N., Mag\u0026iacute;, L., Vallicrosa, G., Gili, J.-M., \u0026amp; Gori, A. (2021b). Involving fishers in scaling up the restoration of cold-water coral gardens on the Mediterranean continental shelf. Biological conservation, 262. https://www.sciencedirect.com/science/article/pii/S0006320721003530.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMontseny, M., Linares, C., Viladrich, N., Capdevila, P., Ambroso, S., D\u0026iacute;az, D., Gili, J. M., \u0026amp; Gori, A. (2020). A new large-scale and cost‐effective restoration method for cold‐water coral gardens. Aquatic Conservation: Marine and Freshwater Ecosystems, 30(5), 977\u0026ndash;987. https://doi.org/10.1002/aqc.3303.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMunro, C., \u0026amp; Munro, L. (2003). Climate change impacts on seafan populations. Reef Research, RR Report 6/2003 RR 08. https://www.marine-bio-images.com/wp-content/uploads/2020/05/Report-RR-08-Jun-2003-Climate-pdf.pdf.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMunro, L. (2004). Determining the reproductive cycle of Eunicella verrucosa. Reef Research, RR Report 3/2003 ETR 07. https://www.marine-bio-images.com/RR_Eunicella_PDFS/Report_RR12Jul2004reproductive%20cycle%20pdf.pdf.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eNajafpour, B., Dorafshan, S., Heyrati, F. P., Can\u0026aacute;rio, A. V. M., \u0026amp; Power, D. M. (2020). Comparativeo ontogeny of the digestive tract of \u003cem\u003eOncorhynchus\u003c/em\u003e Mykiss Salmo \u003cem\u003eTrutta caspius\u003c/em\u003e triploid hybrids to their Parental Species. Aquaculture Nutrition, 27(2), 427\u0026ndash;438. https://doi.org/10.1111/anu.13196.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eO\u0026rsquo;Connor, E., Hynes, S., Chen, W., Papadopoulou, N., \u0026amp; Smith, C. J. (2020). Investigating Societal Attitudes Toward Marine Ecosystem Restoration. Restoration Ecology, 29(S2). https://doi.org/10.1111/rec.13239.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eO\u0026rsquo;Neil, K. L., Serafin, R. M., Patterson, J. T., \u0026amp; Craggs, J. R. (2021). Repeated ex situ spawning in two highly disease susceptible corals in the family Meandrinidae. Frontiers in Marine Science, 8, 669976. https://doi.org/10.3389/fmars.2021.669976.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eOkubo, N., Mezaki, T., Nozawa, Y., Nakano, Y., Lien, Y. T. K., Fukami, H., Hayward, D. C., \u0026amp; Ball, E. E. (2013). Comparative Embryology of Eleven Species of Stony Corals (Scleractinia). PLoS One, 8(12), e84115.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eOrth, R. J., Lefcheck, J. S., McGlathery, K., Aoki, L. R., Luckenbach, M. W., Moore, K. A., Oreska, M. P. J., Snyder, R. A., Wilcox, D. J., \u0026amp; Lusk, B. (2020). Restoration of Seagrass Habitat Leads to Rapid Recovery of Coastal Ecosystem Services. Science Advances, 6(41). https://doi.org/10.1126/sciadv.abc6434.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eOsman, E. O., Suggett, D. J., Attalla, T. M., Casartelli, M., Cook, N., El-Sadek, I., Gallab, A., Goergen, E. A., Garcias-Bonet, N., Glanz, J. S., Pereira, P. H., Ramirez-Sanchez, M., Santoro, E. P., Stead, A., Yoder, S., Benzoni, F., \u0026amp; Peixoto, R. S. (2024). Spatial Variation in Spawning Timing for Multi-Species Acropora Assemblages in the Red Sea. Frontiers in Marine Science, 11. https://doi.org/10.3389/fmars.2024.1333621.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eOunanian, K., Carballo-C\u0026aacute;rdenas, E. C., Tatenhove, J. P. M. v., Delaney, A., Papadopoulou, N., \u0026amp; Smith, C. J. (2018). Governing Marine Ecosystem Restoration: The Role of Discourses and Uncertainties. Marine Policy, 96, 136\u0026ndash;144. https://doi.org/10.1016/j.marpol.2018.08.014.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePandolfi, J. M., Bradbury, R. H., Sala, E., Hughes, T. P., Bjorndal, K. A., Cooke, R. G., McArdle, D., McClenachan, L., Newman, M. J., \u0026amp; Paredes, G. (2003). Global trajectories of the long-term decline of coral reef ecosystems. Science, 301(5635), 955\u0026ndash;958. https://doi.org/10.1126/science.1085706.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePaxton, C. W., Baria, M. V. B., Weis, V. M., \u0026amp; Harii, S. (2016). Effect of elevated temperature on fecundity and reproductive timing in the coral Acropora digitifera. Zygote, 24(4), 511\u0026ndash;516. https://doi.org/10.1017/S0967199415000477.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePendleton, L. H., Comte, A., Langdon, C., Ekstrom, J. A., Cooley, S. R., Suatoni, L., Beck, M. W., Brander, L., Burke, L., Cinner, J. E., Doherty, C., Edwards, P., Gledhill, D. K., Jiang, L. Q., Hooidonk, R. J. V., Teh, L., Waldbusser, G. G., \u0026amp; Ritter, J. (2016). Coral Reefs and People in a High-Co2 World: Where Can Science Make a Difference to People? PLoS One, 11(11), e0164699. https://doi.org/10.1371/journal.pone.0164699.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePetersen, D., Wietheger, A., \u0026amp; Laterveer, M. (2008). Influence of different food sources on the initial development of sexual recruits of reefbuilding corals in aquaculture. Aquaculture, 277(3), 174\u0026ndash;178. https://doi.org/10.1016/j.aquaculture.2008.02.034.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePetersen, L.-E., Kellermann, M. Y., Fiegel, L. J., Nietzer, S., Bickmeyer, U., Abele, D., \u0026amp; Schupp, P. J. (2023). Photodegradation of a Bacterial Pigment and Resulting Hydrogen Peroxide Release Enable Coral Settlement. Scientific Reports, 13(1). https://doi.org/10.1038/s41598-023-30470-w.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePikesley, S. K., Godley, B. J., Latham, H., Richardson, P. B., Robson, L. M., Solandt, J.-L., Trundle, C., Wood, C., \u0026amp; Witt, M. J. (2016). Pink sea fans (Eunicella verrucosa) as indicators of the spatial efficacy of Marine Protected Areas in southwest UK coastal waters. Marine Policy, 64, 38\u0026ndash;45. https://doi.org/10.1016/j.marpol.2015.10.010.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePlucer-Rosario, G., \u0026amp; Randall, R. H. (1987). Preservation of rare coral species by transplantation and examination of their recruitment and growth. Bulletin of Marine Science, 41(2), 585\u0026ndash;593.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePollock, F. J., Katz, S. M., Water, J. A. J. M. v. d., Davies, S. W., Hein, M. Y., Torda, G., Matz, M. V., Beltran, V. H., Buerger, P., Puill-Stephan, E., Abrego, D., Bourne, D. G., \u0026amp; Willis, B. L. (2017). Coral Larvae for Restoration and Research: A Large-Scale Method for Rearing \u003cem\u003eAcropora Millepora\u003c/em\u003e Larvae, Inducing Settlement, and Establishing Symbiosis. Peerj. https://doi.org/10.7717/peerj.3732.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePossingham, H. P., Bode, M., \u0026amp; Klein, C. J. (2015). Optimal conservation outcomes require both restoration and protection. PLoS biology, 13(1), e1002052. https://doi.org/10.1371/journal.pbio.1002052.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePratchett, M. S., Bay, L. K., Coker, D. J., Cole, A. J., \u0026amp; Lawton, R. (2012). Effects of Coral Bleaching on Coral Habitats and Associated Fishes. 59\u0026ndash;67. https://doi.org/10.7882/fs.2012.012.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePrice, D. M., Robert, K., Callaway, A., lacono, C. L., Hall, R., \u0026amp; Huvenne, V. (2019). Using 3D Photogrammetry From ROV Video to Quantify Cold-Water Coral Reef Structural Complexity and Investigate Its Influence on Biodiversity and Community Assemblage. Coral Reefs, 38(5), 1007\u0026ndash;1021. http://doi.org/10.1007/s00338-019-01827-3.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eProuty, N. G., Fisher, C. R., Demopoulos, A. W., \u0026amp; Druffel, E. R. (2016). Growth rates and ages of deep-sea corals impacted by the Deepwater Horizon oil spill. Deep Sea Research Part II: Topical Studies in Oceanography, 129, 196\u0026ndash;212. https://doi.org/10.1016/j.dsr2.2014.10.021.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRakka, M., Maier, S., Van Oevelen, D., Godinho, A., Bilan, M., Orejas, C., \u0026amp; Carreiro-Silva, M. (2021). Contrasting metabolic strategies of two co-occurring deep-sea octocorals. Scientific Reports, 11(1), 10633. https://doi.org/10.1038/s41598-021-90134-5.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRakka, M., Orejas, C., Sampaio, I., Monteiro, J., Parra, H., \u0026amp; Carreiro-Silva, M. (2017). Reproductive biology of the black coral Antipathella wollastoni (Cnidaria: Antipatharia) in the Azores (NE Atlantic). Deep Sea Research Part II: Topical Studies in Oceanography, 145, 131\u0026ndash;141. https://doi.org/10.1016/j.dsr2.2016.05.011.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRandall, C. J., Negri, A. P., Quigley, K. M., Foster, T., Ricardo, G. F., Webster, N. S., Bay, L. K., Harrison, P. L., Babcock, R. C., \u0026amp; Heyward, A. J. (2020). Sexual production of corals for reef restoration in the Anthropocene. Marine Ecology Progress Series, 635, 203\u0026ndash;232. https://www.int-res.com/abstracts/meps/v635/p203-232.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRibes, M., Coma, R., \u0026amp; Gili, J.-M. (1999). Heterogeneous feeding in benthic suspension feeders: the natural diet and grazing rate of the temperate gorgonian Paramuricea clavata (Cnidaria: Octocorallia) over a year cycle. Marine Ecology Progress Series, 183, 125\u0026ndash;137. https://doi.org/10.3354/meps183125.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRibes, M., Coma, R., Rossi, S., \u0026amp; Micheli, M. (2007). Cycle of gonadal development in Eunicella singularis (Cnidaria: Octocorallia): trends in sexual reproduction in gorgonians. Invertebrate Biology, 126(4), 307\u0026ndash;317. https://doi.org/10.1111/j.1744-7410.2007.00101.x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRobinson, J. P. W., Wilson, S. K., Jennings, S., \u0026amp; Graham, N. A. J. (2019). Thermal Stress Induces Persistently Altered Coral Reef Fish Assemblages. Global change biology, 25(8), 2739\u0026ndash;2750. https://doi.org/10.1111/gcb.14704.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRoik, A. K., R\u0026ouml;thig, T., Roder, C., Muller, P. J., \u0026amp; Voolstra, C. R. (2015). Captive Rearing of the Deep-Sea Coral \u003cem\u003eEguchipsammia fistula\u003c/em\u003e from the Red Sea Demonstrates Remarkable Physiological Plasticity. Peerj, 3(e734). https://doi.org/10.7717/peerj.734.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRos, Z. D., Dell\u0026rsquo;Anno, A., Morato, T., Sweetman, A. K., Carreiro-Silva, M., Smith, C. J., Papadopoulou, N., Corinaldesi, C., Bianchelli, S., Gambi, C., Cimino, R., Snelgrove, P. V. R., Dover, C. L. V., \u0026amp; Danovaro, R. (2019). The Deep Sea: The New Frontier for Ecological Restoration. Marine Policy, 108. https://doi.org/10.1016/j.marpol.2019.103642.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRossi, S., Bramanti, L., Gori, A., \u0026amp; Orejas, C. (2017). Marine animal forests: Springer Nature.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSakai, Y., Yamamoto, H. H., \u0026amp; Maruyama, S. (2024). Long-Term Aquarium Records Delineate the Synchronized Spawning Strategy Of \u003cem\u003eAcropora\u003c/em\u003e Corals. Royal Society Open Science, 11(5). https://doi.org/10.1098/rsos.240183.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSartoretto, S., \u0026amp; Francour, P. (2012). Bathymetric distribution and growth rates of Eunicella verrucosa (Cnidaria: Gorgoniidae) populations along the Marseilles coast (France). Scientia Marina, 76(2), 349\u0026ndash;355. https://doi.org/10.3989/scimar.03262.16B.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSciascia, R., Guizien, K., \u0026amp; Magaldi, M. G. (2022). Larval dispersal simulations and connectivity predictions for Mediterranean gorgonian species: sensitivity to flow representation and biological traits. ICES Journal of Marine Science, 79(7), 2043\u0026ndash;2054. https://doi.org/10.1093/icesjms/fsac135.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSheehan, E., Rees, A., Bridger, D., Williams, T., \u0026amp; Hall-Spencer, J. (2017). Strandings of NE Atlantic gorgonians. Biological conservation, 209, 482\u0026ndash;487. https://doi.org/10.1016/J.BIOCON.2017.03.020.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSheehan, E. V., Stevens, T. F., Gall, S. C., Cousens, S. L., \u0026amp; Attrill, M. J. (2013). Recovery of a temperate reef assemblage in a marine protected area following the exclusion of towed demersal fishing. PLoS One, 8(12), e83883. https://doi.org/10.1371/journal.pone.0083883.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSmith, C. J., Verdura, J., Papadopoulou, N., Fraschetti, S., Cebrian, E., Fabbrizzi, E., Monserrat, M., Drake, M., Bianchelli, S., Danovaro, R., Malak, D. A., Ballesteros, E., Benjumea Tesouro, T., Boissery, P., D\u0026rsquo;Ambrosio, P., Galobart, C., Javel, F., Laurent, D., Orfanidis, S., \u0026amp; Mangialajo, L. (2023). A Decision-Support Framework for the Restoration of Cystoseira Sensu Lato Forests. Frontiers in Marine Science, 10. https://doi.org/10.3389/fmars.2023.1159262.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSneed, J. M., Sharp, K. H., Ritchie, K. B., \u0026amp; Paul, V. J. (2014). The chemical cue tetrabromopyrrole from a biofilm bacterium induces settlement of multiple Caribbean corals. Proc Biol Sci, 281(1786). https://doi.org/10.1098/rspb.2013.3086.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSorek, M., \u0026amp; Levy, O. (2014). Coral spawning behavior and timing. In Annual, lunar, and tidal clocks: Patterns and mechanisms of nature's enigmatic rhythms (pp. 81\u0026ndash;97). Tokyo: Springer.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSuggett, D. J., \u0026amp; van Oppen, M. J. H. (2022). Horizon scan of rapidly advancing coral restoration approaches for 21st century reef management. Emerging Topics in Life Sciences, 6(1), 125\u0026ndash;136. https://doi.org/10.1042/ETLS20210240.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSun, Z., Hamel, J.-F., Edinger, E., \u0026amp; Mercier, A. (2010). Reproductive biology of the deep-sea octocoral Drifa glomerata in the Northwest Atlantic. Marine Biology, 157, 863\u0026ndash;873. https://doi.org/10.1007/S00227-009-1369-9.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSun, Z., Hamel, J., \u0026amp; Mercier, A. (2009). Planulation of deep-sea octocorals in the NW Atlantic. Coral Reefs, 28(3), 781. https://doi.org/10.1007/s00338-009-0505-9.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSun, Z., Hamel, J. F., \u0026amp; Mercier, A. (2011). Planulation, larval biology, and early growth of the deep-sea soft corals Gersemia fruticosa and Duva florida (Octocorallia: Alcyonacea). Invertebrate Biology, 130(2), 91\u0026ndash;99. https://doi.org/10.1111/j.1744-7410.2011.00229.x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSuzuki, G., Okada, W., Yasutake, Y., Yamamoto, H., Tanita, I., Yamashita, H., Hayashibara, T., Komatsu, T., Kanyama, T., \u0026amp; Inoue, M. (2020). Enhancing coral larval supply and seedling production using a special bundle collection system \u0026ldquo;coral larval cradle\u0026rdquo; for large-scale coral restoration. Restoration Ecology, 28(5), 1172\u0026ndash;1182. https://doi.org/10.1111/rec.13178.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTebben, J., Motti, C. A., Siboni, N., Tapiolas, D. M., Negri, A. P., Schupp, P. J., Kitamura, M., Hatta, M., Steinberg, P. D., \u0026amp; Harder, T. (2015). Chemical mediation of coral larval settlement by crustose coralline algae. Scientific Reports, 5(1), 10803. https://doi.org/10.1038/srep10803.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTheodor, J. (1967). Contribution a l'\u0026eacute;tude des Gorgones (VII): Ecologie et comportement de la planula. Vie et Milieu, 291\u0026ndash;302. https://hal.sorbonne-universite.fr/hal-02951553.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eToh, T. C., Ng, C. S. L., Peh, J. W. K., Toh, K. B., \u0026amp; Chou, L. M. (2014). Augmenting the post-transplantation growth and survivorship of juvenile scleractinian corals via nutritional enhancement. PLoS One, 9(6), e98529. https://doi.org/10.1371/journal.pone.0098529.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTonra, K. J., Wells, C. D., \u0026amp; Lasker, H. R. (2021). Spawning, embryogenesis, settlement, and post-settlement development of the gorgonian Plexaura homomalla. Invertebrate Biology, 140(2), e12319. https://doi.org/10.1111/ivb.12319.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTorres, G., Melzer, R., Spitzner, F., Šargač, Z., Harzsch, S., \u0026amp; Gimenez, L. (2021). Methods to study organogenesis in decapod crustacean larvae. I. larval rearing, preparation, and fixation. Helgoland Marine Research, 75(1), 1\u0026ndash;21. https://doi.org/10.1186/s10152-021-00548-x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eVan Oppen, M. J., Gates, R. D., Blackall, L. L., Cantin, N., Chakravarti, L. J., Chan, W. Y., Cormick, C., Crean, A., Damjanovic, K., \u0026amp; Epstein, H. (2017). Shifting paradigms in restoration of the world's coral reefs. Global change biology, 23(9), 3437\u0026ndash;3448. https://doi.org/10.1111/gcb.13647.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eVan Oppen, M. J., Oliver, J. K., Putnam, H. M., \u0026amp; Gates, R. D. (2015). Building coral reef resilience through assisted evolution. Proceedings of the National Academy of Sciences, 112(8), 2307\u0026ndash;2313. https://doi.org/10.1073/pnas.1422301112.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eVan Woesik, R., Lacharmoise, F., \u0026amp; K\u0026ouml;ksal, S. (2006). Annual cycles of solar insolation predict spawning times of Caribbean corals. Ecology Letters, 9(4), 390\u0026ndash;398. https://doi.org/10.1111/j.1461-0248.2006.00886.x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eVieira, F. A., Greg\u0026oacute;rio, S. F., Ferraresso, S., Thorne, M. A., Costa, R., Milan, M., Bargelloni, L., Clark, M. S., Canario, A. V., \u0026amp; Power, D. M. (2011). Skin healing and scale regeneration in fed and unfed sea bream, Sparus auratus. BMC Genomics, 12, 1\u0026ndash;19. https://doi.org/10.1186/1471-2164-12-490.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eViladrich, N., Bramanti, L., Tsounis, G., Coppari, M., Dominguez-Carri\u0026oacute;, C., Pruski, A., \u0026amp; Rossi, S. (2021). Estimations of free fatty acid (FFA) as a reliable proxy for larval performance in Mediterranean octocoral species. Mediterranean Marine Science. https://doi.org.10.12681/mms.27151.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eViladrich, N., Linares, C., \u0026amp; Padilla-Gami\u0026ntilde;o, J. L. (2022). Lethal and Sublethal Effects of Thermal Stress on Octocorals Early Life‐history Stages. Global change biology, 28(23), 7049\u0026ndash;7062.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003evon Koch, G. (1887). Die Gorgoniden des Golfes von Neapel und der angrenzenden Meeresabschnitte: Erster Theil einer Monographie der Anthozoa Alcyonaria. In Fauna und Flora des Golfes von Neapel und der angrenzenden Meeres-Abschnitte / hrsg. von der Zoologischen Station zu Neapel (Vol. 9). Berlin: R. Friedlander \u0026amp; sohn.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWahab, M. A., Ferguson, S., Snekkevik, V. K., McCutchan, G., Jeong, S. Y., Severati, A., Randall, C. J., Negri, A. P., \u0026amp; D\u0026iacute;az-Pulido, G. (2023). Hierarchical Settlement Behaviours of Coral Larvae to Common Coralline Algae. Scientific Reports, 13(1). https://doi.org/10.1038/s41598-023-32676-4.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWaller, R. G. (2005). Deep-water Scleractinia (Cnidaria: Anthozoa): current knowledge of reproductive processes. In A. Freiwald \u0026amp; J. M. Roberts (Eds.), Cold-water corals and ecosystems (Vol. Erlangen Earth Conference Series, pp. 691\u0026ndash;700). Berlin, Heidelberg: Springer.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWaller, R. G., Goode, S., Tracey, D., Johnstone, J., \u0026amp; Mercier, A. (2023). A review of current knowledge on reproductive and larval processes of deep-sea corals. Marine Biology, 170(5), 58. https://doi.org/10.1007/s00227-023-04182-8.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWatling, L., France, S. C., Pante, E., \u0026amp; Simpson, A. (2011). Biology of deep-water octocorals. Advances in marine biology, 60, 41\u0026ndash;122. https://doi.org/10.1016/B978-0-12-385529-9.00002-0.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWatling, L., \u0026amp; Norse, E. A. (1998). Disturbance of the seabed by mobile fishing gear: a comparison to forest clearcutting. Conservation biology, 12(6), 1180\u0026ndash;1197. https://doi.org/10.1046/j.1523-1739.1998.0120061180.x.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWeinberg, S., \u0026amp; Weinberg, F. (1979). The life cycle of a gorgonian: Eunicella singularis (Esper, 1794). Bijdragen tot de Dierkunde, 48(2), 127\u0026ndash;140. https://doi.org/10.1163/26660644-04802003.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eWilliams, S. L., Ambo-Rappe, R., Sur, C., Abbott, J. M., \u0026amp; Limbong, S. R. (2017). Species Richness Accelerates Marine Ecosystem Restoration in the Coral Triangle. Proceedings of the National Academy of Sciences, 114(45), 11986\u0026ndash;11991. https://doi.org/10.1073/pnas.1707962114.\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eZelli, E., Qu\u0026eacute;r\u0026eacute;, G., Lago, N., Di Franco, G., Costantini, F., Rossi, S., \u0026amp; Bramanti, L. (2020). Settlement dynamics and recruitment responses of Mediterranean gorgonians larvae to different crustose coralline algae species. Journal of Experimental Marine Biology and Ecology, 530, 151427. https://doi.org/10.1016/j.jembe.2020.151427.\u003c/span\u003e\u003c/li\u003e\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"coral-reefs","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"core","sideBox":"Learn more about [Coral Reefs](http://link.springer.com/journal/338)","snPcode":"338","submissionUrl":"https://submission.nature.com/new-submission/338/3","title":"Coral Reefs","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"Springer Hybrid","inReviewEnabled":true,"inReviewRevisionsEnabled":false},"keywords":"broadcast spawning, pelagic larval duration, settlement competency, octocorallia","lastPublishedDoi":"10.21203/rs.3.rs-5741857/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-5741857/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eThe widespread decline of coral-dominated ecosystems due to human disturbances has highlighted the urgent need for active habitat restoration. Coral restoration using sexually produced individuals instead of clonal fragments is essential to reduce impacts on donor populations and promote genetic diversity, which is vital for adaptability to environmental changes. However, for most coral species, particularly those in temperate and deep-water (\u0026gt;\u0026thinsp;50 m), critical knowledge of reproduction and larval ecology for ex situ sexual propagation is lacking. To address this gap, in this study, we provide the first report of spawning of the octocoral \u003cem\u003eEunicella verrucosa\u003c/em\u003e in the North-East Atlantic and describe details on larval development and settlement. The annual reproductive timing in South-West Portugal was determined from samples collected as fisheries bycatch from a single population source and monitored for comparison across distinct durations and conditions. The species exhibited split-spawning over about one month (mid September \u0026ndash; mid October), with 3 major events approximately every 2 weeks. Spawning patterns suggest lunar periodicity but shifted between colonies kept in distinct conditions. Oocytes were positively buoyant and developed into swimming larvae after 3 days. Settlement trials using substrates like natural rock, CCA, and gorgonian skeleton, showed larvae behaviour testing the substrates about two weeks post-spawning, and settlement activity continuing over three months. Fully developed recruits were observed after one month, with sclerite production starting before tentacle development. New settlement continued for up to three months, indicating a prolonged competency period. This study provides crucial data for coral restoration efforts using \u003cem\u003eex situ\u003c/em\u003e sexual propagation of this vulnerable species.\u003c/p\u003e","manuscriptTitle":"Reproductive phenology and sexual propagation of the pink sea fan Eunicella verrucosa Pallas, 1766 for coral restoration","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-01-28 16:44:56","doi":"10.21203/rs.3.rs-5741857/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2025-04-09T23:41:13+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-02-27T15:01:54+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-02-08T19:56:13+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-02-08T17:09:41+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"212569656740460954634754577978373694758","date":"2025-01-22T17:28:27+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"97895057882137650784077319373214999115","date":"2025-01-19T09:09:20+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"52697620017665607487187017186359108353","date":"2025-01-16T14:14:49+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-01-09T23:55:46+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-01-07T02:14:15+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-01-03T07:58:25+00:00","index":"","fulltext":""},{"type":"submitted","content":"Coral Reefs","date":"2024-12-31T11:55:14+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"coral-reefs","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"core","sideBox":"Learn more about [Coral Reefs](http://link.springer.com/journal/338)","snPcode":"338","submissionUrl":"https://submission.nature.com/new-submission/338/3","title":"Coral Reefs","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"Springer Hybrid","inReviewEnabled":true,"inReviewRevisionsEnabled":false}}],"origin":"","ownerIdentity":"50a36a27-edb6-4cee-bbbf-612bc6f3414c","owner":[],"postedDate":"January 28th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[],"tags":[],"updatedAt":"2025-07-28T16:04:27+00:00","versionOfRecord":{"articleIdentity":"rs-5741857","link":"https://doi.org/10.1007/s00338-025-02705-x","journal":{"identity":"coral-reefs","isVorOnly":false,"title":"Coral Reefs"},"publishedOn":"2025-07-21 15:58:23","publishedOnDateReadable":"July 21st, 2025"},"versionCreatedAt":"2025-01-28 16:44:56","video":"","vorDoi":"10.1007/s00338-025-02705-x","vorDoiUrl":"https://doi.org/10.1007/s00338-025-02705-x","workflowStages":[]},"version":"v1","identity":"rs-5741857","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-5741857","identity":"rs-5741857","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}