Abstract
Corals at very early life stages face high mortality bottlenecks, limiting natural stock-recruitment and the success of restoration and aquaculture strategies reliant on sexual reproduction. Nutrient availability and quality, particularly lipids, are critical for larval survival, dispersal, and settlement, yet the specific lipids influencing early-stage fitness remain unclear. Since measuring lipid depletion in non-surviving larvae is impractical, we developed a targeted supplementation approach to identify nutrients that enhance larval fitness. Supplementing Acropora spathulata larvae with nanoparticle-encapsulated triacylglycerols and sterols significantly improved swimming distance and speed, indicating enhanced dispersal potential. Supplementation with sterols, which are essential for cell structure and metamorphosis, also increased settlement rates. Furthermore, juveniles developed from larvae fed with sterols and omega-3-rich fish oils demonstrated superior performance under controlled and thermal stress conditions during the first six months, while those supplemented with Calanus oil also enhanced survival under elevated temperatures. Lipid profiling revealed key sphingolipids, glycerophospholipids, and fatty acids likely responsible for the improved fitness of fed larvae. Our approach highlights the critical role of lipid composition and availability during coral early life stages and demonstrates the potential of targeted nutritional supplementation to address nutrient deficiencies, enhance coral resilience, and support restoration and aquaculture efforts.
Introduction
Many corals sexually reproduce by releasing gametes into the water column, which mix in the surface waters and develop into small planktonic larvae. To become adult colonies, larvae must continue to develop, locate suitable substrate for settlement, and successfully metamorphose into juvenile corals; these steps all require adequate nutrition to support high energy-consuming processes and complex tissue remodelling1,2. Nutrient availability and quality can therefore significantly influence coral larval survival, growth, dispersal, and settlement success1,3,4,5. Coral larvae are primarily composed of lipids, particularly energy-rich triacylglycerols, phospholipids, and wax esters, which help fuel development and other vital cellular processes5. For example, fatty acids are important for coral thermal plasticity6, while sterols regulate metabolic homoeostasis, developmental processes, and immune responses, including those involved in the stability of the endosymbiosis with dinoflagellates in the family Symbiodiniaceae7,8. However, the specific nutrient profiles required to maximise survival and growth across coral life stages remain poorly understood9,10,11.
Intensifying anthropogenic stressors have driven global declines in reef-building corals, leading to reduced live coral cover and disruptions in key processes essential for reef recovery, such as reproduction rates12. Nutritional imbalance is a key driver of coral mortality, and reduces immune function and fecundity in surviving colonies following environmental stress13. Nutrients, particularly lipids14, in adult corals may be redirected away from gametogenesis and towards metabolic processes essential to withstand environmental stress and thus impair gamete health and hence reproductive success15,16. Environmental factors such as nutrient availability3 (e.g., nitrogen and phosphorus1), salinity17, pH18, and temperature19 can each affect larvae behaviour, settlement and survival20. Consequently, suboptimal environmental conditions inevitably exacerbate coral mortality post-settlement, creating significant imbalances in the stock-recruitment relationship21. Identifying which nutrients and how nutrient availability help regulate reproductive success is therefore key to resolving the metabolic demands governing early life processes in corals and hence advance sexual reproduction-based restoration efforts in the face of escalating environmental pressures11.
Inability to determine if larvae that fail to settle or survive are deficient in specific nutrients hinders our understanding of the impact of nutrition on coral reproductive success. An alternative approach is to draw from nutritional science, where supplements are often used to identify beneficial nutrient interactions, which themselves offer effective intervention tools. For example, carbohydrate supplementation increases the reproductive success of the endangered hihi bird (Notiomnystis cincta)22. Coral larvae rely heavily on maternally acquired nutrients and have limited energy reserves4,5,23,24, yet are capable of obtaining nutrients from the surrounding environment by taking in dissolved nutrients and organic matter1,3,4,25,26, free amino acids27, and small particulates via ciliary currents and mucociliary transport through an oral pore24,28,29. Feeding therefore provides an opportunity to supplement energy reserves and obtain new nutrients not maternally inherited30, which is particularly important in environments where larvae are deployed for restoration purposes and where the nutrient demands and availability might differ from the source reef. Heterotrophic feeding of adult corals is known to enhance survival and growth31,32, improve tissue condition33, and boost photosynthetic rates33,34. For coral larvae raised in aquaria, the provision of exogenous nutrients such as homogenised artemia, can improve settlement rates and immediate post-settlement survival30. However, whether exogenous nutrients also enhance other key physiological aspects – such as larval swimming capacity, long-term survival and environmental resilience – remains unexplored. Coral larvae typically undergo significant lipid-depletion during early development5,26,35,36,37, making lipids logical targets to investigate the effect of exogenous nutrient provision. Given the size of the larvae oral pore (~0.1–2 mm) and primary dissolved organic matter feeding capabilities, nano-delivery of food could therefore conceivably improve uptake rates and even passively diffuse through tissues. Nanoparticles can be utilised as carriers for targeted nutrient delivery to corals as they offer advantages such as efficiency through high surface area-to-volume ratios, which, if caught by mucus strings, could be easily consumed38. A targeted feeding approach utilising nanotechnology to create specific lipid-based food compositions could illuminate the metabolic processes underpinning coral development while optimising delivery methods and reducing the costs associated with supplying exogenous nutrients to support early coral life stages. Here, we show that providing nanoparticle-encapsulated lipid supplements enhances coral larval swimming capacity, settlement, survival and resilience to thermal stress. Our findings underscore the potential of targeted nutritional supplementation to improve coral offspring resilience and offer a pathway to enhance aquaculture outputs and restoration efforts via nutritional supplementation during vulnerable early life stages.
Results and Discussion
Coral larvae actively feed on lipid nanoparticles
While phosphatidylcholine lipid nanoparticles (LNPs) offer a relatively low-cost and biodegradable option for targeted nutrient delivery, stability and uptake are key initial considerations. Lipid nanoparticles containing commercial mixes of Triglycerides (TG), fish oil, Calanus oil, phospholipids, or plant sterols (Table 1, Supplementary Figs. 3 and 4, Supplemental Data 13 and 14), as well as phosphatidylcholine LNPs without additional lipids (henceforth “empty LNPs”) were created and the zeta potential (the electrochemical equilibrium at the particle-liquid interface that determines particle stability) and size measured. These were selected as exogenous lipid supplements as they represent a broad range of lipid subclasses, and include lipids previously identified as major components of corals. Phospholipids include Phosphatidylethanolamine (PE) and phosphatidylcholine, which support membrane structure and fluidity39, Phophotidylinositol (PI) is a precursor to signalling molecules involved in symbiosis40,41, Phosphatidylserine (PS) contributes to membrane dynamics and apoptosis signalling, while phosphatidylglycerine (PG) may aid in Symbiodiniaceae stress response42. Wax esters are important for coral larvae buoyancy5 and triacylglycerols are important for energy metabolism5. Fish oils contain high amounts of omega-3 fatty acids, known to be translocated in the symbiosis and important for coral health43, and sterols are hypothesised to function in signalling processes vital to coral health and resilience35. Most LNPs had a zeta potential between ±15-30 mV (except for sterol LNPs), indicating moderate stability. Empty LNPs exhibited a significantly lower zeta potential compared to triacylglycerols (TG), fish oil, and Calanus oil-filled LNPs (Tukey HSD; p < 0.001, df = 5; Supplementary Data 1), and although not significant, empty LNPs also had a lower zeta potential than phospholipid LNPs (Tukey HSD; p = 0.271, df = 5; Supplementary Data 1). As expected, empty LNPs were significantly smaller than all other LNPs (237.27 ± 5.49 nm, Tukey HSD; p < 0.001, df = 5; Supplementary Data 1). Sterol LNPs had a lower zeta potential than all other LNPs (Tukey HSD; p < 0.001, df = 5; Supplementary Data 1) and were below the ±15 mV threshold (−12.13 ± 0.29 mV), suggesting that these particles were unstable and prone to agglomeration. Consistent with this notion, sterol LNPs exhibited the largest mean size (609.43 ± 45.5 nm), supporting the presence of particle aggregation. Although sterol LNPs were less stable over time due to lower zeta potential and larger size, they remained suspended and visibly intact during the feeding period, allowing effective ingestion by larvae. However, long-term stability of nutrient supplements must therefore be considered if LNPs are to be used as a delivery method. This includes the ability of the particles to maintain their size, structure and dispersion over time, which is critical for the consistent delivery of the contents to the target site (e.g., organism, cell). Instability can lead to aggregation and premature release, reducing efficacy and potentially causing unintended environmental effects.
To determine whether nanoparticles could be actively ingested rather than passively absorbed, we analysed the lipid profiles of larvae exposed (fed) and unexposed (unfed) to phosphatidylcholine LNPs prior to oral pore formation. No significant differences were observed between treatments at the individual lipid level (PERMANOVA, FDR = 0.3, F-value = 1.52, R² = 0.28; Supplementary Data 2, Supplementary Fig. 2) or at the lipid subclass profile level (PERMANOVA, FDR = 0.5, F-value = 1.31, R² = 0.25; Supplementary Data 2, Supplementary Fig. 2). Further T-test analysis supported these findings, as no individual lipid or subclass differed between the two treatments (T-test, p > 0.05; Supplementary Data 2). Consequently, larvae do not appear to passively absorb lipid nanoparticles from the surrounding water at this life stage.
Lipid supplementation alters coral larvae lipid profiles
Once an oral pore had been established (~48 h post fertilisation) and confirmed by microscopy, Acropora spathulata larvae were either unfed, or exposed to one of the six lipid feeding treatments, using the lipid nanoparticles containing commercial mixes of TG, fish oil, Calanus oil, phospholipids, or plant sterols (Table 1, Supplementary Figs. 3 and 4, Supplemental Data 13 and 14), as well as empty LNPs as an additional control to ensure that the nanoparticle delivery vehicle itself does not induce unintended effects, either positive or negative. Lipid mixes were added to yield a final concentration of 0.3 mg mL−1, based on previous data44. After two days of food exposure, lipid uptake was confirmed via Liquid Chromatography-Mass Spectrometry. A total of 567 individual lipids from 71 lipid subclasses were compared between treatments (Supplementary Data 2&3). The overall lipid profile of unfed larvae was significantly distinct to all fed larvae (PERMANOVA, F-value = 31.00, R2 = 0.8, p < 0.001; Fig. 1A, Supplementary Data 3), with differentiation driven by several long chain fatty acids (FA 40:5, FA 40:7, Fig. 1A) and diglycerides (e.g., DG 32:2, DG32:4, DG 30:4, DG42:6 and DG 42:11, Fig. 1A). Sterol and TG were further distinct from all other feeding treatments, but not each other (pairwise PERMANOVA; Supplementary Data 4). These two treatments were distinguished by several triglycerides (TG 30:0, TG 39:0, TG 44:0, Fig. 1A), and sphingomyelins (SM 38:5; O3 and SM 40:3;O3, Fig. 1A). All other fed treatments were similarly clustered, except empty LNPs and phospholipids (pairwise PERMANOVA, FDR = 0.045; Supplementary Data 4). Of the 567 individual lipids, 537 were significantly different between at least one feeding treatment and unfed larvae (Fisher’s LSD; Supplementary Data 5). Given the absence of lipid profile changes prior to oral pore formation, the observed differences in lipid profiles following pore development likely reflect active feeding, coupled with selective uptake and metabolic processing of supplemented lipids.
PCA illustrates differences in individual (A) and lipid subclass (B) lipid compositions among larvae subjected to difference feeding treatments. Full lipid composition data are provided in Supplementary Data 3-8.
At the subclass level, unfed larvae were significantly distinct to all fed larvae (PERMANOVA, F-value = 12.585, R2 = 0.78, p < 0.001; Fig. 1B, Supplementary Data 6). Sterol and TG were further distinct from larvae fed with empty LNPs, phospholipids, and fish oil, but not larvae fed with Calanus oil (pairwise PERMANOVA; Supplementary Data 7). Of the 71 lipid subclasses, 62 were significantly different between at least one feeding treatment and unfed larvae (Fisher’s LSD; Fig. 2, Supplementary Data 8). Sterols are known to support key cellular processes in corals, including metabolic balance, immune function, development and organismal cell structure35, roles that may help explain the physiological changes observed in the sterol-fed larvae. Sterol levels are tightly regulated in eukaryotic cells by balancing uptake, catabolism, storage, and efflux45. Corals, unable to synthesise sterols, must therefore acquire them through translocation from Symbiodiniaceae or heterotrophic feeding46. As demonstrated in our current experiments, aposymbiotic coral larvae can acquire sterols from external sources and substantially increase sterol stores, such as the subclass acyl hexosylceramides (AHexCer), which were more abundant in larvae fed with sterols than in other treatments (Fisher’s LSD; Fig. 2, Supplementary Data 8). Sterol enrichment through feeding could therefore potentially enhance sterol-dependent cellular processes even before the establishment of endosymbiosis.
Lipid subclass text colour refers to lipid class. Raw abundance data and statistical significance data (Fisher’s LSD) are provided in Supplementary Data 6 and Supplementary Data 8. Data were log-transformed and mean-centred.
Phospholipids-fed larvae had the highest relative abundance of most lipid classes. Phospholipids are important components of membrane structure47,48, and represent a major class of membrane lipids in corals39. Given their ubiquitous structural role, the higher abundance may be due to accumulation in the larval membranes, while other lipid classes are perhaps catabolised, such as TGs, or incorporated into more complex lipid structures, such as free fatty acids. Glycerolipids, such as triglycerides (TGs, OxTG and EtherTG) and the galactolipids monogalactosyldiacylglycerol (MGDG) and digalactosyldiacylglycerol (DGDG), were significantly more abundant in all fed treatments compared to unfed larvae (Fisher’s LSD; Fig. 2, Supplementary Data 8). These lipids serve as metabolic energy reserves and contribute to membrane structure and lipid-mediated signalling, especially in photosynthetic systems39. Meanwhile, most lipid subclasses were significantly lower in unfed larvae than in all fed treatments, suggesting that larvae relying solely on maternally inherited stores are potentially nutritionally deficient, which likely contributes to the high mortality naturally observed at this stage21.
Notably, the only lipid subclass more abundant in unfed larvae was ether-linked phosphatidylinositols (EtherPI) (Fisher’s LSD; Fig. 2, Supplementary Data 8). Ether-linked phospholipids are common in marine invertebrates, including the sea anemone Exaiptasia diaphana39, and may contribute to membrane stability due to their tightly packed alkyl chains49. Higher EtherPI abundance in unfed larvae could represent a compensatory response to maintain membrane integrity in nutrient-limited conditions or a physiological adaptation in larvae with limited resources.
Enhanced swimming capacity
A critical aspect of coral larvae dispersal and long term survival is their capacity to find suitable substrate for settlement50. Habitat selection affects both early post-recruitment survival and the likelihood of surviving to the adult (reproductive) stages51,52. Once the coral larva has metamorphosed, it will usually remain attached to the substrate for its entire life, and will only be able to alter its habitat conditions through asexual growth and propagation. While a high proportion of endogenous hydrophobic wax esters facilitates passive dispersal via surface currents53, coral larvae can swim using cilia propulsion54, using chemical cues to find an optimal location51. Swimming has a high-energetic cost54; therefore, the nutritional status of the coral larvae can determine their swimming capacity, affecting both their speed and distance covered. Swimming capacity, therefore, ultimately impacts reef recruitment and the potential for healthy, reproductively active neighbouring reefs to naturally restore nearby degraded reefs50,52.
Larvae fed with TGs and sterols swam on average 2.8 times faster and covered distances 2.9 times larger than unfed larvae or those fed with fish oil or empty LNPs (Table 2; ANOVA Fishers LSD, FDR < 0.001 and FDR < 0.005, respectively, Supplementary Data 9). In addition, larvae fed with phospholipids, TGs and sterols had a maximum speed 2.8 times faster than unfed larvae and larvae fed with empty-LNPs (Table 2; ANOVA Fishers LSD, FDR < 0.001, Supplementary Data 9). All other supplements had no significant effect on the swimming capacity of these larvae (Table 2).
Triglycerides are critical energy reserves for corals, which they typically obtain from their algal symbionts5, but since A. spathulata larvae are aposymbiotic, they rely entirely on external sources to replenish energy reserves. Given the high energy demands of swimming, our study shows that bolstering baseline energy reserves in coral larvae enhances both swimming speed and distance. This is supported by the lipid profile data showing that TG subclasses are significantly more abundant in TG- and sterol-fed larvae (Figs. 1 & 2). One possible explanation for the enhanced TG in sterol-fed larvae is that sterol supplementation may support membrane fluidity or integrity35, indirectly supporting metabolic processes involved in TG synthesis. Additionally, sterols may influence signalling pathways or gene expression related to lipid metabolism, such as upregulation of enzymes responsible for TG biosynthesis45. Further investigation into lipid regulatory pathways in coral larvae is required to clarify these mechanisms.
Sterols, primarily derived from phytoplankton consumption or symbiont translocation55, are also crucial for key physiological processes, including cell signalling and structure. Coral larvae use cilia for movement and chemical sensing54, likely through key signalling pathways, such as the Hedgehog (Hh) cell differentiation pathway, where sterols play vital structural and signalling roles56. Cilia possess a unique lipid composition57, and are enriched with a diverse group of oxidised sterols, or oxysterols, that bind to and activate central signalling molecules57,58. Based on our data, we hypothesise that the unique lipid composition in sterol-fed larvae may enhance cilia function and, by extension, swimming capacity.
Phospholipids also play essential roles in maintaining membrane structure47,48, which could support ciliary propulsion in larvae and potentially enhance the maximum swimming speed as observed here. Additionally, phosphoinositides (PIs), phosphorylated derivatives of membrane phosphatidylinositols, are pivotal in coral symbiotic signalling40,59, but also play a significant role in cilia biology60. Together, these findings suggest that phospholipid composition may be a key determinant of larval mobility and symbiotic functioning, highlighting their dual importance in both dispersal capacity and long-term coral health.
Feeding coral larvae increases settlement rate
The proportion of larvae that successfully settled varied across treatments (ANOVA, F = 4.81, FDR < 0.01; Fig. 3, Supplementary Data 10). Larvae fed with TG, fish oils, phospholipids, or empty LNPs exhibited settlement levels similar to those of unfed larvae (Fisher’s LSD, FDR > 0.05; Fig. 3, Supplementary Data 10). In contrast, larvae fed with sterols exhibited the highest settlement, with nearly half (47%) settling by day 5 (Fisher’s LSD, FDR < 0.01; Fig. 3, Supplementary Data 10) and significantly higher than for unfed larvae and those fed on Calanus oil, empty LNPs, or TG.
Fisher’s LSD post hoc group is shown (a-c, FDR > 0.05; Supplementary Data 10).
Sterols and their derivatives, such as oxysterols and steroid hormones, act as signalling lipids that regulate metabolic homoeostasis, development, organismal structure, and immune responses through G protein-coupled receptors (GPCR) and nuclear receptor activation7. A drug screen targeting GPCR signalling pathways has previously confirmed involvement of these pathways in larval settlement behaviour and metamorphosis53. Our data suggests that the increased sterol availability from supplements may have promoted GPCR signalling and enhanced settlement and metamorphosis. Meanwhile, larvae fed with wax ester-rich Calanus oil showed the lowest settlement, even compared to unfed larvae, with only 6% successfully settling (Fisher’s LSD, FDR < 0.01; Fig. 3, Supplementary Data 10). Wax esters are known to influence coral larval dispersal by affecting buoyancy61,62, which helps larvae remain entrained in surface currents63. As aposymbiotic larvae metabolise wax esters during development5, they become more negatively or neutrally buoyant, enabling them to interact with the substrate and potentially find suitable settlement sites. However, if larvae retain excessive buoyancy, they may struggle to settle, potentially explaining the reduced settlement rate observed in those fed with Calanus oil.
Enhanced survival and thermal resilience of juvenile corals
Once coral larvae settle and metamorphose into benthic juvenile polyps, their nutrient requirements change23,64. Most coral mass spawning events occur in spring, with coral juveniles often experiencing the highest temperatures of the year a few months after settlement. Nutrition is a key factor in coral resilience to environmental challenges, including thermal stress13, therefore early-life nutrition could significantly influence coral juveniles growth and survival. Juveniles generally exhibit higher metabolic activity and growth rates than larvae64, shifting from reliance on stored energy to greater utilisation of symbiont-derived nutrients and heterotrophic feeding16,64. Under thermal stress, however, symbiont-derived nutrition may be disrupted13, leading juvenile corals to rely more heavily on stored energy reserves or heterotrophic feeding65. These sources could become even more crucial if climate change impacts to seawater chemistry affect the availability of key nutrients in the global ocean66,67. Thus, robust nutrition during the larval stage may provide coral juveniles with the resources needed to endure the challenging early months on the reef.
Following larval settlement, all recruits were exposed to homogenous mix of homologous strains of Symbiodiniaceae to establish a symbiosis. Half of the juveniles were maintained at a control temperature (28 °C), while the other half were exposed to 31˚C, a temperature exceeding those recorded by the Davies Reef Weather Station (DRWS) that resulted in mass bleaching on the Great Barrier Reef during the 2020 summer heatwave (equivalent to +3 ˚C of the local reef maximum monthly mean (MMM))68. Survival was monitored at regular intervals over six months.
Juveniles derived from all larval feeding treatments showed significantly higher survival rates over time compared to those from unfed larvae (Generalised Linear Mixed Model (GLMM): survival ~ (food and temperature treatment) + tank, p < 0.001; Supplementary Data 11). Under control temperatures, juveniles initially fed with sterols and fish oils exhibited 2.3- and 2-fold higher final survival, respectively, compared to unfed larvae (Fisher’s Exact, FDR < 0.05; Fig. 4, Supplementary Data 10 & 11).
The total survival of juvenile corals under control (solid-coloured lines) and increased temperatures (dashed-coloured lines) following feeding treatments as larvae (colours) is shown. The red dotted line indicates the start of temperature ramping. Asterisk highlights Fisher’s exact significance (* p < 0.05) of juvenile survival between fed and unfed larvae, within each temperature treatment (control and heat stress). Data are provided in Supplementary Data 11 and Supplementary Data 12.
When comparing temperatures within each feeding treatment, survival rates were significantly higher under control conditions, except for Calanus oil, which showed higher survival under elevated temperatures (GLMM p < 0.001; Supplementary Data 11). However, endpoint survival was statistically indistinguishable between control and heat-exposed juveniles in any of the fed groups (Fisher’s Exact Test, FDR = 1.00; Supplementary Data 11 & 12), suggesting that larval feeding may support thermal resilience in coral recruits. Furthermore, while unfed larvae also showed no significant difference in final survival between control and elevated temperatures (Fisher’s Exact, FDR = 1.00; Fig. 4, Supplementary Data 11 & 12), they had the lowest overall survival across all groups, particularly under heat exposure. Lastly, the rate of mortality for unfed recruits differed significantly between temperature conditions (GLMM p < 0.001; Supplementary Data 11). Together, these findings highlight a heightened vulnerability when nutritional support is lacking during the larval stage, and suggest that robust early-life nutrition can help equip coral juveniles with the resources needed to survive the critical early months on the reef. It is also possible that the low irradiance used to reduce competitive stress from algal overgrowth during recruit rearing69 may have also dampened the combined effects of heat and light stress typically seen under high-irradiance conditions70. Future studies could test higher light levels or include physiological indicators (e.g., photochemical efficiency, ROS production) to better understand light–temperature interactions during early coral development in fed larvae.
Under increased temperatures, larvae fed with sterols, fish oils, and Calanus oil showed 1.8- to 2.1-fold higher final survival relative to the heat-exposed unfed group (Fisher’s Exact, FDR < 0.05; Fig. 4, Supplementary Data 11 & 12). Recruits of fish-oil fed larvae had higher survival at each time point compared to unfed larvae, under both increased and control temperatures (Fisher’s Exact, FDR < 0.05; Fig. 4, Supplementary Data 11 & 12). Similarly, recruits of sterol-fed larvae had 1.5-times higher final survival than those receiving empty LNPs under control temperatures (1.5-fold increase, Fisher’s Exact, p < 0.05; Fig. 4, Supplementary Data 11 & 12). Analysis of survival-over-time analysis revealed no significant difference in overall survival trajectories between sterol- and fish oil-fed larvae under control (GLMM, FDR = 0.439, Supplementary Data 12) or increased temperatures (FDR = 0.117, Supplementary Data 12), suggesting these supplements may confer comparable metabolic benefits. Supporting this interpretation, the lipid subclass compositions of the commercial fish oil and sterol mixtures indicated similar biochemical profiles (Supplementary Fig. 3, Supplementary data 13). Meanwhile, larvae fed with phospholipids, TGs and empty LNPs showed no significant increase in final survival compared to unfed larvae under control or increased temperatures (Fisher’s Exact, FDR = 1.00; Fig. 4, Supplementary Data 11 & 12).
Sphingolipid subclasses sphingomyelin (SM) was more abundant in larvae fed with sterol supplements than in other treatments (Fig. 2, Supplementary Data 6). This pattern may reflect the selective uptake or retention of sphingolipid subclasses present in the sterol commercial mix (Supplementary Fig. 3, Supplementary Data 13). Sphingolipids form crucial components of cell membranes across all eukaryotes and play key roles in cell signalling71. The sphingolipid metabolic pathway produces various lipids that regulate diverse physiological processes, such as apoptosis, cell survival, inflammation, and autophagy, through activation of specific GPCRs72,73,74. A critical aspect of sphingolipid metabolism is the balance between pro-survival (sphingosine-1-phosphate, S1P) and pro-apoptotic (sphingosine and ceramide) sphingolipids75,76. This “sphingolipid rheostat” controls cell fate through the interconversion of these metabolites, guided by metabolic flux and the activity of key enzymes, such as sphingosine kinase (SPHK) and S1P phosphatase (SGPP)76. Evidence suggests that sphingolipid signalling regulates the onset of the cnidarian-Symbiodiniaceae symbiosis by supporting host cell survival75,77. Therefore, the elevated concentrations of these subclasses in sterol-fed larvae, if sustained, may contribute to enhanced coral juvenile fitness and survival. Furthermore, sterol transporters, like Niemann-Pick Type C2 (NPC2), are highly upregulated in symbiotic cnidarians and are localised around symbiosomes, essential for establishing the Symbiodiniaceae symbiosis55,78,79. These juvenile corals were colonised with homologous symbionts one week after settlement, raising the possibility that sterol supplementation supported host cell survival and improved symbiotic function by selectively increasing key sterol-derived metabolites. As a functional symbiosis is essential for optimal juvenile nutrition65,80, this may have ultimately contributed to improved coral growth and survival.
Larvae fed with sterols, fish oils and Calanus oils exhibited a relatively higher abundance of several energy-rich glycerolipid subclasses, including triacylglycerols, following feeding (Fisher’s LSD; Fig. 1 & 2, Supplementary Data 8). Mitochondrial bioenergetics involves energy production through fatty acid oxidation, beginning with the hydrolysis of TG into fatty acids (FAs) for degradation and energy production81, supporting central metabolic processes under both control (in sterol and fish oil fed larvae) and increased temperatures (in sterol, fish oil and Calanus oil fed larvae). Meanwhile the glycerol can be re-esterified into TG or incorporated into phospholipids, a process that requires metabolic energy, and may support coral host membrane structure and if shared with the resident Symbiodiniaceae, could help support the thylakoid membranes of the thermally stressed endosymbionts39. The role of phospholipids in thermal resilience is likely influenced by their specific fatty acid composition, such as unsaturation levels, rather than total phospholipid content alone. This notion is further supported by the higher relative abundance of several TGs and glycerophospholipids in the profiles of Fish oil and Calanus oil-fed larvae (Fisher’s LSD; Figs. 1 & 2, Supplementary Data 5 & 8). Although TG-fed larvae also showed elevated levels of triacylglycerols, these did not translate into significantly improved long-term survival, suggesting that energy reserves alone may be insufficient to confer resilience without other bioactive lipids or supporting compounds. Both Fish oil and Calanus oil-fed larvae were significantly more enriched with omega-3 polyunsaturated fatty acids (PUFAs) than unfed larvae, including eicosanoids (FA 22:2, CAR 22:6, Fig. 1A), and eicosapentaenoic acid (FA 20:5, Fisher’s LSD, FDR < 0.01; Supplementary Data 5), which are frequently detected as upregulated in Symbiodiniaceae under heat stress6, and thus considered important for coral thermal plasticity. These findings reinforce the critical role of PUFAs during coral early development, especially given that corals cannot synthesise these essential fatty acids de novo and must obtain them through diet or symbionts. In addition, 22-carbon oxylipin products (e.g., FA22:5; O4) were also significantly more abundant in Calanus oil and fish oil-fed larvae (Fisher’s LSD, FDR < 0.01; Supplementary Data 5). Oxylipins constitute an expansive group of signalling lipids created by the oxidation of PUFAs, and initial evidence suggests they play a crucial role in the stability of the symbiosis82. Symbiodiniaceae are considered an important source of antioxidant lipids that can enhance thermal resilience43; however, supplementation of these lipids during the aposymbiotic larval stage could enhance stores and support the metabolic stability of the symbiosis and health of the coral, even if the symbiosis is disrupted80. Although such supplements are likely metabolised quickly during the larval stage, our data suggests the settled juvenile corals continue to experience clear metabolic benefits even six months after supplementation. Collectively, our study underscores the importance of optimal nutrition in reducing mortality during these vulnerable early life stages.
Conclusion
Our results show that coral larval lipid profiles play a crucial role in their health, development, and survival, underscoring the importance of both the quality and quantity of nutrients available for coral reproductive success. High-quality lipid nutrition, including essential fatty acids and specific lipid classes such as sterols, supports energy reserves, swimming capacity, metamorphosis, and stress resilience. Lipid reserves fuel early development stages, helping larvae successfully establish new coral colonies and subsequently withstand environmental pressures juveniles. Furthermore, variations in lipid composition, particularly in sphingolipids and glycerolipids, can influence larval settlement rates, growth, and symbiotic relationships, ultimately shaping coral population dynamics and resilience. While the commercial sterol mix used in this study likely contained additional lipid classes, such as triacylglycerols and phospholipids, which may contribute to some of the observed improvements through synergistic or additive effects, sterol-fed larvae consistently exhibited unique physiological benefits and lipidomic signatures. Future work should incorporate purified or factorial lipid formulations to resolve the individual versus interactive roles of specific lipid classes in supporting coral early-life fitness. Nonetheless, our results demonstrate the importance of sterols in the fitness and success of coral early life stages (Fig. 5). While fish oil and TG supplementation provided metabolic benefits, only sterol-fed larvae showed significant improvements across multiple fitness metrics including swimming, settlement, and long-term survival, suggesting that sterols support developmental processes not fully compensated for by energy reserves or PUFAs alone. Several studies have highlighted the potential for sterols to play important roles in coral symbiosis and signalling35,40,79, but this presents the first evidence of critical roles in development and early-life fitness.
Supplementation with sterols and triacylglycerols enhanced larval swimming and settlement, while sterol and fish oil improved recruit survival under both control and, along with Calanus oil, heat stress conditions. Together, these effects highlight how lipid enrichment can enhance early-life performance and support long-term reef fecundity.
Optimising lipid intake is clearly vital for enhancing coral reproductive outcomes, particularly in the face of changing ocean conditions. While reef restoration efforts that focus on enhancing larval supply offer a promising pathway for the recovery of degraded reefs21, they are limited by low post-settlement survival rates. Additionally, these efforts have yet to fully integrate strategies that improve dispersal dynamics or enhance the stress tolerance of surviving juveniles. Here, we show that despite forming the least stable nanoparticles, supplementation with sterols during the vulnerable larval stage supports biologically important and ecologically significant fitness parameters. Because corals cannot synthesise sterols independently, they must obtain them through translocation from their microalgal symbionts Symbiodiniaceae or heterotrophic feeding35, thus supplementing with sterol LNPs provides an alternative source for these aposymbiotic larvae that support vital processes for survival during early life stages. As our findings suggest, coral larvae actively consume lipid nanoparticles rather than absorbing them passively, it is possible that the larger average size of sterol LNPs might support the feeding potential and uptake of these LNPs. The next logical step is to optimise the supplementation regime, including the concentration, timing, frequency, composition, and delivery method. To further enhance the benefits of sterol supplementation, future studies could explore alternative encapsulation strategies, such as solid lipid nanoparticles, or direct dosing approaches to improve uptake and bioavailability83. However, it is important to avoid excess loading of exogenous lipids, which could become a metabolic burden and potentially lead to negative effects such as lipid peroxidation84. While such effects are unlikely during early developmental stages – when energy demands are high and both autotrophic and heterotrophic contributions are minimal – they should still be considered, especially under the altered energy requirements and increased lipid peroxidation conditions imposed by thermal stress.
Methods
Coral spawning and larval rearing
Experiments were conducted in December 2022, at the National Sea Simulator (SeaSim) located at the Australian Institute of Marine Science, Townsville, Queensland, Australia. Six colonies of Acropora spathulata were collected from Davies Reef (-18.831, 147.634; Permit no. G12/35236.1), and transported to SeaSim two days prior to the December full moon. Coral colonies were acclimated in outdoor 1000 L flow-through holding tanks and maintained at ambient temperature (27 °C), 35.8 PSU, and a natural light cycle (under ambient solar and lunar photoperiods) until spawning occurred (5 nights after full moon). Egg and sperm bundle collection and subsequent gamete separation were conducted in accordance with previous methods85. Eggs from a single colony were suspended in 200 mL of 27 °C 1 µM-filtered seawater in 4 L larval rearing containers and mixed-genotype sperm inoculated at 106 cells mL−1. Once cell division was observed (~1 hour), 2.8 L of 27 °C 1 µM filtered seawater was added, and 2 L of 1 µM filtered seawater syphoned from the bottom of the containers to dilute residual sperm. A further 2 L of 1 µM filtered seawater was added gently via syphon, and embryogenesis allowed to continue overnight inside a temperature-controlled room set at 27 °C. After 12 hours, ~2.8 L of the water was syphoned and replaced with new 1 µM filtered seawater. When embryogenesis was complete, and early stages of larvae had developed (~24 h post-fertilisation), larvae were filtered through a 150 µM mesh filter, rinsed gently with 1 µM filtered seawater, and resuspended in 3 L of 1 µM filtered seawater. Larvae were counted and diluted to 1 larva per 1.5 mL seawater and held in 500 L larval rearing tanks (homogenous mix of parents across 6 tanks) with air and water flow (total ~333,333 larvae per tank). After 36 hours, 90 L (approximately 60,000 larvae) were collected from one tank. Larvae were gradually rinsed on a 150 µM filter, resuspended to 1 larva per mL, and divided randomly across 28 × 2 L glass jars with aeration, with each jar containing 1.8 L of seawater (approximately 1,800 larvae per jar). Jars were maintained inside a large flow-through water bath set to 26.5 °C to help regulate the water temperature inside each, under at a maximum daily irradiance of 60 µmol photons m−2 s−1 over a 12:12 photoperiod with a sinusoidal ramping profile. Larvae were rinsed on a 150 µM filter after 12 hours, and resuspended to 1.8 L. Larvae were maintained until an oral pore was observed (~60 hours post-fertilisation).
Lipid cocktail nanoparticle preparation
Lipid nanoparticles (small phosphatidylcholine vesicles with an empty space for the entrapment of other lipids for delivery) were prepared using a Lecithin-in-Water emulsion method as described in Yanasarn and collaborators86. For each lipid treatment mix (Table 1), 800 mg of L-α-Lecithin (phosphatidylcholine) was dissolved in 200 mL of 32 °C MilliQ water. The mixture was heated to 55 °C with continuous stirring until a milky consistency was formed. While maintaining agitation, the respective lipid supplement (Table 1) was added to the dissolved lecithin; after which, a 2% v/v solution of Tween-20 was added drop-by-drop. The heater plate was turned off to allow the mixture to slowly cool with continuous stirring. Fifty mL of each lipid nanoparticle mixture (11.1 µg mL−1 final concentration based on preliminary evidence from Boulotte et al.44) was added to the relevant experimental jar once it had reached room temperature.
Larval feeding
Prior to feeding, larvae were rinsed once more on a 150 µM filter and resuspended in 1.8 L of 1 µM filtered seawater. Larvae were allowed to feed for 8 hours, after which larvae were rinsed on a 150 µM filter with 1 µM filtered seawater, jars thoroughly cleaned, and larvae resuspended in 1.8 L of 1 µM filtered seawater with aeration. Approximately 200 larvae were collected from each jar, washed three times with 1 µM filtered seawater using a 106 µm mesh filter and transferred into 2 mL cryovials. Samples were flash-frozen in liquid nitrogen and stored at −80 °C (T1; N = 28 samples, totalling approximately 5600 larvae). This initial collection occurred after the first feeding and served as fallback samples. The remaining larvae were left for 18 hours, after which the lipid feeding was repeated. A second set of approximately 200 larvae were collected from each jar, washed three times using with 1 µM filtered seawater in a 106 µm mesh filter and transferred into a 2 mL cryovial. Samples were flash-frozen in liquid nitrogen and stored at −80 °C until processing for lipidomics analysis (T2; N = 28 samples, approximately 5600 larvae). These T2 samples were used for lipidomics analysis to compare lipid profiles across treatments. An additional 20 larvae were collected and rinsed from each treatment for larvae swimming assessments.
Passive absorbance of lipid nanoparticles
To determine the potential for the passive absorbance (across cell membranes) of the nanoparticles compared to active feeding, larvae were exposed to empty lipid nanoparticles before they developed an oral pore. Larvae were divided into 6 jars (in addition to those used for subsequent feeding) were rinsed and resuspended in 1.8 L of 1 µM filtered seawater as above. N = 3 jars of larvae were exposed to 111 µg mL−1 empty lipid nanoparticles (Treatment 7, Phosphatidylcholine only, Table 1) for 8 hours, rinsed three times and transferred to a 2 mL cryovial. In addition, N = 3 jars were left untreated for 8 hours, rinsed three times and transferred to a 2 mL cryovial. Samples were flash-frozen in liquid nitrogen and stored at −80 °C until processing for lipidomics analysis
Coral larvae swimming assessment
Swimming patterns of n = 20 larvae from each treatment (n = 5 larvae per jar) were recorded for 2 minutes under 4 x magnification following the second lipid incubation and water change. Videos were split into hyperstacks before automated tracking of larvae in Trackmatev7.12.2, a FIJI v4.0.0 plugin87. A Laplacian of Gaussian (LoG) detection algorithm with quality thresholding between 0.02 −0.5 provided initial spot identification. Once the algorithm ran, additional quality and/or contrast filters were applied, if necessary, to reduce the number of false spots. Of the 20 larvae sampled, only those that were identified by the detection software and remained post filtering were used for subsequent analysis. A Simple Linear Assignment Problem (LAP) tracker was applied to the tiff stacks and identified the individual larvae tracks which were manually validated to ensure track continuity. The diameter of the container holding the larvae was used for measurement calibrations. Mean displacement and speed for each larva were extracted from the produced tracks.
Coral larvae settlement and recruit survival monitoring
Larval settlement took place inside 28 separate 4 L plastic containers (one per jar with a FSW flow-through system) containing 55 aragonite settlement plugs per treatment in a plug-holding crate. Each plug was previously biologically conditioned for 3 months in an outdoor tank set up with non-filtered flow-through seawater to allow the development of biofilm and crustose coralline algae (CCA), providing natural settlement cues for coral larvae88,89,90. Approximately 8.5 h after the second lipid mix exposure, 20 mL of homogenously suspended larvae were sampled from each jar. Larval density (larvae/mL) was quantified using a 10 mL Bogorov chamber under 4 × magnification. These estimates were used to calculate settlement success by comparing them to the number of larvae observed to have settled. To initiate settlement, water flow was halted, and 300 mL from each jar was transferred into the corresponding settlement containers. The settlement containers (with plugs) were placed inside a large flow-through water bath set to 26.5 °C to help regulate the water temperature inside each tank. One week after settlement, flow through water was turned off, the water level dropped to 1 L and 10 mL of a homogenous solution of Symbiodiniaceae cultures SCF082 (UTS-D, Durusdinium trenchii) and SCF055_01.10 (Hetero-M, Cladocopium goreauii) at 1.0 × 10⁶ cells/mL was added, for a final concentration of 10000 cells/mL in the tanks. After ~6 hours, the flow through was turned back on. Although symbiont densities were not quantified in this study due to the destructive nature of the required sampling methods, all recruits were uniformly exposed to Symbiodiniaceae 7 days post-settlement (7 days prior to the first survival measurement). Successful uptake was visually confirmed by the characteristic darkening of recruit tissues, which typically occurs within 48 following exposure91. Two weeks post-settlement, high-resolution macro-photographs of settled coral spat on plugs were taken with a Nikon D810 to assess the initial settlement and survival rate (Fig. 3). Subsequently, all settlement plugs containing settled spats were labelled, randomly mixed, and divided into 28 × 50 L tanks for longer-term survival monitoring. Recruits were held in flow-through aquaria that received three turnovers of filtered seawater (nominal 0.1 µm) per day soured directly from the adjacent coastal waters near Cape Ferguson, Queensland.
Coral larvae lipid extractions and analysis
All steps were performed at 4 °C and using LC-MS grade glassware. Lipid extraction was based on the methyl-tert-butyl ether (MTBE) extraction and phase separation protocol for high-throughput lipidomics92. Approximately 100 coral larvae were transferred to filter paper to remove residual seawater and homogenised in 100% methanol (at −20 °C) and 5 µL of EquiSPLASH LIPIDOMIX mass spec standard (Avanti Polar Lipid 330707), using a Chattaway spatula. Samples were incubated on ice for 10 minutes, and vortexed three times during this time. 1000 µL of MTBE was added, samples vortexed for 30 s and shaken for 1-hour at 1000 rpm at 4 °C on a rotisserie platform. To prepare for phase separation, 250 µL of ultra-pure grade water (4 °C) was added, samples incubated on ice for 10-minutes and centrifuged at 1000 × g for 10 minutes at 4 °C. 900 µL of the organic top layer was collected, dried under a nitrogen steam, and stored at −80 °C until analysis. Sample blanks (N = 2) were undertaken alongside each run.
Prior to analysis, samples were resuspended in 100 µL 2:1 isopropanol methanol (IPA: MeOH) and transferred to autosampler glass vials with a 125 µL glass insert. A pooled sample was created to assist with compound analysis by combining 5 µL of each sample into a single tube and 100 µL of this pooled sample was transferred to an autosampler vial with a 125 µL glass insert. 5 µL injections of each sample were processed via Liquid Chromatography Mass-Spectrometry (Thermo Orbitrap LC-MS) in positive and negative ion mode. Each sample was run as per the methodology described in Roper et al. 93 in positive mode under acidic chromatographic conditions, and in negative mode with neutral chromatographic conditions using an Agilent UPLC system and Waters ACQUITY UPLC CSH C18 Column (130 Å, 1.7 µm, 2.1 mm × 150 mm). The column oven was set to 65 °C for both methods. The positive mode method had a flow rate of 0.4 mL min-1 with mobile phase A consisting of 60:40 ACN : Water + 10 mM Ammonium formate + 0.1% Formic acid and B consisting of 90:10 IPA : ACN + 10 mM Ammonium formate + 0.1% formic acid. For Heated electrospray ionization (H-ESI) and data-dependent acquisition (DDA) parameters, see Supplementary Data 15 and 16. The negative mode method had a flow rate of 0.4 mL min−1 with mobile phase A consisting of 60:40 ACN: Water + 10 mM Ammonium acetate and B consisting of 90:10 IPA : ACN + 10 mM Ammonium acetate. Both methods used the same gradient of the following solvent B: 0.00 min 30%, 2.00 min 30%, 2.50 min 50%, 13.00 min 85%, 13.50 min 99%, 15.00 min 99%, 15.10 min 30%, 18.00 30%. Separated lipids were then ionised into the source of a Thermo Q-Exactive Plus mass spectrometer. Data was acquired via DDA topN, and the scan range was set to 200–1200 m/z, with the difference between the methods listed in Supplementary Data 15 and 16. Every 9 samples a solvent blank and a QC were injected.
Raw lipid spectral data were exported to .raw files and processed in MS-Dial (version 5.2)94 for peak alignment, blank correction, and lipid identification against the LipidMaps LipidFinder (V2) and Structure Database (LMSD). Peak heights for each lipid were exported and normalised to the relative peak height of the internal standards for each mode, followed by normalisation to egg protein content following Matthews et al. 95. Briefly, the remaining extracts were removed, 1 mL 0.2 M NaOH added to the larvae cell debris pellet, and samples vortexed and incubated at 98 °C for 20 mins. Samples were centrifuged at 3000 g for 10 mins. Total protein content of the extracted larvae tissue was then quantified using the Bradford assay (Bradford, 1976). Triplicate 10 μL aliquots of each sample were pipetted into a 96-well plate alongside six serial dilutions of bovine serum albumin (Sigma) of known concentrations: 1, 0.5, 0.25, 0.125, 0.06, 0.031 and 0 mg mL−1. 250 μL of Bradford Reagent was added to each well, resulting in a colorimetric assay of protein concentration within the samples. The 96-well plate was gently agitated on a rotating platform for 10 minutes prior to being analysed on a spectrophotometer at 595 nm (Tecan Spark Microplate Reader). The protein concentration of each sample was calculated from the equation of the protein standard curve. A total of 945 lipids were confirmed. Any lipid with a relative standard deviation (RSD) within a treatment greater than 40% was removed, leaving 567 individual lipids. Data was arranged into all lipids and lipid sub-class (using LipidMaps classifications96) data for further analysis.
Survival analysis
Each tray of recruits was reared in 30 L aquaria receiving flow-through water filtered to 1 µm at a rate of 0.4 L min−1 (providing one turnover per hour). Lighting was provided by custom LED panels (325 W Gen. II multi-chip), at a maximum daily irradiance of 60 µmol photons m−2 s−1 over a 12:12 photoperiod with a sinusoidal ramping profile. This light level was chosen to simulate shaded reef microhabitats and is consistent with recent findings from Ramsby et al. 69, who demonstrated that low-light conditions significantly improved coral spat survival and minimised harmful algal overgrowth. Seawater temperature was maintained at 28 °C in control aquaria, while treatment aquaria were incrementally ramped to 31 °C over one month, at a rate of 0.2 °C per day for 4 consecutive days followed by a 5-day break, with this pattern repeated until the target temperature was reached (Supplementary Fig. 1). Upon reaching the target temperature of 31 °C, coral recruits were then held at this temperature for 6 months (N = 2 tanks/trays per treatment, 53 plugs per tray, total N = 106). High-resolution photographs were taken at pre-determined intervals: 3- (pre ramp), 5-, 9-, 13-, 17-, 21-, and 25-weeks post-settlement to monitor survivorship of coral juveniles over time. Coral recruit survival was assessed by counting the number of plugs containing at least one recruit during each monitoring period. This method was employed to avoid losing survivorship data due to the fusion of some neighbouring coral recruits on the plugs over time.
Statistical analyses
Swimming total distance, mean, and max speed were averaged for each treatment and compared using one-way ANOVA and Tukey Post-hoc in R (v4.4.1) using R Studio (version 2024.09.0 + 375) with the assistance of “broom”97.
Lipid statistical comparisons were conducted in Metaboanalyst (v6.098). Lipid relative abundances were log-transformed and mean-centred, and following confirmation of normality (Shapiro-Wilk test) and homogeneity of variance (Bartlett tests), the data were visualised using Principal Component Analysis (PCA). The significance of treatment grouping separation was analysed using a permutational analysis of variance (PERMANOVA), and the significance of individual lipids and classes was analysed using a one-way ANOVA. All tests were adjusted for multiple comparisons using a Bonferroni false discovery rate.
Recruit survival was visualised using a Kaplan-Meier survival curve using the R packages “survival”99, “survminer”100 and “ggplot2”101. End point survival (number of surviving recruits divided by number of recruits at the first count) was compared using a Fisher’s Exact test with Bonferroni correction (using R packages “dplyr”102 and “rcompanion”103). This analysis evaluates survival proportions at the conclusion of the experiment only, regardless of when deaths occurred. In addition, survival over time was assessed using a generalised linear mixed effect model (R packages “lme4”104, “emmeans”105, and “lmerTest”106) with survival, lipid supplement and temperature as fixed effects, and tank as random effect. The survival-over-time model incorporates both the timing and rate of mortality, answering the questions of whether the pattern or rate of mortality differs over time. The variance for the tank effect was very low (0.0009 + 0.03), suggesting there was no tank effect present.
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
Raw Mass spectral data is available at Metabolomics Workbench (study ID 6054). Other raw data are available at https://doi.org/10.5281/zenodo.15681195107.
Code availability
R scripts used are available at https://doi.org/10.5281/zenodo.15681195107.
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Acknowledgements
This research was carried out with the support of Pure Ocean (Awarded to JLM, DJS) and a University of Technology Sydney Chancellors Fellowship (awarded to JLM).
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J.L.M., D.H., C.H., N.B., Mv.O., J.-B.R. and D.J.S. designed the study; J.L.M., N.B., L.L.M., M.P., D.H., Z.C., and C.R. performed the experiments; J.L.M., H.K., M.P. analysed the data; J.L.M. wrote the article with contributions from M.P., D.H., Z.C., C.H., N.B., J.-B.R. and D.J.S.
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Matthews, J.L., Bartels, N., Kish, H. et al. Sterols are key to coral larvae survival, swimming capacity, and thermal tolerance. Commun Biol 8, 1494 (2025). https://doi.org/10.1038/s42003-025-08965-1
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DOI: https://doi.org/10.1038/s42003-025-08965-1
