(C) PLOS One This story was originally published by PLOS One and is unaltered. . . . . . . . . . . Coloring coral larvae allows tracking of local dispersal and settlement [1] ['Christopher Doropoulos', 'Csiro Oceans', 'Atmosphere', 'St Lucia', 'George Roff'] Date: 2022-12 Quantifying patterns of dispersal and settlement in marine benthic invertebrates is challenging, largely due the complexity of life history traits, small sizes of larvae (<1 mm), and potential for large-scale dispersal (>100 km) in the marine environment. Here, we develop a novel method that allows for immediate differentiation and visual tracking of large numbers of coral larvae (10 6 to 10 9 ) from dispersal to settlement. Neutral red and Nile blue stains were extremely effective in coloring larvae, with minimal impacts on survival and settlement following optimization of incubation times and stain concentrations. Field validation to wild-captured larvae from the Great Barrier Reef demonstrates the efficacy of staining across diverse taxa. The method provides a simple, rapid (<60 minutes), low-cost (approximately USD$1 per 10 5 larva) tool to color coral larvae that facilitates a wide range of de novo laboratory and field studies of larval behavior and ecology with potential applications for conservation planning and understanding patterns of connectivity. Funding: This work was supported by CSIRO Oceans & Atmosphere awarded to CD ( https://www.csiro.au/en/about/people/business-units/oceans-and-atmosphere ), and the Moving Corals Subprogram awarded to CD ( https://gbrrestoration.org/program/moving-corals/ ) that is part of the Reef Restoration and Adaptation Program (RRAP, https://gbrrestoration.org/ ). RRAP is funded by the partnership between the Australian Government’s Reef Trust and the Great Barrier Reef Foundation. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. To overcome these limitations, we developed and validated a novel method for coloring coral larvae that allows for differentiating among sources and species. Through differential vital staining of coral larvae, our method allows for direct tracking of local dispersal through to settlement of larvae onto coral reef substrates. Additionally, the method directly facilitates visual differentiation among larval cohorts and between coral species by assigning multiple colors, facilitating parentage assignment and allowing for de novo studies of behavior and ecological interactions in both laboratory and field studies. We initially tested four vital stains (neutral red, Nile blue, calcein blue, and alizarin red), followed by a series of laboratory experiments to optimize the visual efficacy of the two most successful stains (neutral red and Nile blue) across a range of stain concentrations and larval incubation times that minimized any adverse impacts on larval survival and settlement. Our method was then validated in the field by coloring wild-caught larvae from natural coral spawn slicks and tracking colored larvae to settlement on reef substrates. The method provides a rapid, simple, nontoxic, and low-cost (approximately USD$1 per 10 5 larva) approach with potential to differentiate cohorts and directly quantify fine-scale dispersal in large numbers (10 6 to 10 8 ) of coral larvae. The severe impacts from climate change to coral reefs worldwide in the 21st century has resulted in major population loss and reduced recovery potential [ 2 ]. Consequently, research focuses have shifted towards large-scale restoration efforts that can mitigate against future disturbances and repopulate damaged reefs [ 10 – 14 ]. Of these interventions, larval reseeding is among the most promising large-scale approaches to regenerate depleted coral populations and reestablish breeding populations [ 10 , 15 – 17 ], largely due to exceptionally high reproductive output of corals (as high as 10 6 to 10 7 eggs per colony; [ 18 ]). This approach captures larvae from wild slicks or from gametes released from gravid colonies in aquaria, cultures the larvae until they are competent, and then those competent larvae are released onto reefs to settle over a period of 24 to 120 hours [ 10 , 15 – 17 , 19 ]. Despite the potential for larval reseeding in large-scale restoration, quantifying the impact of the approach at local to regional scales is complex as reseeded larvae are indistinguishable from natural background larval settlement. Increases in the frequency and intensity of anthropogenic disturbances over the past century has led to widespread fragmentation and habitat loss throughout the world’s oceans [ 1 , 2 ]. In marine ecosystems, the persistence and recovery of populations following disturbance is strongly dependent on the dispersal of propagules from adjacent habitats [ 3 , 4 ]. As such, quantifying the spatial patterns of connectivity among habitats is of key importance for management planning and conservation of marine ecosystems [ 3 , 5 , 6 ]. To date, a range of indirect methods including simulation modelling [ 6 ], elemental fingerprinting [ 4 ], and population genetic approaches [ 7 ] have been used to infer patterns of connectivity across a range of temporal and spatial scales. However, for many important habitat-forming benthic marine invertebrates such as corals, sponges, and bivalves, direct quantification of larval dispersal has remained a major challenge [ 5 ]. Benthic marine invertebrates have complex bipartite life histories [ 4 ] with high reproductive output (>10 6 per individual), small sizes (typically <1 mm), and low survival (<1%) [ 3 , 5 ], requiring the tracking of millions of larvae for direct quantification of dispersal. This challenge is further complicated by complex oceanographic currents [ 6 ] and extended pelagic duration phases of upwards of 100 days [ 8 ], resulting in dispersal ranging from 10 −1 to 10 3 km [ 9 ]. Visual assessment of wild coral larvae under light microscopy confirmed the presence of a diverse multispecies larval assemblage (size range: 150 to 650 μm), ranging from cream to pink to red in coloration. Nile blue staining of these wild-captured larvae was highly effective, with >98% of larvae showing discernible staining effects ( Fig 5 ). Following larval release onto a lagoonal patch reef, settlement of stained larvae was detected on settlement tiles to validate the application. While the common, smaller cream-colored larval species were effectively stained entirely blue in color, the larger less common “red” larval species appear to be stained blue in the outer ectodermal layers only, with the underlying, red-pigmented endodermal layers remaining partially visible ( Fig 5 ). While our lab-based experiments on small numbers (n < 100) of cultured larvae clearly reveal the potential of coloring larvae during dispersal and settlement, the applicability of the staining procedure for tracking broadscale dispersal of large numbers (n > 10,000) of larvae in natural environments required (i) validation of coloration against natural diverse coral larvae collected from wild spawn-slicks and (ii) a field validation of larval settlement within a natural coral reef environment ( Fig 5 ). Developing larvae were collected from wild coral spawn slicks adjacent to Lizard Island (northern Great Barrier Reef) and cultured in larval pools on the reef. After 6 days larval development, approximately 10,000 larvae were subsampled from the culture pool (estimated 1.5 million total) and stained with Nile blue (1,000 mg l −1 for 60 minutes) to contrast against the natural colors of coral larvae. For probability of settlement, the filled proportion of the circle indicates the proportion of surviving larvae. Differences in larval survival between treatments shown against controls (dash line), where no mortality was observed (100% survival). Inset notation for probability of settlement indicates significant differences from control within each species at an α of 0.05 ( ns = no significant difference, * = p < 0.05, ** p < = 0.01, *** = p <0.001). Images supplied by authors. Data underlying this Figure can be found at https://doi.org/10.25919/4rry-xg84 . Under the refined staining procedures, the visual effect of larval staining was optimized across the diverse range of taxa ( Fig 3 ). Survival was consistently high (>80%) with no difference among species and treatments ( Fig 4 ), whereas larval settlement varied among species and treatments (χ 2 = 46.2, df = 6, p < 0.0001; Fig D in S1 Text ). For A. anthocercis, larval settlement was high (>75%) across all treatments and did not differ from controls with the exception of a single treatment ( Fig 4 ). Larval settlement of P. sinensis in the neutral red stain was significantly higher in one treatment (10 mg l −1 for 20 minutes), yet significantly lower in the other treatment (100 mg l −1 for 10 minutes), indicating that higher stain concentrations may reduce larval settlement. Larval settlement of P. sinensis in the Nile blue stain was significantly higher in both treatments than in the control ( Fig 4 ). Both stained or unstained (control) larvae of C. aspera and D. favus failed to settle for the duration of the experiment, suggesting the larvae were not competent and/or unresponsive to the crustose coralline algae cue. Following the initial success of neutral red and Nile blue stains, we conducted a second experiment to refine the staining procedure to include a wider range of corals: Acropora anthocercis, Coelastrea aspera, Dipsastraea favus, and Platygyra sinensis, collected from the central Great Barrier Reef. These taxa are functionally distinct [ 21 ] (tabular growth form: A. anthocercis, massive growth forms: C. aspera, D. favus, P. sinensis) and phylogenetically distant [ 20 ] (family: Acroporidae and family: Merulinidae). Based on the outcomes of the first experiment, we reduced the incubation times to 5 to 30 minutes and concentrations to 1 to 100 mg l −1 for larvae stained with neutral red, and 60 to 120 minutes and 1 to 1,000 mg l −1 for larvae stained with Nile blue. In contrast to A. spathulata, larvae of P. daedalea were considerably more robust, with >75% survival ( Fig 2 ) even at longer incubation times (31 hours versus 24 hours; Fig 2 ). Similar to A. spathulata, neutral red and Nile blue resulted in strong staining at intermediate and high concentrations (10 and 100 mg l −1 ; Fig 2 ), while Alizarin red and calcein blue failed to color larvae. In contrast to A. spathulata, larval settlement of P. daedalea was considerably higher, with >70% settlement rates across all stains and concentrations ( Fig 2 ). Trials of mixed red and blue P. daedalea larvae placed with preconditioned settlement tiles showed that larval sources can be readily distinguished by utilizing their coloration (Fig C in S1 Text ). Larval settlement for A. spathulata differed among treatments (χ 2 = 227.49, df = 18, p < 0.001) and was consistently low ( Fig 2 ), with most treatments resulting in <20% settlement and more than half exhibiting less than 10% settlement (Figs 2 and A in S1 Text ). No significant differences were observed in the probability of settlement between control and stained larvae with the exception of neutral red, where settlement was significantly lower than the controls due to lower initial survival rates (Fig B in S1 Text ). For probability of settlement, the filled proportion of the circle indicates the proportion of surviving larvae, and colors indicate the strength of the larval stain at each stage (larval stage and settled larvae; inset key indicates none, light, medium, or strong staining). Letters in bars indicate pairwise differences in probability of larval survival for A. spathulata. When a p value exceeds α = 0.05, then two means have at least one letter in common. Images supplied by authors. Data underlying this Figure can be found at https://doi.org/10.25919/4rry-xg84 . Larval staining of A. spathulata and P. daedalea was highly effective using neutral red and Nile blue, with larvae and newly settled corals easily distinguishable compared to their natural counterparts ( Fig 1 ). Survival of A. spathulata larvae was reduced at longer incubation times and higher concentrations of dyes (χ 2 = 238.75, df = 18, p < 0.001). The visual effect of the neutral red was most obvious at higher concentrations (10 and 100 mg l −1 ) yet led to significantly reduced larval survival even at short incubation periods (Figs 2 and A in S1 Text ). At the lowest concentration (1 mg l −1 ), neutral red achieved light staining after 6 hours, and medium levels of staining after 12 hours of incubation with >60% larval survival. Nile blue achieved the most consistent staining effect of all stains, with light stains observed at the lowest concentration (1 mg l −1 ) after a single hour of incubation ( Fig 2 ). Survival was high in Nile blue across all concentrations ( Fig 2 ) and did not differ from controls (unstained larvae) after 12 hours (Fig B in S1 Text ). Alizarin red and calcein blue failed to have any measurable visual effects on larvae regardless of concentration and incubation time (Fig A in S1 Text ). Larval survival did not differ from controls at the 12-hour time point (Fig B in S1 Text ), and survival decreased with increasing incubation time in treatments ( Fig 2 ). To establish the potential for staining of coral larvae, an initial factorial experiment was conducted using four stains (neutral red, Nile blue, Alizarin red, and calcein blue). By varying stain concentrations (1, 10, 100 mg l −1 ) and incubation times (1, 6, 12, 24 hours) while quantifying larval survival and settlement, we aimed to test the effects of visual staining while reducing the potential for immediate and latent toxic effects on coral larvae. As corals exhibit high patterns of diversity, our initial experiments included two common yet phylogenetically [ 20 ] and functionally [ 21 ] distinct species of coral: Acropora spathulata (family: Acroporidae, corymbose growth form) and Platygyra daedalea (family: Merulinidae, massive growth form). Both species were collected at Heron Island (southern Great Barrier Reef) and larval staining was conducted 5 days after spawning once the larvae had developed their sensory and motility systems. Discussion We outline an optimized methodology that allows for differentiating larval cohorts and direct tracking of dispersal and settlement of broadcast spawning corals. Importantly, the method is validated in both controlled laboratory experiments and in a natural field environment across a range of functionally [21] distinct and phylogenetic distant lineages [20] of corals. To our knowledge, this study represents the first direct tracking of coral larvae from pelagic stage to benthic settlement on a coral reef, facilitating a range of de novo studies from elucidating small-scale patterns of larval settlement at the scale of millimeters to tracking dispersal at the scale of meters to kilometers. To be effective in differentiating among larval cohorts, the proposed method should be (i) direct and easily detectable and (ii) low in toxicity. Coloring coral larvae with vital stains allows for rapid and simple visual differentiation among larval cohorts, with Nile blue and neutral red stained larvae clearly visible from natural larvae with the naked eye (Figs 1, 3, 5, A, and C in S1 Text) despite the small size of coral larvae (300 to 900 μm; [22]). The peak emission spectra of neutral red (610 to 630 nm; [23]) and Nile blue (650 to 670 nm; [24]) are distinct from the spectral signatures of green fluorescent proteins in Acropora millepora larvae (510 to 520 nm; [25]), highlighting their potential use as fluorochromes in cellular imaging or cytometry applications. For example, combing our method with large-particle flow cytometry of live larvae [26] would enable sorting of larval cohorts in experiments to assign parentage, or allow rapid separation to recover experimentally colored larvae from mixed wild cultures in large-scale field experiments. Following refinement of concentration and exposure times, our results indicate that for two vital stains (neutral red and Nile blue), coloring coral larvae has minimal direct (reduced larval survival) or indirect (latent effects on settlement and metamorphosis) impacts, with no clear differences observed from controls. At higher concentrations and incubation times, toxicity differed among coral taxa: larvae from the family Acroporidae exhibited greater sensitivity to neutral red than Merulinidae. The toxicity of neutral red to Acroporidae larvae is counter to that reported from other benthic marine invertebrate larvae (e.g., oysters) that exhibit greater sensitivity to Nile blue [27] and are unaffected by neutral red [28]. While calcein blue and alizarin red have been successfully used in staining adult benthic invertebrates [27–29], as well as mineral deposits found within brooded coral planulae [30], the absence of visual staining in spawning coral larvae due to a lack of calcium binding potential limits their application to post-settlement life history stages (Fig 1) following the onset of early skeletal formation [31]. To be effective in directly tracking larval dispersal, the proposed method must be (i) procedurally simple and rapid; (ii) easily detectable in field settings; (iii) easily scalable to large numbers (106 to 109) of larvae; (iv) nontoxic in the marine environment; (v) widely available; and (v) cost effective. From a procedural perspective, the protocol is simple and rapid (<60 minutes incubation), allowing for application in remote field locations where laboratory facilities are unavailable. In terms of detection, larvae can be detected following release by sampling with plankton tows during the pelagic stage [32] and directly on reef substrates post-settlement using settlement tiles [33] or relatively inexpensive (