Unified methods in collecting, preserving, and archiving coral bleaching and restoration specimens to increase sample utility and interdisciplinary collaboration

Specimen/Sample provenance and metadata

Museums, research aquariums, and private collections have standard protocols for documenting the history, or origin, of individual specimen (Smithsonian Institution, 2006; National Science & Technology Council, Interagency Working Group on Scientific Collections, 2009; National Academies of Sciences, Engineering, & Medicine, 2020). Researchers and practitioners can optimize the use of their data and samples by rigorously cataloguing, and formally documenting as many experimental (e.g., temperature ramp rate, light level, flow), biological (e.g., coral color, morphotype, taxonomy, provenance), and environmental (e.g., depth, nutrient concentrations, reef type) variables as possible (Grottoli et al., 2021) because these data provide needed context for each collection. We refer to these descriptive, contextual data as metadata.

Representative samples can also be properly ‘vouchered’ with a museum for long-term preservation, and such specimen can have important applications for a wide range of future work from these biological collections. First, if such samples include both tissue and the taxonomically informative skeleton, they can provide a taxonomic reference in the event that cryptic species are discovered, or to assign identity of the samples with future changes to taxonomy. Such vouchered samples also provide invaluable reference barcodes for databases that are becoming increasingly important as environmental DNA (eDNA) approaches become commonplace. Likewise, techniques change through time and questions that would have been impossible to address from such samples a couple of decades ago have become common place today with the advent of high-throughput and single-cell sequencing. Finally, even in cases where there is no obvious need to preserve the samples, the value of having historical samples has been showcased repeatedly in the field of epidemiology and emerging zoonotic disease research, where natural history collections have been integrated with host-pathogen research to resolve pathways of transmission (Thompson et al., 2021). The questions that will be answered by historical samples may yet be unknown, but it is only possible to address them if the samples are collected, vouchered and properly maintained.

Sample provenance also includes the documentation of how and where samples and their resulting data and metadata are physically and digitally stored. Growing recognition of the value of historical data and appreciation for FAIR (findability, accessibility, interoperability, and reusability) data standards (Wilkinson et al., 2016) is inspiring the efforts to archive sample data and metadata in ways that facilitate reuse and ensure archived data is available to future researchers (Zerbino et al., 2018; Davis et al., 2019; Percie du Sert et al., 2020). For example, the Genomic Observatories Metadatabase (Deck et al., 2017), stores metadata archives permanently linked to -omics resources stored at the National Center of Biotechnology Information’s (NCBI, Bethesda, MD, USA) and the National Science Foundation’s Data Management Office (BCO-DMO) serve as repositories where samples and associated metadata are linked to the researchers who produced those studies and can be contacted about collaboration or specimen sharing.

Last, in regard to data provenance, many funding agencies have specific data management and dissemination requirements (e.g., BCO-DMO at the National Science Foundation, GenBank at the National Center for Biotechnology Information, Environmental Data Service at the Natural Environment Research Council). However, relevant details concerning these samples and their province legacy data are often overlooked by researchers. For example, a recent sampling of the Sequence Read Archive (SRA) of GenBank found that only ~14% of all archived specimens associated with a sequencing project included both collection year and site as basic metadata that would be required for the reuse of archived genomic data in future studies (Toczydlowski et al., 2021). As the culture of global research and reef conservation and restoration have moved toward more open and collaborative models, there is growing pressure from funding bodies, journals, management agencies, and researchers alike to provide these data in open-access formats (Sibbett, Rieseberg & Narum, 2020), and develop community-wide cyberinfrastructure that facilitates the discovery and reuse of material samples (e.g., iSamples; Davies et al., 2021). Such consolidated efforts stand to benefit the advancement and accessibility of the field of coral bleaching research and restoration science and effort as a whole.

Specimen/Sample collection considerations

There is a myriad of possible techniques for collecting, processing, and archiving most coral specimens (for more details see Supplemental Materials). However, unique differences among coral taxa including individual colonies, their morphotypes, tissue thicknesses, skeletal density, and variation in life history demand special consideration as these variables may affect the biology and chemistry of collected coral samples and could dictate the applicability of many downstream procedures. Additionally, colony and specimen/sample size as well as species-specific variation can affect how corals respond to and recover from stress (Brandt, 2009; Thomas & Palumbi, 2017; Álvarez-Noriega et al., 2018; Levas et al., 2018). The quantity of available sample material can also affect what downstream techniques are possible. Improving the precision of measurements of colony and specimen size is an active area of research (Table S1) with the advent of new technological developments such as 3D laser scanning and photogrammetry (House et al., 2018; Vivian et al., 2019; Zawada, Dornelas & Madin, 2019). Information about the original size of the parent colony or outplant specimen can provide helpful information for interpreting resulting data because size has been shown to be an important bleaching predictor (Álvarez-Noriega et al., 2018).

Collection permits may also restrict the number of samples that can be collected, which can affect the types of analytical methods that are possible downstream and how much excess material may or may not be available for archiving and future research. Further, some agencies restrict the use of specimen for explicitly defined goals within a given project or on a specific permit and thus may be unavailable for alternative end goals. Researchers should be aware of such agency-based limitations and enquire with providers on any downstream use restrictions. Last, developmental stage can have significant impacts on which methods are suitable and practical for any methodological pipeline. For example, the amount of material required for some analyses may be prohibitive when working with coral larvae or gametes, but easily performed on adult tissues. Thus, the types of research questions that can be addressed will vary depending on the life stage of the specimen and dictate the types of downstream analyses and collaborations that are most productive.

Temperature and sample storage considerations

When collecting and preserving coral bleaching and restoration specimens for short and long-term use, documenting a sample’s temperature history is critical (see Box 1 on freezing and cryopreservation). In general, altered temperatures can cause rapid state changes in live specimen physiology, microbiology, and geochemistry. Many subcellular and cellular processes can change within minutes to hours when corals undergo shifts in ambient temperature (Hillyer et al., 2017a), and swift sample processing is important to capture those responses. Once samples are preserved, temperature can further influence the integrity of each sample for some types of analyses. For example, cells lyse if samples are too cold, thus making them unsuitable for imaging of intact cells. Each scientific discipline (Omics, Physiology, and Microscopy and Imaging) has guidelines for optimal preservation temperatures suitable to ensure the integrity for their analytic process (see Fig. 2). The duration of storage for these specimens can also dictate ideal archiving temperature conditions. If samples are intended to be stored for tens of years (e.g., in coral gamete biobanks), cryopreservation and downstream restoration, rapid-freezing in liquid nitrogen, and storing at −80 °C are the safest holding temperatures. If the tissues or cells can tolerate freeze-drying, and the final packaging is vacuum-sealed, then such specimens can be maintained for many years at room temperature.

However, coral bleaching and restoration research is often conducted in locations where adequate freezing agents and materials (e.g., liquid nitrogen, dry shippers, or even ice) may not always be available or reliable. Although not all methods require temperature stabilization, many do (Fig. 2). Therefore, if possible, all researchers should record above and below water (1) the transport holding temperature, (2) any altered temperatures during transport, and (3) the duration of transport. For example, if live or dead specimens were removed from an offshore reef, transported to shore, and placed in new containment, the method and duration of transport as well as the temperature of any onshore activities (e.g., freezer storage, water temperature manipulation) should be documented and included in the sample metadata.

Specimen handling and sterility considerations

There is increased interest in how the coral holobiont microbiome (i.e., Symbiodiniaceae, bacteria, viruses, and other microscopic eukaryotes) responds to, and may be involved in, preventing or exacerbating coral bleaching and/or increasing or reducing restoration success. Many ecological and physiological bleaching studies can be easily paired with Symbiodiniaceae analyses (e.g., cell densities, gene sequencing) through shared samples, but the potential for coral bacterial and/or viral analyses is severely compromised when sterile collection tools (e.g., gloves, bone-cutters) and sterile receptacles (bags or tubes) are not used. The use of aseptic handling techniques during coral collection and processing is a relatively small and inexpensive change in the methodological pipeline that can enable additional downstream microbiome analyses (Fig. 3). For example, a suitable aseptic technique in the field may be as simple as wearing nitrile gloves when handling corals and using sterile receptacles, such as Whirl-Pak sample bags, and minimizing cross-contamination by replacing with clean equipment or sterilizing in between handling different specimens. For microbiome work, additional sampling of the environment (e.g., water and sediments) can provide information about sources of potential contamination. Importantly, while aseptic techniques are ideal for many downstream applications, it is impractical if not impossible to maintain underwater and in some handling situations.

Caveats and considerations for methodologies, accuracy, and usability

In each discipline there may be recommended and, in some cases, well benchmarked standard operating protocols for each individual method discussed below and in the Supplemental. However, many of the methodological pipelines discussed below which are suitable for some aspects of coral-bleaching and restoration research but have not yet been fully evaluated in terms of their accuracy and precision. Therefore, deviations from standard procedures for a given discipline could potentially result in data that are inaccurate, uninterpretable, or unusable. It is important to consider the potential caveats when using any non-standard procedure in field and laboratory work. Yet, as research techniques improve and additional methods and protocols are confirmed as having high precision and accuracy, more of the potential pipelines discussed below may be employed with confidence in any given discipline. For example, using chemically fixed (e.g., in formaldehyde) samples for genomic-based analysis has been considered non-standard in the past, but new work shows that these preserved specimens can be reliably used to gain insight into various aspects of coral biology, retrospectively (Greene et al., 2020).

Freezing or fixation methods dictate most methodological pipelines

Instantaneous freezing or ‘rapid freezing’ in liquid nitrogen upon initial collection followed by ultra-cold storage (e.g., −80 °C) maximizes the number of possible downstream analyses (supports ~64% of 36 methods; Fig. 2 blue and green cells row 7). Analyses that could concurrently or sequentially be conducted after specimen rapid-freezing and cold storage fell primarily within the Omics and Physiology disciplines, while rapid freezing is inappropriate for most tissue Microscopy and Imaging because it alters tissue integrity (see Box 1). Freezing post-collection using −80 °C and more conventional −40 °C or −20 °C freezers were also suitable for several procedures within the Omics- and Physiology-based methods (supports ~83% of 18 Omics- and Physiology analytics), except for some RNA-based analytics, which always require immediate rapid freezing or preservation (e.g., in RNAlater, TRIzol).

Within the Microscopy and Imaging discipline, preserving in paraformaldehyde followed by refrigeration allowed for the greatest number of downstream analytics (70% of all Microscopy and Imaging techniques, row 48), including some Omics methods. However, few if any of the Physiological methods could be conducted on samples stored in these aldehydes.

Methodological considerations are needed to determine the suitability of some collecting, preserving and archiving sample pipelines

A few analyses, including metabolomics, quantification and identification of mycosporine-like amino acids (MAAs), soluble lipid analysis, and histology techniques stood out as highly restrictive in their requirements for initial and secondary storage methods. Such methodological limitations could be due to stringency in storage requirements or, as suggested by the large number of unknowns in Fig. 2, due to insufficient testing of potential alternative methods. Thus, we summed the number of ‘unknown cells’ to determine which methods had the most uncertainty in terms of how samples could be collected, preserved, and archived. Numerous methods had many ‘unknowns’ (Fig. 2, row 60) limiting our ability to find suitable additional pipelines to access outside of their standard procedures. For example, biomass quantification and Raman spectroscopy each respectively had 51% and 83% unknowns for the different possible methodological pipelines we tracked.

Below we discuss considerations specific to each major discipline: Physiology, Omics, and Microscopy and Imaging, given these methodological differences. Furthermore, we add more details about standard operating procedures for each of the major downstream analyses within Fig. 2 and throughout the Supplemental Material. While not an exhaustive list, we aimed to give researchers enough information to consider how to collect, preserve, and archive their specimens for many potential applications. We also recognize that methods are continuously evolving with the advent of new technologies. It is likely that newer, better methods will eventually become available and, thus, future researchers should take steps to confirm that additional procedures have not become available following the publication of this work.

Sample collection

In general, the use of aseptic techniques is strongly encouraged across all coral bleaching and restoration research collections, allowing for the most complete set of downstream analyses, including the characterization of the microbiome (which includes the coral virome) for which aseptic technique is necessary to ensure further analytical accuracy and integrity (Fig. 2). However, Omics methods targeting the coral host and/or Symbiodiniaceae do not require sterile tools and receptacles unless total Symbiodiniaceae community diversity is being examined in high resolution. Additionally, reagents and materials can contaminate samples with off-target cellular materials, foreign nucleic acids, viral particles, and exogenous chemicals. Moreover, compounds or enzymes that degrade, damage, or alter macromolecules and biological compounds (e.g., proteases, R/DNAses) can disrupt or inhibit many molecular processes (e.g., cations and polymerase chain reaction) needed to create Omic datasets or intermediate sample products (e.g., sequencing libraries). Thus, care should be taken to avoid the use of non-sterile materials and/or reagents that are not certified as molecular biology grade.

Preservation for short term storage

Rapid freezing and storage at −80 °C is the optimal collection and storage technique for most Omics work such as genomics, metagenomics, proteomics, and RNA-Seq methods (see below). While RNA-based methods (e.g., transcriptomics) are notoriously sensitive to initial collection and storage conditions and require rapid freezing or immediate storage in salt buffers, DNA is more stable and thus can be collected and preserved in a variety of conditions. Several methods (e.g., DNA- and RNA-based host, symbiont, and microbiome approaches) can also be used when corals are initially preserved in salt buffers, some aldehydes, or DMSO (Gaither et al., 2011; Gray, Pratte & Kellogg, 2013; Hernandez-Agreda, Leggat & Ainsworth, 2018). These buffered specimens can be stored short-term at a variety of temperatures because the compounds stabilize the nucleic acids (Hopwood, 1975; Seutin, White & Boag, 1991; Dawson, Raskoff & Jacobs, 1998; Douglas & Rogers, 1998; McKenzie, 2019). For metabolomics, rapid freezing in liquid nitrogen (which inhibits metabolic processes) and storage at –80 °C is optimal and generally considered best practice, especially if subsequent separation of host and symbiont is desired (Lohr et al., 2019) as storing samples in methanol immediately restricts analyses to holobiont metabolome due to the potential for salt contamination (Hillyer et al., 2017b). Rapid freezing and −80 °C storage is optimal for samples targeting protein analysis or comprehensive proteomics, although samples can also be rapidly processed (via crushing in liquid nitrogen or airbrushing/water-picking) and preserved in a protein extraction buffer containing protease inhibitors for short- and long-term storage (e.g., Barshis et al., 2010; Tisthammer et al., 2021). However, for epigenetics, specific sub-applications have different sample preservation requirements (see Fig. 2). All in all, the approaches available for preserving specimens for Omics work are limited by the requirements of the most stringent aspect of the molecules under study (see Supplemental Material for details).

Processing

The amount of material necessary for each downstream Omics procedure varies significantly and according to the target input/quality requirements of each analysis (e.g., ~1 mg of extracted protein for proteomics). Procedures that use amplification steps (e.g., metabarcoding of the bacterial and archaeal 16S rRNA, Symbiodiniaceae ITS2, or eukaryotic 18S) will require little material (~1 cm²). Genomics, metagenomics, transcriptomics, epigenetics, proteomics, and metabolomics will require more starting material (e.g., >1 cm²) with typically higher quality standards. Single coral fragments are frequently analyzed with multiple different, complementary procedures (e.g., genomics, transcriptomics, and metabarcoding).

Archiving for long term storage

Almost all macromolecules in specimens are more stable long-term when ultra-frozen and many of them can be stored at freezing temperatures indefinitely if in the appropriate fixative or buffer. However, some methods such as Assay for Transposase-Accessible Chromatin using sequencing (ATAC-Seq)-based epigenetics require unusual storage conditions, such as cryopreservation in liquid nitrogen.

Caveats

Delays or alterations in sampling, processing, and preserving any specimen for these Omics techniques may alter the accuracy and precision of the resulting analysis. However, many alternative preservation and storage methods (e.g., formaldehyde) for Omics work remain untested or are incompletely benchmarked, thus it is unknown how these methods may affect downstream analyses. Future Omics research could scrutinize how these under-studied pipelines may alter a specimen’s true biological and chemical composition in the short-term and when stored for long periods of time.

Sample collection

In contrast to several of the microbial analyses described above (see Omics section), sample sterility and aseptic conditions are not an absolute requirement for downstream physiological assays. As a general guide, a coral ramet height >1 cm or tissue area >1 cm² are minimal requirements. However, larger fragment sizes (i.e., several cm²) are generally desirable to minimize edge effects associated with tissue heterogeneity and avoiding tissue damaged in the sampling process. Larger fragments also allow for multiple laboratory analyses and facilitate better cross-comparisons among corals and studies because stress conditions may decrease the amount of a given variable (e.g., lipids), making it harder to measure in very small specimens. In addition, a small sample size (1 cm²) may provide enough material for one analysis (e.g., endosymbiont chlorophyll a), whereas considerably more material is required for other analyses (e.g., coral tissue-based isotopes that can require four times this amount). When combining these requirements with the added benefit of long-term archiving for later potential use, investigators will often want to double their number of ramets when possible. Large fragments or ramets also have the benefit of providing a more representative sample and minimizing fragment edge effects associated with sampling, as well as any positional effects within the coral fragment itself (e.g., top vs side of a branch or coral-mound). If the sample is not immediately processed at the time of collection, immediate rapid freezing in liquid nitrogen or immediate freezing at −80 °C followed by storage in liquid nitrogen or at −80 °C are ideal. Unless the goal is cryopreservation, in many cases freezing at −20 °C or colder is suitable for several physiological analyses.

Preservation for short term storage (days-months)

Storage at −40 °C or colder for days to months is typically suitable for all physiological analyses, though some methods have additional requirements (Fig. 2). Storage at −20 °C for up to several months is also acceptable for many, but not all, analyses. In general, storage in liquid preservatives or fixatives (e.g., methanol, formalin, etc.) is considered either not acceptable or the efficacy of such preservatives is unknown when considered for many types of physiological analyses. Freeze-drying was noted as a suitable method for storing samples for physiological assays and is beginning to be used in coral bleaching studies (Wall et al., 2021; Pupier et al., 2021; Baumann et al., 2021). Freeze-drying is especially conducive to isotopic analysis (Wall et al., 2020) and provides for easy storage and transport as freeze-dried samples can be stored at room temperature. Notably, these samples should be stored in the dark.

Processing

For some physiological analyses, the processed subsample or specimen will be completely consumed and thus not available for long-term archiving. For example, a coral subsample which is ground and burned in a muffle furnace for ash-free dry weight biomass quantification cannot subsequently be used for chlorophyll concentration analysis. Notable exceptions include lipid extracts, skeletal material prepared for elemental analyses, and cryopreserved samples, all of which can be archived long-term for additional downstream analyses. Nevertheless, given the desire for technical replication and repeatability, many investigators typically collect coral fragments large enough to have surplus materials that remains unprocessed and may be placed into a long-term archive for potential use in other downstream analyses (summarized in Fig. 3). A strategically designed workflow could therefore use one suitably sized fragment for multiple methods.

Archiving for long term storage

Future physiological analyses are possible with frozen or freeze-dried fragments, ground whole coral, and frozen tissue homogenates and isolated fractions (e.g., host and microbial fractions), dried skeletal material, and cryopreserved tissues. Except for analyses that rely on live samples (e.g., cryobiology of reproductive cells and nanofragments; see section above on cryopreservation), all non-skeletal physiological analyses are possible with material that has been stored long-term at −80 °C or freeze-dried and then stored between −80 °C and −40 °C. However, the efficacy and accuracy of using material that has been in long-term storage between −20 °C to −40 °C is currently unknown. Due to protein degradation, denaturation, or other decomposition, physiological analyses are not possible on coral material that has been stored long-term at 4 °C or warmer. In most cases, long-term storage of more than a year is reasonable at temperatures of −80 °C or less, but it is unknown if some analyses could be reliably performed on material that was archived for more than 10 years (see above for cryopreservation where the efficacy of long-term storage is established, e.g., sperm storage >10 years). In this regard, more study is required to determine the maximum duration that samples can be archived for each storage method (i.e., liquid nitrogen, −80 °C, and freeze-drying) and still be suitable for physiological analyses. An exception to this rule is with the coral skeleton (usually in the form of cores, cross-sections, ramets, or ground powder), which is best stored dry at room temperature or refrigerated at 4 °C indefinitely so long as no future analysis of the skeletal organic matrix is intended.

Caveats

For all but the skeletal isotopic and elemental analyses, the subsamples for each individual physiological analysis are drawn from a representative, homogeneous mixture of either ground coral or tissue blastate that is collected from a larger fragment in both height and surface area than what is needed to make the specific measurement. In addition, obtaining a larger sized fragment or subsample than what is strictly needed to conduct the analysis is recommended to obtain a representative sample, to minimize edge effects potentially associated with the way the fragment was cut, and any positional effects within the coral fragment itself (e.g., top vs side of branch or mini mound).

Sample collection

Typically, coral samples collected from the field for Microscopy and Imaging should not be rapidly frozen, but instead placed in temporary storage, preferably in a cooler (with some fresh sea water), and out of direct sunlight. Microscopy and Imaging techniques differ from other methods in the requirement that samples must be as intact as possible (e.g., cells not lysed, corallites undamaged, or lacking any fragment alteration). A variety of imaging techniques such as 3-D photogrammetry and CT-scanning for surface area must be conducted on intact colonies or fragments prior to any additional analyses on ground skeleton or blastates. Similarly, photographs for coral color (e.g., to assess bleaching by loss of pigment) ideally are collected while the animal is alive before any other alteration to the colony has taken place and is thus best conducted underwater alongside a white standard or color chart. The need for intact skeletons and tissue extends to a variety of other Microscopy and Imaging techniques at smaller scales. Histology, XRF-scanning, Raman 2-D mapping, Nanoscale secondary ion mass spectrometry (NanoSIMS), and scanning electron microscopy (SEM) all require tissue or skeleton in their original shape or arrangement, on the scale of the analysis. For instance, using SEM to describe corallite morphology is possible on broken skeletal fragments, as long as those fragments are intact at the appropriate scale (e.g., an entire corallite, which may be <1 mm to tens of mm depending on the coral species). There are, however, a few methods that can use ground skeleton or tissue (e.g., Raman spot measurements, geochemistry), which can be conducted on altered or broken samples as long as they are suitably preserved. Investigations concerning the localization of nucleic acids, proteins, or microbes require immediate fixation upon collection.

Preservation for short term storage

The short-term preservation of samples for Microscopy and Imaging varies with the objectives of the research. For analyses requiring tissue fixation, the type of fixative used varies greatly between researchers and applications, and there is no single best method. There are several commercial preservative kits available (e.g., Bouin’s, Z-fixed (buffered aqueous zinc formalin), Glutaraldehyde/Paraformaldehyde solution mix), and many preservative types that can be prepared (e.g., formalin, glutaraldehyde, formaldehyde, methanol, ethanol, salt buffer). The proportion of each fixative (e.g., paraformaldehyde and glutaraldehyde) and time of fixation can significantly affect the accessibility of epitopes for immunolocalization (for both light and electron microscopy) and may require extensive optimization for each target and tissue type. Over-fixation is of particular concern for small samples like gametes and larvae. Techniques targeting nucleic acids (e.g., fluorescence in situ hybridization (FISH)) also require special handling and electron microscopy (EM) grade or molecular grade (RNAse/DNAse-free) reagents (Wada et al., 2016). The temperature and time of preservation varies with the preservative used and the type of sample collected (e.g., fragment, larvae, gamete).

By contrast, some preservation methods are simply unacceptable as they directly induce alterations that significantly affect the capability of researchers to adequately investigate the sample. Examples of this are histological and electron microscopy artefacts in coral tissue integrity and structure induced by freezing, changes in skeletal structure induced by chemical exposure, or the deterioration or alteration of nucleic acids when improperly cooled or stored in particular compounds or at incompatible temperatures.

For geochemistry-based methods, it is highly likely that any chemicals that are added to coral skeletons will change their chemical nature, therefore coral skeletons are best stored dry at room temperature or refrigerated at 4 °C when conducting geochemical focused imaging.

For imaging or light-based methods of Symbiodiniaceae quantification (e.g., hemocytometry, coulter counter, flow cytometry), short term sample preservation (e.g., of a tissue blastate) is recommended at 4 °C without fixation for two reasons: (1) freeze/thaw cycles can lyse symbiont cells, and (2) fixation can alter cell counts. However, coral fragments are commonly stored frozen (−20 °C to −80 °C) prior to tissue homogenization. Tissue homogenates can be stored frozen prior to analysis over the short term with the caveat that cell counts may be affected, and multiple freeze-thaw cycles are best avoided. Additionally, although cleaning skeletons (e.g., with hydrogen peroxide or sodium hypochlorite bleach) have potential to alter the ratios of certain isotopes (Grottoli et al., 2005; Holcomb et al., 2015), there is sparse information available on the effects of preservatives on coral skeletons. However, we suggest that adding any form of chemical preservative prior to geochemical analyses should be done with caution because these chemicals could alter the composition of coral skeletons by adding contaminants or causing partial dissolution.

Archiving for long term storage

Derivatives samples following application Microscopy and Imaging methods of coral skeletons are usually solid and exist in the form of cores, thin sections, or powder, and are best stored dry at room temperature or refrigerated at 4 °C. These can be kept indefinitely, although the true shelf life of each of these has not been thoroughly benchmarked for all downstream analyses. Coral tissues for histology-based methods are typically stored as fixed tissues embedded in paraffin blocks or sections mounted on microscopy slides and can be stored at room temperature or 4 °C. For immunolocalization and fluorescence in situ hybridization (FISH), these blocks can retain their quality indefinitely, whereas any generated thin sections from these blocks can deteriorate much more quickly (Wakai et al., 2014; Alamri, Nam & Blancato, 2017). Thin sections treated with dyes should be stored in the dark, whereas skeletons and tissue blocks are typically not light sensitive.

Derivatives of coral skeletons, in particular skeleton cores and thin sections, can be used for various downstream analyses, including skeletal imagining (e.g., using CT scanning, X-ray, and XRF) and skeleton geochemical analysis (e.g., Raman spectroscopy, isotopic and elemental analysis). Tissue-containing derivatives can be used for similar downstream analysis. There are no derivatives from the symbiont count techniques (i.e., hemocytometry, flow cytometry, automated cell counter), as the small sub-sample volume is typically consumed by these methods, though any remaining original sample may be available for additional analyses depending on its storage method (i.e., tissue blastate, Symbiodiniaceae pellet). Digital imagery produced from many of these techniques (e.g., electron microscopy, CT scanning, X-ray, coral color analysis), may be used for different image analysis techniques downstream.

While most fixed tissue derivatives can be stored at 4 °C or room temperature for long periods of time, stained (i.e., dyed) samples should be stored in the dark, and there is little known about the long-term preservation of samples for imaging. Long-term storage of coral tissues or homogenates for symbiont quantification that have not been chemically fixed is possible at −20 °C to −80 °C for >1 year if subsequential cell counts are performed with a hemocytometer, but not recommended at −20 °C due to Symbiodiniaceae cell degradation. Coral skeletons should be stored in a dry location at room temperature, and properly curated with metadata on collection dates, locations, water depths, etc. (Reich et al., 2012), and ideally with unique accession numbers.The preservation of photographic imagery, particularly those that document reef conditions during the previous century (e.g., Shinn & Kuffner, 2017), is also an important community goal.