A Semi-Automated Workflow for the Cryopreservation of Coral Sperm to Support Biobanking and Aquaculture

Coral reefs globally are experiencing a loss of coral species, populations, and genetic diversity due to ocean warming and acidification caused by climate change, diminishing the viability of these critical habitats and impacting the species that they support^1,2. The most effective approach to secure genetics long-term is cryopreservation and biobanking, which permits the frozen storage of living samples at cryogenic temperatures in liquid nitrogen indefinitely³. The development in 2012 of methods for the cryopreservation of coral sperm⁴ enabled the biobanking of genetics from these species for the first time and led to the development of the first biorepository for coral genetics in 2012⁵. Since then, this cryopreservation protocol has been further refined⁶ and used to secure the genetics of over 50 species of coral globally, with 30 of those species coming from the Great Barrier Reef in Australia⁷. Cryopreserved coral sperm can generate healthy corals that develop normally and have been used to facilitate assisted gene flow experiments in Australia⁸ and the Caribbean⁹. While technologies are currently under development to enable the cryopreservation of complex tissue types such as larvae¹⁰ and adult coral tissues^11,12, cryopreservation of coral sperm is currently the most established tool available for routine biobanking of coral genetics.

As the impacts on coral populations have increased, several countries have initiated large-scale programs to support reef restoration and adaptation (e.g., the Reef Restoration and Adaptation Program [RRAP] in Australia¹³) or to secure remaining imperiled coral populations (e.g., Florida Coral Rescue in the USA¹⁴). In the context of these programs, cryopreservation can be thought of as an enabling technology, supporting research and large-scale coral production in addition to securing existing genetic diversity and preventing extinctions. Sperm cryopreservation can enable greater control over reproduction between populations that are physically or temporally separated and can permit genetic management of broodstock to select for desirable traits such as thermal tolerance or disease resistance⁸. To date, coral sperm biobanking has been undertaken on a relatively small scale in support of biodiversity management⁷, so some level of upscaling will be required if such biobanking is to meet its potential within these larger reef restoration programs. As with all reef restoration efforts based on coral reproduction, the main impediment to increasing biobanking efforts is the limited period during which coral gametes are available, since spawning in most reef-building species coincides with the full moon in late spring and early summer¹⁵, meaning that gametes are only available for cryopreservation and biobanking on a few nights each year. Moreover, in regions where mass spawning occurs, e.g., the Great Barrier Reef, there are typically multiple species spawning simultaneously or within a few hours of each other on the same night¹⁵. Enhancements to the sperm cryopreservation pathway are therefore required to increase the scale and efficiency of processing to maximize biobanking capacity during these brief annual spawning windows while ensuring the integrity of samples and associated metadata.

Cryopreservation and biobanking of coral sperm involve several key steps, from the collection of gametes during spawning to the accession of samples into the biorepository and database (Figure 1). The process begins with the separation of sperm from eggs (in hermaphroditic species) or the collection of sperm from the water column (in gonochoric species), followed by the assessment of sperm motility and concentration. Sperm are then combined with cryoprotectant (10% v/v final concentration, dimethyl sulfoxide, DMSO) in filtered seawater (FSW) and cooled at −20 °C/min in a custom-designed cooling device⁴. Early iterations of this process relied on visual assessment of sperm motility and concentration via phase-contrast microscopy and hemocytometer counts, printing, and labeling tubes as samples were processed for cryopreservation, hand-pipetting sperm samples into cryovials, and cooling on a specially designed floating rack¹⁶. The application of computer-assisted sperm analysis (CASA)¹⁷ and the development of a 3D-printed cooling device⁶ have improved the efficiency and reliability of coral sperm cryopreservation, but the sperm sample processing pathway has remained largely the same since its inception. While this approach is suitable for processing samples from a moderate number of colonies and species on a spawning night, for large volumes and numbers of colonies (e.g., individual samples from >10 colonies at a time), there are bottlenecks in the cryopreservation pathway that impede processing of the samples in a timely manner (i.e., within 2 h of sperm release) without degradation of sample fertility potential. Although large-scale fluid handling systems for cryovials are available (e.g., Thomas et al.¹⁸), they are typically designed for large batch processing (i.e., multiple racks of cryovials) and are not suitable for transport and use in the field, so they are not cost-effective for this application. Therefore, the present study aimed to improve the efficiency of coral sperm cryopreservation by introducing portable, inexpensive automation equipment and methods at key steps in the processing pathway to maximize the number of samples that could be effectively cryopreserved on a spawning night.

The methods described herein can be used to process and cryopreserve sperm from both hermaphroditic and gonochoric coral species. A general introduction and primer on coral spawning and gamete collection for sperm cryopreservation for both reproductive modes can be found at the Smithsonian's National Zoo & Conservation Biology Institute Coral Cryopreservation Training Course¹⁹. The described methods were primarily developed using gametes from Acropora millepora colonies collected from the Keppel Island group on Woppaburra Sea Country and from Davies Reef on Bindal Sea Country, on the Great Barrier Reef in Australia. All collection and use of corals and gametes were undertaken with the free prior and informed consent of the Traditional Custodians of the relevant Sea Countries. The reagents and equipment used for this study are listed in the Table of Materials.

1. Pre-spawning - system checks and preparation of sperm activation solution and cryodiluent

1. Confirm that the barcode scanner is charged, active, and set to enter spreadsheet data as horizontal cell entry. Consult the barcode reader user manual to customize this setting.
2. Prepare coral sperm activation concentrated stock solutions for CASA.
   NOTE: Disposable gloves should be worn when handling chemicals for preparation of the activation solution.
   1. Prepare 30% bovine serum albumin (BSA) stock solution (5 mL) by dissolving 1.5 g of BSA in 4 mL of sterile purified tissue culture water with the aid of a tube warmer set to 37 °C or by holding the tube in cupped hands, with periodic gentle swirling (do not shake vial).
   2. Once in solution, make up to the final volume (5 mL) using tissue culture water, label the tube with the date, and mark "30% BSA in H₂O".
   3. Store in the refrigerator (4 °C) for up to 2 weeks.
   4. Prepare 60 mM caffeine stock solution (25 mL) by dissolving 0.2913 g of caffeine in 24 mL of filtered sea water (FSW).
   5. Once in solution, make up to the final volume (25 mL) using FSW, label the tube with the date and mark "60 mM caffeine in FSW".
   6. Store in the refrigerator for up to 1 month.
      NOTE: The maximum solubility of caffeine in FSW is approximately 71 mM.
3. Prepare sperm activation working solution for sperm samples at ~10⁹ sperm/mL (0.9% BSA + 12 mM caffeine in FSW).
   1. For 10 mL working solution, combine 2.0 mL of 60 mM caffeine stock solution + 0.3 mL of 30% BSA stock solution + 7.7 mL of FSW.
   2. Label the tube with date and "sperm activation working solution - 0.9% BSA + 12 mM caffeine".
   3. Prepare on each day of use, store at room temperature (19-30 °C), and discard the unused solution at the end of the day.
4. Prepare cryoprotectant solutions (cryodiluents). Ensure that disposable gloves are worn when handling the cryoprotectant DMSO.
   ​NOTE: FSW is prepared with varying cryoprotectant concentrations depending on the initial sperm concentration within a sample and the desired final sperm concentration of cryopreserved aliquots. When preparing the cryodiluent, ensure that FSW is aliquoted into the 50 mL tube prior to the addition of DMSO.
   1. Collect raw seawater from the tank system in which corals are held prior to spawning (~35 ppt salinity).
   2. Filter raw seawater using a bottle-top filtration system attached to a vacuum pump with a 0.2 µm non-pyrogenic membrane filter fitted.
   3. For a sperm concentration ≥2 × 10⁹ sperm/mL, prepare 20% DMSO in FSW by combining 10 mL of DMSO + 40 mL of FSW in a 50 mL tube.
   4. For a sperm concentration <2 × 10⁹ sperm/mL, prepare 30% DMSO in FSW by combining 15 mL of DMSO + 35 mL of FSW in a 50 mL tube.
   5. Label the tube with the date and DMSO concentration, i.e., 30% DMSO or 20% DMSO.
   6. Prepare on each day of use, store at room temperature (19-30 °C), and discard the unused solution at the end of each day.
      NOTE: Mixing of DMSO and FSW creates an exothermic reaction, and the cryodiluent should be made up well in advance of use to allow cooling to room temperature. FSW can be stored at 4 °C for preparation of the cryodiluent to counterbalance this heat generation.
5. Prepare thermocouple datalogger.
   1. To measure the approximate cooling rate of each cryopreservation run, place a cryovial containing 10% DMSO in FSW with a thermocouple probe inserted through the lid in a sample slot of the cryopreservation rack alongside the sperm samples.
   2. To make a thermocouple vial, poke or melt a small hole in the cap of the barcoded cryovial and insert a K-type or T-type thermocouple through the opening. Push the thermocouple probe tip down to a level such that it will sit in the middle of the sample when the vial is filled, keeping the probe tip centered in the vial to avoid contact with the inside vial wall. Seal the cap and hold the thermocouple probe in place with hot glue.
   3. Wearing disposable gloves, fill the barcoded cryovial with a volume of 10% DMSO in FSW equal to the volume of the sperm sample to be cryopreserved.
   4. Prepare fresh 10% DMSO in FSW and refill thermocouple vials daily.

2. Sperm preparation and assessment

1. For hermaphroditic species (e.g., Acropora species), collect gamete bundles from the surface of the water using a 3 mL transfer pipette and place them into a labeled 50 mL tube at a ratio of 5 mL of gamete bundles over 5 mL of seawater (10 mL total volume).
2. For gonochoric species, collect sperm from close to the polyp mouth using a 3 mL transfer pipette and place the sample into a labeled 50 mL tube, then proceed directly to step 2.8.
   NOTE: Disposable gloves are not necessary for the collection and handling of gamete samples but may be worn if preferred. Hands should be washed thoroughly with fresh water, or gloves changed between each gamete sample to avoid cross-contamination.
3. For hermaphroditic species, agitate the sperm-egg bundles by gently swirling the sample by hand until the bundles have broken apart.
   NOTE: Some Montipora species have a toxin in the eggs, so they must be allowed to fall apart slowly with minimal agitation to avoid damaging sperm.
4. Let the sample sit for 2 min in a tube rack to allow eggs to float to the surface and sperm to settle (eggs will form a pink/brown layer at the surface, and individual eggs will be visible; see Figure 1i).
5. To separate the sperm and remove contaminating eggs, gently aspirate the sperm from the bottom of the tube using a clean 3-mL transfer pipette.
6. Transfer the sperm sample to a 100 µm cell strainer sitting atop a new sterile 50 mL centrifuge tube. The sample should flow easily through the filter. Tap the filter to encourage sample flow, and if necessary, a clean transfer pipette can be used to collect any residual sample from the underside of the filter. Remove the filter, then cap the tube loosely to allow air exchange.
7. Label the filtered sperm sample with colony ID and move the sample to the sperm assessment workstation.
8. Open the coral biobanking auto-datasheet file (Supplementary File 1). In the sperm assessment tab, input the coral colony metadata (columns A-H).
9. Prepare sperm for CASA assessment.
   1. Mix the sperm sample well and remove a 10 µL aliquot to an empty 1.8 mL microcentrifuge tube.
   2. Immediately add 0.390 mL of sperm activation working solution dropwise over 10-20 s, gently mixing sperm into the solution. This gives a dilution ratio of 1:39 (sperm: solution, v/v) (or dilution factor of 40), which provides an appropriate concentration for CASA assessment (~50 × 10⁶/mL) if raw sperm concentration is ~2 × 10⁹ sperm/mL. Invert the tube 5 times and flick the tube lid prior to opening to settle the solution into the bottom of the tube.
      NOTE: Sperm concentration for CASA should be in the range of 8-50 × 10⁶/mL to permit accurate sperm tracking¹⁷. Additional sperm activation working solution can be added to the raw sperm aliquot to reduce the concentration if needed. Recalculate the dilution rate and enter this value into the spreadsheet. In cases where many samples must be processed simultaneously, the 1.8 mL microcentrifuge tubes can be pre-loaded with 0.390 mL of sperm activation working solution with the 10 µL aliquot of sperm sample added directly to it, then mixed well by inverting the tube 10× and flicking the tube lid prior to opening.
10. Assess sperm motility and concentration of activated sample using computer-assisted sperm analysis (CASA) as described in Zuchowicz et al.¹⁷ by placing 4 µL of activated sperm suspension on a Makler counting chamber²⁰ or by visual assessment and hemocytometer count under phase contrast microscopy as described in Hagedorn et al.⁴.
    NOTE: The Makler chamber and coverslip must be cleaned well with 70% ethanol, then distilled water, and wiped with a lint-free tissue wipe between samples.
11. Input the sperm dilution factor used for CASA assessment and enter CASA outputs for sperm concentration (million/mL), total motility (%), and progressive motility (%) into the auto-datasheet.
    NOTE: Additional CASA metrics, including average path velocity (VAP), straight-line velocity (VSL), and curvilinear velocity (VCL) can optionally be included in the auto-datasheet or retrieved from saved CASA files later.
12. Write the sperm concentration on the filtered sperm sample tube and transfer the sample to the cryopreservation workstation in the same order as of bundle break-up and processing.

3. Sperm dilution and cryoprotectant equilibration

1. At the cryopreservation workstation, open the same coral biobanking auto-datasheet file that the sperm assessment workstation is using and select the Cryopreservation tab. Check the sperm sample tube label and identify the corresponding entry by checking Colony ID (column C). If not accessing the same auto-datasheet file simultaneously between workstations, manually enter data into the Colony ID (column C) and sperm concentration (column F) in the auto-datasheet.
2. Using a serological pipette, measure the sperm sample volume by drawing the sample into the pipette and expelling it back into the same tube; enter the value in column E.
3. Check the auto-calculated column I for the required cryoprotectant concentration and column H for the volume of cryodiluent to be added (Table 1).
4. Start a timer for 10 min and, using a serological pipette, immediately begin adding the cryodiluent, dropwise, with constant gentle mixing of the sample, ensuring that disposable gloves are worn for DMSO handling. 
5. Record cryodiluent addition time in column P.

Table 1: Cryodiluent concentration and dilution ratios for cryopreservation of coral sperm, based on assessment of total sperm concentration in the sample to be cryopreserved.

4. Sperm cryopreservation

1. Read column K to determine how many cryovials to fill. This is auto-calculated from the total volume of sperm and the desired volume per vial (1 mL in most cases). During the 10 min incubation period, proceed through steps 4.2 to 4.6.
2. Uncap sterile barcoded cryovials and set the caps on a clean/sterile surface.
3. Optional: Write the run # on each cryovial using a permanent marker; this can aid in quick sample identification for accessioning into the biobank.
4. Using a serological pipette and an aliquoting pipettor set to the desired sample volume (1 mL), aliquot samples into uncapped barcoded cryovials and recap vials.
5. Place one thermocouple vial and all full sample vials onto an empty cryo rack at room temperature. Ensure that the barcoded surface is facing outwards on all vials.
6. If necessary, place additional dummy cryovials containing 10% DMSO in FSW in any empty spaces on the cryo rack to ensure that all spaces are occupied.
7. Enter the run number into the auto-datasheet (column U) and scan the cryo rack serial number (QR code) into column V.
8. Place the cursor in the correct cell of column Z and scan in the thermocouple vial, followed by all cryovial tubes that are loaded on the rack (column AA onwards). Avoid tilting the rack on its side to ensure the sample does not enter the thread of the vial.
9. While scanning, cross-check that data is being entered into the correct cell and that all cryovials have been scanned once.
10. Set aside loaded and scanned racks for the remainder of the cryoprotectant equilibration period and prepare the cooling vessel for cryopreservation.
11. Fill the cryo rack cooling vessel with liquid nitrogen to the appropriate pre-determined level necessary for cooling at approximately −20 °C/min.
    NOTE: Nitrogen should only be used in well-ventilated areas as there is a risk of asphyxiation. Closed-toe shoes, safety eyewear/face shields, and cryo gloves must be worn when handling liquid nitrogen. Refer to the institutional guidelines for specific information on liquid nitrogen handling and use.
12. At the end of the 10 min cryoprotectant equilibration period, connect the thermocouple probe to the datalogger and begin recording, then gently place the full cryo rack into the cooling vessel and apply the lid.
13. Record the start time in column Q.
14. Once the thermocouple vial reads −80 °C or below on the datalogger, quench the samples in liquid nitrogen by transferring the cryo rack insert to a liquid nitrogen bath in a separate vessel such as a polystyrene box.
15. Remove the cryovials from the rack and transfer them to a dry shipper for transport to the biorepository.

The steps outlined in this protocol build on the original methods for coral sperm cryopreservation described by Hagedorn et al.⁴ and subsequent refinement by Zuchowicz et al.⁶, providing key improvements to increase the efficiency of sperm processing for cryopreservation and biobanking. The use of barcoded cryovials and an aliquoting auto-pipettor simplifies the sperm packaging procedure by removing the need to print and hand-label cryovials, and reduces the strain of hand-pipetting large numbers of samples. Importantly, these improvements also reduce the time needed for sample preparation for cryopreservation, from around 8-10 min to under 5 min. Together with the use of CASA, these improvements reduce the number of personnel required for efficient biobanking of coral sperm, from 4 to 6 in the original methods, down to as few as two people using the current protocol. Additionally, the use of barcoded cryovials means that each tube has a unique identifier, so each can be tracked within the database with greater resolution than the previous method using batch-printed labels.

Field testing of the protocol across two spawning events during the 2022 Great Barrier Reef spawning season in Australia resulted in the cryopreservation and biobanking of 389 samples from 57 colonies across 5 species by two operators (one person for CASA, one person for cryopreservation), with one additional person assisting with bundle break-up and sperm separation as needed. During the November 2022 spawning event on Konomie (North Keppel Island) in Queensland, Australia, 150 samples from 24 colonies of Acropora millepora were cryopreserved by two operators over a 2 h period. The efficient processing and cryopreservation of this volume of samples from such a large number of coral colonies was enabled primarily by the efficiencies gained in sample preparation time and the simplified transfer of metadata between workstations.

Further testing of the workflow during spawning in December 2023 determined that, for coral sperm, the Makler counting chamber provided more accurate motility and concentration data compared to commercially available fixed-coverslip single-use chamber slides commonly used with CASA systems. The Makler counting chamber was validated for CASA using commercially available latex microbeads at a known concentration with a 4 µL sample volume and the standard CASA settings¹⁷ used for coral sperm. These analyses showed consistent counts with low variability (44.5 ± 0.7 million/mL) that fell within the expected concentration range (46.0 ± 7.0 million/mL) provided by the manufacturer (Figure 2). A comparison of CASA data for Acropora millepora collected using the Makler counting chamber and the fixed-coverslip chamber slides found a significant difference in sperm concentration counts between the two chamber types (P = 0.002; Figure 2), with the fixed-coverslip chamber slides registering less than half of the concentration determined using the Makler counting chamber. This trend was also observed in sperm samples from two other coral species (data not shown). Testing during December 2023 also found no significant difference in the post-thaw concentrations of total, motile, or progressively motile sperm in samples cryopreserved using either a 1:1 dilution with 20% DMSO or a 1:2 dilution (sperm: cryodiluent) with 30% DMSO (Figure 3).

Figure 1: The workflow used for semi-automated processing of coral sperm for cryopreservation and examples of mobile equipment used in the field. (A) Coral gamete bundles are collected at a ratio of 5 mL bundles over 5 mL of FSW and broken up to separate eggs and sperm (i). Colony metadata are entered into the auto-datasheet (ii), and the isolated sperm sample is assessed using computer-assisted sperm analysis (CASA) (iii). Sperm quality parameters are added to the auto-datasheet to calculate the addition of either 20% or 30% DMSO in FSW (iv), and the diluted sample is aliquoted into barcoded cryovials (v). The cryovials are loaded onto the cryo rack and scanned into the auto-datasheet (vi), then cooled at a rate of −20 °C/min to −80 °C (vii). The samples are then quenched in liquid nitrogen and transferred to dry shippers for transport to the Taronga CryoDiversity Bank (viii). (B) Example of the CASA (left) and cryopreservation (right) stations set up at a remote field site in the classroom at the Konomie Environmental Education Centre. (C) Example of the lab-based high-capacity cryopreservation station set up to run multiple cryo racks at the Australian Institute of Marine Science National Sea Simulator. Please click here to view a larger version of this figure.

Figure 2: Comparison of two different chamber slide options for measurement of coral sperm concentration. (A) Comparison of total sperm concentration measurements in Acropora millepora (n = 3 individuals) using the Makler counting chamber and commercially available fixed coverslip chamber slides. Columns show mean ± SEM.; asterisks indicate a significant difference (P = 0.002). (B) Validation of the Makler counting chamber using commercially available latex microbeads at a known concentration and standard CASA settings for coral sperm. Please click here to view a larger version of this figure.

Figure 3: Post-thaw concentrations of total, motile, or progressively motile sperm in cryopreserved samples. Post-thaw comparisons of total, motile, and progressively motile sperm concentrations using either 20% or 30% DMSO in FSW to achieve the final 10% DMSO concentration for cryopreservation. Samples from n = 3 individuals were split into 2 aliquots each, with one aliquot cryopreserved with a 1:1 addition of 20% DMSO and the second aliquot diluted to <2 × 10⁹/mL using FSW for cryopreservation with a 1:2 addition (sperm: cryodiluent) of 30% DMSO. Columns show mean ± SEM. Please click here to view a larger version of this figure.

Supplementary File 1: Coral biobanking auto-datasheet file. Please click here to download this File.

The semi-automated processing pathway described in this protocol permits the efficient processing and cryopreservation of coral sperm to secure genetics from threatened species and support reef restoration and adaptation efforts. The motivation for the development of this protocol was the lack of existing systems suited to the throughput requirements of coral sperm cryopreservation and the use of cryovials, since high-throughput processing systems for sperm cryopreservation are typically based on sample packaging in 0.25 mL or 0.5 mL French straws^21,22. By comparison, cryovials are generally either used at a small scale for low throughput processing (e.g., cryopreservation of laboratory samples for research^23,24), or in high-throughput processing systems for bulk samples using expensive, non-portable equipment (e.g., cell culture processing for industry^18,25). We also investigated the potential for using an auto-decapping system to streamline the removal and replacement of cryovial caps, but systems were only available for individual cryovials or for entire cryovial racks, so they did not provide a cost-effective solution. Currently, there are several groups globally who are using the cryopreservation protocol devised by Hagedorn et al.⁴ to secure the genetic diversity of corals, and it is important that this work continues to expand to more reefs around the world. Therefore, a major consideration in the development of the current protocol was the need to utilize cost-effective and accessible technologies that could be readily implemented by these other groups, and which would not be cost-prohibitive for new groups wanting to begin coral sperm cryopreservation.

A key component of the described protocol is the improved handling of sample metadata via linked spreadsheets in Microsoft Excel. Data input is generally straightforward, but it must be noted that editing information in the auto-datasheets should only be done by deleting and re-entering information, as quick functions (e.g., Ctrl + C, Ctrl + V) to edit data will potentially affect simultaneous inputs by other users and can cause problems with data linking between spreadsheets. An important metadata component is a unique identifier (namely, colony ID) that is linked to the donor colony and that is attached to the sample at all stages of the processing pathway. It is essential that sample tubes be clearly labeled with the colony ID at the time of bundle collection, and that this information be accurately transcribed onto any new tubes into which the sample is transferred during preparation (e.g., during filtering to separate eggs and sperm, or for cryodiluent addition). Although the protocol enables the automatic transfer of sample quality information from the CASA operator to the cryopreservation workstation, issues can be encountered when internet or Wi-Fi coverage is poor. If data transfer delays are encountered, it is recommended that the two workstations work offline and maintain separate auto-datasheets that can be reconciled afterward. The key information required by the cryopreservation operator is the colony ID and sample concentration, so it is recommended that the CASA operator write the sample concentration on the sample tube as a backup to ensure that this information is at hand for cryopreservation preparation if there is a delay in the upload or download of metadata between computers.

The choice of barcode scanner and barcoded cryovials used for this protocol can be varied to suit budget and product availability; however, there are some key elements that should be considered in their selection. The barcode scanner should have the capacity for customized settings, specifically the ability to change the data input specifications and direction of data entry. The auto-datasheets used for this protocol use horizontal entry, but on some occasions (e.g., accessioning into the biobank or for other laboratory uses) vertical entry may be required, so it is important that this feature be customizable. While the protocol can be used with both 1D and 2D barcodes, it is recommended that the cryovials selected have a human-readable component (e.g., 1D barcodes typically include a unique number) to permit cross-checking of sample entry during cryopreservation. In addition to sample entry, the barcode scanner can be used to automate the input of some metadata fields by creating and printing out QR codes for information that is repeated across samples (e.g., species names, dates, and reef locations) prior to spawning. This information can then be quickly and easily entered into the auto-datasheets by scanning with the barcode scanner. Moreover, by attaching serial number barcodes to each cryo rack and thermocouple probe, it is also possible to link each cryopreservation run with a specific set of equipment within the database, which is useful for quality control and to identify components that require repair or replacement.

The limiting factors in processing are often the time spent waiting for bundles to break up and the time required to filter and separate the sperm from the eggs prior to CASA and cryopreservation. Where possible, it is recommended to process samples in the order that bundles break up; however, further gains in efficiency can be made through the strategic management of sample processing orders. For example, if there are 5 or fewer cryovials per colony due to low sample volumes (i.e., less than 3 mL sperm per colony) or low sperm concentration requiring a 2:1 dilution with cryodiluent (i.e., less than 2 × 10⁹/mL), then it is better to add cryodiluent to two colony samples at the same time so that they can be run together on the same cryo rack (total # slots available = 11), rather than running them separately with dummies filling the empty spaces. Additionally, it is possible to run multiple cryo racks simultaneously (for samples >6 mL volume) provided that care is taken to ensure that all processes can be completed within the 10 min cryodiluent equilibration time, which can further increase sample throughput. However, when running multiple cryo racks or combining multiple colonies on a single cryo rack, care must be taken to ensure that cryopreservation metadata is assigned to the correct sample in the auto-datasheet, especially if samples are cryopreserved in a different order to their CASA assessment.

In addition to the development of the semi-automated workflow, the present protocol description also provides two methodological comparisons related to sperm concentration that aim to improve sperm analysis and cryopreservation outcomes. In general, collecting 5 mL of gamete bundles over 5 mL of seawater (total volume 10 mL) will result in a sperm concentration at or above 2 × 10⁹ cells/mL, but there are occasions when the sperm concentration may be lower due to species differences or bundles breaking up during collection. The use of a higher concentration DMSO cryodiluent (30% v/v) reduces the amount by which the sperm are diluted to help minimize batch-to-batch variation in the concentration of sperm in the cryovial. Importantly, the use of 30% DMSO to achieve the final DMSO concentration does not impact post-thaw concentration or motility parameters, as shown by the representative data in Figure 3. The second methodological comparison provides an alternative to the single-use fixed-coverslip chamber slides typically used for CASA. The main challenge with using commercially available slides for the analysis of coral sperm is that they can impact the accuracy of motility assessment due to sperm sticking to the slide coating. The use of the activation solution overcomes this issue in many but not all samples, so it is still recommended to perform a separate CASA analysis for motility using a plain slide to ensure reliability and consistency. The use of a Makler counting chamber overcomes the need to analyze concentration and motility separately, and potentially improves the accuracy of concentration measurements (Figure 2), so it is recommended for use with the current protocol. Given this discrepancy in concentration measurements, a finding that has been reported previously²⁰, it is important always to record slide details in the database alongside sperm quality data and, wherever possible, to be consistent in the type of chamber slide used to minimize batch-to-batch variation and help to ensure reliable calculations of sperm: egg ratios for fertilizations.

The semi-automated process described herein provides a standardized and efficient pathway for cryopreservation and biobanking of sperm from threatened coral species while retaining sample biosecurity and quality. The protocol described is easily transferrable and relatively inexpensive to implement in programs around the world that are working to secure existing coral diversity using cryopreservation, which will be essential to prevent extinctions and maximize the resources available for reef restoration efforts now and in the future.