Field Collection and Laboratory Maintenance of Canopy-Forming Giant Kelp to Facilitate Restoration

Kelp tissues used as described in this protocol were collected and overseen by the California Department of Fish and Wildlife under permit S-202020004-20205-001.

1. Preparation of facilities and materials

1. Ensure that kelp culturing facilities can maintain temperature (10-15 °C), provide full-spectrum light (0-180 µmol photons m^-2 s^-1), and filter aeration (0.2 µm pore size). Use incubator systems with a built-in outlet or an access port for wires and tubing, lights, and an air source (Figure 1). If an incubator system is not within the project's scope, budget, or intended scale, use water baths tempered by cool, natural seawater or a chiller (Figure 2). Refer to the Table of Materials for specific details.
   1. Place a thermometer in the growth media or use a temperature gun to ensure the temperature is between 10-18 °C.
      NOTE: Rearing temperatures are site- and season-specific³⁰.
   2. Program full-spectrum lights to a photoperiod of 12 h light: 12 h dark by changing the timing settings on the light source or by utilizing a mechanical timer. Measure the light intensity with a waterproof photosynthetically active radiation (PAR) quantum meter below the surface near the gravel and adjust using a dimmable light source or by layering cellophane (in the case of vegetative gametophyte culturing, see Section 9) or mesh over the light source (for details on light intensity adjustments, see Section 6.3).
   3. Ensure proper aeration by using air pumps³¹ one day after sporulation. Use filters (0.2 µm pore size) to reduce air-borne bacterial contamination.
      NOTE: For 'green gravel' culturing, aeration pressure must be sufficient to circulate water in all culture containers while not disturbing the attachment of early-stage kelp to seeded substrates. If bulking gametophyte biomass (see Section 9.2) are present, aeration pressure must be sufficient to maintain gametophytes suspended in the culture media.
2. Sterilize materials and stations. Prepare these ahead of time (see Table of Materials).
   1. Clean surfaces using 70% isopropyl alcohol. Handle reproductive sori tissue and clean collection equipment outside the 'green gravel' nursery, if possible.
   2. Use the following sterilization methods: rinse using a lab-grade detergent followed by a thorough rinse with distilled water, soak in a diluted bleach solution (according to manufacturer directions) followed by a thorough rinse with distilled water, and autoclave using appropriate settings (glassware or instruments). After sterilization, materials can be stored in a sealed container or wrapped with foil.
   3. Scrub and clean containers with the lids to be used for culturing using a lab-grade detergent, followed by a thorough rinse with distilled water.
      NOTE: Lidded culture containers will help reduce the evaporation of growth media. Leave lids slightly open to allow for air exchange, or use a check valve to reduce airborne contamination. If lidded containers are not available, seal culture containers with a thermoplastic such as paraffin film and make 2-3 perforations. If larger tanks are being used, use anti-evaporation covers made from transparent plastic.
   4. Ensure that gravel have a textured or slightly pitted surface since gametophytes are more likely to be retained on substrates with a high rugosity^32,33. Scrub and rinse gravel until the water runs clear to remove any dust or debris. Soak gravel in a 10% diluted bleach solution for at least 24 h and rinse with filter-sterilized seawater (see Section 2.1). Alternatively, after scrubbing and rinsing, soak gravel for 1 week in de-ionized (DI) water³².
      NOTE: Ideally, locally harvested substrate is used to reduce contamination of the restoration site. Alternatively, aquarium-grade gravel is recommended. Avoid calcareous substrates such as limestone, which can lead to tissue bleaching and subsequent mortality of transplanted kelps³².

2. Preparation of growth media

1. Filter and sterilize seawater according to the following methods, depending on resource availability. Calculate the volume of filter-sterilized seawater needed to refresh culture containers each week (see Section 7) and schedule this filtration/sterilization task accordingly. Store large batches of filter-sterilized seawater in dark containers for up to 6 months at 8-10 °C. If refrigeration is not available, store in a dark, cool area.
   1. Filter water using a vacuum filtration system with a pore size of 0.55-1 µm. Turn the vacuum source off before all the water is pulled through to avoid damaging the filter, and pour the filtered water into a dedicated sterile container. For larger volumes, use a flow-through filtration system. For example, run seawater through a series of three pleated filters (10 µm, 5 µm, and 1 µm) arranged from largest to smallest pore size.
      NOTE: If natural seawater is not accessible, artificial seawater can be prepared. Alternatively, natural seawater can be purchased from aquarium stores, in bulk and is often filtered, sanitized, and pH balanced. Media enrichment is still necessary for these options.
   2. Sterilize filtered seawater using UV and/or autoclaving methods. Connect flow-through systems to an aquarium UV light at a flow rate recommended by the manufacturer. Autoclave seawater in autoclave-safe glassware with slightly open lids or covered with foil and on a liquids cycle (121°C; 1-2 PSI, 15-30 min depending upon the volume of liquid³⁴.
      NOTE: Autoclaving filtered seawater is recommended for the early stages of culture.
2. Enrichment of filter-sterilized seawater with nutrients and vitamins is critical to M. pyrifera growth. Provasoli Enriched Seawater media (PES) is a widely used medium designed for algal cultures³⁵. Purchase this media from algal culture centers. Preparations of PES and additional vitamins for M. pyrifera growth are described in³⁴.
   1. Enrich every 1 L of filtered seawater with 20 mL of PES. Alternatively, use industrial-level culturing media.
   2. Store enrichment solutions according to the manufacturer's recommendations. Enrich filter-sterilized seawater when growth media is needed to avoid degradation of enrichment solutions.

3. Field collection

1. Determine the timing of sporophyll collections to mimic the natural reproductive cycle of local M. pyrifera populations. Consult local experts (e.g., kelp researchers, managers, ecologists, citizen scientists, dive groups) to ensure appropriate timing for sporophyll collection.
2. Obtain the necessary permits for kelp tissue collection that meet local laws and regulations. This can be a time-consuming part of the culturing process and must be incorporated into project timelines.
3. By self-contained underwater breathing apparatus (SCUBA) to select 3-5 sporophyll blades from 10-15 fertile M. pyrifera individuals with visible sori, spaced at least 2 m apart. Select clean and intact sporophylls, if possible, with little to no fouling or degradation. Store sporophyll blades separately according to the parent individual from this point forth.
   NOTE: Sporophylls grow in a dense "skirt" at the base, above the holdfast of the adult kelp, and can be identified by their lack of gas-filled pneumatocysts¹. Mature sorus tissue is often slightly raised and darker in color than surrounding tissue¹.
4. Transport sporophyll blades in dark collection bags to avoid overexposure to sunlight, with minimal seawater from the site to keep blades wet, and store in coolers at approximately 12 °C until arrival at the culturing space. Ensure that samples are not in direct contact with ice.
   NOTE: Sporophylls can be shipped to or from other locations.
   1. Rinse sporophylls with seawater. Wrap blades, collected from a single M. pyrifera individual, in moist paper towels soaked in seawater and again in aluminum foil to avoid light penetration and additional desiccation³⁶. This method of storage is commonly known as the "burrito method".
   2. Place these packages in a cooler with ice, with a protective barrier such as recycled bubble wrap or cardboard. Prepare the cooler for overnight shipment. Ensure that someone is available to receive the shipment and place the packages in refrigerated conditions.

4. Sporulation

1. If possible, process sporophylls in a temperature-controlled environment between 10-15 °C and away from any other cultures. Prepare and sterilize instruments and stations ahead of time. Wear protective gloves when handling kelp tissue to reduce contamination.
2. Optionally store sporophylls for 12-48 h in refrigerated conditions, encouraging spore release from sorus tissue³⁷. To store, use the "burrito method" described in Section 3.3.
3. Select ripe sorus tissue and cut it into 25 cm² sections using sterile scissors. Select 1-2 clean sori sections from 10-15 individual kelp parents, to promote genetic diversity.
   NOTE: If stored, optionally find evidence of partial sporulation on the paper towels, indicating the presence of fertile sorus tissue. Sorus tissue is often slightly raised and darker in color than surrounding tissue.
4. To clean, gently scrub both sides of the sorus tissue in one direction only with a sterile gauze damped with filter-sterilized seawater. If needed, scrape the sorus tissue gently with a sterile razor blade to fully remove fouling. Submerge the sori section in a freshwater bath for 30 s to 1 min and rinse with filter-sterilized seawater.
   NOTE: Refresh the freshwater bath and sterilize the materials in use when handling different sori sections from different individuals to reduce cross-contamination.
5. Submerge each sori section in filter-sterilized seawater tempered to 10-15 °C within a sterile 50 mL centrifuge tube. Place tubes at 4-12 °C in the dark to sporulate for a maximum of 4 h. If a refrigerator is unavailable, store it in a low-light, cool area.
   NOTE: Alternatively, sori sections can be sporulated in a single, sterile container.
6. Using a compound microscope and hemocytometer, observe the spore density of 3-4 samples every 30 min up to 4 h. Change pipette tips between samples. If densities are at least 10,000 spores mL^-1 (see Section 5.1.1), move on to the next step. If a sori section produces no spores after 4 h, discard the sample. Spores can settle within hours after the release but may be observed swimming in a circular motion.
7. Remove each sori section from the tubes with sterile tweezers. Combine the resulting spore solutions into a single, sterilized container and quantify the final combined density.

5. Inoculation

1. Calculate the final volume of spore solution needed for inoculation. Ensure the final concentration is approximately 500-1,000 spores mL⁻¹ in culture containers.
   1. To calculate the concentration of the combined spore sample from counts of the center grid of the hemocytometer, divide the count by 10^-4 mL (representing the volume of solution viewed in the hemocytometer).
   2. To determine the volume of the spore solution to add to each container, determine the amount of growth media needed to submerge substrates within culture containers.
   3. To find the total number of spores in each container, multiply this seawater volume by the desired concentration.
   4. To determine the total volume of spore solution to be added, divide the total amount of spores by the concentration of spores per mL in the spore solution.
2. Place sterile glass slide(s) within culture containers to monitor kelp development. Include at least 30 slides distributed randomly across culture containers for sufficient monitoring (see details in Section 7).
3. Inoculate the calculated volume of spore solution into the culture container using a sterile pipette tip that contains substrates submerged in growth media. Close the container and gently stir to distribute spores. Seal and place the container into the culture system.

6. Rearing conditions

1. Set the temperature between 10-15 °C based on the temperature at the deployment site.
2. After 1 day, provide light aeration with a filtered air source.
3. Set full-spectrum LED lights for aquatic plants to a 12 h light: 12 h dark cycle, with light intensities ranging between 0-180 µmol photon m^-2 s^-1:
   1. Set the light intensity to 5-10 µmol photon m^-2 s^-1 from 0-1 day and increase to 20 – 30 µmol photon m^-2 s^-1 through the end of 1 week.
   2. From this point on, increase the irradiance by 10-20 µmol photon m^-2 s^-1 every 3-4 day until reaching an irradiance of 180 µmol photon m^-2 s^-1 at the end of 6 wk.
   3. Continue to rear cultures at 180 µmol photon m^-2 s^-1 through the end of 8 wk, or when sporophytes have reached approximately 1-2 cm in length.

7. Monitoring

1. Monitor at least two random glass slides daily/every other day for the first two weeks to assess development.
   1. To monitor, handle the slide with sterilized tweezers and place it in a clean Petri dish containing enough sterilized seawater to submerge the glass slide. Do not return glass slides to cultures after being removed to avoid cross-contamination.
   2. Use a compound or inverted microscope at 40-400x magnification to observe early-stage kelps. Track the development with the following timeline (see Figure 3for examples of developmental life history stages).
      NOTE: Settled spores are observed at 0-1 d. Spores can germinate within a few hours, as demonstrated by the formation of a germ tube. Germination is typically observed at 1-2 d. Early gametophytes are typically observed at 1-4 d. Gametogenesis, the process by which cells undergo division and differentiation to form male and female gametes, is typically observed within the first two weeks. Female cells are 5-7 times larger than males. Male gametophytes grow thin, filamentous branches, whereas females are more round or ovoid in shape. Females typically produce eggs or ova within 2-3 week. Sperm released from the males swims to the females and fertilizes the eggs, resulting in the formation of diploid zygotes. Having the right inoculation density will ensure successful reproduction by proximity^38,39. Fertilized eggs develop into embryonic sporophytes. Sporophytes are typically observed within 2-4 week. The zygote undergoes rapid cell division, resulting in the growth of 1-2 cm blades within approximately 6-8 week.
2. After two weeks, monitor at least two random glass slides 1-2 times weekly for healthy growth and contamination until sporophytes reach 1-2 cm in size.
   NOTE: Healthy growth is characterized by golden-brown (as opposed to green or transparent) coloration. There are several quantitative metrics that can be observed on glass slides with an inverted microscope, including survivorship, germination rate, vegetative development, reproductive maturity and fecundity, and sex ratio⁴⁰.
3. Assess contamination by bacteria, fungi, ciliates, and diatoms with a microscope. Remove isolated contamination. Control early signs of diatom contamination with germanium dioxide (GeO₂) (see Section 8.3) treatment.

8. Maintenance

1. Adjust light conditions according to Section 6.3.
2. Every week, change growth media to replenish the necessary nutrients and minerals for M. pyrifera growth.
   1. Chill fresh growth media to the appropriate temperature. Ensure temperature does not exceed 15 °C during this process.
   2. Siphon media out of the culturing containers to avoid disturbing seeded substrates. Let the media drain until the container is nearly empty. Refresh media immediately to minimize desiccation. When refilling growth containers, tilt them slightly so that media runs down the side of the culturing container to disturb substrates minimally.
   3. Randomly rearrange container or tub positions during weekly media changes to account for differences in light irradiance.
      NOTE: See Supplementary File 1 for a calendar to track activities and expectations for Macrocystis cultures. It indicates the timing of adjustments to light and aeration, as well as weekly media changes.
3. Optionally, control diatom contamination with a treatment of germanium dioxide (GeO₂). Add 0.3-0.5 mL of 250 mg/mL GeO₂ to each 1 L of seawater added to the seeded substrates to reduce widespread diatom contamination.
   NOTE: GeO₂ may inhibit algal gamete production. Apply a treatment of GeO₂ in the short window after germination and before peaks of egg and sperm production (1-7 d) and/or after egg fertilization and sporophyte observations (>21 d), followed by a media change 48 h after to remove the chemical. These timelines may vary given culture conditions, so monitoring life stage development with microscopy is the best way to assess the timing of GeO₂ application. If diatom contamination persists in culture containers and overgrowth onto early-stage kelps is observed, consider re-seeding the substrates.

9. Giant kelp vegetative gametophyte culturing

1. Propagate gametophyte cultures in vegetative conditions year-round to reduce dependency on seasonal sporophyll collection from the natural reef.
   1. Store gametophyte cultures according to the source population in flasks filled with growth media at 4-12 °C in red light at an intensity of 5-20 µmol photon m^-2 s^-1 in a 12 light: 12 dark cycle.
   2. Provide constant aeration and change media every 2-6 months.
2. To bulk-up biomass of gametophytes that have been growing asexually for use in 'green gravel' seeding, increase the aeration to suspend gametophytes, increase the frequency of media changes to weekly, and fragment gametophytes every two weeks.
   1. Suspend gametophyte biomass in the culture flask by shaking or stirring and scrape the sides of the culture flask with a sterile tool to dislodge attached gametophytes, if necessary.
   2. Pour the suspended gametophytes into a sterile blender or coffee grinder and pulse the gametophyte solution for 1-2 s approximately 5-15 times, depending on biomass concentration, until no clumped masses are visible.
3. To induce reproduction for 'green gravel' seeding, fragment gametophytes, as explained above. Then, inoculate substrate and increase full-spectrum LED light from 5-20 to 45-60 µmol photon m^-2 s^-1 (+10 µmol photon m^-2 s^-1 daily for photo-acclimation), then increase by 10-20 µmol photon m^-2 s^-1 every 3-4 d until reaching an irradiance of 180 µmol photon m^-2 s^-1.

10. Deployment

1. After 6-8 wk of laboratory culturing, ensure that juvenile sporophytes are 1-2 cm long and ready for deployment (Figure 4). Refresh growth media in culture containers 24 h before deployment.
2. Obtain the necessary permits for gravel deployment that meet local laws and regulations. This can be a time-consuming part of the culturing process and must be incorporated into project timelines.
3. Transport 'green gravel' in trays covered with towels soaked in seawater to keep the kelp hydrated. Place trays in insulated coolers with ice, ensuring they are not in direct contact with ice. Make sure that the 'green gravel' is tightly packed to avoid rolling of the substrates and sporophyte detachment during transportation.
   NOTE: Depending on the space availability, substrates can also be transported in their culture containers or tubs to reduce handling.
4. Transport 'green gravel' for up to 6 h in a shaded cooler. Deployment should be timed to avoid the most direct sunlight. If deploying from a boat, utilize a shaded structure to avoid direct sun during the deployment process.
5. Carefully scatter 'green gravel' from the surface onto the reef below or via SCUBA when trialing at new sites and at small scales.