A.1 Isolating the sori

Once you have the kelp containing the sori, you will work to isolate the tissue from the rest of the plant following the steps below.

Isolating the sorus tissue

• Work to minimize contamination and fouling of the sorus tissue

• Avoid using fouled tissue overgrown with epiphytes (throw it away)

• Wear gloves

• Work with your sorus tissue away from your culture area

• Identify and isolate sorus tissue

• Identify the sorus tissue on your blades

• Use a knife or razor blade to cut out the sorus from the rest of the blade

• Set tissue aside, keep cool and moist

• Remove excess fouling with a blade or scraper if materials permit. Throw away if excessively fouled
• Take care in the scraping as you may damage the sorus
• If no fouling, you may not need to scrape
• Clean tissue
• Remove kelp mucus and other debris by firmly wiping the tissue with a paper towel
• Wipe until dry, do not reuse paper towel between tissues
• Disinfect tissue
• Create a 3% iodine solution
• May be done in a beaker, bucket, or tub
• Carefully immerse the sorus tissue for ~30 seconds
• Use tweezers
• Avoid using other disinfectants (bleach, peroxide, alcohol)
• Rinse the tissue using chilled, filtered sea water
• Rinse until no color from the iodine remains
• Dry using a clean paper towel
• Rub gently and use a new piece of paper towel for each tissue
• Tissue storage
• Place tissue in dry paper towel (or newspaper), with multiple pieces on the top and bottom
• Do not let sorus pieces touch
• Refrigerate at 10°C for 12–24 hours (or what is optimal for the species)
• Ensure correct temperature is maintained
• If no refrigeration is available, store in a low light, cool, dry place

A.2 Spawning the sorus

While the sori are chilling, you can prepare the receptacle for the spore release. A beaker is a useful piece of equipment to use for creating the spore solution, but any clean container will work. It is also best practice to use chilled clean seawater (similar in temperature to the seawater the plant was sourced from). If you are creating a gametophyte suspension you can also add nutrients to promote spore growth. All temperatures and nutrient concentrations below are presented as general guidelines and may need to be modified for specific species. If you wish, you can use only chilled seawater.

Culturing beaker

• Create a mixture of (the ratio provided below is specific for Laminaria digitata and Saccharina latissima):
• 1000 mL chilled seawater (sea temperature)—autoclave to sterilize optional
• 9 mL PES—Provasoli’s Enriched Seawater (PES) culture nutrients—optional
• 0.8 mL germanium dioxide—optional for gametophytes
• 0.9 mL vitamins (optional)
• Adding sorus
• Add sori to the beaker
• Cut into smaller pieces if necessary
• Pieces should be entirely submerged
• Use multiple or larger beakers/containers to avoid over crowding
• Aim for > 20 pieces from different individuals to ensure genetic diversity
• Keep records of species used and trial conditions
• Occasionally stir the sori with a sterile stirrer or swirl the beaker
• Spawning
• You may be able to see a cloudy plume form in the beaker when spawning occurs (if not visible by eye, it is also possible to observe spawning by dropping 1-2 drops of the solution on a microscope slide and using microscope)
• This process may take over an hour
• Try to keep the temperature cool (somewhat species specific, but cooler is often better)
• Calculate stock density (optional)
• Acquire materials (Microscope, hemocytometer, pipettes, cleaning materials)
• See other materials for calculating stock density
• Aim to achieve stocking densities of > 100,000 spores/ mL

A.3 Creating a gametophyte solution

Below are the key points from the cultivation protocol created for farming Laminaria digitata in Ireland. As before, the specifics (temperatures, wavelengths, durations) are reference points and will likely change based on species and region.

A.3.1 Creation of gametophyte cultures

• Gametophyte cultures can be developed from the spore suspensions and maintained for extended periods to increase their volume through vegetative growth under red light and adequate nutrient and temperature conditions.
• To start the process, add further sterilised seawater to the spore suspension to fill the vessel and add sufficient nutrient medium for the volume of culture used.
• Disperse the nutrients throughout the culture by swirling the vessel.
• To enhance growth the developing culture can be aerated by inserting a pre-sterilised glass tube into the culture vessel connected to an air-supply via tubing and an air filter.
• The vegetative development of the gametophytes requires the installation of either single or double strip lighting, such as T5 Lindas, with fluorescent bulbs in a moisture-proof housing that emit relatively low levels of heat.
• The gametophytes need to be maintained at this life stage and this stasis can be achieved by exposing to them to red light or storing the male and female gametophytes separately (they need to be manually separated using tweezers and a dissecting scope). The light units need to be covered in red cellophane to create the required red-light conditions necessary to maintain propagules in their gametophyte stage and allow for vegetative growth. Red (660 nm) hydroponic grow lamps also work.
• The light intensity (PAR) needs to be measured with a meter calibrated in μmol m-2 s-1 aiming for a range of 15-20 μmol m-2 s-1 at the surface of the glassware.
• The light source needs to be fitted with a time switch set to long days; 16 to 24 hours of light per day are sufficient.
• The cultures should be kept at a constant temperature similar to natural seawater temperatures.

A.3.2 Maintenance of gametophyte cultures

• To increase the volume of the gametophyte cultures over the next 3 to 6 months, cultures need to be kept in motion via constant air supply and the nutrient medium needs to be exchanged every 10 to 14 days.
• To maintain the cultures without growth the medium can be changed every 2-3 months and aeration and motion is not required.
• Keep a constant temperature around ambient seawater for the maintenance of gametophyte cultures during vegetative growth.
• Note on contamination: In general, once a culture is contaminated it can take a lot of time to get it clean again. You should take a clean portion of the sample and move it into a new clean culture often. Ciliates and nematodes can be impossible to remove, and it is often better to throw away cultures where these are present.

A.3.3 Induction of reproduction

• Before the maintained gametophyte cultures can be used for seeding and development of sporophyte cultures, reproduction must be induced.
• Refresh the nutrient medium in the culture flasks to ensure that sufficient levels are available for sporophyte development.
• Cover the fluorescent lighting with blue cellophane to trigger sporophyte development.
• If measurable, optimal irradiance at the surface of the glassware is 15-20 μmol m-2 s-1 but values outside this range may also work.
• Set the time switch to equal light and dark periods, i.e., 12:12 hours of light to dark.
• Keep the temperature as before and provide continued aeration of the cultures.
• Maintain the culture flask(s) in these conditions until reproductive structures can be observed under a microscope.
• These will either be the developing unfertilized eggs still attached to the female gametophyte, or the fertilized egg/developing sporophyte.
• The reproductive state of the culture is assessed by following egg development, as it is much more difficult to observe the smaller male reproductive structures.
• Once many reproductive structures are observed you can prepare the culture to add to the substrate. Substrates may be seeded by mixing the gametophyte solution and the vector in the same container or by creating a spray to apply to the vector.

A.4 Further reading

¹ Flavin, K., Flavin, N., Flahive, B., 2013. Kelp Farming Manual: A Guide to the Processes, Techniques, and Equipment for Farming Kelp in New England Waters.

² Rolin, C., Inkster, R., Laing, J., Hedges, J., & McEvoy, L. (2016). Seaweed Cultivation Manual. Shetland Seaweed Growers Project 2014, 16.

³ Merrill, J.E., Gillingham, D.M., 1991. Bull kelp cultivation handbook. [National Coastal Resources Research and Development Institute], [Portland, Or.].