Method Article

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

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DOI:

10.3791/66092

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June 7th, 2024

 ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  , 
1Department of Ecology and Evolutionary Biology, University of California, Irvine, 2Department of Ecology and Evolutionary Biology, University of California, Santa Cruz, 3Facultad de Ciencias Marinas, Universidad Autónoma de Baja California, 4Facultad de Ciencias, Universidad Autónoma de Baja California, 5Instituto de Investigaciones Oceanológicas, Universidad Autónoma de Baja California, 6The Nature Conservancy

In This Article

Summary

This protocol describes the field collection and regular laboratory maintenance of substrates seeded with canopy-forming giant kelp for use in restoration trials to address the success and limitations of the 'green gravel' technique in field settings.

Abstract

Canopy-forming kelps are essential foundation species, supporting biodiversity and providing ecosystem services valued at more than USD$500 billion annually. The global decline of giant kelp forests due to climate-driven ecological stressors underscores the need for innovative restoration strategies. An emerging restoration technique known as 'green gravel' aims to seed young kelps over large areas without extensive underwater labor and represents a promising restoration tool due to cost-effectiveness and scalability. This video article illustrates a protocol and tools for culturing giant kelp, Macrocystis pyrifera. It also provides a resource for further studies to address the successes and limitations of this method in field settings. We outline field and laboratory-based methods for collecting reproductive tissue, sporulating, inoculating, rearing, maintaining, and monitoring substrates seeded with early life stages using the 'green gravel' technique. The protocol simplifies and centralizes current restoration practices in this field to support researchers, managers, and stakeholders in meeting kelp conservation objectives.

Introduction

Canopy-forming kelps (brown macroalgae in the order Laminariales) are globally important foundation species, dominating coastal rocky reefs in temperate and Arctic seas1. These kelps form structurally complex and highly productive biogenic habitats known as kelp forests that support taxonomically diverse marine communities2. Kelp forests worldwide provide many ecosystem services to humans, including commercial fisheries production, carbon and nutrient cycling, and recreational opportunities, with a total estimated value of USD $500 billion per year3.

Despite their substantial value, kelp forests face growing anthropogenic pressures in many regions3. Climate change presents one of the most significant threats to kelps due to long-term ocean warming combined with the increasing frequency of temperature anomalies3,4,5,6,7. Increased ocean temperatures are associated with nutrient limitation8, while exposure to heat stress above physiological thresholds can result in mortality9. In combination with variable regional local stressors7, kelp populations are globally declining by approximately 2% per year10 with significant losses and persistent shifts to alternate community states in certain regions6,11,12,13,14. Natural recovery of kelp populations alone may not be sufficient to reverse the extent of current and projected losses15,16,17,18, underscoring the importance of active restoration.

Current kelp restoration efforts can use a combination of methodologies to re-establish these important foundation species on coastal rocky reefs3,19. Methodologies chosen to address site-specific concerns depend on geographic context, the specific impediments to kelp recovery, and the social-ecological context11. Understanding the connections and interdependence of social-ecological systems is the key, and interventions that engage local institutions and garner support from local communities enhance the likelihood of successful restoration efforts20.

In addition to climate change, herbivore pressure or interspecific competition drives, declines, or suppresses the recovery (e.g., by sea urchins13, herbivorous fish21,22, turf algae9,23, or invasive algae24). Restoration may focus on the removal of these biotic stressors25, although these methods require substantial resources and continuous maintenance11. To catalyze kelp species recovery, there have been efforts toward a direct seeding approach, for example, weighing mesh bags filled with fertile kelp blades to the benthos that release zoospores into the environment26. This method, however, is time-intensive and requires technical underwater installation and removal. Other cases focus on transplanting large quantities of whole adult donor plants, which may compromise closely associated and vulnerable donor populations and are often limited to small scales due to reliance on continual transplantation27.

For regions, where kelp spore limitation may be impeding kelp forest recovery due to habitat fragmentation, a relatively new kelp restoration approach called the 'green gravel' technique, was introduced. The technique was successfully trialed at theFlødevigen Research Station in southern Norway28 and represented a promising option for restoration due to cost-effectiveness and scalability. The workflow of this technique is as follows: (1) a spore solution is created from fertile tissue collected from reproductive adult kelps in the field and then seeded onto small substrates, such as gravel; (2) early-stage kelps are reared in laboratory-controlled abiotic conditions on substrates; (3) substrates with visible sporophytes are deployed in the field on specific reefs as 'green gravel', where sporophytes continue to grow. Note that typical transplantation efforts of adult individuals require laborious and cost-inhibitive underwater installation by divers, and the 'green gravel' technique uses simple deployment from the surface28.

The 'green gravel' technique is currently being tried out by members of numerous international working groups29 across different environments and several laminarian kelp species. This protocol describes the required facilities, materials, and methods for tissue collection, sporulation, seeding, rearing conditions, regular maintenance, and monitoring of early-stage kelp prior to deploying this restoration technique in the field using the giant kelp, Macrocystis pyrifera. This protocol is a valuable resource for researchers, managers, and stakeholders seeking to provide insight into the successes and limitations of this method with M. pyrifera in different field settings.

Protocol

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 rugosity32,33. 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 proximity38,39. 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.

Results

The 'green gravel' restoration technique is still in the piloting phase, with limited outplant survival data for other species28, and no published data yet for Macrocystis pyrifera. Using the field collection and laboratory maintenance outlined in this protocol, we tested the significance of site-specific rearing conditions for two distinct donor kelp populations before hypothetical 'green gravel' deployment (Figure 5). Reproductive kelp tissue was collected in California (USA) from cooler K1 (Santa Cruz 36.60167°N, 121.88508°W) and warmer K4 (San Diego, 32.85036°N, -117.27600°W) populations and reared at two temperatures: (1) 12 °C (the standard culturing temperature for seaweed aquaculture, and the mean winter SST for K1), and (2) 20 °C (the mean summer SST for K4, and a 4 °C heatwave for K1). All glass slides used for monitoring kelp life stage development were marked with a standardized grid, and high-resolution images were captured using this grid as a reference to enable the observation of fixed fields through time using an inverted microscope and camera (N = 5 images per sample, 2.479 mm x 1.859 mm).

After 24 d post sporulation, gametophytes were counted from microscope images (N = 300 images from 60 samples). To test for differences in gametophyte counts, generalized linear mixed effects models were employed with Poisson distribution using the function glmmTMB() in package glmmTMB41, and pairwise comparisons were conducted with emtrends() from package emmeans42in R. Our results illustrate that the response of gametophytes to thermal variability was different between K1 and K4 populations (t = 2.7, p = 0.007), where temperature did not have an effect for the warmer K4 population (estimate = -0.01, standard error [SE] = 0.01, confidence interval [CI] = [-0.03, 0.01]), but did have an effect for the cooler K1 population (estimate = -0.06, SE = 0.02, CI = [-0.10, -0.03]) (Figure 6A), suggesting a possible adaptive divergence in thermal tolerance traits. Kelp gametophytes are often depicted as a resistance stage43, meaning they produce an all-purpose phenotype that is stress tolerant and relatively insensitive to environmental variability. However, these results indicate that thermal variability imposes a significant pressure at this early stage.

After 32 d post sporulation, visible sporophytes with lengths greater than approximately 1 mm were counted on the entirety of each 2.5 cm by 7.5 cm glass slide (N = 72 total samples). To test for differences in visible sporophyte counts, generalized linear mixed effects models were employed with Poisson distribution using the function glmmTMB() in package glmmTMB and pairwise comparisons were conducted with emtrends() from package emmeans in R. Our results illustrate that the response of sporophytes to thermal variability is similar between K1 and K4 differentiated populations (z = 0.92, p = 0.36), where the temperature had an effect for the warmer K4 population (estimate = -0.66, SE = 0.04, CI = [-0.74, - 0.57]), as well as the cooler K1 population (estimate = -0.85, SE = 0.13, CI = [-1.10, -0.60]) (Figure 6B). Samples reared at 20 °C grew few visible sporophytes (mean ± SE = 0.4 ± 0.2) compared to those reared at 12 °C (mean ± SE = 82.4 ± 9.8). This result suggests that sporophyte production is more sensitive to temperature than the gametophyte stage, and that site-specific culturing temperatures must not exceed 15 °C to achieve sporophyte development as outlined in the protocol.

Controlled environment setup with jars for gas exchange analysis in plant respiration experiment.
Figure 1: Diagram of 'green gravel' incubator system. (A) Red light source for vegetatively bulking gametophyte cultures. (B) Access port for electrical wires and tubing, leading to an external outlet. (C) Structure to block full-spectrum light out of the red-light section. (D) A 'green gravel' culturing section. (E) full-spectrum light sources. (F) Tubing lines connected to an external filtered air source. (G) Check valves to reduce airborne contamination. (H) Individual culture containers that minimize contamination. Please click here to view a larger version of this figure.

Solar water heating system setup diagram; solar panels, water tanks, fluid circulation process.
Figure 2: Diagram of 'green gravel' water bath system. (A) Chiller with submerged pump (in I). (B) 20 gallon tub for water bath. (C) Drain to recirculate water bath. (D) Valve for recirculation of water bath. (E) Light source. (F) 2.5 L 'green gravel' container with transparent lid and aeration opening. (G) Aeration source. (H) Pipes that recirculate water with use of submerged pumps. (I) Water bath receiver from/to chiller from/to tubs with submersible pumps. (J) Acrylic cover to minimize water bath evaporation. (K) Mesh shade to adjust light intensity. Please click here to view a larger version of this figure.

Life cycle of gametophyte and sporophyte stages; microscopy diagram shows development times.
Figure 3: Macrocystis pyrifera development. Developmental life history stages of Macrocystis pyrifera from laboratory growth trials. Please click here to view a larger version of this figure.

Marine growth progression; field deployment timeline in a laboratory to sea environment; 120 days.
Figure 4: 'Green gravel' seeded with Macrocystis pyrifera. 'Green gravel' seeded with Macrocystis pyrifera is cultured in the laboratory until sporophytes reach 1-2 cm. 'Green gravel' is then deployed and continues to grow in the field. Please click here to view a larger version of this figure.

Microalgae growth over 40 days at 12°C, 20°C in diagram, showing temperature effect on biomass.
Figure 5: Experimental time series. Example images from a time series following the experimental growth and development of Macrocystis pyrifera gametophytes and sporophytes originating from two populations collected in California (USA) and cultured at two different temperatures. K1 = Santa Cruz, K4 = San Diego. Please click here to view a larger version of this figure.

Bar chart comparing gametophyte and sporophyte density at 12°C and 20°C for K4 and K1 types.
Figure 6: Representative Results. Macrocystis pyrifera life stages observed for K1 Santa Cruz San Diego K4 Santa Cruz populations of origin cultured at constant thermal conditions of 12 and 20 °C. Error bars, mean ± 1 SE. Asterisk (*) denotes statistically significant differences (p < 0.05). (A) Gametophytes at day 24 (N = 300 total images from 60 samples). (B) Visible sporophytes at day 32 (N = 72 samples, within a standardized 2.5 cm by 7.5 cm area). Please click here to view a larger version of this figure.

Supplemental File 1. Please click here to download this File.

Discussion

Anthropogenic climate change is a growing threat to the health of the world's oceans44,45,46,47,48, resulting in major disturbances and biodiversity loss49,50,51,52. To accelerate the restoration of degraded ecosystems, the United Nations has declared 2021 through 2030 the "UN Decade on Ecosystem Restoration," coinciding with the "UN Decade of Ocean Science for Sustainable Development", which aims to reverse the deterioration in ocean health53. In line with this global call to action, the Kelp Forest Alliance has launched the Kelp Forest Challenge to restore 1 million hectares and protect 3 million hectares of kelp forest by the year 204054. Marine restoration is undervalued55, and kelp ecosystems receive considerably less attention than habitats such as coral reefs, mangrove forests, and seagrass meadows56. Restoration of degraded ecosystems has been shown to be effective in re-building marine ecosystems but can cost on average between $80,000 - $1,600,000 per hectare, with median total costs likely to be two to four times higher57. Current and projected losses call upon developing scalable, feasible, and cost-effective kelp restoration methodologies as urgent conservation interventions.

Current kelp restoration efforts use a combination of methodologies to address site-specific drivers of kelp loss, including transplantation of adult kelps, direct seeding of zoospores and/or gametophytes, grazer control, and installation of artificial reefs11. However, these methods require substantial resources and have limited scalability. Typical transplantation of adult kelps requires the laborious deployment of artificial materials or structures on the benthos, by divers. Bottom-up interventions to re-establish coastal rocky reefs, such as controlling competitors and grazers, are also restricted by labor costs as they rely on the manual underwater removal or exclusion of these biotic stressors11. The 'green gravel' technique overcomes these limitations with simple deployment from the surface, requiring no underwater installment or technical knowledge and scalability at relatively low costs28. This innovative approach provides a promising restoration tool, urging extensive trials across diverse locations and environments to unlock its full potential32.

While successful restoration efforts with 'green gravel' have been documented in sheltered fjords in Norway using the sugar kelp, Saccharina latissima26, this technique is still in the piloting phase for Macrocystis pyrifera in the Eastern Pacific. Additional trials are needed to address the expected survivorship of M. pyrifera outplants within its range. In wave-exposed conditions typical of M. pyrifera growth, smaller gravel may be more prone to movement and abrasion, leading to damaged outplants. Furthermore, positive buoyancy provided by gas-filled pneumatocysts of M. pyrifera may lead to 'green gravel' outplants being effectively carried away from the restoration site, and thus, gravel size and weight are important factors to explore for this species. In a recent pilot study (May 2022; Ensenada, Baja California, Mexico), preliminary success in the field with M. pyrifera has been observed, indicated by haptera attachment to surrounding substrate and growth of juveniles reaching 1.2 m in length after two months in the field (Figure 4). This demonstrates a clear opportunity that has yet to be explored in utilizing 'green gravel' for M. pyrifera in the Eastern Pacific. This video showcases the 'green gravel' technique with M. pyrifera and is a valuable resource that simplifies and centralizes existing practices in the culturing phase of restoration to support studies that address successes and limitations in different field settings.

With the 'green gravel' technique, many smaller, individual gravel units can be seeded at a scale that may increase the probability of success compared to more common transplantation approaches with adult plants. However, the key scalable aspect of this technique is its simple deployment from the surface, which can facilitate the restoration of large areas by boat. For field settings where the deployment of small gravel is not suitable, this protocol can be adapted to transplant M. pyrifera on a wide range of substrates, including larger gravel or even small boulders, string that can be tied to natural or deployed underwater anchors, or tiles that can be bolted or glued using marine epoxy to the seafloor in more exposed conditions. These deployment adaptations will not change the facilities needed for M. pyrifera culturing but will subsequently increase the cost of deployment.

Anthropogenic disturbances and climate change are currently overcoming the capacity for natural populations to adapt. This poses significant challenges to traditional conservation efforts that restore ecosystems to their historic states58,59,60,61,62,63. Thus, conservation frameworks have expanded to include anticipatory management considering resilience and adaptive capacity64. Anticipatory management to address climate change is being implemented for tree species in forest ecosystems65 and has been proposed for further restoration efforts to enhance the evolutionary potential of outplants66,67. Although these strategies are inherently easier to manipulate in terrestrial environments, several studies are beginning to explore their application in marine environments62,68,69,70. For example, coral reefs are threatened by numerous anthropogenic stressors that have resulted in unprecedented declines71,72. In response to the losses of these important foundation species, active restoration and assisted adaptation techniques are increasingly advocated to conserve remaining coral reefs and their associated functions62,73,74. One technique involves translocating individuals within their current species distribution range to increase tolerance to heat stress75. Regarding the restoration of canopy-forming kelps, 'green gravel' has a customizable framework to explore assisted adaptation techniques such as translocation of resilient genotypes to vulnerable areas, non-genetic manipulation such as hybridization, or acclimatization of individuals to environmental stress62 with outcomes aimed towards obtaining more resistant strains for restoration programs76,77.

Harnessing local support to enhance restoration endeavors is crucial to sustain kelp ecosystem conservation success. Engaging local stakeholders can increase local buy-in for restoration needs6,50 and promote coastal stewardship that could subsequently result in increased funding and longevity of kelp ecosystem protection. As with all other kelp restoration methodologies, structured decision-making frameworks integrating diverse ecological, socio-economic, and conservation objectives will help achieve optimal outcomes for kelp ecosystems and the communities they support11.

Disclosures

The authors have nothing to disclose.

Acknowledgements

This work was funded by the California Sea Grant Kelp Recovery Research Program R/HCE-17 to JBL and MESB, a National Science Foundation Research Traineeship award DGE-1735040 to PDD, The Nature Conservancy, Schmidt Marine Technology Partners, Sustainable Ocean Alliance, Tinker Foundation to AP-L, and The Climate Science Alliance Baja Working Group to RBL and JL. We thank Steven Allison, Cascade Sorte, Samantha Cunningham, Sam Weber and Caitlin Yee at the University of California, Irvine; Mark Carr, Peter Raimondi, Sarah Eminhizer, Anne Kapuscinski at the University of California, Santa Cruz; Walter Heady and Norah Eddy at The Nature Conservancy; Filipe Alberto and Gabriel Montecinos at the University of Wisconsin, Milwaukee; Jose Antonio Zertuche-González, Alejandra Ferreira-Arrieta, and Liliana Ferreira-Arrieta at the Universidad Autónoma de Baja California; Luis Malpica-Cruz, Alicia Abadía-Cardoso, and Daniel Díaz-Guzmán from MexCal; the MexCalitos divers Alejandra Reyes, Monica Peralta, Teresa Tavera, Julia Navarrete, Ainoa Vilalta, Jeremie Bauer, and Alfonso Ferreira; and Nancy Caruso for technical advice. We thank the Instituto de Investigaciones Oceanológicas, Universidad Autónoma de Baja California for providing facilities used to develop the water bath system. We thank Ira Spitzer for underwater and drone video content.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
0.22 µm filtersMiliporeSCGPS05RENatural seawater sterilization
1 L glass bottlesAmazonB07J6JP4D1Natural seawater sterilization
1 µm filters (water + air)AmazonB01M1VWUWLNatural seawater sterilization
1'' PVC 90-Degree ElbowHome Depot203812125Option 2 - Medium scale - Water bath systems
10 µm filtersAmazonB00D04BG56Natural seawater sterilization
20 µm filtersAmazonB082WS9NPHNatural seawater sterilization
3x5mm tubingAmazonB0852HXPN6Option 1 Small scale - Incubator
4x4'' Sterile GauzeAmazonB07NDK8XM3Sporulation
4x6mm tubingAmazonB08BCRV1FYOption 1 Small scale - Incubator
5 µm filtersAmazonB082WS9NPHNatural seawater sterilization
50 mL falcon tubingAmazonB01M04HGPJSporulation
8x10mm tubingAmazonB01MSM3LLZOption 1 Small scale - Incubator
Air filtersThermo FisherMTGR85010Option 1 Small scale - Incubator
Alcohol lampAmazonB07XWD9WWCSporulation
Ammonium iron(II) sulfate hexahydrate ACS reagent, 99%Sigma215406-100GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Aquarium Grade GravelAmazonB07XRCKFBJOption 1 Small scale - Incubator
Biotin powder, BioReagent, suitable for cell culture, suitable for insect cell culture, suitable for plant cell culture, 99%SigmaB4639-100MGChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Boric Acid, 99.8%, 10043-35-3, MFCD00011337, BH3O3, 61.83, 500gThermo Fisher5090113707Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Calcium D-Pantothenate,ge98.0% (T),C9H17NO5,137-08-6,25g,D-Pantothenic Acid Calcium Salt, P0012-25G 1/EAThermo FisherP001225GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Check valvesAmazonB08HRZR4MMOption 1 Small scale - Incubator
Clear tubing 3/8'' - 10 ftAmazonB07MTYMW13Option 2 - Medium scale - Water bath systems
COBALT(II) SULFATE HEPTAH-100G, WARNING - California - Cancer Hazard, 93-2749-100G 1/EAThermo Fisher5090114752Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Compound microscope with cameraOMAXM83EZ-C50SMonitoring
Culture flaskThermo Fisher07-250-080Option 1 Small scale - Incubator
Culture lightAmazonB07RRRPJ63Option 1 Small scale - Incubator
Culture stoppersAmazonB07DX6J7QDOption 1 Small scale - Incubator
Drainage connectorAmazonB00GUZ6CV0Option 2 - Medium scale - Water bath systems
EDTA CAS Number: 6381-92-6 Molecular Formula: C10H14N2O8Na2- 2H2O Molecular Weight: 372.2Thermo Fisher50213299Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Eisco Safety Pack Graduated Cylinder Sets Class A, ASTM, Capacity: 10 mL, 25 mL, 50 mL, Graduations: 0.2 mL, 0.5 mL, 1.0 mL, Borosilicate 3.3 Glass, Autoclavable: Yes, Class: Class A, Graduated: Yes, Tolerance: 0.10 mL, 0.17 mL, 0.25 mLThermo FisherS81273Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Eisco Safety Pack Graduated Cylinder Sets Class A, ASTM, Capacity: 50 mL, 100 mL, 250 mL, Graduations: 1.0 mL, 1.0 mL, 2.0 mL, Borosilicate 3.3 Glass, Autoclavable: Yes, Class: Class A, Graduated: Yes, Tolerance: 0.25 mL, 0.50 mL, 1.0 mLThermo FisherS81275Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Eisco Safety Pack Volumetric Flask Sets - Class A, ASTM, Capacity: 10 mL, 25 mL, 50 mL, Borosilicate 3.3 Glass, Autoclavable: Yes, Class: Class A, Closure Material: Glass, Closure Size: Stopper Number: 9, 9, 13, Closure Type: Penny Stopper, Graduated: YeThermo FisherS81271Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Filter holderAmazonB07LCKBKCTNatural seawater sterilization
Fisherbrand Graduated Cylinders, Capacity: 500 mL, Graduations: 5 mL, Borosilicate Glass, Autoclavable: Yes, Limit of Error: +/-4.0 mL, Recommended Applications: Education, Subdivision: 5 mL, S63460 1/EAThermo FisherS63460Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
FLEXACAM C1 CameraLeicaFLEXACAM C1Monitoring
Folic acid, C19H19N7O6, CAS Number59303, vitamin m, pteroylglutamic acid, vitamin b9, folvite, folacin, folacid, pteroyllglutamic acid, pteglu, folic acid, folate, 25g, 100781, CHEBI:27470, Yellow to Orange, 2004190, 441.41, OVBPIULPVIDEAOLBPRGKRZSANThermo FisherAAJ6083314Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Free Standing 20 Gallon Utility SinkAmazonB094TLH19LOption 2 - Medium scale - Water bath systems
GERMANIUM DIOXIDE 99.99 10GRThermo FisherAC190000100Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Glass Graduated Cylinders, Class A Round Base, Eisco, For Use With: Measuring liquids, Capacity: 1000 mL, Graduations: 10 mL White, CH0344OWT 1/EAThermo FisherS88442Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Glass slidesAmazonB00L1S93PSOption 2 - Medium scale - Water bath systems
Glycerol phosphate disodium salt hydrate isomeric mixtureSigmaG6501-100GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Growth containers -3.4 Qt- 3.25 Lt transparent containers with transparent lidContainer store#10014828Option 2 - Medium scale - Water bath systems
Growth lightAmazonB086R14MFWOption 1 Small scale - Incubator
HemocytometerAmazonB07TJQDKLJSporulation
Hinged plastic jarsSKS Bottle & Packaging40280125.01SOption 1 Small scale - Incubator
Inositol research grade, USP/NF For bacteriology. Optically inactive. Tested for its suitability in tissue culture. Size - 100G Storage Conditions - +15 C TO +30 C Catalog Number - 26310.01 CAS 87-89-8Thermo Fisher50247745Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Instant Ocean - 50 GAmazonB000255NKAOption 1 Small scale - Incubator
Inverted Microscope Leica DMi1LeicaDMi1Monitoring
Iron(III) chloride hexahydrate ACS reagent, 97%Sigma236489-100GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Licor Ligth Meter Data LoggerLicorLI-250AMonitoring
Light/temperature HOBO data loggerAmazonB075X2SWKNMonitoring
Lights 150WAmazonB0799DQM9VOption 2 - Medium scale - Water bath systems
Manganese sulfate monohydrate meets USP testing specificationsSigmaM8179-100GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Medium size rocks 2-3 inch, 20 poundsHome Depot206823930Option 2 - Medium scale - Water bath systems
Nicotinic Acid, 99%, C6H5NO2, CAS Number59676, daskil, apelagrin, acidum nicotinicum, akotin, 3carboxypyridine, niacin, 3pyridinecarboxylic acid, nicotinic acid, pellagrin, wampocap, 250g, 109591, CHEBI:15940, 1.4, 2004410, 293 deg.C (559 deg.F), 123.11,Thermo FisherAAA1268330Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
p-Aminobenzoic acid 99.82% 4-aminobenzoic acid, C7H7NO2, CAS Number: 150-13-0, 25g, 0210256925 1/EAThermo FisherICN10256925Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
PCV cementAmazonB001D9WRWGOption 2 - Medium scale - Water bath systems
Plastic water valveAmazonB0006JLVE4Option 2 - Medium scale - Water bath systems
Plastic water valveAmazonB07G5FY7X1Option 2 - Medium scale - Water bath systems
Precision scale 1mgAmazonB08DTH95FNMaterials to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Pump for filtered airAmazonB0BG2BT9RXOption 1 Small scale - Incubator
PVC tubing 1x24''Home Depot202300505Option 2 - Medium scale - Water bath systems
Quantum Light meterApogee InstrumentsMQ-510Monitoring
Refrigerated IncubatorThermo Fisher15-103-1566Option 1 Small scale - Incubator
Rubber GrommetsAmazonB07YZD22ZPOption 1 Small scale - Incubator
Salinity refractometerATCB018LRO1SUMonitoring
Shade mesh 6x50 ftHome depot316308418Option 2 - Medium scale - Water bath systems
Sodium Nitrate ge 99.0% Nitric Acid, Sodium Salt, NNaO3, CAS Number: 7631-99-4, 500g, 1/EAThermo FisherBP360500Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Soldering for aeration openingAmazonB08R3515SFOption 2 - Medium scale - Water bath systems
Spray isporopyl alcoholAmazon‎ B08LW5P844Sporulation
Stainless steel sissorsAmazonB07BT4YLHTSporulation
Stainless steel trayAmazonB08CV33YXGSporulation
Stainless steel twizzersAmazonB01JTZTAJSSporulation
Stir BarsAmazonB07C4TNKXBMaterials to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Submersible circulation pump 400 GPHAmazonB07RZKRM13Option 2 - Medium scale - Water bath systems
Submersible Spherical Quantum SensorWaltzUS-SQS/LMonitoring
Temperature gunInfrared Thermometer 749B07VTPJXH9Monitoring
Thiamine hydrochloride BioReagent, suitable for cell culture, suitable for insect cell culture, suitable for plant cell cultureSigmaT1270-25GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Thymine 99% 2, 4-Dihydroxy-5-methylpyrimidine, C5H6N2O2, CAS Number: 65-71-4, 25g, 157850250 1/EAThermo FisherAC157850250Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Transparent Acrylic sheet 24x48 inchHome Depot202038048Option 2 - Medium scale - Water bath systems
Tubing water circulation 1''x10 ftAmazonB07ZC1PSF3Option 2 - Medium scale - Water bath systems
UV light for natural seawater sterilizationAmazonB018OI7PYSNatural seawater sterilization
Vacum pumpAmazonB087XBTPVWNatural seawater sterilization
Vitamin B12 BioReagent, suitable for cell culture, suitable for insect cell culture, suitable for plant cell culture, 98%SigmaV6629-100MGChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Volumetric Flasks, Class A Glass, Eisco, with Polypropylene Stopper, Graduated, White printed markings, Capacity: 1000 mL, CH0446IWT 1/EAThermo FisherS89446Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Volumetric Flasks, Class A Glass, Eisco, with Polypropylene Stopper, Graduated, White printed markings, Capacity: 500 mL, CH0446HWT 1/EAThermo FisherS89445Chemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment
Water Chiller 200-600GPMAmazonB07BHHP71COption 2 - Medium scale - Water bath systems
Y-splitters for 4x6mm tubingAmazonB08XTJKFCHOption 1 Small scale - Incubator
Zinc sulfate heptahydrate BioReagent, suitable for cell cultureSigmaZ0251-100GChemicals to create Provasoli’s Enriched Seawater (PES) and vitamins for media enrichment

References

  1. Schiel, D. R., Foster, M. S. The Biology and Ecology of Giant Kelp Forests. , Univ of California Press. (2015).
  2. Smale, D. A., Burrows, M. T., Moore, P., O'Connor, N., Hawkins, S. J. Threats and knowledge gaps for ecosystem services provided by kelp forests: a northeast Atlantic perspective. Ecol Evol. 3 (11), 4016-4038 (2013).
  3. Eger, A. M., et al. The value of ecosystem services in global marine kelp forests. Nat Comm. 14 (1), 1894(2023).
  4. Bennett, S., Wernberg, T., Arackal Joy, B., de Bettignies, T., Campbell, A. H. Central and rear-edge populations can be equally vulnerable to warming. Nat Comm. 6 (1), 10280(2015).
  5. Jueterbock, A., et al. Climate change impact on seaweed meadow distribution in the North Atlantic rocky intertidal. Ecol Evol. 3 (5), 1356-1373 (2013).
  6. Rogers-Bennett, L., Catton, C. A. Marine heat wave and multiple stressors tip bull kelp forest to sea urchin barrens. Sci Rep. 9 (1), 15050(2019).
  7. Krumhansl, K. A., et al. Global patterns of kelp forest change over the past half-century. PNAS. 113 (48), 13785-13790 (2016).
  8. Zimmerman, R. C., Kremer, J. N. Episodic nutrient supply to a kelp forest ecosystem in Southern California. J Mar Res. 42 (3), 591-604 (1984).
  9. Rothäusler, E., et al. Physiological performance of floating giant kelp Macrocystis pyrifera (phaeophyceae): Latitudinal variability in the effects of temperature and grazing. J Phycol. 47 (2), 269-281 (2011).
  10. Wernberg, T., Krumhansl, K., Filbee-Dexter, K., Pedersen, M. F. Chapter 3 - Status and trends for the world's kelp forests. World Seas: An Environmental Evaluation (Second Edition). , 57-78 (2019).
  11. Eger, A. M., Layton, C., McHugh, T. A., Gleason, M., Eddy, N. Kelp restoration guidebook: Lessons learned from kelp restoration projects around the world. TNCKelp Forest Alliance. , (2022).
  12. Filbee-Dexter, K., et al. Marine heatwaves and the collapse of marginal North Atlantic kelp forests. SciRep. 10 (1), 13388(2020).
  13. Filbee-Dexter, K., Scheibling, R. E. Sea urchin barrens as alternative stable states of collapsed kelp ecosystems. Mar Ecol Prog Ser. 495, 1-25 (2014).
  14. Filbee-Dexter, K., Wernberg, T. Rise of turfs: A new battlefront for globally declining kelp forests. BioSci. 68 (2), 64-76 (2018).
  15. Assis, J., Araújo, M. B., Serrão, E. A. Projected climate changes threaten ancient refugia of kelp forests in the North Atlantic. Glob Change Biol. 24 (1), e55-e66 (2018).
  16. Davis, T., Champion, C., Coleman, M. Ecological interactions mediate projected loss of kelp biomass under climate change. Divers Distrib. 28 (2), 306-317 (2021).
  17. Goldsmit, J., et al. Kelp in the eastern Canadian arctic: Current and future predictions of habitat suitability and cover. Front Mar Sci. 18, 742209(2021).
  18. Ling, S. D., Cornwall, C. E., Tilbrook, B., Hurd, C. L. Remnant kelp bed refugia and future phase-shifts under ocean acidification. PLoS One. 15 (10), e239136(2020).
  19. Eger, A. M., et al. Global kelp forest restoration: past lessons, present status, and future directions. Biol Rev. 97 (4), 1449-1475 (2022).
  20. Waylen, K. A., Fischer, A., McGowan, P. J., Thirgood, S. J., Milner-Gulland, E. J. Effect of local cultural context on the success of community-based conservation interventions. Biol Consv. 24 (4), 1119-1129 (2010).
  21. Vergés, A., et al. The tropicalization of temperate marine ecosystems: climate-mediated changes in herbivory and community phase shifts. Proc Royal Soc. B. 281 (1789), 20140846(2014).
  22. Zarco-Perello, S., Wernberg, T., Langlois, T. J., Vanderklift, M. A. Tropicalization strengthens consumer pressure on habitat-forming seaweeds. Sci Rep. 7 (1), 820(2017).
  23. Worm, B., Lotze, H. K. Chapter 21 - Marine biodiversity and climate change. Climate Change (Third Edition). , 445-464 (2021).
  24. Félix-Loaiza, A. C., Rodríguez-Bravo, L. M., Beas-Luna, R., Lorda, J., de La Cruz-González, E., Malpica-Cruz, L. Marine heatwaves facilitate invasive algae takeover as foundational kelp. Botanica Marina. 65 (5), 315-319 (2022).
  25. Miller, K. I., Blain, C. O., Shears, N. T. Sea urchin removal as a tool for macroalgal restoration: A review on removing "the spiny enemies". Fron Mar Sci. 9, 831001(2022).
  26. Westermeier, R., et al. Repopulation techniques for Macrocystis integrifolia (Phaeophyceae: Laminariales) in Atacama, Chile. J Appl Phycol. 26, 511-518 (2014).
  27. Layton, C., et al. Kelp forest restoration in Australia. Fron Mar Sci. 7, 74(2020).
  28. Fredriksen, S., et al. Green gravel: a novel restoration tool to combat kelp forest decline. Sci Rep. 10 (1), 3983(2020).
  29. Projects of the Green Gravel Action Group. Green Gravel Action Group. (n.d.) . , Available from: https://www.greengravel.org/ (2024).
  30. Fain, S. R., Murray, S. N. Effects of light and temperature on net photosynthesis and dark respiration of gametophytes and embryonic sporophytes of macrocystis pyrifera. J Phycol. 18 (1), 92-98 (1982).
  31. Westermeier, R., Patiño, D., Piel, M. I., Maier, I., Mueller, D. G. A new approach to kelp mariculture in Chile: production of free-floating sporophyte seedlings from gametophyte cultures of Lessonia trabeculata and Macrocystis pyrifera. Aquac Res. 37 (2), 164-171 (2006).
  32. Alsuwaiyan, N., et al. A et al. Green gravel as a vector of dispersal for kelp restoration. Fron Mar Sci. 9, 910417(2022).
  33. Falace, A., Kaleb, S., De La Fuente, G., Asnaghi, V., Chiantore, M. Ex situ cultivation protocol for Cystoseira amentacea var. stricta (Fucales, Phaeophyceae) from a restoration perspective. PloS One. 13 (2), e0193011(2018).
  34. Redmond, S., Green, L., Yarish, C., Kim, J., Neefus, C. New England seaweed culture handbook. , (2014).
  35. Provasoli, L., McLaughlin, J. J. A., Droop, M. R. The development of artificial media for marine algae. Arch Mikrobiol. 25, 392-428 (1957).
  36. Navarro, D., Navarro, D. E. California Kelp Forest Restoration: Science Activity Guide for Teachers. , (2006).
  37. Alsuwaiyan, N. A., et al. A review of protocols for the experimental release of kelp (Laminariales) zoospores. Ecol Evol. 9 (14), 8387-8398 (2019).
  38. Lüning, K., Müller, D. G. Chemical interaction in sexual reproduction of several Laminariales (Phaeophyceae): release and attraction of spermatozoids. Z. Pflanzenphysiol. 89 (4), 333-341 (1978).
  39. Müller, D. G., Maier, I., Gassmann, G. Survey on sexual pheromone specificity in Laminariales (Phaeophyceae). Phycologia. 24 (4), 475-477 (1985).
  40. Vieira, V. M., Oppliger, L. V., Engelen, A. H., Correa, J. A. A new method to quantify and compare the multiple components of fitness-a study case with kelp niche partition by divergent microstage adaptations to temperature. Plos One. 10 (3), e0119670(2015).
  41. Brooks, M. E., et al. glmmTMB balances speed and flexibility among packages for zero-inflated generalized linear mixed modeling. The R Journal. 9 (2), 378-400 (2017).
  42. Russell, L. emmeans: estimated marginal means, aka least-squares means. R package version. 1 (2), (2018).
  43. Ladah, L. B., Zertuche-González, J. A. Survival of microscopic stages of a perennial kelp (Macrocystis pyrifera) from the center and the southern extreme of its range in the Northern Hemisphere after exposure to simulated El Niño stress. Mar Biol. 152, 677-686 (2007).
  44. Halpern, B. S., et al. A global map of human impact on marine ecosystems. Science. 319 (5865), 948-952 (2008).
  45. Halpern, B. S., et al. Spatial and temporal changes in cumulative human impacts on the world's ocean. Nat Comm. 6 (1), 1-7 (2015).
  46. Halpern, B. S., et al. Recent pace of change in human impact on the world's ocean. Sci Rep. 9 (1), 11609(2019).
  47. Micheli, F., et al. Cumulative human impacts on Mediterranean and Black Sea marine ecosystems: assessing current pressures and opportunities. PloS One. 8 (12), e79889(2013).
  48. Portner, H. -O., et al. IPCC, 2022: Summary for policymakers. IPCC. , (2022).
  49. Butchart, S. H. M., et al. Global biodiversity: Indicators of recent declines. Science. 328 (5982), 1164-1168 (2010).
  50. Rocha, J., Yletyinen, J., Biggs, R., Blenckner, T., Peterson, G. Marine regime shifts: Drivers and impacts on ecosystems services. Phil Trans Roy Soc. B. 370 (1659), 20130273(2015).
  51. Worm, B., et al. Impacts of biodiversity loss on ocean ecosystem services. Science. 314 (5800), 787-790 (2006).
  52. Worm, B., Lotze, H. K. Marine biodiversity and climate change. Climate Change (Third Edition). , Chapter 21 445-464 (2021).
  53. Waltham, N. J., et al. UN decade on ecosystem restoration 2021-2030-What chance for success in restoring coastal ecosystems. Fron Mar Sci. 7, (2020).
  54. Kelp Forest Challenge. Kelp Forest Alliance. , Available from: https://kelpforestalliance.com/ (2024).
  55. Gordon, T. A. C., Radford, A., Simpson, S. D., Meekan, M. G. Marine restoration projects are undervalued. Science. 367 (6478), 635-636 (2020).
  56. Morris, R. L., et al. Key principles for managing recovery of kelp forests through restoration. BioScience. 70 (8), 688-698 (2020).
  57. Bayraktarov, E., et al. The cost and feasibility of marine coastal restoration. Ecol Appl. 26 (4), 1055-1074 (2016).
  58. Breed, M. F., et al. Priority actions to improve provenance decision-making. BioScience. 68 (7), 510-516 (2018).
  59. Breed, M. F., et al. The potential of genomics for restoring ecosystems and biodiversity. Nat Rev Genet. 20 (10), 615-628 (2019).
  60. Gurgel, C. F. D., Camacho, O., Minne, A. J. P., Wernberg, T., Coleman, M. A. Marine heatwave drives cryptic loss of genetic diversity in underwater forests. Curr Biol. 30 (7), 1199-1206 (2020).
  61. Hobbs, R. J., Higgs, E., Harris, J. A. Novel ecosystems: implications for conservation and restoration. Trends Ecol Evol. 24 (11), 599-605 (2009).
  62. van Oppen, M. J. H., Oliver, J. K., Putnam, H. M., Gates, R. D. Building coral reef resilience through assisted evolution. PNAS. 112 (8), 2307-2313 (2015).
  63. Perring, M. P., et al. Advances in restoration ecology: Rising to the challenges of the coming decades. Ecosphere. 6 (8), (2015).
  64. Coleman, M. A., et al. Restore or redefine: Future Trajectories for Restoration. Fron MarSci. 7, 237(2020).
  65. O'Neill, G. A. Assisted migration to address climate change in British Columbia: recommendations for interim seed transfer standards. Ministry of Forests and Range, Forest Science Program. , Victoria, B.C. (2008).
  66. Broadhurst, L. M., et al. Seed supply for broadscale restoration: maximizing evolutionary potential. Evol App. 1 (4), 587-597 (2008).
  67. Vitt, P., Havens, K., Kramer, A. T., Sollenberger, D., Yates, E. Assisted migration of plants: Changes in latitudes, changes in attitudes. Biol Cons. 143 (1), 18-27 (2010).
  68. Buerger, P., et al. Heat-evolved microalgal symbionts increase coral bleaching tolerance. Sci Adv. 6 (20), 2498(2020).
  69. Chakravarti, L. J., van Oppen, M. J. H. Experimental evolution in coral photosymbionts as a tool to increase thermal tolerance. Fron Mar Sci. 5, (2018).
  70. van Oppen, M. J. H., et al. Shifting paradigms in restoration of the world's coral reefs. Global Change Biology. 23 (9), 3437-3448 (2017).
  71. Harborne, A. R., Rogers, A., Bozec, Y. -M., Mumby, P. J. Multiple Stressors and the Functioning of Coral Reefs. Ann Rev Mar Sci. 9 (1), 445-468 (2017).
  72. Hughes, T. P., et al. Climate change, human impacts, and the resilience of coral reefs. Science. 301 (5635), 929-933 (2003).
  73. Anthony, K., et al. New interventions are needed to save coral reefs. Nat Ecol & Evol. 1 (10), 1420-1422 (2017).
  74. Darling, E. S., Côté, I. M. Seeking resilience in marine ecosystems. Science. 359 (6379), 986-987 (2018).
  75. van Oppen, M. J. H., Puill-Stephan, E., Lundgren, P., De'ath, G., Bay, L. K. First-generation fitness consequences of inter-populational hybridization in a Great Barrier Reef coral and its implications for assisted migration management. Coral Reefs. 33 (3), 607-611 (2014).
  76. Coleman, M. A., Goold, H. D. Harnessing synthetic biology for kelp forest conservation1. J Phycol. 55 (4), 745-751 (2019).
  77. Liboureau, P., Pearson, G. A., Barreto, L., Serrao, E. A., Kreiner, A., Martins, N. Effects of thermal history on reproductive success and cross-generational effects in the kelp Laminaria pallida (Phaeophyceae). Mar Ecol Prog Ser. 715, 41-56 (2023).

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Giant Kelp RestorationGreen Gravel TechniqueMacrocystis PyriferaKelp CulturingSporophyll CollectionSpore InoculationSubstrate SeedingTemperature Control
Field Collection and Laboratory Maintenance of Canopy-Forming Giant Kelp to Facilitate Restoration
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