Land and Food Systems, Faculty of
DSpace
UBCV
Wessler, Leah
Master of Science - MSc
University of British Columbia
Ocean acidification (OA), driven by upwelling and climate change, can negatively impact the ecological and economic contribution of marine calcifiers along coasts worldwide. OA interferes with calcification, particularly in early life stages, causing mortality, reduced growth, and morphological abnormalities in shellfish such as the Pacific oyster (Magallana gigas). This issue is gaining traction as climate change intensifies, placing shellfish in wild populations and farms alike at risk. Macroalgal photosynthesis by seaweed such as Pacific dulse (Devaleraea mollis) has been proposed to provide small-scale OA refuges, but few controlled experiments quantify this effect, and none have focused on larval shellfish. This study examines the potential for Pacific dulse to mitigate OA and its effects on Pacific oyster larvae. Under continuous light for 23 days, the presence of dulse resulted in a consistent increase in seawater aragonite saturation state by 0.1-0.9, and pH by 0.1-0.5 units, depending on OA condition. Newly fertilized oysters were reared for 48 hours in the absence or presence of dulse under treatments corresponding to ambient (pH 7.8, 450 μatm COâ), future OA (pH 7.6, 800 μatm COâ), and future OA + upwelling (pH 7.4, 1200 μatm COâ) seawater conditions. Dulse fully mitigated OA effects on larval size that ranged from decreases of 5% to 10%. Under the future OA + upwelling treatment, dulse presence reduced the odds of underdeveloped oyster larvae at 14 hours post fertilization (hpf), and larvae with hinge abnormalities at 24 hpf, by over 50%. Dulse induced minor changes to immune response gene expression at 48 hpf. These findings highlight the benefits of seaweed when adjacent to organisms sensitive to OA. These findings will be particularly useful for shellfish farms, habitat restoration efforts, and ocean stewardship practices as a potential mitigation strategy under the changing climate.
DULSE SEAWEED Devaleraea mollis MITIGATES EFFECTS OF OCEAN ACIDIFICATION ON LARVAL PACIFIC OYSTERS Magallana gigas by Leah Wessler B.A., Colorado College, 2017 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Applied Animal Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) October 2025 © Leah Wessler, 2025 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the thesis entitled: DULSE SEAWEED Devaleraea mollis MITIGATES EFFECTS OF OCEAN ACIDIFICATION ON LARVAL PACIFIC OYSTERS Magallana gigas submitted by Leah Wessler in partial fulfilment of the requirements for the degree of Master of Science in Applied Animal Biology Examining Committee: Dr. Andrea Y. Frommel, Assistant Professor, Faculty of Land and Food Systems, UBC Supervisor Dr. Christopher D.G. Harley, Professor, Zoology, UBC Supervisory Committee Member Dr. Colin J. Brauner, Professor, Zoology, UBC Additional Examiner Additional Supervisory Committee Members: Dr. Marina A.G. von Keyserlingk, Professor, Faculty of Land and Food Systems, UBC Supervisory Committee Member iii Abstract Ocean acidification (OA), driven by upwelling and climate change, can negatively impact the ecological and economic contribution of marine calcifiers along coasts worldwide. OA interferes with calcification, particularly in early life stages, causing mortality, reduced growth, and morphological abnormalities in shellfish such as the Pacific oyster (Magallana gigas). This issue is gaining traction as climate change intensifies, placing shellfish in wild populations and farms alike at risk. Macroalgal photosynthesis by seaweed such as Pacific dulse (Devaleraea mollis) has been proposed to provide small-scale OA refuges, but few controlled experiments quantify this effect, and none have focused on larval shellfish. This study examines the potential for Pacific dulse to mitigate OA and its effects on Pacific oyster larvae. Under continuous light for 23 days, the presence of dulse resulted in a consistent increase in seawater aragonite saturation state by 0.1-0.9, and pH by 0.1-0.5 units, depending on OA condition. Newly fertilized oysters were reared for 48 hours in the absence or presence of dulse under treatments corresponding to ambient (pH 7.8, 450 μatm CO2), future OA (pH 7.6, 800 μatm CO2), and future OA + upwelling (pH 7.4, 1200 μatm CO2) seawater conditions. Dulse fully mitigated OA effects on larval size that ranged from decreases of 5% to 10%. Under the future OA + upwelling treatment, dulse presence reduced the odds of underdeveloped oyster larvae at 14 hours post fertilization (hpf), and larvae with hinge abnormalities at 24 hpf, by over 50%. Dulse induced minor changes to immune response gene expression at 48 hpf. These findings highlight the benefits of seaweed when adjacent to organisms sensitive to OA. These findings will be particularly useful for shellfish farms, iv habitat restoration efforts, and ocean stewardship practices as a potential mitigation strategy under the changing climate. v Lay Summary Ocean acidification (OA) occurs during climate change when seawater absorbs excess CO2 gas in the air from burning fossil fuels. This chemical change in the ocean negatively impacts marine organisms, thereby threatening our global ecosystems and food security. Fortunately, seaweed utilizes CO2 during photosynthesis, providing a sustainable tool to combat OA. This study examined whether the presence of seaweed could mitigate OA and its negative effects on larval oysters through removal of CO2 from the surrounding seawater. Oyster larvae were reared with or without seaweed in three seawater treatments corresponding to current and two projected future levels of OA. Findings indicate that dulse presence is able to help oyster larvae overcome OA effects. This knowledge emphasizes the importance of seaweed as a potential mitigation tool for shellfish farms and ecosystem restoration projects as climate change intensifies. vi Preface Findings arising from the research described in this thesis were derived from experiments and analysis conducted at Cedar Aquaculture Research and Development (CARD) in Nanaimo, BC and The University of British Columbia in Vancouver, BC. A flow-through experimental aquaculture system was designed, constructed, and maintained at CARD by the present author under the guidance of Dr. Andrea Frommel, Dr. Nina von Keyserlingk, Dr. Chris Harley, Dr. Jennifer Clark, Dr. Tim Green, and George Triska. All experiments were designed and conducted in the aquaculture system by the author, under the guidance of the aforementioned mentors, with help from labmates Ben Niemann and Virginie Chalifoux and from students Nirav Parmar and Olivia Ballard from Vancouver Island University. Oyster broodstock were collected under permit XR 419 2023. Gene expression sample analysis was conducted by the author at the Vancouver Island University Centre for Shellfish Research under the guidance of Dr. Timothy Green and Dr. Ben Sutherland. Microscopy and image analysis to assess oyster larvae size and morphological condition was conducted by the author at the UBC Vancouver Bioimaging facility under the guidance of Dr. EunKyoung Lee. Dissolved inorganic carbon in seawater samples was analyzed by Dr. Jian Guo, and total alkalinity of seawater samples was analyzed by the author, in The University of British Columbia Earth, Ocean, and Atmospheric Sciences laboratory. Generative AI (ChatGPT) was used in the production of this thesis to troubleshoot while coding in R, as one of multiple resources for understanding statistical terminology, and as a thesaurus. vii Table of Contents Abstract ................................................................................................................................... iii Lay Summary ...........................................................................................................................v Preface ..................................................................................................................................... vi Table of Contents .................................................................................................................. vii List of Tables ............................................................................................................................x List of Figures ......................................................................................................................... xi List of Abbreviations ............................................................................................................ xii Acknowledgements .............................................................................................................. xiii Chapter 1: Introduction ..........................................................................................................1 1.1 Ocean acidification (OA) .......................................................................................... 1 1.1.1 General trends ................................................................................................... 1 1.1.2 Local variability ................................................................................................ 1 1.1.3 Carbonate chemistry: emphasis on aragonite saturation state .......................... 2 1.1.4 OA impact on organisms .................................................................................. 3 1.1.5 OA impact on shellfish industry ....................................................................... 4 1.2 Potential solutions for OA ........................................................................................ 4 1.2.1 Alkalinity enhancement .................................................................................... 4 1.2.2 Large-scale carbon uptake through photosynthesis .......................................... 5 1.2.3 Small-scale carbon uptake through photosynthesis .......................................... 6 1.2.3.1 In the field ..................................................................................................... 6 1.2.3.2 In experimental systems ................................................................................ 6 viii 1.3 Knowledge gaps ........................................................................................................ 7 1.4 Research focus .......................................................................................................... 8 Chapter 2: Dulse mitigates effects of ocean acidification on larval oysters ......................10 2.1 Introduction ............................................................................................................. 10 2.2 Methods................................................................................................................... 16 2.2.1 Experimental setup .......................................................................................... 16 2.2.2 Broodstock origin, fertilization, and larval rearing ......................................... 17 2.2.3 Seawater chemistry ......................................................................................... 17 2.2.4 Oyster condition .............................................................................................. 19 2.2.5 Gene expression .............................................................................................. 22 2.2.6 Statistical analysis ........................................................................................... 23 2.3 Results ..................................................................................................................... 25 2.3.1 Seawater chemistry ......................................................................................... 25 2.3.2 Oyster condition .............................................................................................. 29 2.3.2.1 Size .............................................................................................................. 29 2.3.2.2 Development and morphology .................................................................... 30 2.3.3 Gene expression .............................................................................................. 33 2.4 Discussion ............................................................................................................... 34 2.4.1 OA effects on oyster larvae ............................................................................. 35 2.4.2 Dulse effects on OA ........................................................................................ 37 2.4.3 Dulse effect on larvae ..................................................................................... 38 2.4.4 Larger context ................................................................................................. 41 2.5 Conclusion .............................................................................................................. 44 ix Chapter 3: Conclusions .........................................................................................................46 3.1 Overview of findings .............................................................................................. 46 3.2 Limitations of the research ...................................................................................... 47 3.3 Contributions to the field ........................................................................................ 49 3.3.1 Restoration applications .................................................................................. 49 3.3.2 Shellfish hatchery applications ....................................................................... 50 3.3.3 Context within the seafood industry ............................................................... 50 3.4 Future directions of research ................................................................................... 52 References ...............................................................................................................................54 Appendices ..............................................................................................................................77 Appendix A Experimental setup details ............................................................................. 77 Appendix B Spawning details ............................................................................................. 79 Appendix C RNA extraction details ................................................................................... 80 x List of Tables Table 1. Observed developmental stages and abnormal morphologies of oyster larvae that were used to identify underdeveloped or abnormal D-stage (first shell stage) larvae ............ 21 Table 2. Meter-measured pH on experiment day 2 in oyster beakers and carbonate chemistry parameters on day 3 in dulse absent/present tanks ................................................................. 28 Table 3. Average dulse weight in grams throughout 23 days in tumble culture under continuous light ....................................................................................................................... 81 Table 4. Average pH in oyster beakers measured with a hand-held meter ............................. 82 Table 5. Target and reference M. gigas genes examined in the present study through RT-qPCR, including ion regulation, shell formation, and immune response genes ..................... 83 Table 6. Statistical summary for pairwise comparisons of estimated marginal means (EMMs) of oyster larvae size ................................................................................................................ 85 xi List of Figures Figure 1. Experimental timeline ............................................................................................. 18 Figure 2. Delta pH between paired dulse absent/present tanks sharing a header tank during experiment week 1 and 3, recorded every 5 minutes .............................................................. 26 Figure 3. Average aragonite saturation state measured in dulse absent/present tanks on the same weekday and time across three weeks ........................................................................... 27 Figure 4. Estimated Marginal Means (EMMs) of oyster larvae shell diameter (µm) at two sampling timepoints: 24 and 48 hours post fertilization ......................................................... 29 Figure 5. Bar graphs of average percent abnormal larvae with respect to four condition categories: development, tissue, and two hinge morphological characteristics ...................... 31 Figure 6. Average change driven by dulse presence in relative gene expression across nine M. gigas target genes .................................................................................................................... 33 Figure 7. Experimental aquaculture system built for dulse-oyster co-culture research .......... 78 xii List of Abbreviations ANOVA â analysis of variance AT â total alkalinity Ca²âº â calcium ion CaCOâ â calcium carbonate CARD â Cedar Aquaculture Research and Development COâ²â» â carbonate DIC â dissolved inorganic carbon EMM â estimated marginal mean HâCOâ â carbonic acid HCOââ» â bicarbonate HPD â highest priority density Hpf â hours post fertilization OA â ocean acidification OAE â ocean alkalinity enhancement pCOâ â partial pressure of gaseous COâ in seawater Ωarag â aragonite saturation state Ωcalcite â calcite saturation state xiii Acknowledgements I extend a mountain of gratitude to my supervisor Dr. Andrea Frommel, and my Supervisory Committee members Dr. Nina von Keyserlingk and Dr. Chris Harley, for their invaluable encouragement and guidance. My deepest gratitude to labmates at the Aquaculture and Climate Change Lab, in particular Ben Niemann and Virginie Chalifoux, for their myriad forms of support, including assistance with running experiments on Vancouver Island. To Nirav Parmar, Olivia Ballard, Eloise Pons, and Karyna Howell, I am also deeply grateful for your enthusiastic help with running experiments. I am immensely grateful to Roger Paquette for providing me the opportunity to conduct research at Cedar Aquaculture Research and Development (CARD), and to those who supported me through many months living at CARD, tinkering with and piloting my experimental system, including Jessica Driver, Holli Desrocher, Don MacQuarrie, Renee Dekleer, George Triska, Ken Shepherd, Willie Callies, Lawrence Lambert, Mark Charles, and Leticia. To Cascadia Seaweed, and Dr. Jennifer Clark in particular, I extend gratitude for providing industry-grade Pacific dulse, along with equipment and invaluable guidance in culturing the dulse. I send immense gratitude to Angela Fortune at Nova Harvest for providing me with oyster seed for my research. I am deeply grateful to Dr. Tim Green, Dr. Ben Sutherland, and students Korrina Gilchrist, Marissa Wright-LaGreca, and all others at Vancouver Island University for taking me under their wing during my time on Vancouver Island, providing me with broodstock, teaching me how to spawn and sample larval oysters, and welcoming me into their lab while providing endless guidance for gene expression analysis. Finally, I extend immense gratitude to my xiv Examining Committee members Dr. Andrea Frommel, Dr. Colin Brauner, and Dr. Chris Harley for their thoughtful questions and feedback during my defense. 1 Chapter 1: Introduction 1.1 Ocean acidification (OA) 1.1.1 General trends Gaseous carbon dioxide (CO2) builds in the atmosphere due to the burning of fossil fuels, and the current concentration is higher than at any other time in the last two million years (IPCC, 2019). A third of the excess CO2 in the atmosphere has been absorbed by the ocean (Friedlingstein et al., 2019; Gruber et al., 2019), driving a cascade of chemical changes in seawater worldwide known as ocean acidification (OA) (Hurd et al., 2020). The term directly refers to an increase in hydrogen ions in the ocean, which leads to an increase in acidity caused by a decrease in pH. Since the Industrial Revolution, the average pH of ocean surface water has declined from approximately 8.2 to 8.1, corresponding to a 30% increase in acidity (i.e., hydrogen ion concentration) (Raven et al., 2005). Significant shifts in ocean carbonate chemistry are projected to continue throughout the century (Olson et al., 2025; Peña & Fine, 2024). A simulation of future pH values projects that under RCP 8.5, annual mean surface pH will decrease by 0.21 pH units by 2070 (Peña & Fine, 2024). 1.1.2 Local variability Severity of OA varies depending on local variability in ocean conditions (IPCC, 2019; Olson et al., 2025). For example, western coasts across the globe undergo seasonal upwelling, during which deep water off the continental shelf with high concentrations of CO2 is pulled to the surface (Feely et al., 2008; Reum et al., 2016). Oceans at higher latitudes tend to also have higher CO2 concentrations and other carbonate chemistry parameters that are associated 2 with OA (IPCC, 2019; L.-Q. Jiang et al., 2015). For example, upwelling events along the northwestern coast of North America have resulted in consistent surface pH levels as low as 7.6 since the 1980s (Marliave et al., 2011), which corresponds to a hydrogen ion concentration that is ~200% lower than the current global average at a pH of 8.1 (NOAA, 2025). OA projections for the Pacific coast of BC indicate that by 2070, consistent pH values of 7.4 may be common (Evans et al., 2019; Holdsworth et al., 2021). Organisms living in habitats with naturally higher CO2 levels and fluctuations may be more adapted to OA conditions (Caillon et al., 2023; Stillman & Paganini, 2015). However, chemical thresholds for organism biochemical success may also be surpassed more quickly within these areas as OA progresses, such as undersaturated aragonite states (Bednaršek et al., 2019; Crook et al., 2013; García-Ibáñez et al., 2021; Yamamoto-Kawai et al., 2025). 1.1.3 Carbonate chemistry: emphasis on aragonite saturation state Much of the discourse around OA focuses on changes to levels of pH or carbon dioxide in seawater, but multiple carbonate chemistry parameters change under OA (Hurd et al., 2020). Absorbed CO2 reacts with seawater to form carbonic acid (H2CO3), an unstable acid that dissociates into bicarbonate (HCO3-) and a hydrogen ion. Hydrogen ions from this reaction can bind to carbonate (CO32-), forming more bicarbonate and removing available carbonate ions from the seawater (Hurd et al., 2020). Meanwhile, organisms like coral and shellfish form shells and skeletons through the calcification process by binding calcium ions (Ca2+) with carbonate to form either aragonite or calcite (CaCO3). The removal of carbonate from seawater through the process of OA 3 decreases the available building blocks for calcification. The chemical relationship between formed shells or skeletons (aragonite or calcite) and the availability of their building blocks (calcium and carbonate ions) is called the aragonite or calcite saturation state (Ωarag or Ωcalcite). A saturation state â¥ 1 favors calcification, and a saturation state < 1 favors the dissolution of calcium structures (Hurd et al., 2020). Therefore, tracking changes in Ωarag along with CO2, pH, and other carbonate chemistry parameters in seawater is necessary for a complete understanding of OA effects on ocean ecosystems. 1.1.4 OA impact on organisms Shifts in ocean chemistry driven by OA can interfere with biological activity of marine organisms, such as reproduction, growth, metabolism, immune response, and behavior in both fish and invertebrates (Barry et al., 2014; Bressan et al., 2014; Frommel et al., 2014, 2016, 2025; Kong et al., 2022; Kroeker et al., 2010; Ou et al., 2015; Padilla-Gamiño et al., 2022; Waldbusser et al., 2014; Q. Wang et al., 2016). Calcifying organisms such as shellfish can be sensitive to current levels of ocean acidification, which can cause shell dissolution and mortality in ecologically and commercially important species (Barton et al., 2012; Bednaršek et al., 2017, 2020; Miller et al., 2016; Osborne et al., 2020). Early life stage parameters of shellfish appear to be particularly vulnerable to OA, such as fertilization, larval survival, development, growth, and timing of feeding initiation (Barros et al., 2013; Bates et al., 2025; Waldbusser et al., 2014, 2015; Watson et al., 2009). For example, OA treatments (pH 7.8 and 7.4) decreased fertilization rate and larval viability of Pacific oyster Magallana gigas compared to the control treatment (pH 8.1), and larvae were smaller with more abnormalities in OA treatments at 48, 96, and 144 hours post fertilization (Barros et al., 2013). Such effects 4 compound as shellfish develop past the larval stages, reducing their competitive capability and survival (Gazeau et al., 2010; Kurihara et al., 2007). 1.1.5 OA impact on shellfish industry Along the North American West Coast from Oregon to British Columbia (BC), OA sensitivity in shellfish has contributed to production failures in multiple bivalve hatcheries (Barton et al., 2015; Clements & Chopin, 2017). Although hatcheries for early bivalve life stages occur in land-based aquaculture systems, they are not safe from ocean acidification as most hatcheries source their saltwater directly from the ocean. Despite the economic and ecological importance of marine organisms, research on optimal solutions to mitigate the impacts of climate change remain understudied (Froehlich et al., 2022). 1.2 Potential solutions for OA 1.2.1 Alkalinity enhancement Ocean alkalinity enhancement has been proposed as an efficient way to combat atmospheric CO2 accumulation and OA, which can involve buffering seawater by releasing finely ground alkaline material into the oceans (Eisaman et al., 2023; Gattuso et al., 2021). Increasing ocean alkalinity would theoretically enhance CO2 uptake by the sea surface and increase its buffering capacity, lowering ocean acidity and raising Ωarag and Ωcalcite (Bednaršek et al., 2025). Some oyster hatcheries have already turned to this OA mitigation strategy, adding soda ash to incoming seawater during upwelling events (Barton et al., 2015; Clements & Chopin, 2017; Gimenez et al., 2018). However, soda ash buffering during oyster larvae development has been shown to have a carryover effect on the immune system during 5 subsequent life stages, causing high mortality during viral infections in juvenile oysters (Mackenzie et al., 2022). The potential for negative effects of seawater alkalinity enhancement on marine organisms, particularly those that the enhancement proposes to benefit, underlines the importance of seeking alternatives for ocean carbon uptake. 1.2.2 Large-scale carbon uptake through photosynthesis Blue Carbon, a term defined in 2009, refers to the 55% of all biological carbon that is stored by photosynthesizing marine ecosystems such as mangrove forests, seagrass meadows, and tidal marshes (Nellemann & Corcoran, 2009). This storage process is called carbon sequestration: it begins with plants utilizing sunlight to convert CO2 into oxygen and living tissue full of carbon (photosynthesis), and continues when tissues are buried in ocean sediments for long-term storage of carbon that can last for millennia (Mateo et al., 1997). The contribution to Blue Carbon that marine algae provide is currently being investigated due to their immense capacity for photosynthesis and high growth rates (Bhuyan et al., 2021). However, limited data and certainty exists regarding the carbon sequestration potential of algae (Pessarrodona et al., 2023). Monitoring and modeling the fate of carbon taken up by algae to identify whether itâs released back into the atmosphere or stored long-term as Blue Carbon proves difficult and complex (Hurd et al., 2022; Nishihara et al., 2025). The capacity for primary producers such as algae to uptake CO2 from seawater through photosynthesis is well studied and understood, however, and potentially offers short-term, local refuges from increasing OA conditions (Pessarrodona et al., 2023). 6 1.2.3 Small-scale carbon uptake through photosynthesis 1.2.3.1 In the field Studies show potential for seagrass and macroalgae (seaweed) to actively ameliorate ocean acidification. Seawater pH within seagrass beds can be 25% or more above that of non-vegetated areas, even past daylight hours and across years (Ricart, Ward, et al., 2021), and a 40% increase in growth rate has been observed in oysters when reared adjacent to high seagrass biomass (Ricart, Gaylord, et al., 2021). Kelp forests and farms have been shown to alter the seawater carbonate chemistry, and oysters within a kelp farm have exhibited reduced shell dissolution and faster growth compared to the same species grown outside the kelp farm (Murie & Bourdeau, 2020; Peabody et al., 2019; Pfister et al., 2019). However, OA mitigation is not always apparent, such as when adult mussels are adjacent to fucus and seagrass beds (Wahl et al., 2018). Field studies also come with challenges in quantifying the contribution of seaweed to seawater chemistry, such phytoplankton blooms and variability in CO2 exchange between the air and ocean surface driven by temperature, wind and tidal fluctuations (Murie & Bourdeau, 2020; Pfister et al., 2019; Van Dam et al., 2021). Furthermore, the potential for seaweed to mitigate OA effects on sensitive larval shellfish in the field is unknown due to the challenge of studying planktonic larvae in the field. 1.2.3.2 In experimental systems Studies in mesocosms and laboratories allow us to examine challenging organism life stages, and to model complex OA, organism, and ecosystem relationships in the field by providing data from a controlled environment in which most variables are known such as flow rate, seawater volume, and organism densities. In one mesocosm experiment, gracilaria seaweed 7 ameliorated high pH/low CO2 conditions driven by adjacent scallop respiration (Z. Jiang et al., 2014). Another mesocosm study showed that kelp can mitigate negative effects of OA on adjacent oysters (Z. Jiang et al., 2022). Fucus seaweed did not mitigate effects of OA on mussel calcification in a mesocosm, but was able to mitigate the OA effects on both seawater pH and mussel calcification during a laboratory experiment, depending on seaweed density (Wahl et al., 2018). Land-based laboratory studies help us understand the nuanced relationships within ecosystems, and assist with building models on large-scale impacts of OA. Studies in land-based systems are also directly relevant to aquaculture facilities such as shellfish hatcheries. Previous literature shows that in land-based systems, effects of OA on calcifiers can be mitigated when calcifying foraminifera is incubated with their algal hosts (Doo et al., 2020), juvenile bivalves are in the presence of kelp (Young et al., 2022), adult bivalves are co-cultured with ulva seaweed (Young & Gobler, 2018), and when juvenile abalone is grown with dulse seaweed (Hamilton et al., 2022). However, none of these studies examine sensitive larval life stages, nor directly apply to potential solutions regarding North American West Coast hatchery failures. 1.3 Knowledge gaps It is still relatively unknown how differences between seaweed species and densities affect OA mitigation in sensitive life stages (Clements & Chopin, 2017). One gap in understanding the relationship between OA-sensitive organisms and seaweed in co-culture is the lack of studies on larval life stages in the field or in land-based systems, which may be an inflection 8 point in organism success under OA stress. Most larvae enter pelagic plankton communities and are therefore not necessarily in close proximity to seaweed, but spawning, fertilization, embryonic, and settlement life stages can occur in shallow, coastal habitats (Geffen, 2009; Jablonski & Lutz, 1983; Swearer et al., 2019). To date, no seaweed co-culture studies have been conducted within the context of shellfish hatchery failure along the northwestern Pacific coast of the US and Canada. Only one previous study has examined a species of seaweed (Pacific dulse) that is already farmed in land-based aquaculture in North America (Hamilton et al., 2022). The potential for seaweed to serve as an OA refuge for sensitive organisms in both ecosystem and industry contexts requires further attention. 1.4 Research focus Dulse seaweeds have been harvested as food and medicine for centuries by First Nations in Canada (Krzus, 2019) and by people in northern Europe (Tasch, 2016). The red seaweed is an anti-inflammatory and antioxidant nutraceutical (Millan-Linares et al., 2019), rich in iron, potassium, calcium, and iodine (Krzus, 2019). Currently, dulse is not widely eaten in North America, but has been identified as an important modern food and supplement source (Jibri et al., 2016). Dulse seaweeds are high in protein, showing potential as an alternative protein source (Mendez & Kwon, 2022) such as vegan bacon (Tasch, 2016). Present-day economic importance of dulse includes use as feed, biofilter, and potential OA buffer for red abalone, a high market value seafood (Demetropoulos & Langdon, 2004a, 2004b; Hamilton et al., 2022). 9 Farmed dulse is a particularly good fit for use in shellfish hatcheries as the crop is produced in land-based aquaculture through its entire life cycle, whereas most farmed kelp is only in land-based aquaculture during initial growth stages. Experiments for this project were conducted on Vancouver Island, where shellfish aquaculture is a staple industry, and where OA may have already contributed to shellfish farm collapse, although evidence has not been peer-reviewed (CBC, 2014). OA may be exacerbated along the southern coast of BC during upwelling events, which can coincide with the timing of hatchery spawns (Reum et al., 2016). Vancouver Harbor pH has historically ranged between 7.8 and 8.1, but the pH has dropped below 7.6 quite often beginning in the 1970âs (Marliave et al., 2011). Considering current upwelling trends and future projections of OA for the BC coastline, southern BC could experience regular drops in surface pH to around 7.4 by 2070 (Evans et al., 2019; Holdsworth et al., 2021; Peña & Fine, 2024). This research investigated the potential for the red seaweed Pacific dulse (Devaleraea mollis) to mitigate effects of OA larval Pacific oysters (Magallana gigas) in land-based co-culture. OA was expected to decrease larval oyster size, increase percent larvae exhibiting underdevelopment or morphological abnormalities, and change expression of genes involved with OA-sensitive functions in the larvae (ion regulation, shell formation, and immune response) during the first 48 hours post fertilization. Dulse was expected to increase the seawater pH and Ωarag in all treatments, thereby ameliorating OA conditions. Dulse influence on seawater chemistry was expected to mitigate any size decrease, abnormality increase, or gene expression change induced by OA in the larval oysters. 10 Chapter 2: Dulse mitigates effects of ocean acidification on larval oysters 2.1 Introduction Atmospheric carbon dioxide (CO2) levels continue to surge higher than at any other time in the last two million years due to the burning of fossil fuels (IPCC, 2019). The ocean has absorbed roughly a third of anthropogenically produced CO2 through the chemical process of ocean acidification (OA): gas exchange between air and the ocean surface triggers a host of chemical changes in seawater such as declines in pH, Ωarag, and Ωcalcite (Friedlingstein et al., 2019; Gruber et al., 2019; Hurd et al., 2020). Ocean surface pH has decreased by ~0.1 units since the Industrial Revolution â corresponding to a 30% increase in hydrogen ion concentration â and is projected to drop by another 0.2 pH units in the next 50 years if high fossil fuel burning rates continue (Holdsworth et al., 2021; Peña & Fine, 2024; Raven et al., 2005). Geographic variability in ocean conditions can exacerbate severity of OA (IPCC, 2019; Olson et al., 2025). Western coasts of continents and oceans at higher latitudes are particularly impacted by climate change due to upwelling and tidal mixing: ocean currents pull cold, deep water with high CO2 concentrations onto the continental shelf, exacerbating surface seawater OA conditions to well above those predicted globally for the end of the century (Evans et al., 2019; IPCC, 2019; L.-Q. Jiang et al., 2015; McNeil & Sasse, 2016). High latitude western coasts across the globe, such as the North Salish Sea, could experience regular drops in surface pH to 7.4 and another month of the year with undersaturated Ωarag 11 conditions within the next 50 years (Evans et al., 2019; García-Ibáñez et al., 2021; Yamamoto-Kawai et al., 2025). Many biological activities of marine fish and invertebrates deteriorate under OA seawater conditions, particularly in calcifying organisms such as corals and shellfish (Barry et al., 2014; Bressan et al., 2014; Frommel et al., 2014, 2016, 2025; Kong et al., 2022; Kroeker et al., 2010; Ou et al., 2015; Padilla-Gamiño et al., 2022; Waldbusser et al., 2014; Q. Wang et al., 2016). The ecosystem-wide consequences when such effects are compounded within population and food web scales are unknown and complex (Gaylord et al., 2015). Therefore, modeling ecosystem change as OA intensifies will be vital for the seafood industry, restoration efforts, and coastal communities. Such modeling relies on experimental data on the organismal level that elucidates how species are impacted by specific carbonate chemistry parameters, and what mitigation strategies may be effective. In shellfish, the reduction in pH and Ωarag associated with OA can cause shell dissolution, mortality, feeding and immune response disruption, and microbiome shifts (Kong et al., 2022; Miller et al., 2016; Waldbusser et al., 2015; Q. Wang et al., 2016). Shellfish reefs along coasts of Australia are declining compared to their historical baseline due in part to OA (Gillies et al., 2020). The annual economic deficit due to OA on the European shellfish industry is projected to reach 1 billion USD by 2100 (Narita & Rehdanz, 2017). Along the northern Pacific coast of North America, OA sensitivity in shellfish has led to production failures in bivalve hatcheries, where pH has consistently declined to 7.6 units in certain areas 12 since the 1980s during upwelling events, and seasonal Ωarag undersaturation is already common (Barton et al., 2015; Evans et al., 2019; Marliave et al., 2011). Hatchery failures are partly attributed to how early life stages of shellfish are particularly vulnerable to OA. Undersaturated Ωarag have been found to impact larval shell development and growth in the Mediterranean mussel, California mussel, and Pacific oyster (Barros et al., 2013; Waldbusser et al., 2014, 2015; Watson et al., 2009). In some shellfish such as Pacific oysters, OA vulnerability is particularly acute during the first two days after fertilization when larvae are forming their first shell (Barros et al., 2013; Waldbusser et al., 2013). Such effects compound as shellfish develop past the larval stages, reducing their competitive capability and survival (Gazeau et al., 2010; Kurihara et al., 2007). Research on effective solutions to mitigate climate change impacts on marine organisms such as shellfish remains limited, despite their economic and ecological significance (Clements & Chopin, 2017; Froehlich et al., 2022). Potential OA mitigation strategies include both geochemical and ecosystem-based approaches. To combat the accumulation of atmospheric CO2 and resulting OA, ocean alkalinity enhancement (OAE) has been proposed as a strategy to increase oceanic COâ uptake, strengthen buffering capacity, and raise Ωarag and Ωcalcite through the addition of alkaline material to seawater (Bednaršek et al., 2025; Eisaman et al., 2023; Gattuso et al., 2021). However, benefits may not outweigh the costs: OAE during larval development in oyster hatcheries has been shown to cause high mortality during viral infections in subsequent juvenile life stages (Mackenzie et al., 2022). Focusing instead on fostering 13 marine photosynthesizers can provide a natural, worldwide climate change mitigation strategy through long-term storage of carbon in ocean sediments (Blue Carbon) and short-term alleviation of OA through CO2 uptake during photosynthesis (Macreadie et al., 2019; Pessarrodona et al., 2023). Potential seaweed contributions to Blue Carbon are currently being investigated, given their immense capacity for utilizing sunlight to convert CO2 into oxygen and biomass (Bhuyan et al., 2021). To date, limited data exists on Blue Carbon from macroalgal forests on a global scale, despite evidence that such forests are carbon sinks (Pessarrodona et al., 2023). Irrespective of their contribution to Blue Carbon, seagrass and seaweed can provide short-term and small-scale chemical refuges from OA through the uptake of CO2 (Pessarrodona et al., 2023; Ricart, Ward, et al., 2021). However, quantification can be challenging due to an array of confounding factors, such as phytoplankton blooms and variability in CO2 exchange between the air and ocean surface driven by temperature, wind and tidal fluctuations (Clements & Chopin, 2017; Han et al., 2017; Z. Jiang et al., 2014; Manzello et al., 2012; Murie & Bourdeau, 2020; Pfister et al., 2019; Ricart, Ward, et al., 2021; Van Dam et al., 2021). These refuges require research attention to evaluate possible compounding large-scale benefits for marine organisms and populations, particularly in regions with extended day length in the higher latitudes that allows for net positive photosynthesis. Few studies have clearly quantified how OA refuges provided through photosynthesis may mitigate effects of OA on marine calcifiers. Of these, only one study has direct applicability to land-based aquaculture such as shellfish hatcheries. To date, no research includes the 14 larval stage of shellfish, which can be particularly sensitive to OA and therefore may be an inflection point in organism success under OA stress. Some field and mesocosm studies on seagrass or seaweed OA refuges show a negative or lack of effect on marine calcifiers (Listiawati & Kurihara, 2023; Wahl et al., 2018), while other findings suggest that refuges benefit calcification and growth (Bergstrom et al., 2019; Z. Jiang et al., 2014, 2022; Peabody et al., 2019; Ricart, Gaylord, et al., 2021). Confounding variables that may obscure quantification of OA mitigation by marine photosynthesizers include fluctuating currents, daylight intensity and duration, and organism densities (Clements & Chopin, 2017; Peabody et al., 2019). Furthermore, seagrass OA refuges only benefit the relatively small number of species that thrive in seagrass meadows, are only potentially relevant to ocean aquaculture if meadows are adjacent to farms in an area with low water movement, and are not relevant to land-based aquaculture. Controlled experiments with seaweed can account for confounding variables and apply to a larger number of species, providing clear quantification of OA refuge effects with wide applicability that includes seaweed ecosystem modeling and land-based aquaculture. Previous controlled, land-based experiments on seaweed mitigation of OA show benefits provided by fucus for adult mussles (Wahl et al., 2018), kelp or ulva for various juvenile shellfish species (Young et al., 2022; Young & Gobler, 2018), and dulse for juvenile abalone (Hamilton et al., 2022). However, only one of these studies is directly applicable to land-based industrial or restoration aquaculture. Fucus is not widely cultivated, kelp is not cultivated on land, and though ulva is cultivated on land, the shellfish that were paired with 15 ulva are not typically cultivated on land during their juvenile stage. In contrast, Pacific dulse (Devaleraea mollis) is produced in land-based aquaculture throughout its entire life cycle. Dulse is therefore a particularly good fit for use in land-based co-culture, where it has already been shown to mitigate OA effects of current seawater conditions on juvenile abalone size (Hamilton et al., 2022). Studies on dulse culture have primarily focused on maximizing growth instead of capacity for OA mitigation, so dulse has not previously been tested under 24-hour continuous light to evaluate its potential for stable OA mitigation without nightly carbonate chemistry fluctuations (Demetropoulos & Langdon, 2004a; Dittrich et al., 2025). This present research investigates the potential for OA mitigation benefits of Pacific dulse (Devaleraea mollis) to extend to land-based hatcheries for larval Pacific oysters (Magallana gigas) within the context of both current and future OA conditions. Pacific oysters are native to the Northwestern Pacific Ocean but are farmed in six continents and are the most globally cultivated shellfish in tonnes of produced seafood (FAO, 2025). Larval response under OA to seaweed presence may have population-wide implications and applies directly to current climate-related obstacles in aquaculture. The objectives of the current study were to 1) determine the impact of dulse under continuous light for 23 days on seawater carbonate chemistry at three levels of OA: ambient (pH 7.8, 450 μatm CO2), OA (pH 7.6, 800 μatm CO2), and OA + upwelling (pH 7.4, 1200 μatm CO2), and 2) to determine the potential of dulse to mitigate effects of OA on oyster larvae condition (size, development, morphology) and expression of genes involved with OA-sensitive functions (ion regulation, shell formation, and immune response), during the first 48 hours of larval development. 16 2.2 Methods 2.2.1 Experimental setup The experimental system was located at Cedar Aquaculture Research and Development in Nanaimo, Vancouver Island, BC, Canada (see Figure 7 and experimental setup details in Appendix). CO2 was manipulated with an iks Aquastar system (iks ComputerSysteme GmbH, Karlsbad, Germany) in flow-through header tanks (62 L, < 4 L/min, 10-14°C) with three replicates for each pH target level. Seawater treatment groups were based on three OA scenarios: 1) ambient seawater (pH ~7.8, not manipulated), 2) projected OA near Vancouver Island in 50 years (pH 7.6), and projected OA during upwelling events (pH 7.4) (Evans et al., 2019; Holdsworth et al., 2021; Marliave et al., 2011). Henceforth âpH treatmentâ refers to the target pH manipulated in header tanks for each OA scenario, not the actual pH under which oysters were cultured. Each header tank gravity fed into paired 40L tanks, one with 400g (10 gram/L) Pacific dulse (Devaleraea mollis) in tumble culture, and one with seawater only (18 tanks, 40L, < 2 L/min, 10-15°C). Seaweed density was consistent with industry standards for dulse farming (J. Clark, personal communication, 2022). Tanks with dulse received continuous light at 280-350 photon/m2*s, and tanks with only seawater were in darkness to limit microalgae growth. Dulse was added into the system five days before oyster fertilization for acclimation to the 24-hour continuous light, and cultured for 23 days to monitor the extended capacity for dulse to maintain OA mitigation without diel fluctuations (see Figure 1). Despite lack of fertilizer, dulse weight increased weekly by 30-50% (see Table 3 in Appendix), so biomass was removed on experiment days 3 and 12 to maintain the desired density of 10gram/L. 17 Oyster larvae were cultured in a static system within covered beakers (18 beakers, 3 L, 25°C, n = 3 replicates for each OA x dulse treatment combination), filled with seawater from one of 18 corresponding experimental tanks. All tanks and beakers were covered to limit CO2 off-gassing, and surfaces were cleaned every few days to avoid bacterial or microalgal growth. 2.2.2 Broodstock origin, fertilization, and larval rearing Pacific oyster (Magallana gigas) broodstock were collected in July 2024 from a naturalized population in the intertidal zone surrounding Cedar Aquaculture Research and Development. Oysters were strip spawned into ambient seawater at room temperature (22-25°C) and crossed to create two family lines (see spawning details in Appendix). The family exhibiting a higher number of polar bodies was selected for the experiment, and added to static culture beakers at a density of 30 embryos/ml. At 24 hours post fertilization, a full water exchange was conducted by rinsing larvae onto a 35µm screen, cleaning the oyster beaker, and returning larvae within 1-2 minutes to their beaker filled with fresh, preemptively warmed seawater from the corresponding experimental tank. Oyster culture procedures were adapted from common hatchery practices (FAO, 2004). As larvae were terminally sampled at 48 hours post fertilization, an age that can be reached without initiation of feeding, no larvae feed was added to the static culture (see Figure 1). 2.2.3 Seawater chemistry Oyster beaker pH was measured with a hand-held meter at multiple timepoints (see Table 4 in Appendix), yet beaker volumes were insufficient to sample for lab analysis of carbonate 18 chemistry parameters. Seawater was therefore sampled from the dulse absent/present tanks for carbonate chemistry analysis; samples were taken on experiment days 3, 10, and 17, poisoned with mercuric chloride, and stored in amber glass bottles until lab analysis (see Figure 1). Dissolved inorganic carbon (DIC) and total alkalinity (AT) were analyzed through carbon coulometry and automated acid titration, respectively. Ωarag, Ωcalcite, and pCOâ were calculated from DIC and AT with CO2SYSv3.0 (Pierrot et al., 2021) using solubility and dissociation constants from Lueker et al. (2000), Dickson (1990), Perez & Fraga (1987), and Lee et al. (2010). Figure 1. Experimental timeline. Dulse seaweed was acclimated to seawater treatments for five days before fertilization of oyster larvae, and was held under continuous light for 23 days. Complete water changes from experimental tanks to static oyster beaker cultures were conducted directly before adding fertilized larvae to treatment beakers and at 24 hours post fertilization. Larval oysters were sampled at 14, 24, and 48 hours post fertilization. Water samples were taken from dulse absent/present tanks on experiment days 3, 10, and 17. Dulse in 9 of 18 experimental tanks Oyster larvae in beakers Water change from experimental tanks to oyster beakers Dulse receiving continuous light Experimental tank water sampled for chemical analysis Larvae subsample fixed for morphological analysis Larvae sampled for RT-qPCR 19 2.2.4 Oyster condition Oyster larvae were sampled for visual analysis at three stages of early development: pre and early D-stage larvae at 14 hours post fertilization (hpf), when hatcheries usually initiate feeding at 24 hpf, and well-established D-stage larvae at 48 hpf (Min et al., 2023) (See Figure 1). To sample, beakers were thoroughly mixed to ensure homogenous larval density, 12ml was pipetted onto a 35µm screen rinsed into a 1.5ml microcentrifuge tube. Tubes were centrifuged at 2680 x g / 7000 RPM for 5 minutes, and seawater was replaced with 200µl of 10% buffered formalin for larval fixation. Fixed larvae were pipetted into multi-well imaging plates and photographed using an Evident SpinSR IXplore spinning disk confocal microscope with a 20x objective and 2x Hamamatsu ORCA-Fusion BT back-thinned Gen-III sCMOS camera with CellSens Dimension software. All images were processed blind to the treatment in which the larvae were reared. Development stage and morphological category were scored and size was measured using size-calibrated images in Fiji (ImageJ) (Schindelin et al., 2012). Size was defined as the widest shell diameter parallel to the hinge. D-stage larvae with hinges at an angle or perpendicular to the plane of focus were not measured. Only D-stage larvae categorized as normally developed were measured, because abnormally developed larvae are often smaller and may conflate shell size findings (Waldbusser et al., 2015). At 14 hours post fertilization, egg, morula, and blastula larval stages were considered abnormal (underdeveloped), and gastrula and trochophore stages were considered normally developed (see Table 1). At 24 hours post fertilization, all larvae that had not formed a shell (had not reached D-stage) were considered abnormal (underdeveloped). Abnormal D-stage morphological categories 20 included protruded tissue, convex hinge, and rounded hinge, adapted from Barros et al. (2013) and Wright-LaGreca et al. (2022). 21 Table 1. Observed developmental stages and abnormal morphologies of oyster larvae that were used to identify underdeveloped or abnormal D-stage (first shell stage) larvae. Normal timescales of each developmental stage were adapted from Vogeler et al. (2016). Any larvae developing outside of the normal timescales listed below for each stage was considered abnormally developed. Categories of abnormal D-stage morphologies adapted from Barros et al. (2013) and Wright-LaGreca et al. (2022). hpf: hours post fertilization Observed developmental stages Egg (0-1 hpf) Embryo, morula, and blastula (2-6 hpf) Gastrula and early trochophore (7-14 hpf) Trochophore (15-23 hpf) D-stage (24+ hpf) Observed abnormal D-stage morphologies Protruded tissue Convex hinge Rounded hinge (& protruded tissue) 22 2.2.5 Gene expression At 48 hpf, all remaining larvae were concentrated into a 1.5 ml microcentrifuge tube as described above and stored at -80°C until gene expression analysis through RT-qPCR. RNA was extracted with the Monarch Total RNA Miniprep Kit (New England Biolabs), following manufacturer instructions for samples of tissue under 10mg with slight modifications (see RNA extraction details in Appendix). RNA (50ng) was reversed transcribed using the iScript Select cDNA Synthesis Kit (Bio-Rad) with random primers per manufacturer instructions. Target genes and primers were selected from previous studies on M. gigas larvae (Min et al., 2023; Wright-LaGreca et al., 2022) and adults (Q. Wang et al., 2016) (see Table 5 in Appendix). Three ion regulation and three immune response genes were selected that had previously shown expression level change in oysters under OA. The shell formation genes were recently characterized in larvae, so effect of OA was unknown. Efficiency of primers was tested with two serial dilutions (4fold and 5fold) with five logs of dynamic range beginning with a 1:4 or 1:10 dilution, respectively, of neat cDNA pooled from one sample from each treatment (N=6). PCR efficiency was between 90-110% with a dynamic range of â¤ 1:10 to â¥ 1:1250, unless otherwise noted (see Table 5 in Appendix). Neat cDNA was then diluted 20-fold for qPCR. SsoAdvanced Universal SYBR Green Supermix (Bio-Rad) was used for triplicate 10 µl qPCR reactions in a CFX384 Touch Real-Time PCR Detection System (Bio-Rad) with a thermocycling profile from Wright-LaGreca et al. (2022). NTC and -RT controls showed negligible amplification. Target genes normalized to elongation factor 1-alpha (EF1α, GenBank #AB122066) after comparing primer efficiency of three reference gene candidates from Huan et al. (2017) (see Table 5 in Appendix). Relative 23 gene expression was calculated using the 2-ÎÎCq adapted from Schmittgen & Livak (2008). Mean ÎCq in the ambient, dulse absent treatment provided the baseline for calculating ÎÎCq(A) for statistical modeling. ÎCq from the dulse absent treatment of each tank pair replicate provided the baseline for calculating ÎÎCq(B) to graph change in log2-transformed relative expression due to dulse. 2.2.6 Statistical analysis Statistical analysis was conducted using R software (R Core Team, 2024). All models included header tank as a random factor, or oyster beaker when data was collected at multiple timepoints, and all full models included OA and dulse treatments as interactive fixed factors. Model assumptions were evaluated using ShapiroâWilk tests and DHARMa scaled residuals (for GLMMs), with formal checks for residual uniformity, outliers, and overdispersion. Log-likelihood ratio tests were performed to compare reduced and full model fit. Tukey-adjusted pairwise comparisons of the estimated marginal means (EMMs) were performed to identify whether OA treatments affected the dependent variable when dulse treatment was held constant, or conversely, whether dulse affected the dependent variable when OA treatment was held constant. The two sets of contrasts were considered two separate families. An α-level of P < 0.05 was used for assessing statistical significance, and marginal significance (0.05 < P < 0.1) was noted. Aragonite saturation state was analyzed with a linear mixed model that included factors listed above and sample week as a fixed factor in both reduced and full models. A Type III Analysis of Variance test with Satterthwaite's method was performed on the full model. 24 Three outliers out of 72 samples were removed that had either a pCOâ or Ωarag z-value outside of the acceptable range (|z| â¥ 3). Shell diameter was analyzed using a generalized linear mixed model with a Gaussian distribution and log link because reduced linear mixed model residuals were not normally distributed. In addition to factors listed above, models included time as a fixed factor. To address heteroscedasticity, the model allowed dispersion to vary by hour, treatment, and dulse. A Type III Analysis of Deviance Wald ÏÂ² test was performed on the full model. Four outlier measurements out of 321 were removed that had a shell diameter outside of the acceptable range (|z| â¥ 3) when grouped by sampling time (either 24 or 48 hours post fertilization). Proportion data on percent larvae underdeveloped or with abnormal morphology was analyzed for each timepoint using a Bayesian binomial regression model to accommodate overdispersion and complete separation in the data. Priors used for the intercept and fixed factors were (-1.47, 0.75) and (0, 0.7), respectively, and exponential (1) for the random factor intercept standard deviation. The model ran 8000 post-warmup samples spread across 4 chains (4000 iterations per chain including 2000 warmup iterations). Convergence was assessed using the GelmanâRubin statistic, with all parameters satisfying Râ1. Reduced and full models were compared using leave-one-out cross-validation with refitting to address high Pareto k values. The model with a larger ÎELPD was favored, and considered strongly credible if ÎELPD / SE â¥ 2. We evaluated planned contrasts with Bayesian posterior inference, considering an effect supported if the 95% credible interval for the odds ratio (OR) excluded 1. 25 Relative gene expression was analyzed using ÎÎCq(A) data in a linear mixed model that included factors listed above. Sampling order, which was randomly distributed across treatments, was found to be a significant confounding predictor of expression level, and was therefore added into reduced and full models as a fixed effect. A Type III Analysis of Variance test with Satterthwaite's method was performed on the full model. 2.3 Results 2.3.1 Seawater chemistry Dulse photosynthesis increased pH continuously under 23 days of continuous light in all seawater treatments (see Table 2 and Figure 2). The largest increase occurred in the ambient seawater treatment, which was consistently 0.5 pH units higher with dulse. Increases in OA and OA + upwelling treatments fluctuated between 0.2 and 0.4 pH units higher with dulse. Fluctuations in ÎpH were likely due to periodic dulse weighing and removal to reestablish a density of 10g/L (see Figure 2), water sampling events (see Figure 2), fluctuations in ambient light, and system cleaning events. Aragonite saturation state was affected by OA treatments (F(2, 11.54) = 81.91, p < 0.001), dulse (F(1, 11.55) = 66.14, p < 0.001), and sampling day (F(1, 33.0) = 4.34, p = 0.045). An OA treatment Ã dulse interaction was also detected (F(2, 11.54) = 4.47, p = 0.037). Across three weekly sampling points, dulse increased Ωarag by 0.77 ± 0.12 under the ambient treatment (p < 0.0001), by 0.62 ± 0.12 under the OA treatment (p < 0.001), and by 0.28 ± 0.12 under the OA + upwelling treatment (p < 0.05) (see Figure 3). 26 Figure 2. Delta pH between paired dulse absent/present tanks sharing a header tank during experiment week 1 and 3, recorded every 5 minutes (or periodically, represented by data points). Line and point color (green, yellow, red) represents OA treatment (ambient (pH ~7.8), OA (7.6), & OA + upwelling (7.4)), respectively. Dotted rectangle indicates the oyster experiment time period, solid bars indicate periodic dulse weighing and removal to reestablish a density of 10g/L, and striped bars indicate seawater sampling events.â«½ 27 Figure 3. Average Ωarag measured in dulse absent/present tanks on the same weekday and time across three weeks, on experiment days 3, 10, and 17 (see Figure 1). Circle color (green, yellow, red) represents CO2 treatment (ambient, OA, OA + upwelling, respectively). Transparent circles represent tanks without dulse, and filled circles represent tanks with dulse. Grey dots represent raw values. Circles below line at x = 1 represent unsaturated Ωarag; n = 3 replicates for each OA x dulse treatment combination 28 Table 2. Meter-measured pH ± SD on experiment day 2 in oyster beakers and carbonate chemistry parameters ± SD on day 3 in tanks with (+) or without dulse (â). AT and DIC were analyzed in water samples and the remaining parameters were calculated using CO2SYSv3.0 (Pierrot et al., 2021). For meter-measured pH in oyster beakers at additional timepoints, see Table 4 in Appendix; n = 3 replicates for each OA x dulse treatment combination 1Measured with a meter during 48 hpf larvae sampling 2Calculated using AT and DIC from water sample lab analysis 3High standard deviation and discrepancy between measured and calculated pH due to a measurement of 7.94 in one tank Treatment Oyster beakers Dulse absent/present tanks OA Dulse Measured pH day 21 Calculated pH day 32 pCOâ (µatm) AT (µmol/kg-SW) DIC (µmol/kg-SW) Ωarag Ωcalcite Ambient â 7.89 ± 0.04 7.9 ± 0.01 497 ± 26 1996.4 ± 3.9 1813.9 ± 16.6 1.48 ± 0.05 2.36 ± 0.08 + 8.05 ± 0.07 8.08 ± 0.07 311 ± 26 1998.7 ± 2.9 1878.2 ± 6.3 2.12 ± 0.14 3.38 ± 0.22 OA â 7.75 ± 0.173 7.64 ± 0.02 822 ± 46 2005.4 ± 1.6 1944.9 ± 4.63 0.99 ± 0.05 1.58 ± 0.08 + 7.83 ± 0.09 7.82 ± 0.08 529 ± 31 2004.2 ± 3.1 1893.0 ± 9.1 1.42 ± 0.07 2.26 ± 0.11 OA + upwelling â 7.58 ± 0.05 7.58 ± 0.02 1006 ± 91 2007.4 ± 0.6 1967.5 ± 9.4 0.85 ± 0.07 1.35 ± 0.11 + 7.69 ± 0.05 7.70 ± 0.02 874 ± 66 2005.6 ± 1.1 1952.5 ± 9.0 0.94 ± 0.06 1.49 ± 0.09 29 2.3.2 Oyster condition 2.3.2.1 Size OA and OA + upwelling treatments decreased larval oyster shell size. Dulse fully ameliorated the shell size decreases in all CO2 treatments and timepoints, and even increased shell size in ambient seawater (see Figure 4). Shell size was affected by OA treatments (ÏÂ²(2) = 15.7, p < 0.001), dulse (ÏÂ²(1) = 11.48, p < 0.001), and hpf (ÏÂ²(1) = 95.43, p < 0.001). No interaction was detected for OA treatment x dulse (ÏÂ²(2) = 2.0, p = 0.36). Figure 4. Estimated Marginal Means (EMMs) of oyster larvae shell diameter (µm) at two sampling timepoints: 24 and 48 hours post fertilization. Error bars indicate lower and upper 95% confidence limits. Circle color (green, yellow, red) represents CO2 treatment (ambient, OA, OA + upwelling, respectively). Transparent circles represent treatments without dulse, and filled circles represent treatments with dulse. Brackets show significant pairwise comparisons: â denotes 0.05 < p < 0.1 (marginally significant); * denotes p < 0.05; ** denotes p < 0.01; *** denotes p < 0.001. hpf: hours post fertilization; n = 3 oyster beaker replicates for each OA x dulse treatment combination 30 At 24 hpf, the estimated marginal means (EMMs) of oyster larvae were 7.7% and 5.8% smaller in the OA + upwelling treatment compared to the ambient and OA treatments, respectively (see Table 6 in Appendix for 95% confidence limits and p-values of the pairwise comparisons). A similar pattern occurred when dulse was present: larvae were 4.2% and 4.5% smaller in the OA + upwelling treatment compared to the ambient and OA treatments, respectively. Dulse increased larval size by 5.8% and 7.2% in the OA and OA+ upwelling treatments, respectively. A marginally significant increase of 3.2% was seen with dulse in the ambient treatment (p = 0.08). At 48 hpf, the EMMs of larvae in the OA + upwelling treatment were 7.7% and 5.8% smaller compared to the ambient and OA treatments, respectively. Dulse increased larval size by 5.8% and 7.2% in the OA and OA+ upwelling treatments, respectively. Although dulse fully mitigated OA effects on larval size, larvae with dulse in the OA + upwelling treatment were 4.2% and 4.5% smaller compared to the ambient and OA treatments with dulse, respectively. 2.3.2.2 Development and morphology OA and OA + upwelling treatments had no apparent effect on percent abnormal larvae with respect to development or morphology at any timepoint. Dulse decreased percent underdeveloped larvae and larvae with at least one hinge abnormality in the OA + upwelling treatment at one of the two sampling timepoints (see Figure 5). 31 Figure 5. Bar graphs of average percent abnormal larvae ± SE with respect to: (A) underdevelopment, (B) protruded tissue, and (C) larvae with at least one hinge abnormality (D and E). Graph panels are separated by hours post fertilization (hpf) with columns and type of morphology examined with rows. Bar color (green, yellow, red) represents ocean acidification condition (ambient, OA, and OA + upwelling treatments, respectively). Light bars represent larvae without dulse, and dark bars represent larvae with dulse. Black circles in top left panel corners show full models that outperformed null models (ÎELPD/ SE â¥ 2); n = 3 replicates for each OA x dulse treatment combination. C A B D E 32 At 14 hpf, the dulse x OA + upwelling treatment interaction effect on percent underdeveloped larvae was credibly negative, providing evidence that dulse decreased abnormal development in the OA + upwelling treatment to 0.6% ± 0.6% compared to 2.9% ± 2.9% in the ambient treatment (β = â1.46 ± 0.55, 95% CrI: â2.54 to â0.42; RÌ = 1.00). Furthermore, posterior inference testing within the OA + upwelling treatment showed that dulse credibly reduced the odds of underdeveloped larvae by ~51% (OR = 0.487, 95% CrI 0.399â0.741). Credible difference between treatments did not extend to 24 hpf, nor to percent larvae with protruded tissue at either timepoint. Although OA did not have a credible effect on percent larvae with a convex or rounded hinge, abnormal hinges tended to increase with OA intensity at 24 hpf. More specifically, the average percent with a convex hinge was 16.7% ± 16.7%, 26.2 ± 14.5%, and 40% ± 10% in the ambient, OA, and OA + upwelling treatments when dulse was absent, respectively. Similarly, the average percent with a rounded hinge was 8.3% ± 8.3%, 13.9% ± 7.3%, and 22.2% ± 14.7% in the ambient, OA, and OA + upwelling treatments without dulse, respectively. This tendency did not persist in the presence of dulse. Instead, posterior inference testing within the OA + upwelling treatment showed that dulse credibly reduced the odds of larvae with at least one hinge abnormality by ~51% (OR = 0.492, 95% CrI 0.403â0.748). Credible difference between treatments did not extend to 48 hpf. 33 2.3.3 Gene expression None of the CO2 conditions affected gene expression in larval oysters, but dulse presence induced slight shifts in expression of immune function genes (see Figure 6). Figure 6. Average change driven by dulse presence in relative gene expression ± SE across nine M. gigas target genes. Circle color (green, yellow, red) represents seawater treatment (ambient, OA, and OA + upwelling, respectively). Y-axis scale represents the difference in log2 transformed relative oyster gene expression between oysters in dulse absence and presence treatments paired with the same header tank. Grey dots represent raw Îlog2 values; * denotes significant pairwise comparison between treatments with and without dulse 34 The full NKA (1) expression model was not a better fit over the reduced, but exploratory pairwise comparisons of EMMs indicated that dulse presence tended to decrease NKA (1) expression in larval oysters by 20.5% (p = 0.083, marginally significant). There were no differences noted between treatments in expression of the other ion regulator genes NKA (2) and slc4 (3), or shell formation genes POU2F1, Sox5 and Pax6. The full Def expression model was not a better fit over the reduced, but exploratory model testing suggested a marginally significant interaction for OA treatment x dulse (F = 2.9, p = 0.96, Type III ANOVA). Under ambient and OA + upwelling treatments, dulse decreased average Def expression by almost 50%, whereas under the OA treatment, dulse increased average Def expression by over 100%. The full BPI expression model was a significantly better fit than the reduced (ÏÂ² = 5.7, p = 0.047). Model testing for BPI expression showed a marginally significant interaction between OA and dulse treatments (F = 5.7, p = 0.07, Type III ANOVA). Pairwise comparisons of EMMs showed that dulse increased BPI expression by 109.73% in the ambient treatment (p < 0.05). Gene expression was not affected by OA treatments when dulse was absent. 2.4 Discussion Seaweed mitigation of OA effects on adjacent organisms is a recent and growing area of research â most studies have been published within the past 5 years (Doo et al., 2020). Therefore, a limited number of species combinations and life stages have been examined. 35 This study adds insight into OA effect mitigation between a new species pair (Pacific oyster M. gigas and Pacific dulse D. mollis), and is the first to examine OA mitigation with seaweed in the larval stage of an OA-sensitive organism. Pacific dulse fully mitigated OA-induced decreases in Pacific oyster larvae size, and showed potential for affecting larvae development, morphology, and immune-related gene expression despite the lack of OA effects. 2.4.1 OA effects on oyster larvae In the absence of dulse, oyster size generally decreased with pH and Ωarag. As seen in previous studies, the OA + upwelling treatment (pH 7.4) decreased larvae size relative to the ambient treatment (Barros et al., 2013; Waldbusser et al., 2013; Watson et al., 2009). In contrast, no difference in size between ambient and OA (pH 7.6) treatments was apparent, unlike results from Barros et al. (2013) that showed a decrease in M. gigas larvae size under pH treatments of both 7.4 and 7.8. However, ambient pH was 8.1 in southern Portugal compared to the ambient pH of 7.8 along eastern Vancouver Island, Canada. In the present study, findings suggest that oyster larvae in BC adapt to local seawater conditions â including OA levels that mimic current local upwelling events â but canât always adapt to OA levels that are projected for future upwelling events. This aligns with previous work indicating that oysters along the western coast of North America can develop normally as long as Ωarag remains above 1, even at low pH or high pCOâ (Waldbusser et al., 2014). Oyster larvae development and morphology showed little response to OA treatments. This finding is in contrast with previous literature showing that OA increased abnormal development rates in larval oysters, mussels, and crabs, and increased morphological 36 abnormalities in larval oysters (Barros et al., 2013; Barton et al., 2012; Gazeau et al., 2010; Kurihara et al., 2007; Miller et al., 2016; Waldbusser et al., 2015; Watson et al., 2009). Again, this indicates that oysters in the present study exhibited some potential resilience to OA when other stressors such as disease were absent. However, nonsignificant trends suggested that hinge abnormalities may have increased as pH decreased. Across each development and morphological category in this study, the percentage of abnormal larvae tended to decrease as time passed. Perceived improvements in experimental populations were likely due to mortality of larvae in poor condition. For example, larvae may be particularly sensitive to OA when their first shell is forming around 24 hpf, as shown in Waldbusser et al. (2013). Early abnormalities may have led to mortality between sampling timepoints, while normal larvae that developed past 24 hpf may have partially adapted to OA conditions. However, whether abnormality declined across time due to the mortality or recovery is challenging to distinguish without reliable mortality findings. In the absence of dulse, no difference in gene expression was observed between OA conditions. In contrast, Wright-LaGreca et al. (2022) reported that OA increased expression of ion regulation gene NKA (1) in larval M. gigas. This discrepancy may reflect differences in experimental procedures, such as a lower Ωarag and/or sampling at a timepoint with higher OA sensitivity in the aforementioned study (Ωarag = 0.6 and sampling at 24 hpf), compared to the present study (Ωarag = 0.85 or 0.99 and sampling at 48 hpf). Consistent with the present findings, however, Wright-LaGreca et al. (2022) found no OA-induced changes in the expression of ion regulation genes NKA (2) and slc4 (3), nor 20 additional target genes. 37 OA did not alter the expression of shell formation genes targeted in this study (POU2F1, Sox5, Pax6), despite OA effects on larvae shell size and hinge abnormality trends. These genes were first characterized in Min et al. (2023) for their role in early shell establishment, and their expression is localized at the growing shellâs edge. Further validation of these genes and their effect on shell formation will help determine whether their expression levels are useful markers of successful growth and morphological development. OA also had no apparent effect on expression of immune response genes Def, Prp, and BPI, which differs from previous findings on adult M. gigas showing that OA downregulated expression in the gill and upregulated expression in the digestive gland, leading authors to link OA with immune dysfunction (Q. Wang et al., 2016). Targeted sampling of immune organs in adults may have captured gene expression more accurately; the absence of detectable OA effects in the present study may reflect tissue-specific expression in larvae that is diluted when whole larvae are sampled. 2.4.2 Dulse effects on OA The amelioration of low pH, high pCOâ, and undersaturated Ωarag by seaweed during daylight has previously been documented, with effects often extending into nighttime hours (Z. Jiang et al., 2014; Leal et al., 2024; Wahl et al., 2018; Young & Gobler, 2018). However, diel fluctuations in photosynthesis did not apply to this study, as dulse was held under continuous light and oysters were held in static cultures that received daily water changes from dulse absent/present tanks. Here the data showing dulse amelioration of low pH, high pCOâ, and 38 undersaturated Ωarag included evidence of previously undocumented, continuous 24-hr low pH amelioration extending for at least three weeks. The magnitude of dulse-induced carbonate chemistry change was lowest in the OA + upwelling condition and highest in the ambient condition, which aligns with previous findings indicating that seaweed may become less effective at ameliorating undersaturated Ωarag as OA intensifies (Leal et al., 2024). However, considering that dulse mitigated OA effects in even the most extreme OA condition, and ameliorated undersaturated Ωarag as low as 0.5 in particular replicates (see Figure 3), OA conditions may never reach levels that would render dulse ineffective at OA mitigation. Indeed, calcifier growth depends on maintaining Ωarag above 1 (Hurd et al., 2020), and in almost all cases the Ωarag fell above 1 when dulse was present. Interestingly, the magnitude of dulse-induced change in carbonate chemistry increased in all seawater treatments as the experiment progressed, suggesting that variables other than level of pCOâ may impact the effect of dulse on water chemistry. Such variables may include change in water temperature, or the microscopic epiphyte community on the dulse, driven by shifts in the incoming seawater. 2.4.3 Dulse effect on larvae Dulse presence increased oyster larvae size under all seawater treatments. These findings align with previous literature on size increase in juvenile and adult shellfish when adjacent to seagrass, kelp, or dulse (Hamilton et al., 2022; Peabody et al., 2019; Ricart, Gaylord, et al., 2021). Results also showed that dulse fully mitigated negative effects of OA and OA + upwelling conditions, increasing size to or above that of the ambient treatment at both 24 and 39 48 hpf, which aligns with findings on seaweed mitigation of OA size or growth effects on calcifying plankton and juvenile and adult bivalves (Doo et al., 2020; Z. Jiang et al., 2014; Wahl et al., 2018; Young et al., 2022; Young & Gobler, 2018). However, the benefit of seaweed on size in the ambient treatment shown here is not always seen in other studies (Doo et al., 2020; Young et al., 2022). Results from the present study showing benefits of dulse on larvae size even under the ambient seawater treatment suggest that the seaweed not only promotes normal growth through altering seawater carbonate chemistry, but potentially through indirect mechanisms as well. Alternative modes of action could include the dulse-driven increase in dissolved oxygen (DO): the level of DO available to bivalves is closely linked to their energy budgets, and therefore, their scope for growth (Sui et al., 2016). Low DO can negatively affect bivalve growth and survival, particularly when combined with OA (Gobler et al., 2014; Gu et al., 2019; Y. Wang et al., 2011). Further research may elucidate whether high DO from seaweed presence could, alternatively, benefit bivalve growth and survival. Positive effects of dulse presence extended to larvae development and morphology. Despite the lack of clear OA effects, dulse presence decreased the percentage of larvae that were underdeveloped or had 1-2 hinge abnormalities in the OA + upwelling condition. Trends showed that larvae in the presence of dulse were either as normal or more normal in the OA + upwelling condition compared to the ambient condition with respect to development and shell morphology. This is the first study on the effect of seaweed presence on shellfish 40 larvae, but similar findings from an experiment with urchin larvae showed that seagrass presence decreased abnormalities under low pH (Ravaglioli et al., 2024). Neither OA nor dulse presence had an effect on percent larvae with protruded tissue. Bacterial infection can often lead to protruded tissue in oyster larvae (Estes et al., 2004); therefore, findings suggest that neither OA, dulse, nor dulse epiphytes were creating visually apparent differences in inflammation response. However, conditions in the present study did not include a pathogen challenge, so treatment effects on oyster disease susceptibility remain unclear. OA has been known to suppress immune response pathways in larval M. gigas (Dineshram et al., 2021), and effects of seaweed presence on adult M. gigas disease susceptibility range from beneficial to detrimental, depending on seaweed species (Dugeny et al., 2022). Therefore, future research that includes a pathogen challenge is necessary to evaluate the potential for dulse to mitigate OA-induced immune suppression. In a few instances, dulse presence induced ambiguous or negative changes to oyster larvae. At 48 hpf, dulse significantly upregulated expression of immune response gene BPI in the ambient seawater treatment. BPI has been linked to initial oyster defense against bacterial infection, with consistent levels of background expression (Gonzalez et al., 2007). The dulse-induced upregulation in BPI expression could be in response to some sort of shift in the environmental or internal microbiome that only occurred when dulse was present in the ambient treatment, or could be a sign that dulse could dysregulate oyster immune response. In Mackenzie et al., (2022), buffering seawater with soda ash during M. gigas larval stages led to disease susceptibility in the subsequent juvenile stage. One parallel with the present 41 study when dulse was present in the ambient condition was an unnaturally high Ωarag compared to the control. Further investigation would be necessary to determine whether a high Ωarag compared to regional conditions may affect locally adapted larval oysters. Such Ωarag conditions may also relate to the trend seen at 24 hpf, though non-significant, suggesting that dulse increased percent larvae with a convex hinge in the ambient treatment. Future directions of OA mitigation research would benefit from investigating a potential optimal Ωarag range for Pacific Northwest bivalves that incorporates Ωarag > 1 as the previously-established lower threshold of the range (Waldbusser et al., 2014). If an optimal Ωarag range is quantified for M. gigas larvae, benefits of dulse presence in a hatchery could be maximized by modulating seaweed density or light intensity, depending on the carbonate chemistry of incoming seawater. In coastal ecosystems, stewarding seaweed habitat as an OA refuge does not involve the risk of exceeding any potential Ωarag ceiling for shellfish, as seaweed draws Ωarag down at night during respiration. Conversely, OA mitigation strategies such as OAE could overshoot a potential optimal carbonate chemistry range for shellfish; if an upper threshold exists for Ωarag, OAE would risk surpassing it. 2.4.4 Larger context With wild fisheries continuing to decline â in part due to climate change â and finfish monoculture challenged politically across multiple continents, a niche is opening with potential for growth in seaweed and shellfish farm industries (Ertör & Ortega-Cerdà , 2015; Lam et al., 2016; Weitzman & Bailey, 2019). Bearing in mind that climate change also impacts shellfish farming, studies like the present highlight avenues of OA mitigation that 42 will aid in future success (Barton et al., 2015; Clements & Chopin, 2017). These findings encourage examining aquaculture cultivation strategies that integrate seaweed and shellfish together, such as hatchery co-culture, integrated multi-trophic aquaculture, and traditional clam gardens (Cox et al., 2024). Merits of farming seaweed in land-based shellfish hatcheries could extend beyond OA mitigation to include farm product diversification or supplementation of labor-intensive shellfish larvae diets with single cell detritus derived from the seaweed (Alberto et al., 2020). Benefits of farming seaweed in open-ocean shellfish farms extend beyond OA mitigation to include nutrient recycling, reduction of hypoxia, dampening of wave energy, protection of shorelines, ocean carbon removal, and habitat enhancement for wild species (Chopin, 2013; Chopin et al., 2012; Duarte et al., 2017; Troell et al., 2009; Zhu et al., 2025). This study highlights the value of seaweed presence independent of the debate over whether macroalgae contribute to Blue Carbon. Consideration of seaweeds within the context of ecosystem resilience to climate change should extend beyond the complex question of whether or not carbon fixed by seaweed photosynthesis is sequestered long-term (Hurd et al., 2022; Pessarrodona et al., 2023). Evaluating the species-specific potential for seaweed to provide an OA refuge in a coastal ecosystem is an important consideration: in the field, kelp has been shown to benefit nearby shellfish (Peabody et al., 2019), but fucus has not been seen to increase growth in nearby mussel growth (Wahl et al., 2018). As dulse inhabits a similar ecological niche to fucus, it may show a similar lack of benefit to adjacent shellfish outside of an aquaculture context. However, since the pH in all compared microhabitats was ~ 8.0, OA conditions may have not been the limiting factor for mussel growth in Wahl et al., 43 (2018). As OA represents only one aspect of climate change, it is also important to consider interactive effects of OA and other climate change stressors; for example, not only can ocean warming directly exacerbate effects of OA on many marine organisms, it could also indirectly exacerbate effects by threatening OA refuges through negatively impacting seaweed (Jian et al., 2025). Although findings here only apply to seawater chemistry affected by seaweed under daylight conditions, OA buffering by seaweed has been shown to extend into hours of darkness, and some shellfish can even upregulate calcification during hours when seaweed is photosynthesizing (Wahl et al., 2018). Coastal habitats in which OA conditions can be exacerbated due to natural patterns in carbonate chemistry such as those closer to the poles are also areas with long daylight hours in spring and summer (L.-Q. Jiang et al., 2015). Therefore, organisms with life history stages sensitive to OA that may occur in spring or summer may benefit from seaweed OA refuges, particularly in intertidal zones with low rates of seawater exchange. For example, both Atlantic and Pacific herring spawn, lay eggs, and develop across larval stage in intertidal areas with upwelling at higher latitudes when days are longer, and are sensitive to OA (Frommel et al., 2014). Therefore, some herring may be particularly positioned to benefit from OA refuges provided by seaweed amid multiple anthropogenic stressors including OA, especially because some herring eggs are laid directly on the surface of seaweed (Geffen, 2009). The potential for crossover between seaweed OA refuges and sensitive larvae is expansive due to the range in OA effect parameters and affected species such as embryonic and larval 44 mortality, development, larval swimming ability, settlement and post-settlement processes, and organ damage in shellfish, echinoderms, fish, arthropods, cnidarians, and bryozoans (Espinel-Velasco et al., 2018). Effects can even begin before larval stages, such as during sex determination, gametogenesis, reproductive timing, mating behavior, fertilization, and fecundity of global marine invertebrates across multiple phyla (Padilla-Gamiño et al., 2022). Bearing this in mind, ecological modeling of climate change effects and coastal restoration planning would benefit from considering seaweedsâ capacity for providing OA refuges regardless of how small their spatial and time scales may be, as such scales may occur at inflection points for organism and population success. Findings from the present study provide incentive for seaweed restoration and stewardship as a leading OA mitigation strategy instead of extractive, potentially harmful, and less ecosystem-based strategies such as ocean alkalinity enhancement: most alkaline material production requires mining, and buffering seawater may impair marine organisms (Eisaman et al., 2023; Mackenzie et al., 2022). 2.5 Conclusion The magnitude of benefit that seaweed provides with regard to OA is debated, especially when the conversation revolves around whether seaweed is a key source of Blue Carbon. However, the importance of seaweed as OA intensifies may be more nuanced than carbon sequestration, but no less significant at ecosystem-level scales. These findings provide evidence that proximity to dulse provides advantages to M. gigas larvae, such as providing an OA refuge. These advantages may extend to other calcifiers and organisms sensitive to OA that naturally share habitat space with seaweed or can be cultivated together in sea farms or 45 land-based aquaculture. These results highlight the importance of considering seaweed during habitat stewardship and future OA ecosystem effect modeling, for reasons other than carbon credits. Every organism shielded from OA by proximity to seaweed, even over short-term or seasonal scales, has a better chance for climate change resilience. When such interactions occur at organism inflection points and are scaled up to the ecosystem level, effects of seaweed presence may be detectable not only in the chemistry of the seawater flowing through but in the long-term success of organisms and populations within an ecosystem. 46 Chapter 3: Conclusions 3.1 Overview of findings Dulse presence strongly influenced seawater carbonate chemistry, decreasing pCOâ and increasing pH and Ωarag across all experimental treatments. These shifts indicate that dulse can counteract unfavorable carbonate conditions, including Ωarag undersaturation that are otherwise stressful for calcifying organisms. In treatments without dulse, oyster development and shell morphology showed little difference between ambient and acidified conditions. This pattern suggests that larvae may tolerate certain levels of OA when additional stressors such as pathogen outbreaks or heat stress are absent, or alternatively, that all treatments imposed comparable levels of physiological challenge. The lack of strong responses to OA, particularly under the moderate OA treatment (pH 7.6), aligns with earlier findings showing that oysters can often maintain normal development when Ωarag remains above one, even under elevated pCOâ (Waldbusser et al., 2014). Larval oysters consistently showed improved condition when dulse was present, reflected in enhanced growth, development, and shell morphology, along with subtle shifts in immune gene expression. Dulse fully offset the negative impact of OA on larval size, and most oysters co-cultured with dulse under both OA and OA + upwelling conditions matched or exceeded the performance of those in ambient seawater without dulse. In several cases, even larvae in ambient seawater benefited from dulse relative to those in dulse-free ambient treatments. These patterns point to two possibilities: that current ambient chemistry may already place larvae near the edge of their tolerance range, and/or that dulse provides benefits extending beyond OA mitigation. The importance of such potential benefits becomes 47 increasingly more relevant as ocean acidification continues to progress beyond levels that shellfish populations can naturally withstand. 3.2 Limitations of the research The lack of reliable data on oyster density and mortality detracted from understanding whether decreases in abnormal larvae across time were due to individuals adapting to conditions, or just the mortality of abnormal larvae. The study also only provided a snapshot of effects, and did not provide insight into longer term effects of seaweed presence on oyster larvae. One caveat when applying these findings to shellfish hatchery practices is that this study maintained oyster larvae in static culture, whereas commercial hatcheries typically employ flow-through systems. However, dulse was in a flow-through system, so findings on changes to water chemistry due to dulse presence are very applicable to industry. Carbonate chemistry parameters were analyzed in water samples from the dulse absent/present tanks, which supplied water changes to the oyster beakers, but beaker volumes were insufficient for water sampling. Therefore, although meter-measured pH in beakers stayed relatively close to target treatment values, carbonate chemistry parameters in beakers could have undergone undetected shifts that affected the oyster larvae. Future research would benefit from pooling seawater within treatment replicates for water sample analysis that would provide a general idea of beaker carbonate chemistry profiles. 48 A limitation in applying these findings to coastal habitats is that the dulse was maintained under continuous light rather than a natural dayânight cycle, and, as noted previously, the oysters were held in static culture. Fluctuations in carbonate chemistry parameters due to diel light cycles would greatly affect OA amelioration by seaweed and the resulting OA effect mitigation for oyster larvae. However, organisms from intertidal areas are usually quite adapted to large fluctuations in seawater parameters (Sanford & Kelly, 2011), and a bivalve has previously been shown to shift calcification to daylight hours only when in the presence of seaweed (Wahl et al., 2018). Furthermore, many OA-sensitive life stages of organisms within high-latitude Northern Pacific coastal habitats, such as the farmed Pacific oyster larval stage, occur during the summer when extended daylength allows for prolonged photosynthesis. Gene expression data would have benefited from more replicates or more larvae overall, as RT-qPCR is challenging for larval oysters. Due to their microscopic size, a large population is necessary for analysis. The oyster populations in experimental beakers were only just sufficient for one RNA extraction, leaving no replicate samples available in the event that any step of the extraction or RT-qPCR process was suboptimal. Luckily, gene expression analysis was able to move forward using high quality RNA, but n = 3 replicates all receiving seawater from different tanks led to relative expression data with high between-replicate variability. 49 3.3 Contributions to the field Marine algae mitigation of OA effects on calcifying organisms is a recent and growing area of research â most studies have been published within the past 5 years (Doo et al., 2020). Therefore, a limited number of species combinations and life stages have been examined. This study adds insight into OA effect mitigation between a new species pair (Pacific oyster M. gigas & Pacific dulse D. mollis), and is the first to examine OA mitigation with seaweed in the larval stage of an OA-sensitive organism. 3.3.1 Restoration applications This study can also apply to shellfish restoration efforts that include the rearing of threatened organisms through sensitive life stages before outplantation, such as the Pinto abalone along the west coast of North America (Bates et al., 2025; Sowul et al., 2022). Dulse seaweed may be particularly applicable in this context for OA effect mitigation, as it is fed to juvenile and adult abalone and therefore already grown at abalone rearing facilities (Hamilton et al., 2022). The red seaweed has also already been shown to mitigate effects of OA on farmed juvenile abalone (Hamilton et al., 2022). Efforts in seaweed restoration and stewardship in coastal habitats could also bolster OA-sensitive organism populations. Kelp has the capacity to alter carbonate chemistry in ocean habitats and benefit adjacent organisms on short-term and seasonal scales (Frieder et al., 2012; Koweek et al., 2017; Peabody et al., 2019). These scales may align with periods of sensitivity within the lifecycle of kelp forest organisms, such as spawning events and larval development during summer months, and therefore kelp density could in theory influence the 50 success of entire generations of species within a habitat through short-term seawater chemistry alterations. 3.3.2 Shellfish hatchery applications Land-based co-culture studies can benefit shellfish hatcheries both in industry and restoration spheres. The present findings are the first with application potential for shellfish hatcheries in the aquaculture industry that are already impacted by OA and other climate-driven stressors. OA mitigation through seaweed co-culture may be more environmentally sustainable than using alkaline materials (e.g., soda ash); seaweed is a regenerative resource, whereas soda ash derives from the extractive process of mining brines or trona ore (USGS, 2023). Considering the risk to oyster health of buffering seawater with soda ash (Mackenzie et al., 2022), shellfish yield may increase in hatcheries contending with OA if seaweed could buffer the seawater instead. Seaweed-buffered water has even been shown to change disease resistance by altering the larval microbiome, depending on the type of seaweed (Dugeny et al., 2022). In a co-culture system, seaweed may also benefit: shellfish could have fertilizing effects on seaweed through nitrogen production in their waste (Hargrave et al., 2021). Currently some farmed seaweed is fertilized with inorganic fertilizers, contributing to production costs (Ogawa & Fujita, 1997). Seaweed co-culture could also provide an additional income source to hatcheries through crop diversification. 3.3.3 Context within the seafood industry The production of sustainable seafood is a rising focus of debate, with the rise of climate change stressors like OA that are both a product of and a risk to the food industry (Froehlich 51 et al., 2022). A sharp decline in fish populations worldwide has been forecasted, with 35% loss in annual fisheries revenue predicted by 2050 from climate change alone (Lam et al., 2016). Alternatives to wild-caught seafood include finfish net pen farming, but fish monoculture is politically challenged in multiple countries (Ertör & Ortega-Cerdà , 2015; Weitzman & Bailey, 2019). Shellfish aquaculture has the potential to supply healthy and climate-friendly food products while also alleviating pressure on wild populations (Peabody et al., 2019), but is also threatened by climate change. This study underlines the potential for integrated aquaculture with seaweed and shellfish as an OA-adapted seafood production technique as wild fish populations decline and OA intensifies. Benefits of seaweed as a crop extend past their role in combating OA impacts: they require no freshwater or pesticides and little fertilizer or land to thrive, and extract excess nutrients from seawater they inhabit (Xiao et al., 2017). This makes them an ideal candidate for regenerative aquaculture, removing excess CO2 from the water and replacing it with oxygen, while providing nutritious food for humans and other organisms. When farmed in the ocean, the physical structure of seaweed and bivalve communities have the potential to reduce wave action, stabilize sediment, and provide habitat for wild species with fisheries value (Barrett et al., 2022). Ocean warming is causing declines in algal productivity in many regions (Jian et al., 2025), but the same upwelling events that exacerbate OA also bring cold and nutrient-rich water into coastal waters that seaweed thrives in (Feely et al., 2008; Reum et al., 2016). Therefore, coastal habitats with upwelling, such as the pacific northwest of North America where this study was conducted, are ideal candidates for OA amelioration through seaweed farming and restoration. Despite the myriad of benefits that seaweed and 52 bivalve farming provides, and the potential for seaweed farms to alleviate OA stress on other commercially important species, research such as this study on mitigation effects of seaweed on marine calcifiers remains scarce. 3.4 Future directions of research According to the 2012 Blue Ribbon Panel, we must look to phytoremediation strategies such as algae farming as first step in combating OA in the future (Peabody et al., 2019). The benefits of seaweed with regard to ocean acidification (OA) are debated when the conversation revolves around seaweed sequestering carbon on a global scale. However, the importance of seaweed as OA intensifies may be more nuanced, but no less significant, than large-scale carbon chemistry. These findings provide evidence that proximity to dulse provides significant advantages to Pacific oyster larvae in both current and projected seawater conditions. These advantages may extend to other calcifiers and organisms sensitive to OA in both industry and restoration contexts. Long-term comparisons of the compatibility of particular seaweeds and other co-cultured organisms will be important in the context of seaweed OA mitigation. Outside of the OA context, impact of seaweed on the oyster microbiome and immune function can range from beneficial to detrimental, depending on seaweed species (Dugeny et al., 2022). The long-term effects of dulse presence on Pacific oyster larvae may warrant further analysis in the event that delayed negative impacts on the microbiome and related immune function occur such as with soda ash buffering (Mackenzie et al., 2022). The microbiome of microalgal feed has also been found to greatly influence the larval microbiome (Vignier et al., 2021); therefore, 53 longer-term studies that include microbiome analysis would allow for an examination of potential interactive effects between microalgal feed and seaweed co-culture on oyster larvae microbiome. Examining potential advantages of seaweed presence on larval stages of other organisms may continue to emphasize the importance of even short-term or diel OA refuges. Investigating potential in-situ OA refuges represents a promising research direction, particularly in environments such as tide pools, sheltered bays, and sounds with limited water movement, and where seaweed and associated organisms occur in close proximity. When such interactions are scaled up to the ecosystem level, effects of seaweed presence may not always be detectable in the chemistry of the seawater flowing through but in the long-term success of calcifier populations and organisms that rely on them. 54 References Alberto, P.-R., Gabriela, M.-A., Regina, E.-G., Gabriela, M.-C., Dariel, T.-R., & Cristina, E.-F. (2020). Seaweed single cell detritus effects on the digestive enzymes activity and microbiota of the oyster Crassostrea gigas. Journal of Applied Phycology, 32(5), 3481â3493. https://doi.org/10.1007/s10811-020-02167-4 Barrett, L. T., Theuerkauf, S. J., Rose, J. M., Alleway, H. K., Bricker, S. B., Parker, M., Petrolia, D. R., & Jones, R. C. (2022). Sustainable growth of non-fed aquaculture can generate valuable ecosystem benefits. Ecosystem Services, 53, 101396. https://doi.org/10.1016/j.ecoser.2021.101396 Barros, P., Sobral, P., Range, P., Chícharo, L., & Matias, D. (2013). Effects of sea-water acidification on fertilization and larval development of the oyster Crassostrea gigas. Journal of Experimental Marine Biology and Ecology, 440, 200â206. https://doi.org/10.1016/j.jembe.2012.12.014 Barry, J. P., Lovera, C., Buck, K. R., Peltzer, E. T., Taylor, J. R., Walz, P., Whaling, P. J., & Brewer, P. G. (2014). Use of a Free Ocean CO2 Enrichment (FOCE) System to Evaluate the Effects of Ocean Acidification on the Foraging Behavior of a Deep-Sea Urchin. Environmental Science & Technology, 48(16), 9890â9897. https://doi.org/10.1021/es501603r Barton, A., Hales, B., Waldbusser, G. G., Langdon, C., & Feely, R. A. (2012). The Pacific oyster, Crassostrea gigas, shows negative correlation to naturally elevated carbon dioxide levels: Implications for nearâterm ocean acidification effects. Limnology and Oceanography, 57(3), 698â710. 55 Barton, A., Waldbusser, G., Feely, R., Weisberg, S., Newton, J., Hales, B., Cudd, S., Eudeline, B., Langdon, C., Jefferds, I., King, T., Suhrbier, A., & McLauglin, K. (2015). Impacts of Coastal Acidification on the Pacific Northwest Shellfish Industry and Adaptation Strategies Implemented in Response. Oceanography, 25(2), 146â159. https://doi.org/10.5670/oceanog.2015.38 Bates, E. H., Crim, R. N., Bouma, J. V., Toft, J., & Padilla-Gamiño, J. L. (2025). Exposure of larval pinto abalone to ocean acidification and warming negatively impacts survival, settlement, and size. Scientific Reports, 15(1), 15797. https://doi.org/10.1038/s41598-025-96605-3 Bednaršek, N., Feely, R. A., Beck, M. W., Alin, S. R., Siedlecki, S. A., Calosi, P., Norton, E. L., Saenger, C., Å trus, J., & Greeley, D. (2020). Exoskeleton dissolution with mechanoreceptor damage in larval Dungeness crab related to severity of present-day ocean acidification vertical gradients. Science of The Total Environment, 716, 136610. Bednaršek, N., Feely, R. A., Howes, E. L., Hunt, B. P. V., Kessouri, F., León, P., Lischka, S., Maas, A. E., McLaughlin, K., Nezlin, N. P., Sutula, M., & Weisberg, S. B. (2019). Systematic Review and Meta-Analysis Toward Synthesis of Thresholds of Ocean Acidification Impacts on Calcifying Pteropods and Interactions With Warming. Frontiers in Marine Science, 6, 227. https://doi.org/10.3389/fmars.2019.00227 Bednaršek, N., Feely, R., Tolimieri, N., Hermann, A., Siedlecki, S., Waldbusser, G., McElhany, P., Alin, S., Klinger, T., & Moore-Maley, B. (2017). Exposure history determines pteropod vulnerability to ocean acidification along the US West Coast. Scientific Reports, 7(1), 1â12. 56 Bednaršek, N., van de Mortel, H., Pelletier, G., García-Reyes, M., Feely, R. A., & Dickson, A. G. (2025). Assessment framework to predict sensitivity of marine calcifiers to ocean alkalinity enhancement â identification of biological thresholds and importance of precautionary principle. Biogeosciences, 22(2), 473â498. https://doi.org/10.5194/bg-22-473-2025 Bergstrom, E., Silva, J., Martins, C., & Horta, P. (2019). Seagrass can mitigate negative ocean acidification effects on calcifying algae. Scientific Reports, 9, 1932. https://doi.org/10.1038/s41598-018-35670-3 Bhuyan, Md. S., Islam, Md. N., Sharif, A. S. M., Islam, Md. M., Islam, Md. S., & Hoq, Md. E. (2021). Seaweed: A Powerful Tool for Climate Change Mitigation That Provides Various Ecological Services. In Md. N. Islam & A. van Amstel (Eds.), Bangladesh II: Climate Change Impacts, Mitigation and Adaptation in Developing Countries (pp. 159â192). Springer International Publishing. https://doi.org/10.1007/978-3-030-71950-0_5 Bressan, M., Chinellato, A., Munari, M., Matozzo, V., Manci, A., Marceta, T., Finos, L., Moro, I., Pastore, P., Badocco, D., & Marin, M. G. (2014). Does seawater acidification affect survival, growth and shell integrity in bivalve juveniles? Mar Environ Res, 99, 136â148. Medline. https://doi.org/10.1016/j.marenvres.2014.04.009 Caillon, C., Pernet, F., Lutier, M., & Di Poi, C. (2023). Differential reaction norms to ocean acidification in two oyster species from contrasting habitats. Journal of Experimental Biology, 226(23), jeb246432. https://doi.org/10.1242/jeb.246432 CBC. (2014). Acidic ocean deadly for Vancouver Island scallop industry. 57 Chopin, T. (2013). Aquaculture, Integrated Multi-trophic (IMTA). In Encyclopedia of Sustainability Science and Technology (Vol. 12, pp. 542â564). https://doi.org/10.1007/978-1-4614-5797-8_173 Chopin, T., Cooper, J. A., Reid, G., Cross, S., & Moore, C. (2012). Open-water integrated multi-trophic aquaculture: Environmental biomitigation and economic diversification of fed aquaculture by extractive aquaculture. Reviews in Aquaculture, 4(4), 209â220. https://doi.org/10.1111/j.1753-5131.2012.01074.x Clark, J. (2022). Dulse [Personal communication]. Clements, J. C., & Chopin, T. (2017). Ocean acidification and marine aquaculture in North America: Potential impacts and mitigation strategies. Reviews in Aquaculture, 9(4), 326â341. Cox, K. D., Davies, H. L., Millard-Martin, B., Black, M., Hessing-Lewis, M., Smith, N. F., Juanes, F., & Dudas, S. E. (2024). Ancestral and contemporary intertidal mariculture practices support marine biodiversity in the northeast Pacific. Communications Earth & Environment, 5(1), 351. https://doi.org/10.1038/s43247-024-01515-3 Crook, E. D., Cohen, A. L., Rebolledo-Vieyra, M., Hernandez, L., & Paytan, A. (2013). Reduced calcification and lack of acclimatization by coral colonies growing in areas of persistent natural acidification. Proceedings of the National Academy of Sciences of the United States of America, 110(27), 11044â11049. https://doi.org/10.1073/pnas.1301589110 Demetropoulos, C. L., & Langdon, C. J. (2004a). Enhanced production of Pacific dulse (Palmaria mollis) for co-culture with abalone in a land-based system: Nitrogen, 58 phosphorus, and trace metal nutrition. Aquaculture, 235(1â4), 433â455. https://doi.org/10.1016/j.aquaculture.2003.09.010 Demetropoulos, C. L., & Langdon, C. J. (2004b). Pacific dulse (Palmaria mollis) as a food and biofilter in recirculated, land-based abalone culture systems. Aquacultural Engineering, 32(1), 57â75. Dickson, A. G. (1990). Thermodynamics of the dissociation of boric acid in synthetic seawater from 273.15 to 318.15 K. Deep Sea Research Part A. Oceanographic Research Papers, 37(5), 755â766. Dineshram, R., Xiao, S., Ko, G. W. K., Li, J., Smrithi, K., Thiyagarajan, V., Zhang, Y., & Yu, Z. (2021). Ocean Acidification Triggers Cell Signaling, Suppress Immune and Calcification in the Pacific Oyster Larvae. Frontiers in Marine Science, 8. https://doi.org/10.3389/fmars.2021.782583 Dittrich, M. C., Meyer, L. A., Stekoll, M. S., Kelley, A. L., & Umanzor, S. (2025). Indoor cultivation protocols for Devaleraea mollis and Palmaria hecatensis from Alaska: Effects of temperature, light, and nutrients. Journal of Applied Phycology. https://doi.org/10.1007/s10811-025-03594-x Doo, S. S., Leplastrier, A., Graba-Landry, A., Harianto, J., Coleman, R. A., & Byrne, M. (2020). Amelioration of ocean acidification and warming effects through physiological buffering of a macroalgae. Ecol Evol, 10(15), 8465â8475. PubMed-not-MEDLINE. https://doi.org/10.1002/ece3.6552 Duarte, C. M., Wu, J., Xiao, X., Bruhn, A., & Krause-Jensen, D. (2017). Can seaweed farming play a role in climate change mitigation and adaptation? Frontiers in Marine Science, 4, 100. 59 Dugeny, E., de Lorgeril, J., Petton, B., Toulza, E., Gueguen, Y., & Pernet, F. (2022). Seaweeds influence oyster microbiota and disease susceptibility. J Anim Ecol, 91(4), 805â818. Medline. https://doi.org/10.1111/1365-2656.13662 Eisaman, M. D., Geilert, S., Renforth, P., Bastianini, L., Campbell, J., Dale, A. W., Foteinis, S., Grasse, P., Hawrot, O., Löscher, C. R., Rau, G. H., & Rønning, J. (2023). Assessing the technical aspects of ocean-alkalinity-enhancement approaches. State of the Planet, 2-oae2023, 1â29. https://doi.org/10.5194/sp-2-oae2023-3-2023 Ertör, I., & Ortega-Cerdà , M. (2015). Political lessons from early warnings: Marine finfish aquaculture conflicts in Europe. Marine Policy, 51, 202â210. https://doi.org/10.1016/j.marpol.2014.07.018 Espinel-Velasco, N., Hoffmann, L., Agüera, A., Byrne, M., Dupont, S., Uthicke, S., Webster, N., & Lamare, M. (2018). Effects of ocean acidification on the settlement and metamorphosis of marine invertebrate and fish larvae: A review. Marine Ecology Progress Series, 606, 237â257. https://doi.org/10.3354/meps12754 Estes, R., Friedman, C., Elston, R., & Herwig, R. (2004). Pathogenicity testing of shellfish hatchery bacterial isolates on Pacific oyster Crassostrea gigas larvae. Diseases of Aquatic Organisms, 58, 223â230. https://doi.org/10.3354/dao058223 Evans, W., Pocock, K., Hare, A., Weekes, C., Hales, B., Jackson, J., Gurney-Smith, H., Mathis, J. T., Alin, S. R., & Feely, R. A. (2019). Marine CO2 Patterns in the Northern Salish Sea. Frontiers in Marine Science, 5. https://doi.org/10.3389/fmars.2018.00536 FAO. (2004). The hatchery culture of bivalves: A practical manual. https://www.fao.org/4/y5720e/y5720e00.htm 60 FAO. (2025). Magallana gigas. Cultured Aquatic Species Information Programme. Food and Agriculture Organization of the United Nations. https://www.fao.org/fishery/en/culturedspecies/magallana_gigas Feely, R. A., Sabine, C. L., Hernandez-Ayon, J. M., Ianson, D., & Hales, B. (2008). Evidence for Upwelling of Corrosive âAcidifiedâ Water onto the Continental Shelf. Science, 320(5882), 1490â1492. https://doi.org/10.1126/science.1155676 Frieder, C., Nam, S., Martz, T., & Levin, L. (2012). High temporal and spatial variability of dissolved oxygen and pH in a nearshore California kelp forest. Biogeosciences, 9(10), 3917â3930. Friedlingstein, P., Jones, M. W., OâSullivan, M., Andrew, R. M., Hauck, J., Peters, G. P., Peters, W., Pongratz, J., Sitch, S., Le Quéré, C., Bakker, D. C. E., Canadell, J. G., Ciais, P., Jackson, R. B., Anthoni, P., Barbero, L., Bastos, A., Bastrikov, V., Becker, M., â¦ Zaehle, S. (2019). Global Carbon Budget 2019. Earth System Science Data, 11(4), 1783â1838. https://doi.org/10.5194/essd-11-1783-2019 Froehlich, H. E., Koehn, J. Z., Holsman, K. K., & Halpern, B. S. (2022). Emerging trends in science and news of climate change threats to and adaptation of aquaculture. Aquaculture, 549, 737812. https://doi.org/10.1016/j.aquaculture.2021.737812 Frommel, A. Y., Akbarzadeh, A., Chalifoux, V., Ming, T. J., Collicutt, B., Rolheiser, K., Opie, R., Miller, K. M., Brauner, C. J., & Hunt, B. P. V. (2025). High sensitivity to ocean acidification in wild out-migrating juvenile Pacific salmon is not impacted by feeding success. Ecological Applications, 35(5), e70058. https://doi.org/10.1002/eap.70058 61 Frommel, A. Y., Maneja, R., Lowe, D., Pascoe, C. K., Geffen, A. J., Folkvord, A., Piatkowski, U., & Clemmesen, C. (2014). Organ damage in Atlantic herring larvae as a result of ocean acidification. Ecological Applications: A Publication of the Ecological Society of America, 24(5), 1131â1143. https://doi.org/10.1890/13-0297.1 Frommel, A. Y., Margulies, D., Wexler, J. B., Stein, M. S., Scholey, V. P., Williamson, J. E., Bromhead, D., Nicol, S., & Havenhand, J. (2016). Ocean acidification has lethal and sub-lethal effects on larval development of yellowfin tuna, Thunnus albacares. Journal of Experimental Marine Biology and Ecology, 482, 18â24. https://doi.org/10.1016/j.jembe.2016.04.008 García-Ibáñez, M. I., Bates, N. R., Bakker, D. C. E., Fontela, M., & Velo, A. (2021). Cold-water corals in the Subpolar North Atlantic Ocean exposed to aragonite undersaturation if the 2 °C global warming target is not met. Global and Planetary Change, 201, 103480. https://doi.org/10.1016/j.gloplacha.2021.103480 Gattuso, J.-P., Williamson, P., Duarte, C. M., & Magnan, A. K. (2021). The Potential for Ocean-Based Climate Action: Negative Emissions Technologies and Beyond. Frontiers in Climate, 2. https://doi.org/10.3389/fclim.2020.575716 Gaylord, B., Kroeker, K. J., Sunday, J. M., Anderson, K. M., Barry, J. P., Brown, N. E., Connell, S. D., Dupont, S., Fabricius, K. E., Hall-Spencer, J. M., Klinger, T., Milazzo, M., Munday, P. L., Russell, B. D., Sanford, E., Schreiber, S. J., Thiyagarajan, V., Vaughan, M. L. H., Widdicombe, S., & Harley, C. D. G. (2015). Ocean acidification through the lens of ecological theory. Ecology, 96(1), 3â15. https://doi.org/10.1890/14-0802.1 62 Gazeau, F., Gattuso, J.-P., Dawber, C., Pronker, A. E., Peene, F., Peene, J., Heip, C. H. R., & Middelburg, J. J. (2010). Effect of ocean acidification on the early life stages of the blue mussel Mytilus edulis. Biogeosciences, 7(7), 2051â2060. https://doi.org/10.5194/bg-7-2051-2010 Geffen, A. J. (2009). Advances in herring biology: From simple to complex, coping with plasticity and adaptability. ICES Journal of Marine Science, 66(8), 1688â1695. https://doi.org/10.1093/icesjms/fsp028 Gillies, C. L., Castine, S. A., Alleway, H. K., Crawford, C., Fitzsimons, J. A., Hancock, B., Koch, P., McAfee, D., McLeod, I. M., & zu Ermgassen, P. S. E. (2020). Conservation status of the Oyster Reef Ecosystem of Southern and Eastern Australia. Global Ecology and Conservation, 22, e00988. https://doi.org/10.1016/j.gecco.2020.e00988 Gimenez, I., Waldbusser, G. G., & Hales, B. (2018). Ocean acidification stress index for shellfish (OASIS): Linking Pacific oyster larval survival and exposure to variable carbonate chemistry regimes. Elem Sci Anth, 6, 51. Gobler, C. J., DePasquale, E. L., Griffith, A. W., & Baumann, H. (2014). Hypoxia and Acidification Have Additive and Synergistic Negative Effects on the Growth, Survival, and Metamorphosis of Early Life Stage Bivalves. PLOS ONE, 9(1), e83648. https://doi.org/10.1371/journal.pone.0083648 Gonzalez, M., Gueguen, Y., Destoumieux-Garzón, D., Romestand, B., Fievet, J., Pugnière, M., Roquet, F., Escoubas, J.-M., Vandenbulcke, F., Levy, O., & others. (2007). Evidence of a bactericidal permeability increasing protein in an invertebrate, the Crassostrea gigas Cg-BPI. Proceedings of the National Academy of Sciences, 104(45), 17759â17764. 63 Gruber, N., Clement, D., Carter, B. R., Feely, R. A., van Heuven, S., Hoppema, M., Ishii, M., Key, R. M., Kozyr, A., Lauvset, S. K., Lo Monaco, C., Mathis, J. T., Murata, A., Olsen, A., Perez, F. F., Sabine, C. L., Tanhua, T., & Wanninkhof, R. (2019). The oceanic sink for anthropogenic CO2 from 1994 to 2007. Science, 363(6432), 1193â1199. https://doi.org/10.1126/science.aau5153 Gu, H., Shang, Y., Clements, J., Dupont, S., Wang, T., Wei, S., Wang, X., Chen, J., Huang, W., Hu, M., & Wang, Y. (2019). Hypoxia aggravates the effects of ocean acidification on the physiological energetics of the blue mussel Mytilus edulis. Marine Pollution Bulletin, 149, 110538. https://doi.org/10.1016/j.marpolbul.2019.110538 Hamilton, S. L., Elliott, M. S., deVries, M. S., Adelaars, J., Rintoul, M. D., & Graham, M. H. (2022). Integrated multi-trophic aquaculture mitigates the effects of ocean acidification: Seaweeds raise system pH and improve growth of juvenile abalone. Aquaculture, 560. https://doi.org/10.1016/j.aquaculture.2022.738571 Han, T., Shi, R., Zhanhui, Q., Huang, H., Liang, Q., & Liu, H. (2017). Interactive effects of oyster and seaweed on seawater dissolved inorganic carbon systems: Implications for integrated multi-trophic aquaculture in Chinese coastal waters. Aquaculture Environment Interactions, 9. https://doi.org/10.3354/aei00246 Hargrave, M. S., Ekelund, A., Nylund, G. M., & Pavia, H. (2021). Filtration and fertilisation effects of the bivalves Mytilus edulis and Magallana gigas on the kelp Saccharina latissima in tank culture. Journal of Applied Phycology, 33(6), 3927â3938. https://doi.org/10.1007/s10811-021-02553-6 64 Holdsworth, A. M., Zhai, L., Lu, Y., & Christian, J. R. (2021). Future Changes in Oceanography and Biogeochemistry Along the Canadian Pacific Continental Margin. Frontiers in Marine Science, 8. https://doi.org/10.3389/fmars.2021.602991 Huan, P., Wang, H., & Liu, B. (2017). Assessment of housekeeping genes as internal references in quantitative expression analysis during early development of oyster. Genes Genet Syst, 91(5), 257â265. Medline. https://doi.org/10.1266/ggs.16-00007 Hurd, C. L., Beardall, J., Comeau, S., Cornwall, C. E., Havenhand, J. N., Munday, P. L., Parker, L. M., Raven, J. A., & McGraw, C. M. (2020). Ocean acidification as a multiple driver: How interactions between changing seawater carbonate parameters affect marine life. Marine and Freshwater Research, 71(3), 263. https://doi.org/10.1071/MF19267 Hurd, C. L., Law, C. S., Bach, L. T., Britton, D., Hovenden, M., Paine, E. R., Raven, J. A., Tamsitt, V., & Boyd, P. W. (2022). Forensic carbon accounting: Assessing the role of seaweeds for carbon sequestration. J Phycol, 58(3), 347â363. Medline. https://doi.org/10.1111/jpy.13249 IPCC. (2019). Special Report on Climate Change and Land. https://www.ipcc.ch/srccl/ Jablonski, D., & Lutz, R. A. (1983). Larval Ecology of Marine Benthic Invertebrates: Paleobiological Implications. Biological Reviews, 58(1), 21â89. https://doi.org/10.1111/j.1469-185X.1983.tb00380.x Jian, Z., Zhong, M., Li, R., Su, H., Rao, Q., Wang, Y., Wang, C., Sun, S., Chen, J., & Xie, P. (2025). Warming Enhances the Effects of Acidification on Aquatic Biota: A Global MetaâAnalysis. https://doi.org/10.1111/geb.70080 65 Jiang, L.-Q., Feely, R. A., Carter, B. R., Greeley, D. J., Gledhill, D. K., & Arzayus, K. M. (2015). Climatological distribution of aragonite saturation state in the global oceans. Global Biogeochemical Cycles, 29(10), 1656â1673. https://doi.org/10.1002/2015GB005198 Jiang, Z., Fang, J. G., Han, T. T., Mao, Y. Z., Li, J. Q., & Du, M. R. (2014). The role of Gracilaria lemaneiformis in eliminating the dissolved inorganic carbon released from calcification and respiration process of Chlamys farreri. Journal of Applied Phycology, 26(1), 545â550. https://doi.org/10.1007/s10811-013-0110-8 Jiang, Z., Jiang, W., Rastrick, S. P. S., Wang, X., Fang, J., Du, M., Gao, Y., Mao, Y., Strand, Ã., & Fang, J. (2022). The Potential of Kelp Saccharina japonica in Shielding Pacific Oyster Crassostrea gigas From Elevated Seawater pCO2 Stress. Frontiers in Marine Science, 9. https://doi.org/10.3389/fmars.2022.862172 Jibri, S. M., Jakada, B. H., Umar, H. Y., & Ahmad, T. A. (2016). Importance of Some Algal Species as a Source of Food and Supplement. International Journal of Current Microbiology and Applied Sciences, 5(5), 186â193. https://doi.org/10.20546/ijcmas.2016.505.020 Kong, N., Han, S., Fu, Q., Yu, Z., Wang, L., & Song, L. (2022). Impact of ocean acidification on the intestinal microflora of the Pacific oyster Crassostrea gigas. Aquaculture, 546, 737365. https://doi.org/10.1016/j.aquaculture.2021.737365 Koweek, D. A., Nickols, K. J., Leary, P. R., Litvin, S. Y., Bell, T. W., Luthin, T., Lummis, S., Mucciarone, D. A., & Dunbar, R. B. (2017). A year in the life of a central California kelp forest: Physical and biological insights into biogeochemical variability. Biogeosciences, 14(1), 31â44. 66 Kroeker, K. J., Kordas, R. L., Crim, R. N., & Singh, G. G. (2010). Meta-analysis reveals negative yet variable effects of ocean acidification on marine organisms. Ecology Letters, 13(11), 1419â1434. https://doi.org/10.1111/j.1461-0248.2010.01518.x Krzus, L. (2019). Pacific Dulse. Kurihara, H., Kato, S., & Ishimatsu, A. (2007). Effects of increased seawater pCO2 on early development of the oyster Crassostrea gigas. Aquatic Biology, 1, 91â98. https://doi.org/10.3354/ab00009 Lam, V. W., Cheung, W. W., Reygondeau, G., & Sumaila, U. R. (2016). Projected change in global fisheries revenues under climate change. Scientific Reports, 6(1), 1â8. Leal, P. P., Uribe, D., Henríquez-Antipa, L. A., Jiménez, C., Hormazábal, L., & Cascales, E.-K. (2024). Evaluating the ability of macroalgae to create a chemical refuge for bivalves under ocean acidification conditions in closed-environment experiments. Journal of Applied Phycology, 36(3), 1561â1575. https://doi.org/10.1007/s10811-023-03163-0 Lee, K., Kim, T.-W., Byrne, R. H., Millero, F. J., Feely, R. A., & Liu, Y.-M. (2010). The universal ratio of boron to chlorinity for the North Pacific and North Atlantic oceans. Geochimica et Cosmochimica Acta, 74(6), 1801â1811. Listiawati, V., & Kurihara, H. (2023). Brown seaweed Nemacystus decipiens intensifies the effects of ocean acidification on coral Montipora digitata. Marine Environmental Research, 191, 106150. https://doi.org/10.1016/j.marenvres.2023.106150 Lueker, T. J., Dickson, A. G., & Keeling, C. D. (2000). Ocean pCO2 calculated from dissolved inorganic carbon, alkalinity, and equations for K1 and K2: Validation based 67 on laboratory measurements of CO2 in gas and seawater at equilibrium. Marine Chemistry, 70(1â3), 105â119. Mackenzie, C. L., Pearce, C. M., Leduc, S., Roth, D., Kellogg, C. T., Clemente-Carvalho, R. B., & Green, T. J. (2022). Impacts of Seawater pH Buffering on the Larval Microbiome and Carry-Over Effects on Later-Life Disease Susceptibility in Pacific Oysters. Applied and Environmental Microbiology, 88(22), e01654-22. Macreadie, P. I., Anton, A., Raven, J. A., Beaumont, N., Connolly, R. M., Friess, D. A., Kelleway, J. J., Kennedy, H., Kuwae, T., Lavery, P. S., Lovelock, C. E., Smale, D. A., Apostolaki, E. T., Atwood, T. B., Baldock, J., Bianchi, T. S., Chmura, G. L., Eyre, B. D., Fourqurean, J. W., â¦ Duarte, C. M. (2019). The future of Blue Carbon science. Nat Commun, 10(1), 3998. PubMed-not-MEDLINE. https://doi.org/10.1038/s41467-019-11693-w Manzello, D. P., Enochs, I. C., Melo, N., Gledhill, D. K., & Johns, E. M. (2012). Ocean Acidification Refugia of the Florida Reef Tract. PLOS ONE, 7(7), e41715. https://doi.org/10.1371/journal.pone.0041715 Marliave, J. B., Gibbs, C. J., Gibbs, D. M., Lamb, A. O., & Young, S. J. (2011). Biodiversity stability of shallow marine benthos in Strait of Georgia, British Columbia, Canada through climate regimes, overfishing and ocean acidification. Biodiversity Loss in a Changing Planet, 10, 24606. Mateo, M. A., Romero, J., Pérez, M., Littler, M. M., & Littler, D. S. (1997). Dynamics of Millenary Organic Deposits Resulting from the Growth of the Mediterranean SeagrassPosidonia oceanica. Estuarine, Coastal and Shelf Science, 44(1), 103â110. https://doi.org/10.1006/ecss.1996.0116 68 McNeil, B. I., & Sasse, T. P. (2016). Future ocean hypercapnia driven by anthropogenic amplification of the natural CO2 cycle. Nature, 529(7586), 383â386. Mendez, R. L., & Kwon, J. Y. (2022). Precipitation and characterization of Pacific dulse (Devaleraea mollis) proteins. Journal of Applied Phycology. https://doi.org/10.1007/s10811-022-02853-5 Millan-Linares, M. C., Martin, M. E., Rodriguez, N. M., Toscano, R., Claro, C., Bermudez, B., Pedroche, J., Millan, F., & Montserrat-de la Paz, S. (2019). Nutraceutical Extract from Dulse (Palmaria palmata L.) Inhibits Primary Human Neutrophil Activation. Mar Drugs, 17(11). Medline. https://doi.org/10.3390/md17110610 Miller, J. J., Maher, M., Bohaboy, E., Friedman, C. S., & McElhany, P. (2016). Exposure to low pH reduces survival and delays development in early life stages of Dungeness crab (Cancer magister). Marine Biology, 163(5), 1â11. Min, Y., Li, Q., & Yu, H. (2023). Characterization of larval shell formation and CgPOU2F1, CgSox5, and CgPax6 gene expression during shell morphogenesis in Crassostrea gigas. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 263, 110783. https://doi.org/10.1016/j.cbpb.2022.110783 Murie, K. A., & Bourdeau, P. E. (2020). Fragmented kelp forest canopies retain their ability to alter local seawater chemistry. Scientific Reports, 10(1), 1â13. Narita, D., & Rehdanz, K. (2017). Economic impact of ocean acidification on shellfish production in Europe. Journal of Environmental Planning and Management, 60(3), 500â518. Nellemann, C., & Corcoran, E. (2009). Blue carbon: The role of healthy oceans in binding carbon: A rapid response assessment. UNEP/Earthprint. 69 Nishihara, G. N., Sato, Y., Eger, A. M., Gallagher, J. B., Hurd, C., Kawai, H., Kuwae, T., & Pessarrodona, A. (2025). Expert opinions regarding the concept of blue carbon in seaweed systems. Phycological Research. https://doi.org/10.1111/pre.70009 NOAA. (2025). Ocean acidification | National Oceanic and Atmospheric Administration. https://www.noaa.gov/education/resource-collections/ocean-coasts/ocean-acidification Ogawa, H., & Fujita, M. (1997). The effect of fertilizer application on farming of the seaweed Undaria pinnatifida (Laminariales, Phaeophyta). Phycological Research, 45(2), 113â116. https://doi.org/10.1111/j.1440-1835.1997.tb00070.x Olson, E. M., John, J. G., Dunne, J. P., Stock, C. A., & Drenkard, E. J. (2025). Potential for Regional Resilience to Ocean Warming and Acidification Extremes: Projected Vulnerability Under Contrasting Pathways and Thresholds. Global Change Biology, 31(7), e70360. https://doi.org/10.1111/gcb.70360 Osborne, E. B., Thunell, R. C., Gruber, N., Feely, R. A., & Benitez-Nelson, C. R. (2020). Decadal variability in twentieth-century ocean acidification in the California Current Ecosystem. Nature Geoscience, 13(1), 43â49. Ou, M., Hamilton, T. J., Eom, J., Lyall, E. M., Gallup, J., Jiang, A., Lee, J., Close, D. A., Yun, S.-S., & Brauner, C. J. (2015). Responses of pink salmon to CO2-induced aquatic acidification. Nature Climate Change, 5(10), 950â955. https://doi.org/10.1038/nclimate2694 Padilla-Gamiño, J. L., Alma, L., Spencer, L. H., Venkataraman, Y. R., & Wessler, L. (2022). Ocean acidification does not overlook sex: Review of understudied effects and 70 implications of low pH on marine invertebrate sexual reproduction. Frontiers in Marine Science, 9, 977754. https://doi.org/10.3389/fmars.2022.977754 Peabody, B., Davis, J., Alin, S., Bednarsek, N., Chadsey, M., Feely, R., Horwith, M., Kiefer, D., Mickett, J., Newton, J., Siegrist, Z., & Toft, J. (2019). Investigating seaweed cultivation as a strategy for mitigating ocean acidification in Hood Canal, WA (pp. 1â12). Paul G. Allen Family Foundation adn U.S. Navy. Peña, M. A., & Fine, I. (2024). Future Physical and Biogeochemical Ocean Conditions under Climate Change along the British Columbia Continental Margin. Atmosphere-Ocean, 62(1), 2â23. https://doi.org/10.1080/07055900.2023.2239186 Perez, F. F., & Fraga, F. (1987). Association constant of fluoride and hydrogen ions in seawater. Marine Chemistry, 21(2), 161â168. https://doi.org/10.1016/0304-4203(87)90036-3 Pessarrodona, A., FrancoâSantos, R. M., Wright, L. S., Vanderklift, M. A., Howard, J., Pidgeon, E., Wernberg, T., & FilbeeâDexter, K. (2023). Carbon sequestration and climate change mitigation using macroalgae: A state of knowledge review. Biological Reviews, 98(6), 1945â1971. https://doi.org/10.1111/brv.12990 Pfister, C. A., Altabet, M. A., & Weigel, B. L. (2019). Kelp beds and their local effects on seawater chemistry, productivity, and microbial communities. Ecology, 100(10), e02798. Pierrot, D., Epitalon, J.-M., Orr, J. C., Lewis, E., & Wallace, D. W. R. (2021). MS Excel program developed for CO2 system calculations â version 3.0. https://github.com/dpierrot/co2sys_xl 71 R Core Team. (2024). R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing. https://www.R-project.org/ Ravaglioli, C., De Marchi, L., Giannessi, J., Pretti, C., & Bulleri, F. (2024). Seagrass meadows as ocean acidification refugia for sea urchin larvae. Science of The Total Environment, 906, 167465. https://doi.org/10.1016/j.scitotenv.2023.167465 Raven, J., Caldeira, K., Elderfield, H., Hoegh-Guldberg, O., Liss, P., Riebesell, U., Shepherd, J., Turley, C., & Watson, A. (2005). Ocean acidification due to increasing atmospheric carbon dioxide. The Royal Society. Reum, J. C. P., Alin, S. R., Harvey, C. J., Bednaršek, N., Evans, W., Feely, R. A., Hales, B., Lucey, N., Mathis, J. T., McElhany, P., Newton, J., & Sabine, C. L. (2016). Interpretation and design of ocean acidification experiments in upwelling systems in the context of carbonate chemistry co-variation with temperature and oxygen. ICES Journal of Marine Science, 73(3), 582â595. https://doi.org/10.1093/icesjms/fsu231 Ricart, A. M., Gaylord, B., Hill, T. M., Sigwart, J. D., Shukla, P., Ward, M., Ninokawa, A., & Sanford, E. (2021). Seagrass-driven changes in carbonate chemistry enhance oyster shell growth. Oecologia, 196(2), 565â576. Medline. https://doi.org/10.1007/s00442-021-04949-0 Ricart, A. M., Ward, M., Hill, T. M., Sanford, E., Kroeker, K. J., Takeshita, Y., Merolla, S., Shukla, P., Ninokawa, A. T., Elsmore, K., & Gaylord, B. (2021). Coast-wide evidence of low pH amelioration by seagrass ecosystems. Glob Chang Biol, 27(11), 2580â2591. Medline. https://doi.org/10.1111/gcb.15594 Sanford, E., & Kelly, M. W. (2011). Local adaptation in marine invertebrates. Annual Review of Marine Science, 3(1), 509â535. 72 Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., Preibisch, S., Rueden, C., Saalfeld, S., Schmid, B., Tinevez, J.-Y., White, D. J., Hartenstein, V., Eliceiri, K., Tomancak, P., & Cardona, A. (2012). Fiji: An open-source platform for biological-image analysis. Nature Methods, 9(7), 676â682. https://doi.org/10.1038/nmeth.2019 Schmittgen, T. D., & Livak, K. J. (2008). Analyzing real-time PCR data by the comparative CT method. Nature Protocols, 3(6), 1101â1108. https://doi.org/10.1038/nprot.2008.73 Sowul, K., Carson, H. S., Bouma, J. V., & Fyfe, D. A. (2022). Washington State Recovery Plan for Pinto Abalone. Washington Department of Fish and Wildlife, Fish Program. Stillman, J. H., & Paganini, A. W. (2015). Biochemical adaptation to ocean acidification. The Journal of Experimental Biology, 218(Pt 12), 1946â1955. https://doi.org/10.1242/jeb.115584 Sui, Y., Kong, H., Huang, X., Dupont, S., Hu, M., Storch, D., Pörtner, H.-O., Lu, W., & Wang, Y. (2016). Combined effects of short-term exposure to elevated CO2 and decreased O2 on the physiology and energy budget of the thick shell mussel Mytilus coruscus. Chemosphere, 155, 207â216. https://doi.org/10.1016/j.chemosphere.2016.04.054 Swearer, S. E., Treml, E. A., & Shima, J. S. (2019). A review of biophysical models of marine larval dispersal. Oceanography and Marine Biology. Tasch, B. (2016). Researchers discover seaweed that tastes like bacon and is twice as healthy as kale. Independent. independent.co.uk/life-style 73 Troell, M., Joyce, A., Chopin, T., Neori, A., Buschmann, A. H., & Fang, J.-G. (2009). Ecological engineering in aquacultureâPotential for integrated multi-trophic aquaculture (IMTA) in marine offshore systems. Aquaculture, 297(1), 1â9. https://doi.org/10.1016/j.aquaculture.2009.09.010 USGS. (2023). Soda Ash Statistics and Information. National Minerals Information Center. https://www.usgs.gov/centers/national-minerals-information-center/soda-ash-statistics-and-information Van Dam, B., Polsenaere, P., BarrerasâApodaca, A., Lopes, C., SanchezâMejia, Z., Tokoro, T., Kuwae, T., Loza, L. G., Rutgersson, A., Fourqurean, J., & Thomas, H. (2021). Global Trends in AirâWater CO2 Exchange Over Seagrass Meadows Revealed by Atmospheric Eddy Covariance. Global Biogeochemical Cycles, 35(4). https://doi.org/10.1029/2020gb006848 Vignier, J., Laroche, O., Rolton, A., Wadsworth, P., Kumanan, K., Trochel, B., Pochon, X., & King, N. (2021). Dietary Exposure of Pacific Oyster (Crassostrea gigas) Larvae to Compromised Microalgae Results in Impaired Fitness and Microbiome Shift. Front Microbiol, 12, 706214. PubMed-not-MEDLINE. https://doi.org/10.3389/fmicb.2021.706214 Vogeler, S., Bean, T. P., Lyons, B. P., & Galloway, T. S. (2016). Dynamics of nuclear receptor gene expression during Pacific oyster development. BMC Developmental Biology, 16(1), 1â13. Wahl, M., Schneider Covachã, S., Saderne, V., Hiebenthal, C., Müller, J. D., Pansch, C., & Sawall, Y. (2018). Macroalgae may mitigate ocean acidification effects on mussel 74 calcification by increasing pH and its fluctuations. Limnology and Oceanography, 63(1), 3â21. https://doi.org/10.1002/lno.10608 Waldbusser, G. G., Brunner, E. L., Haley, B. A., Hales, B., Langdon, C. J., & Prahl, F. G. (2013). A developmental and energetic basis linking larval oyster shell formation to acidification sensitivity. Geophysical Research Letters, 40(10), 2171â2176. https://doi.org/10.1002/grl.50449 Waldbusser, G. G., Hales, B., Langdon, C. J., Haley, B. A., Schrader, P., Brunner, E. L., Gray, M. W., Miller, C. A., & Gimenez, I. (2014). Saturation-state sensitivity of marine bivalve larvae to ocean acidification. Nature Climate Change, 5(3), 273â280. https://doi.org/10.1038/nclimate2479 Waldbusser, G. G., Hales, B., Langdon, C. J., Haley, B. A., Schrader, P., Brunner, E. L., Gray, M. W., Miller, C. A., Gimenez, I., & Hutchinson, G. (2015). Ocean Acidification Has Multiple Modes of Action on Bivalve Larvae. PLoS One, 10(6), e0128376. Medline. https://doi.org/10.1371/journal.pone.0128376 Wang, Q., Cao, R., Ning, X., You, L., Mu, C., Wang, C., Wei, L., Cong, M., Wu, H., & Zhao, J. (2016). Effects of ocean acidification on immune responses of the Pacific oyster Crassostrea gigas. Fish Shellfish Immunol, 49, 24â33. Medline. https://doi.org/10.1016/j.fsi.2015.12.025 Wang, Y., Hu, M., Wong, W. H., Shin, P. K. S., & Cheung, S. G. (2011). The combined effects of oxygen availability and salinity on physiological responses and scope for growth in the green-lipped mussel Perna viridis. Marine Pollution Bulletin, 63(5), 255â261. https://doi.org/10.1016/j.marpolbul.2011.02.004 75 Watson, S.-A., Southgate, P. C., Tyler, P. A., & Peck, L. S. (2009). Early Larval Development of the Sydney Rock Oyster Saccostrea glomerata Under Near-Future Predictions of CO2-Driven Ocean Acidification. Journal of Shellfish Research, 28(3), 431â437. https://doi.org/10.2983/035.028.0302 Weitzman, J., & Bailey, M. (2019). Communicating a risk-controversy: Exploring the public discourse on net-pen aquaculture within the Canadian media. Aquaculture, 507, 172â182. https://doi.org/10.1016/j.aquaculture.2019.04.025 Wright-LaGreca, M., Mackenzie, C., & Green, T. J. (2022). Ocean Acidification Alters Developmental Timing and Gene Expression of Ion Transport Proteins During Larval Development in Resilient and Susceptible Lineages of the Pacific Oyster (Crassostrea gigas). Mar Biotechnol (NY), 24(1), 116â124. Medline. https://doi.org/10.1007/s10126-022-10090-7 Xiao, X., Agusti, S., Lin, F., Li, K., Pan, Y., Yu, Y., Zheng, Y., Wu, J., & Duarte, C. M. (2017). Nutrient removal from Chinese coastal waters by large-scale seaweed aquaculture. Sci Rep, 7, 46613. Medline. https://doi.org/10.1038/srep46613 Yamamoto-Kawai, M., Tsujimoto, H., Zhang, Y., Zimmermann, S., & Williams, W. (2025). Vertical Expansion of Aragonite Undersaturated Waters in the Canada Basin of the Arctic Ocean From 2003 to 2019. Journal of Geophysical Research: Oceans, 130(2), e2024JC021166. https://doi.org/10.1029/2024JC021166 Young, C. S., & Gobler, C. J. (2018). The ability of macroalgae to mitigate the negative effects of ocean acidification on four species of North Atlantic bivalve. Biogeosciences, 15(20), 6167â6183. https://doi.org/10.5194/bg-15-6167-2018 76 Young, C. S., Sylvers, L. H., Tomasetti, S. J., Lundstrom, A., Schenone, C., Doall, M. H., & Gobler, C. J. (2022). Kelp (Saccharina latissima) Mitigates Coastal Ocean Acidification and Increases the Growth of North Atlantic Bivalves in Lab Experiments and on an Oyster Farm. Frontiers in Marine Science, 9. https://doi.org/10.3389/fmars.2022.881254 Zhu, Y., Wang, N., Wu, Z., Chen, S., Luo, J., Christakos, G., & Wu, J. (2025). The Role of Seaweed Cultivation in Integrated Multi-Trophic Aquaculture (IMTA): The Current Status and Challenges. Reviews in Aquaculture, 17(3), e70042. https://doi.org/10.1111/raq.70042 77 Appendices Appendix A Experimental setup details Raw seawater collected at 20m depth near Boat Harbor off Vancouver Island was pumped through a 5µm filter and into the nine header tanks (see Figure 7). The Aquastar system (iks ComputerSystems Gmbh) recorded pH in each header tank every 5 minutes for the duration of the experiments, and maintained pH at programmed thresholds by dosing CO2 gas. All header tanks also received continuous diffused air to control for bubbling effects of gas. The pH of 9 dulse absent/present tanks was also recorded every 5 minutes, limited by the Aquastar computer capacity for pH electrode connections. All 18 paired tanks had wavemakers to continuously circulate water and provide a tumble culture for the dulse. 78 Figure 7. Experimental aquaculture system built for dulse-oyster co-culture research. Water entered the circular 2â PVC system (blue) at 30 psi and 10-15°C, feeding into 9 header tanks (teal, yellow & orange) at < 4 L/min to allow for stabilized water chemistry. Carbonate chemistry in header tanks was manipulated with CO2 gas to represent three CO2 conditions: ambient, OA (medium), and OA + upwelling (high). Treatment locations were rotated clockwise within groups of three header tanks for randomization. Each header tank fed into paired dulse absent/present tanks (green). Oyster beakers with static seawater from corresponding dulse absent/present tanks were randomly placed and heated to 25°C in a water bath (brown). All tanks and beakers were lidded to minimize off-gassing of CO2. 79 Appendix B Spawning details Broodstock were washed and shucked, and gonad samples were examined under a compound microscope to establish sex, remove hermaphrodites, and evaluate gamete condition. Two male and two female oysters with superior gametes were strip spawned into ambient seawater at room temperature (22-25°C) to create two family lines. Gametes were washed through a 200 µm filter, and eggs were counted under a compound microscope in triplicate on a Sedgewick Rafter counting chamber in 100µl seawater to calculate density. Sperm was added to eggs in seawater until 2-3 sperm per egg was observed under the microscope. 80 Appendix C RNA extraction details RNA extraction was performed for RT-qPCR using the Monarch Total RNA Miniprep Kit (New England Biolabs) and following manufacturer instructions for samples of tissue under 10mg, with the following changes: Part 1.3, incubated at 42°C for 15 minutes; part 1.4, spun for 2 minutes at 8,000 x g; part 2.4B, combined 8µl DNase I with DNase I reaction buffer to pipette into each sample; part 2.4C, incubated for 15 minutes at 37°C; part 2.5, added additional wash buffer step before addition of priming buffer; part 2.8, added 35µl nuclease-free water to column and incubated on ice for 5 minutes before centrifuging for elution. RNA concentration and purity were evaluated by measuring absorbance using a BioSpectrometer (Eppendorf). RNA concentrations were > 80 ng/µl, A260/A280 ratios were > 2.0, and A260/230 ratios were â¥ 1.4. No RNA degradation or presence of gDNA was detected with 1% agarose gel electrophoresis. 81 Table 3. Average dulse weight in grams ± SD throughout 23 days in tumble culture under continuous light. After each weighing, biomass was removed to reestablish a density of 10g/L; n = 3 replicates for each OA x dulse treatment combination OA treatment Experiment day -5 Experiment day 3 Experiment day 12 Experiment day 18 Ambient 400 g 540 ± 53 g 665 ± 92 g 625 ± 7 g OA 400 g 640 ± 53 g 683 ± 12 g 620 ± 53 g OA + Upwelling 400 g 587 ± 23 g 683 ± 35 g 637 ± 21 g 82 Table 4. Average pH ± SD in oyster beakers measured with a hand-held meter, with calculated pH from dulse absent/present tank water samples for reference. (â) denotes dulse absent; (+) denotes dulse present; n = 3 replicates for each OA x dulse treatment combination Treatment Oyster beakers Dulse absent/present tanks OA Dulse Experiment day 0 2 hpf Experiment day 0 14 hpf Experiment day 2 48 hpf Experiment day 3 calculated pH Ambient â 7.83 ± 0.02 7.82 ± 0.03 7.89 ± 0.04 7.9 ± 0.01 + 8.07 ± 0.09 8.01 ± 0.10 8.05 ± 0.07 8.08 ± 0.07 OA â 7.48 ± 0.07 7.53 ±0.06 7.75 ± 0.17 7.64 ± 0.02 + 7.81 ± 0.03 7.79 ± 0.04 7.83 ± 0.09 7.82 ± 0.08 OA + upwelling â 7.41 ± 0.01 7.47 ± 0.03 7.58 ± 0.05 7.58 ± 0.02 + 7.57 ± 0.10 7.61 ± 0.08 7.69 ± 0.05 7.70 ± 0.02 83 Table 5. Target and reference M. gigas genes examined in the present study through RT-qPCR. Selections included ion regulation genes from Wright-LaGreca et al. (2022), shell formation genes from Min et al. (2023), immune response genes from Q. Wang et al. (2016), and potential reference genes from Huan et al. (2017). Reference gene elf1α expression was selected to calculate relative expression of target genes, as elf1α primers performed the best. Footnotes refer to performance characteristics of note, such as when a value fell outside the range set in the methods. Gene Protein Function GenBank ID Primers (5â-3â) NKA (1) Ion regulation: Na + ,K + â ATPase XM_011436470 F: ACATCGGACCCATTGAGTAC R: CTGGTTGTGCCGGATGTTC NKA (2)1,4 Ion regulation: Na + ,K + â ATPase XM_034476028 F: GAGTGCACACACCTCAAC R: CAGCCCAACATGGTATGGTG slc4 (3) Ion regulation: HCO3-translocating XM_011441189 F: CATCATCCAGGTCCTCTGTC R: CAGCGAAGCTCTCTCTGC POU2F14 Shell formation XP_034303537.1 F: GTCTGGTTTTGTAACCGTAGA R: GCTGATTAGTGGGTGAGGC Sox51 Shell formation XP_011425377.1 F: TGCCCCTCCCTCTTTACTC R: GGTTTGCCTGCTTCATTCT Pax62 Shell formation XP_011433287.2 F: CCAGAATACGCACCACGAA R: ACCAGGGACTTGGATAGGG Def 2,3 Immune response: antimicrobial peptide ACQ76287.1 F: GCTGGATTTGGGTGTCC R: ACAGTAGCCCGCTCTACAAC Prp1,4 Immune response: antimicrobial peptide ACQ72987.1 F: CCACCATGTTCTCTCGGAGG R: CTTGTAGAACGGGCTAGCAC BPI1 Immune response: bacterial permeability increasing protein AAN84552.1 F: GATAGAAATAGGAATGGACGG R: GTTATAGATCCACGCTGCTCC 84 Gene Protein Function GenBank ID Primers (5â-3â) EF1α Potential reference gene 1 AB122066 F: AGTCACCAAGGCTGCACAGAAAG R: TCCGACGTATTTCTTTGCGATGT ARF1 Potential reference gene 2 HS159238 F: TCAGGACAAGATCCGACCACTGT R: GCAGCGTCTCTAAGTTCATCCTCATTA GAPDH5 Potential reference gene 3 AJ5448861 F: AGAACATCGTCAGCAACGCATCC R: CCTCTACCACCACGCCAATCCT 1 Delta slope = 0.11 â 0.17 2 R2 = 0.948-0.952 3 Outliers taken out to improve R2 4 Dynamic range â¤ 1:10 to â¥ 1:256 5 Poor performance characteristics 85 Table 6. Statistical summary for pairwise comparisons of estimated marginal means (EMMs) of oyster larvae size. Hpf: hours post fertilization; n = 3 replicate oyster beakers for each OA x dulse treatment combination Hpf Treatment comparison pair Treatment held constant % difference between EMMs 95% lower CL 95% upper CL p 24 OA & OA + upwelling Dulse absent -5.8% -10.5% -0.8% < 0.05 Ambient & OA + upwelling Dulse absent -7.7% -12.1% -3.2% < 0.001 OA & OA + upwelling Dulse present -4.5% -8.5% -0.4% < 0.05 Ambient & OA + upwelling Dulse present -4.2% -8.1% -0.1% < 0.05 Dulse absent & present Ambient 3.2% -0.3% 7.0% = 0.08 OA 5.8% 1.9% 9.8% < 0.01 OA + upwelling 7.2% 3.0% 11.6% < 0.001 48 OA & OA + upwelling Dulse absent -5.8% -10.5% -0.8% < 0.05 Ambient & OA + upwelling Dulse absent -7.7% -12.1% -3.2% < 0.001 OA & OA + upwelling Dulse present -4.5% -8.5% -0.4% < 0.05 Ambient & OA + upwelling Dulse present -4.2% -8.1% -0.1% < 0.05 Dulse absent & present Ambient 3.2% -0.3% 7.0% = 0.08 OA 5.8% 1.9% 9.8% < 0.01 OA + upwelling 7.2% 3.0% 11.6% < 0.001
Thesis/Dissertation
eng
Applied Animal Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Frommel, Andrea Y.
Dulse seaweed Devaleraea mollis mitigates effects of ocean acidification on larval Pacific oysters Magallana gigas
Text