Team:Georgia State/Design
Our ultimate goal is to save the corals...
...but how? Where do we start?
We began by first learning about what coral bleaching is and what causes it. After speaking to Alanna Waldman, a marine biology research assistant at Nova Southeastern University, we quickly realized there isn’t just one cause of coral bleaching, which means there isn’t just one way of approaching the problem.
Coral bleaching is when corals are put under some sort of stress such as increased temperature and acidic seawater and, as a result, expel their mutualistic microalgal symbiont, Symbiodinium. This leaves the coral “bleached” and vulnerable to disease. However, coral bleaching is reversible; once the conditions are favorable again, the coral can reuptake the algae. The main problem facing corals today is that these stressors are present for increasingly prolonged periods of time. The longer the corals are without their symbionts, the more likely they are to die by disease or by being overpowered by macroalgae.
Now that we had a better understanding of the problem, we began to brainstorm ways of fixing it. But before we could come up with plans of our own, we had to know more about what is currently being done to address this global phenomenon. So, yet again, we spoke to our friend Alanna Waldman, who is involved in these restoration efforts, to learn more. As of right now, there is little research being done to solve the problem of coral bleaching. There are, however, several ongoing restoration efforts where corals are taken to the lab and their stressor is removed to allow their symbiotic algae to be uptaken before they are replanted in-field. However, some issues involved with this are: one, what’s stopping the corals from being bleached again when another stressor reappears, and two, what can speed up restoration efforts to keep up with the rate of coral bleaching worldwide? As of right now, we are simply trying to keep our heads above water.
At first, we wanted to prevent coral bleaching from happening altogether by simply reducing the stressors present. An early target was reactive oxygen stress. We thought we might increase production of antioxidants by the microalgae to reduce oxidative stress resulting from increased photosynthetic activity seen during warming events. After speaking with Alanna, we revised our initial plan for a more practical one: we will utilize the critical period during a bleaching event for the corals to uptake engineered, bleaching-resistant Symbiodinium. This will be done in the same way current reef restoration is done, however with our modified alage, future bleaching events will be reduced.
Although transformation of foreign DNA into cells is well established for many organisms, this technique has not been as successful in the microalgae we are proposing to modify. We searched for information about transformation of Symbiodinium alage and found only two papers that claimed success:
One from 1998 (Ten and Miller) that utilized silicon carbide whiskers, polyethylene glycol (PEG), and vigorous shaking to introduce foreign DNA into Amphidinium sp. and Symbiodinium microadriaticum with a success rate of ~1 ppm. The other, published in 2015, (Ortiz-Matamoros et al.) utilized glass beads to disrupt the Symbiodinium cells then co-incubated them with Agrobacterium tumefaciens to transform foreign DNA into Fugacium kawagutii (formerly Symbiodinium kawagutii) and S. microadriaticum. We proposed to try the latter method as well as a modified version of a common electroporation method used to transform Picihia pastoris yeast and a recently published method that was used to transform O. marina, another dinoflagellate alga that would ultimately serve as our test model chassis.
Before we could transform anything, we first had to successfully culture these organisms in our lab. We tested different media, light intensities, and temperature to determine the optimal growth conditions for Symbiodinium. As we solidified the best culturing techniques, we also created an algae house that carried these algal cultures in flasks at the optimum light intensity and temperature.
We then started the transformation efforts by attempting to replicate the latest one done by Ortiz-Matamoros et al; however, there was no published record of the plasmids they transformed into the Symbiodinium. So, we reached out to their lab and got in touch with Dr. Tania Islas at the Instituto de Ciencias del Mar y Limnología, UNAM who sent us the pCB302-gfp-MBD plasmid they used in their own experiments and advised us on the best way to remove it from the filter paper they sent it on. They also provided us with the recipe for the ASP-8A media we would need to culture the Symbiodinium in, which we were not able to locate elsewhere.
Although this was a great starting point, the pCB302-gfp-MBD is not a codon-optimized plasmid for dinoflagellate expression but, instead, for plant expression. An additional issue was that after they performed that transformation, the Symbiodinium were left unable to reproduce or photosynthesize. This may have been due to the introduced GFP gene conflicting with cellular processes.
The next step in our project design was to create an optimized dinoflagellate expression plasmid carrying a reporter gene. We found that PhD students Brittany Splecher et. al. at University of Connecticut have made exactly that, the Dino III plasmid carrying a GFP gene, which they transformed into a dinoflagellate relative of Symbiodinium microadriaticum, Oxyrrhis marina. We replaced the GFP gene with that of RFP to see if this would cause the Symbiodinium to be more functional post-transformation. So, we removed the GFP part out of the Dino III plasmid then designed, synthesized, and ligated a codon-optimized RFP part in place of the GFP one.
To test the functionality of our modified Dino III plasmid, we used the electroporation procedures performed by the University of Connecticut to transform it into Oxyrrhis marina, our model organism, which has been successfully and stably transformed and is easy to culture. We have plans in place to send our plasmid to the University of Connecticut lab so that they will transform our plasmid as well. In addition, we also cultured Dunaliella tertiolecta, the food source for Oxyrrhis marina. We then developed a predator-prey model for the Oxyrrhis marina and Dunaliella tertiolecta to see how often and how much Dunaliella to feed into the system to achieve the highest Oxyrhhis growth rate possible.
Alongside this, we tested out other electroporation protocols and settings for Symbiodinium in hopes that something may work. Based on the most successful transformation method for Symbiodinium microadriaticum, our project can take one of two routes: either an electroporation is the best transformation method, and the Dino III plasmid will be used to introduce genes we believe will help Symbiodinium resist bleaching events, or the Agrobacterium-mediated transformation is the most efficient method. In which case, we will modify the pCB302-gfp-MBD plasmid and remove the plant optimized parts and replace them with the dinoflagellate promoter, reporter gene, and terminator found in the Dino III plasmid. Once we prove our new binary plasmid works, we would replace the reporter gene with a gene correlated to coral bleaching resistance.
Once we’ve got our engineered Symbiodinium, we will introduce it to the coral. But how can we test if the coral will even uptake our algae in the first place? When in doubt, talk to the experts. We reached out to the Coral Husbandry Expert at the Georgia Aquarium, Kim Stone, to learn more about corals. We didn’t realize before this talk that getting the coral to uptake the algae would be a grand project in and of itself. We thought that we would either dump our modified symbiodinium into a region of coral reef or directly inject the algae into the corals using a syringe-like tool. Which we quickly learned is NOT even close to what we should do. Stone proposed a much more practical and efficient method. She devised a step-by-step process beginning with purposely bleaching the coral of its original algae in a closed system then slowly allowing the coral to recover enough to potentially uptake our modified algae. We may partner with the GA Aquarium in the future to do this when we're ready.
Another aspect of our project that we took into great consideration is safety. Potentially releasing a genetically modified organism outside of the lab should not be handled irresponsibly and safety should be the number one concern. We looked into other labs that had experience with this and found one at the University of San Diego led by Dr. Johnathan Shurin, who was the first to conduct an EPA-approved outdoor experiment using microalgae. We set up a phone interview to discuss how they were able to make their research possible and what measures they had taken to ensure environmental responsibility and safety. They reported back that we would have to get in touch with the Environmental Protection Agency and would have to fill out a “TERA” form for this process to even begin. They also advised that this is a lengthy process. Although this did not affect our project design, it did give us more future direction and an approximate timeline of our final project.
1. Brittany N. Sprecher, Huan Zhang, Senjie Lin (2019, April 9). Nuclear gene transformation in a dinoflagellate.doi: 10.1101/602821
2. Mario Fernando Ortiz-Matamoros, Tania Islas-Flores, Boris Voigt, Diedrik Menzel, František Baluška, Marco A. Villanueva (2015, July 13). Heterologous DNA Uptake in Cultured Symbiodinium spp. Aided by Agrobacterium tumefaciens. doi:10.1371/journal.Pone.0132693
3. Ten Lohuis, M. R., and Miller, D. J. (1998). Genetic transformation of dinoflagellates (Amphidinium and Symbiodinium): expression of GUS in microalgae using heterologous promoter constructs. Plant J. 13, 427–435. doi: 10.1046/j.1365-313X.1998.00040.x
