One Fish, Two Fish…

Ana Silverio, The University of Texas at Austin

The Approach: In my previous post, I explained how important small fishes are to the food web and how their new found interaction with Gracilaria vermiculophylla came about. Now, measuring something such as diversity and abundance may sound confusing but it’s as simple as one, two, three!

Abundance is the number of individuals per species in an ecosystem and relative abundance is the overall evenness of those individuals. Diversity is more of a measurement of variation or how many different species are counted in a designated area/habitat.

Fine mesh seine net being dragged over the 15-meter transect to capture our fish.
Photo Credit: Norma Salcedo

Now that we understand what we are measuring… what’s next? As mentioned before, the Charleston harbor has been introduced with an invasive species of seaweed, but it has served as a home for the juvenile fish. To measure diversity and abundance we have to take samples from two different sites affected by this invasive species. Luckily, it’s a short stroll over to Grice Beach behind our marine lab to find a section of Gracilaria with 20% coverage for our sparse site and one with 80% coverage for our dense site. After establishing our sample sites, we take a 15-meter transect which we will pull our fine-mesh seine net through at about knee-deep water. We quickly but gently pull the net up to the beach and start sorting through our samples placing the fish in a half-gallon jar while discarding any invertebrates. We repeat this at our second site and voilà we have our samples!

Initial sorting process for our samples
Photo Credit: Norma Salcedo

Are we done yet? Of course not! Once we collect both of our samples from the different patches of Gracilaria, we take them back to the lab to set in preservatives for about a week and begin the sorting process. While we sort each jar, we try to identify each fish down to the lowest classification if possible (in a perfect world we would have all of our critters down to species). After identification is complete, we start our measurements of diversity and abundance by counting our fish. When we are finished counting, we organize our data and use statistical analyses to see if there is a significant difference in diversity and abundance in our two sample sites. We have followed procedures from the past two summers and each time we have sampled this summer to make sure we can compare our data at the end.

And now for the big reveal… Drumroll please! Will we find a difference in diversity? In abundance? In neither or both? Will we finally win a battle against the dreadful pluff mud? Although the last part seems unfortunately unlikely, join me next time to finally find out what secrets Gracilaria has tangled up in the Charleston Harbor!


Special thanks to my mentor, Dr. Harold for his support and guidance throughout this project. Also, to Dr. Podolsky and Grice Marine Lab for giving me the opportunity to conduct this research. This project is supported by the Fort Johnson REU program, NSF DBI-1757899.

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Invisible Neighbors: How Gracilaria Changes Bacterial Communities

Lilia Garcia, Illinois Wesleyan University

The Problem: It only takes a walk along the mudflats to notice large patches of wiry, red seaweed. The seaweed is called Gracilaria vermiculophylla, an invasive organisms that is native to East Asia (SERC, 2019)  The seaweed is hard to miss, but its effects on the ecosystem are not easily seen. This summer I will be studying how Gracilaria affects a bacterial community invisible to the naked eye.

Mudflat with Gracilaria, taken by L. Garcia

According to previous studies, Gracilaria is found to increase the amount of a bacteria called Vibrio (Gonzalez, et al., 2014). This may not mean much at first, since most of us don’t think about microscopic interactions. Bacteria, however, are important in maintaining the health of complex environments like estuaries. They cycle and break down nutrients and organic matter, influencing oxygen, carbon, and nitrogen levels. An increase in one group of bacteria, such as Vibrio, can change these patterns. And like most of us know, bacteria tends to spread easily. There are a few strains, or types, of Vibrio, such as V. vulnificus, V. parahaemolyticus, and V. cholera, that are dangerous to human health. An increase in these strains may cause an increase in disease from swimming or eating infected food.†

Vibrio growing on petri dish, taken by L. Garcia

We known Vibrio levels increase with Gracilaria, but we do not know how this happens. We also don’t know if all Vibrio strains increase together, or if only a few strains grow. To understanding the relationship between Gracilaria and Vibrio, I will record how much total Vibrio and how many strains of Vibrio grow in and away from patches of Gracilaria. In order to preserve its own health, Gracilaria produces compounds that promote or stop organisms from growing around it (Assaw et al., 2018). These are compounds I will test against different strains to study the mechanism Gracilaria uses affect specific Vibrio levels. I want to see how the growth of each strain is affected by different extracts. Will the strains further away from the Gracilaria be unable to grow when exposed to a certain type of extract? Will other strains grow better with the extract?

We tend to think about invasive species on a large scale, assessing the damage it causes to other familiar animals and plants. The ecosystem relies on tiny, cellular organism and studying how bacteria changes leads to a deeper understanding of environmental health. An invisible community is changing as Gracilaria flourishes, and there is a lot left to learn about it. 

Acknowledgements

Thank you to my mentor Dr. Erik Sotka, and our collaborator Dr. Erin Lipp. I would also like to thank Dr. Alan Strand and Kristy Hill-Spanik for their supporting guidance. Lastly, thank you to Dr. Loralyn Cozy (IWU) for preparing me to succeed in the lab. All research is funded by Grice Marine Lab and College of Charleston through the Fort Johnson REU Program, NSF DBI-1757899

References

Assaw S, Rosli N, Adilah N, Azmi M, Mazlan N, Ismail N. 2018. Antioxidant and Antibacterial Activities of Polysaccharides and Methanolic Crude Extracts of Local Edible Red Seaweed Gracilaria sp. Malays Appl Biol. 47(4): 135-144. 

Fofonoff PW, Ruiz GM, Steves B, Simkanin C, & Carlton JT. 2019. National Exotic Marine and Estuarine Species Information System. 

Gonzalez D, Gonzalez R, Froelich B, Oliver J, Noble R, McGlathery K. 2014. Non-native macroalga may increase concentrations of Vibrio bacteria on intertidal mudflats. Mar Ecol Prog Ser. 505: 29-36.

Gracilaria: New Intruder Weeding Through Charleston

Ana Silverio, The University of Texas at Austin

The Problem: Invasive species are animals that enter a new habitat away from their own home and are known for usually bringing about negative effects on natives in the area. Invasive species thrive in new environments when they can adapt to local conditions, and cause troubles in the way it works. With their usual predators not around, chaos can erupt, as they take away from some resources from the animals who call this habitat home (Albins et al 2015). Gracilaria vermiculophylla is a type of seaweed but also an invasive species from Asia and first seen on the Virginia coast. Although it is an invasive species, this seaweed seems to be singing a different song than usual (Nyberg et al 2009). Since it was first seen on the beaches of North America, it has taken a different role by providing a new habitat to local fishes. Gracilaria vermiculophylla is a dark brownish red seaweed with tangled strands that brush up against anything wading through the shallow water. Perfect for smaller fish to hide in. Although this seaweed seems to be bringing good things to the fishes not much is understood about what life was like for them under the waters of Charleston before our new stranger came about so we can’t comment on that part of the story. On the other hand, an interaction is indeed unfolding before our eyes and the story behind our new visitor is a bit fishier than one may think.

Example of a sample site: sparse patch of Gracilaria vermiculophylla on Grice Beach.
Photo taken by: Norma Salcedo

Gracilaria vermiculophylla is hard to miss on the shorelines of Charleston, it can be found in patches when the tide dwindles or on the seafloor. Its branches provide an ideal habitat along with a hiding space for juvenile fish during their vital first years of life and increases their numbers (Munari et al 2015). The preservation of these fishes during their early life stages is important to maintaining a healthy food web that keeps marine life afloat. Food is energy and energy is moved up to some of the biggest fisheries in this country from the very bottom of the smallest animals. It is important to know how the bigger fish’s food source is interacting with its habitat to make sure it’s healthy. Understanding how the interaction is working is a key factor in creating conservation plans and maintaining the ecosystem in good health.

Dense patch of Gracilaria vermiculophylla.
Photo taken by: Norma Salcedo

This summer, my research focus is on untangling Gracilaria vermiculophylla’s ecological relationships with these small fishes for a better understanding how diverse life is underwater. Replicating a design from the past two summers, I am curious to see the differences in diversity and abundances based on different patches of seaweed and if body size plays a significant role. Will more seaweed correlate with more diversity? The past two summers revealed some common patterns between fish diversity and patterns of seaweed patches but also some surprising differences between the two field seasons. Will we have a tie breaker this summer? Stay tuned to find out!


Special thanks to my mentor, Dr. Harold for his support and guidance throughout this project. Also, to Dr. Podolsky and Grice Marine Lab for giving me the opportunity to conduct this research. This project is supported by the Fort Johnson REU program, NSF DBI-1757899.


References

 Albins MA (2015) Invasive Pacific lionfish Pterois volitans reduce abundance and species richness of native Bahamian coral-reef fishes. Mar Ecol Prog Ser 522:231-243. 

Munari, C., N. Bocchi, and M. Mistri. “Epifauna associated to the introducedGracilaria vermiculophylla (Rhodophyta; Florideophyceae: Gracilariales) and comparison with the nativeUlva rigida(Chlorophyta; Ulvophyceae: Ulvales) in an Adriatic lagoon.” Italian Journal of Zoology 82.3 (2015): 436-445.

Nyberg, C. D., M. S. Thomsen, and I. Wallentinus. “Flora and fauna associated with the introduced red algaGracilaria vermiculophylla.” European Journal of Phycology 44.3 (2009): 395-403.

RXR sequenced, now on to imposex

Samera Mulatu, Georgia Southern University

IMG-0640Findings: My experience at the Fort Johnson REU Program was phenomenal! Towards the end of the program, I was able to retrieve the RXR gene sequences for the Eastern mud snail. While working towards this goal, I was able to get a first hand glimpse of the long and hard steps and techniques taken to retrieve DNA sequences. From generating primers, doing dissections, extracting RNA, making cDNA, and even making PCR products, these listed skills are only just a short list of what I learned during this research experience. Retrieving the RXR gene sequences for the mud snail, was a trial and error process. Sequences were sent in at least five times, and four of those five times did not give good results. This was a big lesson for me, and reminded me that science is a trial and error process because all of it is a learning process.

Now that the RXR gene sequence for the Eastern mud snail was retrieved, the next steps in this project would be to use the sequences to place the mud snail in its proper spot on the phylogenetic tree. Also, now that the gene sequences are retrieved they will be used next fall by Edwina Mathis (a graduate at MUSC who’s doing her research in this topic) and Dr. Demetri Spyropoulos to induce imposex in the Eastern mud snail while exposing the snails to TBT, SPAN 80, and DOSS. Afterwards, they will measure changes in isoform expression.

The significance of the results from this study will hopefully show that mud snail imposex is a sensitive indicator of endocrine disrupting compounds in the environment which may impact human health and the health of other organisms in the ecosystem. This is because high imposex rates in mud snail species could possibly be linked to higher levels of contamination found in that site within the Charleston Harbor. Hopefully this study will further future research on EDCs and their effects on different species.

I would like to give a big thank to Dr. Demetri Spyropoulos for guiding me in my research. Also to the Fort Johnson REU Program, NSF DBI- 1757899, for providing me with the funds to complete this project.

Related research

Hotchkiss, A.K, A.G.Leblanc, R.M. Sternberg. 2002. Synchronized expression of Retinoid X Receptor mRNA with Reproductive Tract Recrudescence in an Imposex- Susceptible Mollusc. Environ. Sci Technol. 42: 1345- 1351.

Ravitchandirane, V. S, M.Thangaraj. 2013. Phylogenetic Status of Babylonia Zeylanica (Family Babyloniidae) Based on 18S rRNA GENE FRAGMENT.Annals of West University of Timisoara, ser. Biology. 1(2): 135- 140.

Barron- Vivanco, B.S, D. Dominguez- Ojeda, I.M. Medina- Diaz, A.E. Rojas- Garcia, M.L. Robledo- Marenco. 2014. Exposure to tributyltin chloride induces penis and vas deferns development and increases RXR expression in females of the purple snail (Plicopurpura pansa). Invertebrate Survival Journal. 11: 204-2012.

Horiguchi, T., M. Morita, T. Nishikawa, Y. Ohta, H. Shiraishi. 2007. Retinoid X Receptor gene expression and protein content in tissues of the rock shell Thais clavigeraAquatic Toxicology. 84: 379-388.

 

Journey to the Center of the Pluff

Lauren Rodgers, Rutgers University

Version 2The Approach: In my previous blog post I discussed the importance of iron in ocean ecosystems. Because so many living things rely on iron to live and grow, it is important for us to understand how iron cycles, as it enters the ocean, exits the ocean, and changes from one form to another. Zetaproteobacteria are marine bacteria that rely on iron to create energy for themselves, but in this process, they also turn dissolved iron into solid iron. So these bacteria make rust as they grow. Unfortunately, rust isn’t very good for other organisms, and the Zetaproteobacteria effectually remove iron from the ocean. But still, these organisms are one half of the iron cycle and therefore play an prominent role. With our research, we aim to determine whether these bacteria are present in Charleston’s estuaries, and extrapolate how they might be impacting the local iron cycle.

Now, you most likely have one thing on your mind: How are they going to study all of this!? From our lofty research aims, we must simplify those down to into bite sized goals so we can have a successful summer of sampling.

Our Goals:

  1. Identify whether Zetaproteobacteria can be found in the sediments around Charleston.
  2. Measure the amount of Fe(II) and Fe(III) in the sediments

The first thing that we did in order to accomplish these goals is pick sampling sites. We wanted to sample the sediments for these Zetaproteobacteria, so we chose muddy regions close to tidal rivers that empty into the ocean. We wanted tidal rivers because Zetaproteobacteria live in salty waters, and these rivers mix with salt water from the ocean. We decided to look for these muddy regions along the Ashley River, Wando River, Stono River, and Cooper River, picking easily accessible sites far up the river where the water is fresher, midway down the river where the salt content is at a mid-range, and low down on the rivers, near the ocean, where the water is salty.

 

After identifying the sites that we wanted to sample at, we needed to figure out how to sample. We wanted to sample the mud at different depths, so we decided to use syringes to suck up the mud.

 

Once all of the samples were collected it was time to get back into the lab to analyze the data. In order to confirm the presence of Zetaproteobacteria we conducted PCR, which is a process that tells us if there was any DNA belonging to the Zetaproteobacteria in the samples.

 

 

 

To analyze the iron a ferrozine assay was conducted. In a ferrozine assay, different chemicals are added to the samples, which then turn different shades of purple depending on how much iron is present in them.

 

While we have already completed a lot of the data collection, we still have more to do. In the next few weeks we will focus on collecting the last few samples and analyzing them in the lab. Soon all of the results will be ready for interpretation!


I would like to thank my mentor, Dr. Heather Fullerton, for guiding me through this research. I would also like to thank the National Science Foundation for funding this research as well as the College of Charleston and Grice Marine Lab for their support.

Bacteria in the Ocean? That Eat Iron??

Lauren Rodgers, Rutgers University

Version 2The problem: Have you ever asked yourself, what is iron? It is an element? A rock? Some weird orange-ish substance? Is it the tool that you use to get the wrinkles out of clothes? And what does iron even do? Does it just sit there? Does anything eat it? Can we make things out of it? Iron is one of the most abundant elements on earth, yet not many people know much about the important role it plays in our lives.

Iron is more than just an element, or something found within a rock. It’s a nutrient, something necessary for the growth and metabolism of almost every living organism on Earth (Hedrich & Johnson, 2011). In the ocean, iron is found in two different forms, ferrous iron or Fe(II), which is soluble in water, and ferric iron or Fe(III), which is insoluble in water (Hedrich & Johnson, 2011). Because ferrous iron is soluble it is the form of iron that can be used by most organisms in the water (Hedrich & Johnson, 2011). This ferrous iron, however, is limited in the ocean despite its abundance in the Earth’s crust. In fact, Fe(II) is present only in incredibly small concentrations, making it a major limiting factor of growth for all of the plants and algae in the ocean. This is important because these plants and algae serve as the base of many food chains, so if there is a limitation on the growth of these organisms, it affects every other organism throughout the food chain. Though iron is an extremely important nutrient for many living organisms, it is still not well understood. One of the least understood aspects is how iron specifically cycles through different marine environments. Does it ever change form? Does anything add iron to the ocean? Does anything take iron out of the ocean? These questions bring us to Zetaproteobacteria.

Zetaproteobacteria is a recently discovered class of iron-oxidizing microbes. This just means that the bacteria eat iron in the form of Fe(II) and produce Fe(III) as a waste product (Emerson et al., 2007; Chiu et al., 2017). In fact, these waste products can take on the form of hollow tubes, also called tubular sheaths, or twisted stalks that you can see under the microscope!

 

Zetaproteobacteria were initially described in 2007 near hydrothermal vents, utilizing the large concentrations of Fe(II) that were present in the fluid that spewed from the vents (Emerson et al., 2007).

Iron Mat

Iron mat composed of Zetaproteobacteria on a lava rock near the submarine Loihi volcano. (A. Malahoff, Hawaii, Loihi Volcano, July 1988)

How do Zetaproteobacteria relate to the cycling of iron? 

Zetaproteobacteria, with their role in eating iron and transforming it from its soluble Fe(II) state into its insoluble Fe(III) form may have an important role in the cycling of iron through the environment, functioning as an important source of iron removal.

Since their discovery, Zetaproteobacteria have also been observed in many other habitats, including coastal estuarine habitats with lower levels of iron, similar to that of Charleston, SC. (Laufer et al., 2017; Chiu et al., 2017). Our study will try to identify if these Zetaproteobacteria are present in the muddy soils around Charleston, as well as measure the levels of Fe(II) and Fe(III) in the rivers where these bacteria may be found.

 

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Hopefully, through the study of the distribution of Zetaproteobacteria across the globe, including the chemical characteristics of the different environments that they inhabit, we may get a clearer picture of how iron cycles in aquatic environments and the role that these Zetaproteobacteria play.


I would like to thank my mentor, Dr. Heather Fullerton, for guiding me through this research. I would also like to thank the National Science Foundation for funding this research as well as the College of Charleston and Grice Marine Lab for their support.


References 

Chiu, B. K., Kato, S., McAllister, S. M., Field, E. K., & Chan, C. S. (2017). Novel pelagic iron-oxidizing Zetaproteobacteria from the Chesapeake Bay oxic-anoxic transition zone. Frontiers in Microbiology, 8(JUL), 1–16. https://doi.org/10.3389/fmicb.2017.01280

Emerson, D., Rentz, J. A., Lilburn, T. G., Davis, R. E., Aldrich, H., Chan, C. S., & Moyer, C. L. (2007). A novel lineage of proteobacteria involved in formation of marine Fe-oxidizing microbial mat communities. PLoS ONE, 2(8), e667. https://doi.org/10.1371/journal.pone.0000667

Hedrich, S., Schlömann, M., & Johnson, D. B. (2011). The iron-oxidizing proteobacteria. Microbiology,157(6), 1551–1564.

Laufer, K., Nordhoff, M., Halama, M., Martinez, R. ., Obst, M., Nowak, M., … Kappler, A. (2017). Microaerophilic Fe(II)-oxidizing Zetaproteobacteriaisolated from low-Fe marine coastal sediments – physiology and characterization of their twisted stalks. Applied and Environmental Microbiology, 83(February), AEM.03118-16. https://doi.org/10.1128/AEM.03118-16

Mori, J. F., Scott, J. J., Hager, K. W., Moyer, C. L., Küsel, K., & Emerson, D. (2017). Physiological and ecological implications of an iron- or hydrogen-oxidizing member of the Zetaproteobacteria, Ghiorsea bivora, gen. nov., sp. Nov. ISME Journal, 11(11), 2624–2636. https://doi.org/10.1038/ismej.2017.132

The BMA of Today

Christine Hart, Clemson University

2017-06-22 10.29.36

In previous blog posts I described the sand-dwelling microalgae, also known as benthic microalgae (BMA), which are essential to estuary ecosystems. Not only do they produce the air we breathe and food we eat, they also inform us about the subtle changes that are occurring in our environment. Changes that otherwise may go unnoticed.

How do BMA show these environmental changes? By forming the foundation of estuarine energy, they provide a snapshot of how the estuary is functioning as a whole. If changes occur in BMA patterns, this may indicate changes in the overall ecosystem. BMA are also easily characterized and compared using modern molecular approaches. These qualities make BMA living indicators, or bioindicators, that are important in monitoring future ecosystem health.

BMA become visible in the upper layers of sediment at low tide. Later, they decrease in density—or biomass—as the tide rises. Our project studied the mechanism for the increase of biomass during low tide. Previous studies suggested that the mechanism for biomass increase is vertical migration of BMA from lower layers to upper layers of sediment. We also tested whether BMA growth due to high light exposure contributes to the biomass increase.

Our results indicated that both vertical migration and growth due to sunlight exposure were important to the increase in biomass. This is the first contribution to literature that recognizes a multifaceted approach to BMA biomass changes.

Additionally, we studied in how the biomass increase was connected to patterns in the type of BMA in Charleston Harbor. Previous studies suggested that increasing biomass was connected to changes in the abundance of BMA species; therefore, we expected to see the amount of certain BMA species change based on their exposure to migration and sunlight.

We were surprised by our findings. In this study, we found that BMA did not vary over short time periods (by tidal stage or by exposure to migration and sunlight). Instead, we found that BMA varied spatially and over a period of 6 years. In fact, only one of the dominant species of BMA remained the same from 2011 to 2017 (Figure 1).  The long-term change in community coincides with geological changes in the sampling site (Figure 2).

QualitativeLvM-MS

Figure 1. The relative abundance of each dominant BMA species from 2011 to 2017 is shown immediately after sediment exposure (T0) and 3 hours later (TF). Only one species—Halamphora coffeaeformis—remains dominant in 2017. This is evidence of a dramatic change in the dominant type of BMA in Grice Cove.

These are positive results for the use of BMA as bioindicators. If types of BMA are invariable over short periods of time, measurements of BMA will be more precise. Bioindicators must be capable of showing changes that are occurring on a larger environmental scale; therefore, it would be a good sign if the change in BMA community reflects the changing geological environment (Figure 2). Still, more studies on the temporal and spatial patterns of BMA communities should be conducted before BMA can be used as bioindicators.

Changes in Grice Cove

Figure 2. Aerial view of Grice Cove sampling site over time. The approximate location of the sampling site is shown by the white line. Sampling sandbar has changed over time, possibly contributing to community changes. Source: “Grice Cove” 32 degrees 44’58”N 79 degrees 53’45”W. Google Earth. January 2012 to March 2014. June 20, 2017.

This study contributed new information to the studies of BMA biomass during low tide, and showed that the BMA of today in Grice Cove are significantly different than in previous years.

 

Thank you to my mentor, Dr. Craig Plante, and my co-advisor, Kristina Hill-Spanik, for their support and guidance. This project is funded through the National Science Foundation and supported by College of Charleston’s Grice Marine Laboratory.

 

Literature Cited:

Holt, E. A. & Miller, S. W. (2010) Bioindicators: Using Organisms to Measure Environmental Impacts. Nature Education Knowledge 3(10):8.

Lobo, E. A., Heinrich, C. G., Schuch, M., Wetzel, C. E., & Ector, L. (n.d.). Diatoms as Bioindicators in Rivers. In River Algae (pp. 245-271). Springer International Publishing. doi:10.1007/978-3-319-31984-.

MacIntyre, H.L., R.J. Geider, and D.C. Miller. 1996. Microphytobenthos: the ecological role of
 the “Secret Garden” of unvegetated, shallow-water marine habitats. I. Distribution, abundance and primary production. Estuaries 19:186-201.

Rivera-Garcia, L.G., Hill-Spanik, K.M., Berthrong, S.T., and Plante, C. J. Tidal Stage Changes in Structure and Diversity of Intertidal Benthic Diatom Assemblages: A Case Study from Two Contrasting Charleston Harbor Flats. Estuaries and Coasts. In review.