The BMA of Today

Christine Hart, Clemson University

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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).


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.


One Fish, Two Fish, Red Fish, Killifish

Melanie Herrera, U. of Maryland, College Park

After 9 sampling days, 18 collections, and over 3000 fish, we’ve discovered fishes’ habitat preferences are much more complex than we thought. To recap, our hypothesis predicted fish would prefer dense sites of the invasive seaweed, Gracilaria vermiculophylla, over sites with more open water (thus, less Gracilaria).  We also predicted that dense site would have greater diversity by attracting various types of fish due to its branches that conceal fish from predators.

Our belief that Gracilaria would fulfill the refuge effect, attracting more fish and more diverse species, was supported through the copious amounts of fish found in Gracilaria. Despite more abundance in the dense sites of Gracilaria, more diversity was shown in sparse sites (Figure 1). Among both the dense and sparse sites Atlantic Silversides and Bay Anchovies, Pipefish, and Striped Killifish were the most abundant and common species. While similar species occurred in both habitats, the sparse site had more occurrences of species that were considered rare in dense sites. For example, sparse sites had more occurrences of Spade fish and Florida Pompanos than dense sites. Additionally, sparse sites had species of fish such as leatherjackets and lizardfish that never occurred in dense sites.

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Figure 1: Rank abundance patterns of fish in dense sites (represented by triangles) and sparse sites (represented by circles) of G. vermiculophylla at Grice Cove. The number of fishes were calculated as a logarithm as a measure of relative abundance of fish at each site. Species are ranked from most abundant (1) to least abundant (8-10). Slopes show differences in species evenness amongst sites. Steeper slopes exhibit less species evenness.


Supporting our hypothesis, dense sites did demonstrate more abundance. In total, 2944 fish were collected from the dense sites while 361 fish were caught in the sparse sites. It is predicted that smaller-bodied fish used Gracilaria more as a refuge because of their increased vulnerability to threats as small animals. Lack of abundance in sparse sites could be explained by increased exposure to predators and environmental threats.

Increased use of the dense sites shows Gracilaria does contribute towards housing all types of fish, most importantly economically important fishes. According to the National Marine Fisheries Service’s report on fisheries economic in 2011, the seafood industry alone brings in a minimum of $88 million dollars annually. In order to support this important industry, commercial fisheries can use our research to establish sustainable fisheries by understanding the various habitats that help rear economically important fishes. Our identification of the invasive seaweed’s role on housing fish can be used as a protective measure for these fish in future sustainable management.


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Figure 2: Two of the top three most abundant species collected from dense sites of Gracilaria. (Left: Striped Kilifish; Right: Atlantic Silversides).


Thank you so much to my mentors Dr. Tony Harold and Mary Ann McBrayer for their advice and guidance. This research is funded through the National Science Foundation and College of Charleston’s Grice Marine Lab.


Spelunking for Coral Health

Meagan Currie, Swarthmore College

Sadly for the scientists involved, understanding coral health doesn’t always require a wetsuit, air tank or coral reef access. My work with the coral species Acropora cervicornis – or staghorn coral – usually requires a mundane terrestrial dress-code and a green headlamp as my only light source. Working with coral in the lab is more akin to spelunking than scuba diving. This environment, however, allows researchers to control the conditions surrounding the coral in order to more accurately understand the effects of chemical exposure on coral health.

To refresh, I am looking at the effects of the common chemical nonylphenol, which is used in laundry and dish detergents, is a stabilizer in plastic food packaging, and is one of the chemicals that makes up nonylphenol ethyl oxalates, which are found in pesticides, paints, and personal care products. After looking at the development of sea urchins exposed to nonylphenol, I found that the environmentally relevant values (0.1 ug/L – 50 ug/L) did not have a noticeable effect on embryo health. While this is a good sign in terms of the toxicity of the chemical to marine organisms, I was surprised, as most papers that have investigated the toxicity of nonylphenol have found it to be harmful at these levels to a variety of different organisms. Either my urchins were more robust than I had expected, or the chemical was not dissolving in water evenly when I made up my solutions. I decided, because of this, to expose my coral to a wider range of nonylphenol concentrations to get a better sense of whether or not my chemical was reacting. The coral were exposed to nonylphenol at levels ranging from 1 ug/L to 1000 ug/L, and over 96 hours I measured the effects. Let me outline the ways in which researchers monitor coral health in the lab.

During this experiment I consistently measured three things. The first was how well coral tissue regrows when exposed to a chemical, in this case nonylphenol. To do this, I cut the top of half of my coral fragments off at the beginning of the experiment, and took a picture after staining the tissue left on the top of the coral. Over time, a healthy coral fragment will regrow tissue over this wound. An unhealthy coral will take longer to regenerate the tissue, and so at the end of the 96 hour period I took a second picture of the stained tissue to see whether nonylphenol affected the speed of regeneration over this time.

Control fragment regeneration: time 0 (left) to 96 hours (right)

Another way to measure the health of coral is to measure Pulse Amplitude Fluorometery (PAM). Coral have symbiotic algae called zooenthallae, which provide food and oxygen through photosynthesis to the coral polyps. In turn, the coral release carbon dioxide, and provide shelter for the zooenthallae. PAM exposes the coral to a flash of ultraviolet light, which then causes the zooenthalae living in the coral tissue to emit fluorescence.

PAM Example

By measuring the intensity of this florescence, we can better understand the concentration of zooenthallae in the tissue as well as how well they are photosynthesizing. Each day during the exposure I run PAM when the coral are most sensitive to light exposure, right before the light-cycle of their day begins. If you want to see more of the amazing fluorescent world of corals, watch this video created by the reef conservation group Coral Guardians.

Finally, each day I run a basic physioscore to assess how healthy the coral looks and to document changes over time. This involves three different indicators: the polyps; the coloration; and the tissue of coral. I measure these features on a scale of five, with five being completely healthy and zero being nonexistent. Coral extend their anemone-like polyps when they are healthy. The A. cervocornis is a rich brown when its zooenthallae are still present in the tissue, and its tissue should cover the entire fragment of the coral. Below are two images from my study, the first a healthy control after 96 hours and the second a less healthy fragment exposed to nonylpyhenol.

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Having finished my 96-hour spelunking experiment with the coral, I will now gather my data and try to draw conclusions about the effects that nonylphenol has on photosynthesis, regeneration and overall health of the coral. With luck, these data will help solidify our understanding about nonylphenol in the marine environment and its effects on coral and similar invertebrates.

Cells and Instruments, but no Folsom Prison Blues

Brian Wuertz, Warren Wilson College


In my previous post, “Hiding in plain sight”, I introduced DOSS, a compound that has been recently identified as a probable obesogen. We are especially concerned about the potential of this compound to cause obesity symptoms in developing children through exposure from their mothers. While DOSS is in many products we use daily, such as homogenized milk and makeup products, it is commonly prescribed to pregnant women in the form of Colace stool softener. I am investigating both how much DOSS is in certain places in the body and how it may promote obesity.

One of the main concerns about obesity is that it elevates the risk of developing other diseases such as diabetes or cancer by causing a state of chronic inflammation (Bianchini 2002).  Chronic inflammation in  adipose tissue is regulated by immune cells, including macrophages. Macrophages are immune cells found throughout the body that help to fight against infection by recognizing invading bacteria and engulfing them in a process called phagocytosis, literally meaning to eat the other cells. In addition to phagocytosis macrophages are important regulators of the larger inflammatory response by secreting proteins that tell other cells to initiate or maintain a state of inflammation (Fujiwara 2005). This inflammatory reaction may be induced by DOSS. We have seen evidence of increased inflammation and obesity in mice treated with DOSS, so in order to figure out what causes that I am focusing on macrophages because of the way they regulate inflammation.

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I am isolating macrophages from breast milk samples under this hood in a sterile environment to make sure they are not contaminated with bacteria.

One way to study the inflammatory response of macrophages is to expose them to DOSS and then see if they produce the inflammatory proteins. Instead of trying to measure the secreted proteins, we can measure how much RNA is made in the cell. The RNA is the translator molecule that takes the plan for the protein from the DNA and makes it available for the cell to read and make the right protein. I identified genes for four different inflammatory proteins to measure the RNA so we can test if DOSS causes the macrophages to make more of any of them. I am testing macrophages that I am isolating from human placenta and breast milk tissue because the developing child is influenced by inflammation in the placenta and breast milk. Macrophages in these tissues could be the source of inflammation that influences how the child develops.

Okay so we have talked about cells, but what about the instruments? In my last post I introduced my instrument of choice, but did not call it that. It is not a guitar or a saxophone, but the HPLC, or high performance liquid chromatograph. This is simply a fancy instrument used to separate chemical compounds by forcing them through a tiny filter column filled with tiny beads. Some compounds stick more to the beads than others, so when you flow a liquid through the column the compounds come out of the column at different times. It is essential to separate the compounds in a sample because then you can measure the amounts of individual compounds.

We want to know where DOSS goes in the body, so we need to be able to measure how much of it is in a sample. I am working to get a system up and running to measure the amounts of DOSS in samples from different cells and tissues. We want to be able to measure DOSS in humans and in marine mammals such as dolphins. Dolphins are exposed to DOSS in the COREXIT oil spill dispersal agent that is applied to large and small scale oil spill issues along coastlines and in harbors. Dolphins are an important sentinel species, meaning that they can provide insight into human health issues.

I have to prepare a column and get the right mixture of solvents to make DOSS come off of the column in a timely fashion and in a way that we can measure it. The measurement is actually done with a mass spectrometer, which measures allows us to identify the compound based on how much it weighs. The number of atoms and types of atoms in the compound determine the mass of the compound. This mass is how the instrument measures the compound. The technique I am using is therefore called liquid chromatography mass spectrometry or LC-MS and the instrument is also referred to by LC-MS. Hopefully by the end of the summer I will be able to find beautiful data with this instrument that will make a coherent tune rather than a jumble of notes.


This is the MS part. It measures the mass of the compound and then breaks it apart and measures the mass of the pieces of the compounds and the amount of the compounds.


This is the LC or liquid chromatography part of the LC-MS instrument. Most of the work is figuring out the best solvent system to the sample through the small column with the red tag on it.

Funding for this REU program is generously provided by the National Science Foundation and hosted by the College of Charleston. Dr Demetri Spyropoulos at the Medical University of South Carolina is graciously hosting my research project and providing mentorship.



Bianchini, F., Kaaks, R., and Vainio, H. (2002). Overweight, obesity, and cancer risk. The Lancet Oncology 3, 565–574.
Fujiwara, N., and Kobayashi, K. (2005). Macrophages in Inflammation. Current Drug Target -Inflammation & Allergy 4, 281–286.

Gametes galore!

By Cecilia Bueno, Lewis & Clark College

I am conducting my experiment in two parts: the first looks at how increased salinity affects fertilization success, the other looks at how salinity affects sperm function.

The fertilization experiments start with watching the weather- squirrel treefrogs mate on nights when there has been a lot of rain. We collected frogs from dixie plantation on nights when there had been 0.5 inches of rain accumulation or more. Since squirrel treefrogs mate at night, we would go out around 10:30pm and stay often until 3:00am when we found pairs.


Grassy wetland at Dixie Plantation where several pairs of squirrel treefrogs were found

The frog pairs were taken back to the lab where they were placed in individual tubs with water at 6ppt salinity. When the pair had laid around 200 eggs, we removed them from the tubs and placed them with a new partner. This repeated until the frogs no longer went into amplexus- running from 4:30am to 8:00am.

After the eggs had been laid and the frogs stopped going into amplexus, the eggs were transferred from the large tubs to smaller weigh dishes. These weigh dishes with eggs were placed under the microscope and photographed for counting. I then counted how many eggs in each dish were fertilized. These counts- total eggs and number fertilized- will be used to calculate fertilization success of each of the pairs.

The second part of the experiment focused on sperm function of the frogs. Males from the initial pairs were kept for this experiment, while females and fertilized eggs were released. We created a sperm concentrate using the testes of each male. This sperm concentrate was then added separately to different tubes containing water at different salinity levels. We tested sperm at salinity levels which had previously been shown to have an effect on sperm function- 4ppt, 5ppt, 6ppt, 7ppt and 8ppt- as well as a control of 0.39ppt.


Screenshot of a video of sperm from a squirrel treefrog male

Once the sperm concentrate has been added to a test tube, the sperm is activated so I have to act quickly. The diluted sperm solution is placed under a microscope and video taped.

After the videos have all been taken, they will be analyzing them with the CASA (Computer Assisted Sperm Analysis) software developed by Wilson and Leedy. This software, run on ImageJ, tracks motile sperm and calculates percent motility and average velocity of the motile sperm.

When I have counts on percent motility and average velocity of sperm in different treatments, I will be able to compare the results I get from the sperm experiments to the results from the fertility experiments.

I would like to thank the National Science Foundation for funding this REU program, and the Grice Marine Lab of the College of Charleston for hosting us. In particular, I would like to thank my mentor Dr. Allison Welch for her help and support.

Living Life as a Sea Urchin Momma

Hailey Conrad, Rutgers University

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Me working hard to make my sea urchin babies

For my project I am using the same technique that the father of genetics, Gregor Mendel, used to establish his Laws of Heredity: cross breeding. So, I have to breed and raise a whole lot of sea urchins. For a refresher, I’m trying to determine if there is heritable genetic variation in how sea urchin (specifically an Arbacia punctulata population from Woods Hole, Massachusetts) larvae respond to ocean acidification. To do this, I’m rearing sea urchin larvae in low and high carbon dioxide conditions and measuring their skeletal growth. I’m breeding 3 sea urchin males with 3 sea urchin females at a time, for a total of 9 crosses. To tease apart the impact of genetic variation on just the larvae themselves, I will be fertilizing the sea urchin eggs in water aerated with either current atmospheric levels of carbon dioxide, about 410 parts per million, or 2.5 times current atmospheric carbon dioxide levels, about 1,023 parts per million. Then, I will be rearing the larvae in water aerated with either 409 ppm CO2 or 1,023 ppm CO2. This will give me four different treatments for each cross, giving me 36 samples in total. By fertilizing and rearing them in the same and different levels of carbon dioxide I will be able to see how much of an impact being fertilized in water with a higher carbon dioxide concentration has on larval growth versus just the larval growth itself. It’s important for me to make that distinction because I just want to identify genetic variation in larval skeletal growth, and separate out any extraneous “noise” clouding out the data. I’m rearing the larvae in a larval rearing apparatus. Each of the 36 samples will be placed in jar with water aerated with the correct CO2 treatment. Each jar will constantly have atmosphere with the correct CO2 concentration bubbled in. Each has a paddle in it that is hooked to a suspended frame that is swayed by a motor. This keeps the larvae suspended in the water column. The jars are chilled to 14 C by a water bath.


My larval rearing apparatus

After a 6-day period the larvae are removed from the jars and their skeletal growth is measured. They are preserved with 23% methanol and seawater and frozen.


An Arbacia punctulata pluteus

You’re probably curious how the heck I am able to measure the larva’s skeletons. They’re microscopic! Well, I use a microscope coupled to a rotary encoder with a digitizing pad and a camera lucida. Which, looks like this:

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A microscope coupled to a rotary encoder with a digitizing pad and a camera lucid hooked up to a computer

This complicated-sounding hodge-podge of different devices enables me to do something incredible. I can look through the microscope at the larva, and also see the digitizing pad next to the microscope, where I hold a stylus in my hand. When I tap the pad with the stylus and the coordinates of various points on the anatomy of the plutei that I am tapping at get instantly recorded on my computer! The rotary encoder is the piece attached to the left side of the microscope and it enables me to record coordinates in three dimensions. Then, I can use those coordinates to calculate the overall size of the skeleton. My favorite part of doing science is learning how scientists are able to do the seemingly impossible- like measuring something microscopic.

After I gather all of my data, I will do some statistical analysis to see the affect that the male parents have on the skeletal growth of their offspring. I will not be focusing on the impact that females have on the skeletal growth of their offspring. The quality of the egg itself could be an influencing factor on the size of the offspring, whereas sperm is purely genetic material. Like how I’m trying to isolate the influence of ocean acidification during larval rearing from during the act of fertilization, I am trying to isolate just genetic influences on larval skeletal growth from egg quality. Check back to see how it goes!

Special thanks to the National Science Foundation for funding this REU program, the College of Charleston and Grice Marine Laboratory for hosting me, and Dr. Bob Podolsky for mentoring me!




My Days with the Shrimp

Deanna Hausman, The University of Texas at Austin

me with ray





In “What can baby shrimp teach us about oil spills,” I discussed the problem of UV enhanced toxicity of oil. In other words, the fact that UV rays can cause molecules in oil known as PAHs to become more harmful than they would be otherwise. I also discussed the fact that this summer, I will be studying the effects of oil toxicity on grass shrimp, or Palaemontes pugio, an important estuarine species that cycles nutrients through the food chain. Because oil spills are always complex, and organisms can be exposed to oil in many different ways, from the sediment they walk on to the water they swim through, a variety of experiments are needed to get a better understanding of this issue.


A few of the many PAHs- the compounds in oil that harm marine life Photo from:

The first and simplest of the experiments I conducted was the developmental test. In this test, I basically mixed oil and seawater in a giant blender, then took out the water with the oil dissolved in it. Then, I made several dilutions, creating several concentrations of the oily water. Then, I took 6-well plates and filled them up, and placed a single, 24-hour old shrimp in each well. Then, I put these plates in an incubator under UV and non-UV light, and waited for 4 days. After that, I moved the shrimp into clean water, counted how many died, and am currently monitoring them to see how the oil exposure in early life impacts their ability to grow into healthy juveniles.


Shrimp being monitored after initial oil exposure

Another experiment I conducted essentially followed the same procedure as above, but instead of watching them as they grew, I analyzed the shrimp after their 96-hour oil exposure to see whether the oil affected the concentration of a hormone, known as an ecdysteroid, that controls their molting. Essentially, if the concentrations of this steroid are off in a shrimp it can’t grow properly, so it’s very important!

I’m also conducting an oil sheen test. In this experiment, I place 40 larval shrimp in an aquarium, some caged on the bottom and some swimming freely, and then place an extremely thin oil sheen on top. One aquarium goes under UV light and the other goes under fluorescent light, and after exposure I analyze whether the sheens have had a harmful effect. Whether thin oil sheens are toxic is something that’s not very well understood in this species, so it will be very interesting to see the results.

Finally, I’m conducting an experiment to see what occurs when oil is mixed in with sediment. Essentially, this involves putting sediment from an estuary in a jar, adding oil, and tumbling it around so that the oil is completely mixed in. Then, the sediment is placed into beakers along with water and 24-hour old shrimp, and put under UV and non-UV light for 24 hours, in order to see what mortality occurs. This will perhaps be the most informative experiment, as grass shrimp spend most of their time on the seafloor, so if they’re going to be exposed to oil, it will likely be from the sediment they’re walking on.

In short, I have my hands full this summer! It will be very interesting to see the results. Hopefully, this will increase our knowledge of the harmful impacts oil spills can have to estuarine organisms, and allow NOAA and oil spill analysts to make better predictions of the long-range impacts of oil spills. Ultimately, this may help them make better clean-up decisions.

Thank you to my mentors, Dr. Marie Delorenzo and Dr. Paul Pennington, for their guidance. I’d also like to thank Katy Chung for all her help and expertise. This research is funded through the National Science Foundation.