Larval Phthalate Soup

Samuel Daughenbaugh, DePauw University

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The Approach: In my previous post, I described a group of chemical additives called phthalates and their potential impact on the development of sea urchin larvae. The plastic industry uses several phthalates that vary in chemical structure and toxicity levels. One way phthalates differ in structure is by their size. I am studying the effects of three phthalates with different molecule sizes — DMP (small), DBP (medium), and DEHP (large) — on mortality (lethal effect) and larval skeletal growth (sublethal effect).

My first major challenge was to dissolve the chemicals in seawater. As hydrophobic liquids, phthalates only mix with water molecules at very low concentrations; larger types (longer side chains) are less soluble. By dissolving each chemical in acetone, I am able to get DMP into seawater at 1000 parts per million (ppm), or 0.01%, and DBP and DEHP at 1 ppm. I am testing 5 concentrations of each chemical in addition to an acetone control (no phthalate), and a seawater control (no phthalate or acetone).

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Experimental jars with stirring paddles

Once the chemicals are in solution, I spawn male and female sea urchins via electric voltage and collect their sperm and eggs. Then, I fertilized the eggs and introduce them to experimental jars where they then begin to develop into larvae. Small paddles stir the water to increase the oxygen level and keep the larvae suspended. After growing the larvae for two days, a period before they start to depend on food, I transfer them into small tubes, preserve and store them in a freezer.

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Normal 4-arm pluteus larvae (Photo taken by Jaclyn Caruso)

 

To measure and categorize larvae into different stages of development, I observe them under a microscope that can record landmark points on the larval body in three dimensions. After determining the proportion of individuals that failed to develop to the normal 2 or 4-arm pluteus stage (pictured below), I use the landmarks to calculate the lengths of different skeletal features to determine how much the larvae had grown. At the end of each trial, I will have observed hundreds to thousands of dead larvae and once all of them have been counted and measured, I can begin to analyze the data and learn whether the phthalates are having a significant effect on their development.

Acknowledgements

This project is supported by Dr. Robert Podolsky and the Fort Johnson REU Program, NSF DBI-1757899.

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Life in Plastic, It’s not Fantastic

Samuel Daughenbaugh, DePauw University

2DA71FE7-975A-4AA8-8A78-DF3D1E545F05The Problem: We live in a plastic world. Plastics have saturated all aspects of our daily lives and, as a consequence, have also entered the natural world.  About 8.3 billion metric tons have been produced in the past 60 years, playing a pivotal role in the advancement of modern society (Parker, 2018). Although they are used to create many things we enjoy and benefit from, there are serious consequences for the health of humans and the environment that are associated with their use.

We have found plastics in unexpected places, everywhere from human guts to the most remote locations on earth (Schwabl, 2018; Woodall, 2014). Plastics have a long list of negative effects on living organisms, but their impact in the ocean is of special concern. Pictures of turtles with straws up their noses, bottle caps spilling out of dead bird stomachs, and penguins strangled in plastic beverage rings are often posted on social media sites. Less widely known are the chemical additives that leach from plastics. Phthalates are one such group of additives that pose threats to the health of humans and marine life.

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Current Fort Johnson REU Interns (Julianna Duran not pictured) collecting plastic and sand dollars on Otter Island. (Photo credit: R. Podolsky)

Phthalates have been valuable to the plastic industry because they promote flexibility and durability in many plastics (EPA, 2017). An astounding 470 million pounds of phthalates are used in the United States every year (EPA, 2017). This presents a significant problem because phthalates interfere with the production of important hormones that regulate growth and metabolism in humans and other animals (Boas et al., 2012).

This summer I am exploring the effects of three different phthalates– dimethyl phthalate (DMP), di-n-butyl phthalate (DBP), and di-2-ethylhexyl phthalate (DEHP)–on the larval development of marine invertebrates, using the purple-spined sea urchin (Arbacia punctulata) as a model. Sea urchin larvae float freely in the water column for an extended period of time and, therefore, are vulnerable to many marine pollutants.

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Purple-spined sea urchin (Arbacia punctulata)

Sea urchins are an important model because they are closely related to humans. Both humans and sea urchins use a signaling hormone called thyroxine, which is especially important for growth in early developmental stages (Heyland et al., 2006). Exposure to phthalates can disrupt the production of thyroxine. Additionally, larvae are very important to study because they form the base of food webs. Being at the bottom of the food chain means they feed animals at higher levels, many of which humans rely on for protein. Therefore, understanding how phthalates affect sea urchin growth and metabolism can lead to new insights into how these pollutants directly and indirectly impact human health.

 Acknowledgements

I would like to thank my mentor, Dr. Robert Podolsky, for his continued support, guidance, and encouragement. This project is supported by the Fort Johnson REU Program, NSF DBI-1757899.

References

Boas, M., Feldt-Rasmussen, U., & Main, K. M. (2012). Thyroid effects of endocrine disrupting chemicals. Molecular and Cellular Endocrinology, 355(2), 240-248. 

Environmental Protection Agency (Ed.). (2017). Phthalates. America’s Children and the Environment, 3, 1-19.

Heyland, A., Price, D. A., Bodnarova-Buganova, M., & Moroz, L. L. (2006). Thyroid hormone metabolism and peroxidase function in two non-chordate animals. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution, 306B(6), 551-566.

Parker, L. (2018, December 18). A whopping 91% of plastic isn’t recycled. Retrieved from  http://www.nationalgeographic.com

Schwabl, P. (2018, October). Assessment of Microplastic Concentrations in Human Stool. Conference on Nano and microplastics in technical and freshwater systems, Monte    Verità, Ascona, Switzerland.

Woodall, L. C., Sanchez-Vidal, A., Canals, M., Paterson, G. L., Coppock, R., Sleight, V., . . . Thompson, R. C. (2014). The deep sea is a major sink for microplastic debris. Royal      Society Open Science, 1(4), 140317-140317. doi:10.1098/rsos.140317

Clinking Glasses: Not Just for Toasts

Jaclyn Caruso, Salem State University

Me Aquarium

The Approach: How can sea urchins tell us whether preservatives used in cosmetics are harmful to the environment? Our approach involves the rhythmic clinking of glasses—but we’ll get to that.

As detailed in my previous post, concerns about the use of parabens as preservatives led to the introduction of new, “safer” alternatives, like phenoxyethanol and chlorphenesin. We are testing the effects of these preservatives on the early development of the purple sea urchin, Arbacia punctulata. Sea urchins are useful for laboratory studies because they are easy to rear and they have free-swimming larvae with distinct morphologies, allowing us to test for both lethal and subtle effects of these chemicals.

Because this is a first test with sea urchins and concentrations that induce a response have varied among studies, the concentrations of chemicals in seawater we are using span a large range—1000 to 0.1 parts per million (ppm). Including controls with no chemicals creates 34 jars per trial. Over the summer we will be able to complete several independent trials using different male-female pairs.

To collect sperm and eggs from male and female sea urchins, we apply a mild voltage with electrodes to the top of the sea urchin close to the gonopores, the small openings where gametes come out. Once we gather the sperm and eggs, we dilute them to specific concentrations, combine them, and let nature take its course! After an hour, we add fertilized eggs to each of the jars, which are stirred by glass paddles. The stirring allows larvae to get plenty of oxygen and to avoid settling at the bottom of the jar. The paddles have to be glass because plastics can potentially alter the results (ASTM, 2015). The whole apparatus has a pleasant clinking sound once running!

After two days, we collect the developing stages, preserve them with methanol, and keep them at ‑20°C until they are measured. Finally, the analysis begins! We load a counting chamber with the larvae and use a microscope to count the number of individuals that are in each stage of development. By this point, a larva should have 4 arms—known as the pluteus stage.

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A preserved 4-arm pluteus larva. Scale bar = 0.1 mm. Jaclyn Caruso, 2018.

If larvae in the chemical jars are at earlier life stages compared to the controls, it suggests that the chemicals delay or stop development.

To look at more subtle effects, we are also measuring 20–25 individuals that have reached the larva stage per jar using 10 specific landmarks on the body. The microscope uses a camera lucida to project an image of the larva onto a digitizing pad that can record the location of landmarks in three dimensions. We use these data points to generate 3D models of each of the larvae to measure the size of the skeleton. This technique will allow us to test for sublethal effects of the chemicals that might not be obvious at first glance.

Workspace

The camera lucida arm of the microscope sits over the digitizing pad. Jaclyn Caruso, 2018.


Acknowledgements

Thank you to Dr. Bob Podolsky (CofC) for his mentorship, Dr. Cheryl Woodley (NOAA) for providing her procedures and resources, and Pete Meier (CofC) for teaching me the ropes of setting up aquaria. This project is supported by the Fort Johnson REU Program, NSF DBI-1757899.


References

ASTM (2015) ‘Standard Guide for Conducting Static Acute Toxicity Tests with Echninoid Embryos’, Astm, 131, pp. 1–2. doi: 10.1520/D7385-13.2.

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.

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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.

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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!

 

 

 

Playing with Plutei

Hailey Conrad, Rutgers University

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Me! Photo Credit: Kady Palmer

Ocean acidification is known as climate change’s evil twin. When the pH of ocean water drops, carbonate ions in the water form carbonic acid instead of calcium carbonate. Calcium carbonate is the form of calcium that marine animals that have calcium-based skeletons (like us!) and shells use to build their bones and shells. Having smaller and weaker skeletons or shells impacts their ability to survive. However, some individuals within certain species or populations of species have genes that make them more resistant to ocean acidification. If these individuals are able to pass on these genes to their offspring, then the species has the ability to evolve in response to ocean acidification instead of going extinct. This summer I’m working with Dr. Bob Podolsky in College of Charleston’s Grice Marine Field Station to study the extent to which ocean acidification affects Atlantic purple sea urchins, Arbacia punctulata. We are specifically trying to see if any individuals within a population from Woods Hole, Massachusetts, have any heritable genetic resistance to the negative impacts of ocean acidification. We hypothesize that there will be genetic resistance given that the northern Atlantic coast naturally has lower levels of saturated calcium carbonate, so a population that has evolved to live in that type of environment should have some resistance to lower calcium carbonate levels already (Wang et al 2013). We’re using a basic cross breeding technique to rear Arbacia punctulata larvae to their plutei stage, when they have four main body rods. At this stage they look less like sea urchins than they do like Sputnik!

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A sea urchin pluteus larvae with four body rods

Then, we will look to see if any of the male parents consistently produce male offspring that are more resistant to ocean acidification.  If males like these exist within this population, then the species has the capacity to evolve in response to ocean acidification, instead of going extinct! This is a very big deal, and could potentially be very hopeful. Even if we don’t get the results that we are hoping for, the results of this research could inform policy and management decisions.

Literature Cited:

Wang, Z. A., Wanninkhof, R., Cai, W., Byrne, R. H., Hu, X., Peng, T., & Huang, W. (2013). The marine inorganic carbon system along the Gulf of Mexico and Atlantic coasts of the United States: Insights from a transregional coastal carbon study. Limnology and Oceanography, 58(1), 325-342. doi:10.4319/lo.2013.58.1.0325

Thank you to the National Science Foundation and College of Charleston’s Grice Marine Laboratory for funding my project. And, special thanks to Dr. Bob Podolsky for being a wonderful and supportive mentor!