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!




Playing with Plutei

Hailey Conrad, Rutgers University


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!


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!


A day in the Shrimp Lab


Alessandra Jimenez, Whitworth University

Have you ever wondered what it’s like to be a lab researcher who works with live animals? Through this internship, I am experiencing this firsthand in Hollings Marine Laboratory, along with all the responsibilities involved!

A normal workday in the life of a “shrimp intern” is like this: A big part of it is animal care and maintenance. It starts in the morning with a daily visit to the wet lab, where approximately 80 brown shrimp juveniles are kept in four large tanks with circulating water. After feeding them a round of commercial shrimp pellets, I test the salinity of the water in each tank using a refractometer to make sure that each tank has a certain salinity value: 30 parts per thousand, to be exact. I use dechlorinated freshwater and seawater to adjust this value if needed. Besides salinity, I also need to watch out for harmful levels of ammonia (it’s a part of shrimp waste!), nitrates, etc. In usual circumstances, I conduct a water change (replacing old water with new) once a week in order to dilute these chemicals. For the past couple of weeks, however, I have been conducting water changes daily in order to keep ammonia levels neutral in three tanks. Ah, the life of a caretaker of tons of baby shrimp!


Wet lab. @AlessandraJimenez

Besides animal husbandry, I work on my experiment involving the effects of injection of bacteria on tail flipping (Want to learn more about what I’m doing? click here). I have two shrimp at a time in separate, well-aerated tanks, and they are both from the same treatment group. Shrimp are randomly assigned to one of four treatment groups. These treatment groups are designated according to the treatment type (injection of bacteria or saline) and according to the amount of time between the moment of injection and the tail-flipping procedure (4 or 24 hours). I randomly select two shrimps from the wet lab, weigh them, and keep them in the two experimental tanks overnight so they can get used to the new environment, temperature, etc. The next day, I take each shrimp out of the tank momentarily and quickly inject them with bacteria, or a saline buffer if they are part of the control group. Then, I give them 4 or 24 hours (depending on group type) to rest before conducting the actual tail-flipping experiment. Using a stir-rod (basically, a straight stick), I poke the shrimp lightly to induce tail-flipping, and count how many flips they perform before fatigue. The number of flips here is called ‘initial activity’. Then, I give them 20 minutes to recover in the tank before tail-flipping them again. The number of flips this time is called ‘recovery activity’.


Experimental tanks @AlessandraJimenez

Why tail-flip them twice? Well, we hypothesize that recovery activity will be impaired in bacteria-injected shrimp versus the controls, while initial activity would probably not be. This is based on how recovery from tail-flipping activities involves aerobic (or oxygen-fueled) metabolism. Since bacteria accumulate in the gills of shrimp and block oxygen uptake (want to learn more? click here), it would make sense that recovery activity would be reduced. Stay tuned for results later on!

Works Cited:

Gruschczyk, B., Kamp, G., 1990. The shift from glycogenolysis to glycogen resynthesis after escape swimming: studies on the abdominal muscle of the shrimp, Crangon crangon. J Comp Physiol B, 753-760.

Scholnick, D. A., Burnett, K. G., & Burnett, L. E. (2006). Impact of exposure to bacteria on metabolism in the penaeid shrimp Litopenaeus vannamei. Biological Bulletin, 211(1), 44-49.

Many thanks to College of Charleston for hosting my project, Dr. Karen Burnett and Hollings Marine Laboratory for guidance and work space, and NSF for funding the REU program.