Peptide Puzzle Pieces: Perceiving Peculiarly Produced Proteins

Jackson Eberwein, Sonoma State University

The Approach: In my previous post, I discussed some of the problems about Domoic Acid Toxicosis in California sea lions, closing with the potential benefit of finding a biomarker protein. The first step in finding anything is to take a good look! How does someone look at something as small as a protein, though? When trying to see which proteins are in a sample of blood plasma and count how many there are of each, it can get complicated. The way I am achieving this is by using an instrument called a mass spectrometer. With it, I can predict the identity and amount of each protein in a sample. The mass spectrometer is picky, however. To read my proteins, it wants them to be cut up first. 

Orbitrap mass spectrometer (right) with attached liquid chromatograph (left).

Proteins are similar in structure to a tangle of string. The tangle first has to be unraveled, then it is cut up into small bits called peptides. These peptides are put into the mass spectrometer to be read. The shape of each peptide is pretty unique, and that unique shape is used to detect and measure them. With the help of some very handy computer programs, peptide measurements can be compared to a California sea lion genetic database to predict the protein that each peptide came from and how many of those proteins there might have been in the sample. Once we have the names and amounts of the proteins in the sea lion samples, the protein differences between each sample can be looked at. This is where we look for our biomarker. If there are one or more proteins that appear at consistently different levels in sea lion samples with Domoic Acid Toxicosis than in samples without it, those proteins have potential as good biomarkers! 

Acknowledgements

I would like to thank Dr. Michael Janech, Dr. Benjamin Neely, Alison Bland, The Marine Mammal Center, & College of Charleston. Supported in part by the Fort Johnson REU Program, NSF DBI-1757899.

Fresh stacks of muddy bacteria

Lilia Garcia, Illinois Wesleyan University

The Approach: In my last post, I wrote about Gracilaria, an invasive red seaweed on the coast of South Carolina, and its effect on Vibrio bacteria. My project aims to record the number and strains, or types, of Vibrio growing around Gracilaria and compare it to seaweed-free areas. I will also compare the Vibrio count residing on Gracilaria versus the Vibrio residing on a native seaweed called Ulva to see how an invasive species changes the bacterial community. Lastly, I want to understand how Gracilaria stops the growth of specific Vibrio strains by producing chemical compounds.

Mud samples under Gracilaria, taken by K. Coates

To begin solving my questions, I will go out to collect samples in the mudflats outside of the Grice Laboratory. I will collect tubes of water, clumps of Gracilaria and Ulva, and mud from underneath and 1.5 feet away from Gracilaria. Afterwards, I’ll spread all the samples onto dishes with nutrients specifically used to grow Vibrio. The bacteria grow in spots called colonies, and I will count each spot to see how much Vibrio there is in each sample. I am looking for a different amount of colonies in mud samples collected within or away from Gracilaria patches, and a difference in colony numbers between the Ulva and Gracilaria.

Dishes of unique Vibrio, taken by L. Garcia

A single dish from a mud sample can contain hundreds of colonies, differing in color, shape, size, and texture. Each of these colonies represent a different strain of Vibrio, uncovering the diversity of bacteria at different distances from Gracilaria. I will characterize which unique colonies are dangerous to human health, and whether they are found near or away from Gracilaria.

Zones of inhibition against Vibrio strain, taken by L. Garcia

As previously mentioned, I will also test Vibrio strains against chemical compounds made on the surface of Gracilaria. These compounds are able to control the kind of bacteria that grow around seaweed, changing the microscopic habitat. I will mix Gracilaria with chemicals to remove its surface chemistry, then spot the compounds onto dishes growing Vibrio from my mud samples. I am looking for large clear circles, called zone of inhibitions, that tell me the specific strain of Vibrio cannot grow due to the compound.

Nearly all we know about the ecological and economic impact of Gracilaria focuses on large animals, such as fish. My project zooms in on micro-organisms that have been overlooked. The information I collect will help us understand how invasive Gracilaria is changing bacterial communities not only in the Charleston Harbor, but potentially the entire coast.  Although invisible, bacteria make up the foundation of ecosystems and high Vibrio levels may be dangerous for our health. I look forward to finding the answers to my questions hiding quietly in the mud.

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

Larval Phthalate Soup

Samuel Daughenbaugh, DePauw University

57F1DBA6-B9C6-49AA-B37D-4119CD8D2753

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).

58247769228__EF1A094D-D648-4B03-A368-6C1C68BCF865

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.

pasted image 0

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.

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.

Counting Corals

Jordan Penn, Millersville University

The Approach: In my last post, I discussed that the consequences of habitat-degrading practices (e.g., bottom trawling, dumping of waste, drilling) include the loss of species such as gorgonian corals, which provide structural habitat for other species.

My research seeks to understand the relationship between soft corals and their geological substrate. In other words, our lab want to understand whether or not soft corals are more likely to be present on rocky or sandy sea floors. We are also looking for relationships between the abundance of soft corals at different depths. We are investigating these relationships in order to gain some understanding of where soft corals are most likely to be found. 

Example of a transect with three segments. Image credit: Science X.

In order to assess these potential relationships, first we need to divide the video footage of dives from the ROV (remotely-operated vehicle) Beagle into 15-minute transects containing 3 5-minute segments. We take this step in order to determine the density (number of individuals per square meter of area) of corals at each site as accurately as possible.

Next, I will analyze the video footage, counting each Leptogorgia, Acanthogorgia, Eugorgia, Adelogorgia, and sea pen (these are good model organisms because they are conspicuous in our study site). Along with the number of corals, I will denote the type of substrate that was dominant throughout the 5-minute segment (e.g., rocky bottom, sandy bottom, mixed/coarse bottom).

Finally, I will be able to run statistical analyses on these data to determine average density, the average deviation from the determined average density, and potential drivers of diversity at each site (e.g., does depth/bottom type/something else affect how many corals are present in an area?).


Thank you to the members of the Etnoyer Lab for their guidance and assistance as well as the Grice Lab and College of Charleston for funding this project. This project is supported by the Fort Johnson REU Program, NSF DBI-1757899.


References

NOAA. (2012, April 17). NOAA releases new views of Earth’s ocean floor. Retrieved June 17, 2019, from https://phys.org/news/2012-04-noaa-views-earth-ocean-floor.html NOAA

All Shrimp Go To Heaven

Carolina Rios, New York University

The Approach: In my previous post, I discussed the negative impact that polychlorinated biphenyls (PCBs), long-lived chemical contaminants, can have on marine ecosystems and human health. Currently, I am working to verify a proposed model to estimate the impact that PCB contamination has on benthic marine invertebrates.

One of the issues with this proposed model is that it is based on data generated from the 1970s; and the analytical methods now available to scientists are much more sensitive and precise. To verify this model, we will generate contemporary data by running a series of short (acute) toxicity tests.

Testing

Test setup for grass shrimp (P. pugio)

In these acute toxicity tests, we measure the response of three species of marine invertebrates to PCBs. The three organisms that we are testing are grass shrimp (Palaemonetes pugio), amphipods (Leptocheirus plumulosus), and mysids (Mysidopsis bahia). We will be measuring mortality from PCB contamination. The standard tests that we are running consists of 6 concentrations, ranging from 6.25 ppb (parts per billion) to 420 ppb. It is important that we also have a control, so that we can understand the response of the organisms unaffected by PCBs. For the grass shrimp and amphipods, the test will run for 96 hours and we will renew the PCB solutions every 24 hours. Samples will be taken for chemical analysis at 0 hours, 24 hours, and 72 hours, so as to measure both the loss of PCBs over the 24 hour period, as well as the consistency of dosing. Loss of PCBs can be attributed to PCBs binding to the glassware and differences in dosing can be attributed to user variability. For the mysids, the test will also run for 96 hours, but the dosing solutions will not be renewed after the initial dosing. Samples will be taken at 0 hours, 48 hours, and 96 hours, so as to measure the loss of PCBs over the 96 hour period. For all tests, mortality will be recorded every 24 hours until the end of the test.

Analysis

Solid Phase Extraction apparatus. Dosed samples are within the large reservoirs at the top of the apparatus. PCBs will be isolated on the nonpolar solid phase, which consists of the smaller columns below the reservoirs.

The PCBs from the collected samples will be isolated through solid phase extraction. Solid phase extraction consists of a nonpolar solid phase and a polar liquid phase; similar to how oil cannot be mixed into vinegar, PCBs are not very soluble in water. As PCBs are nonpolar and hydrophobic, they will bind to the solid phase. The PCBs can then be lifted off of the column by running a nonpolar solvent (ethyl acetate) through the nonpolar solid phase. The sample is then analyzed using gas chromatography-mass spectrometry (GC-MS). The essential concept of GC-MS is that molecules will separate based on differences in size. This is how the amount of each individual PCB is determined, which can then be used to calculate an actual concentration. Analysis of the chromatograph can give more accurate concentrations, allowing us to understand how concentrations vary over time. This will give us a better understanding of the relationship between dose concentrations and the mortality response.

Acknowledgements

I would like to thank Dr. Ed Wirth and Brian Shaddrix for their continued guidance and support, as well as my co-mentor Dr. Paul Pennington. Supported by the Fort Johnson REU Program, NSF DBI-1757899.

Cloning our way to a perfect sequence

Kelsey Coates, Duquesne University

The Approach: In my first blog post, “FROM FEMALE TO MALE – MUD SNAILS TELL ALL!,” I described the goal of my research, to sequence isoforms of a hormone receptor called the Retinoid X Receptor (RXR) in the eastern mud snail.  

Mud snails all over a beach at Fort Johnson, SC.

These isoforms have yet to be sequenced in the mud snail! But what exactly is a DNA sequence? DNA is made of building blocks called nucleotides. A DNA sequence is the order of the nucleotides. A sequence like ACG could tell the organisms’ body to do one thing while a sequence like AGC could tell the organisms’ body to do another. A bit of the sequence has already been identified, but there is a gap in the sequence we are still trying to figure out.  

Theoretically, different chemicals or different concentrations of the same chemical can change the relative levels of the RXR isoforms. If this hypothesis is confirmed, mud snails can be used in the future to detect contaminants that affect marine organisms in the Charleston Harbor. Their patterns of isoform expression might suggest which seasonal contaminants are present in the environment where they live. For example, chemical one may trigger isoform A which has sequence ACG while chemical two may trigger isoform B which has sequence AGC.            

So how will we get these sequences? It starts with amplifying the known sequence of the mud snail that surrounds the isoform, including the mysterious gap. Amplification will be done by polymerase chain reaction (PCR) to ensure there are thousands of copies of the DNA to work with. After purification, the sequence is ready to be incorporated into a plasmid along with an antibiotic resistance component. Bacteria, like E. Coli, store their DNA in plasmid form compared to the double-helix form of humans.

Plates of E.Coli in the presence of Ampicillin set in the incubator.

Luckily for us, plasmids are easily manipulated and are reproduced rapidly in bacteria. E. Coli will be grown in the presence of the antibiotic ampicillin with the sequence we cloned into its DNA.  If the sequence is incorporated into the plasmid, the bacteria will have anti-biotic resistance and be able to grow on the ampicillin plates. The bacterial colonies with our plasmid will be PCR amplified. Then, after a final plasmid preparation, the samples from E. Coli can be sent to a lab that specializes in sequencing. Hopefully the lab will identify the gap and we will achieve our goal!

ACKNOWLEDGEMENTS

I would like to acknowledge Dr. Demetri Spyropoulos, Edwina Mathis, Dr. Bob Podolsky, The Fort Johnson REU Program, The Hollings Marine Lab, NOAA, and The Grice Marine Lab. This research was supported by the Fort Johnson REU Program, NSF DBI-1757899.

This Is How We Do It ♫

Julianna Duran, Virginia Tech

2AF7E921-9048-492D-9C03-181A396A1CC7

First and foremost, if you didn’t get the reference in the title please click here!

Now that I have educated you on the topic of music, let’s switch to science.

 The Approach: In my previous post I mentioned that I am studying the lipids of Nile Crocodile and Mozambique Tilapia. So the first thing I did is wrestle the reptile like Steve Irwin and hand catch my fish – just kidding, but imagine how cool that would be! My samples were collected from Lake Loskop, South Africa in 2014. Once they were in my possession, here is what I did.

  1. Sample Preparation
    • The muscle tissue samples I received looked like chicken breasts you buy from the grocery store – except the size of a fat bean. These solid chunks need to be turned into a fine powder for me to analyze them. This was done by freezing the sample in the cryomill machine – where the samples were shaken extremely fast and broken up

      Cryomill

      Cryomill

  2. Extraction
    • Think of what happens when you pour oil in water. They go to different ends and don’t mix, right? (Yes) That is exactly what I’m doing with my samples. We are adding lots of chemicals to break down fats into their building blocks: Fatty Acids! The muscle layer (organic layer) hates touching the chemicals, so I take that out and can use it for my next step!
    • Check out a video I made of one of my extractions
  3. Gas Chromatography
    • This instrument is how I will measure the amount of each fatty acid in my samples.
    • How does it work?
      • The sample is injected into the system and enters a narrow glass column. The sample separates in this column based on its weight and boiling point. The particle encounters a flame at the end of the glass, which detects what specific fatty acid it is. The computer then gets this signal and generates a graph showing a fatty acid profile. Each peak on the graph is a different fatty acid, and the height of the peak indicates how much of it there is in the sample.
      • For help envisioning this process, take a look at this video (I used it when I learned about this instrument!)

        blue

        Chromatogram

Summary:

I will be physically and chemically breaking down my samples, then getting fatty acid profiles for each of my individual species. This is all to see if there is a difference between healthy and diseased species and what lipids are most affected by Pansteatitis!


Supported by the Fort Johnson REU Program (NSF DBI-1757899), Dr. Mike Napolitano, Dr. John Bowden, The College of Charleston, NOAA, and NIST. 


References:

CryoMill. https://www.retsch.com/products/milling/ball-mills/mixer-mill-cryomill/function-features/ (accessed Jun 18, 2019).

How to Train Your Shewanella

Katherine Mateos, Carleton College

The Approach: In my previous post, I introduced my project, investigating the role of Antarctic bacterium, Shewanella BF02, in the cycling of volatile organic sulfur compounds (VOSCs). 

Sterile technique in action Photo Credit: Peter Lee

The first order of business in this effort is keeping the Shewanella alive and happy. In order to do this in the lab, I make a liquid (known in the biology world as “medium”) for the Shewanella to live in. Our medium is designed to resemble Blood Falls in chemical makeup. In particular, it is very salty, and contains iron and sulfate. I am also careful to remove all the dissolved oxygen in the medium, since the Blood Falls water has very little oxygen. In my medium, I am also careful to keep out any bacteria other than my Shewanella. Since microbes are everywhere, including in the air, on my skin, and on the lab bench, I  use a special set of techniques to avoid unwanted bacteria from infecting my samples. 

Membrane Inlet Mass Spectrometer

Once we have a perfect mix of chemicals for Shewanella, I also add my target organic sulfur compounds. Because I want to see if Shewanella changes these added compounds, I keep track of them using a technique called isotope labeling. Isotope labeling is a clever trick, where the target compounds are tagged with atoms that are the tiniest bit heavier than the ones that we usually see. If Shewanella make the labeled compounds into the VOSC products that I am interested in, those products will also have the same tag, making it easy to identify them.

To identify the tiny differences in mass between tagged and untagged molecules, I use a piece of equipment called a mass spectrometer. A mass spectrometer works kind of like a scale and can determine the mass of each molecule. This allows me to detect isotopically labeled VOSC products. If I see isotopically labeled products, I can be pretty sure that the Shewanella are cycling the labeled compound that I added to their medium. 

Thank you to my mentor, Dr. Peter A. Lee, and our collaborators, Dr. Jill Mikucki and Abigail Jarratt, for their guidance in the research process. This project is supported by the Fort Johnson REU Program, NSF DBI-1757899.

Algae Microbiome: The Hidden World

Pressley Wilson, University of South Carolina Aiken

The Approach: In my previous post, I discussed (1) the importance of an organism’s microbiome in relation to its health and (2) the importance of algae in marine ecosystems due to their ability in producing oxygen, removing nitrogen and phosphorus from water, and exchanging inorganic carbon.

Considering the importance of these two variables, this summer I am researching the relationship between the algae microbiome and algae species in One’ula Beach, Honolulu, Hawai’i.  

DNA extraction in progress. Photo credit: Dr. Heather Fullerton.

The objectives of my research project are:

  • Identify relationship between algae microbiome and algae species
  • Identify relationship between the microbiome with algae’s morphology

Prediction

This study will predict there is a variation in the microbiome between algae species, due to the different species characteristics, such as calcification.

Sample Collection

The five algal species of different morphologies were hand-sampled by Dr. Heather Spalding at the intertidal region of One’ula Beach. The algae samples were rinsed with artificial saltwater to remove dirt and loosely associated bacteria. After cleaning, each species were placed into (1) a micro-centrifuge tube with 0.5 mL of RNA-later or (2) a 15 milliliter conical tube with 1.5 mL RNA-later. The RNA-later is a DNA preservation agent, was used to stabilize the algae DNA. The samples were stored at 4°C overnight. This overnight incubation allowed the RNA-later to penetrate the bacterial and algal cells to the DNA. After this incubation all tubes were frozen at -20°C and shipped overnight on dry ice to the College of Charleston, South Carolina, where they were stored in  -80°C freezer until DNA extraction.

Sample Analysis a

A completed gel electrophoresis.

A MoBio Fast DNA Spin Kit was used to extract the DNA from the algal samples. This DNA is then tested by PCR to determine if bacteria are present on the algae. To determine the number of bacteria present, qPCR is used. This molecular biology technique is used to quantify specific genes in a sample.

The PCR samples will be analyzed using gel electrophoresis, a molecular biology procedure that uses an electrical current to separate the components of the sample DNA by size. The qPCR data will be compared to a known DNA standard to determine the number of bacteria in our samples and calculations will be performed using excel.

Acknowledgements

I would like to thank Dr. Heather Fullerton for her guidance and support with this project and Dr. Heather Spalding for her sample collection. This project is supported by the Fort Johnson REU Program, NSF DBI-1757899.