Today is the end of week 3 of this program! It was much more independent than the previous two weeks and I continued to gain valuable, hands-on lab experience. Clark came in over the weekend to wrap up the three-day IPTG induction process we began on Friday. Starting Monday, I worked on creating and analyzing an iodixanol gradient as well as making more SDS-PAGE gels and western blots. On Thursday, we went to A&M to view our proteins under a cryo-electron microscope! This was a really exciting opportunity as these machines cost upwards of $6 million. Then finally, we wrapped up the week with a relatively easy day today, doing transmission electron microscopy and working on reading papers and gaining conceptual knowledge.

Monday – Iodixanol Gradient Viral Pellet Purification

Monday started off with pellet sonication and lysate clarification. I worked mostly independently throughout this procedure. I started off by preparing an ice bucket as well as a water-ice mixture in a different bucket–the sonication bucket. Then, I prepared lysis buffer by pouring sonication buffer, BME, and PMSF into a conical tube and then pouring all of this mixture onto my cell pellet (which Clark made over the weekend). PMSF was added because it keeps the lysate stable, preventing endogenous proteases within the cell from interfering with the lysate, which would cause proteolysis of the sample. BME was added to the lysis buffer because it is a reducing agent that would prevent oxidative damage to the target protein during the purification process. Moreover, reducing agents such as BME also break disulfide bridges in proteins, thus helping partially denature proteins during purification. Once the pellet was submerged in the lysis buffer, I cleaned the sonicator probe by spraying it with DI water, 70% ethanol, and wiping it off, repeating this process once more. Then, I placed the probe into the sonication buffer that is sitting on the cell pellet, placing it very close to or right on top of the pellet. The settings of the sonicator were as follows: 7 minutes, 70% amplitude, pulse on: 01, pulse off: 03. Then, I cooled the floor centrifuge in our lab because it would be used in later steps. Once the cell pellet had completely broken up and there were no floating chunks or clumps of cells, I poured the lysate into a centrifuge tube that is compatible with the floor model centrifuge. Then, I grabbed another centrifuge tube to balance it with the lysate with simple water, very carefully ensuring that it was balanced correctly as unbalanced centrifuge runs can be dangerous at high speeds. The centrifuge was run at 30,000 x g for 45 minutes at 4 degrees. During the spin, I prepared a conical tube to collect the lysate, and once the spin was completed, I immediately decanted the supernatant into the prepared tube. Then, I pipetted sonication buffer onto the pellet, without any BME or PMSF, and resuspended until the buffer was cloudy (the pellet does not have to be completely resuspended). Finally, I took samples of the pellet and supernatant for SDS-PAGE gel and western blot. This marked the end of Monday, a very productive day!

Tuesday – Iodixanol Gradient Viral Pellet Purification

On Tuesday, I started off by preparing an iodixanol gradient. This was done by preparing iodixanol solutions of 10%, 20%, 30%, 40%, and 50% using the sonication buffer. No BME or PMSF was necessary. Then, I took 2 polystyrene ultracentrifuge tubes and added 10% iodixanol into both and poured the lysate into one and poured sonication buffer with BME into the other. Clark then placed the ultracentrifuge tubes into the Sw41-Ti rotor tube holder, balanced them, and spun the samples at 37,500 rpm in the rotor for 2 hours at 4 degrees. During the spin, the iodixanol centrifuge gradient was set up. To do this, I prepared two polystyrene ultracentrifuge tubes, adding 1 mL of 10%, 2 mL of 20%, 2 mL of 30%, 2 mL of 40%, and 2 mL of 50% to each. Then, in one tube, we added 1 mL of the supernatant and in the other tube, we added 1 mL of sonication buffer with BME. The tubes were once again balanced to 10 mg and spun at 37,000 rpm for 22 hours at 4 degrees in the Sw41-Ti rotor. Once this spin was done, on the next day (Wednesday), I used a syringe to remove the bands from the tube by drilling the needle through and simply extracting. I added this snippet to the section for Tuesday so I could logically complete this procedure within one section.

Wednesday – SDS-PAGE and Western Blot

On Wednesday, we ran both an SDS-PAGE gel and a western blot. Starting with the SDS-PAGE gel, I started off by turning on the heat block near the gel running station. Once the temperature reached 98 degrees Celsius, I put the samples of the viral pellet, supernatant, and all 3 bands from the iodixanol gradient onto the heat block. First, I heated it for 2 minutes and vented the tubes by rapidly opening and closing them. Following this, I heated the samples for 8 more minutes. While this was being done, I removed the ladder from the freezer to thaw so I was prepared for the SDS-PAGE. Once the samples were warmed up, I placed them into a small centrifuge and pulsed to get rid of the condensation inside of the tube. I removed pre-made gels (that I made last week) from the fridge, removed the well comb, and set up the gel cassette. Since I was only running one gel, I used a buffer dam. I then placed the gel cassette into the gel running box and poured running buffer into the cassette until it was completely full. Next, I loaded the gel and ensured that all wells had the same total volume to equalize the electrical potential across the entire gel and ensure that it will run straight. I am definitely getting a lot more accurate and efficient with pipetting the samples into the wells–a process that was very difficult for me in week 1! Practice makes perfect holds true. Once the gel was loaded, I ran it at 100 V until the dye fronts tightened up on the separating gel. This took about 15 minutes. Then, I increased the voltage to 180 V until the dye front has run off the gel.

^ The dye fronts tightening up on the separating gel.

For organization’s sake, I will now discuss staining and de-staining the gel and write about the western blot following that, although I was doing these processes simultaneously. This is because both have gaps of time where something has to sit for around 20 minutes, which is ample time to achieve something else.

Moving on, I stained and de-stained the gel with Coomassie blue dye. I started by grabbing an empty pipette tip box that’s used for staining and put it on the gel running bench. I got a green wedge gel-cracker and separated the front and back glass plates. This entire leg of the SDS-PAGE process has to be done very carefully–if any mistake is made, it could result in the gel tearing and the results being lost. The gel cracker was then used to cut the stacking gel off and discard it. I cut the gel into half: both halves were the exact same, each of the samples was doubled across the gel because half of the gel would later be needed for the western blot. Then, the half of the gel that would be stained was removed from the plate (gently, of course) and placed into the empty pipette tip box. Staining buffer was poured into the gel and it was then microwaved. Once it was microwaved, the box was moved to the fume hood to let it cool and vent for a few moments. The staining buffer was then poured into the used staining solution bottle using a funnel and the gel was rinsed using DI water. De-staining buffer and two Kimwipes were placed there to absorb the remaining staining solution. This was left on the rocker until the gel was de-stained, and a couple hours later, Clark labelled and scanned the gel.

Now for the western blot! First I retrieved the western blot transfer buffer from the fridge and put it into the bucket–with this buffer, you have to be extremely cautious because it contains methanol, which is highly toxic if ingested. Then, the western blot transfer buffer was poured into the transfer box (an apparatus used for western blotting) until it was about half full. Some filter pads were then soaked in the buffer. While they were soaking, I grabbed a square plate and filled the bottom with methanol, then placing a blot-sized portion of PVDF into it using forceps as you should not touch the paper–even with gloves. Then, I grabbed the other half of the gel to build a gel “sandwich” with. It was placed into the filter paper on one side of the holder and the activated gel was placed on top of it. Then, the air bubbles were removed and the apparatus was clipped shut. A cooling block was added inside the transfer box. Then, transfer buffer was added up until it reached a fill line. The western blot was run using the power source stored inside the fridge. It was run at 90 V for one hour. 10 mL of 10% milk in TBST was then prepared, and once the transfer was done running, the membrane was removed and put into a square plate with all of the 10% milk. It was allowed to block for 30 minutes on the shaker. 10 mL of 5% milk was then prepared, and 1 microliter of primary antibody was added (anti-6X his antibody). The blot was rinsed with TBST until it ran clear, and the antibody-milk solution was poured onto the blot. Today, for time’s sake, it was only allowed to block for 1 hour at room temperature for a faster result. Then, it was washed down 3 times for 10 minutes each using TBST. I then made 10 mL of 5% milk and added 2 microliters of secondary antibody (cross-absorbed goat anti-mouse antibody). The last TBST wash was poured off the blot and the secondary antibody solution was added. It was then incubated at room temperature on the rocker for one hour. Then, it was washed 3 times for 10 minutes each with TBST once again. Following this, the blot was rinsed with some Milli-Q water and then submerged in developing solution (but not too much). Then, I waited for faint bands to develop. Once this was visible, the reaction was inactivated by rinsing with Milli-Q water. The develop blot was then clipped and placed into the incubator. Once it was dry, it was placed into a plastic sleeve, labelled, and scanned by Clark.

Thursday – A&M Trip!

Today, I rode the 6 AM bus to reach Rice by 7 AM and drive to A&M with Clark, Nora, and Kai. We reached A&M at about 9 and Dr. Gaya Yadav, the Cryo-EM technical director, assisted us throughout the day in imaging our samples. The microscope is a very expensive piece of equipment and thus he was the only one allowed to even be in the same room as it! Since we weren’t allowed to do much today and the day primarily involved a lot of waiting, I decided to read up on Cryo-EM and what it does. Cryo-EM is mainly used to determine the structure of biological macromolecules and macromolecular super complexes. The biological samples to be imaged (which could be proteins, viruses, or small cellular components, but in our case were proteins) are rapidly frozen by plunging into liquid nitrogen, a cryogen. This quick freezing process prevents the formation of ice crystals, which can damage the sample, and instead forms vitreous ice that preserves the sample’s native structure. The frozen sample is then placed into the microscope. An electron beam is passed through the sample and the interactions between the electrons and the sample are captured on a detector, creating high-resolution images. The sample is maintained at cryogenic temperatures (around -180 degrees Celsius) to prevent any structural changes or damage. Thousands of 2D images are collected from different angles, using a technique called single-particle analysis, where individual particles are imaged and then computationally combined to reconstruct the 3D structure. The collected 2D images are aligned and averaged to reduce noise and improve the signal. The resulting 3D structure is analyzed to understand various things regarding the biological molecules, including their molecular architecture, functional mechanisms, and their interactions with each other. This was pretty much all we did today–it was a 12 hour day but I enjoyed it thoroughly and learned a lot from just watching Dr. Yadav work!

^ Low quality image of the room with the Cryo-EM in it (we weren’t allowed to go into the room).

Friday – Transmission Electron Microscopy

Today, we decided to take a relatively easy day and focused on doing transmission electron microscopy of some samples. Once again, this is an expensive piece of equipment that I was not allowed to use, but I learned a lot about its functionality from just watching Clark use it. To prepare TEM samples, they have to be made extremely thin, typically less than 100 nm thin to allow electrons to pas through it. Then, they are stained with heavy metal salts, such as uranyl formate (this part was done by Clark as there is alpha radiation involved), as these metals scatter electrons effectively due to their large nucleuses. The electron microscope uses an electron gun to produce a beam of electrons, and this beam is typically generated by heating a tungsten filament or using a field emission gun, which emits electrons when a strong electric field is applied. The electron beam is then focused and directed onto the sample using a series of electromagnetic lenses, which function similarly to glass lenses in light microscopes but use magnetic fields to manipulate the electron beam. The first lens, called the condenser lens, concentrated the electron beam onto the sample. As the electron beam passes through the thin sample, electrons are scattered by atoms in the sample, and the degree of this scattering depends on the density and thickness of the sample as well as the atomic number of the elements present. Areas where more electrons are scattered appear darker in the final image, while areas with less scattering appear lighter, thus creating a contrast that reveals the internal structure of the sample. After passing through the sample, the electron beam is focused by an objective lens to form an initial magnified images. The final image can be observed directly on the screen or captured digitally for further analysis. These images are then analyzed to study the fine details of the sample’s structure, including atomic arrangements, crystallographic information, and even potential defects.

All of this TEM process was done in a dark room. We got great, visible results for the first grid (we analyzed four) but the rest seemed as if the staining was poor because it was grainy and unclear. Nevertheless, this is how science works, and not every experiment is a success-we will just remake the samples and repeat this process. We already plan on adding detergent and adjusting the pH of the samples (lowering it) to see if that changes anything.

^ The clear image for TEM.

^ Clark, Andy, and I in the TEM room.

^ What it looks like when focusing the microscope.

Overall, this week was a very productive week and I gained a lot more insight into the project and overall day-to-day lab procedures. Moreover, as a high schooler, this internship is really helping me understand what I want to do in the future. I’m now thinking of creating some sort of start-up to propel environmental causes forward, just like how Clark did this entire project to determine how dinoRNAVs impact the health of Symbiodiniaceae, a key coral symbiont. There are hundreds if not thousands of people working on various vaccines and health treatments–which are extremely important–but in the process, we have neglected the health of our planet and environment as a whole, which I want to focus my career on changing.