This marked the last week of this research experience! We did not stop experimentation, however!

Key highlights of this week:

Monday

Today, we began the process of IPTG induction. In this process, a gene coding for the desired protein is inserted into E. coli and placed under the control of the lac operon, a natural regulatory system that controls the expression of certain genes in response to the presence of lactose. IPTG (Isopropyl β-D-1-thiogalactopyranoside) is a synthetic analog of lactose that cannot be metabolized by E. coli. When IPTG is added to the culture, it binds to the repressor protein that normally keeps the lac operon turned off, causing the repressor to release from the DNA. This release allows RNA polymerase to bind to the promoter region of the operon and initiate transcription of the gene, leading to the production of the target protein. IPTG induction is favored for its ability to precisely control the timing of protein expression, ensuring that the bacteria can grow to a sufficient density before protein production begins, which often results in higher yields and more efficient use of resources.

We began by making LB broth and autoclaving it. Once the LB was made, we let it cool. Then, we made overnight cultures in 50 mL tubes using previously made LB, stored at the bench. These cultures were done as per usual: carbenicillin and chlorophyllin were added to LB as selective antibiotics. Then, one colony was picked off of a previously transformed plate of E. coli, transformed with Levin MCP and construct 9337. These two overnight cultures were incubated in a 37 degree shaking incubator overnight.

Tuesday

Two flasks with LB were inoculated with the liquid cultures from the previous day and stored on the shaking incubator in the environmental warm room until the optical density reached 0.5-0.6. This was determined by regular checking of the OD of these two flasks. Once it reached this OD value, we carried out the process of IPTG induction. We first made 3 mL of 1 M IPTG. Then, we pipetted in 1 mL of IPTG into both of the flasks and stored them overnight in a shaking incubator at 16 degrees and 220 RPM. This marked the end of Tuesday’s work. As a side note, a lot of this week went into practicing presentations.

Wednesday

We prepared DNA for sequencing today. It involved a lot of new procedures and also involved PCR. After the plasmid minipreparation, a standard lab procedure, was complete, we tested the cell count using a nanodrop, and when we determined it was sufficient, we sent it across the street for sequencing at the Med Center.

Moreover, we pelleted, lysed, and spun the cells from yesterday. We then took samples of the supernatant and pellet. We also washed our nickel column with 8 M urea and stored the supernatant and beads in a 50 mL conical tube for elution tomorrow.

Thursday

Today, we eluted the protein using 2 concentrations of imidazole: 50 mM and 300 mM. The 50 mM served as a wash, while the 250 mM served as an elution. Using standard Ni-NTA protocol, we washed and eluted the protein, took samples of everything (including the supernatant and pellet from yesterday), and ran an SDS-PAGE gel. The results were overnight, so we will find out tomorrow whether or not it succeeded in purifying the protein an adequate amount.

I am very excited for the presentation tomorrow! Good luck everyone!

This week, I did molecular cloning for the first time! It was a great learning experience.

Monday

Today was focused primarily on reading and poster design. The poster is still a work in progress, but we got valuable results this week that will be going on the poster!

Tuesday

Today, me and Clark began the three-day molecular cloning process to test its functionality. To set up the reaction, we added 10 microliters of 5X Q5 reaction buffer, 1 microliter of 10 mM dNTPs, 2.5 microliters of 10 micro-molar forward primer, 2.5 microliters of 10 micro-molar reverse primer, 1 microliter of template DNA (specially designed), 0.5 microliters of Q5 high-fidelity DNA polymerase, and filled up the remainder of the tube with nuclease-free milli-Q water. This reaction was gently mixed and the liquid was all collected to the  bottom with a quick centrifuge spin. The PCR tubes were then transferred to a PCR machine and the thermocycling procedure began. Initial denaturation was at 98 degrees Celsius for 30 seconds, then 35 cycles were run, and the final extension was at 72 degrees Celsius for 2 minutes. Finally, it was held at 10 degrees Celsius infinitely, then stopped on the next day.

This was a very lengthy procedure. I want to go ahead and explain the principle behind PCR. PCR is the abbreviated form of polymerase chain reaction, which is a laboratory technique for rapidly producing, or amplifying, millions to billions of copies of a specific segment of DNA, which can then be studied in greater detail through agarose gels (which is what we did in our experiments). The PCR process has six steps: initialization, denaturation, annealing, elongation/extension, repeated cycles, final elongation, and final hold. Before I explain the steps, I’d like to explain the components that go into PCR. There are five main ingredients needed for PCR. Polymerases are the first ones; they are enzymes that, when subjected to ambient conditions, are able to assemble new strands of DNA from template DNA and nucleotides. The next component is the template DNA. This is the DNA that the polymerase (typically Taq) will read and copy; this DNA can be genomic, plasmid, or cDNA, and in our experimentation, it is plasmid DNA. Next, PCR requires primers, which are short fragments of synthesized DNA that bind to your specific template DNA. These have to be specially designed, we designed and ordered ours last Friday. The forward primer designates the start of PCR and its sequence is the same as the 5′-3′ template DNA sequence. The reverse primer designates the end of PCR, and its sequence is the reverse complement of the template DNA. Next, you need nucleotides. As the monomers of DNA, nucleotides are necessary for making copies of DNA. For most DNA PCRs, deoxynucleotide triphosphates (dNTPs). Finally, you need buffers. For the purposes of our research, we utilized the Q5 buffer, which helps optimize DNA denaturing, renaturing, and polymerase activity.

Next, I’d like to explain thermocycling. The above ingredients are added to a PCR tube, and the tube is thermo-cycled. The first step is initialization, in which the reaction is heated to 98 degrees in order to activate hot-start polymerases to denature the template DNA. Next is denaturation, in which the DNA and primers are denatured to be able to efficiently and effectively anneal to each other in the next step, which is annealing. In annealing, the reaction’s temperature is rapidly lowered so that the denatured primers can form Watson-Crick base pairs with the template DNA. However, the temperature is not significantly lowered as it still must be high enough that only the most stable, perfectly paired double-stranded DNA structures can form. Usually, the perfect annealing temperature is only a few degrees lower than the melting temperature of the primer pair. Moreover, during the annealing step, the polymerase will bind to the primer/template DNA complex, although it will not start reading until the temperature is raised in the next step. Following annealing, elongation/extension is done and repeated 35 times. The reaction is rapidly heated to approximately 72 degrees, and this is when the polymerase begins reading in the 5′-3′ direction and copying the template DNA in the 3′-5′ direction. The higher temperature during this step reduces non-specific primer/template DNA interactions, which aids in increasing the specificity of the reaction. After steps 2-4 are repeated 35 times, the final elongation is done. The reaction is held at 72 degrees for several minutes to allow polymerases to finish reading whatever strand they are currently on. This step, although optional, can help reduce the number of truncated copies in the final product. The last step is the final hold. This is necessary because, as PCR can often take a few hours, they are often done overnight or when the researcher has stepped away. In this step, the PCR product is held at 4 degrees until the tube is taken out.

Wednesday

On Wednesday, the work started immediately at 8:30 AM. In the morning, we created a 1% agarose gel (a procedure I had already done in previous lab experience) and verified that the overnight PCR had worked. To do this, we ran a molecular ladder as well as the PCR sample through the gel and analyzed the band. Once we were sure the PCR had succeeded (as the band was present), we discarded the gel and moved on to the next steps.

Since we had received some fresh PVDF paper, we switched gears and ran an SDS-PAGE gel and Western blot with the same samples from last week–these are the ones that did not work on the Western blot due to the different type of paper used. Once this Western blot was set up for the 9337 samples, we did the regular washes with 10% milk and primary antibody, concluding this process at the end of Wednesday by putting it on the shaker in the environmental cold room.

Next, we did blunt-end ligation cloning. The blunt-end ligation is a non-directional cloning method, meaning the insert can ligate to the vector in two possible directions. For protein expression, the insert must be in one particular direction, and usually, the desired directional clone can be identified using PCR. In blunt-end cloning, both the vector and the insert contain blunt-ends. During the transient association of the ends of the vector and the insert, the DNA ligase enzyme seals the gaps. The T4 DNA ligase utilizes ATP to make a phosophodiester bond between the 3′ hydroxyl group of one DNA strand and the 5′ phosphate group of another DNA strand.

Some of the advantages of blunt-end cloning are:

• simple design of the primers without any extra bases at the 5′ end of the primer
• does not require complementary sequence between insert and vector, so it is a universal cloning method
• versatile

Some of the limitations of blunt-end cloning are:

• non-directional cloning generates only 50% of inserts with the proper orientation
• both inserts and vectors do not have complementary 3′ or 5′ overhangs, resulting in little chance for association stability between insert and vector, leading to lower recombination efficiency when compared to sticky-end cloning
• prone to vector self-ligation
• can have more than one insert
• generating a clone with multiple inserts with the right orientation is low

When we did blunt-end cloning, we followed the protocol below:

We began by preparing the insert, which is any desired fragment of DNA to be cloned into a vector or plasmid. This was what the PCR step was for. Following this, we also prepared our primers (the same ones we used for the PCR). Then, we did phosphatase reaction of the vector prior to ligation. The PCR amplified fragments and restriction-digested fragments contain a 5′ terminal phosphate, which creates a significant problem in blunt-end ligation. Self-ligation increases the background with just the vector without the insert in it. However, this issue can be minimized by reducing the self-ligation by removing phosphates present at the 5′ terminus in the vector. Thus, phosphatase treatment will effectively reduce the background of empty clones by > 95%. Next, we did the ligation reaction, which is a simple process that was run overnight in the thermo-cycler.

Thursday

When blunt-end ligation was complete, we prepared competent cells for heat shock transformation, a fairly standard lab procedure. Clark got in a bit later today due to a doctor’s appointment, so I spent the start of the day washing and scanning my Western blot from yesterday with secondary antibody and TBST.

When Clark got here, he had already prepared competent cells of E. coli, specifically the DH5-Alpha strain. I have worked with this strain in previous years’ research, which proved helpful. Following this, we ran a standard heat-shock protocol in a 42 degrees water bath for 60 seconds. Following this, we put the cells in recovery. Finally, we streaked plates with the recovered cells and checked them the next day to see if the reaction had worked.

Friday

Our plates had a ton of colonies on them! I successfully edited a bacterial genome!

Today was a fairly simple day. I just cast 6 10% polyacrylamide gels using standard procedures I have described in previous blog posts.

Other than that, I worked on my poster and presented my slides from forever ago on a zoom meeting.

This marked the end of week 9!! I’m super excited for the symposium next week!

This was the first week back after the storm! Since the ultracentrifugation and gradient method failed to produce significant amounts of protein, we have recently been trying a newer method of protein purification: columns and FPLC.

Monday

Today, I ran gravity columns. The previously centrifuged pellet was in the -80 C freezer. I then added our lysis buffer, 8 M urea with BME and PMSF, and sonicated it to resuspend. Once the pellet was resuspended with no floating particles (completely homogeneous), I centrifuged it on the floor model centrifuge to attain the clarified lysate. After spinning it for 45 minutes, I poured the supernatant into a different tube and resuspended the pellet in 1 mL of lysis buffer (without BME and PMSF) for sampling. I took samples of both the pellet and supernatant (60 microliters of sample and 20 microliters of 4X loading dye for the SDS-PAGE gel to be run later). Once samples were taken, I set up the gravity column and pipetted 4 mL of beads into the column. I equilibrated it with water and our 8 M urea buffer–this is a very time-consuming process involving watching the liquid drip into a separate container and pass through the nickel beads. When the nickel beads were equilibrated, I added the supernatant/clarified lysate acquired from the previous centrifugation and stored it in a fridge.

A brief explanation on gravity columns: Column chromatography functions on the principle that solutes of the solution will be adsorbed with the help of a stationary phase and later separated into discrete components. Ion- exchange chromatography is based on electrostatic interactions between charged protein groups, and solid support material (matrix). Matrix has an ion load opposite to that of the protein to be separated, and the affinity of the protein to the column is achieved with ionic ties. We spent today washing and planned to elute tomorrow (Tuesday).

Tuesday

Today we eluted the proteins from the supernatant. I started off by collecting flow-through, which was done by opening the valve of the gravity column and allowing the supernatant from yesterday to pass through the nickel beads. The principle is that, upon allowing the supernatant to flow over the beads, the protein with a His-tag will bind to the nickel beads in the column. After this, varying concentrations of imidazole are poured over the beads and collected in various tubes. Clark mentioned that, historically, he has seen success with 250 mM imidazole. Before doing this step, I first had to make solutions with the concentrations of imidazole we would be testing: 30 mM, 250 mM, 500 mM, and 1 M. This was done using 1 M imidazole and 8 M urea (the same buffer we used for lysis). The amounts of each we would use were calculated based on the equation M1V1 = M2V2. We needed 50 mL of 30 mM, 10 mL of 250 mM, 10 mL of 500 mM, and 10 mL of 1 M. Once these solutions of imidazole and urea were made, they were stored in 50 mL conical tubes on ice. The 30 mM was allowed to flow through the beads, but we weren’t expecting much protein to flow with the 30 mM into the elution tube. Following this, the same steps were repeated for 250 mM, 500 mM, and 1 M. This was a very time-taking process as you need to allow the liquid to slowly flow through the small opening on the bottom of the column.

The columns look similar to the image above. We collected samples of the flow-through, 30 mM, 250 mM, 500 mM, 1 M, and beads using the same 60 microliter and 20 microliter ratio I described for Monday. These samples were placed in the freezer, marking the conclusion of today.

Wednesday

Today, we ran SDS-PAGE gels (3 total) and a western blot. My day started by running 2 SDS-PAGE gels (Clark ran the third one). To do this, I removed the samples from the freezer and placed them into the heating block at 98 C. I briefly ventilated the tubes after 2 minutes and allowed them to thaw for 8 more minutes. I also removed the molecular ladder/marker from the freezer and allowed to thaw on the bench. Following this, I removed the gels from the fridge and set up the electrophoresis apparatus. I then loaded the samples into the gel (the two gels contained the same samples, one was meant to transfer onto a western blot, thus we duplicated it). Once the samples, marker, and loading dye were loaded into the gel, I set it to run at 100 V for 15 minutes until the samples hit the separating gel. Next, I set it to 180 V for 45 minutes until the samples run off the gel. When the timer was up, I removed one gel from the apparatus, separated the glass to extract the gel, and stained and de-stained it. It was stained by pouring staining buffer into an empty pipette tip box (repurposed for staining) and microwaving it with the gel for 1 minute. This was placed in the fume hood to vent and later de-stained with clear, de-staining buffer on the rocker. The other gel was used for a western blot (unsuccessful because we did not have the right kind of blot paper). The standard western blot procedure was followed: a “gel-blot sandwich” was created, surrounded by pads and sponges. This was put into the western blot apparatus with transfer buffer and then left to transfer in the fridge at 100 V for 1 hour. During this time there was a fire drill. After the fire drill, when we returned to labs, we found that the contents of the gel did not transfer onto the blot as we did not see the marker on the blot. However, we know there were volts running through the system, so we are hypothesizing that the reason this did not work is because we ran out of the right kind of PVDF paper (so we used alternate PVDF paper). Our gel, which was still on the rocker, was very clear, which is a positive result– it means we removed most of the irrelevant noise surrounding the protein and have isolated the desired protein to a certain extent. This marked the end of Wednesday.

Thursday

Since we do not have PVDF paper, Clark thought it would be interesting if we did a protein refolding experiment. This was just to learn how to run dialysis rather than to get results, as we do not know how successful our purification was due to the lack of PVDF paper. Thus, this was meant to be a learning experience. Today began by creating two different dialysis buffers (as we wanted to test two). Clark made one (with MOPS) that is light sensitive and thus had to be done covered by aluminum foil to maintain a dark environment. I made the other one with Tris, a commonly used compound in buffers. I added NaCl, Tris, 10% glycerol, and water to a 100 mL beaker and used the magnetic stir plate to mix the solution. Then, I filtered it using a vacuum filtering apparatus and stored it in a clean 1 L bottle in the fridge until use. Following this, we took a break/ate lunch and then began the dialysis experiment, which would happen overnight. We took two 100 mL beakers and filled them with our buffers. Then, we let a dialysis membrane/bag soak in the buffer for 5 minutes to soften. Once it had softened, we filled it with 5 mL of the purified protein from Tuesday and clamped both sides of it closed to make it turgent. Following this, we let it sit in the environmental cold room overnight, stirring on a magnetic stir-plate.

Friday

Today was day 2, the final day, of the protein refolding experiment! We were only able to run FPLC for one of the samples (the one in MOPS buffer). We did this by calibrating and equilibrating the FPLC machine. Then, we filled a carousel as per usual and ran the machine (a 2 hour long process). We did not find significant peaks, meaning that there was minimal protein present, which was expected as we noticed most of our protein precipitated (fell out of solution) sitting in the environmental cold room overnight. This was not the only part of today, however. I also had the opportunity to speak with someone who is outside the field of structural biology: they are in synthetic biology. Their name is Malyn and we got coffee altogether with Clark and discussed their project regarding microbes in soil and horizontal gene transfer. I found it exciting to explore various avenues in synthetic biology as well. It’s just a reminder that there are endless directions you can take biology! This marked the end of today.

This week, I did many new things, especially a protein refolding experiment: something I had been wanting to do for a while. We will be running the second dialysis sample through FPLC on Monday; the sample will not be ruined by Monday.

This week was off to a rough start with Hurricane Beryl flooding much of our neighborhood and leading to power outages throughout the Woodlands area. We had power and so we accommodated several affected family friends for the week, which is the reason this blog post is delayed. On Wednesday, I was unable to conduct any lab work because my mentor’s car was flooded in the storm, preventing him from coming in. Consequently, my efforts on Thursday and Friday were redirected towards an in-depth review of the relevant literature and research papers related to my field, allowing me to deepen my understanding of the topic. This blog post will focus on the learnings attained from the various papers I read.

Wednesday

Citation for Paper 1: Marzonie, M., Nitschke, M. R., Bay, L., Bourne, D. G., Harrison, H. B. (2024). Symbiodiniaceae diversity varies by host and environment across thermally distinct reefs. Molecular Ecology.

Information from Paper 1: The study focuses on the diversity of Symbiodiniaceae, a group of endosymbiotic dinoflagellates, which play a vital role in determining the thermal tolerance of corals across different regions and environments. In my project for this REU, we are studying how dinoRNAVs, an RNA virus, impact Symbiodiniaceae and potentially influence the occurrence of coral bleaching. While the individual effects of host genetics, environmental factors, and thermal disturbances on symbiont communities are well-documented, their combined effects remain less understood, prompting the need for the research in the paper. This research was conducted across a 1300 km stretch in Australia’s Coral Sea Marine Park, aiming to identify the interactive effects of host genetics, environment, and thermal disturbances on Symbiodiniaceae communities. The study involved identifying Symbiodiniaceae to the species level in three coral species: Acropora cf humilis, Pocillopora verrucosa, and Pocillopora meandrina, using genetic markers ITS2 and psbAncr for the symbionts and DArT-seq for the coral hosts. The findings revealed that the genus Cladocopium was predominant among the samples, with specific affiliations: Acropora cf humilis with C3k, Pocillopora verrucosa with C. pacificum, and Pocillopora meandrina with C. latusorum. Multivariate analyses showed that the symbiont communities in Acropora were strongly influenced by local environmental conditions and thermal disturbances, whereas in Pocillopora species, host genetic structure played a more significant role. Among the Pocillopora species, the combined effects of environment and host genetics explained more variation in symbiont communities for P. meandrina compared to P. verrucosa. The study also observed that the bleaching event in 2020 had varying impacts on symbiont communities, aligning with patterns seen in P. verrucosa and A. cf humilis, but not in P. meandrina.

Citation for Paper 2: Benites, F. L., Stephens, T. G., Bhattacharya, D. (2022). Multiple waves of viral invasions in Symbiodiniaceae algal genomes. Virus Evolution, 8(2).

Information from Paper 2: Dinoflagellates from the family Symbiodiniaceae are marine organisms that use sunlight to produce energy and form symbiotic relationships with various hosts, including corals. They havelarge and unique genomes characterized by frequent gene duplications, the expansion of gene families with unknown functions, and significant retroposition events, which are processes where genetic elements are copied and inserted into new locations within the genome. Despite the extensive study of their genomes, the role and extent of horizontal gene transfer (HGT) in the evolution of Symbiodiniaceae are not well understood. In some related species, such as Fugacium kawagutii and Brevioloum minutum, a small percentage of their genes are believed to have originated from prokaryotes through HGT events. Higher levels of HGT have been observed in other dinoflagellates, often associated with significant genomic changes, such as the acquisition of viral nucleoproteins that may play a role in chromatin regulation. All sequenced dinoflagellate genomes contain a family of viral-acquired nucleoproteins known as dinoflagellate-viral nucleoproteins (DVNPs), which are thought to have originated from a large DNA algal virus and may help regulate chromosome structure. The acquisition of the DVNP gene family is linked to massive genomic expansion in dinoflagellates and may have provided immunity against infections by similar viruses. Symbiodiniaceae isolated from corals are known to be actively infected by various viral groups, including giant DNA viruses and single-stranded RNA viruses, which may have implications for coral health (not well understood at the moment). There is evidence suggesting that some viral infections in Symbiodiniaceae are latent, meaning the viruses can persist in the host without causing immediate symptoms, and may become active under stress conditions. My work at Rice focuses on determining the impact of dinoRNAVs, single-stranded RNA viruses, on coral colony health.

Thursday

Citation for Paper 3: Chen, B., Wei, Y., Liang, Y., Yu, X., Liao, Z., Qin, Z., Xu, L., Bao, Z. (2024). The microbiome dynamics and interaction of endosymbiotic Symbiodiniaceae and fungi are associated with thermal bleaching susceptibility of coral holobionts. Applied and Environmental Microbiology.

Information from Paper 3: The study focuses on understanding how the interactions between Symbiodiniaceae and fungi within the coral microbiome affect the coral’s susceptibility to thermal bleaching. The research found that heat-tolerant sub-clades of Symbiodiniaceae, specifically the C3u sub-clade and Durusdinium, dominate the coral communities in high-risk thermal bleaching areas. These heat-tolerant Symbiodiniaceae are believed to contribute to the coral’s ability to withstand higher temperatures, potentially reducing bleaching events. Unlike Symbiodiniaceae, the fungal community associated with corals did not have core amplicon sequence variants, indicating high variability among different coral species. The study found a significant positive correlation between fungal richness, the abundance of animal-plant pathogens, and the percentage of coral thermal bleaching, suggesting that higher fungal diversity and pathogen presence increase bleaching susceptibility. Each coral species exhibited a complex Symbiodiniaceae-fungi interaction network (SFIN), driven by the dominant Symbiodiniaceae sub-clades. Corals with low thermal bleaching susceptibility had SFINs characterized by low complexity and high betweenness centrality, indicating a more resilient and less parasitized microbial network. The study highlights that the extra heat tolerance observed in corals from Huangyan Island may be linked to the high abundance of heat-tolerant Symbiodiniaceae and a resilient microbial network. The findings suggest that managing fungal diversity and pathogen abundance could be crucial in mitigating coral bleaching and promoting coral health under thermal stress.

Citation for Paper 4: Springer, K., Kunzmann, A. (2023). Symbiodiniaceae in and ex hospite have differential physiological responses under different heat stress scenarios. Marine Biology Research, 1-13.

Information from Paper 4: The study focuses on the increasing frequency of cnidarian bleaching, which is the breakdown of the symbiosis between cnidarian hosts and their endosymbiotic dinoflagellates, often due to extreme seawater temperatures linked to various human activities, with seawater warming being a significant factor. The primary aim of the research is to understand whether the thermal tolerance of dinoflagellate symbionts differs when they are inside their host (in hospite) compared to when they are free-living (ex hospite). The study measured the maximum quantum yield of photosystem II (Fv/Fm) and symbiont cell density in seven different cnidarian species and five cultures of isolated endosymbionts. These were subjected to three different temperature conditions: 26°C (control), 30°C, and 34°C for a duration of 21 days. Isolated dinoflagellate cells showed a significant susceptibility to elevated temperatures of 30°C, which was evident from a decrease in their photochemical efficiency and cell density. Additionally, there was a progressive disintegration of cellular structures and loss of pigmentation in all but two cultures during the first week of exposure. The study found that bleaching of coral holobionts at 30°C could be explained by a reduced density of algae cells in the host tissue, with this effect being particularly noticeable in soft corals. Exposure to 34°C resulted in drastic bleaching of stony coral species, anemones, and jellyfish, and even led to the death of soft corals. This indicates that higher temperatures have severe impacts on the symbiotic relationship and the survival of these organisms.

Friday

Citation for Paper 5: Mashini, A. (2022). The Impact of Symbiont Diversity on Cellular Integration and Function in the Cnidarian-Dinoflagellate Symbiosis. Open Access Te Herenga Waka-Victoria University of Wellington. Thesis. https://doi.org/10.26686/wgtn.18739247.

Information from Paper 5: Coral health is closely linked to their symbiotic relationship with phototrophic dinoflagellates from the family Symbiodiniaceae. This symbiotic relationship is crucial for coral survival, but it is threatened by elevated temperatures, which can disrupt the symbiosis and lead to coral bleaching, a phenomenon that is becoming more frequent and severe due to climate change. The primary aim of the research was to understand the cellular processes involved in hosting native versus non-native symbionts in the model symbiotic cnidarian Exaiptasia pallida, commonly known as Aiptasia. The study sought to assess the potential for establishing novel symbiotic partnerships by using a multidisciplinary approach, including proteomics and quantitative immunocytochemistry, with a focus on inter-partner nutritional exchange. A new method for characterizing the proteome of Symbiodiniaceae was developed and used to compare the molecular and metabolic pathways underlying successful symbiosis. The proteome of Breviolum minutum, the native symbiont of Aiptasia, was analyzed under different nutritional conditions, revealing distinct proteomes related to immunosuppression, metabolic integration, and oxidative stress in the symbiotic state. The study further characterized the proteome of B. minutum during host colonization and compared it to the proteome of a non-native symbiont, Durusdinium trenchii, which is thermally tolerant but opportunistic. The proteome of D. trenchii showed a lower abundance of photosynthetic proteins and an upregulation of parasite-like immunosuppression mechanisms, indicating a lesser degree of integration with the host compared to B. minutum. Specific immunofluorescent antibodies were designed to localize and quantify host nutrient transporters in Aiptasia colonized with either native or non-native symbionts. The study found different transporter localization patterns in hosts harboring non-native symbionts, suggesting disrupted nutritional flux and a lesser degree of host-symbiont integration. The findings suggest that the cellular integration necessary for a functional symbiosis with efficient nutritional exchange is not easily replicated with non-native symbionts, which may reduce the likelihood of corals adapting to climate change by changing their symbiont population.

This week primarily involved reading papers and familiarizing myself with literature. Next week will focus more on actual lab work.

Monday – Reading Papers

In Monday, no lab activities were done as Clark was feeling unwell and thus did not come into the lab. I spent the day in the office reading papers about the virus we are trying to identify the structure and function of as well as the most similar known virus. I also read into protein affinity tags, nickel column affinity purification, and protein denaturation with urea. This information will be detailed in the poster presentation as well as the PowerPoint for the July 9 presentation.

Tuesday – Cell Lysis and Lysate Preparation (Gravity Columns)

The preparation began with setting up an ice bucket and creating an ice-water mixture to maintain the temperature of the samples throughout the procedure. The sonication buffer, containing 8 M urea, was taken from the 4°C fridge in the FPLC room to ensure it remained cold. 30 mL of the sonication buffer was poured into a 50 mL conical tube, to which 10.5 µL of β-mercaptoethanol (BME) and 300 µL of phenylmethylsulfonyl fluoride (PMSF) were added. Vortexing the solution was done to ensure the PMSF dissolved properly and the BME was evenly distributed to prevent protein degradation during sonication.

The sonication buffer was then poured over the cell pellet until it was fully immersed. The sonicator probe was cleaned using DI water and 70% ethanol, and then wiped with kimwipes. This step was essential to prevent contamination and ensure accurate sonication. The probe was then placed into the buffer, positioned right above or near the cell pellet to ensure sufficient cell disruption.

Sonication was performed with the sonicator set to 7 minutes at 70% amplitude, with a pulse on/off cycle of 01/03. This setting was chosen to break up the cell pellet without overheating the sample. The goal was to disrupt the cells thoroughly, which was monitored by checking for any remaining cell chunks. If clumps persisted, the probe was moved closer to those chunks for better disruption.

During sonication, the blue-capped rotor was retrieved from the fridge and placed in the floor centrifuge to pre-cool it to 4°C. This pre-cooling step was done to maintain the integrity of the lysate during high-speed centrifugation. Once the sonication was complete and the cell pellet fully broken down, the lysate was poured into a compatible centrifuge tube, double-checked to ensure that no floating cell chunks remained.

The lysate was balanced with an identical tube containing DI water to ensure the centrifuge operated safely and effectively. This balance was checked carefully due to the high-speed spin of 30,000 x g for 45 minutes at 4°C, which could potentially damage the centrifuge or pose safety risks if the tubes were not properly balanced.

Post-spin, the clarified lysate was carefully decanted into a pre-cooled 50 mL conical tube, ensuring any remaining cell debris was left behind. This clarified lysate was kept on ice to prevent protein degradation. A small volume of sonication buffer, minus the BME and PMSF, was added to the remaining pellet, resuspending it until the solution turned cloudy. This resuspension helped to prepare the pellet for further analysis.

Following the centrifugation step, the clarified lysate was applied to a gravity column pre-packed with nickel beads, which are specifically designed for affinity purification of His-tagged proteins. The column had been pre-equilibrated with a binding buffer containing 8 M urea to ensure the conditions were optimal for binding the His-tagged proteins. The lysate was allowed to flow through the column by gravity, ensuring that His-tagged proteins had ample time to interact with the nickel ions on the beads.

After the lysate had passed through the column, the next step was washing the column to remove any nonspecifically bound proteins and other contaminants. This was done using the same binding buffer but with a low concentration of imidazole (typically around 20-50 mM). The imidazole in the washing buffer helps to displace weakly bound contaminants without eluting the target His-tagged proteins. Several column volumes of the washing buffer were passed through the column to ensure thorough cleaning. Each wash step was collected and monitored to check for the presence of proteins, ensuring that most contaminants were removed. It was essential to perform multiple washes to achieve a high purity of the bound His-tagged proteins while keeping them securely bound to the nickel beads.

Once the washing steps were completed, the column needed to be stored properly until the elution step was carried out the next day. The column was sealed securely to prevent any evaporation or contamination and stored at 4°C.

Wednesday – Elution & SDS-PAGE gels

The elution process began with the preparation of several elution buffers containing varying concentrations of imidazole: 30 mM, 250 mM, 500 mM, and 1 M. Each buffer also included 8 M urea to maintain protein solubility and a buffering agent to ensure the pH remained stable. These buffers were needed for selectively eluting the bound proteins from the nickel beads based on their affinity for the imidazole.

The gravity column was allowed to equilibrate at room temperature for a few minutes before starting the elution process. Sequentially, each elution buffer was added to the column. The 30 mM imidazole buffer was used first, followed by the 250 mM, 500 mM, and finally the 1 M imidazole buffer. For each concentration, a 10 mL was used to ensure thorough elution. The eluate was collected in separate microcentrifuge tubes, labeled according to the imidazole concentration used. Elution allows for the separation of proteins based on their binding strength to the nickel beads.

After completing the elution steps, a final wash with 1 M imidazole was performed to ensure any remaining loosely bound proteins were removed. This fraction was collected in a separate tube. To confirm that no protein remained bound to the nickel beads, the beads were transferred from the column to a microcentrifuge tube and washed with DI water. The wash fraction was collected for analysis, ensuring that the entire protein sample was accounted for.

Following elution, we ran an SDS-PAGE gel. The purpose of the SDS-PAGE gel was to visualize the different protein fractions and confirm the presence of the target capsid protein 9337 in the 250 mM imidazole elution fraction. Each elution fraction, including the pellet, supernatant, flow-through, and the final wash from the beads, was mixed with protein loading dye. This dye contained SDS, which denatures the proteins and gives them a uniform negative charge, ensuring they migrate based on size during electrophoresis. The samples were heated briefly to ensure complete denaturation.

The SDS-PAGE gel was set up in the electrophoresis apparatus, and the wells were loaded as follows: two wells with loading dye as controls, one with the protein marker, and others with the pellet, supernatant, flow-through, each elution fraction (30 mM, 250 mM, 500 mM, 1 M imidazole), the final wash from the beads, and a second protein marker in the final well.

Electrophoresis was conducted by applying an electric current, causing the proteins to migrate through the gel matrix. The protein marker allowed for the determination of the molecular weights of the separated proteins. The specific band corresponding to the target capsid protein 9337 was expected in the 250 mM imidazole elution fraction, indicating successful purification.

After electrophoresis, the gel was stained to visualize the proteins: the dye solution binds to the proteins, aiding with visualization. This was followed by the de-staining step, done to remove excess dye. The resulting bands on the gel represented the proteins in each sample, which allowed for the assessment of purity and identification of the target protein, which should have been 33 kD but showed as 40 on our gel (something we haven’t understood yet, but will research more about)

That marked the end of this week. Happy (early) 4th of July everyone!! I hope we all have a fun, relaxing weekend.

This week I worked entirely independently. We realized last week that ultracentrifugation was not working for the capsid protein we were trying to purify, so we shifted from that to FPLC, which I will also detail.

Monday – Reading Papers

In Monday, no lab activities were done as Clark was feeling unwell and thus did not come into the lab. I spent the day in the office reading papers about the virus we are trying to identify the structure and function of as well as the most similar known virus. This information will be detailed in the poster presentation as well as the PowerPoint for the July 9 presentation.

Tuesday – FPLC & Nickel Columns

Rather than detailing step-by-step what I did (like my usual blog posts), I decided I wanted to describe how it works, why it is done, and what kind of results we can obtain from the procedure.

We realized that ultracentrifugation was unsuccessful through our Western blot results, which were consistently unclear following the first one and consistently inconclusive. Thus, we realized that ultracentrifugation was not a viable method for capsid protein purification. From here, we moved on to FPLC to see how pure we can get the protein from this method.

Fast Protein Liquid Chromatography (FPLC) systems are essential for separating and purifying proteins and other biomolecules. These systems operate at pressures around 3,500 psi (24 MPa) and usually include a pump, a UV detector, a conductivity meter, and a fraction collector. The pump ensures a controlled flow of liquid, while the UV detector identifies protein peaks by measuring absorbance, typically at 280 nm. The conductivity meter tracks the buffer’s ionic strength, indicating different species in the elution, and the fraction collector gathers the separated samples for further use or analysis.

Modern FPLC systems, like the NGC™ medium-pressure chromatography system, have extra features such as column switching valves, sample pumps, and buffer blending valves. Column switching valves allow for easy switching between columns, sample pumps enable automatic sample application, and buffer blending valves create precise buffer gradients necessary for protein elution. The NGC system is managed through the ChromLab™ software, which makes it easy to set up and monitor the chromatography process digitally.

In a typical FPLC system, the flow path involves loading the sample either manually by injection into a sample loop or automatically using a sample pump. Advanced systems may also include multi-wavelength detectors, which are particularly useful for proteins tagged with chromogenic or fluorescent labels. These detectors monitor sample elution at various wavelengths, helping to separate the protein of interest from contaminants based on their different absorption properties.

Wednesday – Gravity Columns

Today we ran gravity columns. I will once again explain how these work rather than the exact procedure we ran.

Gravity columns are a simpler and more traditional method for separating and purifying proteins and other biomolecules compared to Fast Protein Liquid Chromatography (FPLC) systems. Operating based on gravity flow, these columns allow liquid to move through the column under the force of gravity rather than pressure. A typical setup includes a glass or plastic column filled with a stationary phase, usually a type of resin or gel. The sample is applied at the top of the column and flows downward due to gravity, with different components separating based on their interactions with the resin or gel.

The main components of a gravity column system are the column itself, the stationary phase, sample application, elution buffer, and a fraction collector. The column is filled with the stationary phase that facilitates the separation process. The sample is manually loaded at the top, and the elution buffer helps move the sample through the stationary phase. As the sample travels through the column, different components separate based on their size, charge, or other properties. These separated fractions are then collected at the bottom for further analysis or use.

The process of using a gravity column is straightforward. The sample is loaded at the top, and the elution buffer is added to help move it through the stationary phase. Different components of the sample travel at different speeds, leading to their separation. While gravity columns are less sophisticated than modern FPLC systems, they are still widely used due to their simplicity and cost-effectiveness. They are particularly useful for preliminary purification steps or in situations where high pressure and advanced detection are not necessary. For more detailed information on gravity columns, you can visit related chromatography resources.

Thursday – SDS-PAGE and Western blot

I started the day by putting the samples on the heat block and heated it for 2 minutes. Then, I vented the tubes by rapidly opening and closing and let it heat for 8 minutes for a total of 10 minutes on the heat block. Next, I removed the samples from the heat block and centrifuged them to remove the condensation from inside the tube. Then, I got the gel out of the fridge, removed the well comb, and set up the well cassette using a buffer dam since I was only running one gel. Following this, I placed the gel cassette into the gel running box, then poured running buffer into the cassette until it was about to overflow. Next, I loaded the gel with 5 microliters of marker, pellet, supernatant, iodixanol supernatant, viral pellet, iodixanol gradient band 1, and iodixanol gradient band 2. Then, I ran the gel at 100V for 15 minutes until the dye fronts had tightened on top of the separating gel. When this happened, I increased the voltage to 180V and ran the gel until the dye ran off the front of the gel.

Next, I stained the SDS-PAGE gel with Coomassie blue. I started by grabbing an empty pipette tip box and separating the front plate and back plate using a wedge gel-cracker. I placed the gel cracker into one of the bottom corners and gently pushed the back end of the gel-cracker down to open the gel. I then cut the stacking gel off and threw it away. To remove the gel from the glass, I gently slid the wedge under one of the corners of the front plate and continued pushing up slowly until the gel dislodged fully from one edge. Once this was done, I placed the gel into the empty pipette tip box. Note: Cut the gel into half and use the other half for the Western blot I will describe in the next section. I then poured the staining buffer onto the gel until it was fully covered with a few millimeters of buffer and microwaved the gel and buffer for one minute with the box unclipped. Once the gel was microwaved, I moved the box to the fume hood using heat resistant gloves, then I opened it and let it vent. Then, in the fume hood, I poured the staining buffer into the used staining solution bottle using a funnel and gently rinsed the gel off with DI water. Finally, I added destaining buffer and two kimwipes to the box and left it on the rocker until the gel is destained. I left it overnight. Finally, I put the gel into a plastic sleeve, labeled it, and scanned it.

While this was happening, I was also running a Western blot. I started this process by pouring the Western blot transfer buffer into the transfer box until it was about half full. I then grabbed some filter papers and soaked them in the box with the transfer buffer. While the filter paper was soaking, I grabbed a square plate, brought it to the gel running bench, and filled the bottom with methanol. Next, I cut a blot-sized portion of PVDF using scissors and got the blue protection paper off the blot using special forceps. I then put the blot into the methanol to activate it. To make the transfer sandwich, I placed the sponge, soaked filter paper, gel (backwards orientation), activated PVDF paper, soaked filter paper, and sponge on top of each other in that order. I then clipped it shut and placed it in the Western blot transfer box. Next, I placed a blue cooling block into the front part of the running box, put the box in the fridge, then filled it up with transfer buffer to the “blotting line” on the front of the transfer box. Next, I put the lid on the transfer box, then ran it at 90V for 1 hour. While the transfer was running, I prepared 10 mL of 10% milk in TBST. After the transfer, I removed the membrane from the sandwich and put it into a square plate with all of the 10% milk. I allowed the blot to block for 1 hour and subsequently prepared 10 mL of 5% milk with 1 microliter of antibody (6x Hist) and left it on the rocker for about a minute to mix the antibody into the solution. While the milk and primary antibody was on the rocker, I poured the 10% milk off of the blot and rinsed it with TBST until the TBST ran clear. Next, I poured the primary antibody and 5% milk solution onto the blot and incubated at room temperature for 1 hour. Once this was done, I washed the blot 3 times for 10 minutes each with TBST and made 10 mL of 5% milk in TBST with 2 microliters of secondary antibody while this was blocking. I let the antibody mixture sit on the rocker for about 1 minute to allow it to mix into the milk solution. I poured the last TBST wash off the blot then added the secondary antibody and incubated at room temperature on the rocker for 1 hour. Once this was done, I began with the next set of 3 TBST washes, once again for 10 minutes each. Next, I poured the TBST off the blot into the sink, rinsed with some Milli-Q water, then poured developing solution over the blot until the blot was covered. This blot yielded significant results which we will be analyzed when Clark returns on Monday. Finally, I clipped the developed blot with forceps and put it in the 37 degrees incubator to dry. Finally, I placed it into a plastic sleeve, labeled it, and scanned it.

Friday – Cell Pelleting for IPTG Induction

Today, I pelleted the cells, the final (Day 3) procedure in the IPTG induction process. The protocol is as follows.

To begin the process, I poured the cultures into 500 mL centrifuge bottles, balanced them, and spun them at 4°C with a relative centrifugal force (RCF) of 4,000 for 10 minutes. While the centrifugation was ongoing, I labeled 50 mL tubes in preparation for harvesting the pellets.

After centrifugation, I poured the media off the pellets back into the original flasks and placed the bottles upside down on a stack of paper towels to remove any excess LB. Using the green spatula near the main lab sink, I scooped out the cell pellets. Before using the spatula, I rinsed it with a little water and dried it with a paper towel.

Next, I transferred the cell pellets into the labeled 50 mL tubes. To ensure the cells were properly pelleted at the bottom, I spun the tubes again at 4°C with an RCF of 4,000 for 3 minutes. After centrifugation, I froze the tubes in a labeled Ziploc bag, completing the preparation for future purification steps.

This week was an exciting week as I transitioned from working mostly independently to entirely independently. It involved a few mistakes along the way, but I have learned a great deal from those mistakes. Also, the tour on Friday 6/21/2024 was really exciting and fascinating as it opened my eyes to how the water we use gets filtered, which I’ll make sure to detail in the section for this day!

Monday – Iodixanol Gradient Viral Pellet Purification

Monday was a very productive day as I was on my feet the entire time barring a short lunch break! I find myself really engaged in the work I am doing here and I feel strongly towards the cause the project was inspired by. I started the day by sonicating the pellet and subsequently clarifying the lysate. To do this, I made lysis buffer by mixing (carefully as some of these chemicals are toxic) 30 mL of sonication buffer with 10.5 microliters of BME and 300 microliters of PMSF. One of the challenges for me during my time at the lab has been something that is probably quite simple and straightforward for most people: converting from microliters to mL and vice versa as well as finding the right pipette to use for each job. However, with practice, this has improved! After dissolving the chemicals needed for lysis buffer together, I prepared the sonicator. This was done by spraying the probe of the sonicator with DI water, 70% ethanol, and then DI water again, wiping in between each step wtih kimwipes, special wipes designed to leave no residue. I then put the sonicator probe into the sonication buffer that was sitting on the cell pellet Clark pelleted over the weekend. Since it was frozen rather than fresh, the sonication process was not as long. The settings of the sonicator were 7 minutes, 70% amplitude, pulse on 01, and pulse off 03. While the sonicator was running, I prepared our lab’s floor model centrifuge by grabbing the right rotor out of the fridge, putting it into the floor centrifuge, closing the lid, and turning it on; this was done to pre-cool the centrifuge to ensure that it is ready to go when the lysing is done. When the cell pellet was completely broken up and there were no traces of floating clusters of cells, I poured the lysate into a centrifuge tube that was compatible with the floor model centrifuge and balanced it with DI water in another tube, carefully ensuring they were precisely balanced to prevent any risk of injury due to unbalanced centrifuge runs. I then spun the tubes at 30,000 x g for 45 minutes at 4 degrees. Immediately after the spin was completed, I removed the clarified lysate and immediately decanted the supernatant into the supernatant tube, leaving it on ice. Then, I pipetted 2 mL of sonication buffer onto the pellet without BME or PMSF, resuspended it, and took samples of both the pellet and the supernatant for SDS-PAGE and Western blot.

Once this process was done, I prepared iodixanol solutions of 10%, 20%, 30%, 40%, and 50% using 60% iodixanol, sonication buffer, and the simple dilution formula–CiVi = CfVf. Then, I pipetted 1 mL of 10% iodixanol into 4 polystyrene tubes that are compatible with the ultracentrifuge. I then slowly pipetted all of the clarified lysate into each of the 4 tubes, 1 mL at a time, ensuring that the iodixanol cushion did not mix with the clarified lysate. Once the supernatant was pipetted, I added sonication buffer with BME to ensure that each tube was filled until the top–as Clark says: “There should be enough space in the tube that you can put the end of your pinky in.” He then spun these tubes in the Sw41-Ti rotor tube holders–I could not do this part as I do not have the necessary training to operate the ultracentrifuge. The tubes were spun at 37,500 RPM in the rotor for 2 hours and 4 degrees. Fun fact: Clark found out 2 hours was sufficient (compared to the 22 hours he used to run it for before) accidentally because he forgot to start the ultracentrifuge once. During the spin, I set up the iodixanol gradient by slowly pipetting 50%, 40%, 30%, 20%, and 10% iodixanol on top of each other in a polystyrene ultracentrifuge tube, ensuring that they do not mix and form clear lines in between each concentration. Then, I set up another tube with the same materials, but since this one would not contain any of the sample, it did not matter whether they mixed or not–it was simply a balance tube. After the centrifuge run, I removed the supernatant from one tube, and since the pellets were hard enough to not move, I poured it into a tube. I then pipetted 1 mL of sonication buffer with BME onto the pellet. I then repeated this steps with the other tubes, resuspending each of them. Finally, I pipetted the resuspended pellets on top of the iodixanol gradient (on top of the 10% iodixanol). It was very important the the iodixanol and resuspended pellet fractions did not mix. I added DI water to the balance tube and Clark spun the gradient at 37,500 RPM for 22 hours at 4 degrees in the same ultracentrifuge rotor used earlier (Sw41-Ti). This marked the end of Monday!

Tuesday – Meetings & SDS-PAGE (attempt)

We had meetings until about 11 so I got back to the lab at around 11:30. When I got to the lab, I immediately set to work on running an SDS-PAGE gel. The lanes were as followed: marker, pellet, supernatant, iodixanol supernatant, viral pellet, band 1 from the gradient, and band 2 from the gradient.

I started the day by turning on the heat block near the gel running station and heating then venting the tubes (by opening and closing). Next, I spun the tubes in the small centrifuge to get out the condensation from inside the tube. I then got an SDS-PAGE gel out of the fridge, removed the well comb, and set up the gel cassette with a buffer dam (since I was only running one gel). I placed the cassette into the gel running box and poured running buffer into it. Next, I loaded the gel with 5 microliters in each of the lanes (mentioned in the previous paragraph), repeating each sample for the Western blot. Then, I ran the gel at 100V for 15 minutes so the dye fronts tightened up on top of the separating gel. Once this was done, I increased the voltage to 180 V then ran it until the dye front ran off the gel. However, I was only supposed to run this for 45 minutes, but I ran it for an hour, so it ran too far off the gel. This resulted in the gel being unusable. I repeated this process on Thursday.

Wednesday – Juneteenth!

Spent the day working on college essays 🙁

Thursday – SDS-PAGE and Western blot

I started the day by putting the samples on the heat block and heated it for 2 minutes. Then, I vented the tubes by rapidly opening and closing and let it heat for 8 minutes for a total of 10 minutes on the heat block. Next, I removed the samples from the heat block and centrifuged them to remove the condensation from inside the tube. Then, I got the gel out of the fridge, removed the well comb, and set up the well cassette using a buffer dam since I was only running one gel. Following this, I placed the gel cassette into the gel running box, then poured running buffer into the cassette until it was about to overflow. Next, I loaded the gel with 5 microliters of marker, pellet, supernatant, iodixanol supernatant, viral pellet, iodixanol gradient band 1, and iodixanol gradient band 2. Then, I ran the gel at 100V for 15 minutes until the dye fronts had tightened on top of the separating gel. When this happened, I increased the voltage to 180V and ran the gel until the dye ran off the front of the gel.

Next, I stained the SDS-PAGE gel with Coomassie blue. I started by grabbing an empty pipette tip box and separating the front plate and back plate using a wedge gel-cracker. I placed the gel cracker into one of the bottom corners and gently pushed the back end of the gel-cracker down to open the gel. I then cut the stacking gel off and threw it away. To remove the gel from the glass, I gently slid the wedge under one of the corners of the front plate and continued pushing up slowly until the gel dislodged fully from one edge. Once this was done, I placed the gel into the empty pipette tip box. Note: Cut the gel into half and use the other half for the Western blot I will describe in the next section. I then poured the staining buffer onto the gel until it was fully covered with a few millimeters of buffer and microwaved the gel and buffer for one minute with the box unclipped. Once the gel was microwaved, I moved the box to the fume hood using heat resistant gloves, then I opened it and let it vent. Then, in the fume hood, I poured the staining buffer into the used staining solution bottle using a funnel and gently rinsed the gel off with DI water. Finally, I added destaining buffer and two kimwipes to the box and left it on the rocker until the gel is destained. I left it overnight. Finally, I put the gel into a plastic sleeve, labeled it, and scanned it.

While this was happening, I was also running a Western blot. I started this process by pouring the Western blot transfer buffer into the transfer box until it was about half full. I then grabbed some filter papers and soaked them in the box with the transfer buffer. While the filter paper was soaking, I grabbed a square plate, brought it to the gel running bench, and filled the bottom with methanol. Next, I cut a blot-sized portion of PVDF using scissors and got the blue protection paper off the blot using special forceps. I then put the blot into the methanol to activate it. To make the transfer sandwich, I placed the sponge, soaked filter paper, gel (backwards orientation), activated PVDF paper, soaked filter paper, and sponge on top of each other in that order. I then clipped it shut and placed it in the Western blot transfer box. Next, I placed a blue cooling block into the front part of the running box, put the box in the fridge, then filled it up with transfer buffer to the “blotting line” on the front of the transfer box. Next, I put the lid on the transfer box, then ran it at 90V for 1 hour. While the transfer was running, I prepared 10 mL of 10% milk in TBST. After the transfer, I removed the membrane from the sandwich and put it into a square plate with all of the 10% milk. I allowed the blot to block for 1 hour and subsequently prepared 10 mL of 5% milk with 1 microliter of antibody (6x Hist) and left it on the rocker for about a minute to mix the antibody into the solution. While the milk and primary antibody was on the rocker, I poured the 10% milk off of the blot and rinsed it with TBST until the TBST ran clear. Next, I poured the primary antibody and 5% milk solution onto the blot and incubated at room temperature for 1 hour. Once this was done, I washed the blot 3 times for 10 minutes each with TBST and made 10 mL of 5% milk in TBST with 2 microliters of secondary antibody while this was blocking. I let the antibody mixture sit on the rocker for about 1 minute to allow it to mix into the milk solution. I poured the last TBST wash off the blot then added the secondary antibody and incubated at room temperature on the rocker for 1 hour. Once this was done, I began with the next set of 3 TBST washes, once again for 10 minutes each. Next, I poured the TBST off the blot into the sink, rinsed with some Milli-Q water, then poured developing solution over the blot until the blot was covered. We usually shake it until we start seeing faint bands, but it didn’t really happen this time. This blot yielded no significant results. Finally, I clipped the developed blot with forceps and put it in the 37 degrees incubator to dry. Finally, I placed it into a plastic sleeve, labeled it, and scanned it.

Friday – Field Trip to Surface Water Plant

Today we went to the City of Sugar Land Surface Water Plant. There are four steps in surface water treatment.

1. Coagulation – Chemicals with a positive charge are added to the water. This positive charge is intended the negative charge of dirt and other dissolved particles in the water. When this happens, the particles bind with the chemicals to form slightly larger particles.
2. Flocculation – This follows coagulation. Flocculation involves the gentle mixing of water to form larger and heavier particles called flocs. Sometimes, water treatment plants will add additional chemicals during this step to help the flocs form.
3. Sedimentation – This step is intended to separate out solids from the water. The flocs settle to the bottom of the water because they are heavier and more dense than water.
4. Filtration – The clear water on top of the flocs are filtered to separate additional solids from the water. The clear water passes through different membrane filters with varying pore sizes that are made from varying materials, including sand, gravel, and charcoal. These filters remove dissolved particles and germs larger than the pores, including dust, parasites, and most bacteria. These filters, which are made out of carbon, also work to remove any bad odors. This water treatment plant used ultrafiltration, in which the water goes through a filter membrane with very small pores. These filters only let water and other small molecules through, such as salts and tiny, charged molecules.
5. Disinfection – After the water has been filtered (step 4), water treatment plants added a few disinfectants (like chlorine) to kill any remaining viruses or bacteria. However, it is also ensured that the water has low levels of chemical disinfectant so that homes and businesses are not harmed by chemical pollutants.

Below are a few pictures from the water plant:

Overall, this was a productive week and I learned a lot of new lab skills as well as valuable information about surface water filtration and why it is necessary!

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.

Today is the end of the second week of the REU program! I gained tons of valuable lab experience this week and started working more individually with less guidance. However, Clark was always there to supervise and answer any questions I had. The first two days involved doing IPTG induction and after that we analyzed the results using SDS-PAGE gels and western blotting.

Monday – IPTG Induction to Induce Protein Expression in E. coli

On Monday, I arrived at around 8:30 AM and we started off the day by making 4 liters of LB broth immediately. This is done, once again, by adding 25 g of LB powder and 1 L of Milli-Q water into an Erlenmeyer flask, mixing it using a magnetic stirrer, and autoclaving the mixture. Once the autoclave was done running (about an hour later), we cooled the flasks by putting them in a bucket with ice and water mixed together as water exchanges temperature faster than gases. Once the flasks cooled down and were safe to touch, we added 1 mL of chlorophyllin and 1 mL of carbenicillin to all four flasks. These are antibiotics and are typically added to cell culture media to prevent microbial contamination and grow select cells that have been engineered with the protein we are looking for. One important note is that all of this process must be done in a sterile field, which is the area immediately surrounding a Bunsen burner. Once the antibiotics were added, liquid culture of E. coli (two of 9337 and two of 10367) were added to each of the four flasks (10 mL per 1 L). Then, the flasks were placed in the environmental warm room to shake at 220 rpm at 37 degrees Celsius. They were shaking until they reached an optical density in the range of 0.5-0.6. The optical densities were periodically checked, the first time being after the 2 hour mark, by measuring the absorbance of the OD600 light using a spectrophotometer. Once they reached the ideal optical density, IPTG was added to each of the four flasks and they were placed in the 16 degrees Celsius shaking incubator overnight.

Tuesday – Protein Sample Extraction from E. coli

Tuesday started off with REU sessions until 12. At around 12:30 PM, I got to the lab and we immediately set to work, pelleting the colonies from yesterday on the centrifuge and pouring the excess supernatant into conical tubes. These were pelleted by centrifugation at 4000 x g for 10 minutes at 4 degrees Celsius. Then, the supernatant was poured off without disturbing the cell and the pellet was resuspended in sonication buffer. I noticed that as the cells were sonicated/lysed, the liquid became more viscous and less turbid. Following the lysis, the lysate was centrifuged at a high speed (10,000-20,000 x g) at the same temperature as before. Samples were taken of the pellets for both 10367 and 9337 as well as the supernatants. The next day, we planned on running SDS-PAGE gels, a process I had become familiar with last week, and Western Blots, a protocol that was completely new to me.

Wednesday – SDS PAGE and Western Blotting

On Wednesday, we immediately set to work at around 8:30 AM making 6 10% SDS-PAGE gels. First, we set up the apparatus by clipping two stacked gel plates to the casting frame and clipping 2 casting frames to each of the three casting stands. Once this was assembled, 1.5 mL microcentrifuge tubes were placed right between the clip on the casting stands to prevent leakage; this is not explicitly stated but is a trick Clark came up with after being in the lab for years. Then, DI water was added between each of the plates just to double heck for leakage. Once the apparatus was set up, the materials needed to make the 10% gel were compiled, but I will not list them here as they are easily accessible on a variety of protocols. Two different mixtures were made: the separating gel and the stacking gel. Under an applied electric field, the stacking gel concentrates the SDS-loaded linear protein molecules while the separating gel separates the proteins on the basis of molecular weight. The separating gel was made first, and all the ingredients were added at first except two: the TEMED and ammonium persulfate. This was because when these ingredients are added, the gel will begin to polymerize and must be immediately poured into the crevice between the glass plates. Thus, these components were only added when the gel was ready to be poured. Upon pouring the separating gel, the apparatus was placed into the incubator at 37 degrees to polymerize. Then, the stacking gel was made with all the same components except the pH of Tris-HCl used was 6.8 for the stacking gel rather than the 8.8 it was for the separating gel. Once the gels were made, SDS-PAGE was run for the samples collected yesterday (I’m not going to repeat how this was done as I mentioned it in last week’s blog post). Once the SDS-PAGE gel was run, Clark noticed a faint band he hadn’t seen before and we ran a western blot to confirm/support the results from the gel. Western blots are an analytical technique used to separate and identify proteins following SDS-PAGE. To run this blot, we prepared transfer buffer and assembled a transfer “sandwich” by soaking the gel and pads in the buffer. Then, to assemble the “sandwich,” we layered in the following order: sponge, pad, gel, PVDF paper, pad, sponge. Then, the proteins were transferred to the membrane in an apparatus that was placed in the fridge at 4 degrees Celsius at 100 V for one hour. Then membrane was then incubated in a 5% milk and TBST solution overnight in the environmental cold room, gently shaking at 4 degrees Celsius.

^ SDS PAGE gel

Thursday – Western Blot Analysis

Yesterday, there was a meeting until 10 AM and I reached the lab at about 10:20 AM. Then, we retrieved and analyzed the results of the western blot. We started by creating 5% milk and TBST solution and added primary antibody to it. Then, we poured this solution into the gel after washing it with TBST and let it incubate at room temperature for 1 hour with gentle shaking. Then, we washed the membrane with TBST three times for 10 minutes each to remove the primary antibody so the secondary antibody can bind to the protein. Then, we created another 5% milk and TBST solution and repeated the same process as we did for the primary antibody but with the secondary antibody. Once these washes were finished, we placed it in the incubator to let it dry and, upon taking it out, noticed bands for 10367 that Clark had never detected before. He will be continuing with molecular cloning for 10367 and I will now primarily be focusing on working with 9337. Once this was finished, we created LB broth and liquid colonies to incubate overnight (same thing we did on Sunday-Monday).

^ Western Blot

Friday – Repeat of IPTG Induction

Today, we repeated the same process we did on Monday with the IPTG induction (LB broth, colonies, antibiotics, IPTG, incubate). This marked the end of the week- a rather tedious week but fascinating and a valuable learning experience nonetheless.

 

Today marks the end of my first week as a student in the START REU program. The first couple days did not involve much hands-on lab experience as it was primarily introductions, basic instructions, and safety training. Although I did not work directly in the lab for the initial portion of this week, it was great getting to meet new people who are just as dedicated to their work as I am and just as excited to have this opportunity. Moreover, I also learned a lot more in-depth details about lab safety, relating to both chemicals and biological materials, that I have not come across in my high school or community college labs. This is, of course, because these institutions do not have abundant resources like Rice does, especially high schools. The second half of the training, relating to hazardous biological materials, was particularly interesting to me as it opened my eyes to how dangerous seemingly safe biological materials can be, such as bacteria and viruses. I will be working with viruses, specifically to determine the structure of an unknown protein within dinoRNAVs, so this lab safety training was both helpful for me and relevant to the project I will be working on for these next nine weeks.

I had the chance to meet my mentor, Clark Hamor, for the second time, with the first time being in early February, and he refreshed my memory on how the Tao lab is organized and lab-specific safety guidelines. Moreover, he has also made these past three days both fascinating through the learning experience and comfortable through the smooth adjustment process, being understanding of schedule-related changes. On Wednesday, which was the first day I got to actually start work in the lab, he taught me how to do certain basic, every-day lab procedures, such as loading and running an SDS-PAGE gel. I had only worked with basic agarose gels and DNA molecules prior to this experience, so it was all new to me. What I understand about SDS-PAGE is that it is a method of electrophoresis that allows protein separation by mass. The medium used in this method is a polyacrylamide-based discontinuous gel. While running, this thin, fragile gel is placed between two glass plates in a slab gel. The concept behind this method is that, when an electrical current is run through the gel, smaller proteins migrate faster due to less resistance from the gel matrix whereas larger proteins move slower, leading to separation of the proteins based on their varying sizes. In the SDS-PAGE method, the use of SDS, which is sodium dodecyl sulfate, and polyacrylamide gel removes the influence of the structure and charge of the proteins, thus they are separated based on their size. After watching him load a few wells, I tried doing the same, and it is a really difficult process when you’re new to it, especially given how thin the pipette tips are. Once we were done with the SDS-PAGE gel and it was de-staining, we moved on to practicing streaking plates with E. coli (but unfortunately left those in the incubator too long as we forgot about them) and also making liquid colonies.

Yesterday, which was a, Thursday was a full day in the lab, and I was there from about 8:30 AM to 5:10 PM. Yesterday, we started off by making four one-liter flasks of LB broth because a large amount of growth medium is needed to grow cells for protein expression. The procedure involved mixing 25 g of Luria Bertani powder and 1 L of purified water into each of the four Erlenmeyer flasks and mixing them (for evenness) using a magnetic stirrer. Following this, we autoclaved each flask to ensure sterility. Then, we followed some steps Clark outlined for protein purification that are specific to his protein (as these steps are very subjective), which involved taking the frozen E. coli cells and putting certain buffers into them, putting the tube into an ice bath, and sonicating it. Following all of this, Clark showed me an ultracentrifuge and taught me how to use it, then told me to go back to the office (for safety purposes) while he ran it. This was pretty much all of day 2, it also involved a lot of reading and concept review/understanding.

^ Me with one flask of LB broth.

^ Me pouring antibiotics into the LB broth as a way to check if the protein has been expressed.

Today, a Friday, I got into the lab around 8:30 AM and Clark went over important, relevant concepts with me, such as lac operons and various protein-related concepts. Then, at around 10 AM, we removed the test tubes from the ultracentrifuge (it was done running around then) and noticed the thin bands the proteins separated into. Clark then showed me how to remove those bands using a syringe, but since they were too close together, it didn’t work out as well for two of them (there were three total). Then, once the liquid removed from the test tubes were put into microcentrifuge tubes of 1.5 mL each, I loaded the wells of an SDS gel with the ladder, loading dye, and the protein samples. One thing I learned from this was that it is important to fill all the wells of the gel (even if it is with loading dye) so that the gel does not run crooked. Then, we ran the gel and went back into the office to read and eat lunch and just discuss things related to the project. After, when the gel was done running, we could tell based on how far the molecules have traveled (they should have come down all the way). After we were done with the gel, we stained and de-stained it and placed it into a pipette holder (box) for sufficient de-staining. Since we left the plates from Wednesday in the incubator for too long, we had to re-streak new plates.

^ The thin bands.

^ Me pipetting loading dye into the wells of the SDS-PAGE gel.

^ The loaded wells, ready to run.

^ Leftover (taken off) polyacrylamide gel named “Blob”

Coming from a background with limited knowledge of advanced topics like virology, proteomics, and general biochemistry, I initially felt a little overwhelmed and uncertain about my ability to understand such complex subjects without prior education. However, Clark has been a great mentor who has been incredibly helpful and made the transition into the program easy and accessible, which helped ease my concerns. I am highly motivated to excel during this internship and am excited to continue learning and enhancing my research skills.