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.