Week 9

This week we completed the experimental procedure I discussed in my previous post, and during this process I had the novel experience of working with the flow cytometry machine. Flow cytometry is a procedure by which cells, suspended in a fluid, are pumped one at a time through an extremely narrow tube. As the cells flow through, they are lit by a laser, and a receptor on the other side records the degree of fluorescence that the cells exhibit. We used this machine to test for yellow and red fluorescence in order to quantify the amount of E2F1 and DNA in the cells, respectively.

Unfortunately, we did not grow enough cells, and so we are repeating the 5 day cell phase experiment next week. We had an adequate amount of cells grown in 10% BGS media, because that quantity of BGS allows cells to go into proliferation. However, we also grew cells in starvation media, which contains much less BGS. Placing cells in starvation media pushes the cells into quiescence, preventing them from dividing. We had hoped that by placing the cells in starvation media the transient color would be present to a higher degree at day three, but forgot to account for the fact that the cells would not divide when we seeded the plates. 

Week 8

                This week was spent planning a second experiment that is highly related to the other, longer term experiment that we are still working on. The question we hope to answer with this experiment is, “Is there a threshold to regulate different cell fates? In other words, do different quantities of E2F1 cause cells to enter quiescence, proliferation, arrest, or apoptosis?” In order to answer this question, we are going to take advantage of the fact that the transient cells that we currently have made using the same plasmid contain 10-100 fold variations in quantity of the plasmid. Transient cells are cells that are expressing the plasmid without having incorporated the plasmid into its own DNA, so that the color fades over the course of cell divisions. In most experiments, transient cells are a problem, and the stable cell line that we are trying to make would be used in its place.

                In this experiment, we are going to make use of flow cytometry. We will stain the DNA with a red fluorescent dye called 7-AD, and the protein E2F1 produces already fluoresces yellow with YFP. Flow cytometry will quantify the level that the cells are glowing, and by comparing the amount of DNA to the amount of E2F1, we can determine what quantities of E2F1 cause cells to divide, be arrested, or die.

Week 7

                This week was spent primarily on troubleshooting. We performed several transfections this week, using various plasmids. Once again, we used uncut DNA and DNA cut with the restriction enzyme ApaLI. We also used a positive control using the plasmid mCherry, which is has a high transfection efficiency due to its small size. The bright red color of mCherry also makes it much easier to detect than the YFP that we are using. The point of the positive control is to make sure that the transfection is working as effectively as it should.

                There was extremely high transfection efficiency with the mCherry positive control, exactly what we expected. The transfection efficiency of the uncut and cut DNA was also higher than previous times, leading us to believe that the problems we had with our transfections last week (virtually 0% transfection efficiency) were a single time occurrence and not a problem with the machine. We will start selecting the uncut and cut DNA transfections this upcoming week, as the transfection was so effective.

Week 6

                This week in the lab, we ran into a major problem. Before drug selection, we checked the cells under a fluorescent microscope, and yellow fluorescence was present. We then checked the cells under a regular microscope during drug selection, and saw patches of drug resistant cells forming. However, when we checked the patches this week under the fluorescent microscope, the patches did not fluoresce like we expected. There were only a few cells fluorescing across 12 p100 plates, and one 6 well plate. Such a small number of fluorescent cells bodes ill for trying to form a single cell colony, as we were hoping for a substantial number of glowing cells to sort using flow cytometry.

                The solution that we determined for this problem is twofold. First, we are going to sort the cells that we have already transfected using flow cytometry anyways. The flow cytometry machine tests for glowing cells and has the capability to sort them, either into a microcentrifuge tube or a 96 well plate. We are hoping that the machine will detect more fluorescence then we were able to see with the microscope. We will sort the glowing cells into a microcentrifuge tube and then replate them and let them grow out before re-plating in a 96 well plate.

                We also decided to perform a second and third transfection, each at two voltages. We performed the transfection with both uncut DNA and DNA cut with MfeI-HF. The voltages used were 1900V and 2100V. No sparks were observed. We hope to get better results than in the previous transfection, and the newly transfected cells will provide a backup plan if the flow cytometry fails.

Week 5

                This week was spent drug selecting the cells. The plasmid that was inserted into the cells contains a gene for resistance of a drug called puromycin. Drug resistance is important in selecting which cells contain a certain plasmid, because electroporation is not nearly one hundred percent effective. Only a few cells take up the plasmid, and we need an effective method for isolating these cells from all the others. Drug selection is a useful tool in this process because all cells that do not contain the resistance gene die off, leaving only the cells containing the plasmid to proliferate and form colonies.

                Maintaining a good density of cells on the plate is very important when attempting to create a stable cell line. The confluence of the cells on the plate should not exceed 80 percent. This is because cells respond to a form of inhibition called density dependent inhibition. This means that cells can only divide when there is room for them on the surface they are attached to; when they run into each other, they stop dividing and go into quiescence. If the plate becomes too confluent, the cells will undergo apoptosis and die. Therefore, when maintaining a cell culture, it is important to make sure the plates are split regularly enough that the plate does not become too confluent.

Week 4

                This week we performed another NEON electroporation on a new plate of REF52 cells. The cell type was the same for this electroporation, as was the plasmid used (YFP-ER-E2F1); the only variation was the restriction enzyme digest that was performed. In the previous electroporation, we only linearized the DNA. This means that only a single cut was performed, turning a circular plasmid into a line of DNA. However, the linearized plasmid codes for unnecessary genes and shorter plasmids have higher transfection efficiency. We cut out an unnecessary drug resistance by performing a digest with two separate enzymes, and then performed an electroporation, hoping that this would result in more cells taking up the plasmid.

                In order to perform a NEON electroporation, cells have to be removed from the plate to which they are attached. A chemical called trypsin is used to cause the cells to detach from the plate. However, detaching the cells in this manner causes stress for the cells, and when trypsin is left on the cells for too long (a process called over-trypsinization) cell death may result. In order to avoid this, it is best to work quickly when trypsinizing cells, to remove as much trypsin as possible, and to make sure that the minimum amount of trypsin necessary is used.

Week 3

                This week in the lab we performed our first electroporation on the REF52 cells. Electroporation is a process for getting either linearized DNA or plasmids into cells. Bacterial cells naturally will take up plasmids through a process called transformation, but mammalian cells, such as the REF52 cells we are working with, are unable to do this. Therefore, in order to get our desired plasmid into the cells, we need to open up the cell membrane in such a way that the plasmid can diffuse across. This is done by shocking the cells with a high voltage. This causes the membrane to become porous, and the large DNA molecules are able to easily enter the cell.

                We will be performing two electroporations because the effectiveness of the electroporation can change when the cell is linearized in different ways. Linearized plasmids, or plasmids that are cut so that the DNA is in a straight line instead of a loop, are able to enter electroporated cells more easily than their circular counterparts. The plasmid that we are working with contains some extraneous information encoded into it, and smaller plasmids are able to diffuse into the cells easier than larger ones. We have performed two separate restriction enzyme digest on the plasmid, and each will be electroporated into the cell to see which one is more effective. One cut only uses the enzyme ApaLI, and merely opens up the plasmid without removing any information. The other cut uses both ApaLI and MfeI-HF. This cut removes an unnecessary drug resistance from the plasmid in order to decrease the number of base pairs in the plasmid.