20.109(S08):Testing cell viability (Day3)
Today you can stagger your arrivals to lab (see today’s “talk” page). Only one group at a time will be able to work on the microscope, and assuming that cell culture setup takes ~ 1.5 hours, you will each have ~25 minutes to spend on the microscope. Please be respectful of your labmates’ time. Reading the protocol in advance will help you work more quickly, and is strongly recommended.
Part 1: Cell preparation and counting by Trypan exclusion
You will test one of your two replicates for each of your ?three? cell samples. The cells in monolayer culture will be removed from the culture dish using trypsin (as we used for the MES cells in Module 1), while the cells in alginate must be isolated with an EDTA-citrate buffer. Otherwise, the cells can be treated the same way. Unless otherwise stated, all manipulations should be done with sterile technique.
- Aspirate the culture medium from each of your samples. Be careful not to suck up the beads with the aspirating pipet (I will explain how best to do this in class – basically, press the pipet against the plastic at all times, and keep it far from the beads by rocking the liquid around).
- Rinse the cells with X mL of warm PBS, then aspirate the buffer.
- Add X mL of trypsin or EDTA-citrate as appropriate, and incubate at 37 °C for 3 min.
- Now recover your cells:
- To the monolayer cells, add X mL of warm complete culture medium, pipet up and down, and transfer to a 15 mL conical tube.
- To the citrate cells, add (?fewer?) mL of medium, and collect as above.
- Spin the cells down at 8000g for 8 min (using the centrifuge that is in the TC room).
- Resuspend in X mL of culture medium for the monolayer, and Y for the alginate samples. You can adjust the volumes a bit depending on the size of the pellet you see, but write down what you use.
- Take 90 μL of each cell sample you want to measure into an eppendorf tube.
- Take the cell count samples out of the hood, and mix each one with 10 μL of Trypan blue solution. This is a toxic material, so please be careful not to spill it.
- Count each sample on a hemacytometer – averaging just two 4x4 squares (or counting only one if you're pressed for time) is fine. Toss your eppendorfs in the beaker labeled Cell Collect - they will be disposed of as both biological and hazardous waste by the teaching faculty.
- After you count the cells, transfer ~ 106 cells to a labeled eppendorf tube (for each sample). These will be used for performing the fluorescence assay. Your remaining cells should be aspirated, and the conical tubes thrown in the biohazard waste can by the sink.
Part 2: Preparation for Live/Dead® fluorescence assay
- Move to the main lab, and spin down your cells at 250 g for 10 minutes in a microfuge.
- Completely aspirate the culture medium from the cells, as it can interfere with staining.
- Resuspend the cells in 200 μL of dye solution (obtained from the teaching faculty).
- The dyes may be mutagenic, and should be handled with care.
- Cover the tube of cells with aluminum foil, and let it sit for 15 minutes.
- Centrifuge the cell solution once more, then take it to the fume hood.
- Remove the entire supernatant with a pipet, and expel it in the conical tube labeled Dye Collection. The dye waste will be disposed of by the teaching faculty. You should also throw the pipet tip in the solid waste container in the fume hood.
- Resuspend the cells in 50 μL of HBSS buffer.
- Mix in 950 μL of 4% glutaraldehyde solution with the cells, and incubate for another 15 minutes.
- Meanwhile, prepare for microscopy.
Part 3: Microscopy
- If you are the first group using the microscope, you will have to turn on the microscope and allow it to warm up for 15-20 min. On the mercury lamp that is next to the microscope, first flip the ‘POWER’ switch. Next, hold the ‘Ignition’ button for about a second, then release. The lamp ready and power indicators should both be lit up now – talk to the teaching faculty if this is not the case.
- Pipet ~10 μL of your first cell sample onto a glass slide, then cover the solution with a round coverslip.
- Place the slide on the microscope by pulling the left side of the metal holder.
- Begin your observations with the 10X objective.
- Turn on the illumination using the button at the bottom left of the microscope body (on the right-hand side is a light intensity slider).
- Next, turn the excitation light slider at the top of the microscope to ‘DIA-ILL’ (position 4).
- Try to focus your sample. However, be aware that the contrast will not be great for your cells, and you might not be able to focus unless you find a piece of debris.
- Whether or not you find focus, after a few minutes, switch over to fluorescence. Your cells will be easier to find this way.
- First, turn the white light illumination off.
- Next, move the excitation slider to “FITC” (position 3). You should see a blue light coming from the bottom part of the micrscope.
- Finally, you must slide the light shield (labeled “SHUTTER”) to the right to unblock it. Now you can look in the microscope, and use the coarse focus to find your cells (which should be a bright green colour), then the fine focus to get a clearer view.
- You can switch the excitation slider over to EthD-1 (position 2) to see the red-stained cells. Some of your cells may appear to be dimly red, but the dead ones are usually obviously bright.
- Be aware that the dyes do fade upon prolonged exposure to the excitation light, so don’t stay in one place too long, and when you are not looking in the microscope, slide the light shield back into place.
- You can try looking at your cells with the 40X objective as well. As you move between objectives and samples, choose a few representative fields to take pictures of.
- Remove one eyepiece from the microscope, and replace it with the camera adaptor. Be sure to keep the light shield in place until you are ready to take the picture!
- Set the camera sensitivity to 800 for best results (unless your cells are really bright).
For next time
-oral: begin proposal wiki page
-essay: write a draft