20.109(S08):Testing cell viability (Day3)

From OpenWetWare

Revision as of 19:01, 30 January 2008 by AgiStachowiak (Talk | contribs)
Jump to: navigation, search
20.109(S08): Laboratory Fundamentals of Biological Engineering

Home        People        Schedule Spring 2008        Assignments        Lab Basics        OWW Basics       
DNA Engineering        Protein Engineering        Biomaterials Engineering              

Contents

Introduction

Protocols

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.

  1. 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).
  2. Rinse the cells with X mL of warm PBS, then aspirate the buffer.
  3. Add X mL of trypsin or EDTA-citrate as appropriate, and incubate at 37 °C for 3 min.
  4. 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.
  5. Spin the cells down at 8000g for 8 min (using the centrifuge that is in the TC room).
  6. 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.
  7. Take 90 μL of each cell sample you want to measure into an eppendorf tube.
  8. 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.
  9. 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.
  10. 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

  1. Move to the main lab, and spin down your cells at 250 g for 10 minutes in a microfuge.
  2. Completely aspirate the culture medium from the cells, as it can interfere with staining.
  3. 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.
  4. Cover the tube of cells with aluminum foil, and let it sit for 15 minutes.
  5. Centrifuge the cell solution once more, then take it to the fume hood.
  6. 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.
  7. Resuspend the cells in 50 μL of HBSS buffer.
  8. Mix in 950 μL of 4% glutaraldehyde solution with the cells, and incubate for another 15 minutes.
  9. Meanwhile, prepare for microscopy. This includes placing your memory card in the camera.

Part 3: Microscopy

When observing your cells under fluorescence excitation, you should work with the room lights off for best results. You can turn on the working lamp at the microscope bench as you set up your samples, and otherwise when you need to see what you are doing.

  1. 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.
  2. Pipet ~10 μL of your first cell sample (start with an alginate sample) onto a glass slide, then cover the solution with a round coverslip.
  3. Place the slide on the microscope, coverslip-side up, by pulling the left side of the metal sample holder.
  4. Begin your observations with the 10X objective.
  5. 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).
  6. Next, turn the excitation light slider at the top of the microscope to ‘DIA-ILL’ (position 4).
  7. Try to focus your sample. However, be aware that the contrast is not 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 minute or two, 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 microscope.
    • 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 also 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/brightly stained.
    • 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 actively looking in the microscope, slide the light shield back into place.
  8. You can try looking at your cells with the 40X objective as well if you have time. As you move between objectives and samples, choose a few representative fields to take pictures of. As a minimal data set, try to get 1-2 fields at 10X of both of your alginate samples.
    • To take a picture, 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 (to avoid photobleaching)!
    • Set the camera sensitivity to 800 for best results, unless your cells are really bright.

Part 4: Research idea discussion

Find a place (across the hall, in a coffeeshop, etc.) to discuss the five research results you found with your lab partner, guided by the instructions below.

Writing a research proposal requires that you identify an interesting topic, spend lots of time learning about it, and then design some clever experiments to advance the field. It also requires that you articulate your ideas so any reader is convinced of your expertise, your creativity and the significance of your findings, should you have the opportunity to carry out the experiments you’ve proposed. To begin you must identify your research question. This may be the hardest part and the most fun. Fortunately you started by finding a handful of topics to share with your lab partner. Today you should discuss and evaluate the topics you’ve gathered. Consider them based on:

  • your interest in the topic
  • the availability of good background information
  • your likelihood of successfully advancing current understanding
  • the possibility of advancing foundational technologies or finding practical applications
  • if your proposal could be carried out in a reasonable amount of time and with non-infinite resources

It might be that not one of the topics you’ve identified is really suitable, in which case you should find some new ideas. It’s also possible that through discussion with your lab partner, you’ve found something new to consider. Both of these outcomes are fine but by the end of today’s lab you should have settled on a general topic or two so you can begin the next step in your proposal writing, namely background reading and critical thinking about the topic.

A few ground rules that are 20.109 specific:

  • you should not propose any research question that has been the subject of your UROP or research experience outside of 20.109. This proposal must be original.
  • you should keep in mind that this proposal will be presented to the class, so try to limit your scope to an idea that can be convincingly presented in a ten minute oral presentation.

Once you and your partner have decided on a suitable research problem, it’s time to become an expert on the topic. This will mean searching the literature, talking with people, generating some ideas and critically evaluating them. To keep track of your efforts, you should start a wiki catalog on your OpenWetWare user page. How you format the page is up to you but check out the “yeast rebuild” or the “T7.2” wiki pages on OpenWetWare for examples of research ideas in process. As part of your “for next time assignment” you will have to print out your wiki page specifying your topic, your research goal and at least five helpful references that you’ve read and summarized.

For next time

-oral: begin proposal wiki page

-essay: write a draft

Personal tools