Talk:20.109(S13):Module 3

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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 hour, you will each have ~20-25 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: Bead preparation for Live/Dead® fluorescence assay

  1. Retrieve your 2 six-well dishes from the incubator.
  2. The teaching faculty counted your beads during a recent media exchange (they are easiest to count in the absence of media). Based on the numbers written on your plate, decide how many beads (1-3 per sample) you can spare for today's assay. Ideally, for the three assays on Day 4 you want at least 45-60 beads total remaining (perhaps 30 or fewer for large beads). Be sure to take your bead(s) from only one of the two wells, just in case you contaminate it.
    • Also take this time to describe bead uniformity in your notebook, as this feature may affect your eventual experimental outcomes. Some groups had more luck than others in keeping bead size consistent between and within their two samples.
    • During a later incubation step, you might also take a look at your plate under the microscope, and focus in on cells within the beads. What is cell morphology and density like in each sample? Are there any cells growing under the beads, as a monolayer on the surface of the plate? Keep in mind that these will compete for media nutrients with the cells inside the beads.
  3. Using a sterile spatula, remove the beads (keeping the two samples separate) to two labeled Petri dishes. Do your best to keep the beads remaining in the culture wells sterile – the cells have to stay alive for 5 more days. Briefly dip the sterile spatula into the well, and immediately return your plate to the incubator, onto the shelf from which you took it.
  4. Within the Petri dish, cut your whole beads in half using a spatula or razor blade.
    • Small beads may be difficult to cut in half – if so, look at the intact bead instead.
  5. Per dish, rinse the beads with 3 mL of warm HEPES buffered saline solution (HBSS).
  6. Aspirate the HBSS - this may be easiest/safest to do with a P1000 - then pipet 200 μL of dye solution right on the beads.
  7. Incubate for 15 min. with the TC hood light off.
  8. 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 into the container on the microscope bench; tips will later be disposed of as solid waste in the chemical fume hood. You do not need to throw any later tips away here, as the dye will then be very dilute.
  9. Rinse the cells with 3 mL HBSS buffer again. Pipet off as much liquid as possible, again into the Dye Collection tube.
  10. Soak in 3 mL of 4% glutaraldehyde solution for 15 minutes.
  11. Pipet off the solution, and then bring your Petri dish to the fluorescent microscope bench in the lab.
  12. For observation, place the half-bead on a glass slide and then cover with a coverslip -- don't press down too hard.
    • You will probably want to look at the beads both flat side up (to see the core) and flat side down (to see the surface), time permitting.
    • You can make a "map" of the beads in your notebook and/or on the white surface of the slide. For example, you might have one bead on the left that is core side up and another on the right that is surface side up.
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