Talk:20.109(S13):Module 3

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(New page: ==Protocols== Half the class at a time will work in the tissue culture room today. Today will be physically and mentally laborious, and you've all been working hard, so spend the rest of ...)
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==Protocols==
==Protocols==
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Half the class at a time will work in the tissue culture room today. Today will be physically and mentally laborious, and you've all been working hard, so spend the rest of the afternoon however you see fit. (Whether that involves the FNT assignment, notebook prep, or a walk in the sunshine - are we still expecting sunshine?)
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Today you can stagger your arrivals to lab (see today’s [[Talk:20.109%28S13%29:Testing_cell_viability_%28Day3%29 | 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.
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===Chondrocyte or stem cell culture===
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===Part 1: Bead preparation for Live/Dead® fluorescence assay===
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Today you will work with primary cells that are directly isolated from bovine knee joints. Recently, your teaching faculty harvested cartilage fragments from two bovine knees, and sequentially digested them in pronase and collagenase enzymes to release the chondrocytes. Each joint typically yields > 50-100M cells. Stem cells were harvested from the bone marrow and grown up from a rare population by extended culture in bFGF (basic fibroblast growth factor). After cell isolation, aliquots of several million cells each were frozen and stored in liquid nitrogen.  
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#Retrieve your 2 six-well dishes from the incubator.  
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#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.  
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====Preparation====
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#*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.
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#*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.
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#Begin by setting up your hoods. Prepare any standard equipment and solutions needed.
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#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.
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#Note that the small beakers are for making a calcium chloride bath (not shared, one per person), and the large are for temporary waste in steps 10-12 below (shared, one per hood).
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#Within the Petri dish, cut your whole beads in half using a spatula or razor blade.
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#If you requested a special reagent or equipment, check with the teaching faculty.
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#*Small beads may be difficult to cut in half – if so, look at the intact bead instead.
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#If you are doing an alternative protocol (e.g., 2D culture or collagen gels), check with the teaching faculty.
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#Per dish, rinse the beads with 3 mL of warm HEPES buffered saline solution (HBSS).
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#Aspirate the HBSS - this may be easiest/safest to do with a P1000 - then pipet 200 μL of dye solution right on the beads.
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====Cell culture====
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#Incubate for 15 min. with the TC hood light off.
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#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.
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#When your hood is ready, thaw your aliquot(s) of frozen cells in the water bath. Avoid immersing the cap of the tube in the bath; just hold the body submerged.  Agitate the vial slightly while you hold it. The cells should thaw in less than 5 minutes.
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#Rinse the cells with 3 mL HBSS buffer again. Pipet off as much liquid as possible, again into the Dye Collection tube.
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#Spray the vial with 70% ethanol and take it into your hood. Using a P1000, add the cells drop-wise into the 15 mL conical containing 9 mL of pre-warmed medium. Spin at 800 g for 8 minutes.  
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#Soak in 3 mL of 4% glutaraldehyde solution for 15 minutes.  
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#Aspirate most of the medium off your cell pellet, then gently resuspend in 1 mL of medium using your P1000. Add 3 mL more of medium per vial, using a serological pipet for the addition and subsequent mixing of the medium and cells. Take 90 μL of cells into an eppendorf tube.
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#Pipet off the solution, and then bring your Petri dish to the fluorescent microscope bench in the lab.
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#Add 10 μL of Trypan blue - '''this is a toxic material, so please be careful not to spill it!''' - to the eppendorf tube, and count your cells. Adjust your culture plan if you do not have as many cells as you expected.  
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#For observation, place the half-bead on a glass slide and then cover with a coverslip -- don't press down too hard.
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#*No need to count all 4 corners today - perhaps count 2, especially if your cell count is high.
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#*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.
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#Separate the cells that will make up your two different cultures into two labeled 15 mL conical tubes. Note that the tubes may not all require the same amount of cells, depending on the cell densities you chose for the two cultures. Double-checking your calculations now may save you having to do an extra centrifugation step later!
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#*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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#*Give any excess cells that you have to the teaching faculty, in case other groups want more cells.
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#Spin down your two conical tubes of cells at 800 g for 8 minutes.
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#Resuspend each sample of cells in the appropriate amount of the type and concentration of alginate that you chose.  
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#Using the syringe that has been prepared for you, very carefully pull up the cells, then release them drop-by-drop into the beaker full of calcium chloride solution (20 mL). Recall that calcium effectively polymerizes the alginate, resulting in small gel beads filled with cells. '''Immediately discard the entire syringe into the RED sharps container (not the mayo jars) - do not try to remove or recap the needle.'''
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#*Don't release too quickly or you will get a glob instead of distinct droplets, and try to match your release rate with your partner's.
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#*Depending on the concentration of alginate that you chose, you may have between ~50-150 beads for 1 mL of alginate solution.
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#Allow the polymerization to proceed for 10 min. at room temperature. Then pour your beads into a 50mL conical tube.  
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#Remove the calcium chloride solution from your beads using a large serological pipet (to better avoid aspirating the beads), and put this solution in the temporary waste beaker in your hood.  
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#*Ask the teaching faculty for tips on avoiding sucking up your beads. Basically, you want to keep the pipet close to the wall of the conical tube, so liquid can still be sucked up but the beads don't have room to be.
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#Now fill the conical tube with sodium chloride (20 mL), and gently invert it for 1-2 min. This is to remove excess calcium from the solution.
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#Remove the NaCl using a fresh pipet, then wash the beads again with fresh NaCl. Finally, wash the beads two times with DMEM culture medium (20 mL each time).
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#For each of your two samples, transfer the beads to the two leftmost wells of a 6-well plate, using a sterile spatula. Try to put approximately equal numbers of beads in the two wells.
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#Finally, add 6 mL of warm culture medium to each of your four sample wells, then put the two well-plates in the incubator.
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The teaching faculty will exchange the culture medium as necessary (every other day).
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Revision as of 10:35, 25 April 2013

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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