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Revision as of 14:19, 18 September 2015

Day by Day Bacterial Protocols Mammalian Protocols



Mammalian Protocols
  1. We Grew Up HEK293-FT mammalian cells from Divya Israni’s cell line.
  2. In order to keep the cells alive, we have to passage them every few day.
  3. Next we count the cells in order to plate them, because we want 50,000cells/mL in order to plate them.
  4. We prepared Tissue Culture Plates to grow HEK293-FT mammalian cells, the chassis for our split protein systems and let them incubate overnight.
  5. We used an Epoch kit to Miniprep the DNA plasmids that are going to be transfected.
  6. We prepare the DNA Transfection Mixture by diluting the DNA plasmids we are going to use to appropriate proportions for our experiments.
  7. Using PEI Mediated Transient Transfection to deliver the prepared DNA to plated HEK293-FT mammalian cells.
  8. Then we let the plates Incubate overnight and a day.
  9. We Prepare the Plates for flow cytometry.
  10. We transfer cells from a 48-Well Plate to a 96-Well Plate.
  11. We perform Flow Cytometry with a Fortessa Flow Cytometer.
Mammalian Cell Culturing
Splitting Protocols
Basic Idea
We need to keep the cells alive and well, so every 3 days you need to “passage” them into a new flask of a 1:10 dilution with new media and such. This is ONLY to keep them alive for later use such as plating and transfecting. Always make sure to work in a specific tissue culture room and to sterilize everything before entering the hood. This is to in general not contaminate the cells.
Materials
  1. Flask with previous passage cells (or initial cell line if first passage) [referred to as flask 1]
  2. New empty flask [referred to as flask 2]
  3. DPBS (buffer to wash away excess media)
  4. DMEM (media for cells to utilize) w/ 25 mL bovine serum, 5 mL streptomyacin, 5 mL L-glutamine, 5 mL sodium something
  5. Trypsin (enzyme to lift cells from bottom of flask)
Steps
  1. Ready waste container with bleach, put anything that touched cells into this
  2. Cells will be adhered to bottom of flask 1 containing media in 37 C, CO2 regulated incubator
  3. Flask should be vented (slightly open) when in incubator
  4. Under microscope cells should look like clouds and take up 70% of the space (this means they are 70% confluent)
  5. Tilt flask 1 so that media is away from bottom (with adhered cells) and remove media using seralogical pipette into the bleached waste bucket
  6. Pipette 4-5 mLs of DPBS into corner of flask 1 away from adhered cells
  7. This is to make sure that no cells are dislodged, which would occur if the reagent was pipetted directly onto the cells
  8. Rock flask back and forth to get media off of cells
  9. Make sure not to get any media in the neck of the tube
  10. Pipette 2.5 mLs of Trypsin into corner of flask 1 away from adhered cells (same motion as DPBS)
  11. Rock back and forth to get trypsin on cells
  12. Vent flask 1 and place in incubator for about 5 minutes (no more than 15 mins) for Trypsin to take effect
  13. Label flask 2 with Cell line, Passage number (first time will be “P0”, second “P1” etc), dilution (concentration of cells to media), initials, and today’s date
  14. FOR A 1:10 DILUTION (good for keeping alive, passaging every 3-4 days)
  15. Add 9 mLs of media to flask 2
  16. Make sure not to splash into neck and that all media is spread across the bottom of flask 2
  17. Look at flask 1 under microscope, should see cells moving in clumps
  18. Add 7.5 mLs of media to flask 1 (to bring total volume of flask 1 to 10 mL as we had already added 2.5 mLs of Trypsin)
  19. Tilt the flask and pipette up and down and across the bottom of the flask to wash the cells into the media (about 15 times)
  20. Try for no bubbles
  21. This will also break clumps, important for single cells to be passaged as they will get proper amount of media and grow better
  22. Put 1 mL of media+cells (for 1:10 dilution) from flask 1 into flask 2 that already has media
  23. Gently rock flask 2 forward and backward then back and forth, making sure not to get anything in neck
  24. Look at flask 2 under the microscope to make sure there are not too many clumps, if so continue to pipette mixture up and down
  25. Vent the flask and place in the incubator anytime the cells are not being worked on in the hood
  26. Your cells are passaged 1:10 and good for 3-4 days!
  27. FY1: If using for plates next day, passage 1:2 (5 mLs cells, 5 mLs media)
Counting Cells Protocol
  1. You will need to count your cells before plating them, even before transfecting them.
  2. Want 50,000 cells/mL for density when plating
  3. Use 1 PCR per flask of cells
  4. If making many plates, mix all flasks together beforehand and ensure single cells only, then do following steps
  5. Aliquot some media using 1 mL pipette from flask 2 into the microtube
  6. Add 10 microliters trypan blue and 10 microliters cells+media to a PCR tube and let them sit for a minute - this stains the cells so they can be counted
  7. Take the half moon sheet for the Countess machine and add 10 microliters of the stain solution into the indented half moon
  8. OR simply pipette onto glass slide and count under microscope, want to finally know cells/mL
  9. Insert into Countess and make sure clean so that there is no fuzziness or it will be counted as dead cells
  10. Want at least 90% of cells to be live
  11. Good number is over 1x10^6
Plating Cells Protocol
  1. Get sterile, tissue cultured 48 well plate
  2. Going to put 250 microliters into each well so total necessary media is 12.5 mLs (round up to 15 for safety)
  3. Math: (50,000 cells/mL)(15 mL) = (live cells/mL from Countess)(X mL) and solve for X
  4. Pipette X mLs of media into reservoir and pipette 15-X mLs of media also into the reservoir
  5. Make sure to mix thoroughly and frequently as to ensure only single cells are plated
  6. Eppendorf large Multichannel settings:(at least 1000 ul multichannel pipette)
  7. Dispense
  8. 250 microliters
  9. Speed 4-5
  10. 4 times drop
  11. tip x x tip x x tip x x tip x x
  12. Look at plate under microscope to make sure that all wells look about the same and that it looks like 50,000 cells/mL
  13. Gently tap plate on all four sides against wall to make sure cells spread out
  14. Incubate at 37 C, CO2 regulated
Transfection Protocol
DNA Mixture
  1. Have spreadsheet made in advance about which plasmid goes into which tube
  2. 8 strip PCR tube holds all of DNA + NACL + PEI
  3. Each PCR tube holds enough for 8 wells = 200 microliters
  4. will only be using enough for 6 wells = 150 microliters to run positive and negatives in triplicate
  5. which is 25 microliters/well
  6. Each tube should get 4 plasmids of 50 ng/ul
  7. This can be adjusted depending on how many plasmids need to transfected, important number is that each well gets 250-300 ng/ul DNA
  8. If tube only needs less than 4 plasmids must use blank plasmid (ps-21)
  9. for example: full integrase control has transfection marker, reporter, full integrase and blank plasmid
  10. for example: dark cell control has blank plasmid, blank plasmid, blank plasmid, blank plasmid
  11. All DNA should be diluted to 50 ng/microliter in advance
  12. Or less if using more plasmids
  13. Add 10 microliters of each 50 ng/microliter plasmid to tube for 4 plasmids total = 2000 ng DNA total
  14. Should have 40 microliters of DNA in each tube
  15. GO TO TC HOOD
  16. Add 60 microliters .15 M NaCl to dilute DNA and bring total volume to 100 microliters
  17. Mix NaCl + PEI to be 42:8 for all tubes
  18. Each tube gets 50 microliters of this mixture, add extra wells to ensure enough
  19. Mix thoroughly in reservoir
  20. Add 100 ul NaCl + PEI to each tube
  21. Mix thoroughly!! PEI is what mediates transfection
  22. Let sit for 15-20 mins
  23. Use large yellow integra pipette (at least 125 ul multichannel pipette)
  24. aspirate 75 microliters
  25. will dispense 25 microliters three different times (into three rows of the negative 48 well plate)
  26. Same protocol for positive plate
  27. Incubate overnight
  28. Incubate all of next day
Transferring from 48 to 96 Well Plate
  1. Tilt plate so that media is aggregated, aspirate 250 microliters (all) of media from 48 well plate using large eppendorf multichannel pipette
  2. Add 50 ul of trypsin to each well (so for 48 well plate need at least 2.5 mL of trypsin in reservoir)
  3. MAKE SURE to move plate north to south and east to west and tip back and forth to make sure that trypsin gets ALL cells off of bottom of plate
  4. Incubate for a 10 mins and make sure to look under the microscope - should see single spheres that move if you tip the plate
  5. DO NOT want cells to come off in sheets, will mean too many cells and likely poor transfection
  6. Add 50 microliters of media to each well (again at least 2.5 mL of media in reservoir)
  7. Use yellow integra multichannel pipette
  8. Mix many times
  9. Aspirate 100 ul and transfer to 96 well plate
  10. Flow cytometry or fluorescence microscopy
Cleaning Up
  1. Pipette bleach into anything with old passages of cells in it (will kill them)
  2. Pipette that into waste container already containing bleach (let sit for about 10 mins) then wash down drain while running water
  3. Flask shut tightly and reservoir, both thoroughly cleaned into biohazardous waste
  4. Wipe down inside of hood with ethanol
  5. Close hood and leave UV light on for 15 mins to sterilize
General Tips
  • 70% ethanol on ANYTHING entering hood, especially gloves
  • Do not push window above the sash level (dictates how much air gets in)
  • Try to keep things in the hood and not take too much stuff in there
  • Always keep incubator at 37 C and 5% CO2
  • Media colors: pink = oxygenated; red = grown; orange - yellow = overgrown
  • Confluence = how close together the cells are; want 70% when passaging or plating
  • HEK203T cells are adhesive so do not move flask too much
  • When in the hood do not pass anything over (eg hand) open containers of cells
  • Labelling new flask for passage: (Lab name) (Cell line)
  • (passage #) (day of the week) (dilution)
  • (date) (initials)