Histone H3, phosphorylated - h3f3a - mitotic cells - from upstate and Tissue culture of Xenopus cell lines S3, XTC (Stukenberg lab): Difference between pages
No edit summary |
(Created page with "Protocol submitted by VGP from Stukenberg lab protocols [http://www.xenbase.org/community/person.do?method=display&personId=1311&tabId=0] link to protocol page [http://people.v...") |
||
| Line 1: | Line 1: | ||
Protocol submitted by VGP from Stukenberg lab protocols [http://www.xenbase.org/community/person.do?method=display&personId=1311&tabId=0] | |||
link to protocol page [http://people.virginia.edu/~djb6t/LabWeb/frogs.htm] | |||
Tissue Culture Care Instructions for Xenopus cell lines: S3, XTC | |||
Media: L-15 Medium (Leibovitz), Sigma-Aldrich product # L4386. This comes in powder form, which we resuspend to a 66% solution (dissolve one 14.7g/L vial into 1.4L ddH2O and pH to 7.6). | |||
New media must be filter sterilized into sterile bottles. We usually use Nalgene 500mL receiver bottles and Store media at 4˚C. | |||
Before use with cells, you must add 10% FBS (fetal bovine serum), 1X Penn/Strep (antibiotic, comes as 100X stock), and 1X Sodium Pyruvate (also a 100X stock). | |||
Notes: Antibiotics are good for about 1 month, so if you’re using your media for longer than that, re-supplement it accordingly. | |||
Getting started: | |||
Keep everything sterile. Don’t use non-autoclaved tips, coverslips, or pipets. Make sure all dishes are sterile. | |||
Warm the media and trypsin (if using) to room temperature. (I do this 30-60 minutes before I have hood time). | |||
Make sure the UV light has been on in the hood you’re about to use. Before use, turn off the UV and turn on the blower. Spray gloved hands with 70% EtOH. Wipe down the surface of the hood with 70% EtOH. Spray down and wipe all bottles, (basically anything going into the hood) with 70% EtOH. | |||
Splitting/Passaging Cells: | |||
Remove media. Wash cells with 3-5mL of sterile Dulbecco’s Phosphate Buffered Saline (PBS). Remove PBS and add 2mL 1XTrypsin (red color) per 100mm plate. Swirl around and tap plates a few times. Let sit a few minutes. Add appropriate amount of fresh complete media to new plates for the dilution you’ve chosen. S3 cells grow slowly, so don’t split them too dilute if you need to use them that same week. | |||
Growing cells on coverslips | |||
Coverslips should be at the very least autoclaved or baked at 250˚C. You can also HCl-wash and poly-lysine coat coverslips. Cells are often happier on poly-lysine coated coverslips, so if you’re not happy with how the cells look, you may want to try these. You can transfer coverslips to tissue culture dishes in two ways, either by tweezer (make sure it’s been cleaned well with EtOH, or by using the suction on a sterile Pasteur pipette (of course if your coverslips are in solution, this won’t work). Rinse coverslips a few times in PBS, and then add an appropriate volume of fresh media to them. Then, split your cells as normal, adding a sufficient density of cells to each well. I’d advise splitting them pretty densely so that you can use them the next day if possible. Lab lore indicates that the cells are more mitotic if you use them the next day. | |||
Finishing up | |||
Remove all your materials from the hood. Make sure you’ve turned off the aspirator, thrown away all pipettes, and plugged back in the pipetaid (if necessary). Wipe down the hood with 70% EtOH. Shut the hood, and turn on the UV light. | |||
Revision as of 14:59, 11 November 2011
Protocol submitted by VGP from Stukenberg lab protocols [1]
link to protocol page [2]
Tissue Culture Care Instructions for Xenopus cell lines: S3, XTC
Media: L-15 Medium (Leibovitz), Sigma-Aldrich product # L4386. This comes in powder form, which we resuspend to a 66% solution (dissolve one 14.7g/L vial into 1.4L ddH2O and pH to 7.6). New media must be filter sterilized into sterile bottles. We usually use Nalgene 500mL receiver bottles and Store media at 4˚C. Before use with cells, you must add 10% FBS (fetal bovine serum), 1X Penn/Strep (antibiotic, comes as 100X stock), and 1X Sodium Pyruvate (also a 100X stock). Notes: Antibiotics are good for about 1 month, so if you’re using your media for longer than that, re-supplement it accordingly.
Getting started: Keep everything sterile. Don’t use non-autoclaved tips, coverslips, or pipets. Make sure all dishes are sterile. Warm the media and trypsin (if using) to room temperature. (I do this 30-60 minutes before I have hood time). Make sure the UV light has been on in the hood you’re about to use. Before use, turn off the UV and turn on the blower. Spray gloved hands with 70% EtOH. Wipe down the surface of the hood with 70% EtOH. Spray down and wipe all bottles, (basically anything going into the hood) with 70% EtOH.
Splitting/Passaging Cells: Remove media. Wash cells with 3-5mL of sterile Dulbecco’s Phosphate Buffered Saline (PBS). Remove PBS and add 2mL 1XTrypsin (red color) per 100mm plate. Swirl around and tap plates a few times. Let sit a few minutes. Add appropriate amount of fresh complete media to new plates for the dilution you’ve chosen. S3 cells grow slowly, so don’t split them too dilute if you need to use them that same week.
Growing cells on coverslips Coverslips should be at the very least autoclaved or baked at 250˚C. You can also HCl-wash and poly-lysine coat coverslips. Cells are often happier on poly-lysine coated coverslips, so if you’re not happy with how the cells look, you may want to try these. You can transfer coverslips to tissue culture dishes in two ways, either by tweezer (make sure it’s been cleaned well with EtOH, or by using the suction on a sterile Pasteur pipette (of course if your coverslips are in solution, this won’t work). Rinse coverslips a few times in PBS, and then add an appropriate volume of fresh media to them. Then, split your cells as normal, adding a sufficient density of cells to each well. I’d advise splitting them pretty densely so that you can use them the next day if possible. Lab lore indicates that the cells are more mitotic if you use them the next day.
Finishing up Remove all your materials from the hood. Make sure you’ve turned off the aspirator, thrown away all pipettes, and plugged back in the pipetaid (if necessary). Wipe down the hood with 70% EtOH. Shut the hood, and turn on the UV light.