Koshland Web Site/Methods
Chromosome Spreads II
  1. Spin 5-20ml of exponentially growing culture (i.e. 10ml O.D.=0.5). I use 5 O.D. of cells.
  2. Resuspend the pellet in 1ml Solution 1and transfer cells into eppendorf tube. Cells can be kept on ice like that until the end of the time course.

    Digestion of the cell wall:
  3. Spin the samples at full speed and resuspend pellet in the Spheroplasting solution:
    300ul Solution 1
    6ul DTT (1M)
    20ul Zymolase (3mg/ml) this is the same stuff used for FISH
  4. Incubate at 37°C, check spheroplasting after 6 minutes by mixing 5ul sample with 5ul 2% sarcosyl. Incubate until 90% of the spheroplasts lyse by contact with sarcosyl (It is done by 10 minutes usually).
  5. Transfer the cells into a 2ml eppendorf tube that already contains 1.5 ml of ice cold Solution 2 then mix by gently inverting. Spin down 8’ 800rpm, suck off supernatant carefully and resuspend into 200-400ul Solution 2. Keep on ice until spreading (O/N is also okay).

    Spreading (acid wash and dry slides ahead of time):Nasmyth used MENZEL SuperFrost slides for spreading.
  6. I just took standard slides, washed them with H2O then placed them in boiling 0.01M HCl for 10 minutes. Remove them from the HCl, rinse with 100% ethanol and let dry on paper towels.
  7. To put cells on slides and "spread" the chromosomes, pipet these solutions in rapid succession on the same spot of the slide (UNDER THE FUME HOOD). Each slide should contain only one strain or condition (HU or Nz arrest).

    Cell Suspension: 20ul
    Fixative: 40ul
    Lipsol (1% in water) 80ul (LIP LTD)
    Fixative: 80ul
    Distribute the drop over the slide by moving a glass rod gently parallel to the slide. Dry at room temperature in the fume hood for 2hr. or O/N. If you don’t want to proceed with the immuno-staining, slides can be stored at –80°C.

    Before proceeding prepare
    a. Humid chambers: make enough chambers so that you have 3 or less slides per chamber.
    b. Make 20 ml Blocking buffer: Thaw out 1 tube of 10% BSA/2X PBS (stored in –20°C) and transfer 10 ml into a 50 ml conical tube. Add 9.6 ml H20 to the conical tube and mix by inversion. Get the Blotto that is in a snap cap tube in the 4°C deli refrigerator and vortex well until there are no large clumps then add 400ul to the conical tube and vortex. This is the Blocking Buffer that you will use below.

  8. Place the slides in a coplin jar containing 1X PBS (the level of PBS should be BELOW the writing on the slides otherwise the writing will wash off). Incubate for 30 minutes at room temperature (NO agitation).
  9. Remove each slide from the humid chamber and drain off the excess liquid then add 400ul Blocking Buffer (only do ONE slide at a time to prevent slides from drying). Incubate for 10 minutes at room temperature in humid chamber
  10. Prepare 400ul Primary Antibody for each slide
    Note: MYC tagged proteins: Take 9E10 Mouse anti-myc antibody and dilute 1:5 in Blocking Buffer.
    HA tagged proteins: Take 16B12 Mouse anti-HA antibody and dilute 1:2500 in Blocking Buffer.
  11. Remove all slides from the humid chamber. Treat one slide at a time. Pour off excess buffer from the slide and immediately add 400ul of the primary antibody. Place slide back in the humid chamber. Repeat until all slides have primary antibody.
  12. Incubate slides for 1 hr at 23°C in the humid chamber.
  13. Pour off the solution and place the slides in a coplin jar containing FRESH 1X PBS (the level of PBS should be BELOW the writing on the slides otherwise the writing will wash off). Incubate for 10 minutes at room temperature (NO agitation).
  14. Prepare 400ul Secondary antibody for each slide.
    Note: If the primary antibodies were from mouse: Take Cy3 conjugated Goat anti-mouse antibodies and dilute 1:3000 in Blocking Buffer. (The secondary is a fluorescent antibody so keep in dark by covering with foil or incubating in a closed drawer).
  15. Remove each slide from the humid chamber and drain off the excess liquid then add 400ul fluorescent secondary antibody and place back into the humid chamber.
  16. Incubate in the dark for 1 hr at 23°C in the humid chamber.
  17. Pour off the solution and incubate in coplin jar as described in step 6.
  18. Remove each slide from the coplin jar, pour off the residual buffer and add 50ul mounting media containing DAPI (100 ng/ml DAPI). Cover with large coverslip and seal with rubber cement.

    If the signal is too weak, try with the third antibody (anti goat-Cy3 from donkey, Jackson, 1:2500)


    Solution 1
    Recipe for 500ml:
    40.1ml K2HPO4
    9.9ml 1M KH2PO4
    109.32g Sorbitol (MWT 182.2)
    250ul 1M MgCl2
    Add 325ml H20 and dissolve, then pH to 7.4 with 3M KOH and bring to 500ml with H20.
    Filter sterilize (do not autoclave).

    Solution 2
    Recipe for 500ml:
    9.76g MES (195.2 FWT)
    1ml 0.5M EDTA
    250ul 1M MgCl2
    91.1g Sorbitol (MWT 182.2)
    Add 375ml H20 and pH to 6.4 using 3M KOH (~10ml).
    Filter sterilize (do not autoclave).

    *20% Paraformaldehyde Stock
    Place 10 grams paraformaldehyde in a 50ml conical tube.
    Bring volume to 40 ml distilled H2O
    Add 0.5ml 1N NaoH and heat to 60°C in H2O bath.
    When dissolved, bring to 50ml with H2O then filter sterilize. Can store at room temperature for 1 year.
    Make 4.25% Sucrose stock and filter sterilize.
    Make 20% Paraformaldehyde (see above).
    Then mix 1 part 20% Paraformaldehyde and 4 parts 4.25% Sucrose.
    Yields 4% Paraformaldehyde and 3.4% Sucrose (Make Fresh each time)

    *This is the Nasmyth method. I also used Sue Biggins method, which is easier to make and it worked.

    One step Fixative
    Add 40 ml H2O to a 50 ml conical tube and warm to 60°C.
    Add 2g Paraformaldehyde
    Add 100 ul 1N NaOH and mix by inversion. Place in H2O bath to warm if necessary.
    When paraformaldehyde is dissolved, add 1.7g Sucrose, mix and bring to 50 ml with H20.
    Filter sterilize and store at 4°C. ( Sue says you can use it for 3 months but I used a solution 6 months old and it worked)

    Blocking Buffer (Bob Skibbens formula)
    5% BSA
    2% BLOTTO
    1X PBS

    *Doug’s method to make 20% paraformaldehyde stock:
    Weigh enough to make a 20% solution (do the rest in a hood). Add water, stir and heat slowly (never boil) Add dropwise from a 1M stock of Na2CO3 until paraformaldehyde dissolves. This makes the solution basic (the final concentration of Na2CO3 should not exceed 4mM). After it is all dissolved, continue heating solution for another hour at 80°C (this drives of CO3 as gas and pH will go down). The final pH is 4-6. This stock can be saved on the bench top for months.

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