I. INTRODUCTION

The analysis of chromosomes is of importance both for practical and theoretical reasons. Many human malformation syndromes have been correlated with distinct chromosomal abnormalities. Using the newer techniques of chromosome banding it is now possible to identify each chromosome and regions within a chromosome. Banding techniques are used to monitor cell lines, characterize neoplastic cells, map chromosomes, study chromosomal abnormalities, to study taxonomy and the evolution of species and today, to hybridize individual gene probes to specific sites on chromosomes (the so-called "Chromosome Painting"). In this exercise you will prepare chromosome spreads from cultured cells and analyze them by conventional staining.
 

PREPARATION OF CHROMOSOME SPREADS FROM CELLS CULTURED IN MONOLAYER
 

II. MATERIALS

1. 2-3 T75 flask cultures of cells in log phase of growth (no more than 3/4 monolayer).

2. Sterile colcemid (10 µg per ml).

3. Sterile hypotonic solution (0.075M KCl [0.56 gm/100ml] in H₂O). (What will this do to the cells if they are placed in this solution?)

4. Two 15 ml conical centrifuge tubes.

5. Pasteur pipettes and rubber bulbs.

6. Sterile 1.0 ml serological pipettes.

7. Sterile 5.0 ml serological pipettes.

8. Trypsin-versene.

9. Acetic acid/methanol fixative (Carnoy's Fixative) (1 part glacial acetic acid and 3 parts absolute methanol). Must be made fresh and kept chilled.

10. Clean microscope slides (1" x 3"). To clean slides soak in 6N HC1 at least 5 min, rinse thoroughly with distilled water and air dry.

11. Diluted Giemsa Stain
 

III. PROCEDURE

1. Three days earlier, you will have planted 2-3 T-75 flasks of cells so that by today the cells will still be in log phase and occupy NO MORE THAN 3/4 monolayer. Divide yourselves in two groups and add colcemid to two of the actively dividing cultures and mix well. The final concentration should be in the range of 0.01 to 0.5 µg per ml (a good concentration is a middle one of 0.1µg/ml). The optimum concentration will vary according to cell type. To maximize mitotic activity, 50% of the medium may be changed 12-18 hrs prior to adding colcemid.

2. Incubate the cultures for 60-75 min at 37C. Cultures may be incubated for a longer time to accumulate additional mitotic figures, however, remember that excessive time in colcemid results in over contraction of the chromosomes and poor chromosome morphology. A short time, 75 min, is recommended if chromosomes are to be banded (you will not be doing this procedure). If chromosomes are to be analyzed by conventional staining a longer period of incubation may be used. Look at the cultures during this time to see the accumulation of mitotic cells.  How will you tell a mitotic cell from one which is not in mitosis?

3. After incubation tap the flasks vigorously to dislodge mitotic cells and distribute the medium from each of the flasks, equally into two 15 ml centrifuge tubes. Question: Why don't you need to use trypsin to remove the mitotic cells from the plastic in this case?

4. Rinse each flask with 4 ml PBS, again with vigorous movement of the  over the cell sheet, and distribute equally to the two centrifuge tubes.

5. If you have had difficulty in removing these cells from the culture vessel in the past, repeat step 4 using versene-trypsin.

6. Harvest cells with 2 ml of trypsin-versene solution. Triturate gently and add equal aliquots of the cell suspension to the two centrifuge tubes.

7. Rinse flask with 2 ml of versene and distribute equally between the two centrifuge tubes.

8. Pellet cells at 75 x G (600 rpm) for 5 min.

9. Remove all but 2 drops of supernatant with a Pasteur pipette.

10. Resuspend cells thoroughly by tapping with fingers then slowly add 1.0 ml of 0.075M KC1 and again disperse cells by tapping with fingers. Carefully pool the two cell suspensions into one tube. Add additional KC1 to 3-5 ml and disperse cells by tapping.

11. Incubate 20 min at room temperature.

12. Add two drops of fresh fixative, mix, and centrifuge as in 8 above.

13. Remove supernatant with a Pasteur pipette and gently disperse cells. Slowly add fresh chilled fixative
(3 methanol : 1 acetic acid) to 0.5 ml and mix gently, then agitate vigorously. Increase volume to 3-5 ml with fixative.

14. Incubate at room temperature for 10-15 min.

15. Centrifuge. Repeat steps 13 and 14 twice more. Use extreme care in dispersing cells as, at this stage, they are vulnerable to lysing prematurely. .

16. Carefully, resuspend cells in 0.5 ml of fresh fixative in order to obtain a hazy white suspension. (If too opaque/thick, add more fixative until this hazy or smoky white suspension is obtained.) The cell density is critical in achieving good spreads and optimum banding patterns. This is approximately  should yield approximately 50-100,000 cells per drop (0.05 ml) or 1-2 x 10⁶ cells per ml.

17. Apply 1 or 2 drops of cell suspension from a Pasteur pipette to the center of a clean slide held at a 45 degree angle. Let the drop disperse and move down the slide. Once it reaches the bottom you may quickly place the slide under the microscope and watch the cell bursts and liberation of the chromosomes, as the film of alcohol evaporates. This is quite exciting to see! So each person in the lab should have a slide to work on.  If the chromosomes spreads are not separate enough (i.e. you have overlapping chromosomes) you may blow gently on the drop immediately after it touches the slide. A test slide should be examined under bright field or phase contrast optics to check the cell density and adequacy of chromosome spreading. Adjust the cell density by the addition or removal of fixative. Spreading may be increased by adjusting the height of the pipette over the slide and by the intensity of blowing. Too vigorous will result in chromosomes all over the slide!!

18. If the slides are to be stained the following day place them in a 60C oven. If not, place them in a slide box, preferably one containing desiccant. Slides should be placed in a 60C oven several hrs prior to staining to ensure their dryness.
 

IV.  CONVENTIONAL STAINING

Place slides in a Coplin jar containing Giemsa stain, (Harleco Cat. No. 620 or Fisher SO-G-28, diluted 1:50 in Sorenson buffer). Stain for 8 min. Remove slides, rinse in distilled water and air dry. Slides may now (but we will not do this) be mounted using Permount or Gurr's neutral mounting medium. The intensity of staining may be adjusted by altering the concentration of stain, time of staining, or pH.
 

V. RESULTS

1. Observe the preparations using high dry and oil immersion objectives. Scan the slide and determine the number of spreads present, the adequacy of spreading, the quality of staining or banding and chromosomal morphology.

2. How many chromosomes are there per cell? Count the number of chromosomes in 6-10 well spread metaphase cells. Interpret your results.

3. Examine a spread and describe the morphology of the chromosomes in that spread. Locate and determine the position of centromeres (primary constrictions). How many chromosomes are telocentrics, acrocentrics, metacentrics? Can you identify satellites or secondary constrictions?

4. What are you seeing on the slide as you look it over? Are you looking at cells?