Preparing the Tissue

Please note, tissue preparation is usually done at a distant site.

1. Cut out a section of the lymph node biopsy (as much as can be spared).

2. Trim off excess fat or connective tissue.

3. Place in sterile culture medium (usually provided by cytogenetics laboratory; Subheading 2., step 1) in a universal bottle or similar container as soon as possible after biopsy.

4. Transport to the appropriate laboratory as fast as is practicable (first class post in the UK in usually adequate; see Note 2).

3.2. Setting Up Cultures

1. Macerate the cut node between two sterile defibrinating sticks in a universal to make a cell suspension (see Note 3).

2. Remove any remaining pieces of tissue to a separate container and spin the cell suspension for 5 min at 200g in a bench centrifuge.

3. Replace the supernatant with fresh medium either at room temperature or 37°C but not still cold from the fridge.

4. Perform a total cell count and viability check. If you have access to an automated cell counter use this for the total cell count. Otherwise mix 50 ^L of cell suspension with 50 ^L of red cell lysis reagent such as Zap-oglobin and count the cells in a hemocytometer. Count the cells in the central square and multiply by 2 to allow for the dilution factor and by 104 to calculate the numbers of cells/milliliter in the original suspension. For the viable count mix with Trypan blue instead of Zap-oglobin. Dead cells will take up this vital dye and appear blue. Calculate the percentage viability, taking care to exclude any red cells (which can be identified by their small size) from the count.

5. Set up cultures at 106 viable cells/mL in 5 mL of culture medium, in flat-sided test tubes. See Table 1 for optimal cultures to set up (see Note 4).

6. For TPA cultures, add 0.1mL TPA and wrap tube in aluminium foil to protect from light.

7. Incubate all cultures at 37°C.

8. For F1 cultures block the cells by adding 0.05 mL FdU and 0.05 mL of uridine sometime between 2 and 4 PM on day 1 or immediately if the cultures are set up later than this (see Notes 4 and 5).

9. For F2 cultures, carry out step 8 between 2 and 4 pm the day after setting up the cultures.

10. For F1 and F2 cultures, release the block by adding 0.05 mL of thymidine between 9 and 11 AM the day after the cells were blocked.

11. For F1 and F2 cultures, start the harvest procedure at 5 h, 45 min after releasing the block.

12. For delayed direct cultures start the harvest procedure any time after the cells have warmed up to 37°C after having been set up (see Note 6).

13. For overnight cultures the harvest procedure can be done at any time the day after setting up.

14. For TPA-stimulated cultures, the harvest should be started as near to 72 h after the culture was set up as is practically possible.

Table 1

Optimum Culture Regimes for Different Types of Lymphoma

Table 1

Optimum Culture Regimes for Different Types of Lymphoma

Known high grade or large cells and/or <80% viable

Low grade or small cells with >80% viability

Known/probable, CLL/SLVL or MCL

Deldir, ON, F1, (F2)

ON, F1, (F2)

ON, F1, (F2), TPA3

Deldir, delayed direct culture, harvested later the day it is set up; ON, overnight culture, harvested the day after it is set up; F1 or F2, FdU-synchronized culture harvested 1 (F1) or 2 (F2) after set-up; TPA3, TPA-stimulated culture harvested 3 d after set-up.

Deldir, delayed direct culture, harvested later the day it is set up; ON, overnight culture, harvested the day after it is set up; F1 or F2, FdU-synchronized culture harvested 1 (F1) or 2 (F2) after set-up; TPA3, TPA-stimulated culture harvested 3 d after set-up.

3.3. Harvesting

1. Add 50 pL of Colcemid to the culture, mix well and return to the incubator for 15 min.

3. Remove the supernatant (see Note 7) and gently resuspend the pellet (Note 8).

5. Return to the incubator for an appropriate time. This is laboratory dependent. It will be of the order of 5-30 min (see Note 9).

7. Remove the supernatant and gently resuspend the pellet. Add freshly prepared fixative with the first 1 mL being added dropwise while gently but thoroughly mixing (see Note 10). Top up to at least 5 mL with fix and leave at -20°C for a minimum of 1 h (see Note 11).

3.4. Slide Making for Cytogenetics

1. Wash the cells in fixative four times by spinning at 200g for 10min and replacing the supernatant with freshly prepared fixative.

2. After the final spin, resuspend the pellet in enough fixative for the suspension to appear very slightly cloudy but not milky (see Note 12).

3. Take a fresh clean slide and drop a single drop of suspension (from either a glass or plastic narrow point pipet) on to the slide (see Note 12).

4. Examine under phase-contrast microscopy as the slide dries. Check for suitable cell density, quality of spreading of the chromosomes and phase darkness of chromosome appearance, and adjust the concentration of suspension and time to dry of the slide as necessary (Notes 12 and 13).

5. Make the number of slides required.

6. Age the slides in an oven at 60°C for 1 h (see Note 14).

7. Store spare suspensions at -20°C (see Note 15).

3.5. Banding Chromosomes (see Note 16)

1. Place slides horizontally on a rack over a sink.

2. Add enough H2O2 to cover the area with cells on, and leave for 1.5 min.

3. Rinse thoroughly with tap water and shake the slide to remove excess water.

4. Add 0.5 mL Wright's stain to 1.5 mL of Wright's buffer in a clean container. Mix rapidly and pour on to the slide (see Note 17).

5. Leave for 2-5 min depending on stain batch (see Note 18).

6. Rinse off gently in running tap water and dry with a hair drier.

7. Examine microscopically under a high dry lens (at least x63).

8. Correct staining if necessary (see Subheadings 3.4.1. and 3.4.2.).

9. Mount the slide using Histomount or similar and examine microscopically under a x100 oil lens.

3.5.1. Correcting Understained Slides

(Bloated, Pale Chromosomes With Only Landmark Bands)

1. Add 0.5 mL of Wright's stain to 1.5 mL of Wright's buffer in a clean container. Mix rapidly and pour on to the slide on a rack over the sink.

2. Leave for an amount of time estimated from the degree of underbanding and the length of the initial staining step.

3. Rinse in tap water and dry with the hair dryer

4. Re-examine with the high dry lens

3.5.2. Correcting Overstained Slides

(Dark Chromosomes, Approaching Solid Staining)

1. Soak the slide for 2 min each in 70% ethanol, 95% ethanol containing 1% HCl and 100% ethanol OR

2. Rinse the slide in 3:1 methanol:acetic acid fixative then soak in fresh fix for 5 min OR

3. Rinse the slide in methanol, then water, then methanol again.

5. Add H2O2 to cover the area with cells on and leave for 1.5 min.

6. Wash off thoroughly with tap water and shake the slide to remove excess water.

7. Add 0.5 mL of Wright's stain to 1.5 mL of Wright's buffer in a clean container. Mix rapidly and pour on to the slide

8. Leave for an appropriate time less than the original staining time, estimated from the extent of overstaining and the length of time in the original stain.

9. Rinse off gently in running tap water and dry with a hair drier.

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