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I. Purpose:

Chorionic villus tissue may be used for prenatal diagnosis of aneuploidy or other structural abnormalities. It is usually done between the 9th and 11th week of gestation. Chorionic villus is not derived from the actual fetus and in rare cases may not have the same karyotype as the fetus, giving false results.

II. Culture Procedure:

A. Aseptic technique must be used when setting up the cultures, preferably under a laminar flow hood. Cultures are grown in a mixture of CHANG and F-10. The media must be fresh (less than 4 days old) and be prewarmed and at pH 7.0-7.5. All cultures are incubated in a wet, 5% CO2 incubator at 37 C. Specimens are processed for both direct and in-situ cultured harvests.

B. Collection flasks are prepared in the lab and given to the physician the day of the procedure. For each flask use 10 ml of complete media and add 2 drops of sodium heparin. Prepare at least one extra flask since it is sometimes necessary for the physician to make two attempts to collect a specimen.

C. For each specimen received label a 100 mm, 60 mm, and 30 mm petri dish with the patient number, patient name, date, and CVS. Add 5 mls of complete media to the 60 mm dish and 2 mls of media to the 30 mm dish. Transfer the specimen to the 100 mm dish, rinsing the flask with extra media if needed to transfer all the tissue. Place the dishes in the incubator for 30 minutes to allow the blood to settle so that the tissue can be viewed more easily.

D. The specimen is cleaned using the stereo microscope; initial sorting at about 1.5 x magnification, then zoom to about 3 x mag for cleaning. Using sterile forceps transfer villus material to the 60 mm dish; villus material is light-colored, usually looking like hydras or lumpy sausages. Maternal material should be left in the large dish; maternal tissue may be bloody clots or medium colored, uniform textured ragged pieces or gloves.
Try to keep the villus material in one section of the dish allowing working room for cleaning. Working in the 60 mm dish using 2 pair of forceps, clean each piece of villus material removing all traces of maternal tissue. Some maternal tissue will need to be "peeled" away from the villus material, look especially for regions of the villus that are stuck together. Discard maternal tissue at the top of the dish and transfer the clean villus to the 30 mm dish. Sometimes tissue may get stuck in the teeth of the forceps, free it with the other forceps so that it does not get stuck back on clean tissue.
Put dish in incubator overnight before continuing the procedure. Any leftover tissue should be transferred back into the flask and saved in case of disaster. Store the forceps in a labeled tube of 70% ethanol for use the next day.

E. The next morning the tissue is processed for in-situ culture, the direct harvest is done as part of this procedure. Label a 30 mm petri dish with patient number, patient name, date, CVS, TRYPSIN + COL. Add 2 ml of 1X Trypsin-EDTA and 2 drops of colcemid working solution. Remove the forceps from ethanol, flame to sterilize, then cool by touching the tips into the dish with trypsin. If the forceps are too hot, they will kill the villus tissue. Transfer the tissue to the trypsin dish, working over a black background makes it easier to see when the tissue is transferred. Incubate the tissue in trysin for one hour at 37 C. Save the forceps in ethanol.
About 10 minutes before incubation is done prepare a collagenase solution of 3 mg collagenase in 3 ml of complete media (collagenase is stored at -20 C and should come to room temp before weighing out). Using a 3 ml syringe and an Acrodisc filter, sterile filter the collagenase solution into a sterile 15 ml conical centrifuge tube. Label another tube for the direct harvest with patient number, direct CVS, and identification letter.

Check the specimen on the inverted scope, cells should be detaching from the outer layers of the villus material. Transfer about 2 ml of the sterile collagenase solution into a new 30 mm dish. Flame a pair of forceps, cool the tips in the collagenase dish, then agitate the villus material to further detach the outer cells. Using the forceps transfer the material to the collagenase dish, the material may be very sticky and may be difficult to remove from the forceps. Using a pipet transfer the material in collagenase back into the centrifuge tube. Label with the patient number, patient name, collagenase and time. Incubate for 1 to 3 hours, mixing occasionally, until the tissue is broken up. Transfer the remaining cells in the trypsin solution into the labeled tube for the direct harvest, rinse the dish with 2 ml of media and add that to the tube. See HARVEST PROCEDURE, DIRECT for further instructions.

When the tissue in collagense is sufficiently broken up label culture dishes, 4 dishes fit on 1 square tray. If there was only a small amount of specimen or if the sample was divided for two people to process, set up 12 coverslip cultures. For large specimens set up 12 coverslips and a T-25 flask. Using flamed forceps place a sterile 20 mm x 20 mm coverslip in each dish; it helps to put a small, 0.01 ml, drop of media in the dish first, this keeps the coverslip in place. Coverslips are set up at 2 or 3 different dilutions to minimize premature overgrowth of cultures. Centrifuge the tissue in collagenase for 6 minutes at 150 x g (900 rpm in Sorval GLC-2B). Remove the supernatant, break up the pellet, resuspend in the appropriate volume of media and plate out 0.2 ml per coverslip, spreading out the liquid without running over the edges.

Dilution Method #1: High and Low
Small sample: Resuspend the pellet in 1.6 ml of media, use 0.8 ml to plate 4 high coverslips. Dilute remainder with 0.8 ml, use all to plate 8 low coverslips.

Large sample: Resuspend the pellet in 3.2 ml of media, use 0.8 ml to plate 4 high coverslips. Dilute remainder with 2.4 ml, use 1.6 ml to plate 8 low coverslips, transfer remaining 3.2 ml to a labeled T-25 flask for back-up culture.

Dilution Method #2: 100%, 50%, and 25%
Small sample: Resuspend the pellet in 1.4 ml of media, use 0.8 ml to plate 4 100% coverslips. Dilute the remainder with 0.6 ml of media, use 0.8 ml to plate 4 50% coverslips. Dilute the remainder with 0.4 ml of media, use all to plate 4 25% coverslips.

Large sample: Resuspend the pellet in 2.8 ml of media, use 0.8 ml to plate 4 100% coverslips. Dilute the remainder with 2.0 ml of media, use 0.8 ml to plate 4 50% coverslips. Dilute the remainder with 3.2 ml of media, use 0.8 ml to plate 4 25% coverslips. Transfer the remainder to a T-25 flask for a back-up culture.

Carefully place the trays of dishes in the incubator overnight. The next day flood the coverslips with 1.0 ml of fresh media, return to the incubator.
On day 3 change the media in the dishes. Gently swirl the dishes to remove un-attached tissue, aspirate off old media, add 2.0 ml of fresh media. Return to incubator, check for sufficient growth day 6.

III. Harvest Procedure, Direct:

A. Centrifuge the tube for 6 minutes at 150 x g.

B. Aspirate and discard all but 0.1 ml of supernatant. Resuspend the pellet by mixing by hand, do not use a mechanical vortexer as this may break the cells. Add 2 ml of prewarmed hypotonic solution and mix gently. Let the tubes sit at room temp for 4 - 5 minutes. Add 2 - 3 drops of fixative to the tube.

C. Centrifuge the tube for 6 minutes at 150 x g.

D. Aspirate and discard all but 0.2 ml of supernatant. Mix by hand to break up the cell pellet. It is important to get the cell pellet completely resuspended, but the cells are fragile and easily broken.

E. Slowly add 2 ml of fixative (one pasteur pipetful) letting it run down the side so that it layers on top of the cell suspension. Mix rapidly by hand. Rinse down the sides of the tube with 2 ml of fixative. Cap the tube. Let it sit at room temp for 15 - 20 minutes.

F. Centrifuge the tubes for 6 minutes at 150 x g.

G. Aspirate and discard all but 0.2 ml of supernatant. Resuspend the cell pellet. Add 2 ml of fixative and mix. Rinse down the sides of the tube with 2 ml of fixative. Cap and let sit at room temperature 15 - 20 minutes.

H. Repeat steps F, G. The culture is now ready to make slides. For best results, slides should be made the same day as the harvest.

IV. Slide Preparation:

A. Use frosted-end slides that have been rinsed with dH2O and stored at 4 C. It may be helpful to warm the slides to room temperature before making slides.

B. Centrifuge the tubes for 6 minutes at 150 x g.

C. Remove all but 0.1 ml of supernatant. Resuspend the cell pellet.

D. Using a pasteur pipet, place 2 or 3 drops of cell suspension near the frosted end of the slide. Spread the cells by tilting the slide and blowing gently. Wipe dry the back of the slide and place it on the slide warmer at 35 - 40 C for 1 - 2 minutes until the slide is dry. It may be necessary to adjust the temperature of the wet slides and time on the slide warmer to maximize cell spreading.

E. Label the slides with the patient number, CVS direct, culture identification letter, and slide number.

F. Allow slides to age overnight at room temperature. Slides are now ready for G, Q, C, or NOR banding.

G. After making slides, add a pipetful of fixative to each tube, cap tightly and store at 4 C for short term or -20 C for long term.

V. Harvest Procedure, In-Situ:

A. Starting on day 6 check the coverslips to see if they are ready for harvest. There should be multiple colonies of 50 - 100 cells on a coverslip. If the colonies are allowed to grow too large they may grow into one another and there will be too much cytoplasm for good banding of the chromosomes. If the colonies are too small, insufficient mataphases may be found.

B. Add 10 mcl Ethidium Bromide working solution to each dish, incubate for 30 minutes. Add 2 drops of colcemid working solution, incubate for another 30 minutes.
C. Ten minutes before end of incubation, start up TECAN harvester so that it will be ready.

D. Harvest using TECAN.

E. Drying conditions are very important for the proper spreading of metaphases. Using the TECAN dry program, remove fixative from the dish. Using the aspirator, remove almost all the fix, going around the edge of the coverslip. Allow to dry in a humid environment, 55 - 60% humidity. If the coverslip dries too rapidly, all the cells will be trapped in the membranes. If it dries too slowly, the chromosomes will float away from the metaphase.

F. Remove the coverslip from the petri dish, keeping it right-side up. It helps to hold the dish with your thumb and middle finger and bend up the bottom of the dish with your index finger so that the coverslip is lifted up. Mount the coverslip on a labeled microscope slide using a drop of mounting media. You can put 2 coverslips on 1 slide.

G. Allow the slides to dry overnight at room temperature. Slides are now ready for G, Q, C, or NOR banding. Metaphases are resistant to trypsinization and also tend to soak up more stain than other types of specimens.

V. Solutions:

Colcemid working solution: 10 mcg/ml Colcemid in Hank's Balanced Salt Solution, store at 4 C.

Complete CHANG/F10 media: 45 ml Chang basal media B, 42 ml Nutrient Mix F-10, 5 ml Chang supplement A, 8 ml Fetal calf serum, 1 ml Penicillin/Streptomycin, solution, 1.1 ml L-Glutamine solution, store at 4 C.

Ethidium Bromide working solution: 2 mg/ml Ethidium Bromide in RPMI-1640, stored in dark.

Fixative: 30 ml Methanol and 10 ml Glacial Acetic Acid, prepared fresh.

Hypotonic solution 0.075 M KCl: 2.8 g Potassium Chloride dissolved in 500 ml dH2O.

1X Trypsin EDTA solution; 10 ml stock trypsin solution (10X trypsin in saline), 0.1 g EDTA (disodium salt), dissolved in 490 ml Hank's Balanced Salt Solution. Sterile filtered, 50 ml aliquots. Store frozen.

VI. Reagents:

Chang media: IRVINE cat# T100-018, liquid and frozen supplement, 100 ml bottle, store supplement frozen, basal media at 4 C.

Colcemid: GIBCO cat# 120-5210. 10 mcg/ml in Hank's Balanced Salt Solution, 10 ml bottle, store at 4 C.

EDTA: SIGMA cat# ED2SS. Ethylenediaminetetraacetic acid, disodium salt, dihydrate, 100 g bottle.

Ethidium Bromide: SIGMA cat# E-8751. 2,7-diamino-10-ethyl-9-phenyl-phenanthridinium bromide, 250 mg bottle.

Fetal calf serum: HYCLONE/STERILE SYSTEMS INC. cat# A-1111-D. defined fetal bovine serum, 100 ml bottle, store frozen.

Hank's Balanced Salt Solution: GIBCO cat# 310-4170. Hank's balanced salt solution, without CaCl2, MgCl2, MgSO4, 500 ml bottle, Store at 4 C.

Glacial Acetic Acid: BAXTER SCIENTIFIC PRODUCTS cat# 9508-2. Acetic acid, Glacial ACS (Aldehyde free), 500 ml bottle.

L-Glutamine: GIBCO cat# 320-5030. L-Glutamine solution 100X (200 mM), 20 ml bottle, store frozen.

Methanol: BAXTER SCIENTIFIC PRODUCTS cat# 3016-1. Methyl alcohol anhydrous AR (ACS)(Absolute) Acetone free, 500 ml bottle.

Nutrient Mix F-10: GIBCO cat# 430-1200. Nutrient Mixture F-10 (HAM) powdered form, prepared to instructions, 10 x 1 liter package.

Penicillin/Streptomycin: GIBCO cat# 600-5140. Penicillin/Streptomycin solution, 10,000 units/ml / 10,000 mcg/ml, 20 ml bottle, store frozen.

Potassium Chloride: COLUMBUS CHEMICAL INDUSTRIES, INC. ACS granular, 500 g lots.

RPMI-1640: GIBCO cat# 430-1800 RPMI-1640 powdered form, prepared to instructions, 10 x 1 liter package.

Trypsin Stock Solution: GIBCO cat# 610-5090. Trypsin 2.5% (10X) in saline. Store frozen.

University of Wisconsin - Madison
Waisman Center Cytogenetics Lab