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
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
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
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
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
- 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
- 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
- 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
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
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
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
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
B. Centrifuge the tubes for 6 minutes at 150 x g.
Remove all but 0.1 ml of supernatant. Resuspend the cell pellet.
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
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.
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.
minutes before end of incubation, start up TECAN harvester so that it will be
D. Harvest using TECAN.
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
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.
Colcemid working solution: 10 mcg/ml Colcemid in Hank's Balanced Salt
Solution, store at 4 C.
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.
working solution: 2 mg/ml Ethidium Bromide in RPMI-1640, stored in
Fixative: 30 ml Methanol and 10 ml Glacial Acetic Acid, prepared
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.
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
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
Fetal calf serum: HYCLONE/STERILE SYSTEMS INC. cat# A-1111-D.
defined fetal bovine serum, 100 ml bottle, store frozen.
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.
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
Penicillin/Streptomycin: GIBCO cat# 600-5140.
Penicillin/Streptomycin solution, 10,000 units/ml / 10,000 mcg/ml, 20 ml bottle,
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.
Solution: GIBCO cat# 610-5090. Trypsin 2.5% (10X) in saline. Store