Array CGH of Microdissected Fresh or Formalin Fixed DNA

Random Prime Amplification of DNA from small tumors (fresh or paraffin)

We generally use 50 nanograms of fresh or paraffin microdissected Taqman-quantitated DNA (see microdissection protocol) for each array hybridization. We have found that some paraffin samples work better if you use 50-100 ng, and some fresh DNAs can be used starting with as little as 10 ng. Quantitation of the paraffin DNA by Taqman is done to standardize the amount of DNA being added to the reactions. It has been our experience fluorometric or absorbance measurements are variable for paraffin DNA, likely due to the poor quality of the DNA and contaminating substances. We generally quantitate fresh DNA by fluorometry, because it is more convenient than PCR based methods. However, fluorometry and absorbance methods do not always agree. The most important aspect of quantitation is consistency and reproducibility.

Note: The Reference DNA is always treated the same as the test DNA, to avoid differential hybridization artifact. Thus, for this protocol the Reference DNA is also amplified. The entire product is then Random Prime Labeled as described below. (Note, we choose to use fresh genomic DNA as the reference, rather than normal DNA extracted from paraffin, in order to reduce the baseline noise of the hybridization.)

1) Random Prime Amplification

Make sure that you amplify enough genomic fresh DNA to use as your reference DNA to be labeled.

Test: Reference:

50-100 ng paraffin or fresh tumor DNA** 50 ng fresh genomic DNA

10 ml 2.5x RP mix (from Invitrogen BioPrime Kit) 10 ml 2.5x RP mix

xx ml ddH2O xx ml ddH2O

22 ml total 22 ml total

Note**Paraffin DNA concentration is measured by Taqman and if DNA concentration is too dilute, excess water is evaporated in speedvac and the DNA is resuspended in 12ml ddH2O. For test DNA extracted from fresh tissue, we use 50 ng (measured by Taqman or Fluorometry).

• Vortex, quickspin. Heat to 100°C 5-10 min. Use thin-walled PCR tubes only. Place on ice immediately for 10 minutes.

• Add 2.5 ml RPA dNTPs (see below for recipe) and 0.5 ml Klenow enzyme (from Invitrogen BioPrime Kit), Vortex gently, quickspin.

Incubate for 2 hours at 37°C. Do not incubate for more than 2 hours.

RPA dNTPS: 200 ml total: 4 ml 100 mM dATP, 4 ml 100 mM dCTP, 4 ml 100 mM dGTP, 4 ml 100 mM dTTP, 2 ml 1 M Tris pH 8, 0.4 ml 0.5 M EDTA pH 8 & 181.6 ml dH₂O

2) Qiaquick purification of amplified DNA product:

After amplification the product is cleaned up to remove unincorporated nucleotides and for buffer exchange into water for the next step of random prime labeling.

· Add 125 ml of PB buffer to RPA reaction (5x volume)

· Place Qiaquick column in 2 ml tube, pipet reaction onto column

· Spin at 12400 x g for 60 sec, 25°C, dump liquid

· Add 750 ml PE buffer to the column

· Spin at 12400 x g for 60 sec, 25°C, dump liquid

· Spin again at 13000 x g for 60 sec, 25°C

· Place column in a clean collecting tube

· Add 34 ml ddH₂0, leave 1 min at room temperature

· Spin at 12400 x g for 60 sec, 25°C. The eluent should be around 32-33 ul.

· Product can be stored at -20°C if needed.

3) Random Prime Labeling

DNA can be labeled by random prime labeling either directly (500 ng) or after a pre-amplification step (see above). For each Test sample labeled with Cy3 a corresponding Reference (gender matched) is labeled with Cy-5. Avoid exposure to light as much as possible for the cy dyes.

· The following is prepared in separate tubes

1) Test: 2) Reference

32.4 ml of the cleaned RPA rxn 32.4 ml of cleaned reference RPA DNA (normal control)

32 ml 2.5x RP mix 32 ml 2.5x RP mix (Bioprime Kit, Invitrogen)

64.4 ml 64.4 ml

• Vortex, quickspin. Heat to 100°C for 10 min. Use thin-walled PCR tubes. Place on ice immediately for 10 minutes.

• Remove from ice, quick spin then add

8 ml RPL dNTPs (see below)

1.6 ml Klenow enzyme (from Invitrogen BioPrime Kit)

6 ml cy3-dUTP (test) or cy5-dUTP (reference) (Amersham/Pharmacia)

15.6 ml total volume

• Vortex gently, quickspin. Incubate for 2 hours at 37°C. Do not incubate the amplified DNA for more than the 2 hours. We do not leave the amplified DNA for longer as we have seen an amplification bias in some regions of the genome when we tried this.

RPL dNTPS: 4 ml 100 mM dATP, 4 ml 100 mM dCTP, 4 ml 100 mM dGTP

2 ml 100 mM dTTP

2 ml 1 M Tris pH 8 (PCR room)

0.4 ml 0.5 M EDTA (PCR room)

183.6 ml dH₂O

200 ml total

4) Sephadex column purification of labeled probe: Purify in Amersham G50 Column

• mix column by vortexing, snap off bottom, loosen cap, place column in 1.5 ml screw cap tall tube, spin 1 min at 735x g.
• Place column in new tube, spin 2 mins at 735 x g. Labeled probe will be in tube. You should be able to see a pinkish tint for the test and a bluish tint for the reference. If it is difficult to see the color when holding the tube over a white piece of paper then the samples probably did not label efficiently. Samples may be stored at -20°C protected from light.

Array CGH Hybridization

DAY 1:

1) Reprecipitate DNA's :

Avoid exposure to light as much as possible for the cy dyes.

• Add the following DNAs to a 1.5 ml centrifuge tube, mixing with pipet:
  100 ug Human Cot-1 DNA (100 ml of 1mg/ml Invitrogen cot DNA)
  all of Cy3 labeled amplified test DNA (~80 ml, which is the volume after the sephadex column of the amplified DNA labeling)
  all of Cy5 labeled amplified reference DNA (~80 ml, which is the volume after the sephadex column of the amplified DNA labeling)
• Add 1/10th volume of room temperature 3M Na Acetate, mixing with pipet.
• Add 2.5 X (original) vol 100% EtOH to ppt DNA, vortex gently.
• Spin 30 mins at 14K rpm (20K x g), 4 °C.

NOTE: while probes are spinning you can begin preparing slides as in step 2. Slides can also be prepared before you precipitate the probe mixes.

• Decant supernatant; blot dry, being careful to avoid DNA pellet. Allow pellet to dry for about 10 minutes.
• Carefully add a 18 ml mixture containing 6 ml yeast tRNA (sigma, 100 ug/ml); 8 ml 20% SDS; 4 ml dH20. Do not resuspend with pipet, allow the pellet to slowly redissolve in the mixture for 5 minutes, then vortex vigorously, quick spin down, let sit for another 5 minutes, vortex vigorously, quick spin down. It is critical that the pellet is resuspended well before the next step.
• Add 42 ml of room temperature MM 1.0, do not resuspend with pipet, vortex vigorously, quick spin down, let sit for another 5 minutes, vortex vigorously, quick spin down. If everything is dissolved then you can continue. It is ok to leave it longer to ensure full resupension of the probe.
• Quickly spin (1 sec) to bring the probe mix to bottom of tube.

2) Slide Preparation:

NOTE: while probes are reprecipitating as above, you can prepare slides. Slides cans also be prepared before you precipitate the probe mixes

• Select slides with arrays that have no marks on the chromium or missing chromium. Array spots should all be nice and round and not touching.
• UV cross link the slide at 1200 mjoules in a stratalinker.
• Mark the array carefully on the top edges with a minimal mark using a diamond pencil (about 2 mm away from outer edges of array). You do not want to etch too deeply or into where the rubber cement border is going to be. Remove any pen marks from the slide, and etch the number on the label side so it is clear which is the top of the slide.
• Using a 22 Ga Needle and a 3 cc syringe carefully make a rubber cement dam around the two arrays. Make sure to leave at least 2 mm from the edge of the array and the rubber cement border. Allow rubber cement to dry at RT for about 10 minutes or more. Protect arrays from dust.

• Place slide on 37 °C slide warmer for a minimum of 30 minutes before adding the probe mix.

3) Hybridization:

• Place slides on 37 °C warmer 30 minutes before adding probes.
• Denature the resuspended probe mix at 70 °C for 10-15 minutes.
• Transfer immediately to a 37 °C water bath and incubate for 30 minutes. (probes can be at 37 °C a little longer if necessary due to handling multiple samples at once. It is best to time the denaturation and 37 °C incubation for the probes in sets of 4 (for two slides).
• Prepare slide incubation chamber. We use a flat rectangular plastic slide holder with the opening on one end. We place a rubber cement dam on the bottom of the plastic container and they can be reused indefinitely. Add 300ul of liquid (we use the HWB wash buffer) in the bottom of the chamber and place chamber on the 37 °C warmer.
• Quick spin down probe mix, resuspend gently with pipet, then apply warm probe immediately onto warmed slide on the hot plate. Spread out the probe mix while adding, and avoid bubbles as much as possible. Repeat for second array, then immediately rock the slide back and forth to spread out the probe mix over the whole array, making sure it reaches all edges of the rubber cement. Sometimes it needs a little help and you can use a pipet tip to gently coach it to the edge.
• Apply probes to second slide, and repeat the spreading out technique, leaving first one on the 37°C warmer. Bubbles from the SDS should disperse.
• It is critical that the probe mix is well dispersed, has minimal bubbles before placing it into the slide chamber. Place slides into chamber taking care to keep the slides level, and to avoid disrupting the rubber cement. Seal the chamber with a layer of rubber cement, then a layer of parafilm. Keep the bottom of chamber as flat as possible, do not fold the parafilm on the bottom.

• Incubate for 2 days at 37 °C in a humid chamber on a slowly rocking platform. Move the chamber to the other end of the rocker 2 times per day to endure equal mixing.

DAY 2: WASHES AND STAINING

• Remove slide from chamber keeping it level, and carefully remove the rubber cement, make sure the array is remaining, moist. If it is starting to dry out then stop and place it into the washes, you can remove the remainder while in the PN step.
• Rinse the slide once with PN, then place into pre-warmed HWB (50% formamide/2XSSC) at 48 °C, Swish slide and then incubate for 15 minutes.
• Wash 1X for 20-30 minutes at 48°C in 2X SSC/ 0.1% SDS. (the longer time is for when there are more than 2 slides). Do not wash more than 4-5 slides at a time.
• Wash 1X for 10 mins at RT in PN, remove any remaining rubber cement during this step, taking care to keep slide fully wet.
• Wash 1X for 10 mins at RT in PN.
• Remove slide from PN, blot back and one side, and leave slide wet, then add 100 ml of RT 0.3ug/ml DAPI in PBS buffered glycerol, and carefully place a 22x50 mm coverslip over arrays, avoiding bubbles. Blot out excess DAPI and PN from the edges onto kimwipes and carefully seal the edges with clear nail enamel.

DAY 3: IMAGING

• Slides are usually left flat, overnight at RT in dark container and imaged the following day.
• Chromium slides must be imaged using a CCD camera set up, typical exposure times on our current in house system, are: DAPI 2000ms; Cy3 45-60,000 ms; cy5 180,000 ms. Clear slides can be imaged on the axon scanner, with typical pmts being for cy3 540, and for cy5 680

NOTES ON array CGH VARIABLES:

Cot-1 DNA: Cot DNA varies from lot to lot, and we measure concentration using fluorometry. Greater than 500 ng/ml is acceptable (1 mg/ml is the expected concentration)

Probe DNA: The amount of probe DNA used can be varied depending on the size of the array. We use all the probe from one reaction, and dissolve it in 60 ml for an array that measures xxxs

Slides: The quality of the slides used is important and should be checked for debris on the slides, smudges, and that the targets are nice and round and not touching each other.

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S. DeVries / F. Waldman

Email questions to Sandy DeVries at the UCSF Cancer Center (devries@cc.ucsf.edu)

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