A/B/C Random Amplification Protocol

Genomic DNA will be randomly primed with a sequence tagged oligonucleotide for 2 cycles.  This will create random genomic products with a specific tag at both ends. These products will then be amplified and labeled with a sequence specific dye primer. The protocol has been optimized for DNA amounts ranging from 1 to 50 ng.  Matt's notes: Template DNA template which exceeds the capacity of Sequenase (in Part A) would carry over and unnecessarily contaminate the PCR reaction in part B.  However part B should be able to handle much larger quantities of sequence tagged substrate than is produced in the Part A reaction.

For two color arrays an equal amount of control DNA should also be amplified in parallel to the experimental DNA.  

<big>Material</big>

DNA fragmentation	Part A:  Random priming	Part B:  Dye-primer PCR	Hybridization reagents
Sau3a (NEB) Sau3a 10x buffer BSA 10 mg/mL	Sequenase, T7 DNA pol, 13 U/µl (USB 70775) 5X Sequenase Buffer Sequenase Dilution Buffer (USB dNTP mix (all 4 dNTPs @ 3 mM) 500ug/ml BSA 0.1 M DTT Primer A (X-1): 40 pmol/µl GTT TCC CAG TCA CGA TCN NNN NNN NN	ThermoPol PCR buffer + MgCl2 (NEB B9004S) 100X dNTPs (20 mM each nucleotide) AmpliTaq polymerase (Applied Biosystems N808-0160) PfuTurbo (Stratagene 60025) Primer B:  100 µM (X-2) Cy5-GTT TCC CAG TCA CGA TC (X-3) Cy3-GTT TCC CAG TCA CGA TC	Human Cot-1 DNA (1 µg/µL) (Roche 1581074, $100/ 500 µg) tRNA (10 µg/µL) Microcon YM-10 42407 G-50 Mini-spin columns (Roche 1814427, $151/ 50 columns) Qia-Quick PCR kit (Qiagen 28104, $72/ 2x 25 columns)

<big>Protocol</big>
Part A.  DNA fragmentation and Priming reactions
1.  Quantify genomic DNA by spectrophotemetry of fluorometry and dilute in T.E. to 40 ng/mL.
2.  Create a restriction digest master mix by multiplying the following recipe x the number of unknown plus control DNA samples (+10% for pipetting error:
Reaction 1:
 
3.  Setup the digest by adding 6 µL of this mix per DNA sample (+ controls) in PCR tubes.

4.  While the genomic DNA is digesting set up master mixes for Reactions I, II, and III by multiplying the following recipe by the number of samples (and controls) + 10% for pipetting error:

Reaction 2	Reaction 3	Reaction 4
2 µL 5X Sequenase Buffer	1 µL     5X Sequenase Buffer	0.3 µL Sequenase
1 µL Primer A (40 pmol/µl)	1.5 µL  3 mM dNTP	0.9 µL Seq. Dilution Buffer
3 µL Total Volume	0.75 µL 0.1 M DTT	1.2 µL Total Volume
	1.5 µL   500 ug/µl BSA
	0.3 µL   Sequenase (13U/µl)
	5.05 µl Total Volume

    
5.  Add 3 µL of Reaction 2 to each tube.  Start the Part A PCR program.  Note: Reaction 3 and 4 mixes will be added as the machine is cycling as explained below:


6.  Let the machine finish heating to 94°C x 2 min. When it then cools to 10˚C add 5 µL of Reaction 3 mix to the sample.  The machine should then slowly ramp to 37°C over 8 min. and then hold at 37°C for an additional 8 min to complete the first cycle.

7.  During the second cycle (again at 10ºC) spike each sample with 1.2 µL of Reaction 4 mix.

8.  When the cycling is complete add 43.8 µL dH2 O to each sample to bring their final volumes to 60 µl.
 
Part B.  PCR amplification with Cy-Dye primers:
The part A reactions produced enough DNA product for 2 part B reactions.  For example 2 arrays could be set up with a dye swap.  The FHCRC human BAC array has internal duplicates so performing duplicate arrays is redundant.  If a single reaction will be performed Cy3 should be reserved for the reference (control) DNA since this fluorochrome has more autoflourescense and a more restricted linear range.  (i.e. curvature at low intensities in log ratio plots.)

1.  Assuming only 1 array will be run per sample then create master mixes for Cy5 primers by multiplying the following recipe by the numbers of samples (+ 10% for pipetting errors).  An equal amount of Cy3 mix should be made for the reference DNA.

Matt’s note:  With this recipe primers and dNTPs are stoichiometrically balanced if the mean product is 250 bp.  The total amount of dNTPs is 15 µg/200 µL.

2.  Divide the each primer mix into 4 PCR tubes (50 µL each).  Start the Round B PCR program and place the samples on the machine when it reaches 94˚C.

3.  To check if the PCR reaction went well run 3 µL on 1% agarose gel. A smear of DNA should be present between 0.5 � 1 kb.  (It can be saved to run with step B.4.d).

Part C.  Preparation of Hybridization Mixture:
Caution:  The product of Part B is a potent source of contamination for future amplifications and should be handled with the same care as any PCR product (gloves, filter tips, benchkote paper).  Consider resticting the work area and equipment while handling this material.

1a.  To remove unincorporated nucleotides use a Sephadex G-50 spin column (1a). Alternatively, a PCR cleanup column (1b).  Sephadex columns can be performed according to Molecular Cloning (Vol.3 A8.29), or with a Roche DNA QuickSpin column.  First equilibrate the column with T.E.  Next pool 4 identical PCR reactions and run them through a single column into an amber 1.5 mL Eppendorf tube.

1b.  As an alternative to sephadex columns clean up the PCR product with QiaQuick PCR cleanup columns.  Follow the manufacturer's directions.   In summary:  Open the PCR tubes and add 250 µL of solution PB to each.  With a P200 pipettor mix the DNA and PB from 2 PCR tubes and combine them in a single QiaQuick column.  (Theoretcally the 4 PCR tubes per sample yield 15 µg of PCR product - based on the amount of dNTPs, and each QiaQuick column has a capacity of 10 µg). Repeat this with an identical PCR sample (labeled with the same dye) to pool the products of two PCR tubes in each column.  Pool two identical PCR reactions (labeled with a single dye) to make 100 µl DNA.  add 500 µl of PB.  Centrifuge for 30" - 60".  Wash with 750 µL PE and spin.  Elute DNA by placing column into an amber 1.5 mL Eppendorf tube, add 50 µL EB and spin.  (Optional - add 25 uL EB to column and repeat spin.)

2. Pool identical sample (labled with same dyes).  Save  a 3 µL aliquot for analysis on a 1% agarose gel.

3.  Add to each pooled PCR product add:

4.  Concentrate samples on a Microcon YM-10 concentrator (MW cutoff ~ 10 kDal, 17 bp).  (Alternatively YM-30 concentrators can be used which have the advantage of being quicker and they eliminate primer dimers - MW cutoff ~ 30kDal, 50 bp).  Spin at 14,000g until volume is ~25 µL.  Spinning to smaller volumes will help to eliminate contaminating small molecules and will speed up the next step.

5.  Pool each experimental sample (Cy3 labeled) with an equivalent amount of control sample (Cy5 labeled) and dry down in speed vac to a volume of <5 µL (not too dry!).