Core sample PCR method

rybicki at rybicki at
Tue Jun 9 04:14:50 EST 1992

I got an excellent response to my post on core sample PCR, so I presumed some
more people may appreciate it too.  It was developed in Steve Howell's lab at
the Boyce Thompson Institute, Cornell University, while I was on sabbatical 
and had time to play.  It may well have appeared in print somewhere, but I 
haven't seen it yet.  Works for us!

                       "CORE SAMPLE" PCR:
   A method to re-PCR unique bands from products of mixed size
                         by: Ed Rybicki
                   Department of Microbiology
                     University of Cape Town
                       PB Rondebosch 7700
                          South Africa
               email (internet) ed at

The products of a PCR reaction - especially when this is done on
eukaryotic genomic DNA, and when using degenerate primers - often
contain a mixture of discrete-sized bands, one of which is the
"right" one, while the others represent products of "non-
specific" priming.  It can be a problem to obtain the correct
band in any state approaching purity while maintaining yield, and
attempting to purify the band by cloning all the reaction
products and then probing the library for the correct DNA can be
extraordinarily tedious.

I have applied a simple "core sampling" procedure - involving
"coring" an agarose sample out of a gel, and using it as template
for another round of PCR - to get around this problem, and obtain
unique bands from initially messy backgrounds.  Of course, having
a visible band of the size expected does help; however, the
technique may be used on faith on "right-sized" invisible bands
if need be.

1.   Run products of a PCR amplification on ordinary 1-2% TBE 
     agarose gel, as two or more replicate lanes.

2.   Cut off 1 lane - flanked by marker DNA if desired, and
     notched to allow re-orientation with remainder of gel - and
     stain in preferred ethidium bromide concentration (I use 50
     ng/ml for 10 min).

3.   View excised stained piece on 254nm UV box for maximum
     sensitivity; notch or stab correct band(s) in sample lane.

4.   Prepare "core samplers": using gloves and sterile scissors
     and cut off about 5mm from the tip of as many sterile yellow
     pipette tips (we use Gilson tips) as you will need for

5.   Align stained marked segment with remainder of gel.  Use
     "core samplers" to stab out one or more cores of agarose
     from the centre of bands of interest, using stabbed/notched
     gel lane as reference: a standard gel should give about 10ul
     per core.  You can store extra cores in the tips, wrapped in
     Parafilm, at 4degC, for ever.

6.   Stain remainder of gel, view and photograph at 254nm to
     ensure correct regions were sampled.


7.   Use core samples as substrate in PCR reactions: I make up
     40ul/reaction of reaction mix, and allow 10ul per core.
     One can also shave off as much/little as one likes from an
     extruded core and put that into a reaction mix.
     Simply add core to mix, (push out of tip with sterile toothpick)
     vortex a little, spin down, cover with mineral oil.  PCR 
     according to taste (not inhibited by presence of a little 
     bromophenol blue or of 50ng/ml ethidium bromide).

8.   At end of PCR: if you allow the tubes to cool down the
     reaction mix will set: 2%-odd agarose diluted 1/5 sets quite
     well!  This is no problem for gel running as you then end
     the PCR on a 10 min 72oC cycle, and load the sample into
     wells of a gel BEFORE submeging the gel: sample will set in
     the wells and not float out.

9.   If you wish to extract DNA, end at 72oC and add 50ul pre-
     warmed phenol / 8-OH-quinoline and vortex, add 100ul
     chloroform / isoamyl alcohol (24:1), vortex, spin: agarose
     should be in the phenol/CHCl3 phase.  ALTERNATIVELY: take
     off mineral oil using 50ul CHCl3, take out plug of
     solidified sample and wash in TE, then put into 0.5ml
     Eppendorf-type vial with some siliconised glass wool at
     bottom, and a small needle hole.  Put little Eppi in big
     Eppi without a lid, and spin 6000 rpm 10 min (a la Heery et
     al., 1990; TIG 6(6):173).  Collect filtrate, clean up by
     phenol/CHCl3 and isopropanol/ammonium acetate ppte (1 vol
     IP, 0.2 vol of 10M soln AmmAc).

I have successfully re-amplified a unique 500bp band from a
background of many bands up to 1.5kb from a cDNA PCR of
cauliflower mosaic virus 35S RNA in total turnip RNA extract, and
a 150bp band from a background of bands going up to 3kb from an
amplification of Arabidopsis total genomic DNA using thoroughly
degenerate primers - in the latter case, to a point where it
could be sequenced directly (using same primers) after a
subsequent amplification after purification from a gel plug as

The applications to RFLPing appear obvious: for example, even if 
you use RAPD primers, you could stab out and amplify polymorphic 
bands and use them as RFLP probes.  The advantage over toothpicking
bits of agarose out are that you can store cores in tips for future 
reference, and thge volume is more controllable (you can also take
a bigger sample...).

| Ed Rybicki, PhD             |    "Now you've got the hang of it    |
| (ed at        | There's nothing you can't do with it |
| Dept Microbiology           |        If you're very into it        |
| University of Cape Town     |         You can't go wrong...."      |
| Private Bag, Rondebosch     |                                      |
| 7700, South Africa          |               -Mad John              |
| fax: 27-21-650 4023         | (Ogden's Nut Gone Flake, Small Faces)|
 _ _______________________________________________________________________
|  Ed Rybicki                     |    Now you've got the hang of it      |
|  Dept Microbiology              | There's nothing you can't do with it  |
|  University of Cape Town        |         If you're very into it        |
|  PB Rondebosch 7700             |         You can't go wrong....        |
|  South Africa                   |                                       |
|  ed at             |              - Mad John               |

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