Seth:Quick Transformant Screen

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Contents

Overview

This allows you to screen a large number of inserts the day after completing a transformation, rather than waiting for overnight cultures and performing many unnecessary minipreps. A small amount of DNA is isolated directly from colonies on your plate. PCR amplification using primers complementary to your plasmid lets you determine the presence and approximate size of inserts. The protocol works best for ligation reactions with known insert size(s).

Materials

  • LB (or other suitable media)
  • Selectable marker of choice
  • Sterile deionized water
  • PCR Reagents
    • dNTPs
    • Thermostable polymerase
    • Buffer
      • 50 mM MgCl2
      • 500 mM KCl
      • 200 mM Tris-Hcl (ph 8.4)
    • Primers
    • Red Juice
      • Cresol Red
      • Sucrose
      • MilliQ Water
      • Note: Red Juice is a loading dye for agarose gels. Our lab adds it to PCRs as it saves a step between performing the reaction and running a sample on a gel. There is no negative impact to including this particular loading dye, and anecdotal evidence that it actually improves PCR results. Other loading dyes may have undesirable effects or completely impede the reaction.
  • Agarose
  • Ethidium Bromide
  • DNA Marker

Procedure

  1. After performing your ligation reactions and transformations, plate the transformed cells on agar plates with an appropriate selectable marker. We typically use plasmids with ampicillin resistance.
  2. Grow plates overnight at 37°C
  3. The next day prepare a 96-well plate by adding 20 μL of MilliQ water to as many wells as you have colonies for.
  4. Pick a colony and resuspend thoroughly in the water by pipetting. Cells will lyse over the next several minutes. Begin setting up your PCR immediately.
  5. Prepare a master mix for PCR.
  6. Use 10μL of your lysed cells for template.
  7. As soon as you have taken your template DNA, add 50μL of LB + ampicillin (or whatever selection you are using) to the remaining cells. Cover the plate with parafilm and place at 4°C.
  8. Place samples in a thermal cycler and set program
    • When using M13, T7, or Sp6 primers we use an annealling temperature of 55°C.
    • Adjust extension time depending on your insert size. Be sure to keep in mind the amplified portion of your vector when determining size.
  9. Pour a 1% agarose gel with ethidium bromide.
  10. Load PCR samples and an appropriate ladder (ie 2 log, 1kb, 100 bp)
  11. Electrophorese gel
  12. Visualize your gel and determine which clones have the desired insert.
  13. In a 14 ml culture tube, innoculate 2ml LB + ampicillin with 5μL of the cells from your 96-well plate.
  14. Grow overnight at 37°C and continue to miniprep.

Notes

  • This protocol is meant to be a time and resource saving method, but is not appropriate for every ligation and

transformation. Experiments that yield a large range of fragment sizes or very large fragments, such as BAC or genomic

digests (shotgun type cloning), may exceed the limitations of PCR or your particular taq. Very large inserts may

necessitate such lengthy extension times that the PCR is no longer a time saving tool. Still, with a pool of fragments we

often find this technique useful as it lets us select one or two clones of each size, or choose the clone with our expected

length product.

  • Because you a priming your PCR off the plasmid, there will be a small fragment in every lane of your gel, even if a clone

had no insert. This also means that when looking on your gel you want to factor in that additional DNA when approximating

fragment length. If a lane is completely empty there may have been a problem with your PCR. A template negative PCR reaction

is always a recommended control.

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