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Lab Notes - Molecular Core Lab

Sybr Green Primer Notes

The most common qPCR reactions incorporate SYBR Green dye into PCR products. Careful primer design is critical. If primer dimers or other non-specific PCR products form, they will incorporate SYBR Green dye. This may lead to inaccurate quantification, especially when detecting sequences of low abundance. Some guidelines for designing primers for qPCR experiments follow:

  1. Choose primers to give a product (amplicon) of 60-300 bp. SYBR Green dye fluorescence depends on the presence of double-stranded DNA. Template regions with obvious secondary structures or long runs of the same nucleotide should be avoided when designing primers. Avoid a 3' terminal T on primers if possible. Thymidine tends to misprime more readily than other bases.
  2. Check the Tm. Primers should anneal to DNA template at 60 C. Usually a Tm of 65 C will be sufficient. Lower annealing temperatures may be acceptable if needed. The Tm can be checked with the OligoAnalyzer from IDTDNA. Keep in mind that use of higher annealing temperatures and short extension times during qPCR may increase the specificity of priming and reduce artifacts.
  3. Check for primer dimers and hairpins in each primer. Primer dimers and hairpins can lead to artifacts and inefficient priming.
  4. If the goal of later experiments is to analyze cDNA, design primers to anneal to transcribed sequences. With eukaryotic DNA, design primers within the same exon. This will allow the use of genomic DNA for controls as described further in point 6. The RNA sample should always be treated with RNase-free DNAse prior to reverse transcription.
  5. BLAST your primer sequences. Run a similarity search of primers against whole organism/non-redundant databases to determine if primers might anneal to other (unwanted) targets.
  6. Run positive and negative controls with qPCR to validate the primers. The positive control can use genomic DNA as a template that has 100% similarity to the two primers. It should produce a single product of the expected size after 40 cycles. This should be verified by running the completed reaction on a gel. Genomic DNA has the added benefit that it can be used as a template for different primer pairs. This information will indicate how efficiently different primer pairs can amplify sequences from the same template. If cloned DNA is available that matches the sequence of the two primers, this can be another useful positive control that will help to establish specificity of priming. The negative (no template) control ideally should not produce any products after 40 cycles. If a product is detected, the same negative control should be included with future reactions to establish when primer artifacts are detected.

Freeze Squeeze

Isolation of DNA from agarose gels.

This procedure is based on (M.R. ISLAM, M. RODOVA,  J. P. CALVET, 2002, A fast and efficient method of DNA fragment isolation from agarose gels without using commercial kits. American Biotechnology Laboratory 20:18).

  1. Prepare fresh TAE buffer and a new agarose gel in TAE buffer. Use a clean electrophoresis chamber. Make the gel just thick enough to hold the DNA sample in a single well. Use the lowest concentration of agarose that will give the needed sample resolution.
  2. Immediately following electrophoresis, transfer the gel and casting tray to a UV light source. Cut out the band of interest with a sterile spatula (flaming works well) and place it in a 1.5 mL tube. Crush the gel slice with the spatula.
  3. Freeze at -80 C.
  4. Thaw the sample at room temperature. Gently flick the tube to make sure that no ice remains. Centrifuge at full speed (20,000 x g or as close as you can get) for six minutes. Remove the supernatant (save) with a pipet tip to a fresh tube.
  5. Crush the gel slice with the pipet tip.
  6. Repeat steps 3 and 4 to recover more DNA from the gel slice.
  7. Pool saved supernatants from steps 4 and 6 and centrifuge at full speed (20,000 x g or as close as you can get) for six minutes.
  8. Remove the supernatant (save) with a pipet tip to a fresh tube.
  9. If there is more than 0.4 mL of liquid, divide it into multiple tubes so there is a maximum of 0.4 mL per tube.
  10. Add 1 uL of glycogen (20 mg/mL, Sigma G-1633, type VIII from slipper limpets) and 1/10th volume of 3 M acetate to each tube. Add 2 volumes of 95% ethanol, mix, and place at -20 C overnight.
  11. Centrifuge at full speed (20,000 x g or as close as you can get) for fifteen minutes.
  12. Decant the supernatant and invert the tube on a paper towel.
  13.  Add 0.5 mL of 70% ethanol and vortex. Centrifuge at full speed (20,000 x g or as close as you can get) for five minutes.
  14. Decant the supernatant and invert the tube on a paper towel.
  15. Dry the pellet in a speedvac.
  16. Resuspend the pellet in 20 uL of water.