DNA Extraction from Bird Feathers

The least invasive sources of DNA are small feathers plucked from a bird’s breast. There are videos on YouTube that will show you how to do this in a way that is least distressing to the bird. For this DNA extraction protocol to work, the feathers need to be freshly plucked and not moulted.

When you have the feathers, use a new scalpel/razor blade on a clean surface (e.g. a cutting mat/non-scratch surface) to prepare one or two small sections from each feather of approximately 2–3 mm in length. If your feathers are very small then you may need sections from two feathers or more per sample, while if they are large then you may benefit from using only half of a cut section (e.g. 1–2 mm³ diameter of feather tip tissue).

• For smaller feathers, sample from the tip of the feather 1, which should have skin cells attached from being recently plucked.
• For larger feathers, there is also a DNA-rich area where the feather barbs start to sprout from the calamus 2 that contains a DNA-rich blood clot (Horváth et al. 2005).

Place the feather section/s in a PCR tube labelled on the top and side in permanent marker. To avoid sample cross-contamination, sterilise your scalpel/razor blade and cutting surface between feathers.

Make sure the feather sections are at the base of the tube before starting the DNA extraction; flick or tap the tube on a hard surface until this happens and then keep tube upright.

Pipette 75 μl of HotSHOT Alkaline Lysis Solution into each labelled PCR tube containing a sample. To do this, set the 20–200 μl adjustable pipette to 75 μl and use the 2–200 μl pipette tips. You can either use a fresh pipette tip each time, or use the same pipette tip to carefully squirt the reagent into the tube without touching the sides of the tube. If your pipette tip does touch anything, discard it and use another fresh pipette tip.

Place the sealed PCR tubes in the thermocycler and use the heat block function to heat tubes at 95^oC for 20 minutes.

Leave tubes to cool and then pipette 75 μl of HotSHOT Neutralising Buffer into each labelled PCR tube containing a sample. Again, you can use a fresh pipette tip each time, or use the same pipette tip to carefully squirt the reagent into the tube without touching the sides of the tube. If your pipette tip does touch anything, discard it and use another fresh pipette tip.

Mix the solutions thoroughly by holding each PCR tube between your thumb and forefinger and inverting the tube several times.

If you have access to cold storage or ice, you may get better results by putting it on ice or in the fridge to cool it rapidly and improve precipitation of suspended tissue.

The PCR tube now contains DNA in neutralised buffer. The crude DNA extract in neutralised HotSHOT buffer should be stable at room temperature while setting up a PCR. The extract should be stored in the fridge for short term storage (e.g. during the day), and frozen overnight for long term storage if needed.

For organised storage, we suggest checking the tubes are well sealed, and storing in a small press-to-seal bag. The sample data (date, sample ID, any important metadata) can be written on a small piece of paper and stored inside with the tubes. The outside of the bag can also be labelled with permanent marker with the date of PCR and any relevant information. Multiple bags can be stored in date order in a box in a freezer. Samples should be stored in separate bags and boxes to PCR products to reduce the possibility of PCR contamination. If using this system it is important to only work with one open bag of samples at a time to avoid confusion between identically numbered 0.2 mL tubes.

Your samples may require dilution prior to PCR due to the presence of PCR inhibitors that were co-extracted with bird DNA. A 10x dilution is usually effective in reducing PCR inhibition, but a 20x–100x dilution may also be needed in some cases. Dilution is best done immediately before PCR because the DNA will degrade if stored only in sterile water.

To dilute your samples by 10x, you will need your DNA extracts (defrosted if previously frozen) (1), an empty PCR tube for each extract to be diluted (2), a permanent marker pen for labelling (3), a 20 μL micropipette (4), 20 μL pipette tips (5), and sterile distilled or PCR grade water for dilution.

The first step is to label each tube in a way that allows you to match these to your DNA extract tubes. One approach would be to label your extraction tubes sequentially, e.g. 1–8, and label your dilution tubes 1d-8d.

Next, for a 10x dilution, pipette 18 μL of sterile distilled water or PCR grade water into each tube. You can use the same pipette tip for all tubes provided it doesn’t touch anything but water and the empty PCR tubes. For a 50x dilution, use 98 μL of sterile distilled water.

Finally, pipette 2 μL of DNA extract into each appropriate tube, using a new pipette tip each time. Once you have added your DNA extract to the water you can mix it by pipetting up and down, or by stirring. Close each tube once it has been used, and place it away from the tubes that have not yet been diluted to avoid confusion.

Your DNA extracts are now diluted and are ready to be used for PCR!

Please note that the diluted DNA extracts are unbuffered and their DNA will degrade much more rapidly than the original extracts. If you need to repeat a PCR it may be best to prepare a new dilution from the original extract.