DNA Extraction using the Dipstick DNA Extraction Kit

Protocol

In this protocol, you will extract a tiny amount of DNA from an animal, fungus, or plant, direct to PCR, using the dipstick DNA extraction method. This should take between 30–60 seconds per sample once you are prepared, excluding a break in the middle of the process to set up your PCR.

Preparing your extraction workflow

Prepare your sample list

In a notebook, lab book, or spreadsheet, make a table listing your samples:

• Include any identifying information needed to match the sample to the extraction tube to the PCR in the future. For example, you could include sample code, species name, and date collected.
• Write down the date of extraction on the table.
• Give each sample an extraction number (e.g. 1–8) in a second column.

A simple numbering system can make it easier to label tubes clearly, and track from samples to extractions to PCR products to results.

Label your extraction and wash tubes

You will use two 1.5 mL centrifuge tubes for each sample: one for crude DNA extraction, and one for washing the dipstick.

Label each extraction tubes on the lid and side using a permanent marker pen. Give each extraction tube an extraction number based on your table (e.g. 1–8).

Label all the wash tubes (one per sample), e.g. with a W on the top. These will be used and then discarded, so they don’t need sample-specific labels.

It’s good practice to label the tubes clearly even if you only have one sample.

Add Extraction Buffer to the extraction tubes

Using a new pipette tip, add 200 μL of extraction buffer into the numbered extraction tubes, resulting in one filled extraction tube per sample.

The amount of extraction buffer can be reduced to concentrate the DNA for smaller samples (e.g. as low as 40 μL), and to dilute the DNA for larger samples (e.g. as much as 700 μL). You can also use larger volumes with smaller samples, but very small samples can be difficult to grind in larger volumes of extraction buffer.

Alternatively, you could put your samples in tubes before you fill the extraction tubes with Dipstick Extraction Buffer. This can be useful if you have a large number of samples or if you want to prepare the samples in advance of the extraction.

However, this will mean that the tubes will no longer be clean, and you’ll need to be careful to avoid cross-contamination when filling the extraction tubes.

You can avoid cross-contamination by carefully squirting the extraction buffer into the tubes without touching the tube with the tip. If you touch the tubes, discard the tip and fit a new one before continuing. Alternatively, use a new tip for each tube throughout.

Add Wash Buffer to the wash tubes

Using a new pipette tip, add 700 μL–1 mL of Wash Buffer into labelled wash tubes, making one wash tube per sample. The amount of wash buffer needs to be enough to wash all of the dipstick handle that is exposed to the extraction buffer during extraction.

Prepare your tissue samples

Using a clean sharp cutting implement (a scalpel or half of a double-edged razor blade), and a clean surface (e.g. a clean piece of paper towel), cut a small piece of tissue for extraction.

The piece of tissue must be large enough to grind efficiently using a micropestle. A piece of around 2 mm²–3 mm³ (if flat) or 1 mm³–2 mm³ is usually a good size to use.

Then transfer the small piece of tissue to the extraction tube. You can either flick it into the tube using the blade you used to cut, or use clean tweezers.

Repeat this process for all of your samples.

You now have a small piece of each of your samples in Dipstick Extraction Buffer, ready to be extracted.

Prepare your PCR reagents

Dipstick DNA extractions transfer DNA directly from the wash step of the extraction to a prepared PCR mix, so you will need to have your PCR mix tubes prepared before you complete the final steps of the extraction process.

In this step, you should ensure your PCR reagents are all present, defrosted, and that every reagent tube is well mixed so they are ready to use later.

You should also label your PCR tubes (e.g. 1–10) on the top and sides of the tubes.

Optionally, you can choose to set up your PCR mix tubes at this stage (instead of in Extracting DNA, Step 3 below), particularly if you’re working with only a few samples, setting up on ice, or using a hot start PCR master mix.

However, if you’re processing many samples then there can be a long delay between the PCR mix being made up and the PCR being run. This can increase the risk of producing non-specific amplification, especially if you are setting up at room temperature using a standard (not hot start) PCR master mix.

Setting up the PCR only once all the samples have been ground into crude extractions can minimise the time between the PCR mix being made up and the PCR being run, and thereby reduce the risks of non-specific amplification.

Extracting DNA

Grind the tissue in the extraction buffer

The objective of this step is to break up the cells in your tissue sample and release the DNA into the solution.

To do this, use the micropestle to crush the sample between the side of the pestle and the tube wall. You can rotate the micropestle when firmly inserted to squash and smear the sample. You can also wiggle the micropestle rapidly when it’s more loosely inserted to apply more force.

Try to avoid pressing the tissue into the bottom of the tube because it may become compacted in the tip of the tube and fail to break apart. If it does get very stuck you can use a pipette tip to dislodge it.

If some pieces of tissue can’t be ground, don’t worry. You should have released enough DNA from the rest of the sample.

The solution should become a hazy mixture of cellular debris, with few or no large solid lumps unless the tissue is very fibrous.

Repeat this step for all of your samples, using a clean micropestle each time.

You now have a crude DNA extract for each of your samples.

To clean plastic or glass micropestles, you can soak them in diluted bleach solution (around a 1:10 dilution of thin domestic bleach) with a drop of detergent, for around 10 minutes, rinse off the bleach with tap water, and dry them with a clean paper towel.

Dilute your crude extract with more extraction buffer

At this point your crude extracts may be quite concentrated. For samples with high concentrations of PCR inhibitors, you may get better results by diluting them with more extraction buffer.

As a general rule of thumb, provided your sample was fresh and around 2 mm³, then it can be useful to dilute the extraction to around 500 μL. If your extractions are still extremely cloudy, are strongly pigmented, or have failed to amplify in a previous attempt, then you could try diluting even more.

Ultimately the best amount of dilution will depend on your sample type and the amount of tissue you extracted from, so some trial and experimentation can be useful when working with new types of samples.

The samples can be left at room temperature while you start the next steps.

For some sample types, leaving the crude extract at room temperature for a time might increase the extraction efficiency.

Prepare your PCR mix

It can be a good idea to pause the extraction at this point and set up your PCR mix, although you can also set up your PCR before you start the extractions.

The PCR mix should be the same as you would set up for any other extraction method. If you are following a Bento Lab protocol, it should contain, per reaction:

• 4 μL of 5x Master Mix
• 2 μL of primer mix
• 12 μL of PCR grade water
• 2 μL of wash buffer and DNA on the dipstick

To make a batch mix you can multiply the volume of each component by the number of PCRs required (including controls). It’s often a good idea to add an extra 10% excess of everything to account for pipetting inaccuracies. You can then split the batch mix into the required number of PCR tubes at 18 μL per PCR. The table below shows suggested volumes for different numbers of PCRs.

Components of PCR Mix	1 rxn	10 rxns (+10%)	24 rxns (+10%)	32 rxns (+10%)
5x PCR Master Mix (μL)	4	44	106	141
Primer mix (μL)	2	22	53	70
PCR grade water (μL)	12	132	317	422

If you are adjusting primer concentrations, or adding additional components such as magnesium chloride, you can increase or reduce the amount of PCR grade water to make up the PCR to a final volume of 20 μL (18 μL excluding the 2 μL added and then removed by the dipstick).

Please check your specific PCR protocol for the exact volumes recommended for your application.

Capture DNA onto the dipstick

Take a dipstick out of the bottle, and dip it in the extraction buffer three or more times. Three dips is usually sufficient, but some users have suggested more can work better.

When removing the dipstick, if there is a drip on the end, wipe this off on the inside lip of the tube to minimise carryover of the extract into the wash buffer.

Wash the dipstick into the wash buffer

Dip the same dipstick in the wash buffer tube three times to remove any liquid on the outside, dipping it all the way to the bottom of the tube and ensuring that the handle of the dipstick is fully washed.

When removing the dipstick, wipe off any drips on the inside lip of the tube to avoid carrying extra liquid and potentially PCR inhibitors into the PCR mix.

Once finished, make sure to discard the 1.5 mL tube of wash buffer that you just used.

Dip the washed dipstick into the PCR mix

Dip the dipstick in the PCR mix three times to release enough clean DNA to act as DNA template for the PCR.

Then close the PCR tube, and discard the dipstick.

Repeat this process for all of your samples, using the appropriate PCR mix tubes.

Your PCR reactions are now ready to run!

Run the PCR program

Your PCRs are now ready to run. You should place the PCR tubes in your thermocycler, make sure all the lids are firmly closed, and run the appropriate PCR program immediately.

Storage of crude DNA extracts

You can store your crude DNA extracts at room temperature in the very short term, for example a few hours until the PCR result is confirmed. This can allow you to repeat the PCR if it failed (e.g. if it was too concentrated) or if you want to examine different DNA regions using the same extract.

You can also store the crude extract tubes in the fridge for a longer period. If you do freeze the tubes, please note that precipitation of the detergent in the extraction buffer may occur, and you may need to heat the extracts in very hot water (e..g for 10 minutes or so) until any precipitates are dissolved, before you can successfully extract from the tubes again.