My apologies for a (very) long absence from this site – I’ve been at my new job for about six months, so I guess I can’t use the new job excuse anymore, but I am at the tail end of about a month of travel. I was in Ireland for a week, which was beautiful (and relatively affordable):
Then I was at Cornell for a week-long training hosted by Ed Buckler’s lab:
(sadly this gorge is not where the actual training took place). Unfortunately I got a cold while in Ithaca, and I’m still not quite over it – if you were also at the training and ended up getting sick, I’m sorry! Anyways, that’s enough about me – if you’re here you’re probably looking for info on DNA extractions.
In a previous post I presented a simple protocol for collecting plant tissue for DNA extraction in a 96-well plate format. Then we went over how to grind the tissue. The next step is of course performing the extraction itself. There are many different considerations to make when choosing a protocol for DNA extraction. Many labs have moved towards kit-based extractions, and for good reason: kit-based protocols offer good reproducibility, require minimal reagent preparation, and minimize cross-contamination. For labs that are working with large numbers of samples, automated platforms can perform thousands of extractions per day while limiting consumable use. Finally, send-away extraction services, where a researcher mails out tissue samples and receives DNA a few weeks later, are becoming increasingly feasible.
Of course, not every lab can pay for a robotic liquid-handling system or mail-order extraction services. With that in mind, I’ll be outlining the method that my lab in graduate school used for performing DNA extractions in a 96-well format. This method is relatively straightforward, and doesn’t require any specialized equipment. It is a modification of a method using cetyl trimethylammonium bromide (CTAB) for cell lysis and chloroform for phase separation .
Warnings and Disclaimers
Sorry – before going any further, I have mention these:
This protocol has a few different options for separating aqueous and organic phases after cell lysis, but they all involve chloroform. Chloroform needs to be respected, but is not especially dangerous on its own. Yes, it acts as an anesthetic when inhaled. Whenever you work with chloroform, you need to be working in a fume hood, and using all necessary personal protective equipment (PPE), including gloves, lab-coat, and goggles. You also will need a waste stream dedicated to used reagents and materials containing chloroform – which is essentially all of the reagents/materials used in the extraction. Keep in mind that chloroform will dissolve many polymers – including nitrile gloves. Therefore, it’s a good idea to change gloves if you splash chloroform on them.
Optionally, you can use a phenol/chloroform mixture for phase separation. Phenol can cause deep chemical burns upon contact with skin or mucous membranes. This is made more dangerous when combined with chloroform’s ability to dissolve nitrile gloves. If you are working with phenol/chloroform mixtures, you should take all the precautions of working with chloroform, but you should also wear a liquid-proof lab coat or apron, and chemical-resistant gloves such as those made of Silver Shield or Viton.
The method that is described here can be prone to cross-contamination, due to the very close proximity of sample tubes to one another. Good technique can minimize this risk, but not eliminate it entirely. The exposure to this risk that you can tolerate will greatly depend upon the downstream application of the isolated DNA. If you will be performing an assay to determine the presence/absence of some genomic feature, then you may have no tolerance for any cross-contamination. On the other hand, an assay for discrimination between different alleles may be less exacting. It may be best to test the influence of cross-contamination empirically, for instance by performing an extraction using a plate of two alternating known genotypes, and then using the DNA in whatever assay will ultimately be performed.
In addition, just like in the protocol on sampling tissue for extraction, I present some catalog numbers for products below. I am not paid by these companies; these are just products that I have successfully used in the past (sometimes after trial and error). Other companies/suppliers may market similar products that are just as effective.
DNA Extraction Protocol
With all of that out of the way, we can go on to the protocol itself:
- 1.2ml microtiter strip-tubes (Corning #4413)
- Strip-caps for 1.2ml tubes above (Corning #4418)
- 96 well deep-well plates (ISC Bioexpress Cat. # P-9641-1)
- Reagent reservoirs for 8-channel multichannel pipets
- 8-channel 1000ul or 1200ul pipet with corresponding tips
- CTAB Extraction Buffer (recipe for 1 liter is given at the end of the protocol):
- 24:1 chloroform:octanol mixture, OR
- 25:24:1 phenol:chloroform:isoamyl alcohol mixture (this should be pH 8.0 for DNA extractions, i.e. Acros Organics #327115000)
- Pure isopropanol
- 75% ethanol (NOT denatured)
- RNase A powder suspended to 10 ng/μl in molecular-grade water (Sigma #R6513)
- Spex SamplePrep Genogrinder 2010
- Tabletop centrifuge with rotor for 96/384 well plates
- Rocking Platform (e.g. VWR # 12620-906)
- Small Laboratory Oven or Water Bath set at 60°C – 65°C
- (optional) laboratory lyophilizer or vacuum chamber
Step 0: Labeling Tubes and Plates
Sorry – I’m very fond of including step zero in my protocols. Anyways, the first thing we need to do is some labeling and organizing. First, we need to label some tube strips:
- If you have an odd number of tube strips in total, you’ll need to add an additional dummy strip (with ball bearings) – this is to properly balance everything when centrifuging. Label this tube strip something like “D” or “X”
- Create a new set of tube strips that are labeled exactly the same as the set of tubes holding the tissue (including the dummy strip if present). Remember that in the previous post we labeled all tube strips with unique identifiers. If you have a dummy tube strip, then this set of tubes also gets a dummy strip.
- Use a multichannel pipet to fill the new set of tubes with 450µL of isopropanol
- [optional] Place the new set of tubes in a -20° C freezer for the time being (with the caps off)
Now we need to label a new set of deep 96-well plates, which will really just be used as heavy-duty racks to hold the tubes. These racks will be used for centrifuging, so label them C-1, C-2, C-3, etc. Then place the centrifuging racks next to the centrifuge, where they will stay. Any tubes that will be spun will get transferred into these racks prior to going into the centrifuge. Remember from the last post that the most important thing is that RACKS THAT WERE USED FOR TISSUE GRINDING NEVER GO INTO THE CENTRIFUGE.
Step 1: Cell Lysis
After following the steps in the previous post on tissue grinding, we should have tubes containing a nice pale green powder. Now we are ready to begin the extraction procedure:
- Add suspended RNase A to CTAB buffer in the ratio of 450µL buffer:0.75µL RNase for each sample. Be sure to make plenty of extra (e.g. multiply the number of total samples by 1.15) due to inefficiencies of using a reagent reservoir for the multichannel pipet. Mix CTAB/RNase solution by inverting several or using a stir bar, depending on the container that is chosen
- Add 450µL of CTAB/RNase mixture to each tube using a multichannel pipet
- Place the tubes into a 60° C water bath or oven with CAPS OFF (let the tubes get up to temperature before capping them – otherwise the caps will all pop off). I like to place the tubes in a water bath for about 10 minutes, take them out, cap them, and then transfer them to a 60° C oven so they dry off.
- Incubate tubes at 60° C for one hour, inverting them several times every 15 minutes
- Transfer tubes to centrifuge racks and give them a quick spin. After the spin, transfer the tubes back to the cheap racks they came in for the next step.
Step 2: Separation
In a fume hood:
- Uncap tubes and add 450µl of chloroform/octanol or phenol/chloroform/isoamyl alcohol mixture to each well
- Recap tubes and mix by inverting several times.
- Place tubes in racks, place a folded paper towel under rack lid (the chloroform-CTAB buffer mixture has much less surface tension than water, and a small amount can leak out around the tube caps), and secure rack lid with rubber band
- Place racks on rocker for 30mins. to 1 hour
- Check paper towel for chloroform leaks. Dry any leaked chloroform off of the outside of tubes with paper towel or kimwipe.
- Place tubes into centrifuge racks, DOUBLE-CHECK RACKS ARE BALANCED, centrifuge plate for 30 min. at 4,000 rpm at room. temp. (20°C – 25°C)
Step 3: Remove supernatant
This is the trickiest part of the extraction. After you remove the tubes from the centrifuge, they should exhibit a nice phase separation, with the greenish/brown/black chloroform and organic “junk” on the bottom, and the clear (or slightly pink/reddish if using phenol) aqueous phase containing the DNA on top. The two phases are usually separated by a somewhat hardened skin.
Now you need to carefully remove the top phase, and place it into the new set of tubes containing isopropanol. Note that you need a clean pipet tip for each sample, so this step can potentially go through a lot of tips. If you have a full-plate pipetting station, you can do this step for an entire plate at once. Otherwise, you’ll need to use the multichannel pipet and go one tube strip at a time.
When you are actually pipetting, move quickly – the aqueous phase can slowly leak out of pipet tips after it is sucked up, so you don’t want any drips falling into the wrong tubes when you are transferring. Otherwise, take your time – it is easy to get mixed up here. It’s easiest if the layout of the new set of tubes containing the isopropanol exactly mirrors the layout of the old set of tubes. Also, you can physically move the old set of tubes to some other rack as you remove the supernatant from each one.
Step 4: Precipitation
Once the aqueous phase is transferred into the new tubes containing isopropanol, cap them and then invert them several times. Hopefully in a few tubes you’ll see some white clumps of DNA floating around. Don’t worry if you can’t see the DNA in all the tubes – just trust that it’s in there. Alternatively, if you see big globs of DNA in every tube, it probably means that you harvested too much leaf tissue.
After a few inversions, place the tubes into the centrifuge racks and then centrifuge them for 30 minutes at 4,000 rpm at 4° C.
Step 5: Rinsing
After the tubes are done centrifuging, uncap them and gently pour the isopropanol out into a beaker or other waste container. The DNA should be firmly stuck to the bottom of the tube in a pellet, so you shouldn’t have to worry about throwing the baby out with the bath water.
Once all the isopropanol is drained out, add 450 μL of 70% ethanol to each tube and recap. Invert the tubes several times (the DNA pellets should now be a brighter white color, and may detach from the bottom of the tube). Then place the tubes into their centrifuge racks, and centrifuge them for 30 minutes at 4,000 rpm and 4° C.
If you suspect that your DNA is particularly dirty for any reason, you can repeat the whole rinse step a second time.
Step 6: Drying & Resuspension
After centrifuging the DNA in ethanol, uncap the tubes and gently pour the ethanol out into your waste container. You may want to also dab the ends of the tubes on a paper towel while they are upside-down to remove some extra ethanol.
One problem with using such narrow tubes is that a fair amount of residual ethanol will remain after decanting – even with the best pouring and dabbing technique. My recommendation is to place all the tubes in a vacuum chamber or lyophilizer (without the refrigeration enabled), which should evaporate any remaining ethanol in a few minutes. If you don’t have this equipment available, the tubes can be air-dried for several hours. DNA is relatively resistant to short-term thermal degradation, so you can even dry tubes in an oven set to 40° C for a short period, if you get really desperate.
After drying, just use a multichannel pipet or pipetting station to add 100 μL of water, Tris buffer, or Tris-EDTA buffer to each tube. Your DNA extraction is finished; cap the tubes, place them in the fridge overnight, and the next day you should have resuspended DNA ready for quantification.
Recipe for 1 liter of CTAB DNA extraction buffer
|1M Tris buffer (pH 8.0)||100mL|
|0.25M EDTA buffer (ph 8.0)||80mL|
|NaCl powder||82 grams|
|CTAB powder||20 grams|
|ddH2O||remainder of 1L (~820 mL)|
Combine all ingredients and mix over moderate heat using a stir bar until CTAB powder goes into solution (stir at a slow pace to avoid foaming).
I won’t go into the details of making the Tris or EDTA buffers here – these are also sold ready-to-use in the required molarity/pH by several vendors.