We do other things too: extracting DNA from cambium

This post is about labwork. Anyone reading our blog might have the idea that being in the Aitken lab is much like being Indiana Jones (or possibly Tarzan) – and sure, we are a good-looking bunch of rugged adventurers. But the truth is that although most lab pictures show our feats of courage ascending massive Sitka spruce, or wandering the wilds of Alaska, we spend a lot of time in the lab.

Joane Elleouet and I tried to extract DNA from dried Sitka spruce cambium for six months or so last year, on and off. We went through a number of protocols, and although we eventually modified one of them enough that it mostly worked, we were amazed at the lack of guidance online for cambium extractions. Surely someone had done this before! Nope. We thought that the world wide web had our back, and then it didn’t.

This is the wetlab, the room that hears the most cursing and cheering. We (mostly Dragana) recently cleaned, which is why it looks so good.

What you’ll find below is an annotated version of a CTAB DNA extraction protocol we got from Kristin Nurkowski (who got it from Marco Todesco at the Rieseberg Lab),  which we have modified for cambial tissue. It’s not perfect, but if you’re having trouble getting DNA out of cambium it may be your best bet. Probably some more improvements can be made. Note also that the suggestions we offer are just what one or both of us thought worked best for our own samples as we extracted them: we haven’t tested them with other batches of samples, and often we haven’t tested each alteration to the original protocol in isolation. But despite our recalcitrant tissue, we managed to scrape a 60-90% success rate, depending on the exact methods and samples (which is actually pretty good). Hopefully you can do as well or better.

When not in the lab or the field, this is what we do: Joane, Ian, Jon, and Pia being productive.

Unfortunately, for those of you who are just reading this for fun, recess is over once you cross the dashed line below.

————————————— Beginning of the protocol —————————————

The protocol has been through a number of modifications, but originally it was from Zeng et al. 2002, Acta Botanica Sinica 44; 694-697. There are a few other papers published that have other methods for extracting DNA from cambium (preserved in different ways) that you could look for in case of difficulty.

PART I: Preparing the buffers

CTAB-free Buffer (100 mL total volume):

20 mL 1M Tris-HCl stock, pH 8.0
10 mL 0.5 EDTA stock, pH 8.0
5 mL 5M NaCl stock
65 mL PCR-clean H2O

3% CTAB Buffer (100 mL total volume):

10 mL Tris-HCl stock
5 mL EDTA stock
30 mL NaCl stock
3 g CTAB
1 g PVP
55 mL PCR-clean H2O
Do not autoclave, the PVP will be destroyed.

High Salt TE (100 mL total volume):

20 mL NaCl stock
80 mL TE buffer

TE Buffer (100 mL total volume):

1 mL Tris-HCl stock, pH 8.0
200 uL EDTA stock, pH 8.0
98.8 mL PCR-clean H2O

PART II: Tissue collection

We used discs of cambium and phloem (since it’s difficult to get just cambium – it’s all stuck together in the inner bark) obtained using a leather punch from the base of Sitka spruce trees and dried in silica gel. We shaved around 20-30 mg (usually 22ish) of tissue from the inside (wood side) of each disc. That’s the region that should contain the (supposedly) DNA-rich cambial tissue.

Our samples weren’t perfect to begin with. Mine were collected in late July on Vancouver Island, after the peak of cambial activity in the spring. Collecting earlier might have improved the yield. Joane’s samples had been dried in silica gel for a year, and it’s possible they became less tractable with time.

Put your tissue into a 2 mL tube with one steel ball bearing. It is really important to weigh tissue into a 2 mL tube because the ball bearing can get stuck in the end of the 1.5 mL tube.

PART III: Grinding

1. Place your samples in a mixer mill block, put it in a Styrofoam box, and pour liquid nitrogen over it (wear insulated gloves!).
2. Assemble the block and fit it into the mixer mill.
3. Grind samples at 30 Hz for 1 minute.
4. Quickly disassemble the block and freeze each part again.
5. Reassemble the block, flipping it 180◦ to ensure even grinding, and grind again at 30 Hz for 1 minute. Usually, redistributing the samples within the block each time helps them grind evenly.
6. Disassemble the block and check each sample. It should look like a fine beige powder. If the sample is not ground properly, freeze everything and grind it again. For our samples, it took as many as ten repetitions of the grinding process.
7. After grinding, quickly add the prepared buffer and start the protocol.

PART IV: Protocol

8. Put some 3%CTAB to warm at 65◦F with loose cap
9. Add 1 ml of CTAB-free buffer (cold) + 6 μl β-mercaptoethanol (mix done beforehand in the fumehood with a bit extra)
10. Mix by inversion (shake each sample quickly), keep 10 minutes in the freezer. While waiting, put some 3% CTAB in the waterbath and some 2-propanol in the freezer
11. Spin at 10.000 g for 10 minutes in the centrifuge (longer may be required, depending on your tissue)
12. Discard the supernatant under the fumehood in a trash beaker
13. (if you use particularly nasty tissue, you can repeat this step)
14. Add 500 μl of pre-warmed 3% CTAB + 5 μl of β-mercaptoethanol in the fumehood
15. Incubate at 65°C for 60 minutes
16. Let the tubes cool down a little bit
17. Add 500 μl of chloroform-isoamylalcohol 24:1 in the fumehood. SUPER CORROSIVE!
18. Vortex
19. Spin at full speed for 15 minutes. Meanwhile, label new 1.5mL tubes
20. Move the aqueous phase to the new tubes (1.5mL) under the fumehood. This must be done exactly right – it’s probably the most important step. The aqueous phase is the liquid above the solid phase in the centre of the tube. Don’t take all the liquid, and don’t disturb the part that’s within a couple mm of the solid-liquid interface. Take around 320 uL from the top surface of the aqueous phase slowly and carefully. Too much will give you an impure sample. Too little will decrease the yield of DNA.
21. Add 250 μl of NaCl 5M, mix fast
22. Add 500 μl of cold isopropanol (2-propanol), mix (have a tube ready in the freezer)
23. Leave at least 20 minutes or overnight at -20°C. You can take a break here if you want!
24. Spin at max speed for 15 minutes
25. Wash pellet in fumehood with 1 ml of cold 80% ethanol (prepare the mix from 100% and PCR-clean water)
26. Spin at max speed for 5 minutes
27. Repeat wash (no need to be under the fumehood anymore)
28. Dry pellet (by putting the tubes in the heatblock at 25 to 37 degrees)

Depending on your tissue, you may need to include the following seven optional steps (labelled 29-35). Otherwise proceed directly to step 36. We found that omitting the optional steps often led to severe RNA contamination. The Nanodrop would say we had lots of very pure DNA, but the Qubit (which is much more reliable) would say we had almost none.

29. Resuspend in 400 μl of high salt TE + 2 μl of RNAse A (sometimes more RNAse is needed)
30. Incubate at 37°C for one hour, mixing occasionally until the pellet dissolves. Sometimes this is a LOT of mixing.
31. Add 800 μl of cold 100% ethanol
32. Spin at max speed for 15 minutes
33. Wash pellet with 1 ml of cold 80% ethanol. If the pellet is invisible, it doesn’t mean the sample failed. Just continue, leaving a bit of liquid at the bottom of the tube with every wash, and be very careful not to touch the part of the tube where the pellet should be when you suck out the liquid to dry the pellet.
34. Spin at max speed for 5 minutes
35. Repeat wash

36. Dry pellet and resuspend in 50 uL elution buffer or TE. Use less if you need higher concentrations. The yield of this procedure was often in the range of 250 – 1250 ng when the optional steps (29-35) were included, and more if they weren’t.
37. Quantification and quality control: we used a Nanodrop and a Qubit. The Nanodrop gives (sometimes wildly) inaccurate DNA concentration readings but it can test for purity. The Qubit gives accurate concentrations, but doesn’t tell you what might be in the sample along with the DNA.


It is important to note the two most hazardous reagents used in the protocol. In addition to being stinky, the B-mercaptoethanol is also toxic. Any supernatant or plastic item contaminated by it must remain in the fume hood and becomes hazardous waste. Do not breathe it in or get it on your skin. The chloroform is also toxic and can burn your skin and lungs. Keep the lid on and it in the fumehood! Any plasticware contaminated with chloroform needs to remain in the fumehood until dry, and then it can be thrown out with regular trash. Liquid waste needs to be stored and disposed of according to your local regulations (but not down the drain!).


The biggest problems with this protocol were:

• A mystery contaminant. There seems to be something in Sitka cambium or phloem that gives low 260/230 ratios on the Nanodrop (i.e. a bad quality score), if it isn’t properly removed. Sometimes it removes well, other times not.
• RNA contamination. We sometimes had lots of RNA in the samples if we skipped steps 29-35. If this isn’t a problem for you, omit them. If it is, include them – but the whole protocol gets more finicky when you do. Getting rid of the RNA while keeping enough DNA to work with (in our case for GBS and sequence capture and then next-gen sequencing) wasn’t always easy.
• Low yields. Sometimes we just didn’t get much DNA. The good news is, this often seemed to be protocol-dependent rather than sample-dependent. When a sample didn’t work out, trying the same sample again often solved the problem.

Some suggestions for troubleshooting:

• Try changing the amount of supernatant you take in step 20. Take more if you have no DNA. Take less if you have a contamination problem.
• Try starting with different amounts of tissue. Use more if you have no DNA. Use less if you have a contamination problem.
• Be careful washing the pellet: don’t lose all or part of it! Maybe spinning at colder temperatures (4°C? 10°C?) would help.
• Consider using RNAse A if you still have a contamination problem after including the optional steps. Consider eliminating the RNAse A steps if you have no DNA.
• Try using your newest tissue, collected whenever cambium is active near you (probably spring).

Final note:

All the extra cautions and advice I added to the protocol might make it seem a bit daunting. But remember, it actually works! The protocol isn’t particularly difficult, and maybe it’s less finicky than my comments make it seem — and after a few repetitions you should be churning out extracted samples no problem. So good luck!


Lodgepole pine DNA extractions

If you’ve read any of my previous posts, you may be aware that I have a truck-load of lodgepole pine samples collected last summer that are destined for SNP genotyping in the near future. I am close to wrapping up all of my DNA extractions (fingers crossed) for sending out, and because I tinkered a bit with the standard extraction protocol, I thought I’d discuss it here. Before I get there though, I have to give huge, gigantic, endless thanks to Kristin Nurkowski and Robin Mellway for all of their help. Most of these adjustments are a direct result of their insight and suggestions. They also taught me the standard protocol, how to use the lab’s robot, and saw me through lots and lots of troubleshooting.

To start at the beginning, I have needle samples collected on silica gel. In the field, my assistants and I tried to take the freshest tissue possible (new needle growth at branch ends), which was then kept on silica gel in coin envelopes within sealed ziplocs. When it was not possible to collect new needles, older ones were taken, and we even sampled some clearly dead trees (brown needles) in the off chance they’d work and some dying trees (recently fallen tree, needles not yet brown and brittle, but clearly dead); I’ll discuss this again a bit later. My collections in the Yukon were much later in the summer, so new growth was not new anymore, and we ran into a couple rainy days here which is a real pain when storing on silica. Other notable points were that these samples then sat in tupperwares in the back of a truck for at most 2 weeks while we continued collecting, so they definitely experienced some warm temperatures, but I have seen no correlation in the time since collection and the quality of DNA.

For the standard extraction, we used Machery-Nagel kits with NucleoSpin filters. These come with two different protocols using either PL1 lysis buffer or PL2 lysis buffer. From an initial test of both protocols and various weights of tissues, I found for my samples that the PL2 protocol worked better. PL1 is based on a CTAB extraction while PL2 is SDS based and has an additional step of a protein precipitation using potassium acetate (PL3). 20mg of tissue (dried) also seemed to be best, with less not giving enough DNA, and more also decreasing in yield. Presumably one could increase the amounts of buffers and reagents used per reaction relative to increase in tissue if more tissue wanted to be used, but at these volumes the filters were filled quite high with supernatant at the binding step, so eventually there is a limit. Except where I have pointed out exceptions, I followed the protocol outlined in the manual and used the volumes given, and just upped my lysis incubation to 45 minutes at 65C and my protein precipitation to 20 minutes at -20C (on ice, and in the freezer).

Initially I stuck with this protocol and it worked well.  I had about 80% success per plate, which I was pleased with considering the quality of some of my tissue samples. And believe it or not, some of the dead needles we had collected yielded DNA with enough to meet the cutoff! After completing eight plates of extractions, we had some issues crop up with our robot, and then extractions after that started failing. It is unclear if it was a product of the robot or a bad set of reagents or something entirely unknown, but it was clear nothing in the protocol had changed. It was also clear that it was not alone due to the quality of samples being used as some were trees from the same sites and provenances as samples that had already worked successfully. I did a lot of troubleshooting at this point to try to get things back up and running and eventually we found that adding PVPP to the PL2 buffer got us back up to an 80% success rate.

I used a 1% PVPP buffer for a while and then increased to 2% which improved it further. Because PVPP is mainly insoluble in liquid, I didn’t see the point in increasing past this as it was clear that there was undissolved PVPP in my buffer. I added 1 gram of PVPP to 50 mL of the PL2 buffer (preheated at 65C) plus 10 ul of antifoaming agent since the PL2 is quite bubbly, and incubated this overnight at 65C with 1 mL of RNaseA being added just before starting the extraction. A shaking incubator might be nice to use here since in the water bath, a lot of the PVPP just settles at the bottom of the tube. Using the PVPP improved my 260/230 from about 1.75 on average before to 2.33 on average.

Additionally, it seemed that the second plate from each extraction was on average coming out with poorer quality — the robot can do two plates at once. The robot is not incredibly fast at pipetting, so with a second plate, it sits there a little longer than the first before it is acted upon. This can be helped in the beginning by not removing the second plate of ground tissue from the freezer until just when the robot is ready to add lysis buffer to it. Because I did not have a very large number of plates to do, I instead began single plate extractions and simply kept an old used plate, filter, and elution plate to balance the centrifuge at the necessary steps. It is worth pointing out here that this was not the most efficient route. Though I have not raced the robot, if I had had access to a multichannel pipette, it is clear that doing this protocol sans robot would have been faster and easier. The robot is by no means a requirement for this extraction protocol, but it does have the added benefit of freeing you up to do other work while it pipettes away. I have even heard that newer models of robots have arms that will move the plates around for you! And the robot will definitely be a huge help for the plate normalization step once I have all my samples.

Some final points to make for anyone following along: from nanodropping my samples and qubiting a subset of them, most had about 40% less DNA per ul than was shown by the nanodrop, and I found no significant correlation of this with either my 260/230 or 260/280, though others in the lab have found that to be related for samples extracted from fresh, frozen tissue under the PL1 protocol, and with less of a decrease in DNA quantity between the two.

For samples that didn’t meet my cutoff criteria in terms of quality and quantity, performing the same extraction protocol a second time was successful about 60% of the time. I would speculate that the first failure in these cases was due to either the robot picking up some of the pellet with the supernatant which happened on rare occasions, or due to differences between different needles collected from the same tree.

And lastly, for those samples that did not succeed the first or second time around, it is now time to perform the more labor-intensive but hopefully cleaner and higher-yielding CTAB extraction protocol! Most of my samples that fall into this category are those I collected from the Yukon, so a product of the tissue being older and additionally not drying as quickly from the rainy day extractions. So for anyone out there planning a field season, I definitely advise lots and lots of extra silica gel and the youngest tissue possible — two already-well-known points that cannot be emphasized enough. If I have any serious alterations to the CTAB protocol, I may update this post or add another, but until then I hope that some of this information might be useful to anyone else out there working with conifers and their pesky secondary compounds!