Bead cleanups

This post, which discussed results published by Rohland and Reich (2012), has been removed at the request of Beckman Coulter legal counsel.

DNA precipitations

Haven't posted in a while, so this will be quick.

People ask me about their protocols on a pretty regular basis.  Even if they don't ask specifically, I often ask to see their workflow so I can tell if I am giving reasonable advice in the context of their whole experiment.  One of the most common things I encounter is the insistence on putting samples in the freezer to "help" with precipitating their precious DNA.  However, this idea should have been laid to rest almost 30 years ago now with the publication of "Ethanol Precipitation of DNA" in Focus (Fall 1985, Vol 7, No 4) by Zeugin and Hartley.

The key findings of this paper are that precipitation is less efficient at low temperatures than worm temperatures, length of incubation time has minimal effect except for very dilute samples, and centrifugation time is the most important factor in planning your precipitation.  The precipitations in this paper were NaOAc/Ethanol precipitations with the final NaOAc concentration being 0.3M (1/10 vol 3M NaOAc) and the final EtOH concentration being 75% (~2.5 vols EtOH).

The authors concluded that cold incubation is not beneficial to the precipitation of DNA and can even be counterproductive if your sample contains only a small quantity of DNA.  They speculate that decreased temperature will increase the viscosity of the solution and inhibit the motion of DNA through the ethanol solution during centrifugation.  I agree with this idea, but I would also speculate myself that by removing energy from the system, you simply slow down the kinetics of precipitation where free cations associate with the ribo-phosphate backbone while ethanol drives water molecules out of the double helix structure resulting in precipitation of DNA in a salt form.  Certainly during centrifugation such interactions will increase, so lately my thinking has been that centrifugation is the only thing that matters much during your precipitation.  This is also supported by their data where they looked at length of centrifugation time on precipitation.  Again, high concentration samples precipitated readily while low concentration samples needed more time, but most concentrations seemed to taper toward maximum recovery at 30 min, which unfortunately, was the longest centrifugation time they tested.

For me, this has resulted in the following being my standard precipitation conditions.  I prefer NaCl over NaOAc because I have some loose data indicating better recovery (not shown).  I continue to use NaOAc during EtOH precips of cycle sequencing reactions.

1) Add NaCl to 0.15M.  No need to be exact here.  Use a 5M NaCl stock and you won't need to add much so the volume compensation is unnecessary.  For instance, if you have a 300uL sample, calculate as follows:  (300uL*0.15M)/5M = 9uL.  So just add 9uL 5M NaCL and save yourself the headache.  It's not worth it!!

2) Add 2.5 vols EtOH.  Many protocols say add 2-2.5 vols EtOH.  I think this is problematic since if you use 2 vols, your final EtOH concentration will be only 66% whereas at 2.5 vols you achieve about 71%.  This may seem subtle, but my experience (admittedly anecdotal here) has taught me 2.5 is always better.

3) Mix sample well, and place directly into the centrifuge.  For samples with decent concentration, 30 min at maximum speed.  For low concentrations, longer is better, so maybe 30 min to 60 min (your choice).

4) Immediately after the centrifuge stops, remove the tubes and decant into the sink (plates can be GENTLY centrifuged inverted on a paper towel at low RPM, low acceleration/deceleration for ~10 sec to remove supernatant as per ABI sequencing protocol).  If you happen to be out of the room when your centrifuge stops, start it again for 5 min or so to ensure your DNA is well-adhered to the wall of your tube.

5) Add 70% EtOH.  For 1.5mL tubes I usually do 500uL, for 96 well plates, I usually add 50uL.  Spin again at max speed, but only 10-15 min is necessary now.

7) Decant again as in step 4.  Dry samples down in a vacuum centrifuge, or place on a heat block (~55C) for 10 min to evaporate any residual ethanol.  If you have left over EtOH and you try to run a gel, your sample will maddeningly just float away.  It can also inhibit downstream enzymatic steps.

8) Resuspend in your favorite solution.  Some people like water, but since the pH of "pure" water is often a bit acidic, depurination is a real threat to your sample.  Some people like TE, but even small amounts of EDTA can inhibit a PCR reaction.  For these reasons my go to solution is Tris-Cl pH 8.5, 10mM.

So, I hope someone finds this useful.  Happy precipitating!!

PCR: not so much on Tuesdays

It's been too long since my last post, but I've been busy.  The semester is over, written and oral comps passed, and I can return to being productive with some tinkering on the side.  I've recently been beset by updating some equipment usage logs which isn't hard, but takes up time I'd rather be using to do other things.

When looking at thermal cycler usage for the spring 2012 semester, one interesting thing pops out: Tuesday instrument usage is markedly lower (~40%) than other weekdays.

The numbers are hours used per day of the week from January through May.

What can account for this?  Impulsively, I think a couple of things contribute.  First, people come to lab on Monday with ambitions for the week and maybe some fresh thoughts contrived over the weekend about some result or another.  Once these results are obtained, perhaps some other steps or reflection on these new results are necessary before proceeding, hence the dip on Tuesday.  I think people then "push" to get the rest of their planned work done across the rest of the week, thus maintaining regular instrument usage, even on Fridays.  A few of us obviously spent some fun weekends in lab, too.  Someone I showed this to suggested that perhaps class schedules impede research on Tuesdays, but since Thursdays are well utilized, I think that seems unlikely, given the Tues/Thurs or M/W/F scheduling of classes here at NAU.

So there it is.  A mostly pointless blog post to help me feel better about not putting anything up for a while.  Maybe something more substantive next time...

tailed 454 amplicon sequencing fail

Our lab has been sending samples off to a lab out of state for 454 sequencing of amplicon pools for almost a year now.  We have so far just used this to assess fungal diversity (ITS region), but in theory could come up with all sorts of applications.  Other labs at NAU have been doing the same thing, but look at various targets to assess various questions of diversity (bacterial, archaeal, fungal, functional gene diversity etc).  Another tool I routinely use is tailed fluorescent primers for labeling microsatellites I amplify from my field studies (seeSchuelke, M. (2000). An economic method for the fluorescent labeling of PCR fragments. Nature Biotechnology, 18, 233-234).  Not too long ago, I was thinking tailed primers would be useful for amplifying samples for 454 samples, but it is very expensive to just test this idea out.  Instead, I used it successfully for doing tRFLPs for QC prior to sending samples off for 454 sequencing.  Later I realized this was discussed in the GS Junior Guide for Experimental Design, though some details and crucially, a discussion of the data produced, was missing.

So I was excited to see a couple of recent publications:

Bybee, S. M., Bracken-Grissom, H., Haynes, B. D., Hermansen, R. a, Byers, R. L., Clement, M. J., Udall, J. a, et al. (2011). Targeted amplicon sequencing (TAS): A scalable next-gen approach to multi-locus, multi-taxa phylogenetics. Genome biology and evolution, 3, 1312-1323.

Daigle, D., Simen, B., & Pochart, P. (2011). High-Throughput Sequencing of PCR Products Tagged with Universal Primers Using 454 Life Sciences Systems. Current Protocols in Molecular Biology, 96, 7.5.1-7.5.14.

The authors carefully go through the steps involved in using tailed primers and detail their results.  So in order to maximize the monetary power of all labs doing this here at NAU, I proposed to try using a set of tailed primer to barcode a sample for 454 amplicon sequencing.  I did not spend a lot of time, but things did not go as smoothly as I had hoped.

Here are the primers I used:
M13f-ITS1F 5'-CGCCAGGGTTTTCCCAGTCACGACCTTGGTCATTTAGAGGAAGTAA-3'
M13r-ITS4 5'-TCACACAGGAAACAGCTATGACTCCTCCGCTTATTGATATGC-3'
LibAf-MID01-M13f 5'-CGTATCGCCTCCCTCGCGCCATCAGACGAGTGCGTCGCCAGGGTTTTCCCAGTCACGAC-3'
LibAr-MID01-M13r
5'-CTATGCGCCTTGCCAGCCCGCTCAGACGAGTGCGTTCACACAGGAAACAGCTATGAC-3'
LibAf 5'-CGTATCGCCTCCCTCGCGCCATCAG-3'
LibAr 5'-CTATGCGCCTTGCCAGCCCGCTCAG-3'

The M13-ITS primers have the ITS primer 3' and the M13 sequence 5'.  The LibA-MID-M13 primers have LibA sequence 5', MID01 in the middle, and M13 sequence 3'.

First amplification (20uL reactions, 0.01U/uL Phusion polymerase, MgCl2 at 2.5mM, M13-ITS primers at 200nM; 30 cycles of 90C 30s, 57C 30s, 72C 1min)

There are 4 samples in the gel (2uL/well).  First four wells are each sample at 1X DNA concentration, 2nd four DNA at 1/20X.  Ladder is (bp) 2000, 800, 400, 200, 100.  Note the significant artifact around 100bp.

I column purified the 1/20X products with Epoch Life Science columns (http://www.epochlifescience.com/Product/SpinColumn/minispin.aspx) and quantified with OD260.  All purified products were around 40 ng/uL.

Here is a gel of the column purified products (10uL/well):

So I co-purified my 100bp artifact, but I continued nonetheless.  To try to reduce the amplification of the non-specific product, I added 2.5uL each purified product to 1000uL 10mM Tris-Cl pH 8.8.  I amplified under same conditions both the raw column purified product and the diluted product using LibA-MID01-M13 primers this time:

First 4 samples are using full-strength column purified DNA, second 4 samples are using the dilution.  The ITS product looks really weak now, and the 100bp artifact has dominated my PCR.  No bueno!

I reran the raw M13-ITS products on a gel and cut out the ITS bands this time.  I was generous in the size of gel slices in case there were products I couldn't see.  Briefly, I put the gel slice in a 1.5mL tube, added 3 2mm steel beads, and beat the gel in the GenoGrinder for 15s at about 28hz.  I spun the gel down briefly, added 200uL Tris-Cl, vortexed, and placed in 65C block for 5 min.  I spun 1 min at 15,000rpm, and pipeted off 100uL of supernatant.  Gel-purified products extruded!

I then redid the LibA tagging using the LibA-MID01-M13 primers on my gel-purified samples (full strength only):

Each 2 wells are 8uL column purified M13-ITS product, then 2uL LibA-tagged product from the last PCR round.  You can see the gel purification made a huge difference in maintaining specificity.  You can also see the subtle gel shift as the LibA tagging incorporates another 50bp or so.  Still, some unwated product remains, particularly now at about 180bp.

So I thought I could change PCR conditions and improve things.  I cut out the excess MgCl2 so that it was back to 1.5mM and used the following PCR conditions: 30 cycles of 90C 30s, 63C 2 min, 72C 15s.  Intial PCR (M13-ITS primers) yielded OK results:

The top comb is from another project, but the bottom comb are the same four samples in a PCR serial dilution (2uL/well).  First 4 samples 1X DNA; 2nd 4, 1/10X DNA; 3rd 4, 1/100X DNA.  I thought perhaps the column cleanup was unnecessary and perhaps carryover primer was causing my 180bp non-specific product in the LibA tailing step so I thought I would use Exonuclease I to clean up the samples this time (just the 1X and 1/10X products).

Using the same altered PCR conditions, I did the LibA tailing step:

(First 4 wells, 1X DNA template, 2nd 4 wells, 1/10X template, 9th well, no DNA template).  That pesky 180bp artifact is still present, so ExoI didn't help in this case.  That, plus the negative control this time mean that the artifact is definitely double-stranded primer-dimer of some sort.  Now I reran the first PCR of this series (M13-ITS primers) on a gel and did another gel extraction (1X and 1/10X products):

You can see the 180bp product is pretty strong still and the ITS product is present, but weak.  I attribute this to the fact that the initial ITS product with this method was pretty weak.  I think it could be partially resolved by adding a few more cycles to the PCR and then doing the gel extraction.  Having your product of interest in molar excess seems crucial to this process.

Chatting with a colleague, he was bothered by the number of PCR reactions needed to generate a 454 amplicon library this way.  I agreed that it would unduly bias your pool so that relative abundances would be almost meaningless and then I thought of another solution.  Put all primers in the same mix, and just do a single amplification!  I didn't have time to redo the initial amplification, even though I probably should have, but I thought I could use the last gel-purified products which used the M13-ITS primers:

These are products just from the 1/10X gel-purified products as template (2uL/well).  I included LibA primers (no tail) at 200nM each as drivers of the desired products along with LibA-MID01-M13 primers at 20nM each to get the reaction going.  As you can see, the 180bp product is quite present, but the ITS products came out much better this time.  Perhaps using a stronger initial M13-ITS product or even including M13-ITS in the whole mix would yield a cleaner amplicon pool.  For now I am out of time for this, and these products are not sequenceable.  We will order a set of barcoded primers specific to the loci we are interested in a gasket our 454 plates until something better comes along or we solve the non-specific product issue.  The Bybee et al (2011) reference stated that such non-specific products used up a third of the reads on their plates.  We can't have that!

High altitude PCR 2: preliminary data!

As promised on Friday, here is some initial data for this idea.  I still don't know if it truly is an effect of altitude or just denature temperature, but here is what I did.  I set up 8 identical reactions (10uL each) to amplify an 800bp product from pUC19 and ran them under three different cycling conditions on the same cycler on the same day (barometer = 30.43 in Hg, corrected boiling point = 93.38C).  I note that there was a lot of condensation present on the upper portion of the wells following Cycle 1, but much less after Cycle 2 or Cycle 3.

Cycle 1
1) 95C 1 min
2) 95C 30s
3) 55C 30s
4) 72C 45s
5) goto 2, 29X

Cycle 2
1) 90C 1 min
2) 90C 30s
3) 55C 30s
4) 72C 45s
5) goto 2, 29X

Cycle 3
1) 90C 1 min
2) 80C 30s
3) 90C 30s
4) 55C 30s
5) 72C 45s
6) goto 2, 29X

I ran 2uL each product on a gel:

Everything looks about the same, but you might notice there are more large (greater than 800bp) non-specific products in the 95C denature and the 80C/90C denature programs than the 90C denature alone.  I think this is partially because 30 cycles may have been too many for this amplicon.  I hope to rerun this analysis with fewer cycles another day.

That said, significance was achieved (ANOVA p = 0.0104)!  Again, I think this is largely due to the added non-specific products.  Here is a nice bar graph showing the results with Tukey's HSD separations denoted. The concentrations were obtained using OD260 method following ethanol precipitation of all products.  Each sample was resuspended in 50uL 1X TE.

What can I take away right now?  90C is adequate for denature of a simple bit of DNA such as pUC19 and I know it avoids boiling my sample at this altitude.  Less condensation appears on the upper portion of the tube walls under a 90C denature, presumably preventing concentration of reagents which can lead to production of non-specific bands.  The effect, regardless of its cause, results in a significant difference in total PCR yield, but presumably more of the product in the 90C denature is the product I want while the other cycles produce unwanted by-products.

Where to go next?  Repeat with fewer cycles, possibly use a qPCR to guide the appropriate number of cycles, measure with a pipet the volume of reaction that remains at the bottom of the well following each cycle next time.  For now, reduce denature step to 90C.  When this particular reaction works a bit better, test it again at different altitudes.  Also maybe test a more complex target.

At the very top of Snowbowl (~11,500ft, if they will loan me an outlet for the day), the boiling point would be 89.2C on the same day, thus boiling all of my reactions.  In the Verde valley (~3,500ft), the boiling point would be 96.9C thus eliminating all boiling for all reactions.

A colleague told me this weekend his new motto is "free Tibet (to do PCR)!"

High altitude PCR

It occurred to me the other day that some unreliability among PCR samples that we experience here in Flagstaff may be attributable to the altitude here (7000ft). I did some googling on high altitude PCR but came up with pretty much nothing. Our lab has 4 Biorad iCyclers, 1 Biorad iCycler for qPCR, 1 BioRad Tetrad (MJ style with 4 blocks), 1 MJ Research PTC-200, and 2 MJ Research PTC-100s.

I first dug into the manuals to try to find anything about altitude correction factors. Nothing. The closest I came was finding that they could be "safely used" at altitudes up to 2000m (~6500ft). That gets almost all the way up to Flagstaff, but says nothing about whether there are any altitude-related issues I should be thinking about.

I then called BioRad tech support and chatted with a rep for a bit on this issue. She was a little amused and quickly regurgitated the ideal gas law (PV=nRT) which would have taken me a little longer to come up with. She pointed out that there are temperature sensors in all of the blocks of the machines we use, so the temperature reported by the cycler is correct as long as you accurately enter the volume of your reaction so it can correctly calculate the sample temperature from the block temperature probes.

I then pointed out that water boils at just under 93C here in Flagstaff (assuming barometric pressure of 23 in. Hg). If we are experiencing high atmospheric pressure (most of the time), the boiling point is slightly higher (closer to 95C) or if we have a storm (low atmospheric pressure) it could go as low as 91C. What this means is that every person here at NAU who follows the prescribed protocol for their PCR is boiling their reactions at every denature step. Since the denature temperature for most programs is still 94-95C, this is just the cusp of the boiling point on most days (it is usually sunny here). It also is not above the magic point at which you will significantly degrade the half-life of your enzyme with each step (about 94-95C). The boiling can still have a significant effect on the reaction by changing the concentration of your buffer (increased concentration of all reagents). Sometimes the moisture is retained due to the heated lid preventing condensation on the upper part of the tube and the lid compression preventing any gaseous escape, but more often than not, your reaction will contain less fluid at the end of the cycle than you began with.

Example: If you perform 10uL reactions and after 35 cycles you have 8uL, your MgCl2 concentration (to pick a reagent) will increase from 2mM to 2.5mM over the course of your cycle. Excess of MgCl2 can contribute to mis-priming and production of non-specific products. Excess KCl can produce unwanted short non-specific products. Excess polymerase can result in all kinds of background (a smeared appearance). Some environmental DNA samples such as are often processed in our lab also contain a lot of PCR inhibitors (e.g. polyphenols). The evaporation can therefore increase the endogenous inhibitor concentration to a point that effectively stops the reaction somewhere along the cycle.

You get the idea. Evaporation BAD!!!(say it like Dana Carvey playing George HW Bush)  Though she couldn't provide me a concrete solution to this concern short of an artificially pressurized laboratory, she offered two suggestions that sounded very good to me:

1) Persons performing PCR at high altitude should reduce their temperature of denaturation according to the local atmospheric pressure. For our altitude, she suggested 90C. I asked if this time should be extended slightly to account for the lower temperature and she said she did not know, but it probably wouldn't hurt...which takes us to her second suggestion.

2) PCR (generally speaking at all altitudes) of GC-rich regions should include a "pre-denature" step of 80C for 1 min to "slow" the denature step to allow for the disentanglement of complex secondary structures and eventual denaturation of the GC-rich region. I should point out that she offered this as a complete alternative to ever adding DMSO to your reaction which, as you may know destabilizes hydrogen bonding, thus allowing efficient denaturation of GC-rich regions, but then also complicates your annealing step. She claimed the 80C for 1 min will solve this problem without messing with your annealing and thus may also be useful for high-altitude PCR.

What I learned: At 7000ft, reduce the annealing temperature to 90C. I can probably keep my denature time at 20-30 sec, but a pre-denature step of 15-30 sec at 80C may improve overall reaction efficiency with no further changes (longer if PCR target region is GC-rich).

Happy high altitude PCRing!!

Reducing laboratory costs

I was chatting with a colleague this morning about cutting lab costs. Rather than go into a lengthy discussion, I thought I would post a quick list of some things I do that bake a big difference in my lab budget.

- Use chemical methods when possible for DNA extractions rather than kits. Kits often cost $2-4 per sample. By chemical method in 2mL tubes, the cost immediately plummets. If you scale down your extraction into 96-well PCR plates, your reagent needs as well as your labor get minimized. Some of us just need a few PCR reactions. Depending on your organism, you may be able to do a rapid alkaline lysis method, further reducing your costs and material demands (e.g. http://www.google.com/url?sa=t&rct=j&q=&esrc=s&source=web&cd=2&ved=0CFsQFjAB&url=http%3A%2F%2Fwww.biotechniques.com%2Fmultimedia%2Farchive%2F00010%2F03344rr04_10933a.pdf&ei=98wQT7uZMam02gXiyIGECg&usg=AFQjCNF1HORNnYJQdbCmzbNYj-QGG0oJTw&sig2=lw4xwfnlpUgXNKVE3PPVrw).

- Reduce your PCR reaction volume. Most PCR kits suggest a reaction volume of at least 25uL, some as large as 100uL. I almost never do more than 10uL. If using a thermal cycler with adequate lid pressure, 6uL is possible in a 96 well plate. I now use 384-well plates and with the smaller wells (less headspace), I get good reactions of 4uL. That leaves me 2uL to run on a gel and another 2uL for capillary analysis. No waste!!

- Use reusable silicone PCR mats (e.g. http://www.phenixresearch.com/products/mpcs-3510-sealing-mat-pressure-fit-lid.asp). These things are great and I have always enjoyed less evaporation than with for instance, microseal B films. You can bleach or autoclave them if you like, but that will reduce their life. I simply throw used mats into a beaker with hot tap water for 10 minutes, rinse with RO water, and dry on a paper towel. I've never had a problem with my NTC reactions. For 384-well plates, try these: http://www.phenixresearch.com/products/smx-pcr384-sealing-mat.asp

- Use PCR plates (96 or 384) rather than strip tubes. Each plate costs about as much as 4 strip tubes last I checked. You don't have to throw out your PCR plate after running just a few reactions either. I keep partially used ones on my bench and just mark off the already used wells with a marker. When I need new wells, I peel back my silicone mat and use the unused wells, and don't bother to clean the mat until every well has been used. Still no problems with my NTCs.

- Agarose gels are reusable. Once or twice, but it can get pretty ugly. Make sure to keep spent gels intact in a sealed container (can hold numerous gels at once). Break the desired size of a gel into a flask for remelting and bring the mass back up where it should have been with RO water (if you had a 80mL gel, it should weigh ~80g). Add some new dye (e.g. ethidium or SYBR), pour, and run as always. You will notice some increased background, but for a quick gel to check some unimportant results, this works fine.

- SYBR safe can be used at 25,000X rather than 10,000X with no real loss of sensitivity.

- BigDye reactions (10uL) work fine with just 0.5uL BigDye. Add 2uL sequencing buffer, and sequence as before, or with this modified protocol I like: http://www.biotechniques.com/multimedia/archive/00003/BTN_A_000112499_O_3096a.pdf

Of course there are other things you can do, but this is all I have time for today...