Sanger sequencing might be the way of the past, but it remains an essential tool for many applications. Researchers can submit samples for processing as raw DNA (requiring PCR amplification), plasmid, or PCR product for sequencing. However, as costs for pretty much everything continue to rise, the only way you can control your Sanger costs is to become proficient at this technique and roll back the amount of bigdye that you use per reaction. List price on the Life Tech website is now about $1100 for 800uL of the stuff, and that doesn't include tax, handling, or dry ice charges. But we need our sequences, and some of us just don't have the budget to produce sequence according to the ABI protocol. Here I present a brief protocol for producing high quality sequences using just 0.2uL bigdye per reaction.
1) PCR a clean product (no extra bands)
2) ExoSAP your reactions:
Combine (adjust volumes to maintain ratios) 50uL H2O, 5uL SAP (1U/ul), 0.5uL ExoI (10U/ul). Add 2uL to each reaction per about 5uL volume, mix and spin down, and run cycler program (37C 40min, 80C 20min, 10C forever). Alternatively, you can add less exosap (1uL perhaps), mix and spin down, then let reactions sit on the bench overnight. In the morning kill the enzymes with 20min at 80C.
3) Prepare sequencing reactions:
First, I do this in 384well plates. This means I have very little headspace into which to evaporate any sample volume, and this presumably keeps my chemistry much more stable than if you were to do this in a 96well plate. That said, I have done many many 5uL reactions in 96well plates with no problems, but I very much prefer 384well plates these days.
Start with a high concentration primer working dilution (20uM is good, 15uM is easier for fewer reactions). For the following calculations, I use these solutions: BigDye v3.1, 5X BigDye sequencing buffer, 50mM MgCl2, 20uM primer.
Each reaction contains:
0.2uL BigDye
1uL Sequencing buffer (final at 1X)
0.15uL MgCl2 (final at 1.5mM extra)
0.75uL primer (final at 3uM)
2uL template
0.9uL H2O
Multiply by the number of samples you have and add 10% for pipetting error. Distribute 3uL of this mixture to each well, and follow with 2uL of template. I prefer to seal PCR plates for any thermal cycling applications with reusable silicone mats (http://www.phenixresearch.com/products/smx-pcr384-sealing-mat.asp; http://www.phenixresearch.com/products/mpcs-3510-sealing-mat-pressure-fit-lid.asp) since microseals gave me some grief many years ago (mostly edge evaporation). You just need to wash these with water. Making yourself crazy with bleach and autoclaving will shorten their life substantially, plus it's pretty much a waste of time. Run the following thermal cycle: 95C 2min; 60 cycles of 95C 10s, 50C 10s, 60C 2min; 10C forever.
4) Retrieve your plate and get ready for cleanup. For 384well plates there is not enough space for an ethanol cleanup, so I use a modification of the Rohland and Reich bead cleanup (http://enggen-nau.blogspot.com/2013/03/bead-cleanups.html). Make a higher percentage PEG solution (25% instead of 18%) with this recipe (see other post for part numbers):
2650uL H2O
50uL 10% Tween-20
100uL 1M Tris (pH 7)
2000uL 5M NaCl
5000uL 50% PEG8000
200uL carboxylated beads
Mix solution very well, and careful pipetting the PEG as it is like honey. Add 15uL to each sequencing reaction. Seal thoroughly with adhesive foil and mix by inversion. Spin solution down gently. Just fast/long enough to get the solution into the bottom of the well. If you see pelleted beads, you need to mix again and spin down more gently. This may take some experimenting with your centrifuge. I use an Eppendorf 5804R with an A-2-MTP rotor, and I let it spin up to about 1000 and hit stop to get things into the wells. Let stand for ~45 min. The precipitation is somewhat time-dependent as well as concentration dependent, so the longer you wait (to a point), the more sequence you will see close to the primer. When your timer goes off, or you think you waited long enough, apply your plate to a magnet stand (http://www.alpaqua.com/Products/MagnetPlates/384PostMagnetPlate.aspx). Tape it in place on either end to keep it from moving. Separation should take about 5 min, but waiting another 5 min doesn't hurt. Now, you can pipette the waste volume out or you can do some inverted centrifuging and save a lot of time and tips in the process. With my centrifuge, 1 min inverted spins on 3 folded paper towels (if plate is full) at 400rpm works well. Very important that acceleration and deceleration are set to 1. Any brown you see on the paper towel afterward is usually residual beads that didn't make it to the magnet. I like to "lube" each well by adding 5uL 70% ethanol before the first spin. This reduces the viscosity and eases the solution from each well. Multichannel repeat dispensing electronic pipettes are very useful here. After the first inverted spin, take the magnet/plate back to your bench. Add 25uL 70% EtOH to each well. No need to wait, take the plate right to the centrifuge and spin inverted again. Repeat the 70% wash twice more. After the third wash, allow beads to dry (~30 min at room temp, or 3 min in vacuum centrifuge at 60C, mode D-AL so rotor does not turn). Note that over-dried beads can be very hard to resuspend, and this translates into samples where the DNA doesn't want to go back into solution. Once dry, resuspend samples in 20uL sterile water. It helps to seal with foil so you can vortex the plate. Samples should look like mud during resuspension. If you see any that look clear with brown flakes, keep vortexing. Once samples have had the appearance of mud for ~2-5 min, place plate back on magnet and transfer 10uL to a 96 well plate for sequencing. A little bead carry over will make no difference. Just spin the plate down hard to pellet the beads before submission. If you need to reinject any samples, you still have 10uL of backup sequence product. Also note that you do NOT need to denature. Cycle sequencing produces only single stranded products. Put them right on the instrument.
5) Enjoy your data!!
A note on sequencer usage, our lab has both a 3130 (4 cap) and a 3730xl (96 cap). They do the same thing, and yet their stock protocols were not equal. I wondered at first if this had to do with something else in the instrument, but then I noticed I got crappy, low-signal peaks on the 3130 when I ran the same product on both instruments. I checked, and it injected samples for less time, and at a lower voltage. Further, it cut off sequences after about 600 bases. Ask your sequencing lab about the module they use. If at all possible, have them set the injection voltage to 1.5 kV and the injection time to 20sec as this fixed all my problems. I also extended the run time on the 3130 from 1200 to 1800s, and now I get 1000 bases of sequence. We have many many runs on both instruments (just replaced the array on the 3130 after 1200+ injections), so this protocol adjustment isn't going to shorten instrument life at all.
And finally, some notes on my sequencing recipe. We had some difficult samples last year and did a lot of troubleshooting. First we did a MgCl2 gradient (on ABI control plasmid) to see how this affected results. Addition of 1.5mM MgCl2 seemed to give an extra 20-40 bases of high quality data. However, it was addition of copious amounts of primer that made the real difference. This allowed us to sequence very dilute samples with high success and get nice long read lengths (900+ bases). We experimented with lower volumes of bigdye and got nice data with as little as 0.05uL/reaction, though the signal dropped off after a few hundred bases. Perhaps more cycles would improve this, but evaporation can become a challenge when you start doing 100 cycles, and then the cycler runs forever and you can't get anything else done. I have also found that faster cycling works OK with Bigdye (try cutting extension time to 1 min and raising temp to 68C), but I am not confident enough to use it as a general protocol yet.
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