2bRAD Library Preparation
Introduction
This protocol is primarily adapted from
Eli Meyer's lab at OSU to prepare sequencing libraries for 2bRAD sequencing. Some adaptations have been made based on my own experience and the resources at my lab, and I try to mention those where applicable. I am using a reduced tag representation approach, where only 1/4 of all sites are sequenced based on selected oligos.
Materials
Procedure
Prepare intact, high-quality genomic DNA samples each containing 100ng-1 ug in 8 uL of nuclease free water (NFW). Must be RNA-free!
Eli Meyer protocol calls for 1.2 ug when using AlfI. Matz protocol calls for 100-200 ng when using BcgI enzyme. I've had success with 900ng and AlfI.
Add 900 ng of DNA to well, cover with Airpore tape, and leave overnight at 37degC to dry down.
Add 8 uL NFW to each well. Let dissolve at 4degC for at least an hour before proceeding.
Prepare digestion master mix, with 4 uL per 8 uL of sample.
Combine 4 uL master mix with each 8 uL DNA sample. Incubate at 37degC for 2 hours. Inactivate at 65degC for 15 min. Hold on ice or at 0degC in thermocycler.
Protocol calls to hold at 37degC for at least 1 hour or as long as overnight.
(Optional) For each sample, load 2 μl digested DNA on a 1% agarose gel alongside a comparable amount of intact DNA from the same sample to verify the effectiveness of the digest.
The original gDNA should appear as a single high-molecular weight band (>10kb) while the digested samples should be visibly degraded. An effective AlfI digestion produces a slight downward shift in the original HMW band and a subtle smear trailing downward from that band. If initial gDNA is degraded, this test is not useful and may be skipped.
In this step adaptors are ligated to the restriction fragments produced above. If reduced tag representation (RTR) is required, selective adaptors must be chosen at this stage.
Make fresh adapters by adding the oligos into separate tubes at final concentration of 2 uM each and leaving at room temperature for 10 minutes. Scale depending on how many samples you'll be doing.
Make Ligation master mix. Want total of 50 uL after mixing digestion and ligation master mix. Hold master mix and samples on ice at all times.
If 10 uL of digested sample, make 40 uL of MM per sample. If 12 uL, make 38 uL and just reduce NFW.
Combine 38 uL MM with 12 uL digested DNA. Incubate at 16degC for 2 hours followed by 10 minutes at 62degC, then put in freezer.
Protocol calls for at least 1 hour or overnight incubation. Meyer protocol does not call for heat inactivation step, Matz protocol does... I chose to do heat inactivation.
Plan and record barcode assignments. Each sample has to be assigned a unique combination of “BC” and “HT” oligos. BC oligos introduce index 1 (“i7”) and HT oligos introduce index 2 (“i5”).
Make new primer aliquots if needed.
Make PCR master mix. This is scaled so you have 77.25 uL per PCR reaction instead of 100 uL like in Meyer protocol. This is because 100 uL did not fit in my wells.
Add 3.5 uL of the appropriate BC oligo to each well. Add 3.5 uL of the appropriate HT oligo to each well.
Add 50 uL of ligation product to the appropriate well.
Amplify using minimum number of cycles determined in test PCR (see original Meyer protocol). I'm using 17 cycles.
In this step the sequencing constructs are gel-purified to eliminate residual genomic DNA and primer dimers.
Prepare SB or TAE gel buffer stock if needed.
http://www.openwetware.org/wiki/SB
Prepare a 2% agarose gel. Use a thick comb so at least 75 uL of PCR product will fit. Tape together combs if needed to get larger well.
Add 1 uL loading dye for every 10 uL of PCR product. I add 7.5 uL loading dye.
Load all but 5 uL of mixture into well and run gel long enough to separate target from non target bands (i.e. ~170 vs ~130 bp). I do at least 1 hour at 110mV.
View the gel briefly (<30 seconds) on a UV transilluminator set at low intensity to verify the presence of target bands and adequate separation of molecular weight standards to resolve bands in the 50 to 200 bp range. Typically ~5 cm run distance (well to dye front) is sufficient.
Cut out the target band (~170 bp) in a narrow gel slice, being careful to avoid any primer dimers (70-90 and ~130 bp) that may be present.
At this point a commercial gel extraction kit can be used, but I've had great success with this method and it is obviously much cheaper.
Transfer each gel slice into a 2 ml microcentrifuge tube and add 40 μl NFW.
Centrifuge tubes 1 min at high speed to bring gel into contact with the water. Optional: incubate at 4°C overnight to maximize yield. Otherwise, proceed directly to freezer.
I always leave overnight.
Freeze at -80°C for at least 30 minutes.
Centrifuge at maximum speed in refrigerated centrifuge (4degC) for 10-20 minutes.
Press gel slice against side of tube using pipette tip, and withdraw supernatant (at least 60 μl should be recovered). If less than 60 μl is accessible, repeat centrifugation. Transfer the supernatant to a new PCR tube or plate.
Finally, the libraries are pooled in equal ratios in an effort to sequence all samples at equal coverage. More recent version of Meyer and Matz protocol uses qPCR to do this. I usually follow the original method of quatifying, but qPCR might lead to more even sequencing coverage which could be important if you have a lot of samples per library.
Quantify DNA concentrations of each sample using Quibit High Sensitivity.
My values are usually ~0.3 ng/mL in 60 uL.
Find the lowest concentration among your samples (excluding any that might have failed based on very low (< 0.01ng/mL) DNA concentration or faint/nonexistent band after PCR). Calculate how many ng DNA is in 40 uL for that sample. Use that value to calculate how many uL to add to pool for all other samples. Add to 1.5 mL tube.
Example: Sample A has the lowest concentration across all samples, with 0.100 ng/uL. So I will add 40uL*0.1 ng/uL = 4ng of DNA from each sample to the final pool. I add 40 uL of Sample A to tube, then for all other samples I divide 4ng by theire concentration to get the volume to add to the tube.
Illumina sequencing typically requires templates in ≤20 μl volume at ≥2 nM concentration. The pooled libraries produced above are typically too dilute for sequencing. To purify and concentrate these libraries, they may be precipitated with isopropanol or using a commercial PCR cleanup kit.
I use Quiagen PCR cleanup kit and spin down the sample multiple times until all of it has passed through the column. I dilute in 50 uL elution buffer.