About this document

This lab protocol is published only to demonstrate how samples libraries were prepared for XXX manuscript. This 2bRAD protocol is based off the original protocol which was published by Wang et al. 2012. The most up-to-date version of this protocol is available through the Matz Lab and associated analysis pipelines can be found on Misha Matz’s GitHub repository.

Digestion

  1. Prepare samples each containing 200 ng of DNA in 4 µL, 50 ng/µL
    • Equalize your input DNA samples well (based on qubit or picogreen, not just nanodrop) and ensure that their OD 260/280 ratio is >=1.8 And 260/230 ration is ~2 or higher.
    • Concentrating can be accomplished by ethanol precipitation, drying under vacuum, or by using a zymo clean and concentrate spin column in low volumes of eluant.
    • Making a 50 ng/µL dilution plate makes it easy to add the sample to the digestion mastermix later.
  2. Prepare a digestion master mix.
    Reagent Volume (µL) Total Volume (µL) 96 rxn+10% error
    NEB Buffer #3 0.6 63.36
    320 µM SAM 0.4 42.24
    BcgI (2 U µl-1) 1.0 105.60
    Total 2.0 211.20

    Note: SAM [S-adenosyl-methionine] comes at 32 mM stock, add 198 uL of NFW to 2 uL aliquots of 32 mM SAM.

  3. Add 2 uL of mastermix to each well, can be accomplished quickly and precisely with a 10 uL electronic pipette.

  4. Use a multichannel pipette to combine 4 uL of the DNA sample with the 2 µl master mix (6 µl total volume).

  5. Cover the plate with PCR film, spin down, and incubate at 37°C in a thermocycler with heated lid for 1 hr.

  6. Inactivate the enzyme at 65°C for 10 min then hold samples at 4°C. Hold samples on ice after this.

Digestion PCR Profile
Temperature Time
37°C 60 min
65°C 10 min
4°C HOLD

Ligation

In this step adaptors are ligated to the restriction fragments produced above.

  1. Prepare double stranded adaptors by combining each pair of primers, Adaptor 1 (5ILL-NNRW, Anti5ill-NNRW) and the 12 different pairs of 3illBC, anti3illBC.
  • For a full plate, prepare Adaptor 1 in PCR tube, mix 60 µl of 5ILL-NNRW (10 µM) with 60 µl of Anti5ill-NNRW (10 µM).
Adapter 1 Component Volume (uL) Total Volume (uL)
5ILL-NNRW 0.5 60
Anti5ill-NNRW 0.5 60
Adapter 1 1.0 120
  • For Adaptor 2, set up 12 PCR tubes, to each tube mix 5 µl of 3illBC(1-12) (10 µM) with 5 µl Anti-ill-BC(1-12) (10 µM).
Adapter 2 Component Volume (uL) Total Volume (uL)
3illBC(1-12) 0.5 5
Anti-ill-BC(1-12) 0.5 5
Adapter 2 1.0 10
  • Incubate at 42°C for 5 minutes then keep at room temperature until ligation.
  1. Prepare 12 master mixes for ligations (one for each barcoded 3’ primer). This recipe is for a single reaction, so scale up as needed.
    Component Reaction Volume (uL)
    NFW 15
    10x T4 ligase buffer w 10 mM ATP 2
    5 μM Adapter 1 1
    5 μM Adapter 2 (Different for each column) 1
    T4 DNA ligase 1
    Total 20
First make an initial master mix:
Component Reaction Volume (uL) Total Volume (uL) for 8 rxn+10% error Total Volume for 12 MM+10% error
10x T4 ligase buffer w 10 mM ATP 2 17.6 232.3
5 μM Adapter 1 1 8.8 116.2
T4 DNA ligase 1 8.8 116.2
Total 4 35.2 464.6
Then into 12 separate 0.2 mL PCR tubes add:
Component Total Volume (uL) for 8 rxn+10% error
Initial Master Mix 35.2
5 μM Adapter 2 (Different for each plate column) 8.8
NFW 132.0
Total 176.0

  1. Use a 100 uL multi-channel pipette to combine 20 µl master mix with digested DNA (~25 µl total volume). Keep on ice while mixing.

  2. Incubate at 16°C for BcgI for 12 hours.

  3. Heat at 65°C for at least 40 min to inactivate the ligase (in a thermocycler with heated lid).
    Ligation PCR Profile
    Temperature Time
    16°C 12 hours
    65°C 40 min
    4°C HOLD

Optional Step: qPCR to ensure amplification success

  1. Prepare a qPCR Plate, for each plate you can run three rows of samples in duplicate and three negative control wells, you will need to run three qPCR plates for every ligation plate.
Component Reaction Vol (uL) Total Volume (uL) for 75 rxn+10% error
NFW 5.53 456.23
SYBR Green Mastermix 7.50 618.75
10 uM TruSeq 0.07 5.78
1 uM any ILLBC Primer 0.70 57.75
10 uM P5 0.10 8.25
10 uM P7 0.10 8.25
Total 14.00 1484.00

  1. Add 14 uL of mastermix to each well, avoid bubbles

  2. Add 1 uL of ligation to each well

  3. Centrifuge plate

  4. Turn on qPCR machine, ensure the correct reaction volume (15 uL) is inputted, follow the 2bRAD template and ensure the plate is inserted correctly.

# of Cycles Step Temperature Acquisition Time
1x Pre-Incubation 95C None 10 min
40x Amplification 95C None 15 sec
60C None 30 sec
72C Single 30 sec
1x Melting Curve 95C None 5 sec
65C None 1 min
97C Continuous
1x Cooling 40C None 10   sec

  1. Rank samples from highest to lowest CT score.

  2. Optional: Select 4 sample ligations with relatively low CT scores and 4 samples with relatively high CT scores to use in a test PCR.

Optional Step: Test PCR

Optional: Test PCR for a set of 8 ligations, recommended to use 4 with low Ct values and 4 with high Ct values if qPCR was conducted:
Reagent Volume (uL) per sample Total Volume (uL) for 8 rxn+10% error
dNTPs 2.5 mM ea 0.4 3.52
H2O 9.7 85.36
10 µM IC1-P5 0.4 3.52
10 µM IC1-P7 0.4 3.52
1 µM ILL-BC oligo 2.4 21.12
10 µM TruSeq_UN oligo 0.3 2.64
10x Titanium buffer 2.0 17.60
Titanium Taq 0.4 3.52
Total 16.0 140.80
  1. Add 16 µl of master mix to 8 strip-tubes, then add 4 µl of ligation.
Amplify as follows:
Amplification PCR Profile
Temperature Time
70°C 30 sec
95°C 20 sec
65°C 3 min X 15 cycles
72°C 30 sec
4°C Continuously
  1. Load 5 µl on a 2% agarose gel alongside LMW ladder or other marker that has 150 and 200 bp bands. Confirm that all samples have a visible band at ~180 bp.

Amplification

  1. Pool ligations by row in strip-tubes, using 6 µl from each well. The 96-well plate is now reduced to 8 ligation pools, each corresponding to the original row. Store the ligations at -20°C.

  2. For each reaction prepare the following master mix:
    Reagent Volume (uL) per sample Total Volume (uL) for 8 rxn+10% error
    dNTPs 2.5 mM ea 1 8.8
    H2O 12 105.6
    10 µM IC1-P5 1 8.8
    10 µM IC1-P7 1 8.8
    10x Titanium buffer 5 44.0
    Titanium Taq 1 8.8
    Total 21 184.4
  3. Set up the 8 PCR tubes and combine:
    Reagent Volume (uL)
    Master mix 21
    Pooled Ligation 20
    1 µM ILL-BC primer Different for each tube 6
    2 uM TruSeq Different for each set of 8 tubes 3
    Total 50
  4. Amplify as follows:
    Amplification PCR Profile
    Temperature Time
    70°C 30 sec
    95°C 20 sec
    65°C 3 min X 15 cycles
    72°C 30 sec
    4°C Continuously

  5. Load 5 µl on a 2% agarose gel alongside LMW ladder or other marker that has 150 and 200 bp bands. Running the gel at 150 V for 30 minutes should result in enough band separation.

  6. Confirm that all samples have a visible band at ~180 bp. You might also see a band below 150 bp, which is an artifact from the carried-over ligase (if this is an issue you can heat inactivate the ligase for longer)
  • If the 180 bp product is visible but barely, add two more cycles to the same reactions, do not continue to amplify the samples if no band is visible at 15 cycles, go back and troubleshoot the samples within that pool.

Now, you can send these pools off for automated size-selection through pippin prep, or can manually size-select and gel-purify.

Optional Step: Gel-purification

  1. Prepare a 2% agarose gel using Sodium Borate buffer. Use a wide comb that can accommodate 30-50 μl, or simply tape together two wells.

  2. Load 30-50 μl of sample (40 μl sample + 10 μl loading dye) alongside LMW ladder. Run gel at low voltage for 90 minutes at 100 V or until bands at 150bp and 200bp will be clearly resolved.

  3. View the gel on a blue-light transilluminator to verify the presence of target band and adequate separation of molecular weight standards to resolve bands at ~180 bp and (possibly) below 150 bp. Photograph.

  4. Cut out target ~180 bp band in a narrow gel slice, avoiding the edges of the lane (i.e., cut out the middle 70-75% of the band). Cut just inside the bottom boundary of the target band to avoid getting anything smaller.

  5. Follow the QiaQuick gel extraction protocol.

Optional Step: Combining libraries for sequencing

Pool libraries in equimolar ratios to try to equalize coverage across sample pools.

  1. Prepare a 1:100 dilution of each pooled library by combining 2 uL of eluted library with 198 uL of NFW, can use multi-channel pipette.

  2. Prepare qPCR mastermix, run each library in duplicate and have 2-3 wells reserved for negative controls.
    Reagent Volume (uL) per sample Total Volume (uL) for 21 rxns (8 rxns in duplicate and three negative controls + error)
    NFW 4.3 90.3
    Sybr Green qPCR MM 7.5 157.5
    10 uM P5 0.6 12.6
    10 uM P7 0.6 12.6
    Total 13.0 273.0
    Diluted PCR Product (1:100) 2.0 NA

  3. Conduct qPCR following previously described qPCR profile and calculate CT values for each sample.

  4. To determine volumes of each library for the combined pool:
  • Rank samples from lowest to highest CT and identify reference sample (sample with the highest CT)
  • Calculate the proportion of each library to sequence as: PL= 2 ^ [CT(sample) – CT (reference)]
  • Calculate the volume of each library to use as: V= PL * 28 uL
  1. Combine libraries and follow Zymo Clean and Concentrator Protocol to concentrate final libraries in ~25 uL of NFW.
  • Note: For PCR products you must use a 5:1 ratio of binding buffer to sample.
  1. Qubit the final pool with a high-sensitivity kit to determine if further concentration is needed.