BLink-seq libraries

This page provides the BLink-seq library preparation protocol in full as a reference. The official PDF is better formatted for printing and using in real time at a laboratory bench.

BLink-seq Library Protocol

BLink-seq (bead-barcoded linked-read sequencing) technology generates an Illumina library of tagmented inserts that are individually barcoded based on the molecule of origin. The combinatorial bead-barcode that is added to each sub- fragment of the original HMW gDNA molecule can be used to infer phasing and linkage of sequenced fragments. The method uses BLink beads: magnetic streptavidin dynabeads coupled to a 2x-biotinylated oligo containing a TruSeq- P5 PCR handle, a variable-length barcode (vBC) to offset the start of the Illumina R1 read, a combinatorial barcode (3 x 12nt BC segments), and the Tn5 ME sequence. BLink beads are loaded with purified Tn5 protein to enable on-bead tagmentation of HMW gDNA. A second Tn5 transposome in solution (ME-B) adds the Nextera-P7 PCR handle. PCR with Illumina UDI primers (TruSeq-i5 and Nextera-i7) generates the final BLink-seq library.

The final insert size distribution is determined by the frequency of on-bead tagmentation (i5 end) and solution-phase tagmentation (i7 end). Pilot experiments are recommended to optimize the insert size distribution to maximize the fraction of the library within the 450-1200bp range. Adjusting both the amount of Tn5 used to load beads (Appendix D) and especially the amount of solution-phase ME-B transposome is key to optimizing insert size. Once beads are saturated, adding more Tn5 to beads typically does not alter the insert size, but adding more ME-B transposome during library prep typically will decrease the insert size distribution. The amount of ME-B transposome required scales with gDNA input. BLink libraries can be sequenced on Illumina instruments with standard sequencing primers and read lengths.

Materials and Equipment Required: See Appendix A

ME-B transposome: To prepare ME-B transposome in solution, see Appendix B

BLink beads: Can be stored in 3 formats:

1. fully loaded with active Tn5 transposomes, ready to use: follow protocol below
2. ME-REV hybridized, ready to load with Tn5: see Appendix D to prepare Tn5-loaded beads
3. Beads containing barcoded oligos, but requiring final stripping and ME-REV hybridization: see Appendix C to strip/hybridize ME-REV, and then Appendix D to prepare Tn5-loaded beads

Input genomic DNA: High molecular weight (HMW) genomic DNA will generate BLink-seq libraries with more linked reads and longer N50 values.

DNA quality matters

If genomic DNA is highly degraded, BLink-seq will offer little value over WGS.

Experiment planning

Typical tagmentation reaction components and volumes:

per sample	Component	Notes
25 μl	2x WASH+PEG buffer	final: 1x WASH, 0.1% Triton, 6.5% PEG6000
5 μl	Tn5-loaded BLink beads	see Appendix C+D if needed
5 μl	DNA	1.5ng total, dilute primary stocks to 0.3ng/ul
14 μl	CS+DMF master mix	final: 1x CutSmart buffer, 10% DMF
1 μl	diluted (e.g. ~0.2μM) ME-B transposome*	See Appendix B if needed
50 μl	total volume in tagmentation reaction

*Once the appropriate amount of ME-B is optimized for a given batch of loaded beads + gDNA samples, ME-B transposome can be added to the CS+DMF master mix for high throughput library prep, e.g. adding 15μl CS+DMF+ME- B master mix per sample

Volumes can be modified but the final concentrations of PEG, Triton, CutSmart buffer, and DMF should stay constant. When increasing the amount of beads per reaction, adjustments to the WASH+PEG buffer concentration can be made. For example, to use 50μL beads in a final 150ul tagmentation reaction volume, modify the plan to use a 3x formulation of WASH+PEG buffer and scale up the amount of the CS+DMF master mix to maintain the final concentration per reaction.

The ratio of DNA:beads can change to meet project needs. A higher ratio of DNA to beads will lead to more barcode clashing because multiple molecules may be tagmented on the same bead and have the same barcode. Pilot experiments: vary the amount of ME-B transposome to optimize the amount needed for a good final library size distribution. Typically the primary 2μM ME-B transposome stock is serially diluted in 50% glycerol storage buffer, such that 1μl of different dilutions are tested in the pilot phase. The optimal amount of ME-B transposome required is related to the amount of DNA, not the volume of beads; if the volume of beads is increased but DNA stays consistent, do not alter the amount of ME-B transposome per reaction.

Library Protocol

Tip

This protocol is designed for PCR strip tubes and reagent addition using multichannel pipettes or liquid handling robotics (e.g. Mantis). Strip tubes with individually hinged caps are recommended instead of fixed plate format to allow each strip to be mixed immediately and completely after each addition.

1. Prepare reagents

recommended use bench space and equipment that is dedicated to pre-amplification steps through step 5.

1X WASH+T buffer

Prepare fresh each day, store at room temperature

5mL is sufficient for ~20 samples, scale up or prepare more as needed

Volume	Reagent
4.45 mL	Nuclease-free water
500 μL	10x WASH buffer
50 μL	10% Triton X-100
5 mL	Total volume

2x WASH+PEG buffer

Prepare fresh each day, store at room temperature.

Prepare excess buffer, e.g. 1.2 * N samples, this buffer is viscous and may need careful handling and optimization of settings on automated liquid handlers.

Vol/sample	Reagent
13.0 μL	Nuclease-free water
5.0 μL	10x WASH buffer
0.5 μL	10% Triton X-100
6.5 μL	50% PEG8000
25.0 μL	Total volume

HMW sample gDNA

Dilute to target concentration in 1x WASH+T buffer (e.g. 0.3ng/μl in min 5μl).

Rotate (or shake gently < 500RPM) at room temperature to mix well. Do not vortex vigorously to avoid shearing.

Long-term storage

While DNA samples can be diluted in advance, long-term storage of low concentration gDNA is not recommended as DNA may adsorb to polypropylene. The Triton X-100 in the 1x WASH+T buffer may help block adsorbtion.

2. Tagmentation reaction

Immediately quench after

The tagmentation reaction needs to be stopped with SDS and Triton. These steps are separated into sections for visual clarity, but quenching needs to be done right after incubation finishes.

1. Preheat a Thermomixer with a 96well adapter to 55°C

2. Add 25μl WASH+PEG per sample to labeled strip tubes

3. Add 5μl Tn5-loaded BLink beads to each tube

   • Note that beads are not washed before use, especially if ME-B duplex oligo is added when beads are loaded with Tn5 protein (see Appendix C).

   Master Mix

   Optionally, instead of adding WASH+PEG and beads separately, prepare a master mix before aliquoting to tubes (e.g. 30μl total when using 5μl beads). This may lead to some bead loss because of the excess required to prepare in master mix format, so we recommend adding separately as indicated above if beads are limiting.

4. Close the strip and flick to mix well

   • The beads should be evenly dispersed before gDNA is added

5. One strip at a time, add 5μl diluted gDNA per sample. Immediately close the strip and flick/invert to mix well

   • Rapid and complete mixing is an important step for assay performance
   • recommended Use 1x WASH+T buffer as a no-DNA negative control

6. Incubate minimum 10 min on rotator at room temperature

   • This pre-incubation step allows DNA to potentially bind to beads before magnesium is added to activate Tn5

7. Prepare CutSmart/DMF master mix at room temperature:

   CutSmart/DMF Master Mix

   Vol/sample	Reagent	Notes
   4 μL	Nuclease-free water
   5 μL	10x rCutSmart buffer	final 1x in tagmentation reaction = 10mM Mg++
   5 μL	DMF	final 10% in tagmentation reaction
   1 μL	ME-B transposome (e.g. 1:10 dilution of 2μM stock) *
   15 μL	Total

   *Before the optimal amount of ME-B is determined, do not include ME-B transposome in the master mix.

   Prepare excess master mix, e.g. 1.1 * N samples, or more if required for liquid handling dead volume

   For large master mixes, a primary stock of 2μM loaded ME-B can also be used; adjust the volume of water accordingly.

   8. Remove the strip-tubes from the rotator. If needed, very briefly pulse-spin the strip tubes to collect the volume at the bottom of the tube but try not to pellet the beads. Flick gently if the beads are not well dispersed.
   9. one strip at a time add 15μl CutSmart/DMF/ME-B Master Mix per sample. Immediately close the strip and flick to mix well.
   10. alternative If ME-B is added separately (e.g. pilot phase), first add 1μl diluted ME-B transposome per sample and mix well. Then add 14μl CutSmart/DMF/ME-B Master Mix per sample and immediately mix again.

   Mix well

   Rapid and complete mixing is an important step for assay performance. Addition of the CutSmart buffer containing magnesium will activate Tn5 transposition.

   11. Incubate samples 30 min, 500RPM on a Thermomixer at 55°C. When finished, proceed immediately with SDS quench.

3. Quench Tagmentation Reaction

12. Prepare 2% SDS
    • use a strip-tube reservoir, if needed; excess can be stored at room temperature
13. When the tagmentation reaction is complete, immediately add 5μl 2% SDS per sample (10% of the tagmentation reaction volume). Immediately close each strip after adding SDS and flick to mix well
14. Incubate samples 7 min, 500RPM on a Thermomixer at 55°C
15. Prepare 10% Triton X-100
    • use a strip-tube reservoir, if needed; excess can be stored at room temperature
16. When the SDS incubation is complete, add 5 μl 10% Triton X-100 per sample to quench SDS (equal volume to the amount of 2% SDS added above). Close each strip and flick to mix well.
17. Pulse-spin each strip and place on a magnetic rack

4. ExoI treatment

This is to remove any untagmented oligos on the beads

18. Prepare ExoI master mix on ice

    ExoI Master Mix

    Vol/sample	Reagent
    17 μL	Nuclease-free water
    2 μL	10x rCutSmart buffer (NEB)
    1 μL	Thermolabile ExoI (NEB)
    20 μL	Total

    Prepare excess master mix, e.g. 1.1 * N samples, or more if required for liquid handling dead volume

19. Prepare a multichannel reservoir with 1xWASH+T buffer

20. Remove the supernatant from the beads (on magnet) and discard. On magnet, add 100μl WASH+T

    • optional If desired, beads can be ‘downsampled’ to generate a library for only a portion of the bead-bound tagmented fragments. Downsampling will generate a library with fewer fragments and lower complexity, but can increase the percent of linked reads at lower sequencing depths. The remaining tagmented beads can be stored at 4°C and processed later if desired.

21. To downsample beads: remove the strip from the magnet, flick or mix to disperse the beads, and transfer the desired portion to a new strip or tube and return to magnet. The equivalent downsampling can also be done at the wash step after ExoI treatment and before PCR.

22. Remove the supernatant from the beads (on magnet) and discard

    • If processing many strips, either close them to prevent the beads from drying out while washing all of the strips, or add the ExoI master mix to each strip before continuing with the next strip.

23. Remove the strip from the magnet and add 20 μl ExoI master mix to each sample. Close the strip and flick to mix. Pulse-spin very briefly to ensure all of the liquid is at the bottom. The beads should remain fully dispersed.

24. Incubate samples in a thermoscycler: 10min at 37°C, then 5min at 65°C (heat kill)

    • If possible, set the lid temperature to 65°C.

4. PCR Setup

The last part of the protocol is to attach Universal Dual Index (UDI) adapters/indices to the DNA via PCR. After this, all that’s left is a final bead cleanup.

25. Thaw hybrid UDI stocks (mix of TruSeq-i5 + Nextera-i7, 5nM each) on ice. Pulse-spin when thawed.
26. Prepare PCR master mix on ice with excess, e.g. 1.1 * N samples, or more if required for liquid handling dead volume.

    PCR Master Mix

    Vol/sample	Reagent
    12.5 μL	2x HiFi MM (NEB #M0541)
    7.5 μL	NF water
    20 μL	Total

27. Program a thermocycler as indicated below, including an initial pause step at 72°C. Start the thermocycler so that the block pre-heats to 72°C.
    • When the ExoI incubation is complete, pulse-spin each strip and place on a magnetic rack.
    • The samples are stable on the magnetic rack at room temperature.

    Caution

    Do not proceed until:

    • the supernatant is clear
    • the PCR master mix is prepared
    • the UDI primers are thawed
    • the PCR program is ready
28. Set a (multichannel) pipet to 100ul. One strip at a time on magnet, add 100μl WASH+T (no need to remove the ExoI reaction first – just add WASH+T to the tube). If the beads come off the magnet, pause until the supernatant is clear. With the same tips, remove the entire supernatant (~120 ul).

    Note

    If processing many strips, close them to prevent the beads from drying out while washing all of the strip

    Optional: Bead Downsampling

    To downsample beads, remove the strip from the magnet after adding WASH+T buffer, and mix to fully disperse the beads. Transfer the desired volume to a new strip and place on a magnetic rack. When the supernatant is clear, remove the supernatant. The remaining tagmented beads can be stored at 4° C and processed later if desired.

29. Move the strips containing the bead pellets to a chiller block or a rack on ice.

    Note

    The PCR mix is not a hot-start formulation, because the tagmented ends need to be extended before the first denaturation step. Keep all PCR reagents and the samples on ice while setting up the PCR reaction, and transfer directly to a pre-heated PCR block to minimize polymerase activity below 72°C.

30. On a chiller block or on ice, add 20 μl 2x HiFi Master Mix per sample.
31. On a chiller block or on ice, add 5 μl hybrid UDI stock (TruSeq-i5 + Nextera-i7, 5nM each) to each sample.

    Note

    Important: Record which UDI wells are used for each sample.

Thermocycler Program

• Initial pause: 72°C preheat and hold at temperature
• Initial incubation: 72°C for 5min extend tagmentation gaps
• Initial denaturation: 98°C for 30sec
• 12* Cycles of:
  • 98°C for 10sec
  • 65°C for 30sec
  • 72°C for 1min
• Final extension: 72°C for 5min

*PCR cycle number may need to be adjusted based on beads and DNA input.

32. Recommended put away all pre-PCR reagents before moving to a different bench for post-PCR cleanup.

5. Post-PCR cleanup and quality checks

33. Prepare fresh 80% ethanol (500μl per sample)
34. Place PCR reactions on post-PCR magnetic rack
35. Set up new strips with 22.5 μl DNA cleanup beads (0.9x)
36. Transfer PCR reaction to new strips containing DNA cleanup beads
37. Vortex to mix well, incubate at least 5min at room temperature
38. Pulse-spin and transfer strip to magnet
39. On magnet, wash 2x with 200 μl fresh 80% ethanol
    • For each wash, pipette up and down (on magnet) to rinse the beads, then leave the ethanol wash on the beads for minimum 30sec
40. After 2nd wash: pulse spin and remove residual wash
41. Allow beads to dry briefly, only until droplets are gone (beads should still be dark and glossy)
42. Remove from magnetic rack and resuspend beads in 23μl 0.1x TE
43. Incubate at least 5 min at room temperature, transfer to magnetic rack until clear
44. Transfer supernatant to a clean, labelled strip tube or plate. This is the final BLink-seq library

Optional: quantify and check the size distribution for all or a subset of samples

• Prepare tubes for Qubit assay: add 198 μl 1x HS dsDNA Qubit reagent, hold in the dark at room temperature until equilibrated.
• Transfer 2μl final library stock to Qubit tubes. Vortex, pulse-spin, and incubate 2 min at room temperature.
• Read and record Qubit measurements.
• Prepare aliquots for size QC, dilute if needed to be in the target concentration range. Final library size can be estimated by gel electrophoresis or using an instrument such as the Fragment Analyzer (Agilent) or equivalent. A successful library will typically have a size range from ~300-1500bp, with 40% or more in the target size range of 450-1200bp. To adjust the size range in the next experiment, alter the amount of ME-B transposome used (more ME-B transposome = smaller insert sizes and vice versa). Libraries with less material in the target size range can still be sequenced, but the amount of linked reads may be lower and/or the insert size may be smaller than ideal (<450bp).

Pooling and sequencing

• Pool final libraries, based on QC metrics (Qubit concentration and size distribution) when available. If the final pool will be size-selected, plan the pool based on molarity estimates for the target size range.
• Recommended: size-select the final pool (450-1200bp) on a Pippin instrument (Sage Science) in order to optimize insert sizes prior to sequencing.
• Sequence on an Illumina sequencer, recommended 2x150 PE reads with dual index reads. Index read lengths should match the UDI primers used.

Analysis

To process your BLink-seq data, use Harpy. Harpy includes several modules for preprocessing reads, aligning reads, calling variants (SNPs and SVs), and more.