TITLE: Low-cost Tagmentation Library Prep for low-coverage Illumina sequencing
AUTHORS: Matthew Farnitano, Andrea Sweigart

[DESCRIPTION]
Tagmentation is a popular approach for preparing Illumina sequencing libraries, but kits can be pricey. Using a bulk-prepped and pre-loaded Tn5 tagmentation enzyme, this protocol is a simple, low-cost and high-throughput procedure for prepping libraries. It is designed for multiplexed low-coverage whole genome sequencing, a useful method for genotyping of many samples at low per-sample cost. Samples are prepped in 96-well PCR plates and combined into multiplexed batches. We have successfully used this protocol with Mimulus guttatus and Mimulus nasutus wild-collected and greenhouse-grown samples.

[STEPS]
SECTION: Before you start
1. Obtain Tn5 tagmentation enzyme, pre-loaded with universal adapter sequences. Enzyme purification and loading guidelines are not provided here: see reference Lu et al. (2017). 
SECTION: Before you start
2. Quantify input DNA. We recommend 96-well fluorescence quantification with a plate reader and a kit such as the Invitrogen Quant-iT dsDNA Assay kit, Catalog #Q33120. 
SECTION: Before you start
3. Dilute DNA to approximately 1 ng/uL per sample in molecular-grade H2O. The required input is 1 uL of diluted DNA; we have had success with less than 1 ng on some samples, but final read coverage is correlated with initial sample concentration. If sample concentration is already relatively low (<5 ng/uL), we typically do not dilute. We dilute samples in batches of similar concentration, erring on the side of higher final concentrations. For example, all samples at initial concentration of 5-15 ng/uL might be diluted by 1/5. 
SECTION: Tagmentation
5. Set up tagmentation reaction
SECTION: Tagmentation
5.2. Create reagent Master Mix:
 

SECTION: Tagmentation
5.3. Add 4 uL Master Mix to each well of diluted DNA in 96-well PCR plate (final volume 5 uL)
SECTION: Tagmentation
5.1. Prepare a 96-well PCR plate with 1 uL diluted DNA (~1 ng/uL) per sample well
SECTION: Tagmentation
6. Run tagmentation reaction in thermocycler:

 
Once temperature has reached 10C, quickly move to the next step. Keep the reaction on ice!
SECTION: PCR
7. Set up PCR Reaction
**Samples and PCR reagents should remain on ice as much as possible during PCR setup**
SECTION: Before you start
4. Before starting tagmentation, pull out the following to thaw:
Tagmentation (TD) buffer
Taq Master Mix (for PCR step)
i5 and i7 indices (for PCR step) -- prepare working dilutions if necessary

i5 indices are in individual tubes at 100 uM -- they do not need to be diluted (optionally, dilute to 10uM). Use a separate i5 index for each plate of 96 samples, one index per plate. 
i7 indices are in a 96-well plate, the stock plate is at 100uM. Make a working dilution plate of 10uM.
Use this same i7 plate for all sample plates, one well per sample. 

NOTE: our i5 and i7 index adapter sequences are included in attached files to this protocol. 

SECTION: PCR
8. Run PCR reaction in thermocycler:

 

After PCR, plates can be held at 4C overnight if necessary. If leaving overnight, do NOT combine samples yet! Once combined, proceed immediately to bead cleanup.
SECTION: PCR
7.1. Prepare PCR Master Mix (one mix per plate):

 

*OneTaq Hot Start 2X Master Mix, New England Biolabs, Catalog #M0484L
**Optionally, dilute stock to 10uM and add 10X listed volume (adjust H2O amounts accordingly)

SECTION: PCR
7.2. Add 0.5 uL i7 adapters, 10uM, (96-well plate format) to each tagmentation sample well using multichannel pipet
SECTION: PCR
7.3. Add 19.5 uL PCR Master Mix to each tagmentation sample well. 
To allow for multichannel pipetting, either move PCR Master Mix to a thin trough, or divide evenly into 8-count strip tubes, 263uL per row of 12 samples.
SECTION: Pooling and bead cleanup
10. Pool 48 samples (cols 1-6, or cols 7-12 from one plate) into a single batch for cleanup.
SECTION: Pooling and bead cleanup
10.1. Pipette 20 uL from each sample well into a single microcentrifuge tube (~960 uL total)
SECTION: Pooling and bead cleanup
10.2. Mix well with a P1000 pipet and transfer half (~480 uL) to a second tube. Proceed with cleanup of both tubes. You will have two tubes per 48-sample batch (4 tubes per 96-well plate). 
SECTION: Pooling and bead cleanup
11. Use SPRI magnetic beads and magnetic tube rack to clean up PCR products. 
SECTION: Pooling and bead cleanup
9. Bring SPRI magnetic beads* to room temperature for cleanup, and resuspend fully using a vortexer.
*We have used "CleanNGS DNA & RNA Cleanup Magnetic Beads" from Bulldawg Bio, Catalog #CNGS050
SECTION: Pooling and bead cleanup
11.1. Add 1.5 times sample volume of SPRI beads to each tube (720 uL SPRI beads per 480 uL sample tube).
Gently pipet up and down 5-10 times to mix thoroughly.
SECTION: Pooling and bead cleanup
11.2. Incubate 5 mins at room temperature on tabletop. 
SECTION: Pooling and bead cleanup
11.3. Place tubes on magnetic tube rack. Wait 5 mins. Beads should collect on side of tube, leaving a clear supernatant. Your DNA will be bound to the magnetic beads at this point.
SECTION: Pooling and bead cleanup
11.4. **WHILE TUBES REMAIN ON MAGNET:
Discard clear supernatant with a pipet, taking care not to disturb beads
SECTION: Pooling and bead cleanup
11.5. **WHILE TUBES REMAIN ON MAGNET:
Pipet 500 uL fresh 70% Ethanol directly onto beads. Let sit for 1 min. (Wash #1)
SECTION: Pooling and bead cleanup
11.6. **WHILE TUBES REMAIN ON MAGNET:
Discard clear supernatant with a pipet, taking care not to disturb beads
SECTION: Pooling and bead cleanup
11.7. **WHILE TUBES REMAIN ON MAGNET:
Pipet 500 uL fresh 70% Ethanol directly onto beads. Let sit for 1 min. (Wash #2)
SECTION: Pooling and bead cleanup
11.8. **WHILE TUBES REMAIN ON MAGNET:
Discard clear supernatant with a pipet, taking care not to disturb beads
SECTION: Pooling and bead cleanup
11.9. **WHILE TUBES REMAIN ON MAGNET:
Let beads air-dry until they have a matte appearance, with slight cracking. If necessary, use a P20 or P200 to remove excess liquid drops without disturbing beads.
SECTION: Pooling and bead cleanup
11.10. Once dry, remove tubes from magnet. 
Pipet 17 uL molecular-grade H2O directly onto beads. Gently pipet liquid back onto beads repeatedly, massaging the beads until they resuspend. You should end up with a single aliquot of uniform brown liquid at the bottom of the tube, with no beads stuck to the side.
SECTION: Pooling and bead cleanup
11.11. Incubate 3 mins at room temperature on tabletop.
SECTION: Pooling and bead cleanup
11.12. Place tubes back onto the magnetic tube rack. Wait 5 minutes for beads to collect on the side of the tube, leaving a clear supernatant. YOUR DNA IS NOW IN THE CLEAR SUPERNATANT.
SECTION: Pooling and bead cleanup
11.13. **WHILE TUBES REMAIN ON MAGNET:
Carefully pipet ~15 uL of the clear supernatant, which contains your DNA, into a fresh labelled tube. Do not disturb or pipet up any of the brown beads. You may need to leave a small amount (1-2 uL) behind with the beads. If the beads are accidentally disturbed, return the liquid to the tube and let the beads settle onto the magnet again before re-trying. 
At this point, the two replicate tubes from each 48-sample batch can be combined, for a final volume of 30 uL per batch of 48 samples. 
SECTION: Pooling and bead cleanup
12. The library prep is now complete. We use a Qubit fluorometer to measure the concentration of each 48-sample batch, then combine equal ng amounts into a single tube to submit for sequencing.