For instructions on how to perform blood extraction using a Chemagic 360-D, see pages 21 – 28 in the Revvity instructions for use document.

• Elute your extracted gDNA using 100 µl of Elution Buffer 7 per sample.

Optional: If struggling with sample viscosity or incomplete resuspension during elution, incubate your samples at 50°C on a thermomixer with gentle agitation 300 RPM to ensure homogeneous resuspension.

Note: Approximately 5–12 µg of gDNA is expected following sample extraction.
Expected Qubit measurements of 50–120 ng/μl.

Seal the plate completely with a PCR plate seal.

This plate serves as the sample shearing plate.

Seal the plate completely with a PCR plate seal.

Secure both plates tightly using the adjustable clamp.

Example of a ~15kb N50 fragmented Chemagic gDNA trace in ProSize, showing the smear analysis lines (red) displaying the FP signal <10kb.​

For optimal performance, NEB recommend the following:

1. Thaw all reagents on ice.

2. Flick and/or invert the reagent tubes to ensure they are well mixed.
   Note: Do not vortex the FFPE DNA Repair Mix or Ultra II End Prep Enzyme Mix.

3. Always spin down tubes before opening for the first time each day.

4. The Ultra II End Prep Reaction Buffer and FFPE DNA Repair Buffer may have a little precipitate. Allow the mixtures to come to room temperature and pipette the buffer up and down several times to break up the precipitate, followed by vortexing the tube for 30 seconds to solubilise any precipitate.
   Note: It is important the buffers are mixed well by vortexing.

5. The FFPE DNA Repair Buffer may have a yellow tinge and is fine to use if yellow.

We recommend adding some excess volume (~10%) when preparing your master mix for multiple samples to account for pipetting volume loss.

Ensure the master mix is fully mixed by pipetting full volume ten times. Do not vortex the mix.

Tip: The prepared master mix can be evenly aliquoted into an 8-tube strip, enabling the use of multichannel pipettes to dispense into your sample plate.

Allow enough for 200 µl of fresh 80% ethanol per sample.

Take care not to disturb the pelleted beads.

Dispose of the bead-containing plate.

• Thaw the reagent at room temperature.
• Spin down the reagent tube for 5 seconds.
• Ensure the reagent is fully mixed by performing 10 full volume pipette mixes.

Note: Do NOT vortex the Blunt/TA master mix or the Salt T4-DNA Ligase.

• Remove the sample plate from the magnet and resuspend each sample pellet in 120 µl L Fragment Buffer (LFB) by pipetting.
• Return samples to the magnet and pellet the beads on the magnet for at least 3 minutes or until the supernatant is clear and colourless (whichever is longer).
• Keep the samples on the magnet and aspirate the supernatant, taking care not to aspirate any beads.

Take care not to disturb the pelleted beads.

Dispose of the bead-containing plate.

Keep a note of the quantification measurements.

1. Line up the flow cell with the connector horizontally and vertically before smoothly inserting into position.
2. Press down firmly onto the flow cell and ensure the latch engages and clicks into place.

1. Set a P1000 pipette tip to 200 µl.
2. Insert the tip into the inlet port.
3. Turn the wheel until the dial shows 220-230 µl, or until you see a small volume of buffer entering the pipette tip.

Note: Library loading volume has been increased to improve array coverage.

1. Align the inlet port cut out of the light shield with the inlet port cover on the flow cell. The leading edge of the light shield should sit above the flow cell ID.
2. Firmly press the light shield around the inlet port cover. The inlet port clip will click into place underneath the inlet port cover.

Importantly, ensure that respective barcodes are correctly assigned to corresponding wells of sample-containing 96-well PCR plate.

The samples are automatically ranked from the highest to the lowest facilitating splitting into four groups (“pooling groups”) of descending output.

1. Enter the flow cell type that you will use for sequencing:

• For PromethION Flow Cells enter the product code FLO-PRO114M

2. Enter the QC Pool Concentration measurement obtained during the "Sample QC Pool preparation" step.

The volumes for each sample to be pooled, and the required volume of Elution Buffer (EB) needed, will display on this tab.

The amount of any Elution Buffer (EB) to add to a pooled group is indicated in the library pooling calculator. A minimum value is stated in order to bring the total volume of the pool to 5% over the total volume needed for the required number of flow cell loads.

Please note, the highest concentration of Pooled Groups tested during kit verification was 11.25 ng/µl (equating to ~360 ng of library per flow cell load). The predicted Pooled Group concentration value is indicated on the library pooling calculator. If the value obtained is above 11.25 ng/µl, increase the amount of Elution Buffer needed (“EB to add” cell).

It should also be noted that a minimum of 100 ng of pooled library per flow cell load is recommended, equating to a final Pooled Group concentration of 3.125 ng/µl (as 32 µl of the Pooled Group is used per load). If the lowest‑output sample(s) within a group is/are particularly low, the subsequent pooling of lower volumes of all other samples could result in a final concentration below this lower limit.

If the predicted Pooled Group concentration value is lower than 3.125 ng/µl, it becomes necessary to omit the lowest-performing sample from the pool. To do this:

• Change the respective “Omit sample?” entry from “N” (No) to “Y” (Yes).
• Continue to omit the lowest-performing sample within a group in this way until the Pooled Group concentration reaches at least 3.125 ng/µl.

Conditional formatting within the library pooling calculator is in place to guide the process of forming the Pooled Group at the correct concentration:

• An amber value indicates that more Elution Buffer may be needed as the concentration rises above the verified range.
• A red value indicates that sample omission is required.

Omission of samples from a pooled group automatically adjusts the default number of flow cells needed for the experiment, and the total volume of library required is adjusted accordingly. For the PromethION Flow Cell (FLO-PRO114M) the number of flow cells to be used must match the number of samples that pass the QC step.

The "Manual Pooling Template" tab in the library pooling calculator summarises the volumes of samples/Elution Buffer (EB) for each Pooled Group.

In separate clean 1.5 ml Eppendorf DNA LoBind tubes combine the indicated volumes of barcoded and adapted samples, and Elution Buffer (EB), for the separate pooled groups.

Ensure the pooled groups are well mixed by pipetting ten times at full volume.

If already stored at 4°C from the running of the QC flow cell, briefly spin down the regents and store on ice.

Note: these volumes include an excess. For preparing a master mix, these volumes can be scaled by the number of flow cells to be loaded.

1. Line up the flow cell with the connector horizontally and vertically before smoothly inserting into position.
2. Press down firmly onto the flow cell and ensure the latch engages and clicks into place.

1. Set a P1000 pipette tip to 200 µl.
2. Insert the tip into the inlet port.
3. Turn the wheel until the dial shows 220-230 µl, or until you see a small volume of buffer entering the pipette tip.

1. Align the inlet port cut out of the light shield with the inlet port cover on the flow cell. The leading edge of the light shield should sit above the flow cell ID.
2. Firmly press the light shield around the inlet port cover. The inlet port clip will click into place underneath the inlet port cover.

1. Close the inlet port.
2. Insert a P1000 pipette into a waste port and remove the waste buffer.

Note: As both the inlet port is closed, no fluid should leave the sensor array area.

1. Set a P1000 pipette to 200 µl
2. Insert the tip into the inlet port
3. Turn the wheel until the dial shows 220-230 µl, or until you can see a small volume of buffer entering the pipette tip.

1. Using a P1000 pipette, take 200 µl of the flow cell wash mix
2. Insert the pipette tip into the inlet port, ensuring there are no bubbles in the tip
3. Slowly twist the pipette wheel down to load the flow cell (if possible with your pipette) or push down the plunger very slowly, leaving a small volume of buffer in the pipette tip.
4. Set a timer for a 5 minute incubation.

1. Using a P1000 pipette, take 200 µl of the flow cell wash mix
2. Insert the pipette tip into the inlet port, ensuring there are no bubbles in the tip
3. Slowly twist the pipette wheel down to load the flow cell (if possible with your pipette) or push down the plunger very slowly, leaving a small volume of buffer in the pipette tip.

1. Ensure the inlet port is closed.
2. Insert a P1000 pipette into a waste port and remove the waste buffer

Note: As the inlet port is closed, no fluid should leave the sensor array area.

If already stored at 4°C from the running of the QC flow cell, briefly spin down the regents and store on ice.

Note: these volumes include an excess. For preparing a master mix, these volumes can be scaled by the number of flow cells to be loaded.

1. Set a P1000 pipette tip to 200 µl.
2. Insert the tip into the inlet port.
3. Turn the wheel until the dial shows 220-230 µl, or until you see a small volume of buffer entering the pipette tip.

2. Using a P1000 pipette, take 500 µl of the priming mix
3. Insert the pipette tip into the priming port, ensuring there are no bubbles in the tip
4. Slowly twist the pipette wheel down to load the flow cell (if possible with your pipette) or push down the plunger very slowly, leaving a small volume of buffer in the pipette tip.

During this time, prepare the library for loading using the next steps in the protocol.

1. Ensure the inlet port is closed.
2. Insert a P1000 pipette into a waste port and remove the waste buffer

Note: As the inlet port is closed, no fluid should leave the sensor array area.

1. Set a P1000 pipette tip to 200 µl.
2. Insert the tip into the inlet port.
3. Turn the wheel until the dial shows 220-230 µl, or until you see a small volume of buffer entering the pipette tip.

2. Using a P1000 pipette, take 500 µl of the priming mix
3. Insert the pipette tip into the priming port, ensuring there are no bubbles in the tip
4. Slowly twist the pipette wheel down to load the flow cell (if possible with your pipette) or push down the plunger very slowly, leaving a small volume of buffer in the pipette tip.

1. Close the inlet port.
2. Insert a P1000 pipette into a waste port and remove the waste buffer.

Note: As both the inlet port is closed, no fluid should leave the sensor array area.

1. Set a P1000 pipette tip to 200 µl.
2. Insert the tip into the inlet port.
3. Turn the wheel until the dial shows 220-230 µl, or until you see a small volume of buffer entering the pipette tip.

1. Align the inlet port cut out of the light shield with the inlet port cover on the flow cell. The leading edge of the light shield should sit above the flow cell ID.
2. Firmly press the light shield around the inlet port cover. The inlet port clip will click into place underneath the inlet port cover.

Instructions for returning flow cells can be found here.