Using the Illumina Nextera XT kit at 1/3 reagents

In a genomics lab, if you need to work with really low input DNA samples you can use the Illumina Nextera XT kit. That being said, the kit is rather expensive and can be adapted to work perfectly fine by using 1/3 reagents for your reactions! You'll get pretty much the same results just make sure you keep the ratio of volumes consistent. It's also got dual indexing and can be used for 96 samples! You can find the original protocol here (https://emea.support.illumina.com/content/dam/illumina-support/documents/documentation/chemistry_documentation/samplepreps_nextera/nextera-xt/nextera-xt-library-prep-reference-guide-15031942-05.pdf) and buy the kit here https://www.illumina.com/products/by-type/sequencing-kits/library-prep-kits/nextera-xt-dna.html

You will need to use 1.25 uL of input/template for this protocol. Make sure that the input is <1 ng. Quantify it with a Qubit fluorometer using the manufacturer's protocol. If it is above that amount, dilute it to be <1ng in 1.25 uL.

After thawing, ensure all reagents are adequately mixed by gently inverting the tubes 3–5 times or pulse vortexing for 3 seconds, followed by a quickspin in a microcentrifuge. Make sure you work on ice or a cold plate.

Prepare the Tagmentation Master Mix by adding 2.5 uL of Tagment DNA Buffer and 1.25 uL of Amplification Tagment Mix per sample. Mix by pulse vortexing for 3 seconds, followed by a quickspin in a microcentrifuge.

  1. Pipette 3.75 uL of the Tagmentation Master Mix into nuclease-free PCR tubes.
  2. Pipette 1.25 uL of your sample into their respective PCR tubes.
  3. Flick the tubes to mix the contents thoroughly and quickspin to collect the contents of the tubes.
  4. Place them in a Thermal Cycler for 5 minutes at 55 ̊C and hold at 10 ̊C. (Should you desire to obtain smaller fragments, you can increase the incubation time to a maximum of 10 minutes)
  5. When the samples reach 10 ̊C, immediately remove them and add 1.25 uL of NT Buffer to each sample. You now have a total of 6.25 uL in each tube. The NT Buffer neutralizes the tagmented samples.
  6. Pipette 3.75 uL of the Nextera PCR Master Mix (NPM) into each tube containing the sample.
  7. Pipette 1.25 uL of N-Series index primer to each sample (If you are multiplexing samples together, be sure to use a unique index primer for each respective sample) For example, the first sample gets index primer n701
  8. Pipette 1.25 uL of S-series Index Primer to each sample (if you are multiplexing samples together, be sure to use a unique index primer for each respective sample). For example the first sample gets index primer S502. Note (Together, the first sample with have 2 indexes, i.e. N701/S502
  9. Flick the tubes to mix the contents thoroughly and spin them down. For each sample you should now have a total volume of 12.5 uL.
  10. Place them in the thermal cycler and perform a PCR amplification using the following program: (17 Cycles is usually safe to do )

72C for 3 mins at 1 cycle

95C for 30 seconds at 1 cycle

**17 cycles for the next 3 steps**

95C for 10 seconds

55C for 30 seconds

72C for 60 seconds

end 17 cycles

72C for 5 minutes

10C HOLD

  1. Prepare your magnetic beads for use by vortexing and calibrating them to room temperature 30 minutes prior to use.
  2. Add 7.5 uL of nuclease-free water to make the volume of the PCR product up to 20 uL.
  3. Add 13 uL of magnetic beads to each sample (0.65X). Mix well by pipetting up and down and incubating the sample-bead mixture at room temperature for 10 minutes.
  4. Place the tube on a magnetic stand for 5 minutes. Carefully transfer the supernatant to a fresh tube without disturbing the beads.
  5. Add 3 uL of properly resuspended beads to the transferred supernatant (0.15x of 20 uL of sample is 3uL). Mix well by pipetting up and down and incubate the sample-bead mix at room temperature for 10 minutes.
  6. Place the tube on a magnetic rack for 5 minutes. Remove and save the supernatant into a fresh tube.

*This is for troubleshooting purposes. Your sample is bound to the beads.

  1. Prepare fresh 80% ethanol (400uL per sample).
  2. While keeping the tubes on the magnet, wash beads with 200 ul of freshly prepared 80% ethanol without disturbing the pellet. With the tube still on the magnet, Remove the 80% ethanol using a pipette and discard. Repeat this step for a total of 2 ethanol washes. Make sure to remove ALL remaining ethanol without uptaking any of the beads.
  3. QuickSpin the tube then place the tube back on the magnet and magnetize for 1 minute at room temperature. Pipette off and discard any residual 80% ethanol. Briefly allow the beads to dry by incubating the tube on the magnet for 2 minutes with the tubes' lids kept open.
  4. Remove the tube from the magnetic rack and add 12 ul of Nuclease-Free water and pipette up and down until beads are in a homogenous suspension.
  5. Place the tube on the magnetic rack and magnetize the beads until the eluate is clear and colourless (the time it takes to appear clear may vary, often takes anywhere between 3-5 minutes).
  6. Remove and retain 12 ul of eluate into a clean PCR Tube. Use caution not to Disturb the beads or carry over into the clean pcr tube.
  7. These are the final, cleaned libraries. To finalize the QC on them, run 1uL of the sample on a DNA HS chip on Agilent's Bioanalyzer using the manufacturer's protocol. Additionally, follow the manufacturers protocol from Invitrogen for quantifying the sample using a Qubit HS DNA assay.

Note: Once looking at the profile, if it is over 900 bp, it means that fragmentation was incomplete. You can reset the Nextera prep but properly quantify the original DNA, and dilute it so that it is less than 1 ng of input into the reaction. This will make the fragmentation more efficient in the initial steps.

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Christopher White4 months ago

This looks interesting.

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