TruSeq Total Transcriptome Library Prep (Legacy)

Day 1

Documentation

TruSeq Stranded Total RNA Sample Prep Guide. This method makes a cDNA library of all RNA molecules present in your sample after rRNA depletion.

the success of this kit is dependent on the ability of Illumina’s rRNA depletion beads to bind rRNAs in your target species. Check this list for compatibility of the mouse/rat/human rRNA ‘riboGold’ depletion reagent with other species.

What you’ll need for Day

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The following items are included in the Illumina kit
  • rRNA removal beads (These are the beads for rRNA depletion) (bring these to room temperature before use)
  • rRNA removal mix
  • rRNA binding buffer
  • Elute, Prime, Fragment High mix
  • First strand synthesis (in a brown tube)
  • Second strand marking mix
  • Resuspension buffer
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The following items are NOT included with the Illumina kit and must be purchased separately
  • RNase-free water
  • Magnetic plate for a 96-well tra
  • 96-well trays
  • 80% ethanol (make this fresh the day of use)
  • Superscript II (Invitrogen)
  • RNAClean XP beads (Beckman-Coulter) (bring these to room temperature before use)
  • AMPureXP beads (Beckman-Coulter)

A few important comments before you start

  • When putting plate into the PCR machine, use the plastic film to seal; if not the samples can overflow into other wells
  • We use the Low Sample ‘LS’ protocol as we are typically working with fewer than 48 samples at a time
  • The recommended starting amount of total RNA is 100 ng - 1 ug, but we usually try to stay away from the extreme ends of this spectrum.
  • Split this protocol into two days, stopping on the first day after you have double stranded cDNA. On second day, do the A-tailing, adapter ligation, PCR, cleanup, and quality control steps.
  • For 12 samples, you will need approximately 6-7 hours on the first day, and 6-7 hours the second day.
  • Keep all reagents on ice unless otherwise stated.
  • We typically remove the reagents from storage during an incubation period in the previous step.
  • Have the beads at room temperature at least 30 minutes prior to use.
  • Aliquot the reagents that arrive in large quantities (such as the resuspension buffer and the bead washing buffer) into 2 mL Eppendorf tubes.
  • We do not perform any of the in-line controls.
  • We do not use the plate barcode stickers that come with the kit.
  • Use only filter pipette tips and clean your area so it is free of RNases.
  • Ensure that the AMPureXP beads are mixed well immediately prior to use.
  • There are many mixing steps in this protocol - I frequently use a multi-channel pipette to help speed up this process, particularly if many libraries are being prepared.
  • We use a Tapestation or a BioAnalyzer to assay the input RNA quality. Ribosomal integrity numbers of 7 and higher are preferred.
  • We quantify the input RNA using a Qubit fluorometer.

Step 1: Deplete rRNA

Dilute the RNA with RNase-free water to a final volume of 10 uL per sample.
Add 5 uL of rRNA binding buffer and 5 uL of rRNA removal mix to each well. Pipette up and down 6 times.
Seal the plate and place the plate in the thermocycler and run the “RNA DENATURATION PROGRAM” (68C for 5 min).
Remove the plate from the thermocycler and leave on the bench at room temperature for 1 minute.
Vortex the rRNA removal beads (Illumina) and add 35 uL of beads to each well of a new plate.
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Don’t skip this step by adding the beads to the sample in the old plate. Adding the sample to the beads will ensure optimal performance.
Remove the adhesive seal from the plate and add the sample (20 uL each) to the beads. Quickly pipette up and down 20 times to mix thoroughly.
Incubate at room temperature for 1 minute.
Place the plate on the magnetic stand for 1 minute
Transfer the supernatant to a new plate.
Vortex the RNAClean XP beads to resuspend. Add 99 ul of beads directly to the ribo-depleted RNA samples from above. Mix gently by pipetting 10 times.
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If your starting RNA sample is degraded, use 193 uL RNAClean XP beads.
incubate at RT for 15 min.
Place the plate on the magnetic stand at RT for 5 min.
Remove and discard the supernatant.
With the plate still on magnet, add 190 ul of fresh 80% EtOH to each well without disturbing the beads.
🔥
The TruSeq manual indicates that 200 uL should be used, however, the maximum volume of our 96-well plates is 200 uL, thus we use less so the wells do not overflow.
Incubate EtOH wash for 30 sec, then discard the supernatant.
Use a p10 pipette to remove residual ethanol. Keep the plate on the magnet at RT until dry, usually less than 5 minutes. When dry, the beads will appear matte and cracked. Remove plate from magnet when all samples are dry.
Remove the plate from the magnet, gently add 11 ul of Elution Buffer to each well. Mix thoroughly by pipetting up and down 10 times. You will have to work to get the dried beads off the side of the well and resuspended in the buffer.
Incubate the plate at RT for 2 minutes.
Place the sample on the magnet for 2 minutes.
Transfer 8.5 uL of the supernatant from each well to a new PCR plate to continue with Step 2 below.
There are ~2.5 uL of supernatant still in old plate with the beads, use this to do RNA High Sensitivity TapeStation to check if the rRNA is depleted.
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If the rRNA is not depleted, go back to Step 1 with RNA samples.

Step 2: fragment RNA

This is a non-enzymatic and uses heat and divalent metal cations (magnesium or zinc) to fragment your RNA. If your starting RNA was degraded, you can shorten the the length of time used in the Elu2Frag Program, or skip the fragmentation step altogether.

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If your RNA is degraded, you may shorten the fragmentation time. See Appendix A in the Illumina TruSeq Total RNA Handbook.
Work with 8.5 uL of the supernatant in the new PCR plate.
Add 8.5 uL Elute, Prime, Fragment High mix to each well. Mix by pipetting up and down 10 times.
Place the wells in the thermocyler and run the Elu2Frag program.
Remove the plate once it reaches 4C, centrifuge briefly, and then proceed immediately through the next steps.

Step 3: 1st strand cDNA synthesis

Remove First Strand Synthesis Act D Mix from -20C and thaw at RT. Centrifuge this reagent at 600 x g for 5 sec.
Add 50 ul SuperScript II to the First Strand Synthesis Act D Mix well. If you’re not going to use the entire well of mix, then add SuperScript at a ratio of 1 ul SuperScript for every 9 ul of First Strand reagent. For example, for 10 reactions, mix 90 ul with 10 ul of SS II. Excess mix can be stored in the -20C freezer.
Add 8 ul of First Strand SuperScript mix to each well containing the mRNA.
Place the plate in thermocycler and run the “Synthesize 1st Strand” program (25C for 10 min, 42C for 15 min, 70C for 15 min, hold at 4C).
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This step takes 40 minutes and is your first large break.

Step 4: 2nd strand cDNA synthesis

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When the first strand synthesis is completed, immediately proceed to the second strand synthesis
Add 20 uL second strand marking mix and 5 uL resuspension buffer to each well. Pipette up and down 6 times.
Centrifuge the plate at 600 x g for 30 sec.
Place plate in thermocycler and run the ‘2nd strand’ reaction - 16C for 1hr, with the lid set to 30C.
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This is your second large break of the day.

Step 5: Reaction clean-up

When the reaction is complete, add 90 ul of well-mixed AMPure XP beads to each well containing 50 ul of double stranded cDNA. Incubate RT for 15 min.
Place the plate on the magnet and let sit for 5 min.
Remove and discard 135 ul per well, being careful not to disturb the beads.
Wash beads on magnet by adding 190 ul of 80% EtOH to each well, without disturbing the beads incubate 30 sec at RT.
Remove and discard the supernatant, repeat the wash step and remove the supernatant again.
Keep the plate on the magnet at RT until dry, usually less than 5 minutes. When dry, the beads will appear matte and cracked. Remove plate from magnet when all samples are dry.
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Use a p10 pipette to remove any residual ethanol from the bottom of the well.
Add 17.5 ul of Resuspension Buffer to each well and pipette up and down 10 time to mix. Incubate for 2 min at RT.
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The beads may be dry and you may have to pipette up and down more than 10 times until the beads are fully resuspended.
Place the plate on the magnetic stand for 5 minutes.
Transfer 15 ul of the supernatant (contains the ds cDNA) to new a new plate.
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This is a safe-stopping place. You can cover the plate and store at -20C until you are ready to continue to the second day of the protocol.
Do not store your DNA at this stage for longer than a week. We usually perform the rest of the protocol the next day.

Day 2

What you’ll need for Day 2

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The following items are included in the Illumina kit
  • A-tailing mix
  • Resuspension buffer
  • Ligation mix
  • Index adapters
  • Stop ligation buffer
  • PCR primer cocktail
  • PCR master mix
📌
The following items are NOT included with the Illumina kit and must be purchased separately
  • Magnetic plate for a 96-well tray
  • 96-well trays
  • 80% Ethanol (make this fresh the day of use)
  • AMPureXP beads (Beckman-Coulter)

Step 6: Adenylate cDNA ends

Add 12.5 ul of thawed A-tailing mix and 2.5 uL of resuspension buffer to each well. Pipette up and down 10 times.
Carry out A-tailing by running the thermocycler ‘ATAIL70’ program.
🛑
This step takes 35 minutes and is the first large break of the day.

Step 7: ligate adapters to ends

Add 2.5 ul of resuspension buffer, 2.5 ul of ligation mix, and 2.5 ul of a unique RNA adapter index to each well, mix well by pipetting.
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Remove the ligation mix immediately before use and return to -20C storage immediately after use.
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I moved individual adapter index tubes to strip tubes to make it easier to add the unique indexes to the wells with a multichannel pipette.
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Make sure to record which sample received which index adapter number.
Seal the plate and incubate at 30C in thermocycler for 10 min using the ‘LIGADAPTER’ program.
Remove the plate from the thermocycler and add 5 ul of stop ligation buffer to each reaction, mix well by pipetting.
Add 42 ul of AMPure XP beads to each reaction, mix well by pipetting, and incubate at RT for 15 min.
Place on magnetic stand for 5 min at RT.
With the plate on the magnetic stand, remove 79.5 ul of supernatant from each sample, being careful not to disturb the beads.
With the plate on the magnetic stand, wash with 190 ul 80% EtOH for 30 sec, then remove the ethanol.
Perform the wash step again.
Use a p10 pipette to remove residual ethanol. Keep the plate on the magnet at RT until dry, usually less than 5 minutes. When dry, the beads will appear matte and cracked. Remove plate from magnet when all samples are dry.
Remove the plate from the magnetic stand and add 52.5 ul of Resuspension Buffer to each well, and mix well by pipetting. Incubate at RT for 2 min
Place the plate on magnet for 5 min.
Recover 50 ul from each well and transfer to a new plate.
Carry out a second cleanup round. Add 50 uL of mixed AMPure beads to each well, mix well, incubate for 15 min at RT.
Place the plate on the magnetic stand for 5 min.
With the plate on the magnetic stand, remove and discard 95 ul of supernatant from each sample.
With the plate on the magnetic stand, wash the beads with 190 ul of 80% EtOH for 30 s, then remove the ethanol.
With the plate on the magnetic stand, wash the beads again with 190 uL of 80% EtOH for 30 s, then remove the ethanol.
Use a p10 pipette to remove residual ethanol. Keep the plate on the magnet at RT until dry, usually less than 5 minutes. When dry, the beads will appear matte and cracked. Remove plate from magnet when all samples are dry.
Resuspend beads in 22.5 ul of resuspension Buffer, mix well by pipetting, and incubate 2 min at RT.
Transfer the plate to the magnetic stand and incubate for 5 min.
Recover 20 ul from each well and transfer to a new plate.

Step 8: PCR amplify library

Add 5 ul of PCR primer cocktail and 25 ul of PCR master mix to each well, place the plate in the thermocycler and run the “PCR” program.
🛑
This is another break for 30 minutes.
Add 50 ul of AMPure beads to each well and mix well by pipetting. Incubate 15 min at RT.
Place the plate on the magnet for 5 min.
Remove and discard 95 ul of supernatant from each well.
Wash the beads 190 ul/well of 80% EtOH, wait 30s, remove the supernatant.
Repeat the ethanol wash step.
Use a p10 pipette to remove residual ethanol. Keep the plate on the magnet at RT until dry, usually less than 5 minutes. When dry, the beads will appear matte and cracked. Remove plate from magnet when all samples are dry.
Remove the plate from the magnetic stand, add 32.5 ul of resuspension buffer and mix well, by pipetting, and incubate for 2 min at room temperature.
Place the plate back on the magnetic stand for 5 minutes.
Remove 30 uL of supernatant from each sample and move it into a new plate. This is the final cDNA product.
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Congratulations! You’re done!…well, almost
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Perform quality control steps - we use a Tapestation to determine the library size and qubit to quantify the libraries.
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We recommend sequencing the libraries within 3 weeks.

Sequencing Guidelines

Normalize and Pool

  1. Quantify each of your libraries on Qubit. For most libraries, using the HS dsDNA Qubit assay with 2uL of input will yield a reading. Record the concentration in ng/uL for each library.
  2. Run your samples on Tapestation with either the D1000 or the HSD1000 assay. Remember to allow Tapestation reagents to sit at room temperature for at least 30 minutes before use. Save your Tapestation results by going to File -> Create Report -> Save as pdf. This file can then be emailed or uploaded to Asana. For the base pair length, we usually use the value of the peak identified by the Tapestation analysis software. This value is shown both on the tracing itself and in the Peak Table for each sample.
  3. Download our nM Conversion Calculator here. Enter the concentration (from Qubit) and the base pair length (from Tapestation) in the appropriate cells and it will give you the nM concentration for each library. Normalize and pool all your libraries to 4, 2, 1, or 0.5 nM in a LoBind microcentrifuge tube. If you need to dilute your libraries, we recommend using at least 2uL to minimize pipetting errors. The example sheet of the calculator provides further detail.
  4. Quantify your pool on Qubit and enter into the calculator sheet to check that your pool is close to the nM concentration you normalized to.

Setup Run in Basespace

  1. Sign into Basespace, then go to the Prep tab, Biological Samples, and select Import Samples on the upper right. Use Illumina’s Sample Import Template to enter information about your samples. The SampleID and Name can be the same, but make sure they are unique for each sample. Species can be left blank. Upload the completed .csv to import your samples.
  2. Continue to Prep Libraries. Select your library prep kit based on the index format used. If you used our plate indexes select “IDT-ILMN TruSeq DNA-RNA UD 96 Indexes.” If you used another index format, you will need to use a different entry for library prep kit. The your project name as the Plate ID. For each sample, check the box next to it on the left, then drag the sample name to the appropriate index well.
  3. Proceed to Pool Libraries. Select all your samples on the left, then drag and drop in the pool on the right. Name the pool your project name.
  4. Continue to Plan Run. Select NextSeq and name your run your project name. Select Single Read or Paired End Read, then enter the cycle numbers based on your selected kit. For example, if you were doing a run using a High Output 75 Cycle kit, you would select Single Read and enter 76 for Read 1 Cycles and 0 for Read 2 Cycles.
  5. Press Sequence to complete planning the run. The run will now be available for selection on the sequencer.

Loading the Sequencer

  • The next step is to dilute and denature the prepared libraries. Illumina’s general guidelines for this on the NextSeq can be found here.
  • Illumina’s system guide for the NextSeq, which covers the sequening workflow, can be found here.
  • Your final loading concentration should be 1.3 - 1.8 pM, with most pools loaded at 1.4 or 1.5 pM.