RNAseq library prep

Protocol for tissue-level RNAseq library preparation, multiplexing, submission and data analysis

(A slightly modified protocol based primarily on that used by Josh Gorham in the Seidman lab)


Between 250ng and 1ug of Total RNA from tissues or cells. This protocol starts with 300ng Total RNA.


A dataset capable of comparing genome-wide expression differences between samples

Work Area Setup

Set up TWO basic work stations:

RNA preparation and Pre-Amp station

RNase-Free zone for preparing total and mRNA

Need a small space (about 1.5×2 feet is sufficient)

Equipment needed is: Vortex, mini centrifuge for strips and tubes, two heat blocks (61°C, and 72°C), Magnetic Particle Collectors MPCs (Bar for 1.5ml tubes and plate for 96well plates) for DynaBeads, and PCR machine. Aluminum PCR tube holder. RNase Zap.

Post-AMP work area

Equipment needed is: Separate Magnets for bead isolation (bar and 96well plate), aluminum PCR tube holder, Pipettes

TapeStation Area

TapeStation and accessories

Double stranded cDNA (dscDNA) PREPARATION (day 1)

Before starting

  • heat blocks to 61oC, and to 72o
  • Place Lysis/Binding Buffer and Buffer B at room temperature for 30min to thaw and re-disolve any ppt in solutions.
  • Get a bucket of ice.
  • Equilibrate a pre-amplification aluminum pcr tube holder on ice
  • RNase Free water (200μl per sample)
  • Ensure there are 100 μls of Oligo-dT-DynaBeads per sample and sufficient SuperScript reagents to process all samples.
  • Thoroughly clean down work surfaces with RNase Zap and cover Eppendorf tube racks with aluminum foil.

Pre-amplification station

mRNA purification


  • Dynabeads mRNA DIRECT kit (Invitrogen, cat. no. 610-12) including “Lysis/Binding Buffer” and “Buffer B”
  • Magnetic Particle Concentrator (MPC)(Dynal, Invitrogen, 123-21D)


Bead Prep

  • Vortex provided oligo-dT Dynabeads at low/medium speed for long time (at least 2-3 minutes) to make a homogeneous solution.
  • Aliquot 110μl to of Dynabeads for each sample to be processed to a 1.5ml Eppendorf tube (i.e. 1.1mls for 10 samples) and further homogenize by pipetting.
  • Capture beads with MPC and wash twice with an equal volume Lysis/Binding buffer
  • Resuspend in an equal volume of Lysis/Binding buffer. Then aliquot 50μl beads into two separate 1.5ml tubes per sample. You should have two sets of Dynabeads each of which has one aliquot for each sample.
  • Add a third set of 1.5ml tubes containing 50μl of Lysis/Binding buffer for each sample


RNA Prep

  • Adjust total RNA volume (using 300ngs (to 1μg) as starting material) to 50μl with H2O, add 50μl of Lysis/Binding buffer and immediately place on 61oC heat block for 2 min.
  • Add heated total RNA directly (pipet relatively slowly) to equilibrated DynaBeads and vortex gently.
  • Bind at room temperature for 15 min. with gentle agitation. (During this incubation period prepare the SuperScript III first strand master mix for the next phase of library preparation.)
  • Capture beads on MPC magnet and discard liquid.
  • Wash twice with 200μl Buffer B. To wash, remove Eppendorf tube from magnetic rack, add the wash buffer B and pipet up and down gently until the dyna-beads are in uniform suspension, place the tube immediately back on the magnetic rack. After second wash beads for each step may smear along the back side of the eppi tube differently from the tighter clump that genearally forms. This is normal and accidentally aspirating beads can generally be avoided with selective (gentle) pipeting.
  • Resuspend in 50μl RNase-Free H20 and place immediatly on ice, then vortex gently to resuspend.
  • CRITICAL STEP!! Only process a couple of samples at a time. Elute by incubating at 72oC for 2 min.; immediately capture on Magnet and transfer eluate (~50μl H2O) to a new tube containing 50μl Lysis/binding buffer and add to second tube of dyna-beads to re-capture.

REPEAT ABOVE PROCESS (note that Josh Gorham uses NEW beads for the second purification, I’ve seen good results with using the same beads-which is recommended by the DynaBeads manual)

Elute second round in 12μl RNase-Free H20 as above (note: Josh uses a hard spin and pipetting up and down before 72oC elution of second round: I have been using the same steps as above and see good results) and place directly in a PCR strip on ice containing 3.5μls of SuperScript III First Strand Master mix (below)


cDNA Synthesis:

SuperScript III cDNA synthesis kit (Invitrogen, 18080-051) containing:

  • 10mM dNTPs
  • 0.1M DTT
  • 10x Reverse Transcriptase (RT) buffer
  • RNase OUT
  • RNaseH
  • SuperScript III reverse transcriptase (SS III)
  • Random hexamers (50ng/μl)
  • 25mM MgCl2
  • 5X Second-strand buffer (Invitrogen, cat, no. 10812-014)

DNA polymerase I (Invitrogen, cat. no. 18010025)

AMPure beads (Agencourt AMPure kit, A29152)

Super Script III First Strand Master Mix

Use all 10μl purified mRNA and add as a mastermix (3.5μl/sample):

  • 5μl – random hexamers
  • 1μl – 10mM dNTPs

Incubate above mixture at 65oC for 5 min. and immediately place on ice. Add as a master mix the following (10μl/sample):

  • 2μl – 10x RT buffer
  • 2μl – DTT
  • 4μl – MgCl2
  • 1μl – RNAse out
  • 1μl – SS III

Mix and incubate at a thermocycler using the following program:

  1. 25oC x 10 min.
  2. 50oC x 50 min.
  3. 85oC x 5 min.
  4. 4oC hold

Second Strand

Add as a mastermix the following (80μl/sample):

  • 52μl – H2O
  • 20μl – 5x Second Strand buffer (sold seperatly at BPF)
  • 3μl – 10mM dNTPs
  • 1μl – RNAse H
  • 4μl – DNA polymerase I (sold separatly at BPF)

Mix and incubate at 16oC for 2.5 hours at a thermocycler, then 4°C overnight (optional), store @-20°C.


Before starting

  • Remove HS D1000 tape and buffer from 4oC and equilibrate to room temperature
  • Equilibrate AMPure beads to RT for 30-60min.
  • Ensure there are sufficient AMPure beads (400μl per sample being processed)
  • Ensure sufficient Tagmentation reagents for needed reactions
  • Decide on indexes for each sample such that no two libraries in a multiplexed mixture will have the same indices.

Purify with AMPure XP beads

  • Mix thoroughly and add 3X volume of AMPure beads in supplied PBS (300μls for Second strand mix)
  • Mix gently and incubate for at least 15min at RT
  • Place on magnet and separate beads from solution
  • Aspirate solution and Rinse 2X with Fresh 70% EtOH –DO NOT disturb beads or remove tubes from magnet. Washes are applied over the surface of the bundled beads. Remove every drop of final rinse.
  • Allow 5-15min for the EtOH to dry from the beads but don’t leave so long that beads begin to “crack” from over drying.
  • Remove from Magnet, Add 22μl of H2O (or 10mM Tris-Acetate or Tris-HCl pH8, or TE as needed), pipet 10-20X to re-suspend beads fully
  • Incubate 2min at 37oC
  • Place samples back on magnet and transfer eluate to new tube once beads have separated cleanly.

Measure cDNA concentration via TapeStation (HS-D1000)

Take 1ng of each sample (note: Josh uses 2ngs input cDNA but the Tagmentation protocol emphasizes using 1ng and I’ve seen good results at that input level), and place in a PCR strip in a total volume of 5μl (balanced with H2O) to prepare samples for the next step.

Store samples @-20°C.

Nextera XT DNA sample prep (Illumina)

Use 1ng cDNA input (in 5μl volume).


  1. Add 10μl TD buffer (Illumina)
  2. Add 5μl cDNA (1ng)
  3. Add 5μl ATM buffer (Illumina)

Incubate 55oC x 10 minutes, cycle to 10oC and immediately add 5μl NT buffer (Illumina) and incubate at RT for 5-10 minutes.


  1. Add 15μl NPM buffer (Illumina)
  2. Add 5μl index 1 (Illumina)
  3. Add 5μl index 2 (Illumina)

Run Thermocycler program:

  • 72oC x 3 minutes
  • 95oC x 30 seconds
  • 12 cycles of
    • 95oC for 10 seconds
    • 55oC for 30 seconds
    • 72oC for 30 seconds
  • 72oC for 5 minutes

Hold at 10oC

PCR clean up with AMPure XP beads (Use a separate Tube of AMPure beads that is dedicated to only POST-AMPLIFICATION work)

50μl PCR product + 100μl AMPure XP beads (pipet up and down) (2x)

Elute in 32μl 10mm Tris Acetate pH 8.0. Measure concentration via TapeStation (note: the concentration of the Amplified libraries is often in the middle between the range of the HS-D1000, and normal D1000 Tapes. Generally the D1000 tape should work when starting with 2ng input, below that, use HS D1000). Create normalized aliquots of each library at 1ng/μl by adding a sufficient amount of H2O to 10μl of library. Create multiplexed library by adding 10μl of each Normalized and uniquely indexed library to a multiplexed library tube. Mix thoroughly and transfer 10μl to PCR tube for sequencing at the Biopolymer Facility at Harvard Medical School.