Stranded mRNA Prep, Ligation (Updated TruSeq)

Stranded mRNA Prep, Ligation (Updated TruSeq)

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Documentation

Illumina's Stranded mRNA Sample Prep Guide. This method makes a cDNA library of the polyadenylated mRNA in your eukaryotic samples of interest. It requires high quality RNA with intact polyA tails.

A few important comments before you start

  • The recommended starting amount of total RNA is 25-1000ng, but we usually try to stay away from the extreme ends of this spectrum and use 100ng input.
  • This protocol can be performed in one day, or split into two or more days at the indicated safe stopping points.
  • For 12 samples, you will need approximately 7 hours if completing the protocol in one day. This estimate is based on an experienced user with a multichannel pipette and vortexing at bead cleanup steps. An inexperienced user or one not using time-saving techniques should plan for more time.
  • Keep all reagents on ice unless otherwise stated. Do NOT put any reagents stored at 4C, especially beads, on ice.
  • Have all 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.
  • 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. Samples can also be mixed by sealing the plate and vortexing as described in the Sample Prep Guide linked above.
  • We use a Tapestation to assay the input RNA quality. RNA Integrity Numbers (RIN) of 7 and higher are preferred.
  • We quantify the input RNA using a Qubit fluorometer.

Purify and Fragment mRNA

The following items are included in the Illumina kit. We recommend kits be stored as received in the original manufacturer’s boxes and not unpacked to organize in a reagent box. When kits go bad, it is much easier to contain the damage if reagents were stored as received.

This kit has 1 box of reagents in the 4C fridge and 2 boxes of reagents in the -20C freezer, each labeled with a green dot saying "mRNA Ligation". Make sure you are using the correct boxes and return all reagents to the box they came from. Do not mix reagents with other boxes.

Prepare the following:

NAMELOCATIONACTION
Bead Binding Buffer (BBB)
4C Fridge
Vortex briefly, leave at RT
Bead Washing Buffer (BWB)
4C Fridge
Vortex briefly, leave at RT
Elution Buffer (EB)
4C Fridge
Vortex briefly, leave at RT
RNA Purification Beads (RPBX)
4C Fridge
Let stand at RT for 30 minutes to bring to RT, vortex and invert thoroughly before use
Elute, Prime, Fragment High Mix (EPH3)
-20 Freezer (cDNA synthesis box, gray sticker)
Thaw at RT, vortex briefly
First Strand Synthesis Act D mix (FSA)
-20 Freezer (cDNA synthesis box, brown tube)
Thaw and keep on ice, vortex briefly before use

Prepare the following non-Illumina materials:

NAMELOCATIONACTIONVOLUME
Ampure XP Beads
4C Fridge
Let stand at RT for 30 minutes to bring to RT, vortex and invert thoroughly before use
174uL per sample for entire prep
Fresh 80% Ethanol (200 proof EtOH + MilliQ water)
RT Bench
Mix fresh each day. Low% EtOH negatively affects cleanups.
1.2mL per sample for entire prep
RNAse Free RT-PCR Grade Water
-20C Freezer
Thaw at RT
As needed for diluting RNA input
Post-Style Magnetic Stand for 96 well plates
RT Bench

Step 1: Capture mRNA

Dilute 25-1000ng (we typically do 100ng) total RNA in nuclease-free water to a total volume of 25uL. Do this in a 96-well PCR plate (if only prepping a few samples, the plate can be cut, but smaller than a half plate can be unstable on the magnet during cleanup steps). We typically normalize inputs across all samples. Different inputs can be used, but samples may require different reagent volumes and cycle numbers in later steps based on input.
Add 25 uL well-mixed RPBX to each well. Gently pipette up and down 10 times. Seal the plate with an adhesive seal.
⚠️

Do not store RNA purification beads on ice! We have observed that the beads are very sensitive to cold temperatures. Beads stored at < 2degC may deteriorate, resulting in poor binding of mRNA, and therefore poor library prep that is heavily contaminated with excess adapter.

Place the sealed plate in the thermal cycler. Go to the "STRANDED LIG" folder and run the "MRNA CAP" program (65C for 5min, 4C for 30sec, 23C for 5min, 23C hold).
Remove the plate, keep it sealed, and centrifuge at 280 x g for 10 sec.
Place on the magnetic stand for 2min.
Discard the supernatant from each well of the plate.
Remove the plate from the magnetic stand and add 100uL BWB to each well. Pipette 10 times to mix.
Place the plate on the magnetic stand for 2 min.
Remove and discard the supernatant from each well.

Use at 20uL pipette to remove residual BWB.
Remove the plate from the magnetic stand.
Add 25uL ELB and slowly pipette to completely resuspend beads.
Seal and centrifuge at 280 x g for 10 sec.
Place the plate in the thermal cycler. Run the program “MRNA ELT” (80C for 2 min, 25 C hold).
Remove the plate when the thermal cycler reaches 25C. Centrifuge the sealed plate at 280 x g for 10 sec.

Step 2: Clean Up and Fragment mRNA

On ice, in a microcentrifuge tube, prepare Fragmentation Master Mix with the following volumes per sample (overage already included):

Fragmentation Master Mix

NameVolume
Nuclease-free water
10.5 uL
EPH3
10.5 uL
Add 25uL BBB to each well. Pipette 10 times to mix.
Incubate at room temperature for 5 minutes.
Place on the magnetic stand and wait 2 minutes.
Remove and discard 50uL supernatant from each well.
Remove the plate from the magnetic stand and add 100uL BWB to each well. Pipette 10 times to mix, fully resuspending the beads.
Place on the magnetic stand and wait 2 minutes.
Remove and discard supernatant. With a 20uL pipette, remove and discard any residual supernatant.
Remove from the magnetic stand.
Pipette Fragmentation Master Mix to mix, then add 19uL to each well, slowing pipetting to mix until beads are fully resuspended.
Incubate at room temperature for 2 minutes.
Centrifuge at 280 x g for 10 seconds. Place on thermal cycler and run the "DEN94 8" program (94C for 8min, 4C hold).
Remove the plate once it reaches 4C and centrifuge at 280 x g for 10 seconds
Remove the adhesive seal and place the plate on a magnetic stand at room temperature for 2 min.
Remove 17 uL supernatant from each well into corresponding wells in a new plate. This contains your purified mRNA.

Generate cDNA

Step 3: First strand cDNA synthesis

Prepare the following:

NAMELOCATIONACTION
Ampure XP Beads
4C Fridge
Let stand at RT for 30 minutes to bring to RT, vortex and invert thoroughly before use
Resuspension Buffer (RSB)
-20 Freezer (cDNA synthesis box, purple sticker)
Thaw at RT, vortex briefly
Reverse Transcriptase (RVT)
-20 Freezer (cDNA synthesis box, pink sticker)
Store until needed. Flick to mix, centrifuge briefly
Second Strand Marking Mix (SMM)
-20 Freezer (cDNA synthesis box, blue sticker)
Thaw on ice, invert to mix, centrifuge briefly
On ice, in a microcentrifuge tube, prepare First Strand Synthesis Master Mix with the following volumes per sample (overage already included):

First Strand Synthesis Master Mix

NAMEVOLUME (uL)
FSA
9
RVT
1
Pipette First Strand Synthesis Master Mix to mix.
Centrifuge at 280 x g for 10 seconds. Add 8uL First Strand Synthesis Master Mix to each well and pipette 10 times to mix.
Seal the plate and place in the thermocycler and run the “FSS” program (25C for 10 min, 42C for 15 min, 70C for 15 min, hold at 4C). Return RVT to the -20 freezer immediately after use.
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This step takes about 40 minutes and is your first large break.

Step 4: Second strand cDNA synthesis

When the first strand synthesis is completed, immediately proceed to the second strand synthesis.

Centrifuge sealed plate at 280 x g for 10 seconds. Add 25 uL SMM to each well. Pipette up and down 10 times.
Seal plate, centrifuge if needed, place plate in thermocycler and run the "SSS" program (16C for 1hr).
Make sure you have Ampure XP beads at room temp and fresh 80% ethanol for the next steps.
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This step takes about 1 hour and is your second large break of the day.

Step 5: cDNA clean-up

When the reaction is complete, add 90 ul of well-mixed AMPure XP beads to each well containing cDNA and pipette to mix (or seal plate and vortex for 1 min. Check that the beads are not stuck to the sides of the wells. You may need to spin down the plate and vortex again). Incubate RT for 5 min.
Place the plate on the magnet and let sit for 5 min.
Remove and discard 130 ul supernatant per well, being careful not to disturb the beads.
Wash beads on magnet by adding 175 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.
Remove any residual supernatant with a p20 pipette. Let the plate stand on the magnet at RT for 2 minutes.
Remove from the magnet and add 19.5 ul of RSB to each well and pipette to fully resuspend the beads. Incubate for 2 min at RT.
Centrifuge at 280 x g for 10 seconds.
Place the plate on the magnetic stand for 2 minutes.
Transfer 17.5 ul of the supernatant (contains the ds cDNA) to new a new plate.
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This is a safe stopping point. You can seal the plate and store at -20C for up to 7 days.

Prepare and Amplify Libraries

Step 6: Adenylate cDNA ends

Prepare the following:

NAMELOCATIONACTION
Anchor Plate
-20 Freezer
Thaw at RT, vortex, centrifuge briefly
Index Adapter Plate
-20 Freezer
Thaw at RT, vortex, centrifuge for 1 minute
A-Tailing Mix (ATL4)
-20 Freezer (Ligation box, green sticker)
Thaw at RT, flick to mix, centrifuge briefly
Stop Ligation Buffer (STL)
-20 Freezer (Ligation box, red cap)
Thaw at RT, vortex, centrifuge briefly
Ampure XP Beads
4C Fridge
Let stand at RT for 30 minutes to bring to RT, vortex and invert thoroughly before use
Resuspension Buffer (RSB)
-20 Freezer (cDNA synthesis box, purple sticker)
Thaw at RT, vortex briefly
Enhanced PCR Mix (EPM)
-20 Freezer (Ligation box, yellow sticker)
Thaw at RT, invert to mix, centrifuge briefly
Ligation Mix (LIGX)
-20 Freezer (Ligation box, red sticker)
Store until needed, flick to mix, centrifuge briefly
Add 12.5 ul of thawed A-tailing mix to each well. Using a p200 pipette, pipette up and down 10 times.
Seal and put on the thermocycler for the "ATAIL" program (37C for 30min, 70C for 5min, 4C hold).
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This step takes about 35 minutes and is another break.

Step 7: Ligate Anchors

Each well of the anchor plate is single-use and contains the same RNA Index Anchors. They can be used in any order. Seal the used anchor wells with foil and label before returning to freezer storage.

Add the following volumes to each well in order, according to sample input:

Order of AdditionReagentVol for Sample Input < or = 100ngVol for Sample Input > 100ng
1
RSB
2.5 uL
0 uL
2
RNA Index Anchors
2.5 uL
5 uL
3
LIGX
2.5 uL
2.5 uL
With a p200 pipette, pipette to mix 10 times.
Seal and run on the thermocycler under the "LIG" program (30C for 10min, 4C hold).
Briefly centrifuge plate. Add 5uL STL to each well, pipetting to mix 15 times.

Step 8: Clean Up Adapter-Ligated Fragments

Add 34 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 67 ul of supernatant from each sample, being careful not to disturb the beads.
With the plate on the magnetic stand, wash with 175 ul 80% EtOH for 30 sec, then remove the ethanol.
Perform the wash step again.
Use a p20 pipette to remove residual ethanol. Keep the plate on the magnet at RT for 2 minutes to dry.
Remove the plate from the magnetic stand and add 22 ul of Resuspension Buffer to each well, and mix well by pipetting to fully resuspend beads.
Incubate at RT for 2 minutes.
Centrifuge at 280 x g for 10 seconds.
Place the plate on magnet for 5 min.
Keeping the plate on the magnet, recover 20 ul from each well and transfer to a new plate.
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This is a safe stopping point. You can seal the plate and store at -20C for up to 7 days.

Step 9: Index and Amplify library

Add 10uL index adapter to each well.
⚠️

Indexes come in a 96 well plate, each well containing a unique set of dual indexes. Although they are unique, certain patterns of index use are recommended to balance color channels during sequencing. See Illumina's Index Adapters Pooling Guide. Generally, moving in columns is fine. Seal the used wells with foil and label before returning to freezer storage. Record which index plate (Set A or B) and which index well was used for each sample, as this is needed to demultiplex after sequencing.

Add 20uL EPM. Pipette 10 times to mix.
Seal and place on thermocycler and run the "PCR" program, with cycle repeats according to the table below.

PCR Amplification

Input Amount (ng)Number of PCR Cycles
25
15
100
13
1000
10
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This is a 30-45 minute break, depending on cycle number

Step 10: Clean Up Library

Add 50 ul of AMPure beads to each well and mix well by pipetting. Incubate 15 min at RT.
Place on the magnet and incubate for 5 minutes.
Remove and discard 90 ul of supernatant from each well.
Wash the beads with 175 ul 80% EtOH, wait 30s, remove the supernatant.
Repeat the ethanol wash step.
Use a p20 pipette to remove residual ethanol.
Keep the plate on the magnet at RT for 2 minutes to dry.
Remove the plate from the magnetic stand, add 17 ul of resuspension buffer and mix well by pipetting.
Incubate for 2 min at room temperature.
Place the plate back on the magnetic stand for 5 minutes.
Remove 15 uL of supernatant from each sample and move it into a new plate. This is the final cDNA product.

Congratulations! You’re done!…well, almost

Perform quality control steps - we use Tapestation to determine the library size and Qubit to quantify the libraries. More detail on QC and sequencing is below.

Libraries can be stored at -20 for up to 30 days.

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Sequencing Guidelines

Normalize and Pool

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.
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.
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.
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

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.
Continue to Prep Libraries. Select your library prep kit based on the index format used. If you used our plate indexes select “IDT-ILMN Nextera UD Index Set (A or B) for Nextera DNA Flex.” 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.
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.
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 72 (for dual 10bp indexes) for Read 1 Cycles and 0 for Read 2 Cycles.
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.
  • The recommended loading concentration is 1.6pM and 5% PhiX spike-in.