Protocol for use with NEBNext Ultra Directional RNA Library Prep Kit for Illumina (E7420)

Symbols
This is a point where you can safely stop the protocol and store the samples prior to proceeding to the next step in the protocol.
This caution sign signifies a step in the protocol that has two paths leading to the same end point but is dependent on a user variable, like the type of RNA input.
Colored bullets indicate the cap color of the reagent to be added

The protocol has been optimized using high quality Universal Human Reference Total RNA. For PolyA mRNA selection, high quality RNA with RIN score >7 (measured by bioanalyzer) is required.


Starting Material:
Total RNA (100 ng–1 μg), purified mRNA (10–100 ng), or ribosomal depleted total RNA (10–100 ng) quantified by bioanalyzer.

The protocol is optimized for approximately 200 bp RNA inserts. To generate libraries with longer RNA insert sizes, refer to Appendix A in the manual for recommended fragmentation times and size selection conditions.

Note: Follow steps in "Preparation of First Strand Reaction Buffer and Random Primer Mix" Section 6.1A in the manual if starting material is total RNA. Perform mRNA isolation, fragmentation and priming using the NEBNext Poly (A) mRNA Magnetic Isolation Module (NEB #E7490). If starting material is purified mRNA or ribosomal depleted RNA, proceed to "RNA Fragmentation and Priming Starting from Purified mRNA or ribosomal depleted mRNA" Section 6.1B in the manual.

 6.1A Preparation of First Strand Reaction Buffer and Random Primer Mix

Prepare the First Strand Synthesis Reaction Buffer and Random Primer Mix (2X) as follows in a nuclease-free tube:

    In a sterile tube or 96-well PCR plate mix the following components:
    ( pink) NEBNext First Strand Synthesis Reaction Buffer (5X) 8 μl
     (pink) NEBNext Random Primers 2 μl
    Nuclease-free water 10 μl
    -----------------------------------------------
    Total Volume 20 μl

    Note: Keep the mix on ice during the mRNA isolation.
mRNA Isolation, Fragmentation and Priming Starting with Total RNA

  1. Dilute the total RNA with nuclease-free water to a final volume of 50 μl in a nuclease-free 0.2 ml PCR tube/plate and keep on ice.
  2. Aliquot 15 μl of NEBNext Oligo d(T)25 beads into a nuclease-free 0.2 ml PCR tube/plate.
  3. Wash the beads by adding 100 μl of RNA Binding Buffer to the beads. Pipette the entire volume up and down 6 times to mix thoroughly.
  4. Place the tube/plate on the magnetic rack at room temperature for 2 minutes.
  5. Remove and discard all of the supernatant from the tube/plate. Take care not to disturb the beads.
  6. Remove the tube/plate from the magnetic rack.
  7. Repeat Steps 3–6.
  8. Resuspend the beads in 50 μl of RNA Binding Buffer and add the 50 μl of total RNA sample from Step 1.
  9. Place the tube/plate on a thermal cycler and close the lid. Heat the sample at 65°C for 5 minutes and hold at 4°C to denature the RNA and facilitate binding of the poly-A mRNA to the beads.
  10. Remove the tube/plate from the thermal cycler when the temperature reaches 4°C.
  11. Place the tube/plate on the bench and incubate at room temperature for 5 minutes to allow the mRNA to bind to the beads.
  12. Place the tube/plate on the magnetic rack at room temperature for 2 minutes to separate the poly-A mRNA bound to the beads from the solution.
  13. Remove and discard all of the supernatant. Take care not to disturb the beads.
  14. Remove the tube/plate from the magnetic rack.
  15. Wash the beads by adding 200 μl of Wash Buffer to the tube/plate to remove unbound RNA. Pipette the entire volume up and down 6 times to mix thoroughly.
  16. Place the tube/plate on the magnetic rack at room temperature for 2 minutes.
  17. Remove and discard all of the supernatant from the tube/plate. Take care not to disturb the beads.
  18. Remove the tube/plate from the magnetic rack.
  19. Repeat Steps 15–18.
  20. Add 50 μl of Tris buffer to each sample. Gently pipette the entire volume up and down 6 times to mix thoroughly.
  21. Place the tube/plate on the thermal cycler. Close the lid and heat the samples at 80°C for 2 minutes, then hold at 25°C to elute the Poly-A mRNA from the beads.
  22. Remove the tube/plate from the thermal cycler when the temperature reaches 25°C.
  23. Add 50 μl of RNA Binding Buffer to the sample to allow the mRNA to re-bind to the beads. Gently pipette the entire volume up and down 6 times to mix thoroughly.
  24. Incubate the tube/plate at room temperature for 5 minutes.
  25. Place the tube/plate on the magnetic rack at room temperature for 2 minutes.
  26. Remove and discard all of the supernatant from the tube/plate. Take care not to disturb the beads.
  27. Remove the tube/plate from the magnetic rack.
  28. Wash the beads by adding 200 μl of Wash Buffer. Gently pipette the entire volume up and down 6 times to mix thoroughly.
  29. Place the tube/plate on the magnetic rack at room temperature for 2 minutes.
  30. Remove and discard all of the supernatant from the tube/plate. Take care not to disturb the beads.
  31. Remove the tube/plate from the magnetic rack.
  32. Wash the beads by adding 200 μl of Tris Buffer. Gently pipette the entire volume up and down 6 times to mix thoroughly.
  33. Place the tube/plate on the magnetic rack at room temperature for 2 minutes.
  34. Remove and discard all of the supernatant from the tube/plate. Take care not to disturb the beads.
    Note: It is important to remove all of the supernatant to successfully fragment the mRNA in the subsequent steps. Caution: Do not disturb beads that contain the mRNA.
  35. Remove the tube/plate from the magnetic rack.
    Note: The following fragmentation conditions used in Step 36 generates RNA fragments ~ 200 nt in length. Refer to the figure belowfor fragmentation conditions to generate RNA > 200 nt.
  36. Elute mRNA from the beads by adding 15 μl of the First Strand Synthesis Reaction Buffer and Random Primer Mix (2X) prepared at the start of the protocol and incubating the sample at 94°C for 15 minutes. Immediately, place the tube/plate on the magnetic rack.
  37. Collect the purified mRNA by transferring the 10 μl of supernatant to a clean nuclease-free PCR tube or plate.
  38. Place the tube/plate on ice.
  39. Proceed to "First Strand cDNA Synthesis" Section 6.2 in the manual.
Modified fragmentation times for longer RNA inserts.
Bioanalyzer traces of RNA as shown in RNA Pico Chip. mRNA isolated from Universal Human Reference RNA (1 μg) using the NEBNext Poly (A) mRNA Magnetic Isolation Module (NEB #E7490) and Fragmented with First Strand Synthesis Reaction Buffer and Random Primer Mix (2X) at 94°C for 5, 10 or 15 minutes. For libraries with RNA insert sizes larger than 300 bp, fragment RNA between 5–10 minutes.
 6.1B RNA Fragmentation and Priming Starting from Purified mRNA or ribosomal depleted mRNA

  1. Mix the following components in a nuclease-free PCR tube/plate:
    Purified mRNA/ribosomal depleted RNA (10–100 ng) 5 μl
     (pink) NEBNext First Strand Synthesis Reaction Buffer (5X) 4 μl
     (pink) Random Primers 1 μl
    -----------------------------------------------
    Final volume 10 μl

    Note: The following fragmentation conditions used in Step 2 generates RNA fragments ~ 200 nt in length. Refer to the figure above to generate RNA > 200 nt.

  2.  Incubate the sample at 94°C for 15 minutes.
  3. Transfer the tube/plate to ice.
  4. Proceed to First Strand cDNA Synthesis
6.2 First Strand cDNA Synthesis
  1.  Dilute Actinomycin D stock solution (5 μg/μl) to 0.1 μg/μl in nuclease free water for immediate use.

    Note: Dilute solutions of Actinomycin D are very sensitive to light. In solution, Actinomycin D tends to adsorb to plastic and glass. For these reasons, unused dilute solutions should be discarded and not stored for further use. However, frozen aliquots of a concentrated stock solution (5 μg/μl) are expected to be stable for at least a month at –20°C.

    To the fragmented and primed mRNA (10 μl from "Preparation of First Strand Reaction Buffer and Random Primer Mix" Step 38 or "RNA Fragmentation and Priming Starting from Purified mRNA or
    ribosomal depleted mRNA" Step 3) add the following components.

     (pink) Murine RNase Inhibitor 0.5 μl
    Actinomycin D (0.1 μg/μl) 5 μl
     (pink) ProtoScript® II Reverse Transcriptase 1 μl
    Nuclease-free water 3.5 μl
    -----------------------------------------------
    Final volume 20 μl

    Note: If you are following recommendations in the figure above, for longer RNA fragments, increase the incubation at 42°C from 15 minutes to 50 minutes in Step 2.

  2.  Incubate the sample in a preheated thermal cycler as follows:
    10 minutes at 25°C
    15 minutes at 42°C
    15 minutes at 70°C
    Hold at 4°C
6.3 Perform Second Strand cDNA Synthesis

  1. Add the following reagents to the First Strand Synthesis reaction (20 μl):
    Nuclease-free water 48 μl
     (orange) Second Strand Synthesis Reaction Buffer (10X) 8 μl
     (orange) Second Strand Synthesis Enzyme Mix 4 μl
    -----------------------------------------------
    Total volume 80 μl
  2. Mix thoroughly by gentle pipetting.
  3. Incubate in a thermal cycler for 1 hour at 16°C, with heated lid set at ≤ 40°C.

6.4 Purify the Double-stranded cDNA Using 1.8X Agencourt AMPure XP Beads


  1. Vortex AMPure XP beads to resuspend.
  2. Add 144 μl (1.8X) of resuspended AMPure XP beads to the second strand least 10 times.
  3. Incubate for 5 minutes at room temperature.
  4. Place the tube/plate on an appropriate magnetic rack to separate beads from supernatant. After the solution is clear (about 5 minutes), carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets.
  5. Add 200 μl of freshly prepared 80% ethanol to the sample while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.
  6. Repeat Step 5 once for a total of 2 washes.
  7. Air dry the beads for 5–10 minutes while the tube is on the magnetic rack with lid open. Caution: Do not overdry the beads. This may result in lower recovery of DNA target.
  8. Remove the tube/plate from the magnet. Elute the DNA target from the beads into 60 μl nuclease-free water. Mix well by gently pipetting up and down several times. Place the tube/plate on the magnetic rack until the solution is clear.
  9. Remove 55.5 μl of the supernatant and transfer to a clean nuclease-free PCR tube/plate.

     Note: If you need to stop at this point in the protocol samples can be stored at –20°C.

6.5 Perform End Prep of cDNA Library

  1. Mix the following components in a sterile nuclease-free tube/96-well PCR plate:
    Purified double-stranded cDNA 55.5 μl
     (green) NEBNext End Repair Reaction Buffer (10X) 6.5 μl
     (green) NEBNext End Prep Enzyme Mix 3 μl
    -----------------------------------------------
    Total volume 65 μl
  2. Incubate the sample in a thermal cycler as follows:
    30 minutes at 20°C
    30 minutes at 65°C
    Hold at 4°C
  3. Proceed immediately to adaptor ligation.

6.6 Perform Adaptor Ligation

Dilute the NEBNext Adaptor for Illumina (15 μM) to 1.5 μM with a 10-fold dilution (1:9) with 10 mM NaCl for immediate use.

  1. Add the following components directly to the End Prep reaction mixture (65 μl) and mix well:
     (red) Blunt/TA Ligase Master Mix 15 μl
     (red) Diluted NEBNext Adaptor 1 μl
    Nuclease-free Water 2.5 μl
    -----------------------------------------------
    Total volume 83.5 μl
  2. Incubate 15 minutes at 20°C in a thermal cycler.

    Note: USER Step is performed during the PCR reaction.

6.7 Purify the Ligation Reaction Using Agencourt AMPure XP Beads

 Note: If you are selecting for larger size fragments (> 200 nt) follow the size selection recommendations in the Table.

  1. To the ligation reaction (83.5 μl), add 16.5 μl nuclease-free water to bring the reaction volume to 100 μl.
    Note: X refers to the original sample volume of 100 μl from the above step.
  2. Add 100 μl (1.0X) resuspended Agencourt AMPure XP Beads and mix well by gently pipetting up and down at least 10 times.
  3. Incubate for 5 minutes at room temperature.
  4. Place the tube/plate on an appropriate magnetic rack to separate beads from the supernatant. After the solution is clear (about 5 minutes), discard the supernatant that contains unwanted fragments (Caution: do not discard the beads).
  5. Add 200 μl of freshly prepared 80% ethanol to the sample while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.
  6. Repeat Step 5 once for a total of 2 washes.
  7. Completely remove the residual ethanol.
  8. Air dry beads for 5–10 minutes while the tube is on the magnetic rack with the lid open. Caution: Do not overdry the beads. This may result in lower recovery of DNA target.
  9. Remove the tube/rack from the magnet. Elute DNA target from the beads with 50 μl nuclease-free water. Mix well on by pipetting up and down several times, and put the tube in the magnetic rack until the solution is clear.
  10. Transfer the 50 μl supernatant to a clean PCR tube/plate. Discard beads.
  11. To the 50 μl supernatant, add 50 μl (1.0X) of the resuspended Agencourt AMPure XP Beads and mix well by gently pipetting up and down at least 10 times.
  12. Incubate for 5 minutes at room temperature.
  13. Place the tube/plate on an appropriate magnetic rack to separate beads from the supernatant. After the solution is clear (about 5 minutes), discard the supernatant that contains unwanted fragments (Caution: do not discard the beads).
  14. Add 200 μl of freshly prepared 80% ethanol to the beads while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.
  15. Repeat Step 14 once for a total of 2 washes.
  16. Completely remove the residual ethanol.
  17. Air dry beads for 5-10 minutes while the tube is on the magnetic rack with the lid open. Caution: Do not overdry the beads. This may result in lower recovery of DNA target.
  18. Remove the tube/plate from the magnet. Elute DNA target from the beads with 22 μl nuclease-free water. Mix well by pipetting gently up and down several times, and put the tube/plate in the magnetic rack until the solution is clear.
  19. Without disturbing the bead pellet, transfer 17 μl of the supernatant to a clean PCR tube/plate and proceed to PCR amplification.

    Note: Be sure not to transfer any beads. Trace amounts of bead carry over may affect the optimal performance of the polymerase used in the NEBNext High-Fidelity 2X PCR Master Mix in the subsequent PCR step.

    Table: Recommended size selection conditions for libraries with insert sizes larger than 300 bp.
  20. LIBRARY
    PARAMETERS
    APPROXIMATE
    INSERT SIZE
    250-
    400 bp
    300-
    450 bp
    400-
    600 bp
    500-
    700 bp
    Approx. Final
    Library Size
    350-
    500 bp
    400-
    550 bp
    500-
    700 bp
    600-
    800 bp
    VOLUME TO
    BE ADDED (μl)
    1st Bead
    Selection
    45 40 35 30
    2nd Bead
    Selection
    20 20 15 15
    Note: Any differences in insert sizes between the Agilent Bioanalyzer and that obtained from paired end sequencing can be attributed to the higher clustering efficiency of smaller sized fragments.

6.8 PCR Amplification

For < 96 samples, follow the protocol "Setting up the PCR reactions (< 96 samples)" Section 6.8A in the manual . For 96 samples, follow the protocol "Setting up the PCR reactions (96 samples)" Section 6.8B in the manual

6.8A Setting up the PCR reactions (< 96 samples)


Note: We recommend using a PCR work-up rack such as the TruSeq Index Plate Fixture (Illumina #FC-130-1005) to assist in properly combining the index primers during the PCR amplification step. Alternatively, 96-well deep well plates can be used and aligned against a PCR plate as in the diagram below.

  1. Ensure that a valid combination of i7 and i5 primers is used. See Appendix A in the manual to verify that correct primer combinations have been selected.
  2. Arrange the index primers in the Index Plate Fixture as follows:
    1. Arrange the  (orange) i7 primers in increasing order horizontally, so that the lowest number i7 index primer is in column 1, second lowest number i7 index primer is in column 2, etc.
    2. Arrange the  (white) i5 primers in increasing order vertically, so that the lowest number i5 index primer is in row A, second lowest number i5 index primer is in row B, etc.
    3. Record their positions on the PCR setup template (see Appendix B in the manual).
  3. Using a multichannel pipette, add 2.5 μl of (white) i5 primers to every column (as needed) of the PCR plate. It is critical to change tips between columns to avoid cross-contamination.
  4. Discard the original i5 white caps and apply new caps to avoid index cross-contamination.
  5. Using a multichannel pipette, add 2.5 μl of (orange) i7 primers to every row (as needed) of the PCR plate. It is critical to change tips between rows to avoid cross-contamination.
  6. Discard the original i7 orange caps and apply new caps to avoid index cross-contamination.
  7. Add 25 μl  (blue) NEBNext High-Fidelity 2X PCR Master Mix to each well that contains primers.
  8. Add 3 μl  (blue) USER enzyme to each well that contains primers.
  9. Add 17 μl adaptor ligated DNA fragments to the corresponding well. Gently pipette up and down 5–10 times to mix. It is critical to change tips between samples to avoid cross-contamination. Record each sample position on the PCR setup template (see Appendix B in the manual).
  10. Seal the plate with Bio-Rad Microseal "A" Film and quickly centrifuge.
  11. Perform PCR according to the cycling conditions:
    CYCLE STEP TEMP TIME CYCLES
    User Digestion 37°C 15 minutes 1
    Initial Denaturation 98°C 30 seconds 1
    Denaturation 98°C 10 seconds 12–15*,**
    Annealing 65°C 30 seconds
    Extension 72°C 30 seconds
    Final Extension 72°C 5 minutes 1
    4°C Hold
    * The number of PCR cycles should be adjusted based on RNA input. If 100 ng total RNA or 10 ng purified mRNA or ribosomal-depleted RNA are the starting input, it is recommended to perform 15 cycles of PCR.
    ** It is important to limit the number of PCR cycles to avoid overamplification. If overamplification occurs, larger molecular weight products (> 500 bp) will appear on the bioanalyzer trace.
  12. Proceed to "Purify the PCR Reaction Using AMPure XP Beads" Section 6.9 in the manual

6.8B Setting up the PCR reactions (96 samples)

Note: We recommend using a PCR work-up rack such as the TruSeq Index Plate Fixture (Illumina #FC-130-1005) to assist in properly combining the index primers during the PCR amplification step. Alternatively, 96-well deep well plates can be used and aligned against a PCR plate as in the diagram below.

  1. Arrange the index primers in the Index Plate Fixture as follows:
    1. Arrange (orange) i7 primers in increasing order horizontally, so that i701 is in column 1, i702 is in column 2, i703 is in column 3, etc.
    2. Arrange the  (white) i5 primers in increasing order vertically, so that i501 is in row A, i502 is in row B, i503 is in row C, etc.
    3. Record their positions on the PCR setup template (see Appendix B in the manual).
  2. Using a multichannel pipette, add 2.5 μl of  (white) i5 primers to every column (as needed) of the PCR plate. It is critical to change tips between columns to avoid cross-contamination.
  3. Discard the original i5 white caps and apply new caps to avoid index cross-contamination.
  4. Using a multichannel pipette, add 2.5 μl of  (orange) i7 primers to every row of the PCR plate. It is critical to change tips between rows to avoid cross-contamination.
  5. Discard the original i7 orange caps and apply new caps to avoid index cross-contamination.
  6. Add 25 μl  (blue) NEBNext High-Fidelity 2X PCR Master Mix to each well that contains primers.
  7. Add 3 μl  (blue) USER enzyme to each well that contains primers.
  8. Add 17 μl adaptor ligated DNA fragments to the corresponding well. Gently pipette up and down 5–10 times to mix. It is critical to change tips between samples to avoid cross-contamination. Record each sample position on the PCR setup template (see Appendix B in the manual).
  9. Seal the plate with Bio-Rad Microseal "A" Film and quickly centrifuge.
  10. Perform PCR according to the cycling conditions:
    CYCLE STEP TEMP TIME CYCLES
    User Digestion 37°C 15 minutes 1
    Initial Denaturation 98°C 30 seconds 1
    Denaturation 98°C 10 seconds 12–15*,**
    Annealing 65°C 30 seconds
    Extension 72°C 30 seconds
    Final Extension 72°C 5 minutes 1
    4°C Hold
    * The number of PCR cycles should be adjusted based on RNA input. If 100 ng total RNA or 10 ng purified mRNA or ribosomal-depleted RNA are the starting input, it is recommended to perform 15 cycles of PCR.
    ** It is important to limit the number of PCR cycles to avoid overamplification. If overamplification occurs, larger molecular weight products (> 500 bp) will appear on the bioanalyzer trace.

6.9 Purify the PCR Reaction Using AMPure XP Beads

Note: X refers to the original sample volume from the above step.

  1. Vortex Agencourt AMPure XP Beads to resuspend.
  2. Add 45 μl of resuspended Agencourt AMPure XP Beads to the PCR reactions (~50 μl). Mix well by gently pipetting up and down at least 10 times.
  3. Incubate for 5 minutes at room temperature.
  4. Place the plate on an appropriate magnetic rack to separate beads from the supernatant. After the solution is clear (about 5 minutes), carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets.
  5. Add 200 μl of freshly prepared 80% ethanol to the sample while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.
  6. Repeat Step 5 once for a total of 2 washes.
  7. Air dry the beads for 5-10 minutes while the plate is on the magnetic rack with the lid open. Caution: Do not overdry the beads. This may result in lower recovery of DNA target.
  8. Remove the plate from the magnet. Elute the DNA target from the beads into 23 μl 10 mM Tris-HCl, pH 8.0 or 0.1X TE. Mix well by gently pipetting up and down several times, and place the plate in the magnetic rack until the solution is clear.
  9. Transfer 20 μl of the supernatant to a clean PCR tube, and store at –20°C.

6.10 Assess Library Quality on a Bioanalyzer (Agilent high sensitivity chip)

  1. Dilute (1:4) library in 10 mM Tris or 0.1X TE.
  2. Run 1 μl in a DNA high sensitivity chip.
  3. Check that the electropherogram shows a narrow distribution with a peak size approximately 300 bp.

    Note: If a peak at ~ 80 bp (primers) or 128 bp (adaptor-dimer) is shown in the bioanalyzer traces; bring up the sample volume to 50 μl exactly with nuclease-free water and repeat the Agencourt AMPure XP Bead clean up step.