Preparing normalized cdna libraries for transcriptome sequencing (Illumina HiSeq) Last updated: Oct 28, 2016 Overview First-strand cdna is synthesized using oligo-dt containing primers and an RNA oligo to introduce known sequences at both ends of each transcript via templateswitching. cdna is amplified using a single primer that binds at each end of the template, taking advantage of the PCR suppression effect to enrich for full-length transcripts. cdna is normalized using a double-stranded DNA specific nuclease (DSN) to enrich the library for transcripts expressed at low levels. cdna is randomly fragmented by sonication (preferred method) or enzymatic degradation with a non-specific nuclease mixture (Fragmentase). Fragmented cdna is repaired and tailed for ligation, then ligated to adaptors to build the sequencing construct. Ligation constructs are amplified to introduce sequencing primer binding sites and sample-specific barcodes. Finally, the barcoded libraries are size-selected using gel purification to select fragments of appropriate length for Illumina sequencing. Before you begin 1. Extract >1 µg total RNA from your experimental subject. To optimize the diversity of genes represented in your transcriptome assembly, it s ideal to include multiple stages, tissues, or treatments in this initial sample. Avoid sampling multiple genotypes if possible since nucleotide diversity complicates assembly. 2. Check the quality by loading 100-200 ng on a 1% agarose TAE gel. Intact RNA will show distinct rrna bands without smearing. The integrity of your initial sample is the single most important factor determining success of your library prep and transcriptome assembly. It s worth spending some time and resources optimizing extractions until your RNA is essentially perfectly intact. 3. Check for presence of genomic DNA on gel a high MW band (>10kb) or a visible smear higher than the LSU rrna band. 4. If genomic DNA is visible on the gel, precipitate with LiCl (resuspending in ~10-12 µl) and check another 100-200 ng on a gel.
5. The procedure begins with 2 µg of RNA in 18 µl water. Concentrate the sample if necessary by precipitation or drying under vacuum. Transfer into a PCR tube, strip, or plate. FS-cDNA synthesis In this step, single-stranded cdna is synthesized using total RNA as a template. The oligo-dt containing primer selects for mrna, incorporating an amplification tag at the 3 end of the cdna. Template-switching activity of the reverse transcriptase is used to incorporate the same tag at the 5 end of each cdna. 1. Combine 2 μl of the oligonucleotide CA1-20TVN (10 μm) with 18 µl RNA. Incubate at 65 C for 3 minutes in a thermocycler, then transfer immediately onto ice. 2. Prepare FS-cDNA synthesis master mix. The following volumes are intended for a single reaction, so multiply these values by the number of reactions (plus ~10% to account for pipetting error): 6 μl Nuclease-free water (NFW) 2 μl dntp (10 mm each) 8 μl 5X first-strand buffer (supplied with enzyme) 2 μl 10 μm CA1-TS-YY oligo (RNA, stored at -80 C) 2 μl Tetro Reverse Transcriptase (Bioline) 3. Add 20 μl of this master mix to the RNA from step 1 and mix thoroughly. 4. Incubate in a thermocycler for 1 hr at 45 C, followed by 85 C for 5 minutes to inactivate the RT, then a 4 C hold. 5. Dilute four-fold by adding 120 µl NFW and store on ice or frozen until the next step.
Amplification In this step, FS-cDNA is amplified using a primer complementary to the tags introduced in FS-cDNA synthesis. This reaction uses the PCR suppression effect to selectively suppress amplification of short templates, effectively enriching for complete transcripts. Its important to check the molecular weight distribution at this stage to ensure complete transcripts are recovered. 1. Test-scale PCR a. Prepare master mix. The following volumes are intended for a single reaction each, so multiply these values by the total number of reactions plus a small additional amount to account for pipetting error. This recipe assumes 5 μl of template, so if you use a different amount of template, adjust the water accordingly. 8.75 μl NFW 0.5 μl dntp (10 mm each) 5 μl 5X Q5 PCR buffer 0.5 μl 10 μm PST-CA1 oligo 0.25 μl Q5 DNA polymerase b. Combine 15 μl of master mix with 10 μl diluted FS-cDNA. c. Amplify in a thermocycler using the following profile: 98 C for 0:30, (98 C for 0:10 60 C for 0:25 72 C for 3:00) x 17 d. After 17 cycles, sample 5 μl. Continue adding cycles in increments of 2, up to 23, and sample 5 μl at each pause. Run all samples on a 1% agarose gel. A smear of cdna (500-2000 bp) should be faintly visible. This is an important quality control checkpoint. If your cdna smear doesn t extend above 1.5 kb, the first-strand cdna is not optimal and the final assembly will be fragmented and 3 biased. Troubleshoot FS-cDNA and PCR until you can produce high molecular weight smears in the earliest cycle numbers producing visible amplification.
If no product is observed within 20 cycles, repeat the test using larger amounts of template. 2. Preparation-scale PCR a. Once the optimum amount of template and number of cycles have been determined, scale up the reaction and amplify the remainder of the cdna, using multiple 50 µl reactions per template if necessary. b. This recipe assumes 20 µl of template for a 50 µl reaction (scaled up directly from the test scale reactions), so if you use more template adjust the water accordingly. Prepare several 50-µl reactions for each FS-cDNA sample. 17.5 μl NFW 1 μl dntp (10 mm each) 10 μl 5X Q5 PCR buffer 1 μl 10 μm PST-CA1 oligo 0.5 μl Q5 DNA polymerase c. Combine 30 μl of master mix with 20 μl diluted FS-cDNA. d. Amplify in a thermocycler using the following profile: 98 C for 0:30, (98 C for 0:10 60 C for 0:25 72 C for 3:00) x N, 72 C for 5:00 (N = optimum cycle number determined above) e. After PCR, check 5 μl of the product on a gel to verify that the reaction worked as expected, BEFORE freezing or purifying the product. f. Purify the cdna in a minimal volume (20-30 µl) and quantify using A260 or a fluorescent DNA assay. You ll need at least 0.6-1.2 µg for the next step. cdna Normalization In this step, the abundance of highlyexpressed transcripts is reduced, enriching for transcripts expressed at lower levels. Keys to success in this reaction: Controlling temperature. Samples are hybridized for a long duration to reach equilibrium, then maintained at the same temperature while additional pre-warmed reagents are added. Thoroughly mixing the DSN enzyme by backpipetting prior to drawing the
DSN for normalization reactions. The enzyme tends to precipitate in storage and needs to be resuspended well before use. 1. For each cdna sample, combine the following reagents in a sterile 0.5 ml tube: 4-12 µl cdna (0.6-1.2 µg of cdna) 4 µl 4X DSN Hybridization Buffer to 16 µl NFW 2. Mix well, spin briefly in a microcentrifuge, then hold on ice. 3. Aliquot 4 µl of the reaction mixture into each of four PCR tubes labeled C, 1, 1/2, and 1/4. 4. Overlay the reaction mixture in each tube with a drop of mineral oil and centrifuge the tubes for 2 min at maximum speed in a microcentrifuge. 5. Incubate the tubes in a thermal cycler at 98 C for 2 min. 6. Incubate the tubes at 68 C for 4-7 h, then proceed immediately to the next step. Do not remove the samples from the thermal cycler before DSN treatment. 7. Shortly before the end of the hybridization procedure, prepare two DSN dilutions in two sterile tubes to final DSN concentrations of 0.5 U/µl and 0.25 U/µl, as follows: a. Prepare a ½ dilution by combining 1.5 µl of DSN Storage Buffer with 1.5 µl of DSN stock solution. Mix by gently pipetting the reaction mixture up and down. b. Prepare a ¼ dilution by combining 1 µl of the ½ dilution with 1 µl DSN Storage Buffer. Mix by gently pipetting the reaction mixture up and down. c. Hold the tubes on ice. 8. Pipette some DSN Master Buffer into a PCR tube and prewarm it at 68 C in the thermocycler for 3-5 min. 9. After the hybridization period, while keeping cdna tubes in the PCR block, add 5 µl of the prewarmed DSN master buffer to each tube containing hybridized cdna and mix by gently back-pipetting. Keep the pipette tip at the bottom of the tube to avoid mixing the aqueous solution with its mineral oil overlay during backpipetting. 10. Incubate the tubes at 68 C for 10 min. 11. Keeping tubes in the PCR block, add DSN enzyme as specified in the following table and mix by gently back-pipetting with the pipette tip at the bottom of the tube. Again, avoid mixing with the mineral oil overlay. Tube Add C 1 µl DSN Storage Buffer 1 1 µl DSN stock ½ 1 µl of the ½ dilution DSN ¼ 1 µl of the ¼ dilution DSN
12. Incubate the tubes in the thermal cycler at 68 C for 25 min. 13. Add 5 µl of DSN Stop Solution, briefly vortex and spin the tubes briefly in a microcentrifuge. 14. Incubate in the thermal cycler at 68 C for 5 min then place on ice. 15. Add 25 µl NFW to each tube. Mix well by vortexing, spin briefly in a microcentrifuge, then place on ice or store at 20 C. Amplification of normalized cdna 1. Prepare a PCR master mix by combining the following reagents in the order shown, for each sample. This is enough for four reactions per sample (described below) 57 µl NFW 20 µl 5X Q5 PCR buffer 2 µl 10 mm dntps 4 µl Primer PST-CA1 (10 µm) 1 µl Q5 DNA polymerase 2. Mix well and spin briefly in a microcentrifuge. 3. Aliquot 4 µl of normalized cdna into appropriately labeled PCR tubes. Add 21 µl of the PCR master mix into the tubes, mix well. 4. Subject the tubes to PCR cycling using the following program: 98 C for 0:30, (98 C for 0:10 60 C for 0:25 72 C for 3:00) x 7 5. Put the Experimental tubes (1, ½, and ¼) on ice. Use the Control tube (C) to determine the optimal number of PCR cycles. After 7 cycles, sample 5 μl. Continue adding cycles in increments of 2, up to 13, and sample 5 μl at each pause. 6. Run all on a 1% agarose gel. A smear of cdna (500-2000 bp) should be visible. 7. When the amount of PCR product stops increasing with an additional cycle, the reaction has reached a plateau. The optimal number of cycles is 1 cycle less than that needed to reach the plateau. 8. Retrieve the seven-cycle experimental tubes from ice, return them to the thermal cycler, and subject them to additional PCR cycles to reach the optimal number indicated in the control cdna experiment. Next, immediately subject the tubes to an additional nine cycles of PCR. In total, the experimental tubes are subjected to X+9 PCR cycles, where X is the optimal number of PCR cycles determined for the control tube. Typically this total (X+9) should be in the range of 16-24 cycles. The fewer the better. 9. Analyze 5 µl of each experimental PCR reaction alongside 5 µl of the control PCR reaction representing the optimal number of PCR cycles on a 1% TAE gel.
Control samples should show bands to some extent, depending on the sample. These indicate highly expressed genes. (See example gel image) Normalized samples (1, ½, or ¼) should be smoother with less visible bands. Identify the optimum enzyme treatment: the highest enzyme concentration producing a smooth smear lacking distinct bands, while maintaining a molecular weight range comparable to the control sample. 10. Once the optimum amount of template and number of cycles have been determined, scale up the reaction and amplify the remainder of chosen normalization treatment, using multiple 50 µl reactions per template if necessary. This recipe assumes 8 µl of template (i.e. scaled up two-fold from the test scale PCR) so if you use more template, adjust the water volume accordingly. 28.5 µl NFW 10 µl 5X Q5 PCR buffer 1 µl 10 mm dntps 2 µl primer PST-CA1 (10 µm) 0.5 µl Q5 DNA polymerase 11. Prepare each reaction by combining 42 µl master mix and 8 µl template, then amplify for the optimum number of cycles identified in test scale reactions (step 8 above). Use the following profile: 98 C for 0:30, (98 C for 0:10 60 C for 0:25 72 C for 3:00) x N, 72 C for 5:00 12. Check 5 µl of the product on a gel before proceeding. 13. Combine all amplified normalized cdna into one tube, and purify by ethanol precipitation, a column-based PCR cleanup kit, or using magnetic beads. Elute or dissolve in 30 µl 10 mm Tris, and quantify using A260 or a fluorescent DNA assay. You should have 4 µg of amplified cdna at this point. cdna fragmentation In this step, the normalized and amplified cdna is broken into shorter fragments suitable for sequencing library preparation. Sonication is preferred, although enzymatic methods (Fragmentase) could be substituted at this step.
1. First, check how each sample behaves under sonication by testing a range of sonication times. a. Prepare a sample of 1.5 µg amplified cdna in a total volume of 40 µl NFW. b. Sonicate in 10 sec bursts for a total of 5 minutes, resting 10 sec on ice between each burst. Sample 5 µl at 0:00, 0:30, 1:00, 2:00, and 5:00 and hold samples on ice. c. Prepare a 2% agarose gel and analyze all samples on the gel to identify the minimum sonication time where the majority of product is in the 200-400 bp range. 2. After identifying the optimum sonication conditions (above), prepare an additional sample of 2.1 µg in 60 µl NFW total volume, and sonicate for the optimum time. 3. Check 5 µl of each sample on a 2% gel to confirm that the optimum size range has been produced. If any samples appear insufficiently fragmented, sonicate again briefly and check an additional 5 µl on a gel. Fragment repair and tailing, and adaptor ligation In this step, the fragmented cdna is first endrepaired to provide suitable ends for T-A ligation. Then additional reagents are added to ligate partially double-stranded adaptors to the fragmented cdna. The structure of these adaptors prevents tandem ligation of multiple adaptors. 1. Prepare a diluted enzyme mixture by combining 0.9 µl T4 DNA polymerase (3 U µl -1 ) 12.5 µl T4 polynucleotide kinase (10 U µl -1 ) 0.5 µl Taq DNA polymerase (5 U µl -1 ) Aliquot this dilution in 5 µl volumes and store at -20 C. Avoid repeated freeze thaw cycles. 2. Combine 30 µl of each fragmented sample with a master mix (10 µl) containing (per sample). 1 µl diluted enzyme mixture (above) 4 µl 10X T4 DNA ligase buffer
2 µl 10 mm dntp 1 µl 10 mm ATP (note this is NOT datp) 2 µl 50% PEG-4000 3. Incubate in the following profile: 12 C for 15, 37 C for 15, 72 C for 20, then 4 C hold 4. Next, anneal double-stranded adaptors. Combine the following: 2 µl 100 µm PE-TOP oligo 2 µl 100 µm HT-BOT oligo 16 µl NFW 5. Heat to 95 C for 30 seconds, then place the adaptor mix on ice. 6. Prepare the following ligation master mix on ice (recipe shown for one sample): 6 µl NFW 1 µl 10X T4 DNA Ligase Buffer 2 µl T4 DNA Ligase (400 U µl -1 ) 1 µl adaptor mix (above) 7. Add 10 µl master mix to the 40 µl repaired cdna. 8. Incubate at 16 C for at least 6 hours to overnight. 9. Purify libraries (ethanol precipitate, column based PCR cleanup, or magnetic beads) and resuspend in 25 µl NFW. Amplification and barcoding In this step, the ligation constructs produced in the previous step are amplified using primers that incorporate sample-specific barcodes. If multiple samples will be sequenced in multiplex on the same lane, you should use dual-indexing regardless of the number of samples you re combining. i.e. each sample should get a unique combination of BC and HT oligos (i7 and i5 indices respectively). Its important to minimize cycle numbers at this step to avoid introducing artifacts.
1. First, identify optimal conditions in a test PCR. 2. Set up the following master mix at room temperature (recipe shown for one sample). For these test scale reactions, the same barcode oligos (BCxx, HTxx) can be used for all samples. 13.25 µl NFW 5.0 µl 5X Q5 buffer 0.5 µl 10 mm dntps 0.5 µl 10 µm BCxx (oligo with barcode 1) 0.5 µl 10 µm HTxx (oligo with barcode 2) 0.25 µl Q5 DNA polymerase 3. Combine 20 µl of mix with 5 µl of template for each sample to be tested. 4. Amplify using the following profile: 98 C x 0:30, (98 C x 0:10 65 C x 0:30 72 C x 0:30) * X cycles 5. Sample 5 µl at 10, 14, 18 and 22 cycles. 6. Analyze on a 1% agarose gel to identify the minimum cycle number required to produce a visible smear. If nothing is visible by X=22 cycles, repeat the test using more template (and obviously less water). 7. Once the optimum template amounts and cycle numbers are determined, set up preparation-scale PCR(s). Amplify all the remaining template in multiple 50 µl reactions (scaled up from previous reactions). 8. Prepare the following master mix (recipe shown for each 10 µl of template). This time it s essential to use unique barcodes (BCxx and HTxx) for each sample. 26.5 µl NFW 10.0 µl 5X Q5 buffer 1.0 µl 10 mm dntps 0.5 µl Q5 DNA polymerase 9. Combine 38 µl of mix with 10 µl of template for each sample to be tested. Add 1 µl 10 µm BCxx (oligo with barcode 1), and 1 µl 10 µm HTxx (oligo with barcode 2). 10. Amplify for the minimum cycle number identified in test scale PCRs, using the following profile: 98 C x 0:30, (98 C x 0:10 65 C x 0:30 72 C x 0:30) * X cycles, 72 for 5:00 Size-selection 1. Pour a 2% TAE agarose gel, using clean buffer with freshly added ethidium bromide for both the gel and the running buffer. Use wide combs to maximize the volume that can be loaded. Frequently you will need to tape two or more wells together.
2. Combine all PCRs from each sample with loading buffer and load the entire volume in a single well. 3. Run the gel until ~20 bp resolution is achieved. 4. To avoid damaging DNA, make sure the UV trans-illuminator is set to low intensity, and never let the gel be exposed to UV for more than ~30 seconds. 5. Excise the region of the smear spanning from 350-550 bp. Be sure to photograph the gel before and after cutting out this region to document what was cut. 6. Incubate the gel slice with 40-100 µl NFW (minimum volume to ensure contact with gel slice) and incubate at 4 C overnight. 7. Freeze samples then centrifuge at maximum speed (room temperature) for 10 minutes. Transfer supernatant containing eluted DNA into a new labeled tube. This is now ready for quantification by qpcr prior to pooling, purification, and sequencing.
Primer and adaptor sequences CA1-20TVN CA1-TS-YY (RNA) CA1 PST-CA1 PE-Top HT-Bot BCxx HTxx AAGCAGTGGTATCAACGCAGAGTACTTTTTTTTTTTTTTTTTTTTVN AAGCAGTGGTATCAACGCAGAGTACYYGGG AAGCAGTGGTATCAACGCAGAGTAC GCGGCTGGTGAAAGCAGTGGTATCAACGCAGAGTAC ACACTCTTTCCCTACACGACGCTCTTCCGATC*T /5Phos/GATCGGAAGAGCACACGTCTGAACTCCAGTCA CAAGCAGAAGACGGCATACGAGATAATCGTGTGACTGGAGTTCAGACGTGTGCTCTTCCGATC AATGATACGGCGACCACCGAGATCTACACCGAGAACACTCTTTCCCTACACGACGCTCTTCCGATCT Buffers and reagents for this protocol These buffers are supplied with the DSN (Evrogen) but can also be prepared from stock chemicals if needed. DSN Stop Solution: 10 mm EDTA DSN Storage Buffer: 50 mm Tris, ph 8.0 DSN Master Buffer: 100 mm Tris, 10 mm MgCl 2, 2 mm DTT, ph 8.0 DSN Hybridization Buffer: 200 mm HEPES, 2 M NaCl, ph 7.5