Documentation

Clontech Pico v2 kit. This uses random hexamer priming, without selection by polyT, to make cDNA. This means that it works very well when your starting material is highly degraded (e.g. fixed cells or tissues). The random priming also means that this kit amplifies cDNA from all transcripts, including highly abundant RNA molecules such as rRNAs. We tend to use this kit if 1) investigators are working with highly degraded starting material, particularly FFPE tissues, or 2) if they really prefer to study total transcriptomes, rather than mRNAs.

The protocol below uses the SMARTer Stranded Total RNA-Seq Kit, however another option for cDNA generation when starting with low-input is the Clontech ‘SMART Seq HT’ for High-throughput single-cell mRNA-seq. This is an excellent kit for preparing cDNA very low amounts of RNA (as little as 1-100 cells). This kit is analgous to the well-known Clontech SMART-v4 kit, but is a faster/abbreviated workflow which combines RT/PCR steps and is roughly 30% lower cost compared to SMART-V4.

What you’ll need

• ZapR v2
• RNase inhibitor (40U/ul)
• SeqAmp DNA Polymerase
• SeqAmp CB PCR Buffer (2x)
• SMART-Seq HT Oligonucleotide
• SMART Pico Oligos Mix v2
• 5x First-strand Buffer
• Nuclease-Free Water
• SMARTScribe RT (100 U/uL)
• ZapR Buffer (10X)
• Tris Buffer (5mM)
• PCR2 Primers v2

• SMART TSO Mix v2
• R-probes v2
• Control Total RNA (1 ug/uL)

Not included in kit:

• Eppendorf LoBind 96-well plates
• Eppendorf LoBind DNA tubes
• AMPure XP Beads (4C)

A few important comments before you start

Input RNA Quality and Quantity:

• Purified total RNA samples cannot be resuspended in TE Buffer or have any EDTA in them. They should be resuspended in nuclease-free water. Any EDTA will interfere with RNA fragmentation and will decrease the efficiency of the reverse transcription.
• Samples must be DNAse I treated.
• If working with high quality RNA (RIN score >4), follow Option 1 which includes fragmentation; the fragmentation time will vary depending in the RIN score.
• If working with FFPE samples or highly degraded RNA, follow Option 2 which does not include fragmentation.

Diluting Control RNA

• To 50 ng/uL
  • 38 uL nuclease-free water
  • 2 uL control RNA
• To 5 ng/uL
  • 45 uL nuclease-free water
  • 5 uL diluted Control RNA at 50 ng/uL
• To 0.25 ng/uL
• 95 uL nuclease-free water
• 5 uL diluted Control RNA at 5 ng/uL

Option 1: With Fragmentation (RIN > 4)

1. Mix the following on ice:

1. Incubate tubes at 94C in preheated, hot-lid thermal cycler for the amount of time recommended below:

1. Immediately place samples on ice for 2 minutes.

2. Prepare enough First-Strand Master Mix for all reactions plus 10% by combining the following on ice:

1. Add 7 uL of the First Strand Master Mix to each PCR strip tube, vortex for about 2 seconds, spin down briefly.

2. Incubate the tubes in a preheated hot-lid thermal cycler with the following program (OPT1_FRAG):

1. Leave the samples in the thermal cycler at 4C until the next step

Option 2: Without Fragmentation (RIN< 4 or FFPE Samples)

1. Mix the following on ice:

2 Incubate tubes at 72C preheated, hot-lid thermal cycler for exactly 3 minutes.

1. Immediately place samples on ice for 2 minutes.

2. Prepare enough First-Strand Master Mix for all reactions plus 10% by combining the following on ice:

1. Add 11 uL of the First Strand Master Mix to each PCR strip tube, vortex for about 2 seconds, spin down briefly.

2. Incubate the tubes in a preheated hot-lid thermal cycler with the following program (OPT2_NOFRAG):

1. Leave the samples in the thermal cycler at 4C until the next step

PCR1- Addition of Illumina Adapters and Indexes

1. Prepare enough PCR1 Master Mix for all reactions plus 10% by combining the following reagents in the order shown below, vortex, then spin down:

1. Add 28 uL of PCR Master Mix to each sample

2. Add 1 uL of each 5’ PCR Primer HT to each sample

3. Add 1 uL of each 3’ PCR Primer HT to each sample

4. Tap PCR strip tubes to mix and spin down briefly

5. Place tubes on preheated hot-lid thermal cycler for the following program: (bold denotes steps to be run for 5 cycles)

Purification of RNA-seq Library using AMPure Beads

1. Allow AMPure XP beads to come to room temperature for about 30 minutes before use. Vortex beads for 2 minutes to mix well. Make sure samples are in a 96 well plate that will fit on our magnetic stand. Prepare fresh 80% ethanol, you will about 400 uL per sample.

2. Add 40 uL of AMPure beads to each sample and pipette 10 times to mix.

3. Incubate at room temperature for 8 minutes to allow the DNA to bind to the beads.

4. Place plate on magnetic stand for 5 minutes or longer until the solution is completely clear.

5. While samples are on magnetic stand, remove and discard supernatant. Do not disrupt beads.

6. Keeping samples on magnetic stand, add 180 uL of freshly made 80% ethanol to each sample without disturbing beads. Wait 30 seconds and carefully pipette out and discard supernatant. cDNA will remain bound to beads during the washing process.

7. Repeat wash step above 1 more time.

8. Let samples air dry for about 1 minute on magnetic stand then remove any excess ethanol with a P20.

9. Air dry samples on magnetic stand for 3-5 minutes at room temperature until pellets appear dry and matte. Once you start to see pellets crack, take off magnetic stand to ensure you do not overdry your samples.

10. Once beads are dry, remove from magentic stand and add 52 uL of Nuclease- free water to cover beads, pipette to mix thoroughly until the beads are resuspended.

11. Incubate at room temperature for 5 minutes.

12. Place plate back on magnetic stand for 1 minute or longer until the supernatant is clear.

13. Pipette 50 uL of supernatant to new plate

14. Add 40 uL of vortexed AMPure XP beads to each sample and pipette to mix thoroughly 10 times.

15. Incubate at room temperature for 8 minutes to allow DNA to bind to the beads. During incubation, proceed to next section.

Depletion of Ribosomal cDNA with ZapR v2 and R-Probes v2

In this section, the library fragments originating from rRNA (18S and 28S) and mitochondrial rRNA (m12S and m16S) are cut by ZapR v2 in the presence of R-Probes v2 (mammalian-specific). These R-probes hybridize to ribosomal RNA and mitochondrial rRNA sequences.

• Must make fresh aliquot of 80% ethanol if doing on different day than previous section, you will need 400 uL per sample.

1. Thaw R-probes v2 (-80C, Box J8) and ZapR Buffer (-20C) at room temperature. Once R-probes v2 thaw, place immediately on ice but keep ZapR Buffer at room temperature. Thaw ZapR v2 (-20C) on ice and keep on ice at all times, return to freezer immediately after use.

2. Preheat PCR machine to 72C

3. Once 8 minute incubation of samples and AMPure XP beads is finished, place samples on magnetic stand for 5 minutes or longer until the solution is completely clear.

4. During the 5 minute incubation, pipette 1.5 uL of R-Probes v2 +10% per reaction into a PCR Strip tube on ice and immediately return R-Probes v2 to -80C freezer.

5. Incubate the PCR strip tube of R-probes v2 at 72C in preheated hot-lid thermal cycler using the following program:

1. The PCR strip tube of R-probes should be held at 4C for anywhere between 2-10 minutes. It must be held for at least 2 minutes but not for more than 15 minutes.

2. Once the 5 minute incubation of RNA samples is finished on magnetic stand, pipette and discard clear supernatant without disturbing the beads.

3. Keeping the samples on the magnetic stand, add 180 uL of freshly made 80% ethanol to each sample without disturbing beads. Wait 30 seconds and carefully pipette out and discard supernatant. cDNA will remain bound to beads during the washing process.

4. Repeat wash step above 1 more time.

5. Let samples air dry for about 1 minute on magnetic stand then remove any excess ethanol with a P20.

6. Air dry samples on magnetic stand for 1-2 minutes at room temperature until pellets appear dry and matte. Once you start to see pellets crack, take off magnetic stand to ensure you do not overdry your samples.

7. While beads are drying, prepare the ZapR Master Mix for all reactions plus 10% by combining the following reagents in the order shown below at room temperature, vortex, then spin down:

1. Take plate off magnetic stand. To each well of dried AMPure XP beads, add 22 uL of ZapR Master Mix and pipette to mix thoroughly until the beads are resuspended.

2. Incubate at room temperature for 5 minutes.

3. Place plate back on magnetic stand for 1 minute or longer until the supernatant is clear.

4. Pipette out 20 uL of clear supernatant, without disturbing beads, to a new PCR strip tube.

5. Incubate PCR strip tubes containing supernatant in a preheated hot-lid thermal cycler using the following program:

• Samples can be held at 4C for up to 1 hour but it is recommended that you proceed immediately to next section.

PCR2- Final RNA-Seq Library Amplification

In this section, the library fragments not cleaved by the ZapR reaction will be further enriched in a second round of PCR. Illumina barcodes have already been added to the libraries so this step includes a single pair of primers that are used for all libraries.

1. Prepare a PCR2 Master Mix for all reactions plus 10% by combining the following reagents in the order shown below, vortex, then spin down:

1. Add 80 uL of PCR2 Master Mix to each tube. Mix by tapping gently and spin down.

1. Place tubes in preheated hot-lid thermal cycler for the following program (PCR2):

(bold denotes steps to be run for 9 to 16 cycles, depending on your input RNA, see the second table for the number of cycles.)

Purification of Final RNA-Seq Library Using AMPure Beads

1. Allow AMPure XP beads to come to room temperature for about 30 minutes before use. Vortex beads for 2 minutes to mix well. Make sure samples are in a 96 well plate that will fit on our magnetic stand. Prepare fresh 80% ethanol, you will about 400 uL per sample.

2. Add 100 uL of AMPure beads to each sample and pipette 10 times to mix. [This is a 1:1 ratio; if you kept each sample separated into 2 wells, add 50 uL of beads. If all 100uL of PCR product is one a single well, add 100uL of beads very carefully because this will completely fill the wells.]

3. Incubate at room temperature for 8 minutes to allow the DNA to bind to the beads.

4. Place plate on magnetic stand for 5 minutes or longer until the solution is completely clear.

5. While samples are on magnetic stand, remove and discard supernatant. Do not disrupt beads.

6. Keeping samples on magnetic stand, add 180 uL (use 180 uL per well regardless of whether samples were split) of freshly made 80% ethanol to each sample without disturbing beads. Wait 30 seconds and carefully pipette out and discard supernatant. cDNA will remain bound to beads during the washing process.

7. Repeat wash step above 1 more time.

8. Let samples air dry for about 1 minute on magnetic stand then remove any excess ethanol with a P20.

9. Air dry samples on magnetic stand for 3-5 minutes at room temperature until pellets appear dry and matte. Once you start to see pellets crack, take off magnetic stand to ensure you do not overdry your samples.

10. Once beads are dry, remove from magentic stand and add 20 uL (10uL per well if samples still split) of Tris Buffer to cover beads, pipette to mix thoroughly until the beads are resuspended.

11. Incubate at room temperature for 5 minutes.

12. Place plate back on magnetic stand for 1 minute or longer until the supernatant is clear.

13. Pipette 18 uL (9 uL per well if samples still split) supernatant to LoBind Tubes. This is the final product. If samples were split, recombine supernatant from both halves into a single tube with 18uL total.

Quality Check of Final Product

1. Run samples on HSD1000 Tapestation Assay.

2. Measure concentration with HS dsDNA Qubit assay.

Sequencing Guidelines

Normalize and Pool

1. If not done already, 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 the library prep kit “TruSeqHT” 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.