Single Cell RNAseq – library prep

Before you start

Overview

Day 1 - thaw samples, enzymatic digestion to remove the unwanted nucleic acids and primer dimers, then second strand synthesis. Linear amplification of the dsDNA made during second stand synthesis that takes place over night (between 13-15 hour incubation).

Day 2 - Check library on tapestation/bioanalyzer, amplified RNA is fragmented and used for second reverse transcription reaction to convert sample back to DNA and to add a read primer binding site to each molecule. Clean up with Ampure XP beads, amplified by PCR. qPCR is performed to determine optimal amount of cycles for the final PCR; once final PCR is finished, library can be QCed by tapestation and Qubit.

Bead clean ups are performed after every enzymatic step

Full protocol can be found here

Reagents

• Nuclease-free water, Life Technologies, cat. no. 10977-023
• Agencourt AMPure XP magnetic beads, Beckman Coulter, cat. no. A63881
• Exonuclease I, ExoI; 20,000 U/mL; NEB, cat. no. M0293L
• FastDigest HinFI restriction endonuclease, HinFI; Thermo Fisher/Life Technologies, cat. no. FD0804
• FastDigest Buffer 10x, ThermoFisher/Life Technologies, cat. no. B64
• NEBNext® mRNA Second Strand Synthesis Module, NEB, cat. no. E6111S
• HiScribeTM T7 High Yield RNA Synthesis Kit, NEB, cat. no. E2040S
• RNA Fragmentation Reagents, Ambion/Life technologies, cat. no. AM8740
• PrimeScriptTM Reverse Transcriptase, Takara Clontec, cat. no. 2680A
• RNaseOUT Recombinant Ribonuclease Inhibitor, RNaseOUT,ThermoFisher Scientific, cat. no. 10777-019
• Deoxynucleotide (dNTP) Solution Mix, 10 mM each; NEB, cat. no. N0447L
• Kapa 2x HiFi Hot Start PCR mix, Kapa Biosystems, cat. no. KK2601
• Eva Green Dye, 20x in water; Biotium, cat. no. 31000-T
• 100 μM PE2-N6 primer (1Cell)
• 5 μM PE1/PE2 primer mixture with indices (1Cell)
• 100 μM Custom read 1, read2, and index primers (1Cell)
• Corning Costar Spin-X tube filters, Sigma-Aldrich, cat. no. CLS8162
• Bioanalyzer RNA and HSDNA chips and reagents or Agilent TapeStation HSRNA and HSD1000screen tapes and reagents (Aglient)

Equipment

• Magnetic racks for PCR and/or 1.5 ml tubes
• Centrifuge (e.g., Eppendorf AG, Cat. No. 5424R)
• Thermo-conductive tube rack (VWR Scientific Cat. No. 95045-464)
• Thermal cycler
• qPCR machine
• Thermomixer or dry bath for 1.5 ml tubes
• BioAnalyzer (Agilent) or Tapestation (Agilent)

RNA Elution Buffer (RE)

• 150 uL of 1 M Tris-HCl pH 7.5
• 3 uL of 0.5 M EDTA
• Add nuclease-free water to bring the final volume to 15 mL
• This can be aliquoted into nuclease-free tubes and aliquots can be stored at 4C short term or -20C long term.

DNA Suspension Buffer (DS)

• 150 uL of 1 M Tris-HCl pH 8.0
• 3 uL of 0.5 M EDTA
• Add nuclease-free water to bring the final volume to 15 mL
• This can be aliquoted into nuclease-free tubes and aliquots can be stored at 4C short term or -20C long term.

Blocking Buffer (BB)

used to block the pipet tips so your solution will not stick to the outside or the inside of your tip, block tip for all steps that you are pipetting just your sample):

• 100 uL of 1 M Tris-HCl pH 8.0
• 2 uL of 0.5 M EDTA
• 500 uL of 10% Tween 20 (vol/vol)
• 500 uL of 2% BSA (wt/vol)
• 8898 uL of Sterile Water Total volume is 10 mL, aliquot into 1.5 mL tubes with approximately 0.5 mL per tube, store at -20C and thaw before library prep

80% Ethanol

• Make fresh each day for Ampure XP bead clean ups

Day 1

Enzymatic Digestion (2hrs)

1. Thaw post-RT libaries on ice
2. Centrifuge at 4C for 5 mins at 19,000 rcf to pellet any cell debris in the tube, this will bring down any material more dense than your samples.
3. Prepare the Digestion Mix. For every 70uL of post-RT material, prepare 100uL of Digestion Mix (1000 cells is approximately equal to 35uL of post-RT material so if you sorted 3000 cells, make 150uL of Digestion Mix)

Volumes for Digestion Mix:

1. Without aspirating HFE-7500 oil from the bottom of the tube, carefully transfer the post-RT material from each sample to its own Costar Spin-X tube filter.

2. Add Digestion mix made in previous step to the top of your old tube with the HFE-7500 oil to wash off any residual sample that may still be on the top of the oil. Then transfer the digestion buffer mix to the Costar-x spin tube.

3. Centrifuge the filter column at 4C for 5 mins at >5000 xg.

4. Discard the filter membrane (this contains the beads and all the particles we wanted to filter out, your sample is in the supernatant in the flow through).

5. Incubate the flow-through sample at 37C for 30 minutes.

6. Purify the reaction by using AmpPure XP beads. At this stage, your products are RNA-cDNA hybrids. This is the general protocol for all AmpPure XP bead clean ups. Pay attention to the ratio in each step of beads: sample, the elution buffer used and the amount of elution buffer in each clean up. These will be different in the subsequent steps.

• Ratio of AMpure XP beads to sample is 1.2X ; The elution buffer used here is nuclease free water at a volume of 19 uL

• Add volume of beads directly to the sample. Vortex to mix well.
• Incubate the sample at room temperature for 5 minutes. Your sample will bind to the beads in this step.
• Briefly spin down tube.
• Place the sample(s) on the magnetic stand for 3-5 minutes until the bead collects on the side of the well and the supernatant is clear.
• Aspirate and discard the supernatant without disturbing the beads.
• Keep the sample(s) on the magent and fill with 80% etoh making sure the bead is completely covered. Remove etoh from sample after 30 seconds.
• Repeat wash step once more.
• After the second wash, spin tube down to get any excess etoh off the bead, aspirate out and discard.
• Dry tubes at 37C with caps off for about 3 minutes or until the beads are dry. They should appear matte and begin to crack when dry.
• Add 19uL of elutent and vortex, spin down, and incubate at room temperature for 5 minutes.
• Return the tubes to the magnetic stand and incubate for 1 minute until supernatant is clear. supernatant now contains your sample.
• Transfer 17uL of the supernatant to a fresh PCR tube.

Second Strand Synthesis and In Vitro-Transciption (IVT)

Time: 16-18 hours

1. Set up Second Strand Synthesis (SSS) on ice:
   • Add 2 uL of 10x SSS buffer to the 17 uL eluate from above in the PCR tube.
   • Mix by vortexing, spin down. Add 1 uL of SSS enzyme and mix gently by pipetting up and down.
   • Incubate the reaction at 16C for 2.5 hours, then 65C for 20 minutes.
2. Make a master mix of the following components by multiplying the volume by the amount of samples you have; add 60 uL to each sample:

1. Pipette to mix, spin plate down, and incubate at 37C for 13-15 hours with the lid heated to 50C.

• Pause point After 15h the IVT reaction can be kept at 4C for two hours, this is programmed into our PCR protocol on our machine.

Day 2

1. Immediately after incubation, purify the reaction by using AmpPure XP beads. This is the general protocol for all AmpPure XP bead clean ups. Pay attention to the ratio in each step of beads: sample, the elution buffer used and the amount of elution buffer in each clean up. These will be different in the subsequent steps.

• Ratio of AMpure XP beads to sample is 1.3X (104uL of beads) ; The elution buffer used here is RNA Elution Buffer at a volume of 21 uL

• Add volume of beads directly to the sample. Vortex to mix well.
• Incubate the sample at room temperature for 5 minutes. Your sample will bind to the beads in this step.
• Briefly spin down tube.
• Place the sample(s) on the magnetic stand for 3-5 minutes until the bead collects on the side of the well and the supernatant is clear.
• Aspirate and discard the supernatant without disturbing the beads.
• Keep the sample(s) on the magent and fill with 80% etoh making sure the bead is completely covered. Remove etoh from sample after 30 seconds.
• Repeat wash step once more.
• After the second wash, spin tube down to get any excess etoh off the bead, aspirate out and discard.
• Dry tubes at 37C with caps off for about 3 minutes or until the beads are dry. They should appear matte and begin to crack when dry.
• Add 21uL of elutent and pipette to mix, spin down, and incubate at room temperature for 5 minutes.
• Return the tubes to the magnetic stand and incubate for 1 minute until supernatant is clear. The supernatant now contains your sample.
• Transfer 9uL of supernatant to a well in a new plate. Transfer 10uL to a LoBind tube and label as “post-IVT back up”.

1. Run HS RNA Tapestation on the post-IVT back up. Then put “post-IVT back up” into the -80C freezer, where it can be stored for up to 6 months.

Fragmentation of Amplified RNA

Time: 20 mins [SKIP THIS STEP AND PROCEED TO REVERSE TRANSCRIPTION USING RANDOM HEXAMERS IF MAJORITY OF SAMPLE IS <1000 NT]

1. Prepare Fragmentation Stop Mix for each sample:

1. Vortex the solution from above and place on ice.

2. Allow the thermalcycler to reach 70C (lid at 100-110C)

3. On ice, add 1 uL of 10X RNA Fragmentation reagent to 9 uL of purified of IVT product from previous step.

4. Spin down plate and fragment on thermalcycler at 70C for about 1-3 minutes, depending on what your post IVT tapestation looked like. Here is how to determine the fragmentation time:

• 1cellbio’s general rule is no fragmentation if the bulk of the sample is below 1000nt, 1-2 mins if the bulk of the sample is between 1000-2000nt, and 3 mins if the majority of your sample is above 2000nt.

1. Cool the fragmentation reaction on ice and immediately add 34uL of Fragmentation Stop Mix (prepared in step 14) to each sample and pipette up and down to mix well. Spin plate very briefly.

2. Incubate samples on ice for 5 minutes.

3. Plate samples on magnetic rack and continue through the AMPure XP bead clean up steps below:

Ratio of AMpure XP beads to sample is 1.3X (104uL of beads) ; The elution buffer used here is RNA Elution Buffer at a volume of 10 uL

• Place the sample(s) on the magnetic stand for 3-5 minutes until the bead collects on the side of the well and the supernatant is clear.
• Aspirate and discard the supernatant without disturbing the beads.
• Keep the sample(s) on the magent and fill with 80% etoh making sure the bead is completely covered. Remove etoh from sample after 30 seconds.
• Repeat wash step once more.
• After the second wash, spin tube down to get any excess etoh off the bead, aspirate out and discard.
• Dry tubes or plate at 37C with caps off for about 3 minutes or until the beads are dry. They should appear matte and begin to crack when dry.
• Add 10uL of elutent and pipette to mix, spin down, and incubate at room temperature for 5 minutes.
• Return the tubes to the magnetic stand and incubate for 1 minute until supernatant is clear. Supernatant now contains your sample.
• Transfer 8uL of supernatant to a new well.

Reverse Transcription using Random Hexamers

Time: 2.5 hours

1. On ice, combine the following:

1. Pipette to mix, spin down. Incubate at 70C (lid at 105C) for exactly 3 minutes and cool immediately on ice.

2. On ice, add the following components to the mix to bring the total volume to 20uL:

1. Mix gently by pipetting and incubate the reaction at 30C for 10 minutes then 42C for 1 hour then 70C for 15 minutes to heat inactivate the enzyme.

2. Add 20uL of nuclease-free water, mix by pipetting, and save 20 uL as “pre-PCR back-up” in a LoBind tube. This back up can be stored at -80C for up to 6 months.

3. Purify the reaction with AMPure XP Beads

• Ratio of AMpure XP beads to sample is 1.2X (24 uL) ; The elution buffer used here is DNA elution buffer at a volume of 12 uL

• Add volume of beads directly to the sample. Pipette to mix well.
• Incubate the sample at room temperature for 5 minutes. Your sample will bind to the beads in this step.
• Place the sample(s) on the magnetic stand for 3-5 minutes until the bead collects on the side of the well and the supernatant is clear.
• Aspirate and discard the supernatant without disturbing the beads.
• Keep the sample(s) on the magent and fill with 80% etoh making sure the bead is completely covered. Remove etoh from sample after 30 seconds. The ethanol should fully cover the bead in this step, in a plate add 180 uL of the 80% ethanol.
• Repeat wash step once more.
• After the second wash, use a P10 to get remaining ethanol out of sample and discard.
• Dry samples at 37C with no cover for about 3 minutes or until the beads are dry. They should appear matte and begin to crack when dry.
• Add 12uL of elutent and pipette to mix.
• Incubate at room temperature for 5 minutes.
• Return the tubes to the magnetic stand and incubate for 1 minute until supernatant is clear. Supernatant now contains your sample.
• Transfer 10 uL of supernatant to a new well.

Diagnostic qPCR

Time: 1.5-2 hours

1. Perform a quantitative PCR (qPCR) to determine the correct number of PCR amplification cycles in the next step. Combine the following components on ice:

1. Perform the following qPCR thermalcycling program:

Eva green uses the same excitation and emission settings as SYBR Green.

Italics denote steps to be run for 2 cycles.

Bold denotes steps to be run for 24 cycles.

1. Set the qPCR threshold within the exponential phase of amplification, but closer to where the signal starts to emerge from the noise to avoid the possibilty of over-amplification.
   • To account for the differences of input material between the qPCR and the PCR amplification of our samples, we will calculate the amount of PCR cycles to amplify our sample with now.
   • The library input is 19 times higher than the qPCR input, therefore the difference corresponds to log2^19 = 4.25 cycles. For example, if the number of cycles determines by the qPCR is 2 + 12.67, the required number of PCR cycles for library amplification is 2 + (12.67 -4.25) = 2 + 8.

Library Amplification

Time: 2 hours

1. Combine the following components on ice:

1. Perform thermal cycling protocol shown in step 29, also shown again below:

Italics denote steps to be run for 2 cycles.

Bold denotes steps to be run for amount of cycles determined by qPCR.

• If working with multiple libraries that require different cycle numbers, set the thermal cycler to the maximum number of cycles needed, then manually remove each individual sample when they have reached the desired cycle number.

• Move the individual libraries to an ice bucket while the rest of the libraries are amplifying.

• Once every library has been amplified for the correct amount of cycles, transfer all libraries back to the PCR machine for the final step, 72C for 5 minutes.

1. Add 25 uL of DNA Elution Buffer to the PCR product.

2. Purify using AMPure XP beads.

• Ratio of AMpure XP beads to sample is 0.7X (35 uL) ; The elution buffer used here is DNA elution buffer at a volume of 22 uL

• Add volume of beads directly to the sample. Pipette to mix well.
• Incubate the sample at room temperature for 5 minutes. Your sample will bind to the beads in this step.
• Place the sample(s) on the magnetic stand for 3-5 minutes until the bead collects on the side of the well and the supernatant is clear.
• Aspirate and discard the supernatant without disturbing the beads.
• Keep the sample(s) on the magent and fill with 80% etoh making sure the bead is completely covered. Remove etoh from sample after 30 seconds. The ethanol should fully cover the bead in this step, in a plate add 180 uL of the 80% ethanol.
• Repeat wash step once more.
• After the second wash, use a P10 to get remaining ethanol out of sample and discard.
• Dry samples at 37C with no cover for about 3 minutes or until the beads are dry. They should appear matte and begin to crack when dry.
• Add 22uL of elutent and pipette to mix.
• Incubate at room temperature for 5 minutes.
• Return the tubes to the magnetic stand and incubate for 1 minute until supernatant is clear. Supernatant now contains your sample.
• Transfer 20 uL of supernatant to a new well.

This eluate is the final sequencing-ready library.

1. Test library by running on the HS D1000. The library should look like a smooth hump starting at 200bp and ending at 1.5kb with an average size between 400 and 600bp. This library distribution will vary from sample to sample.

2. Perform Qubit on samples using the HS DNA assay to get the concentration for pooling and sequencing.

Pause Point: store the libraries at -80C or -20C. They should be stable for at least 1 year.

Sequencing Guidelines

• We sequence on a 75 Cycle High Output NextSeq 500 kit. The run cycles are 36 x 50.

• This library prep kit comes with custom primers for Read 1, Read 2, and Index reads. These are in the library preparation box in the -20C and they are to be spiked into the sequencing cartridge in wells 7, 8, and 9 respectively.

• Add 6ul of each primer to 1.994mL HT1, each in its own 5mL tube.

• When ready to spike into wells of sequencing cartridge, add the 2mL mixture to the appropriate well. (7 = read 1, 8 = read 2 and 9 = index)

• Ensure custom indexes R1, R2, and Index are checked off when programming run on Basespace and ensure these boxes are checked off when viewing run parameters on the sequencer.