This protocol uses the Takara/Clontech SMART-seq HT Plus kit for carrying out RNA-seq when you have very low numbers of cells for input (10’s, 100’s or 1000’s of cells).
- Before starting
- What you’ll need
- A few important comments
- Day 1: Sample preparation
- OPTION A: sorting into media
- OPTION B: sorting into lysis buffer
- Day 2: cDNA and library prep
- RNA extraction
- Clontech SMART-Seq cDNA Synthesis
- CDNA clean-up
- HT PLUS Library Prep
- A few important comments before you start
- Step 1: Adaptors
- Step 2: Amplification
- Step 3: Library Cleanup
- Step 4: Quality Check
- Sequencing Guidelines
- Normalize and Pool
- Setup Run in Basespace
- Loading the Sequencer
Clontech ‘SMART-Seq HT PLUS’ 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 analagous 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. The PLUS version includes both cDNA generation and library preparation all in one kit.
The protocol below uses the SMART-Seq HT kit, however another option for cDNA generation when starting with low-input is the Clontech Pico v2 kit. This has a few important difference from the SMART-Seq HT kit above. First, the Pico v2 kit uses random hexamer priming, rather than 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.
What you’ll need
The following items are included in the Clontech SMART-Seq HT kit and are stored at -20C
- 10x Lysis buffer
- RNase inhibitor (40U/ul)
- SeqAmp DNA Polymerase
- One-step Buffer
- SMART-Seq HT Oligonucleotide
- 3’ SMART-Seq CDS Primer II A
- SMARTScribe Reverse Transcriptase (100 U/µl)
- Nuclease-Free Water
- Elution Buffer (10 mM Tris-Cl, pH 8.5)
- Eppendorf LoBind DNA tubes
A few important comments
- Prior to sorting, stain cells in 10% complete media (RPMI with hepes, 10% FCS, Pen/Strep, NEAA, L-glut, Na-Pyruvate, 2ME, etc.)
- You’ll split this protocol into two days: day 1 will be your sort and cell lysis, day 2 will be RNA isolation and/or cDNA generation, and a modified NexteraXT library prep.
- Use only filter pipette tips and clean your area so it is free of RNases.
- Certain steps, such as 1st-strand cDNA synthesis must be carried out in a PCR clean hood
- Be sure to include the kit positive control and a water-only negative control with each experiment.
- the SMART-Seq HT kit uses a 96-well plate for mixing reactions. Do not use tubes.
- Similar to other Clontech low-input kits, SMART-Seq HT uses a template switching method to produce abundant cDNA directly from as few as 1-100 cells or from 10pg-1ng of total RNA.
- This protocol is very sensitive to variations in volume and other factors. Please make sure the pipettes are calibrated and avoid contamination.
Day 1: Sample preparation
Sorting and handling of samples prior to RNA/cDNA/library prep, is the most important factor that will influence your ability to make high quality sequence-ready libraries.
OPTION A: sorting into media
We recommend Option A if you plan to sort > 3000 cells. Following sorting, RNA is extracted and used as input for cDNA generation.
There is a lot of variability in terms of how sensitive cells are to the sorting process – naïve and effector T cells do great, others (like Treg cells) are way more fragile. For these more challenging cell types, you should plan to sort 3-4 times more cells than you need.
- Always use the 100 uM nozzle on the sorter (rather than 70 uM) and sort at low speed (~5-7K events/sec). This will be more gentle on the cells
- Sort into Lo-bind 96-well plates to prevent cells from sticking to sides (Lo-bind gives tighter pellets after spin down)
- Sort into complete media with 50% serum (RPMI with hepes, Pen/Strep, NEAA, L-glut, Na-Pyruvate, 2ME, etc.)
Check the purity of your sort by taking 5-10 ul and running back throught the cytometer.
- Keep everything cold: cells, sort chamber, collection block, collection tubes, etc.
- Minimize time after sort to lysis. If sorting many samples, spin down in batches – don’t wait until the end if possible.
- Spin down cells. ~10,000 cells usually produce a visible pellet. Use a pipette tip to carefully draw off supe.
- lyse cells by adding 300 ul of buffer RLT (from Qiagen RNeasy kit) with 2-ME added fresh. Vortex to lyse cells.
- Flash freeze tube on dry ice and transfer to a prechilled box at -80C.
OPTION B: sorting into lysis buffer
We recommend Option B if you plan to sort < 3000 cells. You will sort directly into lysis buffer and skip RNA extraction entirely, using the crude cell lysate for input directly into the cDNA reaction.
The 10x lysis buffer used in the next step contains detergent, so avoid creating bubbles when pipetting.
- Prepare 10x reaction buffer from the SMART-Seq HT kit by mixing 19 uL of 10x Lysis Buffer with 1 uL of RNase inhibitor. This is enough for ~20 samples. Scale up as needed, but be sure to maintain 19:1 ratio of lysis buffer to RNase inhibitor
- Prepare 1x reaction buffer by mixing 9.5 uL nuclease-free water with 1 uL of 10x reaction buffer. This is enough to sort one sample, so be sure to scale up as needed.
- Add 5ul of 1x reaction buffer to collection eppendorf tube (or one well of a 96-well plate) and sort directly into this tube or well.
- Immediately after sample is sorted, add an additional 5.5 uL of 1x reaction buffer to tube.
- Store at -80C until ready to begin cDNA synthesis. You can proceed directly from this cell lysate to cDNA preparation with no intermediate RNA isolation step.
Day 2: cDNA and library prep
If preferred, the remaining steps could be split into two days, with RNA extraction and cDNA generation in one day, and the modified Nextera library prep the following day.
- If you chose Option A above, extract RNA using the the Qiagen RNeasy micro kit and store at -80C.
- If you chose Option B above, skip RNA extraction and proceed directly to cDNA synthesis
- Assess RNA integrity using an Agilent Tapestation 4200 and High-sensitivity RNA screentape. Although tapestation is not ideal for estimating concentration of RNA, it can be used in this case as a rough estimate, since you won’t have enough material to quantify using Qubit.
Clontech SMART-Seq cDNA Synthesis
|Kit reagent||Volume per rxn (uL)|
SMART-Seq HT Oligonucleotide
|Kit reagent||Volume per rxn (uL)|
SeqAmp DNA Polymerase
SMARTScribe Reverse Transcriptase
|Input of total RNA (or amount of cells)||Typical number of PCR cycles|
1 ng (or about 100 cells)
100 pg (or about 10 cells)
10 pg (or about 1 cells)
We have found that the recommended 10-11 cycles is not quite enough to amplify 1 ng RNA. We usually do 15 cycles for 1 ng and 17 cycles for 100 pg.
The beads contain your cDNA so any beads expelled will cause you to lose part of your samples.
Your cDNA is in now eluted from the beads, so do not discard supernatant
HT PLUS Library Prep
A few important comments before you start
- You'll need 8uL of cDNA, diluted to a concentration of 0.125 ng/ul - 1.25 ng/ul or a total input of 1 - 10ng. All samples should have the same input. If using less than 8ul of cDNA, bring the volume to 8ul with nuclease-free water.
- If your sample is less than 0.125 ng/ul, you can attempt to move forward with the library prep without dilution. cDNA for negative controls of this kit can have up to 100pg/ul of contamination after 17 PCR cycles. If you have a low concentration sample, judge based on your Qubit concentration and cycles used whether to move forward. Tapestation concentrations are typically inaccurate for this.
- Especially if working with unfamiliar samples, including positive and negative controls is encouraged. Negative controls can be 8ul of undiluted products of a negative control from the cDNA step or 8ul of elution buffer. Positive controls must be included in the cDNA step and products can move forward with dilution just like experimental samples.
Step 1: Adaptors
- Stem-loop adaptors (purple)
- FE Dilution Buffer (white)
- Lib Prep Buffer (blue)
- Rxn Enhancer (red)
- Amplification Buffer (orange)
Library Prep Buffer
Library Prep Enzyme
Safe stopping point. The samples can be stored in the thermal cycler at 4°C overnight, or they can be transferred to –20°C for up to a week.
Step 2: Amplification
PrimeSTAR HS DNA Polymerase
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. Generally, moving in columns is fine. Seal the used wells with foil and label with your initials and date used before returning to freezer storage. Record which index well was used for each sample, as this is needed to demultiplex after sequencing.
Step 3: Library Cleanup
Step 4: Quality Check
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 HSD5000. 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. If a single peak has not been identified but the library is still sequencable, then use the average bp size value.
- 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 as IDT-ILMN TruSeq DNA-RNA UD 96 Indexes. 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 76 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.
- Your final loading concentration should be 1.5 - 1.8 pM, with most pools loaded at 1.6-1.7pM.