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
- Documentation
- What you’ll need
- A few important comments
- Sample preparation
- OPTION A: sorting into media
- OPTION B: sorting into lysis buffer
- Protocol
- Step 1: Generate cDNA from total RNA
- RNA extraction
- cDNA Synthesis
- Step 2: cDNA clean-up
- HT PLUS Library Prep
- A few important comments before you start
- Step 3: Adaptors
- Step 2: Amplification
- Step 3: Library Cleanup
- Quality Checks
- Sequencing Guidelines
- Normalize and Pool
Before starting
Documentation
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.
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 -80°C.
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 -80°C until ready to begin cDNA synthesis. You can proceed directly from this cell lysate to cDNA preparation with no intermediate RNA isolation step.
Protocol
Step 1: Generate cDNA from total RNA
The protocol is generally split into two days, with cDNA generation in one day, and library generation the following day.
RNA extraction
- If you chose Option A above, extract RNA using the the Qiagen RNeasy micro kit and store at -80°C.
- 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 may not have enough material to quantify using Qubit.
- If you have sample to spare, you can try to measure RNA concentration using Qubit, but it is unlikely to be above the limit of sensitivity (2 ng/ul). A more practical approach is the use the concentrations provided by High-sensitivity RNA tapestation run. These aren’t ideal, but they’ll have to do.
- Especially if working with unfamiliar samples, including positive and negative controls is encouraged. Negative controls can be 10.5 of Nuclease-free water. Positive controls must be included in the cDNA step and products can move forward with dilution just like experimental samples.
cDNA Synthesis
Thaw the following reagents as instructed:
Name | Location | Action |
|---|---|---|
One-step Buffer | -20°C freezer | Thaw at room temperature |
10X Reaction Buffer | -20°C Freezer | If made, thaw on ice; if not made, see below |
Nuclease-free Water | -20°C freezer | Thaw at room temperature |
3’ SMARTSeq CDS Primer II | -20°C Freezer | Thaw on ice |
SMART-Seq HT Oligonucleotide | -20°C freezer | Thaw on ice |
RNase Inhibitor (RRI) | -20°C freezer | Thaw on ice |
Elution Buffer | -20°C freezer | Thaw at room temperature |
SeqAmp DNA Polymerase | -20°C freezer | Store until needed |
SMARTScribe Reverse Transcriptase | -20°C freezer | Store until needed |
Make stock of 10x Reaction Buffer if not already made (can be scaled up as needed, we usually make a stock of this and keep it in our -20C):
If you have ample RNA, dilute to 0.3-0.5 ng/µL (this is not detectable on the Qubit, so you cannot quantify after diluting). This will allow you to add 10.5ul of RNA to the cDNA reaction, and not exceed 1 ng.
In the pre-PCR biosafety cabinet, add 10.5 uL of diluted RNA or cell lysate to each well to bring your input to 1 ng. If using less than 10.5 uL of RNA, add nuclease-free water to 10.5 uL. Normalize all samples so they have the same input. Keep plate on ice while working with samples.
To each well, add 1 uL of 10x Reaction buffer.
To each well, add 1 uL of 3’ SMART-Seq CDS Primer II A.
Mix gently by pipetting with a p20 at 11 µL.
Seal plate and centrifuge for 10 seconds.
Before denaturing your samples, prepare a master mix for cDNA synthesis (10% overage has already been included). Keep this on ice until ready to use.
Kit reagent | Volume per rxn (uL) |
|---|---|
Nuclease-free water | 0.77 |
One-Step buffer | 8.8 |
SMART-Seq HT Oligonucleotide | 1.1 |
RNase Inhibitor | 0.55 |
Denature your samples by incubating the plate at 72°C in a preheated, hot-lid thermal cycler for 3 minutes. The thermal cycler programs for this protocol can be found under CHMI → Clontech → 72 3.
Immediately after the 3 minutes, place the samples on ice for 2 minutes.
While these samples are on ice, add the enzymes to your first strand cDNA master mix from above at the following volumes (10% overage has already been included):
Kit reagent | Volume per rxn (uL) |
|---|---|
SeqAmp DNA Polymerase | 0.33 |
SMARTScribe Reverse Transcriptase | 2.2 |
Immediately after the samples are on ice for 2 minutes, pipette 12.5 uL of the First Strand cDNA Master Mix with enzymes into each sample.
Mix the contents by pipetting gently up and down 15 times.
Seal plate and centrifuge.
Determine the number of PCR cycles needed based on your original input amount of cells or RNA
Input of total RNA (or amount of cells) | Typical number of PCR cycles |
|---|---|
1 ng (or about 100 cells) | 14-15 |
100 pg (or about 10 cells) | 17-18 |
10 pg (or about 1 cells) | 18 |
We have found that the recommended 10-11 cycles is not quite enough to amplify 1 ng RNA. We usually do 14-15 cycles for 1 ng and 17 cycles for 100 pg.
Place samples in a heated-lid thermal cycler, preheated to 42°C and run the following program, "HT PLUS CDNA" (bolded steps are to be run for 10 to 18 cycles, depending on your input size, see the second table for the number of cycles.)
While the CDNA reaction is running, take the AMPure or homemade beads, elution buffer (Clontech kit), and the reagents for the Tapestation (High sensitivity D5000 reagents, ladder, and screentape) out of cold storage to ensure these have equilibrated to room temperature before use.
Step 2: cDNA clean-up
After the plate comes out of the thermal cycler, you have cDNA! Now we will go through a bead based clean-up.
Vortex AMPure/homemade beads for 2 mins to ensure the beads are mixed well just before use.
Add 25 uL of AMPure XP or homemade beads directly to PCR products. Mix well by pipetting up and down 20 times.
Incubate the bead-cDNA mixture at room temperature for 8 minutes to allow the cDNA to bind to the beads.
During the 8 minute incubation, prepare a master mix of 80% ethanol by mixing 100% ethanol with sterile water, you will need 400 uL per sample.
Place the samples on the magnetic seperation device for about 6 minutes, until the liquid appears completely clear and there are no beads left in the supernatant.
With the plate still on the magnetic stand, remove the supernatant from your samples without disrupting the beads; you should expect around 45 uL of supernatant. If you accidentally pull up some beads with your supernatant, put it back into the well with the bead and mix thoroughly to resuspend the bead, then wait until the liquid is clear again to remove the supernatant.
The beads contain your cDNA so any beads expelled will cause you to lose part of your samples.
Keep the plate on the magnetic stand and add 180 uL of your 80% ethanol mix to each well. Incubate for 30 seconds, then remove ethanol from well without disrupting the bead and expel.
Repeat the wash step above another time.
Go back into the wells with a P20 to remove any excess ethanol.
Incubate samples at room temperature for approximately 2 minutes until pellet is matte. The pellet should not be shiny, which is an indication there is ethanol still in your sample that will reduce your recovery rate of cDNA. Try to avoid over-drying: if the pellet is cracked it will take a significantly longer time to rehydrate these beads.
Once the beads are dry, remove them from the magnetic stand and add 17 uL of Elution Buffer (from the Takara kit in the -20°C) to your samples and pipette up and down until the beads are completely rehydrated. You will have to pipette up and down many times until the samples has no visible bead clumps in them.
Incubate at room temperature for 2 minutes.
Place stand back on the magnetic stand for around 2 minutes until the solution is completely clear.
Your cDNA is in now eluted from the beads, so do not discard supernatant
Transfer 15uL of supernatant into a new 96 well plate. This is your cDNA.
Run the samples through the Agilent TapeStation High Sensitivity D5000 to check that cDNA was made and amplified. Your profile should resemble the one shown below
Check the concentration of your samples using Qubit.
This is a safe stopping point. You can seal the plate and store at -20°C until Day 2.
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.
Dilute your sample with Elution Buffer (as used for elution in the previous clean-up).
- 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.
Step 3: Adaptors
Name | Location | Action |
|---|---|---|
Stem-loop adaptors | -20°C freezer, purple cap | Thaw on ice |
FE Dilution Buffer | -20°C freezer, white cap | Thaw on ice |
Lib Prep Buffer | -20°C freezer, blue cap | Thaw on ice |
Rxn Enhancer | -20°C freezer, red cap | Thaw on ice |
Amp Buffer | -20°C freezer, orange cap | Thaw on ice |
10X FE | -20°C freezer, white cap | Store until needed |
Lib Prep Enzyme | -20°C freezer, blue cap | Store until needed |
PrimeSTAR DNA Polymerase | -20°C freezer, green cap | Store until needed |
Nuclease-free water | -20°C freezer | Thaw at room temperature |
Unique Dual Indexes | -20°C freezer | Thaw at room temperature |
Allow Ampure XP or homemade beads and HSD1000 Tapestation reagents to equilibrate to room temp for at least 30 minutes before use.
On ice, add 4 µl of Stem-Loop Adapters to each well of a 96-well plate, according to the number of reactions to be performed.
Add a total of 8 µl of freshly diluted cDNA sample to a well containing the 4 µl of Stem-Loop Adapters. If the volume is less than 8µL, dilute with Elution Buffer. All samples must be at the same value of total input within 1- 10 ng (normalized).
On ice, prepare 1X FE by diluting the 10X FE in cold FE Dilution Buffer in a 1:9 ratio (1 part 10X FE to 9 parts FE Dilution Buffer). To allow for greater accuracy pipetting the 10X FE, a minimum of 40 µl of 1X FE Preparation should be prepared, which is enough for 36 rxns. If prepping greater than 36 samples, prepare enough for the number of reactions with 10% overage. Do not save excess 1X FE.
Mix 1X FE gently by pipetting up and down 20-30 times. Spin down and keep on ice.
On ice, prepare the Library Prep Master Mix in the table below, per sample (10% overage has already been included):
Name | Volume |
|---|---|
Library Prep Buffer | 4.4 |
Rxn Enhancer | 3.85 |
Library Prep Enzyme | 2.2 |
1X FE | 1.1 |
Vortex the Mix for 5 sec, then spin down briefly to collect the contents at the bottom of the tubes/plate. If necessary,
vortex for an additional 5 sec and spin down again. Keep on ice.
Discard leftover 1X FE. Do not reuse.
On ice, add 10.5 µl of the Library Prep Master Mix to each well containing stem-loop adators and cDNA and pipette to mix a few times.
Seal plate and mix by vortexing for 5 sec, and then spin down briefly.
Place on the thermal cycler for the "HT PLUS LIB" program (40min at 20°C, 10min at 85°C, 4°C Hold).
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
On ice, prepare the Library Amplification Master Mix, per sample (10% overage is already included):
Name | Volume |
|---|---|
Amplification Buffer | 23.65 ul |
PrimeSTAR HS DNA Polymerase | 1.1 ul |
Vortex and spin down the Amplification Master Mix, then keep on ice while working.
Add 22.5 ul Library Amplification Master Mix to each reaction.
Spin down the index plate for 1 min at 300g and add 5 ul of a different index to each well.
Indexes come in a 96 well plate, and each well contains 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.
Pipette to mix 15 times with a p200 at 45 µL.
Seal the plate, vortex, and centrifuge.
Place on the thermal cycler for the "HT PLUS AMP" program. Use the table below to determine the number of PCR cycles. If using 1ng of cDNA input, we recommend 15 PCR cycles to avoid overamplification.
Step 3: Library Cleanup
This protocol has an option to pool libraries before performing the cleanup. Only pool libraries if you are prepping a very large number or you are very certain your libraries will be uniform. Refer to the SMART-Seq HT Plus User Manual for further information.
Make sure the Ampure XP or homemade beads have been at room temp for at least 30 minutes. Vortex thoroughly before use.
Prepare fresh 80% ethanol. You will need about 400ul per sample.
Add 40ul of Ampure XP or homemade beads to each sample. Pipette to mix 15 times.
Incubate at room temp for 5 minutes.
Place on the magnetic stand for about 2 min, or when liquid is clear.
Remove and discard 80 uL supernatant without disturbing the beads.
Add 190 ul fresh 80% ethanol to each sample.
Wait 30 seconds, then remove and discard the ethanol.
Repeat the ethanol wash.
Briefly centrifuge the samples and place back on the magnet. Wait 30 sec, then remove any residual ethanol with a p20 pipette.
Keep the samples on the magnet until the bead pellets are dry. They will appear matte and no longer shiny. This will usually take 2 min, but if you pooled the sample for cleanup and are using larger volumes it may take longer. If cracks appear in the bead pellets, proceed immediately to the next steps. Over or under drying the pellet will affect library yield.
Remove the samples from the magnet and resuspend in 25 ul nuclease free water. Pipette 15 times to resuspend OR seal the plate and vortex.
Incubate at room temp for 5 minutes.
Place back on the magnet for 2 min or until the liquid is clear.
Transfer 23 ul of supernatant to a new plate. These are your purified libraries. Store at -20°C.
Quality Checks
Perform quality control steps :
Tapestation to determine the library size
Qubit to quantify the libraries
Libraries can be stored at -20°C for up to 30 days.
Sequencing Guidelines
Normalize and Pool
Quantify each of your libraries on Qubit. For most libraries, using the HS dsDNA Qubit assay with 2 uL of input will yield a reading. Record the concentration in ng/uL for each library.
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.
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 2 nM in a LoBind microcentrifuge tube. If you need to dilute your libraries, we recommend using at least 2 uL to minimize pipetting errors. The example sheet of the calculator provides further detail.
Quantify your pool on Tapestation and Qubit and enter into the calculator sheet to check that your pool is close to the nM concentration you normalized to.