Both kit versions yield sequences close to the 3’ end of transcripts. The difference is in the location of the Read 1 linker sequence. If it is located in the 5´ part of the second strand synthesis primer (QuantSeq Forward (FWD), Cat. No. 015), NGS reads will be generated towards the poly(A) tail (Figure 1). This version is recommended for gene expression analysis.

Figure 1 | Read orientation for QuantSeq FWD (Cat. No. 015).

With QuantSeq Reverse (REV, Cat. No. 016) the Read 1 linker sequence is located on the 5’ end of the oligodT primer and a Custom Sequencing Primer (CSP, included in the kit) is required for sequencing in order to start the read directly at the 3´ end (Figure 2). Based on this the exact 3’ UTR can be pinpointed.

Figure 2 | Read orientation for QuantSeq REV with CSP (Cat. No. 016).

The upgraded protocol contains a more streamlined protocol with a shortened RNA removal step. Subsequent second strand synthesis and purification steps are also adjusted. Barcodes in the i7 Index Plate have been renamed (7001-7096, formerly BC01-96) and rearranged for a better nucleotide balance for sequencing when only few samples are multiplexed. With this new set-up all indices are now different from Illumina barcodes as former BC05 was replaced. With the QuantSeq FWD HT Kit (015.384) Lexogen furthermore provides a high-throughput version with optional dual indexing (i5 and i7 indices) allowing up to 384 barcode combinations (QuantSeq 3’ mRNA-Seq FWD HT User Guide).

For QuantSeq REV (Cat. No. 016) the Read 1 linker sequence is located at the 5’ end of the oligodT primer. Here a Custom Sequencing Primer (CSP Version 2, included in the kit) is required to achieve cluster calling on Illumina machines. It covers the poly(T) stretch and replaces the Multiplex Read 1 Sequencing primer.
PROVIDE THE RELEVANT INFORMATION TO YOUR SEQUENCING FACILITY ALONG WITH THE CSP!

HiSeq 2000, HiSeq 2500 (CSP Version 2 added on cBot)
CSP Version 2 should be provided in a tube strip at 0.5 µM final concentration in a volume of 120 µl (final concentration 0.5 µM, to be diluted in HT1 = Hybridization buffer). Take 0.6 µl of 100 µM CSP Version 2 and add 119.4 µl of HT1 buffer per sequencing lane. Place the 8-tube strip into the cBot position labeled primers.
HiSeq 2500 (CSP Version 2 replaces HP10 in cBot Cluster Generation Reagent Plate)
Alternatively, CSP Version 2 can be placed directly into the cBot Cluster Generation Reagent Plate. ATTENTION: The standard Illumina Multiplex Read 1 Sequencing Primer solution HP10 (for V4 chemistry located in row 2) provided in the cBot Cluster Generation Reagent Plate has to be REMOVED first! The Illumina V4 chemistry cBot Cluster Generation Reagent Plate only has 8 rows filled. A simple trick is to have the empty rows facing towards you, this way if you want to use a CSP in lane 1, you have to remove the HP10 solution from well 1 (first one on the far left) of the 2nd row, rinse the well a couple of times with HT1 and then add the diluted CSP Version 2. For this take 1.25 µl of 100 µM CSP Version 2 and add 248.75 µl of HT1 buffer per sequencing lane. The CSP should be at 0.5 µM final concentration in a volume of 250 µl (final concentration 0.5 µM, to be diluted in HT1 = Hybridization buffer). ATTENTION: Do not add the CSP to the Standard Illumina Multiplex Read 1 Sequencing Primer = HP10 solution! Always use fresh HT1 and add the CSP / HT1 dilution to the empty and rinsed well.

HiSeq 2500 – Rapid Run
Add 12.5 µl of 100 µM CSP Version 2 to 2487.5 µl HT1 = Hybridization buffer, resulting in a total volume of 2.5 ml and a final CSP concentration of 0.5 µM. In a rapid run, both lanes will use the same sequencing primer. It is not possible to run the two lanes with different sequencing primers.

MiSeq
Clustering is performed on the machine, not on the c-Bot. The MiSeq uses a reservoir of 600 µl with 0.5 µM sequencing primer final concentration, i.e., 3 µl of 100 µM CSP Version 2 in 597 µl HT1.

HiSeq 3000, HiSeq 4000 (CSP Version 2 replaces HP10 in cBot Cluster Generation Reagent Plate)
Usage of a custom sequencing primer is currently not supported on HiSeq 3000 and 4000 machines. A work around as described for the HiSeq2500 (CSP Version 2 REPLACES HP10 in the cBot Cluster Generation Reagent Plate) is possible though. ATTENTION: Do not add the CSP Version 2 to the HP10 solution! A primer mixture would result in low clusters calls and the resulting reads would be contaminated by poly(T) stretches. Always use fresh HT1 and add the CSP Version 2 / HT1 dilution to the empty and rinsed well.

The QuantSeq protocol is optimized for shorter reads (SR50, SR100) and yields mean library sizes of about 335 – 456 bp, with mean insert sizes of 203 – 324 bp. To generate longer libraries, use the QuantSeq-Flex First Strand Synthesis Module (Cat. No. 026), see QuantSeq 3’ mRNA-Seq User Guide, Appendix H, p.30.

The kit uses total RNA as input, hence no prior poly(A) enrichment or rRNA depletion is required. The amount of total RNA needed for QuantSeq depends on the poly(A) RNA content of the sample in question. This protocol was tested extensively with various cell cultures, animal and plant tissues, yeast, fungi, drosophila and human reference RNA. Typical inputs of 500 ng total RNA generate high quality libraries. For mRNA-rich tissues (such as kidney, liver, and brain) input material may be decreased to 50 ng without adjusting the protocol. However, for most efficient detection of low abundant transcripts RNA inputs from 500 ng to 200 ng are recommended.

*All libraries are prepared with external barcodes. Linker sequences are 122 bp including the 6 nt long external barcodes.

QuantSeq REV is only recommended for input amounts greater than 10 ng of total RNA. If you are limited in terms of input, we recommend using QuantSeq FWD. By choosing longer sequencing read lengths, you are also able to read into the 3’ ends.

For QuantSeq REV we recommend not to use input amounts lower than 10 ng total RNA. When using less than 10 ng of total RNA input please follow the recommendations indicated in the table below:

The number of cycles for your endpoint PCR depends on the type of the RNA (tissue, organism), the RNA quality, and the RNA input amount. The reference values given in Appendix C, p.21 are based on Universal Human Reference RNA input and the mRNA content of other RNA sources might differ. To be on the safe side and to prevent under- or overcycling of your sample, we recommend performing a qPCR first. Therefore, we offer a PCR Add-on Kit for Illumina (020.96) with 96 additional PCR reactions. Use 5 µl of P7 Primer (7000) instead of an i7 index in step 27 of the single indexed PCR protocol. Dilute the double-stranded library from step 24 to 19 µl by adding 2 µl of Elution Buffer (EB) in order to have enough template for qPCR and endpoint PCR. Add 1.7 µl of the cDNA into a PCR reaction containing 7 µl PCR Mix, 5 µl P7 Primer 7000, 1 µl Enzyme Mix E from the PCR Add-on Kit, and SYBR Green I (or an equivalent fluorophore, to be provided by the user) to a final concentration of 0.1x (diluted in DMSO). Make the total reaction volume up to 30 µl with EB. Conduct at least 35 cycles to make sure the amplification reaches the plateau. Afterwards take the fluorescence value where the plateau is reached and calculate where the fluorescence is at 50 % of the maximum (see Fig. 3). The value where the fluorescence reaches the maximum (plateau) is taken (15388) and the fluorescence at 50 % of this values (7694) shows which cycle number is optimal for the endpoint PCRs. For the sample in Fig. 3 this would be 15 cycles when using 1/10^th of your sample. If the optimal cycle number lies within two values, it is recommended to always round up to the higher number in order to get more yield. As in the endpoint PCR 10x more cDNA will be used compared to the qPCR, three cycles can be subtracted from the determined cycle number, hence in this example 12 cycles should be used for the endpoint PCR. This is the cycle number you should use for the endpoint PCR using the remaining 17 µl of the template.

Figure 3: Calculation of the number of cycles for the endpoint PCR

Once the number of cycles for the endpoint PCR is established for one type of sample, you can use it in the following experiments. For higher yields you can increase the fluorescence level of the endpoint PCR up to 80 % without overcycling your sample.

In the table below you can see some recommended cycle numbers for the endpoint PCR using 500 ng total RNA input of different RNA sources.

Yes, low quality and FFPE samples can be used with QuantSeq. Some minor protocol modifications are required though. These recommendations are indicated in the table below:

The quality of RNA from FFPE tissues can vary greatly. We recommend measuring the DV₂₀₀ value (the percentage of RNA greater than 200 nt in length) in addition to RIN values, as RIN values become less meaningful for highly degraded samples.

Libraries prepared from FFPE RNA input typically require different (often higher) PCR cycle numbers than those prepared with high quality RNA input (see table below for examples). The DV₂₀₀ value is also not always a reliable predictor of the required number of PCR cycles needed.

When preparing libraries for comparative gene expression profiling, all libraries that will eventually be compared should be amplified using the same number of PCR cycles. To prevent under- or overcycling the libraries, we therefore strongly recommend performing a qPCR assay to determine the optimal number of PCR cycles for the set of samples to be processed within each experiment.

Lexogen’s QuantSeq kit is a library preparation protocol designed to generate sequence-ready Illumina-compatible libraries from polyadenylated RNA within 4.5 hours. When carrying out the protocol for the first time, please allow for more time and read the entire User Guide first.

QuantSeq libraries are intended for a high degree of multiplexing. Barcodes are introduced as standard external barcodes during the PCR amplification step. With the up to 96 external i7 barcodes (i7 Index Plate, 7001-7096) included in the kit and the additionally available four external i5 indices (5001-5096, i5 Dual Indexing Add-on Kit, Cat. No. 047), up to 9,216 samples can be multiplexed and sequenced per lane on an Illumina flow cell.

ATTENTION: QuantSeq REV cannot be multiplexed with other library preps as the Custom Sequencing Primer is needed for sequencing.

With QuantSeq REV and the Custom Sequencing Primer it is possible to exactly pinpoint the 3’ end during Read 1. The reads generated during Read 1 reflect the cDNA sequence, and are therefore in a strand orientation opposite to the genomic reference.

For QuantSeq REV (Cat. No. 016) we do not recommend using TopHat2, since there is hardly any need to search for junctions. Nearly all sequences will originate from the last exon and the 3’untranslated region (UTR). In case of no detected junction, TopHat2 may run into difficulties. Hence, Bowtie2 or BWA can be used for mapping in this case.
More information on the data analysis can be found here.

While single read sequencing does not require any trimming using QuantSeq REV (Cat. No. 016), paired-end sequencing may require the first 12 nucleotides of Read 2 to be trimmed. Alternatively, a less stringent aligner STAR Aligner could be used with the number of allowed mismatches being set to 16 for paired-end reads.
In case of adapter contamination detection it is crucial to trim these sequences (e.g., cutadapt, trim-gallore, or bbduk) in order to align the reads

The QuantSeq 3’ mRNA-Seq REV kit is appropriate for HiSeq 2000/2500, HiSeq 3000, HiSeq 4000, GAIIX, and MiSeq. We do not recommend to sequence QuantSeq REV libraries on NextSeq 500 or NextSeq 550 Illumina platforms. The polyadenylation signal located at the 3’ end, which is quite similar between transcripts, will result in a reduced complexity at the beginning of the reads leading to a lower sequencing quality.

Single-read 50 (SR50) sequencing runs are suitable for both QuantSeq versions. Although, for QuantSeq REV it is favorable to use longer runs (SR100), since starting the read exactly at the 3`UTR will also show the polyadenylation signal, which covers up to 25 nt from the transcription end. As the polyadenylation signal is quite similar for all transcripts, it is better to have longer reads >SR50 available for mapping in order to increase the number of uniquely mapping reads.

If you want to determine the optimal number of cycles for your endpoint PCR using dual indexing, you can still use the PCR Add-on Kit according to the instructions (see FAQ 1.7). The single indexing PCR (i7 only) of the PCR Add-on Kit and the dual indexing PCR (i5 and i7) run with the same efficiency, hence there is no need to exchange any solutions.

For QuantSeq REV it is essential to use the CSP (provided with the QuantSeq kit) for Read 1 sequencing. Make sure that your sequencing facility is aware of this. When handing in QuantSeq REV libraries for sequencing, please include the CSP and the information on how to use the CSP. The CSP should never be mixed together with the standard Illumina Read 1 Sequencing primer. A primer mixture would result in low clusters calls and the resulting reads would be contaminated by poly(T) stretches.

Transcripts may have different and not yet annotated 3’ ends, which might be mistaken for internal priming events of the oligodT primer, when in fact those are true 3’ ends. Artificial spike-in transcripts such as the SIRVs or the ERCC spike-in transcripts only have one defined 3’ end, this provides the only true measure to determine internal priming. If true internal priming is detected, this could be a result of mis-priming during reverse transcription, for instance if the temperature before or during reverse transcription was too low. As outlined in the general section of the User Guide, unless explicitly mentioned, all steps should be carried out at room temperature between 20 °C and 25 °C. In particular, for First Strand cDNA Synthesis (p.11): Do not cool the FS2/E1 mastermix (step 3) and have the RNA/FS1 samples at 42 °C when adding the FS2/E1 mastermix (step 4) to avoid mishybridization. Mix properly by pipetting. Do not forget to shortly spin down the samples at room temperature before and after adding the FS2/E1 mastermix. Centrifugation should not be carried out at 4 °C, always spin down at room temperature! Raising the reaction temperature to 50 °C for reverse transcription can be used for additional prevention of mis-priming.

• First Strand cDNA Synthesis (p.11-12):
  1. At step 3 pre-warm the FS2 / E1 mastermix for 2 – 3 minutes at 42 °C while the RNA / FS1 samples are denaturing for 3 minutes at 85 °C – Do not cool the mastermix on ice!
  2. After the RNA / FS1 samples have cooled to 42 °C, spin these down briefly and then immediately return to the thermocycler and hold at 42 °C.
  3. Add the pre-warmed FS2 / E1 mastermix to the RNA / FS1 samples on the thermocycler at 42 °C (step 4) and mix properly. Any drop in temperature at this point can cause mishybridization! Seal the plate or tubes and begin the 42 °C incubation.

  NOTE! Spin down the samples at room temperature before and after adding the FS2 / E1 mastermix.

• If step is skipped (low input or degraded samples i.e. ≤10 ng, or FFPE samples):
  1. Prepare your RNA samples in 5 μl volumes.
  2. Prepare a mastermix containing 5 μl FS1, 9.5 μl FS2, and 0.5 μl E1, mix well, spin down, and pre-warm at 42 °C on a thermocycler for 2 – 3 minutes.
  3. Bring your RNA samples to room temperature while the mastermix is pre-warming.
  4. Spin down the pre-warmed FS1 / FS2 / E1 mastermix and add 15 μl to each RNA sample. Quickly mix, seal the plate or strip-tubes, spin down briefly at room temperature, and then commence the 42 °C incubation for 15 minutes (or 1 hour for low input RNA (≤ 10 ng)).

1.28
• Proceed immediately to the RNA removal after the reverse transcription is complete! (step 4-5). The previous safe stopping point after reverse transcription (step 4) has been removed. Do not place the samples on ice, and do not store samples at -20 °C at this point! Cooling the samples below room temperature at this point can cause mishybridisation! Best practice handling would be as follows:
  1. After the 42 °C incubation is complete spin down the plate/tubes briefly and place at room temperature.
  2. Immediately add the RNA Removal Solution (RS, thawed at room temperature) to the samples, mix well.
  3. Briefly spin down the plate / tubes at room temperature, then place on the thermocycler to commence the 10 minute incubation at 95 °C (step 6).

  NOTE! To minimise temperature drops at this point the reactions can also be kept at 42 °C while the RNA Removal Solution (RS) is added: Briefly spin down the samples after step 4 and place them back on the thermocycler at 42 °C, remove the sealing foil / tube caps, add the RNA Removal solution to the samples, mix, re-seal the plate / tubes, quickly spin down, and place back on the thermocycler block and re-start the program for the 95 °C incubation.

A second peak between 1000–9000 bp is an indication of overcycling. The library prep has been very efficient and a lot of cDNA was generated. Hence, the PCR ran out of primers and template started to denature and reanneal improperly. This results in longer, bulky molecules that migrate at a lower speed on the Bioanalyzer chip or gels. This can interfere with exact library quantification if relying solely on the Bioanalyzer results. Therefore, a qPCR assay for exact library quantification should be used additionally if such a high molecular weight peak occurs.
For future QuantSeq library preps on similar samples reduce your PCR cycle number accordingly to prevent overcycling. Overcycling may lead to a distortion in gene expression quantification and hence should be avoided.

A carryover of Purification Beads (PB) results in a peak around and beyond the upper marker of the Bioanalyzer. Make sure not to transfer any beads after the final elution in step 41 or 42 for single or dual indexing PCR respectively. Leave approximately 2 µl of the eluate on the beads and do not try to transfer the complete sample, as this will lead to bead carryover. Put your samples once again on the magnet and incubate for 5 minutes. Transfer 15-17 μl of the supernatant into a fresh PCR plate.

The PCR Add-on Kit for Illumina (Cat. No. 020) includes a Reamplification Primer that can be used to add some PCR cycles for your undercycled libraries (QuantSeq 3’ mRNA-Seq User Guide, Appendix F: Library Reamplification, p.27). In general, as QuantSeq is intended for a high degree of multiplexing, undercycled libraries can still be used for preparing a lane mix. The lane mix may need to be concentrated if many libraries of the lane mix were undercycled. Please note that currently only single indexed libraries can be reamplified. If you need to reamplify dual indexed libraries, please contact info@lexogen.com.

Universal Human Reference RNA (UHRR, Agilent) is a good positive control, most of the reference values given in the User Guide are also based on UHRR input.

For PE sequencing use QuantSeq REV (Cat. No. 016). We do not recommend paired-end sequencing for QuantSeq FWD (Cat. No. 015), as the quality of Read 2 would be very low due to the poly(T) stretch at the beginning of Read 2.

To generate longer libraries, use the QuantSeq-Flex First Strand Synthesis Module (Cat. No. 026). More information can be found in the QuantSeq 3’mRNA-Seq FWD User Guide, Appendix H: Modulating Insert Sizes, p.30.

The loading amounts indicated below are provided a guideline for sequencing QuantSeq FWD/REV libraries in pure pools for each instrument. Multiplexing QuantSeq with other external library types is not recommended (see FAQ 1.33). We therefore cannot provide optimal pooling or loading amount guidelines for lanemixes that include QuantSeq libraries in diverse multiplexing setups.
Loading amounts may still need to be adjusted to obtain optimal cluster densities for your respective instrument, or if sequencing chemistries are changed. Inaccuracies in library quantification can affect the loading amount required. Please ensure accurate quantification of lane mixes and lane mix dilutions when preparing for loading.
Unless otherwise indicated the values given in the table below are based on customer experiences. For further inquiries regarding loading amounts please contact info@lexogen.com.

^* PhiX cannot be sequenced in QuantSeq REV runs due to the use of a custom sequencing primer which does not bind to PhiX.
^** Please inquire at info@lexogen.com.
¹ Loading amounts are based on customer-supplied feedback for optimal sequencing quality on HiSeq 4000.
² The values for NextSeq 500 / 550 are recommended loading amounts and have been evaluated by Lexogen to ensure optimal cluster densities of 200 – 260 K/mm² are achieved. We have determined that loading amounts below 2 pM may result in lower cluster densities and reduced index read quality.
³ Loading amounts for NovaSeq Standard workflow are based on user experience using S2 flow cells, and should be relevant for all flow cell types. This range is further calculated according to Illumina’s recommendation to use 1.5x the loading concentrations for HiSeq 4000. The calculated loading amounts shown fall within the Illumina recommended range for PCR-amplified library pools (300 – 600 pM).
⁴ Loading amounts for NovaSeq Xp workflow are based on customer-supplied feedback from sequencing of SENSE mRNA V2 libraries. These concentrations also relate to customer-validated loading amounts used for HiSeq 4000 instruments.

In general, we recommend processing a minimum of 8 samples, using a complete set of eight i7 indices for multiplexing (e.g., 7001-7008). However, if fewer barcodes are required care should be taken to always use indices which give a well balanced signal in both lasers (red and green channels) for each nucleotide position. All columns (1-12) and rows (A-H) fulfill these criteria. The individual libraries within a lane should be mixed at an equimolar ratio to ensure this balance. An evaluation tool to check the color balance of index subsets is available under Support Tools.

Yes! All QuantSeq FWD and REV kits ordered after November 2016 are supplied with a code to access the QuantSeq data analysis pipelines on the Bluebee® Genomics Platform.

The code supplied with the QuantSeq REV Kits (Cat. No. 015) enables only the REV data analysis pipeline to be selected. Do not use Activation codes from QuantSeq FWD kits for analysis of QuantSeq REV data!

Each code contains an equal number of pipeline runs as reactions provided in the kits (e.g. You get 96 data analysis runs with a 96 prep QuantSeq REV Kit (Cat. No. 016.96), meaning you can analyze sequencing data for 96 .fastq files).
If you start the wrong pipeline for your data you can abort or stop the run before it is completed without losing your allocated runs. If the run is completed then additional runs will need to be purchased (Cat. No. 091).

QuantSeq FWD libraries prepared from blood using Globin Block (RS-GBHs or RS-GBSs, Cat. No. 070 and 071) should be analysed using the standard REV pipeline for data analysis.

Analysis pipelines for QuantSeq (FWD, FWD-UMI, and REV) are currently available for the following species:

New species can be added upon request. Please contact bioinfo@lexogen.com if your species of interest is not listed.

All QuantSeq FWD and REV kits ordered after November 2016 are supplied with an activation code to access the QuantSeq Data Analysis pipelines on the BlueBee® Genomics Platform.

The activation codes are printed on a sticker that is attached to the side of the small reagent box (stored at -20 °C) inside the main QuantSeq Kit box (see Figure 4). Activation codes for additional runs can be purchased from Lexogen via the webshop. Activation codes registered after September 20, 2018 are valid for two years. The input file size is limited to 1.5 GB per fastq(.gz) file. If you have larger input files or for further inquiries, please contact info@lexogen.com.

The count summary files from the results of the data analysis pipeline on BlueBee® contain mapped read counts for Ensembl gene IDs. Ensembl gene IDs can be converted into other gene ID types, or additional annotations added using the BioDBnet tools: https://biodbnet-abcc.ncifcrf.gov/db/db2db.php. Simply input the list of Ensembl gene IDs, select the input format as “Ensembl Gene ID”, and then select the output format(s) desired.

The UMI Second Strand Synthesis Module (Cat. No. 081.96) is only compatible with QuantSeq FWD Library Prep. This is because the UMI-containing random primers in this module contain the partial p5 adapter sequence, which is the same as the partial adapter included on the oligo(dT) primers used for QuantSeq REV Library Prep. Hence, using the UMI Module with QuantSeq REV will not result in an amplifiable library. If you are interested in including Unique Molecular Identifiers for QuantSeq REV Library Prep please contact us at info@lexogen.com.

Yes, the i5 Dual Indexing Add-on Kits can be used for dual indexing of QuantSeq REV libraries. Please refer to the i5 Dual Indexing Add-on Kits Instruction Manual (047IM109) for details of dual-indexed library amplification and purification.
For further information about the i5 Dual Indexing Add-on Kit for QuantSeq/SENSE please see the QuantSeq REV online FAQs 4.1 onwards.

We do not recommend multiplexing Lexogen libraries with libraries from other vendors in the same sequencing lane. Though this is possible in principle, specific optimization of index combinations, library pooling conditions, and loading amounts may be required, even for advanced users. Sequencing complex pools that include different library types at different lane shares may have unpredictable effects on sequencing run metrics, read quality, read outputs, and/or demultiplexing performance. Lexogen assumes no responsibility for the altered performance of Lexogen libraries sequenced in combination with external library types in the same lane (or run).

Due to size differences, libraries prepared with the Lexogen Small RNA-Seq Library Prep Kit (or any other small RNA library prep kit) should not be sequenced together with QuantSeq or SENSE libraries. Please refer to the sequencing guidelines for each library type (library adapter details, loading amounts to use, and use of custom sequencing primers, etc), which are provided in our Library Prep Kit User Guides, and online Frequently Asked Questions (FAQs).