TITLE: Library pooling and quality control for Illumina sequencing
AUTHORS: Katherine Smollett, Lily Tong, Jenna Nichols, Kirsty Kwok, Kyriaki Nomikou, Ma. Jowina Galarion, Daniel Mair, Ana Filipe

[DESCRIPTION]
The ability to multiplex many samples on the same run makes Illumina sequencing a powerful and affordable tool for many researchers. When pooling samples for sequencing it is important that they are pooled at equal molarity to prevent over or under representation of any individual sample in the same run. Care also has to be taken in deciding how many samples can be combined to achieve the required sequencing depth, a factor that is further complicated in viral sequencing where the majority of the reads may originate from the host and only a small proportion corresponding to viral reads.

Also of major importance is the final pool quality control (QC) to ensure that all contaminating adapter dimers are removed, generate accurate size distribution and quantification of the final pool. Accurate QC ensures accurate loading on the sequencer, overestimation of the pool molarity can cause under clustering resulting in fewer reads and can even cause run failure. Underestimation of pool molarity causes over clustering, lowering the read quality and risking run failure. 

Adapter dimers are small fragments containing full length adapter sequences which can bind to the flow cell and cluster. Due to their small size, they cluster more efficiently than the longer library fragments and so reduce the library-specific read depth as well as causing over clustering which reduces the data quality possible run failure. In addition free adapter can be incorporated into the library clusters resulting in index hopping and incorrect assignment of library barcodes.

Here we describe our standard workflow for determining the appropriate sequencing depth and pooling by equal molarity of Illumina sequencing libraries, along with our procedure for pool QC.

[BEFORE_START]
It is recommended that individual libraries are cleaned up and QC'd following the protocol Library clean up and quality control for Illumina sequencing. Each Illumina library should have an accurate size and quantification and be free from contaminants.  

[STEPS]
SECTION: Pooling
1. Before pooling the samples determine the number of samples per pool. The estimated reads per sample will depend on the required depth, genome length and expected percentage of on-target reads. The number of samples per pool will also depend on the type of Illumina sequencer/cartridge used and the read length. 

You can use the attached calculator to determine the samples/cartridge and how many pools are required for your experiment.

  

SECTION: Pooling
2. For each pool calculate the volume required so that each library is present in equal quantities. 

 

Library molar concentration can be calculated from the library fragment size and mass concentration as follows:

  

You can use the attached calculator to determine the pooling volumes.
 
  

You may be required to pool sub-pools instead of individual libraries. If so an additional weighting by the number of samples per sub-pool is required. See the attached calculator.

  

SECTION: Pooling
3. Using a fresh 1.5 mL DNA LoBind tube carefully add the required volume of each library.

 


4. Optional: Concentrate pool using 1.4X Ampure XP.

Equilibrate Ampure XP beads to Room temperature and vortex well to mix.
  


4.1. Measure the total volume of the pool and add 1.4X volume of Ampure XP beads and mix well.
4.2. Incubate at Room temperature room temperature for 5 min
4.3. Place on a magnetic rack for 5 min  until beads and solution have fully separated.

 

4.4. Carefully add 200 µL without disturbing the beads.
4.5. Incubate at Room temperature for  30 s.

4.6. Place on magnetic rack for 1 min  until the beads and solution have fully separated.

4.7. Keeping on the magnet and carefully remove supernatant without disturbing the beads.
4.8. Repeat wash with 200 µL.

4.9. Remove all traces of ethanol.

 

4.10. Keeping on the magnet air dry for up to 3 min.

 

4.11. Add the required volume of 10 mM Tris pH 8.0, remove from the magnet and mix well to fully suspend the beads.

4.12. Incubate at Room temperature for at least  2 min to elute the DNA.
4.13. Place back on magnet until beads and solution have fully separated.
4.14. Transfer the supernatant containing the pool DNA to a fresh 1.5 mL DNA LoBind tube. 

SECTION: Pool QC
5. It is recommended that library quantification is performed using a fluorometric method (e.g. Qubit) rather than an absorbance based method (e.g. Nanodrop) as it is more specific, sensitive and accurate.

 

SECTION: Pool QC
5.1. Prepare 0.5 mL thin-walled PCR tubes, including 2 tubes for standard solutions and 3 for each pool. 
Label the tube lids and not the sides.

 

SECTION: Pool QC
5.2. Prepare the Qubit dsDNA High Sensitivity master mix for the total number of samples and standards with an excess.
 
 

  
 

 


SECTION: Pool QC
5.3. Aliquot the Qubit dsDNA High Sensitivity master mix into the assay tubes as follows:

Standards 190 µL 
Samples 199 µL 

SECTION: Pool QC
5.4. Add 10 µL to each standard assay tube.
SECTION: Pool QC
5.5. Add 1 µL to each sample assay tube.
SECTION: Pool QC
5.6. Vortex assay tubes and briefly centrifuge.
SECTION: Pool QC
5.7. Incubate at Room temperature for 2 min.
SECTION: Pool QC
5.8. Select dsDNA high Sensitivity assay on the Qubit Fluorimeter and press "Read Standards".
SECTION: Pool QC
5.9. Insert Standard 1 and 2 into the sample chamber when prompted, close the lid and and press "Read Standard".
SECTION: Pool QC
5.10. Select "Run samples" and select the sample volume as 1 µL.
SECTION: Pool QC
5.11. Insert a sample tube into the sample chamber, close the lid and press "Read tube".
SECTION: Pool QC
5.12. Record the concentration of the Qubit sample in ng/μL.
SECTION: Pool QC
5.13. For each pool ensure all readings are within the same range.
 

SECTION: Pool QC
5.14. Calculate the average pool concentration from the replicates ensuring any outliers are removed.
SECTION: Pool QC
6. It is recommended that the libraries are visualised using capillary electrophoresis, we describe visualisation with a TapeStation and High Sensitivity D5000 ScreenTape. Alternatives such as the BioAnalyzer or Fragment Analyzer can also be used. 

The purpose is to provide a size for the library fragments to give accurate molar quantification and to determine the quality of the library.

 

SECTION: Pool QC
6.1. Ensure that the D5000 ScreenTape and Reagents are equilibrated to Room temperature at least 30 min before use, vortex and briefly centrifuge.
  
  


 

SECTION: Pool QC
6.2. Prepare a dilution of each pool to approximately 1 ng/μL.

SECTION: Pool QC
6.3. In fresh PCR strip tubes prepare the ladder assay tube as follows:
 
 

 
 
SECTION: Pool QC
6.4. Prepare the sample assay tubes as follows:

 
 
 

 
 
SECTION: Pool QC
6.5. Spin down, using IKA vortexer mix at 2000 rpm, 1 min then spin down again.


SECTION: Pool QC
6.6. Load the assay tubes and ScreenTape into the TapeStation instrument.
SECTION: Pool QC
6.7. Select the required sample/ladder positions in the TapeStation software and click "start".
 

SECTION: Pool QC
6.8. Analyse the results, the pool should generate a similar trace to the individual libraries. 

If needed follow the same troubleshooting guidelines as in the protocol Library clean up and quality control for Illumina sequencing.

SECTION: Pool QC
6.9. Determine the pool fragment peak size in bp.

 

SECTION: Pool QC
6.10. Calculate the pool molar concentration using the fragment size (Step 6.9) and mass concentration (Step 5.14).