In the last tutorial, we mainly talked about the steps involved in the library preparation, let's move on to the next: library quantification.
Library Quantification
It is crucial to measure library concentration so that dilutions and calculations can be made accurately. Two main approaches for this are utilised – qPCR and fluorometric quantification methods such as Qubit.
qPCR measures the library concentration using primers that anneal to the P5 and P7 regions of the adapters. This means it can also indicate if the structure of the library is correct, as no amplification during the process could suggest that these regions don' t exist within your library. Use triplicates for each sample and at least two separate dilutions. A positive control, such as a previously sequenced library, is highly recommended. Novogene' s minimum requirement for library concentration is 2nM.
Fluorometric quantification methods use fluorescence-based dyes that selectively bind to either double-stranded DNA, single-stranded DNA or RNA. They are a more accurate method for the quantification of libraries with a broad range of fragment sizes, however they can overestimate the concentration as it measures all dsDNA within the pool – if any dimers or other unwanted dsDNA is within the system it will be picked up and quantified. It is also recommended to use a positive control with these methods.
Library Quality Control
Bioanalyzer instruments produce a trace that indicates the fragment size distribution of the library. They are best used for quality control of libraries with a narrow fragment size distribution as their accuracy decreases as size range increases. This method can show if the library fragments are too small or too large, if beads have carried over from the size selection step, or if PCR artefacts are present in the mixture. It also shows if the sample contains fragments at all, as samples can sometimes gets lost throughout the process, for example, during bead clean-up.
The trace will also show characteristic peaks for adapter and primer dimers if they are present. Adapter dimers contain the full adapter sequence that will be able to bind to the flow cell and generate clusters and sequencing data. They are caused by insufficient starting material at the beginning of library preparation, poor quality starting material or inefficient bead clean-up. Primer dimers don' t contain the adapter sequence so can' t bind to the flow cell and won' t be sequenced. Therefore, adapter dimers cause more issues. They use up sequencing capacity resulting in lower coverage of actual fragments and can negatively impact the sequencing data, sometimes causing a run to stop prematurely. They must be removed with additional clean-up steps with beads or gel purification.
Library normalisation and pooling
Once the libraries have been quantified and quality controlled, the concentration needs to be normalised before they can be pooled. The standard C1V1 = C2V2 calculation can be used to work out the dilution factor that will result in uniform concentration across the libraries that are to be pooled. Then add equal volumes of the libraries you wish to pool to a microcentrifuge tube and pipette 10 times to mix thoroughly.
Sequencing complications
Index hopping occurs when a high volume of samples are multiplexed together, causing the incorrect assignment of a library to a different index, aligning the fragment with the incorrect sequencing template.
How to avoid:
• Remove free adapters during prep. • Store libraries at -20°C • Only pool libraries right before sequencing • Use unique dual indexing strategy – allows for better identification.
Index dropout is a rare phenomenon which appears as an unusually small number of reads assigned to a sample. It is not always clear what causes it but some possibilities are:
• Contamination • Insufficient starting DNA concentration • Interference between adapter sequences and index sequences or read sequences. • Insufficient nucleotide diversity
An additional sequencing run is usually required in this case, once the library has been cleaned up of concentrated.
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