NGS Library Molarity & Equimolar Pooling Calculator (Qubit ng/µL to nM)

Core Formula

Library molar concentration is determined jointly by the Qubit mass concentration and the average fragment length:

c (nM) = c_mass (ng/µL) × 10^6 / (660 × L_bp)

• c_mass: mass concentration from Qubit fluorometric quantification (dsDNA HS kit), in ng/µL
• L_bp: average library fragment length from Bioanalyzer or TapeStation, in bp, including adapters
• 660: average molecular weight of dsDNA per base pair, in g/mol

Derivation: dsDNA molecular weight = 660 × L g/mol; converting ng/µL to g/L requires multiplying by 10⁻³; dividing by molecular weight gives mol/L, then multiplying by 10⁹ converts to nM, which simplifies to the formula above.

Worked example (Lib-A, 5.20 ng/µL, 385 bp): 5.20 × 10⁶ / (660 × 385) = 5,200,000 / 254,100 ≈ 20.5 nM.

Why Do You Need Both Mass Concentration and Fragment Length?

Molarity measures the number of molecules, not mass. Two libraries both at 5 ng/µL — the 200 bp library contains roughly twice as many molecules as the 400 bp library. Pooling by equal mass gives the shorter library twice the template molecules, resulting in a two-fold difference in sequencing depth — the most common cause of failed equimolar pooling.

On “Average Fragment Length”

You must use the complete library length including adapters, i.e., the average peak from the Bioanalyzer electropherogram, not the insert size. Illumina standard TruSeq adapters add approximately 120 bp to the insert (Y-adapter total on both ends): a 250 bp insert library has a complete fragment of approximately 370 bp. Enter the number reported by the instrument directly — no manual addition or subtraction needed.

Equimolar Pooling Protocol

The standard workflow has two steps:

1. Dilute all libraries to the same working concentration W (typically 10 nM) using TE buffer or 10 mM Tris-HCl (pH 8.0):

V_TE = V_stock × (c / W − 1)

2. Take equal volumes from each diluted library and combine — this gives the equimolar pool.

If a library’s stock concentration is already below the target working concentration and cannot be diluted to it, lower the target working concentration until all libraries can reach the same level.

Illumina Loading Concentration Reference

After pooling, further dilute to the loading concentration (after alkaline denaturation to single-stranded DNA):

Follow the Denature and Dilute Guide for each instrument; the table above is for reference only.

Common Mistakes

Using NanoDrop instead of Qubit: NanoDrop relies on A260 absorbance and cannot distinguish target dsDNA from free nucleotides or RNA carry-over, typically overestimating by 10–50%, leading to lower-than-intended loading concentration and insufficient cluster density.

Entering insert size instead of library fragment length: Systematically underestimates library concentration by ~30%, causing shorter-fragment libraries to be over-represented when pooling by molar ratio.

Applicability

• For dsDNA libraries only (Illumina TruSeq, TruSeq Nano, Nextera, and other dsDNA library prep products)
• 660 g/mol/bp is an approximation averaged across all four bases; for libraries with extreme GC bias (< 30% or > 70% GC) the error is approximately 1–2%, negligible for pooling decisions
• When the fragment length distribution is broad (e.g., ATAC-seq with multiple peaks), use the mode of the main peak rather than the arithmetic mean for more accurate results
• Not applicable to ssDNA libraries (small RNA sequencing) or direct RNA quantification