Description
Library quantification kit for Illumina
GenieQuant Library Quantification Kit for Illumina 2.0 is an upgraded qPCR kit for accurate absolute quantification of Illumina sequencing libraries. It measures only amplifiable, adapter-ligated molecules, helping you optimise loading and cluster density. Version 2.0 chemistry improves robustness and reproducibility across library types.
The kit uses dye-based (SYBR Green I) qPCR. A standard curve is generated from six DNA standards, and the concentration of each diluted library is calculated from it by absolute quantification. The GenieQuant SYBR qPCR Master Mix 2.0 uses an antibody-mediated hot-start DNA polymerase and premixes amplification reagents, primers and universal ROX in one tube, giving strong specificity, high amplification efficiency, good GC compatibility, high sensitivity and rapid amplification, which suits high-throughput quantification. Any library at >0.0002 pM carrying sequences complementary to the P5/P7-type primers can be quantified, whatever the library type, construction method or Illumina instrument, with average fragment lengths up to 1 kb. The kit can also detect library contamination in work areas used for Illumina library preparation. For research use only.
Key Features
| Universal ROX | Works on all qPCR platforms without adjusting ROX concentration. | |
| Broad library compatibility | Quantifies any Illumina library with compatible adapter sequences, up to 1 kb average fragment length. | |
| All-in-one premix | Amplification reagents, primers and ROX are combined in a single tube. | |
| Wide dynamic range | Quantifies libraries from 20 pM down to 0.0002 pM. | |
| Lot-to-lot consistency | All reagents undergo rigorous quality control and functional testing. |
Specifications
| Method | Dye-based (SYBR Green I) qPCR, absolute quantification against a 6-point DNA standard curve |
| Platform | Illumina sequencing libraries |
| Quantifiable range | 20–0.0002 pM (5.5–0.000055 pg/μl; 12 × 106 - 12 × 101 copies/μl) |
| Fragment length | Average fragment length up to 1 kb |
| DNA standard size | 452 bp |
| Reaction volume | 20 μl (10 μl possible) |
| Primers | Primer 1: 5'-AATGATACGGCGACCACCGA-3'; Primer 2: 5'-CAAGCAGAAGACGGCATACGA-3' |
| Shelf life | 2 years |
Components
| Components | 500 rxns |
| GenieQuant SYBR qPCR Master Mix 2.0 | 4 × 1.25 ml |
Protect from light after opening. Stable through 30 freeze-thaw cycles, or for up to 2 months at 2 ~ 8°C (protected from light) for short-term repeated use. Also required: GenieQuant Library Quantification Kit for Illumina DNA Standard 1–6 and GenieQuant Library Dilution Buffer.
Storage
Store at -30 ~ -15°C.
- Prepare dilution buffer
- Use GenieQuant Library Dilution Buffer, or a self-prepared diluent (10 mM Tris-HCl, pH 8.0 at 25°C, 0.05% Tween 20). Never dilute libraries in water.
- Dilute the libraries
- Depending on the expected concentration, 1:1,000–1:100,000 dilutions may be appropriate. Include at least one additional 2-fold dilution of each library.
- Prepare dilutions fresh for each assay, keep them on ice and discard after use.
- Thaw reagents
- Thaw the master mix and DNA Standards 1–6, mix by inverting, centrifuge briefly and keep on ice.
- Set up reactions
| Components | Volume |
| GenieQuant SYBR qPCR Master Mix 2.0 | 10 μl |
| DNA Standard 1–6, diluted library or ddH2O (NTC) | 4 μl |
| ddH2O | To 20 μl |
- Dispense DNA Standards from lowest to highest concentration (Standard 6 to Standard 1), using a fresh tip for each.
- For 10 μl reactions, keep standards and libraries at 4 μl and use 5 μl of master mix, with consumables compatible with your instrument.
- Run qPCR
| Stage | Step | Reps | Temperature | Time |
| Stage 1 | Pre-denaturation | 1 | 95°C | 30 sec |
| Stage 2 | Cycles | 35 | 95°C 60°C | 10 sec 30 sec |
| Stage 3 | Melt curve* | 1 | 95°C 60°C 95°C | 15 sec 60 sec 15 sec |
*Melt curve collection varies between instruments; use the default settings.
- Analyse the data
- Set the baseline manually to cycles 1–3 (Standard 1 typically has a Ct of 7–9).
- Replicates should differ by ≤0.2 cycles; discard an outlier replicate before averaging.
- Define the effective Ct range from the NTC: if Ct (NTC) > Ct (Standard 6) + 3, use Standards 1–6; if it lies between Ct (Standard 5) + 3 and Ct (Standard 6) + 3, use Standards 1–5; if between Ct (Standard 4) + 3 and Ct (Standard 5) + 3, use Standards 1–4. Use at least four standards. If Ct (NTC) < Ct (Standard 4) + 3, the assay is seriously contaminated: replace all components and repeat.
- Plot Ct (y-axis) against Log[pM] (x-axis). R2 must be ≥0.99 and the slope between -3.1 and -3.6 (90%–110% efficiency); otherwise repeat the experiment.
- Use only library dilutions whose Ct falls within the effective range.
- Size-adjusted concentration (pM) = [452 bp / average library fragment length (bp)] × concentration of diluted library (pM).
- Original library concentration (nM) = average size-adjusted concentration (pM) × dilution factor / 1,000.
| DNA standard | Molar concentration | Mass concentration | Log[pM] |
| DNA Standard 1 | 20 pM | 5.5 pg/μl | Log[20] |
| DNA Standard 2 | 2 pM | 0.55 pg/μl | Log[2] |
| DNA Standard 3 | 0.2 pM | 0.055 pg/μl | Log[0.2] |
| DNA Standard 4 | 0.02 pM | 0.0055 pg/μl | Log[0.02] |
| DNA Standard 5 | 0.002 pM | 0.00055 pg/μl | Log[0.002] |
| DNA Standard 6 | 0.0002 pM | 0.000055 pg/μl | Log[0.0002] |
These are not final reaction concentrations. Keep standard and library volumes identical; no conversion is needed.
Q1: What if amplification efficiency is outside 90%–110%?
- If Ct (NTC) - Ct (Standard 6) <3 or Ct (Standard 6) - Ct (Standard 5) <3.1, efficiency above 100% may indicate contamination: check melt curves to identify whether the standards or library DNA are contaminated, set the effective Ct range from the NTC and discard contaminated Ct values. The default baseline may also delay the Ct of Standard 1; set the baseline manually to cycles 1–3. Poor pipetting accuracy is another cause.
Q2: What if R2 is below 0.99 or replicates are not reproducible?
- This usually reflects poor pipetting accuracy. Make sure all reagents are thoroughly mixed before use.
Q3: Why are my standards not spaced correctly?
- Ct (Standard 6) - Ct (Standard 5) <3.1 indicates contamination; check melt curves. Ct (Standard 2) - Ct (Standard 1) <3.1 suggests a background subtraction problem; set the baseline to cycles 1–3. A ΔCt >3.6 indicates poor efficiency: mix reagents thoroughly, confirm component concentrations and cycling protocol, and protect the master mix from strong light, which reduces fluorescence and delays Ct values.
Q4: Why do different library dilutions give inconsistent results?
- If ΔCt between 2-fold dilutions is outside 0.9–1.1, or concentrations from different dilutions differ by more than 10%, the causes may be poor pipetting accuracy, insufficient mixing, a library that is hard to amplify (extremely GC- or AT-rich, or average fragment length >1 kb) or a degraded library. Prepare fresh dilutions for each assay, keep them on ice and discard after use.
Q5: What if library dilutions fall outside the standard curve?
- If Ct (library) < Ct (Standard 1), the library is not diluted enough (common with over-amplified libraries): increase the dilution factor and repeat. If Ct (library) > Ct (Standard 6), the library is over-diluted or library preparation failed; a typical library diluted about 1/10,000 should not exceed the Ct of Standard 6. Decrease the dilution factor and repeat.
Q6: Why do the standards amplify but my library does not, or only very late?
- The library may lack adapter sequences that the quantification primers bind to; the initial dilution may be grossly wrong (decrease the dilution factor and repeat); or the library may be degraded (prepare fresh dilutions for each assay).
Q7: Is a double peak in the standards' melt curve a problem?
- No. Different regions of the 452 bp standard melt at different temperatures, giving a characteristic double peak. Raised tails on Standards 1–3 are also normal because their high concentration prevents complete melting at the Tm.
