Best Practices

Best Practices

Avoid Cross-Contamination

  • Open only one adapter tube at a time.
  • When using a kit that contains a DNA Adapter Plate (DAP), clean the bottom of the 96-well PCR plate or eight-tube strip used to pierce the foil seal of a DAP with a sterile 70% Ethanol wipe.
  • Pipette carefully to avoid spillage.
  • Clean pipettes and change gloves between handling different adapter stocks.
  • Clean work surfaces thoroughly before and after the procedure.

Avoid Potential DNA Contaminants

  • Incorrect DNA quantitation might result from DNA contamination caused by interference from superfluous nucleic acids in a sample (e.g., RNA, small nucleic acid fragments, nucleotides, single-stranded DNA), excess proteins, or other contaminating materials.
  • DNA quality might also affect the quantity of usable DNA in a sample. If the DNA is damaged (e.g., heavily nicked or containing extensive apurinic/apyrimidinic sites), then many of these fragments might fail during library preparation.
  • High molecular weight dsDNA derived from host genomes can also interfere with accurate quantitation. Bacterial artificial chromosomes (BACs) and other bacterially-derived plasmids usually contain a small percentage of the chromosomal DNA from the host cells, despite the best purification efforts. These sequences may ultimately give rise to unwanted clusters on a flow cell lane. However, this contamination can be accurately quantified by analyzing aligned reads generated during sequencing against known bacterial sequences and subtracting these out. High molecular weight contamination may also be estimated prior to library preparation using qPCR assays designed to target unique chromosomal markers.

Temperature Considerations

  • Keep libraries at temperatures ≤ 37°C, except where specifically noted.
  • Place reagents on ice after thawing at room temperature.
  • Avoid elevated temperatures, particularly in the steps preceding the adapter ligation, except where specifically noted.
  • When processing more than 48 samples manually, Illumina recommends processing the plate on a bed of ice whenever possible, especially during the enzymatic steps (when using the End Repair Mix, A-Tailing Mix, and Ligation Mix). A large number of samples processed at room temperature may result in uneven catalytic activity, which can lead to reduced quality of the end product.
  • DNA fragments that have a high AT content are more likely to denature into single strands than GC-rich fragments, which can result in an increased probability of creating a bias in the sequencing coverage.
  • Take care not to denature the library prior to the agarose gel electrophoresis process, because single-stranded DNA has a different migration rate.

Handling Liquids

  • Small differences in volumes (±0.5 µl) can sometimes give rise to very large differences in cluster numbers (~100,000).
  • Small volume pipetting can be a source of potential error in protocols that require generation of standard curves, such as PicoGreen assays or qPCR, or those that require small but precise volumes, such as the Agilent Bioanalyzer.
  • If small volumes are unavoidable, then due diligence should be taken to ensure that pipettes are correctly calibrated.
  • Make sure that pipettes are not used at the volume extremes of their performance specifications.
  • Take care with solutions of high molecular weight double-stranded DNA (dsDNA). These can be viscous and not evenly dispersed, resulting in aliquot measurements that are not representative of the true concentration of the solution.
  • Prepare reagents for multiple samples simultaneously to minimize pipetting errors, especially with small volume enzyme additions. Pipette once from the reagent tubes with a larger volume, rather than many times with 1 μl volumes. This method allows you to aliquot in a single pipetting movement to individual samples and standardize across multiple samples.

Handling Master Mix Reagents

  • Minimize freeze-thaw cycles. If you do not intend to consume the reagents in one use, dispense the reagent into aliquots after the initial thaw and refreeze the aliquots in order to avoid excessive freeze-thaw cycles. However, if you aliquot, you may not have enough reagents for the full number of reactions over multiple uses.
  • Add reagents in the order indicated and avoid making master-mixes containing the in-line controls.
  • Take care while adding ATL (A-Tailing Mix) and LIG (Ligation Mix) due to the viscosity of the reagents.

Handling Magnetic Beads

Follow appropriate handling methods when working with SPB or AMPure XP Beads:

  • Prior to use, allow the beads to come to room temperature.
  • Do not reuse beads. Always add fresh beads when performing the procedures.
  • Immediately prior to use, vortex the beads until they are well dispersed. The color of the liquid should appear homogeneous.
  • When performing a low sample protocol:
    • After adding the beads to the reaction, mix the solution gently and thoroughly by pipetting up and down 10 times, making sure the liquid comes in contact with the beads and that the beads are resuspended homogeneously.
    • Pipetting with the tips at the bottom of the well and not pipetting the entire volume of the solution helps prevent the solution from foaming. Excessive foaming leads to sample loss, because the foam is not transferred out of the plate efficiently.
  • When performing a high sample protocol, after adding the beads to the reaction, seal the plate and shake the plate on a microplate shaker at 1,800 rpm for 2 minutes. Repeat, if necessary, until the color of the mixture appears homogeneous after mixing.
  • Take care to minimize bead loss which can impact final yields.
  • Change the tips for each sample.
  • Let the mixed samples incubate at room temperature for the full duration specified in the protocol to ensure maximum recovery.
  • When aspirating the cleared solution from the reaction plate and wash step, it is important to keep the plate on the magnetic stand and to not disturb the separated magnetic beads. Aspirate slowly to prevent the beads from sliding down the sides of the wells and into the pipette tips.
  • To prevent the carryover of beads after elution, approximately 2.5 μl of supernatant are left when the eluates are removed from the bead pellet.
  • Always prepare fresh 70% and/or 80% ethanol, as required in the protocol. Ethanol tends to absorb water from the air, therefore, fresh 70% and/or 80% ethanol should be prepared for optimal results.
  • Be sure to remove all of the ethanol from the bottom of the wells, as it may contain residual contaminants.
  • Keep the reaction plate on the magnetic stand and let it air-dry at room temperature to prevent potential bead loss due to electrostatic forces. Allow for the complete evaporation of residual ethanol, as the presence of ethanol will impact the performance of the subsequent reactions. Illumina recommends at least 8 minutes drying time for the SPB beads and 15 minutes drying time for AMPure XP Beads, but a longer drying time may be required. Remaining ethanol can be removed with a 10 μl pipette.
  • Use the Resuspension Buffer (RSB) for DNA elution.
  • Avoid over drying the beads, which can impact final yields.
  • When performing a low sample protocol, resuspend the dried pellets using a single channel or multichannel pipette.
  • When performing a high sample protocol, resuspend the dried pellets by shaking.
  • When removing and discarding supernatant from the wells, use a single channel or multichannel pipette and take care not to disturb the beads.
  • To maximize sample recovery during elution, incubate the DNA/bead mix for 2 minutes at room temperature before placing the samples onto the magnet.