Best Practices

Best Practices

Avoid Cross-Contamination

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

Handling RNA

RNA is highly susceptible to degradation by RNAse enzymes. RNAse enzymes are present in cells and tissues, and carried on hands, labware, and dust. They are very stable and difficult to inactivate. For these reasons, it is important to follow best laboratory practices while preparing and handling RNA samples:

  • When harvesting total RNA, use a method that quickly disrupts tissue and isolates and stabilizes RNA.
  • Wear gloves and use sterile technique at all times.
  • Reserve a set of pipettes for RNA work. Use sterile RNAse-free filter pipette tips to prevent cross-contamination.
  • Use disposable plasticware that is certified to be RNAse-free. Illumina recommends the use of non-sticky sterile RNAse-free microfuge tubes. A set of these tubes should be designated for this protocol and should not be used for other lab work.
  • All reagents should be prepared from RNAse-free components, including ultra pure water.
  • Store RNA samples by freezing. Avoid extended pauses in the protocol until the RNA is in the form of double-stranded DNA (dsDNA).
  • Use a RNAse/DNAse decontamination solution to decontaminate work surfaces and equipment prior to starting this protocol.

Temperature Considerations

  • Keep libraries at temperatures ≤ 37°C, except where specifically noted in a protocol.
  • Place reagents on ice after thawing at room temperature.
  • When processing more than 48 samples manually, Illumina recommends processing the plate on a bed of ice whenever possible, especially during enzymatic steps (when using the 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.
  • mRNA 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.
  • Temperature is less of an issue after the adapters have been ligated onto the ends of the ds cDNA.

Handling Liquids

Good liquid handling measures are essential, particularly when quantifying libraries or diluting concentrated libraries for making clusters.

  • 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.
  • When adapting this protocol for automation or robots, process >16 samples to minimize reagent loss due to dead volume.

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.
  • Some RNA kits contain First Strand Synthesis Mix Act D (FSA) which contains Actinomycin D, a toxin. Personal injury can occur through inhalation, ingestion, skin contact, and eye contact. Dispose of containers and any unused contents in accordance with the governmental safety standards for your region. Please refer to the product material safety data sheet (MSDS) for detailed environmental, health, and safety information.

Handling Magnetic Beads

Follow appropriate handling methods when working with AMPure XP and RNAClean 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 15 minutes drying time, but a longer drying time may be required. Remaining ethanol can be removed with a 20 μl pipette.
  • Use the Elution Buffer (ELB) for RNA 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.