Molecular-grade reagents and consumables are free of nucleases, reducing the risk of DNA degradation. 

When DNA is stored in regular tubes, some of the DNA sticks to the inner walls of these tubes, making in inaccessible. This can create confusion in PCR results, especially when dealing with diluted samples containing very low DNA concentrations. In contrast, low-bind siliconized tubes have smoother inner walls, so charged DNA molecules have less of a chance to attach to the tube’s inner surface.

Additionally, using tubes not certified as nuclease-free increases the risk of DNA degradation. This becomes, again, particularly problematic when working with low DNA concentrations.

Molecular-grade water is a reliable choice for general PCR applications. PCR-grade water is a premium product marketed explicitly as DNA-free. PCR-grade water can be particularly valuable for specialized applications that demand the utmost assay sensitivity. In such cases, ensuring no unwanted amplification is crucial when running non-template controls (NTCs). Ideally, NTCs should either not produce any amplification or, if they do, the amplification should consistently occur much later than the samples at the lower limit of quantitation (LLOQ).

Note: Even PCR-grade water can become contaminated with human DNA. We advise users to perform their own assessment of PCR-grade water products to ensure they meet their applications’ specific requirements.

The PCR chemistry plays a pivotal role in determining the results of analytical tests. Furthermore, differences in the temperature cycling patterns among various PCR instruments can influence the assay’s performance.

As a result, users can potentially prevent unexpected results in sensitive PCR-based assays by taking a series of steps. These steps include verifying and—if necessary—optimizing, qualifying, and ultimately validating published assays in their specific laboratory settings.

Yes. Users should conduct their own studies to determine the appropriate storage time and conditions. 

Even when using low-binding tubes, a certain amount of DNA can still adhere to the inner walls of these containers during storage. This occurrence can confound PCR results, particularly when working with highly diluted samples with very low DNA concentrations.

To reduce DNA binding to the tube walls, users can employ “filler nucleic acids,” such as double-stranded DNA or oligo- or polynucleotides, in the diluent. These filler nucleic acids minimize DNA adhesion without interfering with downstream applications that target the specific gDNA of interest. First, they prevent target DNA from sticking on the inner walls of the storage tubes, hence rendering target DNA accessible for downstream applications. Second, they play a crucial role in safeguarding nucleic acids from degradation by acting as a substrate for potentially contaminating nucleases, further preserving the integrity of the stored DNA. 

Our team has successfully utilized Poly(A) when testing the performance of genomic DNA reference material via PCR.

1. Prepare the work surface in a Biological Safety Cabinet (BSC) by decontaminating with 10% bleach (or Bleach-Rite^® [Current Technologies]) and then 70% alcohol.
2. Decontaminate pipettors with 10% bleach or Bleach-Rite^®. Spray 10% bleach onto a Kimwipe^® (Kimberly-Clark Corporation) and wipe down the pipettors to decontaminate. It’s important to use lint-free wipes to avoid any potential contamination.
3. Spray 70% alcohol onto a Kimwipe^® and wipe down the pipettors with 70% alcohol to protect them from damage from the bleach.
4. Thaw samples at room temperature, at 4°C, or on ice. Keep the samples on ice or at 4°C until they are used. DNA samples will be stable at room temperature throughout the process of setting up a PCR experiment. If the process takes more than 2 hours, storage at 4°C or on ice is preferred, though degradation is unlikely to occur. When diluting large numbers of samples, work in small batches (≤ 12 at a time). Leave other samples frozen at -20°C or at 4°C if they will be diluted later in the same day.
5. Prepare clearly labelled siliconized or low-retention microcentrifuge tubes for each dilution in the series.
6. Use the following formula to calculate the volume to be transferred from the stock vial to the dilution vial: C1 x V1 = C2 x V2
   C1 = stock concentration
   V1 = volume to be transferred from the stock vial to the dilution tube
   C2 = desired concentration of diluted sample
   V2 = final volume of the diluted sample
   The appropriate volume of diluent to add can be calculated by subtracting V1 from V2.
7. Add the volume of diluent (V2-V1) to each tube in the dilution series; when expelling liquid from the pipette, DO NOT press the plunger past the first stop. The first stop indicates that the intended volume has been expelled. Any excess liquid should be discarded with the pipette tip.
8. Add the volume (V1) of neat (undiluted) sample to the first dilution tube; again, take care to NOT to press the pipette plunger past the first stop so that the sample does not become overly concentrated. Place the pipette tip just barely into the diluent liquid when dispensing, as some excess volume could be adhered to the tip exterior. Discard the pipette tip; be sure to use new pipette tips for each dilution.
9. Vortex the dilution tube for 3 pulses of 5 seconds each.
10. Spin down the tube for ~5 seconds in a mini centrifuge (an extra tube containing diluent can serve as a balance if necessary).
11. Add volume V1 of sample from the first dilution tube to the next tube in the series, using only the first stop of the pipette plunger and just barely submerging the tip into the diluent as before. Discard pipette tip.
12. Vortex the next dilution tube for 3 pulses of 5 seconds each.
13. Spin down as before for ~5 seconds (an extra dilution tube with diluent can serve as a balance).
14. Continue until the dilution series is complete.