Dynabeads Nucleic Acid Purification Support—Troubleshooting

Please review the following possibilities for why your Dynabeads magnetic beads are not pelleting:

• The solution is too viscous. 
• The beads have formed aggregates because of protein-protein interaction. 

Try these suggestions:

• Increase separation time (leave tub on magnet for 2–5 minutes) 
• Add DNase I to the lysate (~0.01 mg/mL) 
• Increase the Tween 20 concentration to ~0.05% of the binding and/or washing buffer. 
• Add up to 20 mM beta-mercaptoethanol to the binding and/or wash buffers.

The streptavidin molecule is covalently attached to the surface of the beads; under normal, recommended conditions, negligible leakage is detected (less than 0.2% of total attached streptavidin after 2 months at 37°C). However, it should be noted that not all of the four streptavidin subunits are covalently coupled to the beads. Typically, one or two of the subunits are covalently coupled. Streptavidin is like other proteins; if heated, it can denature and dissociate into subunits. If Dynabeads Streptavidin magnetic beads are, for instance, boiled, some of the streptavidin subunits may be released (as monomers or aggregates) from the beads. The covalently bound streptavidin subunits will not be affected by such treatment. When streptavidin is bound to biotin, the streptavidin-biotin complex is more stable than the unbound streptavidin molecule.

Yes, you can still use them. To test stability at low temperature in-house, the Dynabeads M-280 Streptavidin beads were placed at -20 degrees C for 24 hrs and then transferred to room temperature (15-25 degrees C) for 72 hrs. This freeze/thaw cycle was repeated 4 times and the beads were stored at 2-8 degrees C for 60 months. The biotin binding capacity was then tested and was seen to be within our specifications.

The binding capacity of streptavidin-coupled Dynabeads magnetic beads is fragment length-dependent. Reduced binding capacity for large DNA or RNA fragments may be due to steric hindrance. For large DNA or RNA fragments (greater than 2 kb in size), we recommend using Dynabeads KilobaseBINDER Kit. Here are some suggestions for optimizing binding capacity.

• Salt concentration affects the binding efficiency of biotinylated nucleic acids to Dynabeads Streptavidin magnetic beads. Optimal binding conditions for biotinylated DNA fragments (up to 1 kb) are achieved at 1 M NaCl (final concentration), 25°C and 15 minutes of incubation. Longer DNA fragments should be immobilized overnight. Biotinylated antibodies should be immobilized in PBS buffer, pH 7.4, supplemented with 0.1% BSA. 
• Ensure that your sample does not contain an excess of free biotin, as the free biotin will bind Dynabeads Streptavidin magnetic beads much more rapidly than larger biotinylated molecules. Biotinylated oligonucleotides should be recovered by reverse-phase HPLC or FPLC to remove free biotin from the sample. 
• We also recommend a titration to optimize the quantity of beads used for each individual application, since both the size of the specific molecule to be immobilized and the biotinylation procedures affect the binding capacity of the beads.

Maintain a high pH and a low salt concentration, which will help to maintain the negative charge on both the nucleic acids and the beads.

The background might be caused by nonspecific binding to the BSA on the bead surface. Alternatively, high background might be caused by nonspecific binding to streptavidin. Increasing either the pH or the salt concentration might help reduce the binding. Dynabeads M-270 Streptavidin magnetic beads might be a better alternative; these beads are not coated with BSA and are hydrophilic, as they are based upon carboxylic acid chemistry.

Dynabeads M-270 Streptavidin magnetic beads and Dynabeads MyOne Streptavidin C1 magnetic beads have a negatively charged surface. The surface charge of the beads may in some samples cause the beads to float or become sticky or aggregate. The stickiness may be due to electrostatic interactions between the beads or between the beads and the tube wall. Usually we recommend washing the beads in a nonionic detergent like Tween 20 detergent before doing the experiment. The problem is usually reduced or eliminated by simply adding Tween 20 detergent to a final concentration of up to 0.1% to the beads, followed by resuspension and washing in buffer without the detergent. An incubation in the Tween 20 solution may be needed, e.g. 5–10 minutes at room temperature on a roller. In addition, we recommend using siliconized tubes. This treatment will most likely reduce the electrostatic potential of the beads.

We do not recommend this as streptavidin becomes hydrophobic and aggregates during denaturation.

Ribosomal RNA is effectively eliminated by reextracting the mRNA from the eluate. Reuse the same Dynabeads Oligo(dT)25 beads that were used for the original isolation. Wash the beads twice in Washing Buffer B. Dilute the eluted mRNA with 4 times its volume of Lysis/Binding Buffer, then add the beads. Incubate with mixing at room temperature for 3–5 minutes, then continue with the Direct mRNA Isolation Protocol.

PCR may be inhibited by inhibitors, or an insufficient number of PCR cycles might have been used. For inhibitors, ensure that the washing buffer is brought to room temperature for use, and add the buffer more vigorously. You can also ensure that the supernatant is completely removed at each washing step, or introduce an additional washing step.

We recommend eluting the DNA in Resuspension Buffer, water, or low ionic strength buffer by incubation at 65 degrees C for >5 mins. The complex must be fully resuspended before elution. The elution buffer can be pre-heated to 65 degrees C. After incubation, place tube in magnetic stand for 30 secs and transfer supernatant to a clean tube. To determine the elution efficiency, you can run some of the eluted DNA on an agarose gel. Alternatively, you can perform a PCR with the Dynabeads to determine if there is residual DNA remaining on the Dynabeads. Using more pre-heated elution buffer would be helpful to improve the elution efficiency.

High PCR background could be caused by too high a concentration of template DNA, primers, Mg2+, or dNTPs. To fix this, reduce the amount of template DNA, primers, enzyme, and Mg2+ used, elute the DNA prior to PCR amplification, and/or try performing hot-start PCR. High background can also be caused by contamination. Ensure that the supernatant is completely removed at each washing step to avoid carryover.

We suggest checking the white blood cell count, and/or increasing the sample quality and quantity.

You can try to pipette for longer, or move the tube rapidly over an uneven surface, e.g., an iron thread tube rack, to achieve a shaking motion. You can also try to use a tip with a smaller aperture.

Here are some suggestions you can try:

• If fragmentation of the complex was observed during the washing steps, reduce the force used to add wash buffer. 
• Elution, if performed, may have been inefficient. Ensure that all Washing Buffer is removed prior to addition of Resuspension Buffer. Increase the force used to homogenize the complex at the resuspension step. 
• Increase sample quality and quantity.

Here are some suggestions you can try:

• Red cell lysis may have been incomplete; increase the incubation time or repeat the red cell lysis step. 
• Add Washing Buffer more vigorously. 
• Ensure that the supernatant is completely removed at each washing step. 
• Use less starting material for PCR.

The biggest cause of yellowing is the addition of isopropanol when Lysis/Binding Buffer is used to make Wash Buffer 1. The use of poor quality isopropanol (not molecular grade) will cause rapid yellowing. High quality alcohols do not cause this.