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We are dedicated to your success. Get back on track. View our expert recommendations for commonly encountered problem scenarios.
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The most common problem related to DNA solubilization occurs when the DNA pellets are overdried. It is very important not to dry pellets longer than 5 minutes. The use of vacuum suction devices to remove the wash solutions may cause overdrying of DNA pellets. Vacuum suction draws air through the pellet and almost always will overdry the DNA pellets. Avoid removing the wash solutions with any type of vacuum suction device and limit the drying time to <5 minutes. If you follow our simple recommendations below, you can avoid many nucleic acid solubility problems.
Remove droplets of ethanol from the wall of the test tube with a sterile cotton swab. Additional ethanol can be removed by touching the pellet with a sterile capillary pipette tip. Excess ethanol will be drawn inside the pipette by capillarity. Residual ethanol that may remain in the DNA pellet will not be harmful. You can usually eliminate DNA solubility issues by adding either TE buffer or 8 mM NaOH to the pellet before all of the ethanol has evaporated. The DNA pellets will become clear after a 5–10 minute incubation, as they begin to rehydrate. In order to solubilize the DNA completely, the solution must be pipetted up and down before removing an aliquot for quantitation.
DNA pellets that are overdried can be solubilized but it may be necessary to put them into the refrigerator and pipet them periodically until they become clear and go into solution.
Polysaccharides are water-soluble and they will partition into the aqueous phase with the RNA. Also, RNA and DNA pellets that contain contaminants tend to solubilize more easily than pellets that are very pure if they are not overdried. Pellets that do not solubilize in 8 mM NaOH will not solubilize in a phenol/chloroform solution, either.
Here are some possible causes for low yield/DNA degradation:
Typically, low absorbance is due to phenol contamination. You should include additional washes with 0.1 M sodium citrate in 10% ethanol. It's not unusual for residual phenol from the extraction to remain, and the A260/A280 ratio of the extracted material would show a higher than expected A280. We recommend a second ethanol precipitation to remove remaining phenol. This will also remove any excess salt. If the tube smells like phenol after the procedure is done, precipitate the DNA again. It is important to do this, as phenol inhibits downstream enzymatic reactions.
If the aqueous phase was removed completely and ethanol was added to the samples, it will remain on top of the TRIzol® Reagent due to ethanol’s lower density. If the samples were centrifuged without mixing the two liquids, the ethanol will remain on top of the TRIzol® Reagent after centrifugation, the DNA will remain at the interface, and the TRIzol® Reagent will be localized to the red organic fraction on the bottom. If the ethanol was not mixed properly, proceed with mixing the samples, then centrifuge and continue to step 1 of the DNA isolation protocol.
If 70% ethanol was added accidentally, it may be possible to get a small volume of water on top of the organic fraction. Since the wash solutions that are used in the protocol do not exceed 30% water, you would expect to see no more than 30% of 0.3 mL (90 µL) of water on top of the organic fraction. You can try removing and discarding the water before proceeding with the isolation. DNA yield may be decreased.
This could also happen if the phase separation was not complete during the RNA isolation step. This can occur because the chloroform was not adequately mixed or if the samples were not centrifuged at the proper g-force or for the required period of time or at the correct temperature. The net result is that significantly less than 600 µL of the RNA aqueous phase will be recovered from the sample. Phase separation problems usually occur when the chloroform is mixed in the tube by vortexing. Due to the large difference in density between TRIzol® Reagent and the organic phase, the solutions are never mixed completely and only a portion of the aqueous phase will be recovered. When the ethanol is added and the samples are remixed sufficiently, the phase separation will go to completion and water could appear on top of the sample.
Yes, this happens due to the dye in the reagent, and seems to be dependent on the volume of the stored reagent. The color change does not affect its performance.
No, the 8 mM NaOH will not affect the DNA integrity. In fact, DNA is most stable at slightly alkaline pH (>7). You will find that the isolated DNA does not resuspend well in water and has even worse solubility in Tris buffer. (Water often has a pH of lower than 7 due to dissolved CO2 from the air. This slightly acidic water will actually cause degradation of your DNA.) The pH of the 8 mM NaOH is ~9 and can be easily adjusted with TE or HEPES once the DNA is in solution. (Over time, the solution becomes neutral upon exposure to air from dissolved carbon dioxide.)
Consider the following if you have a low A260/A280 ratio:
You can try incubating samples resuspended in 8 mM NaOH at 37°C overnight to resuspend the DNA. You can also try incubating at 45°C for 15 minutes.
If the cells or tissue were washed with phosphate buffer solutions prior to DNA isolation, the phosphate may have been carried over and be inhibiting restriction enzymes. We recommend adding DNAzol® Reagent to the DNA solution and reprecipitating with 0.5 volumes of 95% EtOH. Wash twice with 95%, dry briefly, and resuspend in 8 mM NaOH.
It is possible to see two phases after addition of ethanol if the amount of DNAzol® Reagent was too low. Add more DNAzol® Reagent and continue.
The color is due to cells lysing before the DNAzol® Reagent is added. The color is caused by hemoglobin. This contaminant will cause problems during PCR, and must be removed before the ethanol wash step to prevent them. The following are possible reasons for the contamination:
If your PCR reactions have been left open, and your PCR product is quite small, then evaporation can cause a size exclusion effect, reducing your yield or even completely removing your product.
It is possible to increase size of the smallest species purified by increasing the volume of clean-up buffer used, so that the large primers do not remain for the final elution. You will have to carry out a series of titrations to optimize this size exclusion process.
ChargeSwitch® coated beads are inert and will not affect PCR. We have found that volumes of as much as 10 µL of beads have no deleterious effects on regular PCR when spiked in. Excessive quantities greater than ~10 µL of beads will start to inhibit PCR. Some specific applications may be affected by beads, including real-time reactions or MALDI-TOF. In these applications, repeat the final elution binding as described in the protocol.
Buffers have been optimized (through use of millimolar quantities of salts) to avoid the binding of proteins. Our validation studies have indicated that no protein binding occurs.
This can occur for several reasons:
There are several reasons low yields can occur:
Here are some suggestions for your experiments:
Multiple bands can occur due to high heat generated during electrophoresis or by running the gel too fast. The isolated DNA can then appear as multiple bands when the eluted DNA is analyzed on a gel. Denaturation can also occur in AT-rich DNA during the 50°C incubation to dissolve the gel slices. If this happens, solubilize the gel at 37°C for 20 to 30 minutes with repeated vortexing.
There are several options to rescue your DNA obtain good PCR results including diluting your DNA sample, using specific enzymes, and/or using an additive. The methods are described below:
- Dilute your DNA sample 10-100 times prior to PCR which will dilute out the contaminants while the PCR signal in many cases will remain strong.
- Use “tough” enzymes that were specifically designed and optimized to be effective in harsh environments, and work in the presence of high levels of inhibitors, for instance, TaqPath qPCR Master Mix, CG Reagent.
- Use a BSA additive for the PCR reaction. In most cases, if used at appropriate concentration, BSA can rescue your sample.
You can also try repeating the purification with less starting material or increased volume of S1-Lysis buffer. Additionally, after the addition of the S3- Cleanup buffer, you can try incubating the sample for 10 minutes on ice before centrifugation.
Low yield could be caused by inefficient lysis and/or low levels of DNA in the sample. Please try heating samples at 95 degrees C for 5-10 minutes instead of 65 degrees C for 10 minutes after adding S2-Lysis Enhancer. Bead beat for a longer time or at a higher power setting. You can also try to perform the experiment with more starting material, but do not exceed what we specify in the protocol. However for some challenging samples, too much starting material can also result in low yield. In this case, please try less starting material and/or increase the volume of S1-Lysis Buffer.
For some samples, including soil and stool samples, it can be difficult to withdraw 400 μL supernatant while avoiding debris. If less than 250 μL of supernatant is transferred, add S1-Lysis buffer to bring the volume to 400 μL, before adding S3-Cleanup Buffer.
For Research Use Only. Not for use in diagnostic procedures.