Genomic DNA Purification Support—Troubleshooting

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:

• The sample was not fully homogenized or lysed. If any solid material remains after chloroform is added, this indicates that DNA yield may be poor, as DNA will remain trapped in the unhomogenized material. You can remove unhomogenized material by passing the TRIzol® homogenate (prior to phase separation) through a polypropylene filter cloth. 
• The final DNA pellet was not fully redissolved. It can take several hours to resuspend the DNA. Some incubation at 37°C between pipettings will help. Also make sure that it is not too concentrated or it won't go back into solution. If the DNA is not fully redissolved, it will be lost during the final centrifugation when removing the gel-like material. 
• The tissue was not IMMEDIATELY processed or frozen after removal from the animal or other source. 
• Samples were homogenized with a high-speed homogenizer. DNA shearing can happen. 
• If expected yield is <10 μg, there are limitations to the physical action of precipitation that would lead to low yields. A microcarrier (glycogen, tRNA) may be included in the homogenization and/or wash steps, or samples may be pooled to increase the expected yield. 
• If you are only interested in quantitative DNA recovery, we suggest resuspending in freshly made 40 mM NaOH instead of 8 mM NaOH; this will hydrolyze DNA but will facilitate solubilization. Also, do not centrifuge after homogenization (prior to adding chloroform in the initial step), as some of the DNA will precipitate during this centrifugation. If the pellet is slippery, the speed of the first centrifugation may be increased to 5,000 x g; this will make the pellet more difficult to solubilize, and it may need to be vortexed and heated to 50°C.

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:

• The correct amount of DNAzol® Reagent may not have been used. If DNAzol® Reagent was added to a cell pellet, make sure that the volume of reagent was 20 times that of the cell pellet. 
• There may have been a problem in pipetting away the viscous supernatant from the DNA pellet, leading to contamination with protein. The DNA may be extracted again with DNAzol® Reagent or extracted with phenol to remove the protein. 
• In some samples dissolved in water, the ratio may be low due to the acidity of the water or the low ion content in the water. The ratio may go up if the sample is dissolved in TE and the spectrophotometer is zeroed with TE (or 1 to 3 mM Na2PO4, pH ~8.0). The molar extinction coefficient of nucleotides is given at neutral pH, suggesting that the absorbance at 260 nm would be highest at neutral pH. DNA is not stable under acidic conditions, so degradation may occur if the DNA is left in this condition for too long.

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:

• Inadequate or ineffective anticoagulants can result in clot formation; typically, this will give DNA a speckled appearance. Remove them by centrifugation before precipitating the DNA.
• The walls of the tube were not adequately washed with the DNAzol® BD wash solution. Trace amounts of protein on the wall of the tube can cause contamination if this residue is dislodged during the ethanol wash steps.

Wipe the magnetic rods with a soft cloth or tissue paper soaked in a mild detergent solution, soap solution, or alcohol.

The protocol will not start without the tip combs inserted into the tip comb holder.

If the starting material is too viscous, the magnetic rods will not be able to collect the particles. Dilute the sample and check that the sample has been properly homogenized and lysed.

There is a pause function if you hit "stop" in the middle of a run. If you hit start again, the run continues. If you hit stop twice, the run stops completely. There is no way to begin a run in the middle of a protocol.

This happens sometimes, but it will not affect the yield because the sample has been released from the particles.

A potential cause is that the plastic tip comb has warped slightly due to its design. This is why they are packed in pairs, so that they maintain their shape. Flex the backbone of the tip comb and try again.

Turn the machine off, and inspect it for any visible damage. Make sure the correct head is being used for the protocol being run. If the machine shows no visible damage, turn it back on. This should reset the machine. If the problem immediately occurs again, turn the machine off and gently move the magnetic head apparatus to the center of its travel path, then turn the machine on again. If the problem persists, the head may need to be realigned by a service call.

Yes, please order Cat. No. 4489112.

Please follow these recommended steps:

1. Press the ESC key on the keypad to return off the main screen. 
2. Select 1 to enter the manual screen. 
3. Select 2 to return the tips to the holder. The instrument will also move all axes to the original position.
4. Turn off the machine, remove the card, reinsert the card, restart the instrument, and run the protocol without reagents present. 

Should the same error code still appear, try the following: Turn off the machine, remove the card, reinsert the card, restart the instrument, and under the manual menu, choose option 2, return tips. Then run the protocol without reagents.

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:

• The incorrect ratio of gel solubilization buffer to agarose mass was used. When the agarose concentration is below 2%, use 300 μL of gel solubilization buffer for every 100 mg of gel. When the agarose concentration is above 2%, use 600 μL of gel solubilization buffer for every 100 mg of gel. 
• The solubilization step was carried out below 50°C. 
• The sample was not vortexed thoroughly every 3 minutes. 
• The solubilization step was not carried out long enough. 
• The mass of the agarose gel slice was greater than 100 mg. In this case, mincing the agarose will accelerate solubilization.

There are several reasons low yields can occur:

• Loading more than 400 mg of agarose. This will decrease the performance of the cartridge. 
• An incorrect ratio of solubilization buffer to gel was used. Use 30 μL of buffer per 10 mg of gel when the agarose is less than 2%. Use 60 μL of buffer per 10 mg of gel when the agarose concentration is above 2%. 
• Ethanol may not have been added to the wash buffer? Ethanol is necessary to keep DNA bound to the silica membrane. 
• The bottle with buffer W9 was not kept tightly closed when not in use. If evaporation reduces the ethanol content in buffer W9, recovery of DNA will be increasingly poor. 
• The gel was not completely solubilized before it was added to the column. Make sure to dissolve at 50°C and to mix every 3 minutes. 
• The DNA was not completely eluted. TE buffer prewarmed to 65–70°C can increase elution yield. 
• The DNA was supercoiled. To elute supercoiled DNA from the membrane, the ethanol concentration in the wash buffer must be 50 55%. Supercoiled DNA will not be eluted from the membrane using the standard Wash Buffer that contains 70% ethanol. 
• The DNA was supercoiled. Supercoiled DNA will not be eluted from the membrane using the Wash Buffer from the kit.

Here are some suggestions for your experiments:

• Many enzymes, including restriction endonucleases and ligases, are inhibited by small amounts of agarose and by perchlorate contaminants. This is not normally a problem. If it is, however, you can use the DNA without repurification by increasing the amount of enzyme used in the digest or ligation or by increasing the digestion incubation time. Also, you may use less DNA in your digest or ligation with the same total volume and the same amount of restriction enzyme. 
• There may have been residual ethanol in the eluted fragment. Be sure to thoroughly centrifuge to remove the wash buffer, discard the wash buffer, and use a fresh tube to collect the eluted DNA. For applications that are very sensitive to ethanol, let the open spin column stand for 15 minutes at room temperature to let any excess ethanol evaporate. 
• The washing steps may not be as efficient as they should be. Under these circumstances, there may be trace amounts of perchlorate in the eluate. To avoid this, extend the centrifugation times to 5 minutes and wash 2 times with wash buffer. 
• Be sure to perform the optional wash step if you are using higher concentrations of agarose or are adding more than 250 mg to the cartridge. If applications are sensitive to EDTA, elute with water (pH 7.5–8.5), or with 10 mM Tris, pH 8.0 without EDTA.

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.