Tips for sequencing DNA

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Claribel Malone
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1 Sequencing tips and trouble shooting from website Tips for sequencing DNA Use clean DNA The cleanliness of the DNA is the most important factor in the success of automated DNA sequencing. The DNA should be free of proteins, RNA, polysaccharides and genomic DNA. This can best be achieved by using either a commercial plasmid miniprep kit, or by sequencing a PCR amplified fragment. Use small sequencing reaction volumes The high expense of the DNA sequencing chemistries (BigDye) has encouraged many researchers to reduce the amount used from that recommended by ABI (ie 8 µl in a 20 µl reaction volume). The most common approach used is to dilute the sequencing chemistry. This unfortunately has the side effect of causing substrate limitations the DNA polymerase (ie the nucleotide concentration falls below the Km of the DNA polymerase), leading to short reads and "ski sloped" traces (ie traces which show a very rapid drop in raw channel signal intensity). If you wish to save on BigDye then it is much better to reduce the reaction volume rather than BigDye concentration. For more information on how you can reduce BigDye usage without dilution please visit our dlute SEQ DNA sequencing reagent page. Don't use too much DNA in the sequencing reaction This is one of the most common mistakes made. You should aim to use no more than 25 ng of template times the length of the template in kilo bases. For example, if you are sequencing a 3 kb plasmid with a 2 kb insert, you should aim to use 25ng x (3kb + 2kb) = 125 ng in total. Improving plasmid miniprep kits template quality The sequence quality obtainable from most commercial plasmid miniprep preparation kits can be increased by performing a final ethanol precipitation of the purified plasmid

2 DNA. Using this step can remove leftover salts and wash buffer that may not have been removed fully during the kit DNA purification process. Sequence PCR products rather than plasmid templates Polymerase chain reaction templates tend to suffer less from sequencing problems than plasmids do. There are a number of reasons for this. Firstly, PCR products are less likely to contain DNA polymerase inhibitors than plasmid DNA after all if a DNA sample contains DNA polymerase inhibitors then it will not yield a PCR product. Secondly, PCR products are linear so that they are much easier to denature fully without template "snap back" occurring. The avoidance of template snap back can be a great help with high G+C templates and PCR products can be purified for sequencing very easily using ExoI/SAP treatment. Finally, PCR allows modified nucleotides to be incorporated into the PCR template during amplification which can help to overcome secondary structure problems. Dry the template DNA before sequencing One of the simplest ways of reducing the volume of a sequencing reaction is to dry the DNA template down in the sequencing wells before adding the sequencing chemistry and primers. Drying will not effect the DNA, and it allows the reaction volume to be significantly reduced, saving money and improving the quality of the resulting sequencing reaction. The easiest way to accomplish this is to put the DNA in the tube or tray and then place in a PCR machine set to 80 C with the lid open. The DNA should dry in around 10 minutes. You can also add the primer at the same time and dry it down with the DNA. Just remember not to dry down large volumes containing reaction inhibitors like EDTA! Linearize high G+C plasmids before sequencing High G+C (>60%) plasmids can be very difficult to sequence because the supercoiled template can rehybridize (the "snap back" problem) before the sequencing DNA polymerase can complete the full extension. Digestion of the plasmid template with a restriction enzyme before sequencing can help over come this - just remember to not select a

3 restriction enzyme that cuts between the sequencing primer binding site and the region you want to sequence! Use a different sequencing clean-up protocol A common cause of sequencing problems in the reaction clean-up. While many users are able to successfully clean up their reactions by ethanol precipitation, it is also the cause of many problems. Clean-ups by ethanol precipitation requires very precise ethanol concentrations and centrifugation times are used. If the ethanol concentration is too high then the left over sequencing chemistry (BigDye) is precipitated along with the sequencing products, resulting in dye blobs at the start of the trace. If the ethanol concentration is too low then a failed reaction (no signal) results. The leeway between these two outcomes is only 2.5%! If you are having difficulties with your sequencing reaction clean-up you may want to consider trying the phenol/butanol sequencing clean-up method * or purchasing a commercial clean-up kit. Increase the number of sequencing thermo cycles The intensity of sequencing signal is proportional to the number of thermo cycles performed. If you are finding that your signal intensity is just under the acceptable level, then increase your number of cycles can help. For example, if your current protocol use 25 cycles try using 35 or even 45 cycles. You can use up to 99 cycles without any problems. This is very a simple way of increasing signal strength. If you generally cycle your sequencing reaction overnight then this will not impact on the time required (ie rather than sitting at 4 C for hours the reaction will be generating more template).

4 *Overview This protocol is a robust method that will give you the best quality sequencing clean-up. It is more complex than a simple ethanol precipitation so should only be used when you are having clean-up difficulties. 1. Description of Sequencing Clean-up Process Phenol extraction and butanol precipitation of sequencing reactions. 2. Chemicals involved in Phenol/Butanol Cleanup Process Phenol CAS# butanol CAS# Potential Hazards Inhalation of toxic vapors - phenol Inhalation of harmful vapors - butanol Skin contact with toxic and harmful liquids - phenol and butanol are harmful and readily absorbed through skin. Skin burns - phenol burns skin Fire hazard - butanol is flammable, phenol is combustible. 4. Personal Protective Equipment Wear a lab coat, enclosed shoes, safety glasses or goggles and nitrile gloves for eye and skin protection. 5. Engineering/ Ventilation Controls Use phenol and butanol in the fumehood. 6. Special Handling Procedures and Storage Requirements Avoid using phenol and butanol near an open flame. Recap bottles immediately after use. Store source bottles of butanol in the flammables cupboard. Working stocks of solutions are to be kept in glass bottles and labeled correctly. 7. Spill and Accident Procedures Skin Exposure: rinse the affected skin with plenty of water while removing contaminated clothes and shoes. Rinse for at least 15min. Seek medical attention.

5 Eye exposure: Wash eyes for at least 15min, lifting the upper and lower eyelids. Seek medical attention immediately. Small Spills: Notify others in the area of a spill. Do not attempt to clean up if you feel unsure of your ability to do so or if you perceive the risk to be greater than the normal laboratory operations. Cover spill with absorbent material (kitty litter) supplied in the Spill Kit. When absorbent is removed, wash contaminated area. Large Spills: Notify others in the area of a spill. Turn off ignition sources in area. Evacuate area and post entrance ways to spill area. Restrict persons from the spill area until the clean up is complete. Remain in the area in a safe location to assist with response by emergency services. 8. Decontamination Procedures After cleaning up spill with absorbent material, dispose of contaminated material in the appropriate solid waste container. Wash area of spill with water. 9. Waste Disposal Procedures Disposal of the accumulated waste in the appropriate waste disposal container. Phenol/Butanol Sequencing Cleanup Protocol Materials Sequencing reactions that are 8ul volume in 0.2mL tubes MilliQ water Phenol 1-Butanol 0.5ml tubes Protocol 1. Set up the appropriate number of 0.5ml tubes labeled as per the numbering system. Leave one set empty to be used in step 4. Add 300µl of butanol to the other set to be used in step 8.

6 2. To each sequencing reaction add 30µl of MilliQ water using the Eppendorf P50 multi channel pipette if cleaning a 96 well tray or a P200 if working with a smaller set of sequencing reactions. 3. Add 25µl of phenol to each tube using the Eppendorf P50 multi channel pipette if cleaning a 96 well tray or a P200 if working with a smaller set of reactions. 4. Transfer the samples to 0.5ml tubes using a P200 pippetor. 5. Vortex samples until cloudy. Do this by tilting the tube on its side and pulsing it against the vortex for 2 seconds twice. If the sample is still not cloudy at this point, repeat vortexing until it is cloudy. 6. Centrifuge in a table top centrifuge at rpm for 3min. 7. Remove the top layer (aqueous phase) to a fresh 0.5ml tube containing 300µl of butanol using a single channel P200. This step should be done immediately after the centrifuge has stopped and requires careful pipetting as you only want to take the top layer and not the white interface or the lower phenol layer. Try and take about 36µl of the top phase, but if there are white bits as you get closer to the interface, then leave them behind. If the sample is still cloudy after centrifugation then repeat the vortex step and then recentrifuge for 3min. 8. Vortex samples with butanol well. Do this by tilting the tube on its side and pulsing it against the vortex for 10 seconds twice. The butanol needs to be completely mixed in with the sample, if you see droplets at the bottom of the tube then vortex it until the sample and butanol are completely mixed. 9. Centrifuge in a table top centrifuge at 12,000 rpm for 10min. Position the tubes with the edge of the cap facing out, this will ensure you know the orientation of the DNA pellet. 10. Remove the supernatant from the sample. After the centrifugation step 10. The pellet is positioned in the bottom of the edge side of the tube. Remove the supernatant by briefly tilting the tube with the edge facing upwards. Do not jostle the tubes too much as you don't

7 want to disturb the pellet. Tilting the tube removes most of the supernatant, but to remove the remaining supernatant see step Briefly centrifuge the samples by holding the pulse of the centrifuge until it accelerates to max speed then stop the centrifuge. This is enough to bring down the remaining supernatant. 12. Remove the remaining supernatant from the bottom of the tube using an single channel P20. Position your pipette at the opposite corner of the bottom of the tube to the pellet. Do not disturb the pellet. 13. Place tubes in the 80 deg.c drying oven to remove any traces of butanol that are remaining. This should only take 5-10min. 14. Add 1ul (2ul if only loading half) of sequencing loading dye to the sample and vortex for 30s. 15. Store sample in the -20 deg.c. Hints It is a good ideal to aliquot out small lot of phenol in 0.5ml tubes and store in the -20 deg.c freezer. Always take care when working with phenol!

Failed DNA sequencing reaction Identifying failed DNA sequencing reactions The trace chromatogram has noisy or messy sequence peaks with low quality scores.

8 Failed DNA sequencing reaction Identifying failed DNA sequencing reactions The trace chromatogram has noisy or messy sequence peaks with low quality scores. Can have peaks that look real and have high quality scores, especially when the trace has been base called using the KB base caller. No or little signal in the raw data channels except for leftover dye at the beginning of the trace. Signal strength in the raw channel usually below 100. The trace sequence does not match either the expected sequence or other sequences in GenBank. Figure 1. Example of a failed reaction trace. (A) Processed channel data. (B) Raw channel data at the same base location. Causes of failed DNA sequencing reactions Poor quality DNA. Very common when sequencing plasmid miniprep templates. Loss of the reaction during clean up. This can be a particular problem when using ethanol precipitation clean up protocols. Too much template DNA. Excess template can kill the sequencing reaction. Wrong primer used. More common than you might think! Bad water. The water used contains an sequencing inhibitor. Degraded or failed synthesis primer. Oligonucleotide synthesis is chemically complex and primer synthesis failures is fairly common. Dead sequencing chemistry. Can occur if the BigDye chemistry is stored under the wrong conditions or is freeze-thawed too many times. Either the Taq DNA polymerase or dye labeled nucleotides can have degraded.

9 Blocked capillary. Every trace using a particular capillary fails. Can be identified by tracking trace quality on a trace by trace basis. Solving DNA sequencing reaction failures Poor quality DNA. The best way of avoiding this problem is to not sequence plasmid DNA and sequence a PCR amplified fragment of the plasmid insert. If this is not possible then it is recommended that a plasmid miniprep kit is used. One tip is to perform a final ethanol precipitation on the kit purified plasmid DNA. This often solves problems with the quality of the template. Loss of sequencing reaction during clean up. This can be avoided by not using an ethanol precipitation protocol to clean up the sequencing reaction. There are a number of kits that work very well, unfortunately they can be very expensive. One tip for avoiding loss of the reaction DNA pellet when using the ethanol protocol is to add 1µl of a 20 mg/ml solution of glycogen (Sigma G-1508) to the sequencing reaction before adding the ethanol. This helps make the pellet visible and the glycogen does not seem to interfere with the injection of the sequencing fragments onto the sequencers capillaries. Too much template DNA was used. This can be avoid by checking the concentration of the template on an agarose gel before sequencing. This will also allow you to see the purity of the template DNA and if there is a significant amount of contaminating genomic DNA or RNA present. Do not rely on a spectrophotometer reading to calculate the template concentration. Wrong primer. This is simple to solve, but can be difficult to detect. Check the sequence of the primer and template to make sure that the primer binding site is present. This can be a particular problem with some "universal" forward and reverse primer sequences which do not work with some common plasmids. Do not trust other people's working stock solutions (especially those of your unreliable your lab colleagues!) and make your own. It might take 5 minutes longer, but it will save you a lot of future headaches. Bad water. Inhibitors can end up in lab water stocks that can kill DNA sequencing reactions. If you think this may be a problem then throw out the water and use a fresh stock - remember water is cheap. Degraded primer. Don't use old diluted primer stocks. Store the primers in 10mM Tris/ 0.1 mm EDTA (ph 8.5) rather than water. Don't use other peoples stocks. If you have any doubt about how the primer quality through it out and make up a fresh working solution from the primer stock.

10 Failed oligonucleotide synthesis. If you suspect that the primer is poor quality either have it presbyters or check in a polymerase chain reaction (PCR). Alternatively if you have a control template that you know should work with the primer then this can be a good way of identifying primer problems. Dead sequencing chemistry. This is a relatively rare problem, however, if a batch of BigDye chemistry has not been used for some time, or there is any doubt about how it has been stored, then it is advisable to perform a control sequencing reaction before undertaking a large number of experimental reactions. Many problems with dead chemistry can be prevented by storing the BigDye chemistry in small aliquots and avoiding repeated freeze/thaw cycles. Blocked capillary. Can be identified by tracking trace quality on a trace by trace basis. While this can be done manually we recommend using our QualTrace software. If you suspect that a sequencer capillary is blocked then inform the operator.

RayBio Genomic DNA Magnetic Beads Kit

RayBio Genomic DNA Magnetic Beads Kit Catalog #: 801-112 User Manual Last revised January 4 th, 2017 Caution: Extraordinarily useful information enclosed ISO 13485 Certified 3607 Parkway Lane, Suite 100