Tips DNA Sequencing (Revision Date: 09 July 2007)

Transcription

1 Tips DNA Sequencing (Revision Date: 09 July 2007) 1. PCR product: clone, or directly sequence? 2. How do I clean DNA template for sequencing? 3. How much DNA should I use in a sequencing reaction? 4. Why should I aliquot my original tube of BigDye Terminator mix v3.1? 5. How much BigDye should I use in my reactions? 6. How should I prepare my sequencing reactions? 7. What is the standard sequencing protocol? 8. How do I sequence difficult DNA templates? 9. Why should I include positive controls? 10. How should I set up my positive controls? 11. Why does the location of my samples in a 96-well plate matter? 12. How do I clean my sequenced templates? 13. How do I resuspend my cleaned, sequenced templates? 14. How do I submit my samples to the Genomics Facility? 15. How long does it take to obtain my sequence data? 16. How do I obtain my sequence data? 17. How do I analyze my data? 18. Why did signal intensity initially soar and then plummet after ~150 bp? 19. In the Raw Signal view, why did the A, T, C & G baselines separate? 20. Why is there a low-level signal after the PCR-stop in my sequence data? 21. Why does signal intensity differ between capillaries on an array or different 3130XL s? 1. PCR product: clone, or directly sequence? (Back to Top) Cloning PCR products is both expensive and time consuming. Further, if you are concerned that bases at certain positions in the sequence might be heterozygous within an individual, you will have to sequence ~7-8 clones to ensure capturing that information with cloned product; with direct sequencing, such heterozygosity will usually be visible as two overlapping peaks that are roughly ½ as high as surrounding peaks. Nevertheless, sometimes the data cannot be obtained without cloning. a) Try direct sequencing of purified PCR product (by commercial columns or standard ethanol precipitation) when the sample runs as a clean, single band in a 2% agarose gel. b) Try direct sequencing of gel-purified PCR product when the sample runs as a clean band amongst other bands in a 2% agarose gel. c) If you know that your sequence contains a pure poly- singlebase region (> ~8-10 bases), either clone the product or try direct sequencing in both directions. (Due to strand-slippage during the PCR, sequence data becomes unreadable after the pure-base region.) d) With purified plasmid DNA, stretches of pure poly- single-base regions are not a problem, unless the stretches are extremely long (e.g., ~30-40 bases).

2 2. How do I clean DNA template for sequencing? (Back to Top) The final step in most techniques will involve ethanol. It is crucial that all ethanol be removed from the purified sample prior to preparing the sequencing reactions. Thus, your final step should always be to drive off any residual ethanol by incubating samples at 60 o C (open caps; ~10 min). a) PCR products: Assuming a robust reaction, you can do a standard ethanol precipitation (inexpensive method); otherwise, you probably should use a commercial column. b) Plasmids: Commercial columns give the best template for sequencing. If you use a homemade technique, please consult the Plasmid Prep document on our website. c) Commercial columns: Many protocols finish with a 5-min spin after adding the wash ethanol; however, the ethanol vapor pressure under the column will prevent some ethanol from spinning out of the filter. Thus, spin out the wash ethanol (1 min), dump the ethanol and blot collection tubes on a Kimwipe, and finish with a final spin (5 min). 3. How much DNA should I use in a sequencing reaction? (Back to Top) The most common sequencing error is to use too much DNA (volume or quantity). Excess volume increases the potential for including reaction-killing contaminants into the mix. Excess quantity has two synergistic effects, which can result in dramatically reduced sequence read length: (a) BigDye reagents are exhausted by making very small fragments; and, (b) excessive quantities of small fragments clog the capillary on the sequencer, preventing the injection of longer fragments. Unless your protocols consistently generate appropriate concentrations of DNA for sequencing, you will save time and money by taking steps to quantitate your DNA templates prior to sequencing them. a) Ideally, total DNA in a reaction should be ~2-6 ng for most PCR products and ng for plasmid DNA (~4,500 bp vectors). b) PCR products: Residue from commercial kits may give false A260 readings; e.g., a clean column from the Promega Wizard kit sometimes gives a false reading of ~30 ng/μl. i) Option A: Purify enough PCR product for a valid A260 reading (subtracting value of your false reading), and make serial dilution reference photo down to 1 ng in a clean 2% gel. Compare products to photo to estimate needed dilutions. ii) Option B: Run the equivalent of 1 μl & 3 μl of purified samples in a clean 2% gel; alternatively, run out unpurified PCR product and use your results to determine volumes for eluting your DNA from the column (or for resuspension, if doing an ethanol precipitation). Criteria: If 1-μl band is barely visible and 3-μl band is faint (but distinct), use 1-3 μl of DNA in sequencing reaction. If only the 3-ul band is visible, use more template; if the 1-ul band is bright, use less template. In the figure below, each DNA template has been loaded as described in Option B (5 μl of template were mixed with 20 μl of loading dye (20% Rediload); electrophoresed 5 & 15 μl). Probably all of these templates would be suitable for use with 1-3 ul in a sequencing reaction. However, A is verging on becoming too bright; B is perfect; C is getting somewhat weak; D is getting very weak; and, E would likely do much better if ~6 μl of template were used.

3 In the next figure, the templates are loaded in the same way, except in reverse order (i.e., 3 & 1 ul). In each case, 1-3 μl of DNA could be used in sequencing reactions ( A is perfect) although samples that look like B are verging on being too bright and you might want to consider using ~1 μl rather than 3 μl for such samples. c) Plasmid DNA: Take A260/A280 readings on a random sample of templates, and dilute the DNA accordingly; if readings are not relatively consistent, process all templates. i) Minimize variation: keep bacterial growth periods and processed volumes consistent. ii) Minimize growth time: yields better quality DNA and better sequencing results. Note: Spectrophotometers do not measure the quality of the DNA; thus, it is possible to have good OD s and yet have degraded DNA which cannot be sequenced. In fact, you should be suspicious of any samples that give extraordinarily high DNA readings.

4 4. Why should I aliquot my original tube of BigDye Terminator mix v3.1? (Back to Top) a) BigDye chemistry degrades following 5-10 freeze-thaw cycles. b) Given that typical DNA templates can be sequenced with as little as 0.5 μl of BigDye in a 10 μl reaction, a stock tube of BigDye can nominally generate up to 1600 reactions enough to process over sixteen 96-well plates. c) To prepare the BigDye for to aliquoting, vigorously vortex the thawed BigDye (~ 30 s) and spin it down. Store aliquoted BigDye at -20 o C. 5. How much BigDye should I use in my reactions? (Back to Top) Aside from cloning costs, BigDye is the most expensive reagent in sequencing; thus, use as little as necessary a) You can successfully sequence most templates ( bp reads) with 0.5 μl of BigDye in a 10-ul reaction. Although we bump up the relative concentration of BigDye by reducing the total reaction volume to 10 μl, we call this a 1/16X reaction because ABI s standard 1X reaction uses 8 μl BigDye [in 20 μl]). b) Other Reaction sizes 1/16X reactions are less sensitive to overloading with template DNA and may give better results with difficult DNA templates. Although a 1/48X reaction (100 ng pgem) can give >800 bp of good quality sequence, signal strengths are better at 1/32X. Reactions of 1/64X give good data, but only for ~ bp. (Note: Cleaning sequencing reactions by columns may reduce signal strength). BigDye/96 rxn s Reaction size Primer (3.2 μm) Buffer (2.5X) BigDye mix DNA + H 2 O μl 1/2 1 μl 0.00 μl ul 5 μl μl 1/4 1 μl 2.00 μl ul 5 μl 96.0 μl 1/8 1 μl 3.00 μl ul 5 μl 48.0 μl 1/16 1 μl 3.50 μl μl 5 μl 38.4 μl 1/20 1 μl 3.60 μl μl 5 μl 32.0 μl 1/24 1 μl 3.67 μl μl 5 μl 27.4 μl 1/28 1 μl 3.71 μl μl 5 μl 24.0 μl 1/32 1 μl 3.75 μl μl 5 μl 18.0 μl 1/48 1 μl 3.81 μl μl 5 μl 12.0 μl 1/64 1 μl 3.88 μl μl 5 μl 6. How should I prepare my sequencing reactions? (Back to Top) Normally, it is not necessary to keep the 96-well plate on ice while preparing reactions. a) DNA template: Add to bottom of wells, taking care not to cross-contaminate wells. i) When planning the layout of templates in a 96-well plate, keep in mind how the sequencer will process the plate. That is, it will pick up samples in sets of 16; further, a particular capillary in the array will always process the same relative well location from each set of 16 (e.g., the same capillary processes wells A1, A3, A5, A7, A9, and A11). ii) Spin plate prior to adding any other reagents. b) BigDye: Thaw, vortex vigorously, and place on ice otherwise data quality may suffer.

5 c) Mastermixes: Use them to minimize variation; add to top of wells. i) Singe primer: The mastermix should include everything except DNA templates. ii) Multiple primers (one per reaction!): Consider making an overall mastermix of the Buffer, BigDye, and Water... and then use that to make sub-mastermixes with each primer. If there are too few reactions with each primer, then add the primer to the top of each well instead... taking care not to cross-contaminate wells with the DNA templates already present... and spin plate before adding the mastermix. d) Total volume: At BigDye concentrations of <1/16X, reaction volumes >10 μl significantly reduce signal strength and read length; even 1/16X reactions perform better at 10 μl total volume. e) Final step: Seal plate, spin, and put in the PCR machine. 7. What is the standard sequencing protocol? (Back to Top) a) Cycling parameters: 95 o C initial denaturation (2 ); 25X [95 o C (10 s); 50 o C (5 s); 60 o C (4 )]; 20 o C hold (4 o C hold, if reactions will not be removed upon completion). b) Upon completion, either freeze or immediately clean the samples. 8. How do I sequence difficult DNA templates? (Back to Top) Sometimes, you will encounter templates that defy your standard sequencing techniques. The reactions may fail altogether, or they may work very well initially and then fail at a particular stretch of sequence. If you examine the last ~50 bp, you will likely see that the frequency of G or C was extremely high or that the segment consisted of microsatellites either of which leads to secondary structure, causing the polymerase to fall off. In this case, you can try the following ideas either singly or in various combinations. (Note: This is not the same situation as when your signal intensity initially soars and then plummets after ~150 bp.) a) BigDye: Use more BigDye to help force the reaction through the secondary structure. b) Template denaturation: Pipette the DNA (and possibly the primer as too) into the 96-well plate; seal with caps; denature for 5 minutes at o C; bury plate in ice until ready for use. After preparing the mastermix (iced), briefly spin plate prior to opening caps, set plate back on the ice asap, and add the mastermix. Keep plate on ice until the PCR machine is ready. c) Hot-start: Include a hot-start (5 min, o C) at the beginning of the cycling; further, don t put the plate into the PCR machine until it is hot. d) Denaturation temperature: Increase during cycling from the standard 95 o C to o C. e) Cycles: Increase from the standard 25 cycles (up to a maximum of 50). f) Primer design: Design a new primer closer to the region of difficulty. g) Call ABI: There are special kits available that may help with difficult templates. h) Figure (below): Results of standard sequencing (top) and modified sequencing (bottom) with a difficult template that contained a stretch of high G content. Even with the modified sequencing, the signal strength still dropped precipitously after the zone of secondary structure; nevertheless, adequate signal strength was maintained for the remainder of the sequence. When the same template was sequenced with a primer located closer to the high

6 G zone under the same modified sequencing protocol, there was only a moderate drop in signal strength (figure not shown). 9. Why should I include positive controls? (Back to Top) Including positive controls enables you to troubleshoot any sequencing problems more effectively. a) If your reagents are viable and your technique is good, commercially-prepared pgem 3Zf(+) DNA will provide both good quality sequence reads and read lengths of > bp on a 50-cm capillary with POP7. b) In the event of poor sequencing results, there are five primary culprits to consider: (a) 3130XL malfunction; (b) template; (c) sequencing reagents; (d) technique; and, (e) PCR machine. However, without positive controls run in the same sequencing reaction using the same mastermix, you have no idea where to begin troubleshooting. i) Controls work well: Focus on item b (template); but don t totally exclude other possibilities. ii) Controls work poorly or fail: Template issues are less likely. Item a can usually be eliminated by comparing your results to those of other users or those of sequencing standards that were run at the same time. In that case, focus on items c-e. Please contact the staff in the Genomics Facility for further guidance on this topic.

7 10. How should I set up my positive controls? (Back to Top) a) DNA template: We prefer pgem 3Zf(+) for two reasons: (i) it has a very distinctive raw signal signature; and, (ii) we know that it will work consistently. However, if necessary, other DNA sources can be used as positive controls, and may be preferable if your primers will not work on pgem. b) Primer choice: We prefer M13 (F or R) primer for use with pgem 3Zf(+) sequencing results with T3 or T7 are somewhat less robust. However, please note that, unless the same primer is used for both samples and controls, better results with the controls could be caused two different factors: (i) low quality template; or, (ii) degradation of the primer used to sequence your actual samples. c) Technique: Ensure that the controls are sequenced using the same mastermix as your samples. If different primers are used, we recommend that you initially make a primer-free BigDye Mastermix and split it into sub-mastermixes (i.e., one for each primer). For pgem 3Zf(+), use ng of DNA. Ideally, at least two controls should be prepared for any submission; for large submissions, please consider using at least one control for each set of wells. When submitting >16 samples, please place your controls such that: (i) they will all pass through different capillaries; and, (ii) the controls are split evenly amongst your sets of 16 samples. 11. Why does the location of my samples in a 96-well plate matter? (Back to Top) The 3130XL sequencer processes plates in sets of 16; further, a particular capillary in the array will always process the same relative well location from each set of 16 (e.g., the same capillary processes wells A1, A3, A5, A7, A9, and A11). a) Duplicate samples: If each member is processed by different capillaries, variation in the results is influenced by the relative quality of the capillaries used; by contrast, if processed by the same capillary, variation should be due primarily to differential efficiencies between the sequencing reaction of each well. b) Controls: When processed by different capillaries, controls tells us more about how well the entire array is working; by contrast, when the same capillary is used, variation in the results tells us more about the PCR machine that was used to perform the sequencing reactions. c) Run times: Consolidating samples of similar desired read length into sets of 16 can increase sample throughput by letting us decrease the required run times. For example, if the maximum read length required is 200 bp for samples in A1-H2 and 800 bp for samples in A3-H4, we can set the run module of A1-H2 for ~1/4 the time of the run module for A3-H How do I clean my sequenced templates (see pdf s on website)? (Back to Top) a) Ethanol precipitation is inexpensive, simple, and generates good quality data. b) Commercial columns are suitable, but more expensive; costs can be reduced by reusing columns, after appropriate precautions. Reuse leads to dye terminator peaks in first 30-70

8 bp of sequence. To prevent loss of signal strength, dilute completed reactions to 20 μl prior to loading on column. c) CleanSEQ by Agencourt costs ~$0.40/sample, but yields extremely clean DNA and exceptional sequence reads. Please see us prior to using this method. d) BigDye XTerminator Purification Kit (e.g., Part # ): Costs ~$0.50 to $1/sample. Samples are never dried nor resuspended in formamide. ABI claims complete removal of dye blobs and better sequencing results overall; method requires the addition of only two reagents which can be added sequentially or premixed: * XTerminator Solution Scavenges unincorporated dye terminators and free salts from the post-sequencing reaction. * SAM Solution Enhances the performance of the XTerminator Solution and stabilizes the post-purification reactions. Note: If you use this method, you must notify us of that fact when submitting your samples as we need to associate your samples with a different run module on the sequencer. 13. How do I resuspend my cleaned, sequenced templates? (Back to Top) a) Ensure that samples are completely dry (i.e., all water and ethanol have been removed). b) To wells with samples, add 15 μl of Hi-Di formamide (see pdf Formamide issues ). i) Do NOT create Sets of 16 by putting formamide in blank wells; we may need them! ii) Seal samples; lightly vortex; and, briefly centrifuge (store at 4 o C or freeze). c) With good technique, signal strengths should be well above the minimum threshold when samples are resuspended in formamide. However, if signal strength is a problem even after thoroughly troubleshooting your situation, consider resuspending your samples in water. i) Resuspension in water can increase signal strength by >10X, but the samples must be overlain with mineral oil to prevent oxidation. If you wish to resuspend your samples in water, you MUST have prior approval from the staff of the Genomics Facility. ii) If considering this option, please note that one purpose of resuspending samples in formamide is to maintain the DNA in a denatured state. So far, we have not noticed any problem with using water instead of formamide; however, resuspension in water might be inappropriate if your templates are capable of forming substantial secondary structure. 14. How do I submit my samples to the Genomics Facility? (Back to Top) a) Login to Genomics Facility website to access the Request Form : i) Sample names: Download template; follow the INSTRUCTIONS to fill out all required information; save file to your computer under a new name; and, upload file to website. ii) Other information: Fill out appropriately, including comments (if desired).

9 b) Physical submission: Sealed 96-well plates (preferred); tubes ( 20 samples, in a box). i) Label plates: Date; PI name; and, Submitter name (for tubes, use grid coordinates from Sample name file to label tops of tubes and then label the box with Date, PI, and Name). ii) Place samples in To Be Sequenced box in Genomics Facility refrigerator (Rm. A628). 15. How long does it take to obtain my sequence data? (Back to Top) a) Time required for a standard run module: ~2 hr for a single run (16 samples); ~6 hr for a half-plate (48 samples); and, ~12 hr for a full plate (96 samples). b) Typically, results are available within 1-3 working days of submission. 16. How do I obtain my sequence data? (Back to Top) a) Login to the Genomic Facility website. When your sequences are ready, your submission will include a link to the Results File. b) Results Files will be zipped, reducing ~30 Mb of data from a full plate (96 samples) to ~10 Mb; after 30 days, all files are archived. 17. How do I analyze my data? (Back to Top) a) Results are available as text files and electropherograms. i) Text files are accessible with programs such as DNAStar, BioEdit, or Notepad. ii) Electropherograms are accessible with freeware such as: (1) Sequence Scanner v1.0 by ABI (provides raw data, signal intensity, & more!); (2) BioEdit an excellent alignment tool; and, (3) Chromas LITE. b) The 3130XL s Sequencing Analysis Software can sometimes improve your results; however, we will perform such analyses only after a specific request; use Sequence Scanner v1.0 to determine which sequences are worth having reanalyzed. 18. Why did signal intensity initially soar and then plummet after ~150 bp? (Back to Top) If your samples are resuspended in formamide and the average signal intensity is >1000 (in the Annotation window of Sequence Scanner), then you likely used an excessive amount of DNA template in the sequencing reaction.

10 19. In the Raw Signal view, why did the A, T, C & G baselines separate? (Back to Top) There are several possibilities according to ABI. However, a major culprit is degradation of the dye chemistry... most likely due to multiple freeze-thaw cycles. See Why should I aliquot my original tube of BigDye Terminator mix v3.1 for further details. 20. Why is there a low-level signal after the PCR-stop in my sequence data? (Back to Top) There are at least three possibilities. First, you may have a second PCR product present at a much lower level than the primary product. Second, you might have some trace genomic DNA present in the sequencing reaction. Third, if you reuse your sequencing plates by rinsing out the plates, there might be some residual sequencing reaction from previous runs (see picture). 21. Why does signal intensity differ between capillaries on an array or different 3130XL s? (Back to Top) Aside from varying results from your sequencing reactions themselves (which can occur for numerous reasons) or the cleanup, each capillary in the set of 16 allows different amounts of the fluorescent signal to pass through it. A good guide to the performance of an individual capillary is its peak height in a spatial analysis. If you are concerned that the signal strength of your samples may be very low, please state that in your comments in the Request Form. If possible, the samples will then be run on the array currently having the best spatial peaks.