|Bands start out normal (or even over-sized) but decline rapidly - "ski slope" effect.
||The 'ski-slope' effect is best viewed using a program of ours that "crunches" an entire chromatogram into one panel. Below is an example (actually, a fairly mild example of a ski-slope):
This is not well understood, but here are four possible explanations: (i) salt in the DNA, (ii) too much DNA in the reaction, (iii) an unknown impurity "poisoning" the Taq processivity, or (iv) an unknown contaminant increasing the binding of dyes in the enzyme's active site. The latter effect can arise from free NTP's in the sample, and *perhaps* from a contaminant that disturbs the divalent cation concentration (EDTA, Mg++ etc).
Since February 2001, salt is the most common cause of the 'ski-slope' effect. Capillary electrophoresis instruments such as our ABI Model 3700 sequencers are quite sensitive to the presence of excess salt. It tends to favor detection of smaller fragments over larger ones. Our purification protocols are designed to minimize this problem, but it still occurs at times.
The terminator concentrations are carefully adjusted to statistically favor long extension, and the enzyme is modified to be able to accept bulky dye molecules as substrates. Several of the possible explanations given above for the "ski-slope" effect all work by increasing the statistical likelihood of early termination.
|The sequence is generally good, but there's one place where a huge green (or red or black) peak obscures everything under it. The peak shape is clearly abnormal.
||This is a common artifact of automated sequencing that arises from complexes formed between the sequencing dyes and unknown other components (often contaminants). There are two things that cause this artifact:
First, if our sample cleanup is flawed, we might have left excess unincorporated dyes in the sample. We'll usually catch this, since it's pretty obvious on the gel image. Second, your sample itself may have a contaminant that binds unincorporated dyes.
In either case, you may be able to manually re-call the bases "underneath" the blob-peak. If the Core techs feel this is not possible, and if the "blob" appears to arise from our own processing problem, then they will initiate a no-charge repeat for you automatically.
Here's a typical example of what we call a "dye blob":
|Some bands in the first 100 nucleotides show all four colors, with green being the largest.
||Between roughly April of 1996 and January 1999, excessive amounts of DNA in the reaction caused 'A' residues to display multiple colors. The largest peak was always green, and this only happened in the first ca. 100 nucleotides. If you see this artifact in an old chromatogram from that period, may safely assume the base is an 'A' if your multi-colored peaks fit this description.
NOTE: We have not seen this artifact since changing to a newer type of dye, in early 1999. This section being kept primarily as a courtesy to clients of other Cores that still use these older dyes.
|Your sequence proceeds normally, then the bands abruptly become much smaller.
||Secondary structure in the template is the most likely cause of this problem. The polymerase is presumably unable to progress through some stem-loop form. Are you trying to sequence an siRNA (RNAi) construct? These will almost always exhibit strong sec-structure effects. A couple possible solutions: (i) try resequencing by selecting "Sec. Structure Template" as your DNA type (this usually is the best solution by far!), (ii) try to sequence from another primer at a different position (closer or further); (iii) sequence the other strand.
We maybe able to use special cycling conditions and/or special reagents that help the polymerase to push through this region. We cannot do this routinely, as it ties up a thermal cycler for just a few samples, but contact the Core Director to discuss the possibility.
If all else fails, Contact the Core Director for help.
Here's an example of a secondary structure effect:
|Your sequence proceeds normally, then the bands abruptly vanish.
||This usually happens when the template DNA has simply stopped, for example if it was restricted at a downstream site or if the template was a PCR product. This may also be caused by an extremely stable secondary structure. See the section above for suggestions on how to sequence your template.
|Some peaks seem to be missing. The machine called an 'N'.
||If the peak just *before* the missing one is green, this is normal. The enzyme we use has difficulty adding a 'G' immediately after 'A', with the result that the peak will be much smaller. Check to see if your 'N' has a small black band below it and an 'A' immediately before. If so, it's a 'G'-after-'A' dropout.
The chemistries we use are no longer likely to cause "dropouts" as described in the preceding paragraph. Band intensities are much less variable with these newest dyes.
|Your chromatogram proceeds normally, but the bands become broad and low after only a couple hundred nucleotides (or 50 or 400...).
||The resolution of the gel normally decreases after perhaps 750-850 nucleotides, which is normal for a Model 3730 sequencer. You may be able to get good reads as much as 900 nt out, but only rarely and then only with exceptionally clean template.
If the peak resolution decreases substantially earlier, however, see the next section.
|The technician reported "loss of resolution after [nnn] nucleotides".
||You probably have a contaminant in your template that caused a loss of resolution in our capillary electrophoresis instruments. Please see the page describing the Loss of Resolution artifact for full diagnosis and suggestions.
A typical example of a "loss of resolution" artifact:
|Early on, the sequence is fine:
||Before long, problems are evident:
||Too soon, resolution is lost completely:
|The first 10-20 nucleotides are obscured by huge, trashy-looking peaks, then normal sequence is seen thereafter.
||The most likely explanation is that your primer is formed self-dimers and the 'trash' peaks are from sequencing on itself. All primers should be designed using a computer, in order to avoid such artifacts. Most common primer design programs will avoid primers that form self-dimers. Please see Primer Design.
Alternatively, if your sample is a PCR product, these large peaks may arise from a small PCR product contaminating your main band. You wouldn't even see such a product on an agarose gel, if it is small enough. Cut your PCR product out of the gel to isolate a single band, and try again.
|The first 20-50 nucleotides are fine, but suddenly the chromatogram shows mixed peaks or terrible background.
||We often see this when the template DNA is actually a mixture of two clones that are identical up to the cloning site and diverge thereafter. To avoid this problem, you should always streak out your clones to single colonies to ensure they are completely clonal.
Alternatively, your primer could be sitting down on two independent sites within the construct, and generating identical sequence on those two sites up until the point where the two sequences diverge, whereupon you get the peaks-on-peaks effect. This is common when you're priming inside an insert and you've accidently inserted *two* copies of that insert. Other structural errors can produce this type of effect as well.
Here's an example of two mixed clones, identical in sequence until they hit the cloning site:
|The sequence looks great until it hits a polyA (or polyT), and then the bands rise and fall in waves.
||This is called "polymerase slip". It happens when the growing strand temporarily dissociates from the template, then reassociates at a different spot - say, one nucleotide forward or back from where it started. If this happens often enough (as it will on polyA or polyT templates), every individual band becomes a family of closely-spaced peaks giving a 'roller coaster' look to the chromatogram. Try sequencing in the other direction from the opposite strand, or try another primer either closer or further from the homopolymer region.
The following is an excellent example of 'polymerase slip' on a homopolymeric tract: