• The email comments reported "no bands" or "weak bands". Possible explanations:

• There was no DNA in your tube (or far less DNA than necessary).

  Please note: A weak lane, when repeated, will often be blank, and vice-versa. The two outcomes (weak and blank) are often interchangeable, and which you get is a matter of luck, depending on how noise-free is the lane to which your sample was assigned.

  Did you quantitate the DNA using a spectrophotometer?
  

  Make sure your reading was at least 0.1 AU (or as low as 0.05 might be acceptable if the
  spec was carefully calibrated).

  Warning!! Mini-preps, PCR reactions and gel-eluted fragments usually CANNOT be measured
  with a spectrophotometer unless you own a microliter-scale spec. The amount of DNA is
  simply too low for a reliable reading. For large minipreps, you MAY be able to use a
  spec with a 100 ul cuvette, but make sure you aren't reading through the meniscus! For
  example calculations, see Why can't I use a Spec?'.

  Did you estimate the concentration from a gel?

  If you are sequencing PCR reactions, you will almost always have to use an analytical gel
  to estimate DNA concentration. Gel elution of restriction fragments often must be measured
  the same way.

  Estimation by gel is difficult. If you are not experienced at this, find someone who is more
  experienced to help you. You should run the gel on the *same* tube of DNA you are sending us
  ... i.e. don't gel elute the band and assume 100% recovery.

  Double-check all calculations!!




• There was no primer in your primer tube.

  Double-check all calculations!!! The primer concentrations are specified in pMol/ul - that's
  picomoles per microliter not 'picomolar'. There is a six-orders-of-magnitude difference between
  the two!




• The primer did not interact efficiently with the template.

  Here are some possible considerations:
  • Are you SURE of the identity of your plasmid?
    A very common error is to submit an erroneous plasmid due to a cloning mistake,
    rearranged plasmid or damaged priming site. Another common problem is if another
    plasmid is present which doesn't prime, effectively diluting the specific plasmid.
    I recommend you produce a COMPLETE restriction map, looking for unexplained
    bands or unexpected band sizes. Don't just cut out the insert or PCR a fragment as
    verification of your construct!

  • Did you design the primer from accurate sequence information?
    

    If you used a prior sequencing run to design the primer for this one, make sure you
    weren't reading sequence from a poor area of a gel when you designed this primer. Ask
    the Sequencing Core Director to help if you're not sure the source sequence is valid.

  • Are you sure the priming site is present in *this* template?
    

    For example, make sure you didn't use a sequence from Mus musculus to design a primer
    for a Mus castenius sequencing project.

  • Did you design the primer to function at *our* annealing temperatures?
    

    Because we handle so many samples, we must process them all at consensus cycling conditions.
    We anneal at 50 degrees C. Please see our web page on How To Design Sequencing Primers.

  • Might the priming site have been accidently damaged?
    

    Occasionally, cloning artifacts can create a deletion near the insertion site, or a
    deletion that removes the priming site. Careful restriction mapping can detect the latter,
    and using an alternative primer will usually work in the former situation.

  • Was this template a PCR product? Did it amplify correctly?
    

    Never assume you've got the right PCR product - always test it. If you know there's an
    internal restriction site, cut a portion of the product and check it on a gel. Note that
    sometimes you'll get seemingly-beautiful amplification with just *one* primer acting at
    both ends of an illegitimate site. In sequencing, that primer will generate only multiple,
    superimposed sequences. The other PCR primer may not have participated at all, and in
    sequencing will produce only a blank lane.

• The lane was blank THIS time, but the exact same sample worked fine before!

• Your sample may have been weak before when you say it "worked".
  

  A weak lane, when repeated, will often be blank, and vice-versa. The two outcomes (weak and blank)
  are often interchangeable, and which you get is a matter of luck, depending on how noise-free is
  the lane to which your sample was assigned. Check the signal strength of the previous "good" lane
  and if it was below G=150, you probably just got lucky before.

• The lane was not blank, but was uniformly poor.

• Please consult the following table.

The technician said my sample was weak, but another time it worked perfectly! How can that be?	Your sample probably produced only weak bands before, too, but you were luckier. Please see the next section.
There are bands, but they are weak and noisy, or even uninterpretable.	Check your chromatogram for text like: "Signal G:523 A:428 T:617 C:530". On a printed chromatogram, it's near the top. For electronic chromatogram-viewing programs, look for an 'Info' window. These signal- strength numbers are an indication of the relative fluorescent strength of the bands in your lane. We typically examine just the G signal to simplify comparisons. Good samples will have a G signal of 1000-2000. If your signal strength is below 300, background bands that are normally too low to see will become very evident and will interfere with basecalling. If your signal is below about 80, your peaks may get lost in the background noise, and we will report only a blank lane. See the above section, regarding reports of "no bands". The possible causes are almost the same as those for weak bands. You might get lucky sometimes. Baseline noise varies from lane to lane, run to run, and instrument- to-instrument. If you happen to get a low-noise situation, even a sample with a G signal of 50 will give great sequence. That does NOT mean the sample is OK! The next time, that exact same sample will bomb, because the noise was too high.
There apparently were bands, but the technician reported "poor resolution from the start".	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 diagnosis and suggestions.
The bands are present and strong, but irregularly spaced, or with mixed colors. The technician may have reported "superimposed sequences" or used the phrase "peaks on peaks".	If you see this, you usually have two sequences superimposed on each other. There are several common causes: The sequencing primer binds to two (or more) sites on the template. There are two (or more) templates present. This was a PCR reaction, and you didn't remove the original primers. This was a PCR reaction, and one primer generated *both* ends. This was a PCR reaction, and there is more than one amplified species present. A similar outcome is often seen in which the bands start out fine, but later on become superimposed. This is described further down this page. Here's an example of 'mixed peaks' such as might arise from two or more unrelated templates: Another example, this time with templates that might be related. Note the alignment of the peaks:

• The sequence looks OK in some spots, but not others.

  • Please consult the following table:

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 'siRNA construct' 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:

Please contact the Core Director, Dr. Robert Lyons with any questions you may have.