The quality of the DNA template is one of the most critical factors in automated DNA sequencing. Please make certain that the DNA template is at the proper concentration, free of contaminants, dissolved in distilled water and that the DNA is not degraded. There are several steps in producing good template:


Plasmids:  For plasmid preparations (both large-scale or mini-preps), numerous commercial protocols work in addition to traditional methods. It is essential to avoid genomic DNA, excess RNA and contaminants. Many of our customers use Qiagen products with good success, but other manufacturer’s products also work well. Elute in distilled water or in 1 mM Tris. Do not elute in TE, as the EDTA can interfere with sequencing. Cesium Chloride preps work but be certain there is no Cs or EDTA present in the final sample. An extra ethanol precipitation is helpful to eliminate these contaminants.

PCR Products:  For PCR products, there are two purification options. (1) If the PCR product is exceptionally pure, a commercial ultrafiltration product that separates the PCR products from the primers and enzymes can be used.  (2) Alternatively, the PCR products can be electrophoresed and the desired band can be excised and gel-eluted. Please note that the PCR products must be very pure in order to obtain clean sequence. Extraneous bands may appear low-intensity, but could easily contaminate the sequencing run. Also it is essential to remove all traces of the original PCR primers, as these could produce undesired sequencing products by acting as sequencing primers. (Recall that PCR is a bi-directional exponential reaction, whereas sequencing is a uni-directional linear reaction)

Large Clones (BAC, Cosmid, etc…):  BAC (and other) preps are commonly contaminated with 20%-80% genomic DNA. This excess DNA decreases the effective concentration of the template, so genomic DNA must be avoided. It is imperative that during the alkaline lysis step gentle handling of the samples is maintained. Additionally, during the potassium acetate precipitation step to remove protein/genomic DNA, extremely gentle handling is required. A second phenol/chloroform step is also helpful.  Perform a restriction digest of the DNA and examine the bands on a gel. If any smear in the background is visible, there is too much genomic DNA in the prep. The best sequence has been routinely generated from BACs prepared using Qiagen protocols (Qiatip 500, for example) and with an added phenol-chloroform extraction at the end which has routinely helped “bad” BAC preps.


Provided there is ample template DNA, it is best to determine the concentration by UV absorption. Often a mini-prep or PCR product cannot be reliably quantitated using a spectrophotometer. For success getting accurate spec readings on DNA, please consider the following points:

Readings below 0.05 AU require careful blanking in order to ensure accuracy and many specs are unreliable below 0.05 AU.

Contaminants such as RNA and free nucleotides also absorb UV, affecting the AU value.

Avoid chromosomal/genomic DNA contamination. Upon restriction digestion, genomic DNA contamination is visible as a smear in the background of the lane on a given gel. Any visible smear represents a large percentage of the total DNA present, thus resulting in an artificially inflated UV reading with respect to the template to be sequenced.

If there is not enough DNA to measure spectrophotometrically, then estimate the amount of DNA from a gel by comparing it to known standards. It is good practice to do this even in addition to a spec measurement to verify the result.  Further, if the DNA concentration is unable to be accurately determined, good laboratory practice dictates not proceeding until there is enough DNA to measure accurately.


After successfully determining the template DNA concentration, dilute the template to the final concentration using distilled water. Do not use TE or other EDTA-containing buffers and do not add any divalent cations (i.e. Mg, Ca, Mn) or salts.

Plasmids: Dilute to 50-200ng/ul.  100ng/ul is ideal.

PCR products:  For each 1000bp of PCR product length use 40ng/reaction (40ng/kb). For example, if the PCR product is 1500bp in length, then 60ng will be required for a successful reaction. A concentration of 20-40ng/ul is best.

Large Clones (BAC): For large clones, like BAC clones, large amounts of non-template DNA are present. For this reason, the template concentration must be very high since only a small percentage of the clone will be used as sequencing template. To address this issue, the total concentration of DNA must be increased. The best success is achieved with 0.5-1.0ug/ul for this size clones. BAC samples should be visibly viscous and difficult to pipette at this concentration. The excess DNA in the reaction will bind primer non-specifically, so the primer concentration must be increased accordingly. We suggest 10uM as a good standard primer concentration. Our universal primers are automatically used at the higher concentration.


Special consideration must be given to primers and primer design. For PCR, two primers replicate opposite strands which has a significant effect on the results. Inefficient primers still work for PCR. Most PCR amplifications go beyond the linear part of the reaction, meaning inefficient primers will produce as much product as efficient ones. PCR will amplify the target DNA even if it is only a small proportion of the DNA present. Even if 99% of the plasmid prep is junk DNA, the remaining 1% will amplify just fine but will not sequence. For sequencing, this is not the case. Inefficient primers will produce only weak bands and a resulting poor signal.  The best sequencing primers are efficient at binding to the target site, have little to no secondary binding site issues, do not self-anneal (primer-dimer formation) or have internal hairpins or secondary structure problems, and have a reasonable G/C to A/T ratio.  Please be certain that the sequencing primers meet these minimal standards to ensure a positive sequencing result.


This is an example of a good sequencing reaction.  The peaks are evenly spaced, uniform and clean. The automatic base-caller will make few if any mistakes providing confidence that the sequence is correct.

good sequencing reaction.

This next section discusses trouble shooting examples and solutions of automated DNA sequencing.  When reactions are completed, an automatic email is generated.  This email contains general information about possible causes of poor results. When reactions are completed and results are available, an automatic email message is generated and sent to the designated recipient.  A follow up email from our technical staff is generated for problematic or difficult results and provides trouble shooting suggestions and advice.  A copy of the email follows:

We have already created an account for you so please contact us if you need your login information or have any other questions.

Please refer to our trouble shooting guide for help in interpreting your results.  You can request a copy of this trouble shooting guide if you do not have one.  Here are some of the guidelines for interpreting your results:

Possible reasons reactions gave weak/no signal:
Insufficient DNA template/primer.
Contaminated DNA template/primer.
No priming site for the primer.
Poor primer design.

Possible reasons reactions gave multiple sequences:
Slippage after a homopolymer region.
Heterogeneous DNA templates.
Multiple priming sites for the primer.
GC compression.
Repeated sequences in the DNA template.

Possible reasons reactions gave short sequences:
High G/C content.
Secondary structures.
Poor quantification of template and/or primer.

Here at Retrogen have developed some proprietary protocols that can resolve the problematic templates containing high GC content, dinucleotide repeats, homopolymer regions, and hairpin loops, like siRNA template. Please consult with our technical staff for more information about these services.

Thank you very much for your order.

The follow up email from our technical staff will address specific issue regarding the samples such as “weak signal”, “no signal”, “multiple signal” or “short signal”.  These results are as common as good sequencing results and most often reflect difficulties with the template or primer preparations and concentrations.  The remainder of this section breaks down the common trouble shooting issues that occur.


In the chromatogram below, the peaks are ill-defined and not much higher than background noise.  This is characterized as “weak signal” or “no signal”.


This is the view of one of our Controls.  The signal strengths are circled and the approximate average signal strength is 1440.  Reactions ideally have an average signal strength between 800-1500 although it is possible to have excellent looking sequence with signal strengths much higher or lower.  However, signals below 200 or above 4000 often have serious problems.


On a weak reaction the signal strength is very low (averaging 115).  When signals start to get down below 200 a good clean looking sequence is usually rare.

The most common poor result in sequencing is “weak” or “no” signal.  There are many explanations of this result.

As mentioned previously, DNA preparation is the first consideration.  Contaminated templates, low DNA concentrations, unclean samples and mixed samples can cause this result.  Also, the primer design and concentration weighs heavily in the success of the reaction.  Some examples and suggestions follow:

DNA is low concentration:

How was the DNA prepared? Mini-preps, PCR reactions and gel-eluted fragments

usually cannot be measured with a standard spectrophotometer. Unless you own a

microliter-scale spectrophotometer or  a newer “nano-drop” machine the amount of DNA is simply too low for a reliable reading. For larger scale preps (Midi, Maxi, Mega) the spectrophotometer should give you a reasonable concentration value. Was the DNA concentration estimated from a gel? When sequencing PCR reactions, it is

almost always critical 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. It is good practice to have someone with sufficient experience do the estimation. Please be sure to run the gel on the same tube of DNA you are sending us; gel elution of fragments does not result in 100% recovery.

Primer concentration/design problems:

There was insufficient primer in the primer tube.  Please double-check all calculations.

The primer concentrations are specified in pMol/ul – that’s picomoles per microliter not ‘picomolar’. There is six-orders-of-magnitude between them. We prefer a 10uM solution of primer in a clearly labeled tube.

The primer did not interact efficiently with the template. It is recommended that a

complete restriction map is generated.  This can easily explain cloning issues with regard  to the presence, absence or damage of the priming site.

Did you design the primer from accurate sequence information?  If a prior sequencing run was used to design the primer for this one, make sure the region used is from a stable and

reliable sequence. Common primer design software is available both commercially and academically.

Does the primer function at our annealing temperatures? Since we handle so many

samples, we must process them all at consensus cycling conditions.  We anneal at 50 C.


DNA sequencers based on capillary electrophoresis are sensitive to the presence of certain contaminants that may not affect your sample in any way other than sequencing – you may be able to PCR them, restrict them or even transfect them with no problems. We do not know the exact nature of these contaminants, but we know how they typically get into samples, and we know how to remove them.

A very typical example of a Loss of Resolution (LOR) sample:

Early in the run, we see reasonable resolution:


50 or 60 nt further on, it’s obvious something is wrong:


Another 120 nt, and the resolution is terrible and the sequence is essentially unreadable:


This loss of resolution is most often a result of a contaminant in the sequencing reaction.  Since we use a premixed solution of reagents that works on a control and other samples, the origin of the contaminant is limited to the DNA template sample or the primer sample.  Some successful suggestions for removing the unknown contaminant are:

Perform a phenol-chloroform extraction on your template DNA.

 Perform ammonium acetate-isopropyl precipitation on your template DNA.

Pass your sample through a Sephadex G50 spin column.

Mini-preps are the most common LOR samples, and many sequencing facilities have observed that overloaded Qiagen preps (or similar silica preps) are at fault. When overloaded, these plasmid prep kits leave many impurities in the DNA. To avoid overloading, try at least one of the following:

Grow your minipreps for a shorter time. 12 hours is good, 8 hours is better. Yes, the yield may be reduced, but the DNA sequence template will be better.

Grow smaller bacterial cultures. If the kit says no more than 5mls of lysate, limit yourself to 3 mls. Yes, the yield will be reduced. But again the DNA sequence template will be better.

Other short signals can appear in various ways:

Signals can drop of quickly:


There can be a very gradual signal loss (top heavy data)

At the start :


At 200bps:


At 400bps:


There can be an abrupt drop off:


Or there can be a drop off at GC rich region:


Make sure you inform Retrogen if any template is known to be GC rich so that we can use optimal conditions for these templates.  If these conditions still give short signals contact Retrogen to discuss further methods for improving results.

There are four known causes for the LOR problem. Three of the failure modes are within the sequencing facility itself, and are relatively uncommon. We watch for them all the time. The known causes of the LOR problem are as follows:

Air bubbles in the electrophoresis capillaries. If a capillary is partially blocked by an air bubble, the sample in that capillary will exhibit extremely poor resolution (if it elutes at all). This results in sporadic failures of individual samples out of sets that otherwise gave good results. Retrogen will repeat samples exhibiting sporadic LOR upon request.

Plugged cuvette flow ports.  The flow paths in the instrument can become blocked, in which case an entire set of samples will fail with loss-of-resolution – every sample, including standards. We have not seen this problem at Retrogen.

Overloaded lanes.    We have been told by the sequencer manufacturer that excessive amounts of sample (sequencing termination products) in a capillary can cause a loss of resolution much as we describe.

The last failure mode is very common, and is a characteristic of entire sets of samples. In other words, it is a sample prep problem.

Contaminated samples.   We have time and time again seen entire sets of samples produce LOR, when the rest of the samples in that set (including standards) look fine. Samples such as these, when re-run, always give the same LOR result. When the samples are cleaned up (suggestions noted above) they produce excellent results. We can only conclude that there is a contaminant in the samples that reduces resolution in our machines.

Below is an example (actually, a fairly mild example of a ski-slope):


This is not well understood, but here are four possible explanations:

(1) salt in the DNA

(2) too much DNA in the reaction

(3) an unknown impurity “poisoning” the Taq processivity

(4) an unknown contaminant increasing the binding of dyes in the enzyme’s active site.

The last effect can arise from free NTP’s in the sample or from a contaminant that disturbs the divalent cation concentration (EDTA, Mg++ etc). Buffers such as TE should not be used to resuspend DNA preparations that will be used for sequencing.

Salt is the most common cause of the ‘ski-slope’ effect. Capillary electrophoresis instruments such as our 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” affect 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 Retrogen technicians feel this is not possible, and if the “blob” appears to arise from our own processing problem, will initiate a no-charge repeat for you upon request.

Here’s a typical example of what we call a “dye blob”:


The 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. Is it an siRNA (RNAi) construct? These will almost always exhibit strong sec-structure effects. A couple possible solutions:

1) try resequencing by selecting ‘siRNA construct’ as your DNA type (best solution)

(2) try to sequence from another primer at a different position (closer or further)

(3) 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 offer additional charged services in this regard, but contact Retrogen to discuss these trouble shooting options.

Here’s an example of a secondary structure effect:



The first 10-20 nucleotides are obscured by huge, trashy-looking peaks, then normal sequence is seen thereafter.  The most likely explanation is that the primer has 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.

Alternatively, if the sample is a PCR product, these large peaks may arise from a small PCR product contaminating the main sample. These products are usually not seen on agarose gels since they are such a small amount of the PCR reaction, but the produce signal in a sequencing reaction which make the data difficult to interpret at best. Cut the 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.  When the template DNA is actually a mixture of two clones that are identical up to the cloning site and diverge thereafter, multiple sequences are generated. To avoid this problem, always streak out the clones to single colonies to ensure they are completely clonal.

Alternatively, the 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 priming inside an insert and there have been two copies of that insert accidentally inserted. 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” or “slippage”. It happens when the growing strand temporarily dissociates from the template, then re-associates at a different spot, usually one nucleotide forward or back from where it started. If this happens often enough (it will on poly-A or poly-T 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.

Example of homopolymeric (G-rich) region causing multiple signals:


The following is an excellent example of ‘polymerase slip’ on a homopolymeric tract:


The bands are present and strong, but irregularly spaced, or with mixed colors.  This results when two sequences have been superimposed on each other. There are several common causes:

(1) The sequencing primer binds to two (or more) sites on the template.

(2) There are two (or more) templates present.

(3) This was a PCR reaction, and the original primers were not completely removed.

(4) This was a PCR reaction, and one primer generated both ends of the product.

(5) This was a PCR reaction, and there is more than one amplified species present.

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:



At Retrogen we examine virtually every lane from every sample we sequence. We diagnose the problems whenever we can, and some problems just keep coming up again and again. Our sequence analysts spend considerable time with clients who have had problems and carefully examine their experimental design and their protocols for the cause of the failure. The following table is a compilation of the most common failure modes seen in Retrogen:

Inadequate template concentration:

trying to spec a miniprep plasmid or a PCR product

spec readings too low to be accurate

inexperience at gel estimation of band intensity

failure to check PCR product concentration

spec reading with excessive RNA present

spec reading with excessive genomic DNA present

Overloaded miniprep purification devices (“Loss of Resolution” samples)

Failure to streak clones to single-colony

Calculation error in primer concentration

Multiple priming sites present

Multiple inserts present (gives multiple priming or stem-loop)

Sequencing incorrect PCR products

Failure to fully characterize a new plasmid construct

Secondary structure in clone (siRNA constructs, etc)

Homopolymer tract (e.g. 3′ sequencing on a cDNA clone)

Salt contamination in sample (“ski-slope” effect; typically gel-eluted frags)

Mis-paired primers (cross-species primer design, inaccurate sequence basis)

Free dNTPs present in template

Primer dimers

Primer Tm too low

Mixed-up samples and/or primers


Here at Retrogen we strive to provide a clear and long read from the templates submitted.  We have spent significant effort to make our services streamline and successful for the vast majority of templates available.  It is true that not all samples fit into this generalized criteria and we are dedicated to resolving such issues with individual clients.  Through individual follow-up and customer service, we are able to resolve nearly all sequencing issues and generate quality sequence data.  This is not to say that every single sample from every customer results in perfection, but we strive to achieve that standard.

If this manual has not satisfied or answered the trouble shooting requirements from the sample results, please call or email us at and one of our sequencing analysts will discuss the results, experimental design and provide suggestions for getting those positive results.