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Epicentre Forum 3 (2)
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DNA sequencing is a powerful technique with applications in many scientific
fields. Here we discuss some general sequencing considerations as well
as potential problems that may be encountered while performing DNA sequencing.
We describe possible solutions to these problems that Epicentre scientists
have found useful in the course of their work.
What is the best way to purify plasmids for DNA sequencing?
Plasmid templates for use in DNA sequencing may be isolated by a number
of different methods from a standard alkaline lysis preparation to any
number of commercially available columns and matrices. The choice will
depend on cost and convenience to the user. Regardless of the template
isolation procedure used, for successful DNA sequencing, the resulting
template must be free of large amounts of residual detergent, salt, polyethylene
glycol, and ethanol. These compounds can inhibit DNA polymerases. Avoidance
of "boiling detergent" preps, and careful washing and drying
of DNA pellets should yield template that produces good quality sequence
data.
What about PCR product templates?
Both residual amplification primers and unincorporated nucleotides must
to be removed from PCR product preparations prior to sequencing. Residual
primers can produce spurious ladders when using internal labeling protocols
or compete with the sequencing primer for extension in primer labeling
protocols. Unincorporated nucleotides will alter the dNTP/ddNTP ratios
of the termination mixes. Cleanup can be achieved in a variety of ways
including size exclusion spin columns and membranes or by agarose gel
purification. Minimizing exposure of the PCR product to less than 30
seconds of UV light (regardless of the wavelength) will also improve
the quality of the sequence data.
What is the best labeling method to use?
The choice of labeling method employed will depend on the labels, equipment,
and facilities available, as well as the needs and preferences of the
user. The choices of end labeling vs. internal labeling, radiolabeling
vs. non- radiolabeling, manual vs. automated sequencing, and the use
of 32P, 33P, or 35S radiolabels must
be considered. When cycle sequencing, regardless of the detection method
used, primer end labeling protocols almost always yield better data than
comparable internal labeling protocols.
What is the best cycling profile to use for cycle sequencing?
In most cases, the data produced from a 2-step cycling profile (lacking
an annealing step) will be better than data produced from a comparable
3-step cycling profile (including an annealing step). The goal is to
make primer annealing as stringent and specific as possible to avoid
secondary primer annealing events that could produce significant background
signals. To determine if an annealing step is necessary, calculate the
annealing temperature of the sequencing primer and compare it to the
extension temperature to be used. If the annealing temperature is 5°C
below the extension temperature or higher, then a 2-step cycling profile
can be used. Cycling for more than the recommended number of cycles (30)
can result in an increase in "shadow bands" and higher background.
Cycling for less than the recommended number of cycles may improve sequence
quality for some primer/template pairs.
What sequencing gel composition produces the best results?
We have found that using Long-Ranger™ (FMC) modified acrylamide
gels following the manufacturer's directions gives good results. These
gels show excellent resolution and are much more resistant to tearing.
Do I have to run the sequencing gel the same day as it was poured?
Sequencing gels may be poured and run the same day, or stored and run
the following day. When running the gel the same day as it was poured,
make sure the polymerization reaction has gone to completion before using.
This usually takes at least 2 hours and depends on the amount of TEMED
and ammonium persulfate used. Incompletely polymerized gels will yield
fuzzy bands or wide peaks. If running the gel the day after it was poured,
be sure that the top and bottom gel edges remain hydrated during overnight
storage by wrapping a wet paper towel around the top and bottom of the
gel and then wrapping the wet towel in plastic wrap. Dehydrated gel edges
will make it difficult for the sample to enter the gel and can inhibit
proper migration through the gel.
What can be done to make loading of cycle sequencing reactions
containing mineral oil easier?
When loading 0.4 mm gels, remove your sample by inserting the micropipette
tip through the oil phase into the aqueous layer. When a dimpling of
the aqueous layer is seen, the pipette tip has entered the aqueous phase.
Gently compress the plunger, expelling the air from the pipette tip,
and remove the proper volume of reaction product for loading. When loading,
gently expel the reaction product into the well being careful to stop
compressing the plunger as soon as the last of the reaction product has
left the pipette tip. Do not fully compress the plunger since this may
produce an air bubble which will rapidly rise out of the gel and could
carry reaction product with it.
When loading gels thinner than 0.4 mm, first pipet an aliquot of each
reaction sample onto a piece of Parafilm.® The oil will be attracted
to the Parafilm leaving a relatively oil-free aqueous drop in the center
of the sample. Load the samples from these drops as described above.
Is there anything I can do about leaky gel wells?
If the sequencing apparatus allows, placing a clamping device (ie.,
a "paper-binding" clamp or a "three-finger test tube" clamp)
on the apparatus such that the two glass plates are clamped will alleviate
the leakage. After samples have been run into the gel, the clamp(s) can
be moved around or removed as desired. Caution: Turn off the power supply
before touching the metal clamps.
Sometimes I have difficulty loading multiple samples. What is
the problem?
Hard to load wells can be caused by urea that has diffused out of the
gel into the wells. Samples loaded into a urea-containing well will not
settle properly to the bottom of the well resulting in the dye front
and DNA bands running as a dot(s) rather than as a band(s). Because of
this, it is important to remove the urea from each well just prior to
loading. We use upper reservoir buffer forced through a syringe and needle.
Urea can diffuse back into wells quickly, particularly if some samples
are run into the gel before loading subsequent samples. In this case,
clean out the wells again prior to loading the remaining samples.
How do I prevent problems caused by glycerol present in the samples?
Glycerol present in the reaction samples can cause electrophoresis anomalies
when a borate-based buffering system is used in the sequencing gel. While "glycerol-tolerant" buffers
are available, we have found that simply rinsing out the wells extensively
just after the samples have completely entered the gel alleviates much
of this problem.
What can I do if the gel sticks to both plates when the plates
are pried apart?
The gentle application of a water stream from a squirt bottle will dislodge
the gel from the upper plate. Do not lift the upper plate too quickly;
let the water and gravity do the work.
What is the best way to fix a sequencing gel?
We routinely use a 20% ethanol solution as a fixative rather than the
more common methanol/glacial acetic acid mix. We observe no difference
in data quality from this procedure and the ethanol solution is less
hazardous.
What can be done if bubbles or wrinkles are formed upon transfer
of the gel to drying paper?
To remove either problem, first completely wet the back of the drying
paper. Remove bubbles by gently pushing them off to the side of the gel
with a wet rubber- gloved finger. Apply only enough downward pressure
to just barely touch the gel but not enough to compress it. Difficult
to remove bubbles can be pierced with a needle. Remove wrinkles by filling
the wrinkle with water either from a squirt bottle or a needle and syringe,
and then gently work out as described for bubbles. If none of these suggestions
work, refloat the gel back off of the paper (onto a glass plate in the
fixative) and try again.
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