There are many guides to sequencing available on the
internet and through manufacturers such as Applied Biosystems (maker of BigDye). In an
effort to collate what we feel are some of the more important factors, both in
our own direct experience and through conversations with facility users, we
decided to put our own recommendations and insights into one place. Much of this material is available in other documents
on the DNA Blackbox (http://dna.biotech.wisc.edu; scroll down to "Helpful Documents."), but this document
attempts to combine it all. It is a mix
of procedural information and troubleshooting hints, and is based on examining
each of the components of the sequencing procedure. While sample chromatograms are not present,
an attempt has been made to describe certain diagnostic features of the trace
file. A good visual source of such files
can be viewed at http://Cancer-seqbase.uchicago.edu/traces.html or http://www.roswellpark.org/document_3640_639.html
There are several components in a sequencing
reaction—template, primer, sequencing reagent, and “other additives.” The mix of these reagents then undergoes 3
steps—cycle sequencing, post sequencing cleanup, and analysis. By examining each of these processes,
sometimes in conjunction with controls we provide, we can get you sequencing
successfully right away, or, if you’re having difficulty, figure out why you’re
not getting the data you desire.
Template is a very common problem area. If your chromatogram is blank, has very low
signal, or starts well but gradually dies out, the template should be
examined. For plasmids in the 3-10 kb
range, 0.2 ug is a good amount of DNA to use. More is almost never better. One problem is that accurate quantification
of DNA is not easy. Spectrophotometer
readings will invariably overestimate the amount of template DNA, unless you
have CsCl banded material. That’s because use of almost all miniprep kits on the market will result in some RNA, chromosomal
DNA, and other fluorescent cellular material coming through the purification
procedure that will absorb UV light.
These other chemicals won’t necessarily inhibit
the sequencing reaction, but they will contribute to the A260 reading. As a result, relying on this spec reading
alone will cause you to add less than DNA than you think (midi and maxi prep
kits do a better job, giving a higher plasmid:contaminant ratio).
It’s informative to run your template on an agarose gel, using some
standard of known concentration and estimate the relative fluorescent intensity
following gel staining. We can provide
you with a pGEM standard at 0.2 ug/ul to run with your sample. If you are doing many preps and don’t always
feel like quantifying your DNA, a reasonable rule of thumb for minipreps of a high copy number plasmid, following lysis of 2-3 mls of a dense
overnight culture, is to use 1/10th volume of the eluted sample in
the sequencing reaction (but try this first before doing several dozen).
We distribute primers that recognize the Amp or
The problem of sequence starting out strong then rapidly
dying out is common and is usually attributed to "impurities" in the
prep. We have found that these
impurities can't always be removed by EtOH
precipitation and you may need to re-prep the plasmid.
Larger and/or lower copy number plasmids can be difficult to
sequence. It is tempting to lyse a greater volume of culture for the miniprep column, but a better strategy is to stay with 2-3 mls culture/miniprep column and
do multiple columns (or do a midi or maxiprep). The separate preparations can then be pooled;
it may even be necessary to do an additional cleanup and concentration step
such as ethanol precipitation. Also,
keep in mind that larger templates often require higher amounts of DNA (0.5-1.0
ug) to maintain a good molar ratio of template:primer and reagents. It is better to add a small volume of a
concentrated template; adding 10 ul of template to a 20 ul sequencing reaction
is most likely not going to give you good data.
BACs, PI clones, and lambda phage present their own difficulties,
and it may be necessary to investigate kits from several manufacturers to get
good template. Our experience preparing
such templates is limited, and we don’t have specific product recommendations
(though we do have a BAC prep protocol used successfully by a user years ago on
our web site). Most companies are willing
to provide free samples so you can test out their products by giving them or
their sales rep a call. Since these are
also high MW templates you will need to add more DNA to the sequence reaction,
remembering that small volumes of a concentrated prep generally give better
results. See more info at http://dna.biotech.wisc.edu/documents/BAC_Sequencing_Information.htm
.
PCR fragments seem to sequence either incredibly well or not at all. PCR fragments generally must be cleaned up
following the initial amplification and prior to sequencing to remove the two
amplifying primers and the unused nucleotides (some mol bio jocks have their
reactions fine tuned to the point that following amplification there are no
primers or nucleotides left, and they can just add the product directly into a
sequence reaction, but this is not common).
Common ways to clean up the PCR fragment are listed below, with
advantages and disadvantages for each:
1. Cutting the band out of gel and doing a gel
extraction using any one of a number of kits available. We only recommend this if multiple fragments
are seen following amplification, and you need to cut out the fragment that’s
the “right” size. Disadvantages of thi
2. Performing a kit based column cleanup
protocol. This typically involves
applying the entire PCR to a column, then carrying out washing and elution
steps to generate fragment free of contaminating primers and nucleotides. Thi
3. Enzymatic cleanup of the PCR fragment. Commercially available shrimp alkaline phosphatase and exonuclease
(SAP-EXO) are used to inactivate exces
4. Magnetic bead clean-up—“AMPure”
from Agencourt.
This method is similar to bead cleanups following sequencing (see
below). PCR fragment is bound to
magnetic beads, rinsed, then eluted using a special
magnetic plate (again, the same one used for dye terminator removal). We don’t have a lot of experience with this,
but have heard favorable comments from users who have tried it. It would seem to have all the convenience
advantages of the other magnetic bead cleanup method, including relatively low
cost (it is included on our same UW quote as the dye terminator removal beads)
and ease of sample handling in multiwell format.
Official protocols for sequencing
PCR fragments recommend using 10 ng of PCR
fragment per 100 bp of fragment size. However, it is
usually a pain to attempt quantification of PCR fragments, and it often works
fine to empirically determine how much to add based on the intensity of the
fragment on an agarose gel. PCR fragments
can sequence very well, probably due to the high molar ratio of template:primer and the efficient denaturation of a fragment compared to a plasmid. An amount of DNA that is very bright on a gel
will often sequence too well and resolution of the chromatogram will
suffer. However, if you see any product
at all, even if faint, you will probably get decent data from it in a
sequencing reaction (IF it’s the right fragment). Amplifying primers work well to sequence PCR
fragments, as do internal primers.
Additional discussion of sequencing PCR fragments i
Template composition is a factor that can cause sequencing
problems. GC rich templates will generally sequence fine, unless there
are particular regions of very strong secondary structure. A diagnostic trace pattern for this sort of
template would be a chromatogram that looks great to a certain point, then suddenly dies out.
Methods for dealing with this are discussed below, but basically these
involve the addition of denaturants to the sequence reaction or specially
formulated versions of BigDye. Poly A/ Poly T regions will often
cause difficulty—the chromatogram will look fine up to the polyT
(A) stretch, then either be very noisy peaks under peaks, or just long “rolling
hills” of the four chromatogram colors.
This results from “polymerase slippage” on the poly T(A)
region and is a difficult issue to resolve.
The addition of “reaction enhancers” (see below) may help if the problem
is not severe. Another strategy is to
design “anchored primers,” a poly A or poly T sequence with the final 3’ base
being either G, C, or T (or G, C, and A for a poly T primer). With these primers the poly A/T stretch needs
to be at least 17 bases long, and special conditions for annealing (42°) and
cycling (52°) are recommended. Sometimes
this works well, but it may require the user experimenting with different
conditions. We do have anchored primer mixes available free of charge if you
want to try this. Remember you will lose
information just downstream of the primer, as is typical in sequencing.
Finally, di- and tri-nucleotide repeats
can cause poor data. It's usually pretty
obvious when such sequences are at fault.
Often the use of sequencing reagent enhancers (discussed below) helps a
lot.
Primer problems can also be a
cause sequencing failures, and some of these give characteristic chromatogram
patterns. A blank chromatogram, which is
not that diagnostic of a primer problem specifically, can result from use of
the wrong primer, too low a concentration of primer, or simply bad or degraded
primer. However, assuming you’ve used
one of our control Amp or
http://alces.med.umn.edu/rawprimer.html: Primer design
http://www-genome.wi.mit.edu/cgi-bin/primer/primer3.cgi: Primer design
http://genome-www2.stanford.edu/cgi-bin/SGD/web-primer: Primer design (best one for
sequencing primers)
For information
on MW, and oligonucleotide conversions, check out
http://www.basic.nwu.edu/biotools/oligocalc.html: which we use routinely.
Generally it is tough to design
a primer that doesn’t work. However, it
is easy to miscalculate how much you’re adding so it always is worthwhile
re-checking your calculations. This may
involve re-quantifying your primer by spectrophotometry. A basic rule of thumb is that you should use
about the same amount of primer in a sequencing reaction as you do in a PCR
(5-10 pmol, corresponding to 30-60 ng of an 18 mer). If everything has been examined and by all
accounts a primer should work, you might just have a bad primer. We can examine any primer by mass
spectrometry (for a fee) and tell you if it appears good, i.e., full length, or
not. You might want to talk to the
people who made it to find out about getting a replacement. Our DNA Synthesis facility will generally
re-synthesize any primer made here that should work in a sequencing reaction
but doesn’t. Of course this is based on our analysis of the primer and previous
controls that have been carried out by you.
A “noisy” chromatogram—good
signal strength, but peaks under peaks resulting in numerous ambiguities
(“N’s)—can indicate several primer difficulties such as more than one primer
present, more than one primer binding site present, secondary priming at a
related sequence, or degraded primer.
[Thi
It is also possible to get
secondary primer binding if the primer is very GC rich and is being used to
sequence a GC rich template. In these
cases, you can try raising the annealing temperature and adding a denaturant
such as DMSO (to 5% final concentration), formamide (5%), or betaine (1M final concentration) to increase primer:template specificity. Secondary priming giving peaks under peaks,
or too strong a signal resulting in peaks under peaks, can also occur if the
primer concentration is too high in PCR fragment sequencing. In contrast to sequencing plasmids, where too
much primer shows little effect, adding too much primer to a
PCR fragment sequencing will frequently result in noisy data. It is important to limit primer amount to 5 pmol when sequencing PCR fragments.
Finally, if your primer is
starting to degrade, or if there is a high proportion of n-1 products in your oligonucleotide preparation, you will also see peaks under
peaks since you’re essentially adding multiple primers to the reaction. Primers can last a long time, but they can
also degrade and it’s impossible to set an expiration date that covers all
primers. As noted above, we can analyze
your primer by mass spec as a fee for service, but in the interests of time it
may be worth just having a new one synthesized.
In several years of facility
operation, we have not seen a clear cut demonstration of the ABI BigDye reagent being a culprit in sequencing
difficulties. By “clear cut” we mean a
situation where newly acquired reagent worked, but the existing reagent in the
lab did not in a parallel experiment.
Should any lot of BigDye show difficulties, it
would soon be apparent in many labs, including our own, and we would attempt to
notify all users right away. Like any enzymatic mix, however, BigDye should be handled with care (keep in an ice bucket
during use, don’t leave on benchtop overnight,
etc.). Repeated freeze thaw cycles
should be avoided, though in pilot experiments carried out in the lab, five
freeze thaw cycles led to a barely noticeable decrease in activity. Depending on lab usage, it may be
advantageous to re-aliquot what you get from us to minimize this as an
issue. The mix stores well at –20°C,
though for long term storage (>3 months or so) it can be kept at –80°C.
As stated in our protocol
sheets, the amount of reagent to add to your reaction can vary. The “official” reaction recommended by ABI
uses 8 ul of reagent in a 20 ul reaction, but even they acknowledge 4 ul works
just as well. We have found that 2ul in
a 20 ul reaction also works for most templates, and our standard conditions use
that amount (see http://dna.biotech.wisc.edu/documents/Facility_Procedures.htm). We recommend experimenting with this in your
system so you can get the best data for the least cost. If you decide to try doing parallel reactions
with varying amounts of enzyme, we can give you an “R and D” price break on
those samples. If you're doing a large #
of samples and cost is critical, you might also check out the document "BigDye Dilution Experiment" at DNA blackbox
that pushes the amounts of enzyme pretty low while still getting decent results
(http://dna.biotech.wisc.edu/documents/BigDyeDilution.htm).
There is a specialized version
of the BigDye sequencing reagent that contains dGTP instead of the dITP present
in the normal formulation. This is used
specifically for templates with problem regions of secondary structure. Such regions are manifested by a chromatogram
that looks great up to a certain point, then falls off
precipitously. If you are seeing this
sort of pattern, we can provide you with an aliquot of the dGTP
mix to try. It typically works best in a
mixture with the standard BigDye, at a ratio of 3:1
or 2:2 normal:dGTP (it
varies from template to template), and should also be used in conjuction with 1M Betaine and
elevated extension temperature (68 or 72°).
It may take a few attempts to get good data using this reagent. A problems that can
occur with the use of dGTP is that “compressions” are
evident a
We
provide you with the sequencing buffer shipped by ABI with every enzyme
order. Older versions used a 2.5X
dilution buffer comprised of 200 mM Tri
Other
non-commercial reagents can work well in getting refractory templates to
sequence better. DMSO is a favorite
additive of sequencers across the country.
We go back and forth here adding it.
One issue is we have seen it go bad and actually inhibit the reactions,
so you'll want to keep your stocks fresh. But it can help, and it's cheap.
Stick with 1 ul/ 20 ul reaction (final concentration of 5%). Also, people have reported the addition of
formamide to a final concentration of 5% can help when secondary structure is a
problem (a diagnostic chromatogram for secondary structure in a template is
good looking data that abruptly terminates)--we have no data on this. As mentioned above in the dGTP
secondary structure discussion, a chemical called betaine
can enhance sequencing of these abruptly terminating clones, and can also help
other template related problems, including nucleotide repeats. Betaine is used by
making up a 5M stock, then adding it to a final concentration of 1-2 M (we
typically use 1M though some protocols call for more) in your sequencing
reaction. It's also commercially
available in mol boil grade at 5M concentration from Sigma. Single strand binding protein (SSBP)
available from Promega has been reported to help sequencing of refractory
secondary structure templates, used at a concentration of 1 ug/20 ul
reaction. We have no direct data with this
additive.
In the "Helpful
Documents" section of our web site is an older document entitled “Cycle
sequencing sample protocols” that covers a number of sequencing protocols
successfully used by us over the years. However, over time we've narrowed our
manipulations to a couple of standard cycling protocols. Different protocols seem to work better with
different primer/template concentrations, but we haven't developed definitive
rules. We do give you a set of
recommended conditions to start that has been broadly successful in our
hands. If you get sub-par data with
these conditions you may see an improvement using one of the other protocols. These different conditions are found at
"Facility Procedures" (http://dna.biotech.wisc.edu/documents/Facility_Procedures.htm). However, an important point to note is that
there are a variety of conditions that will work, so if you're having trouble
with your sequencing it might be worth checking other factors first before
playing too much with cycling conditions (check out the document
"Available Controls" at http://dna.biotech.wisc.edu/documents/Available_Controls.htm). Our recommended starting
conditions are as follows:
96° 3’ hot start, then 35 cycles
of 96° 10” 50° 15" 60° 3' followed by one cycle of 72° 7’.
Notes on these conditions: the hot start is PCR holdover, and i
35 cycles may be slight overkill
too, but we do it to maximize signal. In
our facility PCR machine space is not at a premium so it’s usually not a big
deal to run the extra cycles. However,
if space is restricted in your lab, in the interest of lab harmony you may want
to experiment with 25 cycles.
Experiments altering the
annealing temperature have not yielded clearcut
results. As an intuitive rule of thumb,
it'
Finally, the 72° 7’ is another
PCR holdover that should be considered optional. We will often pull reactions out of the
machine by the time they have gone > 30 cycles in the interest of getting
data to our users, and these reactions that have bypassed the final extension
look fine (or if they don’t, some other problem is responsible).
Following cycle sequencing,
reactions must be cleaned up to remove excess dye terminators which don’t
incorporate into DNA. Ideally, all the
excess terminators will be removed and all the extension products will
remain. This can be a common source of
problems for individuals, either in terms of getting too many unincorporated
nucleotides in the sample, or in sample loss during cleanup resulting in lower
signal. Low signal is not necessarily a huge problem because the
instrumentation is sensitive enough to extract useful information. It can mean
that if you are on the threshold of observable signal, any slight decrease will
lead to no data at all—there is less “wiggle room.” Another issue is that low signal generally
doesn’t look good out past 600 bp or so. From the capillary you should ideally get
about 800 bp of readable sequence, and this usually depends
on starting out with strong signal (because of the way the capillary
instruments operate, signal strength will usually fall over the course of the
sequence). Another diagnostic sign of
cleanup problems is the so-called “dye blobs.”
In the capillary they can be seen at different places in the
chromatogram--sometimes early, at about 70, and also at about 300 bp (check out for examples http://Cancer-seqbase.uchicago.edu/traces.html or http://www.roswellpark.org/document_3640_639.html). They appear as off-scale, or very tall,
broad, peaks stretching over what may be other legible peaks below. All colors are sometimes seen, but often they
are comprised of just red (the latter resulting from a breakdown product of the
T terminators). These peaks confuse the basecalling software, and lead to inaccurate sequence
data. Often the true peaks are seen
below and the sequence can be manually edited.
However, depending on the signal strength of the other bases, and the
severity of the dye blobs, all the T-signal can get “sucked up” by the dye
blobs and the chromatogram will appear to have no other T’s (this is a function
of the way the software works). Ideally,
no dye blobs beyond small ones appearing early in the run should be seen.
The
three most commonly used methods of dye terminator removal are size exclusion
(i.e., Sephadex G-50 or G-75, or commercially
available) columns, alcohol (isopropanol or ethanol)
precipitation, and magnetic bead adsorption/elution. Protocols for these techniques are available
at DNA blackbox in the "Helpful Documents"
section (http://dna.biotech.wisc.edu/documents/Non-bead_cleanups.htm). All procedures have been successfully
utilized by our users and by us.
However, in our experience the most reproducible and foolproof method is
magnetic bead cleanup. The
“Finally!”
you’re saying, “something that’s not my fault!”
It’s true, once you drop off your sample it’s up to us to get you back
data that looks good, if possible.
Fortunately most of the problems that occur at our end result in
characteristic trace files, and most can be dealt with fairly easily. The important thing is to let us know when
you see this as soon a
Good signal, but peaks are
shifted relative to each other and basecalling
results in many N’s. This suggests the
data were analyzed using the mobility files from the wrong version of BigDye. We can
easily reanalyze these files with the correct input mobilities.
Trace file starts right up in
data (i.e., no
flatline “leader” sequence), resulting in loss of the
beginning of expected sequence. This is
a software issue; the automatic analysis has just begun basecalling
at an inappropriate point. We can
manually set an earlier analysis beginning point that will recover the full
sequence.
Trace file starts out well, but
turns into “rolling hills” of color, or starts right out with rolling
hills. This is either a capillary or a
sample issue, but since it’s impossible to tell (though if it starts out nice
it’s usually the capillary), we will re-run such samples if we notice this or
if requested (you have a week to tell us).
Charged contaminants in the sample will lead to a pattern of rolling
peaks throughout the trace file. If
several samples from a particular user show the same pattern, it’s not the
capillary and you will need to alter your procedures to avoid the problem in
the future. In the meantime we’re
usually willing to do re-runs.
Trace file looks good but there
are one or several sharp multi-color spikes throughout the
sequence. This indicates bubble(s) in
the capillary and a re-run will help.
Trace file looks good but
neither you nor anyone in the lab works on that organism. You got the wrong sequence, baby! We need to know this right away since it’s a
sure bet someone else got the wrong sequence too. We can often figure out where the mistake was
and get you the correct sequences without a rerun, within a few hours or
less. A re-run to make sure can be done
if requested. If you get sequence that’s
from your organism but doesn’t seem to be what your primer should have given
you, or you weren’t expecting exactly that PCR fragment but it’s
close, please look over all your manipulations before calling. If it’s not one of the software or switched
tube issues, chances are it’s something at your end.
Depending on the traceviewing program you're using to look at your
chromatogram, you can find different information about your run. Click around the different windows and/or
menu choices and see what you come up with.
Of particular use is the relative signal strength--given in arbitrary
fluorescent units for each base. Ideally
you want to be in the high hundreds or low thousands; this information can be
valuable when deciding how to get better data if that's a problem/
We
want everyone to get the best sequencing data possible. While there are instances where fluorescent
sequencing with BigDye just isn’t going to work,
these are rare. Don’t be satisfied with
“just OK” sequencing data. If you have
used our controls, have gone over the relevant parts of this guide, and
continue to have problems, bring your template(s) to us and we’ll try it free
of charge. We don’t want people to waste
time and money on sequencing when there are more important questions to answer.