DNA Sequencing FAQ

Q1) What is the best way to purify my sample and the precautionary steps involved?
Ans: Virtually any commercial DNA clean-up kits will give DNA of sufficient quality for automated sequencing. It is important to quantify the concentration of the samples prior sequencing. For high quality results, the DNA should be free from protein, salts, RNA, phenol or ethanol contamination and most commonly incorporated nucleotides from the PCR products. However, the most common problem arises from excess salts that are present in either the DNA template or primers used in the cycle sequencing reaction. This usually occurs if a user eluted their DNA in a high salt buffer, or upon precipitation of their DNA from a manual prep that failed to remove most of the salts when washing the DNA. For automated DNA sequencing, we recommend keeping the DNA in 10mM Tris-HCI pH8.0 buffer or deionised water.

 

Q2) What are the common issues in the DNA purification of ready-to-load samples?
Ans:

  • Isopropanol-precipitated DNA is not washed with 70% ethanol to remove excess salt. Wash the DNA pellet at least once with 70% ethanol. Residual salt in the final template will interfere with the activity of Taq DNA polymerase, resulting in sequence data which extend fewer than 200 bases from the primer and exhibits a low signal to noise ratio.
  • The template DNA is not dried completely before final suspension in water. To remove residual ethanol, dry the DNA pellets for 5 min in Speed Vac. If air drying, a brief 15 min incubation of the opened tube at 65°C is sufficient to completely dry the DNA. Residual ethanol is detrimental to cycle sequencing resulting in data with drastically reduced signal.

We provide cycle sequencing reaction clean-up service should you feel that this should not be left to inexperienced personnel. We use Agencourt magnetic particles to clean up sequencing reactions.

 

Q3) What happens to my DNA Sequencing order once its sent to 1st BASE?
Ans: Once your order (containing DNA templates and primers) arrive at our facilities, they are all verified with their corresponding order forms and processed in the same manner.
After the samples have been verified to be purified and meet our sample requirement guidelines, we batch and proceed with DNA sequencing reactions. Results are analysed and sent back to you via email with a weblink. Standard orders will be processed and results delivered with 2 working days. Additional time will be required for orders that undergo additional preparation services.

 

Q4) I quantitated the DNA with a spectrophotometer but why did you find the template DNA insufficient? 
Ans: A typical plasmid miniprep produces 5µg in 50µL. To save precious DNA from being wasted in what is viewed as destructive testing on the spectrophotometer, you remove 5µL from the 50µL plasmid prep. This amount is diluted with 500µL of water for the A260 measurement. Assuming all goes well, you get a reading of 0.02A260. Sounds good?

I'm afraid not. What most operators don't realize is that spectrophotometers can only read down to 0.05A260 reliably if they are properly blanked. Taking the above scenario, instead of 5µL, you used 20µL. This would have produced a reading of 0.05A260, barely scraping past the accuracy of the spectrophotometer. Matters become worse since not all plasmid minipreps produce 5µg consistently. If you are dealing with a low copy plasmid, getting 1µg would be most fortunate.

To be sure, assume that A260 readings from spectrophotometers to be suspicious. The above problems that face spectrophotometers are also found in the Nanodrop device since it is basically a spectrophotometer. Nanodrop readings of 2.5ng/µL and below are not reliable. To be sure, it is advisable to run template DNA on an agarose gel with an appropriate size ladder with a known concentration for the corresponding size band (check the ladder PDS for info) for comparison. Gel elution results in massive losses if your template is a PCR product. Perform as many PCRs as you comfortably can so that you can have enough PCR products after gel elution (or it was commonly called as PCR gel extraction).

 

Q5) Why does the DNA concentration measured by 1st BASE differ from mine? My lab is using Nanodrop to quantify DNA.
Ans: The most common device for measuring DNA concentration and quality is a spectrophotomer. However, there are issues with using a spectrophotomer.They are addressed below.

1) DNA concentration is essentially measured using UV absorbance at 260 nm (1A260 = 50 µg/mL) in a 1cm path length cuvette. However, the presence of contaminating molecules interfere with this absorbance.These contaminants are basically nucleotides, RNA, EDTA and phenol which are commonly found in nucleic acid preparations like plasmid preparations.

2) The dynamic range for spectrophotometer is from 0.05A260 onwards to 1A260. This would mean you would need at least 20uL of plasmid prep to get a minimal reading of 0.05A260.

Unfortunately, using 20uL for quantitation is not possible as the DNA preparation obtained is most often low, averaging at 5ug in 50uL. If a low copy plasmid is used, to get even 1ug in 50uL is difficult.

Hence, most often, 5uL is taken from the 5ug of 50uL DNA preparation and this will give the reading 0.02A260, which is not an accurate measurement (recall the dynamic range).

Some laboratories use a Nanodrop type device. Please be reminded that a Nanodrop is still a spectrophotemeter and hence is subjected to the same problems highlighted above. The lower detection limit of the nanodrop is 5ng/uL but it is shown to be unreliable at the lower range.

Based on the above explanation, we would like to advise that a more accurate estimation of double-stranded DNA is to perform an agarose gel analysis. DNA ladders with their accompanying product data sheets provide the amounts of DNA in ng to the corresponding ladder bands. Using this, we can reliably estimate by just comparing the cut plasmid with the corresponding size band from the size ladder.

To perform quality quantification, we have taken precautionary steps to maintain accuracy and consistency. In the 1st BASE labs, our laboratory apparatus are calibrated and maintained regularly to ensure they are working within their specified limits. We quantify a lot of DNA templates, so in addition to the calibration and maintenance, we have standards and controls to our quantification assays. These measured values are then checked against the standard curve to further ensure the results stay within statistical tolerances.

As you see, there are multiple checkpoints implemented on the quantification process alone. As such, we assure you that readings for the samples that are submitted are accurate. 

 

Q6) My sample’s concentration do not fall within your indicated minimum requirement, can I still send them for DNA sequencing? 
Ans: We will try our very best to sequence the best data from our customer’s sample. If your purified samples did not fall within our minimum requirement, please spare us more volume (e.g. double up your sample volume to 20uL per reaction). We could help to concentrate the DNA samples prior cycle sequencing. It is beyond our guarantee if there are other factors that causing low concentration (e.g. inefficiency PCR protocols) or low recovery (e.g. present of other contaminants, disruption of DNA during purification process and etc.) of your purified samples. Sending purified DNA sample that falls within our sample requirement will greatly increase the possibility in getting good sequencing results.

 

Q7) Can I send the unpurified PCR products for direct DNA sequencing?
Ans: Although we highly recommend sending purified PCR products for sequencing, but there are still some customers sending unpurified samples for direct sequencing. If the residues of the PCR primers were depleted after the PCR, the noise background generated during the cycle sequencing may not affect the major base calls. However, the present of the other contaminants (e.g. dNTPs, salts and etc.) could still compromise the quality of the DNA sequencing result to some extents. We do provide DNA Sequencing Sample Preparation with purification service. 

 

Q8) How to examine the good quality of the DNA sample based on the observation on agarose gel?
Ans: Purified PCR products (>400bp) for cycle sequencing should show a distinct and clear single band on 1% agarose gel. For the size < 400bp, you should check them on 2% agarose gel for better resolution.Conformations of a DNA plasmid that has not been cut with a restriction enzyme will move with different speeds (slowest to fastest: nicked or open circular, linearised or supercoiled plasmid) on agarose gel. After the agarose gel stained with Ethidium Bromide, the purified plasmid DNA with sufficient DNA intensity and free of RNA contaminants is ready to go, RNA contaminants usually appeared to be a big white cloud under the bottom of the gel. Although BigDye® reactions can tolerate RNA contamination up to 1ug, but the present of the contaminants could still compromise the quality of the results to some extents. Please note that they are other contaminants, e.g. salts, ethanols and etc. could NOT be detected through agarose gel electrophoresis.  If you extract and purify your plasmid DNA using a commercial kit, the purified plasmid DNA usually is good for cycle sequencing.

Please click for our quick guide to examine good quality DNA samples from Agarose gel.

 

Q9) Although my DNA sample was showing faint band, but the exact same sample worked fine before! The signal in this new reaction is weak and noisy, or even un-interpretable.How can that be?
Ans: A faint band, when repeated, will often be insufficient DNA template. It is probably luck should you manage to get it on the first run, depending on how noise-free is the reaction to which your sample was assigned. Check the signal strength of the previous good reaction and if it was below G=150, you probably just got lucky before.

Check the annotation from the softcopy of your electropherogram, the Average Raw Signal Intensity, example: A(512), C(533), G(634), T(630). 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. The possible causes are almost the same as those for weak DNA bands or no band. Please refer our Technical Support for more details.

You might get lucky sometimes. Baseline noise varies from sample to sample, 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 not work, because the noise is too high. 

 

Q10) Is it necessary to send primers with the template?
Ans: We provide a wide range of universal sequencing primers free of charge, please click to see the list. You could provide your own sequencing primer with the DNA template. For added convenience, we can synthesize your priority sequencing primers. Preferential pricing and priority will be given to DNA sequencing customers. 

 

Q11) How much primer do you need for sequencing?
Ans: We require approximately 10 pmoles (or 1µl of 10µMolar) of primer per sequencing reaction. Please send at least 5 µl per DNA template as some evaporation may occur. The ideal primer length for sequencing is between 18-25 bases with about 50% GC content. Primer should not form primer-dimers or prime at multiple sites within your DNA template.

 

Q12) Do I need to lyophilize my samples before submitting for pick up?
Ans:  It is not necessary to lyophilize or dry your DNA samples. Please submit your DNA in sterile ddH2O at room temperature.

 

Q13) Do I need to submit my samples chilled?
Ans:  No. DNA is stable in water at room temperature for a few days.

 

Q14) What is the turnaround time?
Ans: Upon the receipt of your samples to our DNA sequencing facility, an order confirmation will be sent to you through email. We typically deliver the sequencing result to you within 48 working hours upon the receipt of your samples. For ready-to-load samples, results are typically out in 24 working hours.

 

Q15) Will 1st BASE keep my samples and primers?
Ans: All samples and primers will be discarded after a successful reaction. We will only keep your samples and/or primer for 1 week upon receiving them. Please indicate in DNA sequencing form if you like us to keep your sample or primers for further sequencing reactions. A nominal fee will apply for storing your samples or primers in our laboratories.

 

Q16) How do I get the DNA sequencing result and how do I view them?
Ans: The results will be sent through email via a link to our server to prevent email storage deficiency. You can then download the compressed file in .zip format (for Windows), which has the complete sequencing results with (i) electropherogram(s) in .ab1 file and (ii) fasta sequence in .seq file. The electropherogram is saved as .ab1 file which can be opened and analyzed using various freeware: Chromas, FinchTV, Applied Biosystems Sequence Scanner 1.0 and etc.; while the fasta sequence file can be viewed via notepad.

 

Q17) What quality control is performed for DNA sequencing reactions?
Ans: A control sequencing reaction is performed with every batch of customer samples. We use the sequencing reaction controls to evaluate the quality of the overall reactions. Our control reactions need to achieve Phred40 data for a minimum of CRL to 850 bp in order to meet out QC benchmark. We can send the control sequencing chromatogram to you upon request at no charge.

 

Q18) Does the bacterial strain used affect plasmid quality for DNA sequencing, and how about the choice of its liquid media?
Ans:  In a large extent it does. Generally, bacterial strains carrying the endA1 mutation provide better plasmid quality for sequencing. Known strains that produce reliable sequence data are: XL1-Blue, XL10-gold, DH5 series, DH1 series, DH10 series, c600, TOP10 and NM294. Avoid the following strains: HB101, JM83, JM101, TB1 and TG1. Manufacturers of plasmid purification kits have optimized their kits. In general, all of them have used LB broth as the optimized growth medium. Our advice is to follow their instructions. Rich media like 2X TY, Super Broth or Terrific Broth enables cultures to reach stationary phase in much sooner than expected. Under this condition, the plasmid purification columns will be overloaded and resulting in poor quality plasmid for DNA sequencing.

 

Q19) Which M13/pUC primer should I use for my plasmid?
Ans:

Featured in this picture are the positions of the M13/-pUC sequencing primers on the pUC19 cloning site. The 3' positions differ by 20 bases at most and will not significantly contribute to sequencing read lengths. If your plasmid is a M13/pUC derived plasmid, our in-house supplied M13/pUC primers should fit.

 

Q20) Why choose 1st BASE as your DNA sequencing service provider? 
Ans: We place top priority in our customers' satisfaction. In addition, we are committed to providing the best products, services and support to answer our users' needs and beyond. By choosing us, you gain the following benefits: -

  • Daily collection and processing of samples
  • High quality, long read lengths with fast turnaround
  • Email notification of sequencing results, electropherograms and sequences.
  • Prompt technical support and consultation.
  • Provision of one free re-run for failed reaction

 

Q21) I've been experiencing sequencing failures when using SP6 or T7 promoter primers with 1st Base. What is going on?
Ans: This actually is a well known problem with the SP6 primer sequence and not so much with the quality of the primer we have on our hands or the current sequencing methodology. The SP6 and T7 Promoter primers are using the universal promoter sequences. These sequence goes back to the days when sequencing is still with S-35 labeling with Sequenase. Sequencing technology has advanced but not these sequencing primers. The primers' have notoriously low melting temperatures. Over the years, there have been variations to this sequence which involves extending the 5' or 3' end in an effort to raise the Tm. However, the extensions are not common among the vectors. In effect the SP6 designed for a particular vector works for that vector exclusively. At 1st BASE, we have decided to stick to using the universal SP6 sequence which allows compatibility with all vector sequences bearing the SP6 promoter. This came with a cost. There will be times when priming will be insufficient with some vectors. We have tried working with lower annealing temperatures for sequencing with these primers. It shows mixed results. 1st BASE has longer has SP6 and T7 primers boasting higher melting temperatures. Named SP6Long and T7Long, their sequences are listed below:

SP6Long: 5'- ATTTAGGTGACACTATAGAATAC-3' 
T7Long: 5'-GTAATACGACTCACTATAGGGC-3'

Please note that these primers do not work on all vectors. Should your vector's promoter sequences matches the above, please indicate on your sequencing request form to use these primers. Another option is to design and synthesize longer SP6 or T7 promoter primer sequences with higher melting temperatures that match the vectors you use for sequencing. Email us should you wish to embark on this.

 

Q22) How do I interpret the data?
Ans: Follow this link to our sequencing troubleshooting guides.

 

Q23) How to determine DNA Sequencing Data Quality?
Ans: At 1st BASE, the trace data quality is measured by a series of statistical measurements. These statistical measurements were developed by genome sequencing centres to ascertain DNA sequencing data quality. These are not standards set by 1st BASE but by these very same genome centres. The statistical measurements are embedded into our trace files and can be read by certain trace viewers.

Quality Values (QV)
The Applied Biosystems 3730XL Genetic Analyzer uses Applied Biosystems DNA Sequencing Analysis Software v5.2 with KB Basecaller. The KB basecaller assigns quality values (QV) to each basecall (pure and mixed bases). The QV is calculated using the following equation: QV = -10log10 (PE) where PE is the probability that a basecall is erroneous.

The calculated QV for each base may be visualised on the analysed data as coloured QV bars above each basecall. These QV bars are only visible using the DNA Sequencing Analysis Software or Applied Biosystems Sequence Scanner program. The software defines low quality bases (red QV bars which can be written off as unreliable) as those with QV scores less than 15, medium quality bases (yellow QV bars needing a manual review) with scores 15 to 19 and high quality bases (blue QV bars) those with QV scores greater than 20.

The table below illustrates the QV and the corresponding probability of error (the probability that a base was miscalled). Mixed bases will have lower QV values. Typically high quality mixed bases will have QV in the range 10 to 50. It is shown that these error probabilities are very accurate. The high accuracy makes it an ideal tool to assess the quality of sequences. By looking at individual sequences, failed reactions or low-quality reads can easily be identified.

QV PE QV PE QV PE
1 79% 21 0.79% 41 0.01%
5 10% 25 0.31% 45 0.00%
10 10% 30 0.10% 50 0.00%
15 3.20% 35 0.03% 60 0.00%
20 1.00% 40 0.01% 99 0.00%

The QV is also colour coded and appears above the base that is called. It can only be seen by the following software:

  1. ABI's Sequence Scanner 1.0 (MS Windows)
  2. FinchTV (OSX and Windows)
  3. Staden's Trev (multiple platform)
  4. 4Peaks (OSX)
  5. Edit View (OS9)
  6. Chromas (MS Windows but not the freeware version)

Trace Score
The trace score represents the average QV of the bases in the clear range of the sequence. The Sequencing Analysis program determines the clear range of the sequence by trimming bases from the 5' to 3' ends until fewer than 4 bases out of 20 have QV's less than 20 (these values are adjustable, our settings are given). For example, a trace score of 40 would mean that the probability of the basecall being wrong is 1 in 10,000 or the accuracy of the basecall is 99.99%.

Contiguous Read Length (CRL)
The CRL is the longest uninterrupted stretch of bases higher with QV higher than a specified limit. At 1st BASE, it is 20. In the evaluation of the quality of each base, not only the QV of the base is used but also the QV of the adjacent bases within a specified window size.
Low CRL: <300
Medium CRL: 300-600
High CRL: 600 and above
If the sample is not a PCR reaction, a low CRL corresponds to a failed or poor reaction. 
Please note: The average length of usable data is between 750-850 bases. After 850 bases the peak resolution weakens and base calling errors increase.

Average Signal to Noise Ratio
The signal to noise ratio is the average of the average fluorescence intensity of each of the four dyes used in sequencing divided by the average noise for the corresponding dye.

 

Q24) I believe that my DNA plasmid sample (pET-series vector) was good in quality. The exact same sample worked fine with T7 promoter before but not this time! How can that be?
Ans: Based on our observation from the customers using the plasmid construct with pET-series backbone, it frequently fails to be sequenced by T7 Promoter.

We would like to suggest using an alternative sequencing primer pBRrevBam, which primes at upstream of T7 Promoter on pET-series vector [except pET-17, pET-20 and pET-23].  Although both primers have the priming site on pET-series vector, but the signal strength generated by the pBRrevBam primer is 2 to 3 times higher than T7 promoter. This indirectly proves that pBRrevBam primer is more potent to give higher rate of successful results as compared with T7 promoter in cycle sequencing reactions. For this reason, we recommend pET vectors [except pET-17, pET-20 and pET-23] to be sequenced with pBRrevBam primer to replace T7 promoter. pBRrevBam primer is now available from our FREE universal primer list.

For your reference, below is the comparison of the signal intensity (circled in red) for both pBRrevBam and T7 promoter as sequencing primer. These traces are generated using the same amount of DNA template during standard cycle sequencing reaction, and they are generated by the same Thermocycler and Genetic Analyzer.

 

Q25) My DNA sample band is present and strong, but irregularly spaced, or with mixed colours peak. Why? 
Ans: If you see this, you usually have two sequences superimposed on each other. There are several common causes:

  • Multiple priming sites - the sequencing primer binds to two (or more) sites on the template
  • Multiple templates - there are two (or more) templates present.
  • If this was a plasmid reaction – the plasmid DNA might be sample mixed-prep.
  • If this was a PCR reaction,
    1. you didn't remove the original primers; or
    2. one primer generated both ends; or
    3. there is more than one amplified species present.

 

Q26) My PCR products showed primer-dimmers, is it necessary for me to purify the PCR products using gel extraction?
Ans: There are many ways to sequence purified PCR products. While we have received feedbacks from users that they are satisfied with the results from purifying using only normal column purification, if you are new to sequencing PCR products, we would recommend that you gel-purify the fragments as a safeguard measure. 

Based on our observation, normal column purification will not remove double-stranded DNA completely. Occasionally, there are noises observed in the front part of the sequencing traces due to presence of primer-dimer impurities. From the perspective of downstream application, if the user regards the front part of the noise sequences as acceptable to be trimmed off or excluded from pair wise alignment, this should not affect the overall sequencing reads. However, this method may not be ideal for those who need very clean reads from the beginning of the sequencing traces, especially for PCR products that are shorter than 250bp.