SEQUENCING SERVICE CLIENT SOP
Title: SEQUENCING SERVICE CLIENT SOP / SOP No.20.2.A
Version: 2
Effective Date: 21th August 2014
Sequencing Service Client SOP
1 Clean DNA for setup
2 Measure DNA quantity
3 Sequence Setup
4 Handing in your samples
5 Getting your results
Composed / Hiren Sheth / Research AssistantDate composed / 13.11.2011
Reviewed / Dahlia Saroufim / Research Assistant
Date reviewed / 16.08.2010
Edited / Pavel Bitter / GMG Manager
Edited from / Version 1
Edited because / Name change of facility and updates
Authorized / Deborah Kelly / Quality Manager
Date authorized / 16.08.2014
1 Clean DNA for sequence setup
The quality of your template will influence the quality of the sequence results directly, the cleaner the PCR product or plasmid prep, the better your results.
CLEAN DNA -> good results
PCR products
Residual primers and dNTPs must be removed from PCR reactions prior to sequencing. There are various commercial spin column kits (Qiagen, Invitrogen etc) or magnetic bead systems (Dynabeads, Agencourt) available. Alternatively an Exonuclease I / Shrimp Alkaline Phosphatase (Exo/SAP) digestion can be used.
MinElute PCR Purification Kit (50) / Qiagen / 28004 / $ 197.00PureLink™ PCR Purification Kit / Invitrogen / K3100-01 / $ 134.00
Agencourt AMPure 5 mL Kit / Beckman/
Agencourt / A50850 / $140.00
Poor template quality is the most common cause of sequencing problems. Excess PCR primers, dNTPs, enzyme, and buffer components (from PCR) are the most likely contaminations for PCR products in sequence setup.
The following are characteristics of poor quality templates:
-Noisy data or peaks under peaks
-No usable sequence data
-Weak signal
Plasmid Preps
The quality of your plasmid kit will determine sequence results. So, any salt in the plasmid prep will compete with the plasmid DNA when injecting into the capillary. Any mini-prep carryover reagent will harm the efficiency of the Sequenase enzyme. Commercially available mini prep kits will give good sequencing results.
QIAprep Spin Miniprep Kit (50) / Qiagen / 27104 / $172.00QuickLyse Miniprep Kit (100) / Qiagen / 27405 / $327.00
Purelink Quick Plasmid Mini Prep Kit / Invitrogen / K2100-10 / $127.00
These kits are safe and easy to use but an ”in-house” alkaline lysis followed by precipitation will yield good results as well if made sure that no contaminants are carried over and the precipitation has washed away all residual salts. Potential contaminants include: Proteins, RNA, Chromosomal DNA, Residual organic chemicals, e.g., phenol, chloroform, and ethanol, Residual detergents.
Cleaning Up Dirty Templates
A “dirty” template preparation sometimes can be cleaned up with the following method:
1 Extract the DNA twice with 1 volume of chloroform or chloroform:isoamyl alcohol (24:1 v/v).
2 Add 0.16 volumes of 5M NaCl and 1 total volume of 13% PEG.
3 Incubate on ice for 20 minutes, then centrifuge at maximum speed in a microcentrifuge at 2–6 °C for 20 minutes.
4 Rinse the pellet twice with 70% ethanol.
5 Dry the pellet in a vacuum centrifuge for 3–5 minutes or to dryness.
The host strain used for template preparation can impact template quality. One host
strain may produce better sequencing results for a specific template than another.
If you plan to use a commercial template preparation kit, contact the vendor for
information about host strains that work well with that kit.
2 Measure DNA quantity
In more than 95% of all cases in which clients receive bad sequencing results there is one simple cause: the quantity of the plasmid or PCR product DNA that was put into the sequence reaction was wrong.
The sequence reaction is a delicate balance of input DNA and sequence primer. Its not only the amount of DNA in ng, it is the amount of primer binding sites in the target DNA versus amount of sequencing primer. If there is not enough template the signal of your results will be low. But if there is too much DNA then you will receive lots of short fragments, as the kinetics of cycle sequencing produces short sequences first, only later in cycle sequencing longer sequence fragments are produced. With an excess of DNA all primer will bind in the first few cycles of cycle sequencing and later there is no primer left, so that a very typical result is produced (see trouble shooting section).
It is very important to quantify your DNA!!!!
Agarose gel electrophoresis
There are different ways to quantify your DNA, some are more reliable than others.
An Agarose gel will give you a good idea of the quality of your DNA, whether it is fragmented, multiple PCR products (which will NOT sequence well) or whether you only have a smear of bands which again will not sequence.
The gel will also give you an approx. idea about quantities (the faintest band on a gel equals ~40ng of DNA). You can quantify better if you compare the intensity of bands to a ladder with known concentrations.
Purified DNA should run as a single band on an Agarose gel.
Uncut plasmid DNA can run as three bands: supercoiled, nicked, and linear.
Fluorophotometry
The next step down in precision compared to gels are quick measurements that involve intercalating dyes like SYBRgreen, Hoechst or Picogreen and a fluorometer reading. In the facility we have a Qubit from Invitrogen for clients to use, but other systems work as well. The advantage with fluorescence based systems over simple spectrophotometry is that the result is not influenced by contaminants like Ethanol or RNA and your readout is more accurate.
Keep in mind though, that you can not distinguish primer-dimer or multiple PCR products from your product of interest. You will not see whether your plasmid is supercoiled which is harder to sequence. You will not see whether there are protein contaminations or any other sequence inhibiting contaminants.
Spectrophotometry
The most common used and easiest and cheapest way to quantify is a simple A260 absorbance readout. This method is the least precise compared to the above and will not differentiate between RNA and DNA, the readout doesn’t give you a clue about contaminants which often absorb at the same wavelength increasing your value and making you assume you have more DNA than your actual DNA quantity is.
We recommend a Nanodrop measurement only if you are experienced with DNA preps and are aware of the potential confusion with this measurement. The only advantage of the absorbance reading is an idea about protein contamination: your A260/A280 ratio should be 1.7–1.9. Smaller ratios usually indicate contamination by protein or organic chemicals.
Neither of these methods shows the presence of contaminating salts that can cause
noisy data. If you suspect that your DNA is contaminated with salt, remove the salt
before sequencing. The most efficient method for salt removal are spin columns or ethanol precipitation. If you can, check your DNA with different methods. Yes this is costly but setting up sequences is costly too.
3 Sequence setup
The best way to give sequence reactions to the facility is to mix your DNA with primer in 0.2ml 8 strip tubes (Cat number 3131-00; 0.2mL 8-Strip PCR Tubes with Caps pkt/ 1000; $80.00; Interpath Services Pty Ltd.) and dry the mix until no liquid is left in a thermocycler or heat block.
How much DNA?
Of your quantified DNA add the amount given in the following table. The volume of the DNA is not crucial as it is dried down anyway.
Sequencing Reaction of: / DNA amount / Primer amountPlasmid DNA / 20-40ng / + / 3.2pmol
PCR Product / 10-30ng
mix in one 0.2ml tube in a total volume of up to 20ul
If you are in doubt about how much DNA exactly you have then give us then it is a good idea to prepare a dilution series of your template DNA (give us for example 4 tubes with 10, 20, 30 and 40 ng of your plasmid with the same amount of primer in each tube).
How much primer?
Whilst you adjust the DNA amount, it is recommended to keep the primer concentration constant. Use this seemingly odd amount: 3.2pmol of primer.
This amount of primer can be easily added to the sequence reaction if you have it in a concentration of 3.2pmol/ul and add 1ul of your primer.
For this you can dilute a primer stock of usually 100pmol/ul (=100uM) in the following way:
add 1ul of your stock (100uM) dilution into 31ul water.
Add 1ul of this to your DNA then dry down the mix.
Keep in mind, that a dilution of this primer at 3.2pmol/ul is not very stable if repeatedly frozen and thawed, so throw out this dilution and make a fresh one each time you set up sequences.
Some words to your primer design
The choice of primer sequence can have a significant effect on the quality of the sequencing data obtained. The annealing temp for cycle sequencing is 50C, which is a standard program used on all samples. Adjust your annealing temp therefore at around 55C. If you are using a separate primer for cycle sequencing than for previous PCR amplification you usually get better sequence results. You will receive quality data around 20 Bp further downstream from where your primer binds. The average read length is about 500Bp. Whilst you will receive up to 800 Bp data in your results, you will not be able to use the last 300Bp for much more than sequence confirmation. True SNP mutation detection is not possible in the last 300Bp. To read further into the plasmid or insert or PCR product you will have to use a second primer that binds further downstream.
Some general primer design rules:
-Primers should be at least 18 bases long to ensure good hybridization.
-Avoid runs of an identical nucleotide, especially runs of four or more Gs.
-Keep the G-C content in the range 30–80%, preferably 50–55%.
-primers with Tm>45 °C produce better results than primers with lower Tm.
-Use of primers longer than 18 bases also minimizes the chance of having a
secondary hybridization site on the target DNA.
-Avoid palindromes because they can form secondary structures.
-The primer should be as pure as possible, preferably purified by HPLC.
The following formula can be used for a rough estimate of melting temperature:
Tm = (number of A + T residues) x 2 °C + (number of G + C residues) x 4 °C
What next?
Now there are two choices, you can give your sample to the GMG facility OR you can proceed and do your own cycle sequencing. The advantage for the latter is that you can save some money when you are cleaning up the cycle sequencing reaction yourself. The setup costs are more or less the same in the facility or when you setup the cycle sequencing. Obviously you take the risk, if you setup and cleanup your samples. The average saving in a sequence and cleanup that you perform versus the GMG facility usage is $1/sample but you have to consider your labour time as well.
GMG facility usage
If you want the GMG facility to perform the BigDye reaction and Cleanup for you (with the high quality and safety of standardized performance) please go to step 4 “Handing in your samples”
Do it yourself
The reaction is composed of the sequence buffer and the actual 3.1 BigDye reaction mix which contains the enzyme and the labelled dNTPs. The 3.1 BigDye reaction mix can be purchased from the GMG.
The reaction mix is light and temperature sensitive but can be handled at room light and kept without ice for several hour without loss of performance. Make a Mastermix according to the following table:
REAGENT QUANTITY
BigDye Reaction Mix (2.5x) 1 µl
5x Sequencing Buffer 1.5 µl
Primer 3.2 pmol/ul (use 1ul)
Template see table above
Deionized Water add to 10 µl Total Volume
For most applications the reagent mix can be used at this concentration without any deterioration in sequence quality. However in some instances it may be necessary to use a higher concentration of BigDye, we will advise you on this after the first failed results.
Cycling Programme
In the cycling program are three variables. The annealing temperature at 50°C can be slightly raised if your target has secondary structure (only do so after first failed attempt). The extension step at 60°C can be shortened if you want to save time and amplify PCR products or short plasmid sequences, the calculation is: each 10 Bp of your sequence need 1 second to be amplified, so 4min amplifies ~2400Bp. The third variable is the amount of cycles. We recommend to start with 30 cycles, but if your sequence reactions come back as too strong (have a look at the raw data) you might want to reduce to 25 cycles.
Perform an initial denaturation
96°C for 1 min
Repeat the following for 30 cycles:
96°C for 10 sec
50°C for 5 sec
60°C for 4 min
Hold at 10°C until ready to cleanup
Alternatively freeze at -20°C for days (although freezing will reduce data quality slightly, so if you can avoid freezing process immediately).
a) Cycle sequence clean-up methods
Before a sequence reaction can be run on the instrument the unincorporated didesoxynucleotides need to be removed. There are different methods how to do that, the cheapest one is Ethanol precipitation. Below you will find protocols for the most common precipitation methods. There are also a number of commercial kits available (spin columns, magnetic beads, Sephadex) which can be used for low or high throughput (96 well format).
The GMG Facility however uses the BigDye-XTerminator kit for purification of sequencing samples which is recommended by ABI. This ensures reliable high quality sequence cleanup.
The EDTA / Ethanol Precipitation method is recommended for sequencing plasmids. This method produces the cleanest consistent signal and is particularly good at removing unincorporated dye-labelled terminators. However, it may cause some loss of small molecular weight fragments.