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| qPCR Assay Design & Optimization |
Do I need to calibrate my instrument to detect your Pulsar® 650 dye?
Yes, an important aspect of analyzing a FAM/Pulsar 650 duplexed assay is calibration-compensating for any signal crosstalk outside of their designated channels. This is achieved by a CC file loaded into your LightCycler® computer. Dual-labeled probe users have the flexibility to create this file in one of three ways:
1. download a previously-saved CC file according to your instrument model
2. generate an assay-specific CC file using the Dual-labeled probes directly
3. generate a universal CC file using T10 calibration dyes (inquire about
availability)
Unfortunately, Molecular Beacons cannot be used directly to generate an
assay-specific CC file, and so these users are limited to the other methods.
Directions for each of these methods, as well as the CC files, are available
for download from our website and can be found at:
/download/calibration.asp
For additional calibration questions, a FAQ clearly explaining calibration
of the LightCycler® has been kindly provided by our partner, Fluorescentric.
http://www.fluoresentric.com/
How are fluorescent labels on dual-labeled probes accounted for when measuring the absorbance?
Biosearch calculates the extinction coefficient on labeled oligos by adding a correction factor to the extinction coefficient to account for each fluorescent label. The extinction coefficient found on the certificate of analysis accompanying any oligo purchased from Biosearch takes any dyes into consideration. Using this extinction coefficient with an OD reading at 260 should give you an accurate absorbance value; this is the way that Biosearch measures the absorbance during QC. If you require further information, we can provide you with details on what values we use for the correction factors of the dyes, but in order to simply obtain an accurate absorbance value, the extinction coefficient listed on the certificate of analysis should be sufficient.
If you need us to re-send the certificate of analysis for any of your oligos, please contact our Technical Support department at techsupport@biosearchtech.com
How do freeze thaw cycles affect oligonucleotides?
Oligonucleotides should be subjected to minimum number of freeze-thaw cycles. Therefore, we recommend that you prepare aliquots and store microvials each having sufficient material for a day’s set of experiments and freeze at -20° C or -80° C.
“The repetitive freezing/thawing of an oligonucleotide strand can lead to the undesirable decomposition of a sample through the loss of a terminal phosphate group, a base or the entire oligonucleotide unit. This observation relates to the fact that during the freezing/thawing process the oligonucleotide and buffer components isolate from the solvent system. As a result this isolation increases the percentage of similar intermolecular attractions and bond breakages.”
Analysis of the degradation of oligonucleotide strands during the freezing/thawing processes using MALDI-MS
(Analytical Chemistry, Volume 72, Number 20, Pages 5092-5096, 2000.)
How do I calibrate my instrument for the CAL Fluor Dyes?
In order to successfully use dual-labeled probes containing CAL Fluor® and/or Quasar® dyes for real-time PCR and quantitative PCR (qPCR) multiplex assays, certain real-time PCR instruments need to be calibrated to recognize the pure dye spectra. Spectral calibration is critical for multiplexed assays to differentiate overlapping fluorescent signals from one another. The document below with give users step-by-step instructions for calibrating their instruments:
/assets/BTI_Spectral_Calibration_Instructions.pdf
How do I determine the amount of diluent to add to my probe to prepare a stock concentration?
We recommend preparing a 100 µM stock solution of your probe. To prepare a 100 µM stock of reconstituted probe: Multiply the “total nmol” value found on Certificate of Analysis by 10. The resulting number will be the volume of diluent (in microliters) to add to your probe. Once resuspended in the appropriate volume of dilution buffer, the probe will be at a 100 µM stock solution.
Tip: Be sure to make aliquots of working concentration of probe to prevent freeze thaw cycles that may degrade the probe. We recommend freezing probes either at -20 ºC or -80 ºC. Probes and primers can be stored in this state for over one year.
How do I determine which design mode to use in RealTimeDesign?
The RealTimeDesign (RTD) Software offers three different modes to accommodate a range of needs with different users in mind:
Express Mode is designed with simplicity in mind. This mode does not require any input from the user other than sequence submission and label selection. By traversing three parameter sets of decreasing stringency, this mode proposes assays that have been confirmed to amplify with good performance characteristics.
Custom Mode is designed for the advanced user who wishes to inspect and exclude certain oligo candidates from each stage of the design process. To facilitate fine-tuning, custom mode provides access to the same parameter sets traversed in Express Mode.
Batch Mode, while similar to Express Mode, allows users to initiate a design run (against as many as 10 target sequences), and then close the browser. An email will be sent alerting the user that design has completed, at which point the proposed assays can be inspected from the Design Run History page.
How do I enter in a SNP, MNP or InDel sequence into RealTimeDesign?
Users must select the “SNP Genotyping” application in order to have the RealTimeDesign (RTD) Software design Single Nucleotide Polymorphisms (SNPs), Multi-Nucleotide Polymorphisms (MNPs) or Insertions/Deletions (InDels). Users need to annotate their input sequences in one of the following formats:
Single Nucleotide Polymorphism: [C,T] where C and T represent the single base pair mismatch. Alternatively, users can use the IUPAC code for the SNP represented in the following list:
Nucleic acid codes
code description
A Adenine
C Cytosine
G Guanine
T Thymine
U Uracil
R Purine (A or G)
Y Pyrimidine (C, T, or U)
M C or A
K T, U, or G
W T, U, or A
S C or G
B C, T, U, or G (not A)
D A, T, U, or G (not C)
H A, T, U, or C (not G)
V A, C, or G (not T, not U)
N Any base (A, C, G, T, or U)
Multi-Nucleotide Polymorphisms:
[CA/GT] where CA and GT represent the multi-nucleotide mismatch
Insertions/Deletions:
[CA/--] where CA represents the insertion
How do I find the primers and probe sequences after I design them?
The RTD software automatically stores all primer and probe designs for users. To retrieve the previously primer and probe designs, first log into the RTD Software and then go to the “Main Menu”. Then go to the “Design Run History”. You should now see all of the oligos designed starting from the newest listed first. Clicking on “Details” will give you the sequence information.
If your designs are not listed, it is possible that they might be archived. To retrieve archived designs, click on the checkbox “Show Archived Runs”.
How do I know what fluorophore to pick for my dual-labeled probe?
The choice of fluorescent reporter to label your dual-labeled probe depends upon your instrument optics and also the degree of multiplexing you wish to achieve.
If your assays will be amplified separately and not combined together into a multiplexed arrangement, it would be recommended to label each with FAM. FAM is the most commonly used fluorophore and is nicely detected by all real-time PCR instruments.
The optic capabilities of the instrument principally determines the degree of multiplexing and which fluorophores should be used. To determine this, visit our multiplexing qPCR applications webpage.
/assets/bti_bhq_selectionchart.pdf
How do I order primers and probes after I design them?
Once the RTD Software generates a primer and probe design, users can click on “Order Now”. You will then be given an option to select the synthesis scale (200 nmol, 50 nmol and 1 umol). Clicking on “Submit” will take you to a checkout screen that allows users update, delete or place any notes for each oligo design. To finalize the order, click “checkout” to email the order to Biosearch. You will be able to enter in shipping and billing information as well as review your order.
How do I quantitate oligonucleotides by spectrophotometer?
Protocol for the Quantitation of Oligonucleotides by Spectrophotometer:
1. Add an aliquot of the resuspended oligonucleotide to a final volume of 1000 µl of water or appropriate diluent.
2. Vortex or pipette up and down for 15 seconds.
3. Read the absorbance of this dilution at 260 nm (A260).
4. Use the formula below to calculate the molar concentration (use the nmol/ OD260 from the Certificate of Analysis in your calculation) of oligonucleotide in your stock solution.
Concentration in nmol/ml = A260 × nmol per OD × dilution factor*
(* the dilution factor is determined by 1000/amount of resuspended oligo added for the dilution, i.e, if you added 50 µl of resuspended oligo to 950 µl of water to read the absorbance, you dilution factor would be 1000/50 = 20)
If you have to calculate molar concentration and the nmol/ OD260 value is not readily available, the following formula will also work.
Concentration in nmol/ml = A260 × dilution factor × 10^6 / Molar Extinction Coefficient
How many PCR reactions will I be able to run with my probe or primer?
The number of reactions you can run with a given amount of probe or primer is dependent on a few factors, such as input concentration of the primer or probe and the final yield of the synthesis product, which can be variable. If you plan to use your primer for PCR, 1 nmol of product will provide for approximately 100 PCR reactions (when the primer is used at a 200 to 250 nmol concentration in a 50 µL reaction). As for probes, 1 nmol of product is usually enough to run 100-300 reactions but again, this depends on the concentration of probe you are using in your reaction.
I am having a problem with background drift with my Dual-labeled probe, what could be happening?
There are many potential problems that can cause the background to drift upwards in your Dual-labeled probe. It would be good to check if your primers for 3' overlap. But a larger issue is DTT carry-over from your reverse transcription reaction. DTT will degrade the Black Hole Quenchers and cause the fluorescent signal to increase. DTT is present in some reverse transcription mastermixes, so it is best to check this, especially if you are doing a one step reaction.
I am not getting any signal generation with my Dual-labeled probe. What could be going on here?
If this is probe sequence that had not been used before or is a published sequence, there are a few things that can be done to determine probable cause.
1) Double check the sequence of the probes and primers on the tube label and Certificate of Analysis. Are the bases correct? Are the fluorophore and quencher modifications correct?
2) Analyze primer sets with SYBR Green chemistry or run failed real time PCR reaction on a agarose gel and check for correct amplification product size. This will determine if the primers are working as expected.
If these two basic checks have passed and the probe is still not working. There are a few things that could be going on here:
1) Mastermix components: When making your mastermix, could it have been possible that one of the components (i.e. polymerase or the probe itself was left out?). It’s always a good idea to repeat a failed experiment, just to make sure it was not due to a simple mistake as missing an important ingredient in the mastermix.
2) One of the most common causes of failed Real Time PCR, is poor probe design. How long is the probe? What is the Tm of the probe? Dual-labeled probes and Molecular Beacons should usually bind higher than the annealing temperature of thermal cycling. If probe Tm's are too low, the probe will not hybridize resulting in no signal generation.
Tip: It is also recommended to avoid designing a probe with a G at the 5’ end. The G can quench the fluorophore and result in poor signal generation results.
I am performing Real Time PCR and for my negative controls I am getting products. What could be going on here?
The most likely cause of negative controls coming up positive is contamination. Here are a few suggestions to prevent contamination:
1) Aliquot your probe and primers into experiment size volumes. (Highly recommended: This will not only decrease chances of contamination but minimize freeze thaw cycles that can be degrade oligonucleotide quality.
2) Use separate work areas for PCR set-up, DNA/Template Addition, and Amplification product handling.
3) Clean PCR work areas regularly with 70% ethanol and UV irradiation.
4) Pipettes: Use 70% ethanol to clean micropipettes regularly. Use sterile and filtered pipette tips to minimize aerosols. It’s recommended to dedicate pipette tips for Pre-PCR set-up and Post-PCR analysis.
5) Consider using Uracil-N-Glycosylase (UNG): An enzyme that reduces contamination by removing uracil from single and double stranded DNA. This product can be ordered through Roche Applied Science (Catalog# 11775367001)
It’s usage as indicated on package insert:
“Uracil DNA glycosylase can be used with dUTP to eliminate PCR "carry over" contaminations from previous DNA synthesis reactions. To make PCR products suspectible to degradation, dTTP has to be substituted by dUTP in the PCR reaction mix. Subsequent PCR reactions mixes must be pretreated with UNG prior to PCR to degrade uracil-containing DNA. Native DNA does not contain uracil so that the sample is not degraded by this procedure
If contamination is suspected: Replace all reagents and stock buffers. Thoroughly clean PCR preparative areas.
I am using a BHQ labeled probe, how do I adjust my thermal cycler’s settings to account for the quencher?
BHQ labeled probes can be used on any PCR instrument. Set-up varies between instruments, but generally, if there is an option for picking a dye name for a quencher, pick "none" or “dark quencher,” this option is offered so the built-in algorithm will subtract fluorescence associated with the dye-name entered. Since the BHQs have no native fluorescence there is no quencher-fluorescence for which to account. Probes made with BHQ dyes exhibit extremely low background fluorescence, enabling enhanced detection sensitivity!
If I have my own primer designs, can RealTimeDesign (RTD) design only the probe for me?
Users can use Custom Mode and Anchored Oligos in order to manually input the Forward Primer and Reverse Primer. The RTD Software will then use the set parameters to design the dual-labeled probe.
Note: Biosearch does not recommend that users use their own primers. The RTD Software has certain default parameter settings that are used to design the most optimal primer and probe set. Using primers designed outside of the RTD Software may not produce the most optimal design which could effect PCR efficiency.
If I left my oligos sitting on the lab bench over the weekend, are they still ok to use?
Dry oligonucleotides are quite stable, however oligos in solution are not as stable. The stability of your oligos in solution will depend upon the quality of solution used to resuspend the oligos. For most purposes, resuspended oligos should still work well, even if left at room temperature for a couple of days. However, keep in mind that to ensure optimum activity and safeguard maximum performance, dual labeled probes should always be protected from light to avoid photo-bleaching.
If I order a dual-labeled probe with an internal BHQ modification, what will be at the 3’ end of the probe?
For all internally modified BHQ probes there will be a phosphate group synthesized on the 3’ end to prevent extension of the probe, unless otherwise specified by the customer.
Is there any way of designating a specific region on my sequence where RealTimeDesign can design my primers, probes and assays?
RealTimeDesign users can restrict the primer, probe and assay designs to a specific sequence location(such as an intron spice site or exon-exon boundary) by placing a “~” character at a point within the sequence marking the desired location.
Example:
CAAAGGGTTGCAC~AAGATGGATGATCG
What are Tandem Repeats and Mask Tandems in RealTimeDesign?
When using Custom Mode, Tandem Repeats is the toggle that instructs the RealTimeDesign (RTD) Software to automatically mask or unmask the tandems repeats that the software has identified on the inputted sequence. Tandem Repeats are defined by the software as all identified mono, di, tri, tetra and penta nucleotide repeats. Masking Tandems converts the repeat bases to an N. N is the IUPAC ambiguity code that represents any of the the four nucleotide bases A, T, C or G. The RTD Software will eliminate all primers and probes that contain a certain number of ambiguity codes. Unmasking tandems repeats will allow the RTD Software to consider the repeat regions within the inputted sequence with designs primers and probes.
Note: Custom Mode will allow the user to determine which tandem repeat to mask by checking or unchecking the box next to each repeat region. Users can also choose to “Mask All” or “Unmask All” as well.
What does the Overall Rank score in RealTimeDesign represent?
The overall rank is the ordering of primers, probes and assays based on the scoring function. All scoring functions are a composite measure of how far the overall distance of the primer, probe or assay results varies from an ideal primer, probe or assay. All design rules are taken into account including all minimum and maximum constraints. In other words, the rank is based on how closely the primer, probe or assay design matches the ideal values for each parameter setting. The Higher the rank (based on a maximum rank of 100), the more optimal the primer, probe or assay considering the design rules. Users can view the ideal conditions by clicking on “Edit Parameters”.
Note: the theory behind the rank score is that combinations of the highest-ranking primers and probes will result in assays that have a rank very close to the mathematical optimum. The results are assays that are very sensitive and specific.
What is the difference between the FAM-BHQ ValuProbe priced at $95 and the $150 FAM-BHQ probe?
The delivery amount for the Valuprobe is 10 nmole, while the standard $150 probe has a minimum delivery of 10 nmole but averages closer to 20 nmole. The other difference is the level of purification. The Valuprobe is purified using Reverse Phase HPLC so that residual fluorescent contamination and other synthesis impurities are removed. The $150 probe is purified by dual HPLC (anion exchange HPLC followed by reverse phase HPLC) which ensures the absolute highest quality in the manufacture of BHQ-quenched dual label probes.
The level of purification is dependent on the researchers preference for purification stringency. It’s recommended to use dual HPLC purified probes for multiplex reactions. For more detailed information on the probe quality comparisons, please contact techsupport@biosearchtech.com
What is the Failure Count Data in RealTimeDesign?
During the primer and probe design process, the RealTimeDesign (RTD) Software uses a variety of parameter sets (e.g. Amplicon Length, Primer Tm, GC Percent) in order to design primers and probes. Each parameter set has a minimum and maximum setting. If the design falls outside of these parameters, a number will show up in the Failure Count Data indicating how many designs failed for that particular parameter. Editing the Parameters can help to alleviate the failure count.
What is your recommended method for reconstituting and storing oligos?
To reconstitute: Centrifuge the tube for a few seconds to collect the DNA in the bottom of the tube. Carefully open, add an appropriate volume of TE buffer and close the tube. Allow the oligo to rehydrate for several minutes and vortex for 15 seconds. It is recommended that oligos be reconstituted at concentrations greater than 10 µmolar in TE (10mM Tris-HCl, pH 8.0, 1mM EDTA) and stored at -20° C. Stability (when kept at -20° C) of lyophilized material is >1 year and the stability of reconstituted material may vary depending upon storage conditions.
* Note: Many dye-labeled oligos are sensitive to light. Light exposure is kept to a minimum during synthesis and shipping. Upon receipt, please store in the dark.
To prepare a 100 µM stock of reconstituted probe: Multiply the “total nmol” value found on Certificate of Analysis by 10. The resulting number will be the volume of diluent (in microliters) to add to your probe. Once resuspended in the appropriate volume of dilution buffer, the probe will be at a 100 µM stock solution.
For example: If you probe has a total nmol of 20 nmol (as indicated on the certificate of analysis)
(20)(10)= 200 microliters is need to reconstitute probe to get 100 µM stock solution.
When I received my Molecular Beacon probe I also received a complement, what is this for?
Biosearch supplies complements free of charge with each Molecular Beacon probe order. The complement is a short oligonucleotide complementary to the loop region of the probe. The complement can be used as a positive control to validate the probe’s functionality and fluorescence.
Where can I find information that explains the differences between each of the probes you offer?
We offer a number of different qPCR probes for your convenience. We offer Dual-labeled, Molecular Beacons, Scorpions™, Amplifluors® and Plexors™ for your specific assays. For detailed information about how these probes work please watch our Probe Formats Animation (Flash). If you have any further questions, please do not hesitate to contact us by e-mail at: techsupport@biosearchtech.com
Why is my BHQ-Pulsar probe not detected in a singleplex reaction on the Lightcycler?
If you are using a Lightcycler® 2.0 the most likely explanation is an instrumental issue. The Lightcycler instrument seeks FAM fluorescence from each capillary during the optics optimization procedure. In a singleplex reaction with only a BHQ-Pulsar 650 probe the instrument will not detect the capillary. To overcome this instrumental issue, FAM calibration dye needs to be spiked into any capillary that is lacking a FAM probe (Pulsar 650 singleplex reactions). We recommend spiking this dye to a final concentration of 10nM. A FAM calibration dye should have been included with your first order of BHQ-Pulsar probe. Please contact us if we can provide you with additional FAM calibration dye.
If you are using a Lightcycler® 1.2, which does not seek out FAM fluorescence during the optical optimization procedure. Simply raising the pre-set seeking temperature can resolve the detection of the BHQ-Pulsar Probe. Try raising the seek temperature between the ranges of 50° C to 95° C until an adequate signal is detected.
I am a beginner at Real Time PCR, do you have any information to help me design my Dual-labeled assay?
There are a few reference articles and resources for information on designing Dual-labeled assays. These articles have general recommendations on oligonucleotide design, cycling parameters and set-up suggestions.
http://dorakmt.tripod.com/genetics/realtime.html
Here’s the link to another helpful resource:
http://www.gene-quantification.info/ This site offers many of great links to articles on Real Time PCR.
Another great resource is the Yahoo! QPCR listserver:
http://groups.yahoo.com/group/qpcrlistserver/
Also, note that we now offer a user-friendly assay software called Real Time Design. It’s available on our website, free of charge. For information please inquiry at support@biosearchtech.com
Does Biosearch have software available for designing probes and primers?
In early fall of 2005 we launched our RealTimeDesign software for designing primers and probes. RealTimeDesign is free, easy to use and can be used directly through your web browser. The software offers a choice of two modes, an express mode with pre-set parameters and custom mode in which the parameters can be adjusted by the researcher. You can access our RealTimeDesign software by visiting www.qpcrdesign.com
What are the differences between the parameter settings in your primer and probe design software (RealTimeDesign)?
The Most Restrictive parameter set dictates very stringent guidelines for assay design. The resulting assays propose oligo sequences with tight melting temperatures, short amplicon lengths, and no stable mis- alignments within and between the primers. While the robust PCR reaction can tolerate rule breaking without performance impact, this parameter set errs on the conservative side of caution.
The Less Restrictive parameter set continues to adhere to the strict dogma of Dual-labeled design. While, the rules governing primer lengths and mis-alignments are slightly relaxed, there is no perceptible impact upon amplification performance. Rules demanding the close proximity between probe and primer continue to be enforced.
The Least Restrictive parameter set permits a maximum amplicon length of 100 bases, a maximum probe length of 30 bases, primers that differ in their melting temperatures by 1.5 degrees, and probes that anneal 20 bases distant from the forward primer. This parameter set continues to favor primers with a more stable 5' end, resulting in high-performing assays: amplification efficiencies > 95% across 7 orders of magnitude of starting copy number.
The Primer-Dimer Partition offers the same scrutiny as the Less Restrictive parameter set, but offers a more sophisticated method to screen against primer-dimers. This increased sophistication comes at an increased computational cost, as RealTimeDesign identifies and weighs ALL mis-alignments and secondary structures. These are then expressed within an equilibrium to determine the overall propensity to form ANY mis-hybridization, rather than simply the most significant.
How is the Tm calculated in Biosearch’s Real Time Design Software?
Our design software, RealTimeDesign (RTD) calculates TM’s using the unified nearest neighbor parameters established by Santa Lucia. Tm’s calculated on RTD may have discrepancies when compared to other design software's provided by ABI, Exiqon, and IDT. RealTimeDesign’s Tms are in very good agreement with the software programs provided by John SantaLucia and Michael Zuker. These programs are available for free over the internet:
SantaLucia: http://ozone3.chem.wayne.edu
Zuker: http://www.bioinfo.rpi.edu/applications/hybrid/
Is there a formula for calculating the efficiencies of singleplex PCR reactions?
For a singleplex reaction the efficiency of PCR is calculated as follows:
Efficiency= 10^(-1/slope) - 1
(slope is derived from a graph of Ct values vs. log starting concentration) A slope of -3.32 indicates an amplification efficiency of 100%
