REAL-TIME AMPLIFICATION ON THE ROTOR-GENE

Transcription

1 REAL-TIME AMPLIFICATION ON THE ROTOR-GENE Overview of the chemistries and optimizations 1

2 Table of Contents 1 Introduction Real-Time Detection of Amplification Products Real-Time Chemistries Intercalating dye (Sybr-Green I) Dual-Labeled Probes FRET Probe System Amplifluor TM Amplifluor TM SNP Genotyping System Amplifluor TM Direct Gene System Molecular Beacon Probes Scorpion probes Eclipse probes Light-up probes Optimizations for SG, Dual Labeled- and FRET Probes SYBR-GREEN I (SG) SG concentrations Primer concentrations MgCl 2 concentrations Temperature and Times Making primer dimers invisible Some further suggestions for testing or improving SG reactions Dual labeled probes Primer and probe design guidelines for dual labeled probes Guidelines for primers and probes Guidelines for probes Guidelines for primers The cycling parameters How to optimize primer and probe concentration Further suggestions regarding dual labeled probes Multiplex reactions with dual labeled probes Primer and probe design for multiplex reactions including optimization Mutation detection with dual labeled probes (allelic discrimination) Storage recommendations from Biosearch Technologies (USA) Analysis of quantitative data

3 3.3.1 Absolute quantitation Relative quantitation Relative quantitation using two standard curves Comparative Ct method A new mathematical model for relative quantitation analysis (Pfaffl) A new mathematical model for relative quantitation analysis (Liu & Saint) Housekeeping Genes RT-Amplification FRET probes Design of FRET probes and primers Probes for FRET analysis Primers for FRET analysis Primers and Probes for FRET analysis How to optimize FRET probes What is the advantage of using asymmetric primers? Before starting to optimize a FRET assay Optimization Storage of FRET Probes Summary of primer, probe design and related programs Primer design programs FRET probe design Probe design programs Related programs Trouble shooting rex/rea files Initial denaturation step Cycling profile Melt curve analysis Gain settings General setup conditions Experiment settings Other helpful tips Preventing contamination How to calculate gene copy numbers of the human genome More numbers

4 1 Introduction 1.1 Real-Time Detection of Amplification Products The reproducible quantitation of amplification products has long been the goal for many scientists and researchers. The traditional process requires the end-point analysis of amplification products via gel electrophoresis. This method allows for the identification of target and competitor product sizes, estimation of purity, and subjective measuring of band intensities. However, the reproducibility of amplification end products is highly variable due to limiting reagents, which compound the difficulties with this process. It is the exponential phase of amplification that provides the most useful and reproducible data. There is a quantitative relationship between the amount of starting target DNA and the amount of amplification product during the exponential phase of a cycling program. This is the very basis for Real-Time Amplification. Aided by the help of DNA intercalating dyes and probe specific chemistries, the study of the amplification process has improved by quantum leaps as a result of real-time detection. Today s real-time instruments are comprised of a fluorometer and a thermal cycler for the detection of fluorescence during the cycling process. A computer that communicates with the real-time machine collects fluorescence data. This data is displayed in a graphical format through software developed for real-time analysis. Figure 1: The raw data is plotted on a graph of fluorescence vs. cycle number. 4

5 Fluorescent data is collected at least once during each cycle of amplification allowing for real-time monitoring of amplification. A user is able to determine which samples are amplifying on a cycle-by-cycle basis. This instant data allows them to see how individual samples amplify in relation to known standards, positive controls and negative controls. Not only is the user able to monitor the whole reaction during the amplification process, but they can truly optimize their protocols based on the information they receive. This leads to increased sensitivity, specificity and efficiency. After raw data is collected, the analysis can begin. The software for the real-time instrument normalizes the data to account for differences in background fluorescence. Once normalization is complete, a threshold level can be set. This is the level at which fluorescence data is analyzed. Figure 2: The normalized data is plotted on a log scale graph of fluorescence vs. cycle number. The number of cycles it takes for a sample to reach the threshold level is the Ct-value (threshold cycle). The threshold is set at a level where the rate of amplification is the greatest during the exponential phase. This allows for the most accurate and reproducible results. If standards with corresponding concentrations are run, a linear regression analysis produces a standard curve from which the concentration of unknown samples can be determined. 5

6 2 Real-Time Chemistries There are several types of detection chemistries that are being used for real-time amplification. They include: 2.1 Intercalating dye (Sybr-Green I) Intercalating dyes, such as Sybr-Green I, bind to double stranded DNA. Figure 3: Sybr-Green I (SG) a) SG (S) will not bind to ssdna and the intensity of fluorescent signal is low. b) SG (S) binds to dsdna; the fluorescent signal intensity (E) increases. SG is a fluorescent dye that binds to the minor groove of the DNA double helix. The unbound dye exhibits little fluorescence in solution, but upon binding to double-stranded DNA the fluorescence is enhanced. This is utilized in real-time amplification. As DNA is amplified during an amplification reaction, the dye binds to amplified products and the fluorescent signal is increased. The increase is analyzed against the background fluorescence level. Multiple molecules of fluorescent dye bind to the dsdna based on amplicon length. Intercalating dyes can simply be added to the amplification reaction tubes along with all the other traditional amplification components: water, buffer, MgCl 2, dntp s, Taq-Polymerase, primers and template. This is an easy and cost effective approach to real-time detection, as it does not require the design of sequence specific probes and new primer sets. Intercalating dyes are not sequence specific and bind to any dsdna including nonspecific products and primer dimers. Therefore, it is necessary to differentiate between target and artifact signals. Intercalating dyes allow for the melting of amplification products at the end of a run. This is called the melt curve analysis. During the melt curve, the real-time machine continuously monitors the fluorescence of each sample as it is slowly heated from a temperature below the melting point of the products to a temperature above the melting point of the products. The user sets this range. Amplification products will melt at different temperatures based on their lengths and G/C content. As products melt, a decrease in fluorescence is realized and measured by the 6

7 instrument. By taking the differential of the melt curve, the melting peaks can be calculated. The melting peaks reflect the products amplified during the reaction. These peaks are analogous to the bands on an electrophoresis gel. Therefore, it is the melting curve data that allows for the qualitative monitoring of products at the end of a SG run. 2.2 Dual-Labeled Probes Dual labeled probes are short oligos with a fluorescent reporting dye attached to the 5 - end and a quencher molecule attached to the 3 -end. Because the probes are only bp long, the reporter dye and quencher are in close proximity to each other and little fluorescence is detected. During the cycling process, Taq DNA polymerase extends from each primer. The DNA polymerase has an exonuclease activity that cleaves the downstream probe as it extends. As the probe is degraded, the reporter dye is separated from the quencher. Figure 4: Dual labeled probes a) Energy (E) emitted by the donor (D) fluorophore is absorbed/quenched by the acceptor (A) fluorophore. b) The polymerase exonuclease activity separates the D fluorophore from the A fluorophore by hydrolysis resulting in an increase in fluorescent signal. With every cycle of amplification an increase in reporting dye is detected by the real-time instrument due to the cleavage of the probes. Because the reporting dye is cleaved, melt curve analysis is not possible. Dual labeled probes offer higher specificity compared to intercalating dyes. There are two reasons for this. First, dual labeled probes are sequence specific and only bind to complimentary regions. The second reason is that the dual labeled probe for each amplified copy releases only one molecule of fluorescent dye. Due to the increased specificity of dual labeled probes this detection method works well for low copy number templates. 7

8 2.3 FRET Probe System FRET probes rely on the transfer of energy from one fluorescent dye to another. Two separate sequence specific oligos are fluorescently labeled. The upstream probe has a donor molecule on the 3 -end and the downstream probe has an acceptor molecule on the 5 -end. The probes are designed so that they hybridize adjacently to each other on the target sequence and bring the donor and acceptor fluorophores in close proximity (see figure 5). Once the probes are hybridized, the donor and acceptor fluorescent molecules are in close proximity to one another. This allows for transfer of energy from the donor to the acceptor fluorophore, which emits a signal of a different wavelength. Either the decrease in the fluorescence of the donor or the increase in fluorescence of the acceptor can then be detected. Therefore, only when both probes are bound is fluorescence detectable. FRET probes do allow for melt curve analysis. They are extremely useful for genotyping, SNP detection, and other mutation detections. Figure 5: FRET Probes a) Resonance energy (E) transfer is low when the probes are not hybridized. b) Hybridization of the probes brings the donor (D) and acceptor (A) fluorophores into close proximity resulting in increased resonance energy transfer. Figure 6: Melt curve analysis 8

9 2.4 Amplifluor TM Amplifluor TM SNP Genotyping System Intergen Discovery Products introduced a fast, simple and reliable method for genotyping single nucleotide polymorphisms (SNP). This system employs an exclusive dual signal process that uses two fluorescently-labeled Amplifluor primers to distinguish heterozygous from homozygous alleles. This method is based on multiplex allele-specific PCR using two Amplifluor primers labeled with either fluorescein (FAM) or sulforhodamine (SG) for detection. Discrimination of alleles in a single closedtube is achieved with a pair of allele-specific pair of primers having different tail sequences. Depending on the genotype of the sample, a mixed signal of FAM and SR fluorescence is observed for a heterozygote while a single signal (either fluorescein or sulforhodamine) is observed for a homozygote. Figure 7: Assay scheme. The reaction steps for an A/G SNP are shown. (FL) fluorescein, (SR) Sulforhodamine 9

10 Principle: Profile on the Rotor-Gene Denature Cycling (40 repeats) 95ºc, 180 secs Step 1; 95ºc, hold 20 secs Step 2; 55ºc, hold 20 secs, acquiring to Cycling A (FAM, SR) Step 3; 72ºc, hold 20 secs FAM: ex: 470 nm/ det 510 nm SR: ex: 585 nm/ det 660 nm Sulforhodamine (SR) is excited at 585nm and detetced at 660nm. A 4-Channel machine is therefore required to perform those runs. Figure 8: Allelic discrimination analysis on the Rotor-Gene Shown are the results of the Amplifluor SNP Genotyping System used on the Rotor- Gene. A dilution series for each genotype (CC, GC, GG) has been run and analyzed. The heterozygotes (green curves) come up in both channels, whereas the CC genotypes only come up in the FAM channel and the GG genotypes only come up in the SR channel. Using the allelic discrimination analysis in the Rotor-Gene software, all results are displayed in one channel. An additional table tells exactly the genotype of each sample. This newly developed Amplifluor SNP Genotyping System works very well together with the Rotor-Gene. 10

11 2.4.2 Amplifluor TM Direct Gene System Amplifluor TM Direct Gene Systems are based on molecular energy transfer from an excited fluorophore to an acceptor moiety resulting in quenching of fluorescent emission. Target-specific Amplifluor TM primers consist of a 5 intracomplementary sequence labeled with fluorophore (fluorescein) and an energy acceptor 4-(dimethylamine)azo benzene sulfonic acid (DABSYL). The sequence 3 to the hairpin is the target specific primer region. Unincorporated Amplifluor TM primers have a low fluorescence due to the close proximity of the quencher and the fluorophore. Further information could be obtained from: Figure 9: Principal of Direct-labeled Amplifluor Direct Gene System. During the first cycle, Amplifluor Primer 1 anneals to the specific first strand cdna target and is extended by Taq-Polymerase. During the second cycle, the Amplifluor Primer 1 extension product serves as a template for Primer 2 (reverse primer). As Primer 2 is extended by Taq-Polymerase, the Amplifluor hairpin primer is unfolded and a fluorescent signal is generated. The fluorescent signal generated during subsequent amplification cycles increases proportionally to the amount of amplified product. Note: the complementary DNA strand is not shown in the schematic until fluorescence is generated. 11

12 Figure 10 shows the quantitation data, the standard curve and the results in a table. Results: Excellent results have been obtained with the Amplifluor TM Direct Gene System. A dilution series has been run in duplicates starting from 5 x x 10 0 copies. The average difference between Ct-values of duplicates was 0.12, with the maximum separation being 0.36 (five copies) and a minimum of The standard curve shown in the left bottom graph shows an R-value of Molecular Beacon Probes A fourth detection method is Molecular Beacons (MB). These probes are based on a hairpin structure design with a reporter fluorescent dye on one end and a quencher molecule on the other end. The hairpin structure causes the MB probe to fold when not hybridized. This brings the reporter and quencher molecules in close proximity with little to no fluorescence being emitted. However, when the MB hybridizes to the template, the hairpin structure is broken and the reporter dye is no longer quenched. At this point, the real-time instrument detects fluorescence. A melt curve analysis is also possible with Molecular Beacons. 12

13 Figure 11: Molecular Beacon Probes The hairpin structure causes the MB to fold when not hybridized, bringing quencher and fluorophore dyes in close proximity causing quenching of fluorescence. When hybridized, the fluorophore and quencher are separated resulting in increased fluorescence. The number and types of applications that benefit from real-time detection technology are far reaching. Some of the beneficiaries include organism identification, diagnostic testing, gene expression studies, mutation detection, SNP detection, genotyping and micro array confirmation. Each of the different chemistries provides the user with certain advantages depending on the application. It should be noted that there are other isothermal amplification systems for the detection of nucleic acids based on fluorescent chemistries that have also benefited from real-time technology. 2.6 Scorpion probes Scorpion probes differ from the specific detection methods discussed so far. As opposed to dual-labeled probes or molecular beacons, the Scorpions mechanism of probing is intramolecular. Their structure is as follows: the hairpin loop is linked to the 5 end of a primer via a PCR stopper. After extension of the primer during the amplification, the specific probe sequence is able to bind to its complement within the same strand of DNA. This hybridization event opens the hairpin loop so that fluorescence is no longer quenched and an increase in signal is observed. Unimolecular probing is kinetically favorable and efficient. Covalent attachment of the probe to the target amplicon ensures that each probe has a target in the near vicinity. Enzymatic cleavage is not required, thereby reducing the time needed for signaling compared to dual labeled probes, which must bind and be cleaved before an increase in fluorescence is observed. Scorpions primers have successfully been used for mutation detection and quantification, having the specificity and melt curve analysis of the Molecular Beacons with additional speed and efficiency. 13

14 2.7 Eclipse probes A new probe system that uses the minor groove binder (MGB) technology is Eclipse. These are short linear probes that have the minor groove binder and quencher on the 5 end and fluorophore on the 3 end. This is the opposite orientation to dual labeled probes and it is thought that the minor groove binder prevents the exonuclease activity of the Taq-Polymerase from cleaving the probe. Quenching occurs when the random coiling of the probe in the free form brings the quencher and fluorophore close to one another. The probe is straightened out when bound to its target and quenching is decreased, leading to an increase in fluorescent signal. The minor groove binder works together with the quencher to improve quenching, reducing the background level of fluorescence. As with minor groove binder dual labeled probes, short probes can be used and allelic discrimination is good. Sensitivity tests have shown that the probes can detect down to a few copies, when asymmetric amplification is used, and that amplification is linear over a wide range of concentrations. These properties make Eclipse probes suitable for quantitative and qualitative amplification. For more details see MGB probes can be designed using the MGB Eclipse Design Software: Light-up probes The light-up probe is a peptide nucleic acid (PNA) oligonucleotide to which the asymmetric cyanine dye thiazole orange (TO) is tethered. It combines the excellent hybridization properties of PNA and the large fluorescence enhancement of TO upon binding to DNA. When the PNA hybridizes to target DNA, the dye binds and becomes fluorescent. Free probes have low fluorescence, which may increase almost 50-fold upon hybridization to complementary nucleic acid. * O B * O B DNA Charge -1 - O O P O O * N O H N * PNA Charge 0 Figure 12: The graph above shows the chemical structures of DNA and PNA. 14

15 3 Optimizations for SG, Dual Labeled- and FRET Probes 3.1 SYBR-GREEN I (SG) To be able to optimize a SG reaction it is necessary to pre-optimize the conditions of an amplification reaction (MgCl 2, dntp, Taq-Polymerase concentrations etc). The result should be one single band on an agarose gel. The optimization of a SG assay basically depends upon four factors: SG concentration (3.1.1.) Primer concentration (3.1.2.) MgCl 2 -concentrations (3.1.3.) Temperature and Times (3.1.4.) SG concentrations To obtain an optimal SG concentration, a dilution series containing different dilutions of SG should be performed. As outlined above (2.1.), the principle of SG is to intercalate into the dsdna which explains why SG might interfere with the amplification reaction. An excessively high SG concentration can lead to an inhibition of the reaction. A low SG concentration on the other hand may not provide enough SG to label the amplicon sufficiently. The optimal SG concentration therefore has to be a compromise. The recommended SG concentration could range between a final concentration of 1:5000 to 1:100,000. Frequently used dilutions range between 1:10,000 to 1:60,000. 1:30,000 Figure 13 a 1:50,000 1: :60,000 1:125,000 1:400,000 15

16 b c 1:30,000 1: :50,000 1:60,000 Figure 13 shows the results of a SG optimization assay. The concentration of SG ranged from 1:30,000 to 1:400,000. The raw data (a) shows that the first four dilutions (1:30,000, 1:40,000, 1:50,000, 1:60,000) exhibited the highest increase in fluorescence, whereas the higher dilutions (1:125,000, 1:250,000 1:300,000, 1:400,000) showed a decrease in fluorescence. The melt data (b) gave similar results. In figure (c) the quantitation data shows that the lowest Ct-value belongs to samples 1:40,000, 1:50,000 and 1:60,000. 1:50,000 was therefore chosen for further experiments. The aim of the optimization assay is to obtain a SG concentration that gives the lowest Ct-values for a given sample concentration. An optimized assay does not show any primer dimers. If primer dimers are produced, the SG concentration with the highest Ctvalue should be chosen for the assay, thus it is recommended to include NTCs in the optimization process. 16

17 3.1.2 Primer concentrations The optimization of primer concentrations should be performed using various concentrations of forward and reverse primers. The range of the primer concentration should be between 50nM and 300nM, however, exceptions are possible. This dilution series has to be performed with a fixed amount of DNA template, but unlike an optimization reaction with a dual labeled probe (see 3.2) should also be performed with a non template control (NTC). The optimal primer concentration for a given reaction with a DNA template should result in a low Ct-value with a high increase in fluorescence (5 to 50 times), whereas the same reaction without DNA (NTC) should have a high Ct-value. The fact that amplification occurs with a NTC is due to the formation of primer dimers or non-specific product. Higher concentrations of primers may result in the amplification of non-specific products. The melt curves for both reactions should also be obtained. For the reaction performed with DNA it should result in the formation of one single peak, with no additional peaks at lower melt temperatures. Additional peaks at lower melt temperatures could be due to excessive amount of primers in the reaction. The amount of primers should therefore be reduced. The choice to use purified versus non-purified primers for real-time applications ultimately depends on the individual reactions, as there may be variables that contribute to the degree of difficulty of the amplification. From a quality control aspect, desalted HPLC purified primers are recommended. There is more control on such variables as salt concentrations and presence of truncated primers. While primer purity may not influence amplification of the amplification product, it may however contribute to when the NTC's are first seen in the thermal cycling profile. Depending on the purity of the primers, this may contribute to larger variations in Ct-values. Certainly, purified primers provide better specificity and thus may provide increased sensitivity. This is especially important for SG reactions as non-specific product contributes to the SG signal in positive control samples MgCl 2 concentrations As outlined above, SG reactions detect all double stranded DNA. In contrast to reactions with dual-labeled probes, where non-specific products or primer dimers can be tolerated, in SG reactions, it is very important to amplify the specific product. This can be achieved by reducing the amount of MgCl 2 in the reaction. Several aspects of the amplification are affected by MgCl 2 concentration in a reaction. These include DNA polymerase activity, which can affect specificity. The dntps and templates bind magnesium and reduce the amount of free magnesium necessary for enzyme activity. Similarly, for each primer pair and template, optimal magnesium concentrations vary. Greater yields of amplification product can be achieved with higher concentration of free magnesium, albeit this can also increase non-specific amplification. An ideal way to determine optimal MgCl 2 concentration is a titration from 1.5mM to 5mM in 0.5mM increments. 17

18 3.1.4 Temperature and Times Another way of optimizing the reaction is to choose the optimal temperature for the amplification. The three important temperatures are: denature-, annealing- and extension. For the denature temperature we recommend 95 C, for the extension temperature it is generally recommended to use 72 C, although this could vary depending on what enzyme is used. Check the manufacturer s recommendations. The annealing temperature could have a dramatic influence on the reaction. To find out the optimal annealing temperature check with the company which synthesized the primers. Additional optimization might be required, which could be achieved by testing different annealing temperatures. Generally, the higher the temperature, the more specific the reaction. The times used in an amplification reaction are also important. Recommendations for the optimal extension times with SG are hard to predict. Genomic DNA is supposed to have a longer denature time than plasmid DNA, for a shorter amplicon it is sufficient to have shorter extension times compared to a longer amplicon. Given an amplification reaction with cdna and an amplicon length of bp we recommend the following profile: Figure 14 shows a typical temperature profile for a SG run. As mentioned above, this profile is a general suggestion and the annealing temperature has to be adjusted. In summery, to achieve an efficient SG amplification reaction, optimal SG, primer and MgCl 2 concentration as well as the temperature and the times of the profile need to be determined. 18

19 3.1.5 Making primer dimers invisible The following chapter describes how it is possible to obtain reproducible data on the Rotor-Gene, despite the formation of primers dimers. Although the above described optimization steps have to be performed, a feature within the Rotor-Gene software allows for the reduction of the amount of primer dimers seen in the raw data screen and the quantitation screen. Once the melt temperatures for the gene of interest and the primer dimers are determined, a fourth step in the temperature profile could be introduced. By pressing + in edit profile, simply add an additional temperature step to the profile and acquire data above the melting temperature of primer dimers but below the amplicon melting temperature. This additional step should be performed for 15sec. Additional formation of non-specific product is unlikely, since the temperature is too high. When data is acquired at both temperatures, it is possible to determine the differences of those results. Cycling A Cycling B primer dimers amplification products Figure 15: The red curves represent four different DNA concentrations, whereas the bluish samples represent the appropriate NTCs. In cycling A and B, data has been acquired at 72 C and 84 C, respectively. Whereas the NTC could still be observed in Cycling A, non-specific amplification cannot be seen in Cycling B. The melt curves show the DNA products on the right side and the NTCs on the left side. 19

20 3.1.6 Some further suggestions for testing or improving SG reactions The following citation is an excellent publication about optimization of Sybr-Green I on the Rotor-Gene. By simply optimizing the Sybr-Green I MasterMix the author's have been able to detect copies of mrna and dsdna Biotechniques 2002 Apr;32(4):790-2, Evaluation of a homemade SYBR green I reaction mixture for real-time PCR quantification of gene expression. Karsai A, Muller S, Platz S, Hauser MT. University of Agricultural Sciences, Vienna, Austria. Depending on the purity of the DNA inhibitors may prevent the amplification from working. One way to test for inhibitors of reactions, is to spike the reaction mix with a known target template and appropriate primers. In order to perform a melt curve after a SG run, introduce a hold at the same temperature the melt curve is started (app. 60 sec). Melt curves can be repeated. It is possible to re-run a melt curve the day after it was originally performed. Added is an address that lists several primer pairs tested previously with SG. A typical profile for a SG run is as follows: Denature: check the enzyme that is used for the reaction (2 min, 5 min, 10 min or 15 min) Cycling: 95 C/5-20sec, annealing temperature/20sec, 72 C/20sec Hold: 72 C/60sec, Melt: 72 C - 99 C, on Melt A (See for further details) Depending on the type of Rotor-Gene, data can be acquired in the following channels: Emission: 470nm detection: 510nm dual channel / multi channel Emission: 470nm detection: 585hp multi channel machine SG is originally supplied as a 10000x solution. It is recommended to aliquot the SG into 0.2ml/0.5ml tubes after the first thaw. This way, the SG seems to be quite stable as a 20x in 1x TE at -20 degree Celsius. Storing SG in water reduces the life time of SG, it should rather be stored in DMSO. Remember when storing in DMSO that the amplification conditions could change due to the slightly increased DMSO content. For further recommendation see the above citated publication. 20

21 The ability to detect low copy numbers increases with the yield and quality of the mrna and subsequent cdna. High quality dntp's can make a difference in amplifications, especially in the last 5 to 10 cycles. Impure dntp's may also contain contaminating dndp's or dnmp's. A source of SG is Molecular Probes. Melt curve analysis using SG, differentiates between a) the size of the product and b) the G:C content of the product. Single nucleotide polymorphisms cannot be detected. It is recommended to use HPLC purified primers. Although there may not be much of a difference regarding amplification of the amplicon, there could be a big difference regarding the NTC. The optimal length of an amplicon is bp, although assays could also be performed with longer amplicons. The melt temperature of an amplicon can vary and depends on various factors. The melt temperature can be influenced by the ph of a particular buffer, different Taq- Polymerases, SG and MgCl 2 concentrations and others. When comparing melt curves make sure you compare identical conditions. One of the major disadvantages with SG is the non-specific amplification of primer dimers. We recently tested a QuantiTect SYBR Green PCR Kit from Qiagen with exceptional results. Replicates have come up very close together and the NTC did not show up at all. This kit may be very useful when running low copy numbers with SG. A protocol for this assay is available on our Web page and can be found under Commercially Available Kits. Results have shown that the optimal Sybr-Green I concentration might be influenced by the DNA concentration. It could be of interest to run a lower and a higher DNA concentration when optimizing reactions with SG. 21

22 3.2 Dual labeled probes The optimization of a dual labeled probe assay is easier to perform than a SG optimization. The most important thing, which has to be optimized, is the primer/probe concentration and its ratio. The optimal MgCl 2 concentration is usually around 4 to 5mM (final concentration), although in commercially available Master Mixes, MgCl 2 concentrations up to 7.5mM (final concentration) can be found. One of the most critical factors is the design of fluorogenic probes. The guidelines below summarize some suggestions that should be considered when designing primers and a dual labeled probe Primer and probe design guidelines for dual labeled probes Characterize sequence of interest. Choose amplicon with minimal secondary structure. This is important, since secondary structures could effect the efficiency of the reaction. In any real-time application it is desirable to obtain a 100% efficiency of the amplification reaction. This means that each time a cycle is completed twice as many amplicons should be present in the reaction. If the secondary structure is thermodynamically more stable than the oligo target hybridization, hybridization of the target will be disfavored. Secondary structure could also prevent polymerase read through. To check for tandem repeats the following web page might be useful: A program to test for secondary structures is called mfold. Make sure to include the right salt conditions (~50mM), MgCl 2 ( mM) and annealing temperature (~60ºC?). All other setting are as per default Beware of any stem loops and secondary structure with a G value which is negative. They will affect your probe signal. If an amplicon secondary structure is unavoidable, the primer annealing temperature should be increased. Do a BLAST search or a similar analysis to determine the probe specificity: Guidelines for primers and probes 1) Select the probe first and design the primers as close as possible to the probe without overlapping it. 2) The length of the amplicon should not exceed 400 bp. Ideal would be between bp. The shorter the amplicon, the easier an efficient amplification is achieved. Shorter amplicons also increase the consistency of the assay. 3) Keep the GC content between 30 and 70%. G/C-rich sequences are susceptible to nonspecific interactions that may reduce reaction efficiency. 4) Avoid sequences of identical nucleotides, especially G's (no more than four G s in a row). Again, efficiency and reproducibility could be influenced. 5) Test primers and probes against each other to avoid dimer formation and stem loop structures. 6) It is advisable to keep primers and probes 17 bases or longer. The chance of finding a random primer binding sites becomes more significant, when probes shorter. 22

23 Guidelines for probes Design the probes before designing the primers. The T m of the probe should be between 68 and 70 degrees. If the probe design is done by eye, scan the gene of interest for G/C rich regions. Avoid probes with a guanine residue at the 5'-end. A G at the 5 -end of the probe has a quenching effect that could even be maintained after cleavage. Select the strand that gives the probe more C's than G bases. More G s than C s in the probe, seem to reduce the reaction efficiency. The probe should be as short as possible without being longer than 30 nucleotides. Check the probes for DNA folding and secondary structures. The last five bases on the 3 -end should contain no more than two C s and/or G s. It might lead to non-specific product formation Guidelines for primers 1) The T m for the primers should be between 58 and 60 C. This is important, since those assays are usually run at an annealing temperature of 60 C. This is where the required 5 exonuclease has its highest activity. Make sure that the Tm s of the primers are fairly similar. 2) The end of the primer (last five nucleotides) should not have more than 2 G s and C s. 3) Place primers as close as possible to the probe without overlapping The cycling parameters Once the probes and primers are designed, the cycling conditions are always identical. After the initial denaturation time (depending on the Taq-Polymerase), the assay needs to be run for 15 to 20 sec at 95 C and for 60 sec at 60 C. For some assays an extension of 45 sec seems to be enough. Data has to be acquired at annealing temperature How to optimize primer and probe concentration For an assay with a dual labeled probe, optimal performance is achieved by selecting the primer and probe concentration that provides the lowest Ct-value and the highest increase of fluorescence compared to the background. The primer concentrations could be optimized by spanning an initial concentration range of 25nM 900nM. There are several possibilities of how forward and reverse primer concentrations could be combined. The probe concentrations could be optimized by spanning an initial concentration range of 10nM - 250nM. 23

24 The following three pages give an example on how to setup such a dilution series. The below concentrations are just examples, other concentrations of primers and probes could be used. Make up a Master Mix containing everything except for the probe and primers. Master Mix: Stock Final µl Per Master Mix Concentration Concentration Reaction (x34) water X buffer 10x 1x MgCl 2 50 mm 3 mm dntps 2.5 mm 0.2 mm 2 68 Forward Primer * * 5 Reverse Primer * * 5 Probe # # 1 Taq-Polymerase 5U/µl 0.05U/µl DNA 1 34 Total 25 Find below an example of concentrations for a primer/probe dilution series: * Primer stock concentrations are 4.5, 1.5 and 0.25µM Primer final concentrations are 900, 300 and 50nM (See Pt 4 below) # Probe stock concentrations are 6.25, 3.75 and 1.25 µm Probe final concentrations are 250, 150 and 50nM (See Pt. 2 below) Divide the Master Mix into 3 x 140µl and add the 10µl of probe at concentrations of 6.25, 3.75 or 1.25µM. Aliquot 15µl of the three Master Mixes into reaction tubes. To the reaction tubes, add 5µl of each primer to give the final reaction concentrations. The primer stock concentrations are 4.5, 1.5 or 0.25µM. 24

25 water µl 10 x buffer 85 µl MgCl2 51 µl dntp 68 µl Taq 8.5 µl DNA 34 µl Figure 16 add 140 µl to each of the three tubes add 10 µl 6.25 µm probe add 10 µl 3.75 µm probe add 10 µl 1.25 µm probe add 15 µl to each of the nine tubes 4.5µM REVERSE 1.5µM PRIMER + 5µl 0.25µM 4.5µM 1.5µM 0.25µM 4.5µM 1.5µM 0.25µM 4.5µM 1.5µM 0.25µM + 5µl FORWARD PRIMER 25

26 Figure 17: a) Figure 17a shows the raw data of the above described dilution series. Three different probe concentrations and nine different primer concentrations were combined. b) Double-click the RGicon to see the rex file. primer&probe matrix.rex Figure 17b shows the quantitation data. The curve with the lowest Ct-value (arrow) was chosen for further analysis (brown curve on the left side)(primer&probe matrix.rex). Three probe concentrations should be combined with the nine forward and reverse primer concentrations. The 27 reactions should be run on the Rotor-Gene with the above 26

27 described cycling parameters. The result of such a run is shown in the Figure 17. a) shows the raw data and b) shows the data after quantitation. The curve with the lowest Ct value and the highest amplification should be chosen for further experiments. In our case the brown curve with a cycle number of approximately 31 was chosen for further analysis. Note that reactions with the highest fluorescence increase are not necessarily the highest increase above background. Also, the highest increase above background does not always give the lowest Ct-value Further suggestions regarding dual labeled probes A general guideline for primer and probe concentration is a final concentration between 200 and 250nM of probe and 800 to 1000nM of primers. Most assays will work with these concentrations, however, to find the optimal concentration, the above outlined procedure should be followed. Due to the variation in the predicted melt temperature of most primer and probe design programs, it could be useful to do the optimization on three different annealing temperatures (58 C, 60 C, 62 C). The concentrations of the primer may also influence the melt temperature. A high primer concentration may need an anneal temperature at approximately two degrees higher. Programs for designing primers and probes are as follows: Primer3 is a very reliable program. Although this program has several advantages, unfortunately there is no option to design probes with no G at the 5' end. A suggestion is to take off the G at the 5 end and check if the melt temperature is still 8 to 10 C above the melt temperatures of the primers Multiplex reactions with dual labeled probes Multiplex reactions can be performed on the Rotor-Gene due to its capability to detect multiple dyes. Up to four different dyes can be used in one reaction vessel. The recommended dyes used on the Rotor-Gene are FAM, JOE, ROX and Cy5 (for more details about fluorophores used on the Rotor-Gene see our FAQ). The most common multiplex assay application is relative quantitation of gene expression. One probe can be labeled with FAM for detection of a housekeeping gene and the other probes can be labeled with different fluorophores detecting one, two or three different genes in the one reaction vessel. Running four assays in a single tube reduces both the 27

28 running costs and the dependence on accurate pipetting when splitting a sample into two to four separate tubes. To generate an accurate multiplex assay, it is important to ensure that the amplification of one target does not dominate the other target. A domination of one target may prevent the less abundant one from amplifying efficiently. This can lead to inaccurate results or may even inhibit detection of the less abundant target completely. This could be avoided by limiting the concentration of the primers used to amplify the more abundant species. If it is not known which target is the less abundant, it should be determined by running both targets in separate tubes. To determine primer limiting concentrations, several reactions with a final concentration of 20nM to 100nM should be run. The aim is to reduce the increase in fluorescence over the background without influencing the Ct-value. A reaction with a dual-labeled probe needs to have a higher MgCl 2 concentration than traditional amplifications. The higher MgCl 2 concentration is required because the 5-3 exonuclease domain of the Taq-Polymerase prefers a high MgCl 2 concentration. Since the 5-3 exonuclease activity is needed for a dual labeled probe, a higher MgCl 2 (minimum 3.5mM, normally 5mM) is unavoidable Primer and probe design for multiplex reactions including optimization a) Probe and primer design The Tm of all primers should be within 1 C. The Tm of all probes should be within 1 C. Length of amplicons should be small and of similar size. A recommended length is under 100bp and within 5bp of each other. G/C content of the amplicon should be similar - typically within 2%. Avoid placing primers and probes in regions with high secondary structure. b) Consideration before starting Optimize primer and probe concentrations individually. Make sure that the reagent and thermal reaction conditions are kept constant for all targets. Choose a MgCl 2 concentration that is in excess. A final concentration of 5mM to 6mM is recommended. The dntp concentration should be in excess, typically 0.8mM for dntps that include dttp and 1mM for dntps that include dutp. Addition of of 6-8% glycerol can destabilize secondary structure and may help to optimize the assay. 28

29 c) Optimization process for multiplex reactions The amount of DNA used for an assay should guarantee a good signal; - typically around 100ng. For each primer/probe set use around 200nM of probe. Perform a primer optimization assay similar to the pipetting scheme shown in Figure 16, including NTCs. The assay could be performed with replicates. Select a primer concentration that gives the lowest Ct and the highest increase in fluorescence. If the target is a housekeeping gene or a high expresser, use the lowest Ct and the lowest increase in fluorescence. d) Primer Limitation If one target is present in much higher quantities than the other, perform a primer limitation of that target in order to prevent competition for the reagents. A target is considered to be in high abundance over another when the Ct values are greater than 5 Ct's apart when using the same initial starting template. Keep the ratio of the reagents the same, as determined in the primer optimization step and do two fold serial dilutions of the primer concentration. Choose the concentration that gives the lowest increase in fluorescence, but does not shift the Ct value. The information in chapter was mainly taken from a protocol compiled by Renee Horner ( Dual-labeled probe design checklist: b Amplicon length (ideal would be between 75 and 150 bp)? Both primers and probes should close together, but not overlapping. Probes close to the forward primer? Both primers should have the same Tm (between 58 and 60 degree)? Tm of probes ~10 C higher than Tm of primers? Probes and primers between bp? 20-80% G/C, try for ~50%? More C s than G s in probes? No G at the 5 end of probes (quenches fluorescence)? Last 5 bases at 3 end of probes have less than 3 G s and/or C s? Avoid stretches of nucleotides more than 3? Probe not complementary with primers? 29

30 3.2.3 Mutation detection with dual labeled probes (allelic discrimination) To be able to detect mutations with dual labeled probes, two probes with different colors need to be designed. One probe has to be designed to detect the wild type sequence and is labeled with FAM, whereas the other probe detects the mutation and is labeled with JOE. The difference between the probes is therefore usually only one bp. Since the difference in the probes is so minimal, it is recommended to use quite stringent reaction conditions. To analyze the data on the Rotor-Gene go to Analysis and chose Allelic discrimination. Using this analysis, the results of both channels are displayed on one screen. Curves with no markers are the result from CH1 (or FAM channel), curves with circles are the result from CH2 (or JOE channel). Up to four different probes (two mutations) can be analyzed together. After editing the genotype names and adjusting the Threshold above background, the results are displayed in a table below the graph. Figure 18 shows a typical result of such an assay. Figure 18 shows the analysis of six samples, two wild types (red), two mutations (blue) and two heterozygous (green). The red curves come up in CH1 (no markers), the blue curves in CH2 (circles) and the green curves come up in both Channels. The green curves have therefore been called heterozygous. If necessary, a quantitation of those samples could also be performed. 30

31 3.2.4 Storage recommendations from Biosearch Technologies (USA) Dual-labeled probes are shipped lyopholized and can be stored in this state at -80 C for at least one year. Before use, make a stock solution by resuspending the dry probe in aqueous solution to a known concentration and then aliquot into microvials and store at - 20 C. Only the required number of aliquoted microvials should be removed from the freezer, placed on ice and covered until needed. For daily use, probe stock solutions can be further diluted to an appropriate working concentration with HPLC grade water, TE buffer or 10mM Tris (ph 7.5). Aliquots can then be thawed for use as needed. To store probes long term in solution, 0.01% Tween 20 and 0.01% Gelatin should be added to the solution before storing at -80 C. Probes stored in this manner will be stable for one year or more without degradation. To ensure optimal activity and safeguard maximum performance, fluorescent probes should always be stored away from light. Sarstedt provides 2 ml screw capped amber tubes (Cat. No.: ). 31

32 3.3 Analysis of quantitative data There are basically two ways of analyzing quantitative data: a) Absolute Quantitation b) Relative Quantitation Depending on the application and the aim of the analysis, the researcher has to decide how to analyze quantitative data. The following two chapters will give a brief overview of the difference between both methods and how they could be used on the Rotor-Gene Absolute quantitation Absolute quantitation refers to an analysis where unknown samples are compared to a standard curve. A standard is a known DNA sample whose absolute concentration is known. There are many ongoing discussions of what kind of standard has to be used for a standard curve. The ideal would be a DNA sample that was extracted in the same way as the unknown samples. Unfortunately, this is not always possible. Alternatively, the amplicon being studied can be cloned, or a synthetic oligonucleotide used. There are several criteria for absolute standards. The standard must be amplified using the same primers as the gene of interest and must amplify with the same reaction efficiency. The standards must also be quantified accurately. This can be carried out by reading the absorbance at A260, although this does not distinguish between DNA and RNA, or by using a fluorescent nucleic acid stain such as RiboGreen. Once a copy number value has been obtained for a sample it must be normalized to something. Often number of cells or total RNA is used. This requires accurate measurement of input RNA, again either by reading the absorbance or using a nucleic acid stain. Be aware that the accuracy of the absolute quantitation assay is entirely dependent on the accuracy of the standards. There are several ways of analyzing absolute quantitation data on the Rotor-Gene. Beside the standard curve, the R-value, the slope, the intercept and the efficiency of the standard curve is displayed. It is possible to import a standard curve from a previous run. It is also possible to import a standard curve from a previous run and adjust it to a standard (or replicates of the same standard) in the current run. This function is based on the assumption that the slope is more reproducible than Y intercept. A single standard or a standard of replicates will therefore be sufficient for a standard curve. Using this method make sure that the slope is checked regularly, especially when changing reagents. In order to normalize data, the normalizer target is used to correct for variations in input template quantity. The normalizer is typically a housekeeping gene for expression studies or gene of known copy number for zygosity testing. In this method of analysis, the absolute quantities of the target of interest and the normalizer in an unknown sample are determined from the standard curve. In order to determine normalized quantity of the 32

33 target of interest in the unknown sample, the values obtained from the standard curve for the copies of target of interest and the copies of normalizer target are divided. It is possible to run several standard curves either in different channels or in the same channel. Running different standard curves in one channel is mainly for those users who would, for example like to analyze several different genes with SG or the same fluorophore. In the Rotor-Gene software, several different sets of standard curves can be used in the same run. Note: When data is compared over several experiments, the same threshold setting needs to be used. Set the threshold manually. Also remember that the standard curve can degrade. It is recommended to make up standard curves fresh. If unsure, compare old and freshly made up standard curves Relative quantitation The term relative quantitation is used when two or more genes are compared to each other with the result being a ratio. No absolute number is detected. An endogenous control or a housekeeping gene is normally compared to a gene of interest. There are several ways of doing relative quantitation: Relative quantitation using two standard curves The main difference between absolute and relative quantitation is that absolute quantitation uses a known input amount, eg copy numbers, which results in an output amount of copy numbers. Relative quantitation uses relative numbers (eg fold differences) where the results are given as a ratio. The result is the comparison between the ratios. In real-time relative RT-PCR, a standard curve is generated from a dilution series. The sample used for the standard curve is the reference sample. This sample can be a RNA sample, genomic DNA, cdna, or even a cloned DNA, as long as the amplification target (amplicon) is present. It is also important that the reaction efficiency of the reference sample reflects the reaction efficiency of the unknown sample. The units used to describe the dilution series are relative and based on the dilution factor of the standard curve. Units could be equivalent mass amounts (ng, mg etc) or simply fold differences. Typically, an endogenous control is included in the assay. Endogenous controls (or socalled housekeeping genes) are genes, which are not influenced by the treatment Example: if plants are exposed to different light sources or animals treated with different drugs, ideally the endogenous control should not be affected by that treatment in any way. The level of expression of the endogenous control should remain the same. The function of the endogenous control is to correct for sample to sample variations in RT- PCR efficiency, errors in sample quantitation and RNA (DNA) loading amounts. 33

34 A sample therefore has to be analyzed twice, firstly with the gene of interest and secondly with the endogenous control. Two standard curves have to be run to obtain two values (endogenous control and gene of interest) for the one sample. The samples can either be run in separate tubes or as a multiplex reaction. If a multiplex reaction is used the standard curve should also be run as a multiplex reaction due to possible changes in the reaction efficiency (vs a reaction with only one fluorophore). The relative values automatically calculated by using the standard curves are then divided by each other. For each experimental sample, the relative abundance value obtained (not the Ct-value) is divided by the value derived from the control sequence in the corresponding amplification. The analysis of such an experiment is shown below. A RT amplification of the htert and the GAPDH gene was performed using a standard curve for each gene. The values calculated for the unknown samples have been obtained from the columns "calculated concentration" in the Rotor-Gene software. Data was exported to Excel using the export function. No. Name unknown samples Type Calc. Conc. htert Calc. Conc. GAPDH Normalized htert / GAPDH Relative value if needed 1 No.1 htert Sample No 2 htert Sample No.3 htert Sample No.4 htert Sample No.5 htert Sample No.6 htert Sample No.7 htert Sample No.1 GAPDH Sample No 2 GAPDH Sample No.3 GAPDH Sample No.4 GAPDH Sample No.5 GAPDH Sample No.6 GAPDH Sample No.7 GAPDH Sample 6447 The normalized data has been obtained by dividing No.1 htert / No.1 GAPDH which resulted in a normalized value of No unit is given to this value. The ratios of all other samples have been obtained in the same way. If needed, values can then be compared to a so-called calibrator sample. A calibrator sample in general is a sample, which was not treated in any way. In our experiment 34

35 sample No.1 was arbitrarily designated as the calibrator sample (abundance set to 1x) and the normalized values for the remaining samples are expressed as x-fold of 1x. Relative quantitation using two standard curves is performed when the reaction efficiencies of the different targets involved in the assay are different. If the reaction efficiencies are similar (slope difference less than 0.1) than the comparative Ct method can be used. See below for further details Comparative Ct method This method enables relative quantitation of template and increases sample throughput by eliminating the need for standard curves when looking at expression levels relative to an active reference control (normalizer). For this method to be successful, the dynamic range of both the target and reference should be similar. A sensitive method to control this is to look at how Ct (the difference between the two Ct values of two amplifications for the same initial template amount) varies with template dilution. If the efficiencies of the two amplicons are approximately equal, the plot of log input amount versus Ct will have a nearly horizontal line (a slope of <0.1). This means that both amplifications perform equally efficiently across the range of initial template amounts. If the plot shows unequal efficiency, either the previously described standard curve method should be used for quantitation of gene expression or the reactions have to be optimized to obtain similar efficiencies. The dynamic range should be determined for both (1) minimum and maximum concentrations of the targets for which the results are accurate and (2) minimum and maximum ratios of two gene quantities for which the results are accurate. The following example shows a so-called validation experiment performed on the Rotor- Gene (Figure 19). Three replicates of each dilution using standards for the gene of interest and the housekeeping gene in two different channels have been amplified. The serial dilution should reflect the range of expression you anticipate. If uncertain, do a dilution series over several orders of magnitude. The target that is used should be the sample you plan on using as your calibrator sample. The relative quantity of your unknowns will be calculated in terms of this sample. The numbers of standard curves to be run equals the number of targets you are looking at. FAM Gene of interest JOE housekeeper Figure 19 35

36 The table below shows the results of both dilution series. The replicates in each channel were averaged and the differences calculated (last column). Average Replicates Log of Given Conc. Difference FAM JOE GOI-HKG 100 fold GOI-HKG 50 fold GOI-HKG 20 fold GOI-HKG 5 fold GOI-HKG 2 fold GOI-HKG 1 fold GOI = Gene Of Interest, HKG = housekeepig gene The slope and the intercept of the curve were calculated in Excel: y = x where m is the slope and b is the intercept. Given that the slope in the example is < (+/-) 0.1 the assay can be analyzed using the comparative Ct-method. Running the target and endogenous control amplifications in separate tubes and using the standard curve method requires the least amount of optimization and validation. However, running the reactions as a multiplex reaction is more reliable, since the same amount of DNA (same tube) is quantified through two different probes. For data analysis, one of the samples has to be chosen as a normalizer for each comparison to be made (eg wild type, untreated etc). a) If replicates are run, which is recommended, calculate the average first. This gives the average Ct-value for each particular sample. b) Calculate the difference between the gene of interest and the housekeeping gene. The result is the Ct-value [ Ct = Ct (target) - Ct (normalizer)]. c) Calculate the difference between the normalizer and a sample. The result will be the Ct-value. d) The last step in quantitation is to transform these values to absolute values. The formula for the comparative expression level = 2 - Ct. It is a good idea to set up an Excel spread sheet with all the formulas needed. Export the data from the Rotor-Gene results tables into Excel and the data will be calculated automatically. 36

37 ( Ct) = [meanct "FAM" (calibrator sample) meanct "JOE" (calibrator sample)] [meanct "FAM" (patient sample) meanct "JOE" (patient sample)] The above example uses different fluorophores for each gene. This is not absolutely necessary, as two different Sybr-Green I reactions could be analyzed in the same way. For further details on such an assay performed on the Rotor-Gene, see the following publication: Genetic Testing, 2003 (in press): A Rapid and Definitive Test for CMT1A/HNPP Using Multiplexed Real-Time PCR. Lorentzos P, Kaiser T, Kennerson ML, Nicholson GA. Molecular Medicine Laboratory, Clinical Sciences Building, Concord Hospital, Concord, New South Wales, 2139, Australia. The following web address describes this method in detail: A new mathematical model for relative quantitation analysis (Pfaffl) A further method of analyzing quantitative data is a new mathematical model recently published by Pfaffl (2001). This method requires neither standard curves to be run in the experiment (compare ), nor do the reaction efficiencies of the reactions to be similar (compare ). Although the method is also based on reaction efficiencies, only the crossing point deviations of unknown samples versus control samples are needed to exactly describe the relationship between several genes. This makes data analysis very simple. The Pfaffl analysis method is described in the following formula: R = (E target ) CP (control sample) target (E reference ) CP reference (control sample) 37

38 R describes the relative expression ratio between the target and the reference gene. It is the ratio between the housekeeper and the gene of interest of a particular sample. In the final analysis this ratio will be compared to other ratios of other samples. The ratios (R s) of the samples will therefore give the relationship between the relative expressions of different samples. E is the reaction efficiency of the target or the reference. To obtain the reaction efficiency of a given sample, a dilution series (standard curve) needs to be run before the actual experiments. Once the standard curve is obtained, go into the quantitation analysis window of the Rotor-Gene software and use the auto-find threshold function to determine the line of best fit. If the threshold is set manually, make sure that it is set in the exponential phase of the reaction. The reaction efficiency of the run is given in the report. The reaction efficiency in the Rotor-Gene software is calculated as follows: = (10 (-1/slope) ) 1. Only the exponential phase of the amplification reaction is used for the determination of the reaction efficiency. (Other software packages calculate the efficiency using the formula 10 (-1/slope). We refer to this value as the amplification value. The optimal amplification value = 2.) The number obtained from the report is used for the above formula. The optimal reaction efficiency of a reaction is E = 1. The CP is the CP deviation of a control minus a sample, either for the target or the reference. The definition of the CP-value is the crossing point of the threshold with the samples. It therefore equals the Ct-value used in the Rotor-Gene software. To obtain the Ct-values simply move the threshold manually, little above the take-off points of the reactions. To better see the take-off points of a reaction, switch to linear scale in the quantitation window. If replicates for the analysis are run, simply use the average of the controls and samples for the calculation. Exponential phase Figure 20: The above experiment shows the linear scale of a quantitation result with standards run in duplicates. A dilution series was performed with the following dilutions: 20x, 10x, 4x, 2x, and 1x. mean Ct between blue and orange samples Using the zoom function of the software the meanct-value between the duplicates is shown. This is the value used in the above described formula. 38

39 The advantages of the "Pfaffl method" are obvious. It only depends on the Ct and the reaction efficiency. Several genes, as well as results for multiplex reactions could be analyzed and calculated. More details about the above described method as well as the original paper are found under the following web address: This web page also contains a software package that automatically calculates the desired ratios A new mathematical model for relative quantitation analysis (Liu & Saint) This new quantitative method of data analysis is described in the following publications: - Weihong Liu and David Saint. Validation of a quantitative method for real time PCR kinetics. Biochemical and Biophysical Research Communications 294 (2002) Weihong Liu and David Saint. A new quantitative method of real time reverse transcription polymerase chain reaction assay based on simulation of Polymerase Chain Reaction kinetics. Analytical Biochemistry 302, (2002). The analysis methods described in rely on standard curves, although the comparative Ct-method and the "Pfaffl" method only require initial runs using standard curves. Reaction efficiencies are calculated from these standard curves and used for analysis. The reaction efficiency is therefore an average value of several samples used in a standard curve. It doesn't take into account that the reaction efficiencies could vary over different runs or even from sample to sample within the same run. With the quantitation method described by Liu and Saint the reaction efficiency of each sample is calculated, which makes this method very accurate. In addition, there is no need for a standard curve or a validation experiment Housekeeping Genes Housekeeping genes are present in all nucleated cell types since they are necessary for basic cell survival. The mrna synthesis of these genes is considered to be stable in various tissues, even under experimental treatment. Quantitative gene expression assays are typically referenced to an endogenous (internal) control gene to account for differences in DNA/RNA load. The amount of RNA assayed may fluctuate due to differences in tissue mass, cell number, experimental treatment or RNA extraction efficiency. The following publication is an excellent example of how to test if your housekeeping gene suits your needs. Schmittgen TD, Zakrajsek BA. Effect of experimental treatment on housekeeping gene expression: validation by real-time, quantitative RT-PCR. J Biochem Biophys Methods 2000 Nov 20;46(1-2): There are several housekeeping genes which are commonly used: b-actin, GAPDH, cyclophilin, 18S rrna, 28S rrna (reflects natural rrna degradation more than 18S), phosphoglycerokinase, b 2 -microglobulin, b-glucronidase, hypoxanthine ribosyl transferase, transferrin receptor and others. Be aware that housekeeping genes or internal controls may be influenced by assay conditions. Proper validation of internal control genes is necessary when designing quantitative gene-expression studies. 39

40 3.3.4 RT-Amplification Many different applications using a RT step before the actual amplification have been performed on the Rotor-Gene. Both SG and dual labeled probes have been used for this purpose. One of the most recent reviews regarding Absolute quantification of mrna using real-time reverse transcription polymerase chain reaction assays is written by S. A. Bustin. ABSTRACT The reverse transcription polymerase chain reaction (RT-PCR) is the most sensitive method for the detection of low-abundance mrna, often obtained from limited tissue samples. However, it is a complex technique, there are substantial problems associated with its true sensitivity, reproducibility and specificity and, as a quantitative method, it suffers from the problems inherent in PCR. The recent introduction of fluorescence-based kinetic RT-PCR procedures significantly simplifies the process of producing reproducible quantification of mrnas and promises to overcome these limitations. Nevertheless, their successful application depends on a clear understanding of the practical problems, and careful experimental design, application and validation remain essential for accurate quantitative measurements of transcription. This review discusses the technical aspects involved, contrasts conventional and kinetic RT-PCR methods for quantitating gene expression and compares the dierent kinetic RT-PCR systems. It illustrates the usefulness of these assays by demonstrating the significantly dierent levels of transcription between individuals of the housekeeping gene family, glyceraldehyde-3-phosphate-dehydrogenase (GAPDH). Journal of Molecular Endocrinology (2000) 25, The full text of this review can be downloaded in Acrobat PDF format: 40

41 3.4 FRET probes Design of FRET probes and primers To design FRET probes and primers, several programs are available. A recommended program for designing FRET probes can be found under the following address: Figure 21: Meltcalc is an easy to use program. FRET probe design is available within minutes Probes for FRET analysis A good rule of thumb is that the 5 labeled probe (eg. Cy5, BHQ) should have at least a five to seven degree higher melt temperature than the 3 labeled probe (eg. FAM, JOE). Typically, Tm differences between the donor and the acceptor probe can be 10 degree or greater. It is a good idea to start the design with a longer probe sequence and cut it down until the Tm is optimal. If there are stretches of Gs and Cs, try to use these for the 5 labeled probe, as a higher melt temperature is desired. An ideal Tm for the donor probe would be ~ 55 C and for the acceptor probe ~ 65 C but this certainly depends on the sequence. More important is the Tm difference between the probes. Design the probe against the mutant allele. This way you make sure that you distinguish between the mutant and everything else, which will be usually wild type. In some minor cases a second mutation can occur in the same region. Avoid the mismatch T:G. The probe should be design on the opposite strand. The 3 labeled probe (eg. FAM, JOE) should overlap the mutation. The distance between the probes should be between two and 5 bp s. The probes should not be longer than ~25 bp. 41

42 Primers for FRET analysis The recommended length of the amplicon should be bp. Primers should have a Tm between 55 C and 60 C but below the Tm's of the probes. A recommend G:C contents of primers should be between 35 and 65%. The length of the primers should be between bp s. A recommended primer design program (free of charge) is found at: The meltcalc software does not provide primer design capabilities. Primer Tm's should therefore be checked with the meltcalc software. Minor differences between programs can occur Primers and Probes for FRET analysis Check primer and probe sequences for hairpin or primer dimer formation. This is important since it could reduce the efficiency of the assay quite significantly How to optimize FRET probes What is the advantage of using asymmetric primers? This technique is utilized when it is desirable to amplify more of one strand relative to the other. Doing this minimizes binding competition between the probes and the strand that shares their sequence. This facilitates the binding of more probe, increases FRET and produces a higher signal. This is why the ratio of primers has to be optimized Before starting to optimize a FRET assay Check the primer and probe concentrations on a Spectrophotometer. Test purity of primers and probes by doing an amplification in a Thermo Cycler or on the Rotor-Gene. Make up a Master Mix with all reagents needed for an amplification including DNA but without primers and probes. Set up the following reactions: only forward primer (no reverse primer and no probes), only reverse primer (no forward primer and no probes), only donor probe (no primers and no acceptor probe), only acceptor probe (no primers and no donor probe), forward/reverse (no probes), forward/donor probe (no reverse primer no acceptor probe), forward/acceptor probe (no reverse primer no donor probe), reverse/donor probe (no forward primer no acceptor probe), reverse/acceptor probe (no forward primer no donor probe), donor probe / acceptor probe (no primers). 42

43 Analyze on an agarose gel. The result of this assay should be a single band in only one reaction (FOR/REV). If any other combination results in a single product (for example from REV/Cy5), it could be that the Cy5 probe is not blocked properly with a Phosphate group at the 3 -end and acts as a primer. This could lead to false results. Check with the company producing the oligo s. Depending on how many cycles are run, the other combinations should result in either no band or a smear Optimization Once it is determined that the primers and probes produce the correct product size and are not contaminated, the following steps should be performed. Initially choose a homozygote DNA sample with a concentration of 10 to 100ng. Test the reactions with a heterozygote sample when the assay is optimized. Choose a MgCl 2 concentration optimal for the primers, which is usually obtained from the supplier of the primers. If the concentration is not known start with a final concentration of approximately 3mM. MgCl 2 can seriously influence the assay. A MgCl 2 dilution series can be performed once the assay is nearly optimized. Choose an annealing temperature appropriate for your primers. Keep the probes at a final concentration of 400nM each. Check the orientation of the probes in relation to the primers. Do a dilution series with one primer. If the forward primer and the probes are designed in the same orientation, keep the reverse primer at 400nM and do forward primer dilution series from 400nM down to 4nM. This should be carried out in approximately steps. The raw data results (Figure 22) should be curves decreasing in the FAM channel and if Cy5 is attached to the acceptor probe curves increasing in a channel setup with the following filters: 470nm - 610hp. Figure 22: Shown are the raw data of a FRET analysis using probes labeled 3' with FAM and 5' with Cy5. Data have been acquired on FAM (right) and a channel setup with the following filters: 470nm - 610hp (high-pass)(left). If a heterozygous sample has been used two distinguishable peaks should be seen after the melt curve analysis (one peak for a homozygous sample). If you do not see peak(s) in 43