Performing a realtime RTPCR experiment

In general, any experiment involving the PCR that begins with converting an RNA template to cDNA will be more challenging than those that begin with DNA templates. For that reason, the following discussion is centered on RT-PCR. However, the basic principles and discussion concerning PCR below would still apply to those starting with DNA.

Undoubtedly, the most important part of any RT-PCR experiment is the isolation and purification of the RNA template before the experiment is initiated. There are many methods for the isolation of nucleic acids. Most are variants on a few basic isolation themes tailored to meet the special

Figure 1.10

Graphical representation showing the mechanism of action for an hybridization probe pair. Two oligo probes bearing a single dye each, one with a fluorescein dye at the 3' end and the other with a rhodamine dye at the 5' end. When the two oligos anneal to a complementary template, the fluorescein dye is excited by the light source in the instrument and transfers its energy to the rhodamine dye via FRET. FRET can only occur when the two dyes are in close proximity. The instrument is set to detect the rhodamime signal.

needs of the particular organism and/or target nucleic acid of interest (Sambrook and Russell, 2001; Ausubel, 2001). There are a large number of kits on the market for RNA or DNA isolation from different organisms, tissue or cellular sources. One important factor is to minimize the amount of DNA that is carried over with the RNA during isolation. This is important because not all real-time assays cross exon junctions. Even if the assay does cross an exon junction, pseudogenes may be amplified by the assay anyway as they are spliced but imperfect copies of the original transcript reinserted into the genome. For many transcripts, there are one or more pseudogene copies in the mammalian genome. Of course for many organisms, e.g., those of viral and microbial origin, no exons exist. One sure way around this problem in all cases is to DNase I treat each total RNA preparation. Again, there are kits available for this process but it is not that hard to perform this task yourself, as described in Protocol 1.1. Be aware that DNase I treatment on a column as part of the RNA isolation procedure will not be sufficient if your real-time assay has an amplicon shorter than 100 bases. For the larger amplicons, found in SYBR® Green I or other primer-based assays that are 200-250 bases in length, treatment with DNase I on the column will suffice.

RNA from tissue culture cells can be isolated using most available methods. Isolating RNA from tissues, however, can be more problematic. Again potential complications are dependent upon the tissue involved. We have found that homogenization of the tissue using a Polytron or bead-beater device in a phenol-based guanidinium solution (Chomczynski, 1993) works best for rapid degradation of the cellular structure and inhibition of endogenous RNases. Regardless of the source, it is critical to minimize not only DNA but also protein carryover during RNA isolation. The proteins that most commonly carry over in any nucleic acid isolation procedure are nucleic acid binding proteins. These DNA or RNA binding proteins can be disruptive to the RT reaction and in particular, the following PCR. We have found that binding the RNA to a column matrix and washing away protein and other PCR contaminants results in the cleanest RNA for real-time PCR. Column-based kits are available from many manufacturers. However, we have developed a method that couples the superior disruption of a phenol-based guanidinium solution with a column-based final purification of the RNA (Protocol 1.2).

There are examples where this standard isolation method will not suffice. Many plants and some microbes contain carbohydrates that will co-purify with nucleic acids. In these instances, methods based on CTAB (Cetyltrimethylammonium Bromide) can be used to isolate RNA or DNA (Liao et al., 2004). Another method for further purifying RNA is precipitation using LiCl (Lithium Chloride) instead of salt plus alcohol (Liao et al., 2004). Isolation of RNA from small sample sizes, such as those from laser capture microscopy, can require special handling (Gjerdrum et al., 2004). Again, there are several kits on the market for this purpose.

There is nothing about the reactants used in setting up an RT reaction and following PCR that is substantially different for analysis by real-time PCR from those reactions designed for analysis using a gel. The only differences are the concentrations of some of the reactants and the inclusion of a fluorescent moiety for detection in a real-time reaction. There are two critical elements to setting up any real-time PCR experiment, the uniformity of the components in the reaction and the accuracy and care with which each component, particularly the sample, is added to the reaction.

It is important that all the components required for the reaction be uniform from well to well if the results from multiple wells are to be compared. To accomplish this goal, the concept of a master mix is critical. Master mixes are, as the name implies, a combination of all the reactants required for the RT reaction or the PCR minus the variable in the experiment, usually the RNA, cDNA or DNA sample. Thus, all the reactants, primers and probe (if used) are combined, mixed well and dispensed in an equal volume into all the wells of a plate ensuring that each reaction is starting with the same concentration of the basic reactants.

There are many commercially available kits on the market for both the RT step and the subsequent PCR as either one-step or two-step reactions. As the names imply, a one-step kit has reactants that will perform the RT reaction and then the PCR within the same tube with no further manipulation necessary by the investigator. In a two-step reaction, the reverse transcrip-tase reaction is run in one tube and then all or some of the resulting cDNA is used in the polymerase chain reaction, which is performed in a second plate, tube or capillary. The advantage of a one-step method is a savings in set-up time. However, there is always a compromise between the two reactions as neither can be truly optimized. Further, all the template in a one-step reaction will be used for that one reaction. Depending on how the kit is formulated, a no-reverse transcriptase control may not possible. This important control detects DNA contamination and ensures that low-level transcript measurements are legitimate. A two-step kit takes more hands-on time, as the cDNA is set up as a separate reaction and then introduced into the PCR. In a two-step protocol, a no-reverse transcriptase control is easily performed on each sample. Both the RT and the PCR can be optimized individually and only a portion of the cDNA made need be used in any one PCR. There are times when the use of one or the other assay system makes sense and it is up to the investigator to make that decision. In general, if the same results can be obtained with a one-step method compared to the same template in a two-step procedure, the time saved and convenience using the one-step kit makes a lot of sense. If this is not the case or the template is limiting, a two-step method should be used.

In principle, kits for real-time PCR are a major convenience for the investigator. They are also a source of a readily available and consistent master mix. However, they also add substantially to the cost of performing realtime PCR. Our lab has been making master mixes by hand since 1996, when there were no kits available. We continue to do so today. Formulas for master mixes for a two-step RT and a probe-based PCR or SYBR® Green I PCR are in Protocols 1.3, 1.4, and 1.5.

For purposes of the following discussion, a two-step method will be used as an example. After distributing the RT master mix into the number of wells required for the experiment, the samples are pipetted into wells following an appropriate pattern for the experiment. This pipetting step is THE most critical part of the experiment. A slight difference in RT master mix volume from well to well will have a negligible effect on the quantitative outcome of the experiment. In contrast, a very small difference in sample volume among replicates for the same sample will have measurable effects seen in the final data set. For this reason, recently calibrated adjustable pipettors and a steady hand are the key. If the cDNA for each sample is pipetted into replicate wells for the PCR, this step is the second most critical pipetting step as errors here will be multiplied by the any errors made in the initial template pipetting step. For this reason, in the examples shown the PCR master mix is added directly to the entire cDNA reaction to speed up assay setup and to avoid further errors in pipetting.

A good way for new investigators to gauge their pipetting skills is to make a 7-log dilution series of a DNA template and then pipette the dilutions, in triplicate, into a plate bearing PCR master mix in multiple wells. There is nothing more humbling than seeing the results of this experiment for the first time. Often, the replicates are not uniform and the points of the subsequent standard curve rarely fall onto a single straight line. There are two places for pipetting error in this experiment, making the dilution series itself and placing the diluted samples into the individual wells. A mistake in either or both will be seen in the final data. With practice, the curves will become uniform and straight.

Once the PCR plate, tube or capillary sets have been filled with all the reactants, it is ready to be run on a real-time instrument. The set up of the sample positions will be dependent upon the real-time instrument and software. Although they all look different, the methodology of defining the positions and quantities for the standards, unknown samples and controls is pretty much the same for all instruments. One thing that is instrument-specific is the thermocycling conditions. Instruments that use air to heat the reactions tend to cycle much faster than instruments that use metal blocks. There are new fast-blocks for some instruments in an attempt to match the faster air heated systems. The important thing is to use the cycling conditions that match your instrument, the reagents and the assay components, mostly the primers. The first temperature is the melting step. A one-minute melt at 94-95°C is sufficient for assays with short amplicons (<300 bases) using standard Taq. If a hot-start Taq is used, up to 15 minutes will be required for enzyme activation depending on the activation system used for the enzyme. For genomic DNA, at least 3 minutes is a safe time to obtain a good melt for the PCR. Next comes the annealing temperature, which will depend primarily upon the Tm of the PCR primers, their concentration and the MgCl2 concentration in the master mix. A good starting temperature range is 55-60°C. If primer-dimers or inappropriate PCR products are a problem, raise the annealing temperature in 2°C increments until the problem subsides or the reaction stops working. It may be necessary to design a better primer pair if this does not solve the problem. All probe-based assays can be run using 2-step cycles. That is from 94-95°C to 60°C and back to 94-95°C. Good cycling times for these reactions are 12-15 seconds at the high temperature followed by 30 seconds at the lower temperature, usually for 40 cycles. For dye- or primer-based assays, such as SYBR® Green 1, a 3-step cycle is generally used. Here, the cycle stays at the annealing temperature for only 2 seconds and then goes up to 72°C (the optimal temperature for Taq DNA polymerase) for 30 seconds for each cycle (94-95°C, 12-15 seconds; 55-60°C, 2 seconds and 72°C, 30 seconds). Collecting data at the higher temperature helps to melt out primer-dimers.

This practical introduction to running a real-time PCR experiment covered the basics of setting up a reaction. There is, of course, much more to know about running and analyzing the data from a real-time experiment. Those topics will be covered in succeeding chapters of this book.

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