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Quantitative measurement of the levels of mRNA expression using real-time reverse transcription polymerase chain reaction (RT-PCR) has long been used for analyzing expression differences in tissue or cell lines of interest. This method has been used somewhat less frequently to measure the changes in gene expression due to perturbagens such as small molecules or siRNA. The availability of new instrumentation for liquid handling and real-time PCR analysis as well as the commercial availability of start-to-finish kits for RT-PCR has enabled the use of this method for high-throughput small-molecule screening on a scale comparable to traditional high-throughput screening (HTS) assays. This protocol focuses on the special considerations necessary for using quantitative RT-PCR as a primary small-molecule screening assay, including the different methods available for mRNA isolation and analysis.
Phenotypic small-molecule screening, in which a biological system is probed for a change of interest independent of a known target, has been shown to be valuable in the generation of hits for successful lead and drug development (Swinney and Anthony, 2011). When a pathway of interest is to be targeted, a common technique is to use an engineered system such as a reporter gene assay. In such a system, any molecule that modulates a component of the pathway will change the expression level of an easily measured protein such as luciferase or beta-galactosidase. Some disadvantages of this method are the time required to create the modified cell line and the potential for the reporter construct to lead to artifacts. By contrast, a method such as quantitative RT-PCR (qRT-PCR) that directly measures levels of the native mRNA expression is faster to develop and more biologically relevant. Until recently, cost and throughput limited the use of real-time qRT-PCR as a method for screening the tens or hundreds of thousands of compounds typical of primary HTS. As described in this protocol, there are now standard reagents and instruments available that allow the miniaturization and rapid execution of real-time PCR in a robust, cost-effective manner, compatible with high-throughput small-molecule screening. The choice of commercial kits depends in part on the type of cells being analyzed. Protocols for isolating and analyzing cDNA from adherent and suspension cells are described, as well as a third option that bypasses a separate cDNA isolation step and uses cell lysates directly in the qRT-PCR analysis.
The essential piece of equipment for qRT-PCR is the real-time thermal cycler, which is a thermally controlled multi-well block mounted below a detection system that can simultaneously measure the fluorescence signal in all wells of a multiwell assay plate. Instruments are available in 96, 384, and 1536 format; for large-scale HTS, miniaturization to 384 or 1536 is advisable for throughput and cost reasons. Some of the 96 and 384 well instruments available are sold by Applied Biosystems/Life Technologies/Invitrogen (7900, Viia 7), Bio-Rad (CFX line), and Roche (LightCycler 480) and are common in many labs, core facilities, and academic screening centers. In addition, Roche offers the 1536 LightCycler, which enables sub-uL reactions on a scale compatible with ultra-high throughput screening (uHTS (Schlesinger et al., 2010)). Depending on the throughput desired, integration with an automated plate handler or screening system may also be an advantage.
The degree of miniaturization and number of compounds to be screened also dictate whether some form of automated liquid handling may be necessary (Rudnicki and Johnston, 2009). At a minimum, a multichannel pipette is required. Ideally, a bulk reagent dispenser such as the Combi Multidrop (Thermo) and a multichannel pipetting station (96 or 384 tip head) such as the Beckman Multimek or CyBio Vario are used for reagent and cDNA transfer. Unlike qPCR analysis of many different genes from a small number of cDNA preparations (Arany 2008), HTS of small molecules requires repeated analysis of the same gene or small set of genes, typically 2 or 3 genes. For this reason, the detection reagents for the qPCR step can be distributed from a master mix to the plate using a bulk reagent dispenser, but the individual wells with different cDNA samples must be transferred using separate tips with a multichannel pipette or a 96- or 384-tip pipetting robot. For further miniaturization, especially to 1536 format, more specialized instruments are required, including sub-uL bulk dispensers (e.g. Thermo Combi-nL or Biotek BioRaptr) for use with mastermixes and an acoustic dispenser (e.g. Labcyte Echo or EDC ATS) to transfer cDNA samples.
As with all highly parallel experiments such as HTS, sample management is essential. Up to three whole-plate transfers are required for qRT-PCR analysis, so it is essential to track the relationships between the different source and destination plates very carefully. Ideally a laboratory informatics management system (LIMS) database is used to record the transfers; at a minimum all plates should be uniquely labeled, such as with a barcode, and all plate identities should be tracked in a spreadsheet.
The goal of high-throughput compound screening using qRT-PCR as a detection format is to obtain an accurate measurement of the effect of small molecules on the mRNA expression levels of a gene or genes of interest. To detect variation due to nonspecific effects, the expression level of the gene or genes of interest are normalized to the expression level of a control gene, typically a general metabolic or “housekeeping” gene such as glyceraldehyde 3-phosphate dehydrogenase (GAPDH) or TATA-binding protein (TBP) (Schmittgen 2008). The choice of the control gene depends on the expression level of the gene(s) of interest. Ideally, a housekeeping gene with expression levels similar to the target gene(s) is used. The calculations for normalization and data analysis will be discussed below.
qPCR relies on the generation of a fluorescent signal correlated with the amount of PCR product present after each round of thermal cycling. The fluorescent signal can be generated by a general intercalator (Sybr green) that measures all DNA present. Alternatively, a fluorophore-quencher pair attached to a DNA sequence specific for the gene of interest is digested by the DNA polymerase during amplification, releasing the fluorophore and generating a signal. Thus, there are two methods for fluorescent quantification of both a control gene and one or more target genes. Using Sybr green, it is necessary to run a separate qPCR reaction for each gene to be quantified, as the intercalator does not distinguish between the PCR products generated either from each template sequence or from non-specific amplification of other sequences. The readings from the separate plates are then used for normalization. Using hydrolysable probes with different fluorophores for each gene sequence provides additional specificity to the target sequences and enables both genes to be analyzed in the same reaction, as nearly all real-time PCR instruments have the capability to monitor multiple excitation/emission wavelength channels. Multiplexing using multicolor hydrolysis probes is easier and faster in execution and provides more consistent normalization to the control gene. Design of primer-probe sets that perform well together is more difficult than simply identifying PCR primer pairs for use with Sybr green detection, but the up-front investment in assay design is well worth the effort. In addition, although probes are more expensive than unmodified PCR primers, the same probe set is used for the entire screen. Therefore, unlike genotyping experiments where many probes are required at a high cost, the reagents for small-molecule HTS can be purchased or synthesized in bulk at considerable savings.
The difference between the three protocols presented herein (Figure 1) involves the method used to isolate the mRNA and generate cDNA for analysis. The appropriate protocol depends primarily on the cell type being analyzed (adherent or suspension) and by the desired workflow timing. The first method requires a washing step prior to cell lysis to remove cell media that can interfere with cDNA generation, and is therefore only appropriate for adherent cells. While it is possible to remove cell culture components from suspension cells using filter plates or by centrifuging cells, attempting to use such methods in multiwell formats introduces unacceptable variation in the experiment; therefore an alternate protocol using mRNA capture plates that is more ideal for suspension cells is described. Kits also exist that generate cDNA in situ prior to real-time analysis rather than isolating cDNA in a separate step. Overall, the two-step adherent protocol is currently the most cost effective and has the fewest time constraints, as the cDNA is isolated as an intermediate product and can be stored prior to transfer to multiple analysis plates.
As suspension cells cannot be reliably washed in multiwell plates, an alternative protocol is necessary for suspension cells such as those derived from blood lineages. For this protocol, the turbocapture kit from Qiagen can be used as directed for suspension cells to generate cDNA.
For more streamlined HTS using qRT-PCR, kits are available that allow cell lysates to be directly analyzed in the qPCR instrument. This requires that the RT step occur in the real-time PCR instrument. As with the two-step method above, this is only appropriate for adherent cells as wash steps are required.
This protocol is adapted from the Roche Real-time Ready kit for 384 well format plates, volumes should be adjusted accordingly for 96 well format:
All reagents described are specific to the cell culture requirements of the cells to be used or are included in the commercial kits described. The manufacturer’s recommendations with respect to storage and expiration should be followed.
Shortly after the introduction of PCR in 1983, it was recognized that the initial amount of template could be quantitated by stopping the reaction at regular cycle intervals and measuring double-stranded DNA levels by methods such as absorbance or gel quantitation (VanGuilder et al. 2008). Over a decade later, methods were described for generating a signal in situ during amplification that could be read concurrent with thermal cycling (Heid et al. 1996).
While initial quantitative PCR methods were used for determining viral loads, the majority of the applications of the method have been for measuring expression levels of a number of genes in different biological samples such as diseased vs. non-diseased cells or various tissue types, generating transcriptional profiles characteristic of the underlying biology (Derveaux et al., 2010). It serves as a complement to expression microarrays and is often used to validate the latter, offering lower throughput but greater sensitivity and reliability (Ding and Cantor 2004, Arikawa et al. 2008). More recently it has been used to detect other cellular nucleic acids, including non-coding RNAs such as miRNAs (Benes and Castoldi 2010).
Due to the cost of designing detection assays for each separate gene, real time qRT-PCR is often not used in high-throughput format for genotyping. However, applications in which it is desirable to measure the response of a small number of genes to many different treatments are well suited for high-throughput analysis using this technique. In addition to changes in gene expression caused by compound treatment, described here and elsewhere (Hakamatsuka and Tanaka 1997, Maley et al. 2004, Wagner and Arany 2009), any system in which a nucleic acid can be generated as a biomarker or a surrogate for another analyte (such as a protein) can also be analyzed by qPCR (Swartzman et al., 2010). Thus, in addition to describing the biology of a natural or perturbed biological system, it can also be used to detect natural or artificial constructs for diagnostic or screening uses.
The most important factor for a successful high-throughput gene expression assay is the design of the primers and probes used to detect the gene(s) of interest through the qRT-PCR reaction. Fortunately, bioinformatics resources and commercial products released in the last 5 years have made this a much less daunting task. In particular, many of the same companies that sell the instrumentation and consumables for real-time PCR are happy to assist with the design of assays to maximize the use of their products. Bio-Rad, Applied Biosystems/Life
Technologies/Invitrogen, and Roche all offer pre-designed packages of primers and/or probes. These sets are often referred to as assays and are sold as a complete package for real-time analysis. In addition, working with public genome databases, these same sites will design an assay based on any known sequence or gene, optimizing primer sequence and amplicon length. Most assays are available in the two formats described in this protocol, primers alone for use with Sybr green or primer-probe sets for use as Taqman assays. Additional formats, such as molecular beacons or FRET probes, are also available and may have an advantage for specific applications, such as when a loss of signal rather than a gain of signal is desired during amplification.
Another key part of assay design is selection of a proper control gene for normalizing against effects such as cytotoxicity or variation in cell plating. Ideally a gene unrelated to any pathway affected by the target gene should be used, such as the housekeeping genes TBP, GAPDH, or various rRNA sequences. In addition, it is best to use a control gene that amplifies in roughly the same number of cycles as the target gene, as this allows more robust normalization. In extreme cases where target genes are very highly expressed or underexpressed and the target and control signals are not both in the linear detection range of the assay, one of the assays can be adjusted by limiting one of the reagents such as a primer (known as primer-limiting) to artificially change the Cq value by reducing the efficiency of each round of PCR.
It is also ideal to measure the control gene and the target gene in the same reaction plate by using multiplexed probes with differentially emitting fluorophores. This reduces any variability between the control and target genes that is due to pipetting error or unequal treatment (light or temperature exposure) when separate plates are needed for each gene, as is the case when using Sybr green.
Prior to any small molecule high-throughput screen, extensive assay development is required, and is especially critical in an assay as sensitive as qRT-PCR. All steps of a protocol, such as cell culturing, seeding, washing, and reagent transfers should be validated to minimize variation due to technical limitations. Untreated control plates should be run to determine the variability of the measurements and the corresponding sensitivity of the assay. When properly designed, these controls can also serve to test other technical aspects such as consistency of plate positioning (avoiding rotations) and recording transfers among the multiple plates that are generated in the course of an experiment.
In addition to technical sensitivity as measured by standards such as the percent coefficient of variation, in phenotypic assays it is important to establish a threshold for a biologically relevant response. Ideally a small molecule positive control can be used to both establish this biological response during assay development and to monitor the biological consistency during HTS. In the absence of a small molecule control, a genetic control, such as an overexpression vector or siRNA, can be used to establish assay sensitivity. However, genetic controls suffer from the limitation that the effect seen is often orders of magnitude greater than the response that can be expected from a small molecule, making comparison to screening results difficult. Finally, in the absence of either of these controls, a technical control such as a known stress inducer or cytotoxic agent can be used as a technical control simply to track and confirm the execution of reagent transfers, even though there is no relationship of this type of control to the desired biology.
Overall, the design and validation of the assay is key for successful small-molecule screening by real-time qRT-PCR. This type of assay is among the most expensive used for small-molecule screening, and while the biological relevance of the data can provide a good return on this investment, it is easy to waste a large amount of expensive reagents if experiments are not properly planned and tested prior to screening.
The ΔΔCq method described above is used to relate two differently treated samples, in this case compound and mock treatment, where each sample has a target gene compared to a control gene (Schmittgen and Livak, 2008). The data for the mock treatments is typically normally distributed due to experimental error, and single point values can be compared to this distribution for determining likelihood of a significant effect due to compound treatment.
The next step following hit calling is typically retesting of the compounds in the assay at a range of dilutions to construct a concentration response curve. There are two additional considerations in concentration-response tests compared to the primary HTS. First, the goal is not to measure the significance of an outlier, but to observe the biological effect of the compounds. This requires converting an exponential measurement given by Cq to a linear scale that represents the actual relative change in copy number. This conversion is done by the calculation: Fold change = 2ΔΔCq (Table 2.) These values should then be plotted against the tested concentration values for curve fitting (Figure 3.)
The conversion Fold change = 2ΔΔCq (or alternatively 2−ΔΔCq depending on whether a fold repression or fold induction is being described) assumes a perfect PCR efficiency; that is, it assumes that each round of PCR exactly doubles the amount of DNA product during the logarithmic PCR expansion. In reality, PCR efficiencies are usually near 1.9 but can be as low as 1.7 (Tuomi et al. 2010). If more precision is desired, a standard curve can be generated for the qPCR assay to be used by making a serial dilution of a sample PCR template and measuring the actual fold amplification observed in each round of PCR, which can be used in the above calculation (Figure 4.)
The second consideration when trying to fit concentration-response curves is deciding whether the maximal effect can be determined. In the hypothetical graph in Figure 3, the response plateaus at high compound concentration, enabling the determination of an EC50 (50% of maximal effect) for the curve. More typically, either due to complex biology or weakly active compounds, it is not possible to determine the upper asymptote for the fitted curve, and the maximal effect may be very different for various compounds. In these cases, it is more appropriate to determine a threshold of N-fold induction or repression that represents a statistically and biologically significant effect, and set this ECN as the concentration at which the compound achieves the desired threshold. This allows more robust comparison of compounds with a wide range of biological effects and enables better selection for compounds to carry forward for further development.
The greatest difficulty in small-molecule screening is high variability, which can be observed by high variation in the mock treatment wells. This reduces the sensitivity of the experiment and increases the rate of false positives. In qRT-PCR experiments, it has been shown that with careful technique, the qPCR and RT generation together represent less than 20% of the total variation between replicate analyses (Kitchen et al. 2010). The majority of variability comes from biological variability, including sample preparation. In the protocol presented here, this includes cell culture of the cell line to be used, plating and treatment of the cells, and washing and lysis prior to cDNA generation. It is important to have cultured cells as consistent as possible across all compound tests, including minimizing passage number variation and exposure to variable environmental conditions. Plating of cells should be done in manageable batches to reduce variability due to plating order. Washing parameters should be set to minimally disturb the cells, and lysis should be optimized for the cells used to ensure complete reaction. While replicates may help determine the extent of variability seen in treatment wells, it is costly to run biological replicates (treating two separate plates of cells with the same compounds) as the cDNA generation accounts for the majority of the cost of the assay. Real-time qPCR replicates from the same cDNA well can confirm the robustness of the analysis step, but as mentioned above this is not expected to reveal much variability and is time consuming to perform for an entire HTS campaign.
Very high Cq values are indicative of poor mRNA isolation or cDNA generation and are problematic because very small amounts of contamination or variability can have a large effect on the results. If values are too low, confirm the efficiency of the PCR reaction and consider re-optimizing the assay design or even selecting a different target gene. Adjustment of cell number may have a small effect but is unlikely to improve low signals, as a 2-fold increase in cell seeding will only have ~1 cycle effect on Cq.
Finally, it is essential to include controls for non-amplification to ensure that there is no contamination. If no-template controls are showing signals that are within several cycles of experimental wells, the data cannot be accepted as valid. Contamination can be controlled by the same methods used for standard PCR, including isolating and cleaning a workspace and never bringing post-amplification materials into the PCR setup area.
It is expected that in a compound library of appropriate size or design, some amount of biological effect will be seen, if only due to cytotoxic compounds. Typically a validation library containing a sample of bioactive compounds is run early in the screen to confirm the presence of this effect. Figure 2 illustrates a typical assay plate with a range of Cq values due to compound treatment, and similar small-molecule high-throughput screening results have been reported and analyzed previously (Arany et al., 2008). Overall, the active well rate for compounds showing the desired change in target gene expression is highly dependent on the biological system under investigation. It is entirely possible that a particular gene is intractable to specific modulation by small molecules, and that any putative hits will be removed as artifacts in subsequent follow up assays. As with all discovery projects, the researchers needs to make an informed decision about the return on investment of additional screening or follow up.
The largest investment of time is in the assay development and validation but is essential to avoid wasting time and money in downstream failures. Several rounds of mock runs are usually required to get low-noise consistent results through the entire process of cell culture, lysis, cDNA generation, and qPCR analysis. This will take several weeks of iteration.
Depending on the choice of protocol and number of compounds tested, the small-molecule HTS can be staged in various ways. Using a two-step protocol to isolate cDNA has the advantage of allowing the separate execution of each phase of the assay rather than multitasking, which adds to the total project time but can make execution more consistent. The only time-sensitive step in the protocol is the lysis treatment, which can ruin the nucleic acid sample if run too long. For large numbers of multiwell plates it is advisable to have at least two scientists working together to execute the protocol, as subsequent steps can back up and interfere with the key timing step. It is important to recognize other potential bottlenecks as well including standard PCR blocks for the RT step, robotic pipetting throughput, and real-time instrument availability. Proper planning will enable optimal use of these resources
Even though the real-time qPCR analysis step takes 1–1.5 hours per plate, it is now possible to screen tens or hundreds of thousands of compounds using real-time qRT-PCR thanks to miniaturization to 384- or 1536-well format. If an assay is robust and appropriate instrumentation is available, thousands of compounds can be screened each day during a campaign, enhancing the potential for discovery of specific small-molecule modulators of the target gene of interest.
Internet Resources with Annotations
Bio-rad gene expression gateway: http://www.biorad.com/genomicsProvides useful tutorials on experimental design and detection formats
Search of existing qPCR detection reagents and bioinformatics tools for designing custom assays.
Life technologies/Applied Biosystems designer for custom Taqman probes: https://www5.appliedbiosystems.com/tools/cadt/
Life technologies Taqman assays and assay designer.