Real-Time PCR: Understanding C_t

Real-time PCR, also called quantitative PCR or qPCR, can provide a simple and elegant method for determining the amount of a target sequence or gene that is present in a sample. Its very simplicity can sometimes lead to problems by overlooking some of the critical factors that make it work. This review will highlight these factors that must be considered when setting up and evaluating a real-time PCR reaction.

Factors that can influence C_t

C_t (threshold cycle) is the intersection between an amplification curve and a threshold line (Figure 1B). It is a relative measure of the concentration of target in the PCR reaction. Many factors impact the absolute value of C_t besides the concentration of the target. We will discuss the most common template-independent factors that can influence C_t and describe how to evaluate the performance of a real-time PCR reaction.

Figure 1, above, shows several parameters of the real-time reaction amplification plot. The exponential phase in Figure 1B corresponds to the linear phase in Figure 1C. The threshold must be set in the linear phase of the amplification plot in Figure 1C. The C_t value increases with a decreasing amount of template. However, artifacts from the reaction mix or instrument that change the fluorescence measurements associated with the C_t calculation will result in template-independent changes to the C_t value. Therefore, the C_t values from PCR reactions run under different conditions or with different reagents cannot be compared directly.

The fluorescence emission of any molecule is dependent on environmental factors such as the pH of a solution and salt concentration. Figure 2 shows the raw fluorescence data of an Applied Biosystems TaqMan® probe in the background of two different master mixes. Note that the fluorescence intensity is higher in Master Mix A even though the target, probe, and ROX dye concentrations are the same in both cases.

The resulting ΔRn value will, therefore, vary as shown in Figure 3. Note that the baseline fluorescence signals, in a template-independent factor, are different for the two master mixes (Figure 3A). Variations in the C_t value do not reflect the overall performance of the reaction system (Figure 3B). Master mixes with equivalent sensitivity may have different absolute C_t values.

The Rn value is calculated as the ratio of the fluorescence of Applied Biosystems™ FAM™ Dye divided by the fluorescence of ROX dye. Therefore, a lower amount of ROX dye would produce a higher Rn value assuming fluorescence signal from FAM dye is unchanged. This would lead to an increase in baseline Rn and subsequently a smaller ΔRn as well as a different C_t value. The new C_t value obtained by lowering the level of ROX dye has no bearing on the true sensitivity of the reaction, but can have other unintended consequences. Low concentrations of ROX dye can result in increased standard deviation of the C_t value, as shown in Figure 4. The greater the standard deviation, the lower the confidence in distinguishing between small differences in target concentration (see the precision section below).

The efficiency of a PCR reaction can also affect C_t. A dilution series amplified under low efficiency conditions could yield a standard curve with a different slope from one amplified under high efficiency conditions. In Figure 5, two samples (X and Y) amplified under low and high efficiency conditions show different C_t values for the same target concentration. In this example, although the high-efficiency condition (the blue curve in Figure 5) gives a later C_t at high concentrations, it results in better sensitivity at low target concentrations. The PCR efficiency is dependent on the assay, the master mix performance, and sample quality. Generally, an efficiency between 90 and 110% is considered acceptable. the other, could be valuable in concluding that there is less template in the first sample, assuming all other factors such as instruments, reagents, and assays are equal. However, this is not true if different instruments, reagents, primers and probes, or reaction volumes are involved in producing the two C_ts. Therefore, the absolute C_t value comparison is only meaningful when comparing experiments using the same reaction conditions as defined above.

In order to compare two reactions where a condition is changed (for example two different master mixes or two different instruments), the following parameters must be evaluated.

Dynamic range

To properly evaluate PCR efficiency, a minimum of 3 replicates and a minimum of 5 logs of template concentration are necessary. The reason for this suggested level of rigor is illustrated in Figure 6, which demonstrates the possible mathematical variation of slope or efficiency obtained when testing dilutions over 1 log vs. 5 logs. Thus, even if the assay is 100% efficient, a range from 70 to 170% can be obtained when testing a dilution series of a single log, due to the standard deviation in one dilution. For the same number of dilutions or replicates on a 5-log range, the potential artifact is only ±8 %. That means for 94% efficiency on a 5-log range, the assay would have a range of 88% to 100% efficiency. To accurately determine the efficiency of a PCR reaction, a 5-log dilution series must be performed. A slope of –3.3 ±10% reflects an efficiency of 100% ±10%. A PCR reaction with lower efficiency will have lower sensitivity.

R² value

Another critical parameter to evaluating PCR efficiency is R², which is a statistical term that indicates how good one value is at predicting another. When R² is 1, the value of Y (Ct) can be used to accurately predict the value of   (Figure 7A). If R² is 0, the value of X cannot be predicted from the value of Y (Figure 7B). An R² value >0.99 provides good confidence in correlating two values.

Precision

The standard deviation (square root of the variance) is the most common measure of precision. If many data points are close to the mean, the standard deviation is small; if many data points are far from the mean, the standard deviation is large.

In practice, a data set with a sufficient number of replicates forms an approximately normal distribution. This is frequently justified by the classic central limit theorem which states that sums of many independent, identically distributed random variables tend towards the normal distribution as a limit. As shown in Figure 8A, approximately 68% of the values are within 1 standard deviation of the mean, 95% are within two standard deviations, and 99.7% lie within 3 standard deviations.

If a PCR is 100% efficient, the C_t difference between two successive concentrations in a 2-fold dilution is 1 (Figure 8B). To be able to quantify a 2-fold dilution in more than 99.7% of cases, the standard deviation has to be ≤0.167. The greater the standard deviation, the lower the ability to distinguish between 2-fold dilutions. To be able to discriminate between a 2-fold dilution in more than 95% of cases, the standard deviation has to be ≤0.250 (Figure 8C).

Sensitivity

Any system capable of effectively amplifying and detecting one copy of starting template has achieved the ultimate level of sensitivity, regardless of the absolute value of the C_t.

As described earlier, efficiency is a key factor in determining the sensitivity of a reaction (Figure 5). Another important consideration when detecting very low copy numbers is that normal distribution of template is not expected. Instead, a Poisson distribution is followed, which predicts that in a large number of replicates containing an average of one copy of starting template, approximately 37% should actually have no copies, only 37% should contain one copy, and 18% should contain two copies (see Figure 9). Thus, for a reliable low copy detection, a large number of replicates are necessary to provide statistical significance and to overcome the Poisson distribution limitation.

Efficiency, R², precision, and sensitivity are used to determine performance of a PCR reaction when comparing different reaction conditions. For a rigorous evaluation, all factors listed in Table 1 must be evaluated together. In addition to these factors, proper experimental controls (such as no template control, no RT control) and template quality must be evaluated and validated.