Hello Abinet Legesse

There is no difference between Ct and Cq value. These values are all the same, just they are with different names. Ct means threshold cycle and Cq means quantification cycle. In order to standardize qPCR nomenclature, the MIQE (minimum information for publication of quantitative real-time PCR experiments) guidelines recommends the use of the more generic quantification cycle (Cq) term.

Real-time PCR usually quantifies the absolute amount of a target sequence or compares relative amounts of a target sequence between samples. Despite the fact that real-time PCR fluorescent dyes and probes should be sequence-specific, a considerable amount of background fluorescence occurs during most real-time PCR experiments. Thus, it is critical to bypass this background signal in order to glean meaningful information about your target. This issue is addressed by two values in real-time PCR namely, the threshold line and the Cq value.

The threshold line is the point or level of detection at which a reaction reaches a fluorescent intensity above background levels. Before you conduct PCR, the software in your cycler will set a threshold level. This is literally a line in your graph that represents a level above background fluorescence, that also intersects your reaction curve somewhere at the beginning of its exponential phase.

The Cq value is the PCR cycle number at which your sample’s reaction curve intersects this threshold line. This value tells how many cycles it took to detect a real signal from your samples. Real-Time PCR runs will have a reaction curve for each sample and therefore many Cq values. Your cycler’s software calculates and charts the Cq value for each of your samples.

Cq values are inverse to the amount of target nucleic acid that is in your sample, and correlate to the number of target copies in your sample. Lower Cq values (typically below 28 cycles) indicate high amounts of the target sequence. Higher Cq values (above 38 cycles) mean lower amounts of your target nucleic acid. However, high Cq values may also indicate some problems with the target or the PCR set-up.

Best.

To add to this already excellent answer, you can also calculate Cq value via regression.

Threshold methods are to a certain extent arbitrary: you can drag the threshold line up and down and (provided you remain within the exponential phase) the data for all samples within a given run will ostensibly remain entirely comparable (22 vs 23 is the same difference as 23 vs 24). The specific threshold isn't a fixed value, but you must apply the same threshold to all samples within that run.

Regression approaches instead use non-linear curve fitting (for example, the second derivative of the amplification curve, where the rate of change of the rate of change was maximal), and are not arbitrary: there is a single value for each trace.

In my experience, individual wells sometimes approach reaction saturation at sliiiiightly different rates, and thus unless the threshold is set very close to the lowest levels of detection, your thresholding will necessarily measure a degree of variation in reaction efficiency, and you'll see greater variation between replicate Cq values. Regression analysis should not be subject to the same constraints and (again, in my experience) gives more consistent Cq values.

Dear Malcolm Nbre and John Hildyard

Thanks for your elaborative (detailed explanation ) regarding ct vs cq value. I run RT- qPCR for a total of 45 cycles. What I faced is difficulty in determining the lower copy targets ( cq value 41 & 42 ) as positive or negative.

Negative, unless your PCR is incredibly inefficient.

As a rule of thumb, Cq=35 corresponds to 1 target molecule. With efficiency variation (and the innate variation in exactly how Cq is determined) the actual value for "one target molecule" may be anywhere from ~32 to ~38, but if you get Cqs in the 40s, that's primer dimers or instrument noise.

What you should see, if you do a dilution series, is:

• At high [target] or high [cDNA]: non-linear behaviour caused by reaction saturation/inhibitory effects. You have too much cDNA, use less.
• At lower [cDNA], very consistent Cq values across replicate wells, with mean Cq values that increase by ~3.32 for every ten-fold dilution. This should span a large dynamic range (6-7 logs, easily).
• At very low [cDNA], increasing variation between replicate wells: this is stochastic partitioning effects. When you only have ~10 molecules per ul, it's quite easy to have one well with 14 molecules, one with 8, and one with 11, and your Cq values will reflect this. If you need to quantify such low levels accurately, you will need more replicate wells to sample adequately -you'll still use the mean Cq, but of 5-6 replicate wells rather than 3.
• At zero cDNA, either no amplification at all, or sporadic late amplification that represents mispriming or primer dimers. The melt curve will usually reflect this.

Thanks sir!