Quantitative Polymerase Chain Reaction Using a Recombinant DNA Internal Standard and Time-Resolved Fluorometry

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Abstract
Using a 308 bp DNA fragment (target DNA) as a template, we have synthesized an internal standard (IS) that is of the same size and uses the same primers as the target but differs by a 26 bp centrally located sequence. We then designed quantitative polymerase chain reaction (PCR) assays in which the target DNA is coamplified with a constant amount of IS (20 000 molecules). The presence of IS compensates for the reaction-to-reaction variability of the amplification efficiency. The PCR products are assayed by two distinct hybridization protocols. The first approach (QPCR-1) requires that specific probes be immobilized onto microtiter wells, followed by hybridization with digoxigenin-labeled PCR product. In the second protocol (QPCR-2), PCR product is captured onto the wells and hybridized with digoxigenin-tailed specific probes. In both assays, the hybrids are detected using an anti-digoxigenin−alkaline phosphatase conjugate and 5‘-fluorosalicylphosphate as substrate. The hydrolysis product forms a highly fluorescent complex with Tb3+−EDTA, as measured by time-resolved fluorometry. The ratio of the fluorescence values obtained for the amplified target DNA and IS is linearly related to the number of target DNA molecules present in the sample prior to amplification. The linear ranges are 1000−200 000 molecules for QPCR-1 and 2000−200 000 molecules for QPCR-2. The CVs ranged from 3.4 to 9.7%.