Quantitation of CYP1A1 and 1B1 mRNA in polycyclic aromatic hydrocarbon-treated human T-47D and HepG2 cells by a modified bDNA assay using fluorescence detection

The quantitation of mRNA, essential for assessing mechanisms of enzyme regulation, is normally carried out using reverse transcriptase–polymerase chain reaction (RT–PCR). An alternative method uses a signal-amplification nucleic acid probe assay, which measures RNA directly by the QuantiGene Expression Kit and incorporates branched DNA technology from Bayer and luminometer-based readings of a chemilumigenic alkaline phosphatase substrate. To broaden the utility of this assay, we investigated substitution of a fluorescent substrate, 2 ′-(2-benzothiazol)-6 ′-hydroxybenzothiazole phosphate and a fluorometer, and applied the method to quantitation of CYP1A1 and 1B1 mRNA in human T-47D and HepG2 cells following induction by benzo[a]pyrene (B[a]P) and dibenzo[a,h]anthracene (DB[a,h]A). The fluorescence response increased linearly for 200 min without photobleaching and increased linearly ( r 2=0.997) up to at least 0.2 μg total RNA. The data revealed that at 0.5 and 1.0 μM inducing agent, the induction of CYP1A1 mRNA in HepG2 cells by DB[a,h]A exceeded that by B[a]P by 18- and 6-fold, respectively. In T-47D cells B[a]P induced CYP1A1 mRNA by 23-fold and CYP1B1 mRNA by 3.9-fold. A B[a]P cocontaminant in the environment, arsenite, did not affect B[a]P-induced levels of CYP1A1 or 1B1 mRNA in these cells. The modified analytical system provides a rapid-throughput, reproducible, and less labor-intensive method than RT–PCR for quantifying cellular mRNA levels.