We present a new method for quantification of mRNA, in which the limitations of the current quantitative PCR methods can be overcome. A known amount of a synthetic RNA standard differing from the mRNA to be quantified by a single nucleotide is reverse-transcribed and amplified together with the mRNA template using a biotinylated primer. The biotinylated PCR product is immobilized on a streptavidin-coated solid support and denatured. The ratio between the two amplified sequences is determined by separate "mini-sequencing" reactions, in which a detection step primer annealing immediately adjacent to the site of the variable nucleotide is elongated by a single labeled dNTP complementary to the nucleotide at the variable site. The ratio between the incorporated labels accurately determines the ratio between the two sequences in the original RNA sample. We applied this method to quantify the mRNA of human aspartylglucosaminidase (AGA) in tissues and cultured cells. AGA is a lysosomal enzyme participating in the degradation of glycoproteins. A mutation in the AGA gene abolishes the enzyme activity and leads to aspartylglucosaminuria (AGU), a recessively inherited metabolic disorder. The mRNA quantification revealed that the normal and mutant genes are expressed at similar levels in kidney, liver, and cultured fibroblast, whereas the amount of AGA mRNA in normal placenta and brain is significantly higher than that found in the corresponding samples from AGU patients. The method presented here is generally applicable for PCR-based quantification of rare mRNAs and DNA as well.