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.