A colorimetric assay for NADPH-dependent, mercuric ion-specific oxidoreductase activity was developed to facilitate the investigation of mercuric reductase gene expression in polluted aquatic ecosystems. Protein molecules extracted directly from unseeded freshwater and samples seeded with Pseudomonas aeruginosa PU21(Rip64) were quantitatively assayed for mercuric reductase activity in microtiter plates by stoichiometric coupling of mercuric ion reduction to a colorimetric redox chain through NADPH oxidation. Residual NADPH was determined by titration with phenazine methosulfate-catalyzed reduction of methyl thiazolyl tetrazolium to produce visible formazan. Spectrophotometric determination of formazan concentration showed a positive correlation with the amount of NADPH remaining in the reaction mixture (r² = 0.99). Mercuric reductase activity in the protein extracts was inversely related to the amount of NADPH remaining and to the amount of formazan produced. A qualitative nitrocellulose membrane-based version of the method was also developed, where regions of mercuric reductase activity remained colorless against a stained-membrane background. The assay detected induced mercuric reductase activity from 10² CFU, and up to threefold signal intensity was detected in seeded freshwater samples amended with mercury compared to that in mercury-free samples. The efficiency of extraction of bacterial proteins from the freshwater samples was (97 ± 2)% over the range of population densities investigated (10² to 10⁸ CFU/ml). The method was validated by detection of enzyme activity in protein extracts of water samples from a polluted site harboring naturally occurring mercury-resistant bacteria. The new method is proposed as a supplement to the repertoire of molecular techniques available for assessing specific gene expression in heterogeneous microbial communities impacted by mercury pollution.