Urea-induced denaturation of human phenylalanine hydroxylase

Human phenylalanine hydroxylase was expressed and purified from Escherichia coli as a fusion protein with maltose-binding protein. After removal of the fusion partner, the effects of increasing urea concentrations on enzyme activity, aggregation, unfolding, and refolding were examined. At pH 7.50, purified human phenylalanine hydroxylase is transiently activated in the presence of 0-4 M urea but slowly inactivated at higher denaturant concentrations. Intrinsic tryptophan fluorescence spectroscopy showed that the enzyme is denatured through at least two distinct transitions. The presence of phenylalanine (L-Phe) shifts the transition midpoint of the first transition from 1.4 to 2.7 M urea, whereas the second transition is unaffected by this substrate. Apparently the free energy of denaturation was almost identical for the free enzyme and for the enzyme-substrate complex, but significant differences in dDeltaG(D)/d[urea] (m(D) values) were observed for the first denaturation transition. In the absence of substrate, a high rate of non-covalent aggregation was observed for the enzyme in the presence of 1-4 M urea. All three tryptophan residues in the enzyme (Trp-120, Trp-187, and Trp-326) were mutated to phenylalanine, either as single mutations or in combination, in order to identify the residues involved in the spectroscopic transitions. A gradual dissociation of the native tetrameric enzyme to increasingly denatured dimeric and monomeric forms was demonstrated by size exclusion chromatography in the presence of denaturants.