Tatiana K. Shokhireva

The effect of mutation of F87 on the properties of CYP102A1-CYP4C7 chimeras: altered regiospecificity and substrate selectivity

CYP102A1 is a highly active water-soluble bacterial monooxygenase that contains both substrate-binding heme and diflavin reductase subunits, all in a single polypeptide that has been called a “self-sufficient enzyme.” Several years ago we developed a procedure called “scanning chimeragenesis,” where we focused on residues 73–82 of CYP102A1, which contact approximately 40% of the substrates palmitoleic acid and N-palmitoylglycine [Murataliev et al. (2004) Biochemistry 43:1771–1780]. These residues were replaced with the homologous residues of CYP4C7. In the current work, that study has been expanded to include residue 87. Phenylalanine 87 of wild-type CYP102A1 was replaced with the homologous residue of CYP4C7, leucine, as well as with alanine. The full-sized chimeric proteins C(73–78, F87L), C(73–78, F87A), C(75–80, F87L), C(75–80, F87A), C(78–82, F87L) and C(78–82, F87A) have been purified and characterized. Wild-type CYP102A1 is most active toward fatty acids (both lauric and palmitic acids produce ω-1, ω-2, and ω-3 hydroxylated fatty acids), but it also catalyzes the oxidation of farnesol to three products (2, 3- and 10,11-epoxyfarnesols and 9-hydroxyfarnesol). All of the F87-mutant chimeric proteins show dramatic decreases in activities with the natural CYP102A1 substrates. In contrast, C(78–82, F87A) and C(78–82, F87L) have markedly increased activities with farnesol, with the latter showing a 5.7-fold increase in catalytic activity as compared to wild-type CYP102A1. C(78–82, F87L) produces 10,11-epoxyfarnesol as the single primary metabolite. The results show that chimeragenesis involving only the second half of SRS-1 plus F87 is sufficient to change the substrate selectivity of CYP102A1 from fatty acids to farnesol and to produce a single primary product.

Native N-terminus nitrophorin 2 from the kissing bug: similarities to and differences from NP2(D1A).

The first amino acid of mature native nitrophorin 2 is aspartic acid, and when expressed in E. coli, the wild‐type gene of the mature protein retains the methionine‐0, which is produced by translation of the start codon. This form of NP2, (M0)NP2, has been found to have different properties from its D1A mutant, for which the Met0 is cleaved by the methionine aminopeptidase of E. coli (R. E. Berry, T. K. Shokhireva, I. Filippov, M. N. Shokhirev, H. Zhang, F. A. Walker, Biochemistry 2007, 46, 6830). Native N‐terminus nitrophorin 2 ((ΔM0)NP2) has been prepared by employing periplasmic expression of NP2 in E. coli using the pelB leader sequence from Erwinia carotovora, which is present in the pET‐26b expression plasmid (Novagen). This paper details the similarities and differences between the three different N‐terminal forms of nitrophorin 2, (M0)NP2, NP2(D1A), and (ΔM0)NP2. It is found that the NMR spectra of high‐ and low‐spin (ΔM0)NP2 are essentially identical to those of NP2(D1A), but the rate and equilibrium constants for histamine and NO dissociation/association of the two are different.

Assignment of the 1H NMR resonances of protein residues in close proximity to the heme of the nitrophorins: similarities and differences among the four proteins from the saliva of the adult blood-sucking insect Rhodnius prolixus.

The nuclear Overhauser effects (NOEs) observed between heme substituent protons and a small number of nearby protein side chain protons in the water-elimination Fourier transform NOE spectroscopy (WEFT-NOESY) spectra of high- and low-spin wild-type nitrophorin (NP) 2 and its ligand complexes have been analyzed and compared with those observed for the same complexes of wild-type NP3. These assignments were made on naturally abundant isotope samples, with the most useful protein side chains being those of Ile120, Leu122, and Leu132 for NP2 and NP3, and Thr121, Leu123, and Leu133 for NP1 and NP4. It is found that the NOEs observed are identical, with extremely similar protein side chain proton chemical shifts. This is strong evidence that the structure of NP3, for which no X-ray crystal structures are available, is essentially identical to that of NP2, at least near the heme binding pocket. Similarly, the NOEs observed between heme substituents and protein side chains for NP1 and NP4 also indicate that the structures of the protein having both A and B heme orientations are very similar to each other, as well as to the proteins with major B heme orientation of NP2 and NP3. These A and B connectivities can be seen, even though the two heme orientations have similar populations in NP1 and NP4, which complicates the analysis of the NOESY spectra. The histamine complex of wild-type NP2 shows significant shifts of the Leu132 side chain protons relative to all other ligand complexes of NP1–NP4 because of the perturbation of the structure near Leu132 caused by the histamine’s side chain ammonium hydrogen bond to the Asp29 side chain carboxylate.Graphical abstract

Linear correlation between 1H and 13C chemical shifts of ferriheme proteins and model ferrihemes.

The (1)H{(13)C} HMQC experiment at natural-abundance (13)C provides a very useful way of determining not only (1)H but also (13)C chemical shifts of most heme substituents, without isotopic labeling of the hemin. This is true both in model low-spin ferriheme complexes and in low-spin ferriheme proteins, even when the proton resonances are buried in the protein diamagnetic region, because the carbon shifts are much larger than the proton shifts. In addition, in many cases, the protohemin methyl cross peaks are fairly linearly related to each other, with the slope of the correlation, δ(C)/δ(H), being approximately -2.0 for most low-spin ferriheme proteins. The reasons why this should be the case, and when it is not, are discussed.

Dimerization of Nitrophorin 4 at Low pH and Comparison to the K1A Mutant of Nitrophorin 1

Nitrophorin 4, one of the four NO-carrying heme proteins from the salivary glands of Rhodnius prolixus, forms a homodimer at pH 5.0 with a Kd of ∼8 μM. This dimer begins to dissociate at pH 5.5 and is completely dissociated to monomer at pH 7.3, even at 3.7 mM. The dimer is significantly stabilized by binding NO to the heme and at pH 7.3 would require dilution to well below 0.2 mM to completely dissociate the NP4-NO homodimer. The primary techniques used for investigating the homodimer and the monomer–dimer equilibrium were size-exclusion fast protein liquid chromatography at pH 5.0 and 1H{15N} heteronuclear single-quantum coherence spectroscopy as a function of pH and concentration. Preparation of site-directed mutants of NP4 (A1K, D30A, D30N, V36A/D129A/L130A, K38A, R39A, K125A, K125E, D132A, L133V, and K38Q/R39Q/K125Q) showed that the N-terminus, D30, D129, D132, at least one heme propionate, and, by association, likely also E32 and D35 are involved in the dimerization. The “closed loop” form of the A–B and G–H flexible loops of monomeric NP4, which predominates in crystal structures of the monomeric protein reported at pH 5.6 but not at pH 7.5 and which involves all of the residues listed above except D132, is required for dimer formation. Wild-type NP1 does not form a homodimer, but NP1(K1A) and native N-terminal NP1 form dimers in the presence of NO. The homodimer of NP1, however, is considerably less stable than that of NP4 in the absence of NO. This suggests that additional aspartate or glutamate residues present in the C-terminal region of NP4, but not NP1, are also involved in stabilizing the dimer.

Erratum to: Assignment of the 1

Correction to the caption of Fig. 3, Table 1, and the text. Further research [Abriata LA, Zaballa M-E, Berry RE, Yang F, Zhang H, Walker FA, Vila AJ, submitted to Inorg Chem] has shown that the small peak seen at 2.6 ppm of the F1 dimension of the WEFT-NOESY spectrum of NP2(D1A) (top right in Fig. 3) decreases in intensity slowly as a function of the amount of time the protein sample has been in D2O. The spectrum shown in Fig. 3 top right was recorded approximately 1 week after the protein had been dissolved in D2O containing phosphate buffer at pH* 7.0. The intensity of the peak at 2.6 ppm is about 10 % that of the peak at -2 ppm. However, a similar sample left in D2O for 16 months showed this peak at 2.6 ppm to have only *1 % of the intensity of the peak at -2 ppm. Thus this cross peak is that of an exchangeable proton, and cannot be that of the a-CH of His57. By analogy, those of wt NP2 and wt NP3 also shown in the top portion of that figure, and also the 1D NOE shown in the bottom portion of that figure, at similar chemical shifts of 2–3 ppm in each case, are likely exchangeable protons and thus cannot be assigned to the a-CH proton of His57 in these high-spin ferriheme proteins. In the work quoted above by Abriata et al., the a-CH proton of His57 of native N terminus NP2 containing C, N labeled histidine was shown by H{C} HMQC to be at -1.4 ppm. Since native N terminus NP2 and NP2(D1A) have all other proton chemical shifts the same within *0.2 ppm [Berry RE, Muthu D, Shokhireva TK, Garrett SA, Zhang H, Walker FA (2012) Chem Biodiv 9:1739], we should expect to see a cross peak at -1.4 ± 0.2 ppm for the a-CH of His57 in Fig. 3; however, because of the proximity of the cross peak for the b-CH at about -2 ppm, which is expected to be more intense, that a-CH cross peak is not observed by WEFT-NOESY methods. In the paper by Abriata et al. referenced above, Supporting Information Figure S4 presents the raw data which confirm that the WEFT-NOESY cross peak in the spectrum of NP2(D1A) at 2–3 ppm is that of an exchangeable proton. The same work of Abriata et al., quoted above, shows that the cross peak at 2.90 ppm in the H–C plane of the HNCA spectrum of high-spin native N terminus NP2 is that of the backbone amide proton of His57, which is only 2.51 Å from the closest b-CH proton, well within the distance expected for observing NOE cross peaks; in contrast, the a-CH proton is 2.86 Å from that same b-CH proton (PDB file 2EU7, of NP2(D1A)-NH3).