Iron(II)-and 2-oxoglutarate-dependent (Fe/2OG) oxygenases generate iron(IV)-oxo (ferryl) intermediates that can abstract hydrogen from aliphatic carbons (R-H). Hydroxylation proceeds by coupling of the resultant substrate radical (R•) and oxygen of the Fe(III)-OH complex ("oxygen rebound"). Non-hydroxylation outcomes result from different fates of the Fe(III)-OH/R• state; for example, halogenation results from R• coupling to a halogen ligand cis to the hydroxide. We

LolO, a 2-oxoglutarate-dependent nonheme Fe oxygenase, catalyzes both the hydroxylation of 1-exo-acetamidopyrrolizidine (AcAP), a pathway intermediate in the biosynthesis of the loline alkaloids, and the cycloetherification of the resulting alcohol. We have prepared fluorinated AcAP analogues to aid in continued mechanistic investigation of the remarkable LolOcatalyzed cycloetherification step. LolO was able to hydroxylate 6,6-difluoro-AcAP (prepared from N,O-protected 4-oxoproline) and then cycloetherify the resulting alcohol, forming a difluorinated analogue of N-acetylnorloline and providing evidence for a cycloetherification mechanism involving a C(7) radical as opposed to a C(7) carbocation. By contrast, LolO was able to hydroxylate 7,7-difluoro-AcAP (prepared from 3-oxoproline) but failed to cycloetherify it, forming (1R,2R,8S)-7,7-difluoro-2hydroxy-AcAP as the sole product. The divergent LolO-catalyzed reactions of the difluorinated AcAP analogues provide insight into the LolO cycloetherification mechanism and indicate that the 7,7-difluorinated compound, in particular, may be a useful tool to accumulate and characterize the iron intermediate that initiates the cycloetherification reaction.

In mammals, iron(II)‐ and 2‐oxoglutarate‐dependent (Fe/2OG) dioxygenases have roles in oxygen and body‐mass homeostasis, connective tissue synthesis, and control of transcription and epigenetic inheritance.1 Related enzymes in plants, fungi, and bacteria enable diverse biosynthetic pathways to valuable natural‐product drugs by catalyzing halogenation, epimerization, desaturation, cyclization, ring‐opening/expansion, and endoperoxidation reactions; some of these enzymes even catalyze multiple reaction types within the same pathway!2 Almost every enzyme in this class initiates its reaction by using a common oxoiron(IV) (ferryl) intermediate, first demonstrated by the Penn State group in 2003,3,4 to abstract a hydrogen atom from its target substrate (Figure 1). The iron(II)‐chelating co‐substrate, 2OG, is oxidatively decarboxylated to a succinate ligand in formation of the ferryl complex. To mediate an outcome other than hydroxylation, an enzyme must avoid what can be a facile “oxygen rebound,” Groves term for the coupling of the substrate radical with the iron‐coordinated oxygen just after it abstracts the hydrogen. By direct biophysical observation and “metallomimicry” of intermediates, we have rationalized several of these outcomes in terms of the alternative fates of the substrate radical and the structural features of the enzyme that avert oxygen rebound to enable these alternative fates.5‐11 Very recently, we explained how the most unusual Fe/2OG oxygenase discovered to date, the microbial ethylene‐forming enzyme (EFE), branches from the canonical pathway even before the ferryl intermediate is formed (Figure 2),12,13 leading to global fragmentation of 2OG to three carbon dioxide equivalents and ethylene, a reaction that requires but does not transform the amino acid L‐arginine. The elucidation of the unusual “radical‐polar‐crossover” mechanism of EFE13 significantly expands the paradigm of 2OG‐assisted dioxygen activation pathways and suggests unexpected uses of EFE in biotechnology. 1. Hausinger, R. P. Crit. Rev. Biochem. Mol. Biol., 2004, 39, 21–68. 2. Bollinger, J. M., Jr., et al. 2015 in RSC Metallobiology Series No. 3. R.P. Hausinger and C.J. Schofield, eds. pp. 95‐122. Royal Society of Chemistry, Washington, D.C. 3. Price, J. C., et al. Biochemistry,2003,42,7497‐7508. 4. Price, J. C., et al. J. Am. Chem. Soc., 2003, 125, 13008‐13009. 5. Matthews, M. L., et al. Proc. Natl. Acad. Sci. USA, 2009, 106, 17723‐17728. 6. Chang, W.‐c., et al. Science 2014, 343, 1140‐1143. 7. Martinie, R. J., et. al. J. Am. Chem. Soc. 2015,137, 6912‐6919. 8. Martinie, R. J., et al. Inorg. Chem. 2017, 56, 13382‐13389. 9. Dunham, N. P. et al. J. Am. Chem. Soc. 2018, 140, 7116‐7128. 10. Dunham N. P., et al. J. Am. Chem. Soc. 2019, 141, 9964‐9979. 11. Pan, J., et al. J. Am. Chem. Soc.2019, 141, 15153−15165. 12. Copeland, R. A., et al. J. Am. Chem. Soc. 2021, 143, 2293‐2303. 13. Copeland, R. A., et al. Science2021, 373, 1489‐1493.