The research in the Cameron laboratory focuses on cytochrome P450 catalyzed metabolism.  Cytochrome P450 are best known as being the family of enzymes responsible for the breakdown of most pharmaceutical drugs. Additionally, P450 are important in key biological functions such as steroid and bile acid biosynthesis and oxidation of unsaturated lipids to numerous signaling molecules.

History: Cytochrome P450s were first reported in 1964 [1] and were reported to be the heme containing pigment protein responsible for the color of liver.  The name cytochrome P450 is derived from cytochrome + pigment + 450 where 450 refers to the distinctive UV absorption spectra maximum at 450 nm when reduced and adducted with carbon monoxide. Cytochrome P450 are heme-containing membrane associated proteins that are involved in oxygen activation. All cytochrome P450 have a heme prosthetic group that is liganded to the sulfur of a cysteine residue and have one plane of the heme solvent accessible.

Further experimentation revealed multiple P450 enzymes (human genome encodes fifty-seven P450s), primarily localized at the endoplasmic reticulum membrane, capable of
catalyzing the oxidation of a wide range of foreign molecules. Because of the extreme number of substrates oxidized and overlapping substrate specificity, it is not possible to name individual P450 enzymes for the reactions they catalyze. Instead, the naming convention is based upon sequence homology with enzymes assigned to families and sub families. While some enzymes within the same subfamily have similar reactivity (e.g. CYP3A4 and CYP3A5), others have little in common with respect to the reactions they catalyze (e.g. CYP2C8, CYP2C9, CYP2C18, and CYP2C19). To aid the confusion, numbering is usually, but not always based on the order a particular enzyme was sequenced irrespective of species. For example, humans only have a single P450 that is in the 2A family, but it is designated 2A6 (2A1, 2, and 3 are rat enzymes and 2A4, and 5 are mouse P450).

Reactive metabolites: Exposure to foreign chemicals from the environment is ubiquitous and the incredible structural diversity of these xenobiotics makes it impossible to evolve specialized enzymes to safely eliminate each foreign compound. Approximately a dozen P450s are exceptionally non-specific, capable of oxidizing thousands of substrates [3-6]. This high degree of non-specificity is achieved via harnessing the oxidative potential of reactive oxygen species, stabilized within the active site of the P450 [7-9]. These same organic molecules could have been degraded directly through reaction with the hydroxyl radical, an exceptionally strong oxidant; however, this mechanism cannot be controlled and would lead to excessive cellular toxicity. P450s “sequester” and control access to the oxidative species, thus decreasing the oxidation of unintended biomolecules. Because the P450 oxidative species is so strong, substrates can often be oxidized at more than one position and the metabolite profile has as much to do with the presentation of the molecule to the activated oxygen as it does the “easiest” site of oxidation based on redox potential [10-13]. This often leads to the catalysis of multiple metabolites from a single P450 enzyme or sequential metabolism giving rise to secondary and tertiary metabolites. Depending on the nature of the substrate, oxidations catalyzed by P450 can generate electron deficient metabolites which in turn oxidize cellular components. There are certain chemical moieties that are more prone to the generation of reactive metabolites and there have been several excellent reviews that have discussed “structural alerts” and correlated these to the formation of reactive metabolites and/or hepatotoxicity [14-17].

Work within the Cameron lab has examined the formation of chemically reactive metabolites generated by the metabolism of multiple pharmaceuticals, particularly kinase inhibitors, to elucidated the mechanism of reactive metabolite formation and to explain the associated toxicity. Formation of reactive metabolites were demonstrated in human liver samples. Additionally, generation of reactive metabolites in lung samples were found to be greater in smokers than non-smokers for some of these drugs.

In addition to demonstrating the mechanism of metabolic bioactivation, analogs were synthesized to demonstrate structural modifications that would prevent the formation of the reactive metabolite. Additional studies within the lab revealed mechanisms of cellular toxicity and protective pathways within the cell to mitigate the toxicity due to reactive metabolites. Five representative publications from the lab include: 

“Characterization of dasatinib and its structural analogs as CYP3A4 mechanism-based inactivators and the proposed bio-activation pathways.” Li X, He Y, Ruiz CH, Koenig M, Cameron MD. Drug Metabolism and Disposition, 2009, 37(6):1242-1250.  PMCID: PMC3202349.

“Bioactivation of the EGFR Inhibitor Gefitinib: Implications in pulmonary and hepatic toxicities.” Li X, Kamenecka TM, Cameron MD. Chemical Research in Toxicology, 2009 Oct;22(10):1736-42. PMID: 19803472.

“Cytochrome P450-mediated bioactivation of the epidermal growth factor receptor inhibitor erlotinib to a reactive electrophile.” Li X, Kamenecka TM, Cameron MD. Drug Metabolism and Disposition, 2010, Jul;38(7):      1238-45.  PMCID: PMC3202369.

“Potential role of a quetiapine metabolite in quetiapine-induced neutropenia and agranulocytosis.” Li X, Cameron MD. Chem Res Toxicol. 2012 May 21;25(5):1004-11. PMID: 22506851.

“P450 3A-Catalyzed O-dealkylation of Lapatinib Induces Mitochondrial Stress and Activates Nrf2.” Eno MR, El-Gendy BE, Cameron MD. Chem Res Toxicol. 2016 May 16;29(5):784-96. PMID:26958860.


CYP3A4 and CYP3A5 are responsible for the metabolism of approximately 40% of all prescription medications. CYP3A4 is ubiquitously expressed, appreciable CYP3A5 levels are present in only about 20% of Caucasians with the highest incidence of 70% in individuals from sub-Sahara Africa. While the two enzymes have been shown to have similar substrate specificity, there are known clinical differences linked to CYP3A5 genotype and no molecular tools were available to differentiate the activity of CYP3A4 and CYP3A5 in biological samples. The lack of proper chemical tools to differentiate the activity of CYP3A4 and CYP3A5 has led to the long-standing, yet erroneous, convention of treating the two enzymes as if they were one, sometimes referred to as the non-existent enzyme CYP3A. The Cameron lab was the first to discover and optimize tools to differentiate these two enzymes, publishing on the optimization and structural refinement of a CYP3A4 inhibitor with >1000-fold selectivity over CYP3A5.

“Imidazopyridines as selective CYP3A4 inhibitors.” Song X, Li X, Ruiz CH, Yin Y, Feng Y, Kamenecka TM, Cameron MD. Bioorg Med Chem Lett. 2012 Feb 15;22(4):1611-4. PMID: 22264486. “Discovery of a highly selective CYP3A4 inhibitor suitable for reaction phenotyping studies and differentiation of CYP3A4 and CYP3A5.” Li X, Song X, Kamenecka TM, Cameron MD. Drug Metab Dispos. 2012 Sep;40(9):1803-9. PMID: 22696420

Additionally, we discovered a highly selective CYP3A5 substrate which can clearly differentiate CYP3A5 activity from CYP3A4 even in samples such as primary human hepatocytes.

“Characterization of T-5 N-oxide Formation as the First Highly Selective Measure of CYP3A5 Activity.” Li X, Jeso V, Heyward S, Walker GS, Sharma R, Micalizio GC, Cameron MD. Drug Metab Dispos. 2014 Mar 42(3):334-42. PMCID: PMC3935135.

Both compounds have generated significant interest and are being utilized in drug metabolism, drug safety, and pre-clinical IND groups within the pharmaceutical industry.

The compounds can be purchased from Toronto Research Chemicals (although they are ridiculously expensive, note: we do not receive funds from the sale of any of these compounds). The selective CYP3A4 inhibitor is SR-9186 and the catalog number is C962345. The CYP3A5 selective substrate is T5 (or T1032) and is catalog number A622635. The selective N-oxide metabolite generated from CYP3A5 mediated oxidation of T5 is catalog number A622640. Full synthetic protocols for all compounds are published in the above references.

Currently, we are exploring beneficial CYP3A4 catalyzed reactions and the deleterious outcomes associated with long-term ablation of CYP3A4 activity. This is particularly relevant in HIV patients taking protease and integrase inhibitors as part of their HAART treatment where boosting agents are utilized that inactivate CYP3A4 to improve the oral bioavailability and pharmacokinetics of the protease and integrase inhibitors.

1.         Omura, T. and R. Sato, The Carbon Monoxide-Binding Pigment of Liver Microsomes. Ii. Solubilization, Purification, and Properties. J Biol Chem, 1964. 239: p. 2379-85.

2.         Berka, K., et al., Membrane position of ibuprofen agrees with suggested access path entrance to cytochrome P450 2C9 active site. J Phys Chem A, 2011. 115(41): p. 11248-55.

3.         Bu, H.Z., A literature review of enzyme kinetic parameters for CYP3A4-mediated metabolic reactions of 113 drugs in human liver microsomes: structure-kinetics relationship assessment. Curr Drug Metab, 2006. 7(3): p. 231-49.

4.         Niwa, T., et al., Comparison of kinetic parameters for drug oxidation rates and substrate inhibition potential mediated by cytochrome P450 3A4 and 3A5. Curr Drug Metab, 2008. 9(1): p. 20-33.

5.         Wilkinson, G.R., Cytochrome P4503A (CYP3A) metabolism: prediction of in vivo activity in humans. J Pharmacokinet Biopharm, 1996. 24(5): p. 475-90.

6.         Thummel, K.E. and G.R. Wilkinson, In vitro and in vivo drug interactions involving human CYP3A. Annu Rev Pharmacol Toxicol, 1998. 38: p. 389-430.

7.         Ogliaro, F., et al., Searching for the second oxidant in the catalytic cycle of cytochrome P450: a theoretical investigation of the iron(III)-hydroperoxo species and its epoxidation pathways. J Am Chem Soc, 2002. 124(11): p. 2806-17.

8.         Ortiz de Montellano, P.R., Cytochrome P-450 catalysis: radical intermediates and dehydrogenation reactions. Trends Pharmacol Sci, 1989. 10(9): p. 354-9.

9.         Popescu, D.L., et al., Thermodynamic, electrochemical, high-pressure kinetic, and mechanistic studies of the formation of oxo Fe(IV)-TAML species in water. Inorg Chem, 2010. 49(24): p. 11439-48.

10.       Cameron, M.D., et al., Cooperative binding of acetaminophen and caffeine within the P450 3A4 active site. Chem Res Toxicol, 2007. 20(10): p. 1434-41.

11.       Cameron, M.D., et al., Cooperative binding of midazolam with testosterone and alpha-naphthoflavone within the CYP3A4 active site: a NMR T1 paramagnetic relaxation study. Biochemistry, 2005. 44(43): p. 14143-51.

12.       Hummel, M.A., et al., Effector-mediated alteration of substrate orientation in cytochrome P450 2C9. Biochemistry, 2004. 43(22): p. 7207-14.

13.       Kirchmair, J., et al., Computational prediction of metabolism: sites, products, SAR, P450 enzyme dynamics, and mechanisms. J Chem Inf Model, 2012. 52(3): p. 617-48.

14.       Dowty, M.E., et al., Drug design structural alert: formation of trifluoroacetaldehyde through N-dealkylation is linked to testicular lesions in rat. Int J Toxicol, 2011. 30(5): p. 546-50.

15.       Lewis, D.F., C. Ioannides, and D.V. Parke, A combined COMPACT and HazardExpert study of 40 chemicals for which information on mutagenicity and carcinogenicity is known, including the results of human epidemiological studies. Hum Exp Toxicol, 1998. 17(10): p. 577-86.

16.       Kalgutkar, A.S. and J.R. Soglia, Minimising the potential for metabolic activation in drug discovery. Expert Opin Drug Metab Toxicol, 2005. 1(1): p. 91-142.

17.       Kalgutkar, A.S., et al., A comprehensive listing of bioactivation pathways of organic functional groups. Curr Drug Metab, 2005. 6(3): p. 161-225.

18.       Evans, W.E. and M.V. Relling, Pharmacogenomics: translating functional genomics into rational therapeutics. Science, 1999. 286(5439): p. 487-91.