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Occurrence of flavin-containing monooxygenases in non-mammalian eukaryotic organisms1
Comparative Biochemistry and Physiology Part C 121 (1998) 185 – 195
Review
Occurrence of flavin-containing monooxygenases in non-mammalian eukaryotic organisms1 Daniel Schlenk * Department of Pharmacology and En6ironmental Toxicology Program, Research Institute of Pharmaceutical Sciences, School of Pharmacy, Uni6ersity of Mississippi, Oxford, MS 38677, USA Received 19 January 1997; received in revised form 24 July 1997; accepted 6 August 1997
Abstract Flavin-containing monooxygenases (FMOs) have been identified in various organisms from bacteria to humans. However, because of the importance of these enzymes in the biotransformation of xenobiotics, the majority of studies have focused almost entirely upon the mammalian forms of the enzyme. Consequently, this review is an attempt to document the occurrence of FMO expression (mRNA, proteins, activities) in non-mammalian species in an attempt to provide insight about its putative physiological and toxicological roles. Activity indicative of FMO has been observed in numerous invertebrate species but corresponding proteins or transcripts have not been identified. There is a significant gap of information pertaining to insects, echinoderms, avian, reptilian and amphibian species. Significant homology of structure and function is observed in lower vertebrates. Evidence is provided primarily from studies with piscine forms of the enzyme suggesting a possible osmoregulatory role of FMOs, especially in euryhaline species of fish. © 1998 Elsevier Science Inc. All rights reserved. Keywords: Flavin-containing monooxygenase; Trimethylamine; Osmoregulation; Bioactivation; Biotransformation; Evolution
1. Introduction Monooxygenases typically transfer one atom from molecular oxygen utilizing a host of co-substrates such as pterins, copper, iron-containing heme (such as cytochrome P450s) and flavins. Flavin-containing monooxygenases (FMOs; E.C.1.14.13.8) catalyze the four-electron reduction of dioxygen with two electrons derived from a reduced nicotinamide cofactor and two electrons derived from substrate [39]. FMOs occur in several tissues of multicellular organisms and have been * Tel.: +1 601 2325150; fax: +1 601 2325148; e-mail: [email protected] 1 This article was invited by Guest Editors Dr John J. Stegeman and Dr David R. Livingstone to be part of a special issue of CBP on cytochrome P450 (Comp. Biochem. Physiol. 121 C, pages 1 – 412, 1998).
observed in a host of animals including bacteria and humans. The following review will attempt to focus on the mechanism, occurrence, toxicological and physiological significance of FMOs in non-mammalian eukaryotic organisms.
1.1. Mechanism Because of its toxicological and pharmacological importance, the majority of information regarding FMO such as nomenclature and catalytic mechanism has been derived from mammalian studies [24,25,32,39, 62,63]. By understanding the mechanism of FMO catalysis, a better insight into the substrate specificity and role of FMO in xenobiotic biotransformation can be attained. From steady-state and stopped flow kinetic studies with the purified mammalian and bacterial enzymes, it has been determined that the enzyme forms a
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Michaelis complex with NADPH which leads to a rapid reduction of the enzyme-bound flavin. The reduced enzyme-NADP + complex then binds oxygen which adds to the 4a-position of the flavin to generate a hydroperoxy flavin which is common to all of the known aromatic flavin hydroxylases, as well luciferase and cyclohexanone monooxygenase enzymes [5,39]. One significant difference between FMOs and other monooxygenases bearing flavin or other prosthetic groups, is that the oxygenatable substrate for FMO is not required for dioxygen reduction by NADPH [63]. Thus, since FMO resides in the cell as a highly reactive 4a-hydroperoxyflavin form, any soft nucleophile that can make contact with this potent monooxygenating agent will be oxidized. The product formed by oxygen transfer from the hydroperoxyflavin to the nucleophile is released immediately. Because the energy required to drive the reaction is present in the enzyme before it encounters the xenobiotic, precise fit usually required to lower the energy of activation of an enzyme catalyzed reaction, is not necessary such that FMOs catalyze the oxidation of compounds as dissimilar as iodide, boronic acids, phosphines, most functional groups bearing sulfur or selenium, a host of synthetic and naturally occurring amines and hydrazines, as well as aromatic aldehydes with adjacent hydrogen acceptors through Baeyer–Villiger chemistry which originally described the oxidation of ketones by peracids to esters [4,9,39,63].
acterization of FMO activity have utilized compounds or incubation conditions that inhibit P450 or FMO in mammals and assumed their actions in non-mammalian systems are similar. However, mammalian FMO1 can be inactivated by heating microsomal incubations at 45°C for 5 min [62], whereas heating invertebrate samples at 45°C tends to destroy P450 as well as any other enzyme. Even in mammalian systems, it is difficult to find substrates that specifically inhibit FMO without also the inactivation of P450. For example, several invertebrate characterization studies have used methimazole as an inhibitor or substrate for FMO in microsomal preparations [6,28–30,34,44]. However, in rat liver microsomes, methimazole is bioactivated by FMO and subsequently inactivates P450 [12,26,27]. Likewise, methimazole also inhibits peroxidase activity as well [13,36] which co-occurs in microsomal and cytosolic fractions used to measure invertebrate and mammalian FMO [44,45,51]. In summary, the inhibition of a particular FMO-catalyzed reaction by methimazole should not be used as exclusive evidence of FMO activity. Clearly, multiple FMO inhibitors that are structurally diverse should be used in in vitro incubations, especially with invertebrate animals. If possible, studies that correlate FMO activity with FMO protein or mRNA expression using homologous probes from mammals are more appropriate. Unfortunately, mammalian probes only seem to recognize vertebrate proteins and mRNAs [47].
1.2. Location and diagnostic substrates
1.3. Substrate specificity
FMOs are typically membrane-bound enzymes found in the smooth endoplasmic reticulum of the cell, although cytosolic forms predominate in bacteria and unicellular organisms [1,39]. In multicellular organisms, FMO activity is normally greatest in the digestive gland/hepatopancreas/liver of organisms [62,63]. However, tissue distribution of activity and expression may vary depending on the life history of the organism. For example, in euryhaline fish (wide-salinity tolerance), FMO expression and activity is greater in the gill and kidney than the liver, whereas, stenohaline fish (narrow salinity tolerance) tend to have higher levels of FMO in the liver [54]. This phenomenon and its relationship to physiological function will be discussed below. FMO resides in the same intracellular organelle and requires the same cofactors as the cytochrome P450 monooxygenase (NADPH and oxygen) [62,63]. In addition, several substrates used to assay for FMO are also substrates for P450 [62,63]. Thus, it is often necessary to verify in vitro enzyme activity through inhibition of P450 as well as other oxygenases that may contribute to the oxygenation of the substrate. Since little is known regarding specific P450 inhibitors or FMO substrates in non-mammalian species [58], studies involving the char-
In non-mammalian systems few substrates for FMO have been identified compared to the mammalian enzymes (Table 1). Agents that have been identified as substrates in non-mammalian systems include thioethers (aldicarb), thiocarbamates (eptam, thiobencarb), thiocarbamides (methimazole, thiourea), thioamides (thiobenzamide), tertiary amines (trimethylamine, N,N-dimethylaniline), substituted hydrazines (dimethyl and diphenylhydrazine) and aryl-amines (2aminofluorene (2-AF), 2-aminoanthracene (2-AA)). Substrate specificity has been determined directly using spectrophotometric assays or measurement and identification of oxygenated metabolites. In addition, several groups have measured FMO activity indirectly by the Ames assay using putative FMO inhibitors or pH optima to, respectively, eradicate or enhance responses [1,2,4,16,38,40,41,43 –47,54,59]. Trimethylamine (TMA) has been, by far, the most thoroughly used substrate among non-mammalian species to identify FMO activity [2,4,16,59]. However, the most extensive survey of TMA oxidase activity used crude tissue homogenates to measure activity, rather than the microsomal fraction resulting from ultracentrifugation [4]. Consequently, most studies were not performed with a focus on FMO
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Table 1 Invertebrate and vertebrate species in which FMO activity, protein, or mRNA have been observed Species
mRNA
Invertebrates Cunningamella elegans Calanus finmarchius Purified enzyme Trypanosoma cruzi Purified enzyme
Protein
FMO1 two subunits: 52 kD each
Mytilus edulis Digestive gland Carcinus maenus Digestive gland Mytilus gallopro6incialis Digestive gland Crassostrea gigas Viscera Gill Cryptochiton stelleri Digestive gland
Geodia cydonium Whole sponge Tethya aurantium Whole sponge Verongia aerophoba Whole sponge Pellina semitubulosa Whole sponge Fish Cyprinus carpio Liver Morone saxatilis Liver Gadus morhua liver L. limanda Kidney Squalus acanthias Liver
FMO1: 1.4 kb
FMO1: 1.1 kb FMO2: 50 kD
Raja erinacea Liver Ginglymostoma cirratum Liver Oryzias latipes Liver Gill
FMO1: 57 kD
Activity nmol min−1 mg−1
Reference
Pyrilamine N-oxidase: 2.5
Schlenk unpublished
TMA oxidase: 85
[59]
DMA oxidase: 114
[1]
DMA (indirect) DMA oxidase: 0.4-0.7
[29] [33]
2AAF (indirect)
[34]
2-AAF (indirect)
[28]
2-AF (indirect) DMA oxidase: 0.024 90.01; 0.007 9 0.028
[44] [45]
2-AF (indirect) Dimethyl-hydrazine phenyl-hydrazine methimazole DMA oxidase: 0.05 90.02
[41]
2-AA (indirect)
[30]
2-AA (indirect)
[30]
2-AAF (indirect)
[30]
2-AAF (indirect)
[30]
AF, 2-AAF (indirect)
[28]
Thiobencarb S-oxygenase: 108 9 5 TMA oxidase (0.192)
[8] [4]
TMA oxidase: 1.7-5 90.15
[2]
TMA oxidase: 0.07
[2]
TMA oxidase: 0.00007 DMA oxidase: 0.0018 0.639 0.27
[2,47] [16] [46]
TMA oxidase: 0.00009 DMA oxidase: 0.009
[16]
TMA oxidase: 0.105 DMA oxidase: 0.0173
[18]
DMA oxidase: 0.276 Thiourea oxidase: 1.15 90.04
[48] Schlenk and El-Alfy unpublished
D. Schlenk / Comparati6e Biochemistry and Physiology, Part C 121 (1998) 185–195
188 Table 1 (continued) Species
mRNA
Protein
Activity nmol min−1 mg−1
Reference
[18]
FMO1 56-59 kD 56-59 kD 56-59 kD
TMA oxidase: 0.135 DMA oxidase: 0.024 DMA oxidase 0.456 90.343 0.067 9 0.026 0.121 90.109 DMA oxidase
[54,38]
Negaprion bre6irostris Liver Platichthys flesus Gill Liver Kidney Scophthalmus maximus Liver Gill Oncorhyncus mykiss Liver
FMO1 2.5 kb 3.0 kb
FMO1: 3.0 kb
FMO1: 56 kD
FMO1: 3.7 kb
FMO1/FMO2 57/61 kD
Kidney
57/61 kD
Gill
57/61 kD
Intestine
Carcharinus falciformus Liver
FMO2: 50 kD FMO1: 1.1 kb
Anguilla japonica Liver Kidney Poecilia reticulata Whole body Mustelus californicus Liver Leptocottus armatus Liver Porichthys notatus Liver Ophiodon elongatus Liver Hippoglossus stenolepis Liver Psettichthys melanostictus Liver Alosa sapidissima Liver Salmo gairdneri Lepomis cyanellus Liver Lepomis macrochirus Liver
[54]
0.24 90.026 0.186 9 0.015 DMA oxidase: 0.45 Thiourea oxidase: 0.845 Methimazole oxidase: 0.85 TMA oxidase: 0.058 DMA oxidase: 0.5 Thiourea oxidase: 0.51 90.15 Methimazole oxidase: 1.25 DMA oxidase: 0.05 Thiourea oxidase: 0.09 9 0.01 Methimazole oxidase: 0.55 DMA oxidase: 0.1 Thiourea oxidase: 0.1 Methimazole oxidase: 0.35
[40] [48] [49] [4] [49]
DMA oxidase: 5.16 90.89 Methimazole (indirect) TMA oxidase 0.09 0.04
[46] Schlenk unpublished [11]
TMA oxidase: 0.02
[11]
TMA oxidase: 0.285
[4]
TMA oxidase: 0.28
[4]
TMA oxidase: 0.278
[4]
TMA oxidase: 0.07
[4]
TMA oxidase: 0.173
[4]
TMA oxidase: 0.07
[4]
TMA oxidase: 0.045 TMA oxidase: 0.058
[4] [4]
TMA oxidase: 0.042
[4]
TMA oxidase: 0.022 DMA oxidase: 0.152 Thiourea oxidase: 0.255
[4]
[49]
[49]
D. Schlenk / Comparati6e Biochemistry and Physiology, Part C 121 (1998) 185–195
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Table 1 (continued) Species
mRNA
Protein
Micropterus salmoides Liver Aplodinotus grunniens Liver Reptiles Alligator mississippiensis Liver Chryemys pictapicter Liver Avian Gallus domesticus Liver
Activity nmol min−1 mg−1
Reference
Thiourea oxidase: 0.076
[48]
Thiourea oxidase: 0.38
[48]
FMO1: 55 kD
Schlenk and Winston unpublished
FMO1: 55 kD
Schlenk and Stegeman unpublished
FMO1: 55 kD
Amphibia Rana pipiens Liver Rana catesbeiana Liver
0.155
Schlenk unpublished [4]
TMA oxidase: 0.045
[4]
TMA oxidase: 0.108
[4]
(indirectly), implies measurement other than metabolite identification and quantification.
and because of the crude assay techniques, it is quite likely false negatives may have been observed.
2. FMO diversity in non-mammalian species As only three sequences have been identified from a non-mammalian organism (Caenorhabditis elegans) [61], the nomenclature for FMO isoform identification has been created totally based on the mammalian literature with five gene families identified (labeled FMO1– 5) encoding five unique proteins with differing substrate specificities, tissue distributions and regulatory mechanisms [32]. However, recent studies in two laboratories utilizing reverse-transcriptase polymerase chain reaction (RT-PCR) have identified products from eel (Anguilla sp.; Dolphin C, unpublished) and a 850-bp fragment from rainbow trout (Oncorhynchus mykiss) liver (Schlenk D, unpublished). The latter was recognized by a 1.6-kb FMO1 cDNA by Southern blot analyses under stringent conditions. Sequencing of the 850 bp fragment revealed that the product was 97 and 88% homologous to rabbit liver FMO1 and pig liver FMO1, respectively (El-alfy and Schlenk, submitted). Similar RT-PCR products were obtained from Tilapia and Stripped bass (Morone sp.) hepatic RNA, but have yet to be sequenced (Schlenk D, unpublished). Northern blot analyses with RNA from several tissues of smooth dogfish (Squalus acanthias), rainbow
trout (O. mykiss), stripped bass (Morone saxatilis), Atlantic flounder (Platichthys flesus), and turbot (Scophthalmus maximus) probed with a 1.6-kb FMO1 cDNA under non-stringent conditions indicated RNA bands ranging from 1.1 to 3 kb (Table 1). In addition, there were no significant differences between males and females in hepatic FMO1-like mRNA in the turbot (S. maximus) [38]. Northern analyses using FMO1 and FMO2 cDNAs in channel catfish (Ictalurus punctatus) failed to show hybridization even under extremely nonstringent hybridization conditions (i.e. room temperature; Schlenk D, unpublished data). FMO genes are clearly present in several non-mammalian organisms. Recent advances in recombinant DNA methodologies, should allow identification and classification of these genes in the near future. Such studies will allow a better understanding of the evolution and regulation of FMO. Although fish have primarily been examined for expression of FMO RNA, several other phyla have been examined at the protein level by Western blot analyses using antibodies raised against the mammalian proteins (primarily FMO1 and FMO2). The overall trend is that structurally related proteins have been found exclusively in vertebrates. Invertebrates that have been examined include the barnacle (Balanus sp.), gumboot chiton (Cryptochiton stelleri ), the western oyster (Crassostrea 6irginica) and the sea anemone (Anthopleura elegantissima; Schlenk D, unpublished). However, numerous vertebrate species have been shown to express
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microsomal proteins that range from 50 to 60 kD that are recognized by polyclonal antibodies to FMO1 or FMO2. Non-mammalian vertebrates that have shown structurally similar microsomal proteins include the reptilians American alligator (Alligator mississippiensis), the eastern painted turtle (Chrysemys picta picta) and the avian chicken (Gallus domesticus; Table 1). In addition, numerous fish species have been examined and have been shown to express FMO1- or FMO2-like proteins. Included are sharks (S. acanthias and Carcharhinus falciformis), and teleosts with proteins from the rainbow trout (O. mykiss), Atlantic flounder (P. flesus) and turbot (S. maximus) being the best characterized [38,40,42,46,54]. In contrast to mammals, there does not appear to be immunologically distinct isoforms in fish, since antibodies raised against FMO1 and FMO2 recognize the same proteins in all tissues of the few species examined [38,42,46,54]. However, antibodies raised against FMO2 do appear to show less nonspecific binding and stronger signals in Western blots in most fish species [38,42,46,54]. Differences in number and size of microsomal proteins in various tissues recognized by the mammalian antibodies have also been observed. For example, kidney and gill of the Atlantic flounder expressed two FMO2-like proteins, while the liver expressed a single form [54]. Earlier studies in rainbow trout demonstrated two isoforms in several tissues including gill, kidney and liver [42,49]. More recent studies indicate the presence of perhaps three or four isoforms in liver (Schlenk et al., submitted manuscript). In addition, expression of the hepatic isoforms in trout increases with the age of the animal and there does not appear to be any sexual differences in activity but some differences may be present in isoform content between male and female animals [42]. In flounder as well as turbot, expression of hepatic FMO protein was not sexually dimorphic with males and females having similar levels of expression [38,54]. In contrast to these species above, there are many fish species that do not appear to express FMO1- or FMO2-like proteins [4,48]. Common features of these species include: (i) predominantly freshwater origin; (ii) possessing relatively low levels of trimethylamine or TMAO; and (iii) unable to tolerate significant salinity alterations. Without exception, vertebrate animals, such as the little skate (Raja erinacea) [4] or the channel catfish (I. punctatus) [55] lacking an FMO1 or FMO2like protein also fail to demonstrate FMO activity. As discussed above, few compounds have been identified as FMO substrates in non-mammalian species. Prior to the isolation and characterization of FMO from mammals, unless an enzyme had been purified, monooxygenase activities were ascribed to the term ‘mixed function oxidase’ or prefaced by the agent undergoing the oxidation. For example, TMA oxidase has been identified as an enzyme activity since studies
showed the formation of trimethylamine oxide (TMAO) in goldfish (Carassius auratus) and eels (Anguilla japonica) from TMA [21,22]. An association between TMA oxidase and FMO was not observed until 1973, when Goldstein and Dewitt-Harley [16] demonstrated a similarity between TMA oxidase and FMO enzymes in the nurse shark liver (Ginglymostoma cirratum). The first comprehensive review of in vitro TMA oxidase activity was performed by Baker et al. in 1963 [4] in which one bird, one reptile, three amphibians, four elasmobranchs, 15 saltwater teleosts, three anadromous fish taken from fresh water, six freshwater teleosts, four bacteria, three unicellular microbes, one hydra, one planaria, one annelid, one mollusc, four arthropods and 17 plant species were examined. FMO activity has subsequently been observed in several species not demonstrating appreciable activity in these earlier studies. These discrepancies may be explained by more sensitive assays and subcellular fragments used in the more recent studies. Examples include the smooth dogfish (S. acanthias) and rainbow trout (O. mykiss) [40,46]. Likewise, Baker et al. [4] failed to observe activity in any invertebrate species examined. In studies with the insects Heliothis 6irescens and Musca domestica oxidation of DMA was not observed (Hodgson E, personal communication). However, studies with the marine copepod (Calanus finmarchicus), the gumboot chiton (Cryptochiton stelleri ), the western oyster (Crassostrea gigas), the mussels (Mytilus edulis and Mytilus gallopro6incialis), the marine sponges (Geodia cydonium, Tethya aurantium, Verongia aerophoba and Pellina semitubulosa) and the fungus (Cunninghamella elegans) indicate activity consistent with flavin-containing monooxygenases (Table 1). The enzyme was partially purified from C. finmarchicus and was shown to be a cytosolic enzyme of approximately 130 kD with a Km for TMA of 0.3 mM [59]. A cytosolic form of DMA N-oxygenase was also isolated from the protozoan Trypanosoma cruzi having a molecular weight of 100 kD consisting of two 52 kD subunits [1]. The form of FMO isolated from T. cruzi had a pH optimum of 8.0 for the oxidation of NADPH, a Km of 56 mM and a Vmax of 114 nmol min − 1 mg − 1. The enzyme also appeared to have lower apparent affinity constants for sulfur-containing compounds than DMA with respective Km values for disulfoton and methimazole of 0.22 and 0.5 mM, whereas DMA had a relatively high Km of 1 mM [1]. Although TMA oxidase and other FMO catalyzed activities have been observed in several invertebrates, the majority of studies have focused on the characterization of activity in vertebrate fishes [47]. As mentioned above, Goldstein and Dewitt-Harley [16] initially characterized TMA oxidase from the liver of the nurse shark (G. cirratum). For the first time in a non-mammalian species, it was noted that FMO enzyme activity
D. Schlenk / Comparati6e Biochemistry and Physiology, Part C 121 (1998) 185–195
was localized in the microsomal fraction of the cellular homogenate, was dependent upon NADPH and unaffected by azide or carbon monoxide. Reaction rates were optimal at pH 9.0 and inhibited by DMA and chlorpromazine (which are two prototypical FMO substrates). The Michealis affinity constant for TMA was calculated to be 1.5 mM. Studies in the coelacanth Latimeria chalumnae also contained an ‘elasmobranchlike’ hepatic TMA oxidase [18]. It is interesting to note that coelacanths are physiologically similar to elasmobranchs having relatively high concentrations of TMAO and urea in tissues [60]. The next characterization of TMA oxidase was reported by Agustsson and Strom [2] in the cod (Gadus morhua). The hepatic enzyme activity was shown to have a lower pH optimum of 8.2, and Km (11 mM) than the nurse shark enzyme. TMA oxygenase was also observed in the kidney, but not the liver or other tissues of the teleost flatfish Limanda limanda and Pleuronectes platessa [2]. Following these studies, TMA oxygenase activity was measured in several other fish species, but not well characterized. TMA oxidase was observed in the liver of guppy (Poecilia reticulata) and the eel (A. japonica) in studies focused on the source and function of TMA and TMAO in euryhaline fish [11]. Although TMA oxidase has been observed in all of these organisms and correlates with FMO-like protein expression, TMA oxidase have never been purified from a nonmammalian vertebrate. At approximately the same time TMA oxidase was being examined in guppy and eel, studies were underway characterizing FMO activity in the rainbow trout [40]. Hepatic FMO activity (DMA N-oxygenase) in trout was microsomal, NADPH-dependent, competitively inhibited by equimolar concentrations of TMA or aldicarb and non-competitively inhibited by methimazole [40]. The optimum pH for DMA N-oxygenation was 9.2 and was extremely sensitive to temperature and unaffected by co-incubation of n-octylamine [40]. As noted above, there were no significant sexual differences in hepatic FMO activity as measured by DMA N-oxygenase or by methimazole S-oxidase [42]. Each of these activities were observed in gill, kidney, heart and intestine microsomal preparations [49]. Although activity was highly variable among individuals with as much as 100% variation observed from hatchery raised trout, DMA oxygenase directly correlated with hepatic proteins recognized by anti-FMO1 and anti-FMO2 [49]. In addition, contrasting studies by Daikoku et al. [11], intraperitoneal treatment with TMA failed to change expression of FMO protein or DMA N-oxygenase activity in rainbow trout [42]. Likewise, intraperitoneal treatment of turbot or flounder with TMA failed to increase hepatic FMO activity [38,52]. In the early survey studies of Baker et al. [4] smooth dogfish (S. acanthias) failed to show TMA oxidase
191
activity. However, in recent years, smooth dogfish have been shown to have significant levels of hepatic FMO activity that is competitively inhibited by TMA or thiobenzamide and non-competitively inhibited by methimazole [46]. The consistent relationship of inhibition kinetics between trout [40] and smooth dogfish indicates significant structural homology in active sites between the two species. DMA N-oxygenase of the smooth dogfish shark was sensitive to temperature with a 76% loss of activity with a reduction of temperature from 25 to 15°C, and a 99% loss when incubations occurred at 45°C. The optimal pH for DMA N-oxygenase was 9.6 and the maximum velocity was calculated to be 1.3 nmol min − 1 mg − 1 with an apparent Km of 44 mM [46]. To further examine the phylogenetic distribution of FMO in fish, FMO activity, protein and mRNA was characterized in liver microsomes from the turbot (S. maximus) [38]. Hepatic DMA N-oxygenase was optimal at pH 8.8, possessed an apparent Km of 88 mM and was inhibited by methimazole and TMA. In comparative studies with the flounder (P. flesus), it was observed that turbot possessed significantly greater levels of FMO activity in the liver relative to the flounder which was barely above detection [54]. However, the flounder had significantly elevated levels of FMO activity, protein and mRNA in the gill relative to the turbot. Comparison of the enzymatic properties of the P. flesus gill and S. maximus liver enzymes indicated dramatic differences in Km between gill and liver, but were both inhibited by equimolar concentrations of TMA. Gill microsomal activity in each species was unaffected by the mammalian FMO2 substrate, n-octylamine. FMO activity of the gill from another euryhaline fish, the Japanese medaka (Oryzias latipes) is also significantly higher than the hepatic enzyme (Schlenk and El-alfy, submitted manuscript). The differential expression of FMO in euryhaline versus stenohaline fish indicate a potential relationship with osmoregulatory function.
3. Physiological significance Indeed, as mentioned above, a large number of organisms do not possess FMO activity. An early hypothesis for the scattered distribution of the enzyme suggested a random deletion of the gene coding for the enzyme [17]. Other hypotheses include a specific role in chemical defense particularly in sponges and molluscs [30] as well as catalyzing the oxidation of phytoplankton-derived dimethylsulfide to the S-oxide by bivalve molluscs [58]. Several endogenous substrates have been observed in mammals: primarily thio-ether conjugates as well as methionine, cysteamine and TMA [15,62,63]. In fact, a genetic polymorphism has been described in humans that lack FMO3 activity and are unable to
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N-oxygenate odorous TMA to non-odorous TMAO [14]. The disease has been named ‘fish odor syndrome’ due to the excretion of unmetabolized TMA across the skin of the individual [3]. Based on the distribution of the enzyme in fish species, another possible physiological function for FMO may be osmoregulation [47]. In addition to the differential expression of the enzyme in the osmoregulatory organ of the gill in the euryhaline flounder P. flesus [54], other reasons for this hypothesis include the direct correlation of FMO with concentrations of the osmolyte, TMAO. Like FMO expression, TMAO is found in relatively large amounts in marine fishes, but in considerably smaller amounts in freshwater fish [19,23]. The average body fluid levels of TMAO in 72 marine teleosts ranged from 0 to 143 mg TMAO (100 g wet wt.) − 1 with a mean value of 54.9 926.1 mg TMAO (100 g wet wt.) − 1, whereas 21 freshwater teleosts had values that ranged from 0 to 17 mg TMAO (100 g wet wt.) − 1 with a mean of 7.595.4 mg TMAO (100 g wet wt.) − 1 [19,23]. It has been shown that marine and euryhaline fish utilize TMAO as an intracellular osmolyte in the marine environment [60]. A direct correlation between TMAO and salinity was observed in the euryhaline flounder, Pleuronectes flesus [31] and the eel Anguilla anguilla [10,11]. Lastly, other evidence supporting an osmoregulatory role for the enzyme is the regulation of enzyme expression by salinity alteration in euryhaline fish. Recent studies with the Atlantic flounder (P. flesus) have shown that when saltwater adapted flounder are placed in freshwater diluted seawater for as short a period as 1 week, more than 90% reduction of hepatic and gill FMO activity was observed [52,53]. In other studies with Japanese medaka, raising the salinity from 0.15 to 2.0% induced FMO activity more than four-fold in the gill after 24 h (Schlenk and El-alfy, submitted manuscript). In support of these studies, Daikoku et al [11] observed a two to ten-fold increase in hepatic TMA oxidase activity when guppies or eels were placed from low to high salinity. In contrast, saltwater adapted stripped bass (M. saxatilis) actually had 23% lower levels of putative FMO-catalyzed S-oxidase activity than freshwater stripped bass [7]. Although discrepancies are present, there appears to be a significant association between FMO and osmoregulation, especially with the gill enzymes. Little is known regarding the regulation of FMO, even in mammalian systems [56,57]. Consequently, how salinity alterations modify FMO expression is unclear. Preliminary studies with sex steroids such as testosterone and 17b-estradiol in juvenile and sexually mature rainbow trout show significant reduction of hepatic FMO activity [49] which parallels expression of Na + K + ATPase and subsequent saltwater adaptation [35]. Current studies in our laboratory are focusing on the effects of osmoregulatory
hormones and effectors on FMO expression at low and high salinity in euryhaline fish species in order to better understand the role FMO may have in osmoregulation in fish.
4. Toxicological significance Although the physiological significance of FMOs is unclear, FMOs clearly play an important role in the toxicity of various heteroatom containing xenobiotics in mammalian and non-mammalian species. The first indication of a toxicological role for FMO in nonmammalian organisms was reported in studies with the mussel M. edulis [29]. Microsomes as well as postmitochondrial fractions of the digestive gland were shown to oxidize NADPH in the presence of various FMO substrates such as methimazole, DMA, 2-AA and 2-AF at a pH optimum of 8.4 [29]. Postmitochondrial fractions of the digestive gland of Mytilis gallopro6incialis were also shown to activate aminoanthracene to mutagenic metabolites at pH 8.4 through an NADPH-dependent pathway [6]. Since mutagenicity of aminoanthracene was inhibited by co-incubation with methimazole, it was concluded that N-oxidation was occurring through an FMO-catalyzed pathway [6]. However, as discussed above, it should be noted that methimazole is not exclusively selective for FMO and may be activated by FMO to a cytochrome P450 inhibiting compound, the sulfinic acid. This indeed is apparent in studies suggesting that FMO is responsible for the catalysis of 2-acetylaminofluorene (2-AAF) to mutagenic metabolite in mussel digestive gland microsomes which was inhibited by co-incubation by methimazole [28]. It is unlikely that FMO would catalyze the N-oxygenation of an acetylated amine based on the electron-withdrawing properties of the adjacent carbonyl which greatly reduces the nucleophilicity of the amine and would subsequently reduce the likelihood of attack of the hydroperoxyflavin moiety of FMO [62,63]. One possible scenario may be the deacetylation of 2-AAF to 2-AF which may be N-oxygenated by FMO [20]. However, acetylases are typically cytosolic, not microsomal, and could only have occurred if microsomes were contaminated with cytosol [37]. Moreover, coincubation with acetylase inhibitors failed to reduce microsomal activation of 2-AAF to mutagenic metabolites [28]. Indeed, in similar studies using the postmitochondrial fraction from digestive glands of M. edulis, 2-AAF as well as other aromatic amines were also activated to mutagenic metabolites [34]. In contrast to the microsomal studies with M. gallopro6incialis which found no change with acetylase inhibitors, mutagenicity of 2AAF with M. edulis digestive gland was reduced by the acetylase inhibitor, paraoxon [34]. In addition, further characterization of other FMO substrates and their
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metabolites were carried out with M. edulis confirming the presence of FMO activity in digestive gland microsomes [33]. Studies with the Western Oyster (C. gigas) also indicated FMO catalyzed oxidation of the aromatic amine 2-AF to N-oxygenated metabolites which were measured directly and found to be reduced by co-incubation of methimazole [44]. However, other enzymatic co-oxidative pathways were also involved in the N-oxygenation of 2-AF by oyster visceral mass microsomes [44]. Four species of marine sponges (G. cydonium, T. aurantium, V. aerophoba and P. semitubulosa) were also able to catalyze the activation of aromatic amines to mutagenic metabolites through a NADPH-dependent pathway that was optimum at 8.4, again indicating characteristics consistent with FMO [30]. The ability to bioactivate aromatic amines to mutagenic metabolites suggests that organisms capable of such biochemistry should have higher incidences of genotoxic responses such as cancer. However, it is unknown whether FMOcontaining organisms are more or less susceptible to aromatic amine-induced carcinogenesis. Although little is known regarding the relationship between aromatic amine effects and FMO expression in non-mammalian organisms, a considerable amount of evidence suggests that fish that have high levels of FMO are significantly more susceptible to thioether pesticides than fish lacking FMO [48]. For example, in a series of in vitro and in vivo studies in rainbow trout, it was shown that one or more FMOs were responsible for the bioactivation of the thioether pesticide, aldicarb, to the more potent cholinesterase inhibitor, aldicarb sulfoxide [43,50]. Trout are approximately 100 times more sensitive to aldicarb toxicity than channel catfish which do not express FMO [48]. In vivo metabolism studies indicate that channel catfish are capable of converting aldicarb to the less toxic metabolite, aldicarb sulfone, which appears to be catalyzed via a cytochrome P450 pathway [48]. In addition, studies examining the effect of salinity of aldicarb toxicity in the Japanese medaka showed a direct effect of lethality and salinity when fish were exposed to 1 mg l − 1 aldicarb (Schlenk and El-Alfy, submitted manuscript). As noted above, a direct correlation was observed between salinity and FMO expression and activity in the same species. Consequently, it has been hypothesized that salinity increases the toxicity of aldicarb by inducing expression of FMO(s) that catalyze the bioactivation of aldicarb to the more toxic aldicarb sulfoxide. Clearly, further in vitro biotransformation studies are necessary to verify an increase in aldicarb sulfoxide production with salinity. FMOs as well as other monooxygenases have also been shown to catalyze the S-oxygenation of Eptam (ethyl N,N-dipropylthiocarbamate) in hepatic microsomes from fresh and saltwater stripped bass (M. saxatilis) [7]. The S-oxide of Eptam was shown to be an efficient carbamylating agent of protein thiols indicating
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potential toxicity of this metabolite to freshwater fish [7]. Similar studies were performed with the herbicide thiobencarb (p-chlorobenzyl N,N-diethylthiocarbamate) from hepatic microsome obtained from the same species and indicated that FMOs may also catalyze S-oxygenation to the S-oxide which was also a more efficient carbamylating agent than the parent compound reacting with thiol and amine nucleophiles [8].
5. Summary and future studies Although relationships have been observed between FMO expression and activity, catalytically active FMO has yet to be purified or cloned from a non-mammalian organism. Structural and genetic similarities appear to be present in vertebrates (i.e. mammalian antibody and cDNA probes recognize respective proteins and mRNAs that correlate with activity). However, the protein structure apparently is significantly different in invertebrates (mammalian probes do not recognize proteins or mRNAs), even though comparable enzyme activities are observed. Because the proteins have yet to be conventionally purified or expressed through recombinant DNA methodologies, the substrate specificities and molecular evolution of individual isoforms are unclear. In addition, until it can be shown that specific substrates are metabolized by isolated enzymes, correlative studies with bands of protein and mRNA as well as indirect studies using multiple inhibitors will have to suffice in characterizing enzyme expression in various non-mammalian organisms. Clearly, the isolation of catalytically active enzymes from non-mammalian organisms would allow the examination of multiple xenobiotics as substrates and provide a better understanding of the physiological and toxicological roles of FMO.
Acknowledgements I wish to thank all of the collaborators that provided samples, antibodies and discussions regarding the phylogenetic distribution of FMO. Key personnel include Abir El-Alfy, Drs John Stegeman, Gary Winston, David Williams, Ron Hines and David Livingstone. I would also like to thank the Marine Biological Association of the United Kingdom for their support of much of this research.
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Review
Occurrence of flavin-containing monooxygenases in non-mammalian eukaryotic organisms1 Daniel Schlenk * Department of Pharmacology and En6ironmental Toxicology Program, Research Institute of Pharmaceutical Sciences, School of Pharmacy, Uni6ersity of Mississippi, Oxford, MS 38677, USA Received 19 January 1997; received in revised form 24 July 1997; accepted 6 August 1997
Abstract Flavin-containing monooxygenases (FMOs) have been identified in various organisms from bacteria to humans. However, because of the importance of these enzymes in the biotransformation of xenobiotics, the majority of studies have focused almost entirely upon the mammalian forms of the enzyme. Consequently, this review is an attempt to document the occurrence of FMO expression (mRNA, proteins, activities) in non-mammalian species in an attempt to provide insight about its putative physiological and toxicological roles. Activity indicative of FMO has been observed in numerous invertebrate species but corresponding proteins or transcripts have not been identified. There is a significant gap of information pertaining to insects, echinoderms, avian, reptilian and amphibian species. Significant homology of structure and function is observed in lower vertebrates. Evidence is provided primarily from studies with piscine forms of the enzyme suggesting a possible osmoregulatory role of FMOs, especially in euryhaline species of fish. © 1998 Elsevier Science Inc. All rights reserved. Keywords: Flavin-containing monooxygenase; Trimethylamine; Osmoregulation; Bioactivation; Biotransformation; Evolution
1. Introduction Monooxygenases typically transfer one atom from molecular oxygen utilizing a host of co-substrates such as pterins, copper, iron-containing heme (such as cytochrome P450s) and flavins. Flavin-containing monooxygenases (FMOs; E.C.1.14.13.8) catalyze the four-electron reduction of dioxygen with two electrons derived from a reduced nicotinamide cofactor and two electrons derived from substrate [39]. FMOs occur in several tissues of multicellular organisms and have been * Tel.: +1 601 2325150; fax: +1 601 2325148; e-mail: [email protected] 1 This article was invited by Guest Editors Dr John J. Stegeman and Dr David R. Livingstone to be part of a special issue of CBP on cytochrome P450 (Comp. Biochem. Physiol. 121 C, pages 1 – 412, 1998).
observed in a host of animals including bacteria and humans. The following review will attempt to focus on the mechanism, occurrence, toxicological and physiological significance of FMOs in non-mammalian eukaryotic organisms.
1.1. Mechanism Because of its toxicological and pharmacological importance, the majority of information regarding FMO such as nomenclature and catalytic mechanism has been derived from mammalian studies [24,25,32,39, 62,63]. By understanding the mechanism of FMO catalysis, a better insight into the substrate specificity and role of FMO in xenobiotic biotransformation can be attained. From steady-state and stopped flow kinetic studies with the purified mammalian and bacterial enzymes, it has been determined that the enzyme forms a
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Michaelis complex with NADPH which leads to a rapid reduction of the enzyme-bound flavin. The reduced enzyme-NADP + complex then binds oxygen which adds to the 4a-position of the flavin to generate a hydroperoxy flavin which is common to all of the known aromatic flavin hydroxylases, as well luciferase and cyclohexanone monooxygenase enzymes [5,39]. One significant difference between FMOs and other monooxygenases bearing flavin or other prosthetic groups, is that the oxygenatable substrate for FMO is not required for dioxygen reduction by NADPH [63]. Thus, since FMO resides in the cell as a highly reactive 4a-hydroperoxyflavin form, any soft nucleophile that can make contact with this potent monooxygenating agent will be oxidized. The product formed by oxygen transfer from the hydroperoxyflavin to the nucleophile is released immediately. Because the energy required to drive the reaction is present in the enzyme before it encounters the xenobiotic, precise fit usually required to lower the energy of activation of an enzyme catalyzed reaction, is not necessary such that FMOs catalyze the oxidation of compounds as dissimilar as iodide, boronic acids, phosphines, most functional groups bearing sulfur or selenium, a host of synthetic and naturally occurring amines and hydrazines, as well as aromatic aldehydes with adjacent hydrogen acceptors through Baeyer–Villiger chemistry which originally described the oxidation of ketones by peracids to esters [4,9,39,63].
acterization of FMO activity have utilized compounds or incubation conditions that inhibit P450 or FMO in mammals and assumed their actions in non-mammalian systems are similar. However, mammalian FMO1 can be inactivated by heating microsomal incubations at 45°C for 5 min [62], whereas heating invertebrate samples at 45°C tends to destroy P450 as well as any other enzyme. Even in mammalian systems, it is difficult to find substrates that specifically inhibit FMO without also the inactivation of P450. For example, several invertebrate characterization studies have used methimazole as an inhibitor or substrate for FMO in microsomal preparations [6,28–30,34,44]. However, in rat liver microsomes, methimazole is bioactivated by FMO and subsequently inactivates P450 [12,26,27]. Likewise, methimazole also inhibits peroxidase activity as well [13,36] which co-occurs in microsomal and cytosolic fractions used to measure invertebrate and mammalian FMO [44,45,51]. In summary, the inhibition of a particular FMO-catalyzed reaction by methimazole should not be used as exclusive evidence of FMO activity. Clearly, multiple FMO inhibitors that are structurally diverse should be used in in vitro incubations, especially with invertebrate animals. If possible, studies that correlate FMO activity with FMO protein or mRNA expression using homologous probes from mammals are more appropriate. Unfortunately, mammalian probes only seem to recognize vertebrate proteins and mRNAs [47].
1.2. Location and diagnostic substrates
1.3. Substrate specificity
FMOs are typically membrane-bound enzymes found in the smooth endoplasmic reticulum of the cell, although cytosolic forms predominate in bacteria and unicellular organisms [1,39]. In multicellular organisms, FMO activity is normally greatest in the digestive gland/hepatopancreas/liver of organisms [62,63]. However, tissue distribution of activity and expression may vary depending on the life history of the organism. For example, in euryhaline fish (wide-salinity tolerance), FMO expression and activity is greater in the gill and kidney than the liver, whereas, stenohaline fish (narrow salinity tolerance) tend to have higher levels of FMO in the liver [54]. This phenomenon and its relationship to physiological function will be discussed below. FMO resides in the same intracellular organelle and requires the same cofactors as the cytochrome P450 monooxygenase (NADPH and oxygen) [62,63]. In addition, several substrates used to assay for FMO are also substrates for P450 [62,63]. Thus, it is often necessary to verify in vitro enzyme activity through inhibition of P450 as well as other oxygenases that may contribute to the oxygenation of the substrate. Since little is known regarding specific P450 inhibitors or FMO substrates in non-mammalian species [58], studies involving the char-
In non-mammalian systems few substrates for FMO have been identified compared to the mammalian enzymes (Table 1). Agents that have been identified as substrates in non-mammalian systems include thioethers (aldicarb), thiocarbamates (eptam, thiobencarb), thiocarbamides (methimazole, thiourea), thioamides (thiobenzamide), tertiary amines (trimethylamine, N,N-dimethylaniline), substituted hydrazines (dimethyl and diphenylhydrazine) and aryl-amines (2aminofluorene (2-AF), 2-aminoanthracene (2-AA)). Substrate specificity has been determined directly using spectrophotometric assays or measurement and identification of oxygenated metabolites. In addition, several groups have measured FMO activity indirectly by the Ames assay using putative FMO inhibitors or pH optima to, respectively, eradicate or enhance responses [1,2,4,16,38,40,41,43 –47,54,59]. Trimethylamine (TMA) has been, by far, the most thoroughly used substrate among non-mammalian species to identify FMO activity [2,4,16,59]. However, the most extensive survey of TMA oxidase activity used crude tissue homogenates to measure activity, rather than the microsomal fraction resulting from ultracentrifugation [4]. Consequently, most studies were not performed with a focus on FMO
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Table 1 Invertebrate and vertebrate species in which FMO activity, protein, or mRNA have been observed Species
mRNA
Invertebrates Cunningamella elegans Calanus finmarchius Purified enzyme Trypanosoma cruzi Purified enzyme
Protein
FMO1 two subunits: 52 kD each
Mytilus edulis Digestive gland Carcinus maenus Digestive gland Mytilus gallopro6incialis Digestive gland Crassostrea gigas Viscera Gill Cryptochiton stelleri Digestive gland
Geodia cydonium Whole sponge Tethya aurantium Whole sponge Verongia aerophoba Whole sponge Pellina semitubulosa Whole sponge Fish Cyprinus carpio Liver Morone saxatilis Liver Gadus morhua liver L. limanda Kidney Squalus acanthias Liver
FMO1: 1.4 kb
FMO1: 1.1 kb FMO2: 50 kD
Raja erinacea Liver Ginglymostoma cirratum Liver Oryzias latipes Liver Gill
FMO1: 57 kD
Activity nmol min−1 mg−1
Reference
Pyrilamine N-oxidase: 2.5
Schlenk unpublished
TMA oxidase: 85
[59]
DMA oxidase: 114
[1]
DMA (indirect) DMA oxidase: 0.4-0.7
[29] [33]
2AAF (indirect)
[34]
2-AAF (indirect)
[28]
2-AF (indirect) DMA oxidase: 0.024 90.01; 0.007 9 0.028
[44] [45]
2-AF (indirect) Dimethyl-hydrazine phenyl-hydrazine methimazole DMA oxidase: 0.05 90.02
[41]
2-AA (indirect)
[30]
2-AA (indirect)
[30]
2-AAF (indirect)
[30]
2-AAF (indirect)
[30]
AF, 2-AAF (indirect)
[28]
Thiobencarb S-oxygenase: 108 9 5 TMA oxidase (0.192)
[8] [4]
TMA oxidase: 1.7-5 90.15
[2]
TMA oxidase: 0.07
[2]
TMA oxidase: 0.00007 DMA oxidase: 0.0018 0.639 0.27
[2,47] [16] [46]
TMA oxidase: 0.00009 DMA oxidase: 0.009
[16]
TMA oxidase: 0.105 DMA oxidase: 0.0173
[18]
DMA oxidase: 0.276 Thiourea oxidase: 1.15 90.04
[48] Schlenk and El-Alfy unpublished
D. Schlenk / Comparati6e Biochemistry and Physiology, Part C 121 (1998) 185–195
188 Table 1 (continued) Species
mRNA
Protein
Activity nmol min−1 mg−1
Reference
[18]
FMO1 56-59 kD 56-59 kD 56-59 kD
TMA oxidase: 0.135 DMA oxidase: 0.024 DMA oxidase 0.456 90.343 0.067 9 0.026 0.121 90.109 DMA oxidase
[54,38]
Negaprion bre6irostris Liver Platichthys flesus Gill Liver Kidney Scophthalmus maximus Liver Gill Oncorhyncus mykiss Liver
FMO1 2.5 kb 3.0 kb
FMO1: 3.0 kb
FMO1: 56 kD
FMO1: 3.7 kb
FMO1/FMO2 57/61 kD
Kidney
57/61 kD
Gill
57/61 kD
Intestine
Carcharinus falciformus Liver
FMO2: 50 kD FMO1: 1.1 kb
Anguilla japonica Liver Kidney Poecilia reticulata Whole body Mustelus californicus Liver Leptocottus armatus Liver Porichthys notatus Liver Ophiodon elongatus Liver Hippoglossus stenolepis Liver Psettichthys melanostictus Liver Alosa sapidissima Liver Salmo gairdneri Lepomis cyanellus Liver Lepomis macrochirus Liver
[54]
0.24 90.026 0.186 9 0.015 DMA oxidase: 0.45 Thiourea oxidase: 0.845 Methimazole oxidase: 0.85 TMA oxidase: 0.058 DMA oxidase: 0.5 Thiourea oxidase: 0.51 90.15 Methimazole oxidase: 1.25 DMA oxidase: 0.05 Thiourea oxidase: 0.09 9 0.01 Methimazole oxidase: 0.55 DMA oxidase: 0.1 Thiourea oxidase: 0.1 Methimazole oxidase: 0.35
[40] [48] [49] [4] [49]
DMA oxidase: 5.16 90.89 Methimazole (indirect) TMA oxidase 0.09 0.04
[46] Schlenk unpublished [11]
TMA oxidase: 0.02
[11]
TMA oxidase: 0.285
[4]
TMA oxidase: 0.28
[4]
TMA oxidase: 0.278
[4]
TMA oxidase: 0.07
[4]
TMA oxidase: 0.173
[4]
TMA oxidase: 0.07
[4]
TMA oxidase: 0.045 TMA oxidase: 0.058
[4] [4]
TMA oxidase: 0.042
[4]
TMA oxidase: 0.022 DMA oxidase: 0.152 Thiourea oxidase: 0.255
[4]
[49]
[49]
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Table 1 (continued) Species
mRNA
Protein
Micropterus salmoides Liver Aplodinotus grunniens Liver Reptiles Alligator mississippiensis Liver Chryemys pictapicter Liver Avian Gallus domesticus Liver
Activity nmol min−1 mg−1
Reference
Thiourea oxidase: 0.076
[48]
Thiourea oxidase: 0.38
[48]
FMO1: 55 kD
Schlenk and Winston unpublished
FMO1: 55 kD
Schlenk and Stegeman unpublished
FMO1: 55 kD
Amphibia Rana pipiens Liver Rana catesbeiana Liver
0.155
Schlenk unpublished [4]
TMA oxidase: 0.045
[4]
TMA oxidase: 0.108
[4]
(indirectly), implies measurement other than metabolite identification and quantification.
and because of the crude assay techniques, it is quite likely false negatives may have been observed.
2. FMO diversity in non-mammalian species As only three sequences have been identified from a non-mammalian organism (Caenorhabditis elegans) [61], the nomenclature for FMO isoform identification has been created totally based on the mammalian literature with five gene families identified (labeled FMO1– 5) encoding five unique proteins with differing substrate specificities, tissue distributions and regulatory mechanisms [32]. However, recent studies in two laboratories utilizing reverse-transcriptase polymerase chain reaction (RT-PCR) have identified products from eel (Anguilla sp.; Dolphin C, unpublished) and a 850-bp fragment from rainbow trout (Oncorhynchus mykiss) liver (Schlenk D, unpublished). The latter was recognized by a 1.6-kb FMO1 cDNA by Southern blot analyses under stringent conditions. Sequencing of the 850 bp fragment revealed that the product was 97 and 88% homologous to rabbit liver FMO1 and pig liver FMO1, respectively (El-alfy and Schlenk, submitted). Similar RT-PCR products were obtained from Tilapia and Stripped bass (Morone sp.) hepatic RNA, but have yet to be sequenced (Schlenk D, unpublished). Northern blot analyses with RNA from several tissues of smooth dogfish (Squalus acanthias), rainbow
trout (O. mykiss), stripped bass (Morone saxatilis), Atlantic flounder (Platichthys flesus), and turbot (Scophthalmus maximus) probed with a 1.6-kb FMO1 cDNA under non-stringent conditions indicated RNA bands ranging from 1.1 to 3 kb (Table 1). In addition, there were no significant differences between males and females in hepatic FMO1-like mRNA in the turbot (S. maximus) [38]. Northern analyses using FMO1 and FMO2 cDNAs in channel catfish (Ictalurus punctatus) failed to show hybridization even under extremely nonstringent hybridization conditions (i.e. room temperature; Schlenk D, unpublished data). FMO genes are clearly present in several non-mammalian organisms. Recent advances in recombinant DNA methodologies, should allow identification and classification of these genes in the near future. Such studies will allow a better understanding of the evolution and regulation of FMO. Although fish have primarily been examined for expression of FMO RNA, several other phyla have been examined at the protein level by Western blot analyses using antibodies raised against the mammalian proteins (primarily FMO1 and FMO2). The overall trend is that structurally related proteins have been found exclusively in vertebrates. Invertebrates that have been examined include the barnacle (Balanus sp.), gumboot chiton (Cryptochiton stelleri ), the western oyster (Crassostrea 6irginica) and the sea anemone (Anthopleura elegantissima; Schlenk D, unpublished). However, numerous vertebrate species have been shown to express
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microsomal proteins that range from 50 to 60 kD that are recognized by polyclonal antibodies to FMO1 or FMO2. Non-mammalian vertebrates that have shown structurally similar microsomal proteins include the reptilians American alligator (Alligator mississippiensis), the eastern painted turtle (Chrysemys picta picta) and the avian chicken (Gallus domesticus; Table 1). In addition, numerous fish species have been examined and have been shown to express FMO1- or FMO2-like proteins. Included are sharks (S. acanthias and Carcharhinus falciformis), and teleosts with proteins from the rainbow trout (O. mykiss), Atlantic flounder (P. flesus) and turbot (S. maximus) being the best characterized [38,40,42,46,54]. In contrast to mammals, there does not appear to be immunologically distinct isoforms in fish, since antibodies raised against FMO1 and FMO2 recognize the same proteins in all tissues of the few species examined [38,42,46,54]. However, antibodies raised against FMO2 do appear to show less nonspecific binding and stronger signals in Western blots in most fish species [38,42,46,54]. Differences in number and size of microsomal proteins in various tissues recognized by the mammalian antibodies have also been observed. For example, kidney and gill of the Atlantic flounder expressed two FMO2-like proteins, while the liver expressed a single form [54]. Earlier studies in rainbow trout demonstrated two isoforms in several tissues including gill, kidney and liver [42,49]. More recent studies indicate the presence of perhaps three or four isoforms in liver (Schlenk et al., submitted manuscript). In addition, expression of the hepatic isoforms in trout increases with the age of the animal and there does not appear to be any sexual differences in activity but some differences may be present in isoform content between male and female animals [42]. In flounder as well as turbot, expression of hepatic FMO protein was not sexually dimorphic with males and females having similar levels of expression [38,54]. In contrast to these species above, there are many fish species that do not appear to express FMO1- or FMO2-like proteins [4,48]. Common features of these species include: (i) predominantly freshwater origin; (ii) possessing relatively low levels of trimethylamine or TMAO; and (iii) unable to tolerate significant salinity alterations. Without exception, vertebrate animals, such as the little skate (Raja erinacea) [4] or the channel catfish (I. punctatus) [55] lacking an FMO1 or FMO2like protein also fail to demonstrate FMO activity. As discussed above, few compounds have been identified as FMO substrates in non-mammalian species. Prior to the isolation and characterization of FMO from mammals, unless an enzyme had been purified, monooxygenase activities were ascribed to the term ‘mixed function oxidase’ or prefaced by the agent undergoing the oxidation. For example, TMA oxidase has been identified as an enzyme activity since studies
showed the formation of trimethylamine oxide (TMAO) in goldfish (Carassius auratus) and eels (Anguilla japonica) from TMA [21,22]. An association between TMA oxidase and FMO was not observed until 1973, when Goldstein and Dewitt-Harley [16] demonstrated a similarity between TMA oxidase and FMO enzymes in the nurse shark liver (Ginglymostoma cirratum). The first comprehensive review of in vitro TMA oxidase activity was performed by Baker et al. in 1963 [4] in which one bird, one reptile, three amphibians, four elasmobranchs, 15 saltwater teleosts, three anadromous fish taken from fresh water, six freshwater teleosts, four bacteria, three unicellular microbes, one hydra, one planaria, one annelid, one mollusc, four arthropods and 17 plant species were examined. FMO activity has subsequently been observed in several species not demonstrating appreciable activity in these earlier studies. These discrepancies may be explained by more sensitive assays and subcellular fragments used in the more recent studies. Examples include the smooth dogfish (S. acanthias) and rainbow trout (O. mykiss) [40,46]. Likewise, Baker et al. [4] failed to observe activity in any invertebrate species examined. In studies with the insects Heliothis 6irescens and Musca domestica oxidation of DMA was not observed (Hodgson E, personal communication). However, studies with the marine copepod (Calanus finmarchicus), the gumboot chiton (Cryptochiton stelleri ), the western oyster (Crassostrea gigas), the mussels (Mytilus edulis and Mytilus gallopro6incialis), the marine sponges (Geodia cydonium, Tethya aurantium, Verongia aerophoba and Pellina semitubulosa) and the fungus (Cunninghamella elegans) indicate activity consistent with flavin-containing monooxygenases (Table 1). The enzyme was partially purified from C. finmarchicus and was shown to be a cytosolic enzyme of approximately 130 kD with a Km for TMA of 0.3 mM [59]. A cytosolic form of DMA N-oxygenase was also isolated from the protozoan Trypanosoma cruzi having a molecular weight of 100 kD consisting of two 52 kD subunits [1]. The form of FMO isolated from T. cruzi had a pH optimum of 8.0 for the oxidation of NADPH, a Km of 56 mM and a Vmax of 114 nmol min − 1 mg − 1. The enzyme also appeared to have lower apparent affinity constants for sulfur-containing compounds than DMA with respective Km values for disulfoton and methimazole of 0.22 and 0.5 mM, whereas DMA had a relatively high Km of 1 mM [1]. Although TMA oxidase and other FMO catalyzed activities have been observed in several invertebrates, the majority of studies have focused on the characterization of activity in vertebrate fishes [47]. As mentioned above, Goldstein and Dewitt-Harley [16] initially characterized TMA oxidase from the liver of the nurse shark (G. cirratum). For the first time in a non-mammalian species, it was noted that FMO enzyme activity
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was localized in the microsomal fraction of the cellular homogenate, was dependent upon NADPH and unaffected by azide or carbon monoxide. Reaction rates were optimal at pH 9.0 and inhibited by DMA and chlorpromazine (which are two prototypical FMO substrates). The Michealis affinity constant for TMA was calculated to be 1.5 mM. Studies in the coelacanth Latimeria chalumnae also contained an ‘elasmobranchlike’ hepatic TMA oxidase [18]. It is interesting to note that coelacanths are physiologically similar to elasmobranchs having relatively high concentrations of TMAO and urea in tissues [60]. The next characterization of TMA oxidase was reported by Agustsson and Strom [2] in the cod (Gadus morhua). The hepatic enzyme activity was shown to have a lower pH optimum of 8.2, and Km (11 mM) than the nurse shark enzyme. TMA oxygenase was also observed in the kidney, but not the liver or other tissues of the teleost flatfish Limanda limanda and Pleuronectes platessa [2]. Following these studies, TMA oxygenase activity was measured in several other fish species, but not well characterized. TMA oxidase was observed in the liver of guppy (Poecilia reticulata) and the eel (A. japonica) in studies focused on the source and function of TMA and TMAO in euryhaline fish [11]. Although TMA oxidase has been observed in all of these organisms and correlates with FMO-like protein expression, TMA oxidase have never been purified from a nonmammalian vertebrate. At approximately the same time TMA oxidase was being examined in guppy and eel, studies were underway characterizing FMO activity in the rainbow trout [40]. Hepatic FMO activity (DMA N-oxygenase) in trout was microsomal, NADPH-dependent, competitively inhibited by equimolar concentrations of TMA or aldicarb and non-competitively inhibited by methimazole [40]. The optimum pH for DMA N-oxygenation was 9.2 and was extremely sensitive to temperature and unaffected by co-incubation of n-octylamine [40]. As noted above, there were no significant sexual differences in hepatic FMO activity as measured by DMA N-oxygenase or by methimazole S-oxidase [42]. Each of these activities were observed in gill, kidney, heart and intestine microsomal preparations [49]. Although activity was highly variable among individuals with as much as 100% variation observed from hatchery raised trout, DMA oxygenase directly correlated with hepatic proteins recognized by anti-FMO1 and anti-FMO2 [49]. In addition, contrasting studies by Daikoku et al. [11], intraperitoneal treatment with TMA failed to change expression of FMO protein or DMA N-oxygenase activity in rainbow trout [42]. Likewise, intraperitoneal treatment of turbot or flounder with TMA failed to increase hepatic FMO activity [38,52]. In the early survey studies of Baker et al. [4] smooth dogfish (S. acanthias) failed to show TMA oxidase
191
activity. However, in recent years, smooth dogfish have been shown to have significant levels of hepatic FMO activity that is competitively inhibited by TMA or thiobenzamide and non-competitively inhibited by methimazole [46]. The consistent relationship of inhibition kinetics between trout [40] and smooth dogfish indicates significant structural homology in active sites between the two species. DMA N-oxygenase of the smooth dogfish shark was sensitive to temperature with a 76% loss of activity with a reduction of temperature from 25 to 15°C, and a 99% loss when incubations occurred at 45°C. The optimal pH for DMA N-oxygenase was 9.6 and the maximum velocity was calculated to be 1.3 nmol min − 1 mg − 1 with an apparent Km of 44 mM [46]. To further examine the phylogenetic distribution of FMO in fish, FMO activity, protein and mRNA was characterized in liver microsomes from the turbot (S. maximus) [38]. Hepatic DMA N-oxygenase was optimal at pH 8.8, possessed an apparent Km of 88 mM and was inhibited by methimazole and TMA. In comparative studies with the flounder (P. flesus), it was observed that turbot possessed significantly greater levels of FMO activity in the liver relative to the flounder which was barely above detection [54]. However, the flounder had significantly elevated levels of FMO activity, protein and mRNA in the gill relative to the turbot. Comparison of the enzymatic properties of the P. flesus gill and S. maximus liver enzymes indicated dramatic differences in Km between gill and liver, but were both inhibited by equimolar concentrations of TMA. Gill microsomal activity in each species was unaffected by the mammalian FMO2 substrate, n-octylamine. FMO activity of the gill from another euryhaline fish, the Japanese medaka (Oryzias latipes) is also significantly higher than the hepatic enzyme (Schlenk and El-alfy, submitted manuscript). The differential expression of FMO in euryhaline versus stenohaline fish indicate a potential relationship with osmoregulatory function.
3. Physiological significance Indeed, as mentioned above, a large number of organisms do not possess FMO activity. An early hypothesis for the scattered distribution of the enzyme suggested a random deletion of the gene coding for the enzyme [17]. Other hypotheses include a specific role in chemical defense particularly in sponges and molluscs [30] as well as catalyzing the oxidation of phytoplankton-derived dimethylsulfide to the S-oxide by bivalve molluscs [58]. Several endogenous substrates have been observed in mammals: primarily thio-ether conjugates as well as methionine, cysteamine and TMA [15,62,63]. In fact, a genetic polymorphism has been described in humans that lack FMO3 activity and are unable to
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N-oxygenate odorous TMA to non-odorous TMAO [14]. The disease has been named ‘fish odor syndrome’ due to the excretion of unmetabolized TMA across the skin of the individual [3]. Based on the distribution of the enzyme in fish species, another possible physiological function for FMO may be osmoregulation [47]. In addition to the differential expression of the enzyme in the osmoregulatory organ of the gill in the euryhaline flounder P. flesus [54], other reasons for this hypothesis include the direct correlation of FMO with concentrations of the osmolyte, TMAO. Like FMO expression, TMAO is found in relatively large amounts in marine fishes, but in considerably smaller amounts in freshwater fish [19,23]. The average body fluid levels of TMAO in 72 marine teleosts ranged from 0 to 143 mg TMAO (100 g wet wt.) − 1 with a mean value of 54.9 926.1 mg TMAO (100 g wet wt.) − 1, whereas 21 freshwater teleosts had values that ranged from 0 to 17 mg TMAO (100 g wet wt.) − 1 with a mean of 7.595.4 mg TMAO (100 g wet wt.) − 1 [19,23]. It has been shown that marine and euryhaline fish utilize TMAO as an intracellular osmolyte in the marine environment [60]. A direct correlation between TMAO and salinity was observed in the euryhaline flounder, Pleuronectes flesus [31] and the eel Anguilla anguilla [10,11]. Lastly, other evidence supporting an osmoregulatory role for the enzyme is the regulation of enzyme expression by salinity alteration in euryhaline fish. Recent studies with the Atlantic flounder (P. flesus) have shown that when saltwater adapted flounder are placed in freshwater diluted seawater for as short a period as 1 week, more than 90% reduction of hepatic and gill FMO activity was observed [52,53]. In other studies with Japanese medaka, raising the salinity from 0.15 to 2.0% induced FMO activity more than four-fold in the gill after 24 h (Schlenk and El-alfy, submitted manuscript). In support of these studies, Daikoku et al [11] observed a two to ten-fold increase in hepatic TMA oxidase activity when guppies or eels were placed from low to high salinity. In contrast, saltwater adapted stripped bass (M. saxatilis) actually had 23% lower levels of putative FMO-catalyzed S-oxidase activity than freshwater stripped bass [7]. Although discrepancies are present, there appears to be a significant association between FMO and osmoregulation, especially with the gill enzymes. Little is known regarding the regulation of FMO, even in mammalian systems [56,57]. Consequently, how salinity alterations modify FMO expression is unclear. Preliminary studies with sex steroids such as testosterone and 17b-estradiol in juvenile and sexually mature rainbow trout show significant reduction of hepatic FMO activity [49] which parallels expression of Na + K + ATPase and subsequent saltwater adaptation [35]. Current studies in our laboratory are focusing on the effects of osmoregulatory
hormones and effectors on FMO expression at low and high salinity in euryhaline fish species in order to better understand the role FMO may have in osmoregulation in fish.
4. Toxicological significance Although the physiological significance of FMOs is unclear, FMOs clearly play an important role in the toxicity of various heteroatom containing xenobiotics in mammalian and non-mammalian species. The first indication of a toxicological role for FMO in nonmammalian organisms was reported in studies with the mussel M. edulis [29]. Microsomes as well as postmitochondrial fractions of the digestive gland were shown to oxidize NADPH in the presence of various FMO substrates such as methimazole, DMA, 2-AA and 2-AF at a pH optimum of 8.4 [29]. Postmitochondrial fractions of the digestive gland of Mytilis gallopro6incialis were also shown to activate aminoanthracene to mutagenic metabolites at pH 8.4 through an NADPH-dependent pathway [6]. Since mutagenicity of aminoanthracene was inhibited by co-incubation with methimazole, it was concluded that N-oxidation was occurring through an FMO-catalyzed pathway [6]. However, as discussed above, it should be noted that methimazole is not exclusively selective for FMO and may be activated by FMO to a cytochrome P450 inhibiting compound, the sulfinic acid. This indeed is apparent in studies suggesting that FMO is responsible for the catalysis of 2-acetylaminofluorene (2-AAF) to mutagenic metabolite in mussel digestive gland microsomes which was inhibited by co-incubation by methimazole [28]. It is unlikely that FMO would catalyze the N-oxygenation of an acetylated amine based on the electron-withdrawing properties of the adjacent carbonyl which greatly reduces the nucleophilicity of the amine and would subsequently reduce the likelihood of attack of the hydroperoxyflavin moiety of FMO [62,63]. One possible scenario may be the deacetylation of 2-AAF to 2-AF which may be N-oxygenated by FMO [20]. However, acetylases are typically cytosolic, not microsomal, and could only have occurred if microsomes were contaminated with cytosol [37]. Moreover, coincubation with acetylase inhibitors failed to reduce microsomal activation of 2-AAF to mutagenic metabolites [28]. Indeed, in similar studies using the postmitochondrial fraction from digestive glands of M. edulis, 2-AAF as well as other aromatic amines were also activated to mutagenic metabolites [34]. In contrast to the microsomal studies with M. gallopro6incialis which found no change with acetylase inhibitors, mutagenicity of 2AAF with M. edulis digestive gland was reduced by the acetylase inhibitor, paraoxon [34]. In addition, further characterization of other FMO substrates and their
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metabolites were carried out with M. edulis confirming the presence of FMO activity in digestive gland microsomes [33]. Studies with the Western Oyster (C. gigas) also indicated FMO catalyzed oxidation of the aromatic amine 2-AF to N-oxygenated metabolites which were measured directly and found to be reduced by co-incubation of methimazole [44]. However, other enzymatic co-oxidative pathways were also involved in the N-oxygenation of 2-AF by oyster visceral mass microsomes [44]. Four species of marine sponges (G. cydonium, T. aurantium, V. aerophoba and P. semitubulosa) were also able to catalyze the activation of aromatic amines to mutagenic metabolites through a NADPH-dependent pathway that was optimum at 8.4, again indicating characteristics consistent with FMO [30]. The ability to bioactivate aromatic amines to mutagenic metabolites suggests that organisms capable of such biochemistry should have higher incidences of genotoxic responses such as cancer. However, it is unknown whether FMOcontaining organisms are more or less susceptible to aromatic amine-induced carcinogenesis. Although little is known regarding the relationship between aromatic amine effects and FMO expression in non-mammalian organisms, a considerable amount of evidence suggests that fish that have high levels of FMO are significantly more susceptible to thioether pesticides than fish lacking FMO [48]. For example, in a series of in vitro and in vivo studies in rainbow trout, it was shown that one or more FMOs were responsible for the bioactivation of the thioether pesticide, aldicarb, to the more potent cholinesterase inhibitor, aldicarb sulfoxide [43,50]. Trout are approximately 100 times more sensitive to aldicarb toxicity than channel catfish which do not express FMO [48]. In vivo metabolism studies indicate that channel catfish are capable of converting aldicarb to the less toxic metabolite, aldicarb sulfone, which appears to be catalyzed via a cytochrome P450 pathway [48]. In addition, studies examining the effect of salinity of aldicarb toxicity in the Japanese medaka showed a direct effect of lethality and salinity when fish were exposed to 1 mg l − 1 aldicarb (Schlenk and El-Alfy, submitted manuscript). As noted above, a direct correlation was observed between salinity and FMO expression and activity in the same species. Consequently, it has been hypothesized that salinity increases the toxicity of aldicarb by inducing expression of FMO(s) that catalyze the bioactivation of aldicarb to the more toxic aldicarb sulfoxide. Clearly, further in vitro biotransformation studies are necessary to verify an increase in aldicarb sulfoxide production with salinity. FMOs as well as other monooxygenases have also been shown to catalyze the S-oxygenation of Eptam (ethyl N,N-dipropylthiocarbamate) in hepatic microsomes from fresh and saltwater stripped bass (M. saxatilis) [7]. The S-oxide of Eptam was shown to be an efficient carbamylating agent of protein thiols indicating
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potential toxicity of this metabolite to freshwater fish [7]. Similar studies were performed with the herbicide thiobencarb (p-chlorobenzyl N,N-diethylthiocarbamate) from hepatic microsome obtained from the same species and indicated that FMOs may also catalyze S-oxygenation to the S-oxide which was also a more efficient carbamylating agent than the parent compound reacting with thiol and amine nucleophiles [8].
5. Summary and future studies Although relationships have been observed between FMO expression and activity, catalytically active FMO has yet to be purified or cloned from a non-mammalian organism. Structural and genetic similarities appear to be present in vertebrates (i.e. mammalian antibody and cDNA probes recognize respective proteins and mRNAs that correlate with activity). However, the protein structure apparently is significantly different in invertebrates (mammalian probes do not recognize proteins or mRNAs), even though comparable enzyme activities are observed. Because the proteins have yet to be conventionally purified or expressed through recombinant DNA methodologies, the substrate specificities and molecular evolution of individual isoforms are unclear. In addition, until it can be shown that specific substrates are metabolized by isolated enzymes, correlative studies with bands of protein and mRNA as well as indirect studies using multiple inhibitors will have to suffice in characterizing enzyme expression in various non-mammalian organisms. Clearly, the isolation of catalytically active enzymes from non-mammalian organisms would allow the examination of multiple xenobiotics as substrates and provide a better understanding of the physiological and toxicological roles of FMO.
Acknowledgements I wish to thank all of the collaborators that provided samples, antibodies and discussions regarding the phylogenetic distribution of FMO. Key personnel include Abir El-Alfy, Drs John Stegeman, Gary Winston, David Williams, Ron Hines and David Livingstone. I would also like to thank the Marine Biological Association of the United Kingdom for their support of much of this research.
References [1] Agosin M, Ankley GT. Conversion of N,N-dimethylaniline to N,N-dimethylaniline-N-oxide by a cytosolic flavin-containing enzyme from Trypanosoma cruzi. Drug Metab Dispos 1987;15:200 – 3.
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