Weed Science
`
`cambridge.org/wsc
`
`Herbicide Metabolism: Crop Selectivity,
`Bioactivation, Weed Resistance, and
`Regulation
`
`Symposium
`Cite this article: Nandula VK, Riechers DE,
`Ferhatoglu Y, Barrett M, Duke SO, Dayan FE,
`Goldberg-Cavalleri A, Tétard-Jones C, Wortley
`DJ, Onkokesung N, Brazier-Hicks M, Edwards
`R, Gaines T, Iwakami S, Jugulam M, Ma R
`(2019) Herbicide metabolism: crop selectivity,
`bioactivation, weed resistance, and
`regulation. Weed Sci 67:149–175. doi:
`10.1017/wsc.2018.88
`
`Received: 2 July 2018
`Revised: 5 November 2018
`Accepted: 6 November 2018
`
`Associate Editor:
`Patrick J. Tranel, University of Illinois
`
`Key words:
`Crop tolerance; cytochrome P450;
`glutathione; glutathione S-transferase;
`herbicide safener; natural phytotoxin;
`oxylipin
`
`Author for correspondence:
`Vijay Nandula, USDA-ARS, 141 Experiment
`Station Road, Stoneville, MS 38776.
`(Email: vijay.nandula@ars.usda.gov)
`
`Vijay K. Nandula1, Dean E. Riechers2, Yurdagul Ferhatoglu3, Michael Barrett3,
`Stephen O. Duke4, Franck E. Dayan5, Alina Goldberg-Cavalleri6, Catherine
`Tétard-Jones6, David J. Wortley7, Nawaporn Onkokesung6, Melissa Brazier-
`Hicks6, Robert Edwards6, Todd Gaines5, Satoshi Iwakami8, Mithila Jugulam9
`and Rong Ma10
`
`1Research Plant Physiologist, Crop Production Systems Research Unit, USDA-ARS, Stoneville, MS, USA,
`2Professor, Department of Crop Sciences, University of Illinois, Urbana, IL, USA, 3Former Graduate Student and
`Professor, Department of Plant and Soil Sciences, University of Kentucky, Lexington, KY, USA, 4Research
`Leader, Natural Products Utilization Research Unit, USDA-ARS, University, MS, USA, 5Professor and Assistant
`Professor, Department of Bioagricultural Sciences and Pest Management, Colorado State University, Fort
`Collins, CO, USA, 6Research Associate, Research Fellow, Research Associate, Senior Research Associate, and
`Professor, School of Natural and Environmental Sciences, Newcastle University, Newcastle upon Tyne, UK,
`7Boult Wade Tennant, London, UK, 8Assistant Professor, Kyoto University, Kyoto, Japan, 9Associate Professor,
`Department of Agronomy, Kansas State University, Manhattan, KS, USA and 10Assistant Professor, Department
`of Plant Sciences, University of Idaho, Moscow, ID, USA
`
`Abstract
`Several grass and broadleaf weed species around the world have evolved multiple-herbicide
`resistance at alarmingly increasing rates. Research on the biochemical and molecular
`resistance mechanisms of multiple-resistant weed populations indicate a prevalence of
`herbicide metabolism catalyzed by enzyme systems such as cytochrome P450 monoox-
`ygenases and glutathione S-transferases and, to a lesser extent, by glucosyl transferases. A
`symposium was conducted to gain an understanding of the current state of research on
`metabolic resistance mechanisms in weed species that pose major management problems
`around the world. These topics, as well as future directions of investigations that were
`identified in the symposium, are summarized herein. In addition, the latest information on
`selected topics such as the role of safeners in inducing crop tolerance to herbicides, selectivity
`to clomazone, glyphosate metabolism in crops and weeds, and bioactivation of natural
`molecules is reviewed.
`
`Introduction
`
`A common mode of tolerance to herbicides in agronomic crops is by metabolism brought
`about by enzyme systems such as cytochrome P450s (CYPs), glutathione S-transferases
`(GSTs), and glucosyl transferases (GTs). These enzymes, as well as cofactors such as reduced
`glutathione (GSH), are activated by certain chemicals called safeners (Riechers et al. 2010).
`Safeners are applied in combination with herbicides to provide tolerance in grass crops such as
`wheat (Triticum aestivum L.), rice (Oryza sativa L.), corn (Zea mays L.), and grain sorghum
`[(Sorghum bicolor (L.) Moench.] against certain thiocarbamate, chloroacetamide, sulfonylurea
`(SU), and aryoxyphenoxypropionate (AOPP) herbicides applied PRE or POST. Metabolism of
`herbicides usually occurs in three phases: a conversion of the herbicide molecule into a more
`hydrophilic metabolite (Phase 1); followed by conjugation to biomolecules such as glu-
`tathione/sugar (Phase 2); and further conjugation/breakup/oxidation reactions with sub-
`sequent transport to vacuoles or cell walls, where additional breakdown or sequestration
`occurs (Phase 3).
`The next and most important phase after the confirmation of herbicide resistance in a weed
`population is the deciphering of the underlying resistance mechanism(s), which can greatly
`determine the effectiveness of resistance management strategies. One of
`the common
`mechanisms of resistance is metabolic deactivation, whereby the herbicide active ingredient is
`transformed to nonphytotoxic metabolites (Yu and Powles 2014).
`An immediate and urgent challenge for weed scientists is to understand and characterize
`the basis of metabolic resistance to sustain the limited herbicide portfolio and develop inte-
`grated weed management strategies. Metabolic resistance research in weeds has mostly been
`
`© Weed Science Society of America, 2019.
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`Nandula et al.: Review of herbicide metabolism
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`limited to grass species such as rigid ryegrass (Lolium rigidum
`Gaudin), blackgrass (Alopecurus myusuroides Huds.), and
`Echinochloa spp. However, dicot species such as waterhemp
`[Amaranthus tuberculatus (Moq.) J. D. Sauer] and Palmer
`amaranth (Amaranthus palmeri S. Watson) have evolved
`resistance to herbicides with different mechanisms of action
`by enhanced metabolic degradation. Thus, both grass and
`dicot weed species that develop metabolic herbicide resistance
`can pose a severe management challenge.
`The main objective of this symposium was to gain an under-
`standing of current research on metabolic resistance in weeds by
`revisiting the history of related research, including crop tolerance;
`reporting recent advances; and identifying future research
`opportunities. This report is not an exhaustive all-encompassing
`review of herbicide metabolism in crops and weeds; it is a com-
`pilation of papers presented at a symposium during the 2018
`Weed Science Society of America annual meeting.
`
`Complex Signaling, Defense, and Detoxification Pathways
`in Safener-treated Grain Sorghum Shoots
`
`Induction of herbicide-detoxification enzymes catalyzing Phase I
`to III metabolic reactions by safeners is well documented
`(Cummins et al. 2011; Theodoulou et al. 2003; Zhang et al. 2007)
`and has been reviewed extensively in recent years (Kraehmer et al.
`2014; Riechers and Green 2017; Riechers et al. 2010). However,
`identification of signaling genes and pathways leading to safener-
`induced herbicide metabolism has remained mostly elusive.
`Recent research findings have indicated oxidized lipids, or oxy-
`lipins, play an important role in plant defense responses to abiotic
`and biotic stresses (Hou et al. 2016; Mueller and Berger 2009) and
`may also play a key role in safener-mediating signaling (Brazier-
`Hicks et al. 2018; Matsumoto et al. 2015; Riechers et al. 2010;
`Skipsey et al. 2011). In addition to oxylipins, plant hormones such
`as salicylic acid and jasmonic acid (JA) regulate many plant
`responses to pathogen attack or herbivore injury (Gao et al. 2015;
`Koo 2018; Larrieu and Vernoux 2016) and may also function in
`safener-regulated responses (Behringer et al. 2011).
`Although a precise signaling cascade has yet to be established
`for safener-regulated induction of herbicide detoxification in
`cereal crops, several new hypotheses and research areas have
`recently emerged involving oxylipins and other signaling mole-
`cules that will be the subject of future studies. In addition to
`unraveling the complex signaling pathways that lead to the
`induction of enzymes involved in herbicide detoxification, recent
`research has also shown that tissue- and cell-specific expression of
`these enzymes may also play an important role in safener
`mechanisms of action in cereal crops (reviewed by Riechers et al.
`2010) and may potentially explain why dicot plants do not
`respond to safener treatments with increased crop tolerance
`despite increased gene and protein expression (DeRidder and
`Goldsbrough 2006). These two topics will be the focus of the
`following sections.
`
`Oxylipin Involvement in Safener-mediated Signaling
`
`A key finding from research in the mid-2000s was that several
`classes of oxylipins (Mosblech et al. 2009; Mueller 2004) are
`detected in plants following exposure to stresses, and subsequent
`work demonstrated that oxylipins induce the expression of plant
`defense and detoxification genes that mimic safener-induced
`genes and proteins (Loeffler et al. 2005; Mueller et al. 2008;
`
`Riechers et al. 2010; Zhang et al. 2007). Two major categories of
`oxylipins have been detected in plants (Cuyamendous et al. 2015;
`Durand et al. 2011; Mosblech et al. 2009; Mueller and Berger
`2009): (1) phytoprostanes and phytofurans, which are categorized
`based on their nonenzymatic formation via interaction of reactive
`oxygen species with α-linolenic acid (ALA); and (2) enzymatic
`conversion of ALA to 12-oxo-phytodienoic acid (OPDA) and
`subsequent ß-oxidation to yield JA (Figure 1). Interestingly, the
`enzyme catalyzing conversion of OPDA to 3-oxo-2-(2-pentenyl)-
`cyclopentaneoctanoic acid (OPC-8:0, the precursor of JA), OPDA
`reductase (OPR), has been frequently identified in transcript- or
`protein-profiling studies of plant responses to stress (Okazaki and
`Saito 2014; Taki et al. 2005; Yan et al. 2012), including safener-
`treated plants and tissues (Riechers et al. 2010; Rishi et al. 2004;
`Zhang et al. 2007).
`investigated possible links between
`Recent
`research has
`oxylipin-mediated defense signaling and safener mechanism of
`action. The tau-class AtGSTU19 enzyme catalyzed the conjuga-
`tion of GSH to OPDA (Dixon and Edwards 2010), leading to a
`reduction in GSH reactivity. As mentioned earlier, OPR enzymes
`reduce the double bond in the cyclopentenone ring of OPDA,
`resulting in an analogous reduction in reactivity (i.e., electro-
`philicity) but also leading toward biosynthesis of JA (Mueller and
`Berger 2009). Root cultures from Arabidopsis mutants defective in
`
`Figure 1. Representative structures of oxidized lipids (oxylipins) formed in plants.
`Two classes of oxylipins are generated from α-linolenic acid as substrate; either
`nonenzymatically formed (A, generalized phytofuran; or B, phytoprostane) via
`interaction with reactive oxygen species or enzymatically synthesized (C, jasmonic
`acid). For more details on structures and biosynthetic pathways see Cuyamendous
`et al. (2015), Durand et al. (2011), and Mosblech et al. (2009).
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`fatty-acid desaturation (fad3-2/fad7-2/fad8), which are impaired
`in forming the oxylipin precursor ALA, demonstrated a decreased
`ability to respond to safener treatment when AtGSTU24 expres-
`sion was measured and compared with expression in wild-type
`Arabidopsis (Skipsey et al. 2011). Because these fad mutants
`accumulate linoleic acid (18:2) instead of ALA (18:3), they are
`unable to synthesize OPDA or phytoprostanes from ALA sub-
`strate released via lipase activities (Christeller and Galis 2014).
`The decreased ability of these mutant lines to respond to safener
`treatment via induction of GST expression is consistent with a
`link between safener-regulated responses and endogenous oxyli-
`pin signaling.
`Based on the literature regarding oxylipin-regulated gene
`expression (Mueller and Berger 2009) and recent results with fad
`mutants in Arabidopsis (Skipsey et al. 2011), it was postulated that
`certain oxylipins may not only rapidly induce genes involved in
`herbicide detoxification pathways but may also confer safener
`activity in cereals (Brazier-Hicks et al. 2018; Riechers et al. 2010).
`To directly test this hypothesis, a series of compounds modeled
`on oxylipin structures were chemically synthesized and tested for
`biological activity as herbicide safeners in rice (Brazier-Hicks et al.
`2018), in comparison with the commercial rice safener fenclorim.
`Three of the 21 compounds tested rapidly induced GST expres-
`sion in Arabidopsis, but only showed minor whole-plant safening
`activity against pretilachlor herbicide in rice seedlings. In addition
`to possible species-specific differences in responses to these
`potential crop-safening compounds (Brazier-Hicks et al. 2018),
`metabolic pathways and turnover rates of oxylipins (Dueckershoff
`et al. 2008) may differ significantly from those of commercial
`safeners in tissues of cereal crop seedlings (Miller et al. 1996;
`Riechers et al. 2010), therefore requiring further investigation.
`
`Organ-, Tissue-, and Cell-Specific Expression of Safener-
`induced Detoxification Enzymes
`
`As described previously, although the precise signaling pathway
`(s) that regulate gene expression within herbicide detoxification
`pathways have not been elucidated, previous research demon-
`strated that tau-class GST proteins and GST enzyme activities
`involved in herbicide detoxification are highly expressed in the
`outermost cells of wheat seedling coleoptiles after safener treat-
`ment (Riechers et al. 2003). Interestingly, similar results were
`found in safener-treated sorghum coleoptiles using the same tau-
`class wheat GST antiserum (Figure 2). Additional research
`examining stress-responsive gene expression in Arabidopsis cell
`cultures (Mueller et al. 2008) and protein abundance in leaves
`(Dueckershoff et al. 2008) showed that oxylipins (such as phy-
`toprostanes or OPDA)
`trigger detoxification and defense
`responses in a manner similar to safener treatments. Current
`experiments were designed to test the hypothesis that safeners
`
`Figure 2. Tissue distribution of glutathione S-transferase (GST) proteins in a cross
`section of etiolated grain sorghum seedlings, probed with an antiserum raised
`against the tau-class TtGSTU1 protein from wheat (Riechers et al. 2003).
`(A)
`Unsafened (DMSO only) seedling, no primary antiserum (negative control);
`(B)
`unsafened (DMSO only) seedling, probed with a 1:500 dilution of primary antiserum
`raised against TtGSTU1; (C) seedling treated with 10 μM fluxofenim safener for 12 h,
`probed with a 1:500 dilution of primary antiserum raised against TtGSTU1. Red
`arrows in C mark the massive accumulations of immunoreactive GST proteins in the
`outermost coleoptile and epidermal cells. Abbreviations: CL, coleoptile; LP, inner leaf
`primordia.
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`and phytoprostanes induce GST activity and the expression of
`genes related to plant defense and detoxification in sorghum
`shoot coleoptiles in an analogous manner (Riechers et al. 2018). A
`cryostat-microtome sectioning method was developed to extract
`high-quality RNA from the outermost cells of frozen coleoptiles
`(excluding leaf tissues) for transcript profiling to enrich for
`safener- and phytoprostane-responsive mRNAs at different time
`points after treatment (Riechers et al. 2018). Current localization
`experiments are using an antiserum raised against a specific phi-
`class sorghum GST isozyme (SbGSTF1) to further investigate
`tissue-specific expression of different GST subclasses (Labrou
`et al. 2015) in safener-treated grain sorghum seedling tissues (as
`shown in Figure 2).
`Initial RNA-seq results have identified >10-fold increases in
`transcripts of several detoxification genes,
`including multiple
`GSTs, CYPs, and GTs, in safener-treated seedlings compared with
`untreated controls (unpublished data). Moreover,
`transcripts
`encoding proteins related to plant development and defense were
`highly upregulated by safener, such as enzymes involved in lipid
`signaling (including OPRs), hormone-related processes (i.e.,
`synthesis of benzoic acid and salicylic acid), or auxin metabolism
`and homeostasis. Transcripts encoding biosynthetic enzymes
`possibly involved in chemical defense mechanisms in roots (Cook
`et al. 2010) and shoots (Busk and Möller 2002; Halkier and Möller
`1989) of sorghum seedlings were also strongly induced by safener
`treatment in coleoptile tissues (unpublished data). These results
`indicate that safeners may be utilizing signaling pathways and
`enzymatic mechanisms related to generating allelochemicals
`(Baerson et al. 2005) or other defense chemicals against abiotic or
`biotic stresses, as well as upregulating enzymes with the putative
`function of preventing autotoxicity from these chemicals in sor-
`ghum seedlings (Bjarnholt et al. 2018).
`
`Future Research Directions
`
`Ongoing analyses using bioinformatics and comparative gene
`expression approaches are aimed at further mining these RNA-
`seq data to provide additional insight into how transcriptional
`responses are reprogrammed in sorghum coleoptiles following
`safener treatment (unpublished data). An emerging hypothesis
`is that safeners regulate a specific, coordinated, and rapid
`defense and detoxification response in cereal crop seedlings,
`which includes both up- and downregulation of gene expres-
`sion. This research helps to elucidate the yet-to-be discovered
`mechanisms that trigger specific detoxification responses rela-
`ted to safener-regulated protection of cereal crops and, as
`mentioned previously, may also provide insights into the per-
`plexing question of why safeners do not protect dicot crops
`from herbicide injury (DeRidder and Goldsbrough 2006; Rie-
`chers and Green 2017).
`In summary, herbicide safeners are unique organic molecules
`used for crop protection. Safeners increase the success of com-
`mercializing new herbicides by providing a chemical tool to
`enhance crop tolerance and/or crop–weed selectivity for active
`ingredients that otherwise might be removed from primary
`screens due to inadequate crop safety (Riechers and Green
`2017), therefore providing an alternative to creating genetically
`modified crops (Goodrich et al. 2018; Kraehmer et al. 2014).
`Furthermore, safeners may expand the utility of existing herbi-
`cides that do not exhibit adequate crop tolerance without a
`safener as well as expand our basic knowledge of plant responses
`to abiotic stresses.
`
`Contributions of Metabolism to Clomazone Activity and
`Selectivity
`
`Clomazone (Figure 3), a 3-isoxazolidinone, was initially intro-
`duced by FMC Corporation in the 1980s for weed management in
`soybean [Glycine max (L.) Merr.] (Chang et al. 1987). Since that
`time, use of clomazone (also known in the literature as FMC
`57020 and dimethazone) expanded to several additional crops
`(Anonymous 2018). Clomazone
`injury manifests
`itself as
`bleaching of new leaves (Duke and Kenyon 1986). However,
`attempts to tie the clomazone mechanism of action to inhibition
`of phytoene desaturase or steps in the cytoplasmic isoprenoid
`biosynthesis pathway were unsuccessful (Croteau 1992; Lutzov
`et al. 1990; Weimer et al. 1992).
`Seeking to expand the uses of clomazone shortly after its
`commercialization, FMC explored the use of safeners. Naphthalic
`anhydride
`seed treatment afforded some protection from
`clomazone injury to corn, but this system was never commercially
`developed. However, the organophosphate insecticides phorate
`and disulfoton could protect cotton (Gossypium hirsutum L.)
`from clomazone injury (Culpepper et al. 2001). This is still a
`commercial practice. The clomazone label (Anonymous 2018)
`contains specific language regarding use of the insecticides to
`protect cotton from clomazone damage: “Do not apply Command
`3ME Herbicide to cotton unless disulfoton or phorate organo-
`phosphate insecticide is applied in-furrow with the seed at
`planting time” and “Failure to apply either disulfoton or phorate
`insecticides with Command in accordance with in-furrow label
`use directions can result in crop phytotoxicity (bleaching) and/or
`stand reduction.”
`
`Phorate Effects on Clomazone Injury and Metabolism
`
`A series of experiments were initiated, in cooperation with FMC,
`to understand the mechanism of organophosphate safening of
`cotton from clomazone, and the results were originally published
`in two articles (Ferhatoglu et al. [2005] and Ferhatoglu and
`Barrett [2006]). Briefly, the experimental system employed was to
`place 7-d-old cotton seedlings into hydroponic solution with or
`without clomazone and with or without phorate. The chlorophyll
`and carotenoid content of the leaves emerging after the beginning
`of the treatment was measured 6 d after the start of the experi-
`ment. Complete experimental details are in Ferhatoglu et al.
`(2005).
`Clomazone (100 nM) reduced the levels of both chlorophyll
`and carotenoids in the new cotton leaves approximately 80%
`(Figure 4). Phorate (50 μM) partially reversed this reduction,
`while 0.5 and 5 μM phorate were ineffective.
`
`Figure 3. Structures of clomazone, 5-OH clomazone, and 5-keto clomazone.
`
`(cid:18)(cid:38)(cid:45)(cid:37)(cid:35)(cid:38)(cid:24)(cid:27)(cid:28)(cid:27)(cid:1)(cid:29)(cid:40)(cid:38)(cid:36)(cid:1)(cid:31)(cid:42)(cid:42)(cid:39)(cid:41)(cid:15)(cid:6)(cid:6)(cid:45)(cid:45)(cid:45)(cid:5)(cid:26)(cid:24)(cid:36)(cid:25)(cid:40)(cid:32)(cid:27)(cid:30)(cid:28)(cid:5)(cid:38)(cid:40)(cid:30)(cid:6)(cid:26)(cid:38)(cid:40)(cid:28)(cid:5)(cid:1)(cid:18)(cid:32)(cid:30)(cid:32)(cid:22)(cid:38)(cid:39)(cid:1)(cid:4)(cid:1)(cid:23)(cid:21)(cid:18)(cid:16)(cid:2)(cid:41)(cid:1)(cid:18)(cid:32)(cid:30)(cid:32)(cid:42)(cid:24)(cid:35)(cid:1)(cid:18)(cid:28)(cid:41)(cid:34)(cid:42)(cid:38)(cid:39)(cid:1)(cid:19)(cid:32)(cid:25)(cid:40)(cid:24)(cid:40)(cid:46)(cid:3)(cid:1)(cid:38)(cid:37)(cid:1)(cid:9)(cid:11)(cid:1)(cid:20)(cid:24)(cid:40)(cid:1)(cid:9)(cid:7)(cid:8)(cid:14)(cid:1)(cid:24)(cid:42)(cid:1)(cid:8)(cid:10)(cid:15)(cid:7)(cid:7)(cid:15)(cid:8)(cid:13)(cid:3)(cid:1)(cid:41)(cid:43)(cid:25)(cid:33)(cid:28)(cid:26)(cid:42)(cid:1)(cid:42)(cid:38)(cid:1)(cid:42)(cid:31)(cid:28)(cid:1)(cid:17)(cid:24)(cid:36)(cid:25)(cid:40)(cid:32)(cid:27)(cid:30)(cid:28)(cid:1)(cid:17)(cid:38)(cid:40)(cid:28)(cid:1)(cid:42)(cid:28)(cid:40)(cid:36)(cid:41)(cid:1)(cid:38)(cid:29)(cid:1)(cid:43)(cid:41)(cid:28)(cid:3)(cid:1)(cid:24)(cid:44)(cid:24)(cid:32)(cid:35)(cid:24)(cid:25)(cid:35)(cid:28)(cid:1)(cid:24)(cid:42)
`(cid:31)(cid:42)(cid:42)(cid:39)(cid:41)(cid:15)(cid:6)(cid:6)(cid:45)(cid:45)(cid:45)(cid:5)(cid:26)(cid:24)(cid:36)(cid:25)(cid:40)(cid:32)(cid:27)(cid:30)(cid:28)(cid:5)(cid:38)(cid:40)(cid:30)(cid:6)(cid:26)(cid:38)(cid:40)(cid:28)(cid:6)(cid:42)(cid:28)(cid:40)(cid:36)(cid:41)(cid:5)(cid:1)(cid:31)(cid:42)(cid:42)(cid:39)(cid:41)(cid:15)(cid:6)(cid:6)(cid:27)(cid:38)(cid:32)(cid:5)(cid:38)(cid:40)(cid:30)(cid:6)(cid:8)(cid:7)(cid:5)(cid:8)(cid:7)(cid:8)(cid:12)(cid:6)(cid:45)(cid:41)(cid:26)(cid:5)(cid:9)(cid:7)(cid:8)(cid:13)(cid:5)(cid:13)(cid:13)
`
`4
`
`

`

`Weed Science
`
`153
`
`Table 2. Induction of [14C]clomazone metabolism in corn microsomes by seed
`treatment with naphthalic anhydride (0.5% w/w), seedling treatment with
`ethanol (10% v/v), or a combination of the two.
`
`Clomazone metabolite elution timea
`
`Treatment
`
`12.6 min
`
`15.4 min
`
`23 min
`
`None
`
`pmol metabolite − 1 mg microsomal
`protein − 1 min − 1
`
`229 ± 13
`
`55 ± 10 a
`
`3 ± 3 a
`
`Naphthalic anhydride
`
`244 ± 22
`
`117 ± 27 b
`
`73 ±2 b
`
`Ethanol
`
`261 ± 31
`
`106 ± 38 b
`
`15 ± 15 a
`
`Naphthalic anhydride plus ethanol
`
`207 ± 21
`
`23 ± 23 a
`
`42 ± 5 c
`
`a Mean ± SD. Means within a column followed by different letters are significantly different
`at P ≤ 0.05.
`
`were produced in the microsomes (Table 2). Naphthalic induced
`activity for the metabolites eluting at 15.4 and 23 min but not at
`12.6 min (Ferhatoglu et al. 2005). The metabolite eluting at
`12.6 min was not NADPH dependent, so it is not a product of
`CYP activity. Production of the metabolite eluting at 23 min was
`totally inhibited by phorate, while the production of the meta-
`bolite at 15.4 was unaffected. This showed that there were two
`NADPH-dependent clomazone metabolism activities present in
`the corn microsomes, presumably CYP mediated, and that one
`was sensitive to phorate inhibition while the other was not. The
`clomazone metabolite standards 2-chlorobenzyl alcohol and
`5-OH clomazone, supplied by FMC, eluted at 15.4 and 23 min,
`respectively. Therefore, the phorate sensitive activity is presumed
`to be the production of 5-OH clomazone from clomazone.
`The 5-OH clomazone can also cause bleaching in cotton
`seedlings, reducing both chlorophyll and carotenoid levels in the
`plants (unpublished data). The 5-OH clomazone was approxi-
`mately 10% as toxic as clomazone, which is consistent with data
`presented by Chang et al. (1987). However, phorate was ineffec-
`tive as a safener for 5-OH clomazone.
`
`Bioactivation of Clomazone
`
`From this information, a working hypothesis was formed that
`phorate inhibited the CYP responsible for the conversion of
`clomazone to 5-OH clomazone (Figure 3), but phorate was
`ineffective in preventing the formation of the actual toxicant, 5-
`keto clomazone (Figure 3). This hypothesis was based on the
`metabolic pathway for clomazone in soybean (El-Naggar et al.
`1992), which has multiple pathways for clomazone degradation,
`including the formation of 5-keto clomazone. In addition, 5-keto
`clomazone is phytotoxic (Chang et al. 1987). Finally, with the
`discovery of the plastidic isoprenoid pathway (Lichtenthaler 1999;
`Lichtenthaler et al. 1997), it was possible to show that 5-keto-
`clomazone, but not clomazone or 5-OH clomazone, inhibits plant
`1-deoxy-D-xylulose-5-phosphate synthase (DXP synthase; Fer-
`hatoglu and Barrett 2006), the first step in this pathway.
`
`Clomazone Selectivity Is Complicated
`
`In summary, for clomazone to be active, it must be bioactivated to
`its 5-keto clomazone metabolite to be phytotoxic at its site of
`action, DXP synthase, the first step in the chloroplastic isoprenoid
`biosynthesis pathway. The first step in the conversion of clomazone
`to 5-keto clomazone is the CYP-catalyzed formation of 5-OH
`clomazone. Organophosphate insecticides such as phorate inhibit
`
`Figure 4. Effect of phorate on chlorophyll and carotenoid levels in new leaves of
`cotton seedlings treated with 100 nM clomazone for 6 d.
`
`Phorate and other organophosphate insecticides are known
`inhibitors of CYPs (Baerg et al. 1996; Diehl et al. 1995; Kreuz and
`Fonne-Pfister 1992; Mougin et al 1991). They can act as herbicide
`synergists by blocking the CYP-mediated detoxification of an
`active herbicide molecule (Ahrens 1990; Chample and Shaner
`1982).
`To test whether phorate affected clomazone metabolism in
`cotton plants, the roots of cotton seedlings were incubated for 8 h
`in [14C]clomazone with or without 50 μM phorate; this was fol-
`lowed by a 16-h chase period. The phorate reduced clomazone
`metabolism in the shoots, but not the roots (Table 1). The phorate
`treatment had no effect on the unextracted radioactivity. Phorate
`also reduced clomazone metabolism in excised cotton shoots fed
`[14C]clomazone with or without phorate through the cut stem
`(Ferhatoglu et al. 2005).
`
`Clomazone Metabolism in Microsomes
`
`Isolated microsomes are an experimental system that can be used
`for in vitro studies of pesticide, including herbicide, metabolism
`by plant CYPs. While cotton microsomes with the capacity to
`metabolize herbicides had been isolated