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# Glucosinolate-Containing Seed Meal as a Soil Amendment to Control Plant Pests

2000-2002

J. Brown, M.J. Morra University of Idaho Moscow, Idaho Subcontr act Report

NREL/SR-510-35254

Jul y 2005

# Glucosinolate-Containing Seed Meal as a Soil Amendment to Control Plant Pests

2000-2002

J. Brown, M.J. Morra University of Idaho Moscow, Idaho

NREL Technical M onitor: K. Shaine Tyson

Prepa red u n d e r Sub c ontra ct No. XC0

- 9 2909 5-0 1 

### Subcontr act Report

NREL/SR-510-35254

Jul y 2005

National Rene w a ble Energy Laborator y
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## Contents

I. Introduction ............................................................................................................... ......... 3
II. Glucosinolates ............................................................................................................ ....... 4

A. Structure .................................................................................................................. .....................................4
B. Occurrence................................................................................................................. ...................................4 C. Hydrolysis Reactions ........................................................................................................ .............................5 D. Hydrolysis Products........................................................................................................ ..............................6 III. Pesticidal Activity of Glucosinolate Hyrolysis Products .............................................. 8

A. Herbicid al Ac tivity

......................................................................................................... ...............................8 B. Fungicidal and Bact ericidal Activities...................................................................................... ...................12
C. Allelochemical Effects on In sects and Other Invertebrates .................................................................. ......17 IV. Management Considerations........................................................................................ 19

A. Product Formation.......................................................................................................... ............................19
B. Product Activ ity and Fate .................................................................................................. .........................20
1. Li fetimes.................................................................................................................. ................................20
2. Vola tilizatio n ............................................................................................................. ..............................21 3. Organi c Matte r ............................................................................................................. ...........................21 4. Temperature................................................................................................................ .............................22

5. Water ...................................................................................................................... .................................22
6. Soil Texture ............................................................................................................... ..............................23 7. pH......................................................................................................................... ...................................23
8. Microbi al Ac tivity

......................................................................................................... ..........................24
C. Additional A llelochem icals ................................................................................................. .......................25
D. Glucosinolat e Concentrations............................................................................................... ......................26 E. Timing..................................................................................................................... ....................................27
F. Additiona l Bene fits ........................................................................................................ .............................28 V. Crushing and Extraction Technology ........................................................................... 29 VI. Allergenic Reactions...................................................................................................... 30 VII. Registration ............................................................................................................. ..... 33 VII. Genetics and Interspecific Hybridization.................................................................. 34 A. Taxonomy, Genomic Cons titutions, and Re lationships......................................................................... .....34
B. Oil and Seed Meal Char acteristics and Inheritance .......................................................................... ..........34 C. Interspecific and Interg eneric Crosses in Bra ssicaceae

..................................................................... .........36 IX. Recommendations ......................................................................................................... 38
X. Literature Cited ........................................................................................................... ... 39

## Tables

Table 1. Naturally Occurring Glucosinolates

................................................................................................ 72 Table 2. Effects of Aqueous Extracts from Glucosinolate-Containing Tissues on Other Plants

.......... 83 Table 3. Influence of Naturally O ccurring Isothiocyanates on Microorganisms

...................................... 85 Table 4. EPA Registrations for Various Isothiocyanates

............................................................................ 92



## Figures

Figure 1. Generalized structure of glucosinolates

....................................................................................... 93 Figure 2. Glucosinolate degradation pathway. Enzyma tic hydrolysis results in production of glucose followed by HSO 4 - . At least seven other products hav e been observed. Oxazolidinethione and
SCN - are thought to be formed from unstable R-N=C=S intermediates.

.......................................... 94 Figure 3. Relationship between B r assica species as described by U (1935)

......................................... 95

2

## Glucosinolate-Containing Seed Meal as a
Soil Amendment to Control Plant Pests

I

. Introduction

Plants m a y produce com pounds that directly or indi rectly affect their biological environm ent. These com pounds fall within a broad category of com pounds called allelochem icals, and are exclusive of food that influences growth, hea lth, or behavior of other organism s (W hittaker and Feeney 1971). One reason for interest in allelochem icals is their potential for use in alternative pest m a nagem e nt system s. Using plant-produced allelochem icals in agricultural and horticultural practices could m i nim i ze synthetic pesticide use, reduce the associated potential for environm ental contam ination, and contribute to a m o re sustainable agricultural
system

.

Glucosinolates, com pounds that occur in agronom ically im portant crops, m a y represent a viable source of allelochem ic control for various soil-borne plant pests. Extensive studies of several hundred species of the Brassicaceae (Cruciferae) fam ily indicate that virtually all

m a y be capable of producing glucosinolates (Kjaer 1976). Insecticidal, nem a ticidal, fungicidal, and phytotoxic effects are often associat ed with tissues of cruciferous plants. As such, glucosinolates are com m only considered to be ultim ately responsible for pest suppression.

In this review we consider what is known a bout glucosinolates and the factors involved in potential application of glucosinolate-containing tissues in pest control strategies. W e will
focus on the use of seed m eal, but supporting work involving the use of other parts of the plant will be presented when necessary. However, glucosinolates are not biologically active, but are the precursors for the form ation of a va riety of potential allelochem icals. Much of the past em phasis and available literature have f o cused on isothiocyanates (ITCs), apparently
the m o st im portant of those allelochem icals. Any delivery system designed to take advantage of this chem istry m u st consider the fact that glucosinolates m u st be enzym a tically hydrolyzed and that m u ltiple pesticidal com pounds m a y be produced. As such, we will first provide an overview of glucosinolate chem istry and the relevant hydrolytic reactions. The

m o st recent and com p rehensive reviews concer ning glucosinolates are those of Brown and Morra (1997), Fahey et al. (2001), and Rosa et al. (1997).

3

### II. Glucosinolates

A. Structure

Ettlinger and Lundeen in 1956 (Fig. 1) described the general structure for glucosinolates. Found in dicotyledonous plants, they are a class of organic anions usually isolated as potassium or sodium salts (Larsen 1981), but occasionally in other form s. For exam ple, p

hydroxybenzyl glucosinolate is isolated as a salt com p lex with sinapine; an organic cation
derived from choline (Challenger 1959; Kjaer 1960; Van Etten and Tookey 1979). Features

com m on to the class are a ß-D-thioglucose m o iety, a sulfate attached through a C=N bond (sulfonated oxim e ), and a side group (designate d R) that distinguishes one glucosinolate
from another. More than 100 different R groups, and thus glucosinolates, have been
identified or inferred from degradative products (Table 1).

B. Occurrence

Glucosinolates are especially abundant am ong fam ilies of the order Capparales: Tovariaceae, Resedaceae, Capparaceae, Moringaceae, and Brassicaceae. Fam ilies outside the order exhibit occasional occurrence and include the Caricaceae, Euphorbiaceae, Gyrotem onaceae, Lim n athaceae, Salvadoraceae, and Tropaeolaceae fam ilies (Fenwick et al. 1983). A recent literature review provides a com p rehensive list of all species known to contain glucosinolates (Fahey et al. 2001).

Glucosinolate types in plant species are highly va riable. For exam ple, the m a in glucosinolate in radish seed ( Raphanus sativus ) is 4-m e thylsulphinyl-3-butenyl glucosinolate, while

m

u stard seed ( Brassica juncea ) is dom inated by propenyl glucosinolate. Cabbage seed ( Brassica oleracea ) contains m a inly propenyl and 2- hydroxy-3-butenyl glucosinolate; rapeseed ( Brassica napus ) contains 4 m a jor glucosinolates: 2-hydroxy-3-butenyl, 3-butenyl, 4-pentenyl, and 2-hydroxy-4-pentenyl (Sang et al. 1984). Sim ilar differences in glucosinolate types are observed when com p aring vegetative plant parts (Hill et al. 1987; Josefsson 1967b).

Num e rous authors have shown that glucosinolate concentrations vary within plants of a single species, and that fluctuations occur with plant age (Carlson et al. 1987; Chong and Bible 1974; Clossais-Besnard and Larher 1991; Elliott and Stowe 1971; Freer et al. 1989; Griffiths et al. 1991; Johnston and Jones 1966; Josefsson 1967a; Josefsson and Appelqvist 1968; Kondo et al. 1985; Macfarlane-Sm ith and Griffiths 1988; Paxm an and Hill 1974).
Additional environm ental factors such as spacing, m o isture regim e , and nutrient availability affect concentration (Freem an and Mossade ghi 1973; Josefsson 1970a, 1970b; Ju et al. 1982; Louda et al. 1987; MacLeod and Nussbaum 1977; Mailer and Pratley 1990). Glucosinolate type and quantity also vary within tissues of an individual plant. The m a jor glucosinolate in
radish seed is not detected in radish leaves or roots; propenyl glucosinolate is found in all B. juncea tissues (Sang et al. 1984).

4C. Hy droly s is Reactions

Toxicity is not attributed to intact glucosi nolates, but instead to biologically active products such as ITCs, organic cyanides, oxazolidinethiones (OZTs), and ionic thiocyanate (SCN - ) released upon enzym a tic degradation by m y rosinase (thioglucoside glucohydrolase, EC 3.2.3.1) in the presence of water. Degradation also occurs therm a lly or by acid hydrolysis (Kjaer 1976; MacLeod et al. 1981).

Myrosinase is not properly identified as a single enzym e , but rather as a fam ily or group of
sim ilar-acting enzym e s. Multiple form s of the enzym e s exist, both am ong species and within a single plant (Bones and Slupphaug 1989; Falk et al. 1992, 1995a; Lenm an et al. 1990, 1993a; Xue et al. 1993), and all perform a sim ilar function (Björkm a n 1976). Although their genetic sequences are sim ilar to other ß-glycosidases (Lenm a n et al. 1993b), m y rosinases are fairly specific toward glucosinolates (Durham and Poulton 1990). These enzym e s cleave the sulfur-glucose bond regardless of either the enzym e or substrate source. However, the particular enzym e and glucosinolate subs trate influence reaction kinetics (Bones 1990; MacLeod and Rossiter 1986). In addition to plants, t he enzym e s have been discovered in the
bacterium Enterobacter cloacae and the fungus Aspergillus sydowi (MacLeod and Rossiter 1986). Myrosinase-like activity has also been observed in soils (Borek et al. 1996a; Chae and Tabatabai 1983).

Myrosinase and glucosinolates are separated from each other in intact plant tissues; however, the precise nature of the arrangem e nt has not been easy to determ ine. Evidence suggesting

m y rosinase is a cytosolic enzym e associated with m e m b ranes, perhaps surrounding a vacuole containing glucosinolates (Lüthy and Matile 1984; Thangstad et al. 1990), has been supplanted by that obtained using m o re precise m e thodologies. Glucosinolates are probably contained in vacuoles of various types of ce lls (Grob and Matile 1979; Pocock et al. 1987). In contrast, m y rosinase is contained only within structures, called m y rosin grains, of specialized m y rosin cells that are distributed am ong other cells of the plant tissue (Höglund et al. 1992; Thangstad et al. 1991). As glucosi nolate vacuoles do not appear to be present
within m y rosin cells, intercellular rather than intracellular separation occurs. Disrupting cellular tissues allows glucosinolates and m y rosinase to m i x, resulting in the rapid release of glucosinolate degradation products. Myrosinase activity and glucosinolates are preserved in cold-pressed m eal and are no longer physically separated. Thus, adding water im m e diately results in the production of the hydrolysis products, including isothiocyanate, without the
need for additional tissue m aceration.

Enzym a tic degradation is influenced by a num ber of variables, and reviews are available (Chew 1988a; Fenwick et al. 1983; McDanell et al. 1988; McGregor et al. 1983; Sørensen 1990; Tookey et al. 1980; Van Etten and T ookey 1979; Van Etten and W o lff 1973; Van Etten et al. 1969). In the presence of m o isture, glucosinolate hydrolysis produces glucose, sulfate, and an additional aglucon product(s) from the rem a ining N, C, S, and R groups (Figure 2). The latter product is proposed to go through at least one unstable interm ediate,
the ultim ate form controlled by reaction conditions including pH, R-group, and the presence
of m e tal ions such as Fe +2 . In addition, sm aller proteins or coenzym e s accom pany

5m y rosinase and m a y alter the product profile (Falk et al. 1995b; Taipalensuu et al. 1996). Several possible products that exhibit som e form of biological activity m a y be form ed (Fig. 2). The products are all interrelated in that they are isom ers or result from further fission of
the isom ers.

D. Hy droly s is Products

ITCs have historically been considered the “norm a l” products of glucosinolate breakdown. They are often volatile with pungent flavors or odors. The presence of propenyl ITC in

m

u stards and horseradish is responsible for m u ch of the flavor; thus, ITCs are som e tim es called m u stard oils. Form ation requires that the initial unstable aglucon interm ediate undergo a Loessen rearrangem e nt to the R-N=C= S configuration. Isothiocyanates are quite reactive, although less so than the related isocyanates (R-N=C=O). A few com m e rcially
available soil fum i gants depend on the activity of m e thyl ITC either as the parent com pound or as produced from precursors such as sodi um N-m e thyldithiocarbam a te or tetrahydro-3,5-
dim e thyl-2 H -1,3,5-thiadiazine-2-thione. Because of known toxicities, ITCs are often considered likely candidates f o r pesticidal activity.

The ability of ITCs to form cyclic thione com pounds (Hughes 1975) is believed to occur in a spontaneous intram olecular reaction when the ITC contains a ß hydroxyl group on the side chain (R-group) (Fig. 2). The goitrogenic OZTs so form ed are m a jor products from rapeseed ( B. napus ).

Nitrile character is com m on to four additional products. Form ing a nitrile (R-C ≡ N, also known as an organic cyanide), which does not require rearrangem e nt, involves sulfur loss
from the m o lecule. Nitrile form ation is favored over ITC at low pH, but occurs in som e crucifers at a pH where ITC is norm a lly the dom inant product (Gil and MacLeod 1980). The presence of Fe +2 or thiol com pounds increases the likelihood of nitrile form ation (Hasapis and MacLeod 1982; Uda et al. 1986a, 1986b). Epithionitrile form ation requires the sam e conditions as for nitriles, plus term inal unsaturation of the R-group and the presence of an epithiospecifier protein. The epithiospecifier protein possesses a rare property in that it is an

enzym e cofactor that allosterically directs an enzym e to yield a different product (MacLeodand Rossiter 1985; Petroski and Kwolek 1985; Petroski and Tookey 1982). Thiocyanate (RS-C ≡ N) is som e tim es produced, particularly in m e m b ers of the Alyssum

, Coronopus, Lepidium , and Thlaspi fam ilies (Fenwick et al. 1983). Factors controlling organic thiocyanate form ation are not well understood (Larsen 1981).

SCN - production from glucosinolates is contro lled by the presence of specific R-groups. Evidence suggests the anion is a resonance hybrid with greater charge on the S (Hughes 1975; W itczak 1986); however, charge can be localized on either the sulfur (

- S-C ≡ N) or the nitrogen (S=C=N

- ), depending on the environm ent (Beekhuis 1975). Indole and 4- hydroxybenzyl glucosinolates yield SCN

- thought to arise from a highly unstable ITC
interm ediate. SCN

- is form ed from indole glucosinolates over a wide pH range, whereas 4- hydroxybenzyl glucosinolates is typically thought to yield SCN

- only at a m o re basic pH.

W e have recently determ ined that 4-OH benzyl isothiocyanate was not stable even at pH

6values of 3.0 (Borek and Morra, 2003). The half- life decreases with an increase in pH from 3.6 h at pH 3.0 to less than 5 m i n at pH 7.0.

Indolylic com pounds also result during SCN - production from indole glucosinolate hydrolysis. 3-Indolylm e thanol, the im m e diate product, m a y form ascorbigen or 3,3’- diindolylm e thane (McDanell et al. 1988). 3-Indolylacetonitrile, indole acetic acid, and
dioxindoles m a y be form ed under conditions favoring nitrile form ation (McDanell et al. 1988; Monde et al. 1991). Indolylic com pounds in these plant tissues are also form ed by routes other than glucosinolate degradati on (Monde et al. 1991; Takasugi et al. 1988).

7

### III. Pesticidal Activity of Glucosinolate Hy roly sis Products

A. Herbicidal Activity

Glucosinolate-containing plants have a reputa tion for affecting successive or nearby plant

com m unities. Various negative or inhibitory ef f ects can occur. At one tim e dyer’s woad ( Isatis tinctoria ) cultivation was restricted because of a negative influence on following crops (Elliott and Stowe 1971; Fenwick et al. 1983). Yields of legum es (such as clover) and, to a lesser extent, grass are reduced following a Brassica crop (Cam pbell 1959; Kutácek 1964). Cam pbell (1959) showed that pieces of B. oleracea roots placed in a petri dish with seeds of clover ( Trifolium repens ) inhibited germ ination, suggesting direct involvem e nt of

com pounds from the Brassica plants. Flax yields decrease if species of Camelina are present as weeds (Grüm m e r and Beyer 1960). Brassica herbage reduced stand establishm ent of five crop species m o re than twice as m u ch as barley ( Hordeum vulgare ) am endm ents (Vera et al. 1987). Sim ilarly, canola stubble was m o re inhibitory to wild oat ( Avena sterilis ) em ergence than wheat or pea (Jones 1992). Stands of wild Brassica nigra appeared to inhibit

germ ination of annual grasses (Bell and Muller 1973), and broccoli ( B. oleracea ) residues

am ended to soil reduced germ ination and growth of lettuce for 10-21 d (Patrick et al. 1963).
Researchers in Sweden dem onstrated that Sinapis alba seed m eal applid to the soil suppresses weed em ergence (Ascard and Jonasson 1991; Johansson 1992; Johansson and Ascard 1994; Oleszek et al. 1994). For exam ple, m u stard m e al or “cake,” applied in rows between cole crops with the intent of attracti ng enem ies of the cabbage root fly also reduced

num bers of several annual weeds (Ascard and Jonasson 1991).

Results with respect to plant inhibition are not always clear, however, and som e tim es appear inconsistent or contradictory. For exam ple, heavy rape residues ( Brassica campestris L.) reduced wheat, barley, and oat growth in a field trial, but soil N concentrations m easured under heavy residues were less than under light residues. Differences in soil nutrient status were thus possibly responsible for the effect (Horricks 1969). Sim ilar studies using rape straw and chaff residue at 8,970 kg ha

- 1 showed reduced em ergence of brom egrass and alfalfa, and reduced growth of barley, brom egrass, and alfalfa, but reductions were not significantly different than in controls using wheat straw (W addington 1978; W a ddington and Bowren 1978). The effect was am eliorated by applying inorganic nitrogen f e rtilizer; however, subsequent studies indicate that nitrogen lim itations do not fully explain the inhibition (Mason-Sedun and Jessop 1988; Mason-Sedun et al. 1986).

Many variables contribute to differences in phytotoxicity. The Brassica species or cultivar

m a y control the level of phytotoxicity (Mas on-Sedun et al. 1986), although differences are not always dram atic (Vera et al. 1987) and m a y be influenced by the plant growth stage. Fresher plant tissues are m o re effective inhibitors than straw or stubble that has been stored or left after harvest. Aqueous extracts of fresh wild m u stard ( B. campestris ) tissues produce a stronger inhibition of growth than those from aged tissues (Jim énez-Orsornio and
Gliessm an 1987). Green Brassica herbage reduced stand establishm ent m o re than Brassica straw and chaff, which caused only sporadic reductions (Vera et al. 1987). Rape stubble

8residues were less or no m o re effective than pea or wheat residues in inhibiting wheat

germ ination and growth (Jessop and Stewart 1983). The greatest inhibition of wheat occurred with freshly dried Brassica residues, an effect that declined with older residues (Mason-Sedun et al. 1986). Microbial degradation of the tissues m a y produce com pounds apart from glucosinolates that are associated with plant inhibition or stim ulation, further

com p licating data interpretation (Harper and Lynch 1982; Lovett and Duffield 1981; Lovett and Jackson 1980). The target plant species also vary in tolerance to the allelochem icals (Kasting et al. 1974; Oleszek 1987). Tom a toes planted in cabbage-am e nded soil seem particularly sensitive, wilting within 24 h (Ram irez-Villapudua and Munnecke 1988). Suppression of germ ination m a y in part be related to seed size, with sm aller seeds being

m o re susceptible (Vera et al. 1987).

Many studies concerning the effects of glucosi nolate-containing plant tissues on plant growth
have focused on water-soluble com pounds as the responsible agents. Aqueous extracts of
various Brassica and Camelina reduce seedling growth or inhibit germ ination (Table 2). Negative effects are not consistent in all species, and som e plants show no change or are actually stim ulated. There are not always correlations between germ ination inhibition and growth inhibition. Aqueous extracts had little effect on tom a to germ ination, but severely inhibited growth just after germ ination (Gressel and Holm 1964). Depending on plant part, growth stage, or other conditions, extracts from the sam e plant species m a y be inhibitory or
stim ulatory to the target species (Jim énez-Orsornio and Gliessm an 1987).

Intact glucosinolates are water soluble, but of several that have been tested for biological effects, only 3-indolylm e thyl glucosinolate has produced inhibition of plant growth (Bialy et al. 1990; Kutacek 1964). High concentrations of 3-indolylm e thyl glucosinolate in woad m a y be responsible for its inhibition of other plants, possibly by converting som e of this glucosinolate to auxins (Andersen and Muir 1966; Elliott and Stowe 1971; Kutacek 1964; Stewart 1939). Ascorbigen, another possible water-soluble degradation product of this glucosinolate, inhibited germ ination and gr owth of clover, but only at relatively high concentrations (10 -2 -10 -3 M) (Kutacek 1964).

The water-soluble glucosinolate degradation produc t m o st studied in relation to plant growth and herbicidal potential is SCN - . The herbicidal form ulation am itrole-T consists of a m i xture of am ino-1,2,4-triazole and NH 4 SCN. Thiocyanate salts such as KSCN and NH 4 SCN can be
used as selective herbicides and defoliants by varying the concentrations. Oxalis and spurge
in lawns are killed at SCN - concentrations between 1.7-14 g L -1 (0.17-1.4%), and about 11.2 kg ha -1 is used to defoliate cotton (Beekhuis 1975). At 112 kg ha -1 control of poison ivy, poison sum ac, and poison oak is at least as favor able as control by 2,4-D or sodium arsenite. At 897 kg ha -1 SCN

- is a tem porary soil sterilant (Beekhuis 1975). SCN - is produced by B.

napus and S. alba m eals and thus it is of possible im portance in pest control. SCN - is not expected to be a signficant product from B. juncea m eal.

At rates closer to what m i ght be encountered in natural situations (500 μM SCN - ), percent

germ ination of 39 dif f e rent species and cultivars did not dif f e r f r om controls, but growth of 22 species was adversely affected (Stiehl and Bible 1989). Early growth inhibition of som e plants was also observed in bioassays by Park et al. (1983), while in hydroponic cultures Ju

9et al. (1983) found com p lete kill of tobacco at < 5 μg m L -1 and of beans at concentrations over 25 μg m L -1 . Cabbage growth was severely inhibited, though not com p letely killed, at 100 μg m L -1 . SCN - accum u lated in plant tissues and toxicity positively correlated with tissue
SCN - concentrations (Stiehl and Bible 1989; Ju et al. 1983). Squash seedlings had low SCN concentrations in their tissues, suggesting that som e plants m a y degrade or do not readily take up the anion (Stiehl and Bible 1989). W u and Bassler (1969) found that SCN

inactivates cotton chloroplasts by inhibiting O 2 release. Ionic thiocyanate-induced chlorosis

m a y be related to SCN - interactions with iron (Ju et al. 1983). Additional interesting effects
of SCN - on plants include increased yield (6%–15%) when seed potatoes are coated with
0.02% SCN

- and induced apple reddening by SCN

- sprays (Beekhuis 1975; Dustm a n and Duncan 1940).

Other water-soluble glucosinolate degradation products likely play im portant roles in altering seed germ ination and plant growth. W a ter soluble com pounds from B. napus defatted seed

m eal severely inhibited lettuce seed germ ination and those from B. napus leaf tissue

com p letely inhibited germ ination (Brown and Morra 1995, 1996). In contrast, aqueous extracts of root tissue had only a m i nor influence on germ ination. Several organic water soluble glucosinolate degradation products fr om the tissues were found by back-extracting aqueous extracts of the seed m eal with CH 2 Cl 2 and a CaCl 2 solution. W a ter-soluble products were particularly high in the seed m eal and included 5-vinyl OZT, 5-propenyl OZT, 1-cyano3-butene, 3,4-epithiobutyl CN, 2-hydroxy-3,4-epith iobutyl CN, 4-m e thylsulfinylbutyl ITC,
and 5-m e thylsulfinylpentyl ITC. In addition, SCN - is found in water extracts of seed m eal and is an expected product in other tissues in lesser am ounts (Brown and Morra 1991; Brown et al. 1991). These water-soluble glucosinol ate products are probably involved in the inhibition of germ ination by seed m eal tissue. However, inhibition of lettuce seed

germ ination by water-soluble com pounds from leaf and stem tissues could not be explained entirely by the total am ount of identified glucosinolate degradation products (Brown and Morra 1996). This suggests either high toxicity for specific nonvolatile ITCs found in leaf
and stem tissues or the participation of unidentified com pounds.

Plant tissue extract phytotoxicity correlates with tissue am endm ent phytotoxicity, but residues are generally m o re effective inhibitors of seed germ ination than aqueous extracts (Mason-Sedun and Jessop 1988; Mason-Sedun et al. 1986). The greater inhibitory effect of residues suggests that som e thing other than water-soluble toxins influences germ ination. Volatile ITCs are probably the m a jor allelochem icals responsible for inhibitory effects on plants, although som e have disputed the practical significance of these com pounds in vivo (Bell and Muller 1973; Choesin and Boerner 1991).

Plant toxicity is illustrated by m e thyl ITC, of ten studied because of its use as a com m e rcial
soil fum i gant. Methyl ITC is usually generated in situ from a precursor (e.g.
dithiocarbam a te), but m a y be applied directly (Miller 1988). The fum i gant Vapam

, which
releases m e thyl ITC, was the only one of four soil fum i gants found to be toxic to strawberry plants when applied as a drench (Eide 1959). Residues of m e thyl ITC m u st be degraded before planting com m e nces because of phytotoxi city (Sirons 1973). Methyl ITC inhibits growth and germ ination of weeds such as pigweed, dandelion, lam b squarter, berm uda grass,

10purslane, crabgrass, quackgrass, spurge, fleab ane, barnyard grass, fall panicum

, com m on ragweed, stinkgrass, henbit, and carpetweed (Beekhuis 1975; Teasdale and Taylorson 1986).

Many weed seeds are killed while dorm a nt, a capability distinguishing ITCs from other herbicides. Activity in the gas phase for propenyl ITC as well as m e thyl ITC was
dem onstrated by com p lete or severe germ ination inhibition of corn, soybeans, wheat, rapeseed, cucum b er, carrot, alfalfa, dandelion, redroot pigweed, and barnyard grass at concentrations of 1 ppm in the headspace of airtight containers (Vaughn and Boydston 1995). Although exact m echanism s of germ ination inhibition by ITCs are not known, evidence suggests interference with protein synthesis and possibly processes involved in the
form ation of phosphorylated sugars (Leblová -Svobodová and Koštír 1962). Benzyl ITC is a potent inhibitor of plant enzym e activity (Lykkesfeldt and Møller 1993).

ITCs express variable phytotoxicities, a property related to the rem a inder of the m o lecular structure. Benzyl ITC inhibits germ ination and growth of several weeds (Dale 1986; Powell and Spencer 1988; W o lf et al. 1984) at a rate lo wer than m e thyl ITC (Teasdale and Taylorson 1986). Two extrem es of effectiveness were observed in wheat bioassays at ITC concentrations of 500 ppm ; butyl ITC had no e ffect on germ ination, shoot, or root growth while phenylethyl ITC com p letely inhibited germ ination (Bialy et al. 1990). Propenyl,

m e thyl, ethyl, phenyl, and benzyl ITCs were interm ediate with respect to inhibition.

In addition to studies with pure com pounds, organic solvent extracts of papaya shown to contain benzyl ITC inhibited velvetleaf, but not corn, at 4x10 -4 M (W olf et al. 1984).
Sim ilarly, CHCl 3 extracts of m u stard inhibited seedling growth of barley, rye, broccoli, and radish (Jim énez-Orsornio and Gliessm an 1987). Barley and rye were m o st affected; inhibition of radish was not statistically significant. Ethyl acetate extracts of Rorippa indica
contained 8-m e thylsulfinyloctyl, 9-m e thylsu lfinylnonyl, 10-m e thylsulfinyldecyl, and the corresponding sulfonyl ITCs and inhibited lettuce germ ination by 90% (Yam ane et al. 1992). Only 10-m e thylsulfonyldecyl ITC was without activity toward germ ination and growth.

Inhibitory effects with pure com pounds and orga nic extracts also occur in studies involving
volatiles from glucosinolate-containing plant tissues. Volatiles produced from Brassica tissues were shown to inhibit germ ination and growth of lettuce, wheat, and barnyard grass ( Echinochloa crusgalli ) (Oleszek 1987). In a separate study, 22 volatile glucosinolate products consisting of ITCs and nitriles were identified in the headspace of cham bers containing B. napus seed m eal, root, and leaf and stem tissues (Brown and Morra 1995, 1996). Tissues producing those volatiles inhibited germ ination in bioassays, whereas no
inhibition was observed when the sam e tissues were treated to hydrolyze glucosinolates and

rem ove volatile glucosinolate products before bioassays were initiated. Com p arison of headspace volatiles and extracts from the intact and hydrolyzed tissues revealed that ITCs
were the m o st abundant products of intact ti ssues. The presence of other, nonglucosinolate derived volatiles could not explain inhibition since the sam e com pounds were present in the headspace of intact and hydrolyzed tissues. Som e volatile glucosinolate products have also been observed in the headspace of vessels containing B. campestris (1-m ethylpropyl and 4- pentenyl ITC), B. juncea (1-m ethylpropyl and propenyl ITC), B. nigra (propenyl ITC), and S.
alba (3-butenyl ITC and phenylethyl CN) (Tolls ten and Bergström 1988). Propenyl ITC has

11been detected in headspace of cabbage (Chin and Lindsay 1993) and m e thyl thiocyanate as a volatile product from broccoli (Hansen et al. 1992).

The ef f ects of volatile ITCs are short-lived. Benzyl ITC was very ef f ective at inhibiting weed germ ination if the seeds were placed on fresh m e dia; however, if seeds were placed on

m e dia two days old, very little difference was observed from the control (Dale 1986). In seed germ ination bioassays, 50 % of the m e thyl ITC disappeared within 1-2 d and the

rem a inder was alm o st com p letely gone in 5-7 d (Teasdale and Taylorson 1986).

Many of the overall effects on crop plants can be considered beneficial. Tarahum ara Indians
in northern Mexico claim that there are fewer weeds in plots after m u stard is grown (Jim enez-Orsornio and Gliessm an 1987). Ram i rez-Villapudua and Munnecke (1987) noted that plots treated with cabbage am endm ents and solar heating were weed free for m onths. Potato yield was increased and weed biom ass was reduced 50%-96 % after rapeseed was incorporated into the test plots (Boydston and Hang 1995). Palouse wheat farm ers of the
northwestern United States have claim e d higher yields after rapeseed was included in the rotation. In Australia, Kirkegaard et al. ( 1994) m easured greater wheat yields following

Brassica crops com p ared to other break crops. Other beneficial effects m a y result from penetration of the soil by large taproots and suppression of pathogens. However, one potential benefit of glucosinolate-containing plants is weed control by inhibiting seed

germ ination and seedling growth. To take advantage of allelochem ic effects in weed suppression, we m u st consider the m echanism s of plant suppression, characterize the

com pounds and chem istry involved in this suppression, target weed species, and avoid negative effects on the crop. W e m u st especially consider the potential negative consequences of phytoxicity when targeting pests of horticultural crops.

B. Fungicidal and Bactericidal Activities

Glucosinolate products m a y greatly influen ce fungal and bacterial populations. Although products were not identified, in the presence of m y rosinase several alkyl and aryl glucosinolates were cytotoxic to Salmonella typhimurium (Tiedink et al. 1991). ITCs are

am ong the m o st potent products and are suspected to be the m a jor inhibitors of m i crobial activity. Benzyl ITC is som e tim es used as an antibiotic to treat inf ections of the respiratory and urinary tracts (Mennicke et al. 1988). Bacter iostatic, bactericidal, and fungicidal effects
of m a ny ITCs are well docum ented (Table 3).

In m a ny cases, m i croorganism s are differentially inhibited by the sam e ITC. For exam ple, although m e thyl ITC is active against a broad range of organism s, dose responses vary

trem endously am ong species. Munnecke et al. (1962) estim ated the range of toxicity to
Rhizoctonia as 110-1500 μg m e thyl ITC generated h -1 ; Myrothecium , 20-490 μg h -1 ; and Pythium , 10-24 μg h -1 . In general, gram -negative bacteria are less susceptible than gram

positive bacteria to ITCs; som e m i croorganism s avoid toxicity by degrading the inhibitory

com pound (Fenwick et al. 1983; Sm elt et al. 1989).

12The toxicity and range of activity also vary with changes in the ITC R-group. Greater toxicity is often related to increased volatility (Lewis and Papavizas 1971; Munnecke and Martin 1964). Substituents in the R-group govern solubility, which in general is negatively correlated with activity (Drobnica et al. 1967a, 1967b); however, reasons for differences in toxicity are not always clear. Recent studies of Manici et al. (2000) indicate that ITCs produced from thiofunctionalized glucosinolates such as glucoiberin and glucoerucin are

m o re fungitoxic against Pythium irregulare and Rhizoctonia solani than alkenyl glucosinolate products such as propenyl ITC. This research group proposes that species such

as Eruca sativa

, Iberis amara , and Rapistrum rugosum m a y be m o re effective in controlling fungal pathogens because they contain thiofunctionalized glucosinolates.

Glucosinolate products with nitrile character are biologically active, but effects are m o re lim ited than f o r ITCs. The yeast Nematospora sinecaula is inhibited by butyl thiocyanate at 40 μg m L -1 (Holley and Jones 1985) and m odified thiocyanates are used to control slim e

m o ld in the paper industry (Beekhuis 1975; W ood 1975). However, tests against various

m i croorganism s showed that 2-200 tim es as m u ch thiocyanate was required to obtain equivalent growth inhibition com p ared to the corresponding ITC (Virtanen 1965; W a lker et al. 1937). Propenyl CN did not inhibit Peronospora at concentrations > 1000 μg m L -1 (Greenhalgh and Mitchell 1976). A propenyl cyanide concentration approxim a tely 40 tim es that of propenyl ITC was required for equiva lent inhibition, but benzyl CN inhibited Nematospora at < 50 μg m L -1 (Holley and Jones 1985). Solutions of propenyl glucosinolate
with m y rosinase were m o re effective at inhibiting Leptosphaeria maculans at pH 7 (where
ITC should be the dom inate product) than at pH 4 (where the proportion of nitrile product is increased) (Mithen et al. 1986). The sam e authors showed m ild inhibitory effects of 3- indolylacetonitrile to L. maculans . 3-Indolylacetonitrile com p letely inhibited growth of
Penicillium chrysogenum at 200 μg g -1 (Sm i ssm an et al. 1961).

Sim ilarly, other products inhibit m i crobial activ ity and growth, though not to the extent of ITCs. OZTs have little effect on the growth of propionic bacteria (Rutkowski et al. 1972) and the fungal pathogen L. maculans (Mithen et al. 1986), but express som e activity against Phoma lingam (Schnug and Ceynowa 1990) and Aphanomyces euteiches f. pisi (Sm o linska et al. 1997a). 3-Indolylm e thanol is quite inhibitory to L. maculans, but ascorbigen and diindolylm e thane are only m ildly inhibitory (Mithen et al. 1986).

Like OZTs, SCN

- has little effect on propionic bacteria (Rutkowski et al. 1972) and the fungal pathogen L. maculans (Mithen et al. 1986). Am m onium thiocyanate inhibits bacterial growth in soil but stim ulates fungi at concentrations >250 μg g -1 (Sm ith et al. 1945). SCN inhibits nitrite form ing organism s (Sm ith et al. 1945) and SCN - -containing am itrole-T inhibits nitrification (Van Schreven et al. 1970). Although the effects of SCN

- are lim ited,
SCN

- often works synergistically with other chem icals. For exam ple, use with certain peroxides produces bacteriocidal solutions, although neither is effective alone (Beekhuis 1975). Sim ilarly, SCN

- was m o re efficient in lysing cells when com b ined with other anions or lysozym e s than when any agent was used alone at the sam e concentrations (Pollack et al. 1983).

13Activity of aqueous plant extracts against pathogens or other organism s has been observed. Aqueous extracts of m u stard ( B. campestris ) inhibited Verticillium chlamydosporium (Owino et al. 1993) and those of cabbage root inhibited Glomus mosseae spore germ ination (Vierheilig and Ocam po 1990). B. napus seed m eal extracts inhibited A. euteiches (Sm o linska et al. 1997a), but slightly enhanced growth of a Propionibacterium (Rutkowski et al. 1972). Buffered seed hom ogenates of Brassica hirta ( S. alba ) inhibited growth of Nematospora by m o re than 99% (Holley and Jones 1985). Glucosinolates extracted from
Brassica kaber and hydrolyzed in aqueous solution with m y rosinase inhibited Glomus
intraradices (Schreiner and Koide 1993b).

Suppression appears to be related, at least in part, to the solubility of com pounds produced by glucosinolates in the tissue. 4-Hydroxybenzyl ITC has greater solubility than m a ny ITCs
and would be the m o st prom inent glucosinolate product expected in B. hirta and B. kaber extracts. Several water-soluble glucosinolate products present in aqueous extracts of B.

napus were inhibitory to A. euteiches (Sm o linska et al. 1997a). In contrast, water extracts of cabbage tissues did not suppress Aphanomyces root rot of peas (Papavizas 1966). The
dom inant ITC in this tissue has low water solubility.

Volatiles f r om tissues or organic solvent extracts are of ten highly inhibitory toward

m i croorganism s. These fractions are also m o st likely to contain highly active ITCs. Extracts
of B. juncea (in which propenyl ITC is the m o st prom inent product) m a de with 70% ethanol and added to growth m e dia at 0.025%-0.8% were in several cases m o re inhibitory to bacteria than equivalent am ounts of propenyl ITC (Kanem aru and Miyam o to 1990). Sm aller am ounts of other ITCs were also present in the extract s, possibly accounting for the greater toxicity. The authors suggest using m u stard as a food preservative. Extracts (CHCl 3 ) of B. kaber roots expressed antifungal activity toward Cladosporium cucumerinum and Glomus

etunicatum (Schreiner and Koide 1993a). Volatiles from B. kaber were not inhibitory to G.
intraradices , m o st likely because relatively nonvolatile 4-hydroxybenzyl ITC is the m a jor glucosinolate product in these tissues. As m i ght be expected from its glucosinolate profile, B. nigra volatiles were inhibitory toward this organism (Schreiner and Koide 1993b).
Volatiles f r om B. nigra and B. juncea tissues severely inhibited Fusarium sambucinum (Mayton et al. 1996), from B. napus and B. juncea roots suppressed Gaeumannomyces graminis (Angus et al. 1994), and from B. napus m eal com p letely inhibited A. euteiches (Sm o linska 1997b). In all cases, volatile ITCs were im plicated as the responsible agents.
This ef f ect of volatiles f r om plant tissues on m i croorganism s has been term ed
“biofum i gation” (Angus et al. 1994; Kirkegaard et al. 1993).

Studies of pathogen inhibition by glucosinolat e-derived allelochem icals have also been described for soils. Cabbage vapors passed through soil colum n s reduced Rhizoctonia solani contained within that soil (Lewis and Papavizas 1974). Volatiles em anating fromcabbageam ended soil that were passed through tubes containing Pythium ultimum and Sclerotium rolfsii cultures inhibited growth of the cultures, especially when warm ed to 38 o C (Gam liel and Stapleton 1993). Approxim a tely one-third the incidence of Fusarium infection of oat crowns was observed when soils were covered with heavy B. campestris residue com p ared to light residue applications (Horricks 1969). Sealed vessels containing soil am ended with leaf
and stem tissue of nine cruciferous species m a rkedly reduced populations of Fusarium

14oxysporum com p ared to untreated soil and soil am ended with alfalfa, wheat straw, chicken

m a nure, or steer m a nure (Ram irez-Villapudua and Munnecke 1988). Inhibition of Fusarium
was dim i nished or not detectable when ca bbage-am e nded soils were left uncovered, but
tissue am endm ent in com b ination with solarization was m o re ef f e ctive than either
solarization or am endm ent alone (Ram irez-Villapudua and Munnecke 1987).

Detailed investigations of the toxicities of individual isothiocyanates to growth stages of Fusarium oxysporum pathogens in forest nurseries have been conducted in the laboratory. Bioassays with four F. oxysporum isolates were conducted using sealed containers in which 0.3 μL of 2-propenyl, ethyl, buty, phenylethyl, benz yl, or phenyl isothiocyanate was allowed to volatilize. Propenyl and ethyl isothiocya nates were the m o st fungistatic of those

com pounds tested to m y celial growth. Sporula tion and conidia viability were not affected by any of the isothiocyanates. However, the sam e concentrations of propenyl and ethyl isothiocyanates that only inhibited m y celial growth, com p letely suppressed conidia and
chlam ydospore germ ination of all isolates. Othe r glucosinolates releasing ethyl, benzyl, and phenethyl isothiocyanates were also fungitoxic to F. oxysporum conidia and chlam ydospores. Changes in populations of the pathogen resulting from a green m a nure crop will only be
achievable if conidia and chlam ydospores are targeted, since decreased populations resulting
from the direct inhibition of F. oxysporum m y celial growth seem unlikely. W e concluded that the pathogenic F. oxysporum isolates typically infesting nursery soils in northern Idaho
will be m o st suppressed by species of plants containing glucosinolates that release the highest concentrations of propenyl isothiocyanate such as Brassica carinata

, B. nigra , and B. juncea (Sm o linska et al., 2003).

Although the allelochem icals from glucosinolates are inhibitory to m a ny organism s, incorporated organic m a tter m a y sim u ltaneously stim ulate m i crobial populations by providing increased am ounts of carbon substrate. Incorporated B. napus residues were less
stim ulatory than corn residues to Fusarium root diseases in a forest nursery (Jam es, personal

com m unication 2002), but resulted in m o re disease than fallow treatm e nts (Jam es et al. 1996). This is a m a jor obstacle to using m eal am endm ents to control fungal pathogens in
soil. Form ulation technologies or am endm ent strategies m u st be developed to ensure that pathogen populations are not stim ulated.

One of the m o re thoroughly studied pest inhib ition system s includes the use of cruciferous

am endm ents to control Aphanomyces root rot of peas and beans. The results indicate the
successes and difficulties com m only encountered in research concerning the biological activity of glucosinolate-containing crops toward plant pests. In greenhouse studies of soils infested with the fungus, Papavizas (1966) tested leaf and stem am endm ents of several cruciferous plants, of which cabbage was the m o st effective. Controls contained no

am endm ents. About 70% of the control plant roots were infected with Aphanomyces; soils

am ended with as little as 0.5% dried cabbage leaves showed approxim a tely a tenfold reduction in root infection. The effects were still observed on plantings m a de 15 weeks after

am endm ent. Interestingly, although Aphanomyces root rot was clearly suppressed, som e

am ended soils actually had increases in Rhizoctonia root rot.

15W e suspected that volatile com ponents were the responsible agents, so we tried to analyze headspace vapors above am ended soils using aqueous trapping reagents and GC (Lewis and Papavizas 1970). No ITCs were detected in this case, but in contrast to decom posing corn, the vapors included CH 3 SH (thiom ethane), (CH 3 ) 2 S (dim ethyl sulfide), and (CH 3 ) 2 S 2 (dim ethyl disulfide). Bioassays showed that these vapors were toxic to Aphanomyces

, though m u ch less so than ITC vapors which in som e cases were 1000 tim es m o re inhibitory.
Furtherm ore, these vapors could be trapped in aqueous solutions, which suppressed root rot only slightly (Lewis and Papavizas 1971; Papavizas and Lewis 1971).

Additional studies of m i crobial inhibition focuse d on conditions encountered in the field. In containerized plantings both inside and out side greenhouses, am endm ents of Brassicaceae tissues significantly reduced Aphanomyces root rot disease beyond fallow and wheatam ended treatm e nts by 32%-38% (Chan and Close 1987). W h en Brassicaceae crops were grown in soil instead of being introduced as am endm ents, reductions averaged 41%. Davis (1988) found in field studies that wheat in rotation had very little effect on Aphanomyces

, and that a greater reduction was observed when rapeseed varieties were used. However, neither cropping with rapeseed nor am ending with rapeseed tissues resulted in reductions

m o re significant than with oats. In other field studies, sim ilar disease reductions to those observed with rapeseed also occurred with oats and corn (Papavizas and Lewis 1971; Parke and Rand 1989). Root rot severity was reduced in field tests using a white m u stard ( S. alba )

green m a nure but no difference in yield was observed com p ared to fallow, perhaps because of reduced pea em ergence in white m u stard treated plots (Muehlchen et al. 1990). However, continuing a second cycle of peas and white m u stard resulted in reduced root rot severity and increased yield in spite of a sim ilar reduction in pea em ergence. Careful study of the variables influencing allelochem ical efficacy is needed to consistently and effectively control
pests.

B. napus m eal has been used for pest control in a lim ited num ber of studies. Anti-fungal activity against Cylindrocladium parasiticum has been reported (Bhardwaj et al. 1996). The investigators applied 1, 2, or 3 t ha -1 of B. napus m eal to soil and com p ared C. parasiticum control in peanuts to control by Vapam

. The University of Idaho supplied the m eal to the investigators. All three rapeseed m eal rates reduced disease incidence of Cylindrocladium black rot of peanuts. A m a xim u m 70% reduction in disease incidence (com pared to an untreated control) occurred in the 3 t ha -1 treatm e nt of peanut cultivar susceptible to the disease. Although glucosinolate concentration of the m eal was not m e ntioned, the concentration was likely less than 50 μ mo l g -1 and thus substantially below what is contained
in m u stard m eals.

Attem p ts have been m a de to control other fungal pathogens using m eal. Dandurand et al. (2000) used B. napus cv. “Dwarf Essex” m eal in laboratory experim e nts to control
Sclerotinia sclerotiorum and A. euteiches . A 100% reduction in carpogenic germ ination and
a 33% reduction of m y celiogenic germ ination of S. sclerotiorum occurred in soil am ended
with Dwarf Essex m eal. A low glucosinolate B. napus m eal (Stonewall) reduced carpogenic

germ ination by 44% and had no effect on m y celiogenic germ ination, indicating that glucosinolate hydrolysis products were critical to suppression. A. euteiches root rot of pea was reduced 77% by Dwarf Essex m eal and wa s unaffected by low glucosinolate Stonewall

16m eal. In all experim e nts m eal was not hom ogeneously incorporated into the soil, but was placed on the bottom of the bioassay container and separated from the overlying soil by sterile fiberglass cloth.

In sum m a ry, fungal and bacterial pathogens can likely be controlled by glucosinolatecontaining m eals. Previous research was conducted m o st often with B. napus m eal containing glucosinolate concentrations less than 50 μ mo l g -1 tissue. Higher concentration glucosinolate m eals will be m o re effective. Experim e ntation m u st be conducted to determ ine
whether the additional carbon contributed from the m eal exerts a growth-prom o ting effect on the fungal pathogens, thereby decreasing the pesticidal efficacy of the m eal. Am endm ent techniques or form ulations m u st be optim ized to decrease growth prom otion of the pathogen.

C. Allelochemical Effects on Insects and Other Invertebrates

As with other organism s, organic extracts of glucosinolate-containing plant tissues have

m a rked effects on insects. Usually extracts are placed on a m e dium such as filter paper and the solvent is evaporated (controls contain solvent only). Lichtenstein et al. (1964) found root extracts of Brussels sprouts to be very toxic to Drosophila melanogaster , as dem onstrated by 50% m o rtality in 10 m i n. Macerated root tissue was quite toxic also; 50%

m o rtality in 3 h f o r Drosophila and 50% m o rtality in 24 h for the com m on housefly Musca
domestica . Toxicity was in m o st cases strongly correlated with phenylethyl ITC content. Tissues that produced m o rtalities but did not possess phenylethyl ITC produce other ITCs.

Insecticidal activity of several ITCs has been dem onstrated, especially for arom atic

com pounds (Åhm an 1986; Borek et al. 1995b; Chew 1988a; Lichtenstein et al. 1962, 1964; Seo and Tang 1982; W a dleigh and Yu 1988). Borek et al. (1995b) tested the toxicities of six

com m e rcially available ITCs to eggs of the black vine weevil ( Otiorhynchus sulcatus (F.))
and concluded that arom atic ITCs were m o st toxic. Later the sam e group of investigators showed using an expanded set of 12 ITCs and QSAR relationships that ITC polarity was

m o st im portant in explaining observed toxi cities (Borek et al. 1998). Using a linearstructure-activity relationship they predicted that the highest contact toxicities of ITCs to black vine weevil eggs will occur from glucosinolates producing ITCs with higher num bers
of carbon atom s or those bearing sulfinyl, thio, or arom atic m o ieties.

W h itef r inged weevil larvae ( Naupactus leucoloma ) were used as bioassay organism s to test the toxicity of volatile and nonvolatile glucosinolate hydrolysis products (Matthiessen and Shackleton 2000). Com p arisons of the toxicity of two pure ITCs in the headspace of sealed
containers showed that m e thyl ITC vapors we re m o re toxic than phenylethyl ITC across a

tem p erature range of 5°-20°C. The toxicity of volatiles f r om B. juncea m eal (assum e d to be propenyl ITC) were m o re toxic than those from B. napus m eal when equal weights of the
tissues were com p ared. W a ter-soluble products displayed no toxicity to weevil larvae.

In screening products to protect seedling corn from injury by southern corn root worm ( Diabrotica undecimpunctata howardi ), propenyl ITC killed the worm but also prevented the
corn seeds from germ inating (Landis and Gould 1988). Methyl ITC is m a inly used to control

17various fungi that cause dam p ing off and root rot, but is also used to control insects such as
wireworm s and sym philids, and nem a todes (Beekhuis 1975; Mullins and Kirkbright 1987; Sirons 1973; Toba 1984; W o rthing 1987). Prope nyl ITC was the m o st effective of 57
volatile com pounds tested against the wireworm s Limonius californicus and Limonius canus

, with an LC 50 of 2.33 μg m L -1 (Lehm a n 1942). Rapeseed m eal am ended (3% on a weight basis) to soil repelled wireworm s ( L. californicus ), but in uncovered containers did not kill
them (Brown et al. 1991). Maxim u m total ITC content of 301 nm ol g -1 soil decreased rapidly. Subsequent testing using propenyl ITC in the sam e soil showed that wireworm s were killed in lower but sim ilar concentrations in capped vials (LC 50 range 211-238 nm ol g -1 soil) and that propenyl ITC added to soil was m o re persistent than ITC produced from the rapeseed m eal-am e nded soil in the previous test (W illiam s et al. 1993). However, glucosinolate products like ITCs are not always toxic to insects at concentrations found in plant tissues. In fact, ITCs can act as cues or even attractants influencing the behavior of certain insects (Finch and Skinner 1982; Pivnick et al. 1991; Rodm an and Chew 1980; Traynier and Truscott 1991).

Other glucosinolate hydrolysis products have insecticidal properties as well. Organic thiocyanates have been used in insecticides to control weevils in grain and to produce quick knockdown of flying insects such as flies, and have a relatively low im pact on m a m m a ls (Beekhuis 1975; W ood 1975). 3-Indolylacetonitrile, known m o re for its auxin-like activity on plants, also inhibits growth of insects (Sm i ssm an et al. 1961). SCN

- is inef f ective when used alone, but is insecticidal in com b ination with other chem icals (Beekhuis 1975). SCN was toxic to wireworm s only at concentrations m u ch higher than observed in rapeseed m e alam ended soil (McCaffrey et al. 1995).

Invertebrate plant pests other than insects are also potential targets. W i nkler and Otto (1980) found that rotational plantings of rape or m u stard in strawberries checked the spread of som e

nem a todes, particularly Pratylenchus penetrans . Legum inosae, potato, or grass rotations did
not alter nem a tode populations, although green m a nure applications of Legum inosae (peas, beans and vetch), rape, or m u stard reduced nem a tode num bers. Sim ilarly, growing rapeseed in soil and incorporating the tissue as a green m a nure significantly reduced populations of Meloidogyne chitwoodi com p ared to fallow (Mojtahedi et al. 1991), although results were not as clear in a study involving Meloidogyne incognita and Meloidogyne javanica (Johnson et al. 1992). In the presence of m y rosinase, glucosinolates are toxic to Heterodera schachtii
at concentrations of 0.5-5.0 m g m L -1 (Lazzeri et al. 1993). Som e varieties of Brassica crops are being of f e red com m e rcially as “trap crops” f o r nem a todes. Rather than killing the

nem a todes, Brassica plants interfere with nem a tode reproductive cycles. Biochem i cal

m echanism s responsible for nem a tode resistance and suppression, although not well understood (Lazzeri et al. 1993), deserve further attention.

Rapeseed m eal has been used as a soil am endm ent in an attem p t to control nem a todes. Investigators applied 1, 2, or 3 t ha -1 of B. napus m eal to soil and com p ared control of the soybean cyst nem a tode (Heterodera glycines Ichinohe ) in soybean to chem ical control by aldicarb (Bhardwaj et al. 1966). The University of Idaho supplied the m e al to the investigators. The results were am biguous, and acceptable control was not achieved. Higher glucosinolate concentration m eals are thus necessary to control this plant pest.

18

### IV. Management Considerations

The evidence strongly supports the idea that glucosinolate degradation products are biologically active and have considerable potential for use in pest control strategies. However, m a ny factors m u st be considered when these strategies are im plem ented if we are
to take f u ll advantage of this natural pesticidal activity. Release ef f i ciency and release rate
f r om plant tissues, susceptibility of the target species, ef f ectiveness of the particular glucosinolate degradation product, soil texture, organic m a tter, pH, m o isture content, and effectiveness of the products in the aqueous and vapor phases all contribute to pesticide efficacy.

Understanding the processes and fates of the allelochem icals in soil is crucial for m a xim i zing potential benefits in pest control and m i nim i zing potential negative environm ental im pacts. Soils and soil/plant interactions m a y influence which products are form ed and their
effectiveness in the soil environm ent. Although additional work is necessary in this area, enough data have been collected to m a ke som e observations and generalizations concerning the behavior of several glucosinolate products in soil.

A. Product Formation

Conditions affecting product form ation during gl ucosinolate degradation have m o st often
been exam ined in isolated plant tissue system s. For pest control, however, we are particularly interested in what happens in soil. As discussed in previous sections, the plant’s glucosinolate profile and characteristics of the solution in which hydrolysis takes place control qualitative aspects of product form ation. W h en plant tissues are incorporated into soil, the soil’s influence on product form ation m a y be linked as m u ch to its m o isture content
as other soil characteristics. Under drier conditions little soil solution is available to influence the reaction, and the effects of pH and m e tal ions on product form ation could be
determ ined solely by the reaction m i croenvironm ent of the plant tissue. However under

m o re m o ist or wet conditions, the soil solution and soil colloids are m u ch m o re likely to participate in reactions that alter product form ation.

Glucosinolate products such as ITCs, SCN - , OZTs, and to a lesser extent nitriles are produced in soil am ended with plant tissue (Brown et al. 1991, 1994). Nine glucosinolate degradation products—five ITCs, three nitriles, and one OZT—were identified in field soils
extracted after a B. napus plow-down (Gardiner et al. 1999). The m o st abundant products were 2-phenylethyl ITC and benzenepropanitrile. ITCs have also been collected in the rhizospheres of a few plants (Tang and Ta kenaka 1983; Yam a ne et al. 1992), including B.

napus (Kirkegaard et al., 2001).

A m odel system using propenyl glucosinolate and m y rosinase am endm ents was used to investigate product form ation in six soils at two m o isture contents of field capacity (0.033 MPa) and saturation. Propenyl ITC was produced as the dom inant product regardless of soil characteristics (Borek et al. 1994). Only m i nor am ounts of propenyl CN were detected.

19Sinigrin and m y rosinase added to unbuffered aqueous soil extracts resulted in greater proportions of nitrile, and in som e cases nitrile becam e the dom inant product. Glucosinolate hydrolysis produces H + , decreasing solution pH as the reaction proceeds. Although this
enhances nitrile form ation in unbuffered extracts, soils provide a large buffering capacity that in effect prevents nitrile form ation. None of these soils were m o re than m oderately acidic (pH of 5.65-8.3), so glucosinolate hydrolysis in wet acidic soils (pH < 5) m a y result in m o re nitrile form ation. Conditions favoring production of ITCs could be advantageous for pest control since these are typically the m o st biologically active products.

Additionally, product release rates and efficiencies m u st be considered. Although lower in total glucosinolate content, root and shoot tissues of B. napus realeased volatile products

m o re efficiently than did m eal tissue (Brown and Morra 1995, 1996). The condition of the
tissue is also im portant. Fresh tissues m a y release products m o re slowly than those physically disrupted before am endm ent by tissue crushing, air drying, freeze drying, or grinding. Lower m a xim u m concentrations are t hus likely to be produced from fresh tissue, but the period of release is potentially extended. Recent investigations indicate that

m aceration of green tissues is indeed necessary to m a xim i ze ITC release (Morra and Kirkegaard 2002). Such a lim itation does not exist when using m eal am endm ents as soil additives since the seed crushing procedure has resulted in extensive cellular disruption.

B. Product Activity and Fate

1. Lifetimes

Lifetim es of glucosinolate products in the environm ent are generally short, an advantage when considering environm ental im pact. ITCs rem a in in soil for a few days to a few weeks. Even in gas-tight containers, half-lives of m e thyl ITC in several soils were 0.5-50 d at 15 o C (Sm e lt et al. 1989). Concentrations of m e thyl ITC in greenhouse air, where soil had been

am ended with m e tham -sodium , were reduced to less than 0.2% of the original am ount by the third day (Van Den Berg et al. 1992). One week after a m a jor spill of m e tham -sodium into
the Sacram ento River of California, neither m e tham -sodium nor the m e thyl ITC hydrolysis product was detectable (del Rosario et al. 1994). In a germ ination test 80% of the inhibitory capacity of propenyl ITC in soil was gone after 2 weeks (Bell and Muller 1973). Other research showed that the half-life of propenyl ITC in six soils was 20-60 h; propenyl CN had a longer half-life of 80-100 h. The average half- life in all six soils was 48 h for propenyl ITC and 96 h for propenyl CN (Borek et al. 1995a). Biological effects of volatile ITCs should be short-lived, especially in uncovered soils. The potential for negative environm ental consequences is theref ore quite sm all and is illustrated by the f act that no m e thyl ITC
residues were detected in m o re than 200 surface water sam p les (Hogendoorn et al. 1992). However, a short residence tim e places lim its on achieving effective pest control and m a y contribute to the variability observed in the suppression of soil-borne plant pests.

Lifetim es of allelochem icals produced in situ in plant tissue-am ended soil differ from those of the respective pure chem icals am ended to soil without plant tissue. Ninety percent of the

m a xim u m ITC in rapeseed m eal-am e nded soil dissipated within 24 h (Brown et al. 1991),

20whereas only 60% of propenyl ITC am ended in the form of an aqueous phase m i xture to the

sam e soil was lost in 24 h (W illiam s et al. 1993). This is consistent with the finding that an average of 30% propenyl ITC, when added in pure form , was lost in six soils during a 24 h period (Borek et al. 1995a). Accelerated dissipa tion of ITC thus occurs in the presence of plant tissue am endm ents, m o st likely because of sorption to the additional organic carbon.

Lifetim e of SCN - in soil is also relatively short. Losses of SCN - in four soils were 40%-95% over a six-day period (Brown and Morra 1993). In contrast to ITC, however, SCN - produced

in situ in rapeseed m eal-am e nded soil dissipates m o re slowly, having an extended lifetim e in the presence of such tissues (Brown and Morra 1993; Brown et al. 1991). Adding propenyl ITC along with solutions of SCN - to soil inhibited SCN - loss, dem onstrating that

com b inations of glucosinolate products m a y alter the behavior of individual products in soil. In this case, propenyl ITC m o st likely inhibited m i crobial activity and the corresponding

m i crobial degradation of SCN -

.

2. Volatilization

ITC disappears via m a ny routes; volatile losses are one m a jor route. Unsealed bottles had

m

u ch greater losses of m e thyl ITC than sealed bottles (Ashley et al. 1963). Thirty-four percent of the m e thyl ITC in spiked soil was recovered in an air stream passing through it, and nearly all of that within 10 h (Munnecke et al. 1962). Vapor pressures for m e thyl ITC have been reported as 13 and 20.7 m m Hg at 20 o C, relatively low values for a fum i gant (Sirons 1973; Sm elt and Leistra 1974); thus, volatile losses m a y be less of a problem for
ITCs than for som e com m e rcial products. However, volatilization is im portant for even
distribution of fum i gants in soil. Vapor pressu res of various ITCs and nitriles vary widely, so large differences in soil fum i gation resulting from glucosinolate hydrolysis products are expected.

Loss of ITC into the atm o sphere is not desirable, since that am ount of product is not available for pest inhibition. Volatile losses could also be of concern for health and
environm ental reasons. Photodissociation by sun light appears to be one rapid and effective pathway for rem oving m e thyl ITC from the atm o sphere, taking about 41 h (Alvarez and Moore 1994). This process would likely apply to other ITCs; however, questions rem a in as
to the fate and im pact of dissociation products (Geddes et al. 1995).

3. Organic M a tter

Many fum i gants are less effective in soil than in nonsoil system s. This is particularly true of ITCs. Propenyl ITC was about 193 tim es m o re toxic to wireworm s than CS 2 (carbon disulfide) in air, but was only about 29 tim es m o re toxic in a silt-loam soil (Lehm a n 1942).
Sim ilar results were found with m e thyl ITC com p ared to CS 2 and other fum i gants (Matthiesson et al. 1996). Phytotoxicity to wheat from Brassica residues or extracts was generally greater when wheat was grown in sand than in soil (Mason-Sedun and Jessop 1988).

21Sorption of ITCs to soil constituents is an im portant m echanism that decreases ITC effectiveness. Sorption of m e thyl ITC to soil increases with increasing organic m a tter content (Munnecke and Martin 1964; Sm elt and Leistra 1974). As a result, m u ch less m e thyl
ITC is extractable from organic m u ck soils than from m i neral soils (Sirons 1973). Although Ashley et al. (1963) determ ined that the disappearance of extractable m e thyl ITC was slowest in a peat soil am ended with m e tham -sodium , am ounts of extractable ITC could have been controlled by m e tham -sodium hydrolysis rates instead of ITC sorption. As generally
observed for m e thyl ITC, increased propenyl ITC disappearance from soil is correlated with greater organic carbon contents and typically great er nitrogen contents (Borek et al. 1995a). This trend was not apparent for the corresponding nitrile. Faster ITC disappearance likely
results from greater reactivity with nucleophilic groups com m only found in soil organic

m a tter such as alcohols, phenols, thiols, carboxylic acids, and am ines. These types of
reactions m a y also explain reduced ef f i ciency of volatile ITCs released f r om seed m eal tissue

com p ared to other plant tissues (Brown and Morra 1996). Am ino groups in protein, contained in high concentration in rapeseed m eal, readily react with ITCs (Björkm a n 1973; Kawakishi and Kaneko 1987) and reduce m easured volatile concentrations.

4. Temperature

Methyl ITC sorption to soil is not greatly enhanced by increasing the tem p erature from 4 o to
30 o C (Matthiessen et al. 1996), although elevated tem p eratures are correlated with increased

am ounts of ITC in the vapor phase and faster rates of disappearance from soil (Ashley et al. 1963; Borek et al. 1995a; Turner and Corden 1963). Increased tem p erature was also associated with a greater inhibitory effect toward pathogenic fungi in a covered cabbageam ended soil (Gam liel and Stapleton 1993), probably because of m o re thorough fum i gation.
Greater concentrations of volatile thiom e thane, dim e thyl sulf ide, and propenyl ITC were
observed at elevated tem p eratures. Tem p erat ure influences the fate of nonvolatile products
as well. For exam ple, higher tem p erature generally increased disappearance rates of SCN in soil (Brown and Morra 1993).

5. Water

Aqueous solubilities control ITC partitioning in the three-phase soil system . Methyl ITC has reported solubilities in water of 7.6 m g m L - 1 (Sirons 1973) and 8.9 m g m L - 1 (Sm e lt and Leistra 1974) at 20 o C. Estim ates of m e thyl ITC distribution between the vapor, aqueous, and solid phases of soil indicate that m o st of the ITC is in the aqueous phase under m o ist conditions, with a substantial am ount retained in the solid phase (Sm e lt and Leistra 1974). Although ITC in solution still has biological activity, uneven distribution in soil is likely caused by localized concentrations and reduced diffusion (Leistra et al. 1974). This heterogeneity could contribute to variations in effectiveness. W e do not know whether ITCs such as 4-hydroxybenzyl ITC with greater water solubility would have greater pesticidal activity than m o re volatile and less soluble ITCs under wet conditions. Methyl ITC was susceptible to leaching with high m o isture in sandy soil (Frick et al. 1995), therefore, other
ITCs m a y leach in perm eable soils subjected to large water inputs. Henry’s constants m u st theref ore be determ ined f o r the ITCs ofinterest to better predict partitioning within the threephase S soil environm ent.

22Increased water content increases ITC longevity in soil. More m e thyl ITC was m e asured in headspace of soils with lower water contents and rates of disappearance were m o re rapid (Turner and Corden 1963). Although 30% of a pplied m e thyl ITC was lost by volatilization
under a drier m o isture regim e , very little vapor phase m e thyl ITC was lost during a wet regim e (Frick et al. 1995). Increased water content increased the half-life of propenyl ITC (Borek et al. 1995a). Thus continually wet conditions, especially when com b ined with cold

tem p eratures, could result in increased ITC lifetim es and perhaps increased potential for pest inhibition resulting from longer exposure tim es. This could also extend the risks of residual effects on subsequent plantings.

In contrast to ITC, increased soil m o isture content accelerates the disappearance of propenyl CN (Borek et al. 1995a). The fates of ITCs and organic nitriles in soil, at least in the case of propenyl derivatives, appear to be controlle d by different m echanism s . Propenyl CN is
sorbed or reacts m o re quickly at the water phase than at the gas phase, which m a y indicate
that the overall reaction rate f o r nitriles is controlled by the gas/water phase partition
coefficient.

6. Soil Texture

Volatile losses are greatest f r om coarse textured soils, but texture is less im portant than factors such as organic carbon content. In a com p arison of three soils, volatile loss of m e thyl ITC was greatest from a sandy loam (0.67% organic m a tter, 8% clay); least volatilization
occurred from a loam with m o re clay but also greater organic m a tter (2.03% organic m a tter, 25% clay). The finest textured soil, a clay loam (0.55% organic m a tter, 40% clay), had
interm ediate losses of m e thyl ITC com p ared to the other two soils (Munnecke and Martin 1964). Extractable concentrations of m e thyl ITC obtained from sandy, clay, and loam soils

am ended with m e tham -sodium are sim ilar for the first few hundred hours (Ashley et al. 1963). As such, evidence suggests that sorption to soil clay is not a m a jor m echanism of disappearance for ITCs. In contrast, sorption to or degradative catalysis by inorganic soil
constituents m a y play a role in SCN - disappearance from soil, particularly at higher

tem p eratures (Brown and Morra 1993).

7. pH

Soil pH influences the form ation and disappearance of glucosinolate hydrolysis products, although effects on disappearance are not dram a tic. Greater am ounts of m e thyl ITC were trapped in headspace from soils adjusted to a higher than to a lower pH (Munnecke and Martin 1964), and additing lim e to three soils tended to shorten residence tim es of m e thyl ITC in soil (Ashley et al. 1963). No correlation between soil pH and propenyl ITC or propenyl CN disappearance was observed for six soils ranging in pH from 4.35 to 9.10 (Borek et al. 1995a). Typical pH values of agricultural soils are thus not expected to greatly
alter allelochem ical residence tim es.

238. M i crobial Activity

Microbial degradation m a y also be responsib le for the disappearance of glucosinolate hydrolysis products. Soil sterilized with heat and soil extracts sterilized by filtration or heating slowed the disappearance of m e thyl ITC in sealed bottles (Ashley et al. 1963).
Accelerated loss of m e thyl ITC after successive soil applications has been dem onstrated in field and laboratory studies (Sm e lt et al. 1989).

Although m i crobial degradation is suggested by these data, this is not a consistent
observation. For exam ple, attem p ts to sterilize soil by autoclaving or ethyleneoxide

treatm e nt did not change disappearance rates of propenyl ITC (Borek et al. 1995a). The

sam e soil treatm e nts increased propenyl CN half-life only slightly. In contrast to soils that appear to have enhanced degradation, a second treatm e nt of fum i gant to soil did not enhance ITC disappearance (Ashley et al. 1963) and two soils treated at least 13 tim es showed no increase in the ITC disappearance rate (Sm e lt et al. 1989). Sim ilarly, five additions of propenyl ITC or propenyl CN during a period of 10 d f a iled to alter the half -lives of these
chem icals (Borek et al. 1995a). No clear evidence dem onstrates that biotic transform a tion
controls the f a te of ITCs in soil. The relative im portance of m i crobial activity in the degradation of organic nitriles is also uncl ear, although biotic degradation is probable.

Num e rous m i croorganism s have been isolated that degrade aliphatic nitriles and a few that degrade arom atic nitriles (Nawaz et al. 1991, 1992).

Several bacteria have been identified that degrade SCN - (Betts et al. 1979; Happold et al. 1958; Katayam a and Kuraishi 1978; Putilina 1961; Sm ith and Kelly 1988; Stafford and Callely 1969). SCN

- is rapidly broken down, and a flush of growth after breakdown results
from the added nitrogen source (especially for NH 4 SCN). Even af ter tem porary sterilization with 897 kg NH 4 SCN ha -1 m a rked stim ulation of plant growth began to occur after 69 d (Beekhuis 1975). Other investigations showed that autoclaving soils before incubation and adding NaN 3 or propenyl ITC slowed SCN - disappearance (Brown and Morra 1993). Disappearance of SCN - was correlated with organic carbon content of soils when

tem p eratures were at or below 30 o C. This suggests that SCN - losses are related to the

am ount of carbon available to support m i crobial activity rather than SCN - sorption to the organic carbon fraction of the soil.

Results for m e thyl ITC and propenyl ITC discussed throughout this section are not necessarily the sam e as for other ITCs because of the influence of the R group chem istry. Different R groups affect volatility and solubilit y, resulting in altered biological activities. As a consequence, individuals involved in effort s to develop plant varieties for pest control should consider glucosinolate type as well as total glucosinolate concentration.

24C. Additional Allelochemicals

Num e rous other S-containing com pounds, m a inly volatiles, have been identified as secondary products of glucosinolates in crucifer ous tissues. Methanethiol, dim e thyl sulfide,
and dim e thyl disulfide are produced in cabbage-am e nded soils (Gam liel and Stapleton 1993). Although these com pounds possess lower toxicities than ITCs (ITCs > thiols ≅ thiocyanates > sulfides), they are produced in larger am ounts and for m u ch longer periods than propenyl ITC (Gam liel and Stapleton 1993; Lewis and Pa pavizas 1971; Virtanen 1965; W a lker et al. 1937). Greater total am ounts and longer production tim es m a y com p ensate for lower toxicities, increasing the potential im portance of these com pounds in pest inhibition.

Several of these S-containing com pounds m a y result from abiotic or biotic degradation of prim ary glucosinolate hydrolysis products. Tw o water soluble degradation products of 4-

m e thylthio-3-butenyl ITC, one identified as 2-thioxo-3-pyrrolidinecarbaldehyde, were fungicidal (Uda et al. 1993). Som e sm aller m o lecular weight products m i ght be explained by
the sim p le loss of a part of the m o lecule such as m e thanethiol from thioalkyl ITC (Uda et al. 1990). Other com pounds, including COS, H 2 S, and CS 2 are form ed from ITCs by a series of hydrolysis reactions (Bailey et al. 1961; Cha llenger 1959; Dateo et al. 1957). Alternatively,

H 2 S so generated m i ght react with CO 2 and produce COS (Shaw et al. 1980). Aerated and waterlogged soils am ended with SCN - produced COS via an unknown m echanism (Minam i and Fukushi 1981). Only COS was produced in a waterlogged soil am ended with SCN -

, although sam p les were m onitored for H 2 S, CH 3 SH, CH 3 SCH 3 , CS 2 , and CH 3 SSCH 3 (Minam i 1982). Bending and Lincoln (1999) suggest with re spect to fresh tissues that other volatile S

com pounds were likely to be as im portant in pest control as ITCs. Further inform ation regarding reaction pathways would enhance our ability to predict and optim ize pest control efficiency.

Other com pounds besides products of glucosinol ate degradation m a y be biologically active and thus contribute to pest inhibition. For exam ple, S-containing indole com pounds found in B. campestris and B. oleracea are not derived from glucosinolates and act as phytoalexins (Monde et al. 1991; Takasugi et al. 1988). In addition, phenolic com pounds are som e tim es suspected as biologically active agents in cruciferous plants (Kutácek 1964), a possibility
since seed m eals of som e Brassica contain relatively higher phenolic com pound concentrations than other sources (Shahidi a nd Naczk 1992). Given the ubiquitous nature of phenolics, biological activity m a y be related m o re to quality than quantity (Levin 1971; Singleton and Kratzser 1973). Although phenolics ar e often suggested to participate in plant defenses against infection, little work appears to have been done on the potential pesticidal activity of specif i c phenolics in Brassica seed m eals.

Additional inhibitory com pounds m a y be produced by breakdown of the S-containing am ino
acids m e thionine and cysteine. Com pounds such as m e thanethiol, dim e thyl disulfide, and
dim e thyl trisulfide are derive d from the non-protein am ino acid S -m ethyl-1-cysteinesulfoxide (SMCSO) via a m e thyl m e thanethiosulfinate interm ediate (Chin and Lindsay 1994; Marks et al. 1992; Maruyam a 1970; Osterm ayer and Tarbell 1960). The interm ediate is produced

25either enzym a tically by cysteine sulfoxide lyase or nonenzym atically under acidic conditions. SMCSO constitutes 0.25%-2% (dry weight) of the tissue in Brassica plants (Gosden 1979; Marks et al. 1992), which indicates that contributions to pesticidal activity could be significant. Ensiled forage rape ( B. napus ) contained approxim a tely 83% of the original SMCSO content, but only one-tenth the original glucosinolate content (Fales et al. 1987), suggesting SMCSO m a y be less susceptible to rapid degradation and thus rem a in available for longer tim e periods.

In som e cases, m e thanethiosulfinate is proposed to react with H 2 S to f o rm thiols and sulf ides (Chin and Lindsay 1994). Any H 2 S produced from ITCs could thus react with SMCSO products to form yet additional allelochem icals. Optim al pest inhibition m a y require

com b ined interactions of glucosinolate products, SMCSO products, and additional

com pounds produced by reactions between com ponents of these two pools. Various products could be form ed sim u ltaneously or sequentially, with one suite of biologically
active com pounds replacing another during an extended period. Effects on nem a todes have been observed to last several weeks (Santos and Mojtehedi, personal com m unication) suggesting the involvem e nt of other com pounds beyond the active lifetim e of volatile ITCs.

D. Glucosinolate Concentrations

Yield data for m a ny plant/soil/pest system s suggest that glucosinolate concentrations are, or border on being, sufficient for pest control. Effo rts to develop plants with high glucosinolate levels for pest control are worthwhile. Since the traditional approach of plant breeders has been to select for reduced concentrations of glucosinolates, concentration increases are likely. Eventually perhaps, particular strains of plants with specific glucosinolate profiles could be developed for specific targets susceptible to the respective glucosinolate products.

Breeding m a y not be the only m eans to increase product concentration. Som e lim ited increases in resistance have been achieved by applying synthetic ITC precursors to Brassica plants (Dawson et al. 1993; Griffiths et al. 1989) . Methyl jasm onate and salicylic acid, other

com pounds that occur naturally, have been associated with increased levels of certain glucosinolates in leaves when applied to those tissues (Doughty et al. 1995; Kiddle et al. 1994). Perhaps glucosinolates could be applied to plant surfaces in a resistance strategy or to the soil (com bined with m y rosinase) for pest suppression. Although synthetic inputs would
not be elim inated, reductions m i ght be achieved by supplem enting plowed-under plant
tissues with low levels of com m e rcial ITC-producing fum i gants.

Although data are lim ited, we can look quantitatively at the question of glucosinolate concentrations and the potential to control plant pests. The am ount of m e thyl ITC

recom m e nded for soil applications using sodium N-m e thyldithiocarbam ate can be estim ated by assum i ng a 15.2 cm depth of incorporation, soil bulk density of 1.4 g cm -3 , and 100%
conversion to m e thyl ITC (Brown et al. 1991). Calculated values were 517-1294 nm ol

m e thyl ITC g -1 soil depending on the crop and type of control required. These are higher
than the am ount of ITC (301 nm ol g -1 soil) observed in soil am ended with rapeseed m ealam ended at a rate of 3% (w:w) (Brown et al. 1991). To kill 50% of wireworm s ( L.

26californicus ), rapeseed m eal had to be am ended at a rate of 114 g kg - 1 soil (Elberson et al. 1996), too high to be useful. However, the sam e level of control for black vine weevil ( Otiorhynchus sulcatus ) required only 19 g rapeseed seed m eal kg - 1 soil (Borek et al. 1996b).

The previous studies were perform e d with rapeseed m eal having a glucosinolate concentration near 40-50 μm ol g -1 tissue. Com p lete release of glucosinolate to ITC has not been observed (Brown et al. 1991; Brown and Mo rra 1995). Instead, the data indicate that a

m a xim u m of 20% of the glucosinolate will be released as ITC f r om rapeseed m eal (Brown et al. 1991). If we now assum e sim ilar release efficiencies for m u stard m eals that contain glucosinolate concentrations of 250 μm ol g -1 tissue, we can estim ate the am ount of m eal necessary to approach the recom m e ndations application rates for com m e rcially available ITC
fum i gants (517-1294 nm ol m e thyl ITC g -1 soil). A 1% (w:w) m eal am endm ent to soil of a

m eal containing 250 μm ol glucosinolate g -1 tissue would result in the release of 500 nm ol ITC g -1 soil. W e are now at the lower lim it of what is considered necessary for soil sterilization. Doubling the am endm ent rate to 2% (w:w) would increase predicted ITC
release to 1000 nm ol g -1 soil and thus achieve a concentration considered near the upper lim it
of that recom m e nded for com m e rcial fum i gants. Thus, m eals with glucosinolate concentrations in excess of 200 μm ol g -1 tissue m a y indeed be effective in pest control at practical tissue am endm ent rates.

E. Timing

Length of exposure to the allelochem ical is one variable associated with tim ing. Lewis and Papavizas (1971) showed in sealed containers that concentrations failing to kill

m i croorganism s in 2 d m i ght kill them in 4-6 d. Evidence indicates sim ilar results with insects and seeds (Lichtenstein et al. 1962, 1964; Pieczarka and W a rren 1960). Dorm ant seeds m a king up the seed bank in soil and exposed to root exudates over a period of tim e could be m o re susceptible to inhibition than introduced seeds exposed to root exudates for only a short tim e.

Another im portant variable is the tim e when initial exposure occurs; in other words, the tim e when the target organism is introduced to the glucosinolate-containing tissue. Pea seeds planted im m e diately after soil am endm ent with cabbage did not germ inate. Peas planted 3
weeks later showed som e reduction of growth if fresh leaves were used, but not if air-dried leaves were used. Plantings after 7 weeks showed no growth reduction or slight growth
enhancem ent (Papavizas 1966). Only fresh cabbage juice inhibited lactic acid ( Leuconostoc
mesenteroides ) bacterial growth (Kyung and Flem ing 1994). Inhibition of germ ination and plant growth by ITCs was m o st effective when the chem icals were applied to seeds or applied within the first 3 d of the germ ination period (Leblová-Svobodová and Koštír 1962).

Tim i ng m a xim a l glucosinolate product form ation with a susceptible life stage of the target organism m a y be necessary. Glucosinolate con centrations in plants usually start out low,
clim b with fluctuations to som e m a xim u m , then drop off near the tim e of plant senescence (Freer et al. 1989; Griffiths et al. 1991; M acfarlane-Sm ith and Griffiths 1988). Aqueous
extracts of wild m u stard were m o st effective at inhibiting growth when extracts were m a de

27during the bolting and flowering stage (Jim énez-Orsornio and Gliessm an 1987). Coplantings
of Brassica plants with other plants do not inhibit seed germ ination or plant growth (Bell and Muller 1973; Choesin and Boerner 1991), perhaps because plant stages susceptible to inhibition are past by the tim e sufficient glucosinolate products are present. In the case of coplanting with two Brassica species, planting wild m u stard with broccoli resulted in higher broccoli yields, but wild m u stard planted before broccoli inhibited broccoli yield (Jim énezOrsornio and Gliessm an 1987). In a consistent fashion, insects and m i croorganism s m a y be

m o re susceptible at certain stages in their life cycle. Relatively sudden releases and higher concentrations of products from am endm ents tim ed to susceptible stages of an organism ’s life cycle would be m o st effective.

F. Additional Benefits

Other benefits m a y be obtained by using glucos inolate-containing plants in pest control strategies. For exam ple, soil structure is im proved by use of organic am endm ents and high root developm ent like that in oilradish and m u stard is im portant to prevent soil erosion and nitrate leaching. Reports in the literature also indicate that B. napus is efficient at absorbing phosphorus from deficient soils (Grinsted et al . 1982; Hedley et al. 1982b). In addition, plant
tissue am endm ents can serve as nutrient sources and high protein-containing rapeseed m e al

m a y be used as a supplem entary nitrogen source (Johansson and Ascard 1994; Kücke 1993). Carbon to nitrogen ratios of B. juncea

, S. alba , and B. napus m eals range from 7 to 10 (Morra and Johnson-Maynard 2003). Mineralizing the m eal will thus provide a substantial am ount of plant available N and no im m obilization. Biofum igating com post with seed m eal would thus add nitrogen to the m i x and sim u ltaneously reduce unwanted weed seeds and disease pathogens.

28

### V. Crushing and Extraction Technology

Current m e thods of cold pressing the m eal result in residual oil of 5%-10% on a weight
basis. Residual oil m a y help trap hydrophobic ITCs, prevent volatile losses, and increase pesticidal effectiveness. However, residual oil m a y also decrease the release efficiency of
ITC from the m eal into the soil and decrease its effectiveness. The overall im pact of residual
oil in the m eal will vary with the ITC and its respective hydrophobicity. The effect of the oil on pest control has never been determ ined.

Com m e rcial crushing facilities produce tem p eratures in the seed and seed m e al that m a y deactivate m y rosinase. Tem p eratures are increased on purpose to m a xim i ze oil extraction or unintentionally if an extrusion process follows crushing. The m a xim u m tem p erature at which

m y rosinase integrity is m a intained during a com m e rcial crushing procedure has not been
determ ined.

Current research is f o cused on using the m eal as a carrier f o r the pesticide. The m e al will also provide plant nutrients in addition producing ITC for pesticide control. However, the

m eal will also provide a carbon source for fungi and potentially increase pathogen problem s. Extracting the ITC before soil application m a y circum vent this problem

.

Techniques have been developed for extracting propenyl isothiocyanate from B. juncea m eal using a com m e rcially available unit. A com m e rcial enterprise has been started by Mr. Peter Fergeson (Peter_I.M. Ferguson@eol.ieaust.org.au) in NSW Australia to extract propenyl ITC using a unit built by Flavourtech (33 Lenehan Rd., Griffith, NSW 2680, Australia). The ITC
is then m a rketed to Japanese com p anies that use it as a food additive. Using a sim ilar approach for producing pesticides seem s feasible.

29VI. Allergenic Reactions

Although the com pounds are produced by plants, hum an and anim al toxicity is still a concern if alternative pest control strategies are to be im plem ented. Do natural toxic substances in plants have advantages over synthetic pesticides? In contrast to a new synthetic pesticide,

m

u ch inform ation about the toxicology of these products is already known because of the
im portance of cruciferous crops in hum an nutrition and anim al diets.

W h en used as a prim ary food source, glucosi nolate-containing plant m a terial clearly has deleterious effects. Feeding kale to livestock can reduce fertility and induce goiter (Johnston and Jones 1966). Diets of cram be m eal caused early death to rats (Daxenbichler et al. 1966), and in separate tests, rats fed cram be m eal and rapeseed m eal exhibited sym p tom s sim ilar to rats with nitriles in their rations (Van Etten and Tookey 1979). Livestock poisoning has been

com m only reported when anim als are fed excessive cruciferous plant m a terial (Kingsbury 1964).

ITCs are general biocides whose activity results from interaction with proteins. They interact nonspecifically and irreversibly with proteins and am ino acids to form stable products by reacting with sulfhydryl groups (Fenwick et al. 1983; Kawakishi and Kaneko 1985; W a re 1983), disulfide bonds (Kawakishi et al. 1983), and am ines (Kawakishi and Kaneko 1987;

W ood 1975). Reaction with term inal am ino groups is the basis of the well-known Edm a n degradation for am ino acid sequencing of proteins. ITCs inactivate enzym e s in vitro , but the
actual effects in vivo m a y be controlled by accessibility (W ood 1975).

Num e rous studies concerning ITC toxicities have been perf orm e d. Contact of ITCs with
skin and m u cous m e m b ranes (respiratory tract) cause irritation, and m a ny are considered
lachrim a tors. At 20-50 m g kg -1 body weight propenyl ITC produced epithelial hyperplasia,

stom ach ulcers, and m i nor inflam m a tory foci in the livers of dogs, but did not show sim ilar
effects in rats even at 500 m g kg -1 body weight (Hall 1973). Antithyroid effects are attributed to ITCs, although this m a y be caused by m e tabolic conversion of the ITCs to SCN

- . An oral LD 50 in rats for m e thyl ITC is listed as 175 m g kg -1 (W orthing 1987) and 339 m g kg -1 for propenyl ITC (W ood 1975). Subcutaneous LD 50 values are considerably lower at 50

m g kg -1 for m e thyl ITC and 80 m g kg -1 for propenyl ITC (W ood 1975). The LC 50 of m e thyl
ITC at 96 h is 0.13 m g L

- 1 for bluegill and 0.37 m g L

- 1 for trout and carp (W orthing 1987).

Mam m a lian system s m e tabolize and elim inate ITCs fairly rapidly. Less than 5% of a dose of propenyl ITC was retained by rats and m i ce 24 h after adm i nistration (Ioannou et al. 1984).
ITCs adm i nistered both orally and by injection in rats and m i ce are prim arily detoxified by glutathione and excreted as S -(

N

-m ethylthiocarbam oyl)m ercapturic acids (Lam et al. 1993; Ioannou et al. 1984; Mennicke et al. 1983). Det oxification and excretion of benzyl ITC in
hum ans is sim ilar (Mennicke et al. 1988).

ITC attack of DNA and cancer cell induction seem to be unim portant in describing the m ode
of

toxicity. Mutagenicity tests are som e tim es positive, but of ten negative (Fenwick et al.

301983; Ioannou et al. 1984; Musk and Johnson 1993) . Chronic adm i nistration of propenyl
ITC to rats and m i ce caused papillary growths in bladders of m a le rats only (related to m o re concentrated m e tabolite excretion), but did not cause tum o rs (Ioannou et al. 1984). There is in fact evidence that ITCs are anticarcinogenic (Mossoba et al. 1989; Fim ognari et al. 2004). Glucosinolate-containing cruciferous vegetables such as cabbage, broccoli, and Brussels sprouts are correlated with reduced incidence of cancer (Graham et al. 1978). Tum o rs were inhibited in m i ce and rats that were given ITCs both before and after adm i nistering known carcinogens (Fenwick et al. 1983; W a ttenberg 1977, 1981). Short-term chem ical

m echanism s appear to be involved in the suppression, such as high toxicity toward cancer cells (Nastruzzi et al. 1996), but longer term i nduction of enzym e system s that fight cancer in the body are also im plicated (Institute of Food Technologists 1993). A potent inducer of protective enzym e s is 4-m e thylsulfinylbutyl ITC found in broccoli (Zhang, et al. 1992, 1994). Thus, if ITC is consum ed in norm a l dietary levels, it m a y be m o re beneficial than
harm ful.

Toxicological im plications of glucosinolate hydr olysis m u st also include the possibility of
OZT form ation. The m o st com m on OZT, 5-vinyl-2-oxazolidinethione, is often referred to as “goitrin” because of its ability to induce goiter in the thyroid gland. High-protein rapeseed

m eal has lim ited use as a cattle feed, in part because the glucosinolate precursor of goitrin is present in large am ounts. Other inform ation on biological activity is relatively scarce, although OZTs interfere with a few enzym a tic processes. For exam ple, OZTs prevent the
oxidation of trim ethylam ine (Fenwick et al. 1983) and goitrin m oderately inhibits dopam i ne β -hydroxylase (Zenker et al. 1988). OZT toxicities to m i ce reported in the form of LD 50 values range from 1260 to 1415 m g kg -1 (Van Etten and Tookey 1979). There is som e susceptibility to nitrosation in the presence of nitrite (Lüthy et al. 1984), suggesting the possible form ation of carcinogenic com pounds under conditions encountered in the stom ach. General toxicity appears to be relatively low, but f u rther investigation of OZT toxicities

seem s appropriate.

Toxicity of other organic products, including nitriles, epithionitriles, and thiocyanates is likely related to the cyano group. Cyanide salts and gas inactivate certain enzym e system s, especially those involved in cellular respiration such as cytochrom e oxidase (Johnston 1987; Sittig 1985). Sim ilarities in the m e tabolism of organic nitriles and cyanide include increased urinary excretion of SCN - , detection of CN - in tissues, and depressed cytochrom e oxidase activity (Duncan 1991). Toxicities of organic nitriles vary but are generally lower than cyanide salts and hydrogen cyanide. Toxicity seem s particularly pronounced towards the liver (hepatotoxic) and kidney (Dietz et al . 1991; Duncan 1991; Fenwick et al. 1983; Van Etten et al. 1969). The LD 50 in rats for ethyl CN (propanenitrile) is 50-100 m g kg -1 (Sittig 1985); for 1-cyano-2-hydroxy-3-butene, values of 170 m g kg -1 and 200 m g kg -1 have been reported (Fenwick et al. 1983; Tookey et al. 1980). Unsaturated nitriles appear to be less

toxic than saturated nitriles. For exam ple, rat oral LD 50 ’s exceeded 600 m g kg -1 for 3- butenyl CN (4-pentenenitrile) and 720 m g kg -1 for benzonitrile, although acetonitrile (m ethyl CN) is an exception with a reported oral LD 50 in rats of 3030-6500 m g kg -1 (Dietz et al. 1991; Sm iley 1979).

31Thiocyanate and epithionitrile toxicities appear sim ilar to nitrile toxicities. Low m o lecular weight thiocyanates, including benzyl and phenyl, respond to the sam e antidotes for poisoning as does cyanide (W ood 1975). Alkyl thiocyanates are generally m o re active than

arom atic thiocyanate, and toxicity decreases with increasing m o lecular weight (W ood 1975). For 1-cyano-3,4 epithiobutane, m o rtality in rats occurred after ingestion of 180 m g kg -1 (Dietz et al. 1991) and an LD 50 of 109 m g kg -1 has been reported (Fenwick et al. 1983). An

LD 50 range of 178-240 m g kg -1 was estim ated for 1-cyano-2-hydroxy-3,4 epithiobutane (Van Etten et al. 1969).

Although the LD 50 reported for 3-indolylacetonitrile in rats is 255 m g kg -1 , indolyl products such as 3-indolylm e thanol stim ulate enzym a tic system s involved in detoxification of xenobiotics and the inhibition of cancer (Loub et al. 1975). Protective activity, however, is
less dram atic than with ITCs. Results of several experim e nts are som e what m i xed depending on the system exam ined and the specific conditions (McDanell et al. 1988), m a king it
difficult to assess actual im pact. In one study conducted with trout, 3-indolylm e thanol
adm i nistered after exposure to aflatoxin B 1 actually enhanced carcinogenesis. However, 3- indolylm e thanol, as well as several ITCs, OZTs, and nitriles, were not teratogenic in rats (Fenwick at al. 1983).

SCN

- is generally less toxic than either ITCs or organic cyano com pounds. In fact, biological system s often m e tabolically detoxify cyanides by rhodanese-catalyzed conversion
to SCN

- , which is then readily excreted (exogenous sulfur is needed and thus thiosulfate is
adm i nistered as part of the antidote for cyanid e poisoning). Rhodanese is widely distributed in nature (Tabatabai and Singh 1976). As a result, SCN

- was considered nontoxic and was
used for a tim e as a drug to reduce blood pressure. However, side effects of SCN

- caused its use to be dropped.

SCN

- toxicity to rats is sim ilar to benzonitrile with a reported LD 50 for NaSCN of 765 m g kg -1 . Dogs are m o re susceptible than rats or m i ce to SCN

- (W ood 1975). The process of cyanide conversion to SCN

- is apparently in m e tabolic equilibrium , and adding SCN

- to biological system s produces cyanide. However, this alone does not account for its acute toxicity (W ood 1975). SCN

- interacts with proteins, and is listed as the m o st effective anion
in the Hofm eister series for destabilizing protein structure (Voet and Voet 1990; W ood 1975). It m a y act as a noncom petitive inhibitor with m a ny enzym e s, and penetrates cell

m e m b ranes (W ood 1975). SCN

- is a pseudohalogen that acts as a com p etitor with iodine
and thus m a y cause goiter. Unlike goiter caused by OZT, goiter resulting from SCN

- can be corrected by adding iodine to the diet.

Effects of specific glucosinolate degradation products on individual organism s vary and are not always known. In sufficient quantity m a ny of these com pounds are highly toxic. Even with plant tissues, irritation of skin from excessive handling or a negative respiratory
reaction m i ght occur. However, exposure to am ounts encountered in pest control strategies with plant tissues is likely to be m i nim a l. The action of the toxins is direct and “up front.” There do not appear to be serious, m o re subtle, negative long-term im pacts on hum an health such as cancer or birth defects. Indeed, we have probably ingested m a ny of these com pounds in low levels for thousands of years, m o st of the tim e presum ably without detrim ent.

32

### VII. Registration

Products derived from m u stard m eal or m eal-cont aining glucosinolates have been registered for a variety of purposes (Table 4). EPA has granted an “Exem p tion From the Requirem e nt of Tolerance to Isothiocyanate as a Com pone nt of Food Grade Oil of Mustard” (Federal Register 1966). The m o st relevant products in the current discussion include m u stard m eal ( B. juncea ), currently m a rketed in Australia as a snail and slug killer and m eadowf o am ( Limnanthes alba ) m eal m a rketed in the United States as a soil am endm ent in horticulture.
Meadowfoam m eal is m a rketed by Natural Plant Products as Lim n ax TM , AlbaGro TM , and
AlbaAide TM . EPA approved pesticides containing pr openyl ITC as an active ingredient are

m a rketed by Cham pon Natural Products (Cham pon Millenium Chem icals, Inc. 11112 Split Rail Lane, Fairfax Station, VA 22039; Phone: 703-426-8424; E-m a il:
cham pon@ix.netcom .com ; website: www.cham pon.com ). A num ber of other products containing propenyl isothiocyanate as an activ e ingredient have been approved by the EPA for various pests (Table 4). Thus, the precedent for EPA approval of products containing propenyl ITC as an active ingredient has been set. Additional ITCs contained in other m e als
other than B. juncea m o st likely will require greater testing bef o re it can be approved by the
EPA.

33

### VII. Genetics and Interspecific Hy bridization

Canola or rapeseed ( Brassica napus L.) is a m e m b er of the Brassicaceae fam ily. Canola is
the term used to describe Brassicaceae cultivars that contain erucic acid less than 2% of the
total f a tty acid content and less than 30 μ m o l per gram of total glucosinolates in oil-free m eal (Arnoldo et al. 1992). Rapeseed is used to describe oilseed B. napus with high (>40%) erucic acid content in the oil. Traditionally rapeseed cultivars also contained high (80-120

μ m o l per gram of total glucosinolates in oil-free m eal) glucosinolate content in the seed

m eal, although new rapeseed cultivars have seed m eal com p arable to canola.

Early records indicate that canola (rapeseed) was cultivated in India m o re than 3,000 years ago. It was later introduced to China and Japan (Shahidi 1990). The exact center of origin
for B. napus is uncertain; however, it m a y have had m u ltiple areas of origin in Europe within overlapping ranges of the diploid parental taxa ( B. rapa and B. oleracea ) (W arwick and Francis 1994). The m a in phytogeographical zones for B. napus are the Euro-Siberian, and the Irano-Turanian regions (W arwick and Francis 1994). The m a in regions of canola cultivation are parts of Asia (China, India, Indonesia), Europe, the Am ericas (m ainly Canada), parts of Africa, and Australasia.

A. Taxonomy

, Genomic Constitutions, and Relationships

The origins of am phidiploid B. napus

, B. carinata , and B. juncea were suggested by U (1935), based on interspecific hybridizations and chrom o som e counts (Figure 3). This triangle arrangem e nt of species indicates that cultivated canola, B. napus (n = 19, AACC

genom e), originated as a result of natural hybridization between B. oleracea (n = 9, CC

genom e) and B. rapa (n = 10, AA genom e) (U 1935; McNaughton 1976). This has been
confirm e d by later studies using different t echniques, including flavonoid com position (Dass and Nybom 1967) and seed protein serology (Vaughan 1977). Song et al. (1988) used RFLPs to study the genom ic constitution of dipl oid and am phidiploid species, and concluded
that am phidiploid B. napus has evolved through different com b inations of the diploid

m o rphotypes ( B. rapa and B. oleracea ) and that phylogenetic origins m a y be com m on

m echanism s for the natural occurrence of am phidiploids in Brassica , which confirm e d that B. napus is an interspecific hybrid derived from B. rapa and B. oleracea . The authors suggested that the C genom e com ponent is larger in the cultivated B. napus lines, since these plants were closer to the C genom e than the A genom e in the phylogenetic tree.

B. Oil and Seed Meal Characteristics and Inheritance

The m a jor fatty acids of Brassica species oil are palm itic (16:0), stearic (18:0), oleic (18:1), linoleic (18:2), linolenic (18:3), eicosenoic (20:1) and erucic acids (22:1). Som e m u stard species also have relatively high concentrations of ne rvonic acid (24:1). Canola oil m u st, by definition,
not exceed m o re than 2% erucic acid content. In contrast, industrial rapeseed oil is usually very high (40%-50%) in erucic acid content. Laborat ory anim als that eat diets with a high proportion of oil containing high erucic acid levels perform poorly. Also, high erucic acid levels in hum an

34diets can cause abnorm a lity of fat utilization in the heart and skeletal m u scles, with undesirable changes in the tissue of the m yocardium (Roine et al. 1960). Other reports from Canada and Europe have confirm e d these changes and have shown that they do not occur with diets containing canola type oil with a low erucic acid content (Abdellatif et al. 1970). In canola,
reduced erucic acid content is alm o st always associated with high oleic acid content, which is
alm o st as effective in reducing cardiovascular ri sk as is the long-touted polyunsaturated linoleic acid (Ackm a n 1990).

Linolenic acid is very unstable and is easily oxidized to give unpleasant sm elling substances. Canola oil contains 8%-12% linolenic acid, although cultivars can be developed with reduced levels (DeBonte 1994). In contrast, a high percenta ge of linoleic acid in edible oil is desirable (Scarisbrick and Daniels 1986). Canola oil is relatively low in this nutritionally valuable fatty acid, and an increase would im prove its dietary value (Holm e s 1980). Nutritionally, linoleic acid (vitam i n F) is an im portant and essential polyunsaturated fatty acid that also reduces cholesterol
content in the blood and is an essential com ponent of cell m e m b ranes (Scarisbrick and Daniels 1986).

Yellow m u stard ( S. alba ) contains considerably higher erucic acid levels than are suitable as an edible oil. Also, yellow m u stard does not produce high enough erucic acid levels to be valuable for industrial purposes (Carlson and Van Dyne 1992). In addition, yellow m u stard is low in linoleic acid and has higher linolenic acid than would be desirable in edible oil.

Since the early 1960s, m a ny researchers have investigated the inheritance of erucic acid content
in several Brassica species. In B. napus , erucic and eicosenoic acid content are controlled by the genotype of the developing em bryo and not by the sporophyte (Dorrel and Downey 1964; Harvey and Downey 1964; Kondra and Stefanss on 1965; Stefansson and Hougen 1964). Erucic
acid content in B. napus is controlled by a two gene-pair-system acting in an additive m a nner (Downey and Craig 1964; Kondra and Stefanss on 1965; Stefansson and Hougen 1964). Jönsson (1977a, 1977b) found that erucic acid content in B. napus is controlled by a series of m u ltiple alleles. Levels of 5%-10%, 20%-35%, and m o re than 35% erucic acid content are controlled by alleles at one, one or two, and two loci, respectively.

Very few studies have been carried out to exam ine the inheritance of fatty acid profiles in S. alba (Rafiullah 1994). Rafiullah used a 4x4 half dialle l cross to determ ine the inheritance of fatty acid prof ile as quantitative characteristics. The results indicated that erucic and oleic acids had greater additive genetic variation than dom inant genetic variation, and that erucic acid was controlled by additive and nonadditive genes with som e degree of linkage and or epistasis in S.
alba . However, these studies were conducted by chem ically analyzing field open pollinated
seed. This could add serious bias to the resu lts because of outcrossing. Tang et al. (1995) found that erucic acid content in yellow m u stard has a large quantitative com ponent with a few m a jor
dom inant alleles.

Traditional rapeseed cultivars (where canola cultivars were derived) all had relatively high
concentrations of glucosinolates in the seed that adversely af f e cted the quality of the m eal as
livestock feed. W h en glucosinolate com pounds are enzym a tically degraded, they produce toxic

com pounds that can cause m e tabolic disturbances when fed to nonrum inant livestock (Kondra

35and Stefansson 1970). High glucosinolate levels needed therefore to be reduced in rapeseed
cultivars to avoid these detrim ental effects on anim als consum ing the m eal (Josefsson 1972).

A trem endous breeding effort was m a de to devel op low-glucosinolate rapeseed (canola). The cultivar “Bronowsky” was the original, and only, source of low glucosinolates in B. napus and B. rapa . The genes that affect the early stages of biosynthesis of the m a jor glucosinolates apparently control biosynthesis of glucosinolates in Bronowsky. This cultivar had low levels of glucosinolates in 1968 but was unadapted (J önsson 1977a; Josefsson and Appelqvist 1968; Kondra and Downey 1969; Kondra and Stefansson 1970). However, the cultivars “Tower” and “Erglu” were developed relatively quickly by a backcross procedure to com b ine low levels of glucosinolates with low levels of erucic acid (Stefansson and Kondra 1975).

In contrast to canola, yellow m u stard seed has traditionally been selected for condim e nt m u stard powders. Mustard pungency is associated with high content of glucosinolates and their hydrolysis products (Holm e s 1980). Therefore, past breeding efforts in this species have been
aim e d at increasing glucosinolate levels rather than m a king any attem p t to reduce the content of glucosinolates in the seed.

Very few studies have exam ined the inheritance of glucosinolates in yellow m u stard. This is

m a inly because very little genetic variation has been found for this trait within the species.
Glucosinolate content of S. alba seeds produced from the sam e parent fromboth self- and crosspollinations were sim ilar, showing that inheritance of glucosinolates is m a ternally controlled (Kondra and Stefansson 1970; Love et al. 1990). F 2 seed of reciprocal crosses contained both 2- propenyl and 3-butenyl glucosinolates, indicated nuclear genes controlled their form ation. Love et al. (1990) also concluded that the synthesis of both 2-propenyl and 3-butenyl glucosinolates were controlled by m u ltiple additive alleles at the sam e loci. More recently a single recessive allele has been identified (Brown et al. 2002) that com p letely blocked 3-hydroxybenzyl glucosinolate in yellow m u stard seed m eal.

C. Interspecific and Intergeneric Crosses in Brassicaceae

Substantial literature discusses intergeneric cro ss com p atibility within the f a

m ily Brassicacae. B.

napus has been com b ined with several species of Diplotaxis and Eruca by sexual crosses and

em bryo rescue (Batra et al. 1990; Ringdahl et al. 1987), and with various other Brassica species (Glim elius et al. 1990; Jourdan et al. 1989a; Sjodin and Glim elius 1989). Other intergeneric

Brassica hybrids produced include various com b inations of B. juncea

, B. campestris

, Diplotaxis

, Eruca

, Raphanus

, Moricandia , and Trachystoma (e.g. Agnihotri et al. 1990a, 1990b; Bing et al. 1996; Chevre et al. 1997; Kirti et al. 1992; Metz et al. 1997; Sikdar et al. 1990; Takahata and Takeda 1990; Toriyam a et al. 1987a, 1987b). These hybrids generally showed som e degree of fertility in crosses and have a range of chrom o som e num bers in their progeny.

Agnihotri et al. (1990a, 1990b) reported on the production of B. napus × Raphanobrassica
and B. napus × Eruca using controlled crossing followed by em bryo rescue. Lefol and

Darm ency (1993) studied spontaneous and m a nual interspecfic and intergeneric crosses
between B. napus and two weedy relatives, B. adpressa and R. raphanistrum . They were

36able to obtain hybrids that showed superior vegetative developm ent com p ared to the weed parent; the backcrosses to the parents also produced seed. In their studies on interspecific crosses, Nishiyam a et al. (1991) found that m a ny of these crosses produced wrinkled or

em pty seed because of poor endosperm developm ent. However, crosses between B. napus
and B. campestris and between B. napus and B. nigra spontaneously produced a few true F 1 seeds (Nishiyam a et al. 1991). Sucessful intergeneric hybridization between B. napus and S.
alba have recently been reported (Brown et al. 1994, 1997). The result of this latter hybrid

com b ination prom pted further investigation of B. napus x S. alba hybrids with elevated glucosinolate content for use as biofum igants.

Kerlan et al. (1992) studied the potential for interspecific hybrid production between B.

napus and five related species ( B. oleracea L. var capitate

, B. oleracea L. var acephala

, B. nigra L. Koch, B. adpressa L., R. raphanistrum L., and S. arvensis ). Under optim al
conditions with in vitro ovary culture, hybrid plants were obtained in all species cross

com b inations and m o re hybrids were obtained when canola was used as the fem a le.

Eber et al. (1994) reported that spontaneous hybridization between m a le sterile B. napus and B. adpressa and R. raphanistrum where the pollen fertility of hybrids was 0%-30%.

In a study involving six B. napus and four S. arvensis lines (Bing et al. 1995), no hybrids
were obtained from any cross. W h en ovule culture was used, one hybrid from B. napus × S.

arvensis was obtained but was highly sterile. Th e authors concluded that the crossing between and transfer of genes from B. napus to S. arvensis would be rem o te even under
favorable conditions.

37

### IX. Recommendations

Meals with isothiocyanate-producing glucosinol ate concentrations in excess of 200 μm ol g -1

tissue will m o st effectively control a wide variety of plant pests. The target is to produce in
excess of 100 nm ol ITC g -1 soil to approach com m e rcially recom m e nded rates for synthetic ITC pesticides. This should be possible assum i ng glucosinolate to ITC release efficiencies of 20%. Consideration should be given to increasi ng the release efficiency of the ITC from the

m eal using physical m e thods of processing the m eal or through the addition of adjuvants.
Absolute ITC am ounts and release efficiencies m u st be determ ined for the m e als of interest
to confirm that we indeed are producing ITC in excess of 100 nm ol g -1 soil. Meals m u st be tested against specific target organism s a nd phytotoxic effects on the crop plant m u st be
determ ined. Fungal pathogens will be the m o st difficult to control because carbon added to
the soil in the form of m eal m a y enhance growth of the pathogen. Insects, nem a todes, and
weeds will be m o re easily controlled, and com m e rcialization of a final product with acceptable efficacy is likely. Different ITCs expr ess different toxicities as a result of direct
ef

f ects on the plant pest as well as dif f e rential sorption and reaction with soil colloids. Bioassay results obtained in the absence of soil m u st theref ore be conf irm e d within the soil
environm ent. Predictions of ITC pesticidal activity should be approached in the sam e fashion as com m e rcial pesticides by determ ining organic carbon partition coefficients, Henry’s constants, and half -lives. These param e ters will vary with the ITC, and pesticidal activity can be optim ized by selecting ITCs with the greatest potential f o r ef f i cacy. The potential to introduce thiofunctionalized glucosinolates into species by way of breeding
efforts m a y im prove pesticidal efficacy of the m eal. Additional benefits from the m eal such
as N release m a y increase the advantages of using m eal am endm ents in cropping strategies. Effort should be devoted to quantifying the nutrient benefits of m eal am endm ent since such
benefits m a y be just as im portant to the consum er as pesticidal activity.

38X. LITERATURE CITED

Abdellatif, A.M.M. and Vles, R.O. Physical Eff ects of Rapeseed Oil and Canola Oil in Rats. In : Proc. International Conference on Rapeseed and Rapeseed Products, St. Adele, Canada. 1970, pp. 423-431.

Ackm an, R.G. Canola Fatty Acid - An Ideal Mixture for Health, Nutrition and Food Use. In : Canola and Rapeseed Production, Chem istry, Nutrition and Processing. Shahidi, F (Eds). van Nostrand Reinhold, NY. 1990

.

Agnihotri, A., Gupta, B., Lakshm ikum aran, M.S., Shivanna, K.R., Prakash, S., and Jagannathan, V. Production of Eruca-Brassica Hybrids by Em bryo Rescue. Plant Breeding
1990a

, 104 , 281-289.

Agnihotri, A., Shivanna, K.R., Raina, S.N., Lakshim i kum aran, M., Prakash, S., and Jagannathan, V. Production of Brassica napus × Raphanobrassica Hybrids by Em bryo
Rescue: An Attem p t to Introduce Shattering Resistance into B. napus

. Plant Breeding 1990b

, 104 , 292-299.

Åhm a n, I. Toxicities of Host Secondary Com pounds to Eggs of the Brassica Specialist
Dasineura brassicae

. J. Chem Ecol

. 1986

, 12 , 1481-1488.

Al-Shehbaz, I.A. and Al-Sham m a ry, K.I. Di stribution and Chem otaxonom ic Significance of
Glucosinolates in Certain Middle-Eastern Cruciferae. Biochem. Systemat. Ecol

. 1987

, 15

, 559-569.

Alvarez, R.A. and Moore, C.B. Quantum Yield for Production of CH 3 NC in the Photolysis of
CH 3 NCS. Science 1994

, 263 , 205-207.

Andersen, A.S. and Muir, R.M. Auxin Activity of Glucobrassicin. Phys. Plantarum 1966

, 19

, 1038-1048.

Angus, J.F., Gardner, P.A., Kirkegaard, J.A., and Desm archelier, J.M. Biofum igation: Isothiocyanates Released from Brassica Roots Inhibit Growth of the Take-All Fungus. Plant
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71Table 1. NaturallyOccurring Glucosinolates____________________________________________________________________________________________________________________R= trivialnameplantpartaReference.____________________________________________________________________________________________________________________methyl CH3glucocapparinvariousCapparissKjaer and Thomsen 1963avariousCleomesKjaer et al. 1955sKjaer and Thomsen 1963aethyl CH3CH2glucolepdiinLepidiummenziesiisKjaer and Larsen 19542-methylethyl (isopropyl) (CH3)2CH glucoputranjivinBrassica napobrassicarShaw et al. 1989Cochlearia officinalissKjaer and Conti 1953Lunaria biennissKjaer and Conti 1953Moringa peregrinasKjaer et al. 1979propyl CH3CH2CH2Brassica oleraceaelMacLeod and Nussbaum 19771-methylpropyl CH3CH2(CH3)CH glucocochlearinBrassica napobrassicarShaw et al. 1989Brassica campestris(B. rapa)l,rCarlson et al. 1987Dentaria pinnatarDelaveau and Kjaer 19632-methylpropyl CH3(CH3)CHCH2Armoracia lapathifoliarGrob and Matile 1980rGilbert and Nursten 1972BrassicanapussBrown and Morra 1995Conringia orientalislUnderhill and Kirkland 1972Moringa peregrinasKjaer et al. 1979butyl CH3(CH2)3Armoracia lapathifoliarGrob and Matile 1980Capparis flexuosalKjaer and Schuster 19712-methylbutyl CH3CH2(CH3)CHCH2Armoracia lapathifoliarGrob and Matile 1980Dentaria pinnatarDelaveau and Kjaer 19633-methylbutyl CH3(CH3)CHCH2CH2Armoracia lapathifoliarGrob and Matile 1980pentyl CH3(CH2)4Armoracia lapathifoliarGrob and Matile 1980Brassica rapalItoh et al. 1984724-methylpentyl CH3(CH3)CH(CH2)3Armoracia lapathifoliarGrob and Matile 1980Brassica napusl,s,rBrown and Morra 1995,1996Raphanus sativusrKjaer et al. 1978hexyl CH3(CH2)5Brassica napussKondo et al. 1985Raphanus sativusrKjaer et al. 19785-methylhexyl CH3(CH3)CH(CH2)4Armoracia lapathifoliarGrob and Matile 1980Brassica napusl,s,rBrown and Morra 1996allyl CH2CHCH2sinigrinArmoracia lapathifoliarGrob and Matile 1980rBirch et al. 1992lShaw et al. 1989BrassicacarinatasNastruzzi et al. 1996sRöbbelen and Thies 1980Brassica junceasHanley et al. 1983l, s, rSang et al. 1984Brassica napussElfakir et al. 1992rBirch et al. 1992Brassica oleraceaesMinchinton et al. 1982lHill et al. 1987l,rJosefsson1967a,b2-methyl-2-propenyl CH2(CH3)CCH2Peltaria alliaceasDaxenbichler et al. 19913-butenyl CH2CHCH2CH2gluconapinBrassica campestris (B. rapa)sHanley et al. 1983l,rCarlson et al. 1987Brassica oleraceaelShaw et al. 1989rBirch et al. 1992sMinchinton et al. 1982Brassica napussElfakir et al. 1992lKoritsas et al. 1989l, sSang et al. 1984rBirch et al. 1992Brassica napobrassicarShaw et al. 19893-methyl-3-butenyl CH2(CH3)CCH2CH2CapparislinearislKjaer and Wagnières 1965734-pentenyl CH2CH(CH2)3glucobrassicanapinArmoracialapathifoliarGrob and Matile 1980rGilbert and Nursten 1972Brassica campestris (B. rapa)l,rCarlson et al. 1987BrassicaoleraceaesMinchinton et al. 1982Brassica napussQuinsac et al. 1991lKoritsas et al. 1989l, sSang et al. 1984rBirch et al. 1992Brassica napobrassicarShaw et al. 19895-hexenyl CH2CH(CH2)4Armoracia lapathifoliarGrob and Matile 1980Brassica napussKondo et al. 1985Raphanus sativusrKjaer et al. 19782-hydroxypropyl CH3(OH)CHCH2Armoracia lapathifoliarGrob and Matile 19803-hydroxypropyl HOCH2(CH2)2Arabidopsis thalianasHogge et al. 19883-hydroxybutyl CH3(OH)CH(CH2)2Armoracia lapathifoliarGrob and Matile 1980Capparis flexuosalKjaer and Schuster 19714-hydroxybutyl HOCH2(CH2)3Arabidopsis thalianasHogge et al. 1988Capparis flexuosalKjaer and Schuster 19712-hydroxypentyl CH3(CH2)2(OH)CHCH2Armoracia lapathifoliarGrob and Matile 19801-methyl-2-hydroxyethyl HOCH2(CH3)CH glucosysimbrinColuteocarpus vesicariasAl-Shehbaz, Al-Shammary 1987Dentaria laciniatasDaxenbichler et al. 1991EuclidiumsyriacumsAl-Shehbaz, Al-Shammary 1987Torularia torulosasAl-Shehbaz, Al-Shammary 19871-hydroxymethylpropyl CH3CH2(OHCH2)CHColuteocarpus vesicariasAl-Shehbaz, Al-Shammary 19871-ethyl-2-hydroxyethyl HOCH2(CH3CH2)CH glucosisautricinCleome diandrasDaxenbichler et al. 1991SisymbriumaustriacumsKjaer and Christensen 1962a2-hydroxy-2-methylpropyl CH3(OH)(CH3)CCH2glucoconringiinCochlearia officinalissChallenger1959Conringia orientalissChallenger 195974lUnderhill and Kirkland 1972not specEagles et al. 1981ErysimumorientalesKjaer19602-hydroxy-2-methylbutyl glucocleominCleome spinosasKjaer and Thomsen 1962CH3CH2(OH)(CH3)CCH2Dentaria laciniatasDaxenbichler et al. 19914,5,6,7-tetrahydroxydecyl CH3(CH2)2(CH2OH)4(CH2)3Capparis grandisrGaind et al. 1975R-2-hydroxy-3-butenyl CH2CH(OH)CHCH2progoitrinBrassica campestris (B. rapa)sSoluski and Dabrowski 1984(glucorapiferin)l,rCarlson et al. 1987Brassica napobrassicarShaw et al. 1989Brassica napussQuinsac et al. 1991lKoritsas et al. 1989l, s, rSang et al. 1984s Daun1986rBirch et al. 1992Brassica oleraceaesMinchinton et al. 1982lShaw et al. 1989rBirch et al. 1992S-2-hydroxy-3-butenyl CH2CH(OH)CHCH2epiprogoitrinCrambe abyssinicasHanley et al. 1983sWetter and Dyck 1973sBartlet et al. 1994Brassica napuss Daun19862-hydroxy-4-pentenyl CH2CHCH2(OH)CHCH2gluconapoleiferinBrassica napobrassicarShaw et al. 1989Brassica napussElfakir et al. 1992lKoritsas et al. 1989l, s, rSang et al. 1984rBirch et al. 1992Brassica oleraceaerBirch et al. 19923-hydroxy-4-pentenyl CH2CH(OH)CHCH2CH2Armoracia lapathifoliarGrob and Matile 19804-oxoheptyl CH3(CH2)2CO(CH2)3glucocapangulinCapparis angulatasKjaer and Thomsen 1963anot spec.Ettlinger and Kjaer 19685-oxoheptyl CH3CH2CO(CH2)4gluconorcappasalinCapparis salicifoliasKjaer and Thomsen 1963b755-oxooctyl CH3(CH2)2CO(CH2)4glucocappasalinCapparis salicifoliasKjaer and Thomsen 1963a3-methoxycarbonylpropyl CH3OCO(CH2)3glucoerypestrinErysimumaucherianumsAl-Shehbaz, Al-Shammary 1987ErysimumfilifoliumsAl-Shehbaz, Al-Shammary 1987ErysimumrupestresKjaer1960lChisholm19732-methylthioethyl CH3S(CH2)2Armoracia lapathifoliarGrob and Matile 1980(Cochlearia armoracia)3-methylthiopropyl CH3S(CH2)3glucoibervirinArabidopsis thalianasHogge et al. 1988Brassica napussMinchinton et al. 1982Brassica oleraceaesMinchinton et al. 1982lShaw et al. 19894-methylthiobutyl CH3S(CH2)4glucoerucinArabidopsis thalianal,sHogge et al. 1988Brassica campestris (B. rapa)rCarlson et al. 1987Brassica napusrSang et al. 1984rBirch et al. 1992Brassica oleraceaesMinchinton et al. 1982rBirch et al. 1992Eruca sativasKjaer and Gmelin 1955sNastruzzi et al. 1996Raphanus sativusrKjaer et al. 1978Sinapis alba (B. hirta)rBrown and Morra, unp.5-methylthiopentyl CH3S(CH2)5glucoberteroinArabidopsis thalianasHogge et al. 1988Brassica naporbrassicarShaw et al. 1989Brassica campestris (B. rapa)rCarlson et al. 1987Raphanus sativusrKjaer et al. 19786-methylthiohexyl CH3S(CH2)6glucolesquerellinArabidopsis thalianasHogge et al. 1988OchthodiumaegyptiacumrsAl-Shehbaz, Al-Shammary 19877-methylthioheptyl CH3S(CH2)7Arabis laxasAl-Shehbaz, Al-Shammary 1987Arabidopsis thalianasHogge et al. 19888-methylthiooctyl CH3S(CH2)8Arabidopsis thalianal,sHogge et al. 1988769-methylthiononyl CH3S(CH2)9Arabis hirsutasDaxenbichler et al. 199110-methylthiodecyl CH3S(CH2)10Arabis amplexicaulissDaxenbichler et al. 19914-methylthio-3-butenyl CH3SCHCH(CH2)2glucodehydroerucinRaphanus sativussDaxenbichler et al. 1991rVisentin et al. 19923-hydroxy-5-methylthiopentyl CH3S(CH2)2(OH)CH(CH2)2ErysimumhieracifoliumsKjaer and Schuster 1970Armoracia lapathifoliarGrob and Matile 19803-hydroxy-6-methylthiohexyl CH3S(CH2)3(OH)CH(CH2)2ErysimumrhaeticumsKjaer and Schuster 19733-oxo-8-methylthiooctyl CH3S(CH2)5CO(CH2)2Arabis hirsutasKjaer and Schuster 19723-methylsulfinylpropyl CH3SO(CH2)3glucoiberinArabidopsis thalianalHogge et al. 1988Brassica napuss Daun1986Brassica oleracearBirch et al. 1992lHansen et al. 19954-methylsulfinylbutyl CH3SO(CH2)4glucoraphaninArabidopsis thalianal,sHogge et al. 1988(sulphoraphane)Brassica napuss Daun1986Brassica oleracealHansen et al. 1995lHill et al. 19875-methylsulfinylpentyl CH3SO(CH2)5glucoalyssinAlyssumargenteumsKjaer and Gmelin 1956Arabidopsis thalianal,sHogge et al. 1988Berteroa incanasKjaer and Gmelin 19566-methylsulfinylhexyl CH3SO(CH2)6glucohesperinArabidopsis thalianal,sHogge et al. 1988EuclidiumsyriacumsAl-Shehbaz, Al-Shammary 1987Notoceras bicornesAl-Shehbaz, Al-Shammary 19877-methylsulfinylheptyl CH3SO(CH2)7glucosiberinArabis laxasAl-Shehbaz, Al-Shammary 1987Arabidopsis thalianal,sHogge et al. 19888-methylsulfinyloctyl CH3SO(CH2)8glucohirsutinAethionema arabicumsAl-Shehbaz, Al-Shammary 1987Arabidopsis thalianal,sHogge et al. 1988779-methylsulfinylnonyl CH3SO(CH2)9glucoarabinArabis alpinasChallenger 1959Arabis novasAl-Shehbaz, Al-Shammary 1987Drabopsis nudasAl-Shehbaz, Al-Shammary 198710-methylsulfinyldecyl CH3SO(CH2)10glucocamelininvariousCamelinasKjaer et al. 1956b11-methylsulfinylundecyl CH3SO(CH2)11Camelina microcarpasDaxenbichler et al. 19914-methylsulfinyl-3-butenyl CH3SOCHCH(CH2)2glucorapheninBrassica napuss Daun1986rBirch et al. 1992RaphanussativussSang et al. 1984sNastruzzi et al. 19963-hydroxy-5-methylsulfinylpentyl CH3SO(CH2)2(OH)CH(CH2)2ErysimumhieracifoliumbsKjaer and Schuster 19703-hydroxy-6-methylsulfinylhexyl CH3SO(CH2)3(OH)CH(CH2)2ErysimumrhaeticumsKjaer and Schuster 19733-oxo-8-methylsulfinyloctyl CH3SO(CH2)5CO(CH2)2Arabis hirsutasKjaer and Schuster 19723-methylsulfonylpropyl CH3SO2(CH2)3glucocheirolinBrassica oleraceaelShaw et al. 1989Cheiranthus cheiriChallenger1959sNastruzzi et al. 19964-methylsulfonylbutyl CH3SO2(CH2)4glucoerysolinCheiranthus cheiriChallenger1959ErysimumperofskianumsKjaer and Conti 1954not specEagles et al. 19816-methylsulfonylhexyl CH3SO2(CH2)6EuclidiumsyriacumsAl-Shehbaz, Al-Shammary 19878-methylsulfonyloctyl CH3SO2(CH2)8Heliophila amplexicaulissDaxenbichler et al. 19919-methylsulfonylnonyl CH3SO2(CH2)9ArabisturritasDaxenbichler et al. 1991Heliophila amplexicaulissDaxenbichler et al. 199110-methylsulfonyldecyl CH3SO2(CH2)10ArabisturritasDaxenbichler et al. 19914-methylsulphonyl-3-butenyl CH3SO2CHCH(CH2)2Raphanus sativussCole1980783-hydroxy-5-methylsulfonylpentyl CH3SO2(CH2)2(OH)CH(CH2)2ErysimumhieracifoliumbsKjaer and Schuster 19706-methylsulfonyl-3-hydroxyhexyl CH3SO2(CH2)3(OH)CH(CH2)2ErysimumrhaeticumsKjaer and Schuster 1973Benzyl C6H5CH2glucotropaeolinCarica papayacHanley et al. 1983LepidiumsativumsHanley et al. 1983sHasapis and MacLeod 1982Reseda medias,lOlsen and Sørenson 1980Sinapis alba (B. hirta)l,rBrown and Morra, unplCole1976Tropaeolummajuss,lLykkesfeldt and Møller 1993lUnderhill and Wetter 19692-hydroxybenzyl 2-OHC6H5CH2cReseda odorataOlsen and Sørensen 19793-hydroxybenzyl 3-OHC6H5CH2glucolepigraminLepidiumvesicariumsDaxenbichler et al. 1991Reseda medias,lOlsen and Sørenson 19804-hydroxybenzyl 4-OHC6H5CH2glucosinalbinBrassica napussQuinsac et al. 1991sElfakir et al. 1992Sinapis alba (B. hirta)sMinchinton et al. 1982sNastruzzi et al. 1996l,rBrown and Morra, unpSinapis arvensissAl-Shehbaz, Al-Shammary 19873,4-dihydroxybenzyl 3,4(OH)2C6H5CH2Kjaer and Larsen 19733-methoxybenzyl 3-CH3OC6H5CH2Limnanthes douglasiisKjaer1960not spec.Ettlinger and Kjaer 19684-methoxybenzyl 4-CH3OC6H5CH2glucoaubrietinAubrietia hybridal,sKjaer et al. 1956a3,4-dimethoxybenzyl 3,4-(CH3O)2C6H5CH2Heliophila longifoliasDaxenbichler et al. 19913,4,5-trimethoxybenzyl 3,4,5-(CH3O)3C6H5CH2Coronopus squamatussDaxenbichler et al. 19912-benzoyloxyethyl C6H5COO(CH2)2Moricandia arvensissDaxenbichler et al. 1991791-methyl-2-benzoyloxyethyl glucobenzosisymbrinSisymbriumaustriacumsDaxenbichler et al. 1991C6H5COOCH2(CH3)CH21-ethyl-2-benzoyloxyethylglucobenzsisaustricinSisymbriumaustriacumsKjaer and Christensen 1962bC6H5COOCH2(CH3CH2)CH23-benzoyloxypropyl C6H5COO(CH2)3glucomalcomiinArabidopsis thalianal,sHogge et al. 1988MalcolmismaritimasKjaer1960not spec.Challenger 19594-benzoyloxybutyl C6H5COO(CH2)4Arabidopsis thalianal,sHogge et al. 1988sEagles et al. 19815-benzoyloxypentyl C6H5COO(CH2)5Arabidopsis thalianal,sHogge et al. 19886-benzoyloxyhexyl C6H5COO(CH2)6Arabidopsis thalianal,sHogge et al. 19882-(L-rhamnopyranosyloxy)benzyl 2-[(OH)3C6H8O]OC6H4CH2Reseda odoratafOlsen and Sorensen 19794-(L-rhamnopyranosyloxy)benzyl 4-[(OH)3C6H8O]OC6H4CH2Thlaspi perfoliatumsAl-Shehbaz, Al-Shammary 19874-(4’-O-acetyl-L-rhamnosyloxy)benzylMoringa peregrinasKjaer et al. 19794-[(4’-CH3CO)(OH)2C6H8O]OC6H4CH22-phenylethyl C6H5(CH2)2gluconasturtiinArmoracia lapathifoliarGrob and Matile 1980rGilbert and Nursten 1972Brassicacampestris(B. rapa)rCarlson et al. 1987Brassica junceal, s, rSang et al. 1984Brassica oleraceaesMinchinton et al. 1982rBirch et al. 1992Brassica napobrassicarShaw,et al. 1989Brassica napussElfakir et al. 1992lKoritsas et al. 1989l, s, rSang et al. 1984rBirch et al. 1992Reseda medias,lOlsen and Sørenson 1980Sinapis alba (B. hirta)rBrown and Morra, unp.3-phenylpropyl C6H5(CH2)3Armoracia lapathifoliarGrob and Matile 1980804-phenylbutyl C6H5(CH2)4Armoracia lapathifoliarGrob and Matile 19802-hydroxy-2-phenylethyl C6H5(OH)CHCH2glucobarbarinBarbaria vulgarissDaxenbichler et al. 1991Ettlinger and Kjaer 19682-(4-methoxyphenyl)-2-hydroxyethylArabis hirsutasDaxenbichler et al. 1991[4-CH3OC6H4](OH)CHCH2sKjaer and Schuster 19722,2-dimethyl-2-(4-methoxyphenyl)ethylPentadiplandra brazzeanarMigirab et al. 1977[4-CH3OC6H4](CH3)2CCH22-(L-arabinopyranosyloxy)-2-phenylethylSesamoides canescenslOlsen et al. 1981C6H5[(OH)3C5H6O]CHCH23-indolylmethyl C8H6NCH2glucobrassicinBrassica campestris (B. rapa)sSosulski and Dabrowski 1984l,rCarlson et al. 1987lHansen et al. 1995Brassica napobrassicarShaw et al. 1989Brassica napussQuinsac et al. 1991lMacfarlane-Smith et al. 1991l, s, rSang et al. 1984rBirch et al. 1992BrassicaoleraceaelHanley et al. 1983rBirch et al. 1992CamelinaalyssumsBäuerle et al. 1986LepidiumgraminifoliumsBäuerle et al. 1986Raphanus sativuslSang et al. 19844-hydroxy-3-indolylmethyl 4-OHC8H5NCH24-hydroxyglucobrassicinBrassica napobrassicarShaw et al. 1989BrassicanapussQuinsac, et al. 1991l, s, rSang etal. 1984rBirch et al. 1992BrassicaoleraceaesShaw et al. 1989rBirch et al. 19921-methoxy-3-indolylmethyl neoglucobrassicinBrassica campestris (B. rapa)sSosulski and Dabrowski 19841-(CH3O)C8H5NCH2Brassica napobrassicarTruscott et al. 1983Brassica napuslMacfarlane-Smith et al. 199181sElfakir et al. 1992rSang et al. 1984rBirch et al. 1992BrassicaoleraceaelHanley et al. 1983rBirch et al. 1992CochleariaofficianalissBäuerle et al. 19864-methoxy-3-indolylmethyl 4-methoxyglucobrassicinBrassica napobrassicarShaw et al. 19894-(CH3O)C8H5NCH2Brassica napuslKoritsas et al. 1989l, rSang et al. 1984rBirch et al. 1992BrassicaoleraceaelShaw et al. 1989rBirch et al. 1992LepidiumsativumsBäuerle et al. 19861-acetyl-3-indolylmethyl 1-(CH3CO)C8H5NCH2Tovaria pendulasSchraudolf and Bauerle 19861-sulphonate-3-indolylmethyl glucobrassicin-1-sulfonateIsatistinctoriasElliott and Stowe 19701-(OSO3)C8H5NCH2as = seed, young seedling, or inflorescence, l = leaf and/or stem, r = root,bidentification changed toE. virgatumRoth (Kjaer and Schuster1973),ctentative identification.82Table 2. Effects of Aqueous Extracts from Glucosinolate-Containing Tissues on Other Plants

________________________________________________________________________

Plant Inhibition Stimulation or no effect

____________________________________________________________________________ Seedling grow th

B r assica cam pest r is barley ( Hordeum vulgare )

v e t c h ( Vic i a sp. )

r a d i s h ( Raphanus sativus )

w h e a t ( Triticum aestivum )

Brassica juncea w h e a t

(seed tissue) tomato ( Lycopersicum

esculentum )

Brassica napus velvetleaf ( Abutilon theophrasti ) wild mustard ( Brassica kaber )

m o r n i n g g l o r y ( Ipom oea

purpurea )

w h e a t

B r assica nigra Brom us rigidus

Brom us m o llis

Avena fatua

w h e a t

Brassica oleracea r a d i s h l e t t u c e

l e t t u c e ( Lact u ca sat i va )

w h e a t

Cam e lina sat i va flax ( Linium usitatissim u m )

Seed germination

Brassica juncea

(seed tissue) alfalfa ( Medicago sativa ) c a r r o t ( Daucus carota )

(seed tissue) cabbage ( Brassica oleracea ) p e p p e r ( Capsicum frutescens )

83(seed tissue) radish timothy ( Phleum pratense )

(seed tissue) turnip ( B r assica rapa ) t o m a t o

Brassica oleracea w h e a t

___________________________________________________________________________ References: Mason-Sedun and Jessop 1988; Ji m e nez-Orsornio and Gleissm an 1987; MasonSedun et al. 1986; Leather 1983; Bell and Muller 1973; Kutacek 1964; Gressel and Holm

1964; Patrick et al. 1963; Grum m e r and Beyer 1960.

84Table 3. Influence of Naturally Occurring Isothiocy a nates on Microorganisms

________________________________________________________________________
Organism a Concentration Ev aluation criterion

μg mL -1 [M]

____________________________________________________________________________
Methy l ITC

_______________
P. petersonii 20 27x10 -5 inhib. growth & prod.

100 68x10 -5 biomass reduced 93%

P. sherm anii 10 14x10 -5 inhib. growth & prod.

100 68x10 -5 biomass reduced 90%

S. aureus 11 15x10 -5 minimum for inhibition

A. eutiches 0.1 14x10 -7 no zoospore formation

A. niger 23 31x10 -5 50% growth inhibition 4d

50 68x10 -5 100% growth inhibition 4d

110 15x10 -4 100% growth inhibition >14d

C. circinans 40 55x10 -5 100% growth inhibition

G. gram inis 0.1 10x10 -7 minimum for inhibition in hs b

M. sitophila <7 <10x10 -5 100% growth inhibition

P. glaucum 2 27x10 -6 minimum for inhibition

P

. cyclopium 14 19x10 -5 50% growth inhibition 6d

37 51x10 -5 100% growth inhibition 6d

110 15x10 -4 100% growth inhibition >14d

R. oryzae 11 15x10 -5 50% growth inhibition 6d

17 23x10 -5 100% growth inhibition 6d

28 38x10 -5 100% growth inhibition >14d

85Ethy l ITC

_______________
A. niger 17 20x10 -5 50% growth inhibition 4d

122 14x10 -4 100% growth inhibition 4d

C. circinans 40 46x10 -5 no effect

P

. cyclopium >261 >30x10 -4 100% growth inhibition 14d

R. oryzae 261 30x10 -4 100% growth inhibition >14d

Ally l ITC

________________

B. subtilis IFO-13722 0.11 11x10 -7 100% growth inhibition b

B. cereus IFO-13494 0.09 91x10 -8 100% growth inhibition b

E. coli 36 36x10 -5 delay growth 36 h c

E. c o li JCM-1649 0.034 34x10 -8 100% growth inhibition b

P. aeruginosa 27 27x10 -5 delay growth 45 h

P. fragi 16 16x10 -5 delay growth 40 h

P. vulgaris 36 36x10 -5 delay growth 60 h

S. enteritidis JCM-1891 0.11 11x10 -7 100% growth inhibition b

S. aureus 1 10x10 -6 minimum for inhibition

36 36x10 -5 delay growth 33 h

0.11 11x10 -7 100% growth inhbition b

A. alliaceus 60 61x10 -5 biomass reduced 98%

5 51x10 -6 100% growth inhibition

A. eutiches 0.1 10x10 -7 no zoospore formation

A. niger 6 60x10 -6 50% growth inhibition 4d

65 66x10 -5 100% growth inhibition 4d

5 51x10 -6 100% growth inhibition

60 61x10 -5 biomass reduced 60%

B. allii 40 40x10 -5 100% growth inhibition

86B. cinerea 600 61x10 -4 minimum for inhibition of germ.

C. circinans 5 51x10 -6 100% growth inhibition

Cytospora sp. >99 >10x10 -4 100% inhibition of germ

Fusarium sp. >99 >10x10 -4 100% growth inhibition

F. oxysporum NFRI-1011 0.022 22x10 -8 100% growth inhibition b

F. solani IFO-9425 0.034 34x10 -8 100% growth inhibition b

G. gram inis 0.1 10x10 -7 min. for inhibition in hs

G. saubinetti 15 15x10 -5 100% growth inhibition

1 10x10 -6 100% growth inhibition

M. laxa 300 30x10 -4 min. for inhibition of germ.

M. sitophila <10 <10x10 -5 100% growth inhibition

M. piriform is 150 15x10 -4 minimum for inhibition of germ.

P. chrysogenum IFO-6223 0.062 63x10 -8 100% growth inhibiton b

P

. cyclopium 9 90x10 -6 50% growth inhibition 6d

22 22x10 -5 100% growth inhibition 6d

151 15x10 -4 100% growth inhibition >14d

P. expansum 600 61x10 -4 minimum for inhibition of germ.

P. gluacum 9 91x10 -6 minimum for inhibition

R. oryzae 10 10x10 -5 50% growth inhibition 6d

74 75x10 -5 100% growth inhibition 6d

R. stolonifer 600 61x10 -4 minimum for inhibition of germ.

5-Methy l thiopenty l ITC

_____________________

S. aureus 5 29x10 -6 minimum for inhibition

A. niger 17 10x10 -5 50% growth inhibition 4d

35 20x10 -5 100% growth inhibition 4d

66 38x10 -5 100% growth inhibition >14d

87P

. cyclopium 23 13x10 -5 50% growth inhibition 6d

33 19x10 -5 100% growth inhibition 6d

66 38x10 -5 100% growth inhibition >14d

P. gluacum 31 18x10 -5 minimum for inhibition

4-Methy l sulfiny l -3-buteny l ITC

____________________________

B. cinerea 230 13x10 -4 minimum for inhibition of germ.

M. laxa 20 11x10 -5 minimum for inhibition of germ.

M. piriform is 460 26x10 -4 minimum for inhibition of germ.

P. expansum 930 53x10 -4 minimum for inhibition of germ.

R. stolonifer 290 17x10 -4 minimum for inhibition of germ.

Benzy l ITC

______________

E. coli 3 21x10 -6 minimum for inhibition

P. vulgaris 6 42x10 -6 minimum for inhibition

S. lutealis 1.5 10x10 -6 minimum for inhibition

S. aureus 2 13x10 -6 minimum for inhibition

A. eutiches 1 67x10 -7 no zoospore formation

A. niger 2 16x10 -6 50% growth inhibition 4d

27 18x10 -5 100% growth inhibition 4d

57 38x10 -5 100% growth inhibition >14d

Cytospora sp. <75 <50x10 -5 100% growth inhibition

Fusarium s p

. < 1 .5 < 10x10 -6 100% growth inhibition

M. sitophila <1.5 <10x10 -6 100% growth inhibition

P

. cyclopium 6 40x10 -6 50% growth inhibition 6d

30 20x10 -5 100% growth inhibition 6d

57 38x10 -5 100% growth inhibition >14d

P. gluacum 2 13x10 -6 minimum for inhibition

88P. palm i vora 1490 10x10 -3 lethal

R. oryzae 67 45x10 -5 50% growth inhibition 6d

194 13x10 -4 100% growth inhibition 6d

Pheny l ethy l ITC

_______________

S. aureus 4 25x10 -6 minimum for inhibition

A. alliaceus 20 12x10 -5 100% growth inhibition

A. eutiches 1 61x10 -7 no zoospore formation

A. niger 2.6 16x10 -6 50% growth inhibition 4d

3.5 21x10 -6 100% growth inhibition 4d

12 75x10 -6 100% growth inhibition >14d

3 18x10 -6 100% growth inhibition

C. circinans 5 31x10 -6 100% growth inhibition

Fusarium sp. 20 12x10 -5 100% growth inhibition

G. gram inis 0.2 10x10 -7 minimum for inhibition in hs

G. saubinetii 2 12x10 -6 100% growth inhibition

P

. cyclopium 8 48x10 -6 50% growth inhibition 6d

12 76x10 -6 100% growth inhibition 6d

P. glaucum 5 31x10 -6 minimum for inhibition

R. oryzae >122 >75x10 -5 50% growth inhibition 6d

4-Hy droxy b enzy l ITC

___________________

B. cinerea 450 27x10 -4 minimum for inhibition of germ.

M. laxa 90 55x10 -5 minimum for inhibition of germ.

M. piriform is 220 13x10 -4 minimum for inhibition of germ.

P. expansum 90 55x10 -5 minimum for inhibition of germ.

P. glaucum 2000 12x10 -3 minimum for inhibition

R. stolonifer 1800 11x10 -3 minimum for inhibition of germ.

89____________________________________________________________________________ a Bacteria: Fungi:

Bacillus subtilis Aphanom yces eutiches

Bacillus cereus Aspergillus alliaceus

Escherichia coli Aspergillus niger

Propionibacterium petersonii Botrytis allii

Propionibacterium sherm anii Botrytis cinerea

Proteus vulgaris Colletotrichum circinans

Pseudom onas fragi Cytospora

Pseudom onas aeruginosa Fusarium

Salm onella enteritidis Fusarium oxysporum

Sarcina lutealis Fusarium solani

Staphylococcus aureus Gaeum annom yces gram inis

G i

b b e r e l

l a s a u b i n e t i i

M o n i l a s i t o p h i l a

M o n i l i n i

a l a x a

M u c o r piriform i s

P e n i c i l l i u m chrysogenum

P e n i c i l l i u m cyclopium

P e n i c i l l i u m expansum

Penicillium glaucum

P h y t o p h t h o r a palm i v o r

a

R h i z o p u s o r y

z a e

R h i z o p u s s t o l o n i f e r

b headspace concentration

c as compared to control

90References: Angus et al. 1994; Iori et al. 1993; Isshiki et al. 1992; Kanemaru and Miyamoto 1990;

Fenwick et al. 1983; Tang and Takenaka 1983; Rutk owski et al. 1972; Lewis and Papavizas 1971;

Drobnica et al. 1967a; Virtanen 1965; Davis 1964; McKay et al. 1959; Hooker et al. 1943; Walker, et

al. 1937.

91Table 4. EPA Registrations for Various Isothiocy a nates

EPA Registration Number Ty pe Isothiocy a nate

4 9 9

- 1 3 6 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
4 9 9

- 1 3 7 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
4 9 9

- 1 3 8 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
7 3 1

- 1 5 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
7 3 1

- 2 2 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
2139-55 Fungicide, insecticide, miticide methyl
2 1 3 9

- 7 6 F u n g i c i d e

, insecticide, miticide, nematicide,
herbicide methyl

3 0 0 8

- 7 7 f u n g i c i d e m e t h y l
5332-7 repellent vertebrates, herbicide propenyl
5 3 3 2

- 8 r e p e l l e n t

, h e r b i c i d e p r o p e n y l
61966-4 insecticide, miticide, repellent propenyl
61966-5 insecticide, miticide, repellent propenyl
6 1 9 6 6

- 7 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
6 7 0 6 4

- 3 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
70160-1 insecticide, miticide, repellent propenyl
7 0 1 6 0

- 2 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
7 0 1 6 0

- 3 r e p e l l e n t

- v e r t e b r a t e s p r o p e n y l
70160-4 insecticide, miticide, repellent propenyl

92

## O S

H

OH H OH

H HO H C N O S O



## R 3 2 

## CH OH



## H

Figure 1. Generalized structure of glucosinolates

93NO O S

C R

S 3 - D G l u c o s e R N C S

N O H

S R

SC N Fe 2 +

lo w p H R C N + S

R S C N

CH 2 CH ( C H 2 ) N C S

n E p i t h i onit r il e Ni t r il e

T h i o c y anat e I s ot hi oc y anat e

O x az oli d i net hi one

T e r m i nal

uns at ur at i o n 2- Hy dr ox y p- Hy dr ox y benz y l I ndol e

Figure 2. Glucosinolate degradation pathw a y 1

1 Enzymatic hydrolysis results in pr oduction of glucose followed by HSO 4 - . At least seven other products have been observed. Oxazolidinethione and SCN - are thought to be formed from unstable R-N=C=S intermediates.

94S i nap i s arv e n s i s

n= 9
SS S i nap is alba

n= 12
DD B. n i gr

a

n= 8
BB

B

. j unc ea

n= 18
DD B

. c a rinat a

n= 17
BB C C

B

. o l e r ac ea

n=9
CC B

. napu s

n= 19
AA C C B

. rapa

n= 10
AA

Figure 3. Relationship betw een Brassica species as described by U (1935)

95F1146-E(12/2004) REPORT DOCUMENTATION PAGE Form Approved OMB No. 0704-0188

The public reporting burden for this collection of information is estimated to average 1 hour per response, including the time for reviewing instructions, searching existing data sources, gathering and maintaining the data needed, and completing and reviewing the collection of information. Send comments regarding this burden estimate or any other aspect of this collection of information, including suggestions for reducing the burden, to Department of Defense, Executive Services and Comm unications Directorate (0704-0188). Respondents should be aware that notwithstanding any other provision of law, no person shall be subject to any penalty for failing to compl y with a collection of information if it does not display a currently valid OMB control number. PLEASE DO NOT RETURN YOUR FORM TO THE ABOVE ORGANIZATION.

1. REPORT DATE (DD-MM-YYYY) July 2005 2. REPORT TYPE Subcontract Report 3. DATES COVERED (From - To) 2000 - 2002

5a. CONTRACT NUMBER DE-AC36-99-GO10337

5b. GRANT NUMBER 4. TITLE AND SUBTITLE Glucosinolate-Containing Seed Meal as a Soil Amendment to Control Plant Pests: 2000-2002

5c. PROGRAM ELEMENT NUMBER

5d. PROJECT NUMBER NREL/SR-510-35254

5e. TASK NUMBER FC059400 6. AUTHOR(S) K. Shaine Tyson, J. Brown, M. J. Morra

5f. WORK UNIT NUMBER

7. PERFORMING ORGANIZATION NAME(S) AND ADDRESS(ES) University of Idaho Moscow, Idaho 8. PERFORMING ORGANIZATION REPORT NUMBER XC0-9-29095-01

10. SPONSOR/MONITOR'S ACRONYM(S) NREL 9. SPONSORING/MONITORING AGENCY NAME(S) AND ADDRESS(ES) National Renewable Energy Laboratory 1617 Cole Blvd. Golden, CO 80401-3393 11. SPONSORING/MONITORING AGENCY REPORT NUMBER NREL/SR-510-35254

12. DISTRIBUTION AVAILABILITY STATEMENT National Technical Information Service U.S. Department of Commerce 5285 Port Royal Road Springfield, VA 22161

13. SUPPLEMENTARY NOTES NREL Technical Monitor: K. Shaine Tyson

14. ABSTRACT (Maximum 200 Words) Plants may produce compounds that direct ly or indirectly affect their biological environment. These compounds fall within a broad category of compounds called allelochem icals, and are exclusive of food that influences growth, health, or behavior of other organisms (Whittaker and Feeney 1971). One reason for interest in allelochemicals is their potential for use in alternative pe st management systems. Using plant-pro duced allelochemicals in agricultural and horticultural practices could minimize synthetic pestic ide use, reduce the associat ed potential for environmental contamination, and contribute to a more sustainable agricultural system.

15. SUBJECT TERMS glucosinolate; allelochemicals; pest management; pesticide

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a. REPORT Unclassified b. ABSTRACT Unclassified c. THIS PAGE Unclassified 17. LIMITATION OF ABSTRACT

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