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NREL is operated by Midwest Research Institute ● Battelle Contract No. DE-AC36-99-GO10337 

Glucosinolate-Containing Seed 

Meal as a Soil Amendment to 

Control Plant Pests 

2000-2002 

J. Brown, M.J. Morra 

University of Idaho 

Moscow, Idaho 

Subcontract Report 

NREL/SR-510-35254 

July 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 Monitor: K. Shaine Tyson 

Prepared under Subcontract No. XC0-929095-01

Subcontract Report 

NREL/SR-510-35254 

July 2005 

National Renewable Energy Laboratory

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Operated for the U.S. Department of Energy

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by Midwest Research Institute • Battelle 

Contract No. DE-AC36-99-GO10337
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1

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. Herbicidal Activity ........................................................................................................................................8

B. Fungicidal and Bactericidal Activities.........................................................................................................12

C. Allelochemical Effects on Insects and Other Invertebrates ........................................................................17

IV. Management Considerations........................................................................................ 19

A. Product Formation......................................................................................................................................19

B. Product Activity and Fate ...........................................................................................................................20

1. Lifetimes..................................................................................................................................................20

2. Volatilization ...........................................................................................................................................21

3. Organic Matter ........................................................................................................................................21

4. Temperature.............................................................................................................................................22

5. Water .......................................................................................................................................................22

6. Soil Texture .............................................................................................................................................23

7. pH............................................................................................................................................................23

8. Microbial Activity ...................................................................................................................................24

C. Additional Allelochemicals ........................................................................................................................25

D. Glucosinolate Concentrations.....................................................................................................................26

E. Timing.........................................................................................................................................................27

F. Additional Benefits .....................................................................................................................................28

V. Crushing and Extraction Technology ........................................................................... 29

VI. Allergenic Reactions...................................................................................................... 30

VII. Registration .................................................................................................................. 33

VII. Genetics and Interspecific Hybridization.................................................................. 34

A. Taxonomy, Genomic Constitutions, and Relationships..............................................................................34

B. Oil and Seed Meal Characteristics and Inheritance ....................................................................................34

C. Interspecific and Intergeneric Crosses in Brassicaceae ..............................................................................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 Occurring 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. Enzymatic hydrolysis results in production of glucose 

followed by HSO4

-. At least seven other products have been observed. Oxazolidinethione and 

SCN- are thought to be formed from unstable R-N=C=S intermediates...........................................94

Figure 3. Relationship between Brassica species as described by U (1935) .........................................95

2
Glucosinolate-Containing Seed Meal as a 

Soil Amendment to Control Plant Pests

I. Introduction 

Plants may produce compounds that directly or indirectly affect their biological environment. 

These compounds fall within a broad category of compounds called allelochemicals, 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 pest management systems. Using plant-produced allelochemicals in agricultural 

and horticultural practices could minimize synthetic pesticide use, reduce the associated 

potential for environmental contamination, and contribute to a more sustainable agricultural 

system. 

Glucosinolates, compounds that occur in agronomically important crops, may represent a 

viable source of allelochemic control for various soil-borne plant pests. Extensive studies of 

several hundred species of the Brassicaceae (Cruciferae) family indicate that virtually all 

may be capable of producing glucosinolates (Kjaer 1976). Insecticidal, nematicidal, 

fungicidal, and phytotoxic effects are often associated with tissues of cruciferous plants. As 

such, glucosinolates are commonly considered to be ultimately responsible for pest 

suppression. 

In this review we consider what is known about glucosinolates and the factors involved in 

potential application of glucosinolate-containing tissues in pest control strategies. We will 

focus on the use of seed meal, 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 formation of a variety of potential allelochemicals. Much of 

the past emphasis and available literature have focused on isothiocyanates (ITCs), apparently 

the most important of those allelochemicals. Any delivery system designed to take 

advantage of this chemistry must consider the fact that glucosinolates must be enzymatically 

hydrolyzed and that multiple pesticidal compounds may be produced. As such, we will first 

provide an overview of glucosinolate chemistry and the relevant hydrolytic reactions. The 

most recent and comprehensive reviews concerning 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 forms. For example, phydroxybenzyl glucosinolate is isolated as a salt complex with sinapine; an organic cation 

derived from choline (Challenger 1959; Kjaer 1960; Van Etten and Tookey 1979). Features 

common to the class are a ß-D-thioglucose moiety, a sulfate attached through a C=N bond 

(sulfonated oxime), and a side group (designated 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 among families of the order Capparales: Tovariaceae, 

Resedaceae, Capparaceae, Moringaceae, and Brassicaceae. Families outside the order 

exhibit occasional occurrence and include the Caricaceae, Euphorbiaceae, Gyrotemonaceae, 

Limnathaceae, Salvadoraceae, and Tropaeolaceae families (Fenwick et al. 1983). A recent 

literature review provides a comprehensive list of all species known to contain glucosinolates 

(Fahey et al. 2001). 

Glucosinolate types in plant species are highly variable. For example, the main glucosinolate 

in radish seed (Raphanus sativus) is 4-methylsulphinyl-3-butenyl glucosinolate, while 

mustard seed (Brassica juncea) is dominated by propenyl glucosinolate. Cabbage seed 

(Brassica oleracea) contains mainly propenyl and 2-hydroxy-3-butenyl glucosinolate; 

rapeseed (Brassica napus) contains 4 major glucosinolates: 2-hydroxy-3-butenyl, 3-butenyl, 

4-pentenyl, and 2-hydroxy-4-pentenyl (Sang et al. 1984). Similar differences in 

glucosinolate types are observed when comparing vegetative plant parts (Hill et al. 1987; 

Josefsson 1967b). 

Numerous 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-Smith and Griffiths 1988; Paxman and Hill 1974). 

Additional environmental factors such as spacing, moisture regime, and nutrient availability 

affect concentration (Freeman and Mossadeghi 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 major 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). 

4
C. Hydrolysis Reactions 

Toxicity is not attributed to intact glucosinolates, but instead to biologically active products 

such as ITCs, organic cyanides, oxazolidinethiones (OZTs), and ionic thiocyanate (SCN-

) 

released upon enzymatic degradation by myrosinase (thioglucoside glucohydrolase, EC 

3.2.3.1) in the presence of water. Degradation also occurs thermally or by acid hydrolysis 

(Kjaer 1976; MacLeod et al. 1981). 

Myrosinase is not properly identified as a single enzyme, but rather as a family or group of 

similar-acting enzymes. Multiple forms of the enzymes exist, both among species and within 

a single plant (Bones and Slupphaug 1989; Falk et al. 1992, 1995a; Lenman et al. 1990, 

1993a; Xue et al. 1993), and all perform a similar function (Björkman 1976). Although their 

genetic sequences are similar to other ß-glycosidases (Lenman et al. 1993b), myrosinases are 

fairly specific toward glucosinolates (Durham and Poulton 1990). These enzymes cleave the 

sulfur-glucose bond regardless of either the enzyme or substrate source. However, the 

particular enzyme and glucosinolate substrate influence reaction kinetics (Bones 1990; 

MacLeod and Rossiter 1986). In addition to plants, the enzymes 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 arrangement has not been easy to determine. Evidence suggesting 

myrosinase is a cytosolic enzyme associated with membranes, perhaps surrounding a vacuole 

containing glucosinolates (Lüthy and Matile 1984; Thangstad et al. 1990), has been 

supplanted by that obtained using more precise methodologies. Glucosinolates are probably 

contained in vacuoles of various types of cells (Grob and Matile 1979; Pocock et al. 1987). 

In contrast, myrosinase is contained only within structures, called myrosin grains, of 

specialized myrosin cells that are distributed among other cells of the plant tissue (Höglund 

et al. 1992; Thangstad et al. 1991). As glucosinolate vacuoles do not appear to be present 

within myrosin cells, intercellular rather than intracellular separation occurs. Disrupting 

cellular tissues allows glucosinolates and myrosinase to mix, resulting in the rapid release of 

glucosinolate degradation products. Myrosinase activity and glucosinolates are preserved in 

cold-pressed meal and are no longer physically separated. Thus, adding water immediately 

results in the production of the hydrolysis products, including isothiocyanate, without the 

need for additional tissue maceration. 

Enzymatic degradation is influenced by a number 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 Tookey 1979; Van Etten and Wolff 1973; Van 

Etten et al. 1969). In the presence of moisture, glucosinolate hydrolysis produces glucose, 

sulfate, and an additional aglucon product(s) from the remaining N, C, S, and R groups 

(Figure 2). The latter product is proposed to go through at least one unstable intermediate, 

the ultimate form controlled by reaction conditions including pH, R-group, and the presence 

of metal ions such as Fe+2. In addition, smaller proteins or coenzymes accompany 

5
myrosinase and may alter the product profile (Falk et al. 1995b; Taipalensuu et al. 1996). 

Several possible products that exhibit some form of biological activity may be formed (Fig. 

2). The products are all interrelated in that they are isomers or result from further fission of 

the isomers. 

D. Hydrolysis Products 

ITCs have historically been considered the “normal” products of glucosinolate breakdown. 

They are often volatile with pungent flavors or odors. The presence of propenyl ITC in 

mustards and horseradish is responsible for much of the flavor; thus, ITCs are sometimes 

called mustard oils. Formation requires that the initial unstable aglucon intermediate 

undergo a Loessen rearrangement to the R-N=C=S configuration. Isothiocyanates are quite 

reactive, although less so than the related isocyanates (R-N=C=O). A few commercially 

available soil fumigants depend on the activity of methyl ITC either as the parent compound 

or as produced from precursors such as sodium N-methyldithiocarbamate or tetrahydro-3,5-

dimethyl-2H-1,3,5-thiadiazine-2-thione. Because of known toxicities, ITCs are often 

considered likely candidates for pesticidal activity. 

The ability of ITCs to form cyclic thione compounds (Hughes 1975) is believed to occur in a 

spontaneous intramolecular reaction when the ITC contains a ß hydroxyl group on the side 

chain (R-group) (Fig. 2). The goitrogenic OZTs so formed are major products from rapeseed 

(B. napus). 

Nitrile character is common to four additional products. Forming a nitrile (R-C≡N, also 

known as an organic cyanide), which does not require rearrangement, involves sulfur loss 

from the molecule. Nitrile formation is favored over ITC at low pH, but occurs in some

crucifers at a pH where ITC is normally the dominant product (Gil and MacLeod 1980). The 

presence of Fe+2 or thiol compounds increases the likelihood of nitrile formation (Hasapis 

and MacLeod 1982; Uda et al. 1986a, 1986b). Epithionitrile formation requires the same

conditions as for nitriles, plus terminal unsaturation of the R-group and the presence of an 

epithiospecifier protein. The epithiospecifier protein possesses a rare property in that it is an 

enzyme cofactor that allosterically directs an enzyme to yield a different product (MacLeod 

and Rossiter 1985; Petroski and Kwolek 1985; Petroski and Tookey 1982). Thiocyanate (RS-C≡N) is sometimes produced, particularly in members of the Alyssum, Coronopus, 

Lepidium, and Thlaspi families (Fenwick et al. 1983). Factors controlling organic 

thiocyanate formation are not well understood (Larsen 1981). 

SCN-

 production from glucosinolates is controlled by the presence of specific R-groups. 

Evidence suggests the anion is a resonance hybrid with greater charge on the S (Hughes 

1975; Witczak 1986); however, charge can be localized on either the sulfur (-S-C≡N) or the 

nitrogen (S=C=N-), depending on the environment (Beekhuis 1975). Indole and 4-

hydroxybenzyl glucosinolates yield SCN- thought to arise from a highly unstable ITC 

intermediate. SCN- is formed from indole glucosinolates over a wide pH range, whereas 4-

hydroxybenzyl glucosinolates is typically thought to yield SCN- only at a more basic pH. 

We have recently determined that 4-OH benzyl isothiocyanate was not stable even at pH 

6
values 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 min at pH 7.0. 

Indolylic compounds also result during SCN-

 production from indole glucosinolate 

hydrolysis. 3-Indolylmethanol, the immediate product, may form ascorbigen or 3,3’-

diindolylmethane (McDanell et al. 1988). 3-Indolylacetonitrile, indole acetic acid, and 

dioxindoles may be formed under conditions favoring nitrile formation (McDanell et al. 

1988; Monde et al. 1991). Indolylic compounds in these plant tissues are also formed by 

routes other than glucosinolate degradation (Monde et al. 1991; Takasugi et al. 1988). 

7
III. Pesticidal Activity of Glucosinolate Hyrolysis Products 

A. Herbicidal Activity

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

communities. Various negative or inhibitory effects can occur. At one time 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 legumes (such as clover) and, 

to a lesser extent, grass are reduced following a Brassica crop (Campbell 1959; Kutácek 

1964). Campbell (1959) showed that pieces of B. oleracea roots placed in a petri dish with 

seeds of clover (Trifolium repens) inhibited germination, suggesting direct involvement of 

compounds from the Brassica plants. Flax yields decrease if species of Camelina are present 

as weeds (Grümmer and Beyer 1960). Brassica herbage reduced stand establishment of five 

crop species more than twice as much as barley (Hordeum vulgare) amendments (Vera et al. 

1987). Similarly, canola stubble was more inhibitory to wild oat (Avena sterilis) emergence 

than wheat or pea (Jones 1992). Stands of wild Brassica nigra appeared to inhibit 

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

amended to soil reduced germination and growth of lettuce for 10-21 d (Patrick et al. 1963). 

Researchers in Sweden demonstrated that Sinapis alba seed meal applid to the soil 

suppresses weed emergence (Ascard and Jonasson 1991; Johansson 1992; Johansson and 

Ascard 1994; Oleszek et al. 1994). For example, mustard meal or “cake,” applied in rows 

between cole crops with the intent of attracting enemies of the cabbage root fly also reduced 

numbers of several annual weeds (Ascard and Jonasson 1991). 

Results with respect to plant inhibition are not always clear, however, and sometimes appear 

inconsistent or contradictory. For example, heavy rape residues (Brassica campestris L.) 

reduced wheat, barley, and oat growth in a field trial, but soil N concentrations measured 

under heavy residues were less than under light residues. Differences in soil nutrient status 

were thus possibly responsible for the effect (Horricks 1969). Similar studies using rape 

straw and chaff residue at 8,970 kg ha-1 showed reduced emergence of bromegrass and 

alfalfa, and reduced growth of barley, bromegrass, and alfalfa, but reductions were not 

significantly different than in controls using wheat straw (Waddington 1978; Waddington 

and Bowren 1978). The effect was ameliorated by applying inorganic nitrogen fertilizer; 

however, subsequent studies indicate that nitrogen limitations 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 

may control the level of phytotoxicity (Mason-Sedun et al. 1986), although differences are 

not always dramatic (Vera et al. 1987) and may be influenced by the plant growth stage. 

Fresher plant tissues are more effective inhibitors than straw or stubble that has been stored 

or left after harvest. Aqueous extracts of fresh wild mustard (B. campestris) tissues produce 

a stronger inhibition of growth than those from aged tissues (Jiménez-Orsornio and 

Gliessman 1987). Green Brassica herbage reduced stand establishment more than Brassica

straw and chaff, which caused only sporadic reductions (Vera et al. 1987). Rape stubble 

8
residues were less or no more effective than pea or wheat residues in inhibiting wheat 

germination 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 may produce compounds 

apart from glucosinolates that are associated with plant inhibition or stimulation, further 

complicating data interpretation (Harper and Lynch 1982; Lovett and Duffield 1981; Lovett 

and Jackson 1980). The target plant species also vary in tolerance to the allelochemicals 

(Kasting et al. 1974; Oleszek 1987). Tomatoes planted in cabbage-amended soil seem

particularly sensitive, wilting within 24 h (Ramirez-Villapudua and Munnecke 1988). 

Suppression of germination may in part be related to seed size, with smaller seeds being 

more susceptible (Vera et al. 1987). 

Many studies concerning the effects of glucosinolate-containing plant tissues on plant growth 

have focused on water-soluble compounds as the responsible agents. Aqueous extracts of 

various Brassica and Camelina reduce seedling growth or inhibit germination (Table 2). 

Negative effects are not consistent in all species, and some plants show no change or are 

actually stimulated. There are not always correlations between germination inhibition and 

growth inhibition. Aqueous extracts had little effect on tomato germination, but severely 

inhibited growth just after germination (Gressel and Holm 1964). Depending on plant part, 

growth stage, or other conditions, extracts from the same plant species may be inhibitory or 

stimulatory to the target species (Jiménez-Orsornio and Gliessman 1987). 

Intact glucosinolates are water soluble, but of several that have been tested for biological 

effects, only 3-indolylmethyl glucosinolate has produced inhibition of plant growth (Bialy et 

al. 1990; Kutacek 1964). High concentrations of 3-indolylmethyl glucosinolate in woad may 

be responsible for its inhibition of other plants, possibly by converting some 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 germination and growth of clover, but only at relatively high 

concentrations (10-2-10-3 M) (Kutacek 1964). 

The water-soluble glucosinolate degradation product most studied in relation to plant growth 

and herbicidal potential is SCN-

. The herbicidal formulation amitrole-T consists of a mixture 

of amino-1,2,4-triazole and NH4SCN. Thiocyanate salts such as KSCN and NH4SCN 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 sumac, and poison oak is at least as favorable as control by 2,4-D or sodium arsenite. 

At 897 kg ha-1 SCN- is a temporary soil sterilant (Beekhuis 1975). SCN-

 is produced by B. 

napus and S. alba meals and thus it is of possible importance in pest control. SCN-

 is not 

expected to be a signficant product from B. juncea meal. 

At rates closer to what might be encountered in natural situations (500 μM SCN-

), percent 

germination of 39 different species and cultivars did not differ from controls, but growth of

22 species was adversely affected (Stiehl and Bible 1989). Early growth inhibition of some

plants was also observed in bioassays by Park et al. (1983), while in hydroponic cultures Ju 

9
et al. (1983) found complete kill of tobacco at < 5 μg mL-1 and of beans at concentrations 

over 25 μg mL-1. Cabbage growth was severely inhibited, though not completely killed, at 

100 μg mL-1. SCN-

 accumulated in plant tissues and toxicity positively correlated with tissue 

SCN-

 concentrations (Stiehl and Bible 1989; Ju et al. 1983). Squash seedlings had low SCNconcentrations in their tissues, suggesting that some plants may degrade or do not readily 

take up the anion (Stiehl and Bible 1989). Wu and Bassler (1969) found that SCNinactivates cotton chloroplasts by inhibiting O2 release. Ionic thiocyanate-induced chlorosis 

may 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; Dustman and 

Duncan 1940). 

Other water-soluble glucosinolate degradation products likely play important roles in altering 

seed germination and plant growth. Water soluble compounds from B. napus defatted seed 

meal severely inhibited lettuce seed germination and those from B. napus leaf tissue 

completely inhibited germination (Brown and Morra 1995, 1996). In contrast, aqueous 

extracts of root tissue had only a minor influence on germination. Several organic water 

soluble glucosinolate degradation products from the tissues were found by back-extracting 

aqueous extracts of the seed meal with CH2Cl2 and a CaCl2 solution. Water-soluble products 

were particularly high in the seed meal and included 5-vinyl OZT, 5-propenyl OZT, 1-cyano3-butene, 3,4-epithiobutyl CN, 2-hydroxy-3,4-epithiobutyl CN, 4-methylsulfinylbutyl ITC, 

and 5-methylsulfinylpentyl ITC. In addition, SCN-

 is found in water extracts of seed meal 

and is an expected product in other tissues in lesser amounts (Brown and Morra 1991; Brown 

et al. 1991). These water-soluble glucosinolate products are probably involved in the 

inhibition of germination by seed meal tissue. However, inhibition of lettuce seed 

germination by water-soluble compounds from leaf and stem tissues could not be explained 

entirely by the total amount 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 compounds. 

Plant tissue extract phytotoxicity correlates with tissue amendment phytotoxicity, but 

residues are generally more effective inhibitors of seed germination than aqueous extracts 

(Mason-Sedun and Jessop 1988; Mason-Sedun et al. 1986). The greater inhibitory effect of 

residues suggests that something other than water-soluble toxins influences germination. 

Volatile ITCs are probably the major allelochemicals responsible for inhibitory effects on 

plants, although some have disputed the practical significance of these compounds in vivo

(Bell and Muller 1973; Choesin and Boerner 1991). 

Plant toxicity is illustrated by methyl ITC, often studied because of its use as a commercial 

soil fumigant. Methyl ITC is usually generated in situ from a precursor (e.g. 

dithiocarbamate), but may be applied directly (Miller 1988). The fumigant Vapam, which 

releases methyl ITC, was the only one of four soil fumigants found to be toxic to strawberry 

plants when applied as a drench (Eide 1959). Residues of methyl ITC must be degraded 

before planting commences because of phytotoxicity (Sirons 1973). Methyl ITC inhibits 

growth and germination of weeds such as pigweed, dandelion, lambsquarter, bermuda grass, 

10
purslane, crabgrass, quackgrass, spurge, fleabane, barnyard grass, fall panicum, common 

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

Many weed seeds are killed while dormant, a capability distinguishing ITCs from other 

herbicides. Activity in the gas phase for propenyl ITC as well as methyl ITC was 

demonstrated by complete or severe germination inhibition of corn, soybeans, wheat, 

rapeseed, cucumber, 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 mechanisms of germination inhibition by ITCs are not known, 

evidence suggests interference with protein synthesis and possibly processes involved in the 

formation of phosphorylated sugars (Leblová-Svobodová and Koštír 1962). Benzyl ITC is a 

potent inhibitor of plant enzyme activity (Lykkesfeldt and Møller 1993). 

ITCs express variable phytotoxicities, a property related to the remainder of the molecular 

structure. Benzyl ITC inhibits germination and growth of several weeds (Dale 1986; Powell 

and Spencer 1988; Wolf et al. 1984) at a rate lower than methyl ITC (Teasdale and Taylorson 

1986). Two extremes of effectiveness were observed in wheat bioassays at ITC 

concentrations of 500 ppm; butyl ITC had no effect on germination, shoot, or root growth 

while phenylethyl ITC completely inhibited germination (Bialy et al. 1990). Propenyl, 

methyl, ethyl, phenyl, and benzyl ITCs were intermediate with respect to inhibition. 

In addition to studies with pure compounds, organic solvent extracts of papaya shown to 

contain benzyl ITC inhibited velvetleaf, but not corn, at 4x10-4 M (Wolf et al. 1984). 

Similarly, CHCl3 extracts of mustard inhibited seedling growth of barley, rye, broccoli, and 

radish (Jiménez-Orsornio and Gliessman 1987). Barley and rye were most affected; 

inhibition of radish was not statistically significant. Ethyl acetate extracts of Rorippa indica

contained 8-methylsulfinyloctyl, 9-methylsulfinylnonyl, 10-methylsulfinyldecyl, and the 

corresponding sulfonyl ITCs and inhibited lettuce germination by 90% (Yamane et al. 1992). 

Only 10-methylsulfonyldecyl ITC was without activity toward germination and growth. 

Inhibitory effects with pure compounds and organic extracts also occur in studies involving 

volatiles from glucosinolate-containing plant tissues. Volatiles produced from Brassica

tissues were shown to inhibit germination 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 chambers 

containing B. napus seed meal, root, and leaf and stem tissues (Brown and Morra 1995, 

1996). Tissues producing those volatiles inhibited germination in bioassays, whereas no 

inhibition was observed when the same tissues were treated to hydrolyze glucosinolates and 

remove volatile glucosinolate products before bioassays were initiated. Comparison of 

headspace volatiles and extracts from the intact and hydrolyzed tissues revealed that ITCs 

were the most abundant products of intact tissues. The presence of other, nonglucosinolate 

derived volatiles could not explain inhibition since the same compounds were present in the 

headspace of intact and hydrolyzed tissues. Some volatile glucosinolate products have also 

been observed in the headspace of vessels containing B. campestris (1-methylpropyl and 4-

pentenyl ITC), B. juncea (1-methylpropyl and propenyl ITC), B. nigra (propenyl ITC), and S. 

alba (3-butenyl ITC and phenylethyl CN) (Tollsten and Bergström 1988). Propenyl ITC has 

11
been detected in headspace of cabbage (Chin and Lindsay 1993) and methyl thiocyanate as a 

volatile product from broccoli (Hansen et al. 1992). 

The effects of volatile ITCs are short-lived. Benzyl ITC was very effective at inhibiting 

weed germination if the seeds were placed on fresh media; however, if seeds were placed on 

media two days old, very little difference was observed from the control (Dale 1986). In 

seed germination bioassays, 50 % of the methyl ITC disappeared within 1-2 d and the 

remainder was almost completely gone in 5-7 d (Teasdale and Taylorson 1986). 

Many of the overall effects on crop plants can be considered beneficial. Tarahumara Indians 

in northern Mexico claim that there are fewer weeds in plots after mustard is grown 

(Jimenez-Orsornio and Gliessman 1987). Ramirez-Villapudua and Munnecke (1987) noted 

that plots treated with cabbage amendments and solar heating were weed free for months. 

Potato yield was increased and weed biomass was reduced 50%-96 % after rapeseed was 

incorporated into the test plots (Boydston and Hang 1995). Palouse wheat farmers of the 

northwestern United States have claimed higher yields after rapeseed was included in the 

rotation. In Australia, Kirkegaard et al. (1994) measured greater wheat yields following 

Brassica crops compared to other break crops. Other beneficial effects may 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 

germination and seedling growth. To take advantage of allelochemic effects in weed 

suppression, we must consider the mechanisms of plant suppression, characterize the 

compounds and chemistry involved in this suppression, target weed species, and avoid 

negative effects on the crop. We must especially consider the potential negative 

consequences of phytoxicity when targeting pests of horticultural crops. 

B. Fungicidal and Bactericidal Activities 

Glucosinolate products may greatly influence fungal and bacterial populations. Although 

products were not identified, in the presence of myrosinase several alkyl and aryl 

glucosinolates were cytotoxic to Salmonella typhimurium (Tiedink et al. 1991). ITCs are 

among the most potent products and are suspected to be the major inhibitors of microbial 

activity. Benzyl ITC is sometimes used as an antibiotic to treat infections of the respiratory 

and urinary tracts (Mennicke et al. 1988). Bacteriostatic, bactericidal, and fungicidal effects 

of many ITCs are well documented (Table 3). 

In many cases, microorganisms are differentially inhibited by the same ITC. For example, 

although methyl ITC is active against a broad range of organisms, dose responses vary 

tremendously among species. Munnecke et al. (1962) estimated the range of toxicity to 

Rhizoctonia as 110-1500 μg methyl 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 grampositive bacteria to ITCs; some microorganisms avoid toxicity by degrading the inhibitory 

compound (Fenwick et al. 1983; Smelt et al. 1989). 

12
The 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 

more 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 may be more effective in controlling 

fungal pathogens because they contain thiofunctionalized glucosinolates. 

Glucosinolate products with nitrile character are biologically active, but effects are more 

limited than for ITCs. The yeast Nematospora sinecaula is inhibited by butyl thiocyanate at 

40 μg mL-1 (Holley and Jones 1985) and modified thiocyanates are used to control slime 

mold in the paper industry (Beekhuis 1975; Wood 1975). However, tests against various 

microorganisms showed that 2-200 times as much thiocyanate was required to obtain 

equivalent growth inhibition compared to the corresponding ITC (Virtanen 1965; Walker et 

al. 1937). Propenyl CN did not inhibit Peronospora at concentrations > 1000 μg mL-1

(Greenhalgh and Mitchell 1976). A propenyl cyanide concentration approximately 40 times 

that of propenyl ITC was required for equivalent inhibition, but benzyl CN inhibited 

Nematospora at < 50 μg mL-1 (Holley and Jones 1985). Solutions of propenyl glucosinolate 

with myrosinase were more effective at inhibiting Leptosphaeria maculans at pH 7 (where 

ITC should be the dominate product) than at pH 4 (where the proportion of nitrile product is 

increased) (Mithen et al. 1986). The same authors showed mild inhibitory effects of 3-

indolylacetonitrile to L. maculans. 3-Indolylacetonitrile completely inhibited growth of 

Penicillium chrysogenum at 200 μg g-1 (Smissman et al. 1961). 

Similarly, other products inhibit microbial activity 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 some activity against 

Phoma lingam (Schnug and Ceynowa 1990) and Aphanomyces euteiches f. pisi (Smolinska 

et al. 1997a). 3-Indolylmethanol is quite inhibitory to L. maculans, but ascorbigen and 

diindolylmethane are only mildly 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). Ammonium thiocyanate inhibits bacterial 

growth in soil but stimulates fungi at concentrations >250 μg g-1 (Smith et al. 1945). SCNinhibits nitrite forming organisms (Smith et al. 1945) and SCN-

-containing amitrole-T 

inhibits nitrification (Van Schreven et al. 1970). Although the effects of SCN- are limited, 

SCN- often works synergistically with other chemicals. For example, use with certain 

peroxides produces bacteriocidal solutions, although neither is effective alone (Beekhuis 

1975). Similarly, SCN- was more efficient in lysing cells when combined with other anions 

or lysozymes than when any agent was used alone at the same concentrations (Pollack et al. 

1983). 

13
Activity of aqueous plant extracts against pathogens or other organisms has been observed. 

Aqueous extracts of mustard (B. campestris) inhibited Verticillium chlamydosporium (Owino 

et al. 1993) and those of cabbage root inhibited Glomus mosseae spore germination 

(Vierheilig and Ocampo 1990). B. napus seed meal extracts inhibited A. euteiches

(Smolinska et al. 1997a), but slightly enhanced growth of a Propionibacterium (Rutkowski et 

al. 1972). Buffered seed homogenates of Brassica hirta (S. alba) inhibited growth of 

Nematospora by more than 99% (Holley and Jones 1985). Glucosinolates extracted from

Brassica kaber and hydrolyzed in aqueous solution with myrosinase inhibited Glomus 

intraradices (Schreiner and Koide 1993b). 

Suppression appears to be related, at least in part, to the solubility of compounds produced 

by glucosinolates in the tissue. 4-Hydroxybenzyl ITC has greater solubility than many ITCs 

and would be the most prominent 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 (Smolinska et al. 1997a). In contrast, water extracts of 

cabbage tissues did not suppress Aphanomyces root rot of peas (Papavizas 1966). The 

dominant ITC in this tissue has low water solubility. 

Volatiles from tissues or organic solvent extracts are often highly inhibitory toward 

microorganisms. These fractions are also most likely to contain highly active ITCs. Extracts 

of B. juncea (in which propenyl ITC is the most prominent product) made with 70% ethanol 

and added to growth media at 0.025%-0.8% were in several cases more inhibitory to bacteria 

than equivalent amounts of propenyl ITC (Kanemaru and Miyamoto 1990). Smaller amounts 

of other ITCs were also present in the extracts, possibly accounting for the greater toxicity. 

The authors suggest using mustard as a food preservative. Extracts (CHCl3) 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, most likely because relatively nonvolatile 4-hydroxybenzyl ITC is the major 

glucosinolate product in these tissues. As might be expected from its glucosinolate profile, 

B. nigra volatiles were inhibitory toward this organism (Schreiner and Koide 1993b). 

Volatiles from 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 meal completely inhibited A. euteiches

(Smolinska 1997b). In all cases, volatile ITCs were implicated as the responsible agents. 

This effect of volatiles from plant tissues on microorganisms has been termed 

“biofumigation” (Angus et al. 1994; Kirkegaard et al. 1993). 

Studies of pathogen inhibition by glucosinolate-derived allelochemicals have also been 

described for soils. Cabbage vapors passed through soil columns reduced Rhizoctonia solani

contained within that soil (Lewis and Papavizas 1974). Volatiles emanating from cabbageamended soil that were passed through tubes containing Pythium ultimum and Sclerotium 

rolfsii cultures inhibited growth of the cultures, especially when warmed to 38o

C (Gamliel 

and Stapleton 1993). Approximately one-third the incidence of Fusarium infection of oat 

crowns was observed when soils were covered with heavy B. campestris residue compared to 

light residue applications (Horricks 1969). Sealed vessels containing soil amended with leaf 

and stem tissue of nine cruciferous species markedly reduced populations of Fusarium 

14
oxysporum compared to untreated soil and soil amended with alfalfa, wheat straw, chicken 

manure, or steer manure (Ramirez-Villapudua and Munnecke 1988). Inhibition of Fusarium

was diminished or not detectable when cabbage-amended soils were left uncovered, but 

tissue amendment in combination with solarization was more effective than either 

solarization or amendment alone (Ramirez-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, benzyl, or phenyl isothiocyanate was allowed 

to volatilize. Propenyl and ethyl isothiocyanates were the most fungistatic of those 

compounds tested to mycelial growth. Sporulation and conidia viability were not affected by 

any of the isothiocyanates. However, the same concentrations of propenyl and ethyl 

isothiocyanates that only inhibited mycelial growth, completely suppressed conidia and 

chlamydospore germination of all isolates. Other glucosinolates releasing ethyl, benzyl, and 

phenethyl isothiocyanates were also fungitoxic to F. oxysporum conidia and chlamydospores. 

Changes in populations of the pathogen resulting from a green manure crop will only be 

achievable if conidia and chlamydospores are targeted, since decreased populations resulting 

from the direct inhibition of F. oxysporum mycelial growth seem unlikely. We concluded 

that the pathogenic F. oxysporum isolates typically infesting nursery soils in northern Idaho 

will be most suppressed by species of plants containing glucosinolates that release the 

highest concentrations of propenyl isothiocyanate such as Brassica carinata, B. nigra, and B. 

juncea (Smolinska et al., 2003). 

Although the allelochemicals from glucosinolates are inhibitory to many organisms, 

incorporated organic matter may simultaneously stimulate microbial populations by 

providing increased amounts of carbon substrate. Incorporated B. napus residues were less 

stimulatory than corn residues to Fusarium root diseases in a forest nursery (James, personal 

communication 2002), but resulted in more disease than fallow treatments (James et al. 

1996). This is a major obstacle to using meal amendments to control fungal pathogens in 

soil. Formulation technologies or amendment strategies must be developed to ensure that 

pathogen populations are not stimulated. 

One of the more thoroughly studied pest inhibition systems includes the use of cruciferous 

amendments to control Aphanomyces root rot of peas and beans. The results indicate the 

successes and difficulties commonly 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 amendments of several 

cruciferous plants, of which cabbage was the most effective. Controls contained no 

amendments. About 70% of the control plant roots were infected with Aphanomyces; soils 

amended with as little as 0.5% dried cabbage leaves showed approximately a tenfold 

reduction in root infection. The effects were still observed on plantings made 15 weeks after 

amendment. Interestingly, although Aphanomyces root rot was clearly suppressed, some

amended soils actually had increases in Rhizoctonia root rot. 

15
We suspected that volatile components were the responsible agents, so we tried to analyze 

headspace vapors above amended soils using aqueous trapping reagents and GC (Lewis and 

Papavizas 1970). No ITCs were detected in this case, but in contrast to decomposing corn, 

the vapors included CH3SH (thiomethane), (CH3)2S (dimethyl sulfide), and (CH3)2S2

(dimethyl disulfide). Bioassays showed that these vapors were toxic to Aphanomyces, 

though much less so than ITC vapors which in some cases were 1000 times more inhibitory. 

Furthermore, 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 microbial inhibition focused on conditions encountered in the field. In 

containerized plantings both inside and outside greenhouses, amendments of Brassicaceae 

tissues significantly reduced Aphanomyces root rot disease beyond fallow and wheatamended treatments by 32%-38% (Chan and Close 1987). When Brassicaceae crops were 

grown in soil instead of being introduced as amendments, 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 amending with rapeseed tissues resulted in reductions 

more significant than with oats. In other field studies, similar 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 mustard (S. alba) 

green manure but no difference in yield was observed compared to fallow, perhaps because 

of reduced pea emergence in white mustard treated plots (Muehlchen et al. 1990). However, 

continuing a second cycle of peas and white mustard resulted in reduced root rot severity and 

increased yield in spite of a similar reduction in pea emergence. Careful study of the 

variables influencing allelochemical efficacy is needed to consistently and effectively control 

pests. 

B. napus meal has been used for pest control in a limited number 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 meal to soil and compared C. parasiticum

control in peanuts to control by Vapam. The University of Idaho supplied the meal to the 

investigators. All three rapeseed meal rates reduced disease incidence of Cylindrocladium

black rot of peanuts. A maximum 70% reduction in disease incidence (compared to an 

untreated control) occurred in the 3 t ha-1 treatment of peanut cultivar susceptible to the 

disease. Although glucosinolate concentration of the meal was not mentioned, the 

concentration was likely less than 50 μmol g-1 and thus substantially below what is contained 

in mustard meals. 

Attempts have been made to control other fungal pathogens using meal. Dandurand et al. 

(2000) used B. napus cv. “Dwarf Essex” meal in laboratory experiments to control 

Sclerotinia sclerotiorum and A. euteiches. A 100% reduction in carpogenic germination and 

a 33% reduction of myceliogenic germination of S. sclerotiorum occurred in soil amended 

with Dwarf Essex meal. A low glucosinolate B. napus meal (Stonewall) reduced carpogenic 

germination by 44% and had no effect on myceliogenic germination, indicating that 

glucosinolate hydrolysis products were critical to suppression. A. euteiches root rot of pea 

was reduced 77% by Dwarf Essex meal and was unaffected by low glucosinolate Stonewall 

16
meal. In all experiments meal was not homogeneously 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 summary, fungal and bacterial pathogens can likely be controlled by glucosinolatecontaining meals. Previous research was conducted most often with B. napus meal 

containing glucosinolate concentrations less than 50 μmol g-1 tissue. Higher concentration 

glucosinolate meals will be more effective. Experimentation must be conducted to determine 

whether the additional carbon contributed from the meal exerts a growth-promoting effect on 

the fungal pathogens, thereby decreasing the pesticidal efficacy of the meal. Amendment 

techniques or formulations must be optimized to decrease growth promotion of the pathogen. 

C. Allelochemical Effects on Insects and Other Invertebrates 

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

marked effects on insects. Usually extracts are placed on a medium 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 

demonstrated by 50% mortality in 10 min. Macerated root tissue was quite toxic also; 50% 

mortality in 3 h for Drosophila and 50% mortality in 24 h for the common housefly Musca 

domestica. Toxicity was in most cases strongly correlated with phenylethyl ITC content. 

Tissues that produced mortalities but did not possess phenylethyl ITC produce other ITCs. 

Insecticidal activity of several ITCs has been demonstrated, especially for aromatic 

compounds (Åhman 1986; Borek et al. 1995b; Chew 1988a; Lichtenstein et al. 1962, 1964; 

Seo and Tang 1982; Wadleigh and Yu 1988). Borek et al. (1995b) tested the toxicities of six 

commercially available ITCs to eggs of the black vine weevil (Otiorhynchus sulcatus (F.)) 

and concluded that aromatic ITCs were most toxic. Later the same group of investigators 

showed using an expanded set of 12 ITCs and QSAR relationships that ITC polarity was 

most important in explaining observed toxicities (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 numbers 

of carbon atoms or those bearing sulfinyl, thio, or aromatic moieties. 

Whitefringed weevil larvae (Naupactus leucoloma) were used as bioassay organisms to test 

the toxicity of volatile and nonvolatile glucosinolate hydrolysis products (Matthiessen and 

Shackleton 2000). Comparisons of the toxicity of two pure ITCs in the headspace of sealed 

containers showed that methyl ITC vapors were more toxic than phenylethyl ITC across a 

temperature range of 5°-20°C. The toxicity of volatiles from B. juncea meal (assumed to be 

propenyl ITC) were more toxic than those from B. napus meal when equal weights of the 

tissues were compared. Water-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 germinating (Landis and Gould 1988). Methyl ITC is mainly used to control 

17
various fungi that cause damping off and root rot, but is also used to control insects such as 

wireworms and symphilids, and nematodes (Beekhuis 1975; Mullins and Kirkbright 1987; 

Sirons 1973; Toba 1984; Worthing 1987). Propenyl ITC was the most effective of 57 

volatile compounds tested against the wireworms Limonius californicus and Limonius canus, 

with an LC50 of 2.33 μg mL-1 (Lehman 1942). Rapeseed meal amended (3% on a weight 

basis) to soil repelled wireworms (L. californicus), but in uncovered containers did not kill 

them (Brown et al. 1991). Maximum total ITC content of 301 nmol g-1 soil decreased 

rapidly. Subsequent testing using propenyl ITC in the same soil showed that wireworms 

were killed in lower but similar concentrations in capped vials (LC50 range 211-238 nmol g-1

soil) and that propenyl ITC added to soil was more persistent than ITC produced from the 

rapeseed meal-amended soil in the previous test (Williams 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; Rodman 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 impact on mammals 

(Beekhuis 1975; Wood 1975). 3-Indolylacetonitrile, known more for its auxin-like activity 

on plants, also inhibits growth of insects (Smissman et al. 1961). SCN- is ineffective when 

used alone, but is insecticidal in combination with other chemicals (Beekhuis 1975). SCNwas toxic to wireworms only at concentrations much higher than observed in rapeseed mealamended soil (McCaffrey et al. 1995). 

Invertebrate plant pests other than insects are also potential targets. Winkler and Otto (1980) 

found that rotational plantings of rape or mustard in strawberries checked the spread of some

nematodes, particularly Pratylenchus penetrans. Leguminosae, potato, or grass rotations did 

not alter nematode populations, although green manure applications of Leguminosae (peas, 

beans and vetch), rape, or mustard reduced nematode numbers. Similarly, growing rapeseed 

in soil and incorporating the tissue as a green manure significantly reduced populations of 

Meloidogyne chitwoodi compared 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 myrosinase, glucosinolates are toxic to Heterodera schachtii

at concentrations of 0.5-5.0 mg mL-1 (Lazzeri et al. 1993). Some varieties of Brassica crops 

are being offered commercially as “trap crops” for nematodes. Rather than killing the 

nematodes, Brassica plants interfere with nematode reproductive cycles. Biochemical 

mechanisms responsible for nematode resistance and suppression, although not well 

understood (Lazzeri et al. 1993), deserve further attention. 

Rapeseed meal has been used as a soil amendment in an attempt to control nematodes. 

Investigators applied 1, 2, or 3 t ha-1 of B. napus meal to soil and compared control of the

soybean cyst nematode (Heterodera glycines Ichinohe) in soybean to chemical control by 

aldicarb (Bhardwaj et al. 1966). The University of Idaho supplied the meal to the 

investigators. The results were ambiguous, and acceptable control was not achieved. Higher 

glucosinolate concentration meals 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, many factors must be considered when these strategies are implemented if we are 

to take full advantage of this natural pesticidal activity. Release efficiency and release rate 

from plant tissues, susceptibility of the target species, effectiveness of the particular 

glucosinolate degradation product, soil texture, organic matter, pH, moisture content, and 

effectiveness of the products in the aqueous and vapor phases all contribute to pesticide 

efficacy. 

Understanding the processes and fates of the allelochemicals in soil is crucial for maximizing 

potential benefits in pest control and minimizing potential negative environmental impacts. 

Soils and soil/plant interactions may influence which products are formed and their 

effectiveness in the soil environment. Although additional work is necessary in this area, 

enough data have been collected to make some observations and generalizations concerning 

the behavior of several glucosinolate products in soil. 

A. Product Formation 

Conditions affecting product formation during glucosinolate degradation have most often 

been examined in isolated plant tissue systems. 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 formation. When plant tissues are incorporated into 

soil, the soil’s influence on product formation may be linked as much to its moisture content 

as other soil characteristics. Under drier conditions little soil solution is available to 

influence the reaction, and the effects of pH and metal ions on product formation could be 

determined solely by the reaction microenvironment of the plant tissue. However under 

more moist or wet conditions, the soil solution and soil colloids are much more likely to 

participate in reactions that alter product formation. 

Glucosinolate products such as ITCs, SCN-

, OZTs, and to a lesser extent nitriles are 

produced in soil amended 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 most abundant products 

were 2-phenylethyl ITC and benzenepropanitrile. ITCs have also been collected in the 

rhizospheres of a few plants (Tang and Takenaka 1983; Yamane et al. 1992), including B. 

napus (Kirkegaard et al., 2001). 

A model system using propenyl glucosinolate and myrosinase amendments was used to 

investigate product formation in six soils at two moisture contents of field capacity (0.033 

MPa) and saturation. Propenyl ITC was produced as the dominant product regardless of soil 

characteristics (Borek et al. 1994). Only minor amounts of propenyl CN were detected. 

19
Sinigrin and myrosinase added to unbuffered aqueous soil extracts resulted in greater 

proportions of nitrile, and in some cases nitrile became the dominant product. Glucosinolate 

hydrolysis produces H+

, decreasing solution pH as the reaction proceeds. Although this 

enhances nitrile formation in unbuffered extracts, soils provide a large buffering capacity that 

in effect prevents nitrile formation. None of these soils were more than moderately acidic 

(pH of 5.65-8.3), so glucosinolate hydrolysis in wet acidic soils (pH < 5) may result in more 

nitrile formation. Conditions favoring production of ITCs could be advantageous for pest 

control since these are typically the most biologically active products. 

Additionally, product release rates and efficiencies must be considered. Although lower in 

total glucosinolate content, root and shoot tissues of B. napus realeased volatile products 

more efficiently than did meal tissue (Brown and Morra 1995, 1996). The condition of the 

tissue is also important. Fresh tissues may release products more slowly than those 

physically disrupted before amendment by tissue crushing, air drying, freeze drying, or 

grinding. Lower maximum concentrations are thus likely to be produced from fresh tissue, 

but the period of release is potentially extended. Recent investigations indicate that 

maceration of green tissues is indeed necessary to maximize ITC release (Morra and 

Kirkegaard 2002). Such a limitation does not exist when using meal amendments as soil 

additives since the seed crushing procedure has resulted in extensive cellular disruption. 

B. Product Activity and Fate 

1. Lifetimes 

Lifetimes of glucosinolate products in the environment are generally short, an advantage 

when considering environmental impact. ITCs remain in soil for a few days to a few weeks. 

Even in gas-tight containers, half-lives of methyl ITC in several soils were 0.5-50 d at 15o

C 

(Smelt et al. 1989). Concentrations of methyl ITC in greenhouse air, where soil had been 

amended with metham-sodium, were reduced to less than 0.2% of the original amount by the 

third day (Van Den Berg et al. 1992). One week after a major spill of metham-sodium into 

the Sacramento River of California, neither metham-sodium nor the methyl ITC hydrolysis 

product was detectable (del Rosario et al. 1994). In a germination 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 environmental 

consequences is therefore quite small and is illustrated by the fact that no methyl ITC 

residues were detected in more than 200 surface water samples (Hogendoorn et al. 1992). 

However, a short residence time places limits on achieving effective pest control and may 

contribute to the variability observed in the suppression of soil-borne plant pests. 

Lifetimes of allelochemicals produced in situ in plant tissue-amended soil differ from those 

of the respective pure chemicals amended to soil without plant tissue. Ninety percent of the 

maximum ITC in rapeseed meal-amended soil dissipated within 24 h (Brown et al. 1991), 

20
whereas only 60% of propenyl ITC amended in the form of an aqueous phase mixture to the 

same soil was lost in 24 h (Williams 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 dissipation of ITC thus occurs in the presence of 

plant tissue amendments, most likely because of sorption to the additional organic carbon. 

Lifetime 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 meal-amended soil dissipates more slowly, having an extended lifetime 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, demonstrating that 

combinations of glucosinolate products may alter the behavior of individual products in soil. 

In this case, propenyl ITC most likely inhibited microbial activity and the corresponding 

microbial degradation of SCN-

. 

2. Volatilization 

ITC disappears via many routes; volatile losses are one major route. Unsealed bottles had 

much greater losses of methyl ITC than sealed bottles (Ashley et al. 1963). Thirty-four 

percent of the methyl 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 methyl ITC 

have been reported as 13 and 20.7 mm Hg at 20o

C, relatively low values for a fumigant 

(Sirons 1973; Smelt and Leistra 1974); thus, volatile losses may be less of a problem for 

ITCs than for some commercial products. However, volatilization is important for even 

distribution of fumigants in soil. Vapor pressures of various ITCs and nitriles vary widely, 

so large differences in soil fumigation resulting from glucosinolate hydrolysis products are 

expected. 

Loss of ITC into the atmosphere is not desirable, since that amount of product is not 

available for pest inhibition. Volatile losses could also be of concern for health and 

environmental reasons. Photodissociation by sunlight appears to be one rapid and effective 

pathway for removing methyl ITC from the atmosphere, taking about 41 h (Alvarez and 

Moore 1994). This process would likely apply to other ITCs; however, questions remain as 

to the fate and impact of dissociation products (Geddes et al. 1995). 

3. Organic Matter 

Many fumigants are less effective in soil than in nonsoil systems. This is particularly true of 

ITCs. Propenyl ITC was about 193 times more toxic to wireworms than CS2 (carbon 

disulfide) in air, but was only about 29 times more toxic in a silt-loam soil (Lehman 1942). 

Similar results were found with methyl ITC compared to CS2 and other fumigants 

(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). 

21
Sorption of ITCs to soil constituents is an important mechanism that decreases ITC 

effectiveness. Sorption of methyl ITC to soil increases with increasing organic matter 

content (Munnecke and Martin 1964; Smelt and Leistra 1974). As a result, much less methyl 

ITC is extractable from organic muck soils than from mineral soils (Sirons 1973). Although 

Ashley et al. (1963) determined that the disappearance of extractable methyl ITC was 

slowest in a peat soil amended with metham-sodium, amounts of extractable ITC could have 

been controlled by metham-sodium hydrolysis rates instead of ITC sorption. As generally 

observed for methyl ITC, increased propenyl ITC disappearance from soil is correlated with 

greater organic carbon contents and typically greater 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 commonly found in soil organic 

matter such as alcohols, phenols, thiols, carboxylic acids, and amines. These types of 

reactions may also explain reduced efficiency of volatile ITCs released from seed meal tissue 

compared to other plant tissues (Brown and Morra 1996). Amino groups in protein, 

contained in high concentration in rapeseed meal, readily react with ITCs (Björkman 1973; 

Kawakishi and Kaneko 1987) and reduce measured volatile concentrations. 

4. Temperature 

Methyl ITC sorption to soil is not greatly enhanced by increasing the temperature from 4o

 to 

30o C (Matthiessen et al. 1996), although elevated temperatures are correlated with increased 

amounts 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 temperature was also 

associated with a greater inhibitory effect toward pathogenic fungi in a covered cabbageamended soil (Gamliel and Stapleton 1993), probably because of more thorough fumigation. 

Greater concentrations of volatile thiomethane, dimethyl sulfide, and propenyl ITC were 

observed at elevated temperatures. Temperature influences the fate of nonvolatile products 

as well. For example, higher temperature generally increased disappearance rates of SCNin 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 mg mL-1

 (Sirons 1973) and 8.9 mg mL-1 (Smelt and 

Leistra 1974) at 20o

C. Estimates of methyl ITC distribution between the vapor, aqueous, and 

solid phases of soil indicate that most of the ITC is in the aqueous phase under moist 

conditions, with a substantial amount retained in the solid phase (Smelt 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. We do not know whether ITCs 

such as 4-hydroxybenzyl ITC with greater water solubility would have greater pesticidal 

activity than more volatile and less soluble ITCs under wet conditions. Methyl ITC was 

susceptible to leaching with high moisture in sandy soil (Frick et al. 1995), therefore, other 

ITCs may leach in permeable soils subjected to large water inputs. Henry’s constants must 

therefore be determined for the ITCs of interest to better predict partitioning within the threephase S soil environment. 

22
Increased water content increases ITC longevity in soil. More methyl ITC was measured in 

headspace of soils with lower water contents and rates of disappearance were more rapid 

(Turner and Corden 1963). Although 30% of applied methyl ITC was lost by volatilization 

under a drier moisture regime, very little vapor phase methyl ITC was lost during a wet 

regime (Frick et al. 1995). Increased water content increased the half-life of propenyl ITC 

(Borek et al. 1995a). Thus continually wet conditions, especially when combined with cold 

temperatures, could result in increased ITC lifetimes and perhaps increased potential for pest 

inhibition resulting from longer exposure times. This could also extend the risks of residual 

effects on subsequent plantings. 

In contrast to ITC, increased soil moisture 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 controlled by different mechanisms. Propenyl CN is 

sorbed or reacts more quickly at the water phase than at the gas phase, which may indicate 

that the overall reaction rate for nitriles is controlled by the gas/water phase partition 

coefficient. 

6. Soil Texture 

Volatile losses are greatest from coarse textured soils, but texture is less important than 

factors such as organic carbon content. In a comparison of three soils, volatile loss of methyl 

ITC was greatest from a sandy loam (0.67% organic matter, 8% clay); least volatilization 

occurred from a loam with more clay but also greater organic matter (2.03% organic matter, 

25% clay). The finest textured soil, a clay loam (0.55% organic matter, 40% clay), had 

intermediate losses of methyl ITC compared to the other two soils (Munnecke and Martin 

1964). Extractable concentrations of methyl ITC obtained from sandy, clay, and loam soils 

amended with metham-sodium are similar for the first few hundred hours (Ashley et al. 

1963). As such, evidence suggests that sorption to soil clay is not a major mechanism of 

disappearance for ITCs. In contrast, sorption to or degradative catalysis by inorganic soil 

constituents may play a role in SCN-

 disappearance from soil, particularly at higher 

temperatures (Brown and Morra 1993). 

7. pH 

Soil pH influences the formation and disappearance of glucosinolate hydrolysis products, 

although effects on disappearance are not dramatic. Greater amounts of methyl ITC were 

trapped in headspace from soils adjusted to a higher than to a lower pH (Munnecke and 

Martin 1964), and additing lime to three soils tended to shorten residence times of methyl 

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 allelochemical residence times. 

23
8. Microbial Activity 

Microbial degradation may also be responsible for the disappearance of glucosinolate 

hydrolysis products. Soil sterilized with heat and soil extracts sterilized by filtration or 

heating slowed the disappearance of methyl ITC in sealed bottles (Ashley et al. 1963). 

Accelerated loss of methyl ITC after successive soil applications has been demonstrated in 

field and laboratory studies (Smelt et al. 1989). 

Although microbial degradation is suggested by these data, this is not a consistent 

observation. For example, attempts to sterilize soil by autoclaving or ethyleneoxide 

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

same soil treatments increased propenyl CN half-life only slightly. In contrast to soils that 

appear to have enhanced degradation, a second treatment of fumigant to soil did not enhance 

ITC disappearance (Ashley et al. 1963) and two soils treated at least 13 times showed no 

increase in the ITC disappearance rate (Smelt et al. 1989). Similarly, five additions of 

propenyl ITC or propenyl CN during a period of 10 d failed to alter the half-lives of these 

chemicals (Borek et al. 1995a). No clear evidence demonstrates that biotic transformation 

controls the fate of ITCs in soil. The relative importance of microbial activity in the 

degradation of organic nitriles is also unclear, although biotic degradation is probable. 

Numerous microorganisms have been isolated that degrade aliphatic nitriles and a few that 

degrade aromatic nitriles (Nawaz et al. 1991, 1992). 

Several bacteria have been identified that degrade SCN-

 (Betts et al. 1979; Happold et al. 

1958; Katayama and Kuraishi 1978; Putilina 1961; Smith 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 NH4SCN). Even after temporary sterilization 

with 897 kg NH4SCN ha-1 marked stimulation of plant growth began to occur after 69 d 

(Beekhuis 1975). Other investigations showed that autoclaving soils before incubation and 

adding NaN3 or propenyl ITC slowed SCN-

 disappearance (Brown and Morra 1993). 

Disappearance of SCN-

 was correlated with organic carbon content of soils when 

temperatures were at or below 30o

C. This suggests that SCN- losses are related to the 

amount of carbon available to support microbial activity rather than SCN-

 sorption to the 

organic carbon fraction of the soil. 

Results for methyl ITC and propenyl ITC discussed throughout this section are not 

necessarily the same as for other ITCs because of the influence of the R group chemistry. 

Different R groups affect volatility and solubility, resulting in altered biological activities. 

As a consequence, individuals involved in efforts to develop plant varieties for pest control 

should consider glucosinolate type as well as total glucosinolate concentration. 

24
C. Additional Allelochemicals 

Numerous other S-containing compounds, mainly volatiles, have been identified as 

secondary products of glucosinolates in cruciferous tissues. Methanethiol, dimethyl sulfide, 

and dimethyl disulfide are produced in cabbage-amended soils (Gamliel and Stapleton 1993). 

Although these compounds possess lower toxicities than ITCs (ITCs > thiols ≅ thiocyanates 

> sulfides), they are produced in larger amounts and for much longer periods than propenyl 

ITC (Gamliel and Stapleton 1993; Lewis and Papavizas 1971; Virtanen 1965; Walker et al. 

1937). Greater total amounts and longer production times may compensate for lower 

toxicities, increasing the potential importance of these compounds in pest inhibition. 

Several of these S-containing compounds may result from abiotic or biotic degradation of 

primary glucosinolate hydrolysis products. Two water soluble degradation products of 4-

methylthio-3-butenyl ITC, one identified as 2-thioxo-3-pyrrolidinecarbaldehyde, were 

fungicidal (Uda et al. 1993). Some smaller molecular weight products might be explained by 

the simple loss of a part of the molecule such as methanethiol from thioalkyl ITC (Uda et al. 

1990). Other compounds, including COS, H2S, and CS2 are formed from ITCs by a series of 

hydrolysis reactions (Bailey et al. 1961; Challenger 1959; Dateo et al. 1957). Alternatively, 

H2S so generated might react with CO2 and produce COS (Shaw et al. 1980). Aerated and 

waterlogged soils amended with SCN-

 produced COS via an unknown mechanism (Minami 

and Fukushi 1981). Only COS was produced in a waterlogged soil amended with SCN-

, 

although samples were monitored for H2S, CH3SH, CH3SCH3, CS2, and CH3SSCH3 (Minami 

1982). Bending and Lincoln (1999) suggest with respect to fresh tissues that other volatile S 

compounds were likely to be as important in pest control as ITCs. Further information 

regarding reaction pathways would enhance our ability to predict and optimize pest control 

efficiency. 

Other compounds besides products of glucosinolate degradation may be biologically active 

and thus contribute to pest inhibition. For example, S-containing indole compounds 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 compounds are sometimes 

suspected as biologically active agents in cruciferous plants (Kutácek 1964), a possibility 

since seed meals of some Brassica contain relatively higher phenolic compound 

concentrations than other sources (Shahidi and Naczk 1992). Given the ubiquitous nature of 

phenolics, biological activity may be related more to quality than quantity (Levin 1971; 

Singleton and Kratzser 1973). Although phenolics are often suggested to participate in plant 

defenses against infection, little work appears to have been done on the potential pesticidal 

activity of specific phenolics in Brassica seed meals. 

Additional inhibitory compounds may be produced by breakdown of the S-containing amino 

acids methionine and cysteine. Compounds such as methanethiol, dimethyl disulfide, and 

dimethyl trisulfide are derived from the non-protein amino acid S-methyl-1-cysteinesulfoxide 

(SMCSO) via a methyl methanethiosulfinate intermediate (Chin and Lindsay 1994; Marks et 

al. 1992; Maruyama 1970; Ostermayer and Tarbell 1960). The intermediate is produced 

25
either enzymatically by cysteine sulfoxide lyase or nonenzymatically 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 approximately 83% of the 

original SMCSO content, but only one-tenth the original glucosinolate content (Fales et al. 

1987), suggesting SMCSO may be less susceptible to rapid degradation and thus remain 

available for longer time periods. 

In some cases, methanethiosulfinate is proposed to react with H2S to form thiols and sulfides 

(Chin and Lindsay 1994). Any H2S produced from ITCs could thus react with SMCSO 

products to form yet additional allelochemicals. Optimal pest inhibition may require 

combined interactions of glucosinolate products, SMCSO products, and additional 

compounds produced by reactions between components of these two pools. Various 

products could be formed simultaneously or sequentially, with one suite of biologically 

active compounds replacing another during an extended period. Effects on nematodes have 

been observed to last several weeks (Santos and Mojtehedi, personal communication) 

suggesting the involvement of other compounds beyond the active lifetime of volatile ITCs. 

D. Glucosinolate Concentrations 

Yield data for many plant/soil/pest systems suggest that glucosinolate concentrations are, or 

border on being, sufficient for pest control. Efforts 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 may not be the only means to increase product concentration. Some limited 

increases in resistance have been achieved by applying synthetic ITC precursors to Brassica

plants (Dawson et al. 1993; Griffiths et al. 1989). Methyl jasmonate and salicylic acid, other 

compounds 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 (combined with myrosinase) for pest suppression. Although synthetic inputs would 

not be eliminated, reductions might be achieved by supplementing plowed-under plant 

tissues with low levels of commercial ITC-producing fumigants. 

Although data are limited, we can look quantitatively at the question of glucosinolate 

concentrations and the potential to control plant pests. The amount of methyl ITC 

recommended for soil applications using sodium N-methyldithiocarbamate can be estimated 

by assuming a 15.2 cm depth of incorporation, soil bulk density of 1.4 g cm-3, and 100% 

conversion to methyl ITC (Brown et al. 1991). Calculated values were 517-1294 nmol 

methyl ITC g-1 soil depending on the crop and type of control required. These are higher 

than the amount of ITC (301 nmol g-1 soil) observed in soil amended with rapeseed mealamended at a rate of 3% (w:w) (Brown et al. 1991). To kill 50% of wireworms (L. 

26
californicus), rapeseed meal had to be amended at a rate of 114 g kg-1

 soil (Elberson et al. 

1996), too high to be useful. However, the same level of control for black vine weevil 

(Otiorhynchus sulcatus) required only 19 g rapeseed seed meal kg-1

 soil (Borek et al. 1996b). 

The previous studies were performed with rapeseed meal having a glucosinolate 

concentration near 40-50 μmol g-1 tissue. Complete release of glucosinolate to ITC has not 

been observed (Brown et al. 1991; Brown and Morra 1995). Instead, the data indicate that a 

maximum of 20% of the glucosinolate will be released as ITC from rapeseed meal (Brown et 

al. 1991). If we now assume similar release efficiencies for mustard meals that contain 

glucosinolate concentrations of 250 μmol g-1 tissue, we can estimate the amount of meal 

necessary to approach the recommendations application rates for commercially available ITC 

fumigants (517-1294 nmol methyl ITC g-1 soil). A 1% (w:w) meal amendment to soil of a 

meal containing 250 μmol glucosinolate g-1 tissue would result in the release of 500 nmol 

ITC g-1 soil. We are now at the lower limit of what is considered necessary for soil 

sterilization. Doubling the amendment rate to 2% (w:w) would increase predicted ITC 

release to 1000 nmol g-1 soil and thus achieve a concentration considered near the upper limit 

of that recommended for commercial fumigants. Thus, meals with glucosinolate 

concentrations in excess of 200 μmol g-1 tissue may indeed be effective in pest control at 

practical tissue amendment rates. 

E. Timing 

Length of exposure to the allelochemical is one variable associated with timing. Lewis and 

Papavizas (1971) showed in sealed containers that concentrations failing to kill 

microorganisms in 2 d might kill them in 4-6 d. Evidence indicates similar results with 

insects and seeds (Lichtenstein et al. 1962, 1964; Pieczarka and Warren 1960). Dormant 

seeds making up the seed bank in soil and exposed to root exudates over a period of time 

could be more susceptible to inhibition than introduced seeds exposed to root exudates for 

only a short time. 

Another important variable is the time when initial exposure occurs; in other words, the time 

when the target organism is introduced to the glucosinolate-containing tissue. Pea seeds 

planted immediately after soil amendment with cabbage did not germinate. Peas planted 3 

weeks later showed some 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 

enhancement (Papavizas 1966). Only fresh cabbage juice inhibited lactic acid (Leuconostoc

mesenteroides) bacterial growth (Kyung and Fleming 1994). Inhibition of germination and 

plant growth by ITCs was most effective when the chemicals were applied to seeds or 

applied within the first 3 d of the germination period (Leblová-Svobodová and Koštír 1962). 

Timing maximal glucosinolate product formation with a susceptible life stage of the target 

organism may be necessary. Glucosinolate concentrations in plants usually start out low, 

climb with fluctuations to some maximum, then drop off near the time of plant senescence 

(Freer et al. 1989; Griffiths et al. 1991; Macfarlane-Smith and Griffiths 1988). Aqueous 

extracts of wild mustard were most effective at inhibiting growth when extracts were made 

27
during the bolting and flowering stage (Jiménez-Orsornio and Gliessman 1987). Coplantings 

of Brassica plants with other plants do not inhibit seed germination or plant growth (Bell and 

Muller 1973; Choesin and Boerner 1991), perhaps because plant stages susceptible to 

inhibition are past by the time sufficient glucosinolate products are present. In the case of 

coplanting with two Brassica species, planting wild mustard with broccoli resulted in higher 

broccoli yields, but wild mustard planted before broccoli inhibited broccoli yield (JiménezOrsornio and Gliessman 1987). In a consistent fashion, insects and microorganisms may be 

more susceptible at certain stages in their life cycle. Relatively sudden releases and higher 

concentrations of products from amendments timed to susceptible stages of an organism’s 

life cycle would be most effective. 

F. Additional Benefits 

Other benefits may be obtained by using glucosinolate-containing plants in pest control 

strategies. For example, soil structure is improved by use of organic amendments and high 

root development like that in oilradish and mustard is important 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 amendments can serve as nutrient sources and high protein-containing rapeseed meal 

may be used as a supplementary nitrogen source (Johansson and Ascard 1994; Kücke 1993). 

Carbon to nitrogen ratios of B. juncea, S. alba, and B. napus meals range from 7 to 10 (Morra 

and Johnson-Maynard 2003). Mineralizing the meal will thus provide a substantial amount 

of plant available N and no immobilization. Biofumigating compost with seed meal would 

thus add nitrogen to the mix and simultaneously reduce unwanted weed seeds and disease 

pathogens. 

28
V. Crushing and Extraction Technology

Current methods of cold pressing the meal result in residual oil of 5%-10% on a weight 

basis. Residual oil may help trap hydrophobic ITCs, prevent volatile losses, and increase 

pesticidal effectiveness. However, residual oil may also decrease the release efficiency of 

ITC from the meal into the soil and decrease its effectiveness. The overall impact of residual 

oil in the meal will vary with the ITC and its respective hydrophobicity. The effect of the oil 

on pest control has never been determined. 

Commercial crushing facilities produce temperatures in the seed and seed meal that may 

deactivate myrosinase. Temperatures are increased on purpose to maximize oil extraction or 

unintentionally if an extrusion process follows crushing. The maximum temperature at which 

myrosinase integrity is maintained during a commercial crushing procedure has not been 

determined. 

Current research is focused on using the meal as a carrier for the pesticide. The meal will 

also provide plant nutrients in addition producing ITC for pesticide control. However, the 

meal will also provide a carbon source for fungi and potentially increase pathogen problems. 

Extracting the ITC before soil application may circumvent this problem. 

Techniques have been developed for extracting propenyl isothiocyanate from B. juncea meal 

using a commercially available unit. A commercial 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 marketed to Japanese companies that use it as a food additive. Using a similar 

approach for producing pesticides seems feasible. 

29
VI. Allergenic Reactions 

Although the compounds are produced by plants, human and animal toxicity is still a concern 

if alternative pest control strategies are to be implemented. Do natural toxic substances in 

plants have advantages over synthetic pesticides? In contrast to a new synthetic pesticide, 

much information about the toxicology of these products is already known because of the 

importance of cruciferous crops in human nutrition and animal diets. 

When used as a primary food source, glucosinolate-containing plant material clearly has 

deleterious effects. Feeding kale to livestock can reduce fertility and induce goiter (Johnston 

and Jones 1966). Diets of crambe meal caused early death to rats (Daxenbichler et al. 1966), 

and in separate tests, rats fed crambe meal and rapeseed meal exhibited symptoms similar to 

rats with nitriles in their rations (Van Etten and Tookey 1979). Livestock poisoning has been 

commonly reported when animals are fed excessive cruciferous plant material (Kingsbury 

1964). 

ITCs are general biocides whose activity results from interaction with proteins. They interact 

nonspecifically and irreversibly with proteins and amino acids to form stable products by 

reacting with sulfhydryl groups (Fenwick et al. 1983; Kawakishi and Kaneko 1985; Ware 

1983), disulfide bonds (Kawakishi et al. 1983), and amines (Kawakishi and Kaneko 1987; 

Wood 1975). Reaction with terminal amino groups is the basis of the well-known Edman 

degradation for amino acid sequencing of proteins. ITCs inactivate enzymes in vitro, but the 

actual effects in vivo may be controlled by accessibility (Wood 1975). 

Numerous studies concerning ITC toxicities have been performed. Contact of ITCs with 

skin and mucous membranes (respiratory tract) cause irritation, and many are considered 

lachrimators. At 20-50 mg kg-1 body weight propenyl ITC produced epithelial hyperplasia, 

stomach ulcers, and minor inflammatory foci in the livers of dogs, but did not show similar 

effects in rats even at 500 mg kg-1 body weight (Hall 1973). Antithyroid effects are 

attributed to ITCs, although this may be caused by metabolic conversion of the ITCs to SCN-

. An oral LD50 in rats for methyl ITC is listed as 175 mg kg-1 (Worthing 1987) and 339 mg

kg-1 for propenyl ITC (Wood 1975). Subcutaneous LD50 values are considerably lower at 50 

mg kg-1 for methyl ITC and 80 mg kg-1 for propenyl ITC (Wood 1975). The LC50 of methyl 

ITC at 96 h is 0.13 mg L-1 for bluegill and 0.37 mg L-1 for trout and carp (Worthing 1987). 

Mammalian systems metabolize and eliminate ITCs fairly rapidly. Less than 5% of a dose of 

propenyl ITC was retained by rats and mice 24 h after administration (Ioannou et al. 1984). 

ITCs administered both orally and by injection in rats and mice are primarily detoxified by 

glutathione and excreted as S-(N-methylthiocarbamoyl)mercapturic acids (Lam et al. 1993; 

Ioannou et al. 1984; Mennicke et al. 1983). Detoxification and excretion of benzyl ITC in 

humans is similar (Mennicke et al. 1988). 

ITC attack of DNA and cancer cell induction seem to be unimportant in describing the mode 

of toxicity. Mutagenicity tests are sometimes positive, but often negative (Fenwick et al. 

30
1983; Ioannou et al. 1984; Musk and Johnson 1993). Chronic administration of propenyl 

ITC to rats and mice caused papillary growths in bladders of male rats only (related to more 

concentrated metabolite excretion), but did not cause tumors (Ioannou et al. 1984). There is 

in fact evidence that ITCs are anticarcinogenic (Mossoba et al. 1989; Fimognari 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). Tumors were 

inhibited in mice and rats that were given ITCs both before and after administering known 

carcinogens (Fenwick et al. 1983; Wattenberg 1977, 1981). Short-term chemical 

mechanisms appear to be involved in the suppression, such as high toxicity toward cancer 

cells (Nastruzzi et al. 1996), but longer term induction of enzyme systems that fight cancer in 

the body are also implicated (Institute of Food Technologists 1993). A potent inducer of 

protective enzymes is 4-methylsulfinylbutyl ITC found in broccoli (Zhang, et al. 1992, 

1994). Thus, if ITC is consumed in normal dietary levels, it may be more beneficial than 

harmful. 

Toxicological implications of glucosinolate hydrolysis must also include the possibility of 

OZT formation. The most common 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 

meal has limited use as a cattle feed, in part because the glucosinolate precursor of goitrin is 

present in large amounts. Other information on biological activity is relatively scarce, 

although OZTs interfere with a few enzymatic processes. For example, OZTs prevent the 

oxidation of trimethylamine (Fenwick et al. 1983) and goitrin moderately inhibits dopamine 

β-hydroxylase (Zenker et al. 1988). OZT toxicities to mice reported in the form of LD50

values range from 1260 to 1415 mg kg-1 (Van Etten and Tookey 1979). There is some

susceptibility to nitrosation in the presence of nitrite (Lüthy et al. 1984), suggesting the 

possible formation of carcinogenic compounds under conditions encountered in the stomach. 

General toxicity appears to be relatively low, but further investigation of OZT toxicities 

seems appropriate. 

Toxicity of other organic products, including nitriles, epithionitriles, and thiocyanates is 

likely related to the cyano group. Cyanide salts and gas inactivate certain enzyme systems, 

especially those involved in cellular respiration such as cytochrome oxidase (Johnston 1987; 

Sittig 1985). Similarities in the metabolism of organic nitriles and cyanide include increased 

urinary excretion of SCN-

, detection of CN- in tissues, and depressed cytochrome oxidase 

activity (Duncan 1991). Toxicities of organic nitriles vary but are generally lower than 

cyanide salts and hydrogen cyanide. Toxicity seems particularly pronounced towards the 

liver (hepatotoxic) and kidney (Dietz et al. 1991; Duncan 1991; Fenwick et al. 1983; Van 

Etten et al. 1969). The LD50 in rats for ethyl CN (propanenitrile) is 50-100 mg kg-1 (Sittig 

1985); for 1-cyano-2-hydroxy-3-butene, values of 170 mg kg-1 and 200 mg kg-1 have been 

reported (Fenwick et al. 1983; Tookey et al. 1980). Unsaturated nitriles appear to be less 

toxic than saturated nitriles. For example, rat oral LD50’s exceeded 600 mg kg-1 for 3-

butenyl CN (4-pentenenitrile) and 720 mg kg-1 for benzonitrile, although acetonitrile (methyl 

CN) is an exception with a reported oral LD50 in rats of 3030-6500 mg kg-1 (Dietz et al. 

1991; Smiley 1979). 

31
Thiocyanate and epithionitrile toxicities appear similar to nitrile toxicities. Low molecular 

weight thiocyanates, including benzyl and phenyl, respond to the same antidotes for 

poisoning as does cyanide (Wood 1975). Alkyl thiocyanates are generally more active than 

aromatic thiocyanate, and toxicity decreases with increasing molecular weight (Wood 1975). 

For 1-cyano-3,4 epithiobutane, mortality in rats occurred after ingestion of 180 mg kg-1

(Dietz et al. 1991) and an LD50 of 109 mg kg-1 has been reported (Fenwick et al. 1983). An 

LD50 range of 178-240 mg kg-1 was estimated for 1-cyano-2-hydroxy-3,4 epithiobutane (Van 

Etten et al. 1969). 

Although the LD50 reported for 3-indolylacetonitrile in rats is 255 mg kg-1, indolyl products 

such as 3-indolylmethanol stimulate enzymatic systems involved in detoxification of 

xenobiotics and the inhibition of cancer (Loub et al. 1975). Protective activity, however, is 

less dramatic than with ITCs. Results of several experiments are somewhat mixed depending 

on the system examined and the specific conditions (McDanell et al. 1988), making it 

difficult to assess actual impact. In one study conducted with trout, 3-indolylmethanol 

administered after exposure to aflatoxin B1 actually enhanced carcinogenesis. However, 3-

indolylmethanol, 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 compounds. In fact, 

biological systems often metabolically detoxify cyanides by rhodanese-catalyzed conversion 

to SCN-, which is then readily excreted (exogenous sulfur is needed and thus thiosulfate is 

administered as part of the antidote for cyanide poisoning). Rhodanese is widely distributed 

in nature (Tabatabai and Singh 1976). As a result, SCN- was considered nontoxic and was 

used for a time as a drug to reduce blood pressure. However, side effects of SCN- caused its 

use to be dropped. 

SCN- toxicity to rats is similar to benzonitrile with a reported LD50 for NaSCN of 765 mg

kg-1. Dogs are more susceptible than rats or mice to SCN- (Wood 1975). The process of 

cyanide conversion to SCN- is apparently in metabolic equilibrium, and adding SCN- to 

biological systems produces cyanide. However, this alone does not account for its acute 

toxicity (Wood 1975). SCN- interacts with proteins, and is listed as the most effective anion 

in the Hofmeister series for destabilizing protein structure (Voet and Voet 1990; Wood 

1975). It may act as a noncompetitive inhibitor with many enzymes, and penetrates cell 

membranes (Wood 1975). SCN- is a pseudohalogen that acts as a competitor with iodine 

and thus may 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 organisms vary and are 

not always known. In sufficient quantity many of these compounds are highly toxic. Even 

with plant tissues, irritation of skin from excessive handling or a negative respiratory 

reaction might occur. However, exposure to amounts encountered in pest control strategies 

with plant tissues is likely to be minimal. The action of the toxins is direct and “up front.” 

There do not appear to be serious, more subtle, negative long-term impacts on human health 

such as cancer or birth defects. Indeed, we have probably ingested many of these compounds 

in low levels for thousands of years, most of the time presumably without detriment. 

32
VII. Registration 

Products derived from mustard meal or meal-containing glucosinolates have been registered 

for a variety of purposes (Table 4). EPA has granted an “Exemption From the Requirement 

of Tolerance to Isothiocyanate as a Component of Food Grade Oil of Mustard” (Federal 

Register 1966). The most relevant products in the current discussion include mustard meal 

(B. juncea), currently marketed in Australia as a snail and slug killer and meadowfoam

(Limnanthes alba) meal marketed in the United States as a soil amendment in horticulture. 

Meadowfoam meal is marketed by Natural Plant Products as LimnaxTM, AlbaGroTM, and 

AlbaAideTM. EPA approved pesticides containing propenyl ITC as an active ingredient are 

marketed by Champon Natural Products (Champon Millenium Chemicals, Inc. 11112 Split 

Rail Lane, Fairfax Station, VA 22039; Phone: 703-426-8424; E-mail: 

champon@ix.netcom.com; website: www.champon.com). A number of other products 

containing propenyl isothiocyanate as an active 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 meals 

other than B. juncea most likely will require greater testing before it can be approved by the 

EPA. 

33
VII. Genetics and Interspecific Hybridization 

Canola or rapeseed (Brassica napus L.) is a member of the Brassicaceae family. Canola is 

the term used to describe Brassicaceae cultivars that contain erucic acid less than 2% of the 

total fatty acid content and less than 30 μmol per gram of total glucosinolates in oil-free meal 

(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 

μmol per gram of total glucosinolates in oil-free meal) glucosinolate content in the seed 

meal, although new rapeseed cultivars have seed meal comparable to canola. 

Early records indicate that canola (rapeseed) was cultivated in India more 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 may have had multiple areas of origin in Europe within 

overlapping ranges of the diploid parental taxa (B. rapa and B. oleracea) (Warwick and 

Francis 1994). The main phytogeographical zones for B. napus are the Euro-Siberian, and 

the Irano-Turanian regions (Warwick and Francis 1994). The main regions of canola 

cultivation are parts of Asia (China, India, Indonesia), Europe, the Americas (mainly 

Canada), parts of Africa, and Australasia. 

A. Taxonomy, Genomic Constitutions, and Relationships 

The origins of amphidiploid B. napus, B. carinata, and B. juncea were suggested by U 

(1935), based on interspecific hybridizations and chromosome counts (Figure 3). This 

triangle arrangement of species indicates that cultivated canola, B. napus (n = 19, AACC 

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

genome) and B. rapa (n = 10, AA genome) (U 1935; McNaughton 1976). This has been 

confirmed by later studies using different techniques, including flavonoid composition (Dass 

and Nybom 1967) and seed protein serology (Vaughan 1977). Song et al. (1988) used 

RFLPs to study the genomic constitution of diploid and amphidiploid species, and concluded 

that amphidiploid B. napus has evolved through different combinations of the diploid 

morphotypes (B. rapa and B. oleracea) and that phylogenetic origins may be common 

mechanisms for the natural occurrence of amphidiploids in Brassica, which confirmed that 

B. napus is an interspecific hybrid derived from B. rapa and B. oleracea. The authors 

suggested that the C genome component is larger in the cultivated B. napus lines, since these 

plants were closer to the C genome than the A genome in the phylogenetic tree. 

B. Oil and Seed Meal Characteristics and Inheritance 

The major fatty acids of Brassica species oil are palmitic (16:0), stearic (18:0), oleic (18:1), 

linoleic (18:2), linolenic (18:3), eicosenoic (20:1) and erucic acids (22:1). Some mustard species 

also have relatively high concentrations of nervonic acid (24:1). Canola oil must, by definition, 

not exceed more than 2% erucic acid content. In contrast, industrial rapeseed oil is usually very 

high (40%-50%) in erucic acid content. Laboratory animals that eat diets with a high proportion 

of oil containing high erucic acid levels perform poorly. Also, high erucic acid levels in human 

34
diets can cause abnormality of fat utilization in the heart and skeletal muscles, with undesirable 

changes in the tissue of the myocardium (Roine et al. 1960). Other reports from Canada and 

Europe have confirmed 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 almost always associated with high oleic acid content, which is 

almost as effective in reducing cardiovascular risk as is the long-touted polyunsaturated linoleic 

acid (Ackman 1990). 

Linolenic acid is very unstable and is easily oxidized to give unpleasant smelling substances. 

Canola oil contains 8%-12% linolenic acid, although cultivars can be developed with reduced 

levels (DeBonte 1994). In contrast, a high percentage 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 improve its dietary value (Holmes 1980). Nutritionally, linoleic acid 

(vitamin F) is an important and essential polyunsaturated fatty acid that also reduces cholesterol 

content in the blood and is an essential component of cell membranes (Scarisbrick and Daniels 

1986). 

Yellow mustard (S. alba) contains considerably higher erucic acid levels than are suitable as an 

edible oil. Also, yellow mustard does not produce high enough erucic acid levels to be valuable 

for industrial purposes (Carlson and Van Dyne 1992). In addition, yellow mustard is low in 

linoleic acid and has higher linolenic acid than would be desirable in edible oil. 

Since the early 1960s, many 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 embryo and not by the sporophyte (Dorrel and Downey 1964; 

Harvey and Downey 1964; Kondra and Stefansson 1965; Stefansson and Hougen 1964). Erucic 

acid content in B. napus is controlled by a two gene-pair-system acting in an additive manner 

(Downey and Craig 1964; Kondra and Stefansson 1965; Stefansson and Hougen 1964). Jönsson 

(1977a, 1977b) found that erucic acid content in B. napus is controlled by a series of multiple 

alleles. Levels of 5%-10%, 20%-35%, and more 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 examine the inheritance of fatty acid profiles in S. alba

(Rafiullah 1994). Rafiullah used a 4x4 half diallel cross to determine the inheritance of fatty 

acid profile as quantitative characteristics. The results indicated that erucic and oleic acids had 

greater additive genetic variation than dominant genetic variation, and that erucic acid was 

controlled by additive and nonadditive genes with some degree of linkage and or epistasis in S. 

alba. However, these studies were conducted by chemically analyzing field open pollinated 

seed. This could add serious bias to the results because of outcrossing. Tang et al. (1995) found 

that erucic acid content in yellow mustard has a large quantitative component with a few major 

dominant alleles. 

Traditional rapeseed cultivars (where canola cultivars were derived) all had relatively high 

concentrations of glucosinolates in the seed that adversely affected the quality of the meal as 

livestock feed. When glucosinolate compounds are enzymatically degraded, they produce toxic 

compounds that can cause metabolic disturbances when fed to nonruminant livestock (Kondra 

35
and Stefansson 1970). High glucosinolate levels needed therefore to be reduced in rapeseed 

cultivars to avoid these detrimental effects on animals consuming the meal (Josefsson 1972). 

A tremendous breeding effort was made to develop 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 major glucosinolates 

apparently control biosynthesis of glucosinolates in Bronowsky. This cultivar had low levels of 

glucosinolates in 1968 but was unadapted (Jönsson 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 combine low levels of 

glucosinolates with low levels of erucic acid (Stefansson and Kondra 1975). 

In contrast to canola, yellow mustard seed has traditionally been selected for condiment mustard 

powders. Mustard pungency is associated with high content of glucosinolates and their 

hydrolysis products (Holmes 1980). Therefore, past breeding efforts in this species have been 

aimed at increasing glucosinolate levels rather than making any attempt to reduce the content of 

glucosinolates in the seed. 

Very few studies have examined the inheritance of glucosinolates in yellow mustard. This is 

mainly because very little genetic variation has been found for this trait within the species. 

Glucosinolate content of S. alba seeds produced from the same parent from both self- and crosspollinations were similar, showing that inheritance of glucosinolates is maternally controlled 

(Kondra and Stefansson 1970; Love et al. 1990). F2 seed of reciprocal crosses contained both 2-

propenyl and 3-butenyl glucosinolates, indicated nuclear genes controlled their formation. Love 

et al. (1990) also concluded that the synthesis of both 2-propenyl and 3-butenyl glucosinolates 

were controlled by multiple additive alleles at the same loci. More recently a single recessive 

allele has been identified (Brown et al. 2002) that completely blocked 3-hydroxybenzyl 

glucosinolate in yellow mustard seed meal. 

C. Interspecific and Intergeneric Crosses in Brassicaceae 

Substantial literature discusses intergeneric cross compatibility within the family Brassicacae. B. 

napus has been combined with several species of Diplotaxis and Eruca by sexual crosses and 

embryo rescue (Batra et al. 1990; Ringdahl et al. 1987), and with various other Brassica species 

(Glimelius et al. 1990; Jourdan et al. 1989a; Sjodin and Glimelius 1989). Other intergeneric 

Brassica hybrids produced include various combinations 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; Toriyama et al. 1987a, 1987b). These hybrids generally showed some degree of 

fertility in crosses and have a range of chromosome numbers 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 embryo rescue. Lefol and 

Darmency (1993) studied spontaneous and manual interspecfic and intergeneric crosses 

between B. napus and two weedy relatives, B. adpressa and R. raphanistrum. They were 

36
able to obtain hybrids that showed superior vegetative development compared to the weed 

parent; the backcrosses to the parents also produced seed. In their studies on interspecific 

crosses, Nishiyama et al. (1991) found that many of these crosses produced wrinkled or 

empty seed because of poor endosperm development. However, crosses between B. napus

and B. campestris and between B. napus and B. nigra spontaneously produced a few true F1

seeds (Nishiyama 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 

combination prompted further investigation of B. napus x S. alba hybrids with elevated 

glucosinolate content for use as biofumigants. 

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 optimal 

conditions with in vitro ovary culture, hybrid plants were obtained in all species cross 

combinations and more hybrids were obtained when canola was used as the female. 

Eber et al. (1994) reported that spontaneous hybridization between male 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. When ovule culture was used, one hybrid from B. napus × S. 

arvensis was obtained but was highly sterile. The authors concluded that the crossing 

between and transfer of genes from B. napus to S. arvensis would be remote even under 

favorable conditions. 

37
IX. Recommendations 

Meals with isothiocyanate-producing glucosinolate concentrations in excess of 200 μmol g-1

tissue will most effectively control a wide variety of plant pests. The target is to produce in 

excess of 100 nmol ITC g-1 soil to approach commercially recommended rates for synthetic 

ITC pesticides. This should be possible assuming glucosinolate to ITC release efficiencies of 

20%. Consideration should be given to increasing the release efficiency of the ITC from the 

meal using physical methods of processing the meal or through the addition of adjuvants. 

Absolute ITC amounts and release efficiencies must be determined for the meals of interest 

to confirm that we indeed are producing ITC in excess of 100 nmol g-1 soil. Meals must be 

tested against specific target organisms and phytotoxic effects on the crop plant must be 

determined. Fungal pathogens will be the most difficult to control because carbon added to 

the soil in the form of meal may enhance growth of the pathogen. Insects, nematodes, and 

weeds will be more easily controlled, and commercialization of a final product with 

acceptable efficacy is likely. Different ITCs express different toxicities as a result of direct 

effects on the plant pest as well as differential sorption and reaction with soil colloids. 

Bioassay results obtained in the absence of soil must therefore be confirmed within the soil 

environment. Predictions of ITC pesticidal activity should be approached in the same

fashion as commercial pesticides by determining organic carbon partition coefficients, 

Henry’s constants, and half-lives. These parameters will vary with the ITC, and pesticidal 

activity can be optimized by selecting ITCs with the greatest potential for efficacy. The 

potential to introduce thiofunctionalized glucosinolates into species by way of breeding 

efforts may improve pesticidal efficacy of the meal. Additional benefits from the meal such 

as N release may increase the advantages of using meal amendments in cropping strategies. 

Effort should be devoted to quantifying the nutrient benefits of meal amendment since such 

benefits may be just as important to the consumer as pesticidal activity. 

38
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70
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71
Table 1. Naturally Occurring Glucosinolates 

___________________________________________________________________________________________

R= trivial name plant parta 

___________________________________________________________________________________________

methyl CH3 glucocapparin various Capparis s 

various Cleome s 

s 

ethyl CH3CH2 glucolepdiin Lepidium menziesii s 

2-methylethyl (isopropyl) (CH3)2CH glucoputranjivin Brassica napobrassica r 

Cochlearia officinalis s 

Lunaria biennis s 

Moringa peregrina s 

propyl CH3CH2CH2 Brassica oleraceae l 

1-methylpropyl CH3CH2(CH3)CH glucocochlearin Brassica napobrassica r 

Brassica campestris (B. rapa) l,r 

Dentaria pinnata r 

2-methylpropyl CH3(CH3)CHCH2 Armoracia lapathifolia r 

r 

Brassica napus s 

Conringia orientalis l Moringa peregrina s 

butyl CH3(CH2)3 Armoracia lapathifolia r 

Capparis flexuosa l 

2-methylbutyl CH3CH2(CH3)CHCH2 Armoracia lapathifolia r 

Dentaria pinnata r 

3-methylbutyl CH3(CH3)CHCH2CH2 Armoracia lapathifolia r pentyl CH3(CH2)4 Armoracia lapathifolia r 

Brassica rapa l 

72
4-methylpentyl CH3(CH3)CH(CH2)3 Armoracia lapathifolia r 

Brassica napus l,s,r 

1996 

Raphanus sativus r 

hexyl CH3(CH2)5 Brassica napus s 

Raphanus sativus r 

5-methylhexyl CH3(CH3)CH(CH2)4 Armoracia lapathifolia r 

Brassica napus l,s,r 

allyl CH2CHCH2 sinigrin Armoracia lapathifolia r 

r l 

Brassica carinata s 

s 

Brassica juncea s 

l, s, r 

Brassica napus s 

r 

Brassica oleraceae s 

l 

l,r 

2-methyl-2-propenyl CH2(CH3)CCH2 Peltaria alliacea s 

3-butenyl CH2CHCH2CH2 gluconapin Brassica campestris (B. rapa) s 

l,r 

Brassica oleraceae l 

r s 

Brassica napus s 

l l, s r 

Brassica napobrassica r 

3-methyl-3-butenyl CH2(CH3)CCH2CH2 Capparis linearis l 

73
4-pentenyl CH2CH(CH2)3 glucobrassicanapin Armoracia lapathifolia r 

r 

Brassica campestris (B. rapa) l,r 

Brassica oleraceae s 

Brassica napus s 

l l, s r 

Brassica napobrassica r 

5-hexenyl CH2CH(CH2)4 Armoracia lapathifolia r 

Brassica napus s Raphanus sativus r 

2-hydroxypropyl CH3(OH)CHCH2 Armoracia lapathifolia r 

3-hydroxypropyl HOCH2(CH2)2 Arabidopsis thaliana s 

3-hydroxybutyl CH3(OH)CH(CH2)2 Armoracia lapathifolia r 

Capparis flexuosa l 

4-hydroxybutyl HOCH2(CH2)3 Arabidopsis thaliana s 

Capparis flexuosa l 

2-hydroxypentyl CH3(CH2)2(OH)CHCH2 Armoracia lapathifolia r 

1-methyl-2-hydroxyethyl HOCH2(CH3)CH glucosysimbrin Coluteocarpus vesicaria s A

Dentaria laciniata s Euclidium syriacum s ATorularia torulosa s A

1-hydroxymethylpropyl CH3CH2(OHCH2)CH Coluteocarpus vesicaria s A

1-ethyl-2-hydroxyethyl HOCH2(CH3CH2)CH glucosisautricin Cleome diandra s 

Sisymbrium austriacum s 

2-hydroxy-2-methylpropyl CH3(OH)(CH3)CCH2 glucoconringiin Cochlearia officinalis s 

Conringia orientalis s 

74
l 

not spec 

Erysimum orientale s 

2-hydroxy-2-methylbutyl glucocleomin Cleome spinosa s  CH3CH2(OH)(CH3)CCH2 Dentaria laciniata s 

4,5,6,7-tetrahydroxydecyl CH3(CH2)2(CH2OH)4(CH2)3 Capparis grandis r 

R-2-hydroxy-3-butenyl CH2CH(OH)CHCH2 progoitrin Brassica campestris (B. rapa) s 

(glucorapiferin) l,r 

Brassica napobrassica r Brassica napus s 

l l, s, r s r 

Brassica oleraceae s 

l r 

S-2-hydroxy-3-butenyl CH2CH(OH)CHCH2 epiprogoitrin Crambe abyssinica s 

s 

s 

Brassica napus s 

2-hydroxy-4-pentenyl CH2CHCH2(OH)CHCH2 gluconapoleiferin Brassica napobrassica r 

Brassica napus s 

l l, s, r r 

Brassica oleraceae r 

3-hydroxy-4-pentenyl CH2CH(OH)CHCH2CH2 Armoracia lapathifolia r 4-oxoheptyl CH3(CH2)2CO(CH2)3 glucocapangulin Capparis angulata s 

not spec. 

5-oxoheptyl CH3CH2CO(CH2)4 gluconorcappasalin Capparis salicifolia s 

75
5-oxooctyl CH3(CH2)2CO(CH2)4 glucocappasalin Capparis salicifolia s 

3-methoxycarbonylpropyl CH3OCO(CH2)3 glucoerypestrin Erysimum aucherianum s A

Erysimum filifolium s AErysimum rupestre s 

l 

2-methylthioethyl CH3S(CH2)2 Armoracia lapathifolia r 

(Cochlearia armoracia) 

3-methylthiopropyl CH3S(CH2)3 glucoibervirin Arabidopsis thaliana s 

Brassica napus s Brassica oleraceae s 

l 

4-methylthiobutyl CH3S(CH2)4 glucoerucin Arabidopsis thaliana l,s 

Brassica campestris (B. rapa) r 

Brassica napus r 

r 

Brassica oleraceae s 

r 

Eruca sativa s 

s 

Raphanus sativus r Sinapis alba (B. hirta) r 

5-methylthiopentyl CH3S(CH2)5 glucoberteroin Arabidopsis thaliana s 

Brassica naporbrassica r 

Brassica campestris (B. rapa) r 

Raphanus sativus r 

6-methylthiohexyl CH3S(CH2)6 glucolesquerellin Arabidopsis thaliana s 

Ochthodium aegyptiacumr s A

7-methylthioheptyl CH3S(CH2)7 Arabis laxa s A

Arabidopsis thaliana s 

8-methylthiooctyl CH3S(CH2)8 Arabidopsis thaliana l,s 

76
9-methylthiononyl CH3S(CH2)9 Arabis hirsuta s 

10-methylthiodecyl CH3S(CH2)10 Arabis amplexicaulis s 

4-methylthio-3-butenyl CH3SCHCH(CH2)2 glucodehydroerucin Raphanus sativus s 

r 

3-hydroxy-5-methylthiopentyl CH3S(CH2)2(OH)CH(CH2)2 Erysimum hieracifolium s 

Armoracia lapathifolia r 

3-hydroxy-6-methylthiohexyl CH3S(CH2)3(OH)CH(CH2)2 Erysimum rhaeticum s 

3-oxo-8-methylthiooctyl CH3S(CH2)5CO(CH2)2 Arabis hirsuta s 

3-methylsulfinylpropyl CH3SO(CH2)3 glucoiberin Arabidopsis thaliana l 

Brassica napus s Brassica oleracea r 

l 

4-methylsulfinylbutyl CH3SO(CH2)4 glucoraphanin Arabidopsis thaliana l,s 

 (sulphoraphane) Brassica napus s 

Brassica oleracea l 

l 

5-methylsulfinylpentyl CH3SO(CH2)5 glucoalyssin Alyssum argenteum s 

Arabidopsis thaliana l,s Berteroa incana s 

6-methylsulfinylhexyl CH3SO(CH2)6 glucohesperin Arabidopsis thaliana l,s 

Euclidium syriacum s A

Notoceras bicorne s A

7-methylsulfinylheptyl CH3SO(CH2)7 glucosiberin Arabis laxa s A

Arabidopsis thaliana l,s 

8-methylsulfinyloctyl CH3SO(CH2)8 glucohirsutin Aethionema arabicum s A

Arabidopsis thaliana l,s 

77
9-methylsulfinylnonyl CH3SO(CH2)9 glucoarabin Arabis alpina s 

Arabis nova s A

Drabopsis nuda s A

10-methylsulfinyldecyl CH3SO(CH2)10 glucocamelinin various Camelina s 

11-methylsulfinylundecyl CH3SO(CH2)11 Camelina microcarpa s 

4-methylsulfinyl-3-butenyl CH3SOCHCH(CH2)2 glucoraphenin Brassica napus s 

r 

Raphanus sativus s 

s 

3-hydroxy-5-methylsulfinylpentyl CH3SO(CH2)2(OH)CH(CH2)2 Erysimum hieracifoliumb s 

3-hydroxy-6-methylsulfinylhexyl CH3SO(CH2)3(OH)CH(CH2)2 Erysimum rhaeticum s 

3-oxo-8-methylsulfinyloctyl CH3SO(CH2)5CO(CH2)2 Arabis hirsuta s 

3-methylsulfonylpropyl CH3SO2(CH2)3 glucocheirolin Brassica oleraceae l 

Cheiranthus cheiri 

s 

4-methylsulfonylbutyl CH3SO2(CH2)4 glucoerysolin Cheiranthus cheiri 

Erysimum perofskianum s 

not spec 

6-methylsulfonylhexyl CH3SO2(CH2)6 Euclidium syriacum s A

8-methylsulfonyloctyl CH3SO2(CH2)8 Heliophila amplexicaulis s 

9-methylsulfonylnonyl CH3SO2(CH2)9 Arabis turrita s 

Heliophila amplexicaulis s 

10-methylsulfonyldecyl CH3SO2(CH2)10 Arabis turrita s 

4-methylsulphonyl-3-butenyl CH3SO2CHCH(CH2)2 Raphanus sativus s 

78
3-hydroxy-5-methylsulfonylpentyl CH3SO2(CH2)2(OH)CH(CH2)2 Erysimum hieracifoliumb s 

6-methylsulfonyl-3-hydroxyhexyl CH3SO2(CH2)3(OH)CH(CH2)2 Erysimum rhaeticum s 

Benzyl C6H5CH2 glucotropaeolin Carica papaya c 

Lepidium sativum s 

s 

Reseda media s,l Sinapis alba (B. hirta) l,r 

l 

Tropaeolum majus s,l 

l 

2-hydroxybenzyl 2-OHC6H5CH2c Reseda odorata 

3-hydroxybenzyl 3-OHC6H5CH2 glucolepigramin Lepidium vesicarium s 

Reseda media s,l 

4-hydroxybenzyl 4-OHC6H5CH2 glucosinalbin Brassica napus s 

s 

Sinapis alba (B. hirta) s 

s 

l,r 

Sinapis arvensis s A

3,4-dihydroxybenzyl 3,4(OH)2C6H5CH2 

3-methoxybenzyl 3-CH3OC6H5CH2 Limnanthes douglasii s 

not spec. 

4-methoxybenzyl 4-CH3OC6H5CH2 glucoaubrietin Aubrietia hybrida l,s 3,4-dimethoxybenzyl 3,4-(CH3O)2C6H5CH2 Heliophila longifolia s 

3,4,5-trimethoxybenzyl 3,4,5-(CH3O)3C6H5CH2 Coronopus squamatus s 

2-benzoyloxyethyl C6H5COO(CH2)2 Moricandia arvensis s 

79
1-methyl-2-benzoyloxyethyl glucobenzosisymbrin Sisymbrium austriacum s 

 C6H5COOCH2(CH3)CH2

1-ethyl-2-benzoyloxyethyl glucobenzsisaustricin Sisymbrium austriacum s  C6H5COOCH2(CH3CH2)CH23-benzoyloxypropyl C6H5COO(CH2)3 glucomalcomiin Arabidopsis thaliana l,s 

Malcolmis maritima s 

not spec. 

4-benzoyloxybutyl C6H5COO(CH2)4 Arabidopsis thaliana l,s 

s 

5-benzoyloxypentyl C6H5COO(CH2)5 Arabidopsis thaliana l,s 

6-benzoyloxyhexyl C6H5COO(CH2)6 Arabidopsis thaliana l,s 

2-(L-rhamnopyranosyloxy)benzyl 2-[(OH)3C6H8O]OC6H4CH2 Reseda odorata f 

4-(L-rhamnopyranosyloxy)benzyl 4-[(OH)3C6H8O]OC6H4CH2 Thlaspi perfoliatum s A

4-(4’-O-acetyl-L-rhamnosyloxy)benzyl Moringa peregrina s  4-[(4’-CH3CO)(OH)2C6H8O]OC6H4CH2

2-phenylethyl C6H5(CH2)2 gluconasturtiin Armoracia lapathifolia r 

r 

Brassica campestris (B. rapa) r 

Brassica juncea l, s, r Brassica oleraceae s 

r 

Brassica napobrassica r Brassica napus s 

l l, s, r r 

Reseda media s,l Sinapis alba (B. hirta) r 

3-phenylpropyl C6H5(CH2)3 Armoracia lapathifolia r 

80
4-phenylbutyl C6H5(CH2)4 Armoracia lapathifolia r 

2-hydroxy-2-phenylethyl C6H5(OH)CHCH2 glucobarbarin Barbaria vulgaris s 

2-(4-methoxyphenyl)-2-hydroxyethyl Arabis hirsuta s  [4-CH3OC6H4](OH)CHCH2 s 

2,2-dimethyl-2-(4-methoxyphenyl)ethyl Pentadiplandra brazzeana r 

 [4-CH3OC6H4](CH3)2CCH2

2-(L-arabinopyranosyloxy)-2-phenylethyl Sesamoides canescens l 

 C6H5[(OH)3C5H6O]CHCH2

3-indolylmethyl C8H6NCH2 glucobrassicin Brassica campestris (B. rapa) s 

l,r l 

Brassica napobrassica r Brassica napus s 

l l, s, r r 

Brassica oleraceae l 

r 

Camelina alyssum s 

Lepidium graminifolium s 

Raphanus sativus l 

4-hydroxy-3-indolylmethyl 4-OHC8H5NCH2 4-hydroxyglucobrassicin Brassica napobrassica r 

Brassica napus s 

l, s, r r 

Brassica oleraceae s 

r 

1-methoxy-3-indolylmethyl neoglucobrassicin Brassica campestris (B. rapa) s 1-(CH3O)C8H5NCH2 Brassica napobrassica r 

Brassica napus l 

81
s 

r r 

Brassica oleraceae l 

r 

Cochlearia officianalis s 

4-methoxy-3-indolylmethyl 4-methoxyglucobrassicin Brassica napobrassica r 4-(CH3O)C8H5NCH2 Brassica napus l 

l, r r 

Brassica oleraceae l 

r 

Lepidium sativum s 

1-acetyl-3-indolylmethyl 1-(CH3CO)C8H5NCH2 Tovaria pendula s 

1-sulphonate-3-indolylmethyl glucobrassicin-1-sulfonate Isatis tinctoria s  1-(OSO3)C8H5NCH2

a s = seed, young seedling, or inflorescence, l = leaf and/or stem, r = root, b identification changed to E. virgatum 

1973), c tentative identification. 

82
Table 2. Effects of Aqueous Extracts from Glucosinolate-Containing Tissues on Other Plants 

________________________________________________________________________ 

Plant Inhibition Stimulation or no effect 

____________________________________________________________________________

Seedling growth 

Brassica campestris barley (Hordeum vulgare)

vetch (Vicia sp.)

radish (Raphanus sativus)

wheat (Triticum aestivum)

Brassica juncea wheat

 (seed tissue) tomato (Lycopersicum

esculentum) 

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

morningglory (Ipomoea 

 purpurea) 

wheat

Brassica nigra Bromus rigidus

Bromus mollis

Avena fatua

wheat

Brassica oleracea radish lettuce

lettuce (Lactuca sativa)

wheat

Camelina sativa flax (Linium usitatissimum)

Seed germination 

Brassica juncea 

 (seed tissue) 

alfalfa (Medicago sativa) carrot (Daucus carota) 

 (seed tissue) cabbage (Brassica oleracea) pepper (Capsicum frutescens) 

83
 (seed tissue) radish timothy (Phleum pratense) 

 (seed tissue) turnip (Brassica rapa) tomato

Brassica oleracea wheat

___________________________________________________________________________

References: Mason-Sedun and Jessop 1988; Jimenez-Orsornio and Gleissman 1987; MasonSedun et al. 1986; Leather 1983; Bell and Muller 1973; Kutacek 1964; Gressel and Holm

1964; Patrick et al. 1963; Grummer and Beyer 1960. 

84
Table 3. Influence of Naturally Occurring Isothiocyanates on Microorganisms 

________________________________________________________________________ 

Organisma Concentration Evaluation criterion 

 μg mL-1 [M] 

____________________________________________________________________________

Methyl ITC 

 _______________

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

 100 68x10-5 biomass reduced 93% 

P. shermanii 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. graminis 0.1 10x10-7 minimum for inhibition in hsb

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 

85
Ethyl 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 

Allyl ITC 

 ________________

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

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

E. coli 36 36x10-5 delay growth 36 hc

E. coli JCM-1649 0.034 34x10-8 100% growth inhibitionb

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 inhibitionb

S. aureus 1 10x10-6 minimum for inhibition 

 36 36x10-5 delay growth 33 h 

 0.11 11x10-7 100% growth inhbitionb

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

86
B. 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 inhibitionb

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

G. graminis 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. piriformis 150 15x10-4 minimum for inhibition of germ.

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

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-Methylthiopentyl 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 

87
P. 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-Methylsulfinyl-3-butenyl ITC 

 ____________________________

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

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

M. piriformis 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. 

Benzyl 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 sp. <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 

88
P. palmivora 1490 10x10-3 lethal 

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

 194 13x10-4 100% growth inhibition 6d 

Phenylethyl 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. graminis 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-Hydroxybenzyl ITC 

 ___________________

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

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

M. piriformis 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 Aphanomyces eutiches 

Bacillus cereus Aspergillus alliaceus 

Escherichia coli Aspergillus niger 

Propionibacterium petersonii Botrytis allii 

Propionibacterium shermanii Botrytis cinerea 

Proteus vulgaris Colletotrichum circinans 

Pseudomonas fragi Cytospora 

Pseudomonas aeruginosa Fusarium

Salmonella enteritidis Fusarium oxysporum

Sarcina lutealis Fusarium solani 

Staphylococcus aureus Gaeumannomyces graminis 

Gibberella saubinetii

Monila sitophila

Monilinia laxa

Mucor piriformis

Penicillium chrysogenum

Penicillium cyclopium

Penicillium expansum

Penicillium glaucum

Phytophthora palmivora

Rhizopus oryzae

Rhizopus stolonifer

b

 headspace concentration 

c

 as compared to control 

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

Fenwick et al. 1983; Tang and Takenaka 1983; Rutkowski 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. 

91
Table 4. EPA Registrations for Various Isothiocyanates 

EPA Registration Number Type Isothiocyanate 

499-136 repellent-vertebrates propenyl

499-137 repellent-vertebrates propenyl

499-138 repellent-vertebrates propenyl

731-15 repellent-vertebrates propenyl

731-22 repellent-vertebrates propenyl

2139-55 Fungicide, insecticide, miticide methyl 

2139-76 Fungicide, insecticide, miticide, nematicide, 

herbicide 

methyl 

3008-77 fungicide methyl

5332-7 repellent vertebrates, herbicide propenyl 

5332-8 repellent, herbicide propenyl

61966-4 insecticide, miticide, repellent propenyl 

61966-5 insecticide, miticide, repellent propenyl 

61966-7 repellent-vertebrates propenyl

67064-3 repellent-vertebrates propenyl

70160-1 insecticide, miticide, repellent propenyl 

70160-2 repellent-vertebrates propenyl

70160-3 repellent-vertebrates propenyl

70160-4 insecticide, miticide, repellent propenyl 

92
O S

H

OH

OH H

H

HO

H C

N OSO

R

CH OH 2 3

H

Figure 1. Generalized structure of glucosinolates 

93
NO O

S

R C

S 3

-

D Glucose

R N C S

O H N

S

R

SCNFe2 +

low pH

R C N + S

R S C N

CH2 CH (CH 2) N C

S

n

Epithionitrile

Nitrile

Thiocyanate

Isothiocyanate

Oxazolidinethione

Terminal

unsaturation

2-Hydroxy

p-Hydroxybenzyl

Indole

Figure 2. Glucosinolate degradation pathway1

 1 Enzymatic hydrolysis results in production of glucose followed by HSO4

-. At least seven other 

products have been observed. Oxazolidinethione and SCN- are thought to be formed from unstable 

R-N=C=S intermediates.

94
Sinapis arvensis

n=9

SS

Sinapis alba

n=12

DD

B. nigra

n=8

BB

B. juncea

n=18

DD

B. carinata

n=17

BB CC

B. oleracea

n=9

CC

B. napus

n=19

AA CC

B. rapa

n=10

AA

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

95
F1146-E(12/2004) 

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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 

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University of Idaho 

Moscow, Idaho 

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XC0-9-29095-01 

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NREL 

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13. SUPPLEMENTARY NOTES

NREL Technical Monitor: K. Shaine Tyson 

14. ABSTRACT (Maximum 200 Words)

Plants may produce compounds that directly or indirectly affect their biological environment. These compounds fall 

within a broad category of compounds called allelochemicals, 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 pest management systems. Using plant-produced allelochemicals in agricultural 

and horticultural practices could minimize synthetic pesticide use, reduce the associated potential for environmental 

contamination, and contribute to a more sustainable agricultural system. 

15. SUBJECT TERMS

glucosinolate; allelochemicals; pest management; pesticide 

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