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LIBRARY

This is to certify that the

thesis entitled
RESPONSE OF CUCUMBERS (Cucumis sativus L.),
SNAP BEANS (Phasequs vulgaris L.), and NEEDS
TO COVER CROP RESIDUES AND RYE (Secale ceneale L.)

ALLELOCHEMICALS
presented by

William Robert Chase

has been accepted towards fulfillment
of the requirements for

MS degree in Hortichtur‘e

Major professor

DateWO

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MSU Is An Affirmative ActionlEquel Opportunity Institution

 

 

Response or CUCUMBERS (Cucumis eativa L.). SNAP BEANS
(EEO—”$9.! m L. ). mo NEEDS TO covsn CROP RESIDUES

AND RYE (Secale cereale L.) ALLELOCHEMICALS

by

William Robert Chase

A Thesis

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

Master of Science

Department of Horticulture

1990

ABSTRACT

RESPONSE or CUCUMBERS (Cucumis sativa L.), SNAP BEANS

 

(Phaseolus vulgaris L.), AND NEEDS TO COVER CROP RESIDUES
AND RYE (Secale cereale L.) ALLELOCHEMICALS
By

William Robert Chase

Experiments were conducted in 1988 and 1989 to determine the

effectiveness of spring planted oats (Avena sotiva L.), spring barley

 

(Hordeum vulgare L.), and rye for weed control in cucumbers and snap
beans. Although rye produced 20 to 64$ less biomass than cats or
barley, its residues provided superior weed suppression (up to 79%)
compared to ru: residues. Cucumber and snap bean yields were increased
in the presence of the residues in 1988 (a dry year) and slightly
reduced by the residues in 1989 (a wet year).

Microbial transformation of .2(3H)-benzoxazolinone (BOA) an
allelochemical from rye to 2,2'-oxo—1,1’azobenzene (AZOB) occurred
rapidly in four Michigan soils. The soil microbe capable of this

transformation was isolated and identified as Acinetobacter

 

calcoacetius. AZOB, 2,u—dihydroxy-1,4(2H)—benzoxazin-3-one (DIBOA) and
BOA toxicities were assessed singly and in combination on cucumbers,
snap beans, and weed indicators. A208 was generally more toxic to

weeds and less toxic to the craps than DIBOA and BOA.

DEDICATION

This thesis is dedicated to my wife,
Kathy, for her love, patience, and
unceasing understanding; and to my sons,
Nathan and Nicky for providing a
reference point and enduring throughout
these stressful years. Finally to Dr.
Alan R. Putnam for introducing me to
allelopathy and providing me guidance
when he would rather have been at the end

of a fishing pole in Montana.

iii

ACKNOWLEDGMENTS

The author expresses sincere graditude to his major professor, Dr.
Alan R. Putnam for providing the oppountity to advance my knowledge
in Weed Science. Sincere thanks is extended to Dr. Muraleedharan
Nair tku~ his expert laboratory assistance throughtout the research.
Appreciation is also given to Dr. Hugh Price and Dr. Karen Renner for
their assistance as guidance Committee members.

I would also like to acknowledge the technical assistance and
friendship I received from Ms. Ruth Post and Ms. Deborah Thorogood
thoughtout my research. As "square meters and bioassays" is the
bottom line in allelopathy research, I owe many thanks to Les, Jill,
Steve, and Rhonda for their time and energy spent on tedious, but
important measurements. Without their help I would not have been
able to conduct this research.

Finally I would like to acknowledge the Department of
Horticulture for allowing me the opportunity to do ”double duty"

while I was obtaining my Master of Science degree.

iv

Table of Contents

LIST OF TABLES ................................................... vii
LIST OF FIGURES .................................................. viii
LIST OF APPENDICES ............................................... ix
CHAPTER 1 - LITERATURE REVIEW
Introduction ................................................ 1
Conservation Tillage ........................................ 2
Effects of Tillage Systems on Warm - Season Vegetables ...... 6
Allelopathy ................................................. 10
Crop and Weed Response to Allelochemicals ................... 12
Role of Soil Microbes in Allelopathy ........................ 16
Compounds Implicated in Rye Toxicity ........................ 18
Detection of Benzoxazinones and Azoperoxide ................. 22
LITERATURE CITED ................................................. 23

CHAPTER 2 - FIELD AND GREENHOUSE RESPONSE OF CUCUMBERS AND SNAP BEANS
T0 COVER CROPS RESIDUES

ABSTRACT ......................................................... 29
INTRODUCTION ..................................................... 30
MATERIALS AND METHODS
Field Experiments ............................................ 34
1988 ...................................................... 34
1989 ...................................................... 35
Controlled Atmosphere Experiments ............................ 36
Rate and Placement of Residue ............................. 36
Leaching of Residues ...................................... 37
RESULTS AND DISCUSSION
Field Experiments ......................................... 37
Controlled Environment Experiments ....................... #4
Conclusions ............................................... 51
LITERATURE CITED ................................................. 56

Table of Contents Continued
CHAPTER 3: IDENTIFICATION OF THE SOIL MICROBE RESPONSIBLE FOR
THE TRANSFORMATION OF BOA To AZOB AND OUANTIFICATION
OF AZOB IN FIELD SOILS

 

 

ABSTRACT ......................................................... 59
INTRODUCTION ..................................................... 60
MATERIALS and METHODS
General Experimental ..................................... 63
Soil Collection .......................................... 63
Sterilization of Soil .................................... 64
Preparation of Media ..................................... 64
Isolation of Soil Microbes ............................... 65
Separation of Total Soil Microbes into Bacteria and Fungi. 65
Identification of the Microbe ............................ 67
Spray Reagents for TLC Detection ......................... 67
Production and Extraction of A208 from Soil .............. 67
Production of A208 in Liquid Media ....................... 67
Microbial Transformation of BOA to A208 .................. 68
Quantification of BOA and A208 in the Soil Extract ....... 68
In Vitro Transformation studies with Pure
Cultures of Acinetobacter calcoacetius ............... 69
Statistical Methods ...................................... 70
RESULTS and DISCUSSION
Bacterium Isolation ....................................... 70
Spectra Analysis of BOA and A203 .......................... 71
Microbial transformation .................................. 71
Conclusions ............................................... 88
LITERATURE CITED ................................................. 94

CHAPTER 4 - RELATIVE TOXICITY 0F RYE ALLELOCHEMICALS T0 WEEDS AND
VEGETABLE CROPS

ABSTRACT ....................................................... 96
INTRODUCTION ................................................... 97
MATERIALS and METHODS
Spray Reagents for TLC Detection .......................... 99
Source of Allelochemicals ................................. 99
Extraction of DIBOA ....................................... 99
Bioassay setup ............................................ 101
Incubation of Bioassay .................................... 102
Statistical Methods ....................................... 102
RESULTS and DISCUSSION
DIBOA Quantification ...................................... 102
Relative Biological Activity of Allelochemicals ........... 104
Conclusions .............................................. 111
LITERATURE CITED .............................................. 113
vi

LIST OF TABLES

CHAPTER 2:

Table 1: Total Weed Biomass as Influenced by Cover Crops

and Time for 1988 and 1989. ............................. 38
Table 2: Decomposition of Cover Crops During 1988 and 1989 ........ 41
Table 3: Cucumber and Snap Bean Yields as Influenced by Cover

Crops and Weeding in 1988 and 1989 ....................... 42
Table 4: 1989 Temperature and Soil Moisture Comparisons Under

Different Cover Crop Systems ............................. 45
Table 5: 1988 and 1989 Cucumber and Snap Bean Plant Stands as

Influenced by Cover Crop Residue ......................... 46
Table 6: Snap Bean Germination and Growth as Influenced by

Placement of Rye and Poplar Excelsior Residues ........... 50
Table 7: Cucumber Germination and Growth as Influenced by

Placement of Rye and Poplar Excelsior Residues ........... 49

Table 8: Cucumber Germination and Seedling Growth as Influenced by
Periods of Leaching and Microbial Degradation of Rye or

Poplar Excelsiors ........................................ 52
CHAPTER 4.

Table 1. Barnyardgrass root and shoot response to DIBOA, BOA,
and A208 applied singly and in combinations ............ 105

Table 2. Garden Cress germination response to DIBOA, BOA, and
A208 applied singly and in combinations ................ 106

Table 5. Garden Cress root and shoot response to DIBOA, 80A,
and A208, applied singly and in combinations .......... 107

Table 4. Cucumber root and shoot response to DIBOA, BOA, and
A208 applied singly and in combinations ................ 108

Table 5. Snap bean root and shoot response to DIBOA, BOA, and
A208 applied singly and in combinations ................ 110

vii

CHAPTER 1:

Scheme

CHAPTER 3.

Scheme
Figure
Figure
Figure
Figure
Figure

Figure

Figure

Figure

Figure

Figure

CHAPTER 4:

Scheme

10.

LIST OF FIGURES

Scheme for formation of A208 ........................... 21

Isolation, Purification, and Identification of a soil

Microbe Responsible for BOA Transformation to A208 ..... 66
UV spectrum of 2,3-benzoxozolinone (BOA) in MeOH ........ 72
UV spectrum of 2,2’,oxo—1,1’-azobenzene (AZOB) in

MeOH .................................................... 72
UV spectrum of a "fixture of BOA and AZOB (1:1) in

MeOH .................................................... 74
UV absorbance standard curve for varying

concentrations of 80A ................................. 76
UV’ absorbance standard curve for vorying

concentrations of AZOB .................................. 78

Rate of transformation of BOA to AZOB in sterile

soil incubated with Acinetobacter calcoacetius.
Concentration of BOA was 0.1 mg/g soil .................. 81
Rate of transformation of BOA to AZOB in non—sterile

soil incubated with BOA concentration of 0.1 mg/g

of soil ................................................. 83
Rate of transformation of BOA to A208 in non-sterile

soil incubated with BOA concentration of 1.0 mg/g

 

of soil ................................................. 85
A208 concentration in £1 Michigan soils incubated
with 0.1 mg BOA/g soil .................................. 89
BOA concentration in II Michigan soils incubated
with 0.1 mg BOA/g soil .................................. 91

Extraction of DIBOA from 11 day-old greenhouse
grown rye .............................................. 103

viii

APPENDIX 1.

APPENDIX 2.

APPENDIX 3.

LIST OF APPENDICES

Analysis of soil at time of planting for 1988 and 1989 .... 115
Bi-Monthly temperature and precipitation information

for 1988 and 1989 ....................................... 116
Soil laboratory Analysis on four Michigan field soils ..... 117

ix

 

LITERATURE REVIEW

Introduction
At present, there is an increasing concern over monitoring

agriculture’s input and management practices. Since the Environmental
Protection Agency’s (EPA)’S 1984 report to Congress stating that 90%
of the States have identified agriculture as being a nonpoint source of
pollution, the public has been more vocal about what they are willing
to accept as suitable agricultural practices.

0n the other hand, the desire for an increase in crop productivity
has lead farmers to use more inputs such as energy, nutrients, and
pesticide inputs ix: their practices which in turn may adversely impact
the’ natural environment. Increasing energy inputs is usually
correlated with using more tillage practices. This exposes the land to
increasing sediment runoff which carries nutrients and pesticides into
the water systems. Phipps and Crossian (1986) estimated that between
50 to 70% of all nutrients reaching surface waters are principally
nitrogen and phosphorous that originate from agricultural land. A 7-
year study conducted by the National Stream Quality Accounting Network
and the National Water Quality Surveillance Systems evaluated 116
stations for nitrate concentrations and reported 23% to have increasing
nitrogen levels. They associated these increases directly with
agricultural activity.

In 1987, the United States Department of Agriculture reviewed
EPA's priority pesticides to identify pesticides with possible water
pollution qualities according to their usage, amount applied each year,

and a description of their mobility in the soil. 0f the pesticides

?

listed, the majority (88%) were herbicides which were labeled "leacher"
by water and therefore had the potential to enter the water system.
Hallberg (1987) found that in Iowa and Minnesota the herbicides
detected at the highest levels were those applied to the largest
agriculture acres and were leachable herbicides in the triazine and
acetanilide classes. Hall et al. (1989) also reported detecting
simazine (6-chloro-N-N’—diethyl-1,3,5—triazine-2,4-diamine) and
atrazine (6-chloro—N-ethyl-N'-(methy1ethyl)-1,3,5-triazine-2,4-diamine)
at low levels but was unable to detect metolachlor (2-chloro-N-(2-
ethyl-6-methylpheny1)-N-(2-methy1ethyl)acetamide) and cyanazine (2-[[4—
chloro-6—(ethylamino)-1,3,5-triazin-2—yl]amino]-2methylpropanenitrile)
in any wells. However, he also noted that under no-tillage (NT)
conditions herbicide leaching through a well-drained soil matrix was
more pronounced. One could speculate that with increased public
concern for groundwater contamination coupled with the data available
on nutrient leaching and pesticide contamination, safer methods for
food production will be demanded by the public.
Conservation tillggg

During the ”dust bowl” era of the 1930’s, public opinion was
awaken to develop ways to reduce soil erosion caused by wind at the
time of planting on exposed dry soils. With the formation of the Soil
Conservation Service in 1935 and the increased knowledge of ways to
conserve soil loss, NT was evaluated as an alternative cropping system.
Several workers have reported comparisons of conventional tillage (CT)
methods with NT methods. However in the early studies, there was a
problem in that there was no definitive description of NT. For

example, Webster’s strict definition would be no cultivation of the

land whatsoever whereas Young (1982) defines NT as "not disturbing more
than 10% of the surface area and having a protective residue uniformly
distributed over the soil surface the year around”. In Hall et al.
(1989), NT refers to an untilled soil and has no mention of a residue
being present, whereas Knavel et al. (1977 and 1986) and Blevin et al.
(1971) established their NT experiments in either planted ryegrass or
naturally occurring bluegrass. Still another modification is planting
into existing stubble after the harvest of the previous crop such as
wheat (Mascianica et al. 1986). More recently the planned introduction
of cover crop residue into the crop management scheme has provided an
additional approach to NT management (Putnam and Defrank, 1983; Smeda
and Weller 1986). This introduction of a cover crop residue fits under
the scheme put forth by Young (1982).

With the recurrence of "dust bowls days” in the 1950’s and
1980’s there has been renewed interest in using no-tillage to reduce
erosion from wind while at the same time improving water infiltration
and soil organic matter. It is difficult to measure soil losses due
to water erosion because it can vary with the soil type, slope of the
land, and the amount of cover present on the soil surface. For
example, Young (1982) found with conventional tillage when the slope of
the land increased from 3 to 12%, soil losses increased from 3.8 to
27.7 tons per acre per year. In contrast, a no-tillage system with 90$
ground cover on similar slopes last only 0.3 to 2.4 tons per acre per
year. Likewise, Hayes (1982) found that on a 5i slope soil loss could
be reduced by 12 tons per acre per year by adding one ton per acre of
plant residue. Knowing that 1 ton of soil loss per acre is equal to

1/150 of an inch of soil per year and that using plant residues in a

cropping system can greatly reduce erosion has made NT with plant
residue an attractive alternative for conservation-minded farmers.

Additional advantages from NT are improved water infiltration and
soil organic matter. Blevin et al. (1971) reported a 4 to 8% increase
in water content in the top 8.cm of soil when comparing no-tillage to
conventional tillage practices over a two year time period. Glenn and
Welker (1989) evaluated four soil management systems; i: herbicide
strip, cultivated strip, mowed sod, and killed sod and reported that
the sod treatments captured more rainfall than did the herbicide and
cultivated strips. This was attributed to the sod reducing the amount
of runoff and thus increasing the amount of water infiltrating the
soil. This finding led to renewed interest in how increasing water
content of the soil would affect soil temperature. Wall and Stobbe
(1984) showed that there was an average soil temperature decrease of
2°C iJI‘the top 5 cm in NT compared to CT with a surface residue. Also
Derpsch et al. (1986) compared four tillage systems for soil
temperature during the daytime and ranked them from the highest soil
temperature to the lowest at 3 and 6 cm depths. His rankings were in
the order of CT > chisel plow > NT > and permanent cover. Kladivko et
al. (1986) summarized the effects of conservation tillage as improved
soil water content, lower soil temperature, more organic matter, more
water-stable aggregates near the surface, and higher bulk densities
when compared to CT systems.

A specialized approach within NT systems has been the use of plant
residues of cereal grain cover crops. These residues can be beneficial
for improving soil properties but they have also been proven to be

beneficial in interfering with weed growth. Muller (1969) defined the

T

components tof plant interference to include competition for physical
factors such as light, water, and nutrients and also included indirect
effects by the release of breakdown products or chemicals derived from
decaying residues. Molisch (1937) had previously called the latter
allelopathy. It is important to understand what mechanisms of
interference are acting to predict impacts on crop growth. Putnam and
Duke (1974) and Fuerst and Putnam (1983) proposed (1 set protocol to
separate the competitive and allelopathic components.

Several books and reports describe the advantages and
disadvantages of NT for agronomic and horticultural crops (Phillips and
Young, 1973; Young, 1982; and Hayes, 1982). Perennial horticultural
crops lend themselves well to this cultural method (Putnam and Lacy,
1977; Putnam et al., 1983). However, annual vegetable crops have a
wide range of growing conditions and varying seed sizes which has
presented unique challenges for adopting them to NT cover crop systems.
As Putnam and Defrank (1983) generalized, usually large seeded crops

like corn (_Z_e_a mays L.), cucumber (Cucumis sativus L.), pea (Pisum

 

sotivum L.), and snap bean (Phaseolus vulgaris L.) will have an equal
or positive yield response when planted into spring cover crops of

barley (Hordeum vulgare L.), oats (Avena sativa L.), rye (Secale

 

cereale L.), sorghum (Sorghum bicolor L. Monench.), sudangrass (Sorghum

bicolor Sudanense (Piper) Hitchc.) and wheat (Triticum aestivum L.)

 

when compared to bareground . Small seeded vegetables like lettuce
(Lactuca sativa L.), cabbage (Brassica oleracea L.) and carrot (Qggggs
carota L.), generally showed a negative response, ‘while ‘tomato
(Lycopersicon esculentum Mill.) varied from test to test. This could

be due to the fact that larger seeded crops are planted deeper away

from the residues whereas small seeded crops are planted shallow in
close proximity to the toxic residues. Putnam and Defrank (1983)
confirmed their findings by planting cucumber, pea, and snap beans into
spring planted covers with no significant difference in yield when
compared to a conventional tillage system.
Effects of Tillgge Systems on Warm - Season Vegetables

There have been numerous studies conducted with cucumber, snap
bean, lima bean (Phaseolus lunatus L.), sweet corn, summer squash
(Cucurbita pepo L.), transplanted tomato and transplanted pepper
(Capsicum annuum L.) to further define their needs for light, water,
and nutrients in NT systems. Both Williams et al. (1973) and Teasdale
et al. (1983) evaluated snap bean row spacings from 12.7 to 91 cm and
found that as the row spacings were decreased the yield of the snap
beans increased and resulted in decreased weed growth. Teasdale et
al. (1983) concluded that as the rate of canopy closure increases fresh
weed weight decreases by 50 to 67% due to differential shading.
Teasdale suggests his maximum yields and best weed reduction was
obtained within the 15 to 36 cm row spacing whereas wider row spacings
did not intercept enough of the light allowing more earlier—season weed
growth. Their results suggested that for sweet corn, snap bean, and
onion (Allium cepa L.) there is a 2 to 3 week weed-free period needed
after crop emergence to prevent yield losses due to weeds. Mascianica
et al. (1986) double cropped snap beans after harvesting a wheat crop
and studied the effects that stubble heights of 8, 15, 23, 30,and 38 cm
had on the crop morphology and yields. His results suggested that the
shading by 30 and 38 cm-high wheat stubble reduced the yields of the

crop due to shading caused by a reduction of incident light on the

5-?

surface of the soil. He also noted that all stubble heights resulted
in etiolation of plant internodes and that stubble heights of 8 to 23
cm increased the total yields of snap beans. He also concluded that
stubble heights of 15 to 30 cm enhanced the harvest efficiency without
affecting the total yield. This was possible because the interference
of the stubble caused an increase in the elongation of the plant
internodes thereby causing the pod clusters to move upward thus
decreasing pod losses during mechanical harvesting. Mascianica et al.
(1986) also proposed that the stubble provided the snap bean plants
with support thus reducing lodging. Mullins et al. (1988) conducted
experiments to study the effect of cropping systems on snap bean pod
length. His experiments evaluated snap beans planted with 2 row
spacings of 0.46 and 0.92 m and in plots with 3 tillage treatments of
CT, NT, and rotary strip tillage. He concluded from the results that
none of the parameters studied had any effect on pod length and
therefore thought pod length to be strictly an individual cultivar
trait uninfluenced by environment. He also observed poor weed control
when a winter wheat cover crop was removed prior to planting,
particularly in the wider row spacing. This might suggest that the
type of interference the mulch provided was shading the soil which
would have been minimal to the crop in the 2-row spacing.

Generally under NT with cover crops there has been improved
moisture content of the soil. Mascianica et al. (1986) suggested that
the snap bean plant requirement for water is 180 mm per year and that
the growth and pad yields are similar under CT vs NT with adequate
water present. It is when there is a shortage of water which can occur

in a drier growing season that the NT systems have greater yields.

 

Knavel et al. (1977) concurs that there is a greater yield in plant
dry matter in the NT compared to CT in drier years but that CT has
equal or greater yields in wetter growing seasons. Peterson et al.
(1986) also observed that NT treatments generally have wetter and
cooler soils in the spring thus reducing yields due to limited early
growth and development.

Nutrient availablity to the crop has been shown to be
affected by different tillage systems. Mullins et al. (1980) concluded
that tillage did not affect the pH of the soil in the top 10 cm but NT
treatments had higher pH’s in the top 10 to 15 cm depth. Generally he
found under CT systems that the macro-nutrients nitrogen (N),
phosphorous (P), potassum (K), and calcium (Co) were distributed deeper
in the soil but plant uptake was unchanged when comparing CT and NT.
Mullin et al. (1980) found that P and K levels in the soil were not
affected by tillage at the 0 to 5 cm depth but were affected at depths
greater than 10 cm. However, tillage system had no effect on plant
uptake of P and K at any level. Knavel et al. (1977) reported that N,
P, K, and Ca availablity depended on the crop (cucumber, sweet corn,
tomato, pepper), nutrient, tillage system, and time of sampling. For
example, sidedressing with N caused an increase of N in the crap from
the first to second sampling date in NT cucumber, tomato, and corn.
Where there were significant differences between the N content of the
crops due to the cropping systems, it was found to be higher in the CT
system. When N differences were found later in the season, it could
possibly be related to soil moisture changes as reported by Larson et
al. (1970). Skarphol et al. (1987) evaluated snap bean response to N

in NT and CT systems using legume and cereal grain cover crops in 1984

and 1985. His CT treatments consisted of cover crop residues being
incorporated into the soil prior to planting. In 1985, he detected
less nitrate loss possibly because of less leaching and denitrification
and also due to a shorter interval between incorporation of crop
residues prior to planting. the also reported a 14% greater yield in
the NT vs CT treatments for both years and attributed that to an
increase in soil moisture levels. There was no advantage of applying
nitrogen when a legume cover crop was used like hairy vetch (V333

villosa Roth.) or Austrian winter-pea (Pisum sativum ssp. arvense L.

 

Poir.). Knavel et al. (1977) reported that NT crops of cucumber, sweet
corn, tomato, and pepper contain equal or greater levels of P when
compared to CT-grown crops. He and Blevins et al. (1971) offered the
explanation that there is usually greater P uptake in NT systems due
to an increase in soil moisture levels. Knavel et al. (1977) found K
levels in general were not influenced by cultural practices but in some
cases varied in cucumbers and peppers depending on sampling date and
cropping systems. He also found Ca uptake varied among the four
different crops tested, but Ca content was generally lower in NT early
in the season and equal to or greater than that in CT crops by the last
date sampled. Therefore, nutrient availability can vary with the soil
microclimate, tillage system, crops, and time interval sampled in the
growing season.

With the complexity of competitive interaction taking place
involving light, water levels, and nutrient availability it is
difficult to differentiate the responses that are strictly competitive
or allelopathic in field evaluations. Therefore, to determine the

differences, one needs controlled experiments in the greenhouse or

laboratory.
Allelopathy

In more recent years, research has focused on the chemical basis
of plant interference called allelopathy. Allelopathy was first
observed as early as 1832 by de Condolle when he reported ”soil
sickness" that reduced the productivity of the crops. Since then
several articles and books by Tukey (1969) , Rice (1971), Putnam and
Duke (1978), Rice (1984), Putnam and Tang (1986), and Putnam (1988)
have further detailed the development of allelopathy from being an
unexplained occurrence to its current status as a science. Generally
it has been acknowledged that Molisch (1937) first coined the term
allelopathy to mean a "biochemical interaction between all types of
plants including microorganisms " (producing either a detrimental or
beneficial response). Microbial influences may be the result of
phytotoxins produced by microorganisms themselves or the release of
phytotoxins when microorganisms degrade plant residues.
Allelochemicals are therefore compounds that are produced by higher
plants or microorganisms that can affect the growth of other plants.
Whittaker and Feeny (1971) and Rice (1984) further modified Molisch’s
definition of allelopathy to mean the overall mediation of one plant's
growth by another through chemical means. A major obstacle in proving
these allelochemical responses has centered on removing the competitive
component from plant-plant interactions and focusing only an isolation
of allelochemicals.

The diversity of plants has made it difficult to establish a set
protocol for the proof of allelopathy. Fuerst and Putnam (1983) have

developed a set of criteria to prove in a stepwise manner whether or

10

not an allelopathic response is present. Initially the symptoms of
interference must be identified in the natural state. The next step is
to deveIOp supportive assays that can examine factors which might
influence the primary source of toxicity, the method of release, and
the role of microorganisms in the expression of toxicity. To help
support and define the allelochemical response, it is useful to
evaluate several indicator species and record the injury symptoms.
Other major steps in the protocol call for chemical isolation and
identification. Also, it is important to study the release and
movement and to quantify the toxin as a proof that the allelochemical
response is present. This can be confirmed by adding the toxin back
into natural systems and monitoring plant reaction. Drost and Doll
(1980) suggested the importance of confirming the activity of the
isolated chemicals by further testing in field experiments. This
protocol is very expensive, time-consuming and may require cooperation
between scientific disciplines. It is difficult to experimentally
provide a clear cut analogy to what the actual reactions are in natural
ecosystems.

Allelochemicals are present in virtually all plants and can be
found in an array of tissues such as leaves, flowers, fruits, buds,
seeds, stems, and roots (Putnam and DeFrank, 1983). Putnam (1988)
indicated that even though natural product chemists may isolate and
identify new compounds from plants and microorganisms, it is important
to confirm if they are present and released in sufficient quantities to
have an allelopathic effect. Tang et al. (1986) has outlined a
trapping technique using a live plant in its habitat to collect root

exuduates and to evaluate their allelochemical activity. His system

11

uses (1 healthy plant growing in either a basaltic rock or silica sand
medium, continuously circulating nutrient solution and trapping of
chemicals on nonionic resins.
m and Weed Response to Allelochemicals

Ruiz—sifre and Ries (1983) planted sweet corn into 49 day old
sorghum residues which had been divided into treatments of roots,
shoots, and whole plants and compared them to bareground. Of these
treatments, the sorghum shoots showed a 15 to 45% stimulation of corn
seedlings whereas the root residues showed a 20 to 30% inhibition of
seedlings for the first seven weeks, after which growth was essentially
equal to the bareground regime. This perhaps indicated that after
death of the plant there is a release or breakdown of plant material to
form stimulators or inhibitors. In other research, Barnes and Putnam
(1983, 1986) evaluated Spring planted rye compared to bareground and
recorded 0 94¢ reduction in weed biomass. Barnes further partitioned
35 day-old rye into root and shoots and found shoots to be about twice

as inhibitory on germination of lettuce and barnyardgrass (Echinochloa

 

crus-galli (L.)Beauv.) as the roots. Tang et al. (1975) had previously
evaluated rye for the presence of 2(3H)-benzoxazolinone (BOA) and found
8 times more BOA in the shoots than in the roots. Barnes et al. (1987)
further partitioned the rye herbage into water soluble and insoluble
fractions (nut found the soluble fraction to be inhibitory to garden
cress (Lepidium sativum L.). This was in agreement with several other
investigators who have reported the. aqueous fraction of different
herbages to be active on germination (Harper and Lynch, 1982; Guenzi
and McCalla, 1967; Weston and Putnam 1986). Barnes et al. (1987)

proceeded further and partitioned the aqueous rye herbage fraction into

12

acetone, hexane, ethylacetate, diethyl ether, and chloroform fractions
and reported the diethyl ether fraction to be two to ten times more
active than the other organic fractions. In the diethyl ether
fraction, there were two active toxins isolated, 2,4—dihydroxy-1,4(2H)
benzoxazin—3-one (DIBOA) and BOA, polar and non-polar compounds,
respectively. Zungica et cu“ (1983) had previously evaluated rye for
the presence of DIBOA and found it to be present in both cultivated and
wild accessions. He also found that both hydroxamic acids, DIMBOA and
DIBOA, were present in higher concentrations in the younger leaves and
reached their highest concentration after four days. Argandona et al.
(1980, 1981) evaluated rye for hydroxamic acids and found them to be in
highest concentrations from 10—16 days after germination. Nair et al.
(1990) compared two different allelochemicals derived from rye (DIBOA,
80A) run: the microbial transformation product of BOA, 2,2'-oxo-1,1’-

azobenzene (A208), and found corn and soybean (Glycine max L. Merrill)

 

crops to be unaffected while barnyardgrass, redroot pigweed (Amaranthus

retroflexus L.), velvetleaf (Abutilon theophrasti Medik.), garden

 

cress, and tomato were highly sensitive to A208 at concentrations of 5
to 200 ug/ml. Barnes et al. (1987) found dicot species in petri dish
assays to be 30$ more sensitive to rye allelochemicals than monocots.
Putnam (1984) postulated that there can be a positive, neutral or
negative effect of allelochemicals depending on plant species. Ideally
it would be beneficial to be able to harness the positive effects for
crop growth while still maintaining the negative effects on a
particular pest (e.g. selectivity). Barnes and Putnam (1987) indicated
that in 0 NT rye cover cropping system there tends to be greater

activity against weeds near the surface and therefore one possible

13

selectivity scheme would be to place the seeds of a crop below the
active rye residue levels. Putnam and DeFrank (1983) used 7 different
spring planted cover crops prior to planting cucumbers, sweet corn,
peas, and snap beans and monitored good germination, growth, and yield
of these vegetable crops. The same parameters indicated poor response
of direct seeded cabbage and lettuce in the cover crop residues while
seeded tomatoes and carrots were also damaged by some residues. Almeida
(1985) tested 7 different winter cover crop mulches for fresh biomass
production and found large variations. In addition, the cover crops

lupine (Lupinus albus L.), wheat, rape (Brassica napus L.), radish

 

(Raphanus sativus), and oats had total weed biomasses of 300 to 1350
g/m2 with 83 to 100% being grasses, whereas rye and triticale (X
Triticosecale Wittmack.) allowed weed biomasses of 500 and 1100 g/m2
respectively with 64 to 77% being broadleaves. This suggested that
different cover crops are selective in their suppression of weed
growth.

Another possible selectivity mode would be to incorporate the
residue thoroughly with the soil or to plow down the residue.
Christian et al. (1985) compared the effects of the depth of straw
incorporation of three cover crops, wheat, barley and oats, on seedling
establishment. Using a control plot where surface mulch was removed by
burning, he determined that burying the residue 20 cm deep resulted in
a 29% reduction in seedling establishment caused by the cover crop
residues. All of these studies suggested that selectivity may be
achieved by variations in seed size, species type, depth of planting,
and by the choice of a cover crop.

Some workers have attempted correlations between the time

14

intervals of plant decomposition and phytotoxicity. Almeida (1985)
found that although there was decomposition with time, the residue
cover that had the greatest biomass throughout the growing season also
produced the greatest reduction in weed biomass. His ranking from
least to most weed biomass were oats < rye < rape < triticale < wheat <
and < lupine when evaluated at 118 days after desiccation. Smeda and
Putnam (1988) evaluated residues of fall—planted covers of barley, rye,
and wheat desiccated in the spring. They observed that 17 days after
cover crop desiccation, all three covers similarly reduced weed biomass
by 80-903 when compared to a bareground treatment. At 71 days after
desiccation the bareground and barley plots had the same weed biomass,
while wheat and rye still exhibited a 51 to 73% reduction in weed
biomass. Srivastava et al. (1986) evaluated the activity of rotting
wheat straw water extracts and found that the greatest reduction (35-
455) in wheat germination occurred between 8 and 15 days. He related
this toxicity to allelochemical components of the wheat straw because
the treatments cfl’ sterile and non—sterile extracts had essentially the
same results. Kimber (1973) compared rye straw under sterile verses
nonsterile conditions as well as green verses rotting straws. He found
extracts of four day old green straw to be more toxic than the rotted
straw which strongly inhibited wheat growth. Guenzi et al. (1967)
monitored wheat seedling germination following treatments of water
soluble extracts of vmeat, oats, sorghum and corn residues and found
that wheat and cat residues contained no water-soluble toxins after 8
weeks while corn and sorghum residues appeared to release toxic
components for up to 22 weeks. Harper and Lynch (1981) determined by

gravimetric analysis the major components of undegraded oat straw to be

15

he
of
pc

0".

Fe

31.

DE:

7.3% water soluble materials, 35.5% hemicellulose, 42.1% cellulose, and
13.5% lignin. They concluded that although there were seasonal changes
in straw decomposition, hemicellulose and cellulose appear to be
degraded at a fairly constant rate during any time frame measured.
They also observed that acetic acid, the most active allelochemical,
was produced under anaerobic conditions and it’s concentration declined
progressively during straw decomposition. The course of decline in
production of acetic acid parallelled the loss of dry"weight. of
hemicellulose and cellulose. They also concluded that the production
of acetic acid occurred until 21.5% of the straw remained as
polysaccharides. But in practice, no phytotoxic accumulations
occurred in the natural ecosystems once 50% of these compounds had been
decomposed which usually occurred within 50 days following desiccation.
Therefore as stated by Putnam and Duke (1978) and Rice (1984), if the
time interval is long enough, toxins can be liberated by residues
either by decomposition or by leaching with water to eliminate any weed
or crop growth reductions.
Role of Soil Microbes in Allelopathy

Allelochemicals can be released by such processes as
volatilization, root exudation, leaching, microbial transformation, and
decomposition of plant residues (Chapman and Lynch, 1983; Rice, 1984;
and Putnam, 1984). More recently, the involvement of microorganisms to
release phytotoxins from residues or their role in using the residue as
a substrate to produce secondary compounds has been studied (Nair et
al. 1990). One of the early scenarios involved the removal of old
peach trees from orchards and unsuccessful attempts to reestablish a

new peach planting. Proebsting and Gilmore (1941) postulated that upon

16

tree removal, either the roots left in the soil or breakdown products
from the roots were toxic to new trees. They found that combining
peach roots with the soil and planting peach seedlings into the mixture
caused severe inhibition. They proposed that the chemical amygdalin,
an allelochemical derived from the peach root bark was responsible for
the toxicity. However when tested by itself for peach seedling injury,
no activity was observed. But with the addition of the enzyme emulsin
which catalyzes the hydrolysis of amygdalin to glucose, benzaldehyde,
and hydrogen cyanide peach seedlings exhibited severe injury. Patrick
(1955) examined the involvement of microorganisms in the production of
toxic compounds from peach root bark and isolated many microorganisms
from the soil capable of hydrolyzing amygdalin. Based on this, he
proposed that the toxic effects of peach root bark in the soil were
caused by hydrogen cyanide and benzaldehyde produced by microbial
degradation of amygdalin.

Patrick and Kbch (1958) conducted tests on the effects of aqueous

extracts of decaying residues of timothy (Phleum gratense L.), rye,

 

corn, and tobacco (Nicotina tabacum L.) on respiration, oxygen uptake,
and growth of tobacco seedlings. They proposed a correlation between
acidity and toxicity of the decomposing aqueous extracts indicating
that as the residues decompose, the aqueous portion becomes more
acidic. However, they found that if the pH of the extracts were
adjusted to neutrality, the toxicity to tobacco seedlings was retained
without much change. Also they compared the toxicity of extracts of
decomposed residues to extracts of non-decomposed residues mixed*with
soil and confirmed the role of microbes in the toxicity. It appeared

that the plants themselves contained no compounds inhibitory to

17

respiration of tobacco seedlings, but that toxic compounds were formed
during microbial degradation of the residues. Chapman and Lynch (1983)
also studied microbial effects on the toxicity of decomposing plant
residues to barley root growth and concluded that the inhibition was
primarly from the chemical inhibitors.

In general, the total soil microbial population is estimated to be
in the range of 106 to 109 microorganisms per gram of soil (Atlas and
Bartha 1987). Of the microorganisms found in the soil, bacteria and
fungi make up about 90 to 95% of the population. Mishra and Kanaujia
(1968) indicated that studies of microbial populations in soil have
been done on a seasonal or monthly basis since the fluxes of the
population can change rapidly. Tiwari et al. (1987) indicated that
wetting of the soil increases the dissolution of soluble organic matter
and will reflect a rapid increase of bacterial populations during the
first 24 hours. Cochran et al. (1977) generally found no toxicity from
straw during periods of low moisture but within one week after
rainfall, lentil and wheat straws significantly inhibited root
elongation of wheat seedlings. He also observed that production of
toxins from straws was irregular and generally was greater in wet
weather with air temperatures in the range of O to 15°C. He postulated
that during dry periods there could be storage of toxins in the plant
tissue which are released when soil moisture levels increased.
Mishra’s and Cochran's results possibly indicate that with higher
moisture levels there is increased microbial activity resulting in the
production of more phytotoxins.

Compounds Imglicated in Rye Toxicity

As indicated earlier, the allelochemicals present in plants are

18

numerous (uni diverse in their chemical structures. Chou and Patrick
(1976) identified nine compounds in decaying rye residues which
included vanillic, ferulic, phenylacetic, 4-phenylbutryic, p-coumaric,
p-hydroxybenzoic, salicylic, o-coumaric acids, and salicylaldehyde.
These compounds were all found to be inhibitory to germination in
lettuce seed bioassays. Shilling et al. (1985) isolated phenyllactic
acid and hydroxybutyric acid from field grown rye residues and found
them to be inhibitory to redroot pigweed and common lambsquarters
(Chenopodium album L.) root growth at 2 mM concentrations. Barnes et
al. (1987) isolated two hydroxamic acids, DIBOA and BOA from rye
residues, and compared their toxicity to phenyllactic acid and
hydoxybutyric acid. Three monocotyledons and dicotyledons were assayed
at concentrations ranging from 67 to 267 ppm (uni monitored for
germination number, root, and shoot lengths. Among the four
allelochemicals tested there was variation in response of the three
species in the parameters measured. DIBOA and BOA were the most toxic
of the four chemicals to root growth of large crabgrass (Digitaria

sanguinalis (L.)Scop.), barnyardgrass, proso millet (Panicum miliaceum

 

L.), redroot pigweed, tomato, and lettuce at concentrations of 67 to
267 ppm. Queirolo et al. (1981) and Barnes (1981) have suggested that
the benzoxazinones are inhibitory to photophosphorylation in the
chloroplasts which interferes with electron transport. This is similar
to polycyclic urea herbicide activity which in turn is related to the
H-N-C-O grouping (Moreland and Hill (1963)). 80A contains this
chemical grouping and DIBOA has it in a reduced form. Therefore,
chlorosis was a symptom of injury by rye residues on several indicators

(Barnes and Putnam 1983).

19

Virtanen et al. (1957) isolated the cyclic hydroxamic acid, BOA,
from four day-old rye seedlings. It was extracted by boiling the plant
tissue with water and extracting the aqueous portion with diethyl
ether. His unsuccessful attempts to extract BOA from fresh tissue
suggested the presence of a precursor in the tissue. By boiling the
tissue he was able to inactivate all enzymes in the tissue followed by
the extraction of the glucoside precursor, GDIBOA. (Scheme 1). By
treating GDIBOA with a hydrolytic enzyme he was able to obtain the
hydrolysis products glusose and a second precursor, DIBOA. Hofman and
Hofmanova (1969) were able to extract the glucoside precursor from
intact plant tissue using liquid nitrogen. Upon injury to the plant
the glucosidases were released which caused rapid hydrolysis of the
glucoside to DIBOA. Similar scenarios for a methoxylated DIBOA have

been reported to occur in corn, wheat, Job's Tear (Coix lachrymas

 

jobi), and bears breech (Acanthis mollis L.) (Loomis et al., 1957;

 

Koyama et al., 1956; Wolf et al., 1985). These are the only known
instances/of 1,4 benzoxazinone production in higher plants although
their production by microoganisms is fairly common (Tipton et al.,
1967).

Nair et al. (1990) reported the _i_r_1 m production of the
azoperoxide, 2,2,‘oxo-1,1’-azobenzone (AZOB), when BOA was mixed with
soil. He also showed the benzoxazinones produced by rye, DIBOA and 80A,
could rapidly undergo microbial transformation to AZOB, (Scheme 1).
Although intact rye tissue has not been shown to produce AZOB by

itself, the addition of either benzoxazinone to the soil in sufficient

20

3:80

OH

DIBOA—91L;
R O O — glu

Enzyme Hydrolysis 1 OH
I

N a
oxaox
O OH
Chemical Rearrangement ’1 H
A N
>20
' 0
Microbial Metabolism l
n, o — 0 R2

AZOB

N:':N

Scheme 1. Scheme for formation of AZOB proposed by Nair et al . (1990)

'21

quantities has shown rapid production of AZOB. DIBOA and BOA were less
active than AZOB at concentrations of 67 to 266 ppm on garden cress and
barnyardgrass radical elongation. However, AZOB was 8 to 10 times more
herbicidally active than DIBOA and BOA when tested against several
indicator species at 67 ppm.

Detection of Benzoxazinones and AZOB

DIBOA, BOA, and A208 can be detected by thin layer chromatography
(TLC) and spray reagents and quantitative UV—visible spectroscropy
(Hietala and Virtanen, 1960; Nair et al., 1990). These compounds
gave different Rf values on TLC plates with CHCl3-Me0H (6:1) or CH2C12—
EtOAc (9:1) solvent systems and were detected using spray reagents such
as ferric chloride (FeCls) for DIBOA or ceric sulfate (CeSO4) for BOA
(Nair et al., 1990).

The objectives of this study were to 1) determine the
compatibility of spring planted cover crops to cucumbers and snap
beans, 2) to assess response of these crops and weeds to rye
allelochemicals, 3) to quantify the rate of transformation of
benzoxazinones, and 4) to isolate a soil microoraganism responsible for

the biotransformation of BOA to AZOB.

22

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Environment: Annual Policy Reviewu 1986. Washington, 0.0.: The

National Center for Food and Agricultural Policy, Resources for the
Future.

Proebsting, ELL”, and A.E. Gilmore. 1941. The relation of peach root
toxicity to the re-establishing of peach orchards. Proc. Amer. Soc.
Hort. Sci. 38:21-24.

Putnam, A.Ru, 1984c Allelopathic research in agriculture: past
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Putnam, AHR. 1988. .Allelachemicals from plants as herbicides. Weed
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Putnam, A.R. 1988. Allelochemicals: blessings or curses for
agriculture? Comments Agric. and Food Chemistry. 1(4):183-206.

Putnam, A.R. and J. Defrank. 1983. Use of phytotoxic plant residues
for selective weed control. Crop Prot., 2(2), 173—181.

Putnam ,A.R., and M.L. Lacy. 1977. Asparagus Management with No-
tillage. Research Report 339, Michigan State University. 1-10 pp.

26

Putnam, A.R. and W.B. Duke. 1974. Biological suppression of weeds.
Evidence for allelopathy in accessions of cucumber. Science. 18:370—
372.

Putnam, A.R., and W.B. Duke. 1978. Allelopathy in agroecosystems.
Ann. Rev. Phytopathol. 16:431-451.

Putnam, A.R., J. Defrank, J.P. Barnes. 1983. Exploitation of

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68:941-943.

Ruiz-Sifre, G.V., and S.K. Ries. 1983. Response of crops to sorghum
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Rice, E.L. 1971. Some possible roles of inhibitors in old field
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York.

 

Shilling, D.G., R.A. Liebi, and A.D. Worsham. 1985. Rye(Secale
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Tang, CLS., 1986. Putnam, A.R., and 0.8. Tang, eds. The Science of
Allelopathy: Continuous trapping techniques for the study of
allelochemics from higher plants. Chapt. 7:113-131.

 

27

Teasdale, J.R., J.R. Frank. 1983. Effect of Row Spacing on Weed
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Whittaker, R.H. and FLFL Feeny. 1971. Allelochemics: chemical
interactions between species. Science 171:757—770.

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Amer. Soc. Hort. Sci. 98(6):526-529.

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2,4,-dihyroxy-1,4-benzoxazin-3-one and its glucoside from Acanthus
mollis seeds inhibit velvetleaf germination and growth. J. Nat. Prod.
48:59.

Young, H.M.,Jr. 1982. No-Tillage Farming; No-Till Farmer, INC.
Brookfield, WI. 2 ed. 1-202 pp.

Zungica, G.E., V.H. Agrandona, H.M. Niemeyer, and L.J. Corcuera. 1983.
Hydroxamic acid content in wild and cultivated gramineae. Phytochem.
22(12):2665-2668.

28

Chapter 2

FIELD AND GREENHOUSE RESPONSE OF CUCUMBERS AND SNAP BEANS
TO COVER CROP RESIDUES

ABSTRACT
Plant residues (cover crops) have shown promise as a method of
reducing synthetic chemical inputs for weed control. Oats (Avena

sativa L.), spring barley (Hordeum vulgare L.), and rye (Secale cereale

 

L.) were spring planted and allowed to grow for 37 to 45 days before
chemical desiccation with glyphosate, N-(phosphonomethyl)glycine,

followed by planting of cucumbers (Cucumis sativus L.) and snap beans

 

(Phaseolus vulgaris L.). In 1988 and 1989, rye was the most effective
residue reducing weed biomass by 46 to 79% at 35 to 42 days after
planting although it produced 20 to 64% less biomass than the other two
cover crops. In 1988, a drier and warmer year, both vegetables
produced 28 to 59% higher yields with residues compared to bareground,
while in 1989, a wetter and cooler year, the crops grown in residues
showed a reduction in yield of 7 to 33%.

Environmently controlled experiments compared the placement of rye
and poplar excelsior (Populus tremuloides L.) residues placed at the
soil surface cu: 1.3, 2.6, and 3.9 cm depths. Cucumbers and snap beans
planted at 1.3 cm exhibited a 15 to 64% yield reduction depending on
the residue rate and placement. Leaching of rye residues with and
without microbial degradation for four and eight days removed
inhibitory activity against cucumber. However, rye residues that were
not leached provided a 64$ reduction in cucumber germination and 68 to

70$ reduction in seedling biomass when compared to poplar excelsior.

29

 

Chapter 2
INTRODUCTION

No-tillage (NT) has shown promise as a method of reducing
synthetic inputs into cropping schemes. The use of plant residues
(cover crops) in NT systems has been beneficial in reducing erosion,
improving soil moisture content, and interfering with weed growth.
Young (1982) and Hayes (1982) reduced soil erosion by utilizing both NT
and a cover crop with plant residues of 1 ton per acre. The reduction
in soil erosion can be attributed to soil cover which acts to absorb
some of the forces from the rainfall and to impede surface water runoff
that carries soil. Blevins et al. (1971) and Glenn et al. (1989)
generally found that covering the soil surface increased moisture
content in the top 8 cm and attributed this to either improved water
infiltration or reduced water runoff. Cover crops also insulate the
soil and prevent water evaporation. Various investigators reported
improved yields in drier years when decomposing plant residues were
present (Mascianca et al., 1986; Knavel et al., 1977; and Ruiz-sifre
and Ries, 1983). Peterson et al. (1986) reported that soil in NT plots
was generally cooler and wetter in the spring. Wall and Stobbe (1984)
attributed plant growth differences to soil maxima temperatures
throughout the growing season to be an average decrease of about 2°C
with the maximum occurring at about 1300 hours daily.

Another beneficial result from plant residues has been their
ability to interfere with weed growth. Interference includes
competition for physical factors such as light, water, and nutrients

plus chemical effects from the decaying plant material called

30

allelopathy (Muller, 1969). Putnam and Defrank (1983) and Barnes and
Putnam (1983) have attempted to duplicate the physical component of
interference in field settings with the introduction of poplar
(Populus tremuloides L.) excelsior (PE) mulch which degrades slowly.
When comparing the standard check (bareground) to PE there is usually a
higher weed biomass reported in the bareground. Several cover crop
residues however, have reduced weed growth more than the PE.

Williams et al. (1973) and Teasdale and Frank (1983) evaluated
different snap bean row spacings from 13 to 91 cm and generalized that
as the row spacing decreased, the yield of the snap beans increased
while weed growth decreased. Teasdale’s results showed a 50 to 70%
weed biomass reduction due to the shading with the best results
achieved at 15 and 36 cm spacings. His results on corn, snap beans,
and onions indicated the importance of a two to three week weed-free
period after crop emergence to prevent yield losses.

Mascianica et a1. (1986) evaluated the shading that could occur
from residues when planting into a wheat stubble. He concluded that
stubble heights of 30 to 38 cm reduced crop yield due to the reduction
in light recieved by the emerging crop. Although yield reductions
occurred as stubble height increased, fruit quality improved and a
higher percentage of the fruit was recovered at harvest.

Putnam and Defrank (1983) generalized that large seeded craps,

i.e. sweet corn (Zea mays L.), cucumber, peas (Pisum sativum L.), and

 

snap beans, had an equal or enhanced yield when planted into spring

planted cover crops of barley (Hordeum vulgare, oats (Avena sativa L.),

 

 

rye (Secale cerale L.), sorghum (Sorghum biocolor L.), sudangrass

(Sothum biocolor sudanense (Piper) Hitchc.), and wheat (Triticum

31

aestivium L.) compared to bareground. Conversely, yields decreased for

small seeded crops like cabbage (Brassica oleraceae L.) and carrots

 

(Daucus carota L.). Possible reasons for greater tolerance of large
seeded crops could be more vigorous seedlings able to withstand more
variablity in placement of the seed, or deeper planting which places
seed away from the toxic residues.

Plant parts vary in the potential to produce allelochemicals
(Putnam 1988). For example, Barnes and Putnam (1986) reported a 94%
reduction in weed biomass for spring planted rye compared to
bareground. Barnes partitioned 35-day-old rye into roots and shoots
and found shoot residues and extracts to be twice as active as those
from roots on lettuce and barnyardgrass germination. Furthermore,
several investigators performed aqueous partitioning of grass herbages
into water soluble and insoluble fractions and found more activity in
the soluble fractions (Barnes et al., 1987; Harper and Lynch, 1982;
Guenzi et al., 1967; Weston et al., 1986).

Several workers have correlated the time interval of cover crop
decomposition and resulting phytotoxicity. Almeida (1985) found covers
that had the greatest biomass throughout the growing season also
provided the greatest reduction of weed biomass. His ranking of weed
biomass suppression 118 days after desiccation was in the order oats >

rye > rape (Brassica nupus L.) > triticale (5 Triticosecale Whittmack.)

 

) wheat > lupine (Lupinus albus L.). Smeda and Putnam (1988) evaluated
fall planted covers of barley, rye, and wheat for weed control
following desiccation in the spring. At 17 days after cover crop
desiccation, all three residues were statistically equal exhibiting

between 80 to 90$ reduction in weed biomass when compared to a

32

bareground treatment. At 71 days after cover crop desiccation, the
bareground and barley residue had the same weed biomass while wheat and
rye still exhibited 51 to 73% weed biomass reductions. Similarly,
Stivastarva et al. (1986) evaluated water extracts of rotting wheat
straw and found the greatest reduction in wheat germination (35 to 45%)
after straw had rotted for 8 and 15 days. He attributed this toxicity
to allelochemical components of the wheat straw since the sterile and
non-sterile extracts had essentially the same results. Kimber (1973)
compared rye straw under sterile versus nonsterile conditions and green
versus rotting straws. He found cold extracts of four-day-old green
straw to have the greatest inhibition when compared to rotted straw.
Guenzi et al. (1967) monitored water soluble extracts of wheat, oats,
sorghum and corn residues on wheat seedling germination. He found
wheat and oat residues contained no toxic water-soluble toxins after 8
weeks but corn and sorghum residues were observed to have active
extracts for up to 22 weeks. Harper and Lynch (1982) determined that
seasonal changes in oat straw decomposition rates occurred but that the
presence of active allelochemicals parallels the decomposition of plant
residues.

In Michigan and much of the temperate zone, there is a three to
five week period when soils lay fallow prior to planting warm season
vegetables. Cucumbers and snap beans are generally planted after June
1 when soil temperatures reach 15 to 16°C and night temperatures stay
above 10°C. This provides an opportunity to utilize cover crops to
reduce soil erosion and early weed populations. The objective of this
study was to evaluate the response of cucumber and snap beans to three

spring planted cover crops, and to monitor reductions in weed biomass

33

obtained with these cover crop residues.

MATERIALS and METHODS

FIELD EXPERIMENTS. Experiments with five identical treatments were
conducted in 1988 and 1989 at the Horticulture Research Center, E.
Lansing, MI, on a Capac loam (Aeric Ochraqualfs, fine-loamy, mixed)
soil with 0 to 3% slope. Appendix 1 provides the soil analyses from
samples taken at the time of cover crop planting in early May of each
year. ’Heritage’ oats, ’Browers’ spring barley, and ’Wheeler‘ rye were
planted at rates of 143, 133, and 152 kg'ha’1, respectively using a
three-point-hitch, mounted (1.8 meter wide) Moore Unidrill. A control
mulch, PE, was also included for a comparison of the physical impacts
that (I mulch would produceu PE was applied at a rate of 3300 kg'ha"1
within two days after vegetable crop planting. Immediately after PE
application, :1 green plastic netting with square openings of 2.5 X 5.5
cm was laid on top of the mulch to secure it. For comparable purposes,
cover crop treatments were compared to conventional tillage and a slow
degrading mulch of poplar excelsior.

1988. On May 13, plots of 4.5 X 6.0 meters were arranged in a
randomized complete block design having it blocks and planted with the
three cover crops. At 37 days after planting (DAP), all treatments
received glyphosate at 1.5 kg'ha'1 and seven days later paraquat, 1,1'-
dimethyl-4,4’-bipyridinium ion, plus x-77 at 1.1 kg-ho“ plus 0.5% by
volume/volume (vVv). On June 23, ’Flurry (85$) plus Sumpter (15$)‘
cucumbers and ‘Bush Blue Lake 47’ snap beans were planted using 0 Heath
Universal Vacuum Precision Seed Planter. Two rows of each crop were

planted 60 cm apart with seed placement for cucumbers being 1.2 cm and

34

snapbean 0.8 cm within the row. PE was laid 1 DAP and secured with the
netting. Nitrogen fertilizer (55 kg'ha'1) in the for": of ammonium
nitrate» was applied to the soil surface 14 DAP with a hand held
applicator and irrigated with 0.6 cm of water. Throughout the growing
season, random square meter measurements of weed number and total dry
weight shoot biomass for cover crops and weeds were harvested at 21,
32, ("Hi 42 DAP. Cucumbers were destructively harvested on two dates
(44 and 48 DAP), while snapbeans were harvested at 69 DAP. For each
crop, fruit and plant weights were recorded.

1989. On May 2, plots (6.0 X 6.0 m) were laid out in a Latin Square
Design with 5 blocks and planted with the same cover crops and
vegetables as ix: 1988. ‘The plots were sprayed 45 DAP with glyphosate
at 1.2 kg'ha’1. On June 26, cucumbers and snap beans were planted
using the same planter, row spacing, seed spacing, and cultivars as in
1988 except one additional row of each crop was planted. Nitrogen
fertilizer (55 kg'ha'1) was applied to the soil surface at 13 DAP in
the same manner as in 1988. Soil cores were taken 3 DAP using a round
golf course borer (16.2 cm) at a depth of 10 cm to determine soil
moisture content gravimetrically. Soil temperatures were taken at time
intervals of 11, 12, and 15 DAP at a depth of 6.8 cm with a
WEATHERtronics soil thermometer (model 4466). Random square meter
measurements for cover crop shoot biomass were harvested at 10, 41, and
81 DAP and total shoot biomass and number of weeds at 35 and 81 DAP.
On August 1 and 2 the main plots (cover crops) were split into weeded
vs nonweeded. On August 14 all snap bean plots received an additional
hand weeding. Plant and fruit yields were obtained 47 DAP for cucumber

and 81 DAP for snap beans.

35

CONTROLLED ATMOSPHERE EXPERIMENTS: Experiments to study the effects
of placement and decomposition of rye residues on cucumbers and snap
beans where conducted in Conviron (model CMP3244) growth chambers.
Growth conditions in the chambers were 26/23°C, 50% relative humidity,
380 qu“"3s'1 light intensity, and a 16-hour photoperiod. Repeated
experiments were completed with identical treatments arranged in a
randomized complete block design with 3 blocks. Data was analyzed
using analysis of vaiance and means were separated using the LSD test.
Fall-planted Rye residue (’Wheeler', 225 day-old, 4 to 8 leaf, 6
to 11 tillers, 8 to 16 inches in height) stored as whole plants (-20
°C) was lyophilized and ground through a Wiley Mill with a screen size
of 2 mm. Rye and poplar excelsior (5 and 10 g) were placed at
intervals of 1.3 cm on the surface of 12.5 X 18.5 cm rectanglar
styrofoam flats. Spinks loamy sand (Psammentic Hapludalfs, sandy,
mixed, mesic) was added volumetrically using 200 ml of soil for each
1.3 cm of depth added. ‘Flurry’ cucumber and ‘Bush Blue Lake 47’ snap
bean were planted at a 1.3 cm depth in a configuration with 2 rows of 4
seeds per crop. After planting, all flats were placed in 15.5 x 22.5
cm aluminum pans filled with 300 ml of water. The duration of the
experiments was 14 days during which the flats were maintained with
daily surface watering of 100 ml for the first 7 days and 150 ml for
the remaining 7 days. Crops were harvested at the soil surface using a
razor blade to cut the hypocotyl. Plant numbers and total dry weights
of each crop were recorded.
Rate and placement of residue: Rye residue at rates of 5 and 10 grams
was compared to 10 grams of poplar excelsior placed at 0, 1.3, 2.6, and

3.9 cm depths. A no-residue treatment was also included to use as a

36

comparison to the mulch treatments. The data from three duplicate
experiments were combined for analysis.

Leaching of residues Duplicate studies were conducted in the some
environmental setting as previously mentioned using a randomized
complete block design having 3 blocks. Residues were leached by adding
silica sand (50 g) to the bottom of 230 ml styrofoam cups, punching 4
holes for drainage, adding 50 grams of silica sand, and then placing
the residues on top. Microbial decomposition was studied by mixing 10
grams of Spinks loamy sand soil to one set of residue treatments.
Water (200 ml) was added daily to leach the residue for 4 or 8 days.
Residues were then air dried for 24 hours at 30°C and reground to allow
uniform distribution over the cucumber seeds which had been planted 1.3
cm deep. Plants were allowed to grow for 14 days, then harvested by
cutting the plants at soil level. Germination numbers and total weights
were recorded for each treatment. Data from duplicate experiments were
combined for the analysis of variance.

RESULTS and DISCUSSION

Field Experiments. As indicated by the temperature and precipitation
information for the two years (Appendix 2), 1988 had 66$ less rainfall
than 1989 and 26$ more growing degree days (GDD). This data may
explain some of the differences in total biomass of weeds as well as
vegetable yields obtained during the two seasons. In 1988, there was
little weed biomass production at 21 DAP for any of the treatments
(Table 1). However by 42 or 35 DAP, in both years, weed biomass was
significantly reduced when plant residues were compared to no residue.
For both years, the early season weed biomass appeared to be reduced

more with cover crop residues than with PE indicating that they were

37

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38

exhibiting both competitive and allelopathic responses.

Weed biomass increases rapidly with time and as indicated by
Smeda and Putnam (1988). The suppression of *weeds by cover crop
residues decreased between 17 and 71 days after desiccation depending
on conditions. In both years residue treatments significantly reduced
the total weed biomass at 21 to 42 days after planting. This agrees
with previous results of Putnam and Defrank, 1983; Barnes and Putnam,
1986; and Smeda and Weller 1986. The difference between treatments
and years could be related to the different weed spectra, environmental
conditions, and possible concentrations of allelochemicals present from
residues. In both years, common purslane was the predominant weed
species, representing 77 to 78$ of the population in 1988 and 64 to 89$
in 1989 depending on the time sampled. In 1988, other predominante
weeds included redroot. and prostrate pigweeds (16$) and witchgrass
(29$). In 1989, 15$ of the population were miscellaneous weed species
with redroot pigweed (7$) being second in abundance.

With 1989 having more rainfall, it seems likely that
allelochemicals would be leached faster from the residues over a
shorter time interval. Also in 1988, the comparison of PE with cover
crop residues at 42 DAP indicated weed biomass was the highest with the
former treatment. This suggests that the cover crop residues are
supressing weeds other than by a competitive manner, suggesting
allelopathy. When comparing the total weed biomass production at 81
days in 1989 there was excessive weed biomass in all of the residue
treatments, none of which differed from the control. The effect of the
cover crop had been lost probably because the toxins had been leached

from the residues.

39

Total decomposition of the cover crops, at 42 or 81 DAP, was
between 62 to 81$ in both years while PE decomposed only about 15$ over
the duration of the experiments (Table 2). The order from least to
greatest biomass of cover crops was rye < oats < spring barley < PE.
The fact that weed suppression was not related to cover crop biomass in
1988 provides indirect evidence for allelopathy since the cover having
the least biomass had the greatest weed-suppressing activity for both
years.

The yield data of both plants and fruit over the two seasons
indicated several important diferences. Total cucumber plant weight in
1988 for all treatments was less than half that recorded for 1989
(Table 3). Generally higher fruit yields were recorded in 1988. Lower
cucumber plant weights in 1988 could be explained by the higher
temperatures and dryer conditions early in the growing season which
stressed plant growth. Still the plants in the residues were able to
recover and outproduce those in the bareground checks. The higher
fruit yield in 1988, compared to 1989, could be attributed either to
the plant residue retaining more moisture during the early part of the
1988 season or the enhanced phytotoxicity of the residues exhibited
during the early damp part of the 1989 season. These conditions could
have liberated more water soluble allelochemicals that could have
affected crop growth.

Snap bean plant yields were similar for both years but fruit
yields in 1989 were 60$ of those for 1988 in the bareground plots.
This difference could suggest that the plant residues in 1988 provided
a better growing environment by providing improved moisture earlier in

the growing season, while in 1989 there was excessive moisture during

40

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vwazn mwcwn
some demo. some domed
no<m1 neon Ao>nv 1mm: 11111111 mm 111111111 ummu uuuuuuuu mmauntnlnn
smog zoazmma sconce zoazmma
.............. Ao.3-dv---------- ----------Aq.:o-.81------
oanm dew.» Amm.u dew.» o.: ~.u A.m
m. mo-Aa< ~.m.a .mm.m ~lm.a o.o ~.u ..m
z<m ~dm.c dau.m Ndm.o du.o ~.u A.»
nacwas mxnopmwow mom.~ Ao:.m wom.m Au.u ~.m ~.¢
203m Am..: Amo.o dmd.m m.m ~.o ~.o
rmo .om mm.@ :.a
A v 3on3» lam.» .mo.a sacs» ~.m i.m

 

A u n <a~:o «as oo<o1 anon oqwoonm awe 2m.
m I a <a~=m «as so»: cameo" oq (cocoa <m soazmonoa Am m—c:_«poa:n an u-.oA.
u 1 w <o.:e «o1 oo<o1 osoam <m 2030 .w mmozpqwoazo av 1:0.oA.

43

planting time. This higher moisture level not only could have caused
more allelochemicals to be released in a shorter time, but may also
have caused lower soil temperatures unfavorable for bean growth.

In 1989, cucumber yield decreased 19 to 48$ when cover crop
residues were compared to the bareground treatments. This might be
explained by the residues causing either too moist a condition or
temperatures less than ideal during the early season (See Table 4).

The snap bean yield results for 1988 and 1989 show the some
reversals. It appears that the beans and cucumbers responded similarly
to conditions in soil moisture and temperature which might explain the
reduced yields in 1989. In 1989, temperatures were as much as 4°C
higher in bareground plots while moisture levels were up to 18$ higher
in the residue plots (Table 4). If the temperatures were limiting for
cucumber and snap bean growth, they could delay germination of the
plants which would have increased the exposure to toxic components from
the residues. Also the soils under higher moisture conditions might
have been firmer and less friable restricting emergence of dicot crops.
Early stand data for both cucumber and snap bean in 1989 reflected
poorer germination or emergence caused by the residues when compared to
bareground (Table 5). This can be seen when one examines the
progression of plant populations from 11 to 81 DAP. Early stands of
snap beans appeared to be affected less by the residues than the
cucumbers, although snap bean plant weights were about the same for
both years. Fruit yield was greatly reduced in 1989 either due to poor
fruit set or possibly not enough heat units over the same time frame to
produce yields equal to those of 1988.

Controlled Environment Experiments: When evaluating the placement of

44

«>arm a” Ammo qmauowancso can mOAH 3o»mn:wo ooanawumaam canoe Ouwwowoan
oo<me neon m<mno3m.

 

 

 

oo<m1 neon Ac>vv u AA An Am
- $5.3 .............. AcoAA ................
oaem A~.m wo.m u~.A uA.m
m. mo1Am< AA.m no.» No.u uA.o
=<o Au.u uo.m uA.A uA.m
uOUHas mxomHmwow AA.o uo.w u~.m uA.m
zone Ao.o u~.u um.o uA.o
rmo a.omv A.u .o A.» 2m

45

 

A a many «manmwancsom norm: canteen Auoo can Aaoo :ocwm an 0.0 as.

4>arm m" Ammo can Ammo ozocacoe one mace moo: wuazn mnazau an Asa—caaooa
a< oo<m1 neat zompaca.

>. ocncacee swoon mnaaa

 

 

 

Ammo Ammo.
oo<m1 anon Ao>uv a: :m AA Am mu
some zoztomq

.......................... Azo..3-~A-------------------
oanm u.u k.a u.m u.m m.a u.o
m. ma1~a< a.» e.: u.m u.o m.m a.q
=<o a.m e.A u.m ~.o m.o a.~
flanges mxnaAm»01 q.» m.u w.» u.A m.o a.m
zone o.o m.o m.m m.m m.o q.o
-ch.omA u.o ~.m A.m A.e zoos» m.~ u.o

 

A u n <a~=o «on oo<os neon oqaooam can 93o Annosaonpo: awe 2m.
m : a <a~=e qoe so»: oqqoon om (cocoa <w nonsmoama Am mAcaquoaan an a . .oA

46

4>arm mu Ammo and Ammo ococaaoe and mean was: awaan mnozam am uzmcuoaooa
c< oa<o1 neon zomuaco.

m. mzac moo: unann meaaa

 

 

 

Acme AoeoA
oo<o1 neon Ao>vv om AA Am 0A
soon zoazmon
.............................. Azo..3-~A--------1-------

oanm m.m m.q w u Ao.q Q.m
m. aae—a< m.q m.m m.m 6.9 m.m
=<a m.u :.m m.v o.m q.u
10n~a1 mxnawmmos o.m w.u m.A AA.~ Ao.A
zone m.o o.m Ao o AA.¢ Ao.m
rm: A.omv . ~.m w.» A.u zmaaw Ac.» o.»

 

A n n <aAco was oo<os axon oqnmonm aza azo Annmaannuo: awe zm.
m u n <a~=o «as so»: oqqoon as cocoon <m :ozzooao: Am munapqpnazn an a u .oA

47

residue in order to eliminate crop injury, the residues generally need
to be positioned at a distance of 1.3 cm or greater from the seed to
eliminate toxicity (Tables 6 and 7). When rye or poplar excelsior
residues were placed on the soil surface or 1.3 cm depth with the
seeds, snap bean emergence was reduced compared to when the residues
were placed at 2.6 or 3.9 cm depths. However, emergence in all residue
treatments was better than that acheived with bareground. Reduced snap
bean seed germination might be attributed to fluxes in soil moisture
whereas with the residue present the soil moisture content would be
more constant.

Snap bean seedling weights were reduced only when the rye and
pOplar excelsior were placed on the soil surface at the 10 gram rate.
Shoots were more sensitive than roots to the placement of residues when
comparing the 0 and 1.3 cm depth when compared to the 2.6 and 3.9 cm
treatments. Cucumber germination was reduced when rye residues were
placed with seeds at both the 5 and 10 gram rate, while both rates
either at the soil surface or at 1.3 cm depth significantly reduced the
growth of cucumber seedlings. This suggests that a certain level of
rye is needed to liberate enough toxins to reduce cucumber growth. At
the 5 gram rate the residue must be in close contact with the seed to
produce this effect. Also, with the soil surface treatment being the
most active, and shoots being the most sensitive, one might predict
that residues placed on the surface would have the greatest impact on
germinating seeds.

Rye and PE residues (+ or - microbes) were compared 4 and 8
days following leaching. Non-leached rye residues exhibited a 61 to

64$ reduction in cucumber germination and a 66 to 71$ reduction in

48

4>mrm m“ maau moo: newspaanno: can aware: om Haqpcoaooa ex tuaooaoan
on z<o can vanwas mxooAonw zoapacoa

 

 

 

z<o am z<o um
coup: Aoav m a Ao u A0 a m c Ac u Ao c
-- --+--Azo..qu-AA .................. Adonaw ac.qu-AA .......
o ~.A M.» u.~ mam mmfl um:
A.u :.u m.A m.u moA mu: Anuo
~.m u.u m.a :.u VVA Amuu mum
u.o :.A :.m was Ao~u
no s¢m~a=o A.m uAu
HMO A.cmv A.V uou

 

A 1 Dana sensomozn amaze «nos assoc exams—amznm can: :a<»:c nzsea cuoorm.

49

dawn. q“ ococacoa cowanaanuoa can Quote: on Haawcozooa a< v aooaoan
. 0‘ a<o one venues mxocumpos nom»n=ou .

 

 

 

z<o pm z<o am
noon: Anav m c Ao a Ao c m m Ac c A0 a
..... Azo..uoq-AA------ -1--------Aqono~ an.uoquAA------

o :.o ~.a m.m awn Aum «an

A.u :.u u.w N.u mum mmu bmm

~.m m.A m.q N.m rum :Au mm:
u.o N.A V.m muo uvo

:o seepage q.u mam

rmo A.omA . A.am AAm

 

A a cane wonsomoan amaze «was nzwoo extenuaoanm can: =a<pau nzsoo cwoorm.

50

seedling growth when compared to the PE control mulch (N‘ residues
leached for l: to 8 days (Table 8). This suggests that compounds toxic
to cucumbers are present in nonleached rye and could be released
rapidly upon injury of the plant. The rapid reduction in toxicity of
the leached residues suggests that these compounds are relatively water
soluble. Snap bean assays were variable and inconclusive either due to
nonuniform seed germination or a poor growing environment.

Conclusions: There are several environmental factors including
moisture, temperature and desiccation time which can influence the
activity of decomposing plant residues. As these results and others
suggest, there appears to be a time interval of about 25 days after
desiccation when residues can greatly reduce weed growth (Barnes and
Putnam, 1983; Smeda and Putnam, 1988). Harper: and Lynch (1981)
suggested that there are seasonal rate changes in straw decomposition,
and under anaerobic <n~ high moisture situations. greater phytotoxic
activity occurs. This data suggested that after the plant residues are
decomposed to 35 to 70$ of the orginal weight, no weed suppressing
activity remains. As observed in 1989, and previously reported (Glenn
and Welker, 1989; Blevins et al., 1971) plots with residues had higher
soil moisture contents which could have caused toxins to be liberated
quickly. Since 1988 was a drier year, the release of phytotoxins could
have been delayed under low'moisture conditions and thus remained at a
toxic level over a longer period of time (Cochran et al., 1977). The
environmental differences which occurred during the two years of our
studies could help explain the difference in crop and weed response to
allelochemicals. For example, temperatures are generally lowered under

ITT system (Wall and Stobbe, 1984; Derpsch et al., 1986; Peterson et

51

doowo on ococaaos nossuaanuo: ooo mooowpom owotn: om »:wwcooooo o< oosnoom
ow Hoonzpan oao apowocpop ooowooonAoz on s<o os oooaos oxoonmpoa sompocmm.

 

 

 

oo<m a<o A um z<o om
:uuazo..oansAvu:n snucaao.oonuAvnuua
o ~.m q.m m~ nmo
a . u.m w.m ~u~ may
a u.A u.m woo wuu
rmo A.omv o.oa mm
A 1 333 w <35 «2. + on .. 333.com tom 2m. oono oouona 326333
Soon on ooacuooo oono. >HH onroa nonosoonpoom (use zm.

52

al., 1986) which could explain the slower germination of seeds and
lower yields that were obtained in 1989. The greater sensitivity of
shoots to cover crops under slower germination conditions could also
have contributed to the reduced yields in 1989. The mechanisms of
interference occurring under field experiments are difficult to
separate (Fuerst and Putnam, 1983). The cover crops used here have
been reported to have chemical activity, but in the field, this
undoubtedly interacts with other competitive components of
interference. Alemeida (1985) related the greatest chemical activity
of plant residues with those that had the greatest biomass production.
In contrast, our results indicated that the residue with the most
activity was rye which also had the least residue biomass. Therefore,
Alemeida’s results may have reflected light, water, or nutrient
competition effects more than allelopathy.

The majority of weeds present in these studies were broadleaved
species. Alemeida (1985) suggested different residues can be selective
for either grass or broadleaved weeds. Barnes and Putnam (1986), and
Shilling et al. (1985) have reported rye residues to be more active on
dicot than monocot species. This occurred in our 1988 research (a
drier year), while in 1989 rye was the only residue producing a
significant reduction of dicot weeds compared to a bareground control.
Knowing that dicots are more sensitive to residues than monocots makes
it imperative to test dicot crops for safety under a variety of
environments. Results of Putnam and Defrank (1983) and these results
indicate that larger seeded crops are less effected by residues. This
could be because larger seedlings are more vigorous or because of the

placement of the seed below the greatest allelochemical concentration

53

zones. When Christian at al. (1985) removed the residues thxxn the
surface by burning, he removed 29$ of the toxicity observed on the crop
was also removed.

The reduction in both snap bean and cucumber weight when the
residues were placed on the surface can be explained by light
interference as well as allelopathy and can be separated to some extent
by the use of PE. 'These results indicate that shoots are» more
sensitive than roots to rye and the zone of greatest crop activity is
in the upper 1.3 cm of soil. This also explains the reduction in weed
biomass with cover crop residues compared to PE since the majority of
weed seeds germinate in this 1.3 cm zone.

As Putnam and Duke (1978) and Rice (1984) indicated, if the time
interval is long enough, toxins can be liberated from residues and
either be decomposed or leached to eliminate any' weed and crop
reductions. Harper and Lynch (1982), Alemeida (1985), Smeda and Putnam
(1988), Srivastava et al. (1986), Guenzi et a1. (1967) have all
reported residue phytotoxicity at different time intervals. It is
apparent that this time interval difference is related to the different
crop microclimates affecting leaching and the decomposition of the
residues. Ruize-sifre and Ries (1983), Knavel et al. (1977),
Mascianica et al. (1986), Teasdale and Frank (1983), and Williams et
al. (1973) have all shown positive responses to plant residues when
growing NT snap beans under normal to dry growing conditions. If a
vigorous crop can be established and quickly produce a canopy to reduce
the light penetrating to the soil surface, further weed reductions are
apparent. Tiwari et al. (1987) suggested that under drier conditions

there is little toxic substance release from plant material. If this

54

is the case, the reduction in weed biomass earlier in the season can be
attributed more to shading while the later response is due to the
allelochemicals being released. Also in wetter growing seasons,
residues have the potential to cause poorer crop germination, reduced
growth and lower yields either by modifying soil conditions or

releasing more phytotoxins which in turn inhibit crop growth.

55

LITERATURE CITED

Almeida, F.S. de. 1985. Effect of some winter crop mulches on the soil

weed infestation. Proceedings 1985 Brit. Crop Prot.Conf., Weeds.
2:651-695.
Barnes, .JJ’. and A.R. Putnam. 1983. Rye residues contribute weed

suppression in no-tillage crapping systems. J. Chem. Ecol. 9(8):1045-
1057.

Barnes, J.P. and A.R. Putnam. 1986. Evidence for allelopathy by
residues and aqueous extracts of rye (Secale cereale L.) Weed Sci.
34:384-390.

 

Barnes, J.P., A.R. Putnam, B.A. Burke, and A.J. Aasen. 1987.
Isolation and characterization of allelochemicals in rye herbage.
Phytochem. 26(5):1385-1390.

Blevins, R.L., D. Cook, S.H. Phillips, and R.E. Phillips. 1971.
Influence of No-tillage on soil moisture. Agron. J. 63:593-596.

Christian, D.G., B.M. Smallfield, and M.J. Case. 1985. Straw disposal
on heavy clay soils. British Crop Protection Conference - WEEDS. 621-
628.

Cochran, V.L., L.F. Elliott, and R.I. Papendick. 1977. The production
of phytotoxins from surface crop residues. Amer. J. Soil Sci. Soc.
41:903-908.

Derpsch, R., N. Sidiras, and C.H. Roth. 1986. Results of studies made
from 1977 to 1984 to control erosion by cover crops and no-tillage
techniques in Parana, Brazil. Soil and Tillage Research. 8:253-263.

Glenn, D M., and W.V. Welker. 1989. Orchard soil management systems
influence rainfall infiltration. J. Amer. Soc. Hort. Sci. 114(1):10-
14.

Guenzi, W.D., T.M. McCalla, and F.A. Norstadt. 1967. Presence and
persistance of phytotoxic substances in wheat, oat, corn, and sorghum
residues. Agron. J. 54:164-165.

Harper, S.H.T., and J.M. Lynch. 1982. The role of water soluble
components in phytotoxicity from decomposing straw. Plant and Soil.
65:11-17.

Hayes, W.A. 1982. Minimum Tillage Farming. No-Till Farmer, INC,
Brookfield, WI. pp.1-167.

Kimber, R.W. 1973. II. The effect of time of rotting of straw from
some grasses and legumes (M: the growth of wheat seedlings. Plant and
Soil. 38:347-361.

56

Knavel, D.E., J. Ellis, and J. Morrison. 1977. The effect of tillage
systems on the performance and elemental absorption by selected
vegetable crops. J. Amer. Soc. Hort. Sci. 102(3):323-327.

Mascianica, M.P., H.P. Wilson, R.F. Walden, T.E. Hines, and R.R.
Bellinder. 1986. No-tillage snapbean growth in wheat stubble of varied
height. J. Amer. Soc. Hort. Sci. 111(6):853-857.

Muller, C.H. 1969. Allelopathy as a factor in ecological processes.
Vegetatio. 18:348-357.

Peterson, K L., 14A]. Mack, and D.E. Booster. 1986. Effect of tillage

on sweet corn development and yield. J. Amer. Soc. Hort. Sci.
11(1):39-42.
Putnam, A.R. 1988. Allelochemicals: blessing or curses for

agriculture? Comments Agric. and Food Chemistry. 1(4):183-206.

Putnam, A.R. and J. Defrank. 1983. Use of phytotoxic plant residues
for selective weed control. Crop Protection, 2(2), 173—181.

Putnam, A.R., and W.B. Duke. 1978. Allelopathy in agroecosystems.
Ann. Rev. Phytopathol. 16:431-451.

Ruiz-Sifre, G.V., and S.K. Ries. 1983. Response of crops to sorghum
residues. J. Amer. Soc. Hort. Sci. 108(2):262-266.

Rice, E.L. 1984. Allelopathy. 2nd edition. Academic Press, INC., New
York.

Shilling, D.G., R.A. Liebl, and A.D. Worsham. 1985. Rye(Secale
cereale.) and Wheat(Triticum aestivum L.) nmlch: The suppression of
certain broadleaved weeds. and the isolation and identification of
phytotoxins. The Chemistry of Allelopathy: 243-271 pp.

Smeda, R.J., and A.R. Putnam. 1988. Cover crop suppression of weeds
and influence on strawberry yields. Hort Sci. 23(1):132-134.

Smeda, R.J., and 8.0. Weller. 1986. Weed suppression with rye in
transplanted tomatoes. North Central Weed Control Conf. Proc.,
Abstract. 41:27-28.

Srivastava, P.C., N.G. Totey, and 0.M. Prakash. 1986. Effect of straw
extract ("1 water absorption and germination of wheat(Triticum aestivum
L. variety RR-21) seeds. Plant and Soil. 91:143-145.

 

Teasdale, J.R., J.R. Frank. 1983. Effect of row spacing on weed
competition with snapbeans (Phaseolus vulgaris), Weed Sci. 31:81-85.

Tiwari, S.C., B.K. Tiware, and R.R. Mishna. 1987. The influence of

moisture regimes on the popluation and activity of soil microorganisms.
Plant and Soil. 101:133-136.

57

Wall, D.A., E.H. Stobbe. 1984. The effect of tillage on soil
temperature and corn (Zea mays L.) growth in Manitoba. Can. J. Plant
Sci. 64:59-67.

Weston, L.A., B.A. Burke, and A.R. Putnam. 1986. Isolation,
characterization, (nu! activity of phytotoxic compounds from quackgrass
(Agropyron repens (L.) Beauv.). J. Chem. Ecol. 13(3):403-421 (1986)

Williams, C.F., Crabtree, 6., Mock, H.J., Laws, W.D. 1973. Effect of
spacing «m1 weed competition in sweet corn, snapbean, and onion. J.
Amer. Soc. Hort. Sci. 98(6):526-529.

Young, H.M.,Jru 1982. No-Tillage Farming. No-Til Farmer, INC.
Brookfield, WI. 2 ed. 1-202 pp.

 

58

CHAPTER 3
Identification of the Soil Microbe Responsible for the Transformation
of BOA to AZOB and quantification of AZOB in Field Soils

ABSTRACT

Nutrient agar plates enriched with fungicidal and bactericidal
compounds were used to isolate and identify a soil microbe responsible
for the biotransformation of 2(3H)-benzoxazolinone (BOA) to 2,2’-oxo-
1,1’-azobenzene (AZOB). Experiments were conducted to evaluate the
transformation process of BOA to AZOB in the presence of this microbe
under sterile and non-sterile conditions. The quantification of
unreacted BOA and AZOB in field soil was achieved by UV-visible
spectroscopic methods. The yield of AZOB isolated from field soil
showed (1 3 fold increase over reported values. Transformation studies
with soils inoculated with Acinetobacter calcoacetius indicated that
the production of AZOB increased linearly with the concentration of BOA
in sterile soils and showed a quadratic trend in non-sterile soils.
This also established that all soil types studied fix: the
transformation experiments contained either Acinetobacter calcoacetius
or another organism capable of the conversion of BOA to AZOB.

Conversion of BOA to AZOB occurred rapidly in all four soils studied.

59

CHAPTER 3

INTRODUCTION
Numerous allelochemicals have been isolated and identified from
rye (Secale cereale L.) herbage. Chou and Patrick (1976) isolated nine
compounds from decaying rye residues and characterized them to be
vanillic, ferulic, phenyllacetic, 4—phenylbutyric, P-coumaric, P-
hydroxybenzoic, salicylic, O-coumaric acids, and salicylaldehyde. All

nine compounds were found to inhibit lettuce (Lactuca sativa L.) seed

 

germination. Shilling et al. (1985) isolated phenyllactic acid and
hydroxybutyric acid from field grown rye and observed the inhibitory
activity on redroot pigweed (Amaranthus retroflexus L.) and common
lambsquarters (Chenopodium album L.) for germination and root growth.
Barnes et al. (1987) isolated two hydroxamic acids, 2,4-dihydroxy-
1,4(2H)-beenzoxazin-3-one (DIBOA) and 2(3H)-benzoxazoline (BOA), from
rye residues, compared their toxicity to phenyllactic acid and
hydroxybutyric acid and found that the hydroxamic acids, DIBOA and BOA,
exhibited the highest toxicity to both monocots and dicots.

Virtanen et al. (1957) reported the presence of BOA in 4-day old
rye seedlings. The benzoxazinones are proposed to occur in plants as
glucosides and upon injury are released to DIBOA by glucosidase enzymes
(Hofman and Hofmanova, 1968; Zungica et al., 1983). The quantities of
benzoxazinones in the plant have been shown to vary with the species
(Zungica et al., 1983), age (Argandona et al., 1981) and depend on the
part of the plant used (Tang et al., 1975; Barnes et al., 1987). More
recently, Nair et al. (1990) reported the production of an azoperoxide,

2,2’,oxo-1,1'-azobenzene (AZOB) in field soils incubated with BOA.

60

They also showed that the benzoxazinones produced by rye, DIBOA and
BOA, could rapidly undergo microbial transformation to AZOB. This new
allelochemical AZOB exhibited more inhibition to garden cress
(Lepidium sativum L.) (a dicot) root elongation at all rates studied

(67 to 250 ppm) whereas for barnyardgrass (Echinochloa crus-galli L.)

 

(a monocot) more inhibitory activity occurred at 67 ppm. The higher
rates produced toxicity on barnyardgrass similar to the two
benzoxazinones.

Allelochemicals (xx: be released into the environment by processes
such as volatilization, root exudation, leaching, microbial
transformation, and release of toxins from decomposition of plant
residues (Chapman and Lynch, 1983; Rice, 1984; Putnam, 1984). The
involvement of microorganisms to either release or produce phytotoxins
from residues or use the residue as a substrate to produce bioactive
compounds has been well documented. For example, Proebsting and
Gilmore (1941) reported that with soil microbes and peach tree roots
present there was severe inhibition in growth of young peach trees
planted in the peach tree roots-soil mixture. Initially they suspected
that the chemical amygdalin leached from 'the tree roots and *was
responsible for this inhibition. However applying this compound to the
soil did not inhibit the growth of peach trees. Later, Patrick (1955)
examined the involvement of microorganisms in the production of
toxicity from peach root bark and isolated several microorganisms
capable of degrading amygdalin in the soil. Based on these tests, he
proposed the toxic effects of peach root bark in the soil were caused
by hydrogen cyanide and benzaldehyde produced by microbial

decomposition of amygdalin.

61

Investigation of the crops containing benzoxazolinones for their
potential germicidal, insecticidal, and herbicidal activity involved
quantification of these compounds and studies of their transformation
in the environment. Quantification of hydroxamic acids in plant
material or soil was achieved earlier by four methods; tracing CM
isotopes, direct measurement of benzoxazolinone at (a specific
wavelength, spectrofluorometry, and by the use of colorimetric reagents
such as FeC13 or CeSOu (Long et al., 1974; Barnes et al. 1987).
However, the spectroscopic measurement cfl’ benzoxazolinones and
azoperoxide is a convenient method to quantify these compounds in
plant (N‘ soil extracts. By monitoring the optical density (00) of the
benzoxazolinones or azoperoxide, at their A max in MeOH solutions gives
accurate concentrations of these compounds in soil or plant extracts.

Rye is considered an important cover crop in Michigan, other parts
of the United States, and around the world (Putnam et al., 1984; Smeda
and Weller, 1986). Allelopathic effects and weed control activity of
rye residues were attributed partially to DIBOA and BOA (Barnes et al.,
1987). Discovery of AZOB in the field soil containing BOA and its
enhanced allelopathic activity over BOA and DIBOA was additional
chemical support for the strong allelopathic activity demonstrated by
rye residue (Nair et al., 1990). Therefore, further research was
conducted an the transformation of BOA to AZOB in field soils. The
objectives of this study were a) to isolate and identify the soil
microbes capable of transforming BOA to AZOB and b) to study the rate
of transformation of benzoxazolinones by this soil microbe in Michigan

field soils.

62

«71-

1. I

MATERIALS AND METHODS

General experimental: Ultraviolet (UV) and visible absorption spectral
analysis were performed on a Gilford Response II ultraviolet
spectrophotometer. BOA was obtained commercially from the Aldrich
Chemical company, Milwaukee, Wisconsin and was uniformly mixed with
soil using a mortar and pestle before adding water, and vortexing (1
min) on a Scientific Products 88220 Deluxe Mixer at 50 cycles per
second. Column chromatography was performed using a J.T. Baker Flash
Column (20 X 2.7 cm, 250 ml). The silica column used was (Analtech
sovent, 35 to 75 micron particle size, 60 Angstrom pore size) and the
flow rate was 3 ml/min unless otherwise specified. All TLC (250
micron) and preparative TLC were done on Anspec plates (1500 micron).
Homogenization of microbial colonies was done on a Brinkman Polytron
(CH-6010) blender for five minutes.

Soil collection. Spinks loamy sand soil (Psammentic, Hapludalfs,
sandy, mixed, mesic) was collected (November, 1987) from the
Horticulture Research Center field, located on Jolly road in East
Lansing, Michigan. Soil was stored in a cubic yard bin at room
temperature (15-22 0C) for one year prior to use. Three other soils,
Kalamazoo sandy loam (Typic Hapludalfs, fine-loamy, mixed, mesic),
Metea loam (Arenic Hapludalfs, loamy, mixed, mesic) and Oshtemo sandy
loam (Typic Hapludalfs, coarse-loamy, mixed, mesic) were collected in
October and November of 1988 and stored in the same conditions
previously mentioned. The Kalamazoo Sandy loam soil was collected at
the Clarksville Research Station located in Clarksville, MI, Oshtemo
sandy loam from the Sodus Horticulture Research Farm located in Sodus,

MI, and the Metea loam was collected from the Moore Seed Farm located

63

in Elsie, MI. Each of these three sites had two adjacent sampling
areas, one with a gramineae crop grown the same year and another with
no crap for two consecutive years. All soils were seived using an
aluminium sieve (2 X 12 mm opening) prior to conducting the
experiments.

Sterilization of soil: Spinks loamy sand soil was sterilized by
weighing soil into Erlenmeyer flasks and autoclaving (1 h, 15
atmosphere, 120 °C) for 4 successive days. Distilled water used for
the experiments was sterilized by autoclaving (20 min., 120 °C) in 250
ml erlenmeyer flasks (100 ml) or in 50 ml test tubes, each containing
10 ml.

Preparation of media: Yeast maltose glucose (YMG) agar, potato
dextrose agar (PDA), and Nz-Amine agar were used for growth and
identification of soil microbes. YMG was prepared by mixing yeast (4
g), maltose (10 9), glucose (4 g), and bacto-agar (18 g) in distilled
water (1 L) and dispensed into 2 L erlenmeyer flasks (250 ml each) and
autoclaved (20 min, 120 °C at 15 atmosphere). The cooled media (50 - 55
°C, 20 ml) was poured into petri dishes (100 X 15 mm) and allowed to
settle at room temperature. The same procedure was repeated for PDA
agar (PDA 39 g/L) and Nz-Amine agar (NZ-Amine agar 3 g/L, bactoagar 18
g/L) plates. All plates were stored at room temperature.

Antibacterial plates were prepared by dissolving PDA in distilled
water and autoclaved as above. After cooling the medium to 50 - 55 °C,
a solution of chloramphenicol (100 mg dissolved in 10 ml ethanol) was
added and thoroughly mixed. Plates were then poured and stored at room
temperature.

Antifungal Nz-Amine agar plates were prepared according to the

64

procedure described for antibacterial plates with the exception that
Nystation (0.03 g; dissolved ix: 1 ml DMSO) and cycloheximide (0.5 9
dissolved in 10 ml acetone) were added to the cooling Nz-Amine agar.
Plates were poured and stored at room temperature.

Isolation of soil microbes: Spinks loamy sand soil (1 g) was suspended
in sterile physiological saline (9 ml) (0.85$ NaCl in distilled water,
w/v), vortexed (15 min) and allowed to settle (1 h). This soil extract
was tested for lg gitgg transformation of BOA to AZOB by incubating BOA
(10 mg) with sterile soil (100 9), according to the procedure published
earlier (Nair et al., 1990). Cultures were allowed to incubate in the
dark at 26 0C (96 h), and extracted with MeOH (50 ml X 3).
Biotransformation of BOA to AZOB by the soil extract was verified by
TLC comparison with the standard AZOB and by UV-visible spectroscopy.
Separation of total soil microbes into bacteria and fungi. Serial 10-
fold dilutions of the supernatant/soil extract*was prepared in sterile
distilled water. Soil extracts (1 ml each) were pipetted into ten
different autoclaved test tubes, diluted to 10 ml each by sterile
distilled water and ndxed well. Solutions from each tube (10 ul) were
lawned seperately onto YMG plates. Plates at dilution of 10'6 showed
the best separation of fungal and bacterial colonies. .A total of 9
individual colonies were further transferred onto YMG plates
separately and assayed with BOA for the production of AZOB. Colonies
found to be positive for transformation of BOA to AZOB were transferred
to Nz-Amine agar and PDA plates enriched :with antibacterial and
antifungal compounds, respectively, and incubated (72 h). Two fungi and
one bacterium thus obtained were assayed for the transformation of BOA

to AZOB. All fungi were found to be inefficient in the transformation

65

sci]

Sc:

Field Soil (1 g)
I
I 1. stirred with saline (10 ml)
I 2. decant
I
I

 

soil Extract ( transformation assay)

I
I 1. serial dilution in saline
I 2. transfer onto YMG plates
I
I
I

Fully grown microbial colionies (4 fungi and 5 bacteria)
(transformation assay)

I
I
I 1. Suspended in saline

I 2. transfer onto bactericidal and
I fungicidal plates

I
I

 

fungi (2) bacteria (transformation assay)
(negative for I
BOA --- AZOB) 1. bacterium positive for

I
I transformation assay
I 2. identification

I

I

 

Acinetobacter calcoacetius

 

Scheme 1. Isolation, purification, and identification of a soil
microbe responsible for BOA transformation to AZOB.

66

of BOA to AZOB however the bacterium was able to convert BOA to AZOB.
Identification of the microbe: The bacterium which converted BOA
to AZOB ‘was further purified an antifungal Nz-Amine plates.
Morphological and biochemical properties of the bacterium were
performed on a Viteck Auto Microbic System using Vitek GNT gram
negative identification cards.

Spray reagents for TLC detection. DIBOA was detected on thin-layer
plates with ferric chloride (FeCl3) spray reagent consisting of 5%
FeCl3 in 95$ ethanol, acidified with concentrated HCl. A spray reagent
consisting of 1$ ceric sulfate (CeSOu) in concentrated H2804 was used
to detect BOA.

Production and extraction of AZOB from soil. A commercial sample of
BOA (4 g) was mixed thoroughly with Spinks loamy sand (4 kg) with
distilled water (400 ml) and incubated (26 °c, 96 h) in a x 2 L
Erlenmeyer flasks. The soil was extracted with methanol (8 X 500 ml)
and filtered through a Buchner funnel using filter paper (Whatman No.
1). The extract was further filtered through a sintered glass filter
(fine, 4 - 5 micron) and the solvent was removed by rotary evaporation.
A preliminary TLC (toluene: ethylacetate, 5:4) of the crude extract
indicated the presence of unreacted BOA and AZOB. Separation of BOA
from AZOB was achieved by flash column chromatography (260 g, toluene:
ethylacetate, 5:4). The orange band eluted was collected and dried 13
39232 affording dark brown crystals of AZOB (1.18 9).

Production of AZOB in liquid media. Baffle-bottomed Erlenmeyer flasks
(500 ml) containing distilled water (100 ml) were autoclaved and cooled
to room temperature. BOA (80 mg) and freshly homogenized colonies of

Acinetobacter calcoacetius (5.3 X 1010 CFU, 1 ml) were added to the

67

water and incubated (26 °C) under shake and non-shake conditions. The
fermentation broth (96 I1 old) was extracted with EtOAc (3 )( 75 ml).
Shake and non-shake flasks were extracted with EtOAc separately. The
EtOAc layer from shake flasks ‘was dried over anhydrous M950“ and
evaporated _i_n mg (4 day). Purification of AZOB from BOA by TLC
(toluene: ethylacetate, 5:4) gave an orange band, free from BOA, and
was eluted with CHC13:MeOH (1:1) and dried _ifl M affording pure
cyrstals of AZOB (46 am). A similar procedure was followed for the
non-shake flasks but the yield of AZOB was not calculated.

Microbial transformation of BOA to AZOB: Experiments were conducted
to study the rate of BOA transformation by the naturally occurring
microbial population of Acinetobacter calcoacetius in field soil. BOA
(3 and 30 mg each) was mixed with Spinks loamy sand (30 g each)
containing distilled water (3.5 ml) in culture tubes (2.5 X 15.0 cm).
Sterile styrofoam plugs were used to cap the tubes which were then
wrapped with aluminum foil for the duration of the experiment. Culture
tubes were placed in the dark and incubated (26 0C) for periods of 1,
2, 4, 6, 12, and 24 days. At the end of the incubation period, the
soil was extracted with MeOH (3 X 150 ml). The extracts were dried
over anhydrous M9804 and the solvent was removed 1g ygggg. All dried
samples were stored at -20 °C.

Quantification of BOA and A208 in the soil extract: The samples were
analyzed spectrophotometrically for BOA and AZOB. UV-visible spectra
of BOA and AZOB in MeOH gave ll maximum at 273 and 432 nm, respectively
(Figures 1 and 2). In equal mixtures of BOA/AZOB, the maxima were
still distinct (Figure 3). Standard curves for BOA and AZOB were

prepared using known quantities of BOA (from 0.748 to 286.0 ug/100 ml

68

in MeOH) and A208 (from 1.51 to 96.402 ug/IOO ml in MeOH) and by
recording the absorbance at 273 and 432 nm, respectively, for BOA and
AZOB.

Standard solutions of each of the dried soil extracts stored at -
20 °C were prepared by dissolving the extract in MeOH in a 100 ml
volumetric flask. The absorbance was recorded for each solution.
Concentrations of BOA and AZOB in a given sample were obtained from the
standard curve (Figures 4 and 5).

In Vitro transformation studies with pure cultures of Acinetobacter

 

calcoacetius: Cultures of Acinetobacter Calcoacetius for

 

 

transformation studies were prepared from the stock cultures grown on
YMG plates and kept at 4°C. The cultures used in each test were
prepared by transferring a stored culture onto YMG plates followed by
incubation (72 h) before use.

Acinetobacter calcoacetius colonies from eleven fully grown plates
were pooled and suspended in 110 ml of sterile distilled water. A
portion of this stock solution (3.5 ml) was then added to each
treatment of BOA (3 mg) in sterile Spinks loamy sand (30 g) that was
placed in culture tubes (2.5 X 15.0 cm). All treatments were incubated
(26 °C) in the dark for 1, 2, 4, 6, 12, and 24 days and extracted with
MeOH (3 X 150 ml). These soil extracts were quantified for BOA and
AZOB by the spectroscopic procedure described earlier.

Microbial populations were estimated by monitoring the colony
forming units (CFU) of the stock solution. Ten-fold serial dilutions
were made in sterilized distilled water by transferring 1 ml of the
stock culture to 9 ml of distilled water. After uniform mixing, 100 ul

of each dilutions were placed onto YMG plates, sealed with parafilm and

69

incubated (26 °C, 24 h). The CFU of the stock solution was calculated
to be 5.3 x 101°.

Statistical methods. Data obtained in duplicated transformation
studies were subjected to analysis of variance and means were compared
with the least significant different (LSD) test.

RESULTS and DISCUSSION

Bacteriunl isolation: Initial purification of soil extracts containing
total microbial isolates were positive for the conversion of BOA to
AZOB. Partial purification of the total isolates into bacteria and
fungi afforded one bacterium and two fungi positive for the
tranformation of BOA to AZOB. However, final purification of the fungi
on bactericidal plates did not (show transformation of BOA and hence,
they were not investigated further.

The microbe capable of transforming BOA to AZOB in field soil was
identified as (1 gram negative rod-shaped bacterium with a tendency to
form coccoid bodies and grew well in a medium containing O.4$ peptone
and 0.02$ tryptophan, under aerobic conditions. This organism was
capable of using malonate as the sole source of carbon. Decarboxylase
tests of the bacterium were negative with lysine, ornithine and
arginine. Glucosei'was not fermented in the presence of specific
inhibitors, DP300 and p-coumaric and did not produce acid with
raffinose, sorbitol, sucrose, inositol, adonitol, rhamnose, arabinose,
or glucose. Also, it did not hydrolyze esculin or o-nitrophenyl-B-D-
galactopyranoside» Under aerobic conditions, it did not produce acid
with glucose, lactose, maltose, mannitol or xylose. Tests for the
production of H28, urease and plant indican were negative with this

organism. The aforementioned properties are in conformity with those

70

of Acinetobacter calcoacetius described in Bergey’s manual of

 

systematic Bacteriology (1984).

Spectra analysis of BOA and AZOB. Even though analysis of
benzoxazolines were reported earlier (Virtanen and Hietala, 1957; Long
et al., 1974; Nair et al., 1990), these methods are not convenient or
efficient to quantify BOA and AZOB present in soil extracts. The UV-
visible spectra of BOA (Figure 1) and AZOB (Figure 2) gave distinct
absorption maxima at 273 and 432 nm. Since BOA and AZOB gave
characteristic UV-visible absorption maxima, it is easy to calculate
the concentrations of these compounds from their mixtures by monitoring
the optical density (00). This was confirmed from the concentration
curves obtained for BOA and AZOB where OD was read at 273 and 432 nm,
respectivitly. Standard curves were prepared separately by plotting
each compound’s concentration against its OD. The curve obtained from
the mixtures of BOA and AZOB were identical to the curve generated by
plotting concentrations of pure compounds against its 00 (Figure 3).
Concentrations of AZOB and unreacted BOA in the various soil extracts
were obtained directly from the standard curve (Figures 4 and 5).
Microbial transformation. The percentages of AZOB isolated from in
m transformation experiments was greater than previously reported
(Nair et al., 1990). The modified extraction procedure of AZOB from
soil gave yields three times greater (29.8$ vs 9.1$) than previously
reported values (Nair et al., 1990). This could be attributed to the
new solvent system employed for the separation of AZOB from BOA. In
the reported procedure (Nair et al., 1990), HCOOH and NHAOH were used

as part of the developing solvent system and these may have degraded a

71

Figure 1. UV spectrum of 2,3-benoxazolinone (BOA) In MeOH

Figure 2. UV spectrum of 2,2',oxo-1,l'-azobenzene (AZOB) ‘
in MeOH

72

 

4.00

 

ADSORBQNCE

 

 

0.00
200.0

4.00

 

NANOMETERS

 

 

 

   

flISORlflNCE
l

 

 

 

 

I
NANCIETERS

73

F1gure 3. UV spectrum of a mixture of BOA and AZOB (1:1) in
MeOH

74

 

ABSORDGHCE

 

0.00
200.0

 

 

 

A i ‘
HRHOIETERS 500.0

75

Figure 4. UV absorbance standard curve for varying concentrations

of BOA

76

ABSORBANCE

 

 

 

 

 

_
_

mo 30 1.5 moo mwo woo
002823352 A828 3:

77

Figure 5. UV absorbance standard curve for varying concentration

of AZOB

78

ABSORBANCE

 

A.m00.
A.000..U
0.000....
0.000.H
0.5001
0.M00I1.

 

 

 

 

0.000

r _
q _

— p
q —

m0 #0 00 00 A00
OOZOmZHm>jOZ 30:00 3:

79

portion of the A208 isolated. The rate of production of A208 in field
soil was investigated under environmently controlled conditions over a
period of 24 days using sterile and non-sterile soils. In both cases,

Acinetobacter calcoacetius cultures were added (5.3 X 1010) to the

 

soils previously incorporated with BOA. The rate of production of AZOB
was linearly related to the quantity of BOA available in the soil
(Figure 6). At the end of day 12, only 7$ AZOB was detected with a
loss of 12$ BOA. However, the 0.1 mg of BOA/g soil (non-sterile)
experiment showed a quadratic trend in the production of AZOB (Figure
7). Maximum yield of AZOB (9.7 $) was observed during the incubation
period of days 2 to 4 with an 85$ decrease in BOA concentration. This
is simliar to the 9.1$ reported by Nair et a1. (1990).

The recovery of BOA from the soil was 33$ of the total amount
incubated. The remainder of the BOA could have already converted to
AZOB or a portion could have decomposed to products that are not yet
characterized. Also, BOA might have decomposed to other products under
mechanisms which are not yet known. However, a possible mechanism for
the production of AZOB from BOA, monomethoxy AZOB (MAZOB) from MBOA and
dimethoxy AZOB (DIMAZOB) from MBOA was described earlier by Nair et al.
(1990). When the concentration of BOA in field soil was raised to 1.0
mg/g soil, the AZOB production increased linearly to the concentration
of BOA (Figure 8). This indicated that the excess BOA had an impact on
the total microflora in the soil. Usually, microbial populations
present in the soil are low in numbers but explode quickly when a
suitable substrate becomes available (Atalas and Bartha. 1987).
Therefore having observed different trends for the BOA transformation

in non-sterile field soil, it would seem logical that given the

80

Figure 6. Rate of transformation of BOA to AZOB in sterile soil
incubated with Acinetobacter calcoacetius.

Concentration of BOA was 0.1 mg/g soil.

81

 

82

Figure 7. Rate of transformation of BOA to AZOB in non-sterile
soil incubated with a BOA concentration 0.1 mg/g of

soil.

83

Weight (ug)

$0 1...

A00 4..
m0 Ii
00 II

 

 

 

84

Figure 8. Rate of transformation of BOA to AZOB in non-sterile
soil incubated with a BOA concentration of 1.0 mg/g of

soil.

85

Weight (ug)

A000I_
000 I
000 I

.NOOI

 

moo A

N00 I

A00 I

 

 

AN

86

N 0
DIBOA—91“ I
R O O — qu

Enzyme Hydrolysis 1 OH

I
N o
DICBOA
O/\OH

(Acinetobacter calcoacetius)

Microbial MetaboIism \0001
R1 — R2

AZOB
N :7. N

Scheme 2: Proposed scheme for formation of AZOB with

Acinetobacter calcoacetius.

87

appropriate time interval a quadratic trend could be expected at both
rates of BOA (early lag phase followed by linear phase).
Rapid AZOB production did occur in the different Michigan soils

tested and would suggest the presence of Acinetobacter calcoacetius

 

(Scheme 2) or another similiar organism capable of the conversion of
BOA to AZOB (Figures 9 and 10). Ahalysis of various soil parameters
did show differences in nechanical makeup, fertility, organic matter
and cation exchange capacity but did not correlate with the
transformation process (Appendix 3). Whether or not a gramineae crop
was previouly grown did not affect the conversion.

AZOB was produced in large quantities in aqueous shake cultures
for bioassay experiments. This method was convenient, cost effective,
and less time consuming. Extraction cfi‘ AZOB and BOA by EtOAc from
aqueous media was easily performed, and even though the concentration
of AZOB in the non-shake conditions was not evaluated, the rate of
production (Hi AZOB under shake conditions appeared greater. Since the
production of AZOB occurred readily in liquid cultures it is not
neccessary to use soil for the transformation of BOA. Even with a
lower yield from the liquid medium (24$) when compared to the soil
(30.8$), water was a nwme attractive transformation medium for large
scale 13 315:9 production of A208.

Conclusions: ' The experiments with BOA in non-sterile soil also
indicated that the A208 produced might have been used by other soil
microbe(s) as c: substrate thus explaining the quadratic trend observed
ix: Figure ‘7. This data supports the difficulty encountered in the
detection of AZOB from the soil incubated with rye tissue. The

concentration of AZOB started to decline in the experiment with lower

Figure 9. AZOB concentration in 4 Michigan soiis incubated with

0.1 mg BOA/g soii.

89

Weight (ug)

00-

00..

$0L

m0L

A0-

 

>N0w

 

Asvom
OmAAAoBo
XoEBmNoo
mvoxo

 

90

Figure 10. BOA concentrations in 4 Michigan soiis incubated with

0.1 mg BOA/g soii.

91

 

GO)

 

 

 

moo- 263m
...... Omioao
) Amo. II i 1.: AAoAmSmnoo
(MI. /III 0223.
M. So. ..\.. . .
0 cool.
m. IIII .\ ...V../
w \I.\/ ......./
mo- . 11:: .:w
0\ I’ll, .
\ I
O — a _ A
m 0 AN

92

concentrations of BOA (Figure 7) by the end of day 4. Similar
situations in the field can be envisioned where BOA leached from the
rye residues is undergoing rapid transformation to AZOB. This
experiment also suggested that the accumulation of AZOB *will be

difficult to detect in field soils with or without rye residues.

93

LITERATURE CITED

Almeida, F.S. 1985. Effect of some winter crop mulches on the soil weed
infestation. Proc. 1985 Brit. Crop Prot. Conf , Weeds. 2:651-695.

Atalas, R.M., and I2. Bartha. 1987. Microbial Ecology: Fundamentals
and aplications. The Benjanim/Cummings publishing Company, Inc. Menlo
Park, CA. Pp. 71.

 

Barnes, J.P. and A.R. Putnam. 1986. Evidence for allelopathy by
residues and aqueous extracts of rye (Secale cereale L.) Weed Sci.
34:384-390.

 

Barnes, J.P. and A.R. Putnam. 1987. Role of benzoxazinones in
allelopathy by rye (Secale cereale L.). J. Chem. Ecol. 13(4):889-905.

Barnes, J.P., A.R. Putnam, B.A. Burke, and A.J. Aasen. 1987.
Isolation and characterization of allelochemicals in rye herbage.
Phytochem. 26(5):1385-1390.

Chapman, S.J., and J.M. Lynch. 1983. The relative roles of micro-
organisms and their metabolites in the phytotoxicity of decomposing
plant residues. Plant and Soil. 74 457-459.

Chou, C.H. and Z.A. Patrick. 1976. Identification (uni phytotoxic
activity of compounds produced during decomposition of rye and corn
residues in soil. J. Chem. Ecol. 2(3):369-387.

Hietala, P.K. and AMI. Virtanen. 1960. Precursors of benzoxazolinone
in rye plants: precursor I, the glucoside. .Acta Chem. Scand.
14(2):502-504.

Hofman, .J., and O. Hofmanova. 1968. 1,4-Benzoxazinone derivatives in
plants: sephadex fractionation and identification of a new glucoside.
Euro. J. Biochem. 8:109-112.

Nair, M.G., C.J. Whitenack, and A.R. Putnam. 1990. 2,2’-oxo-1,1'-
azobenzene. A microbially transformed allelochemical from 2,3,-
benzoxazoline: I. J. Chem. Ecology. 16(2):353-364.

Long, B.J., G.M. Dunn, and 0.0. Routley. 1974. Rapid procedure for
estimating cyclic hydroxamate (DMIBOA) concentration in Maize. Crop
Sci. 14:601-603.

Patrick, Z.A. 1955. The peach replant problem in Ontario. II. Toxic
substances from microbial decomposition products of peach root
residues. Can. J. Bot. 33:461-485.

Proebsting, E.L., and AnE. Gilmore. 1941. The relation of peach root
toxicity to the re-establishing of peach orchards. Proc. Amer. Soc.
Hort. Sci. 38:21-24.

94

Putnam, A.R., 1984. Allelopathic research in agriculture: past
highlights and potential. The Chemistry of Allelopathy. ACS. Sym.
Series. No. 286.

 

Rice, E.L. 1984. Allelopathy. 2nd edition. Academic Press, INC., New
York.

Shilling, D.G., R.A. Liebl, and A.D. Worsham. 1985. Rye(Secale
cerealeu) and Wheat(Triticum aestivum L.) nmlch: The suppression of
certain broadleaved weeds and the isolation and identification of
phytotoxins. The Chemistry of Allelopathy: 243-271 pp.

Smeda, R.J., and 8.0. Weller. 1986. Weed suppression with rye in
transplanted tomatoes. North Central Weed Control Conference
Proceedings, Abstract. 41:27-28.

Smeda, R.J., and A.R. Putnam. 1988. Cover crop suppression of weeds
and influence on strawberry yields. Hort Sci. 23(1):132-134.

Tang, C.S., S.H. Chang, D. H00, and ICJL. Yanagihara. 1975. Gas
chromatographic determination of 2(3)-benzoxazolinones from cereal
plants. Phytochem. 4:2077-2079.

Virtanen, A.I., P.K. Hietala, and 0. Wahlroos. 1957. Antimicrobial
substances in cereals and fodder plants. .Arch. Biochem. Biophys.
69:486-500.

Zungica, G.E., V.H. Agrandona, H.M. Niemeyer, and L.J. Corcuera. 1983.
Hydroxamic acid content in wild and cultivated gramineae. Phytochem.
22(12):2665-2668.

95

CHAPTER 4

Relative Toxicity of Rye (Secale cereale L.) Allelochemicals
to Weeds and Vegetable Crops

ABSTRACT

Three allelochemicals from rye or its breakdown products were
evaluated for activity on garden cress (Lepidum sativum L.),
barnyardgrass (Echinochloa crus-galli (L.) Beauv.). cucumber (Cucumis

sativus L.), and snap bean (Phaseolus vulgaris I..). 2,4-dihydroxy-

 

1,4(2H)-benzoxazin-3-one (DIBOA), 2(3H)-benzoxazolinone (BOA), and
2,2’-oxo-1,1'-azobenzene (AZOB) were all applied singly at 50, 100, and
200 ppm and in two and three way combinations each at 50 and 100 ppm.
AZOB at 100 and 200 ppm produced 38 to 49$ more inhibition than
DIBOA while combinations of BOA/DIBOA which contained AZOB at 100 ppm
had 54 11> 90$ more inhibition when compared to DIBOA/BOA combinations.
All combinations were slightly antagonistic to barnyardgrass while
several combinations caused c: synergistic response: to garden cress
germination (uni growth. Cucumbers and snap beans exhibited both types
of responses depending on the allelochemical combination and
application rateu Plant—produced benzoxazinones were more inhibitory
to crops inn”: weeds. Therefore, improved selectivity to cucumbers and
snap beans would be expected if there was rapid transformation of the

benzoxazinones to the microbially produced AZOB.

96

INTRODUCTION

Rye (Secale cereale L.) is an annual cereal grain that has shown
potential as a cover crop in a variety of cropping systems (Putnam and
Defrank, 1983; Barnes and Putnam, 1986), because it has been found to
reduce weed biomass, improve soil aggregation, reduce soil erosion and
enhance water penetration and retention (Blevins et al., 1971; Young,
1982). With the renewed interest in lowering energy inputs, reduced
tillage systems employing cover crops will undoubtedly be an important
tool of the future. Maintenance of residues on the soil surface will
be inherent to these reduced tillage systems. Residues can interact
with weeds or crops to potentially reduce the growth of the plants.
This interference can arise from competition for essential plant growth
requirements such as light, water, and nutrients (Knavel et al, 1977;
Teasdale et al., 1983; Mascianica et al., 1986; Skarphol et al., 1987;
Glen and Welker, 1989) or from allelopathic chemicals released into the
environment from decaying plant residues (Shilling et al., 1985; Barnes
et al., 1987; Nair et al., 1990).

Allelopathy is defined as one plant chemically interfering with
the germination, growth, or development of another and has been
implicated in numerous plant interactions (Muller, 1966; Whittaker and
Feeny, 1971; Barnes et al., 1986). Proof of allelopathy requires
detection of small quantities of biologically active compounds. Even
though natural product chemists are able to characterize new plant
compounds, rigorous testing is required to determine their allelopathic
potential (Putnam 1988).

Although there have been numerous studies on toxicity of rye

97

residues (Kimber, 1973; Chou and Patrick, 1976; Putnam and Defrank,
1983; Shilling et al., 1985; Barnes and Putnam, 1986; Barnes et al.,
1987), no one has offered (1 complete explaination of interference
reported by these plant residues. There have been several attempts to
isolate and identify the many compounds within rye showing biological
activity. Virtanen (1957) isolated c: cyclic hydroxamic acid, 2(3H)-
benzoxazoline (BOA), from four-day old rye seedlings. He reported BOA
as being responsible for the chemical resistance of rye to Fusarium
infection. Tang et al. (1975) further quantified BOA present in rye
and found 8 times more BOA in the shoots than in the roots. Chou and
Patrick (1976) identified nine compounds from decaying rye tissue and
found them applied individually' to be inhibitory to lettuce seed
germination. Zungica et al (1983) have evaluated rye for two
hydroxamic acids, 2,4-dihydroxy-1,4(2H)-benzoxazin-3-one (DIBOA) and
BOA, and found them commonly occurring in both cultivated and wild
accessions. Shilling et al. (1985) isolated phenyllactic acid (PLA)
and hydroxybutyric acid (HBA) from field grown rye and found each

allelochemical inhibited redroot pigweed (Amaranthus retroflexus L.),

 

common lambsquarters (Chenopodium album L.), and common ragweed

 

(Ambrosia artemisiifolia L.) growth at concentrations of 2 mM. Barnes
et al. (1987) compared the activity of four rye compounds PLA, HBA,
DIBOA, and BOA and reported DIBOA and BOA to be the most toxic to large
crabgrass (Digitaria sanguinalis (L . ) Scop. ), barnyardgrass

(Echinochloa crus-galli (L.) Beauv.). proso millet (Panicum miliaceum

 

L.), redroot pigweed, tomato (Lycopersicon esculentum Mill.), and

 

lettuce (Lactuca sativa L.) root growth at concentrations of 67 to 250

 

PPM-

98

Nair et al. (1990) isolated a microbially produced 2,2’-oxo-1,1’-
azobenzene from the soil enriched with BOA (Chapter 1, page 21, Scheme
1) and showed that DIBOA, BOA, and AZOB were inhibitory to

barnyardgrass and garden cress (Lepidum sativum L.) at concentrations

 

of 67 to 250 ppm. In these tests, AZOB was more biologically active
than BOA and DIBOA.

Although there have been numerous compounds isolated and
identified from rye tissue, it is difficult to predict the active
concentration of any one compound at any one time as it occurs in
nature. It is possible that they might occur either alone or in
numerous combinations under various environmental and plant conditions.
Since the hydroxamic acids, BOA and DIBOA, and their AZOB breakdown
product exhibited the most activity, the objective of this study was to
evaluate these three known rye allelochemicals singly and in

combinations for their activity on weeds, cucumbers and snap beans.

MATERIALS AND METHODS

Spray Reagents for TLC Detection. DIBOA was detected on thin—layer
plates with ferric chloride (FeCls) spray reactant consisting of 5$
FeC13 in 95$ ethanol, acidified with concentrated HCl. A spray reagent
consisting of 1$ ceric sulfate (CeSOa) in concentrated H280“ was used
to detect BOA.

SOURCE OF ALLELOCHEMICALS. BOA was obtained from Aldrich Chemical
company and AZOB was produced from BOA according to the procedure
described in Chapter 3, page 63.

Extraction of DIBOA. Seedlings of rye (’Wheeler’, 11 days old, 2.2 kg)
were homogenized in a Waring blender with distilled water (4.5 L), and

kept at room temperature (1 h) to ensure enzymatic hydrolysis of the

99

DIBOA-glucoside to DIBOA. After filtering through cheese cloth, the
filtrate was heated in a water bath until the temperature of the
extract reached 70 °C. It was then cooled immediately in an ice bath.
Coagulated components were removed by vacuum filtration using filter
paper (Whatman No. 1). Filtrate was then lyophilized at 5 °C and
stored at -20 °C until further separation.

Lyophilized rye extract (121.91 9) was stirred with HCl in MeOH
(1.5 I” pH 1.0) for 2 hours at room temperature. The mixture was then
filtered through a sintered glass. filter (fine, 4-5 A°) and the
filtrate dried lg ygggg. This crude residue was extracted with CH013
(3 X 100 ml) and the organic extract was evaporated to dryness. The
oily residue (35.3 g) was redissolved in CHCl3 and precipitated*with
hexane (500 ml) and filtered. This procedure was repeated twice and
the combined CHCl3-hexane extract was dried 13 ygggg (12.84 g). A TLC
analysis of this crude product confirmed the presence of DIBOA.
Further purification of DIBOA from the above partially purified mixture
was carried out by vacuum liquid chromatography. .A slurry of column
silica (40-60 mesh size, 260 g) in CHCls-MeOH (4:1) was packed under
vacuum :u: a sintered glass filter (medium, 10-15 A) and washed with
CHCls-MeOH (4:1, 300 ml). The partially purified gum containing DIBOA
(12.84 g) in CHCls-MeOH (4:1, 30 ml) was applied on the silica under
vacuum and eluted with the same solvent system. Four fractions were
collected in 250 ml portions and analyzed by TLC. Fractions 2 and 3,
positive for DIBOA by FeCl3 spray, were combined and evaporated to
dryness (10.94 g). This was purified again by flash column
chromatography (silica, CHCls-MeOH 4:1 v/v). 'The initial fraction (30

ml) was discarded and upon TLC analysis of fractions 1-21 (each 5 ml)

100

were found to be negative for DIBOA. The fractions 22-29 (each 10 ml),
monitored by TLC with FeCl3 spray, were found to contain the largest
quantity of DIBOA. Removal of solvent at reduced pressure afforded a
brown solid (2.43 9). Final purification of DIBOA was achieved by TLC
(silica, CHCl3-MeOH, 4:1) and the resulting product recrystallized from
CHCls-MeOH mixtures, and gave pale yellow crystals of DIBOA (187 mg).

BIOASSAY: Activity of three known compounds associated with rye
(DIBOA, BOA, and AZOB) were assayed on two weed indicators (garden

cress and barnyardgrass) and two crops (cucumber (Cucumus sativus L.)

 

and snap bean (Phaseolus vulgarus L.)). Growth parameters recorded
were seed germination, and root and shoot lengths. All three compounds
were applied alone at 50, 100, and 200 ppm and in two and in three way
combinations each applied at 50 and 100 ppm. The experimental design
was a randomized complete block design with 4 replications.

Stock solutions (1mg‘ml'1 methanol) of purified compounds were
applied to filter paper (Whatman’s No. 1) which lined the petri
dishes. Pure methanol was also included as a control. After
application of the compounds on the filter paper, the solvent was
allowed to evaporate (2 to 3 hrs.) in the dark before seed placement.
In the snap bean assays, the solvent was allowed to evaporate for an
additional 6 to 8 hrs prior to planting.

Weed seeds (15) were uniformly distributed on filter paper (4.25
cm) in petri dishes (60 X 15 mm) after which distilled water (1.5 ml)
was added. Garden cross and barnyardgrass seeds were then incubated
for 72 and 96 hrs respectively. ‘Flurry' cucumber seeds (5) were
uniformly distributed on filter paper (7 cm) in petri dishes (100 X 15

mm) and then distilled water (2 ml) was added. Cucumbers were then

101

incubated for 120 hrs. ‘Bush Blue Lake 47’ snap bean seeds (4) were
uniformly distributed cu: seed germination paper (5 X 5 cm) in Magenta
boxes (6 X 6 X 9 cm) after which distilled water (5 ml) was added. A
cellulose sponge (3.0 X 3.8 X 6.2 cm) saturated with distilled water
was suspended from the sides of the box before attaching the lids. Each
container was individually sealed with parafilm (American Can Co.) and
incubated for 216 hrs.
Incubation of bioassay. After placing the lids onto the containers,
they were randomly placed on trays (36 X 57 cm) that were lined with
damp brown paper toweling. Eoch tray held one to two blocks and was
covered with a clear plastic bag before placing into the incubator at
26 °C for the duration of the study.
Statistical methods. Data obtained in bioassays were subjected to
analysis of variance and means were compared with the least significant
difference (LSD) test. Tables represent the means from duplicate
experiments. Analysis for synergism or antagonism of combination
treatments were conducted according to Colby’s methods (1967).

RESULT AND DISCUSSION
DIBOA quantification: Since DIBOA was not available commercially, it
was neccessary to isolate it from field or greenhouse grown rye tissue.
The modification of DIBOA extraction procedure (Scheme 1) gave a higher
yield of DIBOA than previously reported (Nair et al., 1990). The major
difference in the extraction step was the treatment of lyophilized
extract with HCl in MeOH. Barnes et al. (1986) quantified hydroxamic
acids by colorimetric methods and obtained a 27 fold increase of DIBOA

in greenhouse grown rye shoots over field grown. Our yields were lower

102

Fresh Rye Tissue (2.2 kg)

I
I
I 1. Blended with distilled water (4.5 L)
I 2. Stand for 1 h

I 3. Filtered through cheese cloth

I
I

 

I I

I I
Residue Filtrate
discarded I

I
I 1. Heat 70°C

I 2. Cooled to room temperature
I

I

I

 

 

3. Filtered
I I
Residue Filtrate (121.91 g)
discarded I
I
I 1. Dissolved in HCleeOH (3N, IL)
I 2. Stirred for 2 h
I 3. Filtered
I
I
I I
Residue Filtrate (10.94)
discarded I

I
I 1. Evaporated to dryness

I 2. Extracted with CHC13:MeOH
I (4:1, 250 ml)

I

 

I I
Residue CHCl3IMeOH Extract
discarded I

1. evaporated to dryness
2. extract purified

by chromatography
3. crystallization

I
DIBOA (187 mg)

Scheme 1. Extraction of DIBOA from 11 day-old greenhouse grown rye

103

than Barnes et al. (1986) but were four-fold higher than Nair et al.
(1990). However we did not process all the positive DIBOA containing
fractions. This DIBOA along with BOA and AZOB was used for evaluating
allelopathic activity of these compounds under field and greenhouse
conditions.

Relative biological activity of allelochemicals: Of the three
chemicals tested cu: barnyardgrass, AZOB was the most active, reducing
both root and shoot growth at all three application rates (Table 1).
Combinations of the compounds containing AZOB showed 41 to 67$ more
activity on roots and 20 to 33$ more activity on shoots when compared
to combinations of DIBOA/BOA at the same concentrations. The
combinations showed slightly less activity when compared to expected
values by Colby‘s methods. This would suggest that these compounds are
antagonistic when they are applied together.

For garden cress, all parameters measured were inhibited by the
application cfi’ AZOB (Tables 2 and 3). Germination was reduced by 3 to
4-fold by combinations containing AZOB/BOA. each at 100 ppm. In
addition, AZOB cu“ BOA applied singly showed 2 to 4-fold more activity
on both shoot and root growth when compared to DIBOA. Shoot and root
growth was severely inhibited with combinations of AZOB/BOA or AZOB
alone at 100 ppm. In the above mentioned combinations, the observed
values were always less than the expected, suggesting synergism. The
order of activity on garden cress was AZOB > BOA > DIBOA which is in
agreement with Nair et al. (1990).

For cucumbers, DIBOA was more inhibitory to both root and shoot

growth than either BOA or AZOB when applied singly (Table 4). When

104

 

 

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105

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106

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108

 

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applied singly (n: 50 ppm, none of the chemicals reduced root or shoot
growth, but when concentrations were 100 ppm or greater there were
significant reductions in growth. Again, DIBOA, BOA and all
combinations were more active» with increasing concentrations. The
DIBOA/BOA combinations were 14 to 44$ more inhibitory than the
combinations of BOA/AZOB and appeared ix) be synergistic while the
BOA/AZOB combinations appeared to have antagonistic effects. This
suggested that the most toxic compounds to cucumbers were ones found in
the rye herbage and not the :nicrobially produced AZOB. Irf rapid
conversion of these compounds did occur, one could expect improved
cucumber crop safety in the presence of decomposing plant material.

In the snap bean assay, there was more variation perhaps due to
seed variability or more intricate experimental design. Overall, shoot
growth appeared more sensitive than roots with DIBOA and BOA showing a
rate» response with increasing concentrations (Table 5). For
combinations with 50 ppm of each compound, only slight synergistic
activity was seen, while at 100 ppm, antagonism was evident.
Applications of BOA at 100 and 200 ppm and combinations of BOA/AZOB
showed the greatest inhibition. Both large seeded dicot crops were more
affected by DIBOA/BOA and BOA/AZOB combinations. From previous work
(Putnam (Hui Defrank, 1983 and Skarphol et al., 1987) both cucumber and
snap beans have been shown to respond favorably to cover crop residues.
.A possible explanation is that the large seed crops are located too
deep to be influenced by the residues, or alternately the
concentrations of these chemicals are too low to affect the emergence
of the crops. The effect seen on small seeded weeds such as cress and

barnyardgrass could come from rapid degradation of the benzoxazinones

109

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apnou< one u: nose—noneona.

 

 

 

 

 

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110

from the injured plant material and the more active microbially
produced AZOB. Also the smaller seeds are located closer to the soil
surface where higher concentrations of these compounds might exist.

Conclusions: The benzoxazinones have been reported in cereal grains
and corn plants (Virtanen et cu“ 1957). In addition, the recent
isolation of the microbially produced AZOB may help to account for a
portion of the 88$ of inhibition by rye residues that Barnes et al.
(1986) could not account for. Our results show that both small seeded
species were strongly inhibited by AZOB and garden cress by BOA as
well. These results agree with Barnes et al. (1986) and Nair et al.
(1990) where they reported BOA to be more active on dicot and small
seeded crop species. FRH‘ monocots, there has been allelochemical
activity reported with DIBOA. The finding that AZOB has herbicidal
potential against most small seeded species gives hope that microbially
produced allelochemicals would be useful in weed control.

In general, when comparing the response of the different growth
parameters, root and shoot inhibition were more sensitive indicators of
allelopathic activity than germination inhibition. Only in the case of
garden cress was germination a positive indicator of activity. Neither
of the large seeded crops nor barnyardgrass showed significant
germination inhibition. This indicated that the mode of action for
barnyardgrass, cucumber, and snap bean was not on germination processes
but. rather (N1 early plant development processes. This suggests the
expected field response would be reduced plant growth and not reduction
in number of plants.

Our results not only suggest that these compounds are selectively

allelopathic, but indicated the relative quantities of these compounds

111

are important in determining the expected outcome. Earlier bioassays
with these compounds used small seeded species whereas ours compared
larger seeded crops. It appears that selectivity can be achieved based
on seed size and seed placement. This same principle has allowed the
selective use of synthetic herbicides for decades. The difficulty with

allelochemicals is ix: predict or quantify what might be present at any

particular time in the field.

112

LITERATURE CITED

Barnes, J.P. and A.R. Putnam. 1986. Evidence for allelopathy by
residues and aqueous extracts of rye (Secale cereale L.) Weed Sci.
34:384-390.

 

Barnes, J.P. and A.R. PutnanL 1987: Role of benzoxazinones in
allelopathy by rye (Secale cereale L.). J. Chem. Ecol. 13(4):889-905.

 

Barnes, J.P., A.R. Putnam, B.A. Burke, and A.J. Aasen. 1987.
Isolation and characterization of allelochemicals. in rye herbage.
Phytochem. 26(5):1385-1390.

Blevins, R.L., 0. Cook, S.H. Phillips, and R.E. Phillips. 1971.
Influence of no-tillage on soil moisture. Agron. J. 63:593-596.

Chou, C.H. and Z.A. Patrick. 1976. Identification and phytotoxic
activity of compounds produced during decomposition of rye and corn
residues in soil. J. Chem. Ecol. 2(3):369-387.

Colby, S.R. 1967. Calculating synergistic and antangonistic responses
of herbicide combinations. Weeds. 15:20-22.

Glenn, D.M., and WKV. Welker. 1989. Orchard soil management systems
influence rainfall infiltration. J. Amer. Soc. Hort. Sci. 114(1):10-
14.

Kimber, R.W. 1973. II. The effect of time of rotting of straw from
some grasses and legumes on the growth of wheat seedlings. Plant and
Soil. 38:347-361.

Knavel, D E., J. Ellis, and J. Morrison. 1977. The effect of tillage
systems on the performance and elemental absorption by selected
vegetable crops. J. Amer. Soc. Hort. Sci. 102(3):323-327.

Mascianica, M.P., H.P. Wilson, R.F. Walden, T.E. Hines, and R.R.
Bellinder. 1986. No-tillage snapbean growth in wheat stubble of varied
height. J. Amer. Soc. Hort. Sci. 111(6):853-857.

Muller, C.H. 1969. Allelopathy as a factor in ecological processes.
Vegetatio. 18:348-357.

Nair, M.G., C.J. Whitenack, and A.R. Putnam. 1990. 2,2’-oxo-1,1’-
azobenzene. A microbially transformed allelochemical from 2,3,—
benzoxazoline: I. J. Chem. Ecology. 16(2):353-364.

Putnam, A.R. and J. Defrank. 1983. Use of phytotoxic plant residues
for selective weed control. Crop Protection, 2(2), 173-181.

113

Shilling, D.G., R.A. Liebl, and A.D. Worsham. 1985. Rye(Secale
cereale L.) and Wheat(Triticum aestivum L.) mulch: The suppression of
certain broadleaved weeds and the isolation and identification of
phytotoxins. The Chemistry of Allelopathy: 243-271 pp.

Skarphol, B.J., K.A. Corey, and J.J. Meisinger. 1987. Response of
snap beans to tillage and cover crop combinations. J. Amer. Soc. Hort.
Sci. 112(6):936-941.

Tang, C.S., S.H. Chang, D. H00, and K.H. Yanagihara. 1975. Gas
chromatographic determination of 2(3)-benzoxazolinones from cereal
plants. Phytochem. 4:2077-2079.

Teasdale, J.R., J.R. Frank. 1983. Effect of row spacing on weed

 

competition with snap beans (Phaseolus vulgaris L.). Weed Science.
31:81-85.

Virtanen, A.I., P.K. Hietala, and O. Wahlroos. 1957. Antimicrobial
substances in cereals and fodder plants. .Arch. Biochem. Biophys.
69:486-500.

Virtanen, A.I. and P.K. Hietala. 1960. Precursors of benzoxazolinone
in rye plants: Precusor II, the aglucone. Acta Chem. Scand.
14(2):499-502.

Whittaker, R.H. (uni P.P. Feeny. 1971. Allelochemics: chemical
interactions between species. Science 171:757-770.

Young, H.M.,Jr. 1982. No-Tillage Farming. No-Til Farmer, INC.
Brookfield, WI. 2 ed. 202 pp.

Zungica, G.E., V.H. Agrandona, H.M. Niemeyer, and L.J. Corcuera. 1983.
Hydroxamic acid content in wild and cultivated gramineae. Phytochem.
22(12):2665-2668.

114

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