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This is to certify that the

 

 

 

dissertation entitled
ALLELOPATHIC POTENTIAL OF ASPARAGUS

(Asparagus officinalis L.)
presented by

Anne C. Hartung

has been accepted towards fulfillment
of the requirements for

Ph.D

. degnxin Horticulture

Major professor

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ALLELOPATHIC POTENTIAL OF ASPARAGUS (Asparagus officinalis L.)

By

Anne C. Hartung

A DISSERTATION

Sublitted to
Michigan State University
in partial fulfill-cut of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Depart-cut of Horticulture

1987

©1987

ANNE C . HARTUNG

All Rights Reserved

ABSTRACT
ALLELOPATHIC POTENTIAL OF ASPARAGUS (Asparagus officinalis L.)

By

Anne C. Hartung

Two major problems of asparagus (Asparagus officinalis LJ are
premature decline of fields and inability to replant asparagus after
removal of old fields. Asparagus is believed to have allelopathic
potential and compounds released from its tissues increase the
incidence of Fusarium root rot on asparagus seedlings. Several plant
species were inhibited differently by compounds released from dried
root or fern tissues incubated in sterile or nonsterile soil under
anaerobic or aerobic conditions. Both root and fern tissues were
inhibitory to seedling development under anaerobh: or aerobic
conditions, as well as sterile or nonsterile soil conditions. Soil
microorganisms were not necessary for, and over time, reduced asparagus
tissue toxicity in soil. When excised asparagus roots were treated
with increasing concentrations of water extracts from dried asparagus
root tissues, electrolyte efflux was increased and respiration
decreased. Exposure of asparagus seedlings to similar water extracts
of asparagus root tissue decreased the peroxidase activity.

The responses of microbial populations in soils with dried

asparagus root or fern tissue or planted with asparagus, A; sprengeri,

ANNE C. HARTUNG

and other selected crops were compared. Fusarium spp. and other fungal
populations increase lei3 fold in treatments where asparagus root
tissue was incorporated into soil. Although bacterial CFU's quickly
increased in treatments where fern tissue was incorporated, both
bacterial and fungal populations were reduced in soils where asparagus
plants were grown compared to soils with other crops. The chloroform
extracts of asparagus root tissues were separated using column, thin
layer and high performance liquid chromatography. Ferulic, isoferulic,
malic, citric, and fumaric acids were identified in HPLC fractions by
GC-MS. NMR spectra of compounds isolated using preparative TLC or
soxlet extractions also suggested thatiasparagusic acid and caffeic

acid were present in root tissues.

DEDICATION

This dissertation is dedicated to "The Abbott Bros. Band" of East
Lansing, MI, including those greats, Bob M. Abbott, crack lead guitar,
Dick w. Abbott, soulman, Barbara A. Abbott, Big mamma, and Brian H.
Abbott, the jazz man - not forgetting great backup on bass, vocals, and
guitar from their cousin, Michelle B. Abbott. I could never have made

it without you guys. I tip my hat to you, Mr. Moore and Dr. Johnston.

ii

ACKNOHLEDGEHENTS

There is no way I could possibly acknowledge all those kind souls
who helped me throughout the development of this Thesis. I want to
thank the members of my committee, my major professor, Dr. Alan R.
Putnam for his support at some difficult times during my tenure, as
well as some of the laughs we also shared. Also, Dr. Hugh Price, Dr.
Irvin Widders, and Dr. Matt Zabik gave their time and expertise. I
also owe a debt of gratitude to Dr. Christine Stephens for her belief
in me as a student and her interest in my career.

For structural elucidation of isolated chemicals, I owe many
thanks to the patience and interest of Dr. Basil Burke, Palo Alto,
California, for his interest in me as a student and his genuine
kindness, and his support staff, Wendy Goldsby and Dr. Muralee Nair.

I thank my wonderful and patient husband as well as my beautiful,
intelligent, understanding, tremendous daughter, for their kind and
patient waiting throughout this ordeal.

I thank you Jackie Schartzer because I never would have made it
look like a dissertation without you. I'll never forget your
friendship and support.

But most of all, I thank me because its done, by God, and I<1id

it.

TABLE OF CONTENTS

PACE
LIST OF TABLES ........................... vii
LIST OF FIGURES .......................... ix
CHAPTER 1: LITERATURE REVIEW
INTRODUCTION ........................... l
Asparagus Decline in Michigan .................. l
Allelopathy... . . .. . ................... 5
Allelopathy and Microorganisms.. . . .............. 6
Asparagus as an Allelopathic Plant ................ 8
Isolated and Identifed Compounds from Asparagus Tissue ...... 9
LITERATURE CITED ......................... 13
CHAPTER II: ALLELOPATHIC POTENTIAL OF ASPARAGUS RESIDUES IN
STERILE AND NONSTERILE SOIL
ABSTRACT ............................. 22
INTRODUCTION ........................... 23
MATERIALS AND METHODS ...................... 24
Preparation of Asparagus Plant Material ............. 24
Assessment of Microbial Contamination of Asparagus Tissues. . . . 25
Toxicity of Asparagus Tissues in Soil .............. 26
Preparation of Purified Asparagus Root Extracts ......... 28
Bioassay of Partially Purified Compounds with Indicator
Species ............................. 29
RESULTS AND DISCUSSION ...................... 31
Assessment of Microbial Contamination of Asparagus Tissues. . . . 3l
Toxicity of Asparagus Tissues in Soil .............. 3i
Bioassays of Partially Purified Compounds on Indicator
Species ............. . .......... . . . . . 45
LITERATURE CITED ......................... 48

iv

TABLE OF CONTENTS continued:

PACE
CHAPTER III: CHARACTERIZATION OF INHIBITORY ACTIVITY OF ASPARAGUS
ROOT EXTRACTS ON ASPARAGUS SEEDLINGS
ABSTRACT ............................. 5T
INTRODUCTION ........................... 52
MATERIALS AND METHODS ...................... 53
Preparation of Asparagus Plant Material ............. 53
Preparation of Fusarium Inoculum ................. 53
Inhibition of Asparagus Seedlings by Root Tissue. ........ 54
Biochemical Assays on Asparagus Seedlings ............ 55
Comparison of Inhibitory Activity from Different Aged
Asparagus Plants ........................ 56
RESULTS .............................. 58
Preparation of Asparagus Plant Material ............. 58
Inhibition of Asparagus Seedlings by Root Tissues ........ 58

Physiological and Biochemical Assays on Asparagus Seedlings . . . 58
Comparison of Inhibitory Activity from Different Aged

Asparagus Plants ........................ 63
DISCUSSION ............................ 69
LITERATURE CITED ......................... 75

CHAPTER IV: INTERACTIONS 0F ASPARAGUS ROOT AND FERN TISSUE WITH
MICROBIAL ORGANISMS

ABSTRACT ............................. 78
INTRODUCTION ........................... 78
MATERIALS AND METHODS ...................... 80
The Effect of Asparagus Root or Fern Tissue and Plant Species
on Naturally Occurring Microbial Populations .......... 80
Selective Media Used for Evaluation of Microbial Species in
Greenhouse Experiments ...................... 82
Preparation of Partially Purified Asparagus Root Extracts . . . . 83
Effects of water Extracts on Isolates of Pvthium spp ....... 84
Sensitivity of Bacterial Isolates to Purified Extracts of
Asparagus Root Tissue ...... . ............... 85
RESULTS ............................. 86
Effects of Asparagus Root and Fern Residues and Plant Species
on Naturally Occurring Microbial Populations .......... 86
Effects of Water Extracts on Isolates of Pzthium spp ....... 90
Sensitivity of Bacterial Isolates to Purified Extracts of
Asparagus Root Tissue ...................... 92

TABLE OF CONTENTS continued:

 

PACE
DISCUSSION ............................ 92
Effects of Asparagus Root or Fern Residues and Plant Species
on Naturally Occurring Microbial Populations .......... 92
Effects of water Extracts on Isolates of Pythium spp ....... 94
Sensitivity of Bacterial Isolates to Purified Extracts
of Asparagus Root Tissue .................... 95
LITERATURE CITED ......................... 96
CHAPTER V:ISOLATION AND CHARACTERIZATION OF ALLELOCHEMICALS IN
ASPARAGUS (ASPARAGUS OFFICINALIS L.) ROOT TISSUE
ABSTRACT ............................. TOO
INTRODUCTION ........................... lOl
MATERIALS AND METHODS ....................... lO3
Preparation of Asparagus Plant Material ........... . .lO3
Bioassay Procedure for Evaluation of Isolated Components
of Asparagus Root Extractions .................. l04
Bioassay of Known Compounds and Crude Extracts .......... l05
Purification of Inhibitory Components in Asparagus Root
Extracts: Scheme #1 ....................... l05
Purification of Inhibitory Components in Asparagus Root
Extracts: Scheme #2 ....................... ll2
Purification of Inhibitory Components in Asparagus Root
Extracts: Scheme #3 ....................... ll3
Isolation of Inhibitory Components from Soil ........... ll4
RESULTS .............................. ll4
Purification of Inhibitory Components in Asparagus Root
Extracts: Scheme 1 ....................... ll4
Bioassay of Known Standards and Cl8 Fraction. . . . ...... ll8
Purification of Inhibitory Components in Asparagus Root
Extracts: Scheme 2 ....................... lZl
Purification of Inhibitory Components in Asparagus Root
Extracts: Scheme 3 ..................... 126
Isolation for Inhibitory Components from Soil .......... l33
DISCUSSION ............................ T36
LITERATURE CITED ......................... T39
CHAPTER VI: SUMMARY AND CONCLUSIONS ................

vi

TABLE

LIST OF TABLES
PAGE

CHAPTER 2:

Experimental design for soil bioassays of the

response of indicator seedling species to dried

asparagus root or fern tissues incorporated

into sterile or nonsterile soil. . . . . . . . . . . . . 27

Percent germination and time of evaluation of

indicator species used in petri plate bioassays

with purified asparagus root extract from a

silica gel "flash" column. ..... . . . . . . . . . . 3O

Inhibition of root and shoot length of indicator
species in response to increasing concentrations
of a purified asparagus root extract . . . . . . . . . . 46

CHAPTER 3:

02 uptake after infiltration of asparagus root
tissues with distilled water or asparagus aqueous

extracts, or from submerging excised asparagus
roots in aqueous extracts or distilled water after
five hours . . . . . . . . . . . . . . . . . . . . . . . 66

CHAPTER 4:

Response of bacterial populations in Spinks sandy

loam soil amended with 50 g of dried asparagus

root or fern tissue or planted with asparagus,

snap beans, or sweet corn. . . . . . . . . . . . . . . . 87

Response of fungal populations in Spinks sandy loam

soil amended with asparagus root or fern tissue

or planted with asparagus, snap beans,

or sweet corn. . . . . . . . . . . . . . . . . . . . . . 89

Response of Fusarium spp. populations in Spinks

sandy loam soil amended with dried asparagus root

or fern tissue or planted with asparagus, snap

beans, or sweet corn . . . . . . . . . . . . . . . . . . 9l

vii

LIST OF TABLES Continued:

TABLE PACE
CHAPTER 4:
4 Inhibition of bacterial isolates by isolated
components of asparagus root extracts .......... 93

viii

FIGURE

LIST OF FIGURES

CHAPTER 11:

Response of lettuce, cv. Grand Rapids to

increasing concentrations of dried asparagus root
or fern tissue in sterile or nonsterile soil
incubated under aerobic or anaerobic conditions. . .

Response of curly cress root growth to increasing
concentrations of dried asparagus root or fern
tissue in sterile or nonsterile soil incubated

under aerobic or anaerobic conditions. . . . . . . .

Response of barnyardgrass root length to increasing
concentrations of dried asparagus root or fern
tissue in sterile or nonsterile soil incubated
under aerobic or anaerobic conditions. . . . . . . .

Response of smooth crabgrass shoot length to
increasing concentrations of dried asparagus root
and fern tissue in sterile or nonsterile soil
incubated under aerobic or anaerobic conditions. . .

Percent germination of smooth crabgrass in response
to increasing concentrations of asparagus root or
fern tissue incorporated in sterile or nonsterile
soil and incubated under aerobic or anaerobic
conditions . . . . . . . . . . . . . . . . . . . . .

CHAPTER III:

Root rot ratings of asparagus seedlings (cv. UC147)
grown in soil amended with increasing levels of
dried asparagus root tissue alone or in combination

with F. oxysporum f. sp. asparagi or
F. moniliforme . . . . . . . . . . . . .

 

Electrolyte efflux over time from excised asparagus
root tissue treated with increasing concentrations
of a water extract of asparagus root tissue. . . . .

Peroxidase activity of asparagus seedlings

infiltrated with increasing concentrations of a
water extract of asparagus root tissue . . . . . . .

ix

PAGE
. . 33
. . 35
. 38
. . 4T
. . 43
. . 59
. . 61
. 64

LIST OF FIGURES Continued:

FIGURE PAGE
CHAPTER III:

4 Root and shoot length of curly cress seedlings
treated with increasing concentrations of a water
extract of asparagus root tissue from 5, 12, and
20 yr old plantations ............ . ..... 67

5 Root length of curly cress treated with increasing
concentrations of autoclaved and nonautoclaved
water extract of asparagus root tissues from 5, 12,

or 20 yr old asparagus fields. . . . . . . ....... 70
CHAPTER V:
1 Chemical structures of plant growth regulators

and nematicides isolated from etiolated asparagus
fern tissues and asparagus root tissue . . . . . . . . . 124

2 Flow diagram for the extraction and partitioning of
dried asparagus root tissue. . . . . . . . . . . . . . . 107

3 Flow diagram for the separation of compounds from
the chloroform extract of dried asparagus root
tissue 0 O O O O O O O O O O O O O O O O O O O O O O O O 109

4 Response of curly cress to known concentrations of

solvent extracts from aqueous extract of dried

asparagus root tissue. . . . . . . . . . . ....... 116
5 GC-MS spectrum of peak collected from HPLC

fractionation of dried asparagus root extract
that was inhibitory to cress seed germination
and radicle elongation .............. . . . 119

6 1H NMR of a compound isolated from asparagus root
tissue using preparative TLC, mobile phase
chloroform:methanol 8.5:].5. Compound at Rf=0.07
was scraped from the plate and eluted from the silica

gel using chloroform:methanol 5:1. . . . . ....... 122
7 Mass spectrum of a compound isolated from asparagus

root tissue using preparative TLC, mobile phase

chloroform:methanol 8.5:l.5. . . . ....... . . . . 127
8 1H NMR spectrum of a compound isolated from

asparagus root tissue using soxhlet extraction . . . . . 129

9 1H NMR spectrum of a known standard of
caffeic acid dissolved in deuterated methanol. . . . . . 131

CHAPTER I
LITERATURE REVIEH

INTRODUCTION

Asparagus Decline in Michigan

Asparagus (Asparagus officinalis LJ isauiimportant vegetable

 

crop grown on sandy soils that_are not always considered suitable for
other cash crops. Traditionally, it was assumed that if properly
managed, plantations should stay commercially profitable for greater
than 20 years (Tiedgens, 1924, 1926; Hanna, 1947; Lougheed and
Tiessen, 1985). But, despite increased production due to increased
acreage, yields of asparagus are currently declining, causing great
consternation among asparagus growers. Today fields are being removed
from production after 6 to 8 years due to sparse stands and small spear
size resulting in lowjyields (Takatori and Souther, 1978). In 1978,
the average yield for Michigan was 1,500 pounds per acre whereas in
1980, the state average was 900 pounds per acre (Anon, 1977, 1980).
Today, only isolated commercial fields and experimental plots in
Michigan report high yields.

Reduced yields from declining fields have also been reported for
other production areas of the country. In New Jersey, asparagus
production has dwindled from 30,000 acres to less than 1,000 acres

(Herner and Vest, 1974). California, the largest producer of asparagus

2

in the United States, reports acreage planted to asparagus had
decreased from 44,000 acres in 1974, to 28,000 acres in 1978 (Takatori
and Souther, 1978). Declines in yields have also been reported for
other asparagus growing areas of the world (Lougheed and Tiessen,
1985). In the Netherlands, acreage planted to asparagus decreased from
500 ha to 340 ha in 1970 (VanBakel and Kerstems, 1970).

Most asparagus fields are planted with approximately 10,000 crowns
per acre, but 20% may be lost in the first five years after planting.
In a 1978 field survey of asparagus fields in Michigan, the average
crown population was 3,153 crowns per acre representing an overall 70%
reduction in crown survival (Hodupp, 1983).

In addition to the decrease in longevity and productivity of
etablished fields, experience has demonstrated that asparagus cannot be
replanted in soils where decline forced removal (Tf the crowns
(Tiedgens, 1924, 1926; Hanna, 1947L. Fields replanted immediately
after plowing out crowns produced yields never more than one half the
expected poundage, and when plants were direct seeded into old
plantations, seedling mortality was practically 100% after 2-3 months
(Hanna, 1947).

It is currently hypothesized that Fusarium crown and root rot is
ultimately the causal factor in the decline in longevity and vigor, as
well as inability to replant old asparagus fields (Cohen, 1946; Grogan
and Kimble, 1959; Molot and Simone, 1965; Endo and Burkholder, 1971;
Johnston, et. al. 1971). Decline is believed to be accelerated by
environmental, biological, and physical stresses on the plants that may
predispose them to infection by the Fusarium crown and/or root rot
organisms (Farrish, 1939; Takatori, et. al. 1970, 1974; Hartung and

Stephens, 1983, 1984; Evans and Stephens, 1984, 1985; Evans , et. al.,

1985; Robb, 1984). The Fusarium species implicated in the disease on

asparagus are _F_.oxysporum (Schlecht.) emend. Snyder and Hansen f. sp.

 

asparagi Cohen, and F_._ moniliforme (Sheldon) emend. Snyder and Hansen.

 

The Fusarium spp. are among the most cosmopolitan of the fungi and
are of great economic importance since they play a major role in
reducing yields and quality of many important food crops of the world
(Nelson, et. al., 1981). The fusaria are capable of surviving in the
soil almost indefinitely as chlamydospores or other resting structures
(Nelson et. al., 1937; Booth, 1971; Nyvall and Kommedahl, 1968). They
are facultative parasites which colonize living and non-living host
tissue and are also capable of invading non-host tissues (Alexander,
1961; Hendrix and Nielson, 1958). Many of the modern fusarium diseases
such as corn stalk rot are caused by a complex of organisms aided and
abetted by a plethora of environmental factors and cultural practices
(Toussoun and Patrick, 1963; Nelson, et. al.,1981). Loxysporum is
the most frequently isolated of the fusarium species in soils and is
very active as a saphrophyte (Nelson, et. a1. 1981). F_. moniliforme is
distributed throughout the world but is most common in the warmer
regions. It is a major parasite on several species of the Poaceae,
particularly rice. It is often found in association with other fungal

organisms, particularly Loxysporum, acting in consort with them to

 

produce disease (Hendrix and Nielson, 1958; Booth, 1981; Nelson, et.
a1. 1981).

Fusarium crown and root rot is considered to be the most common
disease of asparagus in the United States (Walker, 1952). The above-
ground symptoms caused by both pathogens include yellowing, stunting

and/or wilting of the ferns. The ferns may die in various stange of

4
elongation (Cohen, 1946, Endo and Burkeholder, 1971). Symptoms on

asparagus,associated with f: oxysporum f. sp. asparagi are elliptical
reddish-brown lesions found at the base of the ferns and vascular

discoloration within stems, roots and crowns. f; monilifbrme causes dry

 

crown rot and brown stem pith discoloration but no vascular
discoloration (Johnston, et. al., 1979L. Tbtal root collapse can occur

when seedlings are inoculated with f: moniliforme. f; moniliforme was

 

 

once considered primarily important in older asparagus plantations, but
recent research indicates this pathogen may also be the primary
pathogen in one-year old crowns (Damicone and Manning, 1985).

Efforts to develop chemical controls for the Fusarium pathogens on
asparagus have not proven successful. Soil fumigation and seed
treatment reduce but do not eliminate decline symptoms (Damicone and
Cooley, 1981; Manning and Vardaro, 1977; Lacy, 1979L. Control is also
confounded by the perennial nature of asparagus and the difficulty of
combatting the pathogen in the soil without damaging the plant. In
crops already studied, the most successful strategy for Fusarium wilt
control has been the development of resistant varieties. Over 26
Fusarium wilt resistant vegetable varieties have been developed through
plant breeding efforts (Mace, et. al., 1981). Asparagus is not
considered to be isogenic and genetic variability found within the
species and availability of germplasm from wild species should produce
some genetic resistance within the species (Tiedgens, 1924, 1926;
Robb, 1984L. However, efforts world wide for the past 20 years of
breeding have not produced any truly resistant varieties (Lougheed and

Tiessen, 1985).

Allelopathy

Plant residues from various sources contribute to the organic
matter component of the soil. These tissues from young, mature and
dead plant parts are ultimately decomposed in the soil by chemical or
microbial breakdown. Thus, chemical components previously bound in the
plant materials are made available fOr microbial and plant uptake. At
any one time, the soil may contain a vast array of chemical components
which may affect plant growth in either beneficial or detrimental ways.

Decomposition of plant residues in the soil associated with the
formation of phytotoxic substances has been widely documented (Borner,
1960; McCalla and Haskins, 1961; Patrick and Koch, 1963; Toussoun and
Patrick, 1963; Schroth and Hilderbrand, 1964; Toussoun, et. al., 1968;
Lindermann, 1970; Patrick, 1971; Putnam and Duke, 1978; Stowe, 1979;
Rice, 1979, 1984). In the early 1900's, DeCandolle (1932) had
suggested that 'soil sickness' was due to crop plant exudates. In the
past 25 years, a large amount of scientific effort has been devoted to
elucidating how root exudates. leachates from living plants, and plant
residues affect plant growth, interference between plant populations
and microbial populations. This biochemical interaction that plants
exert on other plants is called allelopathy. Allelopathy was first
defined as any chemical effect of one plant upon another plant
(Molisch, 1937). Allelopathy can be separated from other forms of

plant interference in that the detrimental effects are exerted through

5

6

the release of a chemical into the environment (Muller, 1969; Fuerst
and Putnam, 1983; Rovira, 1969; Whittaker and Feeny, 1971).
Allelochemical stresses upon crop plants can reduce crop emergence,
growth and yield, as well as resistance to plant pathogens (McCalla and
Haskins, 1970; Linderman 1970; Bell, 1974L. The science of allelopathy
has recently been the focus of several books (Rice, 1984; Thompson,

1985; Putnam and Tang, 1986).

Al lel apathy and Ni croorgani sas

Allelopathic effects appear to be especially important in natural
communities dominated by a single species (Whittaker, 1975). However,
relatively few reports have been published that examine how
a1 lelopathic compounds may affect microorganism, particularly plant
pathogens. Unidentified fungitoxic exudates on leaf surfaces are
believed to offer limited protection against plant pathogens (Barbosa
and Saunders, 1986).

In an examination of the peach tree decline and replant problem,
Chandler and Daniell (1974) found peach seedlings grown in either old
peach soil or in the presence of peach soil leachates were more
susceptible to infection by Pseudomonas syringae than seedlings grown
in control soils or soils from a pecan orchard. They postulated that
toxins from dead peach roots may predispose new trees to bacterial
canker and thus contribute to peach decline. Amygdalin, a cyanogenic
glycoside, was isolated from peach roots and believed to be the primary
source of toxic substances present in the soil (Patrick, 1955). In the
presence of enzymes provided by the microbial population in the soil,

amygdalin is cleaved in two places producing two toxins, hydrogen

cyanide and benzaldehyde. He demonstrated that young peach roots are
extremely susceptible to damage by these compounds while other Prggus
spp. are less affected.

Exposure of tobacco plants to leachates obtained from decomposing
rye and timothy residues increased the susceptibility of both resistant
and susceptible varieties of tobacco to black root rot caused by

Thielaviopsis basicola (Beck and BrJ Ferraris (Patrick and Koch,

 

1963). Using 16 different tobacco varieties ranging from susceptible
to resistant to black root rot and 6 different isolates of'];_basicola,
they showed that exposure to the leachates overcame resistance of the
tobacco to the pathogens. They postulated that the leachates were
damaging the tobacco roots and, therefore, making the plants more
suseptible for infection and colonization by the pathogen. They
hypothesized toxins produced by rye or timothy were responsible for the
breakdown of resistance seen in fields since rye and timothy were often
used in rotation with tobacco. They concluded these toxins may be an
important predisposition factor in the disease syndrome.

Toxic substances produced from decomposing residues of rye,
barley, broccoli, and broad bean greatly enhanced the pathogenesis of
E; solani (Mart.) Sacc. f.sp. phaseoli (Burk) Snyder and Hansen on
beans (Toussoun 1963). Disease enhancement, measured by"lesion
development on bean stems, was greatest using extracts obtained during
the early stages (less than 1 month) of decay of the residues. They
postulated this enhancement was due to an additive effect of the
extract and the pathogen; the extract was preconditioning the roots to
fungal invasion. They noted that root rots are not necessarily caused
by specific pathogens and hypothesized that organisms ordinarily

causing little damage might become more pathogenic if conditions were

8

favorable for pathogen development. Other experiments showed these
toxins had a direct effect on the host cells, altering the cell
permeability. They concluded the resulting increased exudation of
ninhydrin-positive compounds and other substances were readily
available to organisms in the infection court and were mainly
responsible for predisposing the host to infection by pathogenic

organisms.

Asparagus as an Allelopathic Plant

Recent investigations of the causal factors of asparagus decline
have suggested that allelopathic compounds are present in asparagus
root tissue (Laufer and Garrison, 1977; Shafer and Garrison, 1980a,
1980b, 1986; Yang, 1982, 1985; Young, 1984; Hartung and Stephens, 1983,
1984; Hodupp, 1983; Hartung and Putnam, 1985L. Several studies have
shown thataillelochemicals released from both the growing asparagus
plant and from the senescing root tissue are not only allelopathic to
other plant species but are also autoallelopathic or autotoxic (Laufer
and Garrison, 1977; Shafer and Garrison, 1980a, 1986; Yang, 1982, 1985;
Hartung and Stephens, 1983; Hartung and Putnam, 1985). Water extracts
from asparagus seedlings grown from tissue cultured plants inhibited
the growth of asparagus seedlings grown in plastic pouches (Yang,
1982). Also, evidence shows that asparagus tissues incorporated into
soil increase the incidence of asparagus root rot on asparagus
seedlings (Hartung and Stephens, 1983). In the presence of dried
asparagus root tissue incorporated into soil with or without either of
the Fusarium pathogens, asparagus seedlings had more root rot than seen

on control plants. The dry weight of those treatments was also less

than those of control plants. The Fusarium pathogens, in contrast to
other soil-borne microorganisms, appeared insensitive to the
al 1e10pathic substances in the root and fern tissue (Hartung, 1983).
Shafer reported that toxicity from asparagus root tissue decreased in a
soil medium over time but made no attempt to determine if degradation
was primarily through microorganisms (Shafer and Garrison, 1986).
Also, they did not examine any effects from fern tissue.

In studies utilizing an XAD-4 resin trapping procedure originally
developed to recover phenolic and chlorophenoxy herbicides from soil,
compounds inhibitory to asparagus seedlings were collected from healthy
asparagus plants grown in sand over a period of 6 months (Young, 1984).
No differences were noted between 3 varieties of asparagus. In another
report, he found that asparagus seedlings planted in unsterilized
”used” vermiculite and treatments with root tissues alone incorporated
into vermiculite, started to "yellow and wilt" after 21 days but were
normal in controls and treatments with sterilized vermiculite and root
tissues (Young, 1986). He postulated that there is an interaction
between the residues and microorganisms. However, the exact nature of
the ”used vermiculite“cn'its microbial content was not published,

making it difficult to assess the results.

Isolated and Identified Conpounds froo Asparagus Tissue

Several compounds that may contribute to the autotoxic properties
of asparagus have already been identified. Asparagusic acid, a 1,2-
dithiolane, dihydroasparagusic acid and s-acetyldihydroasparagusic acid

were isolated and identified from etiolated asparagus tissues by a

10

Japanese group in 1972 (Kitahara, et. a1, 1972; Yanagawa, et. al.,1972;
Yanagawa, et. al, 1973a). Asparagusic acid inhibited lettuce root and
hypocotyl growth at 6.67 x 10'4 M, and was active on rice, rape,
radish, carrot, and barnyardgrass at similar concentrations. It's
activity closly parallelled that of abscisic acid. The structure and
activity of asparagusic acid was confirmed by synthesis (Yanagawa, et.
al., 1973b; Yanagawa, 1979). Dihydroasparagusic acid and S-
acetyldihydroasparagusic acid were also isolated and their structures
confirmed by synthesis (Yanagawa, 1973a, 1973c). They are reported to
be inhibitory at the same concentration as asparagusic acid. In one
report, these compounds were compared in activity to lipoic acid which
is known to participate in the transfer of acyl groups. Since the
asparagus compounds were similar to lipoid acid in structure, they
examined if lipoic acid could be replaced with asparagusic acid or its
derivatives (Yanagawa, et. al., 1973c). They used Streptococcus
faecalis 10Cl since its growth is stimulated by lipoic acid.
Asparagusic acid did increase growth of the bacteria and also
stimulated the rate of pyruvate oxidation but at concentrations 10,000
times that of lipoic acid. They also examined the effects of
asparagusic acids on the stimulation of pyruvate oxidation in asparagus
mitochondria as compared to lipoic acid (Yanagawa, 1973a). Both
asparagusic acid and dihydroasparagusic acid stimulated pyruvate
oxidation but lipoic acid showed no effect. These effects were
opposite those obtained with §;LE!EELLI§.IOCI- Asparagusic acid is
biosynthesized by a different pathway that that of lipoic acid (Perry,
et.al. 19821. Other research has shown that dihydroasparagusic acid
promoted rooting of mung bean cuttings at low concentrations (10'9

~10'5), and killed roots at higher concentrations (10'4) (Kuhnle, et.

11

al., 1975a). This group also reported that dihdroasparagusic acid

retarded germination of Pisum sativum cv. Early Alaska, but no effect

 

from the compound was noted when plants were grown to first anthesis
from seeds (Kuhnle, et. al. 1975b).

Asparagusic acid has also been isolated from the roots of
asparagus.and found to be inhibitory to nematodes, specifically the

emergence of the second stage larvae of Heterodera rostochiensis and H;

 

glycines, and the second stage larvae of H; rostochiensis and

Meloidogyne hapla, and the larvae and adults of Pratylenchus penetrans

 

and Pratylenchus curvitatus (Takasugi, et. al.,1975).

 

Young has reported that several phenolic compounds are released
from the root system of intact asparagus plants (Young, 1986). Using
an XAD-4 resin column (originally developed to recover phenolic and
chlorophenoxy herbicides) attached to pots of sand where asparagus was
growing, he isolated 3,4-dihydroxybenzoic acid, 2,6-
dimethoxyacetophenone, and 8-(m-hydroxypheny1)propionic acid. However,
he did not report if these compounds were inhibitory to asparagus
seedlings.

Though compounds isolated by the Japanese workers are considerably
toxic to germination and radical elongation of several species, these
compounds occur at extremely low concentrations in the tissues. Since
asparagus possesses large storage roots that are continually dying as
the crown grows, large amounts of asparagus root material may be
present in asparagus fields at any one time. At this point, no one has
isolated a compound or compounds that can reasonably account for all
the inhibitory activity present in asparagus root tissue. Even though

several compounds have been isolated from asparagus tissues or

12

artificial growth media, none have been shown to be active in the
asparagus rhizosphere. In addition, no detailed biochemical work has
been initiated to determine if components in root extracts affect
Specific sites in the asparagus tissue. Also, even though some
research has concentrated on how asparagus tissues are degraded in the
soil, little attention has been devoted to the fate of these components
in natural soil and how microorganisms may be affecting the toxic
substances present in the tissues.

In addition, there is also a need to further investigate how toxic
components of the tissues may be affecting microbial populations in the
soil. If asparagus extracts are inhibitory to the general microbial
population and not to the Fusarium pathogens, this may decrease
competition among microorganisms in the soil, thereby giving the root
pathogens a competitive edge in the agroecosystem where asparagus is
grown. It is the purpose of this dissertation to examine these
questions, and to more fully elucidate the role of allelopathy in the

asparagus decline syndrome.

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eds., John Wiley and Sons, Inc. NY. pp. 317.

CHAPTER II

ALLELOPATHIC POTENTIAL OF ASPARAGUS RESIDUES IN
STERILE AND NONSTERILE SOIL

ABSTRACT

Asparagus, (Asparagus officinalis L.) has been reported to have

 

allelopathic potential and compounds released from its tissues were
hypothesized to increase Fusarium root rot on asparagus seedlings.
Dried asparagus root and fern tissues were incorporated into sterile or
nonsterile soil to determine if microbial populations influenced their
inhibitory activities. Four different indicator species; Lepidium
sativum, cv. Curly Cress, Lactuca sativum, cv. Grand Rapids, Digitaria
ischaemum, and Echinochloa crusgalli were utilized in bioassays.
Species responded differently to compounds released from the root and
fern tissues when grown under aerobic or anaerobic conditions, as well
as sterile or nonsterile soil conditions. Both tissues were inhibitory
to seedling development under both aerobic and anaerobic conditions.
Fern tissue was less inhibitory than root tissue to seedling root and
shoot growth. Microbial populations in the soil generally degraded
tissue toxicity, but in one case slightly increased its inhibitory
activity for a short time. Purified extracts of asparagus root tissues
were inhibitory to nine of ten indicator species. Data from purified

extract experiments agreed with data from soil bioassays.

22

INTRODUCTION

In the past eight years several research papers have reported
asparagus (Asparagus officinalis LJ to be an allelopathic plant
(Laufer and Garrison, 1977; Shafer and Garrison, 1980a; 1980b; 1986;
Yang, 1982; Hartung and Stephens, 1984; Hartung and Putnam, 1985;
Young, 1986). Allelopathy has been defined as the detrimental
influence of one plant on another through the release of chemicals into
the environment (Molisch, 1937).Chemical interference by compounds
released from plants has been identified as an important component of
plant interference in natural and agroecosystems (Borner, 1950; Tukey,
1969; Rice, 1984, 1979; Putnam and Duke, 1978;). Allelochemical
stresses upon crop plants can reduce crop emergence, growth and yield,
as well as resistance to plant pathogens (Bell, 1974; Linderman, 1970;
McCal 1a and Haskins, 1970.). In the case of asparagus, not only are
these compounds thought to be detrimental to other plants, but are also
considered to be autotoxic or autoallelopathic (Yang, 1982; Hartung and
Putnam, 1985; Shafer, 1980b; Young, 1984). Shafer (1986) reported that
toxicity from asparagus root tissue decreased in a soil medium over
time but made no attempt to determine whether microorganisms may have
degraded the compounds released from the root tisssue.

When asparagus was planted in soil amended with sterilized dried
root or rhizome tissue in the presence of Fusarium oxysporum
(Schlecht.) Snyd. and Hans. f.sp. asgaragi Cohen or f_._moniliforme

Sheldon Snyd. and Hans., there was a significant increase in the

23

24

severity'of root rot caused by these two pathogens andaadecrease in
the dry weight of the asparagus crown (Hartung and Stephens, 1984).
These data suggest that chemical components released directly from the
root tissue or altered by microorganisms were either affecting the
asparagus plant directly or interacting with microflora of the
rhizosphere to increase the susceptibility of the asparagus plant to
infection by the disease organisms. Therefore, this research was done
to determine the fate of the compounds released from the asparagus root
tissue in the soil, to determine the influence of soil microorganisms
on the toxicity of the root and fern tissue under both aerobic and
anaerobic conditions, and to assess the effects of purified asparagus
root extracts on seed germination and radical growth of different

seedling species.

MATERIALS AND METHODS

Preparation of Asparagus plant material. A commercial asparagus field
(<20 yr old) in Oceana County, MI was excavated and asparagus crowns
(cv. Martha Washington) were collected. The storage and fibrous roots
were separated from the rhizomes and washed after which all dead and
visably diseased plant material was discarded. The roots were oven
dried at 40 C and ground in a Wiley Mill (mesh screen size 1 mm). Fern
tissue was clipped from the crowns and prepared in the same manner as
the root tissue. All plant tissues were sterilized with propylene
oxide gas (Tuite, 1969). Containers with sterilized tissue were
allowed to exhaust under a fume hood for 24 hr to dissipate all
propylene dioxide. Dried tissues were stored in brown glass bottles at

-20 C until needed for petri plate bioassays or extractions.

25

Assess-ant of microbial contalination of asparagus tissues. To
ascertain that sterilized tissues were free from Fusarium sp., or other
contaminating microorganisms, 1 g of dried tissue was spread evenly
over the surface of a 9 cm petri plate containing Komada's medium
(Komada, 1975), potato dextrose agar (PDA) (39 g PDA, Difco
Laboratories, Detroit, MI, in 1 liter distilled water (0H20)L There
were 4 replica plates per tissue. After one week, plates were assessed
for the number of fungal colonies growing on the medium. Plates were

examined every day for contaminating organisms for ten days.

Toxicity of asparagus tissues in soil. Spinks sandy loam soil
(Psammentic Hapludalf, sandy, mixed, mesic, with 1% organic matter, pH
(L5) collected from a non-agricultural site, was air dried, sifted
through a (L5 mm screen, weighed into 150 9 samples and either
tinsdillated (one hr each time at 120 C; 23 psi) or left nonsterilized.
Sterility of the soil was confirmed by randomly selecting five
different soil samples and plating out two dilutions/soil sample on PDA
and observing for microbial growth over a period of 14 days.

In order to evaluate the fate of the toxic substances present in
asparagus tissues when exposed to uficroorganisms, tissues were
incorporated into sterile and nonsterile soil and exposed to anaerobic
and aerobic conditions. To assess the initial tissue toxicity, a "non-
incubated treatment" was done for each set of experiments. Tissues
were incorporated into the soil and indicator species placed
immediately on the soil surface. Asparagus root or fern tissue
previously sterilized using propylene oxide gas was then mixed
thoroughly with the soil samples at 0.5%, 1% and 2% by weight and

poured into square petri plates (10 x 10 x 1.5 cm). Sterile DHzo was

26

added to each plate at a water potential of -0.01 Bars for each
treatment. Paper toweling cut into 1 cm fragments was also
incorporated into the soil at similar rates to serve as a control for
physical influence of the tissues. Filter papers (Whatman #1) were cut
to cover one half of the surface of the soil and ten seeds each of two
indicator species were placedcnithe filter papers. The plates were
placed vertically in a moist chamber (to induce geotrophic growth) for
72 hr then root and/or shoot length or percent germination were
measured for each indicator species.

To ascertain the effects of microorganisms under aerobic and
anaerobic conditions, two more treatments were included in the
experiment. A similar group of the plates not seeded with the
indicator species was incubated in a moist chamber for 72 hr at 27 C in
the dark and designated the "aerobic treatment". The remaining plates
without the indicator species were placed in a desiccation chamber and
flushed repeatedly with nitrogen to assure all oxygen was removed from
the chamber. The plates were incubated under anaerobic conditions for
72 hr in the dark. This group was designated the “anaerobic
treatment". Both the aerobic and anaerobic treatments were then
planted as before with the same two seed species as the nonincubated
treatment, bioassayed and measured as above (Table 1). Seedling
species used as indicator species were lettuce, Lactuca sativa L. cv.

Grand Rapids, Lepidium sativum L.cvu Curly Cress, smooth crabgrass,

 

Digitaria ischaemum (Schreb.) Muhl., and barnyardgrass, Echinochloa

 

crusgalli (LJ Beauv. These species were chosen for uniformity of

 

germination rates, and as representatives of both monocot and dicot
palants. There were three replications for each treatment, and the data

were subjected to an analysis of variance and trend analysis. The

Table 1.

27

Experimental factors for soil bioassays of the response of
indicator seedling species to dried asparagus root or fern
tissues incorporated into sterile or nonsterile soil. The

design was a randomized complete block with 3 replications.

 

 

Incubation Soil Condition Tissue Level (%)
Not Incubated Sterile Root 0.0
Anaerobic Nonsterile Fern 0.5
Aerobic 1.0

2.0

 

28

experiment was repeated twice and data presented is from one

representative experiment.

Preparation of purified asparagus root extracts. Previously published
data from several laboratories have indicated that water extracts of
asparagus root tissues are inhibitory to seedling germination and
radical elongation (Yang, 1982; Hartung and Stephens, 1984; Hartung and
Putnam, 1985 Shafer and Garrison, 1986). Therefore water extracts from
asparagus root tissue were subjected to chemical extraction to purify
the chemical components implicated in the inhibitory responses. All
fractions throughout the isolation procedure were bioassayed on curly
cress to determine their relative biological activities. Ground
asparagus root tissue was extracted by stirring overnight at 4 C (1:10
VOOt t15$U€=DH20). The particulates were then removed by filtering
through four layers of cheese cloth and centrifuging at 6,000 g for 20
min. The supernatant was decanted carefully from the pellet (the
pellet was discarded) and then precipitated with acetone (4:1 v:v
acetone:sample) overnight at 4 C. The precipitant was discarded and
the liquid concentrated to 1/4 the original volume on a Buchi Rotary
evaporator. This concentrate was extracted with chloroform 3 times
(1:1 v:v). The chloroform fraction was reduced to dryness, weighed,
redissolved in methanol, filter sterilized, and then bioassayed using
curly cress as an indicator. For bioassay, the fraction was dissolved
in methanol at known concentrations and applied to filter paper in 6 cm
petri plates. The methanol was allowed to evaporate, then 1.0 ml of

0H20 and 10 curly cress seeds were added to each plate. The chloroform

‘fraction was further purified using an octadecyl bonded phase solid

29
support in a Baker flash chromatography column (190 mm x 20 mm) eluted
with a step gradient of Acetonitrile 100% to Methanol 100% at 25%
intermediate steps in 100 m1 fractions. Pressure for the column was
provided by laboratory airline at a rate of 0.2 cm/sec. Fractions (25
m1) from the column were collected, spotted on thin layer
chromatography plates (TLC) (Whatman silica gel 60 F-254; mobile phase,
chloroform:methanol, 9:1) then recombined according to similar Rf
values when compared under TLC as above. The fractions were dried
under nitrogen, weighed, and bioassayed. The inhibitory fractions were
combined and separated further on a silica gel solid support in a Baker
flash chromatography column (180 )l 30 mm) using a gradient of
chloroform 100% to methanol 100% in the following order: 100%
chloroform, 98:2, 96:4, 90:10, 85:15, 80:20, 70:30, 40:60, 100%
methanol. Each gradient step was applied to the column in 200 ml
fractions. Air pressure for the column was as above. Fractions of 75
ml each were collected and developed on TLC plates as above. Fractions
with similar Rf values were combined, dried under nitrogen, weighed and
bioassayed on curly cress as above. One inhibitory fraction from the

column was used in bioassays using all the seedling species.

Bioassay of partially purified compounds on indicator seed species. .A
preliminary bioassay was done with ten different seed species to
determine the optimum time of germination and optimum number of seeds
necessary to produce relevant data in the bioassay studies (Table 2).
All seed species were bioassayed at concentrations of 150, 100 and 50
PPM-

For bioassays, the inhibitory fraction was resuspended in

methanol, diluted to the proper concentration, then spotted on Whatman

30

Tank: 2: Percent germination and time of evaluation of indicator
species used in petri plate bioassays with purified

asparagus root extract from a silica gel “flash" column.

 

 

 

 

 

 

 

 

 

 

Estimated Incubation Seeds per
Species Germination(%)a period Petri plate
Lactuca sativa L., 85 72 25
cv Grand Rapids
Raphanus sativus L., 100 72 20
cv Cherrybell
Amaranthus retroflexus L. 40 72 50
Lepidium sativum, 100 72 20
cv curly cress
Portulaca oleracea L. 90 96 25
Lycopersicon esculentum, L. 85 96 25
cv Lafayette
Setaria italica (L.) Beauv. 85 72 20
Digitaria sanguinalis (L.) Beauv. 65 96 30
D. ischaemum (Schreb.) Muhl. 60 96 35
Echinochloa crusgalli (L.) Beauv. 40 96 50

 

 

abased on a sample of 100 seeds/species.

31

#1 filter paper in a 60 x 15 mm sterile petri dish. The methanol was
allowed to evaporate from the filter paper, then 1.5 m1 of DHzo was
added to the plate. The seeds to be tested were then placed in the
dishes and spread evenly around the plates with a glass rod. After the
designated incubation period, seeds were evaluated for root and shoot
length as well as percent germination. Solvent and DHzo controls were
also included in each assay. Due to scarcity of isolated components,
there were two replications per treatment. Data were subjected to
Analysis of Varience and trend analysis if applicable. The sol vent

control was used as the zero level in all trend analysis.

RESULTS AND DISCUSSION

Assessment of microbial contamination of asparagus tissues. After a
period of ten days, no Fusarium spp. or other microbial colonies were
present in asparagus tissue samples sterilized with propylene dioxide

gas.

Toxicity of asparagus tissues in soil. Within each seedling species,
germination and growth that occurred with paper toweling and the no-
tissue controls were not significantly different. Therefore, results
obtained were not attributed to the physical presence of tissue in
soil. As expected, indicator species varied in their response to
asparagus root and fern tissue when these tissues were incorporated
into sterile and nonsterile soil. In almost all instances, the third
order interactions of tissue level x tissue source x soil condition

were significant. For this reason and to provide consistency in

32

presentation, the graphs showing these interactions are depicted
throughout. For lettuce there was highly significant interaction for
the linear influences of tissue level x tissue source x soil condition
(Figure 1). There was also an interaction between treatment type and
soil condition. Initially, root and fern tissues were equally
inhibitory to root growth in the nonincubated treatment. However, root
tissue showed decreased inhibitory activity under nonsterile conditions
in both aerobic and anaerobic incubations as compared to the
nonincubated treatments. Fern tissue was equally'toxic to lettuce
radical growth under all treatment regimes. These data indicate that
lettuce seedlings were less susceptible to components released from the
root tissue than to components present in the fern tissue.

For cress root growth, there was a highly significant interaction
of incubationitsoil condition x tissue level (Figure 2L.Under all
treatments, root and fern tissue under nonsterile soil conditions were
equally toxic to cress root growth. Trend analysis revealed the
response was primarily linear (F=29.45 for linear response). Under
anaerobic and sterile conditions, the inhibitory activity of fern
tissue was greater than root tissue except at the lowest level of
incorporation of tissue into soil. As with lettuce, there was also a
highly significant interaction between soil condition x tissue type x
tissue level. Inhibitory activity from root tissue was apparently not
affected by resident microbial populations in the nonsterile soil
treatments. However, in all treatments, inhibitory'activity’of'the
fern tissue was significantly decreased when exposed to a nonsterile
soil condition and this response was quadratic. Although lettuce and
cress generally responded alike, they did respond differently to the

(lifferent asparagus tissues incorporated into the soil. Cress was most

Figure

1:

33

Response of Lettuce, cv. Grand Rapids to increasing
concentrations of dried asparagus root or fern tissue in
sterile or nonsterile soil incubated under aerobic or
anaerobic conditions. Not incubated treatment served as a
control for initial activity present in the tissues.
Interaction of tissue levels x tissue type x soil
condition; and treatment type x soil condition were

significant at P=0.01.

 

 

34

Root Length of Lettuce,
c.v. Grand Rapids (mm)

I 302.9015
9!... 303203335
OIO moqabfioqzo
Clio «23.203315

 

 

 

 

 

Pm db Nb Pm f0

 

 

 

22 50:33.. 3.90363 >032.“

>mumqmmcm x09 2 non: immcm
6:08 mos

35

Fimue 2: Response of Curly Cress root growth to increasing
concentrations of dried asparagus root or fern tissue in
sterile or nonsterile soil incubated under aerobic or
anaerobic conditions. Interactions of incubation x tissue
source x soil condition (LSD=A); and soil condition x
tissue source x tissue level (LSD=B) were significant at

P=0.01.

36

        

rmU >

ObGH—H H—Hobd
Pom—.1. Hob.—

..m—u m

N (O
O O
F

Root Length of Curly Cress (mm)
3

 

20” 520030

5503020 300* 0a «03 4.00:0

a
I 300090420

0.1.... 10002033050
oilo 0.03.9025
10:36 m03.20:«»0_.=0

 

  

 

 

Pa 3 .0 Po
3.00320

8:80 02..

 

  

 

. _
9m ._ .0
>032."

v.0

37

inhibited by root tissue, whereas lettuce was inhibited more by the
fern tissue.

Cress shoot growth showed similar responses with the fiallowing
exceptions (data not presented). Under anaerobic and nonsterile soil
conditions, inhibitory activity from both root and fern tissue was
greater than in aerobic or nonincubated controls. Under sterile
conditions, fern tissue was more inhibitory than root tissue, while
under nonsterile conditions root tissue was more inhibitory except at
the highest concentrations of tissue incorporation. These data suggest
that inhibitory compounds more active on shoot growth of cress may be
degraded more quickly by the resident aerobic microbial population
for fern tissue. However, compounds from root tissue are not degraded
by the natural microbial populations.

For root growth of barnyardgrass, there was a significant
interaction between tissue type x soil condition x tissue level (Figure
3). No other interactions showed significance. Root tissue showed less
activity under nonsterile soil conditions in both the anaerobic and
aerobic treatments. In the nonincubated treatment, fern tissue was
more inhibitory under nonsterile soil conditions. The data indicate
that compounds present in the fern tissue are more inhibitory to
barnyardgrass root growth, and that inhibitory activity may be greater
upon initial breakdown by microbes but is quickly degraded as time
elapses. Shoot growth for barnyardgrass was also inhibited
significantly by fern tissue but not by root tissue (data not shown).

Since a preliminary experiment indicated that germination and
growth of smooth crabgrass seeds varied considerably more than other

species, 100 seeds were used in bioassays with this indicator species.

Figure 3:

38

Response (If barnyardgrass root length to increasing
concentrations of dried asparagus root or fern tissue in
sterile or nonsterile soil incubated under aerobic or
anaerobic conditions. Non-incubated treatments served as
controls for the initial inhibitory activity present in the
tissues. Interaction of tissue type x soil condition x

tissue level were significant at P=oxn.

 

39

I 300090.20
0.-.... 30002030350
OII.O 303.9050
0......0 30320303420

 

 

 

Barnyardgrass Root Length (mm)

 

 

 

 

 

 

 

 

. _
Pm . .0 Po
20" 520200 2.00.620

>000q0mc0 300” on 30.5 4.00:0
8: 000 m2:

P p
0.0 ._ .o
>0~0§0

Nb

40

Analysis of smooth crabgrass shoot growth showed significant
interactions only between incubation type x tissue level, and soil
condition x tissue level (Figure 4). There was no inhibitory activity
from asparagus tissue on smooth crabgrass except at the higher levels
of root tissue incorporation under anaerobic and sterile soil
conditions or fern tissue under aerobic and sterile conditions.
Therefore, compounds responsible for inhibition of shoot growth can be
degraded by the microbial populations in the soil.

Germination of smooth crabgrass.was also reduced, particularly'
with root tissue (Figure 5). In the nonincubated treatments, root and
fern tissue were equalljrtoxic.to germination. Root tissue did not
inhibit germination under either sterile or nonsterile soil conditions
in the aerobic treatment, but was highly inhibitory under anaerobic
conditions. Fern tissues remained only slightly inhibitory to
germination in both sterile and nonsterile conditions, but did not
inhibit germination greater than 50% under any treatment conditions.
These data suggest that fern tissue is only marginally toxic to
germination of smooth crabgrass and that toxicity is either unaffected
by microbial breakdown in the soil, or that compounds responsible for
the inhibitory response are at low concentrations in the asparagus fern
tissues. Also, toxicity from root tissue, although more inhibitory
initially when compared to fern tissue, is degraded by the microbial
populations in the soil, and therefore loses its toxic properties.

Data from all these bioassays indicate differential response by
plant species to toxic substances released into the soil from both root
and fern tissues of the asparagus plant" Several reports have been
published concerning toxicity from root tissue but little research has

been done concerning activity from fern tissue. Fern residues did not

Figure

4:

41

Response of Smooth Crabgrass shoot growthix>increasing
concentrations of dried asparagus root or fern tissue in
sterile or nonsterile soil incubated under aerobic or
anaerobic conditions. Not incubated plates served as
controls for initial inhibitory activity present in the
tissues. Interaction between incubation type x tissue
level; and soil condition x tissue level were significant

at P=0.05.

42

I 30009025
0.1.. 300" 2030525
0110 30.3.9025
0.110 3032030525

.L
01
I
T

l

 

.5
O
T
9
O
01
o
O
.5

 

 

    

OI

 

_ b

p — . -
Pm . .0 Po Pm . .0 Po 0.0 ._.o No
20» 5030050 >300305 >0..00.o

 

 

 

_

 

 

 

Smooth Crabgrass Shoot Length (mm) ‘

>0omqmmc0 300" 0.. 30.3 100:0
5: com 003

sin;
e in
c or
d 15
1 the
5518
can‘

Figure 5:

43

Percent germination of smooth crabgrass in response to
increasing concentrations of asparagus root or fern tissue
incorporated in sterile or nonsterile soil and incubated
under aerobic or anaerobic conditions. Not incubated
plates were controls for the initial inhibitory activity
present in asparagus tissues. Interactions of incubation
type x soil condition, and tissue type x level were

significant at P=0.05.

44

.a
O
0

Percent Germination 0f Smooth Crabgrass

 

 

 

 

 

I 303.9025
Oil-o 3032030525
Clio 30009025
01110 30002030525

 

 

 

P — _ _ b b p _ L

Pm .b 0.0 Pm ..o P0 0.0 ._.0 Po
20» 30:55: >300~005 .5305

>000..0m:0 300» ca 303 4.00:0
a) com m0...

45

inhibit growth of asparagus seedlings (Yang, 1982). In these studies,
fern tissue was shown to inhibit both dicot species and one monocot
species differently depending on sterility of the soil. Toxic activity
present in both the tissues was also shown to be decreased in some
treatments when exposed to nonsterile soil conditions. These data
suggest several conclusions: a) there are probably several inhibitor
compounds present in the root and fern tissue that may be responsible
for allelopathic properties of asparagus. b)some plant species, e.g.
lettuce possesses more tolerance to the toxic principles in the
tissues, c) some of the al lelochemical activity may be destroyed by
microbial breakdown, or d) increased activity may occur under anaerobic
conditions as shown by the inhibition of lettuce root growth by fern

tissue.

Bioassays of partially purified compounds on indicator seed Species.
Purified fractions from a silica gel flash column on four monocot and
six dicot plant species revealed 150 values.for root length ranging
from 28 ppm for lettuce and common purslane to 130 ppm for Japanese
"111 let (Table 3)- 15olevels for shoot length ranged from 30 ppm for
common purslane to 125 ppm on Japanese millet. Only barnyardgrass was
not significantly inhibited by the purified extract, which agrees with
the lack of sensitivity of this species in soil bioassays. These
results agree with reports concerning other allelochemicals on
barnyardgrass (Barnes and Putnam, 1986; Weston and Putnam, 1986). Both
foxtail millet and tomato had the high 150 levels for root length, both
had significant decreases in percent germination (41% for foxtail

millet, 18% for tomato. Germination of all other species was

46

Table 3: Inhibition of root and shoot length of indicator species in

response to increasing concentrations of a purified

asparagus Y‘OOC EXCT‘BC‘C.

 

 

 

 

 

 

 

 

 

 

I50 (ppm)a

Species Root Length Shoot Length Germinationb

(mm) (mm)
Lactuca sativum 28 37 nsc
Raphanus sativus 36 105 ns
Amaranthus retroflexus 68 80 ns
Lepidium sativum 37 90 ns
Portulaca oleraceae 28 30 ns
Lycopersicon esculentum 120 97 *
Setaria italica 130 125 *
Digitaria sanguinalis 35 55 ns
0. ischaemum 43 68 92 **
Echinochloa crusgalli ns ns ns

 

 

also indicates concentration of asparagus root extract where 50%

inhibition of root or shoot length occurs.

cns=nonsignificant

*significant at P=0.05. no 150 can be calculated at this level.

**significant at P=0.01

47

unaffected except for smooth crabgrass. Smooth crabgrass germination
was completely inhibited at the highest concentrations while root and
shoot length were inhibited by 43 ppm and 68 ppm respectively. Shoot
length 150 levels were always greater than 150 levels for root length
in all species except for tomato. However, tomato root length was only
7.5 mm in controls.

Both root and fern tissues of asparagus contain compounds
potentially inhibitory to a wide range of seedling species. Compounds
important in the inhibitory activity of root tissue can be isolated and
concentrated by solvent extraction and column chromatography from water
extracts of asparagus root tissues. No attempt was made to determine
if such isolation was possible from the fern tissues. Since inhibitory
activity is not rapidly decreased when root and fern tissues are
subjected to microbial breakdown, compounds released into the soil as
plants naturally senesce during their normal life cycle could remain in
the soil solution until leached out by rain or irrigation. The data
suggest that severe inhibition of seedling growth can occur in a
relatively short period of time under nonsterile soil conditions, and
that microorganisms are not required to produce phytotoxic activity.
Also, seedling species and tissues vary in their response to asparagus
toxins suggesting several chemicals, mechanisms of action or mechanisms

of defense against the toxins released by asparagus tissues.

LITERATURE CITED

Barnes, J. B. and A. R. Putnam. 1986. Evidence for al lelopathy by

residues and aqueous extracts of Rye (Secale cereale). Weed

 

Science. 34:384-390.
Bell, A. A., 1974. Biochemical basis of resistance of plants to

pathogens. in: Biological Control 91 Plant Insects and Diseases.

 

F.G. Maxwell and FZS. Harris, eds. Pp. 403-461. Univ. Press of
Mississippi, Jackson.

Borner,lL 1950.The roleirftoxic substances Hithe interaction
of higher plants. Bot. Rev. 16:51-65.

Hartung, AJL and C5L.Stephens. 1984. Effects of allelopathic
substances produced by asparagus on the incidence and severity of
Fusarium crown rot. J. Chem. Ecol. 1163-1174.

Hartung, A. C. and A. R. Putnam. 1985. Extracts of asparagus root
tissue are phytotoxic. Proceedings of the Sixth International
Asparagus Symposium. E.C. Lougheed, and H. Tiessen, eds.
Univeristy of Guelph. Pp. 258-266.

Komada, H. 1975. Development of a selective medium for

quantitative isolation of Fusarium oxysporum from natural soils.

 

Rev. Plant. Plot Res. 8:114-125.
Laufer, G. A., and S. A. Garrison. 1977. The effect of asparagus
tissue on seed germination and asparagus seedling growth. Possible

allelopathic interaction. HortSci. 12:385 (AbstrJ

48

10.

11.

12.

13.
14.

15.

16.

17.

18.

19.

49

Linderman, R. G. 1970. Plant residue decomposition products and
their effects on host roots and fungi pathogenic to roots.
Phytopathol. 60:19-22.

McCalla, T. M., and F. A. Haskins. 1964. Phytotoxic substances
from soil microorganisms and residues. Bacteriol. Rev. 28:181-207.
Molisch, H. 1937. Der Einfluss einer pflanze auf die audere
allelopathie. Jena:Fisher.

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

Rice, E. L. 1984. Allelopathy. Academic Press. N.Y. 422 pp.

 

Rice, E. L. 1979. Allelopathy-An Update. Bot. Rev. 45:15-109.
Shafer, W. E., and S. A. Garrison. 1980a. Effects of decomposing
asparagus root tissues on lettuce, tomato, and asparagus seed
emergence. HortSci. 15:406.(AbstrJ

Shafer, W. E.,.and S. A. Garrison. 1980b. Effects of asparagus
root extracts on lettuce and asparagus seed germination and
growth. HortSci. 15:406-407. (Abstr.)

Shafer, W. E., and S. A. Garrison. 1986. Al lelopathic effects of
soil incorporated asparagus root on lettuce, tomato, and asparagus
seedling emergence.FmrtSci.21(1h82-84.

Tiuite, (L, 1969. Plant Pathological Methods. Burgess Publishing
Co. Minneapolis, Minnesota.

Tukey, H. 8., Jr. 1969. Implications of allelopathy in
agricultural plant science. Bot. Rev. 35:1-16.

Weston, L. A. and A. R. Putnam. 1986. Inhibition of legume
seedling growth by residues and extracts of Quackgrass (Agropyron

repens). Weed Science. 34:366-372.

50

20. Yang, Hsu-Jen. 1982. Autotoxicity of Asparagus officinal is L. J.
Amer. Soc. Hort. Sci. 107:860-862.
21. Young, C. C. 1984. Autointoxication in root exudates of Asparagus

officinalis L. Plant and Soil. 82:247-253.

 

22. Young, C. C. 1986. Autointoxication of Asparagus officinalis L.

in, The Science Q Allelopathy. A. R. Putnam and Chung-Shin Tang,

 

 

eds., John Wiley and Sons, Inc. pp. 317.

CHAPTER III

CHARACTERIZATION OF INHIBITORY ACTIVITY OF ASPARAGUS
ROOT EXTRACTS ON ASPARAGUS SEEDLINGS

ABSTRACT

Asparagus decline, frequently attributed to root rot of asparagus
(Asparagus officinalis 1.) caused by pathogenic Fusarium spp., has been
correlated with the release of toxic chemicals from senescing root
tissue. Greenhouse studies showed root rot was increased when
asparagus seedlings were grown in the presence of increasing amounts of
dried root tissue incorporated into soil with either no pathogen or in
combination with f. oxysporum f. sp. asparagi or f} moniliforme. When
excised asparagus roots were treated with increasing concentrations of
a water extract of dried asparagus root tissues, electrolyte efflux was
increased, peroxidase activity decreased linearly and respiration was
decreased. Active components in the extracts were heat stable. Data
suggest allelochemicals of asparagus have a direct effect on the
asparagus plant and are not necessarily produced by microbial

intervention.

51

INTRODUCTION

Recent investigation of the causal factors of asparagus (Asparagus

officinalis L.) decline has suggested that allelopathic compounds are

 

present in asparagus root tissue (3, 4, 6, 10, 11, 12, 15, 16, 17).
Several studies have indicated that allelochemicals are released from
both the growing asparagus plant and from the senescing root tissue (6.
10, 12,15, 16,17). Toxic activity has also been shown to persist in
soil and affect growth of many species including asparagus (11, 12).
Also, evidence indicates that these compounds increase the incidence of
asparagus root rot caused by Fusarium oxysporum (Schlecht.) Snyd. and
Hans. f. sp. asparagi Cohen, and f} moniliforme Sheldon Snyd. and Hans.
(3). These previous studies did not attempt to quantify the response
of the asparagus plant to the level of root tissue in the soil, nor to
elucidate the mode of action toxic compounds in the root tissue may
have on asparagus. This study was directed at 1) quantifying the
response of asparagus seedlings to root tissue incorporated in the soil
with or without presence of the Fusarium pathogens, 2) developing
meaningful bioassays to determine the physiological and biochemical
responses of asparagus to autotoxic components from the root tissues
and 3) evaluate inhibitory activity and its stability in different aged

root tissues.

52

MATERIALS AND METHODS

Preparation of Asparagus Plant Material. A commercial asparagus field
(20 yr old) in Oceana County, MI was excavated and asparagus crowns
(cv. Martha Washington) were collected and washed. The storage and
fibrous roots were separated from the rhizomes and all dead and
visably diseased plant material was discarded. The roots were oven-
dried at 50 C and ground in a Wiley Mill (mesh screen size=1 mm). All
plant tissues were sterilized with propylene oxide gas (14).
Containers with sterilized root tissue were allowed to exhaust under a
fume hood for 24 hr to dissipate any residual propylene oxide. Dried
sterilized tissue was stored in brown glass bottles at -20 C until used
for chemical isolation or bioassays. The tissue was checked for
microbial contamination by plating 1 g of sterilized tissue from each
container on Komada's Medium (5), a selective medium specific for
Fusarium spp. or potato dextrose agar (PDA) (Difco Laboratories,
Detroit, MI.)f0r otherinicrobial species. 'There were three replica
plates for each container and plates were examined each day for a

period of ten days for the presence of microbial colonies.

Preparation of Fusarium Inoculum. Millet seeds (Setaria italica (L.)

(250 g) and distilled water (DHZO) (100 ml) were placed in a 1 liter

flask and tinsdillated (1 hr each time), then inoculated after cooling

with a 4 mm diameter plug of actively growing mycelium of _F_.

53

54

moniliforme (FM) or f. oxysporum f. sp. sparagi (FDA). The seed

 

 

inoculum was incubated at 26 C and each flask was shaken daily to
facilitate mycelial spread throughout thelnillet seeds. After two
weeks of incubation, the inoculum was air dried, then stored in paper

bags at 26 C until used in soil amendment studies.

Inhibition of Asparagus Seedling Growth by Root Tissue. Previous
research indicated when asparagus is planted in soil amended with
sterilized dried root or crown tissues in the presence or absence of
FDA or FM, there is a significant increase in the severity of Fusarium
root rot and less growth of the asparagus plant (3). Since root tissue
was shown to be the most toxic, a 4 x 3 factorial experiment was
performed using 0, 5, 10, and 20 9 root tissue with either no pathogen
or in combination with 134 or FOA. Dried root tissue was incorporated
into steamed sand (Psammentic Hapludalf, sandy, mixed mesic) with or
without the mil let seed inoculum (8 g/pot), poured into 4 inch clay
pots and three month old hybrid asparagus seedlings (cv. "DC 147”)
planted in the amended mix. Sterilized, uninoculated millet seed was
added in the control treatment. There were 6 plants per treatment.
The inoculated seedlings were placed in a completely randomized design
on a greenhouse bench and watered daily. The plants were fertilized
twice with soluble fertilizer (20:20:20) during the duration of the
experiment. After 8 wks, the plants were harvested and evaluated
visuallyrfOr root rot using a scale of 1-5, where 1=no root rot, and
5=death of the plant. All ratings were assigned without knowledge of
the specified treatment. The data were subjected to analysis of

variance and the means separated by the LSD test.

55

Biochemical Assays on Asparagus Seedlings. Because greenhouse
experiments suggest that the root tissue may be damaging the asparagus
plant directly, three bioassay methods were used to further clarify the
type of damage which might be incurred on the asparagus plant.
Bioassays were developed using water extracts of root tissue to
evaluate the effect of toxic root tissue on electrolyte efflux,
respiration and peroxidase activity. These methods were chosen to
evaluate if asparagus tissue damaged through cellular disruption or
plant energy processes. Preliminary experiments for each assay were
performed to determine optimum concentrations of root extract necessary
for each test. In electrolyte efflux studies, asparagus storage and
fibrous root tissue from 1 month-old asparagus seedlings (cv. UC 147),
was cut into 2 mm sections, weighed into 200 mg fresh weight samples
and placed into 201nl vials. Water extract of dried asparagus root
tissue (2.0 and 5.0 mg/ml) was added to the vial. The root extract had
been previously filter sterilized (Nalgene 0.2 m disposable
filterware) and a 061001284 DHzO control was also included in the
experiment. The vials were vacuum infiltrated by reducing the air
pressure to about 2 cm Hg for 15 min, then the extracts were decanted
from the tissue, and the samples were rinsed three times in deionized
DHZO. Deionized DH20 (10 ml) was then added to the vials and samples
placed on a reciprocal shaker (100 strokes/min). Conductance of the
ambient solutions were determined at intervals with51conductivity
meter equipped with a pipet-type electrode assembly. The conductance
value for root tissue in the deionized DHzo controls was used as a
correction factor to calculate root extract induced electrolyte efflux.

There were 3 replications per treatment and the experiment was repeated

56
2 times. Data presented are the results from one representative
experiment.

In respiration studies, fresh asparagus root tissue was removed
from one year-old crowns (cv. Viking) and cut into 1 cm segments,
separated into 1 9 samples, and immersed in a 300 ug/ml solution of
either a sterilized water extract of asparagus root tissue or DHzO-
The tissue was infiltrated for 15 min under vacuum pressure as above.
Entire roots were also cut from the asparagus rhizome, separated into 1
9 samples, and either submerged for 1 hr in asparagus root extract
(300 pg/ml ) or DH20 in 20 ml vials or left intact in the vials and not
submerged. All solutions were decanted from the tissues. Oxygen uptake
was monitored using a Gilson Respirometer (20 C) at 20 min intervals
over a period of 5 hr. There were 3 replications for each treatment,
and the experiment was repeated 2 times. Data presented are results
from one representative experiment. The data were subjected to an
Analysis of Variance and the means separated by Duncans's Multiple
Range Test.

One month-old seedlings (cv. DC 147) were infiltrated with 0, 10,
100 or 1000 ug/ml solution of asparagus root extract and monitored over
time for peroxidase activity. Peroxidase was assayed by the method of
Ridge and Osborne (8) and expressed as the change in Aug/"11ml 9 dry
weight of asparagus root tissue. There were 3 replications for each
treatment. The data were subjected to an Analysis of Variance and the

means separated by the LS0 test.

Comparison of Inhibitory Activity from Different Aged Asparagus Plants.
Asparagus crowns (cv. Martha Washington) were excavated from

plantations 5, 12, and 20 yrs old and prepared as previously described.

57

Dried root tissue (25 g) from each field was extracted with 500 m1 0H20
overnight at 41L The particulates were removed by straining each
sample through 3 layers of cheesecloth and centrifuging for 20 min at

6,000 g. The samples were then decanted from the pellet and

lyophilized to determine the dry weight, resolvated in DH20 at known
concentrations (0.0, 0.5, 1.0, 2.0 mg/ml) and applied to petri dishes
(60 x 15 cm). Previous studies in this laboratory have shown that

Lepidium sativum L. (cv. Curly Cress) is a fast and reliable seedling

 

for evaluating the inhibitory activity of allelopathic compounds (7L
Therefore, curly cress was used to test inhibition from root extracts
of asparagus tissues of increasing ages. Curly cress seeds (10/dish)
were added to the dishes along with 1.5 ml DHzo, then incubated for 72
hr at 26 C in a growth chamber. There were 3 replications per
treatment and the treatments were arranged in a randomized complete

block design. Root and shoot lengths were measured and 150 levels

determined by interpolation. The data were subjected to an Analysis of
Variance and the means separated by the LS0 test.

Part of each sample was used to determine if the compounds present
in the extract were heat stabile. Each sample (50 ml) was autoclaved
20 min at 120 C at 23 psi then applied to petri plates (60 x 15 cm) at
known concentrations (0.0, 0.5, 1.0, 2.0 mg/ml) and bioassayed using
curly cress as above. Unautoclaved samples at equal concentrations and

controls were also bioassayed as indicated above.

RESULTS

Preparation of Asparagus Plant Material. In allelopathy studies, it
is important to separate the contribution of chemical compounds
produced by plant tissues as they are degraded by resident microbes and
compounds released directly from plant tissues. Therefore it was
important to sterilize plant tissues used in all experiments to remove
the confounding factor of microbes. After asparagus root and fern
tissues were treated with propylene oxide gas, then plated out on
either Komada's medium or PDA, no Fusarial or other microbial colonies

were detected.

Inhibition of Asparagus Seedling Growth by Root Tissues. When root
tissue was incorporated into the soil, increasing levels of tissue
resulted in significantly increased levels of root rot on asparagus
seedlings (Figure 1). When either of the Fusarium pathogens were
incorporated in combination with the root tissue, there was a
significant increase in the level of root rot on the asparagus plant.
Analysis of variance of these data indicated significant main effects

from the root tissue and the pathogens on the asparagus plant.

Physiological and Biochemical Assays on Asparagus Seedlings.
Electrolyte efflux increased at the very highest (5 mg/ml) crude

extract levels (P=0.01) as well as the 2mg/ml level P=0.05 (Figure 2).

58

Figure l:

59

Root rot ratings of asparagus seedlings (cv. UC 147) grown
in soil amended with increasing levels of dried asparagus
root tissue alone or in combination with [Loxysporum f.
sp. asparagi or _F_._moniliforme. Control was soil amended
with sterilized millet seed only' or increasing

concentrations of dried asparagus root tissue.

 

60

F. oxysporum

 

- F. moniliforme
:3 Control

.05%

LSD O

ended

 

 

 

 

 

 

 

Asparagus Root Tissue (g)/5009 Soil

61

Figure 2: Electrolyte efflux over time from excised asparagus root
tissue treated with increasing concentrations of a water

extract of asparagus root tissue.

62

Change in Conductivity (,umhos)

 

-4 N do h 01
O O O O O C
d H
511101 I \ 5
water a U
. II
> P
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3 '° 3
q
-l
q
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Fl- 9 "
CD
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63

Electrolyte efflux was also tested at concentrations similar to those
used in other biochemical assays (data not shown) but no significant
increase in leakage was observed. No attempt was made to determine ion
influx or exchange.

Peroxidase activity was decreased over timerwhen the seedlings
were subjected to increasing concentrations of the crude root extract
(Figure 3).

In respiration experiments, the control tissues ( not submerged in
DH20) exhibited higher rates of 02 Uptake than other treatments
throughout the experiment (Table 1). In tissue subjected to the
aqueous root extracts, respiration was significantly less than in the
DHzO treatments. Infiltration alone significantly reduced respiration
as compared to the control tissue. However, infiltration with the
aqueous root extracts reduced respiration considerably more than
infiltration with DH20. Not suprisingly, submerging the root tissue

in distilled water also reduced its respiration, but not differently

than when the tissue was infiltrated with DH20-

Comparison of Inhibitory Activity From Different Aged Asparagus Plants.
Both root and shoot lengths of cress seed were inhibited by increasing

levels of asparagus root extracts, but root length was consistantly the

more sensitive parameter'( average 150 for root length=1083 ug/ml.
average 150 for shoot length=1644 ug/ml) (Figure 4L. Both root and
shoot bioassays indicated that the tissue from the 5 yr-old plants
contained the most inhibitory activity, whereas the 12 yr-old and 20
yr-old were less active but not significantly different from each other
(LSD=3.60 at 0.01 level for both root and shoot length). Root length

was inhibited the most by the 5 yr extracts over all dilutions but was

Figure 3:

64

Peroxidase activity of asparagus seedlings infiltrated with
increasing concentrations of a water extract of asparagus

root tissue.

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66

Table 1; 02 uptake after infiltration of asparagus root tissues with
distilled water or asparagus aqueous extracts (300 ug/ml), or
from submerging excised asparagus roots in aqueous extracts

or distilled water after five hr.

 

 

Treatment 02 Uptake (ul/g-hr)z
Infiltrated with root extract 46.2 a
Infiltrated with distilled water 64.6 b
Submerged in root extract 39.4 a
Submerged in distilled water 61.1 b
Not submerged 112.0 c

 

2Means with uncommon letters differ at P= 0.05 by Duncan's Multiple

Range Test.

67

Figure 4: Root and shoot length of curly cress seedlings treated with
increasing concentrations of a water extract of asparagus

root tissue from 5, 12, and 20 yr old plantations. A=Root

Length, B=Shoot length. Dotted line=150 IEVEIS-

20

 

 

5 yr.
—'-°' 1 2 yr.
——' 20 yr.

o.o1‘

.5
C

 

Root Length (mm)

 

 

 

 

16

.2
O

 

 

Shoot Length (mm)

 

 

l 31 : 2 l
500 1000 ' 2000

Asparagus Root Extract (149/ ml Dry Weight)

110

hi

6C

69

not significantly different from the 12 and 20 yr-old treatments at the
highest concentration. Measurements of shoot length showed that
activity from the 12 and 20 yr extracts was not significantly different
from each other (150=1950 ug/ml in 12 yr and 2000 ug/ml in 20 yr
treatments). The 5 yr-old tissue extracts had the most inhibitory
activity on shoot growth (150=983 WQIMI).

Autoclaving extracts from the 5 and the 12 yr-old tissue tissue
had no significant effect on inhibitory activity (Figure 5).
Autoclaved extracts from the 20 yr-old tissue showed less activity than
the non-autoclaved extracts at the intermediate level but did not

differ at the highest or lowest levels.

DISCUSSION

Root rot on asparagus is significantly increased when asparagus
seedlings are grown in the presence of increasing levels of dried
asparagus root tissue with or without FDA or FM. Dried fern tissue was
also tested in a similar fashion in prior experiments did not provide
the same increase in disease as was noted with the root tissue (3).
These data suggest that a chemical or physical property of the root
tissue influences the incidence of Fusarium root rot on the asparagus
seedlings. The data also suggests that this is a direct effect of the
residues on the asparagus plants and not an interaction of the residues
and the pathogens. In experiments with sterilized and nonsterilized
vermiculite, Young (18) found that asparagus seedlings planted in
unsterilized "used“ vermiculite and treatments with root tissues alone

started to "yellow and wilt" after 21 days but were normal in controls

70

Fimne 5: Root length of curly cress treated with huncasing
concentrations of autoclaved and nonautoclaved water
extract of asparagus root tissues from 5, 12, or 20 yr old

plantings.

easing
water

yr cl:

71

2O

 
    
 

LSD 10.051001

 

E

5 \

5 ' Not autoclaved
m

5 10 - —— Autoclaved

.1

“ I-

O

O .

m

 

 

Root Length (mm)

 

 

 

500 1 000 2000

Root Length (mm)

 

 

l l

500 1 000 2000

20 Yr.

Asparagus Root Extract (AQI ml Dry Wt)

72

and treatments with sterilized vermiculite and root tissues. He
postulated that there is an interaction between the residues and
microorganisms. However, the exact nature of the "used vermiculite" or
its microbial content was not published, making it difficult to assess
the results. Our data support the hypothesis that root rot on the
crown is increased due to damage to the root system that could be
caused by toxic components released from senescing root tissue rather
than from microbial intervention.

Although asparagus has been reported to release autotoxic
compounds from the root tissues as well as from intact root systems, no
data has been presented concerning the nature of the autoxicity or the
site of.action. 'Three types of assays were used to investigate what
processes could be affectedtn/root extracts; electrolyte efflux to
test for tissue perterbation (1), peroxidase activity, as an increase
in peroxidase as been associated with increased disease resistance (2),
and respiratory activity of young asparagus root tissue. In
electrolyte efflux studies, efflux occurred only at much higher extract
concentrations than*where responses occurred in the other bioassays.
These results suggest that compounds important in tissue perturbations
are at low concentrations, that asparagus tissues are not sensitive to
the compounds in the crude extractcn~other’compounds'hithe complex
mixture of the crude extract could be confounding the experiment.
Peroxidase activity was significantly decreased after 18 hr. Since an
increase in peroxidase is associated with disease resistance, these
data may suggest that the plant's defense mechanisms to combat
infection from its pathogens are reduced in the presence of root

tissue, making the plant more susceptible to disease. Respiration was

73

also significantly reduced in one year-old root tissues indicating that
compounds present in the extracts are damaging the energy-producing
processes of young roots. These data all support the hypothesis that
chemicals present in the water extracts are altering the plant's
biochemical processes.

In experiments to test different aged tissues, it may not be
surprising that tissue from 12 and 20 year-old plants did not vary in
toxicity since the storage root tissues of the asparagus crown is
replaced approximately every 6 years (9, 13). Although plants were
removed from fields of increasing age, tissue present in these fields
may vary in age (9, 13). Although tissue collected from younger
plantings was more toxic than tissue collected from older plantings, it
is difficult to deduce from these experiments if tissue age was the
determining factor. Chemicals may be detoxified or leached from the
tissue as tissues age, but other expermentation is needed to assess the
true effects of tissue age on tissue toxicity. Compounds present in
tissues are heat stable, suggesting that the components may be
difficult to destroy in a natural agroecosystem. Shafer and Garrison
presented data indicating that toxicity is present in the soil for up
to 90 days (12). With their data, it is difficult to determine if this
loss of toxicity is due to microbial breakdown or leaching of the
compounds from the rhizosphere. Data collected in this laboratory
suggests that the compounds important in the autotoxic responses have a
pronounced polar nature and therefore may be $01 vated from the soil
system by leaching (3, 4). We have reported previously that root
tissue did not affect the growth of FDA or FM but will affect other
microorganisms (3). Therefore replant problems in old asparagus

plantations may be attributed to the continual presence of Fusarium

74

pathogens once the crowns have been destroyed rather than long term

persistence of toxins in the soil.

LITERATURE CITED

Bronson,(L R.andIL P.Scheffer. 1977.Heat- and aging-induced
tolerance of sorghum and oat tissues to host-selective toxins.
Phytopath. 67:1232-1238.

Hammerschmidt,R.,E.lL Nuckles,and J.Kuc. 1982. Association
of enhanced peroxidase activity with induced systemic resistance
of cucumber to Colletotrichum langenarium. Physiol. Plant Phys.
20:73-82.

Hartung, A. C., and C. T. Stephens. 1983. Effects of
alle10pathic substances produced by asparagus on the incidence and
severity of Fusarium crown rot. J. Chem. Ecol. 9:1163-1174.
Hartung, A. C. and A. R. Putnam. 1985. Extracts of asparagus
root tissue are phytotoxic. Proceedings of the Sixth International
Asparagus Symposium. E.(L Lougheed, and H. Tiessen, eds.
University of Guelph. p. 258-266.

Komada, H. 1975. Development of a selective medium for
quantitative isolation of Fusarium oxysporum from natural soils.
Rev. Plant. Plot Res. 8:114-125.

Laufer, G. A., and S. A. Garrison. 1977. The effect of asparagus
tissue on seed germination and asparagus seedling growth.

Possible allelopathic interaction. HortSci. 12:385 (Abstr.)

75

7.

10.

11.

12.

13.

14.

15.

76

Lehle, F. R. and A. R. Putnam. 1982. Quantification of
allelopathic potential of sorghum residues by novel indexing of
Richards'Function fitted to cumulative cress seed germination
curves. Plant Physiol. 69:1212-1216.

Ridge, A. and S. Osborne. 1970. Hydroxyproline and peroxidase

activity in cell walls of Pisum sativum: Regulation by ethylene.

 

J. Exp. Bot. 21:843-856.

Robb, A. R. 1984. Physiology of asparagus (Asparagus

 

officinalis) as related to the production of the crop. New

 

Zealand J. Exp. Agri. 12:251-260.

Shafer, W. E. and S. A. Garrison. 1980. Effects of decomposing
asparagus root tissues on lettuce, tomato, and asparagus seed
emergence. HortSci. 15:406. (Abst.)

Shafer, W. E. and S. A. Garrison. 1980. Effects of asparagus
root extracts on lettuce and asparagus seed germination and
growth. HortSci. 15:406-407.(AbstrJ

Shafer, W. E. and S. A. Garrison. 1986. Allelopathic effects of
soil incorporated asparagus roots (H1 lettuce, tomato, and
asparagus seedling emergence. HortSci. 21(1):82-84.A

Tiedjens, V. A. 1926. Some observations on root and crown bud
formation in Asparagus officinalis. Proc. Am. Soc. Hort. Sci.
23:189-195.

Tuite, J. 1969. Plant Pathological Methods. Burgess Publishing Co.
Minneapolis, Minnesota.

Yang, Hsu-Jen. 1982. Autotoxicity of Asparagus officinalis L.

J. Amer. Soc. Hort. Sci. 107:860-862.

77

16. Yang, Hsu-Jen. 1985. Autotoxic and alle10pathic characteristics of

Asparagus officinalis L. Proceedings of the Sixth International

 

Asparagus Symposium. E. C. Lougheed and H. Tiessen., eds.
University of Guelph. p.267-276.
17. Young, C. C. 1984. Autotoxication in root exudates of Asparagus

officinalis L. Plant and Soil. 82:247-253.

 

18. Young. C. C. 1986. Autotoxication of Asparagus officinal is L. In:

The Science pf Al lelopathy. A.R. Putnam and Chung-Shin Tang ed.

 

 

John Wiley and Sons, Inc. 317 pp.

Chapter IV
IMPACT OF ASPARAGUS AND ASPARAGUS PLANT TISSUES ON
SOIL MICROBE POPULATIONS

ABSTRACT

The responses of microbial populations in soils with dried
asparagus root or fern tissue or soils planted with asparagus

(Asparagus officinalis L. cv DC 147), TL. sprengeri, snap beans

 

 

(Phaseolus vulgaris cv. Bush Blue Lake) or sweet corn (Zea mays cv.

 

Calico King) were evaluated. Epggpigm spp. and other fungal
populations increased in treatments where asparagus root tissue was
incorporated into soil. In contrast, bacterial species quickly
colonized fern tissue. Both bacterial and fungal populations were
significantly lower in treatments where asparagus plants were grown but
not.in treatments where snap beans or sweet corn were planted. Nine
isolates of Pythium spp- and 6 of 18 bacterial isolates were inhibited

by purified components from asparagus root tissue.

INTRODUCTION

Plant residues from various sources contribute to the organic
matter component of soils. These tissues from young, mature, senescing

and dead plant parts are ultimately decomposed in the soil by the

78

79

resident microbial populations<n~abiotic factors. Thus, chemical
components bound in the plant materials are made available for
microbial and plant uptake» At any one time, the soil may contain a
vast array of chemical components which may affect plant growth in
either beneficial or detrimental ways.

Asparagus (Asparagus officinalis 1.), an important vegetable crop

 

in Michigan, is often plagued with two major problems; a) crown death
from a root rotting disease caused by f, oxysporum f; sp. asparagi

(FDA), and E_._moniliforme (FM), (1, 2) and b) decline and unprofitable

 

yields from replanted fields. (4, 5, 23). The two problems have been
collectively termed "Asparagus Decline". Recent research suggests
that, although decline is ultimately caused by pathogenic Fusarium
species, asparagus also contains compounds that are autotoxic or
autoallelopathic, and these compounds may be attributing to the
severity of the decline problem (6, 7, 9, 20, 21, 22, 25, 26, 27).
Allelopathy is defined as a direct or indirect harmful affect by one
plant (including microorganisms) on another through the production of
chemical compounds released into the environment (18).

Decomposition of plant products in the soil associated with the
formation of phytotoxic substances has been documented by numerous
reports (3, 11, 12, 14, 15, 23, 24). One mechanism by which
allelopathic compounds may indirectly affect plant growth is by
affecting the microbial populations in the soil leading to an
ecological imbalance of the naturally occuring microflora. This
imbalance can result in an increased level of disease on crop plants.
Tousson and Patrick (23) reported that the incidence of root rot of
bean (Phaseolus vulgaris) caused by F, _s_pl_a_ni_ (Mart.) was increased if

the roots were exposed to extracts from decomposing residues of rye,

80

barley, broccoli, and broad bean. Patrick and Koch (16) reported that
exposure of tobacco to leachates from decomposing rye and timothy
residues increased the susceptibility of both resistant and susceptible

varieties of tobacco to black root rot caused by Thielaviopsis basicola

 

(Beck. and Br.) Ferraris.

Preliminary observations have indicated that microbial populations
in asparagus fields are reduced when compared to microbial populations
in similar soils planted with other crops (Putnam et. al., unpublished
data). Also, we have reported previously that extracts of asparagus
root tissues are inhibitory to Pythium spp. but not to either FOA or FM
(6L. Based on these data, it was hypothesized that toxic substances
released from senescing asparagus tissues may be harmful to some
organisms but not to the pathogenic species of Fusarium, thus creating
a more favorable environment for pathogenesis to occur. The purpose of
this research was a) to determine how asparagus root or fern residues
incorporated in the soil would affect natural microbial populations, b)
to determine if the presence of asparagus plants as compared to other
plants (amt snap beans or corn) could affect populations and diversity
of microbial species in the soil and c) to directly test phytotoxic
extracts of asparagus root tissue for inhibitory activity on isolated

bacterial or fungal Species.

MATERIALS AND METHODS

Impacts of various plant tissues on naturally occuring microbial
populations. Asparagus root and fern tissues collected, ground, and

sterilized as previously reported (6) were incorporated (509/180 9

81

soil) into Spinks sandy loam soil (Psammentic Hapludalf, sandy, mixed,
mesic, with 1% organic matter, pH 6.5) and the soil placed in 15.2 cm
clay pots. In a separate treatment, Shredded poplar excelsior (50 g)
was also incorporated into soil as a control for the physical influence
of organic matter in the soil. Another treatment used a living
asparagus crown ( DC 157, 1 yr old) planted a similar pot. There were 2
plants per treatment and 3 replications of each treatment. The pots
were placed on a greenhouse bench and subirrigated as necessary to
maintain equal moisture levels in all treatments. Unwanted plant
growth was suppressed by hand weeding. Plants were fertilized once a
week with soluble fertilizer (20:20:20) during the duration of the
experiment. Microbial p0pulations were monitored at 1, 4, 8, 12, 18,
and 22 weeks.

In a second experiment, treatments were the same except snap beans

(Phaseolus vulgaris L. cv. Bush Blue Lake) and sweet corn (Zgg.mays cv.

 

Calico King) were included as plant controls and the poplar excelsior
treatment was omitted. Samples were taken at 2, 5, 9, 18, and 22
weeks.

To monitor changes in the microbial populations in the soil over
the duration of the experiment, the soil was first analysed for total
microbial population before the experiment was initiated. The baseline
populations of microbial organisms were determined from three random

samples from the soil. Data for total bacterial (sensu stricto),

 

fungi, actinomycetes, and Fusarium spp. populations were all monitored.
A dilution series was made in normal saline and samples plated on
selective media. To evaluate changes microbial populations over time,
soil samples were taken from each pot by removing the top 2 cm of soil,

then taking 4 samples from each pot with an 8 mm diameter cork borer.

82

The 4 cork borer samples were mixed well, allowed to air dry, then
sieved through a 40 mesh Sieve (425 u) to remove any plant material,
and weighed into 1 9 samples. Each sample was mixed with 9 ml of
Sterile normal saline U185% aqueous NaCl) and vortex-mixed for 30
sec. The particulates were allowed to settle and serial 10-fold
dilutions were made. Aliquots of 0.11nl from each dilution were plated
on media selective for bacteria, fungi, actinomycetes, and Fusarium
spp. The Petri plates were incubated at 25 C for 3 days for enumeration
of bacteria, 4 days for fungi, 7 days for actinomycetes, and 10 days
for Fusarium, then discrete colonies were counted. There were 2
dilutions per sample and 2 replica plates for each dilution. Microbial
counts from the 2 replica plates were averaged to obtain the number of
colony forming units (CFU) per gram of soil. Data were subjected to an
Analysis of Variance for each time period and the means separated by

LSD test.

Selective media used for evaluation of microbial species in greenhouse
experiments. The medium used for bacteria was nutrient agar (Difco
Laboratories, Detroit, MI); 2 ml nystatin, (15 mg/ml in DMSO), and 28
ml cycloheximide, (15 mg/ml in Distilled H20 (Dugon, Solutions were
added to one liter of medium after autoclaving. Fungal p0pulations
were evaluated on Czapeks' agar (30 9 sucrose, 2 g sodium nitrate, 1 g
dipotassium phosphate, 0.5 9 magnesium sulphate, 0.5 g potassium
chloride, and 0.01 g ferrous sulfate in 1 L DHzo amended with rose
bengal (35 mg/L to restrict colonies) and chloramphenicol (100 mg/L to
suppress bacteria). Populations of actinomycetes were evaluated on a

selective medium containing 3 g N.Z.-amine-A (Schefield Chemicals,

83
Norwick, NY) , 18 g bacto agar (Difco Laboratories, Detroit, MI), in 1
1- tap H20; 5 ml bromocreosol green (0.04% in DH20) was added to
restrict the colonies after adjusting the pH to 7.2. Antifungal
antibiotics nystatin and cycloheximide were added asceptical ly as
described above. For estimating populations of Fusarium spp. Komada's
medium was used (8). The medium contained potassium monophosphate, 1.0
9. magnesium sulphate + 7 H20, 0.5 g, potassium chloride, 0.5 g, Fe-Na
EDTA, 0.01 g, L-asparagine, 2.0 g, D-galactose, 20 g, sodium borate +
10 H20, 1,0 g, oxgal l , 0.5 g, PCNB (75%)(Terachlor), 1.0 g, Bacto agar,

15 9. added to 1 L 0f 01120 and steamed for 1 hr. The pH was adjusted
between 3.8-4.0 with 10% phosphoric acid. Chloramphenicol, (0.25 g/L),

was added after the medium had cooled.

Preparation of partially purified asparagus root extracts. Since
other laboratories as well as this one have published that water
extracts of root tissue are toxic to seedling germination and radical
elongation, water extracts from root tissue were subjected to solvent
extraction and thin layer chromatography (TLC) to try and isolate
chemical components that may also be inhibitory to microbial organisms
(6, 25). Ground asparagus root tissue was extracted by stirring
overnight at 4 C (1:10 root tissueonzo), The particulates were then
removed by filtering through four layers of cheese cloth and
centrifuging at 6,000 x g for 20 min. The supernatant was decanted
carefully from the pellet (the pellet was discarded) and then
precipitated with acetone (4:1 v:v acetone:sample) overnight at 4 C.
The precipitant was discarded and the liquid concentrated to 1/4 the
original volumn on a Buchi Rotary evaporator. This concentrate was

extracted with chloroform 3 times (1:1 v:v). The chloroform fraction

84

was reduced to dryness, weighed, redissolved in methanol, and filter
sterilized. This extract was further purified on TLC plates (Whatman
Silica gel 60, 200 0 thickness, mobile phase, hexane:ethyl acetate 1:1
v:v). TLC plates were washed twice in this solvent system and allowed
to dry each time before using in purification procedures. After the
chloroform fraction was develOped on the plates, the edge of each plate
was removed and developed using 5% vanillin in concentrated H2304, Rf
values were determined for separate spots and the remainder of the
plate was scraped accordingly. Individual Spots were eluted from the
silica gel with 100 ml of methanol or chloroform. Samples were
filtered, dried under nitrogen, then weighed and kept at -20 C until
used in bioassays. A duplicate plate with no sample was also developed
in the sol vent system, scraped and eluted with methanol for use as a

control.

Effects of water extracts on isolates of Pythium spp. Previously
published data have indicated that asparagus root tissues are
inhibitory to Pythium spp. but not to Rhizoctonia spp. or either of the
Fusarium root rot organisms that attack asparagus plants (7). m
spp. is a well known soil pathogen and different isolates were used to
test for sensitivity to water extracts of asparagus root tissue.
Thirteen isolates of Pythium spp. were tested for sensitivity to water
extracts of dried asparagus root tissue. Eleven isolates of P.55Lpflmgp
Trow, were collected in Ohio, and one isolate each of P. ultimum and P_.

aphanadermatum (Edson) Fitzs., were collected in Michigan. A water

 

extract of dried asparagus root tissue was sterilized with a millipore

filter U122 1m) to remove any bacterial or fungal contamination. The

85

extract was applied to potato dextrose agar (PDA) (Difco Laboratories,
Detroit, MI) plates (9 cm diameter) ULZ ml) and allowed to dry. Each
plate was then inoculated with a 2 mm agar plug of an actively growing
Pythium isolate. Cultures were allowed to grow for 4 days at 25 C then

visually assessed for hyphal inhibition or stimulation. There were 2

replica plates/isolate and DHzO was added to the control plates.

Sensitivity of bacterial isolates to purified extracts of asparagus
root tissue. Natural'Ly occuring soil bacteria were isolated from
Spinks sandy loam soil using Martin's selective medium for bacteria
(13). The medium contained (L3% trypticase soybroth (BBL Microbiology
Systems, Cockeysville, MD), 50 ppm PCNB, and 15 g bacto agar (Difco
Laboratories, Detroit, MI) in 1 L DHzo, autoclaved for 20 min then
poured into 9 cm petri dishes. Individual colonies were removed from
the selective medium and streaked on an amended nutrient agar. Single
colonies were subcultured on agar slants and Stored at 4 C until used
in bacteria sensitivity tests. The amended nutrient agar medium
contained 1.4 g nutrient broth (Difco Laboratories, Detroit, MI) 5 9
glucose, 5 g yeast extract (Difco Laboratories, Detroit, MI), and 15 g
PACtO agar in 1 L DH20. All bacterial Studies were done using this
medium. Soft agar overlays made from this medium contained 0.15% bacto
agar.

The bacterial suspension in the soft agar overlay was made by
adding 1 ml of sterile normal saline to a 24 hr-old culture in a slant.
vortexing 30 sec to suspend the bacteria in the saline, then using 100
pl of this suspension to inoculate the soft agar overlay.

Individual bacterial isolates were tested for sensitivity to

components eluted from TLC plates. Dried components from TLC plates

86

were redissolved in methanol at 1.0 and 0.5 mg/ml , then 10 ul of each
dilution spotted on amended nutrient agar along with a methanol and a
silica gel plate control. After the methanol had evaporated from the
plate, a soft agar overlay inoculated with a bacterial isolate was
poured over the surface of the plate. Plates were incubated at 25 C
for 48 hrs then examined for zones of inhibition in the bacterial lawn.
There were 18 isolates of bacteria tested for sensitivity to isolated
compounds from asparagus root tissue. All cultures that were sensitive
to the extracts were identified by the Animal Health Diagnostic

Laboratory, PJL Box 30076, Lansing, MI.

RESULTS

Impacts of various plant tissues on naturally'occuring microbial
populations. In all microbial Species tested, poplar excelsior had
little effect on microbial populations, indicating that the physical
presence of non-nutritive tissues did not affect population levels
(data not Shown). Therefore this treatment was deleted from subsequent
experiments.

There were no differences in bacterial populations between pots
containing a living asparagus plant and the control soil in experiment
one (Table 1). In pots containing root tissue, there were significant
increases in the bacterial populations at 4 and 21 weeks, then
populations returned to levels not Significantly different from the
control soil. However, there were highly significant increases in the
bacterial populations in soils amended with fern tissue. Bacterial

populations were Significantly higher for soil amended with fern tissue

87
Table 1: Estimates of bacterial populations (CFU'S)lTlSplnkS sandy
loam soil amended with 50 g of dried asparagus root or fern
tissue or planted with asparagus, snap beans, or sweet corn.
Initial bacterial population in 1 g of unamended soil was
36.8 x 106 cru in Experiment #1 and 37.4 x 106 CPU in

Experiment #2.

 

x106 CFU/g Soil at the

 

 

EXPERIMENT #1 Time of Evaluation (weeks)
Treatment 1 4 8 12 18 22
Control 47.3 bc* 87.0 c 26.5 c 24.5 b 41.5 b 25.0 c

Root Tissue 97.4 bc 136.8 ab 103.5 b 46.0 b 78.0 b 92.1 b
Fern Tissue 150.0 a 150.0 a 150.0 a 150.0 a 133.2 a 140.0 a
Asparagus 72.2 bc 83.4 c 18.4 c 32.5 b 59.4 b 28.7 c

 

x106 CFU/g Soil at the

 

 

EXPERIMENT #2 Time of Evaluation (weeks)
Treatment 2 5 18 22
Control 21.3 b 30.4 bc 21.8 c 11.9 bc

Root Tissue 150.0 a 143.8 a 98.6 a 54.3 a
Fern Tissue 150.0 a 139.0 a 122.4 a 65.6 a
Asparagus 33.8 b 27.2 bc 26.4 be 24.0 b
A, sprengeri 24.6 b 35.8 b 32.4 bc 14.3 bc
Snap Beans 38.0 b 23.0 bc 28.9 be 23.4 b

Sweet Corn 18.6 b 27.2 bc 67.3 b 19.5 bc

 

*All numbers with a letter in common are not significantly different

according to LSD for columns only.

88

after only 1 week compared to the control soil. Bacterial populations
remained high throughout the experiment and at 22 weeks were still 140
x 106 CFU/g soil, whereas control soil populations were 25 x 106 CFU/g
soil. Similar increases in the bacterial populations in soil amended
with fern tissue were also obtained in the second experiment. AS
before, root tissue amended soil again showed increased bacterial
populations, and the populations began to decline as the experiment
progressed. Soil planted with asparagus, Ag sprengeri, snap beans, and
sweet corn showed no differences in bacterial populations when compared
to the control.

The control soil, when considered alone, showed the highest number
of fungal CFU after 4 weeks (26.4 x 104 CFU/g soil). In subsequent
weeks, population counts were less (Table 2). In pots where an
asparagus plant was present in the soil, fungal populations did not
vary from those counted in the control soil except at 8 weeks in
experiment 1. Fungal populations in treatments where fern tissue was
incorporated in the soil were also not different from the control soil
except at 8 weeks in experiment 1. However, when root tissue was
incorporated in the soil, fungal populations increased Significantly
over controls after only 1 week. Population counts were always higher
in treatments amended with root tissue than in any other treatments.

This increase in fungal (Hflrs for root tissue also occurred in the
second experiment. Fungal populations in soil amended with fern tissue
did not differ from controls in either experiment. Fungal populations
in soil where snap beans were planted Showed a significant increase at
9 weeks as compared to the control soil. Fungal populations in the
snap bean treatment at 9 weeks were similar numerically to those in

soil amended with root tissue. However, at 18 and 22 weeks, after snap

Table 2: Estimates of fungal populations (CFUHH in Spinks sandy loam

soil

amended with asparagus root or fern tissue or planted

89

with asparagus, snap beans, or sweet corn.

population in 1 g of unamended soil was 9.1 x 104 CFU in

Experiment #1, and 6.5 x 104 CFU in Experiment #2.

Initial fungal

 

EXPERIMENT #1

x104 CFU/g $011 at the

Time of Evaluation (weeks)

 

 

 

 

 

Treatment 1 4 8 12 18
Control 8.0 bcd* 26.4 bc 1.4 b 1.0 b 7.8 b
Root Tissue 20.3 a 150.0 a 88.7 a 28.3 a 29.2 a
Fern Tissue 1.3 cd 55.8 b 7.0 b 1.1 b 3.0 b
Asparagus 8.8 bc 19.5 c 38.2 b 0.8 b 14.8 ab
x104 CFU/g 5011 at the
EXPERIMENT #2 Time of Evaluation (weeks)
Treatment 2 5 9 18 22
Control 4.0 b 10.8 b 7.3 c 5.8 b 4.0 b
Root Tissue 83.9 a 101.1 a 79.4 a 46.9 a 32.4 a
Fern Tissue 18.3 b 10.4 b 3.4 c 3.3 b 2.6 b
Asparagus 1.5 b 5.5 b 3.8 c 3.3 b 7.0 b
Snap Beans 2.2 b 5.8 b 67.7 ab 13.8 b 6.5 b
Sweet Corn 1.2 b 9.5 b 31.5 bc 6.7 b 6.8 b

 

*Numbers without a letter in common are Significantly different at

P=0Jn.or greater according to LSD for columns only.

90

beans had begun to senesce, populations were similar to levels Similar
in the control soil. In treatments where asparagus was grown in the
soil, populations did not vary from those of the control.

Population counts for Fusarium spp. were significantly higher for
treatments where root tissue was incorporated into the soil than in any
other treatment (Table 3). This was true for all time periods
analysed. All other treatments varied little from control soil. In
treatments where asparagus was grown in the soil, Fusarium populations
increased significantly from other treatments after 18 weeks, but
populations did not reach the levels seen in the root tissue
treatments. This result was not obtained in the second experiment in
which only the treatment where root tissue was incorporated in the soil
Showed a significant increase in Fusarium populations.

For actinomycetes, the population counts in all treatments varied
considerably and data were not consistent from one experiment to the

next. Therefore, no conclusions could be drawn from them.

Effects of water extracts on isolates of Pythium spp. Six of the E.
gltlmgm isolates and the one 3. aphanidermatum isolate did not grow
within 2 cm of the areas where a water extract of asparagus root tissue
had been applied on PDA (data not Shown). With 3 other isolates,
sparse growth of the Pythium hyphae was observed on the area where the
root extract was applied. Hyphal Strands were appressed on the agar
plate and did not exibit the normal fluffy appearance of the rest of

the culture or the control plates.

91

Table 3: Estimates of Fusarium spp. populations (CFU's) in Spinks
sandy loam soil amended with dried asparagus root or fern
tissue or planted with asparagus, snap beans, or sweet corn.
Initial propagule count per 1 g of soil was 20.3 x 103 CFU in

both experiments.

 

x103 CFU/g Soil at the

 

 

EXPERIMENT #1 Time of Evaluation (weeks)
Treatment 4 8 12 18 22
Control 29.1 b* 4.9 b 5.2 b 2.1 c 3.6 0
Root Tissue 93.3 a 79.2 a 133.4 a 64.3 a 67.5 a
Fern Tissue 4.2 b 6.8 b 3.1 b 4.3 c 6.5 c
Asparagus 4.8 b 11.0 b 6.9 b 26.1 b 23.4 b

 

x103 CPU] 9 Soil at the

 

 

EXPERIMENT #2 Time of Evaluation (weeks)
Treatment 2 5 9 18 22
Control 13.3 b 15.8 b 11.8 b 10.1 b 6.5 b
Root Tissue 126.3 a 82.3 a 71.8 a 85.7 a 79.5 a
Fern Tissue 11.5 b 5.8 b 5.0 b 2.7 b 0.3 b
Asparagus 7.8 b 8.2 b 9.0 b 9.6 b 7.8 b
Snap Beans 7.3 b 12.3 b 29.8 b 17.8 b 4.3 b
Sweet Corn 9.9 b 18.7 b 20.6 b 12.0 b 11.2 b

 

*Numbers with no letters in common are Significantly different at

P=0.01 or greater by LSD in columns only.

92

Sensitivity of bacterial isolates to purified extracts of asparagus
root tissue.(Hi18 cultures tested, 8 were inhibited by isolated
components of asparagus root tissue eluted from TLC plates (Table 4).
Components from rf=0.35 and 0.26 have been shown to also contain
compounds inhibitory to seed germination and radical elongation of

Lepidium sativum cv. curly cress (6). Components at rf=0.82, 0.75,

 

and 0.5 did not affect seed germination or radical elongation, but were
Strongly inhibitive of bacterial growth. Isolates of bacteria
particularly sensitive to asparagus root extracts were identified as

8acillg§_spp., and B, circulans.

DISCUSSION

Impacts of various tissues and plant species on naturally'occuring
microbial populations. In all experiments, the coefficient of
variation was high. Data collected in this laboratory and others have
Shown that high variation is not unusual for microbial population
assessments in natural soil. However, despite high variability,
statistical analysis suggested significant differences in several
cases.

Bacterial species were capable of increasing their populations in
soil amended with fern or root tissue to 2-3 fold higher levels when
compared to soils planted with asparagus, snap beans, sweet corn, or
natural soil. Fern tissue was quickly colonized by bacterial Species.
Root tissue was also colonized quickly. Decline of the bacterial
populations occured more quickly in the root tissue than in the fern

tissue. Visual assessment of the samples indicated that the general

93

Table 4: Inhibition of bacterial isolates by isolated components of

asparagus FOOT. extracts.

 

 

Isolate #a Sensitive be of
Microorganisms Active Fractions
2a Bacillus spp. 0.35
3a 8, circulans 0.82
0.80
0.26
7 B. spp 0.35
7a 8_. circul ans 0.26
10 ‘8, circulans 0.35
16 B; circulans 0.35
17 B; circulans 0.35
48 B, circulans 0.82
0.75
0.50
0.35

 

aIsolates collected from Spinks sandy loam soil never before under
cultivation.

be of purified asparagus root component inhibitory to bacterial
growth. Components were eluted from TLC Silica gel plate (developed
with hexane:ethyl acetate 1:1 as the mobile phase) with methanol or

chloroform.

94

species diversity in the soil had changed in both root and fern tissue
treatments. This indicated that tissues contained compounds that
selectively inhibited some bacteria or that certain species of bacteria
were better able to colonize the root and fern tissue than others.
Since root and fern tissue had been surface sterilized before amending
into soil, the change in species population was not caused by
contaminating bacterial Species on the dried tissues.

Both total fungal and Fusarium spp. population counts increased in
number when root tissue was incorporated into soil. Fusarium
populations increased quickly in the soil and remained at significantly
high levels throughout the duration of the experiments. Although total
fungal populations increased, counts began to decline after 4 weeks in
the first experiment and after 5 weeks in the second experiment. In
asparagus plant treatments, fungal populations were observed at levels
similar to those in the control soil. Fungal populations in the snap
bean and sweet corn treatments initially increased, then declined over
time. These data agree with observations of fungal populations in
asparagus plantations, snap bean fields, and corn fields in
agroecosystems in Michigan (unpublished data). These data may suggest
that competitive saphrophytic fungal species may be incapable of

developing in the rhizosphere of the asparagus plant.

Effects of water extracts on isolates of Pythium spp. Water extracts
of asparagus root tissue were capable of inhibiting completely or
depressing hyphal growth of isolates of Pythium spp. These data
suggest that certain soil borne fungal species are sensitive to the

compounds contained in asparagus root extracts, and that these

95

compounds may be affecting the species diversity in soils with an

excess of asparagus root tissue.

Sensitivity of bacterial isolates to purified extracts of asparagus
root tissue. One third of the bacterial isolates collected from sandy
soil were inhibited by purified extracts from asparagus root tissue.
These data suggest that isolated toxins from asparagus root tissues are
capable of reducing the diversity of saphrophytic soil bacteria.

When compared to other plant species, fungal populations are not
as high when asparagus plants are grown in the soil. However, fungal
populations, particularly Fusarium can increase suprisingly quickly on
asparagus root tissues, but not on asparagus fern tissues. Bacteria
colonize fern tissue very quickly and populations remain high in soil
where fern residue is present. This may suggest bacteria quickly
break down fern tissue, but not root tissue. Colonization and
subsequent breakdown of the fern tissue by bacteria may account for the
absense of toxicity in nonsterile soil reported in the literature.
Data presented here suggest that toxins present in asparagus root
tissues are capable of reducing the numbers and diversity of microbial
populations in natural soil. Since the herbaceous perennial asparagus
plant is always producing new tissues and older tissues are continually
dying in the soil, there is always a substantial amount of tissue in
various stages of development in the soil system. Data in these
experiments suggest fungal population and Fusarium populations may
increase in a relatively short time on excess tissue. Other
microorganisms may be detrimentally affected also. This phenomenon may
contribute to some of the problems associated with asparagus decline in

asparagus plantations.

LITERATURE CITED

Cohen, Sylvan. I. and F.D. Heald. 1941. A wilt and root rot of

asparagus caused by Fusarium oxysporum Schlecht. Plant Disease

 

Reporter. 25: 503-509.
Endo, R. M., and E. C. Burkholder. 1971. The association of

Fusarium moniliforme with the crown rot complex of asparagus.

 

Phytopathol. 61: 891 (AbstrJ

Fischer. H-P., and D. Bellus. 1983. Phytotoxicants from
microorganisms and related compounds. Pestic. Sci. 14: 334-346.
Grogan, R. G., and K. A. Kimble. 1959. The association of
Fusarium wilt with the asparagus decline and replant problem in
California. Phytopathol. 49: 122-125.

Johnson, SJL, J.K. Springer, G.D. Lewis. 1979. Epsggigm
moniliforme as a sourse of stem and crown rot of asparagus and its
association with asparagus decline. Phytopathol. 67:778-780.
Hartung, A. C., and C. T. Stephens. 1983. Effects of allelopathic
substances produced by asparagus on incidence and severity of
asparagus decline due to Fusarium crown rot. J. Chem. Ecol.
9:1163-1174.

Hartung, A. C., and A. R. Putnam. Extracts of asparagus root
tissue are phytotoxic. Proceedings of the Sixth International

Asparagus Symposium. E.CL Lockheed andlt.Tiessen, eds. Univ.

of Guelph. pp. 258-266.

96

10.

11.

12.

13.

14.

15.

16.

97

Komada, H. 1975. Development of a selective medium for
quantitative isolation of Fusarium oxysporum from natural soils.
Rev. Plant. Plot Res. 8: 114-125.

Laufer, G. A., and S. A. Garrison. 1977. The effect of asparagus
tissue on seed germination and asparagus seedling growth.
Possible allelopathic interaction. HortSci. 12:385 (AbstrJ
Lehle, F. R. and A. R. Putnam. 1982. Quantification of
allelopathic potential of sorghum residues by novel indexing of
Richards Function fitted to cumulative cress seed germinations.
Plant Physiol. 69:1212-1216.

Lockwood, J. L. 1968. The fungal environment of soil bacteria.
In: The Ecology of Soil Bacteria, an International Symposium. T.
R. G. Gray and D. Parkinson, eds. Liverpool Univ. Press. pp. 65.
Linderman, R. G. 1970. Plant residue decomposition products and
their effects on host roots and fungi pathogenic to roots.
Phytopathol. 60:19-22.

Martin, J. K. 1975. Comparison of agar media for counts of
viable soil bacteria. Soil Biol. Biochem. 7:401-402.

McCal la, T. M., and F. A. Haskins. 1961. Phytotoxic substances
from soil microorganisms and crop residues. Bact. Rev. 28:181-
207.

Patrick, Z. A., 1971. Phytotoxic substances associated with the
decomposition in soil of plant residues. Soil Sci. 13-18.
Patrick, 2. A., and L. W. Koch. 1963. The adverse influence of
phytotoxic substances from decomposing plant residues on

resistance of tobacco to black root rot. Can. J. Bot. 41:746-758.

17.

18.

19.
20.

21.

22.

23.

23.

24.

25.

26.

98

Proebstring, E. L., and A. E. Gilmore. 1941. The relation of
peach root toxicity to the re-establishment of peach orchards.
Proc. Am. Soc. Hort. Sci. 38:21-26.

Rice, E. L. 1984. Allelopathy. Academic Press. N.Y. Pp.422.

 

Rice, E. L. 1979. Allelopathy-An Update. Bot. Rev. 45:15-109.
Shafer, W. E., and S. A. Garrison. 1980. Effects of decomposing
asparagus root tissues on lettuce, tomato, and asparagus seed
emergence. HortSci. 15:406. (Abstr.).

Shafer, W. E., and S. A. Garrison. 1980. Effects of aspargus root
extracts on lettuce and asparagus seed germination and growth.
HortSci. 15:406-407. (Abstr.).

Shafer, W. E., and S. A. Garrison. 1986. Allelopathic effects
of soil incorporated asparagus roots on lettuce, tomato, and
asparagus seedling emergence. HortSci. 21(1):82-84.

Takatori, F., and F. Souther. 1978. Asparagus Workshop
Proceedings. Dept. Plant Sciences, University California,
Riverside, CA. pp 156.

Toussoun, T. A., and Z. A. Patrick. 1963. Effect of phytotoxic
substances from decomposing plant residues on root rot of bean.
Phytopathol. 53:265-270.

Toussoun, T. A., A. R. Weinhold, R. G. Linderman, and Z. A.
Patick. 1968. Nature of phytotoxic substances produced during
plant residue decomposition in soil. Phytopathol. 58:41-45.

Yang, Hsu-Jen. 1982. Autotoxicity of Asparagus officinalis L. J.
Amer. Soc. Hort. Sci. 107:860-862.

Young, C. C. 1984. Autointoxication in root exudates of W
officinalis L. Plant and Soil. 82:247-253.

 

99

27. Young, C. C. 1986. Autointoxication of Asparagus officinalis L.

in, The Science pf Allelopathy. A. R. Putnam and C. S. Tang, eds.

 

John Wiley and Sons, Inc. pp 317.

CHAPTER V

ISOLATION AND CHARACTERIZATION OF ALLELOCHEMICALS IN
ASPARAGUS (ASPARAGUS OFFICINALIS l.) ROOT TISSUE

ABSTRACT'

Isolation and characterization of allelochemicals from asparagus

(Asparagus officinalis LJ were attempted using aqueous extracts of

 

aspragus root tissues. Ferulic, isoferulic, malic, citric, and fumaric
acids were all found to be present in phytotoxic fractions isolated by
HPLC. These acids were active in Curly Cress (Lepidium sativum)
bioassays With 150 levels oflLS mg/ml for ferulic, LA mg/ml for
malic, and 1.6 mg/ml for citric. Isoferulic acid gave no inhibition of
curly cress at any level tested. Bioassays comparing activities of
combined standards and crude extracts at different pH levels showed
that inhibitory activity of known Standards and crude extracts was not
affected by pH. NMR spectra suggested that a derivative of asparagusic
acid or asparagusic acid 3-oxide was present in an inhibitory fraction
isolated from preparative TLC plates. Isolation of inhibitory
compounds using soxhlet extraction of dried asparagus root tissue
revealed caffeic acid to be an additional phytotoxin in the root
tissue. However, bioassays comparing the inhibitory activity of a

caffeic acid standard with the crude caffeic acid fraction Showed pure

100

101

caffeic acid to be less active (150=2.0 mg/ml) than the crude fraction
(150=0.26 mg/ml). This suggested that a derivative of caffeic acid

might be involved.

INTRODUCTION

Asparagus (Asparagus officinalis L.) is a perennial vegetable crop
grown on sandy soils in temperate areas of the EaStern and Western
United States. Asparagus decline, resulting in reductions in
commercial yields and quality, and death of crowns in fields
continuously cropped to asparagus is a well known and well documented
problem on asparagus in many production areas (2, 4, 8, 7). Asparagus
decline is ultimately attributed to the presence of two soil-borne
pathogens, Fusarium oxysporum (Schlecht. ) emend. Snyd. and Hans. f.

Sp. aSparagi Cohen, and F_. moniliforme (Sheld.) emend. Snyd. and Hans.,

 

that once introduced to ecosystem, survive in the soil indefinitely (1,
2, 4, 9). However, other laboratories as well as this one, have
presented evidence that asparagus is an autotoxic or autoallelopathic
plant (5, 6, 7,11,16,17,18, 20, 22). Allelochemicals are defined
as compounds released from one plant (or microorganisms) that affect
the growth of another plant (14). Autotoxicities are considered aS
special cases of allelopathy, and the same compounds may exert
interspecies effects. Compounds released from living or senescing
asparagus root tissue are hypothesized to damage the asparagus root
system, which may decrease the growth of the crown (ll, l6, 17, 18, 20,
22, 23). Injury to the root system from these autotoxic compounds may

also provide infection courts for the entrance of the Fusarium

102

pathogens, therefore enhancing the incidence of disease on the plant
(5). Young (23) has reported that several phenolic compounds are
released from the root system of intact asparagus plants. Using an
XAD-4 resin trapping procedure originally developed to recover phenolic
and chlorophenoxy herbicides from soil, he isolated 3,4-
dihydroxybenzoic acid, 2,6-dihydroxybenzoic acid, 3,4-
dihydroxyphenylacetic acid, 3,4-dimethoxyacetophenone, and b-(m-
hydroxyphenyl)propionic acid. However, he did not report if these
compounds were inhibitory to asparagus seedlings nor did he examine if
senescing tissue possessed any inhibitory compounds that may also be
contributing to the damage in the field. He also did not determine if
these compounds were of plant or microbial origin. Kitahara, et. a1.
(10) isolated three compounds they called growth regulators from
etiolated asparagus ferns, asparagusic acid (1,2-dithiolane-4-
carboxylic acid) with activity comparable to abscisic acid,
dihydroasparagusic acid (8,8'-dimercaptoisobutyric acid), and S-
acetyldihydroasparagusic acid (B-S-acetyl-b'-mercaptoisobutyric acid)
(Figure 1). Though these compounds are considerably toxic to seedling
germination and radical elongation of several seed Species, these
compounds occurred at extremely low concentrations in the tissues and
are quite labile once isolated. Asparagusic acid has also been
isolated from the roots of asparagus and found to be inhibitory to
nematodes, Specifically the emergence of the second stage larve of

Heterodera rostochiensis and H;_glycines and the second stage larvae of

 

H, rostochiensis and Meloidogyne hapla, and the larvae and adults of

Pratylenchus penetrans and Paratylenchus curvitatus (19). A patent is

 

presently held by Sankyo Co. Ltd., Japan, on asparagusic acid and its

derivatives for activity on nematodes and plant growth regulation.

103

Since asparagus possesses large storage roots that are continually
dying as the crown grows, large amounts of asparagus root material may
be present in asparagus fields at any one time. It has been reported
that these root residues contain a considerable amount of toxic
substances that inhibit seed germination and radical elongation of
different seedling species (16, 17, 18), as well as asparagus growth,
peroxidase activity of asparagus seedlings, and respiration of the
crown (7). At this point, no one has isolated compounds that can
account for all the inhibitory activities present in the root tissue.
Also, even though several compounds have been isolated from asparagus
tissues, none have been isolated from or Shown to be active in the
agroecosystems where asparagus is grown. The purpose of this research
was to attempt to isolate compound(s) from asparagus root tissue that
are responsible for the reported allelopathic properties of asparagus
and test their effectiveness in bioassays, and to try and recover these

compounds from the soil.

MATERIALS AND METHODS

Preparation of asparagus plant material. Asparagus crowns (cv. Martha
Washington) were excavated from a research plot (7 yr old) located at
the Horticultural Research Farm, Michigan State University, E. Lansing,
MI. The storage and fibrous roots were separated from the rhizomes and
all dead and visibly diseased plant material was discarded. The roots
were oven dried at 50 C and ground in a Wiley Mill (mesh screen size=1
mm). All plant tissues were sterilized with propylene oxide gas and

checked for microbial contamination as previously reported (6, 7).

104

Containers with sterilized root tissue were allowed to exhaust under a
fume hood for 24 hr to dissipate any residual propylene dioxide. Dried
sterilized tissue was stored in brown glass bottles at -20 C until used

for chemical isolation.

Bioassay procedure for evaluation of isolated components of asparagus
root extractions. Previous studies in this laboratory have shown that

Lepidium sativum L. (cv. Curly Cress) is a fast and reliable seedling

 

for evaluating the inhibitory activity of many allelopathic compounds
(12). Therefore, curly cress was used to test for inhibitory activity
from root extracts throughout each step of all isolation procedures and
for evaluation of activity from known standards of isolated compounds.
All fractions to be assayed were first taken to dryness and weighed.
Separate fractions were then redissolved in methanol (or an appropriate
solvent, depending on solubility of the fraction) and a dilution series
was made. A known amount of each fraction was Spotted on filter paper
(Whatman #1) in glass petri dishes. Evaporated solvent controls and
DH20 (deionized water) controls were always included in each bioassay.
Also, crude extract of asparagus root often included as a control when
appropriate. All fractions were filter sterilized before bioassaying.
The solvent was allowed to evaporate completely from the filter paper
and 1-5 "'1 01’ DHZO was added to the dish. Curly cress seeds (IO/dish)
were added to the dish and distributed uniformly with a glass rod. The
dishes were covered, arranged in a randomized complete block design,
and placed in a moist chamber for 72 hr at 26 C. Root length was
measured and 150 levels determined by interpolation. There were 3

replications per treatment in all cases except when paucity of the

105

compound only allowed for 2 replications or single observations. When
replication was impossible, a standard error was calculated for the
root length of individual seedlings. When there was replication, the
data were subjected to an Analysis of Variance and the means separated

by the LS0 test.

Bioassays of known compounds and crude extracts. When available,
standards of isolated compounds were tested for inhibitory activity and
compared to those of the crude inhibitory fractions from the 018
column. Known standards were bioassayed from 0.025 ug/ml to 2.0 mg/ml
concentrations.

Bioassays comparing the known standards and the active fractions
from the C18 column at different pH levels were done to assure that
inhibition seen in the fractions was not simply a pH effect. The C18
fractions and the known standards were bioassayed at their original pH
as well as the pH of the other fraction. The standards were combined,
the pH determined (pH=3.6), then the solution was separated into two
treatments; one was left at the original pH and the other adjusted to
the pH of the 018 fraction (pH=5.6). The same procedure was done with
the C18 fraction. The combined standards and the crude were bioassayed
at 125, 250, and 500 ppm. The C18 fraction was also bioassayed at pH
7.0 at 25, 50, and 150 ppm. A buffer control and methanol solvent

control were also included in the bioassay.

Purification of inhibitory components in asparagus root extracts:
Scheme #1. Previously published data from several laboratories and
this one have indicated that water extracts of asparagus root tissues

are inhibitory to seedling germination and radical elongation (6, ll,

106

17). Therefore water extracts from asparagus root tissue were
subjected to sequential extraction with solvents to purify the chemical
components implicated in the inhibitory responses. Ground asparagus
root tissue was extracted by stirring overnight at 4 C (1:10, root
t1'SSHJFRDHZO) (Figure 2). The particulates were removed by filtering
through four layers of cheese cloth and centrifuging at 6,000 x g for
20 min. The supernatant was decanted carefully from the pellet (the
pellet was discarded) and then proteinaceous and lipophyllic materials
precipitated with acetone (4:1 v:v acetone:sample) by Slowly stirring
overnight at 4 C. The precipitate was removed from the eluant by
filtering through filter paper (Whatman #4, #1, and #42 respectively)
and bioassayed. The precipitate was discarded and the liquid
concentrated to 1/4 the original volumn on a Buchi Rotary evaporator at
40 C. This concentrate was sequentially extracted with the following
solvent series: chloroform, diethyl ether, dichloromethane, ethyl
acetate and water-saturated n-butanol. The aqueous root extract was
partitioned three times (1:1 v:v), with each of the six solvents. Each
solvent fraction was dried with MgSO4, concentrated to dryness by
rotary evaporation at 40 C, weighed and bioassayed at 10 and 100 ppm.
The data were subjected to an Analysis of Variance and means separated
by the LS0 test. The percent of the original fraction represented by
each solvent extraction was calculated.

The chloroform fraction was further purified by column
chromatography (Figure 3L. The dried chloroform fraction was first
redissolved in methanol, after which a white precipitate formed. The
precipitate was separated from the methanol soluble supernatant by

filtering through filter paper (Whatman #4, #1, and #42 respectively)

107

Figure 2. Flow diagram for the extraction and partitioning of dried

asparagus root tissue.

 

 

108

EXTRACTION PROCEDURE

DRIED ASPARAGUS ROOT TISSUE
1:10

stir slowly at 4 C
overnight

 

V

CENTRIFUGE
20 min at 6000 g

......... Bioassay lyophilized
aqueous fraction

 

V
ACETONE PRECIPITATION
4:1

stir slowly at 4 C
overnight

 

V
FILTER
#4, #1, #42 filter paper

......... Bioassay precipitate
discard

 

V

REMOVE ACETONE BY ROTOEVAPORATION

.......... Bioassay aqueous
fraction
V

PARTITION AQUEOUS EXTRACT

---- extract 6 x each wit
chlorofbrm 2
diethyl ether 1
dichloromethane l
ethyl acetate 1
n-butanol 2

 

V

REMOVE SOLVENTS BV ROTOEVAPORATION

V

BIOASSAV

Figure 3.

109

Flow diagram for the chromatographic separation of
compounds from the chloroform extract of dried asparagus

root tissue.

 

 

 

110

COLUMN CHROMATOGRAPHY QE_CHLOROFORM FRACTION
CHLOROFORM FRACTION

dry, weigh, redissolve in Methanol

 

l
PRECIPITANT METHANOL SOLUBLE
(not active) I

V
C18 FLASH COLUMN
elute with 100 ml each of:
acetonitrile 100%
acetonitrilezmethanol 75:25
acetonitrilezmethanol 50:50
acetonitrilezmethanol 25:75

 

methanol 1001
F I
FRACTIONS 94-8 FRACTION #1-3
(not active) (active)

V

SILICA GEL FLASH COLUMN
elute with 200 ml each of:
chloroform 100%
chlorofbrmzmethanol 98:2
chlorofbrmzmethanol 96:4
chloroform:methanol 90:10
chloroform:methanol 85:15
chlorofbrm:methanol 80:20
chloroform :methanol 70:30
chlorofOrmzmethanol 40:60
methanol 1001

i i

V
FRACTION #1,2,8,9 FRACTION #4,5,6,7
(little activity) (active)

V

HPLC

C18 column
mobile phase: acetonitrilezflPLC-grade H20 1:99
flow rate= 1 ml [min
UV dectector 254 nm and 280 nm

 

 

I v I
7 peaks 5 peaks 5 peaks

1 active 1 active 3 active

111
in a buchner funnel. The methanol soluble fraction was further
purified using an octadecyl (C18) bonded phase solid support in a Baker
flash chromatography column (190 mm x 20 mm) eluted with a Step
gradient of acetonitrile (100%) to methanol (100%) at 25% intermediate
steps in 100 ml fractions. Pressure for the column was provided by
laboratory air line at a rate of 0.2 cm/sec. Fractions (25 ml) from
the column were collected, spotted on thin layer chromatography plates:
(Whatman silica gel 60 F-254; mobile phase: chloroform:methanol 9:1 and
chloroform:methanol 8:2), then recombined according to Similar Rf
values. The fractions were dried under nitrogen, weighed, and
bioassayed at 25, 50, and 100 ppm. The first three fractions were
combined and separated further on a Baker flash chromatography column
(180 x 30 mm) using a silica gel solid support and a mobile phase
gradient of chloroform (100%) to methanol (100%) in the followdng
order: 100% chloroform, 98:2, 96:4, 90:10, 85:15, 80:20, 70:30, 40:60,
100% methanol. Each gradient step was applied to the column in 200 m1
fractions. Air pressure for the column was as above. Fractions of 75
ml each were collected, concentrated, and developed on TLC plates as
above. Fractions with similar Rf values were combined, dried under
nitrogen, weighed, and bioassayed at 25 and 50 ppm. Individual
fractions were then further purified by high pressure liquid
chromatography (HPLC) using a Waters uBondapak C18, 8 mm x 10 cm radial
compression column. The UV detector (Waters 490 multiwave
programmable) measured absorbance at 254 and 280 nm. The solvent
system was acetonitrile:H20, 1:99 at a flow rate of 1.0 ml/min. The
mixture was resolved into 5 distinct areas and 5 fractions were
collected after repeated injections. Three separate fractions from the

silica gel flash columns were further separated in this manner. Each

112

fraction from the HPLC collection was dried under nitrogen, weighed,
and bioassayed at 50 ppm on curly cress. From each of these HPLC
separations, purified samples were sent to Dr. Basil Burke at Arco
Plant Cell Research Institute, Dublin, CA, for spectral evaluation.
HPLC fractions were silylated and then evaluated by GC-Mass
Spectroscopy (GC-MS) using a temperature gradient from 110-250 C at 15
degrees/min. Spectral scans from total ion chromatogram were observed
and a library search done on each Spectrum using the Finnigan NBS
Library Compilation series. Standards of elucidated compounds were
then injected into the GC-MS (150/min on a 30 M DBS column) under the
same conditions to further substantiate the identification of these
compounds by matching GC retention times, as well as comparing Mass

Spectra.

Purification of inhibitory compounds in dried asparagus root tissue:
Scheme #2; The methanol soluble portion of the chloroform fraction
isolated as above was also separated using preparative TLC. Silica gel
(Merck DC-Fertigplatten, 60 F-254 0.5 mm, 20 x 20 cm) plates were first
washed in 85:15 chloroform:methanol, then approximately 600 mg of the
methanol soluble chloroform fraction was streaked near the base of the
plate. In all, 6 plates were used. Plates were run twice then viewed
under UV (254 nm) light. Nine distinct bands were scraped off the
Silica plates and eluted from the Silica gel using chloroform 100%, (or
chloroform:methanol, 5:1 v:v for bands at Rf>0.4) through a fritted
glass filter and collected, dried, weighed, and bioassayed at 30 ppm.
One fraction (Rf=0.53) was rechromatographed on Silica gel plates as

above. Again nine fractions were detected by UV (254 nm) light,

113

scraped and eluted from the plates as above. A portion of five of
these fractions (Rf=0.69, 0.61, 0.52, 0.46, 0.29, and 0.07) were
silylated for GC-MS analysis and the 1H NMR Spectrum of remaining
portion of each of these small fractions were obtained deuterated
methanol on a Varian XL 300 MHz nuclear magnetic resonance Spectrometer

(NMR).

Purification of inhibitory components from dried asparagus root tissue:
Scheme #3. Dried asparagus root tissue (250 g) was extracted with
hexane for 4 hr using a soxhlet extraction device. The hexane was
removed and replaced withinethanol and the tissue further extracted
overnight with methanol. The methanol extract was then acidified by
dilute HCl to pH 2.0 and extracted with 50 ml of ether 5 times. This
ether fraction was then extracted with 75 ml of 5% aqueous sodium
bicarbonate 2 times. The alkaline portion was then acidified with HCl
to pH 2.0, then extracted with 50 ml of ether 3 times. Fractions were
then visualized on TLC after development with mobile phase;
chloroform:methanol 2L5:1.5, and mobile phase; toluenezethyl
formatezformic acid, 5:4:1L. In the last ether fraction, a yellow
compound crystallized on the sides of a round bottomed flask. These
crystals were removed by decanting the ether from the flask, washing
the crystals with hexane, then dissolving the crystals in methanol.
The crystals were prepared with deuterated methanol for structural
elucidation on the NMR.

All fractions from this isolation procedure were bioassayed on
curly cress. The data was subjected to an Analysis of Variance and the
means separated by LSD. A dilution series of the last ether fraction

was bioassayed at 0.025-2.0 mg/ml using curly cress.

114

Isolation of inhibitory components from soil. Soil was collected from
the rhizosphere of 20 and 40 year-old commercial asparagus fields where
asparagus decline had been observed in Oceana County, MI, as well as
adjoining control fields where no asparagus was grown. There was no
possibility of water runoff from the adjacent asparagus field into the
control area, and the soil type was essentially the same as in the
asparagus field. Soil was sieved through a 2000 0 screen to remove
most plant fragments. Soil (4.5 kg) was extracted with MethanolzDHzo
overnight at 4 C. Soil particulates were removed from the
methanol:water solution and the methanol removed from the aqueous
fraction by rotoevaporation. Precipitation with acetone and subsequent
removal and extraction with chloroform was done on the soil extract as
described in the procedure for asparagus tissue extraction. The
chloroform soluble fraction was concentrated, dried under nitrogen,
weighed and bioassayed on curly cress as described above at
concentrations ranging from 32.7 mg/ml to .032 mg/ml. There were three
replications per treatment and the data were subjected to Analysis of

Variance.

RESULTS

Purification of inhibitory components in asparagus root extracts:
Scheme #1. Water extraction of dried asparagus root tissue extracted
approximately 35% of the mass from the original material. After
acetone precipitation, approximately 83% of the water extract remained.
Chloroform extraction accounted for 0.022%, diethyl ether, 0.009%,

dichloromethane, 0.004%, ethyl acetate, 0.011%, and n-butanol, 0.12% of

115

the original dried root material. When bioassayed on curly cress, the
chloroform fraction contained the highest Specific activity ( 150:1755
ppm hithis extraction)(Figure 4). The diethyl ether fraction also
had considerable activity (150=34 ppm). Dichloromethane and ethyl
acetate fractions also contained inhibitory activity but to a lesser
extent than the chloroform and ether fractions (I50=82 and 100 ppm
respectively). Also, at lower concentrations, the dichloromethane and
ethyl acetate fractions were slightly stimulatory. Because of the
greatest inhibitory activity in the chloroform fraction, this fraction
was chosen for further isolation attempts. By refining the isolation
technique, chloroform sometimes removed as much as 0.17% of the
original dried material, but based on a weight for weight basis, 150
levels were aways very similar, usually being less than 40 ppm for each
isolation.

Fractionation of the methanol-soluble portion of the chloroform
fraction on an octadecyl bonded phase soild support flash column and
subsequent bioassay showed the first 3 fractions to be inhibitory
115o=25 ppm). ‘TLC indicated that these fractions were chemically
Similar and they were therefore combined. However, these active
fractions still contained several chemical components and were further
fractionated on a silica gel solid support flash column. Nine
fractions were collected, 4 of which gave 98% inhibition of curly
cress at less than 50 ppm. Fraction #1 and #2 contained some
inhibitory activity at 100 ppm but had no Rf values in common when
developed on TLC plates. When developed on TLC, the four inhibitory
fractions Showed only one common Rf area so were kept separate. 150
levels for these fractions were 35 ppm for fraction #4, 58 ppm for

fraction #5, 18 ppm for fraction #6, and 30 ppm for fraction #7.

Figure

4.

116

Response of curly cress to 10 and 100 ug/ml of solvent
extracts from aqueous extract of dried asparagus root

tissue.

 

 

 

Radicle Length (mm)

—A
D

  

 

117

I 10: IOpg/ml
0100: lOOpg/ml

10 10

f
/

    

10

 
    
 

190

  

100 100
r r

/

4 / / r / /

CH0!5 Ether MeCI2 EtOAc Butanol Control

Solvent Extraction of Root Tissue

  

118

Because the 150 levels were quite similar after this procedure, the
active components were suspected to be either poorly separated or
interacting with the column solid support. However, fractions
contained substantially fewer chemical Species than before indicating
that the silica column provided some clean-up of the mixture.

The components in fraction #6 were separated on HPLC. Recovery of
dry weight from repeated injections for this separation was 67%. Five
separate peak areas were collected and bioassayed on curly cress. Only
one peak area proved to contain inhibitory activity. This component,
when evaluated by GC-MS still appeared to contain at least 12 different
compounds. A library search done for matching spectra, and injection
of known standards at the same conditions on the GC-MS revealed the
presence of fumaric acid, malic acid, isoferulic acid, ferulic acid,
dihydrocitric acid, and citric acid (hydroxyl group position for
citric acid could not be determined), as well as several sugar moieties
(Figure 5L. GC-MS of other HPLC isolated peaks collected under similar
conditions also Showed the presence of the above mentioned acidic

compounds and sugars.

Bioassay of known standards and C18 fraction. When standards of
identified compounds were bioassayed on curly cress, 150 levels were
(L5 mg for ferulic acid, 1.46 mg for malic acid, and 1.58 mg for citric
acid. The 150 level for C18 fraction #1 in this bioassay was 0.029 mg.

There was no difference in the activity seen at pH=5.6 or 3.5 in
the crude fraction 1150 for both treatments=42 ppm). The combined
standards only inhibited curly cress at the 500 ppm level. Inhibition

for the standards was only significant at P=0.05 and no 150 could be

Figure 5.

119

Total ion current of GC-MS of Fraction #3 of peak collected
from HPLC separation of dried asparagus root extract tha
was inhibitory to cress seed germination and radicle
elongation. A, B = silylated peaks; C = Fumaric acid; 0 =
Malic acid; E = Dihydro citric; F = Mannose; G = Isomer of
ferulic acid;ii= Glucopyranose; and.J==Ferulic acid or

isoferulic acid.

 

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121

determined.

There was slight inhibition of the curly cress at the higher
concentrations of the phosphate buffer control in bioassays comparing
the C18 extract at pH=5.6 and 7.0. Therefore, the evaporated methanol

control may be a more appropriate benchmark. Using the methanol

solvent control. the 150 level, the 150 for the crude was 20 ppm.

Purification of inhibitory compounds in dried asparagus root tissue:
Scheme #2. When the compound at Rf=0.07 was subjected to 1H NMR in
deuterated methanol, the resulting Spectrum suggested the structure may
be that of asparagusic acid-l-oxide (1,2-dithiolane-4-carboxylic acid-
1-oxide) or asparagusic acid (1,2-dithiolane-4-carboxylic acid) (Figure
6,1). The Spectrum also reveals several Signals representing
impurities associated with the methods used. The singlet at. 61.22
represents a co-eluate from TLC adsorbents, an artifact usually
observed when small quantities are purified by preparative TLC.
Singlets at 6 3.22 and 4.80 correspond to undeuterated methanol,
partially'deuterated water, and the exchange of the OH group on the
acid moiety of the compound. The spectrum also shows a clear multiplet
at 6 3.98-4.12 ( 1 H, m) that is attributed to H3, an absorption at
6 3.82-3.87 (1 H, dd, J=8 Hz, J=6 Hz), corresponding to H4, absorption
at 6 3.6-3.66 (2 H, J values unable to be determined) corresponding to
H2 and H5, and absorption at 6 3.45-3.54 (1 H, t J=6 Hz) attributed
to “1 (Figure 1). These values agree well with those published by
Yanagawa (21 ), except for a slight downward shift of the ppm values.
This downward shift is probably due to the difference in solvent use to
dissolve the fraction in, and the difference in the type of NMR used to

evaluate the sample. When this compound was silylated, the GC-MS

Figure 6.

122

1H NMR of a compound isolated from asparagus root tissue
using preparative TLC, mobile phase, chloroform:methanol
8.5:1.5. The compound at Rf=0.07 was scraped from the

plate and eluted from the Silica using cholorformzmethanol

5:1.

 

 

123

 

 

 

 

 

1:

 

 

 

 

 

The“

 

 

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m .2

u N 2 33: o

Figure

1:

124

Chemical structures irf plant growth regulators and
nematicides isolated from etiolated asparagus shoot and
root tissue. A = asparagusic acid-l-oxide; B =
asparagusic acid;(2= S-acetyldihydro asparagusic acid;

and D = dihydro asparagusic acid.

 

 

126

showed the silylated methylated molecular ion at m/z=222, and a
fragmentation pattern with a peak at m/z=189 indicating a loss of an SH
group, and m/z=189, a loss of the silylating group and 1 hydrogen,
m/z=129, a rearrangement, and the base peak at m/z=73, loss of the
silylating group (Figure 7). This fragmentation pattern is suggestive
of asparagusic acid, 1,2-dithiolane-4-carboxylic acid or 1 of its other
dirivatives. From data available, it was not possible to distinguish

completely which dirivatives are actually'present.

Purification of inhibitory components from dried asparagus root tissue:
Scheme #3. When a crystallized component from the last ether fraction
was subjected to 1H NMR in deuterated methanol, the spectrum revealed a
Singlet at 1.30, 3.30 and 4.92 due to solvent impurities,
undeuterated methanol and partially deuterated water, respectively.
The other protons of this compound appeared at 7.53 (1 H, d, J=15
Hz), 7.01 (1 H, d, J=2 Hz), 6.92 (1 H, dd, J=8 Hz, J=2 Hz), 6.78 (1 H,
d, J=8 Hz), and 6,21, (1H, d, J=15 Hz) (Figure 8). These data suggest
the structure was that of caffeic acid, 2,4-dihydroxycinnamic acid.
When a standard of caffeic acid was run under the same conditions in
the 1H NMR, the Spectra of the known standard and the purified unknown
from the ether fraction matched exactly (Figure 9).

When the caffeic acid standard was compared to the last ether
extract and the C18 crude active fraction, 150 levels were 1.98 mg,
0.25 mg, and 0.03 mg respectively. Bioassays of all fractions from
this isolation procedure Show inhibitory activity only in the last

ether fraction.

Figure 7.

127

GC-Mass spectrum of a compound isolated from asparagus root
tissue using preparative TLC, mobile phase,
chloroform:methanol 8.5:1.5. Compound at Rf=0.07 was
scraped from the plate and eluted from the silica gel with

chloroform:methanol 5:1.

128

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129

Figure 8. 1H NMR spectrum of compound x isolated from asparagus root
tissue using soxhlet extraction. The crystalline compound

was dissolved in deuterated methanol.

130

 

 

 

 

 

 

 

 

 

 

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131

Figure 9. 1H NMR of a known standard of caffeic acid dissolved in

deuterated methanol.

132

 

 

 

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133
Isolation for inhibitory conponents frol soil. In all bioassays on
curly cress with chloroform extracts from both asparagus fields and the
control soil, no inhibitory activity was noted except at concentrations
of 32 mg/ml. Therefore, there were no differences in inhibitory

activity from the three soils.

DISCUSSION SECTION

Isolation of compounds from asparagus root extracts revealed the
presence of several known acidic compounds that have been previously
reported to be important in allelopathic interactions. Ferulic acid
has been reported as a germination inhibitor produced by Eiflfilifli
glyssum, and present in residues of corn, wheat, sorghum and oats.
This compound has also been isolated from soils under al lelopathic
plants (lSL. Fumaric acid is a well known microbial toxin produced by
Rhizopus spp. in hull rot disease on almond (13). Also, isolations
suggested that asparagusic acid or one of its derivatives were present
in the inhibitory fractions. These are extremely active growth
inhibitors (l9, Zl). Caffeic acid is reported to be fungistatic
against Helminthosporium carbonum in potatoes, and phytotoxic against
many plant species and families (l5). These compounds and the
inhibitory fractions were shown to be active at varying pH levels. The
fact that all inhibitory activity cannot be accounted for by pH has
also been documented in the literature specifically for malic and
citric acid (3). The known standards of several of the acidic
compounds were shown to be inhibitory in our bioassay system but never
to the extent to explain all of the activity of the crude extracts of

asparagus root tissue. Also, inhibitory activity was present in other

134

solvent extracts other than the chloroform extract used for isolation
procedures. These may have interacted in an additive or synergistic
manner with the isolated compounds. Also, the cress seed bioassay may
not reflect all the biological activity'of the active components.YOung
(23) reported isolating several. compounds from asparagus root
exudation. These compounds were not present in any fractions from our
isolation procedures. He did not test for activity of these compounds
on asparagus or any other plant species. Also, since his extracts were
from nonsterile sand, it is difficult to determine if these compounds
were released from the asparagus root system or are products from
microbial species or microbial breakdown of root exudates.

In soil extraction experiments, the methanol:water extraction
procedure may have failed to remove allelochemicals bound tightly to
soil particles. Also, since allelochemicals from asparagus will be
active in the very narrow region of the asparagus root rhizosphere,
isolation attempts from bulk soil may not be representative of chemical
concentrations in that region. Allelochemicals will be active on the
surface of the root, through uptake by the plant and subsequent effects
on the biochemical processes of the plant, or through inhibition of
beneficial rhizosphere microflora. Therefore, concentrations of
.allelochemicals in bulk soil may appear quite dilute.

Our experiments show that asparagus root tissue contains a number
of inhibitory components important in the inhibition of seed
germination and radicle elongation. These toxic components may be

released from asparagus root tissue through exudation or degradation of
the root tissue and may play a role in the alleged allelopathic

activity of the asparagus plant. These data, as well as data presented

l35
elsewhere (4, 5, 6) suggest that the allelopathic properties of
asparagus are not due to the presence of any one chemical compound
acting in isolation from other biotic factors associated with the
asparagus agroecosystem, but instead is due to the presence of many
chemical components that may act differently depending on soils,
environment conditions and microbial populations. Further
clarification of the chemicals released by asparagus root tissue, and
their biological activity on asparagus plants is needed to substantiate

their role in asparagus decline.

LITERAIURE CITED

Cohen, S. 1., and F. D. Heald. 1941. A wilt and root rot of
asparagus caused by Fusarium oxysporum Schlecht. Pflant Disease
Reporter. 25:503-609.

Endo, R. M., and E. C. Burkholder. 1971. The association of
Fusarium moniliforme with the crown rot complex of asparagus.
Phytopathol. 61:891 (Abstr.)

Evanari, M. l949. Germination inhibitors. Bot. Rev. l5(3):l53-
194.

Grogan, R. G. and K. A. Kimble. 1959. The association of
Fusarium wilt with asparagus decline and replant problem in
California. Phytopathol. 99:122-125.

Hartung, A. C” and C. T. Stephens. 1984. Allelopathic
properties of asparagus: Interaction with Fusarium spp. and
bioassay techniques. Phytopathol. 74(7):800 (Abstr.)

Hartung, A. C., and C. T. Stephens. 1983. Effects of
allelopathic substances produced by asparagus on the incidence and
severity of Fusarium crown rot. J. Chem. Ecol. 9:1163-1174.
Hartung, A.(L, and A.FL Putnam. 1985. Extracts of asparagus
root tissues are phytotoxic. Proc. of the Sixth International
Asparagus Symposium. E. C. Lougheed and H. Tiessen, eds. Univ.
Guelph. Pp. 258-266.

Herner, R. C., and G. Vest. 1974. Asparagus workshop
proceedings. Department of Horticulture. Michigan State

University. Pp. 79.

136

10.

11.

12.

13.

14.

15.

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Wiley and Sons, Inc. Pp 317.

CHAPTER VI

SUNNARY AND CONCLUSIONS

Asparagus, like many other perennial crops is afflicted with
autotoxicity and replant problems. Classical studies that have
addressed similar problems reveal that they are complex and may involve
an array of pathogens, pests, nutritional factors and allelochemicals.
The asparagus decline problem also appears to be complex and involves
the crown and root rotting organisms, Fusarium oxysporum fisp. asparagi

and f; moniliforme. These diseases are believed to ultimately cause

 

plant death.

This dissertation indicates that a series of other factors may
interact with the asparagus plant or the pathogens to make the plant
more susceptible to disease. For example, when asparagus root tissues
or extracts are present, young asparagus plants become more susceptible

to infection by f; oxysporum flsp. asparagi and f; moniliforme. Since

 

asparagus root extracts induce electrolyte leakage, decrease peroxidase
activity, and decrease respiration in young asparagus roots, it may be
hypothesized that compounds present in these extracts adversely
affected the cell membranes making them more susceptible to invasion by
the Fusarium organisms. Decreased peroxidase activity in particular,

has been related to increased disease incidence in other species.

139

140

The addition of asparagus root and fern tissues to soil produces
toxicity not only on asparagus, but on a variety of dicotyledon and
monocotyleton indicators. Since asparagus seedlings germinate and grow
so slowly, another bioassay species was needed to quickly assess the
toxicity of asparagus allelochemicals. Curly cress (Lepidium sativum)
proved to be a reliable species for this purpose. When asparagus
tissues or extracts were incubated under aerobfl: or anaerobic
conditions, their toxicity was generally decreased as compared to the
toxicity of freshly prepared tissues or extracts. This indicates that
over the short term, soil microbes tend to be detoxifiers other than
producers of toxins. This is in contrast to other cases of
allelopathy, particularly those involving cyanohydrin compounds, where
microbes are necessary to release the toxins.

Asparagus tissues in the soil had pronounced influences on the
soil microbial dynamics. Most importantly, the Fusarium species
rapidly increased whenever root tissues were present in the soil. In
contrast, Pythium and figglllus species were strongly inhibited by
isolated fractions from asparagus root extracts. The compounds found
to inhibit the microbial species may be the same or different from
those important sseedling inhibition. The data suggest that several
compounds are important in the inhibitor interactions. In general, the
bacterial and fungal populations were considerably lower in soils where
aSparagus grew in contrast to soils containing snapbeans and corn. It
is hypothesized that the compounds released by asparagus shift the
microbial balance in the rhizosphere in favor of the Fusarium
organisms. This may occur through a reduction of their competitors or

because they are better able to utilize the substrates in asparagus

141
tissue. In any event, the asparagus rhizosphere creates an environment
especially suited for growth of Fusarium species.

Several biologically active compounds were isolated from asparagus
root tissues. In any isolation procedure, the investigator must make
arbitrary decisions about which fractions to follow. Some fractions
with lower specific activity may receive lower priority, but should not
be totally ignored. Since the chloroform fraction displayed the
highest specific activity in our fractionations, these compounds were
isolated first. A variety of organic acids were isolated in this
fraction. The TCA cycle acids, citric, malic, and fumaric are commonly
implicated in allelopathy. They are common seed germination and
seedling growth inhibitors in fruits and herbage of many craps.
Release of high quantities of these materials around asparagus roots
could produce a high acidity condition in the rhisozphere. The ferulic
and isoferulic acids found in root tissues have often been implicated
iniallelopathy. The former compound is persistent enough in soil to be
implicated as a toxicant from several crop residues. Ferulic acid is
one of the most frequently mentioned allelochemicals that is isolated
from soil. Soxhlet extraction of asparagus tissue also removed caffeic
acid, which isalknown inhibitor of higher plant and fungal growth.
Caffeic acid is particularly toxic to the root growth of germinating
seedlings and might influence the susceptibility of young asparagus
roots to disease invasion.

The NMR spectra suggested that a derivative of asparagusic acid or
asparagusic acid-i-oxide was present in the root tissue. These
compounds were previously reported to be highly phytotoxic and
nematocidal. Since none of the other isolated chemicals showed

activity high enough to account for the high activity of the semi-

142
purified extracts, these compounds or their analogs may contribute
considerable activity. Further chemistry work should be focussed on
these compounds and how they might interact with the other
allelochemicals from asparagus roots. Once these compounds and their
activities are well characterized serious attempts through trapping or
extraction must be directed toward their isolation from the asparagus
rhizosphere. This will provide more convincing proof of their role in

allelopathy in the asparagus decline syndrome that occurs in the field.