”was...“
‘:51 V
- ‘14..-. a. .
. . r .
9 K
. urn... (.74:

«#55. ¢

.1 L...

3 .51.

I a _.
)1! F»
a,
.-

x
n

In!

‘ 2.01”

If“ -

33!

L .
"Lo

“’3.”

Auwufll.

. .
+7.».E... mu,
.ng u“ T x
. 659111..

 

LIBRARY
Michigan State
University

This is to certify that the
thesis entitled

FUNGAL DISEASE SUPPRESSION IN SOIL AMENDED
WITH COVER CROPS AND OTHER ORGANIC INPUTS

presented by

KANCHAN UDAY DATE

has been accepted towards fulfillment
of the requirements for the

MS. degree in HORTICULTURE

 

 

 

 

12/10/2004

 

Date

MSU is an Affirmative Action/Equal Opportunity Institution

 

 

 

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE DATE DUE DATE DUE

059B“ @309

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

2/05 exam .r p. 1'5

 

—__.

FUNGAL DISEASE SUPPRESSION IN SOIL AMENDED WITH COVER CROPS
AND OTHER ORGANIC INPUTS

By

Kanchan Uday Date

A THESIS
Submitted to
Michigan State University
In partial fulfillment of the requirements
for the degree of

MASTER OF SCIENCE

Department of Horticulture

2004

ABSTRACT

FUNGAL DISEASE SUPPRESSION IN SOIL AMENDED WITH COVER CROPS AND
OTHER ORGANIC INPUTS

By

Kanchan Date

Effect of different soil amendments on suppression of Pythium ultimum, Rhizoctonia
solani and F usarz'um solani was assessed. The two main factors evaluated in growth
suppression were cover crop species (Brassicajuncea L. or Secale cereale L.) and tissue
type (root, shoot or root + shoot). In the first field based experiment, comparison was
done between effect of mustard and rye cover crop residues on potato root and
tuber health. In the field, treatment with mustard residues of any tissue type and
the combination of and rye roots + shoots tissues most effective in controlling R.
solani infection of potato roots. A greenhouse container experiment to determine
the effect of different amounts of mustard root and shoot residue on disease
infection showed that the highest amount of shoot input provided the maximum
disease suppression. Similar results were observed in the laboratory and
greenhouse bioassays where maximum benefit of residue was observed at 24 h.
As time passed, the fungal growth inhibiting effect decreased. In the second field
study, the effect of different amounts of carbon input on overall health of potato
tubers and roots was assessed. It was observed that treatment with highest quality
and quantity of carbon yielded maximum disease suppression benefits. Potato
tuber yields were significantly enhanced by the accumulative effect of rye cover

crop + poultry manure, compared to a bare control.

Dedicated to my grandmother, parents, sister and husband.

iii

ACKNOWLEDGEMENTS

I would like to thank Dr. Sieglinde Snapp; my thesis advisor, guide and mentor, for
giving me the opportunity to work with her. Her motivation, resolute support and
exemplary guidance have been instrumental in helping me attain my academic goals. I
would also like to express my sincere gratitude to Dr. Mathieu Ngouajio and Dr. William
Kirk for their advice, time and for serving on the graduate committee. A special word of
thanks should go to Kitty O’Neil and Bill Quackenbush for all their valuable inputs,
encouragement and help during the course of my research. I wish to acknowledge the
guidance and direction offered by Dr. Sasha Kravchenko in all statistical analysis and to
Dr. Adrien Ndayegamiye for his valuable input. My coworkers in the laboratory:
Shinsuke, Karen, Courtney, Judith, Edgar and Rich deserve special mention.

This work would not have been possible without the help and support from my family:
my parents, my sister Kavita, and my husband Yogaraj. This work was made possible by
their prayers, support, motivation, and belief. Yogaraj has been instrumental in my entire
thesis. Thank you for all your moral support. My heartfelt gratitude goes out to my
friends who have influenced me and the choices I have made in this endeavor of mine. I
would like to thank Belinda, Mohan, Erin, Robert, Adwait, Priya and all others who have

provided me with their support and help during the past two years.

iv

TABLE OF CONTENTS

 

LIST OF TABLES .......................................................................................................... viii

LIST OF FIGURES ......................................................................................................... xi

Chapter 1. Literature review - -- - -- 1
Introduction ............................................................................................................ 1
Taxonomy and external morphology of potato (Solanum tuberosum L.) .............. 2
History and importance of potatoes ....................................................................... 3
Impact of Rhizoctonia solani on potatoes .............................................................. 4
Increasing the soil organic matter .......................................................................... 6
Organic matter mediated disease suppression ....................................................... 7
Improving soil quality using cover crops ............................................................... 8
Improving soil quality using cover crops ............................................................... 9
Decreasing disease incidence using biofumigation ............................................. 11
Biocidal effect of crucifers ................................................................................... 12
Hypothesis ............................................................................................................ 14
Objectives ............................................................................................................ 14
References cited ................................................................................................... 15

Chapter 2. Effect of Oriental mustard (Brassica juncea L.) residues on potato

 

(Solanum tuberosum L.) - 22
Abstract ............................................................................................................... 22
Introduction ......................................................................................................... 24
Materials and Methods ........................................................................................ 3O

Mustard field experiment ........................................................................... 3O
Plot information ............................................................................. 30
Cover crop details .......................................................................... 30
Inoculation of plots ........................................................................ 31
Incorporation of cover crop residues .............................................. 31
Treatments ...................................................................................... 32
Potato variety and fertilization ....................................................... 32
Potato measurements ..................................................................... 33
Analysis for detection of Pratylenchus penetrans and Verticillium
dahliae ............................................................................................ 35
Experimental design and statistical analysis .................................. 37

Greenhouse container study ....................................................................... 38
Treatments ...................................................................................... 39
Mustard growing conditions .......................................................... 39
Greenhouse conditions ................................................................... 40
Experimental setup ......................................................................... 4O
Inoculation ..................................................................................... 4O
Potato plant growth conditions ...................................................... 42

Monitoring root and tuber disease ................................................. 42

Reisolating fungi from roots .......................................................... 43
Experimental design and statistical analysis .................................. 44

Results .................................................................................................................. 46
Mustard field experiment ........................................................................... 46
Fungal infection of potato roots ..................................................... 46

Dry weight of roots, specific gravity, and yield of tubers ............. 49

Microbial and nematology analysis ............................................... 52

Greenhouse container experiment .............................................................. 6O

% Area of diseased root ................................................................. 6O

Tuber infection scoring .................................................................. 63

Reisolating fungi from roots and tuber .......................................... 65

Discussion ........................................................................................................... 66
Conclusion .......................................................................................................... 68
Future work ......................................................................................................... 69
References cited .................................................................................................. 70

Chapter 3. Bioassay to test suppression of Rhizoctonia solani, Pythium ultimum and

 

F usarium solani by mustard and rye tissue - - - -- - - .......... 75
Abstract ................................................................................................................ 75
Introduction .......................................................................................................... 77
Materials and Methods ......................................................................................... 82

Jar Bioassay ................................................................................................ 82
Plants used and growing conditions ................................................ 82
Pathogen culture .............................................................................. 82
Fungal response to plant ................................................................. 83
Dose response curve ....................................................................... 83
Recording fungal growth ................................................................ 85
Testing biocidal activity of mustard ............................................... 85

Container Bioassay ..................................................................................... 86
Container Specifications ................................................................. 86
Soil and treatments .......................................................................... 86
Tubers ............................................................................................. 87
Procedure ........................................................................................ 87

Replications and Statistical analysis ........................................................... 89

Results .................................................................................................................. 91

Jar Bioassay ................................................................................................ 91
Pythium ultimum ............................................................................. 91
F usarium solani ............................................................................ 95
Rhizoctom'a solam' ......................................................................... 97

Jar bioassay for dose response experiment ............................................... 100
Pythium ultimum ........................................................................... 100
F usarium solani ............................................................................ 100
Rhizoctonia solani ......................................................................... 104

Container tuber bioassay ........................................................................... 106

Discussion .......................................................................................................... l 13

vi

Conclusion ......................................................................................................... l 16

 

Future work ........................................................................................................ 116
References cited ................................................................................................. 117
Chapter 4. Effect of organic fertility amendments on soil fungi _ 120
Abstract .............................................................................................................. 120
Introduction ........................................................................................................ 121
Materials and Methods ....................................................................................... 125
Plot Information ........................................................................................ 125
Treatments ................................................................................................. 125
Inoculation of the plots ............................................................................. 126
Potato variety and fertilization .................................................................. 127
Potato harvest and disease measurements ................................................. 127
Results ................................................................................................................ 130
Infection of potato roots by soil fungi ....................................................... 130
Specific gravity ......................................................................................... 135

Dry weight of roots ................................................................................... 138
Yield .......................................................................................................... 142
Discussion .......................................................................................................... 145
Conclusions ........................................................................................................ 146
Future work ........................................................................................................ 146
References cited ................................................................................................. 147

vii

LIST OF TABLES

Table 2.1: Mustard field experiment. Treatments used in the experiment ....................... 32
Table 2.2: Greenhouse container study. Treatments used in the study ............................. 39
Table 2.3: Mustard field experiment. Analysis of variance for % area of diseased root..47
Table 2.4: Mustard field experiment. Planned contrasts for % of diseased root .............. 47
Table 2.5: Mustard Field Experiment. Analysis of variance for dry weight of roots. ...... 49

Table 2.6: Mustard Field Experiment. Analysis of variance for specific gravity and yield
of tubers ............................................................................................................................ 49

Table 2.7: Mustard field experiment. Least square means and standard errors for dry
weight of root .................................................................................................................... 50

Table 2.8: Mustard Field Experiment. Least square means for organic carbon % in the
treatments .......................................................................................................................... 54

Table 2.9: Mustard Field Experiment. Analysis of variance for substrate induced
respiration ......................................................................................................................... 56

Table 2.10: Mustard field experiment. Planned contrasts for substrate induced respiration
........................ ' 56

Table 2.11: Mustard field experiment. Analysis of variance for Pratylenchus penetrans in
May 2004 and September 2004 ........................................................................................ 58

Table 2.12: Mustard field experiment. Planned contrasts for Pratylenchus penetrans in
May 2004 and September 2004 ........................................................................................ 58

Table 2.13: Mustard Field Experiment. Analysis of variance for Verticillium dahliae in
May 2004 and September 2004 ........................................................................................ 58

Table 2.14: Mustard Field Experiment. Planned contrasts for Verticillr'um dahliae in May
2004 and September 2004 ................................................................................................. 59

Table 2.15: Greenhouse container study. Analysis of variance for % area of diseased root
........................................................................................................................................... 62

Table 2.16: Greenhouse container study. Planned contrasts for % area of diseased root. 62

viii

Table 2.17: Greenhouse container study. Analysis of variance for tuber scoring ............ 64

Table 3.1: Container tuber bioassay. Treatments used for bioassay ................................. 87
Table 3.2: Jar bioassay. Analysis of variance for Pythium ultimum ................................ 94
Table 3.3: Jar bioassay. Planned contrasts at 24 h and 48 h for Pythium ultimum ........... 94
Table 3.4: Jar bioassay. Flamed contrasts at 24 h and 48 h for F usarium solani ............ 95

Table 3.5: Jar bioassay. Least square means with standard errors and analysis of variance
at 24 h, 48 h and 120 h for F usarium solani ..................................................................... 96

Table 3.6: Jar bioassay. Planned contrasts at 24 h and 48 h for Rhizoctom'a solani ......... 97
Table 3.7: Jar bioassay. Analysis of variance at 24 h and 48 h for Rhizoctonia solam'....98

Table 3.8: Jar bioassay. Least square means and analysis of variance for Pythium
ultimum, and F usarium solam' in dose response jar bioassay ......................................... 101

Table 3.9: Jar bioassay. Least square means and analysis of variance for Rhizoctonia

solani in dose response bioassay ...................................................................................... 105
Table 3.10: Container tuber bioassay. Analysis of variance for tuber rating .................. 109
Table 3.11: Container tuber bioassay. Planned contrasts between treatments ................ 109
Table 4.1: Rye with amendments field experiment. Treatments used in the study ......... 126

Table 4.2: Rye with amendments field experiment. Analysis of variance for % of
diseased root ..................................................................................................................... 134

Table 4.3: Rye with amendments field experiment. Planned contrasts for % of diseased
root ................................................................................................................................... 134

Table 4.4: Rye with amendments field experiment. Analysis of variance for specific
gravity of tubers ............................................................................................................... 137

Table 4.5: Rye with amendments field experiment. Planned contrasts for specific gravity
of tubers. .......................................................................................................................... 137

Table 4.6: Rye with amendments field experiment. Analysis of variance for dry weight of
roots .................................................................................................................................. 141

Table 4.7: Rye with amendments field experiment. Planned contrasts for dry weight of
roots .................................................................................................................................. 141

ix

Table 4.8: Rye with amendments field experiment. Analysis of variance for yield ........ 144

Table 4.9: Rye with amendments field experiment. Planned contrasts for tuber yield 144

LIST OF FIGURES

Figure 2.1: Scale used by WinRHIZOTM for root infection: (a) 0-25 % infection, (b) 25-
50 % infection, (c) 50-75 % infection, (d) 75-100 % infection ........................................ 36

Figure 2.2: Experimental setup for greenhouse container study ....................................... 41

Figure 2.3: Greenhouse container study. Experimental set-up in greenhouse showing
containers with minirhizotron tubes .................................................................................. 41

Figure 2.4: Greenhouse container study. Scale used for rating tubers on basis of %
infection: (a) 0-25 % infection, (b) 25-50 % infection, (c) 50-75 % infection, (d) 75-100

% infection ........................................................................................................................ 43
Figure 2.5: Mustard field experiment. % area of diseased root ........................................ 48
Figure 2.6: Mustard field experiment. Specific gravity for different treatments .............. 50
Figure 2.7: Mustard field experiment. Yield of fresh tubers in g / plant .......................... 50
Figure 2.8: Mustard Field experiment. Substrate induced respiration in mg COz/kg/h
between 0 - 10 days .......................................................................................................... 55
Figure 2.9: Mustard Field experiment. Substrate induced respiration in mg C02/kg/h
between10-20 days ............................................................................................................ 55
Figure 2.10: Concentration of Pratylenchus penetrans spores/ 100 cm3 soil observed in
May 2004 .......................................................................................................................... 57
Figure 2.11: Concentration of Pratylenchus penetrans spores/ 100 cm3 soil observed in
September 2004 ................................................................................................................ 57
Figure 2.12: Greenhouse container study. % Area of diseased root ................................ 61
Figure 2.13: Greenhouse container study. % Area of infected tubers ............................. 63
Figure 2.14: Re-isolation of Rhizoctonia solani from potato: (a) potato tuber, (b) potato
root .................................................................................................................................... 65
Figure 3.1: Jar bioassay. Experimental set up for bioassay .............................................. 84

Figure 3.2: Jar bioassay. Experimental set up for dose response of fungi to mustard tissue:
(a) 10 g mustard shoot, (b) 5 g mustard shoot, (c) 1 g mustard shoot, ((1) bare ................ 84

xi

Figure 3.3: Container tuber bioassay. Experimental set up for container tuber bioassay:
(1) container used for experiment (2) randomized complete block design in greenhouse

........................................................................................................................................... 88
Figure 3.4: Container tuber bioassay. Scale used for rating tubers .................................. 89
Figure 3.5: Jar bioassay. Growth of Pythium ultimum observed at 24 h and 48 h ........... 92

Figure 3.6: Jar bioassay. Growth of Pythium ultimum on petri dish: (1a) mustard root +
shoot at 24 h, (1b) bare at 24 h, (2a) mustard root + shoot at 48 h, (2b) bare at 48 h ....... 93

Figure 3.7: Jar bioassay. Growth of Rhizoctonia solam' on petri plate observed at 24 h and

48 h. Fungal growth expressed as % area of growth on plate ........................................... 99
Figure 3.8: Dose response of Pythium ultimum to mustard tissue over time ................... 102
Figure 3.9: Dose response of F usarium solani to mustard tissue over time .................... 103

Figure 3.10: Container tuber bioassay. Comparisons between tubers from 2 treatments. a)
tubers in soil with mustard root + shoot as treatment b) Tubers in fumigated soil ......... 107

Figure 3.11: Container tuber bioassay. Tubers showing effect of different treatments: (a)
mustard root + shoot, (b) rye root + shoot, (c) mustard shoot, ((1) fumigated, (e) sterile
soil, (f) Pythium ultimum inoculated soil ......................................................................... 108

Figure 3.12: Container tuber bioassay. Comparison between treatment with rye root +
shoot at a) & days after incorporation of cover crop b) 40 days after incorporation of
cover crop ......................................................................................................................... 110

Figure 3.13: Container tuber bioassay. Tuber Infection for different treatments ............ 11 1

Figure 3.14: Container tuber bioassay. Comparison between tubers placed in inoculated
soil and tubers placed in sterile soil: (a) soil inoculated with Pythium ultimum, (b) soil
inoculated with F usarium solani, (c) soil inoculated with Rhizoctonia solani, (d) sterile
autoclaved soil ................................................................................................................. 112

Figure 4.1: Roots scanned by WinRHIZOTM showing different levels of infection: (a) 0-
25 % infected, (b) 25-50 % infected, (c) 50-75 % infected, ((1) 75-100 % infected ........ 128

Figure 4. 2: Rye with amendments field experiment. Scale used for rating tubers of basis
of % infection: (a) 0- 25 % infection, (b) 25- 50 % infection, (c) 50- 75 % infection, (d) 75-
100 % infection ................................................................................................................ 129

Figure 4.3: Rye with amendments field experiment. % of diseased root for each treatment
in 2003 at 2 harvests: (a) harvest 1, (b) harvest 2 ............................................................ 132

xii

Figure 4.4: Rye with amendments field experiment. % of diseased root for each treatment

in 2004 at 2 harvests: (a) harvest 1, (b) harvest 2 ............................................................ 133
Figure 4.5: Rye with amendments field experiment. Specific gravity for 2003 ............. 136
Figure 4.6: Rye with amendments field experiment. Specific gravity for'2004 .............. 136

Figure 4.7: Rye with amendments field experiment. Dry weight of roots in 2003 at 2
harvests: (a) harvest 1, (b) harvest 2 ................................................................................ 139

Figure 4.8: Rye with amendments field experiment. Dry weight of roots in 2004 at 2
harvests: (a) harvest 1, (b) harvest 2 ................................................................................ 140

Figure 4.9: Rye with amendments field experiment. Yield of fresh tubers in g/plant in
2003 .................................................................................................................................. 143

Figure 4.10: Rye with amendments field experiment. Yield of fresh tubers in g/plant in
2004 .................................................................................................................................. 143

xiii

Chapter 1. Literature Review

Introduction

Vegetables are grown on almost 1.5 million hectares in the US with a total
value of over US$ 9 billion (USDA, 1998). Unfortunately there are many factors
including several soil borne pests and diseases of vegetables that are known to negatively
and substantially affect both the yield and the quality of vegetables (Abawi et al., 2000).
Crop losses caused by plant pests and diseases run into millions of dollars worldwide.
Soil organic matter consists of living, partially to fully decomposed organic materials and
is an important indicator of soil health.

Soil health can be considered a subset of ecosystem health. A healthy ecosystem
is characterized by integrity of nutrient cycles and energy flows, stability, and resilience
to disturbance or stress (O’Neill et al., 1986). Thus, soil health may be associated with
biological diversity and stability. Plant and animal disease outbreaks can be considered as
indicators of instability and poor ecosystem health. Therefore, there is likely also a link
between soil organic matter, the ability of the biological community to suppress plant
pathogens, the p0pulation density of plant pathogens in soil, and ultimately disease
incidence and severity (van Bruggen and Grunwald, 1996). Organic matter is one of the
most important components of soil (Magdoff, 1992). Many biological, physical, and
chemical properties of soils are a function of total soil organic matter. Organic matter
confers many benefits to ecosystems including increasing nutrient availability to plants,
providing a favorable physical condition for plant growth, increasing soil buffering
capacity, stimulating root development, increasing biological diversity, and facilitating a

number of global cycles such as carbon and nitrogen. Current farming practices fail to

replenish the organic matter in the soil after harvest and constitute a very important
problem.

Soil organic matter serves as a primary indicator of soil quality and health for
both scientists and farmers (Romig et al., 1995). For centuries, farmers have consciously
and unwittingly manipulated the ecology of soil by addition or depletion of organic
matter (Bailey et al., 2003). Production changes in the last 50 years have been detrimental
to soil health and water quality, thereby increasing plant diseases and other pest
problems, all within a relatively short period of time (Pimental et al., 1991). Growers
generally respond to the threat of soil borne diseases with generous application of
pesticides. The wide use of pesticides in agriculture has resulted in grower concern about
the environmental impacts and there are also concerns about becoming dependent on one
pesticide

In view of these facts, efforts are now aimed at promoting more sustainable
methods of agriculture. In this study, we focus on two cover crops (Cereal rye and
Oriental mustard) to help improve quality and yield of the subsequent crop (Potato).
Cereal is a widely used winter cover crop by Michigan growers. Cereal rye was chosen
keeping in mind that it produces large quantities of biomass consequently adding more
organic matter to soil. Oriental mustard was selected for its inherent biofumigant
properties.

Taxonomy and External Morphology of Potato

The potato (Solanum tuberosum L.) belongs to the family Solanaceae, with about

90 genera and 2,800 species. Though the family is found throughout the world, it is

especially concentrated in the tropical regions of Latin America (Cornell, 1962). The

potato and all its wild relatives belong to the genus Solanum, which consists of about
2,000 species. The section Tuberariurn (Cornell, 1962), also known as section Petota
(D’Arcy, 1972) classified within this genus includes the tuber-bearing members, of which
the cultivated potato is best known.

The cultivated potato is herbaceous and ranges from 0.5-1.0 meter in height. The
leaves are alternate and irregularly pinnately compound. The inflorescence consists of
several flowers which are pentamerous, actinomorphic, perfect, and have sympetalous
corollas. The fruits are bicarpellate berries and can be absent in many cultivars (Burbank,
1921). Tubers are formed underground from stolons, from which adventitious roots are
developed into a fibrous mass (Burton, 1969).

History and Importance of Potato

The potato originated in South America, specifically in the Andean highlands of
Bolivia, Ecuador and Peru. It was first domesticated almost 6,000 years ago in the area
around Lake Titicaca (Burton, 1948). They were an important source of starch and
carbohydrates and could be stored and transported easily. The Spanish explorers of the
1500’s were the first Europeans to come in contact with the potato. Native Andean
potatoes are cultivated even today and display an extraordinary diversity of taste and
texture.

Today the potato is the world's fourth most important food crop and by far the
most important vegetable. Potatoes are cultivated worldwide on acreage of 17.9 million
hectares. Potatoes are mainly grown commercially in the United States as row crops in
monoculture. Tubers of the selected cultivar are cut into seed pieces, planted, treated with

fertilizer, pesticide, and mechanically harvested (Burton, 1969). Each year about 7

percent of tubers are used as seed; about 48 percent are processed; and about 30 percent
are used as fresh vegetable. In volume and sales dollars, potatoes are Michigan’s leading
produce commodity.

Studies have estimated losses due to plant disease in the four major crops (rice,
wheat, potato and maize) to be between 10 and 16% of potential production which
translates into about. 64 x 109 US$ over the years 1988-90. Despite its global dominance,
the potato suffers tremendous losses due to disease that translate into billions of dollars in
wasted resources and lost sales both in the US and abroad every year.

Impact of Rhizoctonia solani on potatoes

Soil borne pests and diseases are a serious problem to a potato grower. They are
difficult to detect as they exist out of sight and are difficult to measure. Low populations
causing extensive damage (e.g. soil insects), their microscopic size, (e.g. fungal
pathogens, nematodes) and because of resting stages that differ from the active stage (e. g.
fungal pathogens), detection of these pathogens is challenging (Matthiessen et al.,
University of Idaho, Moscow, Idaho, USA). Growers generally respond to the threat of
soil borne diseases with generous application of pesticides such as firmigants e.g. Vapam.
All economically important plants are damaged by one or several diseases that can
greatly limit the yield potential and the quality of the harvested produce (Agrios 1988).
Rhizoctonia disease of potato tubers and roots, often referred to as black scurf is caused
by the fungus Rhizoctonia solani Kuhn (Tsror et al., 2001). It is distributed worldwide,
has physiological strains, has a wide host range, and causes different symptoms on the
same host depending on the time of infection (Lewis et al., 2001). R.solani infects potato

sprouts during the emergence period. Symptoms include delayed emergence, stem canker

and lesion expansion and girdling can seriously debilitate young plants. R. solani attacks
tubers, underground stems, and stolons of potato plants. Early season stress caused by R.
solani can lead to decrease in stem number, decrease in tuber set, reduction in tuber size
and reduced tuber quality.

In temperate production areas, losses due to R. solani are sporadic and occur only
when weather is cold and wet in the weeks following planting. In northern areas, where
growers often must plant in cold soils, R. solani is a more consistent problem. Poor
stands, stunted plants, reduced tuber number and size, and misshapen tubers are
characteristic of the Rhizoctonia disease. R. solani commonly survives on seed tubers but
can also survive and grow on alternate crops. The fungus exists on potato in three
distinct phases. One is parasitic on the potato plant and the other two phases are not
pathogenic as they exist on the potato and provide inoculum capable of later infection.
These two phases are: (1) Black sclerotia present principally on tubers, which serve to
carry the fungus over the winter and on which no spores are produced; and (2) White
superficial myceliurn present on the lower stems which represents the asexual stage of the
fimgus. When R. solani reproduces asexually by means of mycelia and sclerotia, it is
called T hanatephorous cucumeris. Even with the use of seed treatment fungicides and
fumigants, crop loss due to damping-off ranges from 5-10% in the US alone, which still
accounts for a loss of millions of dollars (Gilpatrick, 1979). It is a common occurrence in
most potato producing areas of the world. The saprophytic survival capability of R. solani
in debris of specific crops (Specht and Leach, 1987) and the pathogenicity of specific
strains of R. solani selected for by various preceding crops (Johnston et al., 1994) also

influence the composition of soil populations pathogenic to potatoes and consequently

disease severity in succeeding potato crops. The importance of the Rhizoctonia disease
complex of potatoes has long been debated. It is now considered a major cause of crop
losses in Maine and other potato-producing areas (Johnston et al., 1994).

Soil fumigants should reach pathogens in all physical and biological niches in the
soil. As a result, however, soil fumigants often lead to the eradication of beneficial
organisms, and may bring about a negative shift in the biological equilibrium. This
creates a microbial vacutun, which may lead to the population of pathogens increasing
and causing even more damage than those originally targeted for control (Gamliel et al.,
2000). Microbial diversity in soil is normally assessed as species or genetic diversity
rather than structural and functional diversity. However, these last two measures of
diversity may be more relevant to soil health (V isser and Parkinson, 1992). Soils,
especially those with a low microbial p0pulation are more vulnerable to reinvasion of
pathogens following fumigation. Thus, non-chemical methods of effectively controlling
soilbome diseases are needed. In view of these observations, a number of strategies have
been proposed to combat these potato diseases.

Increasing the Soil Organic Matter

Soil organic matter (SOM) is known to affect soil aeration, structure, drainage,
moisture holding capacity, nutrient availability, and microbial ecology (Davey, 1996).
SOM is important for enhancing water holding capacity of soils; supplies nutrients,
which are crucial for crop production; for protection against erosion; and helps support a
healthy and diverse set of microscopic plants and animals (The State of the Nations
Ecosystem, 2002 report). The amount of carbon (C) input into the soil from crop residues

generally increases SOM (Peterson et al., 1998; Hendrix et al., 1998). SOM stabilizes soil

pH, which plays a central role in nutrient supply and availability for plant uptake
(Campbell et al., 1996). Lack of residues can result in low SOM. In a study by
Nyakatawa et al., 2001, winter rye cover cropping rapidly increased surface SOM. This
increase was attributed to the large (quantities of residues produced by the winter rye
cover crop. Short term benefits of increased surface SOM such as improved soil water
conservation, seedling establishment, crop growth, and yield were clearly visible in this
study.
Organic matter mediated disease suppression

Suppression of soil-borne diseases can be achieved by addition of organic
residues in cropping systems. There are reports of organic amendment-mediated soil
disease suppression since as early as the nineteenth century. In a study by Tepper, 1892,
manure applications were used to control take-all of wheat. Root rot of cotton caused by
Phymatotrichum omnivorum was reduced when the soil was amended with manure
(Pammel, 1980). Considerable progress has been made in the utilization of organic
materials as soil amendments for the control of plant-parasitic nematodes (Singh and
Sitaramaiah, 1970; Muller and Gooch, 1982; Akhtar, 1993). Composting can be used as
an effective and desirable soil organic amendment (Hoitink, 1986; Dick and McCoy,
1993). In addition to increasing organic matter of the soil, amending with composts also
increases soil microbial populations (Pera et al., 1983; Perucci, 1990), which improves
the soil quality. The suppressive activity of compost towards plant pathogens has been
well documented with the majority of success shown in containerized systems (Hoitink,
1986; Nelson and Craft, 1992). In cropping systems amended with organic matter,

suppression occurs because of activation of the indigenous soil microbial community.

Improving soil quality using cover crops

A cover crop is a crop grown as a complementary plant to a cash crop to benefit
the soil in many ways: it can in some circumstances reduce erosion, weed pressure,
insects, plant parasitic nematodes, other pest problems, and can improve soil quality
(Mutch and Martin, 2000). The Soil Science Society of America (1987) defined cover
crops as those used as green manures incorporated into the soil while green or at
maturity, for soil improvement and protection.

Effective nitrogen scavenging fall-sown cover crops should exhibit rapid
germination, aggressive and extensive rooting systems (Sainju et al., 1998), good winter
hardiness, and early spring regrowth (Weinert et al., 2002). Winter cover crops can
improve N cycling and reduce the amount of N below the root zone as shown in a study
of potato based rotations (Weinert et al., 2002). Winter cover crops reduce the potential
for nitrate (N03) leaching by (i) absorbing and storing N in plant tissue during leaching
prone winter months during soil water recharge and (ii) absorbing and transpiring water,
lessening water percolation (Weinert et al., 2002). Cereal and brassica crops display these
characteristics and are well suited for winter cover cropping (Wagger and Mengel, 1988;
Brinsfield and Staver, 1991). These cover crops can accumulate up to 150 kg N ha'1
(Hoyt and Mikkelsen, 1991; Sherman, 1992; Ditsch et al., 1993), with rooting systems
reaching depths of between 80 and 150 cm (Frye et al., 1985; Sarrantonio, 1992).

Green manures and cover crops have been shown to differ significantly in their
suppression of root rot severity and damage to plant growth, as indicated by greenhouse
studies. (Abawi et al., 2000). In addition, differential effects of various cover crops on the

severity of root rot and yield have been observed under field conditions (Abawi et al.,

2000). A number of cover crops and green manures can be effective in suppressing
nematode populations and infections (Halbrendt, 1996; Mojtahedi et al., 1991; Mojtahedi
et al., 1993). Organic amendments, such as incorporation of green plant residues in the
field or compost added to potting mix, can suppress the activity of R. solani (Gorodecki
and Hadar, 1990; Papavizas and Davey, 1960; Tuitert etal., 1998).

Rye has been effectively used as a winter cover crop as it provides sufficient
ground cover to prevent wind erosion. Besides being convenient to grow, rye has a
profusely branched root system which prevents soil compaction in soils that are annually
tilled. The extensive root system also enables it to scavenge nutrients efficiently from the
soil profile. The growth habit of common rye is very competitive. In a study by Creamer
et al., 1997, rye comprised of at least 80% of the above ground biomass just before
desiccation. In a study by Ranells et al., 1997, rye monoculture recovered 39% of the
labeled 15N fertilizer as compared to 19% by the rye-crimson clover biculture and 4% by
the crimson clover monoculture. This study confirmed the superior ability of rye, in
monoculture and biculture, to scavenge residual soil N compared with crimson clover.
Impact of crop residue on Rhizoctonia solani

Plant residues left of or near the soil surface may contribute to suppression of
soilbome pathogens in minimum tillage systems. Disease suppression can be induced by
organic and fertilizer amendments (Baker and Chet, 1982) or practices such as mulching
which stimulates aggressive competition among soil inhabitants in the root zone (Baker
and Cook, 1974; Boosalis et al., 1981; Tu and Findlay, 1986). Organic amendments, such
as the incorporation of green plant residues in the field or compost added to potting mix,

have been shown to suppress the activity of R. solani. (Gorodecki and Hadar, 1990;

Papavizas and Davey, 1960; Tuitert et al., 1998). This suppression has been correlated
with increased antagonistic soil microbial activity (Tuitert et al., 1998), and sterilization
of composts or suppressive field soils has eliminated the suppressive effects (Wiseman et
al., 1996; Gorodecki and Hadar, 1990; Kobayashi and Ko, 1985).

Sustainability of farming systems using short term rotations has become a major
concern in potato production. Research with potato cropping systems has shown that the
frequency and host range of crops in a potato rotation influences potato soilbome disease
incidence and development (Hide and Read, 1991; Honeycutt et al., 1996; Specht and
Leach, 1987). Continuous potato production schemes have been shown to result in 58%
of plants developing stern lesions caused by R. solani, as compared to 12~22% of stems
from potatoes grown with other rotation crops (Honeycutt et al., 1996). Longer rotations
(out of potatoes) are associated with reduced soil inoculum levels of R. solani as
compared to short rotations (Scholte, 1987); the preceding crop and its relative ability to
harbor potato pathogens will also affect the incidence of potato diseases in the succeeding
crops. Thus, when buckwheat is grown prior to potatoes, greater soil population densities
of R. solani and increased stem canker in potatoes developed (Specht and Leach, 1987).
However, oat—potato ( Frank and Murphy, 1977), annual ryegrass—potato (Johnston et al.,
date; Specht and Leach, 1987) or clover—potato (Johnston et al., 1994) sequences were
found to reduce both R. solani inoculum levels in soil and subsequent disease
development in the potato crop.

In soil, the success or failure in the struggle for survival by R. solani is affected by
physical and biological factors. Along with quantity, type and availability of energy

materials, volatiles released as decomposition products from plants considerable affect

10

survival of the pathogen. Plant residues which are low in their C:N ratio when
decomposed in soil result in the release of volatiles which causes pigmentation of R.
solani in culture and reduces survival and saprophytic activity of the pathogen in soil
(Lewis and Papavizas, 1974).
Decreasing disease incidence using Biofumigation

Despite the wide use of pesticides in agriculture, growers are now beginning to
become concerned about environmental impacts and there are also concerns about
becoming dependent on one pesticide. ‘Biofumigation’, using plants that produce toxic
compounds to exert control over pests and diseases in soil, offers a biological alternative
to the use of chemical pesticides. Since the late 19th century, there have been reports of
plant extracts showing biocidal activity. Biofumigation refers to the use of plants as
rotation crops that contain biologically active compounds or green manures that suppress
soil-home pests and diseases in agricultural production systems. Biofumigation offers a
biological alternative for producing fumigant-like chemicals in the soil, providing an
option for suppressing soil-borne pests and diseases, and helping promote soil health.
This technique exploits a plant defensive enzymatic system which is common to some
plant families such as Brassicaceae, Capparidaceae, Moringaceae etc. Over the past
decade, several authors have reported that many compounds extracted from wild,
cultivated, and medicinal plants show fungitoxic activity (Gourinath and Manoharachari,
1991; Rahalison et al., 1993; Yegen et al., 1992).

Glucosinolates are a class of naturally occurring ionic compounds found in plants
usually as the potassium or sodium salt. They have been the subject of much attention

because of their involvement in biofumigation. In biofumigation, plant based

11

glucosinolates are hydrolyzed in field soil to form toxic products like isothiocyanates,
thiocyanates, nitriles, oxazolidinethiones, and ionic thiocyanate (Brown and Morra,
1997). These degradation products may exert a suppressive or controlling effect on a
wide range of soil-borne plant pathogens including R. solani (Lewis and Papavizas,
1974)
Biocidal effect of crucifers

Crucifer tissues have long been known to produce biocidal effects following their
addition to the soil (Angus et al., 1994). Crucifers such as oriental mustard emit volatiles
that inhibit growth of undesirable soil fungi. Field experiments during the early 1990's
demonstrated that wheat crops grew more vigorously following Brassica break crops
such as canola and Indian mustard than other break crops such as linseed or oats (Angus
et al. 1991, Kirkegaard et a1. 1994). Glucosinolate-containing plants in the Brassicaceae
family represent a potential source of allelochemical control for a variety of soil-borne
pests (Fahey et al., 2001). Walker et al., (1937) observed antifungal activity of mustard
oils; Hooker et al., (1943) confirmed the antifungal activity of cruciferous plant extracts
containing allyl and phenethyl isothiocyanates (ITC). Glucosinolates are precursors of
ITC’s which are detrimental to many soil-borne organisms. ITC are released when plants
are physically broken up, and during the break down of residues in the soil. Compounds
from the Cruciferae are used as potentially valid alternatives to controlling soil-borne
pathogens (Luisa et al., 1997). This observed pest suppression is assumed to result from
ITC and possibly other allelochemicals produced from glucosinolates within the tissues
(Morra et al., 2002). Incorporation of Brassica tissues into the soil reduces the fungal

population in the soil. Suppression of Pythium ultimum population in soil observed

12

during the first weeks after incorporation into soil could prevent the rapid increase of P.
ultimum, thus reducing the period in which this pathogen can cause damping-off or root
rot of seedlings and young plants (Lazzeri et al., 2001). Oriental mustard (Brassica
juncea L.), known commercially as Pacific Gold, gave the best combination of traits:
medium height and biomass production, high glucosinolate level, and late flowering
(Bryant, 2003). The oriental mustard soil incorporation treatment, produced 18 % more
marketable fruit yield than non-amended control plots (Harvey et al., 2001). Different
plant species in the Brassicaceae family have different levels of glucosinolate. Plant
tissues exhibit varied responses on fungal colonies depending on the amount of
glucosinolate present in them. A study by Harding et al., (2001) showed that most
Brassica plant materials suppressed fungal growth, but the response varied between plant
parts and fungal species. For example B. juncea meal completely inhibited growth of all
fungi whereas leaf tissue inhibited fungi between 20 — 50%. Studies have demonstrated
the ability of B. juncea to reduce incidence of R. solani. In principle, the utilization of
ITC of glucosinolates may be a good compromise between fungitoxic activity toward

some phytopathogenic fungi and environmental protection.

13

Objectives

The overall objective of this study was to improve quality of potato roots and
tubers and reduce disease incidence using cover crops and other soil amendments.
The specific objectives were;
(1) To study the biocidal effect of Brassica juncea root v/s shoot on the incidence of
Rhizoctonia solani on potato tubers and roots.
(2) To evaluate the dose response of Brassicajuncea (oriental mustard) on R. solani
(3) Noting the overall effect of soil amended with Secale cereale (cereal rye) on potato
tuber yield, quality and root rot disease.

(4) To see influence of soil with different organic amendments on disease.

Overall hypothesis are; (1) Organic amendments reduce disease severity of potato
plants compared to a bare fallow without organic inputs. (2) Organic amendments will
enhance potato health as much as a fumigated control. (3) Organic amendment benefits
will be related to the properties of the cover crop incorporated and also to the quantity of
carbon added. (4) The combination of poultry manure plus rye has the most diverse
substrates and will be associated with the greatest health benefits compared to either

applied alone.

Root-health hypothesis are; (1) Organic amendments enhance root health in
potato plants compared to a bare fallow without organic inputs. (2) Organic amendments
will enhance root health as much as fumigated control. (3) Diversity of substrate will

determine the root health benefit.

14

REFERENCES CITED

Abawi, G. S., and T. L. Widmer. 2000. Impact of soil health management practices on
soilbome pathogens, nematodes and root diseases of vegetable crops. Appl Soil
Ecol 15 (1):37-47.

Agrios, G.N. 1988. Plant Pathology. Academic Press, New York, pp. 803

Akhtar, M. 1993. Utilisation of plant-origin waste materials for the control of parasitic
nematodes. Bioresource Technology 46: 255—257.

Angus, J. F., A. F. Van Herwaarden and G. N. Howe. 1991. Productivity and break crop
effects of winter-growing oilseeds. Aust.J.Exp.Agric. 31 :669—677.

Angus, J. F., P. A. Gardner, J. A. Kirkegaard, and J. M. Desmarchelier. 1994.
Biofumigation: isothiocyanates released from Brassica roots inhibit growth of the
take-all fungus. Plant and Soil 162:107—112.

Bailey, K. L., and G. Lazarovits. 2003. Suppressing soil-borne diseases with residue
management and organic amendments. Soil and Tillage Research 72(2): 169-180

Baker , K. F., and R. J. Cook. 1974. Biological control of plant pathogens. Freeman, San
Frisisco, CA, pp. 433.

Baker, R. F., and I. Chet. 1982. Induction of suppressiveness. In: Schneider, R.W. (Ed.),
Suppressive Soils and Plant Disease. American Phytopathological Society, St.
Paul, MN, pp. 35-50.

Boosalis, M. G., B. Doupnik, and G. N. Odvody. 1981. Conservation tillage in relation to
plant diseases. In: Piementel, D. (Ed.), Handbook of Pest Management in
Agriculture, Vol. 1. CRC Press, Boca Raton, FL, pp. 445—474.

Brinsfield, R. B., and K. W. Staver. 1991. Use of cereal grain cover crops for reducing
groundwater nitrate contamination in the Chesapeake Bay region. In W.L.
Hargrove (Ed.) Cover crops for clean water. Proc. Int. Conf., West Tennessee
Exp. Stn., Jackson. Soil and Water Conserv. Soc., Ankeny, IA, pp. 79—82.

Brown, P. D., and M. J. Morra. 1997. Control of soil-borne plant pests using
glucosinolate containing plants. Advances in Agronomy 61 :167-23.

Bryant, D., 2003. Biofumigation monitored with Brassica cover crops. Western Farm
Press. Viewed on 26th February 2004.
http://westemfarmpress.com/ar/farming_biofumigation_monitored_brassica/index
.htrn

15

Burbank, L. 1921. How Plants are Trained to Work for Man: Grafting and Budding, Vol
2. Collier, New York, pp. 352.

Burton, W. G. 1948. The Potato. lst ed. Butler and Tanner Ltd. London.

Burton, W. G. 1969. Potato. In: Encyclopaedia Britannica, Vol 18, pp. 95-134. Benton,
Chicago et alibi., pp.1197.

Campbell, C. A., B. G. McConkey, R. P. Zentner, F. Selles, and D. Curtin. 1996. Long-
tenn effects of tillage and crop rotations on soil organic C and total N in a clay
soil in southwestern Saskatchewan. Canadian J. Soil Sci. 76:395—401.

Correll, D. S. 1962. The Potato and Its Wild Relatives: Section Tuberarium of the Genus

Solanum, Correll, D. S. (Ed.). Texas Research Foundation. Renner, Texas. pp.
344-352.

Creamer, N. G., M. A. Bennett, and B. R. Stinner. 1997. Evaluation of cover crop
mixtures for use in vegetable production systems. HortScience 32:866—870.

D'Arcy, W. G. 1972. Solanaceae studies 11. Typification of subdivisions in Solanum.
Annals of The Missouri Botanical Garden. 59:262-278.

Davey, C. B.1996. Nursery soil management—organic amendments. In: Landis, T. D.,
Douth, D.B. (Tech. Coordinators), National Proceedings, Forest and Conservation
Nursery Associations, pp. 6—18.

Dick, W. A., and E. L. McCoy. 1993. Enhancing soil fertility by addition of compost. In:
Hoitink, H.A.J., Keener, H.M. (Eds), Science and Engineering of Composting:
Design, Environmental, Microbiological and Utilization Aspects. Renaissance
Publications. Worthington, OH, pp. 622-644.

Ditsch, D. C., M. M. Alley, K. R. Kelly, and Y. Z. Lei. 1993. Effectiveness of winter rye
for accumulating residual fertilizer N following corn. J. Soil Water Conserv.
48:125—132.

Fahey, J. W., A. T. Zalcmann, and P. Talalay. 2001. The chemical diversity and
distribution of glucosinolates and isothiocyanates among plants. Phytochemistry
56:5—51.

Frank, J. A., and H. J. Murphy. 1977. The effect of crop rotation on Rhizoctonia diseases
of potatoes. Am. Potato J. 54:315—322.

Frye, W. W., W. G. Smith, and R. J. Williams. 1985. Economics of winter cover crops as
a source of nitrogen in no-till corn. J. Soil Water Conserv. 40:246—249.

16

Gamliel, A., M. Austerweil, and G. Kritzrnan. 2000. Non-chemical approach to soilbome
pest management-organic amendments. Crop Prot. 19:847—853.

Gilpatrick, J. D. (Ed.), 1979. Contemporary Control of Plant Diseases with Chemicals.
American Phytopathological Society, St. Paul.

Gorodecki, B., and Y. Hadar, 1990. Suppression of Rhizoctonia solani and Sclerotium
rolfivii in container media containing composted separated cattle manure and
composted grape marc. Crop Prot. 92271-274.

Halbrendt, J. M. 1996. Allelopathy in the management of plant-parasitic nematodes.
Journal of Nematology 28:8-14.

Harding, R. and T. Wicks. 2001. In vitro suppression of mycelial growth of potato
pathogens by volatiles released by Brassicae residues.
http://www.sardi.sagov.au/pages/horticulture/patholggv/hort pn_invitrobrassica.
htm:sectID=337&tempID=1 1 1

 

Hendrix, P. F ., A. J. Franzluebbers, and D.V. McCracken. 1998. Management effects on
C accumulation and loss in soils of the southern Appalachian Piedmont of
Georgia. Soil Till. Res. 47:245—251.

Hide, G. A., and P. J. Read. 1991. Effects of rotation length, fungicide treatment of seed
tubers and nematicide on diseases and the quality of potato tubers. Ann. Appl.
Biol. 119: 77—87.

Hoitink, H. A. J. and P. C. Fahy. 1986. Basis for the control of soilbome plant pathogens
with composts. Annu. Rev. Phytopathol. 24:93—114.

Honeycutt, C. W., W. M. Clapharn, and S. S. Leach. 1996. Crop rotation and N
fertilization effects on growth, yield, and disease incidence in potato. Am. Potato
J. 73:45—61.

Hooker, W. J., J. C. Walker, and F. G. Smith. 1943. Toxicity of beta phenyl
isothiocyanate to certain fungi. American Journal of Botany. 30:632-637.

Hoyt, G. D., and R. L. Mikkelsen. 1991. Soil nitrogen movement under winter cover
crops and residues.. In W.L. Hargrove (Ed.) Cover crops for clean water. Proc.
Int. Conf., West Tennesse Exp. Stn., Jackson. 9—11 Apr. 1991. Soil and Water
Conserv. Soc., Ankeny, IA, pp. 91—94.

Johnston, H. W., M. J. Celetti, J. Kimpinski, and H. W. Platt. 1994. Fungal pathogens and
Pratylenchus penetrans associated with preceding crops of clovers, winter wheat,
and annual ryegrass and their influence on succeeding potato crops on Prince
Edward Island. Am. Potato J. 71 :797—808.

17

Kirkegaard, J. A., P. A. Gardner, J. F. Angus, E. Koetz. 1994. Effect of Brassica break
crops on the grth and yield of wheat. Aust. J. Agric. Res. 45:529—545.

Kobayashi, N., and W. H. Ko. 1985. Nature of suppression of Rhizoctonia solani in
Hawaiian soils. Br. Mycol. Soc. 84:691—694.

Lazzeri, L., and L. M. Manici. 2001. Allelopathic effect of glucosinolate containing
plant green manure on Pythium sp. and total fungal p0pulation in soil.
HortScience. 37(7):1283-1289.

Lewis, J. A., and G. C. Papavizas. 1974. Effect of volatiles from decomposing plant
tissues of pigmentation, growth, and survival of Rhizoctonia solani. Soil Science.
18(3): 156-163.

Lewis, J. A., and R. D. Lumsden. 2001. Biocontrol of damping-off of greenhouse-grown
crops caused by Rhizoctonia solani with a formulation of Trichoderma spp. Crop
Prot. 20(1): 49-56.

Magdoff, F., 1992. Building Soils for Better Crops: Organic Matter Management.
University of Nebraska Press, Lincoln, NE, pp 176.

Matthiessen, J ., J. Kirkegaard, and M. Morra. Biofumigation for Soil-home Pest and
Disease Suppression - Current Status and Future Directions. Viewed on 8th
F ebruary2004.
http://www.sardi.sa.gov.au/pages/horticulture/pathology/hort_pn_biosoilstatus.ht
m:sectID=336&tempID=l 1 1

Mojtahedi, H., G. S. Santo, A. N. Hang, and J. H. Wilson. 1991. Suppression of root-knot
nematode populations with selected rapeseed cultivars as green manure. J.
Nematol. 23:170-174.

Mojtahedi, H., G. S. Santo, and R. E. Ingharn. 1993. Suppression of Meloidogyne
chitwoodi with sudan grass cultivars as green manure. Journal of Nematology 25:
303-311.

Muller, R., and P. S. Gooch. 1982. Organic amendments in nematode control. An
examination of the literature. NematrOpica 12: 3 1 9—3 26.

Mutch, D. R., and T. E. Martin. 2000. Cover crops. In Michigan field crop ecology.
Michigan State University. Extension bulletin. E-2646, pp 44-53.

Nelson, E. B., and C. M. Craft. 1992. Suppression of dollar spot on creeping bentgrass

and annual bluegrass turf with compost-amended topdressing. Plant Dis. 76:954—-
958.

18

Nyakatawa, E. 2., K. C. Reddy, and K. R. Sistani. 2001. Tillage, cover cropping, and

poultry litter effects on selected soil chemical properties. Soil Tillage Res. 58:69—
74

O’Neill, R. V., D. L. DeAngeles, J. B. Waide, T. F. H. Allen. 1986. A hierarchical
concept of ecosystems. Princeton University Press, Princeton, NJ, pp 263.

Pammel. L. H. 1980. Cotton root-rot. Texas agricultural experimental station bulletin,
7:1-30.

Papavizas, G. C., and C. B. Davey. 1960 Rhizoctonia disease of bean as affected by
decomposing green plant materials and associated microflora. Phytopathology
50:516—521.

Pera, A., G. Vallani, I. SirenoM. L. Bianchin, and M. de Bertoldi, 1983. Effect of organic
matter on rhizosphere microorganisms and root development of sorghum plants in
two different soils. Plant Soil. 74: 3—18.

Perucci, P., 1990. Effect of addition of municipal solid waste compost on microbial
biomass and enzyme activities in soil. Biol. Fertil. Soils 10:221-226.

Peterson, G. A., A. D. Halvorson, J. L. Havlin, O. R. Jones, D. J. Lyon, and D. L.
Tanaka. 1998. Reduced tillage and increasing cropping intensity in the Great
Plains conserves soil C. Soil Till. Res. 47:207—218.

Pimental, D., L. McLaughlin, A. Zepp, B. Lakitan, T. Kraus, P. Kleinman, F. Vancini,
W.J. Roach, E. Graap, W.S. Keeton, and G. Selig. 1991. Environmental and
economic effects of reducing pesticide use. BioScience 41 :401—409.

Rahalison, L., M. Hamfurger, K. Hostettman, M. Monod, E. Frenk, M. P. Gupta, A. 1.
Santana, A. M. Correa, and A. C. Gonzalez. 1993. Screening for antifungal
activity of Panamanian plants. International Journal of Pharrnacognosy 31(1):68-
76.

Ranells, N. N., and M. G. Wagger. 1997. Nitrogen-15 recovery and release by rye and
crimson clover cover crops. Soil Sci. Soc. Am. J. 61:943-948.

Romig, D. E., M. J. Garlynd, R. F. Harris, and K. McSweeney. 1995. How farmers assess
soil health and quality. J. Soil Water Conserv. 50:229—236.

Sainju, U. M., B. P. Singh, and W. F. Whitehead. 1998. Cover crop root distribution and
its effects on soil nitrogen cycling. Agron. J. 90:511—518.

Sarrantonio, M. 1992. Opportunities and challenges for the inclusion of soil-improving
crops in vegetable production systems. Hortic. Sci. 27:754—758.

19

Scholte, K. 1987. The effect of crop rotation and granular nematicides on the incidence of
Rhizoctonia solani in potato. Potato Res. 30: 1 87-199.

Shennan, C. 1992. Cover crops, N cycling and soil properties in semi-irrigated vegetable
production systems. Hortic. Sci. 27:749—754.

Singh, R. S., and K. Sitaramaiah. 1970. Control of plant-parasitic nematodes with organic
soil amendments. PANS 16: 287—297.

Specht, L. P., and S. S. Leach. 1987. Effects of crop rotation on Rhizoctonia disease of
white potato. Plant Dis. 71 :433—437.

Tepper, J. G. o. 1892. “Take-all” and its remedies. Agricultural Gazetteer New South
Wales. 3 269-72.

Tsror (Lahkim), L., R. Barak, and B. Sneh. 2001. Biological control of black scurf on
potato under organic management. Crop Protection. 20(2)145-150.

Tu, J. C., and W. I. Findlay. 1986. The effects of different green manure crops and tillage
practices on pea root rots. In: British Crop Protection Conference on Pests and
Diseases, vol. 1. British Crop Protection Council Publications, UK, pp. 229—236.

Tuitert, G., M. Szczech, and GI. Bollen. 1998. Suppression of Rhizoctonia solani in
potting mixes amended with compost made from organic household waste.
Phytopathology 88:7 64—773.

van Bruggen, A. H. C., and N. J. Grunwald. 1996. Tests for risk assessment of root
infection by plant pathogens. In: Doran, J.W., Jones, A.J. (Eds), Methods for
Assessing Soil Quality. Soil Science Society of America, Madison, WI, pp. 293—
310.

Visser, S. and D. Parkinson. 1992. Soil biological criteria as indicators of soil quality:
soil micro-organisms. Am. J. Alternative Agric. 7 233-37.

Wagger, M. G., and D. B. Mengel. 1988. The role of nonlegurninous cover crops in the
efficient use of water and nitrogen. In W.L. Hargrove (Ed.) Cropping strategies
for efficient use of water and nitrogen. ASA Spec. Publ. 51. ASA, CSSA, and
SSSA, Madison, WI, p. 115—128.

Walker, J. C., S. Morrel, and H. H. Foster. 1937. Toxicity of mustard oils and related
sulfur compounds to certain fungi. Am. J. Bot. 24:536 541.

Weinert, T., W. L. Pan, M. R. Moneymaker, G.S. Santo, and R.G. Stevens. 2002.

Nitrogen recycling by non-leguminous winter cover crops to reduce leaching in
potato rotations. Agron. J. 94:365-372.

20

Wiseman, B. M., S. M. Neate, KO. Keller, and SE. Smith. 1996. Suppression of
Rhizoctonia solani anastomosis group 8 in Australia and its biological nature. Soil

Biol. Biochem. 28:727—732.

Yegen, O., Berger, and R. Heitefuss. 1992. Investigations on the fungitoxicity of Extracts
of 6 selected plants from Turkey against Phytopahtogenic Fungi Z Pflanzenk

Pflanzen. 99(4):349-359.

21

Chapter 2: Effect of Oriental Mustard (Brassicajuncea L.) residues on potato

(Solanum tuberosum L.) fungal diseases.

ABSTRACT

Cover crops have the potential to significantly suppress Rhizoctonia solani Kuhn,
a common potato fungal pathogen. There is limited knowledge of the processes involved
in cover crop residue interaction with soil-borne diseases. To study this interaction, a
field experiment was conducted in parallel with a container study to determine effect of
cover crop tissues on fungal incidence. In the field experiment, cover crops were
incorporated according to the experimental design and the table stock variety Onaway
was planted 2 weeks later. The seeding rate of rye was 100 kg/ha while that of mustard
was 23 kg/ha. Plants were destructively harvested and roots were recovered to assess
disease presence using WinRhionM to quantify dark versus white root tissue. Soil from
the same field was collected for a large-volume container study This study evaluated
oriental mustard tissues (root, shoot and root plus shoot) with two rates of biomass
application. Incorporation of oriental mustard (Brassica juncea) and rye (Secale cereale)
proved beneficial to the preceding potato (Solanum tuberosum) crop by reducing
Rhizoctonia solani root and tuber symptoms. In the field study, rye roots plus shoots and
mustard roots plus shoots reduced R. solani infection by > 50% when compared to a
fumigated treatment. Rye and mustard roots and shoots on their own were less effective.
Neither treatment had any significant effect on the yield. In the container study the shoots
with the higher rate were the most effective at enhancing root health and suppressed R.

solani infection by 75%. Treatment with mustard roots and shoots applied together at the

22

higher rate reduced symptoms of R. solani by 43%, however the tubers with the least

black scurf were associated with this treatment that included roots and shoots.

23

INTRODUCTION

Cover crops can be considered the backbone of any sustainable annual cropping
system. A cover crop is a crop grown as a complementary plant to a cash crop to benefit
the soil in many ways: it can reduce erosion, weed pressure, insects, and plant parasitic
nematodes, soil fungal diseases, thus improving soil quality (Mutch and Martin 2000). A
cover crop can be used as a green manure. The Soil Science Society of America (1987)
has defined green manures as plant materials incorporated into the soil while green,
flowering or at maturity, for soil improvement and protection.

Advantages of incorporating cover crops into soil have been seen in many studies
(Weinert et al., 2002). It has been hypothesized that cover crops contribute to suppression
of soil diseases by increasing the soil active organic matter content, microbial biomass
and thereby the microbial activity, which in turn supports competition that may suppress
pathogenic organisms (Kuo et al., 1997, Mendes et al., 1999, Workneh et al., 1993).
Green manures and cover crops have been shown to differ significantly in their
suppression of root rot severity and damage to plant growth, as indicated by greenhouse
studies (Abawi et al., 2000). In addition, differential effects of various cover crops on the
severity of root rot and yield have been observed under field conditions (Abawi et al.,
2000)

There are reports of organic amendment-mediated soil disease suppression since
as early as the nineteenth century. Root rot of cotton caused by Phymatotrichum
omnivorum was reduced when the soil was amended with manure (Pammel, 1980).

A number of cover crops and green manures can be effective in suppressing

nematode populations and infections (Mojtahedi et al.,1991; 1993; Halbrendt, 1996).

24

Considerable progress has been made in the utilization of organic materials as soil
amendments for the control of plant-parasitic nematodes but soil borne disease
suppression is less well understood (Singh and Sitaramaiah, 1970; Muller and Gooch,
1982; Akhtar, 1993).

A rapid increase in soil microorganisms occurs after a young, relatively lush
green manure crop is incorporated into the soil; this can include beneficial and parasitic
organisms. The soil microbes multiply to attack the freshly incorporated plant material.
During microbial breakdown, nutrients held within the plant tissues are released and
made available to the following crop. The suppressive activity of soil amendments
towards plant pathogens has been well documented in particular for containerized
systems (Hoitink, 1986 and Nelson and Craft, 1992). In cropping systems amended with
organic matter, it has been hypothesized that suppression occurs because of activation of
the indigenous soil microbial community.

Despite the wide use of pesticides in agriculture, growers are now beginning to
become concerned about environmental impacts. In addition, there are concerns about
becoming dependent on one or a narrow range of chemical formulations. If species or
varieties of cover crops could be used to destroy plant pathogen propagules, thereby
decreasing intensity of plant diseases, this would be a most sustainable alternative to
pesticides (Kazmar., 1995; Reddy and Patrick, 1989; Sequeira, 1962; Shetty, 2000).
Reducing the crop damage caused by soilbome fungal pathogens through soil
amendments with brassica plants have received considerable research attention recently.
Biofumigation, defined as the incorporation of biomass into soil, results in the release of

toxic volatiles that reduce soil pests. Other benefits of biofumigation may include:

25

improved soil texture, increased water holding capacity, and improved soil microbial
community structure, if the biofumigant treatment includes enough organic matter.

The Brassicaceae family is a potential source of potential biofumigation material.
The family consists of approximately 375 genera and 3200 species. The Brassica genus
consists of around 100 species including Brassica juncea commonly called “Oriental or
Indian Mustard”. Family members contain secondary plant metabolites, glucosinolates,
which are believed to be involved in plant defense. Brassicas, including numerous
mustard species, provide not only biomass to the soil but when incorporated they release
glucosinolates, which further break down into isothiocyanates with fumigant properties
similar to metham-sodium. Good controls of Pythium ultimum, F usarium oxysporum f. sp
cumim' and Rhizoctonia solani have been reported by amending the soil with leaves of
Brassica species (Ramirez-Villapudua and Munnecke, 1988; Charron and Sams, 1999;
Marwar and Lodha, 2002).

Brassicas in many studies have also been shown to suppress fungal populations
indirectly. In a field study by Subbarao et al., 1999, broccoli residues suppressed
Verticillium dahliae wilt of cauliflower this effect however was not observed in vitro
(Shetty et al., 2000). This could be attributed to increased microbial populations in the
field study which may have created competition that suppressed V. dahliae microsclerotia
thereby reducing infection.

Glucosinolates and their breakdown products have been the focus of many studies
because of the possibility of using them as natural pesticides (Lichtenstein et al., 1964;
Tsao et al., 1996). However, there is evidence that glucosinolates themselves possess

limited biological activity until they are hydrolyzed (Borek et al., 1994). When tissues are

26

damaged, glucosinolates are enzymatically broken down by myrosinase to produce
nitriles, thiocyanates, isothiocyanates and other products. The breakdown products are
generally relatively small molecules, which make many of them volatile. Isothiocyanates,
the predominant breakdown product, show biocidal activity on fungi (Charron,. and Sams
1999; Harvey and Sams, 2000), bacteria (Delaquis, and Mazza. 1995) and other pests.
They are reported to be highly biocidal to a diverse range of organisms including soil
fungi, insects and germinating seeds (Brown and Morra 1997; Kirkegaard et al., 1994).
Walker (1997) demonstrated that residues of Brassica spp. reduced soil populations of
citrus nematode, Tylenchulus semr’penetrans with humus rape resulting in 81% reduction
of larvae. Soil amended with Brassica leaf tissues can exhibit highly nematicidal effects.
Leaf tissues of Brassicas have been noted to have higher glucosinolate levels than root
tissue which may explain 56-95% reduction in Pratyelnchus neglectus by Brassica shoots
and only 0-48% reduction by roots (Potter et al., 1998). In soil, reduction of Pythium
ultimum and Sclerotium rolfsii by dry cabbage leaf amendment together with solarization
was correlated to release of isothiocyanates and aldehydes (Gamliel and Stapleton, 1993).

Oriental mustard has been effectively used as a biofumigant. Anti-fungal
compounds have been isolated from yellow mustard seeds (Neumann et al., 1996).
Amending soil with mustard and providing heat causes differential effect on soil fungi.
This effect was investigated by Stapleton and Duncan (1998). Heating of soil amended
with Brassica tissues to a maximal temperature of 38°C reduced incidence of propagules
of Pythium ultimum, Sclerotium rolfiii and Meloidogyne incognita. When cruciferous soil
amendments were combined with the sublethal heating regime, nematode galling was

reduced by 95-100%, and recovery of active fungi was reduced by 85-100%.

27

The concentration of glucosinolate is variable in different parts of the mustard
plant. The majority of studies involving biofumigation using mustard as a cover crop
utilize the shoots as controls against fungal diseases. This may be due to higher amounts
of glucosinolates present in shoots than in roots. It also appears that research has not
substantially investigated root tissue impact. Previous studies have shown that shoots of
Brassica oleracea, Brassica nigra and Brassica juncea contain high concentrations of
sinigrin (a precursor of isothiocyanate) as compared to roots (Sang et al., 1984;
Kirkegaard 1998 and Sarwar 1998). However there has been no research we know of that
investigates effect of field grown effect of mustard root versus mustard shoot on soil
fungal disease, which was why we initiated this study.

Rhizoctonia solani, Pythium ultimum, Fusarium solani and other soilbome
diseases have emerged as important problems for potato growers in recent years. R.
solani infection of potato sprouts during the emergence period can delay emergence and
seriously debilitate young plants. It can reduce plant growth and tuber production, and
cause lesions on tubers, seriously affecting tuber quality and marketability. Once the
pathogen becomes established in an area, it will persist indefinitely. Organic
amendments, such as the incorporation of green plant residues in the field or compost
added to potting mix, have been shown to suppress the activity of R. solani. Gorodecki
and Hadar, 1990, in a study found that barn (cow) manure significantly suppresses R.
solani incidence. Compost from two commercial composting facilities was investigated
to see effect on R. solani in potting mixtures. It was noted that both the composts
suppressed growth of R. solani in potting mixtures provided the composts were long-

matured (Tuitert et al., 1998). Papavizas and Davey, 1960 studied effect of corn (Zea

28

mays) residues on R. solani and found that incidence of the pathogen was reduced when
soil was amended with corn residues.

P. ultimum is generally a problem in soils that remain wet for very long periods of
time or soils that harbor stagnant water. It is a very fast growing saprophytic soil fungus.
Pythium spp complete their life cycle within 44 h of inoculation but the populations
decline within 30 days as investigated in a field study by Hancock, 1981. They are not
good competitors in soil. If the soil has already been colonized by other microorganisms,
populations of Pythium have been shown to be reduced (Barton, 1961). Because of this
effect, if the cover crop used can increase the soil microbial populations, a reduced
Pythium infection may be possible. Similarly a study by Park, 1958, reported that
F usarium spp are also not good competitors in soil.

Longer rotations (out of potatoes) generally tend to reduce soil inoculum levels of
fungi more than short rotations (Scholte, 1987). Soils, especially those with a low
microbial population are more vulnerable to reinvasion of pathogens following
fumigation. It is possible that the fumigants used in cropping systems reduce populations
of soil beneficial microbes as well. Because of this non-chemical method of effectively
controlling soilbome diseases are needed.

The objectives of this study were: 1) To evaluate the effect of mustard and rye
cover crop residues on potato root and tuber health. 2) To determine if mustard root or
shoot is more effective in controlling disease incidence on potato roots and tuber. 3) To
determine if quantity of mustard root, shoot or root and shoot combined residues are is

important to disease suppressive activity.

29

MATERIALS AND METHODS
MUSTARD FIELD EXPERIMENT
Plot information

The field experiment was conducted at Sandhill research plot near Michigan State
University in East Lansing, Michigan in 2004. The plot was 53*107 m in area. The soil
was well drained and sandy type. The top soil had been removed 15 years ago and sandy
subsoil remained. It was a grass fallow until continuous potatoes 3 years ago was
initiated. This management of continuous potatoes was a strategy representative of field
potato production conditions and susceptible to potato pathogens. Cover crops were
established in 2002 and then in 2003 for this experiment keeping in mind the
experimental design described below.
Cover crop details

The two cover crops studied were rye (Secale cereale L.) and oriental mustard
(Brassica juncea L.). The variety of mustard used was “Pacific Gold”. This variety was
selected for its medium height, high biomass production, high glucosinolate level, and
late flowering properties. The seeding rate of rye was 100 kg/ha while that of mustard
was 23 kg/ha. The amount of shoot biomass incorporated into the soil was calculated by
using a standard 0.5 x 0.5 m quadrat. The quadrat was randomly thrown to a spot in the
plot, the shoots were cut and the fresh and dry weight was calculated. For the root
biomass; six soil cores per quadrat at 0-8 inches depth were taken. The soil was
composited and wet sieved using 6/64 in mesh. The roots were collected with tweezers.
The roots and shoots were placed in an 80°C oven and dry weights were determined. The

biomass by dry weight was determined to be 1834 (1153) kg/ha for mustard shoot, 370 (i

30

96) kg/ha for mustard root, 2809 (i185) kg/ha for rye shoot, and 589 (:l: 132) kg/ha for
rye root.
Inoculation of Plots

After cover crops were established and before the cover crops were incorporated
into the soil, all the plots were inoculated with 3 potato fungal pathogens (14th June
2004). Pythium ultimum, Rhizoctonia solani and F usarium solani isolates were obtained
from the Department of Plant Pathology at Michigan State University. These three
pathogenic organisms were cultured on potato dextrose media and then inoculated on
millet seed. The amount of inoculum needed for the field was determined on a weight
basis. The number of spores on 1 g of millet seed was determined. The millet seed was
taken in a beaker, mixed with 5ml water and stirred for 5 mins. The solution was filtered
through cheese cloth. 40 micro liters of this filtered solution was loaded on a
hemacytometer. Spores were counted on the hemacytometer grid and were diluted in
water for a final spore count of 1600 spores/ml for (F. solani) was 1600 spores/ml, 2000
oospores/ml for P. ultimurn and as R. solani, does not produce asexual spores or conidia
the inoculum was added at the rate of 490 g millet seed/plot (875 sq ft). Around 25 kg/ha
millet seed was used for inoculating the field for each fungus and for Pythium. The millet
seed was spread evenly on the entire plot. The cover crops were moved and disked in 7
days after inoculum was applied so that the fungal spores had time to establish and grow
in the soil.
Incorporation of cover crop residues

The cover cr0ps were moved based on treatments in each plot (Table 2.1). In

treatments with shoot only, the shoots of an entire plot were razed using a forage

31

harvester and were then moved to the needed plot. The shoots were spread evenly using
rakes and were then disked in. The plots from where the shoots were moved were the
plots which had roots only as treatments. The shoots were moved to the shoot only plot.
Treatments

The treatments were based on previous preliminary experiments. For this study
since we were looking at effect of root versus shoot of cover crops, the treatments were
root, shoot and root plus shoot of rye and mustard respectively. The control was a winter

fallow plot with no treatment at all.

Table 2.1. Mustard Field Experiment. Treatments used in the experiment.

 

 

 

 

 

 

 

 

Trt No. Treatment
1 Mustard Roots + Shoots
2 Mustard Roots
3 Mustard Shoots
4 Furnigated
5 Rye Roots + Shoots
6 Rye Roots
7 Rye Shoots

 

 

 

 

Potato variety and fertilization

The variety of potato used was “Onaway”. Seed pieces (52 g) were planted on
30th June 2004 at 34 in distance within rows and 12 in between rows. Nitrogen fertilizer
was applied in splits of 100 kg N /ha at planting, 50 N kg/ha at hilling and 50 N kg/ha at

tuberization for a final rate of 200 kg N/ha (Snapp et al., 2002). Potassium (0-0-60) and

32

phosphorous (19-17-0) were applied at 200 K20 and 35 P205 kg/ha respectively before
planting. The plot was irrigated using a traveler irrigation system.
Potato measurements
Soil Chycterization - Soil organic C was determined by the Walkley-Black method
(Allison et al., 1965)., and total N was by the Kjeldahl method; the total N was extracted
by digestion in H2804 in a bloc digester, then the N content of the digest was
determined on a Technicon Auto-analyser. Ammonium nitrites and nitrates (NH; and
N03) were extracted with KCl (2N) (Bremner 1982) and measured on a Technicon Auto-
analyser.
Soil Biology - Twice over the season, a series of soil dilution bioassays were done to
study effect of the treatments on soil fungi. Soil was sampled 7 days and 40 days after
cover crops were incorporated. 10 g soil was placed in a sterile flask with 90 ml water
and stirred on a mechanical shaker at 160 rpm for 30 mins. 10 ml of suspension was
withdrawn while in motion and 90 m1 sterile water was added to it. This was shaken for 1
min and another 10 ml was withdrawn. A dilution of 10'4 was obtained. 1 ml of this
suspension was spread over the surface of PDA media and spread using a flamed L-
shaped rod. The plated were incubated for 3-4 days and at the end of the 4th day, fungal
colonies were isolated. The fungi obtained from these tests were isolated and cultured on
selective media to ascertain the kind of fungal species.

Microbial biomass analysis was done once mid-season. Soil microbial respiration
(C02) was used to measure biological activity potential, (Kumar and Goh, 2000). A
quantity of 100 grams of soil, collected in the field after soil had been mixed with

different organic residues, was incubated in plastic containers (500 ml) at 25°C for 20

33

days. A solution of glucose (glucose-demineralized water ratio 1:10) was added in each
soil sample to adjust soil moisture to 85% of the field capacity. Vials containing 10 mL
NaOH (1N) were introduced to trap evolved C02 during incubation. The samples were
incubated for 20 days at what temeprature. After 10 and 20 days, vials containing NaOH
were sampled to measure the C02 content. Soils were also sampled at each period to
measure mineralized N.

The amount of accumulated CO2 was determined by titration of excess NaOH
after precipitation of carbonates by barium chloride (BaCl2). The titration was made by
automatic titrimeter with hydrochloric acid (HCl 0.25N). (Anderson 1982). Mineralized
N was extracted with 2 M KCl solution (sol-solution ratio 1:4) (Bremner 1982), and
measured using a Technicon Autoanalyzer. Mineralized C and N were expressed in mg
kg".

Root monitoring - For determining % of root infection, the potato roots were washed with

 

distilled water and analyzed using the WhinRHIZOTM program (Figure 2.1). Fungi were
reisolated from the infected roots using selective media. For this, the root was washed
thoroughly with sterile water. The infected portion of the root was cut using a scalpel.
The cut section was washed with water and then dipped in ethanol. Before transferring
the section on media, it was washed with water again to leave no trace of ethanol. The
root section was placed in a fungal selective media and the petri plate was incubated for
4-5 days. The tubers were scored visually for infection on a scale of 0-3. The scale was
based on % of infection. 0 was 0-25% infection, 1 was 25-50% infection, 2 was 50-75%
infection and 3 was 75-100% infection (Figure 2.4). The dry weights of the roots and

tubers were obtained.

34

Yield tuber monitoring - Destructive harvest measurements were taken twice over the
entire season. The first measurement was taken one month after planting and the second
two months after planting. At harvest, the potato vines were killed using two herbicides
MatrixTM at 0.98 MM and PoastTM at 1.23 L/ha + 0.9 L crop oil concentrate. The last
measurement was done at the final harvest which was on 11th October 2004. Yield
measurements were taken at final harvest along with tuber infection and size
classification. The tubers were sorted based on their diameter into A (2.5 in), B (1.87 in),
over sized (> 3.25 in) and pick outs (deformed tubers).

Analysis for detection of Pratylenchus penetrans and Verticillium dahliae

Processing and analysing the soil samples for P. penetrans and V. dahliae was done by
the Entomology Department at Michigan State University. The amount of soil used for
the analysis was 100 cc for P. penetrans and 10 g for V. dahliae. The method used for

extraction was sieving and decanting (Barker, 198 5).

35

 

Figure 2.1. Mustard Field Experiment. Scale for root infection used by
Wianon‘“. a: 0-25% infection, b: 25-50% infection, c: 50-75% infection, d: 75-
100% infection.

36

Experimental design and Statistical Analysis

The experimental design was a randomized complete block with four replications.
The data was checked for normality and there was no need to transform it. All data were
analyzed by two way ANOVA to test the effect of the cover crop (Rye or Mustard) and
tissue type (Root, Shoot or Root+Shoot). In cases where significance was observed, least
square means was used to determine the effect of an individual treatment. Planned
contrasts were considered using to differentiate significance of each factor. The SAS glm

procedure was used for analyses and an alpha value of 0.05 was maintained.

37

GREENHOUSE CONTAINER STUDY
Treatments

A study was conducted in a container experiment to determine the effect of
different quantities of mustard residues and tissue type on common potato pathogens. The
tissues that were studied were mustard shoots, mustard roots and mustard roots plus
shoots. For each type, two levels were maintained depending on the dry weight of
incorporated tissue (Table 2.2). The weight of the tissue to be added was determined on
the basis of area of the container in comparison with the weight of dry matter obtained in
field conditions. For mustard shoots, the two levels were fresh weight of shoots
equivalent to (1) 40 g dry weight and (2) 160 g dry weight. For mustard roots, the two
levels were fresh weight of roots equivalent to (1) 17 g dry weight and (2) 70 g dry
weight. For mustard roots+shoots, the two levels were fresh weight of root and shoot
combined equivalent to (1) 57 g dry weight and (2) 230 g dry weight. The tissues were
mixed into the soil at a depth of 0 — 20 cm with a trowel. After incorporation of the
tissues, a waiting period of 10 days was observed. During this period, the containers were
covered with “Saran wrap” to avoid loss of volatile glucosinolates. The experimental set-
up is shown in Figure 2.2 and 2.3. The mustard tissues used for this study were obtained

by growing it in the greenhouse at controlled conditions.

38

Table 2.2. Greenhouse container study. Treatments used in the study.

 

Trt No. Treatment

 

1 Mustard Root Lvl 1

 

2 Mustard Shoot Lvl 1

 

3 Mustard Root + Shoot Lvl 1

 

4 Mustard Root Lvl 2

 

5 Mustard Shoot Lvl 2

 

6 Mustard Root + Shoot Lvl 2

 

7 Bare Soil

 

 

 

 

Mustard Growing conditions

The variety of mustard that was used is commonly called ‘Pacific Gold’. Plants
were grown in 43 x 33 x 15 cm plastic tubs. Seeds were sown at twice the field seeding
rate (22.5 kg/ha) to get more biomass. The soil used was a mixture of sandy soil + 40%
perlite. The soil was collected from a research trial field (Sandhill) in East Lansing,
Michigan. The soil was steamed at 70°C for one hour, and then sieved with a 5 mm sieve
before mixing with perlite. Plants were grown in the greenhouse at Michigan State
University at 28°C. The light was maintained at 16 h daylight period using halogen
lights. The fertilizer used was- Osmocotem, which is a slow release plant food with 14-
14-14 N, P, K. All plants were watered uniformly. Both roots and shoots of the plants
were harvested just before flowering. They were washed with distilled water and cut into
small pieces. A compound sample from a large number (10-15 plants) of plants was used

in the bioassay. The tissue samples were used immediately afler harvest.

39

Greenhouse conditions

The study was done in a greenhouse at Michigan State University. The daylight
was maintained at 16 h days using halogen lights and the temperature in the greenhouse
was constant at 70°F throughout the study.
Experimental setup

The containers had a volume of 75L with a height of 58.4 cm and a diameter of
48.3 cm. The soil was a sandy texture and was collected form top soil of the plot at
“Sandhill” site adjacent to the experimental field at Michigan State University. Soil was
collected at 0-25 cm depth. The soil was steamed at 82°C for 3 h to make sure that it was
free of pathogens. It was then sieved through a 6 mm mesh sieve and mixed with 40%
perlite to prevent compaction. Before the containers were filled with soil, 2
minirhizotron tubes were placed criss-cross to each other with their base touching the
base of the container. The container was then filled with soil to the top.
Inoculation

Each of the containers was inoculated with the potato fungi to be studied. In this
experiment we studied Rhizoctonia solani Kuhn, F usarium solani L., and Pythium
ultimum L. The three isolates were obtained from the department of Plant Pathology at
Michigan State University. Each isolate was grown on potato dextrose agar media till the
desired spore count level was obtained. For this study the spore count was set at 1600
spores/ml for F solani, 2000 spores/ml for P. ultimum and for R. solani, the inoculum
level was 55 g dry inoculum/sq in. After inoculation of soil, it was incorporated with

mustard tissues at desired quantities

4O

 

MiniRhizotron
tube

Potato root

Container

 

 

 

 

 

Figure 2.3 Greenhouse container experiment. Experimental set-up in greenhouse

showing containers with minirhizotron tubes.

41

Potato plant growth conditions

The potato cultivar used was “Onaway”. The seed tubers were surface
sterilized with 5% Clorox bleach and washed thoroughly with distilled water to
avoid any cross contamination. Seed pieces were planted 8 cm deep. Two seed
pieces were planted per container and the more vigourous plant was left in the
container. The plants were watered as needed based on soil moisture content
measured by tensiometers. To maintain soil moisture, soil tensiometers were
installed at 15 cm depth. Fertilizer was applied at field rates. Nitrogen fertilizer
was applied in 3 splits at 200 N kg/ha as recommended. 100 N kg/ha was applied
at planting, 50 N kg/ha at hilling and 50 N kg/ha at tuberization (Snapp et al.,
2002). Potassium (0-0-60) and phosphorous (19-17-0) were applied at 200 K20
and 35 P205 kg/ha respectively before planting.
Monitoring root and tuber disease

Root disease was monitored by taking picture of roots using the Mini Rhizotron
camera in tubes. Pictures were taken nine times over the entire growth season at seven
day intervals. Each time, pictures were taken at 12 depths along each tube. Pictures were
analyzed using BTC-ICAPTM software to observe infection (browning) of roots over time
inside the soil. At the end of the plant growth, plants were harvested and roots were
analyzed for infection. The roots were washed with distilled water and were stored for a
week before analysis in a solution of 5% ethanol. They were analyzed using the
WinRHIZOTM program. The tubers were scored visually for infection on a scale of 0-3.
The scale was based on % of infection. 0 was 0-25% infection, 1 was 25-50% infection, 2

was 50-75% infection and 3 was 75-100% infection (Figure 2.4).

42

 

Figure 2.4. Greenhouse container study. Scale used for rating tubers on basis of %
infection. a: 0 (II-25%), b: l (ZS-50%), c: 2 (SO-75%), d: 3 (75-100%)

Reisolating fungi from roots

Since there were three fungal species used for this study it was important to
determine how many or which of the three pathogens had infected the roots. For this, the
infected portion of roots was cut using a scalpel. This root section was washed in water

and ethanol and cultured on fungal selective media. Since the three pathogens used were

43

P. ultimum, R. solani and F. solani, the selective media used was specific for each

pathogen. The selective media recipes were as follows:

1)

2)

3)

For P. ultimum — 1L water autoclaved with 17 g cornmeal agar, 20 g sucrose, 1mg
ZnCl2, 0.02 mg CuSO4.5H20, 0.02 mg M003, 0.02 mg MnCl2, 0.02 mg
FeSO4.7H20, 10 mg MgS04.7H20, 10 mg CaCl2, 100 mg Thiamine
hydrochloride, 10 mg Benomyl, 100 mg PCNB, 10 mg Rose Bengal, 2 ml RAN
and 1 ml of a 100 ppm dimethomorph stock solution.

For R. solani — 1L water autoclaved with 20 g Agar, 1 g K2HP04, 0.5 g KCl, 0.2 g
NaN02, 0.5 g MgSO4.7H20, 10 mg FeSO4.7H20 and 0.4 g Gallic Acid. After
autoclaving 90 mg F enaminosulf, 50 mg Chloramphenicol and 50 mg
Streptomycin was added to it.

For F. solani — 1L water autoclaved with 20 g agar, 20 g sucrose, 1 g KH2HP04,
0.5 g KCl, 0.375 g Quintozene, 2 g KN03, 0.5 g MgSO4.7H20 and 0.5 g Oxagall.
After autoclaving 70 mg dodine acetate, 600 mg streptomycing and 50 mg

chlortetracycline HCl was added.

Growth of colonies was monitored on the media for a period of 10 days and the species

that grew was identified.

Experimental design and Statistical Analysis

The experimental design was a completely randomized block design with 3

replications. A two way ANOVA was used to determine the significance of the study.

The first factor was the type of mustard tissue used (root, shoot or root+shoot). The

second factor was the rate at which the tissue was incorporated (Ratel or Rate2). Planned

contrasts (root v/s shoot and rate lv/s rate 2) were used to determine the effect of one

44

treatment against another and least square means using a one way ANOVA was used to
determine the best treatment. SAS was used for all the above statistical analyses and an

alpha value of 0.05 was maintained.

45

RESULTS
MUSTARD FIELD EXPERIMENT
Fungal infection of potato roots:

The components of the analysis of variance for % area of diseased root is
presented in Table 2.3. For the 1St and the 2"d harvest, the cover crop (mustard or rye), the
tissue (root, shoot or root + shoot) was significant. While the interaction between cover
crop and tissue was significant only for the 1St harvest. This can be attributed to a
decreased effect of glucosinolates later in the season. Throughout the growing season, the
treatment with roots + shoots was consistently associated with the lowest fungal infection
on roots. Addition of roots and shoots together may have increased the soil microbial
organisms which indirectly created competition for the fungal pathogens. This is
consistent with the findings of Kuo et al., 1997, Mendes et al., 1999 and Workneh et al.,
1993. Although the SIR results did not conclusively show increased microbial respiration
in the treatments with root and shoot together, it is possible that addition of the tissues
together increased the microbial biomass around the potato rhizosphere. Analysis with
the least square means show that the % of disease infection was lowest in mustard roots +
shoots (Figure 2.5). The treatment with rye including both the roots and shoots gave
almost as good results as the mustard treatment. This can be related to the amount of
biomass that was incorporated in the soil. The rye root and shoot biomass getting disked
in the soil was greater than the mustard. It is not yet determined, but this disease
reduction in the rye treatment could also be associated with the allelopathic effect of rye.
Mustard roots and mustard shoots by themselves gave consistent results in both the 1St

and the 2nd harvest.

46

Earlier in the season, planned contrasts for the effect of mustard versus rye
determined that there was a significant difference between the two cover crops (Table
2.4.). As time progressed, this contrast between the two decreased and they showed
almost the same effects. This effect could be due to greater glucosinolates levels in the
mustard treatment when residues were freshly incorporated. Glucosinolates being

volatile, it maybe possible that the effects later subsided.

Table 2.3. Mustard Field Experiment. Analysis of variance for % diseased root area.

 

ANOVA Pr > F

Source of variation df % diseased area

lst harvest 2nd harvest
Cover Crop 1 <0.0001 0.0026
Tissue 2 <0.0001 <0.0001
CoverCrop X Tissue 2 <0.0484 0.2396 NS

 

NS: Not Significant at P<0.05

Table 2.4. Mustard field experiment. Planned contrasts for % of diseased root.

 

Pr > M
Parameter lst Harvest 2nd Harvest
Mustard v/s Rye 0.0033 0.2943 NS
Shoots v/s Roots 0.0653 0.3176 NS

 

NS: Not Significant at P<0.05

47

 

 

 

 

55
l 50 j a 1st Harvest
l 8 45 ’1
a 40 ;
‘6 35 1
o l
.. 301 ch
b
§ 25 ‘
‘g 201
1:5 15 ; cd (1
1"- 10 1
1° 5+ I I
io\° ,
l o- —. .- .
b \ \. N. \. \. O"
l 6‘ A0 X S b x ‘
' {‘0 Q‘ GI ‘ 9"? (if 0"
e°° 9° 0° e"
I o ‘5 ,8
Qt) 0 I
09 1
. _ - _fi , -_ _ 9 ,1
I -, ,. __,_
l 55. ---#~7 #7 - Al
13 50 a2nd Harvest ,1
a 451 11
g 401
‘2 35 .
{B 30 ~ 1
‘3 25 j ‘
g 20 .1
if: 15
.o 10 i
«e 51 j
‘ 0
I ,5ch (29° 0° 6.6290
1 62.0 (e. 53°00 xe“ 50390 xoo‘e“
{(5 e- e5 00x 09 $1 00 l
I of 4° 5* .
l f at ‘
Treatments 4“)

 

Figure 2.5. Mustard Field experiment. % area of diseased root. Treatments with

different letters are statistically different at P<0.05.

4s

Dry weight of roots, specific gravity and yield of tubers:

Dry weights of roots were not significantly different in either the treatments or
the harvests. The specific gravity for all the treatments was almost the same (figure 2.6,
table 2.7). The treatments did not significantly alter the tuber yields (table 2.7). Although
the overall yields for all treatments were lower than expected (figure 2.7.) The analysis of
variance for dry weight of root is given in table 2.5. ANOVA for specific gravity and

yield are given in table 2.6.

Table 2.5. Mustard Field Experiment. Analysis of variance for dry weight of roots.

 

ANOVA Pr > F
Source of variation df Dry wt of root

lst harvest 2nd harvest

Cover Crop 1 0.0192 0.8962 NS
Tissue 2 0.0947 NS 0.1665 NS
CoverCrop X Tissue 2 0.1696 NS 0.0888 NS

 

NS: Not Significant at P<0.05

Table 2.6. Mustard Field Experiment. Analysis of variance for specific gravity and

 

yield of tubers.
ANOVA Pr > F
Specific
Source of variation df Gravity Yield(g/plant)
Cover Crop 1 0.7078 NS 0.5449 NS
Tissue 2 0.2439 NS 0.6854 NS
CoverCrop * Tissue 2 0.8226 NS 0.3659 NS

 

NS: Not Significant at P<0.05

49

Table 2.7. Mustard Field Experiment. LS Means and Standard Errors for specific
gravity and yield of tubers.

 

LS MEANS
Specific Gravity Yield

Fumigated 1.072 (:h 0.002) 354.20 (:1: 32.48)
Rye Root 1.076 (d: 0.003) 442.95 (:1: 66.38)
Rye Shoot 1.076 (at 0.001) 399.60 (i 46.81)
Rye Root + Shoot 1.073 (:t 0.002) 350.00 ($41.17)
Mustard Root 1.075 (i 0.001) 369.80 (i 111.80)
Mustard Shoot 1.077 (i 0.001) 487.75 (:t 46.49)

Mustard Root + Shoot 1.076 (:1: 0.0008) 429.55 (i 34.25)

 

 

_ 1.070
1.077
1.075 -
1.075
1.074 «
1.073
1.072 »
1.071
1.07
1.069

111)

Specific Gravity

 

 

 

 

 

 

 

Figure 2.6. Mustard Field Experiment. Specific Gravity for different treatments.

Treatments followed by the same letter are not significantly different at P<0.05

50

 

 

 

 

 

 

1 cove
‘3 500 .1
1 5 l a a a
. a.
'3, 400 - a
1 c a a
3 300
1c
1 § 200
. b
F; 100-
1';
1 0
b x \ x 6‘ a. x
_ g9 0° 05°00 6°C? <29 600 6°00
‘0" 0 o x 1P o x ‘
«6° Qfi QT 0° 00 63¢ oé I
0% Q V‘s @Q- 1
. e“ 90
j Treatment ‘50

 

Figure 2.7. Mustard Field Experiment. Yield of fresh tubers in g / plant. Treatments
followed by the same letter are not significantly different at P<0.05.

51

Microbial and Nematology analysis

Figures 2.9 and 2.10 show results from the Substrate Induced Respiration (SIR)
at 0-10 days and 10-20 days respectively. At 0 — 10 days, rye roots plus shoots treatment
had the lowest SIR than all other treatments including control fallow. Apparently, the
root+shoot of rye caused a temporary immobilization and minimized SIR from 0-10 days,
presumably due to low quality residues of rye taking time to be decomposed by soil
microorganisms which became more active in the 2"d time period. Between 10-20 days,
the SIR was highest for rye root+shoot (1.64 mg C02/kg/h) and lowest for rye shoots
(1.43 mg C02/kg/h) (table 2.12). SIR overall was lower at 10-20 days.

The disease suppression effect associated with mustard tissues (Figure 2.5) can
be related to microbial activity in the soil which may compete with the soil pathogenic
fungal diseases. Table 2.11 presents the ANOVA for different sources of variation.
Contrasts between rye versus mustard and root versus shoot are given in Table 2.13 and
show no significant treatment effects.

Soils were monitored for presence of root rot nematodes (Pratylenchus
penetrans) and colonies of Verticillium dahliae, which are associated with the potato
disease verticillium wilt. In the first sampling on May 2004 (7 days after tissue
incorporation), treatments from root+shoot residue incorporation did not show presence
of P. penetrans (Figure 2.10). The number of nematodes increased by the second
evaluation date - September 2004. It is possible that reduced number of nematodes for
mustard and rye root+shoot be attributed to maximum effect of the glucosinolates and
increased microbial population early in the season. While later in September, effect due

to the soil amendments reduces. Although the treatments show differences in the spore

52

count for P. penetrans, the threshold level for the pathogen to cause a significant damage
to crop is 150 spores / 100 cc of soil. In the given experiment the levels were very low.
The carbon content was low at the experiment site. The organic carbon % for each
treatment is presented in table 2.8.

Treatment with the mustard root+shoot combination consistently showed lowest
colony p0pulations of Verticillium dahliae in May and September 2004. The
concentration of V. dahliae was very low in May and increased by September. Treatment
with mustard root had the highest colony count in May while rye root has the highest
colony count in September. At 2"d monitoring time period (September 2004), high
variability was seen in the colony count for V. dahliae. This may have resulted in no
significant differences observed in the treatment with root+shoot. Overall low levels of V.
dahliae were seen in this study. This can be as a result of the very sandy nature of the soil
at the field site.

The analysis of variance and planned contrasts for P. penetrans are
presented in Tables 2.11 and 2.12. Analysis of variance and planned contrasts for V.

dahliae colonies are given in table 2.13 and table 2.14 respectively.

53

Table 2.8. Mustard Field Experiment. Least Square Means for Organic

Carbon % in the treatments.

 

Treatment LS Means
Fumigated 0.61 a
Rye Roots 0.56 a
Rye Shoots 0.56 a
Rye Roots + Shoots 0.63 a
Mustard Roots 0.63 a
Mustard Shoots 0.56 a
Mustard Roots + Shoots 0.58 a

 

LS Means with different letters are significantly different
at P<0.05. Least Significance difference = 0.1046

54

 

 

 

 

 

1
1
1
1

Treatment ‘3’
Figure 2.8. Mustard Field experiment. Substrate induced respiration in mg

C02/kg/h between 0 — 10 days. Treatments followed by the same letter are
not significantly different at P<0.05.

 

 

 

 

 

 

1T 3,,,,_,.-__., — 1
1
w: 2‘ 1
x- ab ab b a ab ab ab
2 1
6
o 1-
m
E
l
0 , 1 j
x x x o
1 150° 6°00 Q90 9.,“ 9.29 1
0&9 \x .556 0‘6 \x '
k
55»
Treatment \b

Figure 2.9. Mustard Field experiment. Substrate induced respiration in mg
C02/kg/h between 10-20 days. Treatments followed by the same letter are not
significantly different at P<0.05.

55

Table 2.9. Mustard Field Experiment. Analysis of variance for substrate induced

 

respiration.
ANOVA Pr > F
Source of Variation df C02 released C02 released
from 0—10 days from 10-20 days
Covercrop 1 0.1360 NS 0.9020 NS
Tissue 2 0.2502 NS 0.1935 NS
Covercrop * Tissue 2 0.3792 NS 0.5929 NS

 

NS: Not Significant at P<0.05

Table 2.10. Mustard Field Experiment. Planned contrasts for substrate induced

 

respiration.
Pr > M
Parameter C02 released C02 released
from 0-10 days from 10-20 days
Mustard v/s Rye 0.1203 NS 0.5936 NS
Roots v/s Shoots 0.2290 NS 0.1468 NS

 

NS: Not Significant at P<0.05

56

 

May2004
j_10'
'3
in 8-
1E
1§ 6'
18 4-
I'D
12
(I .
1e 2
1: 01

 

 

 

 

Figure 2.10. Concentration of Pratylenchus penetrans spores/100 cmJ soil observed
in May 2004. Treatments followed by the same letter are not significantly different
at P<0.05.

 

.5
N

a September 2004 a ‘

.A
O
4

nematodes/100 cm 3 soil
-5 0')

M

 

 

 

09 1
Treatment ‘1‘ 3

Figure 2.11. Concentration of Prarylenchus penetrans spores/100 cm3 soil observed
in September 2004. Treatments followed by the same letter are not significantly
different at P<0.05.

57

Table 2.11. Mustard Field Experiment. Analysis of Variance for Pratylenchus

penetrans in May 2004 and September 2004.

 

ANOVA Pr > F

Source of variation if my September
M4. M1
Covercrop 1 0.3097 NS 0.7566 NS
Tissue 2 0.0956 NS 0.1786 NS
Covercrop * Tissue 2 0.5731 NS 0.4997 NS

 

NS: Not Significant at P<0.05

Table 2.12. Components of analysis of variance and planned contrasts for spore
population of Pratjylenchus penetrans in 100 cm3 soil in May and September 2004 in

plots treated with incorporations of mustard or rye shoots only or roots only.

 

 

Pr > t

Parameter Date of evaluation
May2004 September

Mustard v/s Rye 0.9365 NS 0.2898 NS

Root v/s Shoot 1.0000 NS 0.2898 NS

 

NS: Not Significant at P<0.05

Table 2.13. Mustard Field Experiment. Analysis of Variance for Verticillium dahliae
in May 2004 and September 2004.
ANOVA Pr > F

 

Source of variation d_f _M_ay September
29.9.4. MA
Covercrop 1 0.1419 NS 0.2883 NS
Tissue 2 0.2179 NS 0.5142 NS
Covercrop "‘ Tissue 2 0.2179 NS 0.4516 NS

 

NS: Not Significant at P<0.05

58

Table 2.14. Mustard Field Experiment. Planned contrasts for Verticillium dahliae in
May 2004 and September 2004.

 

Pr > t
Parameter M_ay September
are 2.0021
Mustard v/s Rye 0.0449 0.2248 NS
Root v/s Shoot 0.1228 NS 0.8627 NS

 

NS: Not Significant at P<0.05

59

GREENHOUSE CONTAINER EXPERIMENT
% Area of diseased root

% of disease infection on the roots was lower for potatoes grown after
incorporation of cover crop shoots. This is consistent with the findings of Charron and
Sams, 1999, who determined that a 75% decrease in Rhizoctonia solani growth was seen
when subjected to shredded leaves of Brassica juncea. The distribution of glucosinolate
in a mustard plant is not uniform. The amount of glucosinolate was (report number in
units of measurement) in the shoots and (report number) in the roots (Figure 2.12). From
figure 2.12 we can infer that the amount of glucosinolate is higher in the shoots than in
the roots. This is also consistent with the findings of Sang et al., 1984; Kirkgaard 1998
and Sarwar 1998. Disease infection was lowest in the treatments with the higher rate of
tissue incorporation. Reduction in disease in these treatments is attributed to the increased
amount of glucosinolates being added to the soil.

The treatments with mustard roots only were not associated with decrease in
infection. This may be related to two factors. 1) Roots of mustard may not be high in
glucosinolate concentration, 2) Since the amount of root biomass being added into the
soil was almost one third the amount of shoots being incorporated, the low rate might
have had an effect on disease control.

Mustard roots + shoots treatment did not show any added benefit in this
experiment as compared to mustard shoot only. Although in a field study, it would be
impractical to separate the roots and shoots for incorporation. Taking this fact into
consideration, treatment with mustard root + shoot could be used as an effective fungal

disease control.

60

Contrasts between the two rates and roots v/s shoots were shown in Table 2.16.
There was a significant difference measured for the tissue type that was incorporated into
the soil (root, shoot or root+shoot) with P < 0.0003. The rate of tissue incorporation (P <
0.6931) and the interaction between the two (F < 0.4523) was found to be not

significantly different (table 2.15).

 

100

 

, .
90 . a /0 area of root disease

80
70 -
60 .
50
40

% of diseased root

30
20

 

 

 

Bare Root Root Shoot Shoot Rt+Sht Rt+Sht
Rate1 Rate2 Rate1 Rate2 Ratet Rate2

Treatment

 

 

 

Figure 2.12. Greenhouse container study. % area of diseased root. Treatments

followed by the same letter are not significantly different at P<0.05.

61

Table 2.15. Greenhouse container experiment. Analysis of Variance for % area of
diseased root.

ANOVA Pr > F

 

Source of Variation df Pr > F
Tissue 2 0.0003
Rate 1 0.6931 NS
Tissue * Rate 2 0.4523 NS

 

NS: Not Significant at P<0.05

Table 2.16. Greenhouse container study. Planned contrasts for % of diseased root.

 

Pr > |t|

Parameter
Rate 1 v/s Rate 2 0.0737 NS
Shoots v/s Roots 0.3299 NS

 

NS: Not significant at P<0.05

62

Tuber Infection Scoring

The analysis of variance is given in table 2.17. The tubers from the experiment
were rated on basis of the surface infection by the fungi. They were sorted into 4 classes
depending on the % of infection on each tuber. The percent of tubers with sclerotia of R.
solani on the surface area in each of the four classes for each incorporation treatment is
shown in Figure 2.13. The higher the % belonging to the 0-25 % category the better is the

tuber health.

 

 

”Ti-7f- I

  

80% ~

60%

40%:

 

% infected tubers

20% »

 

 

 

 

 

 

 

 

 

 

1 0% 1 1
3 e ‘I~ ‘1- ‘l~ 5+ ‘I~ ‘|~
oft e99 6‘“ e90 6“"
0° Q~ 0°.“ 63‘ 9.01; 6&6

o as 0

Q‘ Q 9 5“ «8‘ Q5):

- . , - iTreéiinfnj‘:
1 a 0-25 % infection :1 25-50 % infection ‘
1 :2! 50-75 % infection I 75-100 % infection 1

%,

Figure 2.13. Greenhouse container study. % Area of infected tubers.

63

Table 2.17. Greenhouse container experiment. Analysis of Variance for tuber

 

scoring
ANOVA Pr > F
Source of Variation df Pr > F
Tissue 2 1.0000 NS
Rate 1 0.9998 NS
Tissue * Rate 2 1.0000 NS

 

NS: Not significant at P<0.05

64

Reisolation of fungi from roots and tubers

The soil was inoculated with Pythium ultimum, F usarium solani and Rhizoctonia
solani. But when the infected portion of roots and tubers was cultured on selective media,
it was observed that the only fungal infection was that by R. solani (Figure 2.14.) R.
solani was isolated from both the roots and the tubers. P. ultimum grows only in very wet
conditions where the soil needs to be damp all the time. This was not the case in our
experiment because of which we did not see any P. ultimum infection. Although the F.
solani inoculum had a very high spore count, we still did not see any infection because of

it.

 

Figure 2.14. Reisolation of Rhizoctonia solani from, a: potato tuber and b: potato

root.

65

DISCUSSION

Suppression of all soil-borne diseases and nematodes in the field experiment was
greatest with mustard root + shoot treatment at both the first and second harvest dates in
May and September. In contrast, in the container study, treatment with mustard shoots at
the higher level exhibited maximum disease suppression. This can be related to growing
condition differences in the field and in the greenhouse. In the field, it is possible that the
roots and the shoots interacted to bring about increase in the soil microbial community
which indirectly created competition for the fungi. This can also be related to some
change in the soil fertility that was caused by the microbes which indirectly resulted in
controlling fungal growth as shown by Bardin et al., 2004. In this container study,
microorganisms associated with pea straws were responsible for concerting nitrate into
volatile ammonia which in turn controlled Pythium damping-off disease in sugar beet.

In the field study, treatment with rye root+shoot also effectively controlled fungal
infection. It is not clear why this treatment may result in disease suppression, but it could
be related to microbial community competition or to some allelopathic effect of rye.
These effects can be compared to a field study by Sumner et al., 1995 who found that
Pythium irregulare and Rhizoctonia solani populations decreased when rye (Secale
cereale) or Brassicas (B. oleracea and B. campestris) were used as cover crops as
compared to when legumes were used as cover crops.

Although not significantly different with most of the other treatments, it was
observed that the dry weight of roots was highest for treatment with rye root+shoot at

harvest 1 and mustard root+shoot at harvest 2.

66

The overall P. penetrans and V. dahliae populations were very low in our field.
Since the field soil was very sandy in nature and the history of potato production was
short, it is not surprising that these populations were low. Interestingly in a study by
Knudsen et al., 2002 who studied disease suppression soil types and found that sandy
soils are more suppressive to disease than clay soils, thus texture shall be considered.

It is possible that reduced number of nematodes for mustard and rye root+shoot
may be attributed to maximum effect of the glucosinolates and increased microbial
population early in the season. The nematicidal property of plant residues was shown in a
study by Guertal et al., 1998 who in a greenhouse study found that Rotylenchulus
reniformis nematode populations were reduced following a rye amendment. There have
been many studies demonstrating nematicidal effect of Brassicas. Potter et al. (1998)
showed that soil amended with leaf tissues of Brassicas was highly nematicidal killing
56-95 % of exposed root lesion nematode, P. penetrans. V. dahliae populations were
consistently lower in both May and September in soil treated with mustard root+shoot.

Longer term studies in our field experiments should provide more insights.

67

CONCLUSION

Taken together data from the two experiments supports the use of mustard as a
cover crop for controlling Rhizoctonia solani in the subsequent potato crop. Although
mustard in this experiment did not produce high biomass, the results were clear.
Treatments with mustard shoots consistently gave better results than treatments with only
roots. Since glucosinolates, the compounds responsible for reduction in general soil-
bome inoculum, are predominant in shoots, greater reduction in symptoms of potato tuber
and root diseases in treatments with shoots is not surprising. The treatments did not
significantly differ in yield but over a longer period of time, the biofumigant property of
mustard is sure to have a favorable effect on the yield. The specific gravity for all the
treatments was almost the same and did not show any significant pattern.

Rye, which is very easy to establish had provided some suppression of disease
infection in this field experiment. This maybe due to rapid increase in the number of
beneficial soil microbes, supported by the cover crop residue substrate, which creates
competition for R. solani in early stages of its growth. This could have an effect on the
overall population of R. solani. However bulk soil measurements of soil microbial
organism activity was moderately suppressed by rye root + shoot and thus was not
correlated to suppressive activity. It is possible that the competition occurs in the potato
root rhizosphere, not in the bulk soil. Further work is needed to determine reason for

reduction in disease when rye roots and shoots together are used as cover crops.

68

FUTURE WORK

Analysis of glucosinolate content in the root and shoot of mustard could provide
insights regarding the superior biofumigant property of shoots over roots.

The best growth stage for incorporating mustard residues can be studied so as to
obtain maximum fungal disease reduction benefit.

Effect of rye and mustard on soil microbial population immediately after
incorporation should be determined, focusing on crop root rhizosphere communities

rather than a bulk soil.

69

REFERENCES CITED

Abawi, G. S., and T. L. Widmer. 2000. Impact of soil health management practices on
soilbome pathogens, nematodes and root diseases of vegetable crops. Appl Soil
Ecol 15 (1):37-47.

Akhtar, M. 1993. Utilisation of plant-origin waste materials for the control of parasitic
nematodes. Bioresource Technology 46: 255—25 7.

Allison, L. E., W. B. Bollen, and C. D. Moodie. 1965. Total carbon In: C.A. Black et al.,
eds. Methods of soil analysis. Agronomy no.9. American Society of Agronomy
and Soil Science Society of America, Madison, WI. pp1346-1365.

Angus, J. F., P. A. Gardner, J. A. Kirkegaard, and J. M. Desmarchelier. 1994.
Biofumigation: isothiocyanates released from Brassica roots inhibit growth of the
take-all fungus. Plant and Soil 162:107—112.

Bardin, S. D., H. C. Huang, and J. R. Moyer. 2004. Control of Pythium damping-off of
sugar beet by seed treatment with crop straw powders and a biological agent.
Biological Control. 29:453-460.

Barker, K. R. 1985. Nematode extraction and bioassays. In: K.R. Barker, C.C. Carter and
J.N. Sasser, Editors, Methodology vol. II, NCSU Graphics, Raleigh pp. 19—35.

Barton, R. 1961. Saprophytic ability of Pythium marnillatum in soils 11. Factors
restricting P. Mamillatum to pioneer colonization of substrates. Trans. Br. Mycol.
Soc. 44:105-118.

Borek V., M. J. Morra, P. D. Brown, J. P. McCaffrey. 1994. Allelochemicals produced
during sinigrin decomposition in soil. J Agric Food Chem. 42:1030—1034.

Bremner, J. M. and C. S. Mulvaney. 1982. Total nitrogen. Pages 595-622 in AL. Page,
R.H. Miller, and DR. Keeney, eds. Methods of soil analysis. 2nd ed. Agronomy
no 9. ASA and SSSA, Madison, WI.

Brown, P. D., and M. J. Morra. 1997. Control of soil-borne plant pests using
glucosinolate containing plants. Advances in Agronomy 61:167-231.

Charron, C. S., S. E. Sams. 1999. Inhibition of Pythium ultimum and Rhizoctonia solani
by shredded leaves of Brassica species. J. Am. Soc. Hort. Sci. 124:462-467.

Delaquis, P. J. and G. Mazza. 1995. Food Technol. Nov. 73-84

70

Gamliel, A., T. J. Stapleton. 1993. Characterization of antifungal volatile compounds
evolved from solarized soil amended with cabbage residues. Phytopathology
83:899-905.

Gorodecki, B., and Y. Hadar. 1990. Suppression of Rhizoctonia solani and Sclerotium
rolfsii in container media containing composted separated cattle manure and
composted grape marc. Crop Prot. 9:271-274.

Guertal, E. A., E. J. Sikora, A. K. Hagan, and R. Rodriguez-Kabana. 1998. Effect of
winter cover crops on populations of southern root-knot and reniform nematodes.
Agriculture, ecosystems and environment. 70:1-6.

Halbrendt, J. M. 1996. Allelopathy in the management of plant-parasitic nematodes.
Journal of Nematology 28:8-14.

Hancock, J. G. 1981. Longevity of Pythium ultimum in moist soils. Phytopathology.
71:1033-1037.

Harvey and Sams. 2000. Methyl Bromide Alternatives Emissions Reductions. Proc. Ann.
Intl. Res. Conf. 92-1

Hoitink, H. A. J. and P. C. F ahy. 1986. Basis for the control of soilbome plant pathogens
with composts. Annu. Rev. Phytopathol. 24293—114.

Honeycutt, C. W., W. M. Clapharn, and S. S. Leach. 1996. Crop rotation and N
fertilization effects on growth, yield, and disease incidence in potato. Am. Potato
J. 73:45—61.

Kazrnar, E. R. 1995. The effect of intercropped oats on Aphanomyces root rot of pea.
Masters thesis, University of Wisconson, Madison.

Kirkegaard, J. A., P. A. Gardner, J. F. Angus, E. Koetz. 1994. Effect of Brassica break
crops on the growth and yield of wheat. Aust. J. Agric. Res. 45:529—545.

Kirkegaard, J. A. and M. Sarwar. 1998. Biofumigation potential of brassicas. 1. Variation
in glucosinolate profiles of diverse field-grown brassicas. Plant Soil 201 :71—89.

Knudsen, M. B., K. M. Larsen, D. F. Jensen, and J. Hockenhull. 2002. Potential
suppressiveness of different field soils to Pythium damping-off of sugar beet.
App. Soil. Ecol. 21:119-129.

Kumar, K. and K. M. Goh. 2000. Crop residues and management practices: effects on
soil quality, soil nitrogen dynamics, crop yield and nitrogen recovery. Advances
in Agronomy 68:198-279.

71

Kuo, S., U. M. Sainju, and E. J. Jellum. 1997. Winter cover crop effects on soil organic
carbon and carbohydrate in soil. Soil Science Society of America Journal. 61 :145-
152.

Lazzeri L., G. Baruzzi, L., Malaguti and L. Antoniacci. 2003. Replacing methyl bromide
in strawberry production. 59(9):983-990.

Lichtenstein E. P., D. G. Morgan, C. H. Mueller. 1964. Naturally occurring insecticides
in cruciferous crops. J Agric Food Chem. 12:158—161.

Mawar, R., S. Lodha. 2002. Brassica amendments and summer irrigation for control of
Macrophomina phaseolina and Fusarium oxysporum f. sp. cumuni in hot arid
regions. Phytopathol. Mediterr. 41 :45-54.

Mendes, I. C., A. K. Bandick, R. P. Dick, and P. J. Bottomley. 1999. Microbial biomass
and activities in soil aggregates affected by winter cover crops. Soil Science of
America Journal. 63:873-881.

Mojtahedi, H., G. S. Santo, A. N. Hang, and J. H. Wilson. 1991. Suppression of root-knot
nematode populations with selected rapeseed cultivars as green manure. J.
Nematol. 23: 1 70-174.

Mojtahedi, H., G. S. Santo, and R. E. Ingham. 1993. Suppression of Meloz’dogyne
chitwoodi with sudangrass cultivars as green manure. Journal of Nematology 25:
303-311.

Muller, R., and P. S. Gooch. 1982. Organic amendments in nematode control. An
examination of the literature. Nematropica 12: 319—326.

Mutch, D. R., and T. E. Martin. 2000. Cover crops. In Michigan field crop ecology.
Michigan State University. Extension bulletin. E-2646, pp 44-53.

Nelson, E. B., and C. M. Craft. 1992. Suppression of dollar spot on creeping bentgrass
and annual bluegrass turf with compost-amended t0pdressing. Plant Dis. 76:954—
958.

Neumann, G. M., Condron R., and Polya G. M. 1996. Purification and mass
spectrometry-based sequencing of yellow mustard (Sinapis alba L.) 6 kDa
proteins. Identification as antifungal proteins. Int. J. Pept. Protein Res. 47:437-
446.

Pammel. L. H. 1980. Cotton root-rot. Texas agricultural experimental station bulletin,
7:1-30.

72

Papavizas, G. C., and C. B. Davey. 1960 Rhizoctonia disease of bean as affected by
decomposing green plant materials and associated microflora. Phytopathology
50:516—521.

Park, D. 1958. The saprophytic status of F usarium oxysporum Schl. causing vascular wilt
of oil palm. Annals of Botany. 22:19-35.

Potter, M. J ., K. A. Davies, and A. J. Rathjen. 1998. Suppressive impact of glucosinolates
in Brassica vegetative tissues on root lesion nematodes (Pratylenchus neglectus).
Journal of Chemical Ecology, 24:67-80.

Ramirez-Villapudua, J., D. E. Munnecke. 1988. Effects of solar heating and soil
amendments of cruciferous residues on Fusarium oxysporum f. sp. conglutians
and other organisms. Phytopathology 78:289-295.

Reddy, M. S. and Z. A. Patrick. 1989. Effect of host, nonhost and fallow soil on
populations of T hielaviopsis basicola and severity of black root rot. Canadian
Journal of Plant Pathology. 11:68-74.

Sang, J. P., I. R. Minchinton, P. K. Johnstone, and R. J. Truscott, 1984. Glucosinolates
profiles in the seed, root and leaf tissue of cabbage, mustard, rapeseed, radish and
swede. Can. J. Plant Sci. 64:77-93.

Sarwar, M. and J. A. Kirkegaard. 1998. Biofumigation potential of brassicas. II. Effect of
environment and ontogeny on glucosinolate production and implications for
screening. Plant Soil 201: 91—101.

Scholte, K. 1987. The effect of crop rotation and granular nematicides on the incidence of
Rhizoctonia solani in potato. Potato Res. 30: 187—199.

Sequeira, L. 1962. Influence of organic amendments on survival of F usarium oxysporum
f. cubense in the soil. Phytopathology. 52:976-982.

Shetty, K. G., K. V. Subbarao, 0. C. Huisman, and J. C. Hubbard. 2000. Mechanism of
broccoli-mediated Verticillium wilt reduction in cauliflower. Phytopathology.
90:305-310.

Singh, R. S., and K. Sitaramaiah. 1970. Control of plant-parasitic nematodes with organic
soil amendments. PANS 16: 287—297.

Snapp, S., D. Smucker and M. Vitosh. 2002. Nitrogen Management for Michigan
Potatoes. Extension bulletin E-2779, New, March. Michigan State University.

Stapleton, J. J. and R. A. Duncan. 1998. Soil disinfestation with cruciferous amendments

and sublethal heating: Effect on Meloidogyne incognita, Sclerotium rolfrii and
Pythium ultimum. Plant Pathol. 47: 737—742.

73

Subbarao, K. V., J. C. Hubbard, and S. T. Koike. 1999. Evaluation of broccoli residue
incorporation into field soil for Verticillium wilt control in cauliflower. Plant
Disease. 83: 124-129.

Sumner, D. R., S. C. Phatak, J. D. Gay, and R. B. Chalfant. 1995. Soilbome pathogens in
a vegetable double-crop with conservation tillage following winter cover crops.
Crop Protection. 14(6): 495-500.

Tsao R., M. Reuber, L. Johnson, J.R., Coats. 1996. Insecticidal toxicity of glucosinolate-
containing extracts from crambe seeds. J Agric Entomol. 13:109—120.

Tuitert, G., M. Szczech, and G.J. Bollen. 1998. Suppression of Rhizoctonia solani in
potting mixes amended with compost made from organic household waste.
Phytopathology 88:764—773.

Walker, G. 1997. Effect of Brassica residues and other organic amendments on
abundance and sex ratio of Tylenchus semipenetrans in soil. Australian Journal of
Experimental Agriculture. 37:693-700.

Weinert, T., W. L. Pan, M. R. Moneymaker, G. S. Santo, and R. G. Stevens. 2002.
Nitrogen recycling by non-leguminous winter cover crops to reduce leaching in
potato rotations. Agron. J. 94:365-372.

Workneh, F ., A. H. C. Van Bruggen, L. E. Drinkwater, and C. Sherman. 1993. Variables

associated with corky root and Phytophthora root rot of tomatoes in organic and
conventional farms. Phytopathology. 83:58 1 -5 89.

74

Chapter 3: Bioassay to test suppression of Rhizoctonia solani, Pythium ultimum and

F usarium solani by Mustard (Brassica juncea L.) and Rye (Secale cereale L.) tissues.

ABSTRACT

Bioassays are needed to assess the suppressive effects of cover crop residues on
soil borne diseases. We experimented with 2 bioassays to study this. The first was a jar
bioassay where we investigated the suppressive effect of 10 g mustard and rye tissues
(root, shoot and root plus shoot) on Pythium ultimum, Fusarium solani and Rhizoctonia
solani. Following this, a dose response bioassay was done to study effect of different
amount of mustard and rye tissue on inhibiting growth of the 3 fungi. Here the dosage
was 10 g, 5 g and 1 g. The tissues were macerated and put inside glass jars. A plug of R.
solani, P. ultimum and F. solani was placed in the center of a 100 mm Petri dish
containing fresh Potato Dextrose Agar. The Petri dish was then inverted over the mouth
of the glass jar containing the above mentioned treatments and fungal growth on the PDA
plate was monitored. This was expressed as % of growth on the Petri dish. The second
bioassay was a tuber bioassay which was done thrice over the entire growing season
based on date of cover crop incorporation. Tubers were placed in 9 * 8 * 2 in clear
containers in soil collected from the individual field treatments presented in chapter 2.
The tubers were scored for fungal infection and suppressive effects of the residue
amended soil were monitored. Mustard residues had markedly suppressive effect on
growth of all three fungi as compared to the no residue control. At 48 h, treatment with
mustard shoot had no P. ultimum growth while the no residue control had 100 % growth
on the petri dish. A similar trend was observed with no residue control showing 100 % (at

48 h) and 8.36 % (at 120 h) growth for R. solani and F. solani respectively, while

75

mustard shoots showed 11.3 % growth for R. solani and 0 % growth for F. solani. The
mustard root+shoot treatment had significant effect in suppressing fungal growth for all 3
fungi as compared to the no residue control. In all three cases, the effect was more
pronounced at 24 h than at 48 h (P. ultimum and R. solani) or 120 h (F solani). For the
dose response bioassay, overall 10 g mustard residues were most effective in controlling
fungal expansion. For P. ultimum, at 48 h, 10 g mustard shoots showed 17.4 % growth as
compared to 56.5 % grth shown by 1g mustard shoot. Similarly trend was seen for
treatment with 10 g mustard shoot which exhibited 18 % growth while 1 g mustard shoot
showed 100 % growth for R. solani at 48 h. Overall as residue biomass was increased, the
suppressive effective increased. For the tuber bioassay, the treatment with mustard root +
shoot had healthiest tubers (rating 0.75) which were comparable to the fumigated control
(rating 1.00). Overall effect of the cover crop and tissue interaction was significant at 7
days incorporation (P < 0.0001) but at 40 days, the interaction was not significant (P <
0.4377). When planned contrasts between different treatments were drawn, it was
observed that there was no significant difference between mustard and firmigated.
Mustard versus rye showed significance (P<0.0001) at both 7 days and 40 days

incorporation.

76

INTRODUCTION

Soilbome plant pathogens survive in the soil for very long periods of time and
cause extensive damage to many crops. For many years the most common practice for
their control was fumigation either before or after cropping. Unfortunately certain
fumigants possess negative attributes, such as health hazards, environmental pollution,
and even potential atmospheric ozone depletion. Taking all these factors into
consideration, there is now increased environmental concern which has triggered
regulatory restriction on the use of soil fumigants.

The induction of disease suppression in soils has been successfully achieved using
organic and fertilizer amendments (Hoitink and Fahy, 1986), or cultural practices, such as
mulching, which it is believed, stimulates aggressive competition for food substrate
between soil inhabitants in the rhizosphere (Baker and Cook, 1974 and Chen et al., 1987),
the most biologically active fraction of the soil (Lynch 1990, and Brussaard et al., 1990).
The disease suppressive activity of manures and composts can also be attributed to the
production of volatile and non-volatile toxic compounds released during plant residue
decomposition, or to changes in the proportion of pathogen-antagonistic microorganisms
in the rhizosphere (De Brito Alvarez et al., 1995 and Chen et al., 1987).

Incorporation of green manure, rotation crop residues, or other organic materials
is frequently recommended to prevent the increase of pathogens in newly cultivated soil
and to produce conditions less favorable to pathogens in established crop land (Butler,
1961). In the US. approximately 1 billion tons of organic and inorganic agricultural
recyclable by-products are generated yearly out of which 400 million tons are crop

residue (Edwards and Someshwar, 2000). These residues can be effectively used for

77

agricultural purpose with great potential benefits. High yields for extended periods of
crop cultivation in areas of China were associated with the use of organic sources of
fertilizer or waste products (Kelman and Cook, 1977 and Shen, 1997). Such amendments
contribute to root health by improving soil structure and reducing the negative impact of
soil-borne pathogens. Davis et al., 2001, examined 100 commercial potato fields for soil
characteristics, disease, and yield and found that the factors most closely related to soil
quality, i.e. organic matter, organic nitrogen and increased nutrient availability, were
associated with reduced Verticillium wilt and higher tuber yields.

There are many factors involved in the mechanism of disease suppression during
decomposition of amendments in soil. One factor of considerable interest is the effect of
volatile compounds evolved during decomposition of amendments in soil on pathogens
(Angus et al., 1994). Secondary products from higher plants represent an enormous
diversity of biologically active compounds that can be exploited as fumigants when plant
residues are incorporated as green manure. Glucosinolates, a group of secondary
metabolites are present in the seeds and vegetative tissue of many plants belonging to the
family Cruciferaceae. Good controls of Pythium ultimum, F usarium oxysporum f. sp
cumini and Rhizoctonia solani have been seen by amending the soil with leaves of
Brassica species. (Ramirez-Villapudua and Munnecke, 1988; Charron and Sams, 1999;
Marwar and Lodha, 2002). In biofumigation, plant based glucosinolates are hydrolyzed in
field soil to form toxic products like isothiocyanates, thiocyanates, nitriles,
oxazolidinethiones, and ionic thiocyanate (Brown and Morra, 1997). Glucosinolates are
usually present in the leaves of Brassica spp. at concentrations that can yield bioactive

catabolites in amounts sufficient to prevent development or spread of certain pathogens

78

(Sang et al., 1984; Kirkegaard 1998 and Sarwar 1998). Studies have been reported as
early as 1937 relating mustard oil and other volatile substances to plant disease resistance
(Walter et al., 1937). Glucosinolates and their break-down products are known to be
involved in conferring resistance to certain pests and diseases in plants (Donkin et
al.,1995, Gidamoustaris and Mithen 1995, Ludwig-Muller et al., 1997). A study by
Papavizas in 1966 determined that several cruciferous amendments when incorporated
into soil reduced Aphanomyces root rot of peas considerably. Field experiments during
the early 1990's demonstrated that wheat crops grew more vigorously following Brassica
break crops such as canola and Indian mustard than other break crops such as linseed or
oats (Angus et al. 1991, Kirkegaard et al. 1994). Crucifers grown as cover crops offer
several advantages. They grow quickly, resist cold fall temperatures, are excellent
nitrogen accumulators, the seeds are relatively inexpensive and they are winter killed.
Most importantly they have been proven to be an effective control against soil diseases
such as scab and rhizoctonia (Lewis and Papavizas, 1974).On'ental mustard (Brassica
juncea L.), a variety known commercially as Pacific Gold, belongs to the family
Cruciferaceae and provides an excellent combination of traits: medium height, high
biomass production, high glucosinolate level, and late flowering (Dan Bryant, Western
Farm Press, 2003). Soil-bome fungal disease suppression is more likely to be improved
by using a variety high in glucosinolate. Oriental mustard is high in its glucosinolate
content.

Macerated tissues of Brassica species have been shown to reduce disease
incidence. Anti fungal activity associated with Brassica residues is often attributed to

release of isothiocyanates from the plant tissue. Isothiocyanates are break-down products

79

of glucosinolates. Brassica juncea completely suppressed activity of Pythium ultimum
and Rhizoctonia solani in a study by Charron and Sams, 1999. Rhizoctonia solani, in a
container study, was also suppressed by cabbage volatiles (Lewis and Papavizas, 1974).
In a study by Pavlica et al., 1978, volatile compounds have also been shown to reduce the
number of fungal propagules. Gamliel and Stapelton (1993) determined that Pythium
ultimum propagules were reduced by >95% when exposed to volatiles from heated
cabbage amended soil.

In this chapter, we studied 2 bioassays which investigated effect of oriental
mustard (for its biofumigant product) and wheeler rye (for its allelopathic property) on 3
soil fungi — Pythium ultimum, Rhizoctonia solani and Fusarium solani. In this study it
was our aim to study effect of different tissues of the 2 cover crops. For this, 2 bioassays
were done to primarily investigate the effect of tissues on potato (Solanum tuberosum L.)
roots and tubers. From the 2 bioassay’s we established a relation between incidence and
infection of fungal diseases and effect of mustard and rye cover crops; their tissues; and
amount of tissue.

In the following study, we experimented with 2 bioassays (jar bioassay and

container tuber bioassay) to test the following hypotheses;

80

Objectives
Jar Bioassay
a) Mustard root+shoot will suppress development of all three fungi.
b) Mustard shoot only will be as effective as the mustard root+shoot in
controlling disease growth.
c) Over time, effect of the treatments will wane.
(1) Rye root+shoot will give noticeable results earlier which will subside over
time.
c) A higher dose of mustard treatment will be more effective at restricting fungal
proliferation.
Container Tuber Bioassay
a) Soil from the mustard field experiment treatments will show similar effects on
tubers.
b) Tubers in sterile soil will show absolutely no fungal infection
c) Tubers placed in soil inoculated with each of the fungi will show very

distinctive infection symptoms.

81

MATERIALS AND METHODS
JAR BIOASSAY
Plants used and growing conditions:

Oriental mustard (Brassicajuncea L.) and Cereal rye (Secale cereale L.) were the
two plants used in this bioassay. The variety of mustard that was used is commonly called
‘Pacific Gold’. Plants were grown in 17 x13 x 6 in. plastic tubs. Seeds were sown at twice
the field seeding rate (22.5 kg/ha) to get more biomass. The soil used was a mixture of
sandy soil + 40% perlite. The soil was collected from a research trial field (Sandhill) in
East Lansing, Michigan. The soil was steamed at 70° C for one hour, and then sieved
with a 5 mm sieve before mixing with perlite. Plants were grown in the greenhouse at
Michigan State University at 28°C. The light was maintained at 16 h daylight period
using halogen lights. The fertilizer used was- Osmocote®, which is a slow release plant
food with 14-14-14 N, P, K. All plants were watered uniformly. Both roots and shoots of
the plants were harvested just before flowering. They were washed with diStilled water
and cut into small pieces. A compound sample from a large number (10-15 plants) of
plants was used in the bioassay. The tissue samples were used immediately after harvest.
Pathogen culture:

The three fungi tested in this bioassay were Rhizoctonia solani, F usarium solani
and Pythium ultimum. All three were obtained from Department of Plant Pathology at
Michigan State University. R. solani and F. solani were cultured on 39 g Potato dextrose
agar (PDA) mixed with 1 L distilled water and autoclaved for 30 mins. 25 g/L ampicillin
(Sigma-aldrich chemicals) was added to the PDA to eliminate bacterial contamination.

Given its sensitivity to ampicillin, P. ultimum was cultured on PDA without ampicillin.

82

The fungi were cultured at room temperature in aseptic conditions under the hood. R.
solani was cultured in dark while the other two were cultured in light.
Fungal response to plant:

Residues from mustard and rye root and shoot and the control were tested. The
residues were chopped with scissors into 2 cm pieces. The amounts of residues used
were- 10 g plant root, 10 g plant shoot, 10 g root + shoot, and control (no plant tissue).
The tissues were macerated using a mortar and pestle and put inside 400 ml glass jars. A
5 mm plug of R. solani, P. ultimum and F. solani was removed from 10 day old cultures
and placed in the center of a 100 mm Petri dish containing fresh PDA. The Petri dish was
then inverted over the mouth of the glass jar containing the above mentioned treatments.
The glass jar and the Petri dish were sealed at the junction using two layers of parafilm.
The whole procedure was done in the hood under aseptic conditions. Fungal growth was
monitored.

Dose Response Curve:

In addition to the above experiment, another experiment was done to see response
of R. solani to different amounts of plant root and shoot. Rye and mustard were grown
and processed in the same way as given above. The treatments used for this experiment
were 10 g, 5 g, and 1 g of plant shoot, 10g, 5 g, and 1 g of plant root and 10 g, 5 g, and 1
g of plant shoot + root and control (no plant tissue). A 5 mm plug was removed from a 10
day old culture of R. solani and placed in the center of a 100 mm Petri dish containing
fresh PDA. The Petri dish was then inverted over the mouth of the glass jar containing

the above mentioned treatments. The glass jar and the Petri dish were sealed at the

83

junction using two layers of Para film. The whole procedure was done in the hood under

aseptic conditions. Fungal growth was monitored.

 

Fungal Plug
Parafilm

Glass jar

Petri dish

Macerated Tissue

 

 

 

 

Figure 3.1. Jar bioassay. Experimental set up for bioassay

 

Figure 3.2. Jar bioassay. Experimental set-up for dose response of fungi to mustard
tissue. Figure shows fungal growth reaction to different amounts of mustard shoots.

a) 10 g mustard shoot, b) 5 g mustard shoot, c) 1 g mustard shoot, d) bare.

84

Recording fungal growth:

Radial mycelial growth of the fungi was recorded at 24-h intervals for 72 h for P.
ultimum and R. solani as the mean of 2 perpendicular diameters. Since the growth rate of
F solani is lower than P. ultimum and R. solani, it was monitored at 24 h, 48 h and at 120
h. Fungal growth was expressed as a percentage of area growth on Petri dishes in control
jars without plant material.

Testing biocidal activity of Mustard

The fimgal plugs were removed from the Petri dishes and were transferred to
fresh PDA media outside the glass jars. Fungal growth was monitored for P. ultimum, R.

solani and F. solani for a period for 48 h.

85

CONTAINER BIOASSAY
Container Specifications

This experiment was done in the greenhouse. The container was a clear plastic
box 9 X 8 X 2 inches in volume. This is a standard size box manufactured by Pactive-
ClearView® Sell Out®. The boxes were drilled with 4 holes on the top to allow air
circulation.

Soil and treatments

Soil was collected from the same plot where the mustard research trial was in
progress (Sandhill). The bioassay monitoring was repeated thrice over the entire growing
season from samples taken on 10‘h June 2004, 22nd June 2004 and 22"d July 2004 to
reflect field conditions during those periods. These time periods were based on the day
when the cover crops were incorporated (15th June 2004). Soil was sampled from
treatment 1 to 7 which are listed in table 3.1. 6 subsamples were taken from each plot
which were composited and used for the study. The soil was sampled at 0-4 in depth.

For the first 7 treatments, soil was collected directly from the corresponding plot
in the field experiment. For treatment 8, soil was collected from an area outside the
research plot. This soil was sterilized by autoclaving at 82 C for 30 min. This was doneto
ensure that the soil was completely free of all pathogens. Treatment 9 was divided into 3
sub treatments. Each sub treatment was inoculated with different fungi. The 3 fungi used
in this experiment were Pythium ultimum, Rhizoctonia solani and F usarium solani. Each
fungus was used from the same batch that was used to inoculate the field. The inoculum

rate used was twice that used in the field to ensure infection in those positive controls.

86

 

Table 3.1. Container tuber bioassay. Treatments used for bioassay.

 

 

 

 

 

 

 

 

 

 

 

 

 

N0 TREATMENTS
1 Rye Root
2 Rye Shoot
3 Rye Root + Shoot
4 Mustard Root
5 Mustard Shoot
6 Mustard Root + Shoot
7 Fumigated
8 Sterile Soil
9 Inoculated Soil (P. ultimum, R.
solani and F. solani)

 

Tubers

The tubers used were from the same batch that was used to plant in the field.
Tubers were washed with distilled water; surface sterilized with 5 % Clorox® bleach and
then again washed thoroughly with distilled water. Care was taken to make sure that
tubers were of equivalent size.
Procedure

The soil from the above-mentioned treatments was put inside the containers, the
volume of soil being 100 cubic in. Each tuber was cut into 2 halves and 2 tubers were
placed on top of the soil in the containers. The cut surface facing the soil and the soil was
compacted around each half tuber to provide maximum contact between the soil and the
tuber (Figure 3.3). The containers were watered everyday to maintain uniform moisture
as judged by moisture condensation on top of lid and on appearance of the soil. The

experiment was monitored for 10 days. Pictures were taken at the end of the 10th day and

87

tubers were rated on the basis of degree of infection. The ratings were 0, 1, 2, 3, 4, and 5

with 0 being no infection at all and 5 being completely rotted (Figure 3.4)

 

Figure 3.3. Container tuber bioassay. Experimental set-up of container tuber

bioassay. (1) container used for the experiment. (2) randomized complete block

design in the greenhouse.

88

 

3 4 I 5

Figure 3.4. Container bioassay. Scale used for rating tubers.

Replications and Statistical analysis
Jar Bioassay

The experimental design was a completely randomized block with three
replications of each combination of fungus and plant residues or control.
For dose response curve experiment: A completely randomized design with four
replications of each treatment was used. A one way ANOVA was used to test growth

response to different amounts of plant tissue.

89

ANOVA with 2 factors (covercrop and tissue type) for each fungal species was
used to test growth response to plant tissue. Since the growth was monitored over time,
repeated measured was used as a way of analyzing the data. To see the effect of
individual treatments, planned contrasts were used. The contrasts were 1) bare versus
mustard 2) bare versus rye and 3) mustard versus rye. Each of these contrasts were
compared on the basis of the hours of incubation.

Container tuber bioassay

The experimental design was a completely randomized block design with 9
treatments and 3 replications each with 4 tubers and 12 observation units. ANOVA with
2 factors (cover crop and tissue type) was used to analyze the data. For calculating the

least square means and planned contrasts were calculated.

90

RESULTS

JAR BIOASSAY

Pythium ultimum — In this experiment 10 g of each tissue and tissue combination was
used to test effect on P. ultimum. At 24 h, mustard root+shoot and mustard shoot only,
completely inhibited growth of fiingi. This effect is attributed to presence of high levels
of volatile glucosinolates released from cut up mustard shoot residues which suppressed
fimgal growth. Mustard root also restricted growth at 24 h but at 48 h, 7.5% (Table 3.2)
of the petri dish was covered with P. ultimum indicating that fungal growth suppression
effect was more temporary with root residues. This could be related to lower
concentration of glucosinolates present in mustard root, which lost anti-fungal properties
by the 2"“I day. Rye root had a similar effect to mustard root in restricting fungus growth
to 0.75% area of the Petri dish covered with mycelia (Figure 3.5). Rye root limited
fungal growth to 8% at the end of 48 h (Table 3.2). In comparison, no residue controls
showed P. ultimum growth on 16% area of the Petri dish at 24 h and 100% growth at 48 h
(Figure 3.6). Treatment with rye shoot and rye root plus shoot did not have as inhibitory
effects as rye roots which apparently contain allelopathic compounds. At 48h, mustard
shoot completely inhibited P. ultimum growth (Table 3.2) and mustard root+shoot limited
fungal expansion to 5% of the petri dish (Figure 3.5). Since the treatment with mustard
root+shoot had 5 g root and 5 g shoot, it is possible that lower amounts of glucosinolates
could have allowed growth of P. ultimum to a certain degree. The ANOVA’s are

presented in table 3.2 and the planned contrasts are shown in table 3.3.

91

 

P. ultimum at 24 h

801

.h
C

N
O
4

 

 

 

% area of Infected plate in cm2
0)
O

O

 

 

 

 

 

I 120 1
100 _ g P. ultimum at 48h 1

1
80 -

601

 

 

 

 

 

 

 

% area of infected plate in cm2

 

 

 

Figure 3.5. Jar bioassay. Growth observed at 24 h and 48 h for Pythium ultimum

Fungal growth expressed as % area of growth on plate.

92

 

Figure 3.6. Jar bioassay. Growth of Pythium ultimum on Petri dish. (la) Mustard
Root+Shoot at 24 h, (1b) Bare at 24 h, (2a) Mustard Root+Shoot at 48 h, (2b) Bare
at 48 h.

93

AN OVA Pr > F
Cover crop

Tissue

Cover crop X Tissue
Time

Cover crop X Time
Tissue X Time

Cover crop X Tissue X Time

Table 3.2 Jar bioassay. Analysis of variance for Pythium ultimum.

< 0.0001
< 0.0001
< 0.0001
< 0.0001
< 0.0001
< 0.0001
< 0.0001

 

NS: Not Significant at P < 0.05

Table 3.3 Jar bioassay. Planned Contrasts at 24 h and 48 h for Pythium ultimum.

 

Pr > |t|

Parameter 24 h
Bare v/s Mustard <0.0001
Bare v/s Rye <0.0001

Mustard v/s Rye <0.0001

48 h

<0.0001
<0.0001
<0.0001

 

NS: Not Significant at P < 0.05

94

F usarium solani - In this experiment 10 g of tissue was used to test effect of residues on
F. solani. The results were observed at 24, 48 and 120 h because F solani is a slow
growing fungus. Mustard root and mustard shoot completely inhibited expression of F.
solani throughout the experiment (Table 3.5), while neither of the rye treatments had a
pronounced effect on inhibiting growth of F. solani as compared to the mustard treatment.
Mustard root+shoot controlled fungal growth to a certain extent initially and succeeded in
limiting its growth substantially after 120 h (Table 3.5). It was observed that no residue
control showed no fungal growth at 24 h; at 48 h, the growth was 0.63 % and at 120 h
fungal expansion had increased to 8.36% (Figure 3.7). The ANOVA’s are presented in

table 3.5, and the planned contrasts in table 3.4.

Table 3.4 Jar bioassay. Planned Contrasts at 24 h and 48 h for F usarium solani.

 

Pr > M

Parameter 24 h 48 h 120 h
Bare v/s Mustard 0.9534 NS 0.3090 NS < 0.0001
Bare v/s Rye 0.4912 NS 0.8256 NS <0.0001

Mustard v/s Rye 0.3739 NS 0.2595 NS 0.0094
NS: Not Significant at P < 0.05

 

95

Table 3.5 Jar bioassay. LS Means with Standard Errors and analysis of variance at

24 h, 48 h and 120 h for F usarium solani.

 

LS MEANS

Bare

Rye Root

Rye Shoot

Rye Root + Shoot
Mustard Root
Mustard Shoot
Mustard Root + Shoot
ANOVA Pr > F
Cover crop

Tissue

Cover crop X Tissue
Time

Cover crop X Time
Tissue X Time

Cover crop X Tissue X Time

24h

0.00 (3: 0.00)
0.12 (c 0.04)
0.33 (c 0.33)
0.72 (e 0.28)
0.00 (e 0.00)
0.00 (c 0.00)
0.10 (c 1.10)

0.8687 NS
0.0011
0.0089
<0.0001
0.9634 NS
0.002
0.0311

48h

2.63 (c 0.14)
0.18 (e 0.02)
0.80 (c 0.80)
0.84 (:1: 0.18)
0.00 (e 0.00)
0.00 (c 0.00)
0.14 (e 0.14)

120 h

8.36 (i 0.58)
0.20 (c 0.07)
1.84 (:l: 1.84)
1.34 (.1: 0.64)
0.00 (:1: 0.00)
0.00 (:1: 0.00)
0.18 (:l: 0.08)

 

NS: Not Significant at P < 0.05

96

Rhizoctonia solani — All treatments used 10 g of tissue in this bioassay to observe residue
tissue effect. At 24 and at 48 h maximum R. solani growth was observed in the no residue
treatment and minimum in the treatment with mustard root+shoot (Figure 3.7). Although
mustard shoot had twice the amount of shoot than in treatment with mustard root+shoot,
mustard root+shoot was more effective at controlling fungal growth. This effect could be
related to some interaction between the roots and the shoots. At end of 48 h, control, rye
root, rye shoot and rye root+shoot showed similar levels of fungal growth — 100% (Figure
3.7). All the mustard treatments were very effective in limiting growth of R. solani,
indicating the potential to use mustard can be an effective tool in controlling R. solani
infection in the field. The ANOVA’s are presented in table 3.7 and the planned contrasts

are shown in table 3.6.

Table 3.6. Jar bioassay. Planned Contrasts at 24 h and 48 h for Rhizoctonia solani.

 

Pr > M

Parameter 24 h 48 h
Bare v/s Mustard <0.0001 <0.0001
Bare v/s Rye <0.0001 <0.0001
Mustard v/s Rye <0.0001 <0.0001

 

NS: Not Significant at P < 0.05

97

Table 3.7 Jar bioassay. Analysis of variance for Rhizoctonia solani.

 

ANOVA Pr > F

Cover crop < 0.0001
Tissue < 0.0001
Cover crop X Tissue < 0.0001
Time < 0.0001
Cover crop X Time < 0.0001
Tissue X Time < 0.0001
Cover crop X Tissue X Time < 0.0001

 

NS: Not Significant at P < 0.05

98

 

R. solani at 24 h

.3
O
O

60

A
O

N
O

 

 

% area of infected plate in cm2
0

 

 

 

 

1 120
{NE 100 1 a a a R. solanr at 48 h 1
1
° 1
.5 80 1
1 *3 C 1
1 a 60 -
1 E 40 — 1
O
1"" e
r: 20 1
1 ‘5 9h hr l
l 3 0- 1
'- \ \. \- \ ‘K.
1 «I «9 o o o 6* o o 1
.\° a” 09° 98° 98° :39 gc° 94° 1
A 0 \x '0‘ «P x l
‘1 Q. Q$ 00 04} é? 06" 1
1 Q. ‘3 ®° 6Q~ '
1 *0 6‘ l
‘ 1
1
l

1 Treatment $59

Figure 3.7. Jar bioassay. Growth of Rhizoctonia solani on petri plate observed at 24

h and 48 h. Fungal growth expressed as % area of growth on plate.

99

JAR BIOASSAY FOR DOSE RESPONSE

Pythium ultimum — The dose response of mustard treatments on P. ultimum showed a
similar trend for all tissues and tissue combinations. Figure 3.8 shows that the infection at
48 h was greater in all treatments than at 24 h. The more the amount of tissue used, the
lesser was the fungal growth. This was true for all treatments (Figure 3.8). The amount of
glucosinolate present in shoots is significantly greater than the amount present in roots.
This explains greater inhibition of fungi in treatments involving mustard shoot. The best
treatment for controlling P. ultimum was 10 g of mustard shoot at 24 and 48 h. This could

be related to this treatment having maximum concentration of glucosinolate.

F usarium solani - Since this is a slow growing fungus, the effects were observed at 24, 48
and 120 h. At all times, 10 g of tissue inhibited fungal growth the most (Figure 3.9). This
can be explained by the fact that more the amount of tissue, greater is the glucosinolate
concentration. Comparing figure 3.9 and table 3.5, it is noticed that the mustard root and
mustard shoot only treatments in this experiment did allow some growth of F. solani,
though this was much lesser than growth in the bare treatment (Figure 3.9). Analysis of

variance for P. ultimum, and F. solani are shown in table 3.8.

100

85 v a a “583% “oz Hmz

 

 

 

 

mz ammo 885 25
X 250:3. X 0:32.
885v mz 825 25 x .5285.
:55 ~83. 25 x 22H
88.9 885 v 22:
mz N83 #86 25225. x 32F
88.? 885 v “582
$56 88o v 2522
.7...— <>cz<
2: no 3 as S _ o o o ”.2 new 2: «.2 SN 23 sfifséesgz
82 S E :3 8.0 E o o c 4.: em new 8.0 med SN 825 2282
8.0 2 mm woo 3o 5 o o o as v.8 2: 3... 2m 3 822382
@7222 m.—
3: mm 3 3: mm m: we mm 3 we mm 3 we 3 3
.82 is fa is 24m
mflSQa. E3mh§§h~ £§§§= 53.2%..”

 

Emu—v EDA— flc latch” .«O NOH< e\e

 

$3233 .3.— 3.3.—mo.—

omec 3 .353. 553.3% can £35.53 555$ .Su 353.3.» he amen—ans 6:5 2:88 9:33 .maoa— S333: nan. .w.m oEfiH

101

1 __ "126—

 

 

 

 

 

 

 

 

 

 

1 Dose Response of P. ultimum to Mustard Root
.3; 100 ~
3 k +
3 80 — 1
8 “E .
5 E “* 1
2 .0. 1-22—2-21 1
5 -D— 24 h 1 1
"\° 2° 1 1+5§IU 1
1 o . , 1
I 10 g 5 g 1 g '
1 _ _*____ _ Weight of_ Root_in_g__ _-__ ___f_ i,
t 100

 

Dose response of P. ultimum to Mustard Shoot 1
43‘— 24 h1 1
12:11"

J

    

% area of infected plate
In en?
8

 

 

 

 

 

1

4o. 1

2° / 1

D— 1

o T ’ 1
10 19

 

9 5 9 ' .
Weight of shoot in g i

 

1

 

Dose Response of P. ultimum to Mustard

100 1 Root+Shoot

    

% area of Infected
plate in cm:

888

N
O

 

 

O

 

*r

10 g 5 g 1 9
Weight of Root+Shoot in g

 

 

Figure 3.8. Dose response of Pythium ultimum to mustard tissue over time.

102

 

 

 

   

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

1 5 T1
Dose Response of F solani to Mustard 1
u 4 . 1 +— 1091 Root “A 1
g u —D—59 1 1
‘2’. 53 q ...‘... 1g 1 3." 1
5' «5 __-_ 1
o
g g2 ~ 1
a a. 1
.\ 1 fl 1
1
, 0 1 1 1
1 24h 48h 120h 1
1 Time _J
1 “ ‘5’ ”’1
Dose Response of F. solani to Mustard 1
1: 4 1 W Shoot 1
8 .. 1 1
33 E 3 1.0.59 1
c o A 19 1
.7. .E " 1
o
a 13?. 2 1
a a 1
o\° 1 “ 1
1 ° 1
1 24h 48h ' 120h 1
L--.__ _ m _____- Tlme _ _ 1
5
Dose Response of F. solani to Mustard
5 4 _ 109 Root+Shoot
C
:3 NE +59 1
U u 3 . - ‘ -~ 19 1
.2 Lw______._1
5 .E 1
E 332 1
2 .- ma 1
N 1 A 1 f4 1
=3
o 1 . 1
24h 48h 120h ;
Time 1

 

 

 

Figure 3.9. Dose response of F usarium solani to Mustard tissue over time.

103

Rhizoctonia solani — In this experiment along with mustard, effect of rye on R. solani
was also studied. In case of mustard tissues, fungal response to dose was most clearly
seen with shoot tissue (Table 3.9). It was less obvious in mustard root + shoot. In mustard
shoots, 10 g of tissue inhibited growth of R. solani best, while 1 g of tissue gave the
poorest result. Except for shoots, in all the other cases by 48 h, R. solani had completely
colonized the entire petri plate. With rye tissues, the trend was not as clear. At the highest
dosage, which is with 10 g of tissue, rye root + shoot provided most growth suppression

at 24 h (Table 3.9). Analysis of variance for R. solani are shown in table 3.9.

104

 

good V Seed V 08E. X

 

 

:30: x Rama.
32:. v 823 v 25 x 2525.
82:. v 52:. v 25 x 225
585 v 52; v 25
good v 886 v 2:522 X 0332.
523 v 88.0 v 2225.
52:. v 885 v 225
2 A .5 <>oz<
2: 2: 2: 25. mm 02 EN 2: 2: 2: w: 2.2 822 + 63:
2.2 22 2: Z 2. N: Sm 2: 2: on: 2: 2: 62m
2: 2: 2: 22 S n2. 2: 2: 2: EA mm 23 83:
mz<22 m:
3: mm w: 3: mm m: 3: mm 3 3: mm m:
a 2. 2 E 2 2V 2 E
3 ME

5:. Eng .8 .332. £=e~ue~3fl he 5.39.6 «e 3.3. .x.

 

22233 new 3:27.9— 33. E .353 um=o~ue~3~ 28 353...; he manta—ans ES 2:3:— suasam unsou— Sammaema 2a.. dam sink.

105

CONTAINER TUBER BIOASSAY

In this experiment, the bioassay provided information about soil-borne disease
presence 3 times over the entire season. The first time, before the incorporation of cover
crops, a baseline was established to see effect of existing soil borne disease presence on
tubers. This showed that there was no difference by rep or by treatment in terms of
preexisting soil borne disease. Second time, the bioassay was carried out 7 days and third
time 40 days after incorporation of cover crop tissues. A similar trend was observed 2"d
and 3rd time (Figure 3.13) with most of the treatments. The treatment with mustard root +
shoot had healthiest tubers (table 3.10). These were comparable to the fumigated control
(figure 3.10). In case of rye, the treatment with root + shoot had moderately healthy
tubers (overall rating 1.25). Tubers from this had slightly more disease than tubers from
fumigated control at 7 days incorporation (rating 1.12), but at 40 days (rating 1.37), the
health of tubers deteriorated with this treatment (figure 3.12). Overall effect of the cover
crop and tissue were significant, at 7 days incorporation but at 40 days, the interaction
was not significant (Table 3.10). When estimates between different treatments were
drawn, it was observed that there was no significant difference between mustard and
fumigated. Fumigated control being better than the rye treatment, significant difference
was observed between them (Table 3.11).

Overall from this experiment, it was observed that there was unknown interaction
between the roots and shoots of the 2 cover crops that enhanced suppression of the fungi.
It is possible that the mixed quality of residues from roots+ shoots with a range of

available carbon and nitrogen increased the soil microbial community which proved to be

106

a greater competition for the fungi. In case of mustard, the glucosinolate released by the
shoots proved to be anti-fungal and inhibited growth and infection of fungi.

Figure 3.13 shows a comparison between the different treatments used in this
bioassay. The sterile soil treatment (Figure 3.11e) provided the healthiest tubers with no
infection. Mustard root + shoot (Figure 3.11a) amended soil had comparable results to the
fumigated control (Figure 3.11d) and sometimes was better than the fumigated control.
When soil was inoculated with different fungi, tuber infection by Pythium ultimum

consistently had high levels of rotten tubers (Figure 3.14).

 

Figure 3.10. Container tuber bioassay. Comparison between tubers from 2

treatments. a) tubers in soil with mustard root+shoot as treatment. b) tubers in soil

which was fumigated.

107

Rill‘ .i -i
l’Vi Hi! \i

  

Figure 3.11. Container tuber bioassay. Tubers showing effect of different
treatments. (a) Mustard root + shoot, (b) Rye Root + Shoot, (c) Mustard Shoot, ((1)

Fumigated, (e) Sterile soil, (f) P. ultimum inoculated soil.

108

Table 3.10. Container tuber bioassay. Analysis of variance for tuber rating. Rating

tubers from 0 — 5, 0 being no infection and 5 being completely destroyed. For scale,

refer to figure 3.8.
ANOVA Pr >F
Covercrop <0.0001
Tissue <0.0001
Covercrop X Tissue <0.0001

<0.0001
<0.0001
0.4377 NS

 

NS: Not Significant at P<0.05

Table 3.11. Container tuber bioassay. Planned contrasts between treatments.

 

CONTRASTS TUBER RATING
Pr > |t|
7 days after 40 days after
covercrop covercrop
incorporated incorporated
Root +Shoot v/s Fumigated 0.5536 NS 0.7692 NS
Mustard v/s Fumigated 0.8339 NS 0.1490 NS
Rye v/s Fumigated <0.0001 <0.0001
Mustard v/s Rye <0.0001 <0.0001
Mustard shoot v/s Rye shoot <0.0001 <0.0001

 

NS: Not Significant at P<0.05

109

 

Figure 3.12. Container Tuber Bioassay. Comparison between treatment with rye
root + shoot at, a: 7 days after incorporation of cover crop and b: 40 days after

incorporation of cover crop.

110

 

Tuber Bioassay 1

 

 

 

 

 

 

1
1 a“ a

1

1

6* 1

o .

J“ 1
9 1
Treatment 2*) 1

Figure 3.13. Container tuber bioassay. Tuber infection for different treatments.
Tuber Bioassay 1 = 7 days after incorporation of tissue. Tuber Bioassay 2 = 40 days

after incorporation of tissue. For scale refer to figure 3.8.

111

    

    
    

man 1 -11"

“in-21%;" rusauum

 

.z:

m‘ruuM

HIV! 1

MI l-tlll *«Hl (d)

EH 1"-

NIH/nu:

 

(c)

Figure 3.14. Container tuber bioassay. Comparison between tubers placed in
inoculated soil and tubers placed in sterile soil. a — soil inoculated with Pythium
ultimum, b — soil inoculated with F usarium solani, c — soil inoculated with

Rhizoctonia solani, d — sterile autoclaved soil.

112

DISCUSSION

There have been studies since as early as 1937 which release of anti-fungal
volatiles from mustard (Walter et al., 1937). These volatiles, termed as glucosinolates are
present in tissues of Brassica spp. at concentrations high enough to prevent development
or spread of certain soil pathogens (Kirkegaard 1998 and Sarwar 1998). The suppressive
effect of the volatiles released from root, shoot and root+shoot of Brassica juncea L. on
P. ultimum, R. solani and F. solani was studied with the aid of the above mentioned
bioassays.

Effectiveness of fungal suppression is dependent on species, age and type of
Brassica tissue (Kirkgaard et al., 1996). B. juncea was chosen as it exhibits the highest
fungicidal activity and also has the highest glucosinolate concentration among other
Brassica spp (B. napus, B. rapa, R. sativus, S. alba) as shown by Smolinska and
Horbowicz, 1999.

In the jar bioassay, the treatment with mustard shoots only was most effective in
suppressing growth of the fungi. Similar result was also observed in the dose response
bioassay. This may be due to the fact that the concentration of glucosinolate differs in
different part of the plant and has been shown to be the highest in leaves (Sang et al.,
1984). Mustard root only at 48 h limited growth of P. ultimum (7.56%) and R. solani
(24.32%) but was not as effective as mustard shoot only. This result can be correlated to
concentration of glucosinolate being higher in mustard shoots than in roots.

Growth of P. ultimum was significantly controlled by treatment with mustard
root+shoot. In this treatment, the growth was 0 % at 24 h and was restricted to 5.54 % by

48 h. A similar trend was observed with R. solani and F. solani also where the growth

113

had slightly increased by 48 h. It is possible that the amount of isothiocyanate being
released by the tissues had reduced as time passed. This is consistent with the findings of
Lewis and Papavizas, 1970 who in a container study determined that volatiles released
from leaf and stem of crucifers were maximum at the 1St week and had completely
stopped by week 4. This is also corroborated by Doughty et al., 1996 who found that
most isothiocyanates were released from Brassica spp for the first 9 days after
incorporation of residue. This same effect was observed in the container tuber bioassay.
Soil collected 7 days after incorporation of mustard residues was better at suppressing
tuber infection than the soil collected at 40 days after incorporation. This may explain
maximum reduction in disease intensity soon after incorporation of residue. Although,
this contradicts findings by Lazzeri et al., 2001, which show that fungal growth is not just
limited to glucosinolate and their hydrolysis derived products and addition of fresh
organic material increased fungal population in their study.

Reduction of fungal population immediately afier incorporation of residue has a
significant effect on reducing the overall fungal infection later in the season. As reported
by Papavizas, 1962, sensitivity of a fungus (R. solani) to Brassica is greater during the
early stage of saprophytic colonization of substrate than after it has already established
itself. Initial reduction in population can considerably lower population later in the season
and can be an important factor of disease suppression in the field.

Rye root had mixed effects on the 3 fungi although it was significantly better than
control with no residue at suppressing fungal growth. It is not determined but this may be
due to some allelopathic effect of the root. Overall for all times monitored, control with

no residue gave the least suppression of disease.

114

In the dose response bioassay, a general trend was observed where more the
residue biomass greater was the disease suppression. This is consistent with the findings
of Charron et al., 1999 who determined that higher the isothiocyanate concentration,
greater was the P. ultimum and R. solani concentration. In the container tuber bioassay,
overall a similar trend was seen at 7 days and 40 days after cover crop was incorporated
with the difference being that the tuber infection had on the whole increased at 40 days
after incorporation. Soil amended with mustard shoot (rating 1.25) was associated with

the healthiest tubers. This treatment was comparable to the fumigated soil (rating 1.00).

115

CONCLUSION

We derived the following conclusions from the results of the jar bioassay and container

bioassay:

1)

2)

3)

4)

5)

Mustard shoots in the bioassay were a very effective tool in
suppressing fungal infection even more than mustard root + shoot. In
field study however, mustard root+shoot proved to be equally effective
against P. ultimum, F. solani and R. solani.

Rye roots and shoots together can minimize intensity of fungal
colonization. This can be attributed to certain allelochemical properties
of rye.

Interaction between root and shoot of either mustard or rye may have
increased beneficial microbial population which competed with the
fungus and can decreased infection.

Higher the amount of mustard tissues incorporated, greater will be the
benefits of disease suppression.

Rye should continue to be used as a winter cover crop and growers
should consider trying mustard if soil borne disease is a major problem

in potato production.

FUTURE WORK

It would be useful to derive at a seeding rate for mustard which will be most

beneficial for decreasing incidence of soil fungal pathogens. Effect of the different tissues

on soil microbial communities can be studied. This can be correlated to disease infection.

116

REFERENCES CITED

Angus, J.F., A.F. Van Herwaarden and G.N. Howe. 1991. Productivity and break crop
effects of winter-growing oilseeds. Aust.J.Exp.Agric. 31 :669—677.

Angus, J.F., P.A. Gardner, J.A. Kirkegaard, and J.M. Desmarchelier. 1994.
Biofumigation: isothiocyanates released from Brassica roots inhibit growth of
the take-all fungus. Plant and Soil 162:107—112.

Baker, K.F., and R.J. Cook. 1974. Biological Control of Plant Pathogens, lst ed.
Freeman, San Francisco.

Brown, P.D., and M.J. Morra. 1997. Control of soil-borne plant pests using glucosinolate
containing plants. Advances in Agronomy 61 :167-23.

Brussaard, L., L.A. Bouwman, M. Geurs, J. Hassink and KB. Zwart. 1990. Biomass,
composition and temporal dynamics of soil organisms of a silt loam under
conventional and integrated management. Neth. J. Agric. Sci. 38: 283—302.

Bryant, D., (Sep 8 2003). Biofumigation monitored with Brassica cover crops. Western

farm press. Viewed on 26th February 2004.
http://westemfarmpress.com/ar/farming_biofumigation_monitored_brassica/ind
ex.htm

Butler, F.C. 1961. Root and foot rot diseases of wheat. Agr. Res. Inst. Wagga Wagga,
N.S.W., Australia. Sci. Bull. No.77. pp. 98.

Charron CS. and CE. Sams. 1999. Inhibition of Pythium ultimum and Rhizoctonia
solani by shredded leaves of brassica species. J.Amer. Soc. Hort. Sci.
124(5):462-467.

Chen, W., H.A.J. Hoitink and AF. Schmittenner. 1987. Factors affecting suppression of
Pythium damping-off in container media amended with composts.
Phytopathology 77:755—760.

Chen, W., H.A.J. Hoitink and OH. Tuovinen. 1987. The role of microbial activity in
suppression of damping-off caused by Pythium ultimum. Phytopathology
78:314—322.

Davis, J.R., O.C. Huisman, D.O. Everson and AT Schneider. 2001. Verticillium wilt of

potato: a model of key factors related to disease severity and tuber yield in
southeastern Idaho. Am. J. Potato Res. 78:291—300.

117

De Brito Alvarez, M.A., S. Gagne and H. Antoun. 1995. Effect of compost on
rhizosphere microflora of the tomato and on the incidence of plant growth-
promoting rhizobacteria. Appl. Environ. Microbiol. 61 :194—199.

Donkin S.G., M.A. Eiteman and PL. Williams. 1995. Toxicity of glucosinolates and their
enzymatic decomposition products to Caenorhabditis elegans . J Nematol
27:25 8- 262.

Edwards, J.H., and A.V. Someshwar. 2000. Chemical, physical, and biological
characteristics of agricultural an forest by-products for land applications. In:
Bartels, J.M., Dick, W.A. (Eds), Land Application of Agricultural, Industrial,
and Municipal By-products. Soil Society of America Series Book 6, pp. 1—62

Gamliel, A. and J. J. Stapelton. 1993. Characterization of antifungal volatile compounds
evolved from solarized soil amended with cabbage residues. Phytopathology.
832899-905.

Giamoustaris, A., and R. Mithen. 1995. The effect of modifying the glucosinolate content
of leaves of oilseed rape (Brassica napus ssp. oleifera) and its interaction with
specialist and generalist pests . Ann Appl Biol 126: 347 363.

Hoitink, H.A.J., and PC. Fahy. 1986. Basis for the control of soilbome plant pathogens
with composts. Annu. Rev. Phytopathol. 24:93—114.

Kelman, A., and R.J. Cook. 1977. Plant pathology in the People’s Republic of China.
Annu. Rev. Phytopathol. 15:409—429.

Kirkegaard, J.A., P.A. Gardner, J.F. Angus, E. Koetz. 1994. Effect of Brassica break
crops on the growth and yield of wheat. Aust. J. Agric. Res. 45:529—545.

Kirkgaard, J. A., P. T. Wong, and J. M. Desmarchellier. 1996. In vitro suppression of
fungal root pathogens of cereals by Brassica tissues. Plant Pathology. 45: 593-
603.

Kirkegaard, J. A. and M. Sarwar. 1998. Biofumigation potential of brassicas. 1. Variation
in glucosinolate profiles of diverse field-grown brassicas. Plant Soil 201:71—89.

Lazzeri, L. and L. M. Manici. 2001. Allelopathic effect of glucosinolate-containing plant
green manure on Pythium sp. And total fungal population in soil. HortScience.
36(7):]283-1289.

Lewis, J. A., and G. C. Papavizas. 1970. Evolution of volatile sulfur-containing

compounds from decomposition of crucifers in soil. Soil Biol Biochem 2: 239-
246.

118

Lewis, J.A., and G.C. Papavizas. 1974. Effect of Volatiles from decomposing plant
tissues of pigmentation, growth, and survival of Rhizoctonia solani. Soil
Science. 18(3): 156-163.

Ludwig-Muller J., B. Schubert, K. Pieper, S. Ihmig, and W. Hilgenberg. 1997.
Glucosinolate content in susceptible and resistant Chinese cabbage varieties
during development of clubroot disease. Phytochemistry 44: 407-414.

Lynch, J .M., 1990. The Rhizosphere. Wiley/Interscience, Chichester, UK.

Mawar, R., S. Lodha. 2002. Brassica amendments and summer irrigation for control of
Macrophomina phaseolina and F usarium oxysporum f. sp. cumuni in hot arid
regions. Phytopathol. Mediterr. 41 :45-54.

Pavlica, D. A., T. S. Hora, J. J. Bradshaw, R. K. Skogerboe, and R. Baker. 1978. Volatile
compounds from soil influencing activities of soil fungi. Phytopathology 68: 758—
765.

Papavizas G.C. 1966. Suppression of aphanomyces Root Rot of Peas by Cruciferous Soil
Amendments. Phytopathology 56: 1071-1075.

Patrick, Z.A., T.A. Tousson and L.W. Koch. 1964. Effects of crop-residue decomposition
products on plant roots. Annu. Rev. Phytopathol. 2:267—292.

Ramirez-Villapudua, J ., D. E. Munnecke. 1988. Effects of solar heating and soil
amendments of cruciferous residues on Fusarium oxysporum f. sp. conglutians
and other organisms. Phytopathology 781289-295.

Sang, J. P., I. R. Minchinton, P. K. Johnstone, and R. J. Truscott, 1984. Glucosinolates
profiles in the seed, root and leaf tissue of cabbage, mustard, rapeseed, radish and
swede. Can. J. Plant Sci. 64:77—93.

Sarwar, M. and J. A. Kirkegaard. 1998. Biofumigation potential of brassicas. II. Effect of
environment and ontogeny on glucosinolate production and implications for
screening. Plant Soil 201: 91-101.

Shen, D. 1997. Microbial diversity and application of microbial products for agricultural
purposes. Agric. Ecosyst. Environ. 62:237—245.

Smolinska, U., and M. Horbowicz. 1999. fungicidal activity of volatiles from selected
cruciferous plants against resting propagules of soil-borne fungal pathogens.
147:119-124.

Walter, J .C., S. Morell, and H. H. Foster. 1937. Toxicity of mustard oils and related sulfur
compounds to certain fungi. Amer. J. Bot. 24:536-541.

119

Chapter 4: Effect of organic fertility amendments on soil fungi.

ABSTRACT

This experiment investigated various soil fertility amendments and their effect on
fungal infection of potato (Solanum tuberosum) roots. The experiment was balanced by
amount of carbon inputs applies to the soil with different organic amendments. The field
experiment was repeated in 2003 and 2004. In 2003, at the 1St harvest, the plot without
any soil amendment (bare) had the highest % of diseased root — 54.85%. The lowest root
infection of 26.64 % was for potatoes grown in soil amended with rye + 5000 lb/a
compost. This soil amendment had the highest amount of carbon input to the soil (2390
lb/a). In 2004, the same pattern of root rot suppression was observed. However in 2004,
treatment effects to root rot response were much more evident. At both monitoring time
periods, rye + 5000 lb/a compost was associated with the lowest fungal infection which
was significantly different than bare. A comparison of treatments was conducted on the
basis of Carbon added. The significance of 0 lb/a Carbon, 1200 lb/a Carbon and 2300 lb/a
Carbon were looked at cumulatively at both years. In 2003, there was no significant
treatment effect for any of the contrasts. But in 2004, root rot was significantly higher 0
lb/a Carbon versus 2390 lb/a Carbon. Overall a consistent pattern was noted. The amount
of Carbon input and the quality of residue affected the % of disease infection. The
specific gravity of tubers and the dry weight of potato roots did not exhibit any particular
changes with the soil amendments. A significant difference in yield was observed
between 0 lb/a Carbon and 1200 lb/a Carbon in 2003 while in 2004 this effect was not
apparent. In 2004, 0 lb/a Carbon versus 2390 lb/a Carbon exhibited significant difference

in the yield.

120

INTRODUCTION

Organic amendments in general are thought to suppress soil pathogens, and have
been long used as method of biocontrol (Cook and Baker, 1983). The surface layer (about
0-15 cm in depth) in agricultural soil is the zone of highest organic C and N pools, soil
microbial biomass and microbial activity (Doran, 1987; Janzen et al., 1992; Van Gestel et
al.,l991), and is consequently important for providing nutrients to plants (Campbell,
1978; Paul, 1984; Woods, 1989). Application of cover crops residues or manure are
possible means of improving nutrient dynamics in the surface layer of intensively
managed crop systems.

Cover cropping may influence other aspects of the agroecosystem, either
positively or negatively for agricultural purposes. The mechanism may be biotic (e.g.
microbial interactions, plant pest interactions) or abiotic (e.g. soil physical factors;
Lockwood, 1988). Cover crops and crop rotations can have various effects on soilbome
diseases. Results of greenhouse tests have shown that green manures of cover crops
differed significantly in their suppression of root rot severity and damage to plant growth
(Abawi et al., 2000). In addition, differential effects of green manures of various cover
crops on the severity of root rot and yield have also been observed under field conditions
(Abawi et al., 2000). Incorporated cover crop residues may provide either a food base
which encourages pathogen growth (Phillips et al., 1971) or conversely, some such as
crucifers may actually decrease soil pathogen populations (Lewis and Papavizas, 1971;
Subbarao et al., 1994b). More complex interactions also occur in which cover crops may
cause one soilbome pathogen to increase while suppressing another (Gardner and

Caswell-Chen, 1994). It has been demonstrated that increasing N inputs, either in organic

121

or inorganic form may increase the severity of root rot, probably due to higher
evaporation and plant water stress (Papendick and Cook, 1974; Cook, 1980)

Animal manures have been used as soil amendments in plant production for
centuries. With an increase in the human consumption of poultry meat and eggs in the
USA, more poultry manure and litter is available for disposal on agricultural fields.
Poultry manures and litters are known to have the potential to provide biological control
of nematodes (Gonzalez and Canto-Saenz, 1993, Kaplan et al., 1992; Kaplan and Noe,
1993; Riegel et al., 1996) and soilbome pathogenic fungi (Homma et al., 1979; Osunlaja,
1990). In some research on control of nematodes and soilbome pathogenic fungi, rates
used in greenhouse and laboratory experiments exceeded 20 MT/ha (Conn and
Lazarovits, 1999; Kaplan and Noe, 1993). There has been limited research on the
influence of environmentally acceptable rates of poultry litters on root diseases and
nematodes.

Frequently litter is removed from poultry houses and spread immediately on
fields. Very little composting is done before spreading. The litter may be high in cellulose
and lignin (Fauci and Dick, 1994). The different litters may have an effect on population
densities of Rhizoctonia solani Kuhn, Pythium spp., and other fungi in soil. Poultry litters
may influence population densities of fungi and bacteria that may be saprophytic
competitors or antagonists of soilbome pathogenic fungi (Riegel and Noe, 2000).

Baker (1965) discusses the different stages in the life cycle of a pathogen where
suppression may take place. At spore germination the soil may have fungistasis activity,
that is, the exogenous energy source may be deficit due to competition from other

microorganisms or suppression of germination can be caused by toxic compounds

122

excreted by the soil microflora. However, after germination the growth of the pathogen
and the process of host infection may also be inhibited by competition, antibiosis or
exploitation (Baker, 1965; Schneider, 1982). Suppression of soil-borne plant pathogens
may be enhanced by the incorporation of organic amendmentsinto soil. Sivapalan et al.,
1993 in a field study showed that high compost rates (120 tons per hectare) increased the
total microbial populations and the number of species of fungi compared to the lower
compost rate (80 tons per hectare). Lumsden et al., 1987 studied soils from the Chinampa
region in Mexico and found that their agroecosystem involves management of high
quantities of organic matter thus maintaining a high organic level in the soil which
stimulated soil microbial activity. This indirectly resulted in higher suppression of
Pythium sps in their soils. Similar results were found by Nitta, 1991 where increased
microbial diversity caused greater disease suppression in soil.

Farming practices such as crop rotation, stubble retention and fertilizer use, all
affect ecological niches available for microorganisms. Disease suppression may be tied to
communities of microorganisms associated with a specific substrate, under certain
environmental and management conditions (Hoitink et al., 1991).

A study by Sumner et al., 2002 investigated the effect of continuous applications
of poultry litter on root diseases, nematodes, and weeds with different tillage practices in
vegetable production in Georgia where they did not detect permanent changes in crop
yield or population densities of soilbome pathogenic or saprophytic fungi and root-knot
nematodes with the long-term use of poultry litters. Rhizoctonia solani is a common
saprophyte in soil and many isolates will grow on cellulose (Papavizas, 1970), and some

may grow on lignin (Sherwood, 1970). But in the same study by Sumner et al., 2002,

123

diseases induced by R. solani were not increased by any of the litter amendments in either
tillage system. In contrast to this, in Nigeria air-dried poultry manure reduced F usarium
stalk rot but increased charcoal rot in corn (Osunlaja, 1990). Though the two diseases
here are different, these investigations show the opposite effect that an organic
amendment treatment can have on different soil fungal pathogens.

Rye has been effectively used as a winter cover crop as it provides plenty of
ground cover and holds the soil in place. Besides being one of the easiest crops to grow,
rye has a profusely branched root system which prevents soil compaction in soils that are
annually tilled. Not only that, the extensive root system also enables it to scavenge
nutrients efficiently from the soil profile. Rye, by nature is very competitive. In a study
by Creamer et al., 1997, rye comprised of at least 80% of the above ground biomass just
before kill.

The overall objective of this study was to improve quality and health of potato
roots and tubers and reduce disease incidence, comparing the effect of rye in combination

with other soil amendments.

124

MATERIAL AND METHODS
Plot information

The field experiment was conducted at Sandhill research plot near Michigan State
University in East Lansing, Michigan in 2004. The plot was 54 x 106 m in area. The soil
was very well drained and sandy type. The top soil had been removed 15 years ago and
was then fallow until continuous potatoes 2 years ago. This made the field more
susceptible to potato pathogens. The cover crop grown was winter rye (Secale cereale).
The seeding rate of rye was 34-40 kg/ha. The amount of biomass incorporated into the
soil was calculated by using a standard 0.5 x 0.5 m quadrat. The biomass by dry weight
was determined to be 2820 kg/ha for rye shoot, and 28 kg/ha for rye root.
Treatments

The experiment was monitored in the year 2003 and 2004. The 2 basic treatments
were 1) rye grown as a cover crop and 2) bare soil. In addition to this, the plots were
amended with different amounts of poultry compost and sugar beet pulp. The treatments
were balanced by the amount of Carbon input. The plots were split plots with or without

each amendment. The treatments are listed in table 4.1.

125

Table 4.1. Rye with amendments field experiment. Treatments used in the study

 

 

 

 

 

 

 

 

No. Treatment
1 Bare (+/- 10,000 lb/a compost)
2 Bare (+/- 5,000 lb/a beet pulp)
3 Bare (+/- 5,000 lb/a compost)
4 F umigated
5 Rye (+/- 5,000 lb/a compost)
6 Rye (+/- 2,500 lb/a beet pulp)
7 Bare (+/- 2,500 lb/a beet pulp)

 

 

 

 

Inoculation of the plots

The first year (2003), the plots were inoculated with Pythium ultimum,
F usarium solani and Sclerotinia sclerotiorum. All three were obtained from the
Department of Plant Pathology at Michigan State University. The spore counts were; 4.5
X 105 spores/ml for P. ultimum, 16 X 105 spores/ml for F. solani and 10 sclerotia/g of
soil for S. sclerotiorum. Even though the field was not inoculated with Rhizoctonia
solani, during analysis, it was seen that the potato roots were infected with R. solani. No
infection by S. sclerotiorum was observed. Hence, the next year (2004), the soil Was
inoculated with P. ultimum, F. solani and R. solani, the inoculum levels being 20 x 107
spores/m1 suspension 16 X 107 spores/ml suspension and 490 g dry inoculum/plot (875 sq
ft) respectively. The fungi were cultured on Potato Dextrose Media and then inoculated
on Millet seed. The amount of inoculum needed for the field was determined on a weight

basis. The millet seed was spread evenly on the entire plot.

126

Potato variety and fertilization

The variety of potato used was “Onaway”. Seed pieces (52 g) were planted on
30th June 2004 at 34 in distance within rows and 12 in between rows. Nitrogen fertilizer
was applied in 3 splits at 200 N kg/ha as recommended. 100 N kg/ha was applied at
planting, 50 N kg/ha at hilling and 50 N kg/ha at tuberization. Potassium (0-0-60) and
phosphorous (19-17-0) were applied at 180 K20 and 40 P205 kg/ha respectively before
planting. The plot was irrigated using a traveler irrigation system.
Potato harvest and disease measurements
Destructive harvest measurements were taken twice each season. The first measurement
was taken a month after planting and the second 2 months after planting. At harvest, the
potato vines were killed using two herbicides MatrixTM at 0.98 L/ha and PoastTM at 1.23
L/ha + 2.2 L crop oil concentrate. The last measurement was done at the final harvest.
Twice over the season, a series of soil dilution bioassays were done to study effect of the
treatments on soil fungi. The fungi obtained from these tests were isolated and cultured
on selective media to ascertain the kind of fungal infection. For determining % of root
infection, the potato roots were washed with distilled water and analyzed using the
WhinRI-IIZOTM program (figure 4.1). Fungi were reisolated from the infected roots using
selective media. The tubers were scored visually for infection on a scale of 0-3. The scale
was based on % of infection. 0 was 0-25% infection, 1 was 25-50% infection, 2 was 50-
75°/o infection and 3 was 75-100% infection (figure 4.2). The dry weights of the roots and
tubers were obtained. Yield measurements were taken at final harvest along with tuber

infection and size classification.

127

 

Figure 4.1. Roots scanned by WinRhizo showing different levels of infection. a) 0-

25% infected b) 25-50% infected c) 50-75% infected d) 75-100% infected.

128

 

Figure 4.2. Rye with amendments field experiment. Scale used for rating tubers on
basis of % infection. a) 0 (0-25%), h) l (ZS-50%), c) 2 (50-75%), (I) 3 (75-100%).

129

RESULTS
Infection of Potato roots by soil fungi

Infection of roots by soil fungi was studied twice each year for 2 years. For both
years the 1St harvest and 2nd harvests were 30 and 75 days after planting potatoes
respectively. In 2003, at the 1St harvest (figure 4.3a), the plot without any soil amendment
(bare) showed highest % of diseased root — 54.85%. While the lowest root infection —
26.64 %, for this harvest was for amendment with rye + 5000 lb/a compost. This soil
amendment added the highest amount of carbon input to the soil. This treatment though
was not significantly different than the other amended treatments but % infection wise
was comparable to fumigated with 29.76 % infection. At the 2“d harvest in 2003, the
effects of the treatments were not pronounced, although rye + 2500 lb/a beet pulp gave
the lowest infection —- 57.94% (figure 4.3b). In all the treatments, by the second harvest
the amount of infection had increased considerably. Results for the ANOVA are listed in
table 4.2.

In 2004, effects of the different treatments were much more evident. At the 1St and
the 2'”d harvest, rye + 5000 lb/a compost consistently was associated with the lowest
fungal infection (figure 4.4a and 4.4b); this was significantly different than bare. At the
1St harvest, bare had the highest °/o of infection -— 46.14 % while rye + 5000 lb/a compost
was 12.64 %. By the 2"d harvest % of infection for rye + 5000 lb/a compost was
contained at 16.02 %. Interestingly this treatment had the highest tuber yield. The
fumigated plot in both years was effective at suppressing infection at the 1St harvest but

by the second harvest, the effect of fumigation had waned.

130

The significance of 0 lb/a Carbon, 1200 lb/a Carbon and 2300 lb/a Carbon were
looked at cumulatively at both years at both harvests. The results are given in table 4.3. In
2003, there was no significance for any of the contrasts. But in 2004, 0 lb/a Carbon
versus 2390 lb/a Carbon was significant at both the harvests (table 4.3). % of fungal
infection on roots at 0 lb/a Carbon versus 1200 lb/a Carbon was significant at the 1St
harvest but the effect reduced considerably by the 2"d harvest.

From the results in 2003 and 2004, it was noticed that the amounts of Carbon
input into the soil made a difference on the % of disease infection. It was also noted that
along with the quantity of Carbon, the quality of carbon and the effect it has on soil

microbes can also influence disease incidence.

131

 

 

_-01b/a --12001b/a --23901b/a
_ a Carbon Carbon Carbon

% of diseased root

 

 

 

 

Treatments Q‘Q‘g Q1)“ (a)

 

 

 

100 --0 lb/a --1200 lb/a -- 2390 lb/a
Carbon Carbon Carbon
80 ab a .313

    

%of diseased root
0')
O

 

 

 

 

40
20
O
K
as gas? of .cf 5°“? (if ,Q‘f
«of good) «i o .3?
'9 @CP x69 ’13?
Treatments GP QSQ (b)

 

Figure 4.3. Rye with amendments field experiment. Graphs show % of diseased root
for each treatment in 2003 at 2 harvests. a) Harvest 1 b) Harvest 2. Treatments

followed by the same letter are not significant at P < 0.05.

132

 

 

60
50 - 0 lb/a Carbon
[a - - 1200 lb/a Carbon

‘6 404 / - 23901b/a Carbon
2
'a
o
to
to
a:
.92
'u
h
0
e\°

 

 

 

A“ 0”
Treatments Q. 8:) (a)

 

 

 

- o lb/a Carbon
60 < - - 1200 lb/a Carbon
- 2390 lb/a Carbon

 

 

 

 

Treatments

 

 

Figure 4.4. Rye with amendments field experiment. Graphs show % of diseased root
for each treatment in 2004 at 2 harvests. a) Harvest l b) Harvest 2. Treatments

followed by the same letter are not significant at P < 0.05

133

Table 4.2. Rye with amendments field experiment. ANOVA showing significance of

factors for % of diseased root.

 

Analysis of Variance Pr > F

% Of diseased % Of diseased

root in 2003 root in 2004
Harvest 1
Factor 0.0449 0.1600 NS
Split 0.0947 NS 0.0013
Factor X Split 0.1174 NS 0.1598 NS
Block 0.0912 NS 0.2736 NS
Block X Factor 0.7533 NS 0.4333 NS
Block X Factor X Split 0.4057 NS 0.0040
Harvest 2
Factor 0.4627 NS 0.4116 NS
Split 0.0135 0.0066
Factor X Split 0.1702 0.2012 NS
Block 0.3739 NS 0.9166 NS
Block X Factor 0.3261 NS 0.2094 NS
Block X Factor X Split 0.1236 NS 0.0117

 

 

NS: Not Significant at P < 0.05

Table 4.3. Rye with amendments field experiment. Planned contrasts for % of

diseased root.

 

 

 

CONTRASTS °/o OF DISEASED ROOT

Pr > |t| 2003 2004
Harvest 1 H_arvest 2 Harvest 1 Harvest 2

0 lb/a C v/s 1200 lb/a C 0.2708 NS 0.3714 NS 0.026 0.3539 NS

0 lb/a C v/s 2390 lb/a C 0.1223 NS 0.2895 NS 0.0004 0.0304

1200 lb/a C v/s 2390 lb/a C 0.9985 NS 0.0752 NS 0.2219 NS 0.2634 NS
NS: Not Significant at P < 0.05

 

134

Specific Gravig

The specific gravities of tubers were measured at the time of final harvest. The
specific gravities in both 2003 and 2004 were not significantly different for any of the
treatments. In 2003, the specific gravity was highest for treatment with rye + 5000 lb/a
compost (figure 4.6) while in 2004, it was highest for the fumigated treatment (figure
4.5).

The ANOVA for the different factors involved in analyzing the data is presented
in table 4.4.

Contrasts between the amounts of carbon deposited in the soil and its effect on the

specific gravity is shown in table 4.5. None of the treatments were significantly different.

135

[-_,,_,, mvi

1.070

 

- 0 lb/a Carbon
1.068 n - - 1200 Ib/a Carbon a
- 2390 lb/a Carbon

 

 

 

1
1
1
1
1
1

0
Treatment @ Que

Figure 4.5. Rye with amendments field raggment.Sp—ecif; GraVfly for 2003.

g

Treatments followed by the same letter are not significant at P < 0.05

 

1.070

 

— 0 We Carbon
1.068 , a a - - 1200 lb/a Carbon
3 - 23901b/aCarbaon

   

 

 

31.066 , a
0
51.062
0
e
¢fihoem
1.058J """
e a as Q Q 3} Q
Q. f as o0 .39 .of 33 (as (as
Q" 00639 e 00 <2per QC? Q52?
0
‘9 (13300 '9ch 096) {-90 {139
0
Treatment qfi $10

 

 

 

Figure 4.6. Rye with amendments field experiment. Specific Gravity for 2004.
Treatments followed by the same letter are not significant at P < 0.05

136

Table 4.4. Rye field experiment. ANOVA for specific gravity of tubers.

 

Analysis of Variance Pr > F
SPECIFIC GRAVITY OF TUBERS

2003 2004
Factor 0.1566 NS 0.0426
Split 0.7353 NS 0.1487 NS
Factor X Split 0.2752 NS 0.0173
Block 1.0363 NS 0.3459 NS
Block X Factor 1.0000 NS 0.0363
Block X Factor X Split 0.3059 NS 0.2383 NS

 

NS: Not Significant at P < 0.05

Table 4.5. Rye with amendments field experiment. Planned contrasts for specific

 

gravity of tubers.
CONTRASTS SPECIFIC GRAVITY OF TUBERS
Pr > |t|
ms. M.
0 lb/a C v/s 1200 lb/a C 0.3316 NS 0.4794 NS
0 lb/a C v/s 2390 lb/a C 0.1480 NS 0.1245 NS
1200 lb/a C v/s 2390 lb/a C 0.9489 NS 0.0517 NS

 

NS: Not Significant at P < 0.05

137

D5! weight of roots

The treatments at both harvests in 2003 and 2004 had no significant effect on the
dry weights of root. Effect of treatments in 2003 and 2004 on dry weights of root is
presented in figure 4.7 and 4.8 respectively.

The ANOVA for the different factors involved in analyzing the data is presented
in table 4.6.

Contrasts between the amounts of carbon deposited in the soil and its effect on the
dry weight of root is shown in table 4.7. Amounts of Carbon input had any no significant

difference between the dry wt of roots.

138

 

 

  

 

 

 

 

 

  

 

 

16- f
14 I-01b/a --12001b/a I-23901b/a
‘ Carbon Carbon Carbon a
:12 y a a a I-I-i-I-C-Z
.3, 10 a :;:;:;:;:;:;
0: 8 -
O
E 5‘
s 4*
2 a
0 .
0 0 c} Q ‘x.
of .ogfb as s80 3‘1 Q06 Q$Q
6“ 0 005° 6‘
‘<‘> 09° 0 Q 962’
e «9 99° on“
0 X
Treatments Qt) qfio (a)
20 , L
18 -01b/a --12001b/a --23901b/a
16 Carbon Carbon Carbon
I» a a a
.514 .................
312 «
E10-
° 8
s 6-
E 4 .
2 -
0
0 0 é Q 9:} Q
of .f’b 8’) «<90 .52“> «080 Q‘f
(0‘ 00 0 0° é}
<<° 9° 6 8’0
9 9° 6° 6°
\9 b9 ‘0 q?)
0 X
Treatments Q$ q:\° (b)

 

 

 

Figure 4.7. Rye with amendments field experiment. Dry weight of roots in 2003 at 2
harvests. a) Harvest 1 b) Harvest 2. Treatments followed by the same letter are not

significant at P < 0.05

139

 

 

a

Drywtofrooting
O —I N (A) A 01 O) \l (I) (D

 

Tre atme nts

 

- 0 lb/a Carbon

- - 1200 Ib/a Carbon

- 23901b/a Carbon
a

  

 

(a)

 

 

 

11

1o _ - 0 lb/a Carbon

 

- - 1200 lb/a Carbon

, - 2390 lb/a Carbon

  

a

 

a

  

 

 

 

Treatments

 

 

 

Figure 4.8. Rye with amendments field experiment. Dry weight of roots in 2004 at 2
harvests. a) Harvest 1 b) Harvest 2. Treatments followed by the same letter are not

significant at P < 0.05

140

Table 4.6. Rye with amendments field experiment. ANOVA for dry weight of roots.

 

Analysis of Variance Pr > F

DE! weight of Dry weight of
root in g in 2003 root in g in 2004

Harvest 1

Factor 0.5338 NS 0.3631 NS
Split 0.2494 NS 0.8913 NS
Factor X Split 0.0434 0.7317 NS
Block 0.5364 NS 0.1735 NS
Block X Factor 0.2820 NS 0.1914 NS
Block X Factor X Split 0.7683 0.2500 NS
Harvest 2

Factor 0.6212 NS 0.7098 NS
Split 0.1028 NS 0.1093 NS
Factor X Split 0.9590 NS 0.5606 NS
Block 0.0571 NS 0.2804 NS
Block X Factor 0.8972 NS 0.6891 NS
Block X Factor X Split 0.0925 NS 0.0060

 

NS: Not Significant at P < 0.05

Table 4.7. Rye with amendments field experiment. Planned contrasts for dry weight

 

of roots.
CONTRASTS DRY WEIGHT OF ROOT IN g
Pr > |t| 2003 2004
Harvest 1 Harvest 2 Harvest 1 Harvest 2
0 lb/a C v/s 1200 lb/a C 0.2440 NS 0.3714 NS 0.3819 NS 0.0747 NS
0 lb/a C v/s 2390 lb/a C 0.8192 NS 0.2895 NS 0.5023 NS 0.2295 NS

1200 lb/a C v/s 2390 lb/a C 0.1480 NS 0.0752 NS 0.1536 NS 0.5565 NS
NS: Not Significant at P < 0.05

 

141

M

The yields were measured at the final harvest and were calculated on basis of
fresh weight of tubers in g/plant. In 2003 and 2004, treatment with rye + 5000 lb/a
compost gave the highest yield at 483.9 g/plant (figure 4.9) and 586.4 g/plant (figure
4.10) respectively. Both in 2003 and 2004, the yield for rye + 5000 lb/a compost was
significantly different than the lowest yield for those years which was fumigated
treatment (yield 341.4 g/plant) for 2003 and 5000 lb/a beet pulp (yield 367.8 g/plant) for
2004.

The ANOVA for the different factors involved in the experiment is presented in
table 4.8.

When contrasts for the effect of amount of carbon inputs in the soil on yield were
calculated, a significant difference was seen between 0 lb/a Carbon and 1200 lb/a Carbon
in 2003 while in 2004 this effect was not observed (table 4.10). In 2004, 0 lb/a Carbon

versus 2390 lb/a Carbon showed significant difference in the yield (table 4.10).

142

 

700

 

--01b/a - -12001b/a 533- 23901b/a
Carbon Carbon Carbon

600

 

 

 

 

 

Treatment: at? Qge’

 

Figure 4.9. Rye with amendments field experiment. Graph shows yield of fresh
tubers in g/plant in 2003. Treatments followed by the same letter are not significant
at P < 0.05

 

 

- 0 lb/a Carbon
600 - - 12001b/a Carbon

Yield in glplant

 

 

06 0 6Q 0‘ 006& Q}?
5 6° 6 6° 6 99 6
63° (1330 \QQ 99° x636 (1330
s“ 0..
Treatment 8‘ Q$

 

 

 

Figure 4.10. Rye with amendments field experiment. Graph shows yield of fresh
tubers in g/plant in 2004. Treatments followed by the same letter are not significant
at P < 0.05

143

Table 4.8. Rye with amendments field experiment. ANOVA for Yield as fresh

weight of tubers in g/plant

 

Analysis of Variance Pr > F

Yield in glplant in 2003

Yield in gzplant in 2004

Factor 0.1088 NS 0.5417 NS
Split 0.4686 NS 0.2919 NS
Factor X Split 0.4358 NS 0.6555 NS
Block 0.2446 NS 0.7824 NS
Block X Factor 0.5719 NS 0.0711 NS
Block X Factor X Split 0.0504 0.0029

 

NS: Not Significant at P < 0.05

Table 4.9. Rye with amendments field experiement. Planned contrasts for tuber

 

yield.
CONTRASTS YIELD or TUBERS
Pr > |t|

M2 300—4 -
0 lb/a C v/s 1200 lb/a C 0.0401 0.3604 NS
0 lb/a C v/s 2390 lb/a C 0.2431 NS 0.0232
1200 lb/a C v/s 2390 lb/a C 0.3789 NS 0.2194 NS

 

NS: Not Significant at P < 0.05

144

DISCUSSION

The disease incidence in both years at both harvests was highest for treatment
without any amendment (bare). In 2003 at harvest 1, the disease suppression was
maximum for treatment with rye+5000 lb/a compost. Similar pattern was observed in
2004 at both harvest periods. This effect might be due to both the quantity and quality of
organic matter added. Rye on its own may have controlled the soil fungal infection to a
certain degree as reported in a field study by Sumner et al., 1995, where populations of
Pythium irregulare and Rhizoctonia solani decreased after amending soil with rye
(Secale cereale) residues. Addition of compost increases the soil microbial population
which creates competition for the fungi. This is consistent with the findings of Pera et al.,
1983 and Perucci , 1990) who reported that in addition to increasing organic matter of
soil, amending with composts also increases the overall microbial population in soil.
Interaction between the rye and compost might have proved beneficial for the overall root
health of potatoes.

It is interesting to note that overall, treatments with highest carbon input (2390
lb/a Carbon) showed the maximum disease suppression. It is possible that the additional
carbon acts as a food source to the microbial community and also plays a role in
improving the overall health of the plant which indirectly causes greater disease
suppression.

Although the specific gravities were not significantly different, in 2003, the
specific gravity was highest for treatment with rye+5000 lb/a compost while in 2004, it

was highest for fiimigated treatment.

145

CONCLUSIONS

The soil amendments in this experiment were balanced by Carbon. Effect of the
different amounts of Carbon on existing and introduced soil fungi was investigated in this
study. Along with the disease incidence, effect on yield, specific gravity was also studied.
In both the years, it was noticed that the amount of Carbon input into the soil made a
difference on the % of disease infection. The quality of residue is also an important factor
to consider when disease incidence is to be reduced. In this experiment, overall, treatment
with rye + 5000 lb/a Carbon gave the best results. This treatment on the whole was
successful in limiting disease infection on potato roots. This effect could be due to many
reasons one of them being increase in beneficial soil microbes which indirectly competes
with the antagonistic soil fungi. Greater amount of soil organic carbon (SOC) and carbon
inputs to soil result in greater land productivity and contributes to favorable soil
properties such as moisture and nutrient retention, which in turn buffer ecosystems from
abiotic stresses (Woomer et al., 1994; Murage et al., 2000). This may result in greater
resistance of plants to soil fungal diseases. Rye may also participate in its ability to
produce certain allelochemicals which could reduce growth of fungi.
FUTURE WORK
It would be useful to study what effect soil organic carbon has on soil fungal populations.
Data on allelopathic cover crops and their effect on fungal infections might be

informative for farmers who use these as winter covers.

146

REFERENCES CITED

Abawi, G.S., and TL. Widmer. 2000. Impact of soil health management practices on
soilbome pathogens, nematodes and root diseases of vegetable crops. Appl Soil
Ecol 15 (1):37—47.

Baker, R., 1965. The dynamics of inoculum, In: Baker, K.F., Snyder W.C. (Eds),
Ecology of Soil-bome Plant Pathogens. Prelude to Biological Control, University
of California Press, Berkeley, CA, pp. 395—404.

Campbell, C. A. 1978. Soil organic carbon, nitrogen and fertility. In: M. Schnitzer and S.
U. Khan (Eds), Soil organic Matter. Developments in Soil Science 8. Elsevier,
Amsterdam, pp. 173-271.

Creamer, N.G., M.A. Bennett, and B.R. Stinner. 1997. Evaluation of cover crop mixtures
for use in vegetable production systems. HortScience 32:866—870.

Conn, K. L., and G. Lazarovits. 1999. Impact of animal manure on verticillium wilt,
potato scab, and soil microbial populations. Can. J. Plant Pathol. 21 :81-92.

Cook, R.J., 1980. Fusarium foot rot of wheat and its control in the Pacific Northwest.
Plant Disease. 64: 1061-1066

Cook, R. J ., and K. F. Baker. 1983. The nature and practice of biological control of plant
pathogens. Am. Phytopathol. Soc., St Paul, MN.

Doran, J. W. 1987. Microbial biomass and mineralizable nitrogen distributions in no-
tillage and plowed soils. Biol. F ertil. Soils, 5:68-75.

Fauci, M. F., and R. P. Dick. 1994. Soil microbial dynamics—short term and long term
effects of inorganic and organic nitrogen. Soil Sci. Soc. Am. J. 58:801—806.

Gardner, J ., and E. P. Casewell-Chen. 1994. Raphanus sativus, Sinapis alba, and
F agopyrum esculentum as hosts to Meloidogyne incognita, Meloidogyne javanica,
and Plasmodiophora brassicae. J. Nematol. 26:759-760.

Gonzalez, A., and M. Canto-Saenz. 1993. Comparisons of 5 organic amendments for the
control of Globera pallida in microplots in Peru. Nematropica 23:133—138.

Hoitink, H.A.J., Y. Inbar, and M. J. Boehm. 1991. Status of compost-amended potting

mixes naturally suppressive to soil-borne diseases of floricultural crops. Plant
Disease 75: 869—873.

147

Homma, Y., C. Kubo, M. Ishii, and K. Ohata. 1979. Effect of organic amendments to soil

on suppression of disease severity of tomato F usarium-wilt. Bull. Shikoku Agric.
Exp. Station 34289—101.

Janzen, H. H., C. A. Campbell, S. A. Brandt, G. P. Lafond, and L. Townley-Smith. 1992.
Light-fraction organic matter in soils from long-term crop rotations. Soil Sci. Soc.
Am. J. 56:1799-1806.

Kaplan, M., J. P. Noe, and P. G. Hartel. 1992. The role of microbes associated with
chicken litter in the suppression of Meloidogyne arenaria. J. Nematol. 24:522—
527.

Kaplan, M., and J. P. Noe. 1993. Effects of chicken-excrement amendments on
Meloidogyne arenaria. J. Nematol. 25:71—77.

Lewis, J. A., and G. C. Papavizas. 1971. Effect of sulfur-containing volatile compounds
and vapors from cabbage decomposition on Aphanomyces euteiches.
Phytopathology. 61:208-214.

Lockwood, J. L. 1988. Evolution concepts associated with soilborne plant pathogens.
Ann. Rev. Phytopathol. 29293-121.

Lumsden, R. D., R. Garcia-E, J. A. Lewis, and G. A. Frias—T. 1987. Suppression of
damping-off caused by Pythium spp. in soil from indigenous Mexican Chinampa
agricultural system. Soil Biol. Biochem. 19: 501-508.

Murage, E. W., N. K. Karanja, P. C. Smithson, and P. L. Woomer. 2000. Differences in
soil properties between productive and non-productive fields identified by
smallhold farmers in the Central Highlands of Kenya. Agriculture, Ecosystems
and Environment 79, pp. 1—8.

Nitta, T. 1991. Diversity of root fungal floras: Its implications for soil-borne diseases and
crop growth. JARQ. 25: 6—1 1.

Osunlaja, S. 0. Effect of organic soil amendments on the incidence of stalk rot of maize.
Plant Soil 1272237—241.

Papavizas, G. C. 1970. Colonization and growth of Rhizoctonia solani in soil. In: J.R.
Parmenter, Editor, Rhizoctonia Solani, Biology and Pathology, University of
California Press, Berkeley. pp. 108—122.

Papendick, R. I., and R. J. Cook. 1974. Plant water stress and development of F usarium
foot rot in wheat subjected to different cultural practices. Phytopathology.
642358-363

Paul, E. A. 1984. Dynamics of organic matter in soils. Plant Soil, 76:275-285.

148

Pera, A., G. Vallani, I. SirenoM. L. Bianchin, and M. de Bertoldi, 1983. Effect of organic
matter on rhizosphere microorganisms and root development of sorghum plants in
two different soils. Plant Soil. 74: 3—18.

Perucci, P., 1990. Effect of addition of municipal solid waste compost on microbial
biomass and enzyme activities in soil. Biol. Fertil. Soils 10:221—226.

Phillips, D. J., A. G. Watson, A. R. Weinhold, and W. C. Snyder. 1971. Damage of
lettuce seedlings related to crop residue decomposition. Plant Dis. Rep., 55:837-
841.

Riegel, C., F. A. Fernandez, and J. P. Noe. 1996. Meloidogyne incognito infested soil
amended with chicken litter. J. Nematol. 28:369—3 78.

Riegel, C., and J. P. Noe. 2000. Chicken litter soil amendment effects on soil microbes
and Meloidogme incognita on cotton. Plant Dis. 84: 1275—1 281.

Schneider, R.W., 1982. Suppressive Soils and Plant Disease, APS Press, St. Paul, MN, 88
PP-

Sherwood, R. T. 1970. Physiology of Rhizoctonia solani. In: J.R. Parmenter, Editor,
Rhizoctonia Solani, Biology and Pathology, University of California Press,
Berkeley. pp. 69—92.

Sivapalan, A., W. C. Morgan, and P. R. Franz. 1993. Monitoring populations of soil
microorganisms during conversion from conventional to an organic system of
vegetable growing. Biol. Agric. Horti. 10: 9—27.

Subbarao, K. V., J. C. Hubbard, and S. T. Koike. 1994. Effects of broccoli residue on
Verticillium dahliae microsclerotia and wilt incidence in cauliflower.
Phytopathology. 84:225.

Sumner, D. R., S. C. Phatak, J. D. Gay, and R. B. Chalfant. 1995. Soilbome pathogens in
a vegetable double-crop with conservation tillage following winter cover crops.
Crop Protection. 14(6): 495-500.

Sumner, D. R., M. R. Hall, J. D. Gay, G. MacDonald, S. 1. Savage, and R. K. Bramwell.
2002. Root diseases, weeds, and nematodes with poultry litter and conservation
tillage in a sweet com-snap bean double crop. Crop protection. 21:963-972.

Van Gestel, M., J. N. Ladd, and M. Amato. 1991. Carbon and Nitrogen mineralization
from two soils of contrasting texture and microaggregate stability: influence of
sequential fumigation, drying and storage. Soil Biol. Biochem. 23:313-322.

Woods, L. E. 1989. Active organic matter distribution in the surface 15 cm of
undisturbed and cultivated soil. Biol. Fertil. Soils, 81271-278.

149

 

Woomer, P. L., A. Martin, A. Albrecht, D. V. S. Resck, and H. W. Scharpenseel. 1994.
The importance and management of soil organic matter in the tropics. In:
Woomer, PL. and Swift, M.J., Editors, 1994. The Biological Management of
Tropical Soil Fertility, Wiley, Chichester, UK, pp. 47—80.

150

 

111111111111111111111111j1