Citation
Suppression of Meloidogyne Spp. by Pasteuria Penetrans

Material Information

Title:
Suppression of Meloidogyne Spp. by Pasteuria Penetrans
Creator:
Fulton, Elke Weibelzahl, 1963-
Publication Date:
Language:
English
Physical Description:
xvi, 152 leaves : ill. ; 29 cm.

Subjects

Subjects / Keywords:
Bacteria, Sporeforming ( lcsh )
Dissertations, Academic -- Entomology and Nematology -- UF ( lcsh )
Entomology and Nematology thesis, Ph.D ( lcsh )
Meloidogyne arenaria -- Control ( lcsh )
Meloidogyne incognita -- Control ( lcsh )
Meloidogyne javanica -- Control ( lcsh )
Nematocides -- Testing ( lcsh )
Plant nematodes -- Control ( lcsh )
Alachua County ( local )
Endospores ( jstor )
Roundworms ( jstor )
Pasteuria ( jstor )
Genre:
bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Thesis:
Thesis (Ph.D.)--University of Florida, 1998.
Bibliography:
Includes bibliographical references (leaves 132-151).
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Elke Weibelzahl Fulton.

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University of Florida
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Resource Identifier:
029226198 ( ALEPH )
39556899 ( OCLC )

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Full Text









SUPPRESSION OF MELOIDOGYNE SPP. BY PASTEURIA PENETRANS


By

ELKE WEIBELZAHL FULTON














A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY


UNIVERSITY OF FLORIDA


1998






























Before you can reach to the top of a tree and understand the buds and flowers,
you will have to go deep to the roots, because the secret lies there.
And the deeper the roots go, the higher the tree goes.

- Nietzsche






























To a man who taught me many of my skills, my father, Armin Willi Paul Weibelzahl.














ACKNOWLEDGMENTS


I am grateful to the efforts of many people, and I am bothered by the omission of many names from these two pages. My thanks go to all who have helped me.

First and foremost, Don W. Dickson, my chairman, took a chance on me and has been patient and generous in providing the financial support, direction, and freedom that I have enjoyed in the course of obtaining my doctorate in his laboratory. Robert McSorley, Dave Mitchell, and Ben Whitty, my committee members, provided tremendous assistance with their kindness and encouragement. Robert McSorley reviewed my manuscripts and dissertation and was most supportive with his statistical advice. Dave Mitchell taught me to consider especially the philosophical aspects of science. By working with Ben Whitty, I developed a major interest in applied nematology. Without the recommendations by Simon Gowen and Richard Sikora to pursue advanced education in tropical and subtropical regions, I would not have discovered my favorable working environment. Thanks go to them as well.

I also wish to express sincere appreciation to Reginald Wilcox and Tom Hewlett, whose helping hands and cheerful spirits made the technical aspects of field and greenhouse projects run smoothly. Jay Harrison deserves recognition for supporting me with his statistical ingenuity. Many thanks go to Steve Lasley and all those computer wizards who are responsible for the knowledge I have gained about computers.


iv








Deserving additional, special mention are John Strayer, who helped me begin;

Charlie Tarjan, who helped me to get through; and Grover Smart, who helped me complete the graduate program at the University of Florida. I am also thankful for the privilege of having worked with the wonderful students they have recruited for the department. Special thanks must be given to Georgina Robinson, Bettina Moser, and Leandro Freitas for their friendship and encouragement throughout the years in graduate school.

I am most grateful to my husband, Michael, our son, Riley, and our daughter, Kyra, all of whom have entered my life in the course of obtaining my doctorate. Their tremendous patience, countless compromises, and never-ending love surrounded me with an ocean of emotional support. Finally, I wish to thank my mother, Anna Weibelzahl, and my faithful siblings and friends on the other side of the Atlantic for their alliance despite the enormous distance between us. They all did a wonderful job in keeping my spirits up.


v









TABLE OF CONTENTS

Page

ACKNOWLEDGMENTS ....................................................................... iv

LIST OF TABLES............................................................................................................... ix

LIST OF FIGURES..................................................................................................... xii

ABSTRA CT........................................................................................................................ xv

CHAPTERS

1 INTRODUCTION ............................................................................................................ 1

Biological Control of Nem atodes .............................................................................. 1
Introduction...... ......................................................................................... 1
Historical Background............................................................................... 2
Taxonom ic Status of the Genus Pasteuria ................................................ 3
Life Cycle of Pasteuria spp...................................................................... 6
Ecology of Pasteuria spp........................................................................ 13
Biological Control of Nem atodes by Pasteuria spp................................. 19
Biological control attributes .......................................................... 19
Natural control............................................................................. 21
Inundative application................................................................. 22
Integrated nem atode m anagem ent ............................................... 23
Useful Methods and Techniques for Studying Pasteuria spp................. 24
Detection of Pasteuria spp. in soils and nematodes.................... 24
Isolation of Pasteluria spp. from soils and nematodes ................ 25
Quantification of endospores in soils, root powder or suspension........................................................................ 26
Culture and preservation of Pasteuria penetrans......................... 27
Attachm ent ................................................................................... 28
Objectives................................................................................................................ 29

2 POPULATION DEVELOPMENT OF MELOIDOGYNE ARENARIA RACE
1 AND PASTEURIA PENETRANS IN A 6.5-YEAR MICROPLOT
STUD Y ................................................................................................................... 30

Introduction ............................................................................................................. 30
M aterials and M ethods ........................................................................................ 31
Results..................................................................................................................... 33
Discussion............................................................................................................... 47


vi








3 USE OF MICROWAVE HEATING IN EVALUATION OF A
MELOIDOGYNE ARENARIA-SUPPRESSIVE SOIL CONTAINING
PASTEURIA PENETRANS AND ITS APPLICATION IN A
SUPPRESSIV E-SOIL TEST.............................................................................. 51

Introduction ........................................................................................................ 51
M aterials and M ethods ......................................................................................... 52
M icrow ave Treatm ent.............................................................................. 52
Suppressive Soil Test. ............................................................................ 54
N em atode Origin..................................................................................... 56
Statistical Analysis................................................................................... 56
Results ..................................................................................................................... 57
M icrow ave Treatm ent.............................................................................. 57
Suppressive Soil Test. ............................................................................ 60
Discussion............................................................................................................... 67

4 POPULATION DEVELOPMENT OF MELOIDOGYNE SPP. AND
PASTEURIA PENETRANS AS AFFECTED BY CULTURAL
PRA CTICES IN TOBACCO .............................................................................. 72

Introduction ......................................................................................................... 72
M aterials and M ethods ......................................................................................... 73
N em atode Populations.............................................................................. 74
Pasteuria penetrans Isolates .................................................................... 74
Laboratory Experim ent............................................................................ 75
Field Experim ent..................................................................................... 75
Results.....................................................................................................................77
Laboratory Experim ent............................................................................ 77
Field Experim ent..................................................................................... 77
Discussion............................................................................................................... 85

5 SUPPRESSION OF MELOIDOGYNE INCOGNITA AND M. JAVANICA BY
PASTEURIA PENETRANS IN FIELD SO IL..................................................... 88

Introduction ......................................................................................................... 88
M aterials and M ethods ......................................................................................... 89
Soil Treatm ents.......................................................................................... 89
Laboratory Experim ents......................................................................... 90
Greenhouse Experim ent......................................................................... 91
Nem atode Origin..................................................................................... 92
Statistical A nalysis................................................................................... 93
Results.....................................................................................................................93
Laboratory Experim ents......................................................................... 93
Greenhouse Experim ent......................................................................... 93
D iscussion............................................................................................................... 99


vii








6 MELOIDOGYNE ARENARIA AND PASTEURIA PENETRANS
POPULATION DENSITY DEVELOPMENT IN METHYL BROMIDE
TREATED SOIL AS AFFECTED BY AN INTERCROPPING
SYSTEM .............................................................................................................. 101

Introduction ........................................................................................................... 101
M aterials and M ethods .......................................................................................... 103
Results................................................................................................................... 109
Discussion............................................................................................................. 124

7 Sum m ary....................................................................................................................... 128

LIST OF REFERENCES................................................................................................. 130

APPEN DIX ...................................................................................................................... 151

BIOGRA PHICAL SKETCH ........................................................................................... 152


viii














LIST OF TABLES


Table page

1-1. Geographic distribution of Pasteuria spp. ............................................................. 13
2-1. Effect of Meloidogyne arenaria alone and in combination with Pasteuria
penetrans, and of a rye, vetch, or bare fallow winter cover crop, on yield
and performance of peanut in the 5b year (fall of 1991) of a 6.5-year
m icroplot experim ent.......................................................................................... 35
2-2. Effect of Meloidogyne arenaria alone and in combination with Pasteuria
penetrans, and of a rye, wheat, or bare fallow winter cover crop on yield and performance of peanut in the 6" year (1992) of a 6.5-year microplot
experim en t ............................................................................................................... 3 6
2-3. Effect of Meloidogyne arenaria alone and in combination with Pasteuria
penetrans, and of a rye, wheat, or bare fallow winter cover crop, on yield and performance of peanut in the 7' year (1993) of a 6.5-year microplot
experim en t............................................................................................................... 37
2-4. The percentage of second-stage juveniles (J2) of Meloidogyne arenaria
infected with Pasteuria penetrans and the average number of endospores
per juvenile in peanut microplots infested with M. arenaria and P.
penetrans in the spring of 1987, and rotated with rye, vetch or wheat, and
bare fallow as winter cropping sequence ............................................................ 42
3-1. Colony forming units (cfu) of soil fungi and attachment of Pasteuria
penetrans endospores to second-stage juveniles (J2) of Meloidogyne
arenaria in untreated soil and soil autoclaved twice for 1.5 hours at 55 kPa,
microwaved for 3 minutes/kg of soil, or air dried for 2 weeks in the
greenh o u se ............................................................................................................... 6 1
3-2. ANOVA table for the effect of soil source, soil treatments, and Meloidogyne
arenaria race 1 inoculum levels on nematode reproduction and fresh root
w eights of peanut cv. Florunner .......................................................................... 62
3-3. Effect of autoclaving, microwaving, and air-drying of soil infested with
Meloidogyne arenaria race 1 alone on nematode reproduction, percentage
of females infected by Pasteuria penetrans, and fresh root weights of
peanut cv. Florunner following inoculation with 0 or 2,000 second-stage
ju ven iles................................................................................................................... 6 3
3-4. Effect of autoclaving, microwaving, and air-drying of soil infested with
Meloidogyne arenaria race 1 and Pasteuria penetrans on nematode
reproduction, percentage of females infected by P. penetrans and fresh root


ix








weights of peanut cv. Florunner following inoculation with 0 or 2,000
second-stage juveniles of M . arenari a race 1 ..................................................... 64
3-5. Effect of autoclaving, microwaving, and air-drying of soil maintained free of
nematodes and Pasteuria penetrans on nematode reproduction, percentage of females infected with P. penetrans and fresh root weights of peanut cv.
Florunner following inoculation with 0 or 2,000 second-stage juveniles of
M eloidogyne arenaria race 1............................................................................... 66
4-1. .Attachment pattern on second-stage juveniles (J2) of Meloidogyne javanica
and M. incognita over three generations of Pasteuria penetrans isolate P110 from M . incognita infecting tobacco ............................................................. 78
4-2. Attachment pattern on second-stage juveniles (J2) of Meloidogyne javanica
and M. incognita over three generations of Pasteuria penetrans isolate P120 from M . javanica infecting tobacco. ............................................................ 79
4-3. Population density development of Meloidogynre spp. in 1992 and 1993 as
determined by the number of second-stage juveniles in the soil, and the
galling indices of roots of two tobacco cultivars treated with two inorganic
nitrogen rates and three autumn cover crop treatments......................................... 81
4-4. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar
history, and their effects on the number of second-stage juveniles (J2) of Meloidogyne spp. in 100 g of soil collected at planting and after the final
harvest in 1992 ................................................................................................... . . 82
4-5. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar
history, and their effects on root-galling indices in 1992 and 1993....................... 82
4-6. Percentage of second-stage juveniles (J2) of Meloidogyne spp. with attached
endospores of Pasteuria penetrans, average number of endospores attached
per juvenile in the soil, and percentage of P. penetrans-infected females collected from two tobacco cultivars in a field treated with two nitrogen
fertilizer rates and three autumn cover crops in 1992 and 1993...........................84
4-7. Interaction between inorganic nitrogen fertilizer levels, tobacco cultivar
history, and cover crops, and their effects on the percentage of second-stage
juveniles (J2) encumbered with endospores of Pasteuria penetrans at
planting in 1992 ................................................................................................. . . 85
5-1. Survival of soil fungi and Pasteuria penetrans in untreated soil or soil
autoclaved twice for 1.5 hours at 55 kPa, microwaved for 3 minutes/kg of
soil, or air-dried for 2 weeks in the greenhouse.................................................. 94
5-2. ANOVA table for the effects of tobacco cultivars, soil treatments, and
Meloidogyne incognita race 1 inoculum levels on nematode reproduction
and plant perform ance. ......................................................................................... 95
5-3. Effect of autoclaving, microwaving, and air-drying on soil suppressiveness to
Meloidogyne spp. and on the expression of root-knot on tobacco cultivar
Coker-371 Gold following inoculation with 0 or 2,000 second-stage
juveniles of M . incognita race 1.......................................................................... 96
5-4. Effect of autoclaving, microwaving, and air-drying on soil suppressiveness to
Meloidogyne spp. and on the expression of root-knot on tobacco cv.


x








Northrup King-326 following inoculation with 0 or 2,000 second-stage
juveniles of M . incognita race 1.......................................................................... 97
6-1. Grouping of 90 microplots based on the mean number of Pasteuria
penetrans endospores attached per second-stage juvenile of Meloidogyne arenaria race 1 in a bioassay on microplot soil conducted in the spring of
19 9 4 ....................................................................................................................... 1 10
6-2. Main effect of differences of groups with different Pasteuria penetrans
endospore densities in microplots intercropped to two cycles of corn and
beans in rotation with peanut on the population development of P.
penetrans and Meloidogyne arenaria race 1 as determined by endospore
attachment bioassay, soil sample extraction, and galling index.............................. 111
6-3. Mean number of Pasteuria penetrans endospores attached per second-stage
juvenile of Meloidogyne arenaria race 1 in bioassays of microplot soil with
different initial endospore densities intercropped with two cycles of corn
and beans in 1994, planted to hairy vetch in the winter of 1994-95, and
cropped with peanut in the summer of 1995. ........................................................ 112
6-4. Percentage of second-stage juveniles of Meloidogyne arenaria race 1 with
endospores of Pasteuria penetrans attached in bioassays of microplot soil
with different initial endospore densities intercropped with two cycles of
corn and beans in 1994, planted to hairy vetch in the winter of 1994-95, and
cropped with peanut in the sum m er of 1995 ......................................................... 116
6-5. Number of second-stage juveniles of Meloidogyne arenaria race 1 per 100
cm3 of microplot soil with different initial Pasteuria penetrans endospore
densities intercropped with two cycles of corn and beans in 1994, planted to
hairy vetch in the winter of 1994-95, and cropped with peanut in the
sum m er of 1995 .................................................................................................... 117
6-6. Number of Pasteuria penetrans endospores attached per second-stage
juvenile of Meloidogyne arenaria race 1 in microplot soil with different
initial endospore densities intercropped with two cycles of corn and beans in
1994, planted to hairy vetch in the winter of 1994-95, and cropped with
peanut in the sum m er of 1995............................................................................... 119
6-7. Percentage of second-stage juveniles of Meloidogyne arenaria race 1 with
endospores of Pasteuria penetrans attached in microplot soil with different initial endospore densities intercropped with two cycles of corn and beans in
1994, planted to hairy vetch in the winter of 1994-95, and cropped with
peanut in the sum m er of 1995............................................................................... 121
6-8. Root galling rates of two intercroping systems with corn and beans rotated
with peanut grown in microplots grouped by different Pasteuria penetrans
endospore densities, and inoculated with Meloidogyne arenaria in the
spring of 19 94 . ...................................................................................................... 12 2
6-9. Spearman correlation coefficients of ranked data obtained from bioassays and
from the analysis of soil extracted from 90 microplots containing varying
population densities of Pasteuria penetrans endospores; plots were
intercropped with two cycles of corn and beans in 1994, planted to hairy


xi








vetch in the winter of 1994-95, and cropped with peanut in the summer of
19 9 5 ....................................................................................................................... 12 3
A-1. Effect of Meloidogyne arenaria (RKN) alone and in combination with
Pasteuria penetrans (RKN + Pp), and of a rye, vetch, or bare fallow winter
cover crop on the ring nematode population density in the falls of 1991 to
1993 of a 6.5-year m icroplot experim ent............................................................... 151


xii














LIST OF FIGURES


Figure page


1-1. Life cycle of Meloidogyne sp. and its bacterial parasite, Pasteuria penetrans............ 9
2-1. Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100
cm3 of soil and the number of endospores attached per juvenile extracted from peanut microplots infested with M. arenaria, and M. arenaria plus
P asteuria penetrans ............................................................................................ 34
2-2. Microplot treatment plan indicating the presence of Pasteuria penetrans
endospores as determined by soil sample analysis in the fall of 1991................. 40
2-3. Microplot treatment plan indicating the presence of Pasteuria penetrans
endospores as determined by soil sample analysis in the fall of 1992................. 41
2-4. Microplot treatment plan indicating the presence of Pasteuria penetrans
endospores as determined by soil sample analysis in the fall of 1993................. 42
2-5. Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100
cm3 of soil extracted from soil in microplots with A) M. arenaria, and B)
M . arenaria plus P. penetrans ........................................................................... 43
2-6. Number of attached endospores per juvenile extracted from soil in microplots
with A) M. arenaria, and B) M. arenaria plus P. penetrans............................. 45
2-7. Log presentation of peanut yield from plots treated with Meloidogyne
arenaria race 1 (RKN), M. arenaria plus Pasteuria penetrans (RKN + Pp), and untreated (Control); and the number of endospores per secondstage juvenile in RKN and RKN + Pp plots....................................................... 46
3-1. Effect of microwave radiation treatment of 1 kg of soil containing Pasteuria
penetrans on A) the attachment of P. penetrans endospores to Meloidogyne
arenaria race 1 and B) the survival of selected fungi as determined by the
num ber of colony-form ing units........................................................................ 57
3-2. Relationship between the soil moisture content and the number of endospores
of Pasteuria penetrans attached per second-stage juvenile (J2) of
Meloidogyne arenaria race 1 after 48 hours exposure at room temperature........ 58 3-3. Effect of soil moisture content and microwave treatment time on the
attachment of Pasteuria penetrans endospores to Meloidogyne arenaria
rac e I ....................................................................................................................... 5 8
5-1. Effect of soil treatments on a soil suppressive to Meloidogyne spp. and the
expression of root-knot on tobacco cultivar Coker 371 Gold following
inoculation with 2,000 second-stage juveniles of M. incognita race 1 ................. 84


xiii








6-1. Cut away illustration of a sweet corn and pole bean intercropping system in
field m icroplots (76 cm diam .).............................................................................. 104
6-2. Cut away illustration of a peanut crop in field iicroplots (76 cm diam.).................. 106
6-3. Relationship between the initial endospore population density and A) the final
number of endospores per juvenile as determined by bioassay, B) the final number of second-stage juvenile (J2) per 100 cm3 of soil, and C) the final
number of endospores per J2 as determined by soil sample extraction................. 113
6-4. Rate of change for A) the number of endospores of Pasteuria penetrans per
juvenile of Meloidogyne arenaria as determined by bioassay, B) the
number of second-stage juveniles (J2)/100 cm3 of soil, and C) the number
of endospores/juvenile, as determined in soil sample extraction based on the
initial endospore population density, and the change over the 1995 season........... 114


xiv












Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

-SUPPRESSION OF MELOIDOGYNE SPP. BY PASTEURIA PENETRANS By

Elke Weibelzahl-Fulton

May 1998

Chairman: Don W. Dickson
Major Department: Entomology and Nematology


Suppression of plant-parasitic nematodes with microbial agents is an alternative or supplemental management tactic that is receiving increased interest among nematologists. One nematode antagonist, the endospore-forming bacterium Pasteuria penetrans, has shown great potential in suppressing field populations of several plant-parasitic nematodes throughout the world. This obligate parasite of mainly root-knot nematodes, Meloidogyne spp., was studied for its potential to suppress M. arenaria on peanut, and M. incognita and M. javanica on tobacco. The objectives were to monitor the population densities of the root-knot nematode and P. penetrans in peanut microplots and in a naturally infested tobacco field, to develop a suppressive-soil test that allows the determination of the role of P. penetrans in nematode-suppressive soils, and to evaluate the effect of cultural practices, such as crop rotation, autumn cover crops, resistance and fertilizer regimes on the abundance of P. penetrans. Within 4 years after inoculation or contamination by P. penetrans, M. arenaria was nearly eliminated from microplot soil, and peanut yields were similar to that of the nematode free control. The P. penetrans-infested soil remained suppressive to M. arenaria for 3 years. Similar results were observed in a tobacco field.


xv








After 4 years of tobacco monoculture, a mixed population of M. incognita, M. javanica, and M. arenaria was suppressed to non-damaging levels. Two P. penetrans isolates were more pathogenic to M. incognita than to M. javanica. Inorganic nitrogen fertilizer rates and host resistance to M. incognita had inconsistent effects on the endospore build-up, which was favored by cultivation of susceptible cover crops. A microwave radiation treatment of

3 minutes/kg of soil containing 6% to 7% water reduced the fungal population in soil samples without impairing attachment of P. penetrans endospores to nematode juveniles. This treatment allowed the separation of nematode suppression by fungi from that caused by P. penetrans. Nematode suppressiveness in microplot and field soil was preserved after microwave and air-drying treatments, but not after soil was autoclaved. The bacterial parasite, P. penetrans, was identified as the main contributor to the suppressiveness of the microplot and the field soil.


xvi














CHAPTER 1
INTRODUCTION


Biological Control of Nematodes



Introduction



Nematologists are continuously unveiling the biological control potential of

Pasteuria penetrans (Thorne, 1940) Sayre and Starr, 1985, a mycelial endospore-forming bacterial parasite of mainly root-knot nematodes. The ecological history of P. penetrans in association with Meloidogyne spp., as well as knowledge about its role in nematode management strategies, is essential for improving the use of the organism as an effective biological control agent. This information also is important for recognition and effective exploitation of soils naturally infested with P. penetrans.

In 1991, soils suppressive to the peanut root-knot nematode, M. arenaria (Neal) Chitwood race 1, were produced in microplots at the University of Florida Agronomy Farm, Green Acres, Alachua County, Florida. Ninety microplots (76 cm in diameter) were established with 30 inoculated centrally with relatively low numbers of P. penetrans and Meloidogyne arenaria, 30 inoculated with M. arenaria alone, and 30 left uninoculated as control plots (Oostendorp et al., 199 1a). The microplots were planted to peanut in the spring of 1987. Within 3 years, yields of plots infected with P. penetrans and M. arenaria increased to a level comparable to the untreated control and the damage from root-knot nematodes decreased, suggesting that the soil became suppressive to the nematode. A


I





2


minimum of 3 years of intensive cropping was required for P. penetrans to amplify to suppressive levels. During the same period, a tobacco field that was naturally infested with Meloidogyne spp. was discovered to contain a soil suppressive to the nematode. This site was located about 300 meters from the microplot site. The bacterial parasite P. penetrans was reported to cause the nematode suppression (Chen et al., 1994).

The goal of this project was to monitor the dynamics of the bacterium in the two sites and further establish the relationship of P. penetrans to suppressive soil.



Historical Background



Biological control of nematodes has been of great interest to nematologists for almost as long as plant-parasitic nematodes have been known to damage crops (Stirling, 1991). Cobb (1920) considered using parasites and predators of nematodes as biological control agents when he suggested transferring predacious nematodes to sugarbeet fields in order to control populations of Heterodera schachtii Schmidt. Thorne (1927), however, questioned the economic benefits of predacious nematodes, and further studies on their potential for biological control of plant-parasitic nematodes came to an end. Linford's (1937, 1939) early efforts to use predacious fungi to control plant-parasitic nematodes stimulated interest in the nematode-trapping fungi in France, the United States, England, and the former USSR (Stirling, 1991). Due to the lack of success of these early experiments, the interest in biological control declined. It was not until the late- 1970s that interest in biological control revived when the potential health and environmental problems associated with the use of some nematicides surfaced (Kerry, 1990; 1993; Stirling, 1991; Thomason, 1987). The suspension of two of the most reliable soil fumigants, DBCP (1,2dibromo-3-chloropropane) and EDB (ethylene dibromide), and the lack of promising new





3


candidate nematicides, led nematologists to re-evaluate their goals in terms of nematode management research. This fact, plus the classical studies that demonstrated fungal antagonists suppressed populations of Heterodera avenae Woll. in Europe (Kerry et al., 1980; 1982) and other studies showing that fungal antagonists also suppressed Meloidogyne spp. (Stirling and Mankau, 1978; Stirling et al., 1979), rekindled interest in biological control. In recent years, some attempts have been made to market various fungal agents for nematode control (Al-Hazim et al., 1993; Timm, 1987), but the products generally have not been accepted.

Unfortunately, there are still no widely used examples of the contrived use of

biological antagonists to control plant-parasitic nematodes. Although there are many likely candidates (Dickson et al., 1994; Jansson, 1988; Kerry, 1987; 1988; Mankau, 1975b; 1980b; Sayre et al., 1988; Stirling, 1988; 1991; Tribe, 1980), the resources devoted to research on biological control of nematodes have been relatively limited (Stirling, 1991). There is optimism, however, that biological control will play an increasingly important role in future nematode management programs.

The search for nematode antagonists has generally centered on predacious and parasitic fungi and bacterial parasites. One such bacterium, P. penetrans, was identified and considered to have great potential for biological control of root-knot nematodes (Dickson et al., 1994). A Pasteuria sp. was first associated with parasites of plantparasitic nematodes when Thorne (1940) reported it as Duboscqia penetrans parasitizing Pratylenchus pratensis. Since the first reports that Pasteuria penetrans parasitized rootknot nematodes (Mankau, 1980a; 1980b; Stirling and White, 1982; Stirling, 1984), the organism became the subject of an increasing number of research projects (Stirling, 1991). Meloidogyne spp. were reported to be suppressed successfully in numerous pot, microplot, and field experiments (Brown et al., 1985; Channer and Gowen, 1988; Chen et al., 1996b;





4


1997c; Daudi, 1990; Dube and Smart, 1987; Jaya Raj and Mani, 1988; Maheswari and Mani, 1988; Maheswari et al., 1987; 1988; Oostendorp et al., 1991a; Stirling, 1984; Tzortzakakis, 1994a; Vargas et al., 1992; Weibelzahl-Fulton et al., 1996). However, limitations in mass production methods of P. penetrans have prevented its commercialization.



Taxonomic Status of the Genus Pasteuria



The genus Pasteuria comprises a group of Gram positive mycelial and endosporeforming bacteria that parasitize bodies of invertebrates, including nematodes (Williams et al., 1989a). The initial studies of the genus Pasteuria date back to 1888, when Metchnikoff described an internal parasite of two Daphnia spp. (water fleas) as Pasteuria ramosa (Metchnikoff, 1888). Partial credit for the discovery of the bacterium also should be given to Cobb (1906), who noted Pasteuria sp. as an internal parasite of nematodes, sketched it, and described it to be "perhaps monads." The taxonomy of this nematode parasite, however, has been subject to continuous confusion ever since Thorne (1940) described the first member of the group and, believing it was a protozoan, named it Duboscqia penetrans. He could not have realized its bacterial nature, mainly because ultrastructural techniques were not available to him and the concept of the prokaryotic cell had not yet been introduced. Thorne's (1940) description and nomenclature persisted for 35 years. Although other scientists worked with the organism and questioned its taxonomic placement (Williams, 1960), it was not until the reexamination by Mankau (1975a), using electron microscopy, that its affinity to bacteria rather than protozoa was revealed. The nematode parasite was renamed Bacillus penetrans (Thorne, 1940) Mankau, 1975.





5


Only a decade ago, Sayre et al. (1983) established a sound basis for the taxonomy of this group of hyperparasites. Detailed comparisons of the genera Pasteuria and Bacillus cleared up the confusion and revealed major similarities between the nematode parasite and Pasteuria ramosa (Sayre and Wergin, 1977; Sayre et al, 1983). Their close relationship became obvious in several distinctive morphological characteristics, such as the dichotomously branched mycelial microcolonies that give rise to fragmentation and sporangia, and finally to endogenous spores formed within the old mother cell wall. Similarities also had been shown at the ultrastructural level in the unique forms and sequences of life stages of the two organisms (Sayre and Starr, 1985). The parasite of plant-parasitic nematodes, previously known as Bacillus penetrans, was then placed in the genus Pasteuria and renamed Pasteuria penetrans. However, the newly named bacterium was by no means a uniform entity. Different isolates of this organism were found to differ in their physical and pathological characteristics (Bird et al., 1990). The description of two new species followed: Pasteuria thornei Starr & Sayre (1988), which parasitized the lesion nematode Pratylenchus brachyurus (Godfrey), and Pasteuria nishizawae Sayre, Wergin, Schmidt & Starr (1991), which parasitized cyst nematodes, Globodera rostochiensis Woll. and Heterodera glycines Ichnohe. A fifth species, which parasitized the pea cyst nematode, Heterodera goettingiana Liebscher, has been reviewed for taxonomic classification (Sturhan et al., 1994). Numerous Pasteuria spp. have endospores distinctly smaller or larger than those previously described (Chen, 1996; Ciancio and Mankau, 1989; Ciancio et al., 1992; 1994; Giblin-Davis, 1990; Giblin-Davis et al., 1990; Jaffee et al., 1985; Noel and Stanger, 1994; Sayre et al., 1985). Some isolates display a cross-genera host range and various biological characteristics (Bhattacharya and Swarup, 1988; Mankau, 1975; Oostendorp et al., 1990; Pan et al, 1993; Sharma and Davies, 1996; Vargas and Acosta, 1990). Thus, Pasteuria spp. are being differentiated by host specificity,





6


developmental characteristics, and shape and size of sporangia and endospores (Sayre and Starr, 1989).


Based on a study of host records and endospore morphometrics of pathotypes

described previously as the P. penetrans group, Ciancio et al. (1994) concluded that host taxonomy and endospore dimensions are of limited value to the definition of Pasteuria spp. This would partially explain a discovery by Davies et al. (1992), who identified different surface proteins of endospores in three P. penetrans populations, all parasites of Meloidogyne incognita. A recent study shows that P. penetrans produces heterogeneous endospores (Davies et al., 1994). These subpopulations of endospores show specificity to various nematode populations.


No definite criteria or genetic data are available to establish whether distinct species of Pasteuria differing in their biology and physiology share common hosts and morphometrics, or whether some or even all of the Pasteuria members should be considered as pathotypes of a unique species, regardless of hosts and morphometrics (Ciancio et al., 1994). The confusion in taxonomy of Pasteuria probably will not be clarified until the bacterial genom properties are elucidated. Axenic cultivation of the bacteria is crucial to understanding the biology and taxonomy of Pasteuria, but artificial cultivation of Pasteuria spp. has not been successful (Bishop and Ellar, 1991; Previc and Cox, 1993; Reise et al., 1988).




Life Cycle of Pasteuria spp.



The life cycle of P. penetrans in root-knot nematodes was first described in detail by Sayre and Wergin (1977). Their observations were confirmed by Imbriani and Mankau





7


(1977) and additional studies have contributed useful information on the life cycle (Davis et al., 1988; Mankau and Prassad, 1977; Sayre, 1988; Sayre and Starr, 1985; Sturhan et al., 1994). Pasteuria penetrans is a density dependent obligate parasite that grows and multiplies within the bodies of plant-parasitic nematodes. Up to 2.5 million nonmotile endospores are produced per female of Meloidogyne spp. (Hewlett and Dickson, 1993) and released into the soil environment upon degradation of the nematode carcass.

The developmental stages of all members of Pasteuria spp. parasitic on nematodes appear to be similar, but there are differences in shapes and sizes of endospores and sporangia among isolates obtained from different nematode genera. The following detailed description and illustration of the developmental stages covers P. penetrans and Meloidogyne sp. and draws attention to other species for notable differences (Fig. 1-1).

Attachment. The organism encounters host nematodes when endospores passively adhere to the cuticle of a migratory stage of the nematode in soil. In the case of Meloidogyne sp., a single second-stage juvenile may have one to several hundred endospores attached to its cuticle (Davies et al., 1991). Mobility and infectivity of the second-stage juveniles (J2) is reduced when somewhere between 7 and 50 endospores are attached to the juvenile (Brown and Smart, 1985; Davies et al., 1988, 1990; Sell and Hansen, 1987; Stirling, 1984). Since attachment can occur readily in soil or aqueous suspension (Slana and Sayre, 1982), there appear to be no requirements for adhesion other than those found on the endospore or nematode cuticle surface.

There is considerable variation in the ability of P. penetrans to attach to and infect species of nematodes, particularly within isolates parasitic on Meloidogyne spp. (Brown and Smart, 1984; Davies et al., 1988; Davies and Danks, 1992; 1993; Oostendorp et al., 1990; Sell and Hansen, 1987; Spaull, 1984; Stirling, 1985; Verdejo-Lucas, 1992). Cuticular variations in ultrastructure and composition between nematode populations

























Fig. 1-1. Life cycle of the root-knot nematode, Meloidogyne sp., with and without its bacterial parasite, Pasteuria penetrans (outer circle). 1) Second-stage juveniles (J2) entering root tip, 2) migrating intercellulary in cortex, 3) J2 establishing feeding sites in the vascular system; germinating P. penetrans endospores, 4) third-stage juveniles, 5) fourthstage juveniles, 6, 7) young females, 8) female with P. penetrans lays no or few eggs, whereas a healthy female forms an egg mass and lays eggs, and 9) infected female carcass degrades and releases mature endospores into the soil. The inner circle illustrates the life cycle of P. penetrans and its various developmental stages. Cross sections and top views of a) mature endospores, b) endospore attached to cuticle of Meloidogyne sp., c) germinating endospore, d) microcolonies formed in the pseudocoelom of the nematode, e) septations in rapidly growing thallus, f) dichotomously branched hyphae with elongated terminal cells, g) fragmented thalli separated from the thallus and visible forespore, h) cell wall separates forespore from parasporium of the egg-shaped sporangium, i) differentiation of spore core and perisporal fibers, j) mature endospores surrounded by exosporium and sporangium, and k) endospores released into soil.





9


92 44




























7 "RN





10


(Davies and Danks, 1992; Reddigari et al., 1986) may be responsible for different levels of attachment. The greatest spore attachment occurs when endospores are exposed to Meloidogyne spp. from which they were originally isolated (Davies et al., 1988; Davies et al., 1994; Oostendorp et al., 1990). Some P. penetrans isolates attached indiscriminately to Meloidogyne spp. other than the original host (Stirling, 1985).

The infective juvenile of Meloidogyne spp. is the only stage that appears to be

parasitized by P. penetrans (Mankau, 1980b), and this could perhaps be attributed to the cuticular surface composition differences observed between the nematode's life stages (McClure and Stynes, 1988). It has been suggested that the surfaces of J2 contain structural carbohydrate recognition units that are probably not collagen, and that these interact with N-acetylglucosamine moieties on the endospore surface that are linked to either glycoproteins or peptidoglycans (Bird et al., 1989; Davies and Danks, 1993; Spiegel et al., 1996). However, it also was suggested that collagens might be involved in attachment (Persidis et al., 1991). Several studies suggest that host specificity is caused by differences in the amount and nature of surface proteins of the endospores (Chen et al., 1997a; Davies et al., 1992, 1994).

Infection and development. Endospore attachment to J2 of Meloidogyne sp. is not necessarily followed by infection (De Silva and Gowen, 1994). The endosporeencumbered J2 penetrate the growing roots of a host plant behind the root cap. In the zone of cellular differentiation, the J2 reside in cortical tissue with their heads in the periphery of the vascular tissue where feeding starts (Hussey, 1985). Thereafter, a germ tube emerges from a central pore at the basal side of the endospore and penetrates the cuticle and hypodermis of the nematode (Sayre and Starr, 1985). There is evidence that germ tubes which do not penetrate the cuticle, emerge between the "rim" of the perisporium and the





11


cuticle of the nematode (Birchfield and Antonopoulus, 1976). Endospore germination appears to be triggered by the onset of feeding (personal observations).

The protoplast of the endospore enters the nematode body through the germ tube and develops into vegetative, spherical colonies consisting of a dichotomously branched, septate mycelium (Sayre and Starr, 1985). These early stages (mycelial colonies) are visible with the light microscope as dense granulation within the nematode pseudocoelom. Fragmentation of the thalli separates the mycelial masses, which are then transported in the fluid of the pseudocoelomic cavity through the body of the host nematode. These fragments give rise to microcolonies of four, eight, or more terminal cells. In advanced developmental stages, an increasing number of colonies with two, three, or four clubshaped, enlarged terminal cells are seen (doublets, triplets, and quartets). These each give rise to a single sporangium in which a single endospore is formed. The process of endospore formation appears typical of that found in other endospore-forming bacteria (Chen et al., 1997b).

As the sporangium enlarges, a septum is formed in the upper third of the mother

cell separating the incipient forespore cytoplasm from the remainder of the cell (Chen et al., 1997b). The septum growing around the forespore finally provides a double-layered membrane that encloses the condensed cytoplasm. The wing-like perisporium and the cortex develop between the two membranes surrounding the forespore. An inner cortex and outer zone, surrounded by an irregular granular epicortical layer develops. The outer spore coat is deposited on the outer membrane and forms the laminar inner coat of the mature endospore. Granular material of the spore mother cell concentrates at the basal side of the endospore and is finally engulfed by the exosporium, which surrounds the entire endospore. Sporangia are of lenticular, almost spheroidal shape, with a round to conical upper part and an irregularly shaped basal part. When and where the endospores are





12


released from the sporangia remains unknown. The germination of a single spore is enough to create infection in a Meloidogyne sp. female (Stirling, 1984). Generally the infection rate of females increases with increasing endospore attachment on the juvenile (Davies et al., 1988; Stirling, 1984).

- Pasteuria penetrans is morphologically different from P. thornei and P.

nishizawae in shape and size of the sporangia and the endospores (Sayre et al., 1988; 1991). Light microscopy revealed that mature endospores of P. penetrans, including parasporal fibers and sporangial wall, are saucer to bowl-shaped, measuring about 4.5 gm in diameter and about 3.6 gm in height. Endospores of P. nishizawae are of similar shape with a broadly elliptical central body, averaging 5.3 gm in diameter and about 4.3 gm in height. Endospores of P. thornei, however, are rhomboidal shaped and smaller in size, averaging 3.5 gm in diameter and 3.1 pm in height. The translucent circular perisporium is easily differentiated from the distinct spore wall. In P. penetrans and P. nishizawae, the spore wall decreases in thickness toward the center of the basal side of the endospore. In P. penetrans, a morphological discreet area has been observed on the basal side of the endospore, which appears to form the germination pore.

Unlike females, males of Meloidogyne spp. seem to become parasitized by P.

penetrans rather rarely (Abrantes and Volas, 1988; Freitas et al., 1996; Hatz and Dickson, 1992; Page and Bridge, 1985). Endospores obtained from seven isolates of P. penetrans did not attach to males of M. arenaria (Freitas et al., 1996), and observations by Hatz and Dickson (1992) suggest that infected males were a consequence of sex reversal because no males with a single gonad were observed to contain endospores.

Some isolates of P. penetrans have been reported to complete their life cycles in J2 of Meloidogyne spp. (Giblin-Davis et al., 1990; Dickson et al., 1994). Other Pasteuria spp. from Tylenchulus semipenetrans Cobb and Heterodera avenae have endospores that





13


are morphologically similar to those of P. penetrans, but they reproduce in J2 and not in females (Davies et al., 1990; Fattah et al., 1989; Kaplan, 1994; Sturhan et al., 1994; Winkelheide, 1993). The isolate specific to T. semipenetrans also reproduce in the male (Kaplan, 1994). Another isolate of P. penetrans was able to reproduce in Pratylenchus scribneri and Meloidogyne spp. (Oostendorp et al., 1990). Pasteuria sp. from Helicotylencus lobus formed mature endospores in the juvenile, female, and male life stages (Ciancio et al., 1992). The life cycle of P. thornei can be completed in any of the juvenile stages and in the adult of its host, Pratylenchus brachyurus (Starr and Sayre, 1988).



Ecology of Pasteuria spp.



Pasteuria spp. are widespread in different biotopes and habitats (Ciancio et al., 1994). Their spores have the morphological and biochemical features of bacterial endospores (Sayre and Wergin, 1977; Williams et al., 1989), and therefore can tolerate environmental extremes. Factors such as soil moisture, soil temperature, soil pore size, organic matter, and clay content are important in the ecology of P. penetrans.

Geographic distribution. Pasteuria spp. are worldwide in distribution and have been reported from many countries (Table 1-1).

Moisture level. Endospores of Pasteuria spp. are resistant to desiccation (Williams et al., 1989b). The water content of soils, however, affects hatching and movement of nematodes (Baxter and Blake, 1969) and, therefore, is likely to influence the efficacy of P. penetrans. Davies et al. (1991) reported soil moisture affects the growth of P. penetrans within developing females. However, P. penetrans was cultivated successfully in a hydroponic solution (Serracin et al., 1997), which suggests that the failure





14


TABLE 1-1. Geographic distribution of Pasteuria spp.


Literature source


North America
Canada U.S.A.


U.S.A., California

U.S.A., Colorado U.S.A., Florida



U.S.A., Georgia U.S.A., Hawai U.S.A., Illinois U.S.A., Louisiana U.S.A., Maryland U.S.A., Oregon U.S.A., South Carolina U.S.A., South Dakota U.S.A., Utah Central America
Colombia Cuba Dominican Republic Haiti
Nicaragua Puerto Rico South America
Bolivia Brazil Peru
Venezuela Europe
Austria Belgium Croatia Denmark England Finland France Germany Greece Hungary Iceland Italy


Gonzales et al., 1987; Sayre and Starr, 1988; Sturhan, 1988 Allen, 1941; Altherr and Deboutteville, 1972; Bernard and Niblack, 1982; Ciancio et al., 1994; Minton and Sayre, 1989; Sayre and Starr, 1988; Sturhan, 1988 Ciancio et al.,1992; Ciancio and Mankau, 1989b; Sayre and Starr, 1988
Sayre and Starr, 1988 Chen, 1996; Esser, 1980; Giblin-Davis et al., 1990; Hewlett et al., 1994; Inserra et al., 1992; Kaplan, 1994; Oostendorp et al., 1990; Sayre and Starr, 1988; Walter and Kaplan, 1990; Weibelzahl-Fulton et al., 1996
Sayre and Starr, 1988; Minton and Sayre, 1989 Ko, 1995; Sayre and Starr, 1988 Noel and Stanger, 1994 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988

Ciancio and Mankau, 1989a Sturhan, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988; Sturhan, 1985; 1988 Vargas and Acosta, 1990; Vargas et al., 1992

Ciancio and Mankau, 1989b; Page and Bridge, 1985; Sturhan, 1988 Dos Santos, 1981; Sayre and Starr, 1988; Sturhan, 1988 Ciancio and Mankau, 1989a; Ciancio et al., 1994 Sayre and Starr, 1988

Sayre and Starr, 1988 Sayre and Starr, 1988 Ciancio et al., 1994 Williams, 1960 Davies et al., 1990; Sturhan, 1988 Sayre and Starr, 1988 Sturhan, 1985; 1988 Sayre and Starr, 1988; Sturhan, 1988; 1985; Steiner, 1938; Vovlas et al., 1993; Sayre and Starr, 1988 Ciancio et al., 1994 Sayre and Starr, 1988; Sturhan, 1988 Abrantes and Vovlas, 1988; Ciancio,1995; Ciancio et al., 1987; 1994; Davies et al., 1990; Roccuzzo and Ciancio, 1991; Walia et al., 1990


Location


Location





15


TABLE 1-1. Con
The Netherlands Poland Portugal Romania Scotland Spain Sweden Switzerland Africa
Algeria Ethiopia Ivory Coast Liberia Malawi Mozambique Nigeria Senegal Sierra Leone Somalia South Africa Tanzania Togo Uganda Zaire Zimbabwe Asia
China India

Iran
Iraq
Israel Japan Pakistan South Korea Sri Lanka U.S.S.R.
Australia
South Australia

Islands
Azores Canary Islands Madeira Islands Malta Mauritius Philippines Samoa Sio Tomd


tinued


Kuiper, 1958; Sayre and Starr, 1988 Ciancio et al., 1994 Abrantes and Vovlas, 1988; Ciancio et al., 1994; Sayre and Starr, 1988 Sayre and Starr, 1988 Verdejo-Lucas, 1992 Sayre and Starr, 1988 Sayre and Starr, 1988; Sturhan, 1988

Ciancio et al., 1994 Ciancio et al., 1994 Sturhan, 1988 Ciancio et al., 1994 Sturhan, 1988 Sturhan, 1988; Sayre and Starr, 1988 Sayre and Starr, 1988; Sturhan, 1988 Mankau, 1980 Ciancio et al., 1994; Sayre and Starr, 1988 Ciancio et al., 1994 Sayre and Starr, 1988; Spaull, 1981; Sturhan, 1988 Madulu et al., 1994; Siddiqi, 1991 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Stubbs and Gowen, 1996

Pan et al., 1993 Bhattacharya and Swarup, 1988; Page and Bridge, 1986; Sharma and Davies, 1996; Sharma and Sharma, 1989; Walia et al., 1990 Barooti, 1989; Maafi, 1993; Sayre and Starr, 1988; Sturhan, 1988 Fattah et al., 1989 Ciancio et al., 1994; Sayre and Starr, 1988 Sayre and Starr, 1988; Sayre et al., 1991a; Sturhan, 1988 Maqbool and Zaki, 1990 Ciancio et al., 1994 Ciancio et al., 1994 Sturhan, 1988; Subbotin et al., 1994

Bird and Brisbane, 1988; Sayre and Starr, 1988; Stirling and White, 1982; Sturhan, 1988

Sayre and Starr, 1988; Sturhan, 1988 Sayre and Starr, 1988; Sturhan, 1988 Sayre and Starr, 1988 Ciancio et al., 1994 Ciancio et al., 1994; Sayre and Starr, 1988; Williams, 1960 Ciancio et al., 1994; Sturhan, 1988 Sayre and Starr, 1988 Ciancio et al., 1994





16


of Davies' system was probably due to poor development of the root system in wet soils. Isolates of P. penetrans have survived for several weeks in dry, moist, and wet soils, and in soils with fluctuating moisture levels without loss of their ability to attach to their nematode hosts (Oostendorp et al., 1990). Their abundance in soils is positively correlated with increasing amounts of annual rainfall (Ko et al., 1995).

Temperature. Attachment, infection, and pathogenesis of Pasteuria spp. is affected by temperature (Ahmed and Gowen, 1991; Freitas et al., 1997; Hatz and Dickson, 1992; Nakasono et al., 1993; Sekhar and Gill, 1990a; Serracin et al., 1997; Serracin-Ulate, 1995; Singh et al., 1990; Stirling, 1981; Stirling et al., 1979, 1990). Stirling (1981) found that temperatures near or above the optimum for the nematode caused P. penetrans to attach at higher rates and to proliferate extensively before the infected female host reached maturity; in contrast, at 20 *C the rate of attachment decreased and females often developed ovaries containing eggs before infection prevented further development.

Endospore densities of P. penetrans in soil increase with an increase of the average annual temperature (Ko et al., 1995). Walker and Wachtel (1988) observed that soil solarization with clear polyethylene increased the rate of infection of Meloidogynejavanica by P. penetrans. The higher soil temperature may have led to an increase in attachment rates because of the likely increase in nematode mobility, thus leading to an increased probability of endospore contact with J2; on the other hand the higher temperature may have simply increased the rate of nematode development, thereby increasing endospore numbers in soil. Recent studies, however, have shown that attachment of endospores to J2 was reduced after endospores were exposed to 40, 50, and 60 *C (Freitas et al., 1997). Attachment occurred following heating of endospores to 80 *C for 30 minutes, but P. penetrans did not develop inside the nematode (Dutky and Sayre, 1978). Although the rate of attachment has been reported to be reduced when endospores were treated at 100 C for





17


at least 30 minutes, attachment was not prevented (Freitas, 1997; Stirling et al., 1986; Williams et al., 1989b). The greatest receptivity of J2 of M. arenaria to endospore attachment occurred when J2 were treated in water at 30 'C and 35 C and then exposed to endospores (Freitas et al., 1997).

- Soil type. Pasteuria penetrans was detected in a wide variety of soil types ranging from pure sand to organic soils (Sturhan, 1985). Observations in laboratory experiments suggest that an increasing sand content might improve endospore attachment (Singh and Dhawan, 1992). Other workers demonstrated that soil texture had no effect on endospore attachment (Hewlett, personal communication). Sandy soils allow endospores to distribute readily with percolating water (Oostendorp et al., 1990).

Organic matter. The organic matter component of soil is unlikely to influence

Pasteuria spp., other than providing fungal antagonists of nematodes with more favorable habitat and food source (Kerry, 1993). Habitat and food sources may influence the biological control attributes of P. penetrans (Dube and Smart, 1987; Maheswari and Mani, 1988). Mode of action of organic amendments against nematodes consists of more than the direct effects on nematophagous fungi. When organic soil amendments in the form of oil cakes were added to P. penetrans-infested soil in a pot experiment, M. javanica was synergistically reduced (Maheswari et al., 1988). Organic amendments can improve soil structure and soil fertility, alter the level of plant resistance, release toxic compounds, and stimulate nematode antagonistic microorganisms (Stirling, 1991). All of these changes by organic amendments may affect soil nematode densities dramatically.

Chemical factors. Pasteuria penetrans was detected in acidic as well as in alkaline soils (Sturhan, 1985). However, the optimum pH for attachment of endospores of Pasteuria spp. is between 7.0 and 8.5 (O'Brian, 1980). The endospore's resistance to chemical compounds allows them to survive nematicide applications (Mankau and





18


Prassad, 1972; Stirling, 1984). Nematostatic field dosages of 1,3-dichloropropene, aldicarb, carbofuran, fenamiphos, and ethoprop had no noticeable effect on P. penetrans (Mankau and Prassad, 1972). Increased attachment by P. penetrans was observed when subnematostatic concentrations of organophosphate or organocarbamate nematicides were applied to soil containing nematodes and endospores (Brown and Nordmeyer, 1985). As reported by Bunt (1987), the low concentration of a nematicide might have increased the random nematode movement in the soil, which would increase the likelihood of contact with bacterial endospores. Freitas (1997) reported that chloropicrin alone or in combination with methyl bromide was highly detrimental to the development of P. penetrans because endospore formation was inhibited.

Host-parasite relationship. Pasteuria penetrans parasitism has been recorded for more than 236 nematode species in 102 genera, including plant parasitic and free living species (Ciancio et al., 1994; Sturhan, 1985; 1988). In several observations the hostparasite relationship is based on cuticular attachment only. Stirling (1991) concluded that such published data should be treated with caution, because endospore attachment does not actually determine parasitism. Because of its obligate nature, the bacterium is unable to reproduce in the absence of the host and would, therefore, be disadvantaged if it eliminated its host. With increasing endospore densities in soils, the infectivity of endosporeencumbered J2 of Meloidogyne sp. has been shown to be reduced, thus leading to the production of fewer endospore-filled females (Stirling, 1991). Endospore densities are likely to stabilize at a certain equilibrium level (Stirling, 1991). Williams (1960) and later Spaull (1984) noted this density-dependent relationship between P. penetrans and its Meloidogyne host.





19


Biological Control of Nematodes by Pasteuria spp.



Biological control attributes

Since the time that Sayre and Starr (1988) reported Pasteuria spp. to be parasitic to most-species of plant-parasitic nematodes, much effort has been made to understand P. penetrans isolates that parasitize the economically important root-knot nematodes. Pasteuria penetrans has many of the attributes required by a successful biological control agent. The bacterium produces no environmental hazards and reduces or prevents reproduction of its host. Also, it reduces the infectivity of endospore-encumbered juveniles to their plant host. Infectivity may be reduced when as few as 15 endospores are attached to the juvenile (Davies et al., 1988). Juveniles are prevented from invading roots when they are each encumbered with 25 to 30 endospores (Stirling, 1984; Stirling et al., 1990).

Host specificity. Isolates of P. penetrans can vary in specificity to different

Meloidogyne spp. (Brown and Smart, 1984; Davies et al., 1988; Davies and Danks, 1992; 1993; Oostendorp et al., 1990; Sell and Hansen, 1987; Spaull, 1984; Stirling, 1985; Verdejo-Lucas, 1992). Field populations of Meloidogyne spp. may be diverse, and it is possible that populations of P. penetrans with a restricted host range will affect nematode species to varying degrees. However, if there is time for natural selection to take place, it may be possible for new strains of Pasteuria spp. to evolve that are better fitted to the local environment or that are more virulent to the local nematode population. Davies et al. (1988), and Channer and Gowen (1992) support this concept by demonstrating that an initially poor host of a P. penetrans isolate became more susceptible to the parasite when exposed to endospores grown on nematodes of its own species. However, individual nematodes escaping parasitism produced a generation with increased resistance to attachment by endospores of that particular P. penetrans isolate. The authors concluded





20


that populations of P. penetrans and Meloidogyne spp. can be genetically heterogeneous with respect to host specificity and susceptibility, respectively. Thus, an isolate of P. penetrans with a wide host range of plant-parasitic nematodes needs to be employed to ensure durable biological control (Channer and Gowen, 1992; Tzortzakakis and Gowen, 1994b). Genetic diversity within a population of P. penetrans may enable the organism to maintain itself in situations in which nematode populations are altered by a change in cropping practice (Stirling, 1991).

Environmental factors. Endospores can be stored in air-dried soil and root material for up to several years without apparent loss in the ability to attach to host nematodes (Mani, 1988; Mankau and Prassad, 1977; Stirling and Wachtel, 1980). Their viability was retained following repeated freezing and thawing (Bird et al., 1990). Since the endospores are relatively small, they can be applied easily to the soil surface and redistributed by percolating water (Oostendorp et al., 1990), which is a particularly useful attribute for a biological control agent (Stirling, 1991).

Obligate parasitism. The apparent obligate nature of parasitism by Pasteuria spp. limits their biological control potential for two reasons. Spores that lose their viability or are consumed by other soil organisms can only be replaced in the presence of host nematodes and susceptible host plants. - More problematic is the lack of efficient technology for the industrial production of the organism (Bishop, 1991; Dickson et al., 1994; Reise et al., 1988; Stirling, 1991). This constraint stimulates efforts in the exploitation of naturally occurring isolates of P. penetrans for control of Meloidogyne sp. in home gardens, small-holdings of subsistence farmers (Gowen and Ahmed, 1990), and horticultural industries (Dunn, personal communication; Giblin-Davis, 1990).





21


Natural control

Plant-parasitic nematodes and their bacterial parasites have co-evolved for a long time (Ciancio et al., 1994). There are few data on the extent of nematode suppression caused by Pasteuria sp. in undisturbed soil or in soils with continuing monoculture. With increasing awareness about the qualities of the hyperparasites, more effort have been invested in long-term field studies, and the processes in which soils become suppressive to nematodes are becoming better understood (Chen, 1996b; 1997c; Dickson et al., 1994; Oostendorp et al., 199 1a; Stirling, 1991; Weibelzahl-Fulton et al., 1996).

Pasteuria sp. may have little short-term impact on nematode populations at low spore densities because only a few nematodes come in contact with the endospores. However, with each infected root-knot nematode female capable of producing up to 2.5 million endospores, endospore densities increase over time, thereby reaching nematode suppressive levels (Oostendorp et al., 1991 a; Chapter 3). Williams (1960) and later Spaull (1984) noted that P. penetrans had little impact on the severity of root-knot nematodes in a natural infestation of both parasites; however, the bacterial parasite may still have been in an early phase of population build-up. Others have reported soils with root-knot nematodes and P. penetrans becoming highly suppressive to the nematode (Bird and Brisbane, 1988; Chen et al., 1994; Dickson et al., 1994; Minton and Sayre, 1989; Stirling, 1984; Weibelzahl-Fulton and Dickson, 1996; Weibelzahl-Fulton et al., 1996). When rootknot nematode and P. penetrans infested soils were tested for their suppressiveness, the rate of reproduction of Meloidogyne spp. was frequently reduced in untreated soils compared to soils in which P. penetrans had been eliminated by autoclaving (Dickson et al., 1994; Weibelzahl-Fulton et al., 1996). Nematode suppression was enhanced by cultivation of susceptible hosts for several consecutive years.





22


In a pot experiment, soil naturally infested with Belonolaimus longicaudatus Rau and a Pasteuria species specific to the nematode was not suppressive to its host nematode for the first 6 months (Giblin-Davis et al., 1990), however, at 12 months post-inoculation there was an increase in the number of Pasteuria-filled nematodes. The nematode density was significantly lowered (Giblin-Davis, 1990).



Inundative application

Despite the difficulties scientists originally faced in working with spore-infested soil, P. penetrans gave promising results against M. incognita in pot experiments (Mankau, 1975b). An improvement of the mass-production technique allowed scientists to produce a spore-laden, easily handled root powder (Stirling and Wachtel, 1980). This provides a means for more extensive testing and further confirmation of the bacterium's potential as a biological control agent. Control of Meloidogyne sp. by P. penetrans was reported from greenhouse tests (Brown and Nordmeyer, 1985; Channer and Gowen, 1988; Maheswari and Mani, 1988, Maheswari et al., 1987; 1988; Raj et al., 1988; Raj and Mani, 1988; Tzortzakakis, 1994a; Vargas et al., 1992), microplot studies (Brown et al., 1985; Chen et al., 1996b; 1997c; Daudi, 1990; Dube and Smart, 1987; Oostendorp et al., 199 1a), and small-scale field experiments (Channer and Gowen, 1988; Stirling, 1984).

Several investigators have provided estimates of the endospore population density needed in the soil to provide nematode control (Brown and Smart, 1985; Chen et al., 1996b; Davies et al., 1989; 1990; Sell and Hansen, 1987; Stirling, 1984). A concentration of 10' endospores/g of soil prevented J2 of Meloidogyne spp. from infecting plant roots when the nematode moved either 4 or 8 cm through the soil (Stirling et al., 1990). Nematodes that moved 2 cm through the soil were infective but they did not produce progeny because of infection by the bacterial parasite. Concentrations as low as 104





23


endospores/g of soil reduced root-knot nematode fecundity (Ahmed and Gowen, 1991; Gowen et al., 1989), and caused suppression of M. arenaria infection on peanut (Chen et al, 1996b). Densities of 10' endospores/g of soil throughout the top 15 cm provides levels of control comparable to a nematicide application (Stirling, 1991). Most recently, Chen et al. (1-996b) demonstrated that adding 10,000 to 100,000 endospores per g of soil to rootknot nematode infested soil provided control of M. arenaria on peanut in the first season. Application of P. penetrans endospores 2.5 cm deep in soil appeared to be more effective for parasitism of M. incognita on tomato than a surface application or an application 5 cm deep (Ahmad et al., 1994).



Integrated nematode management

There have been several attempts to integrate P. penetrans with other nematode

management techniques. Synergistic effects on the management of Meloidogyne spp. were reported from greenhouse experiments in which P. penetrans was combined with organophosphate or organocarbamate nematicides at nematostatic field application rates (Maheswari et al., 1987; Tzortzakakis, 1994a) or at subnematostatic rates as low as 1.5 ppm and 0.25 ppm, respectively (Brown and Nordmeyer, 1985). Successful control of Meloidogyne spp. also may be obtained by combining P. penetrans with one or several other biological control strategies. De Leij et al. (1992) noted that the egg mass-colonizing fungal endoparasite Verticillium chlamydosporium Goddard complimented the suppression of M. incognita by P. penetrans by attacking a different life stage of the nematode, thus giving better results than either of the organisms alone or the nematicide treatment. Similar results were obtained in experiments where Paecilomyces lilacinus (Dube and Smart, 1987; Maheswari and Mani, 1988; Shahzad et al., 1990; Zaki and Maqbool, 1991),





24


Talaromycesflavus, or Bacillus subtilis (Zaki, 1991) were added as additional nematode antagonists, or where oil cakes were added as a soil amendment (Maheswari et al., 1988).

The efficacy of P. penetrans was enhanced with soil solarization applied alone

(Freitas, 1997; Walker and Wachtel, 1989) and in combination with an organophosphate nematicide (Tzortzakakis, 1994a; Walker and Wachtel, 1989). The population density of P. penetrans also was affected by crop rotations and winter cover crops. In the presence of a susceptible host plant, the density of P. penetrans endospores build up in the presence of an increasing Meloidogyne spp. population (Brown et al., 1985; Chen et al., 1994; Madulu et al., 1994; Oostendorp et al., 1991a). A non-host, resistant host, or fallow rotation has been shown to hamper the increase in endospore density (Chen et al., 1994; Madulu et al., 1994; Oostendorp et al., 199 1a). Summer crops were generally more effective than winter cover crops, which is probably explained by the high temperature dependence of both the root-knot nematode and its bacterial parasite.



Useful Methods and Techniques for Studying Pasteuria spp.



Detection of Pasteuria spp. in soils and in nematodes

Pasteuria spp. can be detected readily on or in nematodes using a number of

simple techniques. Endospore-encumbered or endospore-filled migratory nematode stages have to be extracted from the soil. One method for their extraction is to use a centrifugalflotation technique (Jenkins, 1964). Increasing the specific gravity of the sucrose solution used in the centrifugal-flotation technique from 1.14 to 1.22 and 1.26 led to recovery of a higher number of spore-filled bodies of Pratylencus scribneri and B. longicaudatus, respectively (Oostendorp et al., 1991b). Extraction efficacy of endospore-filled Hoplolaimus galeatus (Cobb) Thorne was not increased with the denser sucrose solutions.





25


Nematodes may be examined for adhering or internal endospores with the aid of an inverted microscope at X400 magnification. For easier detection, endospores adhering to the body of a nematode may be stained with Brilliant Blue G (Bird, 1988), or labeled with fluorescein (Charnecki, 1997; Charnecki et al., 1996). Detection of endospores in soil can be increased by adding host nematodes to a soil sample and storing the sample for several days to allow the nematodes to migrate through the soil, or by adding the nematodes and water to the soil and shaking the contents for 24 hours before extraction (Stirling and White, 1982). The attachment of endospores to the nematode body does not indicate infection; evidence of internal parasitism is needed to confirm that the nematode is actually a host. The parasite's presence can be confirmed by squashing females on a glass slide and observing them for the presence of vegetative stages or endospores of the parasite (Hatz and Dickson, 1992). For better recognition of the different developmental stages of the bacterium, specimens can be heat fixed and Gram stained (Cappuccino and Sherman, 1986). A simpler technique by Serracin et al. (1997) used lactophenol and 1% methyl blue stain to dye the bacterial structures inside the nematode body. However, its application did not successfully stain bacterial structures in vermiform J2 of Meloidogyne sp.



Isolation of Pasteuria spp. from soils and nematodes

With the exception of the soil-inhabiting mature endospore, the bacterium's other life stages can only be found inside nematode bodies. In sedentary nematodes, such as Meloidogyne spp., Tylenchulus semipenetrans, Heterodera glycines, H. goettingiana, and Globodera rostochiensis, early life stages of Pasteuria spp. can be found in sedentary juvenile stages and in the adult female (Sayre et al., 1991 a; Sturhan et al., 1994; Oostendorp, personal communication). These nematode life stages can be dissected from enzymatically digested plant tissue (Hussey, 1971). Endospore-filled females of





26


Meloidogyne spp. are usually white, opaque and larger than uninfected females (Hatz and Dickson, 1992). The infected females can be distinguished visually and collected by hand with the use of a stereomicroscope. Migratory life stages of sedentary nematodes and ectoparasitic nematodes parasitized with Pasteuria spp. must be hand-picked from nematode suspensions after they are extracted from soil. By surface sterilizing infected nematodes and squashing them in sterilized water, different life stages of Pasteuria spp. can be observed free of contaminating microorganisms (Williams et al., 1989).



Ouantification of endospores in soils, root powder, or suspensions

The quantification of the bacterial endospores is important, because the efficacy of Pasteuria spp. is density dependent (Stirling, 1991). However, methods for enumeration of Pasteuria spp. in soils have received little attention, and endospore densities in the soil are usually based on the number of endospores attached to soil inhibiting nematode stages. Time and effort to determine the number of endospores of P. penetrans attached to J2 of Meloidogyne spp. may be reduced considerably by using tally thresholds (Chen and Dickson, 1997).

Juveniles of M. javanica were used as probes for P. penetrans (Stirling, 1984; Stirling and White, 1982), and since there is a direct relationship between endospore concentration in soil and the number of endospores attached (Stirling et al., 1990), a bioassay was developed to estimate the relative endospore density in soil (Oostendorp et al., 199 1a). By using a particular probe nematode in the bioassay, only that component of the P. penetrans population that attaches to that nematode is detected.

Endospores also have been extracted from soil by differential-centrifugation (Davies et al., 1990); however, the technique is tedious and difficult to use. The total number of endospores in a particular soil may not accurately represent the suppressivness





27


of the soil; thus according to Stirling (1991), a bioassay is a more useful quantification method than the use of a direct extraction method.

Chen et al. (1996a) observed that the highest estimate of the number of endospores in root material was obtained by suspending machine ground root powder in deionized water and pouring the material onto a 250-jim-pore sieve. Endospores passed through the sieve and were concentrated in water, and the number of endospores was calculated with the help of a hemacytometer. This refined quantification allowed an improvement compared to the previous application of the inoculum by weight (Brown et al., 1985; Dube and Smart, 1987; Raj and Mani, 1988; Stirling, 1984; Zaki and Maqbool, 1991; 1992a; 1992b).



Culture and preservation of Pasteuria penetrans

Methods have been developed for the production of P. penetrans endospores in females of Meloidogyne spp. parasitizing plant roots. This is currently necessary because in vitro cultivation has not been successful (Bishop and Ellar, 1991; Previc and Cox, 1993; Reise et al., 1988). Pasteuria penetrans-encumbered J2 of Meloidogyne sp. must be inoculated onto a susceptible host, such as tomato (Stirling and Wachtel, 1980). Once the bacterium has developed to maturity, the root systems may be harvested, washed, air dried, and ground into a fine endospore-laden powder. Although tomato plants are most commonly used, cucumber, Cucurbita vulgaris L., was shown to yield higher endospore numbers than tomato (Cho et al., 1997). This system may produce a highly variable yield of endospores; however, it is superior to the use of endospore infested soil (Dutky and Sayre, 1978; Mankau and Prassad, 1977). The plant system has been optimized (Sharma and Stirling, 1991), and most recently a hydroponic cultivation system has been reported (Serracin-Ulate, 1995). Verdejo and Jaffee (1988) used a gnotobiotic technique for





28


producing endospores; however, this method requires rigorous aseptic conditions and is therefore not a good procedure for general use.

The time period required for P. penetrans to complete its life cycle has been shown to vary with different isolates (Hatz and Dickson, 1993; Serracin et al., 1997; Stirling, 198 1). Thus, a random determination of the progress of the bacterium's development in a cultivation system can help avoid harvesting prematurely, thus improving the efficiency of the endospore yield. The lack of an efficient technology for the mass-cultivation of Pasteuria spp. endospores is the greatest impediment to the use of this microbe as a biological control agent (Dickson et al., 1994; Stirling, 1991).



Attachment

Endospore-host attachment studies are the first step in establishing the host specificity of Pasteuria spp. and determining their biological efficacies. Most of the techniques rely on nematode movement through soil (Brown and Smart, 1985), water (Channer and Gowen, 1988), or agar (Verdejo and Jaffee, 1988), when each is laden with endospores, or the agitation of a nematode-endospore-water suspension (Ahmed et al., 1990; Bird, 1986; Bird et al., 1990; Davies et al., 1988; Oostendorp et al., 1990). Hewlett and Dickson (1993) consistently achieved attachment of P. penetrans to Meloidogyne spp., H. galeatus, and B. longicaudatus using a centrifuge technique. The method is fast, and allows studies of Pasteuria spp. collected from nematode species that yield relatively few endospores per cadaver.





29


Objectives



The objectives of this dissertation were 1) to monitor the population densities of Meloidogyne spp. and P. penetrans in microplots and in a naturally infested tobacco field; 2) to develop a test for soil suppressiveness that allowed the determination of the role of P. penetrans in suppressing Meloidogyne spp. populations in both microplots and tobacco field plots; and 3) to evaluate the effect of cultural practices, such as an intercropping system with corn and beans in rotation with peanut, autumn cover crops, resistance, and fertilizer regimes on the population development of both M. arenaria and P. penetrans.














CHAPTER 2
POPULATION DEVELOPMENT OF MELOIDOGYNE ARENARIA RACE 1 AND
PASTEURIA PENETRANS IN A 6.5-YEAR MICROPLOT STUDY



Introduction



The peanut root-knot nematode, Meloidogyne arenaria (Neal) Chitwood race 1, is one of the most important soil pathogens of commercially grown peanut (Arachis hypogaea L.) (Minton, 1984). It occurs on peanut in many countries and is especially troublesome in the southeastern United States (Minton, 1984; Porter et al., 1984). Continuing environmental problems associated with the use of nematicides (Thomason, 1987) has resulted in more scientists studying nematode management strategies alternative to chemical control (Kerry, 1990). The use of microbial agents for biological control of plant-parasitic nematodes is an alternative management tactic that is receiving increased interest among nematologists. Among many organisms identified as antagonists of plantparasitic nematodes, the endospore forming bacterium, Pasteuria penetrans, has been demonstrated to have potential for the control of mainly Meloidogyne spp. It has been suggested that P. penetrans may suppress nematode population densities below economic damage levels (Chen et al., 1996b; Dickson et al., 1994; Minton and Sayre, 1989; Stirling, 1991). In order to determine the efficacy of the density-dependent obligate parasite, it is important to understand its development as influenced by its nematode host.

The population densities of P. penetrans and of M. arenaria race I with and

without the nematode antagonist were determined from the spring of 1987 to the fall of


30





31


1989 (Oostendorp et al., 199 1a). During this period, soil suppressive to M. arenaria was produced in microplots. The microplots had been planted continuously to peanut in summer and either vetch (Vicia villosa Roth) or wheat (Triticum avenae L. cultivar 302), rye (Secale cerale L. cultivar Wrens Abruzzi), or bare fallowed during the winter. Pasteuria penetrans, initially applied to the soil in relatively low numbers, was increased to numbers which suppressed M. arenaria. Within 3 years, yield of plots infected with P. penetrans and M. arenaria increased to a level comparable to the untreated control (Oostendorp et al., 1991a). The objective of this study was to continue to monitor the population densities of both M. arenaria race 1 and P. penetrans in these microplots in order to determine their long-term effects on M. arenaria densities.



Materials and Methods



In 1987, 90 microplots (76 cm in diameter), located at the University of Florida,

Green Acres Agronomy Farm, Alachua County, were initially arranged in 10 rows of nine plots each, with a distance of 1.5 m between plots, in a loamy, siliceous, hypothermic Grossarenic Paleudults with 90% sand, 4% silt, 6% clay, and 1.8% organic matter (Oostendorp et al., 199 1a). A split-plot design was used with main plots consisting of three soil treatments: an untreated control, M. arenaria alone (RKN), and M. arenaria plus P. penetrans (RKN + Pp). Three summer-winter cover crop rotations of peanut-rye, peanut-vetch through 1991, with peanut-wheat thereafter, and peanut-bare fallow were subplots. Wheat replaced vetch because the incidence of soilborne diseases on peanut increased following vetch. Each combination of soil treatments and winter cover crops was replicated 10 times. The microplots were initiated by placing 800 cm3 of soil preparation containing uninfected tomato roots, tomato roots infected with M. arenaria





32


race 1, or with M. arenaria race 1 plus P. penetrans in a hole in the center of each microplot receiving the control, RKN, or RKN + Pp treatment, respectively. Monitoring of the bacterium and nematode population densities for this dissertation began in the spring of 1990 and continued through the fall of 1993. Data from the period 1987 to 1989 are included herein to provide a complete data set on the long-term effects of P. penetrans.

Two months before planting in the spring of 1991 and 1993, uninoculated control plots were treated with 977 kg methyl bromide/hectare (98% methyl bromide plus 2% chloropicrin), applied broadcast under a 3-mm polyethylene plastic covering. On 6 May 1990, 5 May 1991, 13 May 1992, and 7 May 1993, three pairs of peanut seeds of cv. Florunner were planted 4-cm deep in an equally spaced pattern in each plot. After emergence, one of each pair of seedlings was removed. A 90-cm-high, wire-mesh fence was placed around each plot to confine the growth of the peanut foliage inside the microplot. Plots were weeded manually. Through 1991, plots were irrigated by overhead sprinklers. Thereafter, a microjet sprinkler system was installed, delivering an amount of water equivalent to 4 mm per day. Every 10 to 14 days, insects and foliar disease pathogens were controlled with esfenvalerate, chloropyrifos, insecticidal soap, chlorothalonil, or liquid sulfur.

In mid-November each year, after harvest of peanut, rye and vetch or wheat were broadcast seeded as winter cover crops, or plots were bare fallowed through the winter. Vetch, wheat, and peanut are hosts for M. arenaria race 1, but rye is a poor host. The soil in each plot was turned using a spade, and leveled between crops.

At harvest (136 to 140 days after planting), peanut plant shoots were evaluated in their appearance, and the root systems, as well as the pods and pegs, were rated for galling. A good stand of plants with a densely closed canopy was rated as 0, and a poor stand of plants with sparse canopy was rated as 5. Galling was rated according to the following





33


scale: 0 = no galls, 1 = 1-10, 2 = 11-20,3 = 21-55, 4 = 56-80, and 5 = 81-100% of roots or pods and pegs galled (Barker et al., 1986). Three to five days after the plants were lifted out of the ground, all pods were removed and placed into paper bags, dried at 60 C until the moisture content was reduced to about 10%, and weighed to determine yield. The number of second-stage juveniles (J2) of M. arenaria /100 cm3 of soil was estimated at harvest of peanut and each of the winter cover crops. Soil samples consisting of a composite of five, 2.5-cm-diameter cores per plot were taken from the top 20 cm of each microplot with the use of a cone-shaped auger. The soil was mixed and processed with a centrifugal-flotation method (Jenkins, 1964). The numbers of J2, the rate of attachment of P. penetrans, and the number of endospores attached per 20 randomly selected J2 was determined using an inverted microscope at X400 magnification.

Data were subjected to analysis of variance (ANOVA). If the attained probability level of the main plot (treatments) or the subplot (cover crop) effects was significant at P < 0.05, the means were separated by Duncan's multiple-range test. If a significant main plot X subplot interaction (P < 0.05) was observed, main plot means for each subplot and subplot means for each main plot were separated by Duncan's multiple-range test. If an insufficient number of J2 or no J2 were extracted from soil of RKN + Pp plots, replicates without J2 were disregarded for the calculation of rate of attachment with P. penetrans and the number of endospores per juvenile, and data were presented in a seperate table .



Results



After the initial 3-year period of the experiment (1987 to 1989), the number of J2 in the soil of RKN + Pp plots decreased, and remained lower than those in the RKN plots (P

0.05) (Fig. 2-1, Tables 2-1 to 2-3). The nematode population density in microplots






34


100000
J2 in RKN - -Endospores/juvenile in RKN
10000 4 -4--J2 in RKN + Pp - --Endospores/uvenile in RKN + Pp


1000
a
a
a
100
a A a

10 , , / -I


1---b -b b b
-. 1 -/ b
0.1
r- CO CO 01 01. 0 0 M N (
-0 CO - , 00 O- OD -1 a,

CL CL 06 0- .

Sam ling date

Fig. 2-1. Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100 cm3 of soil and the number of endospores attached per juvenile extracted from peanut microplots infested with M. arenaria (RKN), and M. arenaria plus P. penetrans (RKN + Pp). Analysis was based on means of 10 replicates, with 20 or fewer J2 observed per replication to determine the number of endospores per juvenile. Means labeled with a different letter are different according to Duncan's multiple-range test (P < 0.05); small letters were used for comparisons of juvenile numbers, capital letters for comparing endospore numbers. Data from 1987 to 1989, were from Oostendorp et al. (1991a).






TABLE 2-1. Effect of Meloidogyne arenaria (RKN) alone and in combination with Pasteuria penetrans (RKN + Pp), and of a rye, vetch, or bare fallow winter cover crop on yield and performance of peanut in the 5" year (fall of 1991) of a 6.5-year microplot experiment.
J2 per % J2 with Plant performance
100 cm3 of attached Endospores Yield Galling index'
soil endospores per juvenile (g/plot) Shoots' Roots Pods
Main effects
Treatment Control l b 0 b 0 b 269 a 2.9 a 0 b 0 b
RKN 677 a 55 a 1.6 a 167 b 2.1 b 2.8 a 2.6 a
RKN + Pp l b 23 a 3.7 a 252 a 2.9 a 0.2 b 0 b
Cover crop Rye 188 17 0.3 b 253 2.7 0.8 0.7
Vetch 253 25 0.1 b 224 2.6 1.2 0.9
Fallow 237 36 4.0 a 212 2.5 1.1 1
Interactions
Rye Control 0 0 0 b 281 2.5 0 0
RKN 564 40 0.7 a 193 2.1 2.5 2.1
RKN + Pp I 10, O.lca 285 3.5 0 0
Vetch Control 2 0 0 b 286 3.3 0 0
RKN 757 76 3 a 133 1.9 3.2 2.6
RKN + Pp 0 O 0"b 254 2.6 0.3 0
Fallow Control 1 0 0 c 241 2.9 0 0
RKN 709 48 1.1 b 176 2.2 3 3
RKN + Pp 2 60' 1 Ia 218 2.5 0.4 0

Means followed by no letters or the same letter within a column of three observations for main effect or 10 observations for interactions were not different according to Duncan's multiple-range test (P < 0.05).
Means of 30 (for main effects) or 10 (for interactions) replicates; 20 second-stage juveniles (J2) per replicate.
There was a treatment X cover crop interaction (P < 0.05) in the number of endospores per juvenile.
a Plant performance rating scale: 0 = good stand of plants with a closed canopy, and 5 = poor stand of plants with sparse canopy.

b Galling index: 0 = no galls, 1 = 1-10, 2 = 11-20, 3 = 21-55, 4 = 56-80, and 5 = 81-100% of roots or pods galled.
C Means of less than 20 J2 per replicate.







TABLE 2-2. Effect of Meloidogyne arenaria (RKN) alone and in combination with Pasteuria penetrans (RKN + Pp), and of a rye, wheat, or bare fallow winter cover crop on yield and performance of peanut in the 6th year (1992) of a 6.5-year microplot experiment.
% J2 with Plant performance
J2 per attached Endospores Yield Galling index'
100 cm3of soil endospores per juvenile (g/plot) Shoots Roots Pods
Main effects
Spring Fall Spring Fall Spring Fall Fall Fall Fall Fall
Treatment Control 0.1 b 103 0 Ohb 0 b 0 c 408 a 3.7 a 0.2 b 0.2 b
RKN 110 a 1,239 a 19 a 36 a 5 a 8 b 190 b 2.1 b 3.2 a 3.2 a
RKN + Pp 2 b I c 36 a 27 a 9 a 22 a 400 a 4.0 a 0 b 0 b
Cover crop Rye 28 b 385 b 18 24 4 13 a 340 3.3 1.0 1.0
Wheat 32 b 476 a 15 23 3 4 b 336 3.4 1.2 1.3
Fallow 53 a 483 a 22 50 8 12 a 322 3.0 1.2 1.2
Interactions
Rye Control 0 0 0 0 0 0 c 404 3.8 0 0
RKN 83 1,154 23 32 6 5 b 210 2.2 3.1 2.9
RKN + Pp 1 2 30c 41c 5c 34'a 405 4.0 0 0
Wheat Control 0.2 220 f 0 f 0 b 400 3.7 0.5 0.5
RKN 93 1,206 19 48 4 8 a 199 2.4 3.2 3.4
RKN + Pp 1.9 0.5 28c 20' 6' 4ca 409 4.1 0 0
Fallow Control 0 90 0 0 0 0 c 419 3.7 0.2 0.2
RKN 156 1,357 15 23 7 10b 161 1.7 3.4 3.4
RKN + Pp 1.6 0.8 50c 20' 16c 27'a 386 3.7 0 0

Means followed by no letter or the same letter within a column of three observations for main effect or 10 observations for interactions were not different according to Duncan's multiple-range test (P < 0.05).

Means of 30 replicates (for main effects) or 10 replicates (for interactions); 20 second-stage juveniles (J2) per replicate.
Plant performance rating scale: 0 = good stand of plants with a closed canopy, and 5 = poor stand of plants with sparse canopy.
b Galling index: 0 = no galls, 1 = 1-10, 2 = 11-20, 3 = 21-55, 4 = 56-80, and 5 = 81-100% of roots or pods galled.
Means of less than 20 J2 per replicate.


Ws






TABLE 2-3. Effect of Meloidogyne arenaria (RKN) alone and in combination with Pasteuria penetrans (RKN + Pp),


or bare fallow winter cover crop on yield and performance of peanut in the 7'" year (1993) of a 6.5-year microplot experiment.
% J2 with Plant performance
J2 per attached Endospores Yield Calling index '
100 cm3of soil endospores per juvenile (g/plot) Shoots' Roots Pods
Main effects
Spring Fall Spring Fall Spring Fall Fall Fall Fall Fall
Treatment Control 0 b 0 b 0 b 8 c 0 b 3 b 218 2.6 0 b 0 b
RKN 79 a 339 a 29 a 91 a 4 a 34 a 204 2.4 1.3 a 0.6 a
RKN + Pp 0 b 3 b 10 a 33 b 2 a 16 a 217 2.9 0 b 0 b
Cover crop Rye 17 b 125 ab 12 42 1.2 b 16 b 210 2.7 0.4 0.2
Wheat 26 b 93 b 14 50 3 a 25 a 229 2.7 0.5 0.1
Fallow 36 a 131 a 13 41 1.2 b 12 b 200 2.5 0.5 0.2
Interactions
Rye Control 0 b 0 b 0 13c 0 0.2cb 212 2.4 0 0
RKN 52 a 352 a 26 92 3 38 a 228 2.9 1.1 0.6
RKN + Pp 0 b 2 b IOC 20c 0.6' 9'ab 190 2.7 0 0
Wheat Control 0 b 0 b 0 10' 0 9cb 222 2.7 0 0
RKN 78 a 278 a 31 100 5 36 a 220 2.2 1.4 0.4
RKN + Pp 0 b l b 10 40c 4c 29'a 244 3.2 0 0
Fallow Control 0 b 0 c 01 0 0 0 c 220 2.7 0 0
RKN 107 a 387 a 30 82 3 27 a 165 2.0 1.5 0.7
RKN + Pp 0 b 6 b iOC 40' 0.3c 10cb 216 2.8 0 0
Means followed by no letter or the same letter within a column of three observations for main effect or 10 observations for interactions were


not different according to Duncan's multiple-range test (P < 0.05).
Means of 30 replicates (for main effects) or 10 replicates (for interactions); 20 second-stage juveniles (J2) per replicate.
There were treatment X cover crop interactions (P < 0.05) in the number of ring nematodes and the number of J2 per 100 cmi soil.

" Plant performance rating scale: 0 = good stand of plants with a closed canopy, and 5 = poor stand of plants with sparse canopy.
h Galling index: 0 = no galls, I = 1-10, 2 = 11-20, 3 = 21-55, 4 = 56-80, and 5 = 81-100% of roots or pods galled.
Means of less than 20 J2 per replicate.


and of a rye, wheat,








inoculated with P. penetrans was nearly undetectable in soil samples from 1991 to 1993. The number of P. penetrans endospores adhering to J2 increased to a high of 17.2 by 1989 and remained relatively consistent during the remaining 4-year period (Oostendorp et al., 199 1a; Fig. 2-1; Tables 2-1 to 2-3). Through this time, 10-66% of the J2 were encumbered with endospores. However, when replicates without J2 were excluded from analysis for RKN + Pp plots, the incidence of endospores per juvenile and attachment to J2 were much higher than those for 20 J2 (Table 2-4). In the fall of 1990, the endospore density of P. penetrans in RKN + Pp plots had amplified to yield an average of 19.5 endospores/juvenile (data not shown). Between 1991 and 1993, the endospore density remained at a level at which 95% to 100% of the recoverable J2 were encumbered with mean numbers of 3 to 72 endospores (Table 2-4).

In the soil of RKN plots, the number of J2 remained high throughout the

experiment (Oostendorp et al., 199 1a; Fig. 2-1, Tables 2-1 to 2-3). In the third year (1989), 2 of the 30 replicates of the RKN plots were contaminated with the bacterial parasite (Oostendorp et al., 199 1a), and P. penetrans endospores were found to have contaminated 16, 29, and all 30 RKN plots in the falls of 1991, 1992, and 1993, respectively (Figs. 2-2 to 2-4, Tables 2-1 to 2-3). In fall of 1993, endospore population densities of RKN plots had increased to averaged 91% attachment rate with 34 endospores/juvenile (Table 2-3).

Numbers of nematodes and endospore attachment levels generally did not vary

with cover crops (P < 0.05) (Fig. 2-5 , Tables 2-1 to 2-3). However, in the spring of 1989, the nematode population densities in RKN plots under vetch slightly exceeded those of other plots (Fig. 2-5 A). Second-stage juvenile numbers in vetch and bare fallowed plots exceeded those in the rye plots in the spring of 1990. However, in the spring of 1992 and spring of 1993, lower numbers of J2 were recovered from plots under rye and wheat than


38





39


TABLE 2-4. The percentage of second-stage juveniles (J2) of Meloidogyne arenaria infected with Pasteuria penetrans and the average number of endospores per juvenile in peanut microplots infested with M. arenaria and P. penetrans in the spring of 1987, and rotated with rye, vetch or wheat, and bare fallow as winter cropping sequence.

% J2 with % J2 with
endospores endospores
Cover crop attached Endospores/juvenile attached Endospores/juvenile
Fall 1991
Rye 100(5) 10(5)
Vetch (0) (0)
Fallow 100(18) 40.2(18)

Spring 1992 Fall 1992
Rye 100(12) 17.8(12) 66.7(17) 32.4(17)
Wheat 100(19) 35.3(19) 66.7(5) 8.5(5)
Fallow 95(16) 21.6(16) 100(8) 44.1(8)

Spring 1993 Fall 1993
Rye 100(1) 6(1) 100(16) 43(16)
Wheat 100(1) 42(1) 100(9) 72(9)
Fallow 100(1) 3(1) 80(62) 20(62)


Means of varying number of replicates. Numbers in parenthesis are the number of nematodes observed.





40


Replicate

S CRNR












2 NR EE HEJE







6 LEF PE PR CFCHRN N



7 CF CV CR NF N R P V P



8 FVN NF PV R PF C CR F




ElHHH H H E


10 CV CR CF NV- NR NF PV P


0% juveniles with 1-99% juveniles 100% juveniles with
endospores attached with endospores endospores attached
attached

Fig. 2-2. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1991. No second-stage juveniles were extracted from non-shaded plots. The first letter indicates the pre-experimental treatment (C = plots free of nematodes and P. penetrans, N = plots inoculated with Meloidogyne arenaria, and P = plots inoculated with M. arenaria plus P. penetrans), and the second letter indicates the winter cover crops (V = vetch, R = rye, and F = fallow).





41


Replicate

1 W R FNW NR F EEE P







3 PF W CR CF



4 CR CF



5 NF NW NR PF PW R CF C CR



6 RNR


7 CFl CWHIE7E




9 CW P F CW CR CF NW NR NFPW P







10 CW CR CF NW NR NF


0% juveniles with 1-99% juveniles 100% juveniles with
endospores attached wxith endospores endospores attached
[ dittached

Fig. 2-3. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1992. No second-stage juveniles
(12) were extracted from non-shaded plots. The first letter indicates the pre-experimental treatment (C = plots free of nematodes and P. penetrans, N = plots inoculated with Meloidogyne arenarna, and P = plots inoculated with M. arenaria plus P. penetrans), and the second letter indicates the winter cover crops (W = wheat, R = rye. and F = fallow).





42


Replicate




















3 R P WCR C CWNR P F N
4 CR CF C R N W P F P








6 PF PW PR CF CW CR NF NW R



7 CFE EU LW








9fl PR FlCW CR CFF 10 CW CR CHR N W P


0 % juveniles with 1-99% juveniles
endospores attached with endospores
attached


100% juveniles with endospores attached


Fig. 2-4. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1993. No second-stage juveniles
(J2) were extracted from non-shaded plots. The first letter indicates the pre-experimental treatment (C = plots free of nematodes and P. penetrans, N = plots inoculated with Meloidogyne arenaria, and P = plots inoculated with M. arenaria plus P. penetrans), and the second letter indicates the winter cover crops (W = wheat, R = rye, and F = fallow).







43








100000
A -- -Bare fallow
9 Rye
---A -- Vetch-wheat


10000






01000

0%
rX a
--- a Na/a
a.a


5 100 b b bb
b
E
z

10



100000
B -0--Bare fallow
Rye
10000 ---A-- Vetch-wheat


0
"1000

E

g 100


k
NV 10


.0
A ------E .-, | ,-I


0.1
N- 00 0 a) 0) 0 0 ;1 - N' C\J M' CI)
CF C N e CO d a e s ) 0) a ra n af






soletace ao s ai icolt wihA) a. reai , ,a, ana, rnai lsP enras Pat
weepatd= anu insme and bar faw, or plate tore or aec (18-9 ad wha
(a cc a CO CO M Ca(
LL C LL L L LL LL LLa . LU.
CO U) U) U) (n (n
Sampling date





Fig. 2-5. Number of second-stage juveniles (J2) of Meloidogyne arenaria race I in 100 cm' of soil extracted from soil in microplots with A) M. arenaria, and B) M. arenaria plus P. penetrans. Plots were planted to peanut in summer and bare fallowed, or planted to rye, or vetch (1987-1990) and wheat (1991-1992) in winter. Means of the cropping sequences at individual sampling dates labeled by no or the same letter are not different according to Duncan's multiple-range test (P < 0.05).





44


rye and wheat than from those under bare fallow (P 0.05). In the fall of 1991, in RKN plots the estimated endospore population density in vetch-wheat plots was highest, and, together with rye plots, exceeded the endospore counts of fallow plots in the fall of 1993 (P < 0.05) (Fig. 2-6 A). From the spring of 1989 till the spring of 1990, the attachment level and the number of endospores per juvenile in RKN + Pp plots under vetch were higher than under bare fallow (P < 0.05) (Fig. 2-6 B). Thereafter, the endospore counts in vetchwheat were frequently lower than in bare fallow plots. In the fall of 1993, the vetch-wheat plots yielded the highest estimated endospore population density. In the fall of 1990 and of 1992, the endospore counts in rye plots exceeded those of bare fallow plots and vetchwheat plots, respectively.

According to main effect means in the spring and the fall of 1993, the attachment rates of 3 and 25 endospores per juvenile across all nematode treatments in wheat plots were higher (P < 0.05) than the levels observed in rye or fallowed plots (Table 2-3). The highest number of 107 J2/100 cm3 of soil in fallowed RKN plots observed in the spring of 1993 was numerically greatest, but yet consistent with the main effect of highest juvenile levels in RKN plots.

Following the initial increase in peanut yield in RKN + Pp plots after 3 years, the

yields remained high and not different from those in control plots for the remaining 4 years (P 0.05) (Fig. 2-7, Tables 2-1 to 2-3). In 1993, there were no differences among yields in any treatments, which is attributed to P. penetrans infesting RKN plots (Fig. 2-6 A, Table 2-3). Meloidogyne arenaria was low or undetectable in control plots throughout the experiment. Throughout the last 4 years, the galling indices for roots and pods in RKN plots were higher than gall ratings of RKN + Pp and control plots (P 0.05) (Tables 2-1 to 2-3).







45


--e- -Bare fal
" Rye
---A-- Vetch-v


40


35






25 30
CL


01
6a 25 15
0 10




E

5


.3 -- U- -


0 0)

0)


Sampling date


0)


0)


0)


a)


COj
M-


CO) U)


Fig. 2-6. Number of attached endospores per juvenile extracted from soil in microplots with A) M. arenaria, and B) M. arenaria plus P. penetrans. Plots were planted to peanut in summer and bare fallowed, or planted to rye, or vetch (1987-1990) and wheat (1991-1992) in winter. Means of the cropping sequences at individual sampling dates labeled by no or the same letter are not different according to Duncan's multiple-range test (P < 0.05). Means of 10 replicates, with 20 or fewer J2 observed per replication.


A


0)

M0
L-


40 35 30 25


20 15 10



5


a)


CL !)

0
CL
0
0 'a
0 a)





E
z


----Bare fallow
-Rye a
- - -A- - Vetch-wheat

a a
a




ab



a '*


a \ ' b
- b
b

,b-


0


0O U)


CC)

CO


a)
00


U)


0)

Ca
LIL


C0 0)


CO
LL


a


~heat b,










a/ b/


B






46












number of endospores


grams/microplot


10001


b


- ~ -


b


I ....f 2 - J


I--k-


a,


C


Control
RKN + Pp
1 Endospores/J2 in RKN


1987


1988


1989


1990


1991


%N


4


/


7_I


a


a


-RKN
-*Endospores/J2 in
RKN + Pp


1992


1993


Cropping season
Fig. 2-7. Log presentation of peanut yield (grams per microplot) from plots treated with Meloidogyne arenaria race 1 (RKN), and Al. arenaria plus Pasteuria penetrans (RKN + Pp). and untreated plots; and the number of endospores per second-stage juvenile (J2) in RKN and RKN + Pp plots. Data from 1987 to 1989 were from Oostendorp et al., 1991a. Means of 10 replicates, with 20 or fewer juveniles observed per replication to determine the number of endospores per J2. Means with the same letter are not significantly different according to Duncan's multiple-range test (P < 0.05).
Bar graphs - peanut yields in grams; line graphs - number endospores per juvenile.


a


100 +


10 +


0.1


I .... -__ U U -...


100


* 10





-1





0.1


.L-


W//


N a


4





47


Throughout the last 3 years of the experiment (1991 to 1993), the population

density development of Criconemella spp. was favored in plots that were fallowed through the winter (P < 0.05) (Appendix). An interaction between treatment and cover crop showed a higher number of ring nematodes in fallowed RKN plots than in fallowed RKN + Pp plots. In the fall of 1991 the highest number of ring nematodes in the soil coincided with the highest number of endospores of P. penetrans attached per juvenile of Meloidogyne arenaria.



Discussion



Populations of P. penetrans, as estimated by endospore attachment to

juveniles, increased over 6.5 years from relatively low levels of endospores added initially or introduced by contamination during the course of the experiment to levels after 3 years that were suppressive to the nematode population. This confirms the work by Oostendorp et al. (199 1a) and Chen et al. (1997c), whereby P. penetrans endospores increased to levels that suppressed root-knot nematodes when introduced at low numbers (approximately 1,000 spores/g of soil). Within four cropping seasons, nematode attachment rates in plots with endospores added increased to nearly 100%, with an average of about 20 endospores per juvenile, and peanut yields reached levels equivalent to those in the nematode-free control.

The population density of M. arenaria in RKN + Pp plots remained nearly

undetectable throughout the last 3 years of this 6.5-year experiment. It was difficult to attain an estimate of endospores per juvenile over the course of the experiment because the number of J2 in many RKN + Pp plots dropped below 20, which was the number used for estimating the endospore density. This was illustrated in the fall of 1991, when a decrease





48


in the number of endospores per juvenile and a decrease in the percentage of J2 with attached endospores were observed in rye and vetch plots. Individual observations revealed a further increase in the endospore population density under either cropping sequence. In 1992 and 1993, the attachment rates of juveniles with P. penetrans were around 100%, and the number of endospores per juvenile frequently exceeded 25. At this rate, root penetration has been shown to be prevented (Davies et al., 1988; Stirling, 1984: Stirling et al., 1990). Nematode suppression for three consecutive years did not affect the ability of P. penetrans endospores to attach to their nematode hosts. However, the experimental design did not allow for the determination of the number of J2 that may have inadvertently entered the microplots each season. Although it has been reported that the suppressiveness of root-knot nematode by P. penetrans is dependant on the density of endospores in the soil (Stirling, 1991), it appears in the present study that P. penetrans increased to a suppressive level after 3 years and remained sufficiently high to prevent further increases in the RKN densities. Pasteuria penetrans remained at a highly suppressive level for 3.5 years following the initial increase to suppressive levels in 3 years. The nematode population density required to maintain effective densities of P. penetrans to continue nematode suppression remains unknown.

In 1991 and 1992, the residual nematode population in RKN plots reduced peanut yields and affected the plant performance. However, in 1993, plant shoots and yields appeared to have been affected by leaf chlorosis and reduced fruiting, apparently as a result of the symptomatic manganese deficiency (Dickson, personal communication; Porter et al., 1984), rather than by nematode infection. There was only slight galling of roots, pods, and pegs by root-knot nematodes in the RKN plots in 1993, which indicates that nematode reproduction was becoming suppressed, and that the P. penetrans population density in the soil was continuing to increase to suppressive levels.





49


With no P. penetrans to interfere, the nematode population density peaked when

peanut was followed by a susceptible crop. Those conditions favored the contamination by and the build-up of endospores of P. penetrans. This observation agrees with those of other scientists (Chen et al., 1994; Madulu et al., 1994; Oostendorp et al., 199 1a), in which the continuous availability of a host plant favored the amplification of the bacterial parasite in its nematode host. However, since the nematode population density in the presence of P. penetrans did not differ among the cropping sequences, it is suggested that nematodes in vetch plots were already partially suppressed in years 2 and 3 of the experiment. Because of the density dependent nature of P. penetrans (Bird and Brisbane, 1988; Chen et al., 1996b; Dickson et al., 1994; Minton and Sayre, 1989; Stirling, 1984), the endospore population development is expected to be influenced by the population density of its nematode host (Sirling, 1991).

The winter cover crop had a consistent effect on the population density of Criconemella spp. Fallowing the peanut microplots increased the number of ring nematodes more than growing rye and wheat or vetch for winter cover. Criconelialla sp. was reported as a host for Pasteuria spp. (Sturhan, 1985; 1988). However, endospores of isolates of Pasteuria spp. from Criconemnella spp.were smaller and are believed to be a different species (Hewlett, personal communication); hence, they do not likely contribute to the suppression of M. arenaria in the peanut microplots.

For a period of 3 years (1991 to 1993), P. penetrans protected peanut from rootknot damage. The presence of a large population of M. arenaria in the RKN plots favored a continuous endospore population increase until the nematode population was detrimentally affected after a period of 4 years (1990 to 1993), which resulted from inadvertent contamination by P. penetrans endospores.





50


The build-up of the endospore density in the soil can be enhanced by supplying the nematode population with a good host plant throughout the year. Thus, although there is still no large-scale production of P. penetrans inoculum in sight, the use of this nematode antagonist in integrated nematode management programs might be feasible. There is a need for further experiments to determine whether the nematode suppression can be achieved in a shorter time span, and whether it can be maintained beyond a period of 3 years.













CHAPTER 3
USE OF MICROWAVE HEATING IN EVALUATION OF A MELOIDOGYNE ARENARIA-SUPPRESSIVE SOIL CONTAINING PASTEURIA PENETRANS AND ITS APPLICATION IN A SUPPRESSIVE-SOIL TEST


Introduction



Pathogen-suppressive soils may be defined as soils in which the pathogen does not establish or persist, establishes but causes little or no damage, or establishes and causes disease (conducive soil), but thereafter the disease is reduced even though the pathogen persists in the soil (Baker and Cook, 1982). Suppressiveness can develop as a result of the buildup of antagonists in response to a high pathogen population (Baker and Cook, 1982), especially in situations where susceptible crops are grown in succession.

The endospore-forming, obligate bacterial parasite of root-knot nematodes,

Pasteuria penetrans (Thorne) Sayre & Starr, is widely distributed in agricultural soils throughout the world (Chapter 1) and has been shown to be effective in controlling rootknot nematodes, Meloidogyne spp., in field or microplot studies (Brown and Smart, 1985; Chen et al., 1994;1996b; 1997c; Dickson et al., 1994; Minton and Sayre, 1989; Oostendorp et al., 1991a). In most cases, the suppression of Meloidogyne spp. by P. penetrans occurred after long-term monoculture of a susceptible host (Bird and Brisbane, 1988; Chen et al., 1994; Mankau, 1980a; Minton and Sayre, 1989; Weibelzahl-Fulton et al., 1996).

Many physical and chemical means have been developed to eliminate or reduce populations of soilborne plant pathogens for phytopathological studies (Mulder, 1979). Nematode suppressiveness of soil has been evaluated previously using soil applications


51





52


of fungicides, such as captafol (Crump, 1987), mancozeb and iprodione (Mertens and Stirling, 1993), benomyl or tachigaren (Qadri and Saleh, 1990), and 40% formaldehyde (Dickson et al., 1994; Kerry et al., 1982; Qadri and Saleh, 1990). Suppression of fungal populations by fungicides is not consistent, thus the interpretation of results can be problematic. Microwave heating of soil for selective periods of time can be useful to kill or reduce populations of soilborne fungal propagules, yet have little effect on populations of prokaryotic organisms (Chen et al., 1995; Ferriss, 1984). The effect of the microwave radiation treatment on microorganisms increases with increasing temperature. The temperature increase depends on treatment time, and on factors such as soil water, clay, and organic matter content (Baker and Fuller, 1969; Ferriss, 1984).

The objectives of this study were to determine if the microwave radiation treatment has the potential to selectively eliminate fungal antagonists from soils containing P. penetrans, and to evaluate the use of the microwave treatment in testing three microplot soils for suppressiveness to M. arenaria.



Materials and Methods



Microwave Treatment



Treatment time. On 12 January 1993, approximately 12 liters of soil, a loamy, siliceous, hypothermic Grossarenic Paleudults with 90% sand, 4% silt, 6% clay, 1.8% organic matter, 6.9% moisture, and a density of 2.78, were collected 0-20 cm deep with a bucket auger (10-cm diameter) from microplots infested with P. penetrans and Meloidogyne arenaria race 1 located at the University of Florida, Green Acres Agronomy Farm, Alachua County. Soil was selectively collected from microplots that were bare





53


fallowed during the winter. The soil was passed through a 5-mm-aperture sieve, and mixed thoroughly. Six subsamples of 1 kg each were placed in open polypropylene bags, leveled to a thickness of approximately 5 cm, and placed centrally in the 18-liter cavity of a microwave oven (1,500 Watts, 2,450 MHz) (Tappan Appliance, Mansfield, OH). The subsamples were either microwave treated for 3, 4, 5, 6, and 8 minutes, or left untreated. The bags remained open through the heat treatment, and were closed before removal from the oven. The soil was allowed to cool to room temperature.

Two hours later, a dilution plating technique was used to assay for selected fungi (Johnson and Curl, 1972). One gram of soil was dispersed in 200 ml of sterilized water and stirred constantly. One milliliter of the soil solution was plated on potato dextrose agar containing 50 mg of chlortetracycline hydrochloride, 100 mg of streptomycin sulfate, and 1 ml of Tergitol NP10 per 1 liter solution (supplements were added to agar at a temperature of < 50 "C) (Steiner and Watson, 1965). Five replicates were plated for each treatment. After 3 days of incubation at room temperature, the fungal colonies were counted.

A soil bioassay with Meloidogyne arenaria race 1 as the host nematode was used to determine the presence of P. penetrans (Oostendorp et al., 199 1a). Forty grams of dry soil from each treatment were placed in each of five, 10-cm-diameter petri dishes. Then, 300 second-stage juveniles (J2) were added in I ml of water to each dish. After 2 days of incubation at room temperature, the J2 were extracted by a centrifugal-flotation method (Jenkins, 1964), and the number of endospores per 20 randomly selected J2 per replicate was determined with the aid of an inverted microscope at X400 magnification.

Soil moisture content. On 8 March 1993, 1.5-kg lots of the dried soil collected on 12 January 1993 were placed into plastic bags, and the soil moisture content was adjusted to 1%, 3%, 5%, or 7% (weight of water per dry soil weight) by adding tap water. After 2 hours, the soil was mixed thoroughly, divided into three subsamples, placed into open





54


polypropylene bags, and was either heated in the microwave oven for 3 or 4 minutes per kilogram of soil or left untreated. The numbers of fungal colonies per gram of soil and endospores per juvenile were determined as described above.



Suppressive-Soil Test



Soil treatments. On 23 April 1993 and 7 May 1993, a total of 120 liters of soil was collected from the same microplots as stated above. For the repeat test, soil was collected on 23 April and 5 June. For each experiment, a subtotal of 40 liters of soil was collected from microplots with M. arenaria inoculum alone (RKN), M. arenaria plus P. penetrans inoculum (RKN + Pp), and a control treatment without nematodes or bacteria (control) (for more information on the soil sources see Chapter 2). The soils were passed through a 5mm-aperture sieve, mixed, and divided into four subsamples of 10 liters each. The subsamples were either autoclaved, microwaved, air dried, or left untreated.

For the autoclave treatment, sterilized 15-cm-diameter clay pots were filled with

soil (ca. 1 liter/pot), covered with aluminum foil, and placed in an autoclave. After the first heat treatment for 1.5 hours at 55 kPa (Ferriss, 1984), the pots were removed from the autoclave. The treatment was repeated after pots and soil had cooled to room temperature (Mulder, 1979).

For the microwave heating treatment, 1-kg lots of soil were placed in even layers in open polypropylene bags and heated in a microwave oven for 3 minutes (Ferriss, 1984) at full power, which caused the soil temperature to rise to about 75 *C.

For the air-drying treatment, 40 liters of soil were collected from the microplots on 7 May and 5 June 1993. The soil was placed in 3- to 5-cm-thick layers on plastic trays and





55


stored in the greenhouse for 2 or 6 weeks before testing. The untreated soil was stored at room temperature, and used within 24 hours after collection.

Laboratory experiment. Two hours after microwaving soil and the last autoclaving treatment, and 2 weeks after the beginning of the air-dried treatment, the number of fungal colonies per gram of soil and endospores per juvenile was determined simultaneously for all four soil treatments. Procedures were applied as described above.

Greenhouse experiments. Treatments were arranged in a 3 X 4 X 2 factorial design that included the following three soil sources: soil from microplots with M. arenaria inoculum alone (RKN), M. arenaria plus P. penetrans inoculum (RKN + Pp), and a control treatment without nematodes or bacteria (control); four soil treatments (autoclaved, microwaved, air-dried, and untreated); and inoculum levels of 0 and 2,000 J2 of M. arenaria race 1. The soil from each treatment was placed into sterilized 15-cmdiameter clay pots (ca. I liter/pot). Treatments were replicated four and five times in experiments 1 and 2, respectively. About 100 ml and 50 ml of water were added to the airdried soil and to all other soil treatments, respectively. Each pot was covered with aluminum foil and held for 3 days in a growth room at 28 to 32 *C and 14 hours of light, or in a greenhouse for experiments 1 and 2, respectively. Meloidogyne arenaria J2 were suspended in 10 ml of water and dispensed equally to each of five holes, 5-cm deep, in the soil of each pot. Five days later, one 7-week-old peanut seedling, Arachis hypogaea L. cv. Florunner, for experiment 1, or one 5-week-old peanut seedling of the same cultivar for experiment 2 was transplanted into each pot. A commercial Rhizobium sp. was added to each pot (McSorley et al., 1992). Every 10 to 14 days, insects and diseases were controlled using esfenvalerate, chloropyrifos, insecticidal soap, chlorothalonil, or liquid sulfur.





56


Experiments 1 and 2 were run for 53 and 55 days, respectively. Plant shoots were cut off at ground level and discarded. The root systems were washed free of soil, excess water was removed with paper towels, and roots were weighed and then stored in plastic bags at 4 *C. The following day, roots were stained with Phloxine B (Dickson and Strubel, 1965), and the numbers of root galls and egg masses were counted. Eggs were extracted with 1.05% sodium hypochlorite (Hussey, 1971) and their number was counted. Twenty globular mature females were picked randomly from each root system, crushed on a slide and observed for infection by P. penetrans with the aid of a compound microscope at X 1000 magnification.




Nematode Origin

The isolate of M. arenaria race 1 used originated from peanut grown on the

University of Florida, Green Acres Agronomy Research Farm, Alachua County. The species and race determinations were confirmed by examination of the perineal pattern, length of J2, and a differential host test (Hartman and Sasser, 1985). The nematode population was cultured in a greenhouse on tomato (Lycopersicon esculentum Mill. cv. Rutgers) from a single egg mass. Eggs of M. arenaria were extracted from infected tomato roots (Hussey and Barker, 1973), hatched by the Baermann method (RodriguezKabana and Pope, 1981) and used as 1- to 4-day-old J2.




Statistical Analysis

The number of endospores per juvenile was transformed with logI0(x + 1) before analysis. The percentages of female infection were transformed to arcsin (Ix) before analysis. All data were subjected to factorial analysis of variance (ANOVA). Means were





57


separated and compared by Duncan's multiple-range test. Regressions were performed to determine the relationship between the microwave treatment time and the survival of P. penetrans and fungal populations.




Results



Microwave Treatment



Treatment time. Population densities of P. penetrans endospores and fungi

decreased with the increasing treatment time (Y= 1.95x2 - 30.3 Ix + 115.67) (Fig. 3-1). Following treatment for 3 minutes, the attachment of endospores was reduced by almost 50% (average of 59 endospores/juvenile). When the treatment time was extended to 4 minutes/kg of soil and 5 minutes/kg of soil, the attachment was reduced to 20 endospores per juvenile and nearly eliminated, respectively. The number of fungal colonies was greatly reduced by all treatment times, as compared to the untreated soil (Y = 1.04x2 12.21x + 33.69). Fusarium sp., Paecilomyces lilacinus, Penicillium sp., Pythium sp., Trichoderma sp., and a variety of unidentified fungi were isolated.

Soil moisture content. The attachment of P. penetrans endospores to J2 increased slightly with increasing soil moisture content (Y = -1875x + 274x + 10.28) (Fig. 3-2). Numbers of endospores attached per juvenile decreased in soils of all tested moisture levels treated for 3 and 4 minutes in the microwave (P ; 0.05 ) (Fig. 3-3). A further decrease in attachment was observed with increased treatment time in soils containing 3, 5, and 7% moisture. These observations validate the numbers observed in the treatment-time study reported above (Fig. 3-2). However, the endospore counts of this study were much lower






58


120


1 2 3 4 5 6 7






B y = 1.0376x2 - 12.209x + 33.688
F'= 0.958












Tm
-0



S12 3 4 5 6 7

Treatment time in minutes


Fig. 3-1. Effect of microwave radiation treatment of 1 kg of soil containing Pasteuria penetrans on A) the attachment of P. penetrans endospores to Meloidogyne arenaria race 1 and B) the survival of soilborn fungi as determined by the number of colony-forming units. Values are means of five replications; 20 randomly selected secondstage juveniles were observed.


C


A
y = 1.9448x2 - 30.307x + 115.67 F=0.9602









00


100


80


- 60 o 40
40

20-


0


-20 35 30 25

20


15 10 5
U
0 -5






59


25 , 20



15 10 5 0


10%


2%


3%


4%


5%


6%


7%


Soil moisture content (weight water/dry soil weight)

Fig. 3-2. Relationship between the soil moisture content and the number of endospores of Pasteuria penetrans attached per second-stage juvenile (J2) of Meloidogyne arenaria race 1 after 48 hours exposure at room temperature. Values are means of two replications; 20 randomly selected J2 were observed.


25,


20 15
L.


10 5


0


o untreated o 3 minutes
* 4 minutes


a






b



1%


a


a


3% 5%
Soil moisture content (weight water/dry soil weight


7 a











b
_E c

7%
)


Fig. 3-3. Effect of soil moisture content and microwave treatment time on the attachment of Pasteuria penetrans endospores to Meloidogyne arenaria race 1. Values are means of two replications; 20 randomly selected second-stage juveniles were observed. Means within each moisture treatment followed by the same letter do not differ at P < 0.05 according to Duncan's multiple-range test.


0













y = -1875x2 + 274x + 10.278
R2 = 0.5466





60


than those of the treatment-time study.

Fungal populations were detected at all moisture levels of the untreated soil only. Population densities increased with increasing moisture content of the soil (Y= 21x + 1; R2= 0.99) (data not shown) and were 1.2 x 104, 1.7 x 104, 2 x 104, and 2.5 x 104 colony forming units (cfu)/g of soil containing 1%, 3%, 5%, and 7% moisture, respectively.



Suppressive-Soil Test



Laboratory experiment. Survival of fungi and P. penetrans in the microplot soil varied with soil sources and soil treatments (P < 0.05) (Table 3-1). Regardless of soil source, populations of fungi were lower in autoclaved or microwaved soil than in air dried and untreated soil (P < 0.05). The fungal populations in soil from RKN plots generally exceeded those in soil from RKN + Pp plots and control soils in untreated or in air dried soils. No P. penetrans endospores were detected in control soil or in any autoclaved soils. The number of endospores attached per juvenile did not differ among the microwaved, airdried, and untreated soils.

Greenhouse experiments. The effects of the soil sources, treatments, and

inoculation levels were similar in both experiments for all data, except the number of eggs as affected by the soil treatments and the female infection rate (Table 3-2). In experiment 2, the number of eggs did not vary with soil treatments, and the single interaction affecting the female infection rate was the soil source X soil treatment interaction.

Reproduction of M. arenaria on peanut, as expressed by the number of root galls, egg masses, and eggs, was consistently higher in autoclaved soil than in microwaved, airdried, and untreated soil infested with M. arenaria alone (Table 3-3) or M. arenaria plus P. penetrans (Table 3-4) after inoculation with 2,000 J2 of M. arenaria. In soil infested





61


TABLE 3-1. Colony forming units (cfu) of soil fungi and attachment of Pasteuria penetrans endospores to second-stage juveniles (J2) of Meloidogyne arenaria in untreated soil and soil autoclaved twice for 1.5 hours at 55 kPa, microwaved for 3 minutes/kg of soil, or air dried for 2 weeks in the greenhouse. Soils contained 6.9% moisture and were collected from peanut microplots infected with M. arenaria inoculum alone (RKN), M. arenaria plus P. penetrans inoculum (RKN + Pp), and a control treatment without added nematodes or bacteria (control).

Cfu of fungi Mean number of P. per gram of penetrans endospores Soil source Soil treatment soil per juvenile

Control Autoclaved 80 b A 0 a

Microwaved 80 b A 0 a

Air-dried 1.1 x 104 a B 0 a

Untreated 1.6 x 104 a B 0 a

RKN Autoclaved 0 b A 0 b

Microwaved 40 b A 1.1 ab

Air-dried 3.4 x 104 a A 8.7 a

Untreated 4.5 x i04 a A 9.5 a

RKN + Pp Autoclaved 80 b A 0 b

Microwaved 0 b A 9.5 a

Air-dried 1.5 x 104 a B 12.4 a

Untreated 1.5 x 104 a B 9.8 a
Values are means of five replicates for the cfu counts and two replicates for the endospore counts; 20 randomly selected J2 were observed per replicate.
The data were transformed with logI0(x + 1) before being subjected to ANOVA. Presented data are untransformed.
Means within columns for each soil source or for each treatment followed by the
same lower case letter or upper case letter,respectively, do not differ at P < 0.05 according to Duncan's multiple-range test.





62


- TABLE 3-2. ANOVA table for the effect of soil source (microplots infected with Meloidogyne arenaria race 1 alone, with M. arenaria race 1 plus Pasteuria penetrans, and without either organism), soil treatments (autoclaved, microwaved, air-dried, and untreated), and M. arenaria race 1 inoculum levels (0 or 2,000 second-stage juveniles) on nematode reproduction and fresh root weights of peanut cv. Florunner.


Number of Number of Number of Females Fresh root
Source of variation galls egg masses eggs infected weight

Experiment 1
Soil source (S) NS
Soil treatment (T) ** NS
Inoculation level (I) * NS
S x T ** NS
S x I ** NS
IxT ** NS
I x T x S *** *** * NS
Experiment 2
Soil source (S) *** *** ** NS
Soil treatment (T) NS * *
Inoculation level (I) * NS
S x T ** *** * NS
S x I ** NS NS
IxT NS NS
IxTxS NS NS

*, * * represents P < 0.05, P < 0.01, and P < 0.001, respectively. NS = nonsignificant at P < 0.05.







TABLE 3-3. Effect of autoclaving, microwaving, and air-drying of soil infested with Meloidogyne arenaria race I alone on the nematode reproduction, percentage of females infected by Pasteuria penetrans, and fresh root weights of peanut cv. Florunner following inoculation with 0 or 2,000 second-stage juveniles (J2). a


Nematode reproduction
Number of root galls Number of egg masses Number of eggs per % females infected Fresh root weight
per root system per root system root system (mean of 20 observations) (g)
Soil Inoculation level Inoculation level Inoculation level Inoculation level Inoculation level
treatment +h - + - + - + - +
Experiment I

Autoclaved 605 a 0 b 522 a 0 b 98,750 a 0 b 0 b d 4.8 a 4.5 a

Microwaved 18 b 0 b 5 b 0 b 1,150 b 0 b I0'a - 3.9 b 4.4 a

Air dried 0 c 0 b 0 c 0 b 40 c 0 - - 3.3 b 4.7 a

Untreated 36 b 10 a 4 b 6 a 560 b 290 a 17'a 77a 3.4 b 4.3 a

Experiment 2

Autoclaved 491 a 0 b 380 a 0 b 116,960 a 0 b 0 b- 4.1 a 2.5 b

Microwaved 0 b 0 b 0 b 0 b 40 b 0 b- - 4.7 a 4.2 a

Air dried 0.8 b 0 b 0 b 0 b 20 b 0 b 30'a - 3.5 a 3.9 b

Untreated I b 5 a 0.8 b 1 a 220 b 200 a 17'a 69'a 3.5 a 3.2 b

Values are means of four replicates in experiment 1 and five replicates in experiment 2; means within columns in each experiment followed by the same letter do not differ at P < 0.05 according to Duncan's multiple-range test.
a The soil in the 6.5-year microplot experiment that was initially infested with M. arenaria alone become infested with P. penetrans endospores in the 4" year.
b + Inoculated with 2,000 J2 of M. arenaria race 1; - No nematodes added.

c Less than 20 observations in at least one replicate.
d - = No observation.









TABLE 3-4. Effect of autoclaving, microwaving, and air-drying of soil infested with Meloidogyne arenaria race I and Pasteuria penetrans on the nematode reproduction, percentage of females infected by P. penetrans, and fresh root weights of peanut cv. Florunner following inoculation with 0 or 2,000 second-stage juveniles (J2) of M. arenaria race 1.


Nematode reproduction
Number of root galls Number of egg masses Number of eggs per % females infected Fresh root weight
per root system per root system root system (mean of 20 observations) (g)
Soil Inoculation level Inoculation level Inoculation level Inoculation level Inoculation level
treatment +A- + - + - + - +
Experiment I

Autoclaved 702 a 0 a 608 a 0 a 137,370 a 0 a 0 b -d 4.6 a 4.5 a

Microwaved 10b 0 a 0 b 0 a 130 b 0 a 100 a - 4.5 a 4.0 a

Air dried 8 b 0 a 0 b 0 a 60 b 0 a 100 a - 3.3 b 4.2 a

Untreated 3 b 0 a 0 b 0 a 10b 0 a 100 a - 3.5 b 3.2 b

Experiment 2

Autoclaved 300 a 0 a 256 a 0 a 72,780 a 0 a 0 b - 3.5 a 4.9 a

Microwaved 6 b 0 a 3 b 0 a 260 b 0 a 77'a - 3.3 a 3.3 b

Air dried 0 b 0 a 0 b 0 a O c 0 a - - 2.0 b 2.7 b

Untreated 0 b 0 a 0 b 0 a 120 b 0 a - - 3.4 a 2.7 b

Values are means of four replicates in experiment I and five replicates in experiment 2; means within columns in each experiment followed by the same letter do not differ at P < 0.05 according to Duncan's multiple-range test.
" + Inoculated with 2,000 J2 of M. arenaria race 1; - No nematodes added.
b Less than 20 observations in at least one replicate.

d - = No observation.





65


with M. arenaria alone of experiment 1, fewer nematodes reproduced in air dried than in microwaved, and untreated soil (P < 0.05) (Table 3-3). This observation was not validated by experiment 2. Nematode reproduction in all three autoclaved soils did not differ from nematode reproduction in microwaved and air-dried soil collected from microplots without nematodes P. penetrans, except in the uninoculated microwave treatment of experiment 2 (P < 0.001, statistics not shown) (Tables 3-3 to 3-5). When no nematode inoculum was added, a very low level of nematode reproduction was observed only in untreated soil from plots infested with M. arenaria alone (P < 0.001) (Table 3-3), and in microwaved soil from ininoculated plots of experiment 2 (Table 3-5).

No females were infected with P. penetrans in all autoclaved soils, and in soil treatments of nematode and Pasteuria-free soil (Tables 3-3 to 3-5). When soil was infested with nematodes, 100% of the females were infected in microwaved, air-dried, and untreated RKN + Pp soils of experiment 1, and 77% were infected in microwaved soil in experiment 2 (Table 3-4). A lower incidence of female infection occurred in the soil from microplots that had become infested with P. penetrans. Ten to thirty percent of females in microwaved, air-dried, or untreated RKN soil were infected with P. penetrans when nematodes were added (Table 3-3). When no nematode inoculum was added, infection rates of females in untreated RKN soil were 69 and 77%. However, females in uninoculated microwaved soil of nematode- and P. penetrans-free microplots were not infected by P. penetrans.

The effects of soil treatment on the fresh root weight were inconsistent (Tables 3-3 to 3-5). Peanut plants cultured in autoclaved or microwaved soil were not different or tended to developed larger root systems than those grown in air dried and untreated soils (P < 0.05). Inoculation level had no effect on the fresh root weight (P < 0.05) (Table 3-2).









TABLE 3-5. Effect of autoclaving, microwaving, and air drying of soil maintained free of nematodes and Pasteuria penetrans on the nematode reproduction, percentage of females infected with P. penetrans and fresh root weights of peanut cv. Florunner following-inoculation with 0 or 2,000 second-stage juveniles (J2) of M. arenaria race 1.


Nematode reproduction
Number of root galls Number of egg masses Number of eggs per % females infected Fresh root weight
per root system per root system root system (mean of 20 observations) (g)
Soil Inoculation level Inoculation level Inoculation level Inoculation level Inoculation level
treatment + - + - + - + - +
Experiment I

Autoclaved 588 a 0 a 513 a 0 a 118,500 a 0 a 0 a d 4.9 a 4.9 a

Microwaved 402 a 0 a 406 a 0 a 130,900 a 0 a 0 a - 4.7 a 4 b

Air dried 376 a 0 a 256 ab 0 a 72,000 a 0 a 0 a - 4.7 a 3.8 b

Untreated 182 b 0 a III b 0 a 31,200 b 0 a 0 a - 4.1 a 3.9 b

Experiment 2

Autoclaved 363 a 0 b 303 a 0 b 88,500 a 0 b 0 a - 3 b 4.3 a

Microwaved 375 a 3 a 306 a 2 a 86,000 a 380 a 0 a Oba 4.3 a 3 b

Air dried 148 b 0 b 118 b 0 b 34,600 b 0 b 0 a - 2.5 b 4.6 a

Untreated 432 a 0 b 382 a 0 b 88,000 a 0 b 0 a - 3.3 b 2.8 b

Values are means of four replicates in experiment I and five replicates in experiment 2; means within columns in each experiment followed by the same letter do not differ at P < 0.05 according to Duncan's multiple-range test.

+ Inoculated with 2,000 J2 of M. arenaria race 1; - No nematodes added.
h Less than 20 observations in at least one replicate.
- d - = No observation.


ON ON





67


When no nematodes were added, no nematode reproduction was noted in all but

the untreated soils of RKN and RKN + Pp soil and in microwaved control soil (Tables 3-3 to 3-5, statistics not shown). Nematode reproduction generally did not differ between inoculated and uninoculated treatments in microwaved, air dried, and untreated soils of RKN and RKN + Pp plots (except in the microwaved and untreated soil of RKN plots), but was higher in all inoculated, control soils and following the autoclave treatment (P <

0.05).



Discussion



An increase in treatment time or in soil moisture content resulted in an increasing effect of microwaving soil on soil microorganisms. The response of soil fungi to an increase in soil moisture content does not agree with Ferriss's (1984) observations on microorganisms. He reported that soil temperature and the effect on reduction of populations of microorganisms correlated positively with MW treatment time and negatively with soil moisture content. Moisture levels approaching field capacity were required to kill spores of root-pathogenic fungi by microwave heat treatment (Baker and Fuller, 1969). The percentages of soil moisture of loamy soils used in Ferriss' (1984) studies exceeded by far the soil moisture contents of soils used in this study. Loamy soils would probably require an extended period of microwave treatment to approach an effect similar to that attained on sandy soils. The different results also may be explained by the fact that sandier soils cool faster than loamy soils after microwaving (Ferriss, 1984); hence, fungal propagules may have been affected by the duration of the elevated temperatures rather than by the temperature peak.





68


Endospores of P. penetrans are reportedly resistant to high temperatures (Stirling et al., 1986; Williams et al., 1989); however, when endospores were exposed to 80 *C for 30 minutes their development was impeded (Dutky and Sayre, 1978). More recent studies showed that attachment was close to minimum at 60 'C or higher, but it was not completely prevented at 100 'C for 5 hours per day over 10 days (Freitas, 1997). In this study, a microwave treatment time of 3 minutes for a 1-kg sample of soil containing nearly 7% moisture heated the soil to 70 to 75 *C for less than 1 minute. Following the microwave treatment, the number of endospores attached per juvenile was reduced to a variable degree, but was not lower than four endospores per juvenile. This effect was tolerable because the germination of a single spore is enough to create infection in the female Meloidogyne (Stirling, 1984); and hence, sufficient to be used to confirm nematode suppression by P. penetrans. One hundred percent infection of females occurred after the microwave soil treatment, indicating that P. penetrans was able to survive the microwave treatment and suppress the nematode population in the pot experiments.

Although methods to quantify facultative and obligate nematophagous fungi in soil have been developed, they do not always give satisfactory results (Dackman et al., 1987). Propagules of saprophytic and nematophagous fungi are of similar structure (Barron, 1992); therefore, they are likely to withstand similar amounts of microwave radiation. If the results of the dilution plating technique were extrapolated to the survival of nematophagous fungi, microwaving soil for 3 minutes per kg of soil had the potential to selectively eliminate this group of microorganisms. This allows for the separation of the antagonistic effects of fungi and the bacterial parasite. Microwaving soil 3 minutes for each kilogram of soil was chosen to be the most appropriate microwave treatment to be used in the suppressive soil test. The treatment does not leave a nematicidal residual effect,





69


as observed with soil applications of 40% formaldehyde (Dickson et al., 1994; Kerry et al., 1982) and benomyl (Hoestra, 1976) treatment.

The numerical difference in attachment of endospores to J2 between the two

microwave studies on water content and treatment time may be due to the fact that the dried soil containing P. penetrans was stored for 8 weeks between the studies, thus affecting the ability of endospores to attach to nematodes. A minimum of 3 days moisture incubation (Brown and Nordmeyer, 1985) before the application of soil treatments and nematode inoculum may have enabled more endospores to attach.

Attachment by P. penetrans was detected on J2 exposed to soil from RKN and

RKN + Pp microplots. Nematode reproduction was consistently inhibited in the presence of viable P. penetrans endospores in these soils. Endospores in both soils existed in densities at which plant infectivity by the J2 was reduced, which protected the root systems from being severely galled. Similar results were found when J2 were encumbered with 15 or more endospores of P. penetrans and used to inoculate tomato (Brown and Smart, 1985; Davies et al., 1988; 1990; Sell and Hansen, 1987; Stirling, 1984). The infection of M. arenaria by P. penetrans resulted in low fecundity. This supports earlier observations that infection by P. penetrans causes sterility or reduced fecundity in females (Mankau, 1980a; Minton and Sayre, 1989); however, the number of isolated females was high enough to statistically confirm 100% infection by P. penetrans only from soils infested with M. arenaria and P. penetrans used in experiment 1. Additional tests with soils containing a more diluted P. penetrans population density may be needed to confirm the nematode suppression by P. penetrans. Temperature has been reported to affect nematode mobility (Bird and Wallace, 1965), attachment of endospores to J2 (Freitas et al., 1997; Hatz and Dickson, 1992; Stirling, 1981), and the development of P. penetrans in vivo (Hatz and Dickson, 1992; Nakasono et al., 1993; Serracin et al., 1997). The constant





70


temperatures of the growth room used for experiment 1 probably favored the development of P. penetrans.

Although in this study the effect of fungi and P. penetrans could not be evaluated separately, an additive nematode suppression would be expected to cause differences in nematode reproduction between the treatments of microwaving or air drying of RKN and RKN + Pp soils. The lowest numbers of root galls, egg masses, and eggs were observed following air-drying of RKN soil in experiment 1 and of RKN + Pp soil in experiment 2. However, since these effects could not be observed when replicated, fungi appeared to play only a minor role in nematode suppression. These results supported the hypothesis that P. penetrans is the main contributor to the suppressiveness in soils from both the RKN and RKN + Pp microplots.

In the suppressive soil test, the microwave treatment was helpful because it reduced fungi that might also be a factor in suppressive soils. However, the endospore density in RKN + Pp soil was so high that it greatly suppressed the plant infectivity by endosporeencumbered J2, and the female infection rate could not be determined. It cannot be ruled out that the nematode suppression was due to other factors than parasitism by P. penetrans; however, compared to the control soil, there were only minor differences between the test results of the RKN and the RKN + Pp soil comprising nematode suppression observed in Chapter 2. Hence, it can be concluded that the suppressiveness of the soils is of similar nature.

At the time the soil for this test was collected from the microplots, nematodes of RKN + Pp plots had been suppressed for 3 years. This study demonstrated that endospores of P. penetrans survived 3 years in the absence of a host and successfully suppressed a reinoculated nematode population. Although it is not known exactly, dormant endospores are probably able to survive for many more years. Pasteuria spp.





71


differ from successful natural enemies of insects, which usually require a constant food supply to maintain their population density (Bennett, 1974). The absence of a host nematode appears to have little impact on the survival of P. penetrans, which may be a leading attribute for the organism in suppression of nematodes under various conditions.














CHAPTER 4
POPULATION DEVELOPMENT OF MELOIDOGYNE SPP. AND PASTEURIA
.PENETRANS AS AFFECTED BY CULTURAL PRACTICES IN TOBACCO Introduction



The endospore-forming bacterial parasite, Pasteuria penetrans (Thorne) Sayre & Starr, is widely distributed in agricultural soils throughout the world (Sayre and Starr, 1988; Stirling, 1991) and contributes to natural and induced nematode control, especially of Meloidogyne spp. (Bird and Brisbane, 1988; Brown et al., 1985; Channer and Gowen, 1988; Chen et al. 1994; 1996b; 1997c; Dickson et al., 1994; Kerry, 1990; Mankau, 1980a; 1980b; Minton and Sayre, 1989; Oostendorp et al., 199 1a; Weibelzahl-Fulton et al., 1996). In most observations, the suppression of Meloidogyne spp. by P. penetrans occurred after long-term monoculture of susceptible hosts in association with Meloidogyne spp. (Bird and Brisbane, 1988; Chen et al., 1994; Mankau, 1980a; Minton and Sayre, 1989; Chapter 2). Pasteuria penetrans has great potential for integration with other cultural or nematode management practices (Brown and Nordmeyer, 1985; Freitas, 1997; Maheswari et al., 1988; Oostendorp et al., 1991a; Stapleton and Heald, 1991; Tzortzakakis and Gowen, 1994a; Walker and Wachtel, 1989). Pathogenicity and virulence of P. penetrans can vary among different bacterial isolates (Channer and Gowen, 1992; Davies et al., 1994; Oostendorp et al., 1990), and knowledge about their biology is required to effectively include the organism into nematode management strategies (Stirling, 1991).

A mixed population of root-knot nematodes, Meloidogyne incognita (Kofoid &

White) Chitwood, M. javanica (Treub) Chitwood, and M. arenaria (Neal) Chitwood, was


72





73


observed to damage tobacco grown in a field at the University of Florida Green Acres Agronomy Farm, Alachua County. Pasteuria penetrans and several species of nematophagous fungi were isolated from the site in 1991 (Chen et al., 1994). Pasteuria penetrans was identified to be suppressive to the mixed populations of root-knot nematodes, and significant changes in the density of Meloidogyne spp. were observed with inorganic nitrogen rates, cover crops, and tobacco cultivars (Chen et al., 1994).

The objectives of this study were to test the pathogenicity of the naturally occurring P. penetrans population to M. incognita and M. javanica in the laboratory, and to determine the effects of nitrogen fertilizer, autumn cover crop, and tobacco cultivars on the population density development of Meloidogyne spp. and P. penetrans in the tobacco field.



Materials and Methods



In the summer 1992, a random sample of approximately 30 roots was collected from the tobacco site. The roots were washed free of soil, cut into 2- to 3-cm pieces and mixed thoroughly. Forty females and accompanying egg masses of Meloidogyne spp. were dissected from randomly selected root pieces, and the females were placed individually on an eye glass in a drop of deionized water. The egg masses were treated with 0.5% NaOCI for 1 minute, washed with sterile water three times, placed individually into 2.5-ml microfuge tubes containing 1 ml of deionized water, and incubated at room temperature. The females were cut open at the neck region to release the body contents, which were then examined for mature endospores of P. penetrans. The nematode species was identified by optical examination of 20 perineal patterns.





74


Nematode Populations



One week after isolation from roots, 5 ml of water containing approximately 250 second-stage juveniles (J2) hatched from egg masses obtained from three Pasteuria-free females of either M. incognita or M. javanica were placed into five 2-cm-deep holes around the stems of two 4-week-old tomato (Lycopersicon esculentum L. cv. Rutgers) plants, and maintained in the greenhouse. After 60 days, the root systems were harvested and washed free of soil. Eggs of M. incognita and M. javanica were extracted from the infected tomato roots (Hussey and Barker, 1973), hatched by the Baermann method (Rodrfguez-Kibana and Pope, 1981), and used for experiments as I- to 4-day-old J2, or inoculated to ten 6-week-old tomato plants, and maintained in the greenhouse.



Pasteuria penetrans Isolates



One week after their isolation from roots, approximately 250 J2 of three Pasteuriafree females of both M. incognita and M. javanica were exposed to mature endospores of P. penetrans harvested from one female of the same species. After 24 hours incubation in water at room temperature (26 0C), the average number of endospores attached to 20 randomly selected J2 was determined with an inverted microscope at X400 magnification. The endospore encumbered J2 were suspended in 5 ml of water and then placed into five 2-cm-deep holes around the stems of two 4-week-old tomato (Lycopersicon esculentum L. cv. Rutgers) plants, and maintained in the greenhouse. After 60 days, the root systems were harvested and washed free of soil.

Root systems were enzymatically digested in Pectinol (Genencor, South San

Francisco, CA) at room temperature for 48 hours. Mature globose females were hand-




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picked from the softened roots, and air dried on an eye glass. After 3 days, the nematode bodies were rehydrated for 8 hours and macerated in a glass tissue-grinder. The number of endospores was counted with the aid of a hemacytometer, and their concentration was adjusted to 1 x 10 endospores per ml water.



Laboratory Experiment



Attachment studies were conducted using the centrifuge technique (Hewlett and Dickson, 1993). A 0.1-ml sample of the endospore-water suspension and 0.1 ml of a suspension of 2,000 J2 of M. incognita or M. javanica per ml of water were placed in a

0.25-ml, previously silanized microfuge tube and centrifuged at 9,500g for 2 minutes using a microfuge. Each attachment study was replicated five times. The content of the microtube was stirred and placed on a glass slide with a pipette. The number of endospores per juvenile was determined for 20 randomly selected nematodes per replicate. Thereafter, the endospore-encumbered nematodes of both species were suspended in 5 ml of water and inoculated to tomato plants as described above.



Field Experiment



The site had been planted to tobacco in the same plots for seven consecutive years. The 3 X 2 X 2 factorial treatment design included: three autumn cover crops (hairy indigo, Indigofera hirsuta L.; forage sorghum, Sorghum spp.; and weed fallow); two inorganic nitrogen fertilizers (89 and 158 kg of ammonium nitrate/ha); and two tobacco cultivars (Coker 371-Gold, which is susceptible to M. incognita, M. javanica, and M. arenaria; and Northrup King K-326, which is resistant to M. incognita but susceptible to M. javanica and





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M. arenaria). Hairy indigo and forage sorghum were planted in August and plowed under in November of each year. Rye was planted over the entire field in early December to serve as a winter cover crop, and plowed under in March. Tobacco seedlings were transplanted on 27 March and on 2 April of 1992 and 1993, respectively. The final harvest was .taken on 4 August and 11 August of 1992 and 1993, respectively. Preplant fertilization included a broadcast application of 1,800 kg of 6-6-18 (N-P-K) and 340 kg of 15-0-14 (N-P-K) of sodium-potassium nitrate per hectare. At the final cultivation, in late April, granular ammonium nitrate was applied adjacent to the plant stems. The plots were arranged in complete randomized blocks, and replicated four times. Each block consisted of one row with a row spacing of 1.2 m and a length of 12 m.

A 12-core soil sample was taken with a cone-shaped auger (2.5-cm diameter) from the root rhizosphere (20 cm deep) of each plot on 2 April, 9 June, and 28 August of 1992, and on 2 April, 24 June, and 14 August of 1993. The soil was mixed, sampled for soil moisture determination (samples of 2 April 1992 and 1993), and processed using centrifugal-flotation method (Jenkins, 1964). The number of J2/100 cm3 of soil, the rate of attachment of P. penetrans, and the number of endospores attached per 20 randomly selected J2 were determined using an inverted microscope at X400 magnification. Root samples were collected after the final tobacco harvest, 20 females were dissected randomly from each sample, crushed on a glass slide and checked for infection by P. penetrans with a compound microscope at X1000 magnification. Effects of the autumn cover crop were determined for the 1992 season only.





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Results



Laboratory Experiment



. Meloidogyne incognita and M. javanica were identified from tobacco root samples, which was thereafter confirmed by Chen et al. (1994). After 24 hours of incubation of the original P. penetrans isolates with J2 of either species at room temperature, 87% of J2 of M. incognita and 82% of J2 of M. javanica were attached, with an average number of 3.8 and 6.7 endospores per juvenile, respectively (data not shown). With the exception of one attachment test conducted with the P. penetrans-isolate from M. javanica, the number of endospores per juvenile of M. incognita was higher than or not different than the number of endospores attached per juvenile of M. javanica (P < 0.05) (Tables 4-1 and 4-2). Attachment of the M. javanica-isolate was higher on M. incognita than on M. javanica after the bacterial parasite was grown on M. incognita for two consecutive generations (P <

0.05) (Table 4-2). However, endospore attachment of the M. incognita-isolate after three generations on M. javanica was either maintained or reduced (Table 4-1). The percentages of J2 attached with endospores were between 72 and 100% (Tables 4-1 and 4-2). In the first and second generation attachment tests, the number of endospores per J2 and the percentage of encumbered J2 followed the same trends. In the third generation attachment tests, the percentages of J2 with attached endospores generally were not different.



Field Experiment



The soil moisture content ranged from 6.1-11.8% soil moisture in 1992, and 6.411.1% in 1993. The number of root-knot nematode juveniles in the soil and the galling









TABLE 4-1. Attachment pattern on second-stage juveniles (J2) of Meloidogyne javanica and M. incognita over three generations of Pasteuria penetrans isolate P-i 10 from M. incognita infecting tobacco.

1st generation 2nd generation 3rd generation
Number of Number of Number of
Test endospores/ % Test endospores/ % Test endospores/ %
nematode juvenile attachment nematode juvenile attachment nematode juvenile attachment
M. incognita" 15.3 a 100 a M. incognitab 9.7 a 94 a M. incognita 4.8 100

M. javanica 3.9 100
M. javanicab 2.0 b 66 b M. incognita 21 a 86
M. javanica 7 b 75
M. javanica" 6.2 b 78 b M. incognita' 6.9 72 M. incognita 8.0 81
M. javanica 3.5 90
M. javanicab 3.2 77 M. incognita 1.8 89
M. javanica 1.4 93
Means followed by no letter within a column of two observations were not different according to Student's t-test. Values are means of five replicates; 20 randomly selected J2 were observed per replicate.
a endospores produced in this nematode population were used in the 2nd generation attachment test.
b endospores produced in this nematode population were used in the 3rd generation attachment test.


-1










TABLE 4-2. Attachment pattern on second-stage juveniles (J2) of Meloidogyne javanica and M. incognita over three generations of Pasteuria penetrans isolate P-120 from M. javanica infecting tobacco.

Ist generation 2nd generation 3rd generation
Number of Number of Number of
Test endospores/ % Test endospores/ % Test endospores/ %
nematode juvenile attachment nematode juvenile attachment nematode juvenile attachment
M. incognita' 4.8 93 M. incognita 17.4 a 100 a M. incognita 20.8 a 100

M. javanica 14.2 b 100
M. javanica' 2.5 b 88 b M. incognita 2.4 91
M. javanica 2.5 84

M. javanicaa 8.3 97 M. incognita" 1.4 b 73 b M. incognita 1.4 74
M.javanica 1.5 72
M. javanicab 10.2 a 100 a M. incognita 12.6 100
M. javanica 8.0 100
Means followed by no letter within a column of two observations were not different according to Student's t-test. Values are means of five replicates; 20 randomly selected J2 were observed per replicate.
a endospores produced in this nematode population were used in the 2nd generation attachment test.
b endospores produced in this nematode population were used in the 3rd generation attachment test.





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indices were generally higher in 1992 than in 1993 (statistics not shown) (Table 4-3). At planting and after harvest in 1992, the root-knot nematode density was higher in plots with the history of the higher rate of inorganic nitrogen than in those that received the low nitrogen rate (P < 0.05). This effect was not observed in 1993. Throughout 1992 and 1993, the population density of J2 in the soil was increased by both tobacco cultivar histories, but no difference was observed between cultivars in 1992 and most of 1993. However, a higher nematode population density under the resistant tobacco cultivar than under the susceptible cultivar was revealed on the last sampling in 1993. The number of J2 was higher with weeds than hairy indigo or sorghum at harvest in 1992. At mid-season of 1992, but not in 1993, an interaction between cultivar and autumn cover crop was found to affect the number of J2 in the soil (Table 4-3). The nematode population density was lowest in NK-326 plots that followed the forage sorghum cover crop (data not shown). This observation was not validated by repeat in 1993.

In 1992, interactions between the inorganic nitrogen fertilizer levels and the tobacco cultivar histories affected the number of J2 in the soil at planting and after tobacco harvest (P < 0.05) (Tables 4-3 and 4-4). At planting and after harvest, the higher rate of ammonium nitrate fertilizer resulted in a greater population density of J2 in the root rhizosphere of NK-326 than in Coker 371-Gold. As opposed to NK 326, the number of nematodes in the root rhizosphere of Coker 371-Gold was not increased by the higher level of fertilizer (Table 4-4).

Root galling of Coker 371-Gold was less than that of NK-326 (P < 0.001 for 1992, P < 0.05 for 1993). In 1992, root galling on tobacco was less in plots following hairy indigo and forage sorghum than in plots following weeds (P < 0.01), but no observations were made in 1993 to validate the effects of the cover crops. The galling indices were not affected by the nitrogen treatments; however, an interaction between the inorganic nitrogen






TABLE 4-3. Population density development of Meloidogyne spp. in 1992 and 1993 as determined by the number of second-stage juveniles (J2) in the soil, and the galling indices of roots of two tobacco cultivars treated with two inorganic nitrogen rates, and three autumn cover crop treatments.

J2 per 100 cm3 of soil
Treatment 1992 1993 Galling index'
Treatment level 2 April 9 June 28 August 2 April 24 June 14 August 1992 1993

Nitrogen 89 kg/ha 8 b 72 308 b 3 20 234 2.9 1.6
158 kg/ha 16 a 41 657 a 5 34 240 2.8 1.9

Cultivar Coker 371-Gold 9 38 473 4 31 138 b 2.4 b 1.2 b
NK-326 15 75 491 4 23 336 a 3.3 a 2.3 a

Cover crop Weeds 11 56 653 a - - - 3.3 a
Hairy indigo 9 44 270 b - - - 2.7 b
Forage sorghum 15 69 372 b - - - 2.6 b

ANOVA Block NS NS NS NS NS NS
Nitrogen (N) ** NS * NS NS NS NS NS
Cultivar (C) NS NS NS NS NS * *** *
Cover crop (Cc) NS NS * **
NxC * NS * NS NS NS * *
NxCc NS NS NS - - - NS
CxCc NS * NS - - - NS
NxCxCc NS NS NS - - - *
Data are means of main effects. The number of J2 were transformed with logo (x + 1) before being subjected to ANOVA. Means within treatments of each date followed by the same or by no letter do not differ at P < 0.05 according to ANOVA for nitrogen and cultivar, or Duncan's multiple-range test for cover crop.
*, **, *** represent P < 0.05, P < 0.01, and P < 0.001, respectively, NS = not significant at P < 0.05.
- = no observation.
a The galling index scale: 0 = no galls, 1 = 1-10, 2 = 11-20, 3 = 21-55, 4 = 56-80, 5 = 81-100% of roots galled.


00




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TABLE 4-4. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar history, and their effects on the number of second-stage juveniles (J2) of Meloidogyne spp. in 100 g of soil collected at planting and after the final harvest in 1992.


Number of J2 per 100 g of soil

2 April 28 August

Cultivar history 89 kg N 158 kg N 89 kg N 158 kg N

Coker 371-Gold 8 a A 9 b A 477 a A 469 b A
NK-326 8 a B 23 a A 140 b B 844 a A

The values were transformed with log,( (x + 1) before being subjected to ANOVA; the same lower case letters in columns or upper case letter in rows for the same sampling dates indicate no significant difference at P < 0.05 according to ANOVA.











TABLE 4-5. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar history, and their effects on the root-galling index in 1992 and 1993.


Root-galling index

1992 1993

Cultivar history 89 kg N 158 kg N 89 kg N 158 kg N

Coker 371-Gold 2.7 a A 2.2 b A 1.0 b A 1.4 b A
NK-326 3.1 a A 3.4 a A 2.2 a A 2.4 a A

The values were transformed with arcsin (Ax) before being subjected to ANOVA; the same lower case letters in columns or upper case letter in rows indicate no significant difference at P < 0.05 according to ANOVA.
a The galling index scale: 0 = no galls, 1 = 1-10, 2 = 11-20, 3 = 21-55, 4 = 5680, 5 = 81-100% of roots galled.





83


fertilizer and the tobacco cultivar history was found to affect the galling indices (P < 0.001) (Tables 4-3 and 4-5). The high rate of ammonium nitrate resulted in more severe root galling on NK-326 than on Coker 371-Gold in both years (P < 0.05) (Table 4-5). This statement is valid for the low level of nitrogen in 1992, but not for 1993. In comparison with a root galling index of 3.8 in the weed fallow plots, the severity of root galling decreased to 2.0 and 1.7 after Coker 371-Gold plots were covered with hairy indigo or forage sorghum, respectively (data not shown). Galling indices differed between the blocks in both seasons (P < 0.001) (Table 4-3).

A high percentage of the J2 had endospores of P. penetrans attached (Table 4-6). The percentage attachment and the number of endospores attached per juvenile remained fairly constant over the six sampling dates. In June and August of 1992, and in August 1993, the percentages of J2 with attached endospores were higher in the root rhizosphere of Coker 371-Gold than in NK-326 (P < 0.05). The autumn cover crops affected the percentage of attached J2 only on the first sampling date of the first year. At that time, more J2 were encumbered with endospores after an autumn cover with weeds or hairy indigo than after forage sorghum. No effect of the inorganic nitrogen fertilizer was observed on the percentage of J2 with attached endospores of P. penetrans or the number of endospores per juvenile. The percentage of females infected by P. penetrans was not affected by the treatments. Variations among blocks affected endospore attachment and the female infection rate at various sampling dates.

An interaction between tobacco cultivar history and autumn cover crop revealed that in the root rhizosphere of NK-326, more J2 were encumbered with endospores after weed and hairy indigo autumn cover than after forage sorghum (P < 0.05) (data not shown). There was an interaction among the nitrogen levels, the tobacco cultivar history, and the autumn cover crops affecting the percentage of J2 attached with






TABLE 4-6. Percentage of second-stage juveniles (J2) of Meloidogyne spp. with attached endospores of Pasteuria penetrans, average number of endospores attached per juvenile in the soil, and percentage of P. penetrans-infected females collected from two tobacco cultivars in a field treated with two nitrogen fertilizer rates, and three autumn cover crops in 1992 and 1993.


Percentage of J2 with attached endospores Number of endospores per juvenile % infected
1992 1993 1992 1993 females
Treatment 2 9 28 2 24 14 2 9 28 2 24 14
Treatment level April June August April June August April June August April June August 1992 1993

Nitrogen 89 kg/ha 24 56 64 66 42 73 0.9 3.1 4.3 1.4 0.4 1.8 69 73
158 kg/ha 23 62 55 29 50 64 1.8 3.4 3.3 0.6 0.6 1.0 52 56

Cultivars Coker-371 Gold 23 67 a 65 a 55 53 78 a 1.3 3.3 4.0 1.1 0.5 2.1 a 66 68
NK-326 24 51 b 54 b 40 39 58 b 1.4 3.1 3.5 0.8 0.4 0.8 b 55 61
Cover
crops Weeds 37 a 63 60 - - - 2.0 3.8 4.3 - - - 59
Hairy indigo 24 a 66 65 - - - 1.1 2.4 4.1 - - - 67
Forage
sorghum 8 b 48 54 - - - 1.0 3.5 3.0 - - - 55

ANOVA Block NS * NS NS ** ** NS NS * NS NS * ** *
Nitrogen (N) NS NS NS NS NS NS NS NS NS NS NS NS NS NS
Cultivar (C) NS * ** NS NS ** NS NS NS NS NS * NS NS
Cover crop
(Cc) ** NS NS - - - NS NS NS - - - NS
NxC NS NS NS NS NS NS NS NS NS NS NS NS NS NS
Nx Cc NS NS NS - - - NS NS NS - - - -
Cx Cc * NS NS - - - NS NS NS - - - -
N x C x Cc NS NS - - - NS NS NS - - - -
Data are means of main effects. The percentages were transformed with arcsin ('x) before analysis and the average numbers of endospores per juvenile were transformed with loglo (x + 1) before being subjected to ANOVA. Means within treatments at each date followed by the same or by no letter do not differ at P < 0.05 according to ANOVA for nitrogen and cultivar, or Duncan's multiplerange test for cover crop.

*, **, * represent P < 0.05, P < 0.01, and P < 0.001, respectively, NS = not significant at P < 0.05.

- = no observation.


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Full Text

PAGE 1

SUPPRESSION OF MELOIDOGYNE SPP. BY PASTEURIA PENETRANS By ELKE WEIBELZAHL FULTON A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 1998

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Before you can reach to the top of a tree and understand the buds and flowers, you will have to go deep to the roots, because the secret lies there. And the deeper the roots go, the higher the tree goes. Nietzsche

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a man who taught me many of my skills, my father, Armin Willi Paul Weibelzahl.

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ACKNOWLEDGMENTS I am grateful to the efforts of many people, and I am bothered by the omission of many names from these two pages. My thanks go to all who have helped me. First and foremost, Don W. Dickson, my chairman, took a chance on me and has been patient and generous in providing the financial support, direction, and freedom that I have enjoyed in the course of obtaining my doctorate in his laboratory. Robert McSorley, Dave Mitchell, and Ben Whitty, my committee members, provided tremendous assistance with their kindness and encouragement. Robert McSorley reviewed my manuscripts and dissertation and was most supportive with his statistical advice. Dave Mitchell taught me to consider especially the philosophical aspects of science. By working with Ben Whitty, I developed a major interest in applied nematology. Without the recommendations by Simon Gowen and Richard Sikora to pursue advanced education in tropical and subtropical regions, I would not have discovered my favorable working environment. Thanks go to them as well. I also wish to express sincere appreciation to Reginald Wilcox and Tom Hewlett, whose helping hands and cheerful spirits made the technical aspects of field and greenhouse projects run smoothly. Jay Harrison deserves recognition for supporting me with his statistical ingenuity. Many thanks go to Steve Lasley and all those computer wizards who are responsible for the knowledge I have gained about computers. iv

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Deserving additional, special mention are John Strayer, who helped me begin; Charlie Tarjan, who helped me to get through; and Grover Smart, who helped me complete the graduate program at the University of Florida. I am also thankful for the privilege of having worked with the wonderful students they have recruited for the department. Special thanks must be given to Georgina Robinson, Bettina Moser, and Leandro Freitas for their friendship and encouragement throughout the years in graduate school. I am most grateful to my husband, Michael, our son, Riley, and our daughter, Kyra, all of whom have entered my life in the course of obtaining my doctorate. Their tremendous patience, countless compromises, and never-ending love surrounded me with an ocean of emotional support. Finally, I wish to thank my mother, Anna Weibelzahl, and my faithful siblings and friends on the other side of the Atlantic for their alliance despite the enormous distance between us. They all did a wonderful job in keeping my spirits up. V

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TABLE OF CONTENTS Page ACKNOWLEDGMENTS iv LIST OF TABLES ix LIST OF FIGURES xiii ABSTRACT xv CHAPTERS 1 INTRODUCTION 1 Biological Control of Nematodes 1 Introduction 1 Historical Background 2 Taxonomic Status of the Genus Pasteuria 3 Life Cycle of Pasteuria spp 6 Ecology of Pasteuria spp 13 Biological Control of Nematodes by Pasteuria spp 19 Biological control attributes 19 Natural control 21 Inundative application 22 Integrated nematode management 23 Useful Methods and Techniques for Studying Pasteuria spp 24 Detection of Pasteuria spp. in soils and nematodes 24 Isolation of Pasteuria spp. from soils and nematodes 25 Quantification of endospores in soils, root powder or suspension 26 Culture and preservation of Pasteuria penetrans 27 Attachment 28 Objectives 29 2 POPULATION DEVELOPMENT OF MELOIDOGYNE ARENARIA RACE 1 AND PASTEURIA PENETRANS IN A 6.5-YEAR MICROPLOT STUDY 30 Introduction 30 Materials and Methods 31 Results 33 Discussion 47 vi

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3 USE OF MICROWAVE HEATING IN EVALUATION OF A MELOIDOGYNE A/?£:A^A/?M-SUPPRESSIVE SOIL CONTAINING pasteuria penetrans and its application in a suppressive-soil test 51 Introduction 51 Materials and Methods 52 Microwave Treatment 52 Suppressive Soil Test 54 Nematode Origin 56 Statistical Analysis 56 Results 57 Microwave Treatment 57 Suppressive Soil Test 60 Discussion 67 4 POPULATION DEVELOPMENT OF MELOIDOGYNE SPP. AND PASTEURIA PENETRANS AS AFFECTED BY CULTURAL PRACTICES IN TOBACCO 72 Introduction 72 Materials and Methods 73 Nematode Populations 74 Pasteuria penetrans Isolates 74 Laboratory Experiment 75 Field Experiment 75 Results 77 Laboratory Experiment 77 Field Experiment 77 Discussion 85 5 SUPPRESSION OF MELOIDOGYNE INCOGNITA AND M. JAVANICA BY PASTEURIA PENETRANS IN FIELD SOIL 88 Introduction 88 Materials and Methods 89 Soil Treatments 89 Laboratory Experiments 90 Greenhouse Experiment 91 Nematode Origin 92 Statistical Analysis 93 Results 93 Laboratory Experiments 93 Greenhouse Experiment 93 Discussion 99 vii

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6 MELOIDOGYNE ARENARIA AND PASTEURIA PENETRANS POPULATION DENSITY DEVELOPMENT IN METHYL BROMIDE TREATED SOIL AS AFFECTED BY AN INTERCROPPING SYSTEM 101 Introduction 101 Materials and Methods 103 Results 109 Discussion 124 7 Summary 128 LIST OF REFERENCES 130 APPENDIX 151 BIOGRAPHICAL SKETCH 152 viii

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LIST OF TABLES Table page 1L Geographic distribution of Pasteuria spp 13 21. Effect of Meloidogyne arenaria alone and in combination with Pasteuria penetrans, and of a rye, vetch, or bare fallow winter cover crop, on yield and performance of peanut in the 5* year (fall of 1991) of a 6.5-year microplot experiment 35 2-2. Effect of Meloidogyne arenaria alone and in combination with Pasteuria penetrans, and of a rye, wheat, or bare fallow winter cover crop on yield and performance of peanut in the 6* year (1992) of a 6.5-year microplot experiment 36 2-3. Effect of Meloidogyne arenaria alone and in combination with Pasteuria penetrans, and of a rye, wheat, or bare fallow winter cover crop, on yield and perfoiTnance of peanut in the 7* year (1993) of a 6.5-year microplot experiment 37 24. The percentage of second-stage juveniles (J2) of Meloidogyne arenaria infected with Pasteuria penetrans and the average number of endospores per juvenile in peanut microplots infested with M. arenaria and P. penetrans in the spring of 1987, and rotated with rye, vetch or wheat, and bare fallow as winter cropping sequence 42 31. Colony forming units (cfu) of soil fungi and attachment of Pasteuria penetrans endospores to second-stage juveniles (J2) of Meloidogyne arenaria in untreated soil and soil autoclaved twice for 1.5 hours at 55 kPa, microwaved for 3 minutes/kg of soil, or air dried for 2 weeks in the greenhouse 61 3-2. ANOVA table for the effect of soil source, soil treatments, and Meloidogyne arenaria race 1 inoculum levels on nematode reproduction and fresh root weights of peanut cv. Florunner 62 3-3. Effect of autoclaving, microwaving, and air-drying of soil infested with Meloidogyne arenaria race 1 alone on nematode reproduction, percentage of females infected by Pasteuria penetrans, and fresh root weights of peanut cv. Florunner following inoculation with 0 or 2,000 second-stage juveniles 63 3-4. Effect of autoclaving, microwaving, and air-drying of soil infested with Meloidogyne arenaria race 1 and Pasteuria penetrans on nematode reproduction, percentage of females infected by P. penetrans and fresh root ix

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weights of peanut cv. Florunner following inoculation with 0 or 2,000 second-stage juveniles of M. arenaria race 1 64 35. Effect of autoclaving, microwaving, and air-drying of soil maintained free of nematodes and Pasteuria penetrans on nematode reproduction, percentage of females infected with P. penetrans and fresh root weights of peanut cv. Florunner following inoculation with 0 or 2,000 second-stage juveniles of Meloidogyne arenaria race 1 66 41. .Attachment pattern on second-stage juveniles (J2) of Meloidogyne javanica and M. incognita over three generations of Pasteuria penetrans isolate P1 10 from M. incognita infecting tobacco 78 4-2. Attachment pattern on second-stage juveniles (J2) of Meloidogyne javanica and M. incognita over three generations of Pasteuria penetrans isolate P120 from M. javanica infecting tobacco 79 4-3. Population density development of Meloidogyne spp. in 1992 and 1993 as determined by the number of second-stage juveniles in the soil, and the galling indices of roots of two tobacco cultivars treated with two inorganic nitrogen rates and three autumn cover crop treatments 81 4-4. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar history, and their effects on the number of second-stage juveniles (J2) of Meloidogyne spp. in 100 g of soil collected at planting and after the final harvest in 1992 82 4-5. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar history, and their effects on root-galling indices in 1992 and 1993 82 4-6. Percentage of second-stage juveniles (J2) of Meloidogyne spp. with attached endospores of Pasteuria penetrans, average number of endospores attached per juvenile in the soil, and percentage of P. penetrans-infccted females collected from two tobacco cultivars in a field treated with two nitrogen fertilizer rates and three autumn cover crops in 1992 and 1993 84 47. Interaction between inorganic nitrogen fertilizer levels, tobacco cultivar history, and cover crops, and their effects on the percentage of second-stage juveniles (J2) encumbered with endospores of Pasteuria penetrans at planting in 1992 85 51. Survival of soil fungi and Pasteuria penetrans in untreated soil or soil autoclaved twice for 1 .5 hours at 55 kPa, microwaved for 3 minutes/kg of soil, or air-dried for 2 weeks in the greenhouse 94 5-2. ANOVA table for the effects of tobacco cultivars, soil treatments, and Meloidogyne incognita race 1 inoculum levels on nematode reproduction and plant performance 95 5-3. Effect of autoclaving, microwaving, and air-drying on soil suppressiveness to Meloidogyne spp. and on the expression of root-knot on tobacco cultivar Coker-371 Gold following inoculation with 0 or 2,000 second-stage juveniles of M. incognita race 1 96 5-4. Effect of autoclaving, microwaving, and air-drying on soil suppressiveness to Meloidogyne spp. and on the expression of root-knot on tobacco cv. X

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Northrup King-326 following inoculation with 0 or 2,000 second-stage juveniles of M. incognita race 1 97 6-1. Grouping of 90 microplots based on the mean number of Pc!5/eMna penetrans endospores attached per second-stage juvenile of Meloidogyne arenaria race 1 in a bioassay on microplot soil conducted in the spring of 1994 110 6-2. Main effect of differences of groups with different Pasteuria penetrans endospore densities in microplots intercropped to two cycles of com and beans in rotation with peanut on the population development of P. penetrans and Meloidogyne arenaria race 1 as determined by endospore attachment bioassay, soil sample extraction, and galling index 1 1 1 6-3. Mean number of Pasteuria penetrans endospores attached per second-stage juvenile of Meloidogyne arenaria race 1 in bioassays of microplot soil with different initial endospore densities intercropped with two cycles of com and beans in 1994, planted to hairy vetch in the winter of 1994-95, and cropped with peanut in the summer of 1995 112 6-4. Percentage of second-stage juveniles of Meloidogyne arenaria race 1 with endospores of Pasteuria penetrans attached in bioassays of microplot soil with different initial endospore densities intercropped with two cycles of com and beans in 1994, planted to hairy vetch in the winter of 1994-95, and cropped with peanut in the summer of 1995 116 6-5. Number of second-stage juveniles of Meloidogyne arenaria race 1 per 100 cm^ of microplot soil with different initial Pasteuria penetrans endospore densities intercropped with two cycles of com and beans in 1994, planted to hairy vetch in the winter of 1994-95, and cropped with peanut in the summer of 1995 1 17 6-6. Number of Pasteuria penetrans endospores attached per second-stage juvenile of Meloidogyne arenaria race 1 in microplot soil with different initial endospore densities intercropped with two cycles of com and beans in 1994, planted to hairy vetch in the winter of 1994-95, and cropped with peanut in the summer of 1995 119 6-7. Percentage of second-stage juveniles of Meloidogyne arenaria race 1 with endospores of Pasteuria penetrans attached in microplot soil with different initial endospore densities intercropped with two cycles of com and beans in 1994, planted to hairy vetch in the winter of 1994-95, and cropped with peanut in the summer of 1995 121 6-8. Root galling rates of two intercroping systems with com and beans rotated with peanut grown in microplots grouped by different Pasteuria penetrans endospore densities, and inoculated with Meloidogyne arenaria in the spring of 1994 122 6-9. Spearman correlation coefficients of ranked data obtained from bioassays and from the analysis of soil extracted from 90 microplots containing varying population densities of Pasteuria penetrans endospores; plots were intercropped with two cycles of com and beans in 1994, planted to hairy xi

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vetch in the winter of 1994-95, and cropped with peanut in the summer of 1995 123 Effect of Meloidogyne arenaria (RKN) alone and in combination with Pasteuria penetrans (RKN + Pp), and of a rye, vetch, or bare fallow winter cover crop on the ring nematode population density in the falls of 1991 to 1993 of a 6.5-year microplot experiment 151 xii

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LIST OF FIGURES Figure page 11. Life cycle of Meloidogyne sp. and its bacterial parasite, Pasteuria penetrans 9 21. Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100 cm^ of soil and the number of endospores attached per juvenile extracted from peanut microplots infested with M. arenaria, and M. arenaria plus Pasteuria penetrans 34 2-2. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1991 40 2-3. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1992 41 2-4. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1993 42 2-5. Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100 cm^ of soil extracted from soil in microplots with A) M. arenaria, and B) M. arenaria plus P. penetrans 43 2-6. Number of attached endospores per juvenile extracted from soil in microplots with A) M. arenaria, and B) M. arenaria plus P. penetrans 45 27. Log presentation of peanut yield from plots treated with Meloidogyne arenaria race 1 (RKN), M. arenaria plus Pasteuria penetrans (RKN + Pp), and untreated (Control); and the number of endospores per secondstage juvenile in RKN and RKN + Pp plots 46 31 . Effect of microwave radiation treatment of 1 kg of soil containing Pasteuria penetrans on A) the attachment of P. penetrans endospores to Meloidogyne arenaria race 1 and B) the survival of selected fungi as determined by the number of colony-forming units 57 3-2. Relationship between the soil moisture content and the number of endospores of Pasteuria penetrans attached per second-stage juvenile (J2) of Meloidogyne arenaria race 1 after 48 hours exposure at room temperature 58 3-3. Effect of soil moisture content and microwave treatment time on the attachment of Pasteuria penetrans endospores to Meloidogyne arenaria race 1 58 5-1. Effect of soil treatments on a soil suppressive to Meloidogyne spp. and the expression of root-knot on tobacco cultivar Coker 37 1 Gold following inoculation with 2,000 second-stage juveniles of M. incognita race 1 84 xiii

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61 . Cut away illustration of a sweet com and pole bean intercropping system in field microplots (76 cm diam.) 104 6-2. Cut away illustration of a peanut crop in field microplots (76 cm diam.) 106 6-3. Relationship between the initial endospore population density and A) the final number of endospores per juvenile as determined by bioassay, B) the final number of second-stage juvenile (J2) per 100 cm^ of soil, and C) the final number of endospores per J2 as determined by soil sample extraction 113 6-4. 'Rate of change for A) the number of endospores of Pasteuria penetrans per juvenile of Meloidogyne arenaria as determined by bioassay, B) the number of second-stage juveniles (J2)/100 cm' of soil, and C) the number of endospores/juvenile, as determined in soil sample extraction based on the initial endospore population density, and the change over the 1995 season 1 14 xiv

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy • SUPPRESSION OF MELOIDOGYNE SPP. BY PASTEURIA PENETRANS By Elke Weibelzahl-Fulton May 1998 Chairman: Don W. Dickson Major Department: Entomology and Hematology Suppression of plant-parasitic nematodes with microbial agents is an altemative or supplemental management tactic that is receiving increased interest among nematologists. One nematode antagonist, the endospore-forming bacterium Pasteuria penetrans, has shown great potential in suppressing field populations of several plant-parasitic nematodes throughout the world. This obligate parasite of mainly root-knot nematodes, Meloidogyne spp., was studied for its potential to suppress M. arenaria on peanut, and M. incognita and M. javanica on tobacco. The objectives were to monitor the population densities of the root-knot nematode and P. penetrans in peanut microplots and in a naturally infested tobacco field, to develop a suppressive-soil test that allows the determination of the role of P. penetrans in nematode-suppressive soils, and to evaluate the effect of cultural practices, such as crop rotation, autumn cover crops, resistance and fertilizer regimes on the abundance of P. penetrans. Within 4 years after inoculafion or contamination by P. penetrans, M. arenaria was nearly eliminated from microplot soil, and peanut yields were similar to that of the nematode free control. The P. penetrans-infested soil remained suppressive to M. arenaria for 3 years. Similar results were observed in a tobacco field. XV

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After 4 years of tobacco monoculture, a mixed population of M. incognita, M. javanica, and M. arenaria was suppressed to non-damaging levels. Two P. penetrans isolates were more pathogenic to M. incognita than to M. javanica. Inorganic nitrogen fertihzer rates and host resistance to M. incognita had inconsistent effects on the endospore build-up, which was favored by cultivation of susceptible cover crops. A microwave radiation treatment of 3 minutes/kg of soil containing 6% to 7% water reduced the fungal population in soil samples without impairing attachment of P. penetrans endospores to nematode juveniles. This treatment allowed the separation of nematode suppression by fungi from that caused by P. penetrans. Nematode suppressiveness in microplot and field soil was preserved after microwave and air-drying treatments, but not after soil was autoclaved. The bacterial parasite, P. penetrans, was identified as the main contributor to the suppressiveness of the microplot and the field soil. xvi

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CHAPTER 1 INTRODUCTION Biological Control of Nematodes Introduction Nematologists are continuously unveiling the biological control potential of Pasteuria penetrans (Thome, 1940) Sayre and Starr, 1985, a mycelial endospore-forming bacterial parasite of mainly root-knot nematodes. The ecological history of P. penetrans in association with Meloidogyne spp., as well as knowledge about its role in nematode management strategies, is essential for improving the use of the organism as an effective biological control agent. This information also is important for recognition and effective exploitation of soils naturally infested with P. penetrans. In 1991, soils suppressive to the peanut root-knot nematode, M. arenaria (Neal) Chitwood race 1, were produced in microplots at the University of Florida Agronomy Farm, Green Acres, Alachua County, Florida. Ninety microplots (76 cm in diameter) were established with 30 inoculated centrally with relatively low numbers of P. penetrans and Meloidogyne arenaria, 30 inoculated with M. arenaria alone, and 30 left uninoculated as control plots (Oostendorp et al., 1991a). The microplots were planted to peanut in the spring of 1987. Within 3 years, yields of plots infected with P. penetrans and M. arenaria increased to a level comparable to the untreated control and the damage from root-knot nematodes decreased, suggesting that the soil became suppressive to the nematode. A 1

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2 minimum of 3 years of intensive cropping was required for P. penetrans to amplify to suppressive levels. During the same period, a tobacco field that was naturally infested with Meloidogyne spp. was discovered to contain a soil suppressive to the nematode. This site was located about 300 meters from the microplot site. The bacterial parasite P. penetrans was reported to cause the nematode suppression (Chen et al., 1994). The goal of this project was to monitor the dynamics of the bacterium in the two sites and further establish the relationship of P. penetrans to suppressive soil. Historical Background Biological control of nematodes has been of great interest to nematologists for almost as long as plant-parasitic nematodes have been known to damage crops (Stirhng, 1991). Cobb (1920) considered using parasites and predators of nematodes as biological control agents when he suggested transferring predacious nematodes to sugarbeet fields in order to control populations of Heterodera schachtii Schmidt. Thome (1927), however, questioned the economic benefits of predacious nematodes, and further studies on their potential for biological control of plant-parasitic nematodes came to an end. Linford's (1937, 1939) early efforts to use predacious fungi to control plant-parasitic nematodes stimulated interest in the nematode-trapping fungi in France, the United States, England, and the former USSR (Stiriing, 1991). Due to the lack of success of these early experiments, the interest in biological control declined. It was not until the late1970s that interest in biological control revived when the potential health and environmental problems associated with the use of some nematicides surfaced (Kerry, 1990; 1993; Stirling, 1991; Thomason, 1987). The suspension of two of the most reliable soil fumigants, DBCP (1,2dibromo-3-chloropropane) and EDB (ethylene dibromide), and the lack of promising new

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3 candidate nematicides, led nematologists to re-evaluate their goals in terms of nematode management research. This fact, plus the classical studies that demonstrated fungal antagonists suppressed populations of Heterodera avenae Woll. in Europe (Kerry et al., 1980; 1982) and other studies showing that fungal antagonists also suppressed Meloidogyne spp. (Stirling and Mankau, 1978; Stirling et al., 1979), rekindled interest in biological control. In recent years, some attempts have been made to market various fungal agents for nematode control (Al-Hazim et al., 1993; Timm, 1987), but the products generally have not been accepted. Unfortunately, there are still no widely used examples of the contrived use of biological antagonists to control plant-parasitic nematodes. Although there are many likely candidates (Dickson et al., 1994; Jansson, 1988; Kerry, 1987; 1988; Mankau, 1975b; 1980b; Sayre et al., 1988; Stirling, 1988; 1991; Tribe, 1980), the resources devoted to research on biological control of nematodes have been relatively limited (Stirling, 1991). There is optimism, however, that biological control will play an increasingly important role in future nematode management programs. The search for nematode antagonists has generally centered on predacious and parasitic fiingi and bacterial parasites. One such bacterium, P. penetrans, was identified and considered to have great potential for biological control of root-knot nematodes (Dickson et al., 1994). A Pasteuria sp. was first associated with parasites of plantparasitic nematodes when Thome (1940) reported it as Duboscqia penetrans parasitizing Pratylenchus pratensis. Since the first reports that Pasteuria penetrans parasitized rootknot nematodes (Mankau, 1980a; 1980b; Stirling and White, 1982; Stirling, 1984), the organism became the subject of an increasing number of research projects (Stirling, 1991). Meloidogyne spp. were reported to be suppressed successfully in numerous pot, microplot, and field experiments (Brown et al., 1985; Channer and Gowen, 1988; Chen et al., 1996b;

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4 1997c; Daudi, 1990; Dube and Smart, 1987; Jaya Raj and Mani, 1988; Maheswari and Mani, 1988; Maheswari et al., 1987; 1988; Oostendorp et al., 1991a; Stirling, 1984; Tzortzakakis, 1994a; Vargas et al., 1992; Weibelzahl-Fulton et al., 1996). However, limitations in mass production methods of P. penetrans have prevented its commercialization. Taxonomic Status of the Genus Pasteuria The genus Pasteuria comprises a group of Gram positive mycelial and endosporeforming bacteria that parasitize bodies of invertebrates, including nematodes (Williams et al., 1989a). The initial studies of the genus Pasteuria date back to 1888, when Metchnikoff described an internal parasite of two Daphnia spp. (water fleas) as Pasteuria ramosa (Metchnikoff, 1888). Partial credit for the discovery of the bacterium also should be given to Cobb (1906), who noted Pasteuria sp. as an internal parasite of nematodes, sketched it, and described it to be "perhaps monads." The taxonomy of this nematode parasite, however, has been subject to continuous confusion ever since Thome (1940) described the first member of the group and, believing it was a protozoan, named it Duboscqia penetrans. He could not have realized its bacterial nature, mainly because ultrastructural techniques were not available to him and the concept of the prokaryotic cell had not yet been introduced. Thome's (1940) description and nomenclature persisted for 35 years. Although other scientists worked with the organism and questioned its taxonomic placement (Williams, 1960), it was not until the reexamination by Mankau (1975a), using electron microscopy, that its affinity to bacteria rather than protozoa was revealed. The nematode parasite was renamed Bacillus penetrans (Thome, 1940) Mankau, 1975.

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5 Only a decade ago, Sayre et al. (1983) established a sound basis for the taxonomy of this group of hyperparasites. Detailed comparisons of the genera Pasteuria and Bacillus cleared up the confusion and revealed major similarities between the nematode parasite and Pasteuria ramosa (Sayre and Wergin, 1977; Sayre et al, 1983). Their close relati-onship became obvious in several distinctive morphological characteristics, such as the dichotomously branched mycelial microcolonies that give rise to fragmentation and sporangia, and finally to endogenous spores formed within the old mother cell wall. Similarities also had been shown at the ultrastructural level in the unique forms and sequences of life stages of the two organisms (Sayre and Starr, 1985). The parasite of plant-parasitic nematodes, previously known as Bacillus penetrans, was then placed in the genus Pasteuria and renamed Pasteuria penetrans. However, the newly named bacterium was by no means a uniform entity. Different isolates of this organism were found to differ in their physical and pathological characteristics (Bird et al., 1990). The description of two new species followed: Pasteuria thomei Starr & Sayre (1988), which parasitized the lesion nematode Pratylenchus brachyurus (Godfrey), and Pasteuria nishizawae Sayre, Wergin, Schmidt & Starr (1991), which parasitized cyst nematodes, Globodera rostochiensis WoU. and Heterodera glycines Ichnohe. A fifth species, which parasitized the pea cyst nematode, Heterodera goettingiana Liebscher, has been reviewed for taxonomic classification (Sturhan et al., 1994). Numerous Pasteuria spp. have endospores distinctly smaller or larger than those previously described (Chen, 1996; Ciancio and Mankau, 1989; Ciancio et al., 1992; 1994; Giblin-Davis, 1990; Giblin-Davis et al., 1990; Jaffee et al., 1985; Noel and Stanger, 1994; Sayre et al., 1985). Some isolates display a cross-genera host range and various biological characteristics (Bhattacharya and Swamp, 1988; Mankau, 1975; Oostendorp et al., 1990; Pan et al, 1993; Sharma and Davies, 1996; Vargas and Acosta, 1990). Thus, Pasteuria spp. are being differentiated by host specificity,

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6 developmental characteristics, and shape and size of sporangia and endospores (Sayre and Starr, 1989). Based on a study of host records and endospore morphometries of pathotypes described previously as the P. penetrans group, Ciancio et al. (1994) concluded that host taxonomy and endospore dimensions are of limited value to the definition of Pasteuria spp. This would partially explain a discovery by Davies et al. (1992), who identified different surface proteins of endospores in three P. penetrans populations, all parasites of Meloidogyne incognita. A recent study shows that P. penetrans produces heterogeneous endospores (Davies et al., 1994). These subpopulations of endospores show specificity to various nematode populations. No definite criteria or genetic data are available to establish whether distinct species of Pasteuria differing in their biology and physiology share common hosts and morphometries, or whether some or even all of the Pasteuria members should be considered as pathotypes of a unique species, regardless of hosts and morphometries (Ciancio et al., 1994). The confusion in taxonomy of Pasteuria probably will not be clarified until the bacterial genom properties are elucidated. Axenic cultivation of the bacteria is crucial to understanding the biology and taxonomy of Pasteuria, but artificial cultivation of Pasteuria spp. has not been successful (Bishop and Ellar, 1991; Pre vie and Cox, 1993; Reise et al., 1988). Life Cycle of Pasteuria spp. The life cycle of P. penetrans in root-knot nematodes was first described in detail by Sayre and Wergin (1977). Their observations were confirmed by Imbriani and Mankau

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7 (1977) and additional studies have contributed useful information on the life cycle (Davis et al., 1988; Mankau and Prassad, 1977; Sayre, 1988; Sayre and Starr, 1985; Sturhan et al., 1994). Pasteuria penetrans is a density dependent obligate parasite that grows and multiplies within the bodies of plant-parasitic nematodes. Up to 2.5 million nonmotile endospores are produced per female of Meloidogyne spp. (Hewlett and Dickson, 1993) and released into the soil environment upon degradation of the nematode carcass. The developmental stages of all members of Pasteuria spp. parasitic on nematodes appear to be similar, but there are differences in shapes and sizes of endospores and sporangia among isolates obtained from different nematode genera. The following detailed description and illustration of the developmental stages covers P. penetrans and Meloidogyne sp. and draws attention to other species for notable differences (Fig. 1-1). Attachment. The organism encounters host nematodes when endospores passively adhere to the cuticle of a migratory stage of the nematode in soil. In the case of Meloidogyne sp., a single second-stage juvenile may have one to several hundred endospores attached to its cuticle (Davies et al., 1991). Mobility and infectivity of the second-stage juveniles (J2) is reduced when somewhere between 7 and 50 endospores are attached to the juvenile (Brown and Smart, 1985; Davies et al., 1988, 1990; Sell and Hansen, 1987; Stirling, 1984). Since attachment can occur readily in soil or aqueous suspension (Slana and Sayre, 1982), there appear to be no requirements for adhesion other than those found on the endospore or nematode cuticle surface. There is considerable variation in the ability of P. penetrans to attach to and infect species of nematodes, particularly within isolates parasitic on Meloidogyne spp. (Brown and Smart, 1984; Davies et al., 1988; Davies and Danks, 1992; 1993; Oostendorp et al., 1990; Sell and Hansen, 1987; Spaull, 1984; Stirling, 1985; Verdejo-Lucas, 1992). Cuticular variations in ultrastructure and composition between nematode populations

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Fig. 1-1. Life cycle of the root-knot nematode, Meloidogyne sp., with and without its bacterial parasite, Pasteuria penetrans ( outer circle). 1) Second-stage juveniles (J2) entering root tip, 2) migrating intercellulary in cortex, 3) J2 establishing feeding sites in the vascular system; germinating P. penetrans endospores, 4) third-stage juveniles, 5) fourthstage juveniles, 6, 7) young females, 8) female with P. penetrans lays no or few eggs, whereas a healthy female forms an egg mass and lays eggs, and 9) infected female carcass degrades and releases mature endospores into the soil. The inner circle illustrates the life cycle of P. penetrans and its various developmental stages. Cross sections and top views of a) mature endospores, b) endospore attached to cuticle of Meloidogyne sp., c) germinating endospore, d) microcolonies formed in the pseudocoelom of the nematode, e) septations in rapidly growing thallus, f) dichotomously branched hyphae with elongated terminal cells, g) fragmented thaUi separated from the thallus and visible forespore, h) cell wall separates forespore from parasporium of the egg-shaped sporangium, i) differentiation of spore core and perisporal fibers, j) mature endospores surrounded by exosporium and sporangium, and k) endospores released into soil.

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9

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10 (Davies and Danks, 1992; Reddigari et al., 1986) may be responsible for different levels of attachment. The greatest spore attachment occurs when endospores are exposed to Meloidogyne spp. from which they were originally isolated (Davies et al., 1988; Davies et al., 1994; Oostendorp et al., 1990). Some P. penetrans isolates attached indiscriminately to Meloidogyne spp. other than the original host (Stirling, 1985). The infective juvenile of Meloidogyne spp. is the only stage that appears to be parasitized by P. penetrans (Mankau, 1980b), and this could perhaps be attributed to the cuticular surface composition differences observed between the nematode's life stages (McClure and Stynes, 1988). It has been suggested that the surfaces of J2 contain structural carbohydrate recognition units that are probably not collagen, and that these interact with N-acetylglucosamine moieties on the endospore surface that are linked to either glycoproteins or peptidoglycans (Bird et al., 1989; Davies and Danks, 1993; Spiegel et al., 1996). However, it also was suggested that collagens might be involved in attachment (Persidis et al., 1991). Several studies suggest that host specificity is caused by differences in the amount and nature of surface proteins of the endospores (Chen et al., 1997a; Davies et al., 1992, 1994). Infection and development. Endospore attachment to J2 of Meloidogyne sp. is not necessarily followed by infection (De Silva and Gowen, 1994). The endosporeencumbered J2 penetrate the growing roots of a host plant behind the root cap. In the zone of cellular differentiation, the J2 reside in cortical tissue with their heads in the periphery of the vascular tissue where feeding starts (Hussey, 1985). Thereafter, a germ tube emerges from a central pore at the basal side of the endospore and penetrates the cuticle and hypodermis of the nematode (Sayre and Starr, 1985). There is evidence that germ tubes which do not penetrate the cuticle, emerge between the "rim" of the perisporium and the

PAGE 27

11 cuticle of the nematode (Birchfield and Antonopoulus, 1976). Endospore germination appears to be triggered by the onset of feeding (personal observations). The protoplast of the endospore enters the nematode body through the germ tube and develops into vegetative, spherical colonies consisting of a dichotomously branched, septate mycelium (Sayre and Starr, 1985). These early stages (mycehal colonies) are visible with the light microscope as dense granulation within the nematode pseudocoelom. Fragmentation of the thalli separates the mycelial masses, which are then transported in the fluid of the pseudocoelomic cavity through the body of the host nematode. These fragments give rise to microcolonies of four, eight, or more terminal cells. In advanced developmental stages, an increasing number of colonies with two, three, or four clubshaped, enlarged terminal cells are seen (doublets, triplets, and quartets). These each give rise to a single sporangium in which a single endospore is formed. The process of endospore formation appears typical of that found in other endospore-forming bacteria (Chenetal., 1997b). As the sporangium enlarges, a septum is formed in the upper third of the mother cell separating the incipient forespore cytoplasm from the remainder of the cell (Chen et al., 1997b). The septum growing around the forespore finally provides a double-layered membrane that encloses the condensed cytoplasm. The wing-like perisporium and the cortex develop between the two membranes surrounding the forespore. An inner cortex and outer zone, surrounded by an irregular granular epicortical layer develops. The outer spore coat is deposited on the outer membrane and forms the laminar inner coat of the mature endospore. Granular material of the spore mother cell concentrates at the basal side of the endospore and is finally engulfed by the exosporium, which surrounds the entire endospore. Sporangia are of lenticular, almost spheroidal shape, with a round to conical upper part and an irregularly shaped basal part. When and where the endospores are

PAGE 28

released from the sporangia remains unknown. The germination of a single spore is enough to create infection in a Meloidogyne sp. female (Stirling, 1984). Generally the infection rate of females increases with increasing endospore attachment on the juvenile (Davies et al., 1988; Stirling, 1984). • Pasteuria penetrans is morphologically different from P. thomei and P. nishizawae in shape and size of the sporangia and the endospores (Sayre et al., 1988; 1991). Light microscopy revealed that mature endospores of P. penetrans, including parasporal fibers and sporangial wall, are saucer to bowl-shaped, measuring about 4.5 p.m in diameter and about 3.6 |im in height. Endospores of P. nishizawae are of similar shape with a broadly elliptical central body, averaging 5.3 |im in diameter and about 4.3 )im in height. Endospores of P. thomei, however, are rhomboidal shaped and smaller in size, averaging 3.5 |im in diameter and 3. 1 |Lim in height. The translucent circular perisporium is easily differentiated from the distinct spore wall. In P. penetrans and P. nishizawae, the spore wall decreases in thickness toward the center of the basal side of the endospore. In P. penetrans, a morphological discreet area has been observed on the basal side of the endospore, which appears to form the germination pore. Unlike females, males of Meloidogyne spp. seem to become parasitized by P. penetrans rather rarely (Abrantes and Volas, 1988; Freitas et al., 1996; Hatz and Dickson, 1992; Page and Bridge, 1985). Endospores obtained from seven isolates of P. penetrans did not attach to males of M. arenaria (Freitas et al., 1996), and observations by Hatz and Dickson (1992) suggest that infected males were a consequence of sex reversal because no males with a single gonad were observed to contain endospores. Some isolates of P. penetrans have been reported to complete their life cycles in J2 of Meloidogyne spp. (Giblin-Davis et al., 1990; Dickson et al., 1994). 0\h&x Pasteuria spp. from Tylenchulus semipenetrans Cobb and Heterodera avenae have endospores that

PAGE 29

13 are morphologically similar to those of P. penetrans, but they reproduce in J2 and not in females (Davies et al., 1990; Fattah et al., 1989; Kaplan, 1994; Sturhan et al., 1994; Winkelheide, 1993). The isolate specific to T. semipenetrans also reproduce in the male (Kaplan, 1994). Another isolate of P. penetrans was able to reproduce in Pratylenchus scribneri and Meloidogyne spp. (Oostendorp et al., 1990). Pasteuria sp. from Helicotylencus lobus formed mature endospores in the juvenile, female, and male life stages (Ciancio et al., 1992). The life cycle of P. thomei can be completed in any of the juvenile stages and in the adult of its host, Pratylenchus brachyurus (Starr and Sayre, 1988). Ecology of Pasteuria spp. Pasteuria spp. are widespread in different biotopes and habitats (Ciancio et al., 1994). Their spores have the morphological and biochemical features of bacterial endospores (Sayre and Wergin, 1977; Williams et al., 1989), and therefore can tolerate environmental extremes. Factors such as soil moisture, soil temperature, soil pore size, organic matter, and clay content are important in the ecology of P. penetrans. Geographic distribution. Pasteuria spp. are worldwide in distribution and have been reported from many countries (Table 1-1). Moisture level. Endospores of Pasteuria spp. are resistant to desiccation (Williams et al., 1989b). The water content of soils, however, affects hatching and movement of nematodes (Baxter and Blake, 1969) and, therefore, is likely to influence the efficacy of P. penetrans. Davies et al. (1991) reported soil moisture affects the growth of P. penetrans within developing females. However, P. penetrans was cultivated successfully in a hydroponic solution (Serracin et al., 1997), which suggests that the failure

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14 Table 1-1. Geographic distribution of Pasteuria spp. Location Literature source North America Canada U.S.A. U.S.A., Cahfomia U.S.A., Colorado U.S.A., Florida U.S.A., Georgia U.S.A., Hawai U.S.A., Illinois U.S.A., Louisiana U.S.A., Maryland U.S.A., Oregon U.S.A., South Carolina U.S.A., South Dakota U.S.A., Utah Central America Colombia Cuba Dominican Republic Haiti Nicaragua Puerto Rico South America Bolivia Brazil Peru Venezuela Europe Austria Belgium Croatia Denmark England Finland France Germany Greece Hungary Iceland Italy Gonzales et al., 1987; Sayre and Starr, 1988; Sturhan, 1988 Allen, 1941; Altherr and Deboutteville, 1972; Bernard and Niblack, 1982; Ciancio et al., 1994; Minton and Sayre, 1989; Sayre and Starr, 1988; Sturhan, 1988 Ciancio et al.,1992; Ciancio and Mankau, 1989b; Sayre and Starr, 1988 Sayre and Starr, 1988 Chen, 1996; Esser, 1980; Gibhn-Davis et al., 1990; Hewlett et al., 1994; Inserra et al., 1992; Kaplan, 1994; Oostendorp et al., 1990; Sayre and Starr, 1988; Walter and Kaplan, 1990; Weibelzahl-Fulton et al., 1996 Sayre and Starr, 1988; Minton and Sayre, 1989 Ko, 1995; Sayre and Starr, 1988 Noel and Stanger, 1994 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Ciancio and Mankau, 1989a Sturhan, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988; Sturhan, 1985; 1988 Vargas and Acosta, 1990; Vargas et al., 1992 Ciancio and Mankau, 1989b; Page and Bridge, 1985; Sturhan, 1988 Dos Santos, 1981; Sayre and Starr, 1988; Sturhan, 1988 Ciancio and Mankau, 1989a; Ciancio et al., 1994 Sayre and Starr, 1988 Sayre and Starr, 1988 Sayre and Starr, 1988 Ciancio et al., 1994 WilHams, 1960 Davies et al., 1990; Sturhan, 1988 Sayre and Starr, 1988 Sturhan, 1985; 1988 Sayre and Starr, 1988; Sturhan, 1988; 1985; Steiner, 1938; Vovlas et al., 1993; Sayre and Starr, 1988 Ciancio et al., 1994 Sayre and Starr, 1988; Sturhan, 1988 Abrantes and Vovlas, 1988; Ciancio, 1995; Ciancio et al., 1987; 1994; Davies et al., 1990; Roccuzzo and Ciancio, 1991; Walia et al., 1990

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15 Table 1-1. Continued "T"! XT 4.1 1 1„ The Netherlands Kuiper, 1958; Sayre and Starr, 1988 Poland * ill r\r\ A Ciancio et al., 1994 Portugal Abrantes and Vovlas, 1988; Romania Ciancio et al., 1994; Sayre and Starr, 1988 Scotland Sayre and Starr, 1988 Spain Verdejo-Lucas, 1992 Sweden Sayre and Starr, 1988 Switzerland Sayre and Starr, 1988; Sturhan, 1988 Africa Algeria Ciancio et al., 1994 Ethiopia Ciancio et al., 1994 Ivory Coast Sturhan, 1988 Liberia Ciancio et al., 1994 Malawi Sturhan, 1988 Hi* 1 Mozambique Sturhan, 1988; Sayre and Starr, 1988 XT* Nigeria Sayre and Starr, 1988; Sturhan, 1988 Senegal Mankau, 1980 Sierra Leone Ciancio et al., 1994; Sayre and Starr, 1988 Somalia Ciancio et al., 1994 South Africa Sayre and Starr, 1988; Spaull, 1981; Sturhan, 1988 Tanzania Madulu et al., 1994; Siddiqi, 1991 Togo Sayre and Starr, 1988 Uganda Sayre and Starr, 1988 Zaire Sayre and Starr, 1988 Zimbabwe Stubbs and Gowen, 1996 Asia China Pan et al., 1993 Inaia Bhattacharya and Swamp, 1988; Page and Bridge, 1986; Sharma and Davies, 1996; Sharma and Sharma, 1989; Walia et al., 1990 T Iran Barooti, 1989; Maafi, 1993; Sayre and Starr, 1988; Sturhan, 1988 T Iraq Fattahetal., 1989 Israel Ciancio et al., 1994; Sayre and Starr, 1988 Japan Sayre and Starr, 1988; Sayre et al., 1991a; Sturhan, 1988 Pakistan Maqbool andZaki, 1990 South Korea Ciancio et al., 1994 Sn Lanka Ciancio et al., 1994 T T o o n U.S.S.R. Sturhan, 1988; Subbotin et al., 1994 Australia South Australia Bird and Brisbane, 1988; Sayre and Starr, 1988; Stirling and White, 1982; Sturhan, 1988 Islands Azores Sayre and Starr, 1988; Sturhan, 1988 Canary Islands Sayre and Starr, 1988; Sturhan, 1988 Madeira Islands Sayre and Starr, 1988 Malta Ciancio et al., 1994 Mauritius Ciancio et al., 1994; Sayre and Starr, 1988; Williams, 1960 Philippines Ciancio et al., 1994; Sturhan, 1988 Samoa Sayre and Starr, 1988 Sao Tome Ciancio et al., 1994

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16 of Davies' system was probably due to poor development of the root system in wet soils. Isolates of P. penetrans have survived for several weeks in dry, moist, and wet soils, and in soils with fluctuating moisture levels without loss of their ability to attach to their nematode hosts (Oostendorp et al., 1990). Their abundance in soils is positively correlated with increasing amounts of annual rainfall (Ko et al., 1995). Temperature. Attachment, infection, and pathogenesis of Pasteuria spp. is affected by temperature (Ahmed and Gowen, 1991; Freitas et al., 1997; Hatz and Dickson, 1992; Nakasono et al., 1993; Sekhar and Gill, 1990a; Serracin et al., 1997; Serracin-Ulate, 1995; Singh et al., 1990; Stirling, 1981; Stirling et al., 1979, 1990). Stirling (1981) found that temperatures near or above the optimum for the nematode caused P. penetrans to attach at higher rates and to proliferate extensively before the infected female host reached maturity; in contrast, at 20 °C the rate of attachment decreased and females often developed ovaries containing eggs before infection prevented further development. Endospore densities of P. penetrans in soil increase with an increase of the average annual temperature (Ko et al., 1995). Walker and Wachtel (1988) observed that soil solarization with clear polyethylene increased the rate of infection of Meloidogyne javanica by P. penetrans. The higher soil temperature may have led to an increase in attachment rates because of the likely increase in nematode mobility, thus leading to an increased probability of endospore contact with J2; on the other hand the higher temperature may have simply increased the rate of nematode development, thereby increasing endospore numbers in soil. Recent studies, however, have shown that attachment of endospores to J2 was reduced after endospores were exposed to 40, 50, and 60 °C (Freitas et al., 1997). Attachment occurred following heating of endospores to 80 °C for 30 minutes, but P. penetrans did not develop inside the nematode (Dutky and Sayre, 1978). Although the rate of attachment has been reported to be reduced when endospores were treated at 100 °C for

PAGE 33

17 at least 30 minutes, attachment was not prevented (Freitas, 1997; Stirling et al., 1986; Williams et al., 1989b). The greatest receptivity of J2 of M. arenaria to endospore attachment occurred when J2 were treated in water at 30 °C and 35 °C and then exposed to endospores (Freitas et al., 1997). • Soil type. Pasteuria penetrans was detected in a wide variety of soil types ranging from pure sand to organic soils (Sturhan, 1985). Observations in laboratory experiments suggest that an increasing sand content might improve endospore attachment (Singh and Dhawan, 1992). Other workers demonstrated that soil texture had no effect on endospore attachment (Hewlett, personal communication). Sandy soils allow endospores to distribute readily with percolating water (Oostendorp et al., 1990). Organic matter. The organic matter component of soil is unlikely to influence Pasteuria spp., other than providing fungal antagonists of nematodes with more favorable habitat and food source (Kerry, 1993). Habitat and food sources may influence the biological control attributes of P. penetrans (Dube and Smart, 1987; Maheswari and Mani, 1988). Mode of action of organic amendments against nematodes consists of more than the direct effects on nematophagous fungi. When organic soil amendments in the form of oil cakes were added to P. penetrans-Mested soil in a pot experiment, M. javanica was synergistically reduced (Maheswari et al., 1988). Organic amendments can improve soil structure and soil fertility, alter the level of plant resistance, release toxic compounds, and stimulate nematode antagonistic microorganisms (Stirling, 1991). All of these changes by organic amendments may affect soil nematode densities dramatically. Chemical factors. Pasteuria penetrans was detected in acidic as well as in alkaline soils (Sturhan, 1985). However, the optimum pH for attachment of endospores of Pasteuria spp. is between 7.0 and 8.5 (O'Brian, 1980). The endospore's resistance to chemical compounds allows them to survive nematicide applications (Mankau and

PAGE 34

18 Prassad, 1972; Stirling, 1984). Nematostatic field dosages of 1,3-dichloropropene, aldicarb, carbofuran, fenamiphos, and ethoprop had no noticeable effect on P. penetrans (Mankau and Prassad, 1972). Increased attachment by P. penetrans was observed when subnematostatic concentrations of organophosphate or organocarbamate nematicides were applied to soil containing nematodes and endospores (Brown and Nordmeyer, 1985). As reported by Bunt (1987), the low concentration of a nematicide might have increased the random nematode movement in the soil, which would increase the likelihood of contact with bacterial endospores. Freitas (1997) reported that chloropicrin alone or in combination with methyl bromide was highly detrimental to the development of P. penetrans because endospore formation was inhibited. Host-parasite relationship. Pasteuria penetrans parasitism has been recorded for more than 236 nematode species in 102 genera, including plant parasitic and free living species (Ciancio et al., 1994; Sturhan, 1985; 1988). In several observations the hostparasite relationship is based on cuticular attachment only. Stirling (1991) concluded that such published data should be treated with caution, because endospore attachment does not actually determine parasitism. Because of its obligate nature, the bacterium is unable to reproduce in the absence of the host and would, therefore, be disadvantaged if it eliminated its host. With increasing endospore densities in soils, the infectivity of endosporeencumbered J2 of Meloidogyne sp. has been shown to be reduced, thus leading to the production of fewer endospore-filled females (Stirling, 1991). Endospore densities are likely to stabilize at a certain equilibrium level (Stirling, 1991). Williams (1960) and later Spaull (1984) noted this density-dependent relationship between P. penetrans and its Meloidogyne host.

PAGE 35

Biological Control of Nematodes by Pasteuria spp. 19 Biological control attributes Since the time that Sayre and Starr (1988) reported Pasteuria spp. to be parasitic to mostspecies of plant-parasitic nematodes, much effort has been made to understand P. penetrans isolates that parasitize the economically important root-knot nematodes. Pasteuria penetrans has many of the attributes required by a successful biological control agent. The bacterium produces no environmental hazards and reduces or prevents reproduction of its host. Also, it reduces the infectivity of endospore-encumbered juveniles to their plant host. Infectivity may be reduced when as few as 15 endospores are attached to the juvenile (Davies et al., 1988). Juveniles are prevented from invading roots when they are each encumbered with 25 to 30 endospores (Stirling, 1984; Stirling et al., 1990). Host specificity. Isolates of P. penetrans can vary in specificity to different Meloidogyne spp. (Brown and Smart, 1984; Davies et al., 1988; Davies and Danks, 1992; 1993; Oostendorp et al., 1990; Sell and Hansen, 1987; Spaull, 1984; Stirling, 1985; Verdejo-Lucas, 1992). Field populations of Meloidogyne spp. may be diverse, and it is possible that populations of P. penetrans with a restricted host range will affect nematode species to varying degrees. However, if there is time for natural selection to take place, it may be possible for new strains of Pasteuria spp. to evolve that are better fitted to the local environment or that are more virulent to the local nematode population. Davies et al. (1988), and Channer and Gowen (1992) support this concept by demonstrating that an initially poor host of a P. penetrans isolate became more susceptible to the parasite when exposed to endospores grown on nematodes of its own species. However, individual nematodes escaping parasitism produced a generation with increased resistance to attachment by endospores of that particular P. penetrans isolate. The authors concluded

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21 Natural control Plant-parasitic nematodes and their bacterial parasites have co-evolved for a long time (Ciancio et al., 1994). There are few data on the extent of nematode suppression caused by Pasteuria sp. in undisturbed soil or in soils with continuing monoculture. With increasing awareness about the qualities of the hyperparasites, more effort have been invested in long-term field studies, and the processes in which soils become suppressive to nematodes are becoming better understood (Chen, 1996b; 1997c; Dickson et al., 1994; Oostendorp et al., 1991a; Stirling, 1991; Weibelzahl-Fulton et al., 1996). Pasteuria sp. may have little short-term impact on nematode populations at low spore densities because only a few nematodes come in contact with the endospores. However, with each infected root-knot nematode female capable of producing up to 2.5 million endospores, endospore densities increase over time, thereby reaching nematode suppressive levels (Oostendorp et al., 1991a; Chapter 3). Wilhams (1960) and later Spaull (1984) noted that P. penetrans had httle impact on the severity of root-knot nematodes in a natural infestation of both parasites; however, the bacterial parasite may still have been in an early phase of population build-up. Others have reported soils with root-knot nematodes and P. penetrans becoming highly suppressive to the nematode (Bird and Brisbane, 1988; Chen et al., 1994; Dickson et al., 1994; Minton and Sayre, 1989; Stirling, 1984; Weibelzahl-Fulton and Dickson, 1996; Weibelzahl-Fulton et al., 1996). When rootknot nematode and P. penetrans infested soils were tested for their suppressiveness, the rate of reproduction of Meloidogyne spp. was frequently reduced in untreated soils compared to soils in which P. penetrans had been eliminated by autoclaving (Dickson et al., 1994; Weibelzahl-Fulton et al., 1996). Nematode suppression was enhanced by cultivation of susceptible hosts for several consecutive years.

PAGE 37

In a pot experiment, soil naturally infested with Belonolaimus longicaudatus Rau and a Pasteuria species specific to the nematode was not suppressive to its host nematode for the first 6 months (Giblin-Davis et al., 1990), however, at 12 months post-inoculation there was an increase in the number of Pasteuria-T\\\td. nematodes. The nematode density was significantly lowered (Giblin-Davis, 1990). Inundative application Despite the difficulties scientists originally faced in working with spore-infested soil, P. penetrans gave promising results against M. incognita in pot experiments (Mankau, 1975b). An improvement of the mass-production technique allowed scientists to produce a spore-laden, easily handled root powder (Stirling and Wachtel, 1980). This provides a means for more extensive testing and further confirmation of the bacterium's potenfial as a biological control agent. Control of Meloidogyne sp. by P. penetrans was reported from greenhouse tests (Brown and Nordmeyer, 1985; Channer and Gowen, 1988; Maheswari and Mani, 1988, Maheswari et al., 1987; 1988; Raj et al., 1988; Raj and Mani, 1988; Tzortzakakis, 1994a; Vargas et al., 1992), microplot studies (Brown et al., 1985; Chen et al., 1996b; 1997c; Daudi, 1990; Dube and Smart, 1987; Oostendorp et al., 1991a), and small-scale field experiments (Channer and Gowen, 1988; Stirling, 1984). Several investigators have provided estimates of the endospore population density needed in the soil to provide nematode control (Brown and Smart, 1985; Chen et al., 1996b; Davies et al., 1989; 1990; Sell and Hansen, 1987; Stirling, 1984). A concentration of 10^ endospores/g of soil prevented J2 of Meloidogyne spp. from infecting plant roots when the nematode moved either 4 or 8 cm through the soil (Stirling et al., 1990). Nematodes that moved 2 cm through the soil were infective but they did not produce progeny because of infection by the bacterial parasite. Concentrations as low as lO'*

PAGE 38

23 endospores/g of soil reduced root-knot nematode fecundity (Ahmed and Gowen, 1991; Gowen et al., 1989), and caused suppression of M. arenaria infection on peanut (Chen et al, 1996b). Densities of 10^ endospores/g of soil throughout the top 15 cm provides levels of control comparable to a nematicide application (Stirling, 1991). Most recently, Chen et al. (l-996b) demonstrated that adding 10,000 to 100,000 endospores per g of soil to rootknot nematode infested soil provided control of M. arenaria on peanut in the first season. Application of P. penetrans endospores 2.5 cm deep in soil appeared to be more effective for parasitism of M. incognita on tomato than a surface application or an application 5 cm deep (Ahmad et al., 1994). Integrated nematode management There have been several attempts to integrate P. penetrans with other nematode management techniques. Synergistic effects on the management of Meloidogyne spp. were reported from greenhouse experiments in which P. penetrans was combined with organophosphate or organocarbamate nematicides at nematostatic field application rates (Maheswari et al., 1987; Tzortzakakis, 1994a) or at subnematostatic rates as low as 1.5 ppm and 0.25 ppm, respectively (Brown and Nordmeyer, 1985). Successful control of Meloidogyne spp. also may be obtained by combining P. penetrans with one or several other biological control strategies. De Leij et al. (1992) noted that the egg mass-colonizing fungal endoparasite Verticillium chlamydosporium Goddard complimented the suppression of M. incognita by P. penetrans by attacking a different life stage of the nematode, thus giving better results than either of the organisms alone or the nematicide treatment. Similar results were obtained in experiments where Paecilomyces lilacinus (Dube and Smart, 1987; Maheswari and Mani, 1988; Shahzad et al., 1990; Zaki and Maqbool, 1991),

PAGE 39

24 Talaromyces flavus, or Bacillus subtilis (Zaki, 1991) were added as additional nematode antagonists, or where oil cakes were added as a soil amendment (Maheswari et al., 1988). The efficacy of P. penetrans was enhanced with soil solarization applied alone (Freitas, 1997; Walker and Wachtel, 1989) and in combination with an organophosphate nematicide (Tzortzakakis, 1994a; Walker and Wachtel, 1989). The population density of P. penetrans also was affected by crop rotations and winter cover crops. In the presence of a susceptible host plant, the density of P. penetrans endospores build up in the presence of an increasing Meloidogyne spp. population (Brown et al., 1985; Chen et al., 1994; Madulu et al., 1994; Oostendorp et al., 1991a). A non-host, resistant host, or fallow rotation has been shown to hamper the increase in endospore density (Chen et al., 1994; Madulu et al., 1994; Oostendorp et al., 1991a). Summer crops were generally more effective than winter cover crops, which is probably explained by the high temperature dependence of both the root-knot nematode and its bacterial parasite. Useful Methods and Techniques for Studving Pasteuria spp. Detection of Pasteuria spp. in soils and in nematodes Pasteuria spp. can be detected readily on or in nematodes using a number of simple techniques. Endospore-encumbered or endospore-filled migratory nematode stages have to be extracted from the soil. One method for their extraction is to use a centrifugalflotation technique (Jenkins, 1964). Increasing the specific gravity of the sucrose solution used in the centrifugal-flotation technique from 1.14 to 1.22 and 1.26 led to recovery of a higher number of spore-filled bodies of Pratylencus scribneri and B. longicaudatus, respectively (Oostendorp et al., 1991b). Extraction efficacy of endospore-filled Hoplolaimus galeatus (Cobb) Thome was not increased with the denser sucrose solutions.

PAGE 40

25 Nematodes may be examined for adhering or internal endospores with the aid of an inverted microscope at X400 magnification. For easier detection, endospores adhering to the body of a nematode may be stained with Brilhant Blue G (Bird, 1988), or labeled with fluorescein (Chamecki, 1997; Chamecki et al., 1996). Detection of endospores in soil can be increased by adding host nematodes to a soil sample and storing the sample for several days to allow the nematodes to migrate through the soil, or by adding the nematodes and water to the soil and shaking the contents for 24 hours before extraction (Stirling and White, 1982). The attachment of endospores to the nematode body does not indicate infection; evidence of intemal parasitism is needed to confirm that the nematode is actually a host. The parasite's presence can be confirmed by squashing females on a glass slide and observing them for the presence of vegetative stages or endospores of the parasite (Hatz and Dickson, 1992). For better recognition of the different developmental stages of the bacterium, specimens can be heat fixed and Gram stained (Cappuccino and Sherman, 1986). A simpler technique by Serracin et al. (1997) used lactophenol and 1% methyl blue stain to dye the bacterial structures inside the nematode body. However, its application did not successfully stain bacterial structures in vermiform J2 of Meloidogyne sp. Isolation of Pasteuria spp. from soils and nematodes With the exception of the soil-inhabiting mature endospore, the bacterium's other Ufe stages can only be found inside nematode bodies. In sedentary nematodes, such as Meloidogyne spp., Tylenchulus semipenetrans, Heterodera glycines, H. goettingiana, and Globodera rostochiensis, early life stages of Pasteuria spp. can be found in sedentary juvenile stages and in the adult female (Sayre et al., 1991a; Sturhan et al., 1994; Oostendorp, personal communication). These nematode life stages can be dissected from enzymatically digested plant tissue (Hussey, 1971). Endospore-filled females of

PAGE 41

26 Meloidogyne spp. are usually white, opaque and larger than uninfected females (Hatz and Dickson, 1992). The infected females can be distinguished visually and collected by hand with the use of a stereomicroscope. Migratory life stages of sedentary nematodes and ectoparasitic nematodes parasitized with Pasteuria spp. must be hand-picked from nematode suspensions after they are extracted from soil. By surface sterilizing infected nematodes and squashing them in sterilized water, different life stages of Pasteuria spp. can be observed free of contaminating microorganisms (WilUams et al., 1989). Quantification of endospores in soils, root powder, or suspensions The quantification of the bacterial endospores is important, because the efficacy of Pasteuria spp. is density dependent (Stiriing, 1991). However, methods for enumeration of Pasteuria spp. in soils have received little attention, and endospore densities in the soil are usually based on the number of endospores attached to soil inhibiting nematode stages. Time and effort to determine the number of endospores of P. penetrans attached to J2 of Meloidogyne spp. may be reduced considerably by using tally thresholds (Chen and Dickson, 1997). Juveniles of M. javanica were used as probes for P. penetrans (Stirling, 1984; Stirling and White, 1982), and since there is a direct relationship between endospore concentration in soil and the number of endospores attached (Stirling et al, 1990), a bioassay was developed to estimate the relative endospore density in soil (Oostendorp et al., 1991a). By using a particular probe nematode in the bioassay, only that component of the P. penetrans population that attaches to that nematode is detected. Endospores also have been extracted from soil by differential-centrifugation (Davies et al., 1990); however, the technique is tedious and difficult to use. The total number of endospores in a particular soil may not accurately represent the suppressivness

PAGE 42

27 of the soil; thus according to Stirling (1991), a bioassay is a more useful quantification method than the use of a direct extraction method. Chen et al. (1996a) observed that the highest estimate of the number of endospores in root material was obtained by suspending machine ground root powder in deionized water and pouring the material onto a 250-|j.m-pore sieve. Endospores passed through the sieve and were concentrated in water, and the number of endospores was calculated with the help of a hemacytometer. This refined quantification allowed an improvement compared to the previous application of the inoculum by weight (Brown et al., 1985; Dube and Smart, 1987; Raj and Mani, 1988; Stirling, 1984; Zaki and Maqbool, 1991; 1992a; 1992b). Culture and preservation of Pasteuria penetrans Methods have been developed for the production of P. penetrans endospores in females of Meloidogyne spp. parasitizing plant roots. This is currently necessary because in vitro cultivation has not been successful (Bishop and Ellar, 1991; Previc and Cox, 1993; Reise et al., 1988). Pasteuria penetrans-encumhcrcd J2 of Meloidogyne sp. must be inoculated onto a susceptible host, such as tomato (Stirling and Wachtel, 1980). Once the bacterium has developed to maturity, the root systems may be harvested, washed, air dried, and ground into a fine endospore-laden powder. Although tomato plants are most commonly used, cucumber, Cucurbita vulgaris L., was shown to yield higher endospore numbers than tomato (Cho et al., 1997). This system may produce a highly variable yield of endospores; however, it is superior to the use of endospore infested soil (Dutky and Sayre, 1978; Mankau and Prassad, 1977). The plant system has been optimized (Sharma and Sliding, 1991), and most recently a hydroponic cultivation system has been reported (Serracin-Ulate, 1995). Verdejo and Jaffee (1988) used a gnotobiotic technique for

PAGE 43

28 producing endospores; however, this method requires rigorous aseptic conditions and is therefore not a good procedure for general use. The time period required for P. penetrans to complete its life cycle has been shown to vary with different isolates (Hatz and Dickson, 1993; Serracin et al., 1997; Stirling, 198 1-). Thus, a random determination of the progress of the bacterium's development in a cultivation system can help avoid harvesting prematurely, thus improving the efficiency of the endospore yield. The lack of an efficient technology for the mass-cultivation of Pasteuria spp. endospores is the greatest impediment to the use of this microbe as a biological control agent (Dickson et al., 1994; Stirling, 1991). Attachment Endospore-host attachment studies are the first step in establishing the host specificity of Pasteuria spp. and determining their biological efficacies. Most of the techniques rely on nematode movement through soil (Brown and Smart, 1985), water (Channer and Gowen, 1988), or agar (Verdejo and Jaffee, 1988), when each is laden with endospores, or the agitation of a nematode-endospore-water suspension (Ahmed et al., 1990; Bird, 1986; Bird et al., 1990; Davies et al., 1988; Oostendorp et al., 1990). Hewlett and Dickson (1993) consistently achieved attachment of P. penetrans to Meloidogyne spp., H. galeatus, and B. longicaudatus using a centrifuge technique. The method is fast, and allows studies of Pasteuria spp. collected from nematode species that yield relatively few endospores per cadaver.

PAGE 44

29 Objectives The objectives of this dissertation were 1 ) to monitor the population densities of Meloidogyne spp. and P. penetrans in microplots and in a naturally infested tobacco field; 2) todevelop a test for soil suppressiveness that allowed the determination of the role of P. penetrans in suppressing Meloidogyne spp. populations in both microplots and tobacco field plots; and 3) to evaluate the effect of cultural practices, such as an intercropping system with com and beans in rotation with peanut, autumn cover crops, resistance, and fertilizer regimes on the population development of both M. arenaria and P. penetrans.

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CHAPTER 2 POPULATION DEVELOPMENT OF MELOIDOGYNE ARENARIA RACE 1 AND PASTEURIA PENETRANS IN A 6.5-YEAR MICROPLOT STUDY Introduction The peanut root-knot nematode, Meloidogyne arenaria (Neal) Chitwood race I, is one of the most important soil pathogens of commercially grown peanut {Arachis hypogaea L.) (Minton, 1984). It occurs on peanut in many countries and is especially troublesome in the southeastern United States (Minton, 1984; Porter et al., 1984). Continuing environmental problems associated with the use of nematicides (Thomason, 1987) has resulted in more scientists studying nematode management strategies alternative to chemical control (Kerry, 1990). The use of microbial agents for biological control of plant-parasitic nematodes is an altemative management tactic that is receiving increased interest among nematologists. Among many organisms identified as antagonists of plantparasitic nematodes, the endospore forming bacterium, Pasteuria penetrans, has been demonstrated to have potential for the control of mainly Meloidogyne spp. It has been suggested that P. penetrans may suppress nematode population densities below economic damage levels (Chen et al., 1996b; Dickson et al., 1994; Minton and Sayre, 1989; Stirling, 1991). In order to determine the efficacy of the density-dependent obligate parasite, it is important to understand its development as influenced by its nematode host. The population densities of P. penetrans and of M. arenaria race 1 with and without the nematode antagonist were determined from the spring of 1987 to the fall of 30

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31 1989 (Oostendorp et al., 1991a). During this period, soil suppressive to M. arenaria was produced in microplots. The microplots had been planted continuously to peanut in summer and either vetch {Vicia villosa Roth) or wheat (Triticum avenae L. cultivar 302), rye {Secale cerale L. cultivar Wrens Abruzzi), or bare fallowed during the winter. Pasteuria penetrans, initially applied to the soil in relatively low numbers, was increased to numbers which suppressed M. arenaria. Within 3 years, yield of plots infected with P. penetrans and M. arenaria increased to a level comparable to the untreated control (Oostendorp et al., 1991a). The objective of this study was to continue to monitor the population densities of both M. arenaria race 1 and P. penetrans in these microplots in order to determine their long-term effects on M. arenaria densities. Materials and Methods In 1987, 90 microplots (76 cm in diameter), located at the University of Florida, Green Acres Agronomy Farm, Alachua County, were initially arranged in 10 rows of nine plots each, with a distance of 1.5 m between plots, in a loamy, siliceous, hypothermic Grossarenic Paleudults with 90% sand, 4% silt, 6% clay, and 1.8% organic matter (Oostendorp et al., 1991a). A split-plot design was used with main plots consisting of three soil treatments: an untreated control, M. arenaria alone (RKN), and M. arenaria plus P. penetrans (RKN + Pp). Three summerwinter cover crop rotations of peanut-rye, peanutvetch through 1991, with peanutwheat thereafter, and peanut-bare fallow were subplots. Wheat replaced vetch because the incidence of soilbome diseases on peanut increased following vetch. Each combination of soil treatments and winter cover crops was replicated 10 times. The microplots were initiated by placing 800 cm^ of soil preparation containing uninfected tomato roots, tomato roots infected with M. arenaria

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32 race 1, or with M. arenaria race 1 plus P. penetrans in a hole in the center of each microplot receiving the control, RKN, or RKN + Pp treatment, respectively. Monitoring of the bacterium and nematode population densities for this dissertation began in the spring of 1990 and continued through the fall of 1993. Data from the period 1987 to 1989 are included herein to provide a complete data set on the long-term effects of P. penetrans. Two months before planting in the spring of 1991 and 1993, uninoculated control plots were treated with 977 kg methyl bromide/hectare (98% methyl bromide plus 2% chloropicrin), applied broadcast under a 3-mm polyethylene plastic covering. On 6 May 1990, 5 May 1991, 13 May 1992, and 7 May 1993, three pairs of peanut seeds of cv. Florunner were planted 4-cm deep in an equally spaced pattern in each plot. After emergence, one of each pair of seedlings was removed. A 90-cm-high, wire-mesh fence was placed around each plot to confine the growth of the peanut foliage inside the microplot. Plots were weeded manually. Through 1991, plots were irrigated by overhead sprinklers. Thereafter, a microjet sprinkler system was installed, delivering an amount of water equivalent to 4 mm per day. Every 10 to 14 days, insects and foliar disease pathogens were controlled with esfenvalerate, chloropyrifos, insecticidal soap, chlorothalonil, or liquid sulfur. In mid-November each year, after harvest of peanut, rye and vetch or wheat were broadcast seeded as winter cover crops, or plots were bare fallowed through the winter. Vetch, wheat, and peanut are hosts for M. arenaria race 1 , but rye is a poor host. The soil in each plot was turned using a spade, and leveled between crops. At harvest (136 to 140 days after planting), peanut plant shoots were evaluated in their appearance, and the root systems, as well as the pods and pegs, were rated for galling. A good stand of plants with a densely closed canopy was rated as 0, and a poor stand of plants with sparse canopy was rated as 5. Galling was rated according to the following

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33 scale: 0 = no galls, 1 = 1-10,2= 1 1-20, 3 = 21-55, 4 = 56-80, and 5 = 81-100% of roots or pods and pegs galled (Barker et al., 1986). Three to five days after the plants were lifted out of the ground, all pods were removed and placed into paper bags, dried at 60 °C until the moisture content was reduced to about 10%, and weighed to determine yield. The number of second-stage juveniles (J2) of M. arenaria /lOO cm"* of soil was estimated at harvest of peanut and each of the winter cover crops. Soil samples consisting of a composite of five, 2.5-cm-diameter cores per plot were taken from the top 20 cm of each microplot with the use of a cone-shaped auger. The soil was mixed and processed with a centrifugal-flotation method (Jenkins, 1964). The numbers of J2, the rate of attachment of P. penetrans, and the number of endospores attached per 20 randomly selected J2 was determined using an inverted microscope at X400 magnification. Data were subjected to analysis of variance (ANOVA). If the attained probability level of the main plot (treatments) or the subplot (cover crop) effects was significant at P < 0.05, the means were separated by Duncan's multiple-range test. If a significant main plot X subplot interaction (P < 0.05) was observed, main plot means for each subplot and subplot means for each main plot were separated by Duncan's multiple-range test. If an insufficient number of J2 or no J2 were extracted from soil of RKN + Pp plots, replicates without J2 were disregarded for the calculation of rate of attachment with P. penetrans and the number of endospores per juvenile, and data were presented in a seperate table . Results After the initial 3-year period of the experiment (1987 to 1989), the number of J2 in the soil of RKN + Pp plots decreased, and remained lower than those in the RKN plots (P < 0.05) (Fig. 2-1, Tables 2-1 to 2-3). The nematode population density in microplots

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34 Fig. 2-1. Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100 cm^ of soil and the number of endospores attached per juvenile extracted from peanut microplots infested with M. arenaria (RKN), and M. arenaria plus P. penetrans (RKN + Pp). Analysis was based on means of 10 replicates, with 20 or fewer J2 observed per replication to determine the number of endospores per juvenile. Means labeled with a different letter are different according to Duncan's multiple-range test {P < 0.05); small letters were used for comparisons of juvenile numbers, capital letters for comparing endospore numbers. Data from 1987 to 1989, were from Oostendorp et al. (1991a).

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35 T3 X 60 c a n tN NO O O NO O 00 -i^ ^ NO — r~— NO o. 00 00 O X U Oi. c u E + O tN u u > tN NO c o + 1^ c o U Qi ai u ctJ ca > o U u at (J u > o — o. ac o ON o + Z 1) p i d r VI o a. O BO ca t3 c 6 E ca 3 ^ Q ^ 2 § s o c ca u D. P u > 3 U 60 C3 T3 C o o u c/) o tN o D. on O T3 C u ca X o d VI c o a. o >» o. o c ca u CA u. ea O. — "a C &o _C3 q. po O o. T3 O an C/3 O o u O >4-> o o o. II >rl 8 •o 1 c ca 00 >i II do c •T3 ca e o ca T3 U d' 00 _o \o >o ca II ea "a. 13 c ca I tN II o" tN V c r: tS o o 00 II o > o u ea o c E o c ea o X s c ea ca u u c C3 c ca tN ca 00 o c ca O es o O. 00 1= tN O tN C ea o -S •o — ~ o 60 ea u

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36 X u o c at c o B. O . a c 5 x: u U O o o Xi — o U X "ca oq o CO S 2 m o ON o O ON TJ— ON ON — NO O — NO 00 Tj— o O X X O ca OO ca U O X ca O NO c o u ca o c 00 6 2 IT) 00 O ca — u = 1 2 "S 60 S .E O "O o CJ o ca T3 O o ca o c u > 3 •™» o 60 ca o u o o D. — o a o c ca o ca o. X CL o CO 1= ca o a c ca o o Q. c ca >. c o c ca CJ •T3 U _o O ca c ca iS O. o C ca -o o o 60 ID O c ca E c ca _o H cn O a. o o o o c ca o* 00 I NO >n II o (N CNl o' ca o O = u Q. •-1 O Ol c ca X ain _jj II E ca o cn cn 60 "3 C 60 ites a O c II o >< u 60 c ca a

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37 (3 a o o. o T3 C W -a 22 D. CO o c/3 u o o o -vo £3 (si PL, C3 d CS O £1 O as d d XT d d d d d d rd CS CS o\ rr(N CS CN' CN CS CN CS r-; o oo CS CS CS — 00 O CTv O (S — ' O — — CN O ti, C^l CS CS CS CS CS rs 00 o o ON r<-i CO CN — OO ON X) o o. a. — CS XI o XI o CS CS (N O o o — O a x3 (N — OO O h b X2 CN X! O O. X) o XJ o 00 — CS 00 XJ o Q. Oh O O X3 o o 00 o XI NO XI o a. CU c o U c E 02 ei2 D. p > o U c o U >> 02 Di c o U at x: c o U Z 2 2 ^ 02 02 o c E 3 O c d •5 VI U " (U O 00 £ i eg t. "5 E o c 3 X2 o 2 U 00 If o c a 03 w S J2 u o c C3 u c c C3 1) c 'c3 E p O o o c CL c > o 00 -o c o u o " c o E C3 o D. 0) O c o x: >^ Q. O C (J O. c O. o 00 «) O O. X •= T3 O C c/) a CO O e poo of II o V) o •a 1 c a 00 >^ II do >n c •o CS c u CS a u 00 _o vo C3 II Tt <0 "O 1 am CS a. II m O T3 d^ c CS 3 1 •o II o o CNl 00 d II u o 1 "d. tl 2i seal c/T per 00 c gall J2 rati no o CS u tJ II an c o CS >< y. Vi cS •a u c of o. 00 c P c CS CS « u CL, o

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38 inoculated with P. penetrans was nearly undetectable in soil samples from 1991 to 1993. The number of P. penetrans endospores adhering to J2 increased to a high of 17.2 by 1989 and remained relatively consistent during the remaining 4-year period (Oostendorp et al., 1991a; Fig. 2-1; Tables 2-1 to 2-3). Through this time, 10-66% of the J2 were encumbered with endospores. However, when replicates without J2 were excluded from analysis for RKN + Pp plots, the incidence of endospores per juvenile and attachment to J2 were much higher than those for 20 J2 (Table 2-4). In the fall of 1990, the endospore density of P. penetrans in RKN + Pp plots had amplified to yield an average of 19.5 endospores/juvenile (data not shown). Between 1991 and 1993, the endospore density remained at a level at which 95% to 100% of the recoverable J2 were encumbered with mean numbers of 3 to 72 endospores (Table 2-4). In the soil of RKN plots, the number of J2 remained high throughout the experiment (Oostendorp et al., 1991a; Fig. 2-1, Tables 2-1 to 2-3). In the third year (1989), 2 of the 30 replicates of the RKN plots were contaminated with the bacterial parasite (Oostendorp et al., 1991a), and P. penetrans endospores were found to have contaminated 16, 29, and all 30 RKN plots in the falls of 1991, 1992, and 1993, respectively (Figs. 2-2 to 2-4, Tables 2-1 to 2-3). In fall of 1993, endospore population densities of RKN plots had increased to averaged 91% attachment rate with 34 endospores/juvenile (Table 2-3). Numbers of nematodes and endospore attachment levels generally did not vary with cover crops (P < 0.05) (Fig. 2-5 , Tables 21 to 2-3). However, in the spring of 1989, the nematode population densities in RKN plots under vetch slightly exceeded those of other plots (Fig. 2-5 A). Second-stage juvenile numbers in vetch and bare fallowed plots exceeded those in the rye plots in the spring of 1990. However, in the spring of 1992 and spring of 1993, lower numbers of J2 were recovered from plots under rye and wheat than

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39 Table 2-4. The percentage of second-stage juveniles (J2) of Meloidogyne arenaria infected with Pasteuria penetrans and the average number of endospores per juvenile in peanut microplots infested with M. arenaria and P. penetrans in the spring of 1987, and rotated with rye, vetch or wheat, and bare fallow as winter cropping sequence. % J2 with % J2 with endospores endospores Cover crop attached Endospores/juvenile attached Endospores/juvenile Fall 1991 Rye 100 (5) 10 (5) Vetch (0) (0) Fallow 100(18) 40.2(18) Spring 1992 Fall 1992 Rye 100(12) 17.8(12) 66.7(17) 32.4(17) Wheat 100(19) 35.3 (19) 66.7(5) 8.5(5) Fallow 95(16) 21.6(16) 100(8) 44.1 (8) Spring 1993 Fall 1993 Rye 100(1) 6(1) 100(16) 43(16) Wheat 100(1) 42(1) 100(9) 72(9) Fallow 100(1) 3(1) 80(62) 20(62) Means of varying number of rephcates. Numbers in parenthesis are the number of nematodes observed.

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40 Replicate 1 2 3 4 5 6 cv CR CF NV NR PV PR PF NR NF NV PR PF PV CR CF CV PR PF PV CR CF CV NR NF NV CR CF CV NR NF NV PR PF PV NF NV NR PF PV PR CF CV CR PF PV PR CF CV CR NF Inv NR 7 CF CV CR NF NV NR PF PV PR o s NV NR NF PV PR PF CV CR CF 9 PV PR PF CV CR CF NV NR 10 CV CR CF NVNR NF PV PR 1 PF 1 ( ( 3% juveniles with jndospores attached ^: a -99% juveniles vith endospores ttached j^H 1 ^^^^1 e 00% juveniles with ndospores attached Fig. 2-2. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1991. No second-stage juveniles were extracted from non-shaded plots. The first letter indicates the pre-experimental treatment (C = plots free of nematodes and P. penetrans, N = plots inoculated with Meloidogyne arenaria, and P = plots inoculated with M. arenaria plus P. penetrans), and the second letter indicates the winter cover crops (V = vetch, R = rye, and F = fallow).

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41 Repli cate 1 CW CR CF NW NR NF PW NR NF. NW PF CR CF CW PR PF CR CF CW NR NF CR CF CW NR NF NW PR PF PW NF NW NR PF PW CF CW CR PF PW PR CF CW CR NF NW CF CW CR NF NW NR PF PW PR NW NR NF PW PR CW CR CF PW PR PF CW CR CF NW NR NF 10 CW CR CF NW NR NF PW PR 0% juveniles with endospores attached 1-99% juveniles with endospores attached 100% juveniles with endospores attached Fig. 2-3. Microplot treatment plan indicating the presence of Pasteuria penetrans endospores as determined by soil sample analysis in the fall of 1992. No second-stage juveniles (J2) were extracted from non-shaded plots. The first letter indicates the pre-experimental treatment (C = plots free of nematodes and P. penetrans, N = plots inoculated with Meloidogyne arenaria, and P = plots inoculated with M. arenaria plus P. penetrans), and the second letter indicates the winter cover crops (W = wheat, R = rye, and F = fallow).

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42 Replicate 0 % juveniles with endospores attached 1-99% juveniles with endospores attached 100% juveniles with endospores attached Fig. 2-4. Microplot treatment plan indicating the presence of Pasteiiria penetrans endospores as determined by soil sample analysis in the fall of 1993. No second-stage juveniles (J2) were extracted from non-shaded plots. The first letter indicates the pre-experimental treatment (C = plots free of nematodes and P. penetrans, N = plots inoculated with Meloidogyne arenaria, and P = plots inoculated with M. arenaria plus P. penetrans), and the second letter indicates the winter cover crops (W = wheat, R = rye, and F = fallow).

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43 Fig. 2-5, Number of second-stage juveniles (J2) of Meloidogyne arenaria race 1 in 100 cm' of soil extracted from soil in microplots with A) M. arenaria, and B) M. arenaria plus P. penetrans Plots were planted to peanut in summer and bare fallowed, or planted to rye, or vetch (1987-1990) and wheat (1991-1992) in winter. Means of the cropping sequences at individual sampling dates labeled by no or the same letter are not different according to Duncan's multiple-range test (P < 0 05)

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44 rye and wheat than from those under bare fallow {P < 0.05). In the fall of 1991, in RKN plots the estimated endospore population density in vetch-wheat plots was highest, and, together with rye plots, exceeded the endospore counts of fallow plots in the fall of 1993 (P < 0.05) (Fig. 2-6 A). From the spring of 1989 till the spring of 1990, the attachment level and the number of endospores per juvenile in RKN + Pp plots under vetch were higher than under bare fallow (P < 0.05) (Fig. 2-6 B). Thereafter, the endospore counts in vetchwheat were frequently lower than in bare fallow plots. In the fall of 1993, the vetchwheat plots yielded the highest estimated endospore population density. In the fall of 1990 and of 1992, the endospore counts in rye plots exceeded those of bare fallow plots and vetchwheat plots, respectively. According to main effect means in the spring and the fall of 1993, the attachment rates of 3 and 25 endospores per juvenile across all nematode treatments in wheat plots were higher (P < 0.05) than the levels observed in rye or fallowed plots (Table 2-3). The highest number of 107 J2/100 cm^ of soil in fallowed RKN plots observed in the spring of 1993 was numerically greatest, but yet consistent with the main effect of highest juvenile levels in RKN plots. Following the initial increase in peanut yield in RKN + Pp plots after 3 years, the yields remained high and not different from those in control plots for the remaining 4 years (P < 0.05) (Fig. 2-7, Tables 2-1 to 2-3). In 1993, there were no differences among yields in any treatments, which is attributed to P. penetrans infesting RKN plots (Fig. 2-6 A, Table 2-3). Meloidogyne arenaria was low or undetectable in control plots throughout the experiment. Throughout the last 4 years, the galling indices for roots and pods in RKN plots were higher than gall ratings of RKN + Pp and control plots {P < 0.05) (Tables 2-1 to 2-3).

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45 Sampling date CM OJ en CD Oi ai 05 OJ Ol CD O) "(^ U_ liQ. Q. CO CO Fig. 2-6. Number of attached endospores per juvenile extracted from soil in microplots with A) M. arenaria, and B) M. arenaria plus P. penetrans. Plots were planted to peanut in summer and bare fallowed, or planted to rye, or vetch (1987-1990) and wheat (1991-1992) in winter. Means of the croppmg sequences at mdividual sampling dates labeled by no or the same letter are not different accordmg to Duncan's multiple-range test {P < 0.05). Means of 10 replicates, with 20 or fewer J2 observed per replication.

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46 gram s/microp lot 1000 number of endospores 100 10 0.1 100 1987 1988 1989 1990 1991 1992 1993 Cropping season Fig. 2-7. Log presentation of peanut yield (grams per microplot) from plots treated witli Meloidogyne arenaria race 1 (RKN), and M. arenaria plus Pasteuria penetrans (RKN + Pp). and untreated plots; and the number of endospores per second-stage juvenile (J2) in RKN and RKN + Pp plots Data from 1987 to 1989 were from Oostendorp et al., 1991a. Means of 10 replicates, with 20 or fewer juveniles observed per replication to determine the number of endospores per J2. Means with the same letter are not significantly different according to Duncan's multiple-range test (P < 0,05). Bar graphs peanut yields in grams; line graphs number endospores per juvenile.

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47 Throughout the last 3 years of the experiment ( 1991 to 1993), the population density development of Criconemella spp. was favored in plots that were fallowed through the winter {P < 0.05) (Appendix). An interaction between treatment and cover crop showed a higher number of ring nematodes in fallowed RKN plots than in fallowed RKN + Pp plots. In the fall of 1991 the highest number of ring nematodes in the soil coincided with the highest number of endospores of P. penetrans attached per juvenile of Meloidogyne arenaria. Discussion Populations of P. penetrans, as estimated by endospore attachment to juveniles, increased over 6.5 years from relatively low levels of endospores added initially or introduced by contamination during the course of the experiment to levels after 3 years that were suppressive to the nematode population. This confirms the work by Oostendorp et al. (1991a) and Chen et al. (1997c), whereby P. penetrans endospores increased to levels that suppressed root-knot nematodes when introduced at low numbers (approximately 1,000 spores/g of soil). Within four cropping seasons, nematode attachment rates in plots with endospores added increased to nearly 100%, with an average of about 20 endospores per juvenile, and peanut yields reached levels equivalent to those in the nematode-free control. The population density of M. arenaria in RKN + Pp plots remained nearly undetectable throughout the last 3 years of this 6.5-year experiment. It was difficult to attain an estimate of endospores per juvenile over the course of the experiment because the number of J2 in many RKN + Pp plots dropped below 20, which was the number used for estimating the endospore density. This was illustrated in the fall of 1991, when a decrease

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48 in the number of endospores per juvenile and a decrease in the percentage of J2 with attached endospores were observed in rye and vetch plots. Individual observations revealed a further increase in the endospore population density under either cropping sequence. In 1992 and 1993, the attachment rates of juveniles with P. penetrans were around 100%, and the number of endospores per juvenile frequently exceeded 25. At this rate, root penetration has been shown to be prevented (Davies et al., 1988; Stirling, 1984; Stirling et al., 1990). Nematode suppression for three consecutive years did not affect the ability of P. penetrans endospores to attach to their nematode hosts. However, the experimental design did not allow for the determination of the number of J2 that may have inadvertently entered the microplots each season. Although it has been reported that the suppressiveness of root-knot nematode by P. penetrans is dependant on the density of endospores in the soil (Stirling, 1991), it appears in the present study that P. penetrans increased to a suppressive level after 3 years and remained sufficiently high to prevent further increases in the RKN densities. Pasteuria penetrans remained at a highly suppressive level for 3.5 years following the initial increase to suppressive levels in 3 years. The nematode population density required to maintain effective densities of P. penetrans to continue nematode suppression remains unknown. In 1991 and 1992, the residual nematode population in RKN plots reduced peanut yields and affected the plant performance. However, in 1993, plant shoots and yields appeared to have been affected by leaf chlorosis and reduced fruiting, apparently as a result of the symptomatic manganese deficiency (Dickson, personal communication; Porter et al., 1984), rather than by nematode infection. There was only slight galling of roots, pods, and pegs by root-knot nematodes in the RKN plots in 1993, which indicates that nematode reproduction was becoming suppressed, and that the P. penetrans population density in the soil was continuing to increase to suppressive levels.

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49 With no P. penetrans to interfere, the nematode population density peaked when peanut was followed by a susceptible crop. Those conditions favored the contamination by and the build-up of endospores of P. penetrans. This observation agrees with those of other scientists (Chen et al., 1994; Madulu et al., 1994; Oostendorp et al., 1991a), in which the continuous availability of a host plant favored the amplification of the bacterial parasite in its nematode host. However, since the nematode population density in the presence of P. penetrans did not differ among the cropping sequences, it is suggested that nematodes in vetch plots were already partially suppressed in years 2 and 3 of the experiment. Because of the density dependent nature of P. penetrans (Bird and Brisbane, 1988; Chen et al., 1996b; Dickson et al, 1994; Minton and Sayre, 1989; Stirling, 1984), the endospore population development is expected to be influenced by the population density of its nematode host (Siding, 1991). r The winter cover crop had a consistent effect on the population density of Criconemella spp. Fallowing the peanut microplois increased the number of ring nematodes more than growing rye and wheat or vetch for winter cover. Criconeinalla sp. was reported as a host for Pasteuria spp. (Sturhan, 1985; 1988). However, endospores of isolates of Pasteuria spp. from Criconemella spp.were smaller and are believed to be a different species (Hewlett, personal communication); hence, they do not likely contribute to the suppression of M. arenaria in the peanut microplots. For a period of 3 years (1991 to 1993), P. penetrans protected peanut from rootknot damage. The presence of a large population of M. arenaria in the RKN plots favored a continuous endospore population increase until the nematode population was detrimentally affected after a period of 4 years (1990 to 1993), which resulted from inadvertent contamination by P. penetrans endospores.

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50 The build-up of the endospore density in the soil can be enhanced by supplying the nematode population with a good host plant throughout the year. Thus, although there is still no large-scale production of P. penetrans inoculum in sight, the use of this nematode antagonist in integrated nematode management programs might be feasible. There is a need for further experiments to determine whether the nematode suppression can be achieved in a shorter time span, and whether it can be maintained beyond a period of 3 years.

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CHAPTER 3 USE OF MICROWAVE HEATING IN EVALUATION OF A MELOIDOGYNE A/?£:A^Ai?M-SUPPRESSIVE SOIL CONTAINING PASTEURIA PENETRANS AND ITS APPLICATION IN A SUPPRESSIVE-SOIL TEST Introduction Pathogen-suppressive soils may be defined as soils in which the pathogen does not establish or persist, establishes but causes little or no damage, or establishes and causes disease (conducive soil), but thereafter the disease is reduced even though the pathogen persists in the soil (Baker and Cook, 1982). Suppressiveness can develop as a result of the buildup of antagonists in response to a high pathogen population (Baker and Cook, 1982), especially in situations where susceptible crops are grown in succession. The endospore-forming, obligate bacterial parasite of root-knot nematodes, Pasteuria penetrans (Thome) Sayre & Starr, is widely distributed in agricultural soils throughout the world (Chapter 1) and has been shown to be effective in controlling rootknot nematodes, Meloidogyne spp., in field or microplot studies (Brown and Smart, 1985; Chen et al., 1994;1996b; 1997c; Dickson et al., 1994; Minton and Sayre, 1989; Oostendorp et al., 1991a). In most cases, the suppression of Meloidogyne spp. by P. penetrans occurred after long-term monoculture of a susceptible host (Bird and Brisbane, 1988; Chen et al., 1994; Mankau, 1980a; Minton and Sayre, 1989; Weibelzahl-Fulton et al., 1996). Many physical and chemical means have been developed to eliminate or reduce populations of soilbome plant pathogens for phytopathological studies (Mulder, 1979). Nematode suppressiveness of soil has been evaluated previously using soil applications 51

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52 of fungicides, such as captafol (Crump, 1987), mancozeb and iprodione (Mertens and Stirling, 1993), benomyl or tachigaren (Qadri and Saleh, 1990), and 40% formaldehyde (Dickson et al., 1994; Kerry et al., 1982; Qadri and Saleh, 1990). Suppression of fungal populations by fungicides is not consistent, thus the interpretation of results can be problematic. Microwave heating of soil for selective periods of time can be useful to kill or reduce populations of soilbome fungal propagules, yet have little effect on populations of prokaryotic organisms (Chen et al., 1995; Ferriss, 1984). The effect of the microwave radiation treatment on microorganisms increases with increasing temperature. The temperature increase depends on treatment time, and on factors such as soil water, clay, and organic matter content (Baker and Fuller, 1969; Ferriss, 1984). The objectives of this study were to determine if the microwave radiation treatment has the potential to selectively eliminate fungal antagonists from soils containing P. penetrans, and to evaluate the use of the microwave treatment in testing three mdcroplot soils for suppressiveness to M. arenaria. Materials and Methods Microwave Treatment Treatment time. On 12 January 1993, approximately 12 liters of soil, a loamy, siliceous, hypothermic Grossarenic Paleudults with 90% sand, 4% silt, 6% clay, 1.8% organic matter, 6.9% moisture, and a density of 2.78, were collected 0-20 cm deep with a bucket auger (10-cm diameter) from microplots infested with P. penetrans and Meloidogyne arenaria race 1 located at the University of Florida, Green Acres Agronomy Farm, Alachua County. Soil was selectively collected from microplots that were bare

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53 fallowed during the winter. The soil was passed through a 5-mm-aperture sieve, and mixed thoroughly. Six subsamples of 1 kg each were placed in open polypropylene bags, leveled to a thickness of approximately 5 cm, and placed centrally in the 18-liter cavity of a microwave oven (1,500 Watts, 2,450 MHz) (Tappan Appliance, Mansfield, OH). The subsamples were either microwave treated for 3, 4, 5, 6, and 8 minutes, or left untreated. The bags remained open through the heat treatment, and were closed before removal from the oven. The soil was allowed to cool to room temperature. Two hours later, a dilution plating technique was used to assay for selected fungi (Johnson and Curl, 1972). One gram of soil was dispersed in 200 ml of sterilized water and stirred constantly. One milliliter of the soil solution was plated on potato dextrose agar containing 50 mg of chlortetracycline hydrochloride, 100 mg of streptomycin sulfate, and 1 ml of Tergitol NPIO per 1 liter solution (supplements were added to agar at a temperature of < 50 °C) (Steiner and Watson, 1965). Five replicates were plated for each treatment. After 3 days of incubation at room temperature, the fungal colonies were counted. A soil bioassay with Meloidogyne arenaria race 1 as the host nematode was used to determine the presence of P. penetrans (Oostendorp et al., 1991a). Forty grams of dry soil from each treatment were placed in each of five, 10-cm-diameter petri dishes. Then, 300 second-stage juveniles (J2) were added in 1 ml of water to each dish. After 2 days of incubation at room temperature, the J2 were extracted by a centrifugal-flotation method (Jenkins, 1964), and the number of endospores per 20 randomly selected J2 per replicate was determined with the aid of an inverted microscope at X400 magnification. Soil moistur e content. On 8 March 1993, 1.5-kg lots of the dried soil collected on 12 January 1993 were placed into plastic bags, and the soil moisture content was adjusted to 1%, 3%, 5%, or 7% (weight of water per dry soil weight) by adding tap water. After 2 hours, the soil was mixed thoroughly, divided into three subsamples, placed into open

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54 polypropylene bags, and was either heated in the microwave oven for 3 or 4 minutes per kilogram of soil or left untreated. The numbers of fungal colonies per gram of soil and endospores per juvenile were determined as described above. Suppressive-Soil Test Soil treatments. On 23 April 1993 and 7 May 1993, a total of 120 liters of soil was collected from the same microplots as stated above. For the repeat test, soil was collected on 23 April and 5 June. For each experiment, a subtotal of 40 liters of soil was collected from microplots with M. arenaria inoculum alone (RKN), M. arenaria plus P. penetrans inoculum (RKN + Pp), and a control treatment without nematodes or bacteria (control) (for more information on the soil sources see Chapter 2). The soils were passed through a 5mm-aperture sieve, mixed, and divided into four subsamples of 10 liters each. The subsamples were either autoclaved, microwaved, air dried, or left untreated. For the autoclave treatment, sterilized 15-cm-diameter clay pots were filled with soil (ca. 1 liter/pot), covered with aluminum foil, and placed in an autoclave. After the first heat treatment for 1.5 hours at 55 kPa (Ferriss, 1984), the pots were removed from the autoclave. The treatment was repeated after pots and soil had cooled to room temperature (Mulder, 1979). For the microwave heating treatment, 1-kg lots of soil were placed in even layers in open polypropylene bags and heated in a microwave oven for 3 minutes (Ferriss, 1984) at full power, which caused the soil temperature to rise to about 75 °C. For the air-drying treatment, 40 liters of soil were collected from the microplots on 7 May and 5 June 1993. The soil was placed in 3to 5-cm-thick layers on plastic trays and

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55 stored in the greenhouse for 2 or 6 weeks before testing. The untreated soil was stored at room temperature, and used within 24 hours after collection. Laboratory experiment. Two hours after microwaving soil and the last autoclaving treatment, and 2 weeks after the beginning of the air-dried treatment, the number of fungal colonies per gram of soil and endospores per juvenile was determined simultaneously for all four soil treatments. Procedures were applied as described above. Greenhouse experiments. Treatments were arranged in a 3 X 4 X 2 factorial design that included the following three soil sources: soil from microplots with M. arenaria inoculum alone (RKN), M. arenaria plus P. penetrans inoculum (RKN + Pp), and a control treatment without nematodes or bacteria (control); four soil treatments (autoclaved, microwaved, air-dried, and untreated); and inoculum levels of 0 and 2,000 J2 of M. arenaria race 1. The soil from each treatment was placed into sterilized 15-cmdiameter clay pots (ca. 1 liter/pot). Treatments were replicated four and five times in experiments 1 and 2, respectively. About 100 ml and 50 ml of water were added to the airdried soil and to all other soil treatments, respectively. Each pot was covered with aluminum foil and held for 3 days in a growth room at 28 to 32 °C and 14 hours of light, or in a greenhouse for experiments 1 and 2, respectively. Meloidogyne arenaria J2 were suspended in 10 ml of water and dispensed equally to each of five holes, 5-cm deep, in the soil of each pot. Five days later, one 7-week-old peanut seedling, Arachis hypogaea L. cv. Florunner, for experiment 1, or one 5-week-old peanut seedhng of the same cultivar for experiment 2 was transplanted into each pot. A commercial Rhizobium sp. was added to each pot (McSoriey et al., 1992). Every 10 to 14 days, insects and diseases were controlled using esfenvalerate, chloropyrifos, insecticidal soap, chlorothalonil, or liquid sulfur.

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56 Experiments 1 and 2 were run for 53 and 55 days, respectively. Plant shoots were cut off at ground level and discarded. The root systems were washed free of soil, excess water was removed with paper towels, and roots were weighed and then stored in plastic bags at 4 °C. The following day, roots were stained with Phloxine B (Dickson and Strubel, 1965), and the numbers of root galls and egg masses were counted. Eggs were extracted with 1.05% sodium hypochlorite (Hussey, 1971) and their number was counted. Twenty globular mature females were picked randomly from each root system, crushed on a slide and observed for infection by P. penetrans with the aid of a compound microscope at XIOOO magnification. Nematode Origin The isolate of M arenaria race 1 used originated from peanut grown on the University of Florida, Green Acres Agronomy Research Farm, Alachua County. The species and race determinations were confirmed by examination of the perineal pattern, length of J2, and a differential host test (Hartman and Sasser, 1985). The nematode population was cultured in a greenhouse on tomato {Lycopersicon esculentum Mill. cv. Rutgers) from a single egg mass. Eggs of M. arenaria were extracted from infected tomato roots (Hussey and Barker, 1973), hatched by the Baermann method (Rodn'guezKabana and Pope, 1981) and used as 1to 4-day-old J2. Statistical Analysis The number of endospores per juvenile was transformed with log^^ix + 1) before analysis. The percentages of female infection were transformed to arcsin (Vx) before analysis. All data were subjected to factorial analysis of variance (ANOVA). Means were

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57 separated and compared by Duncan's multiple-range test. Regressions were performed to determine the relationship between the microwave treatment time and the survival of P. penetrans and fungal populations. Results Microwave Treatment Treatment time. Population densities of P. penetrans endospores and fungi decreased with the increasing treatment time {Y= l.95x^ 30.3 Ix + 1 15.67) (Fig. 3-1). Following treatment for 3 minutes, the attachment of endospores was reduced by almost 50% (average of 59 endospores/juvenile). When the treatment time was extended to 4 minutes/kg of soil and 5 minutes/kg of soil, the attachment was reduced to 20 endospores per juvenile and nearly eliminated, respectively. The number of fungal colonies was greatly reduced by all treatment times, as compared to the untreated soil (Y= l.04x^ \2.2\x + 33.69). Fusarium sp., Paecilomyces lilaciniis, Penicillium sp., Pythium sp., Trichoderma sp., and a variety of unidentified fungi were isolated. Soil moisture content. The attachment of P. penetrans endospores to J2 increased slightly with increasing soil moisture content {Y = -1875x^ -I11 Ax + 10.28) (Fig. 3-2). Numbers of endospores attached per juvenile decreased in soils of all tested moisture levels treated for 3 and 4 minutes in the microwave (P < 0.05 ) (Fig. 3-3). A further decrease in attachment was observed with increased treatment time in soils containing 3, 5, and 7% moisture. These observations validate the numbers observed in the treatment-time study reported above (Fig. 3-2). However, the endospore counts of this study were much lower

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58 120 35 9-5 Treatment time in minutes Fig. 3-1. Effect of microwave radiation treatment of 1 kg of soil containing Pasteuria penetrans on A) the attachment of P. penetrans endospores to Meloidogyne arenaria race 1 and B) the survival of soilbom fungi as determined by the number of colony-forming units. Values are means of five replications; 20 randomly selected secondstage juveniles were observed.

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59 25 « 20 15 0 -f— 1% y = -1875x2 + 274x + 10.278 r2 = 0.5466 2% 3% 4% 5% 6% Soil moisture content (weight water/dry soil weight) 7% Fig. 3-2. Relationship between the soil moisture content and the number of endospores of Pasteuria penetrans attached per second-stage juvenile (J2) of Meloidogyne arenaria race 1 after 48 hours exposure at room temperature. Values are means of two replications; 20 randomly selected J2 were observed. 25 20 •s > 3 u w a U] u o a. cn O •O B (X) 1 5 1 0 5 untreated 3 minutes 4 minutes 1% 3% 5% 7% Soil moisture content (weight water/dry soil weight) Fig. 3-3. Effect of soil moisture content and microwave treatment time on the attachment oi Pasteuria penetrans endospores to Meloidogyne arenaria race 1. Values are means of two replications; 20 randomly selected second-stage juveniles were observed. Means within each moisture treatment followed by the same letter do not differ at P < 0.05 according to Duncan's multiple-range test.

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than those of the treatment-time study. Fungal populations were detected at all moisture levels of the untreated soil only. Population densities increased with increasing moisture content of the soil (Y=2lx+ 1; = 0.99) (data not shown) and were 1.2 x 10^ 1.7 x 10\ 2 x 10^ and 2.5 x lO' colony forming units (cfu)/g of soil containing 1%, 3%, 5%, and 7% moisture, respectively. Suppressive-Soil Test Laboratory experiment. Survival of fungi and P. penetrans in the microplot soil varied with soil sources and soil treatments (P < 0.05) (Table 3-1). Regardless of soil source, populations of fungi were lower in autoclaved or microwaved soil than in air dried and untreated soil (P < 0.05). The fungal populations in soil from RKN plots generally exceeded those in soil from RKN + Pp plots and control soils in untreated or in air dried soils. No P. penetrans endospores were detected in control soil or in any autoclaved soils. The number of endospores attached per juvenile did not differ among the microwaved, airdried, and untreated soils. Greenhouse experiments. The effects of the soil sources, treatments, and inoculation levels were similar in both experiments for all data, except the number of eggs as affected by the soil treatments and the female infection rate (Table 3-2). In experiment 2, the number of eggs did not vary with soil treatments, and the single interaction affecting the female infection rate was the soil source X soil treatment interaction. Reproduction of A/, arenaria on peanut, as expressed by the number of root galls, egg masses, and eggs, was consistently higher in autoclaved soil than in microwaved, airdried, and untreated soil infested with M. arenaria alone (Table 3-3) orM. arenaria plus P. penetrans (Table 3-4) after inoculation with 2,000 J2 of M. arenaria. In soil infested

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61 Table 3-1. Colony forming units (cfu) of soil fungi and attachment of Pasteuria penetrans endospores to second-stage juveniles (J2) of Meloidogyne arenaria in untreated soil and soil autoclaved twice for 1 .5 hours at 55 kPa, microwaved for 3 minutes/kg of soil, or air dried for 2 weeks in the greenhouse. Soils contained 6.9% moisture and were collected from peanut microplots infected with M. arenaria inoculum alone (RKN), M. arenaria plus P. penetrans inoculum (RKN + Pp), and a control treatment without added nematodes or bacteria (control). o • 1 Soil source Soil treatment Cfu of fungi per gram of soil Mean number of P. penetrans endospores per juvenile Control Autoclaved 80 b A Oa Microwaved 80 b A Oa Air-dried 1.1 X 10' a B Oa Untreated 1.6 X lO-'a B Oa RKN Autoclaved Ob A Ob Microwaved 40 b A 1.1 ab Air-dried 3.4x10' a A 8.7 a Untreated 4.5 X 10' a A 9.5 a RKN + Pp Autoclaved 80 b A Ob Microwaved Ob A 9.5 a Air-dried 1.5 X 10' a B 12.4 a Untreated 1.5 X 10' a B 9.8 a Values are means of five replicates for the cfu counts and two replicates for the endospore counts; 20 randomly selected J2 were observed per replicate. The data were transformed with logio(x -I1) before being subjected to ANOVA. Presented data are untransformed. Means within columns for each soil source or for each treatment followed by the same lower case letter or upper case Ietter,respectively, do not differ at P < 0.05 according to Duncan's multiple-range test.

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62 • Table 3-2. ANOVA table for the effect of soil source (microplots infected with Meloidogyne arenaria race I alone, with M. arenaria race 1 plus Pasteuria penetrans, and without either organism), soil treatments (autoclaved, microwaved, air-dried, and untreated), and M. arenaria race 1 inoculum levels (0 or 2,000 second-stage juveniles) on nematode reproduction and fresh root weights of peanut cv. Florunner. Source of variation Number of galls Number of egg masses Number of eggs Females infected Fresh root weight Experiment 1 Soil source (S) *** *** *** *** NS Soil treatment (T) *** *** *** ** NS Tnonilation IpvpI (1^ *** *** *** SxT *** *** *** ** NS S xl *** *** ** NS IxT *** *** *** ** NS IxTxS *** *** *** * NS Experiment 2 Soil source (S) *** *** ** *** NS Soil treatment (T) *** *** NS * * Inoculation level (I) *** *** * NS SxT *** ** *** * NS Sxl *** ** *** NS NS IxT *** *** *** NS NS IxTxS *** *** NS NS *, **, *** represents P < 0.05, P < 0.01, and P < 0.001, respectively. NS = nonsignificant atP< 0.05.

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65 with M. arenaria alone of experiment 1, fewer nematodes reproduced in air dried than in microwaved, and untreated soil (P < 0.05) (Table 3-3). This observation was not validated by experiment 2. Nematode reproduction in all three autoclaved soils did not differ from nematode reproduction in microwaved and air-dried soil collected from microplots without nematodes P. penetrans, except in the uninoculated microwave treatment of experiment 2 {P < 0.001, statistics not shown) (Tables 3-3 to 3-5). When no nematode inoculum was added, a very low level of nematode reproduction was observed only in untreated soil from plots infested with M. arenaria alone (P < 0.001) (Table 3-3), and in microwaved soil from ininoculated plots of experiment 2 (Table 3-5). No females were infected with P. penetrans in all autoclaved soils, and in soil treatments of nematode and Pasteuria-fveQ soil (Tables 3-3 to 3-5). When soil was infested with nematodes, 100% of the females were infected in microwaved, air-dried, and untreated RKN + Pp soils of experiment 1, and 77% were infected in microwaved soil in experiment 2 (Table 3-4). A lower incidence of female infection occurred in the soil from microplots that had become infested with P. penetrans. Ten to thirty percent of females in microwaved, air-dried, or untreated RKN soil were infected with P. penetrans when nematodes were added (Table 3-3). When no nematode inoculum was added, infection rates of females in untreated RKN soil were 69 and 77%. However, females in uninoculated microwaved soil of nematodeand P. penetrans-fvce microplots were not infected by P. penetrans. The effects of soil treatment on the fresh root weight were inconsistent (Tables 3-3 to 3-5). Peanut plants culmred in autoclaved or microwaved soil were not different or tended to developed larger root systems than those grown in air dried and untreated soils (P < 0.05). Inoculation level had no effect on the fresh root weight (P < 0.05) (Table 3-2).

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66 u c O 3 S ° 2-.£ £ D «s i o _o o o a B > 3 C C13 U c T3 1) C c E o o o -a .4O O t, e c != =^ > 2 53 u b 2 E ii o i2 'o 0) c C3 > 1) n c -a .-2 § uj p O 4) t. T3 O §° E £ '3 o 'w o u t« o E o E 60 S 60 O E o 60 E .2 O 0^ o > s E o u c/0 i: XI XI X ca X 03 X 00 as VD 00 CO rl-" fS CS a X CS X X ON lO rn (N c o < C3 O ca O ca O o o' CI C3 O C3 O ca O ca O O o ca O X ca ca ca m VO o O iri •o ca o •a u > ca o ii ca O ca o o o ca o ca o ca ca ca X 00 CN oo o r00 -a ca u CS c o E 'C o X m ca O X O o o in oo' 00 X o ca o ca o o 00 O o o so' oo ca CN X o ca •a > "o o 3 < > ca o Io ii ca O X o o o X o X o ca o X o o o o oo 00 X o X ca ca X ca m 00 CS O o 00 X o ca ca X ca en 00 CS rm •o u ca c u E 'C u o. X o X (J ca u CS U u |: X 2 i .E Q. CO ^ a 3 ca p o ^ M CO ^ n ^ § c Q o o c ca __ c 5 o c " c o c « g-q U O E VI CO Qs =^ CJ u cH U T3 u T3 73 ca CO &> T3 O E u c o z u ej ca 53 CO flj C s u " i ca ca CO to 1) U X 3 ca >• > •a o o CS 1—1 o o o ts" X ca u "5. u. 0) c o ca 5 ^ c o ca > X o o CS c ca X CB > X o o z II

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67 When no nematodes were added, no nematode reproduction was noted in all but the untreated soils of RKN and RKN + Pp soil and in microwaved control soil (Tables 3-3 to 3-5, statistics not shown). Nematode reproduction generally did not differ between inoculated and uninoculated treatments in microwaved, air dried, and untreated soils of RKN and RKN + Pp plots (except in the microwaved and untreated soil of RKN plots), but was higher in all inoculated, control soils and following the autoclave treatment {P < 0.05). Discussion An increase in treatment time or in soil moisture content resulted in an increasing effect of microwaving soil on soil microorganisms. The response of soil fungi to an increase in soil moisture content does not agree with Ferriss's (1984) observations on microorganisms. He reported that soil temperature and the effect on reduction of populations of microorganisms correlated positively with MW treatment time and negatively with soil moisture content. Moisture levels approaching field capacity were required to kill spores of root-pathogenic fungi by microwave heat treatment (Baker and Fuller, 1969). The percentages of soil moisture of loamy soils used in Ferriss' (1984) studies exceeded by far the soil moisture contents of soils used in this study. Loamy soils would probably require an extended period of microwave treatment to approach an effect similar to that attained on sandy soils. The different results also may be explained by the fact that sandier soils cool faster than loamy soils after microwaving (Ferriss, 1984); hence, fungal propagules may have been affected by the duration of the elevated temperatures rather than by the temperature peak.

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68 Endospores of P. penetrans are reportedly resistant to high temperatures (Stirling et al., 1986; Williams et al., 1989); however, when endospores were exposed to 80 °C for 30 minutes their development was impeded (Dutky and Sayre, 1978). More recent studies showed that attachment was close to minimum at 60 °C or higher, but it was not completely prevented at 100 °C for 5 hours per day over 10 days (Freitas, 1997). In this study, a microwave treatment time of 3 minutes for a 1-kg sample of soil containing nearly 7% moisture heated the soil to 70 to 75 °C for less than 1 minute. Following the microwave treatment, the number of endospores attached per juvenile was reduced to a variable degree, but was not lower than four endospores per juvenile. This effect was tolerable because the germination of a single spore is enough to create infection in the female Meloidogyne (Stirling, 1984); and hence, sufficient to be used to confirm nematode suppression by P. penetrans. One hundred percent infection of females occurred after the microwave soil treatment, indicating that P. penetrans was able to survive the microwave treatment and suppress the nematode population in the pot experiments. Although methods to quantify facultative and obligate nematophagous fungi in soil have been developed, they do not always give satisfactory results (Dackman et al., 1987). Propagules of saprophytic and nematophagous fungi are of similar structure (Barron, 1992); therefore, they are likely to withstand similar amounts of microwave radiation. If the results of the dilution plating technique were extrapolated to the survival of nematophagous fungi, microwaving soil for 3 minutes per kg of soil had the potential to selectively eUminate this group of microorganisms. This allows for the separation of the antagonistic effects of fungi and the bacterial parasite. Microwaving soil 3 minutes for each kilogram of soil was chosen to be the most appropriate microwave treatment to be used in the suppressive soil test. The treatment does not leave a nematicidal residual effect,

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69 as observed with soil applications of 40% formaldehyde (Dickson et al., 1994; Kerry et al., 1982) and benomyl (Hoestra, 1976) treatment. The numerical difference in attachment of endospores to J2 between the two microwave studies on water content and treatment time may be due to the fact that the dried soil containing P. penetrans was stored for 8 weeks between the studies, thus affecting the ability of endospores to attach to nematodes. A minimum of 3 days moisture incubation (Brown and Nordmeyer, 1985) before the application of soil treatments and nematode inoculum may have enabled more endospores to attach. Attachment by P. penetrans was detected on J2 exposed to soil from RKN and RKN + Pp microplots. Nematode reproduction was consistently inhibited in the presence of viable P. penetrans endospores in these soils. Endospores in both soils existed in densities at which plant infectivity by the J2 was reduced, which protected the root systems from being severely galled. Similar results were found when J2 were encumbered with 15 or more endospores of P. penetrans and used to inoculate tomato (Brown and Smart, 1985; Davies et al., 1988; 1990; Sell and Hansen, 1987; Stirling, 1984). The infection of M. arenaria by P. penetrans resulted in low fecundity. This supports earlier observations that infection by P. penetrans causes sterility or reduced fecundity in females (Mankau, 1980a; Minton and Sayre, 1989); however, the number of isolated females was high enough to statistically confirm 100% infection by P. penetrans only from soils infested with M. arenaria and P. penetrans used in experiment 1. Additional tests with soils containing a more diluted P. penetrans population density may be needed to confirm the nematode suppression by P. penetrans. Temperature has been reported to affect nematode mobihty (Bird and Wallace, 1965), attachment of endospores to J2 (Freitas et al., 1997; Hatz and Dickson, 1992; Stirling, 1981), and the development of P. penetrans in vivo (Hatz and Dickson, 1992; Nakasono et al., 1993; Serracin et al., 1997). The constant

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70 temperatures of the growth room used for experiment 1 probably favored the development of P. penetrans. Although in this study the effect of fungi and P. penetrans could not be evaluated separately, an additive nematode suppression would be expected to cause differences in nematode reproduction between the treatments of microwaving or air drying of RKN and RKN + Pp soils. The lowest numbers of root galls, egg masses, and eggs were observed following air-drying of RKN soil in experiment 1 and of RKN + Pp soil in experiment 2. However, since these effects could not be observed when replicated, fungi appeared to play only a minor role in nematode suppression. These results supported the hypothesis that P. penetrans is the main contributor to the suppressiveness in soils from both the RKN and RKN + Pp microplots. In the suppressive soil test, the microwave treatment was helpful because it reduced fungi that might also be a factor in suppressive soils. However, the endospore density in RKN + Pp soil was so high that it greatly suppressed the plant infectivity by endosporeencumbered J2, and the female infection rate could not be determined. It cannot be ruled out that the nematode suppression was due to other factors than parasitism by P. penetrans; however, compared to the control soil, there were only minor differences between the test results of the RKN and the RKN + Pp soil comprising nematode suppression observed in Chapter 2. Hence, it can be concluded that the suppressiveness of the soils is of similar nature. At the time the soil for this test was collected from the microplots, nematodes of RKN + Pp plots had been suppressed for 3 years. This study demonstrated that endospores of P. penetrans survived 3 years in the absence of a host and successfully suppressed a reinoculated nematode population. Although it is not known exacdy, dormant endospores are probably able to survive for many more years. Pasteiiria spp.

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differ from successful natural enemies of insects, which usually require a constant food supply to maintain their population density (Bennett, 1974). The absence of a host nematode appears to have little impact on the survival of P. penetrans, which may be a leading attribute for the organism in suppression of nematodes under various conditions

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CHAPTER 4 POPULATION DEVELOPMENT OF MELOIDOGYNE SPP. AND PASTEURIA PENETRANS AS AFFECTED BY CULTURAL PRACTICES IN TOBACCO Introduction The endospore-forming bacterial parasite, Pasteuria penetrans (Thome) Sayre & Starr, is widely distributed in agricultural soils throughout the world (Sayre and Starr, 1988; Stirling, 1991) and contributes to natural and induced nematode control, especially of Meloidogyne spp. (Bird and Brisbane, 1988; Brown et al., 1985; Channer and Gowen, 1988; Chen et al. 1994; 1996b; 1997c; Dickson et al., 1994; Kerry, 1990; Mankau, 1980a; 1980b; Minton and Sayre, 1989; Oostendorp et al., 1991a; Weibelzahl-Fulton et al., 1996). In most observations, the suppression of Meloidogyne spp. by P. penetrans occurred after long-term monoculture of susceptible hosts in association with Meloidogyne spp. (Bird and Brisbane, 1988; Chen et al., 1994; Mankau, 1980a; Minton and Sayre, 1989; Chapter 2). Pasteuria penetrans has great potential for integration with other cultural or nematode management practices (Brown and Nordmeyer, 1985; Freitas, 1997; Maheswari et al., 1988; Oostendorp et al., 1991a; Stapleton and Heald, 1991; Tzortzakakis and Gowen, 1994a; Walker and Wachtel, 1989). Pathogenicity and virulence of P. penetrans can vary among different bacterial isolates (Channer and Gowen, 1992; Davies et al., 1994; Oostendorp et al., 1990), and knowledge about their biology is required to effectively include the organism into nematode management strategies (Stirling, 1991). A mixed population of root-knot nematodes, Meloidogyne incognita (Kofoid & White) Chitwood, M.javanica (Treub) Chitwood, and M. arenaria (Neal) Chitwood, was 72

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73 observed to damage tobacco grown in a field at the University of Florida Green Acres Agronomy Farm, Alachua County. Pasteuria penetrans and several species of nematophagous fungi were isolated from the site in 1991 (Chen et al., 1994). Pasteuria penetrans was identified to be suppressive to the mixed populations of root-knot nematodes, and significant changes in the density of Meloidogyne spp. were observed with inorganic nitrogen rates, cover crops, and tobacco cultivars (Chen et al., 1994). The objectives of this study were to test the pathogenicity of the naturally occurring P. penetrans population to M. incognita and M. javanica in the laboratory, and to determine the effects of nitrogen fertilizer, autumn cover crop, and tobacco cultivars on the population density development of Meloidogyne spp. and P. penetrans in the tobacco field. Materials and Methods In the summer 1992, a random sample of approximately 30 roots was collected from the tobacco site. The roots were washed free of soil, cut into 2to 3-cm pieces and mixed thoroughly. Forty females and accompanying egg masses of Meloidogyne spp. were dissected from randomly selected root pieces, and the females were placed individually on an eye glass in a drop of deionized water. The egg masses were treated with 0.5% NaOCl for 1 minute, washed with sterile water three times, placed individually into 2.5-ml microfuge tubes containing 1 ml of deionized water, and incubated at room temperature. The females were cut open at the neck region to release the body contents, which were then examined for mature endospores of P. penetrans. The nematode species was identified by optical examination of 20 perineal patterns.

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74 Nematode Populations One week after isolation from roots, 5 ml of water containing approximately 250 second-stage juveniles (J2) hatched from egg masses obtained from three Pasteuria-free females of either M. incognita or M. javanica were placed into five 2-cm-deep holes around the stems of two 4week-old tomato (Lycopersicon esculentum L. cv. Rutgers) plants, and maintained in the greenhouse. After 60 days, the root systems were harvested and washed free of soil. Eggs of M. incognita and M. javanica were extracted from the infected tomato roots (Hussey and Barker, 1973), hatched by the Baermann method (Rodn'guez-Kabana and Pope, 1981), and used for experiments as 1to 4-day-old J2, or inoculated to ten 6-week-old tomato plants, and maintained in the greenhouse. Pasteuria penetrans Isolates One week after their isolation from roots, approximately 250 J2 of three Pasteuriafree females of both M. incognita and M. javanica were exposed to mature endospores of P. penetrans harvested from one female of the same species. After 24 hours incubation in water at room temperature (26 °C), the average number of endospores attached to 20 randomly selected J2 was determined with an inverted microscope at X400 magnification. The endospore encumbered J2 were suspended in 5 ml of water and then placed into five 2-cm-deep holes around the stems of two 4-week-old tomato (Lycopersicon esculentum L. cv. Rutgers) plants, and maintained in the greenhouse. After 60 days, the root systems were harvested and washed free of soil. Root systems were enzymatically digested in Pectinol (Genencor, South San Francisco, CA) at room temperature for 48 hours. Mature globose females were hand-

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75 picked from the softened roots, and air dried on an eye glass. After 3 days, the nematode bodies were rehydrated for 8 hours and macerated in a glass tissue-grinder. The number of endospores was counted with the aid of a hemacytometer, and their concentration was adjusted to 1 x 10^ endospores per ml water. Laboratory Experiment Attachment studies were conducted using the centrifuge technique (Hewlett and Dickson, 1993). A 0.1 -ml sample of the endospore-water suspension and 0.1 ml of a suspension of 2,000 J2 of M. incognita or M. javanica per ml of water were placed in a 0.25-ml, previously silanized microfuge tube and centrifuged at 9,500g for 2 minutes using a microfuge. Each attachment study was replicated five times. The content of the microtube was stirred and placed on a glass slide with a pipette. The number of endospores per juvenile was determined for 20 randomly selected nematodes per repUcate. Thereafter, the endospore-encumbered nematodes of both species were suspended in 5 ml of water and inoculated to tomato plants as described above. Field Experiment The site had been planted to tobacco in the same plots for seven consecutive years. The 3 X 2 X 2 factorial treatment design included: three autumn cover crops (hairy indigo, Indigofera hirsuta L.; forage sorghum. Sorghum spp.; and weed fallow); two inorganic nitrogen fertilizers (89 and 158 kg of ammonium nitrate/ha); and two tobacco cultivars (Coker 371 -Gold, which is susceptible to M. incognita, M. javanica, and M. arenaria; and Northrup King K-326, which is resistant to M. incognita but susceptible to M. javanica and

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76 M. arenaria). Hairy indigo and forage sorghum were planted in August and plowed under in November of each year. Rye was planted over the entire field in early December to serve as a winter cover crop, and plowed under in March. Tobacco seedlings were transplanted on 27 March and on 2 April of 1992 and 1993, respectively. The final harvest was .taken on 4 August and 1 1 August of 1992 and 1993, respecdvely. Preplant fertilization included a broadcast applicafion of 1,800 kg of 6-6-18 (N-P-K) and 340 kg of 15-0-14 (N-P-K) of sodium-potassium nitrate per hectare. At the final cultivation, in late April, granular ammonium nitrate was applied adjacent to the plant stems. The plots were arranged in complete randomized blocks, and replicated four times. Each block consisted of one row with a row spacing of 1.2 m and a length of 12 m. A 12-core soil sample was taken with a cone-shaped auger (2.5-cm diameter) from the root rhizosphere (20 cm deep) of each plot on 2 April, 9 June, and 28 August of 1992, and on 2 April, 24 June, and 14 August of 1993. The soil was mixed, sampled for soil moisture determination (samples of 2 April 1992 and 1993), and processed using centrifugal-flotation method (Jenkins, 1964). The number of J2/100 cm^ of soil, the rate of attachment of P. penetrans, and the number of endospores attached per 20 randomly selected J2 were determined using an inverted microscope at X400 magnification. Root samples were collected after the final tobacco harvest, 20 females were dissected randomly from each sample, crushed on a glass slide and checked for infection by P. penetrans with a compound microscope at XI 000 magnification. Effects of the autumn cover crop were determined for the 1992 season only.

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Results 77 Laboratory Experiment . Meloidogyne incognita and M. javanica were identified from tobacco root samples, which was thereafter confirmed by Chen et al. (1994). After 24 hours of incubation of the original P. penetrans isolates with J2 of either species at room temperature, 87% of J2 of M. incognita and 82% of J2 of M. javanica were attached, with an average number of 3.8 and 6.7 endospores per juvenile, respectively (data not shown). With the exception of one attachment test conducted with the P. pene trans-isolate from M. javanica, the number of endospores per juvenile of M. incognita was higher than or not different than the number of endospores attached per juvenile of M. javanica (P < 0.05) (Tables 4-1 and 4-2). Attachment of the M. javanica-isolate was higher on M. incognita than on M. javanica after the bacterial parasite was grown on M. incognita for two consecutive generations (P < 0.05) (Table 4-2). However, endospore attachment of the M. incognita-isolate after three generations on M. javanica was either maintained or reduced (Table 4-1). The percentages of J2 attached with endospores were between 72 and 100% (Tables 4-1 and 4-2). In the first and second generation attachment tests, the number of endospores per J2 and the percentage of encumbered J2 followed the same trends. In the third generation attachment tests, the percentages of J2 with attached endospores generally were not different. Field Experiment The soil moisture content ranged from 6.1-1 1.8% soil moisture in 1992, and 6.411.1% in 1993. The number of root-knot nematode juveniles in the soil and the galling

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78 c a t> C V Ml C/5 i> S o T3 w O c c3 C (D C •T3 C (N O i> " o ^ S •a ^ o C c D C JO e o o C In P C o o oo O o 00 o as oo o o c3 ON o o o CO in I X) p 00 >n 00 X) X) o I . incc ~i o o § — ^ .c -I ^' g •a 3 o c C3 O O ^ C u, 1) "-^ > -o s t .2 ^ ^ o ^ (U o o c ^ S , D X C 03 C a D s o 03 o u l-l 03 O _3 > 3 o 0) T3 O c T3 D C D W) •a c D X (U 3 o3 (U C cn X 3 C o 03 03 3 a, o T3 o c3 •a o o 3 o 1/1 o on O c

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ttachment « 3rd generation Number of endospores/ juvenile Test nematode lachment 2nd generation Number of endospores/ juvenile Test nematode ttachment 1st generation Number of endospores/ juvenile Test nematode o o O — ' o as 00 o o 00 CN d CM -Ih (50 O s; en On 00 O '5 C 60 o o c (N .1 s 53 60 O o c 8 00 00 en X> m CN a s s c 60 r\ 5 60 incc § Q O •S 00 I .3 .1 5 C 60 o c3 d 53 O O q 00 ^ ^ ^ 00 u 3 O B (U -5 .id C3 o '-S o u c Ui rv (U > T3 > (U c« > C "-I .2 a 1^ C/! (U •§ ^ o o c ^ £3 on = >. s-g S c 5 2 o (N k< . > D c a, C § 6 ID o S3 0) _3 >

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80 indices were generally higher in 1992 than in 1993 (statistics not shown) (Table 4-3). At planting and after harvest in 1992, the root-knot nematode density was higher in plots with the history of the higher rate of inorganic nitrogen than in those that received the low nitrogen rate (P < 0.05). This effect was not observed in 1993. Throughout 1992 and 1993, the population density of J2 in the soil was increased by both tobacco cultivar histories, but no difference was observed between cultivars in 1992 and most of 1993. However, a higher nematode population density under the resistant tobacco cultivar than under the susceptible cultivar was revealed on the last sampling in 1993. The number of J2 was higher with weeds than hairy indigo or sorghum at harvest in 1992. At mid-season of 1992, but not in 1993, an interaction between cultivar and autumn cover crop was found to affect the number of J2 in the soil (Table 4-3). The nematode population density was lowest in NK-326 plots that followed the forage sorghum cover crop (data not shown). This observation was not validated by repeat in 1993. In 1992, interactions between the inorganic nitrogen fertilizer levels and the tobacco cultivar histories affected the number of J2 in the soil at planting and after tobacco harvest {P < 0.05) (Tables 4-3 and 4-4). At planting and after harvest, the higher rate of ammonium nitrate fertilizer resulted in a greater population density of J2 in the root rhizosphere of NK-326 than in Coker 371 -Gold. As opposed to NK 326, the number of nematodes in the root rhizosphere of Coker 371 -Gold was not increased by the higher level of fertilizer (Table 4-4). Root galling of Coker 371 -Gold was less than that of NK-326 {P < 0.001 for 1992, P < 0.05 for 1993). In 1992, root galling on tobacco was less in plots following hairy indigo and forage sorghum than in plots following weeds (P < 0.01), but no observations were made in 1993 to validate the effects of the cover crops. The galling indices were not affected by the nitrogen treatments; however, an interaction between the inorganic nitrogen

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81 c o c o JO s D C ^ -S C w -a " (U TO u I— « r<^ On c/j ^ > CS O c o . o ex 2 ^ o 1=8 O o •3 c C t>0 § 'o 0c/3 4-J OQ U a 2 o u, > o o t/3 C o o D 1/3 •a B a X lU T3 60 15 O o O o U O. CM 3 3 < 3 C flj > ON OO o U * C/2 z z z z z OO 00 00 z z z 00 00 z z 00 Z 00 Z 00 Z 00 00 00 OO 00 Z Z Z Z Z 00 00 00 Z Z Z Z * 00 * Z * 00 00 Z Z 00 OO 00 z z z o 5 < > o z < o u U O O 2 3 O Z U u o U o o ^ U U U X X X Z U Z < > Z<2 << 2 > T3 O 4> Z x-2 3 o o o m o o Q VI + =^ :S o ^ c o o -a o o ? o E o !/3 o , E ^ u 3 •a c C u U & 1 X H 1 » ."J E efi hd n's c o 'c3 ea nc E 3 <+O Q o c/3 1/3 4— • c o c C3 i-T E — treat _> "3 c O Data withi n and 1/3 u C c3 o Me nitr o o VI a, 4— > 03 C C3 U c o c 00 z 1) _> o u C/3 o o d> VI Q, •a c c3 O CD VI a, >n o d VI c c . .2 CL, )( * if> K C/3 X o o c 73 O o 00 o' 00 NO in m m CM CO o" CN CM 60 O c II o jj 13 u X (U T3 C c C3 (U X H

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82 Table 4-4. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar history, and their effects on the number of second-stage juveniles (J2) of Meloidogyne spp. in 100 g of soil collected at planting and after the final harvest in 1992. Number of J2 per 100 g of soil 2 April 28 August Cultivar history 89kgN 158 kg N 89 kgN 158 kgN Coker 371 -Gold 8aA 9b A 477 a A 469 b A NK-326 8aB 23 a A 140 bB 844 a A The values were transformed with log,o (;c -I1) before being subjected to ANOVA; the same lower case letters in columns or upper case letter in rows for the same sampling dates indicate no significant difference at P < 0.05 according to ANOVA. Table 4-5. Interaction between inorganic nitrogen fertilizer levels and tobacco cultivar history, and their effects on the root-galling index in 1992 and 1993. Root-galling index" 1992 1993 Cultivar history 89 kgN 158 kgN 89 kgN 158 kgN Coker 371 -Gold 2.7 a A 2.2 b A 1.0b A 1.4 b A NK-326 3.1 aA 3.4 a A 2.2 a A 2.4 a A The values were transformed with arcsin (V;c) before being subjected to ANOVA; the same lower case letters in columns or upper case letter in rows indicate no significant difference at P< 0.05 according to ANOVA. " The galling index scale: 0 = no galls, 1 = 1-10, 2 = 1 1-20, 3 = 21-55, 4 = 5680, 5 = 81-100% of roots galled.

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83 fertilizer and the tobacco cultivar history was found to affect the galling indices (P < 0.001) (Tables 4-3 and 4-5). The high rate of ammonium nitrate resulted in more severe root galling on NK-326 than on Coker 371 -Gold in both years (P < 0.05) (Table 4-5). This statement is valid for the low level of nitrogen in 1992, but not for 1993. In comparison with a root galling index of 3.8 in the weed fallow plots, the severity of root galling decreased to 2.0 and 1.7 after Coker 371 -Gold plots were covered with hairy indigo or forage sorghum, respectively (data not shown). Galling indices differed between the blocks in both seasons (P < 0.001) (Table 4-3). A high percentage of the J2 had endospores of P. penetrans attached (Table 4-6). The percentage attachment and the number of endospores attached per juvenile remained fairly constant over the six sampling dates. In June and August of 1992, and in August 1993, the percentages of J2 with attached endospores were higher in the root rhizosphere of Coker 37 1 -Gold than in NK-326 (P < 0.05). The autumn cover crops affected the percentage of attached J2 only on the first sampling date of the first year. At that time, more J2 were encumbered with endospores after an autumn cover with weeds or hairy indigo than after forage sorghum. No effect of the inorganic nitrogen fertilizer was observed on the percentage of J2 with attached endospores of P. penetrans or the number of endospores per juvenile. The percentage of females infected by P. penetrans was not affected by the treatments. Variations among blocks affected endospore attachment and the female infection rate at various sampling dates. An interaction between tobacco cultivar history and autumn cover crop revealed that in the root rhizosphere of NK-326, more 32 were encumbered with endospores after weed and hairy indigo autumn cover than after forage sorghum (P < 0.05) (data not shown). There was an interaction among the nitrogen levels, the tobacco cultivar history, and the autumn cover crops affecting the percentage of J2 attached with

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84 o B o •S w ^ 8 a. O « m C On «5 « « c D ^ On C a\ 03 t/3 o O > o o n. o c c/3 (u 5 >s a 60 U P o — ^ O c 3 -a > 3 I T3 C o o u C 03 o s 00 T3 C3 a, ^ c I < ^ c o uj 4) c3 -> -9 > C o u o ^ 2 1) M c E OS O a y. O T3 C u CM a o o a « o o c u u N T3 rti ch a ro 'S o O u M C3 C O U u cu 00 §1 0\ c b > u — OS (S 00 O o d ro fo — . ro ro 0\ 00 d — ro -"t tN O in 00 — , VO in CO X) — oo 5 "5b ^ 00 Ov in 00 — c u 00 o O jO vo in vo in ro tN CN 2 "o a rro vo 0) ro O ^ U Z Ov t-~ in vo in in on on 2 Z on on Z Z ro — ; O ro oo in ro cs ro q — o o in vo vo ro vo vo vo ro tN o 60 in oo X) 00 on Z on on on z z z c/3 on on z z z on on z z on on on z z z on on on z z z on » Z J on on Z Z on on on z z z on on z z z * on on on z z z on 2 on on Z Z on ' Z ' on on on on on z z z z z on on on on on z z z z z on on (/) on on z z z z z on ' Z ' on Z ' on Z on on on on on z z z z z on on on on on z z z z z * c/2 on „ , * Z Z * * ,U .2 o a: o 00 > u O 0) CQ Z U U 8 a u (JO** .u u u u XXX z z u C3 3 u > o U o 3 C u cs u. U > u •a o C u E "c « o c 3 Q « x: "a >v ^ H ^ > o 60 <= < s 4i > — I 3 ^ (U o C 60£ C 60 '5 .S ^ -5 ,P x: c O o p (U t« £ ^ 60 ~ Ui P. U 60 3 c o c o T3 u. B o c ou x: H -a p c/5 S .S g 2 e p S S s E ^ S ^ §= ^ 2 O Ui P p > O 5^ :? o d VI c CJ "S 60 O a II 00 Z p > • ^ p p a o o d VI Q, •o c o d VI in o d VI Q, c 2^ * * C > P 1/1 X3 O O c

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85 Table 4-7. Interaction among inorganic nitrogen fertilizer levels, tobacco cultivar history, and cover crops, and their effects on the percentage of second-stage juveniles (J2) encumbered with endospores of Pasteuria penetrans at planting in 1992. Cultivar historv Cover cron Percentage of J2 encumbered with endospores 89kgN 158 kg N Coker 371 -Gold Weed fallow 34 a A 34 a A Hairy indigo 6b A 27 a A Forage sorghum 15 b A 12aA NK-326 Weed fallow 32 b A 46 a A Hairy indigo 46 a A 17bB Forage sorghum 1 c A 4bA The values were transformed with arcsin (Vx) before subjected to ANOVA and Duncan's multiple-range test. The same lower case letters in columns within each cultivar history or upper case letter in rows indicate no significant difference at P < 0.05 according to Duncan's multiple-range test or Student Mest, respectively. endospores at the time of planting (Table 4-6 and 4-7). In NK-326 plots under hairy indigo, the low rate of ammonium nitrate resulted in the highest endospores attachment per juvenile in comparison with weeds and sorghum {P < 0.01); however, at the high rate of fertilizer the percentage of attachment was highest after weed fallow. The trends were not observed in 1993. The highest endospores attachment per juvenile in Coker 371 -Gold was observed at the low rate of nitrogen after weed fallow, but there was no differenc between the nitrogen levels. Discussion Over a time period of 2 years in a long-term field experiment with tobacco, the nematode population densities and root-galling indices declined, whereas the P.

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86 penetrans population densities increased. The relatively high percentages of females with endospores verifies that P. penetrans was established and impacted the root-knot nematode population. The two isolates of P. penetrans were similar or sometimes more virulent to M. incognita than to M. javanica; however this experiment was not repeated for validation. Multiple years of Coker 371 -Gold favored endospore build-up in the soil. These observations are in contrast with results from a preceding season (Chen et al., 1994), in which nematode populations densities in the root rhizosphere of NK-326 were lower than those of Coker 371 -Gold, and the percentage of endospore encumbered J2 was not different or slightly higher in NK-326 then in Coker 371 -Gold plots. This trend may have resulted from the suppression of predominantly M. incognita, and the build-up of mainly M. javanica, which was less susceptible to attachment by P. penetrans endospores under laboratory conditions. Inorganic nitrogen, urea and their analogs are known to have nematicidal effects (Rodnguez-Kabana, 1986; Rodn'guez-Kabana et al., 1981; Spiegel et al., 1982; Talavera et al., 1984), and to inhibit the development of root-knot nematodes in plant roots by preventing the formation of feeding sites (Glazer and Orion, 1984). Results observed in this study suggest that this mechanism may be of little or no importance for tobacco under field conditions. During the 1992 season, but not in the 1993 season, the tobacco cultivar with resistance to M. incognia supported a higher nematode population than the susceptible cultivar when receiving the high level of nitrogen. In early season, effects by the weed fallow and hairy indigo autumn cover crop may enhance the percentage of endospore-encumbered J2 in the soil. This trend was observed in 1992 but could not be validated in 1993. Although hairy indigo is usually planted to suppress root-knot nematode populations, some nematode reproduction can

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87 occur (Dominguez et al., 1985; McSorley et al., 1994) and contribute to the build-up of P. penetrans endospores. There were outcroppings of clay pockets across the field. These small areas hold more water, thereby leading to variations in soil moisture. Although not mapped, its existence probably effected galling indices, the percentage of J2 with attached endospores, and the percentage of females infected with P. penetrans. The development of Meloidogyne spp. is affected by fine textured soils (Windham and Barker, 1986), and the development of both the nematode and P. penetrans was slower when soil moisture was maintained at field capacity (Davies et al., 1991). The efficacy of the bacterial parasite is known to depend on the density of its host nematode (Stiriing, 1991). All this suggests that oxygen depletion in wet soil inhibits root development and respiration, supports the establishment of root pathogens, lowers the reproductive rates of the nematode, and, hence, hampers the increase of the P. penetrans endospore population densities in soil. It is not clear whether the resistant tobacco cultivar induced a shift in the composition of the root-knot nematode population. Supporting laboratory studies suggested that attachment of the M. javanica-isolatc to M. incognita increased after the bacterial parasite was grown on M. incognita for two consecutive generations; thus, the P. penetrans population present in the field appears to be capable of adjusting to a possible change in the nematode population. If there is time for natural selection to take place, it may be possible for new strains of Pasteuria spp. to evolve that are better fitted to the local environment or more virulent to the shifting nematode population. This also was observed by others (Channer and Gowen, 1992; Oostendorp et al., 1990). As previously reported, P. penetrans has maintained itself in situafions where nematode populations were altered by a change in cropping practice (Stirling, 1991).

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CHAPTER 5 SUPPRESSION OF MELOIDOGYNE INCOGNITA AND M. JAVANICA BY PASTEURIA PENETRANS IN FIELD SOIL Introduction Pathogen-suppressive soils are defined (Baker and Cook, 1982) as soils in which the pathogen does not establish or persist, establishes but causes little or no damage, or establishes and causes disease (conducive soil) but thereafter the disease is reduced even though the pathogen persists in the soil. Soils suppressive to plant pathogens may occur naturally as an inherent characteristic of the physical, chemical, or biological structure of a particular soil. Suppressiveness also may be induced by some agronomic practices, such as planting a crop or adding organic or nutritional amendments, which change the microflora. Induced suppressiveness apparently develops as a result of the buildup of antagonists in response to a high pathogen population (Baker and Cook, 1982). The endospore-forming bacterial parasite, Pasteuria penetrans (Thome) Sayre & Starr, is widely distributed in agricultural soils throughout the world (Sayre and Starr, Stirling, 1991; 1988) and contributes to natural and induced nematode control, especially of Meloidogyne spp. (Bird and Brisbane, 1988; Brown et al., 1985; Channer and Gowen, 1988; Chen et al., 1996b; Dickson et al., 1994; Kerry, 1990; Mankau, 1980a; 1980b; Minton and Sayre, 1989; Oostendorp et al., 1991a). The suppression of Meloidogyne spp. by P. penetrans occurred mostly after long-term monoculture of susceptible hosts with 88

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89 Meloidogyne spp. (Bird and Brisbane, 1988; Chen et al., 1994; Mankau, 1980a; Minton andSayre, 1989). In 1994, a site in Alachua County, Florida that was suppressive to Meloidogyne spp. was identified (Chen et al., 1994; Chapter 4). The field had been planted continuously for 7 years to flue-cured tobacco (Nicotiana tabacum L.). No nematicides or other nematode management practices were applied to the site. Root-knot damage was initially severe and caused by a mixed population of M. incognita race 1 (Kofoid & White) Chitwood and M.javanica (Treub) Chitwood. Yield reductions decreased over the years to a point where there was little economical loss (Whitty, personal communication). Endospores of P. penetrans were found in mature females and attached to second-stage juveniles (J2) of both nematode species from samples collected throughout the field, and several species of fungi were isolated from J2 and egg masses (Chen et al., 1994). The objective of this study was to determine the role of P. penetrans in suppressing M. incognita and M.javanica in soil from this tobacco field. Materials and Methods Soil Treatments The field site was located at the University of Florida Green Acres Agronomy Farm, Alachua County. Approximately 90 liters of soil, a loamy, siliceous, hypothermic Grossarenic Paleudults with 90% sand, 4% silt, 6% clay, and 2.5% organic matter, was collected with a bucket auger (10-cm diameter) in March 1993 (greenhouse experiment 1), and in July 1993 (greenhouse experiment 2) from the tobacco field. The soil was passed

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90 through a 5-mm-aperture sieve, mixed, and divided into four subsamples of 22 liters each. The subsamples were either autoclaved, microwaved, air-dried, or left untreated. Autoclave heating treatment. Sterilized, 15-cm-diameter clay pots were filled with soil (ca. 1 liter/pot), covered with aluminum foil, and placed in an autoclave. After the first heat treatment for 1.5 hours at 55 kPa, the pots were removed from the autoclave. The treatment was repeated after pots and soil had cooled to room temperature. Microwave heating treatment. For the microwave heating treatment, 1-kg lots of soil were placed in even layers in open plastic bags and heated in a microwave oven for 3 minutes (Chapter 3) at full power (1,500 W, 2,450 Mhz), which caused the temperamre to rise to about 75 °C. Air-drving treatment. The 22 liters of soil for the air-drying treatment were collected 2 weeks prior to the collection of soil for the autoclaving, microwaving, and untreated soil treatments. The soil was placed in 3to 5-cm-thick layers on plastic trays and stored for 2 weeks in the greenhouse to eliminate or reduce the natural nematode population. Untreated soil: This soil remained as collected from the field. Laboratory Experiments One day after the soil was treated, a dilution plating technique was employed to assay for selected fungi (Johnson and Curl, 1972). One gram of the mixed soil from each soil treatment was dispersed and constantly agitated in 200 ml of sterilized water. Under sterile conditions 1 ml of the soil solution was plated on potato dextrose agar (PDA; Difco

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91 Laboratories, Detroid, MI) containing 100 mg streptomycin, 50 mg chlortetracycline and 1.0 ml of tergitol/liter of medium, respectively (Steiner and Watson, 1965). Six replicates were plated for each soil treatment. After 3 days of incubation, the fungal colony-forming units (cfii)/g of soil were determined. A soil bioassay (Oostendorp et al., 1991a) used Meloidogyne incognita race 1 as the host to determine the presence of P. penetrans. A sample of 50 g of dry soil from each treatment was placed in a petri dish, and replicated six times for the bioassay. The soil water content was adjusted to 100% field capacity to increase the rate of endospore attachment, and the dishes were left uncovered at room temperature (Brown and Nordmeyer, 1985). After 3 days, 500 J2 of the test nematode were added and the moisture level was adjusted to 50% of field capacity. The J2 were extracted by centrifugal flotation 2 days later (Jenkins, 1964), and the number of endospores per juvenile was determined with the aid of an inverted microscope for 20 randomly selected J2 per replicate. Greenhouse Experiment Treatments were arranged in a 2 X 2 X 4 factorial design that included four soil treatments (as listed above), inoculum levels of 0 and 2,000 J2 of A/, incognita race 1/pot, and tobacco cultivars Coker 371 -Gold (susceptible to both M. incognita and M.javanica) and Northrup King K-326 (resistant to M. incognita but susceptible to M.javanica). The soil from each treatment was placed into sterilized 15-cm-diameter clay pots (ca. 1 liter/pot). Treatments were replicated six times in the first experiment and five times in the second experiment. The moisture level was adjusted to approximately 50% of the field capacity. Pots were covered with aluminum foil and held for 3 days in a growth room at

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92 28-30 °C and 14 hours of light. Meloidogyne incognita race 1 J2 were suspended in 10 ml of water and dispensed equally into each of five holes, 5-cm deep, in the soil of each pot. Five days later, one 5to 10-cm-tall tobacco seedling was transplanted into each pot. A 2020-20 (N-P-K) fertilizer was apphed once per week, and insects were controlled as needed using recommended insecticides. The two experiments were run for 45 days and 46 days, respectively. Plant height and fresh shoot weights were recorded. The root systems were washed free of soil, excess water was removed with paper towels, and the roots were weighed and then stored in plastic bags at 4 °C. The following day, roots were stained with Phloxine B (Dickson and Strubel, 1965), and the numbers of root galls and egg masses were counted. Eggs were extracted with 1.05% sodium hypochlorite (Hussey, 1971) and the number of eggs counted. Nematode Origin The Meloidogyne incognita race 1 isolate used in this study originated from tobacco grown on the University of Florida Green Acres Agronomy Research Farm, Alachua County, Florida. Species and race of the nematode population were identified by optical examination of the perineal pattern with the aid of light microscopy, and by the results of a differential host test (Hartman and Sasser, 1985). The nematode was cultured in a greenhouse on tomato (Lycopersicon esculentum Mill. cv. Rutgers) from a single egg mass. Eggs of M. incognita were extracted from infected tomato roots (Hussey and Barker, 1973), hatched on a Baermann (Rodrfguez-Kabana and Pope, 1981) and used as 1to 4-day-old J2 in the bioassay and the suppressive soil experiments.

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93 Statistical Analysis All data were subjected to factorial analysis of variance (ANOVA). Different means were separated and compared with Duncan' multiple-range test or Student's r-test. Results Laboratory Experiments Survival of the fungal populations and P. penetrans in the field soil varied with soil treatments (Table 5-1). Populations of fungi in autoclaved, microwaved, and air-dried soil were different (P < 0.05); however, quantitative survival of fungi in the air-dried soil did not differ from the untreated soil {P < 0.05). The numbers of P. penetrans endospores per juvenile did not differ among the microwaved, air-dried, and untreated soil (P< 0.001), but all endospores were killed in the autoclaved soil (Table 5-1). In the microwaved soil, fungal populations were reduced in comparison to untreated or air-dried soil, whereas the attachment of endospores was not affected. Greenhouse Experiment Suppressive soil treatment results in both experiment 1 and 2 were similar (Table 5-2). hiteractions were detected for cultivars, soil treatments, and inoculation levels (P <0.05); thus, data of the different cultivars are presented separately (Tables 5-3 and 5-4).

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94 Table 5-1. Survival of soil fungi and Pasteuria penetrans in untreated soil or soil autoclaved twice for 1.5 hours at 55 kPa, microwaved for 3 minutes/kg of soil, air-dried for 2 weeks in the greenhouse. Soil treatments Cfu of fungi per g of soif Mean number of P. penetrans endospores attached per 20 J2'' Autoclaved Oc Ob Microwaved 2 X 10^ b 10 a Air-dried 2,7 X lo' a 13 a Untreated 2,4 X lo' a 8a Values are means two experiments; six replicates each. Means within columns followed by the same letter do not differ at P < 0.05 according to Duncan's multiple-range test. a The number of fungal propagules was determined by dilution plating, and is presented as colony-forming units (cfu). b The endospores of P. penetrans were bioassayed on second-stage juveniles (J2) of Meloidogyne incognita race 1.

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95 Table 5-2. ANOVA table for the effects of tobacco cultivars, soil treatments (autoclaving, microwaving, air-drying, and untreated), and Meloidogyne incognita race 1 inoculum levels (0 or 2,000 second-stage juveniles) on nematode reproduction and plant performance. Number Number of egg Number Total of galls masses of eggs plant Shoot Root Total ANOVA per root per root per root fresh fresh fresh plant system system system weight weight weight growth Experiment 1 Cultivar (C) *** *** *** ** ** ** ** Soil treatment (ST) *** *** *** *** *** ** *** Inoculation level (I) *** *** *** NS NS NS NS CxST *** *** *** *** ** *** ** Cxi *** *** *** NS NS NS NS IxST *** *** * ** NS ** I X ST X C *** *** NS * NS ** Experiment 2 Cultivar (C) *** *** *** ** * ** NS Soil treatment (ST) *** *** *** *** *** *** Inoculation level (I) *** *** *** NS NS NS NS CxST ** *** *** *** ** *** ** Cxi *** *** *** NS NS NS NS IxST *** *** *** NS ** NS ** I X ST X C *** NS NS NS ** *. **. *** represent P< 0.05, P < 0.01, and P < 0.001, respectively, and NS = no differences at P < 0.05 according to Duncan's multiple-range test.

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96 X) o o ^ o ^ O U c .~ ^ c " 10 S-S. o eo c 2 o '35 "o a o 5. " rr u s « •a ri c . CO g do o. 00 c §2 5 3 u o c o — = I « is en 4J ' — s 3 S CO _ ^ §J 00 c c a >> 53 ^ ? « 8 c — s: 60 >^ C So .a 5 6 j= > ^ o — O XI D. o o cn 3 2 . o . O <^ J, O *3 — o u o c C3 E Urn a Urn o. c 60 •a u O O 60 C/3 60 "o o o x: C/2 x: ^ « 6{ 1) _ x: es 6£ f2 O O Q. 60 60 U O " XI 3 60 60 o « 2 CO 60 E O ste o CO O O O lU u X) E 3 c u E o u E u (U O. X U es X! XI VO tN so o tN (?s a es C5 CN 00 tN (N tN CS es es es rso Pes es es es rtN es u X x; tN 00 so tN r00 U-) es es cs es rr~ so C sC tN SO es X3 X3 00 OS cS o 3 < x> o tN o OS tN T3 > CS o x; X! tN in 00 a o es tN CN O o o u Q. X es es cS CB Tics SO CN O es OS tN CJs' -ct JO o o o o > _es o o 3 < XI es es CN XI f»1 o O 00 00 XI o o w-i O -a > CS o CJ XI tN es es in tN tN so c<^ so tN OS OS OS so CN OS OS in CS es es Ties CO Tt a so oo IT) CN OO >n >n o es rOs OS OS CN CN oo tN CS u u c O 60 C o u o es m o o VI a, es w o c o -o o _u u E CS x: XI T3 o c 1) X u x: tN <£ 60 C >> 60 c ? o c o 3 a. o c w c u T3 to ^ es C •a o •o T3 CS CO U •o o u c o z II u o es 00 (J o tN »— » o o o CN x: 1) 60 CO »— I U 3 cS > C tS o c 3 Q II . -13 O (L> 60

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97 g S is o > o c a S " o 1 u. T3 S ° S § Q C/3 O O o " o u O i1 c o. o 3 ^ o "55 00 5 .£ Q ,0 U C > cs s .1 ^ J? = o o S c a. o u u bo VI C 3 I3 2 o e: 5 s > O "« s 2 2 a .2 es VI m 60 • c ~" u o „ =a a. I < O u C 3 ttO O U u c C3 E £ U a c S3 60 60 'S o o 60 'S o o 00 60 "u J3 • . « 6£ 1) O 3 T3 O a o o CO E u 3 o VI 00 00 u o E 2 ol 60 O 60 O a. B "« 60 E O ste o >u VI o O o imb a =1 2 c — u D. X a u ah m 00 VO ca XI X3 ts — a a ca (S 00 (S a ca ca ca + ON fN tN cs XI x> ca o 00 ca x> Xl XI + XI XI ts oo On 00 rXI XI ca O ON ri 00 cn XI X3 ca o o XI ts XI o x> ts > o 3 <: XI X3 O — X) m o T3 0) > ca o o ii XI -a ca NO ca ca T3 U c ca ca ca ca to ca ca ca ca CO to Xi ca XI XI >r) ca "o o 3 < rXI o Xl o X) o XI o Xl o o O -a > ca ? o u CJ x: OO NO OO OO fN tN* Tits X5 CJ O NO o ts ts o rts" o >n 00 to o to o o a u u C 60 C o o o ca iri o CD VI Q, o c o -a U E u x: u ts u a 00 c u -a 00 c o c o 3 O. o D. c u 13 O 3 ca c •a CJ o o ea C/5 U T3 O E U c o Z II u CJ ca o CJ o ts o o o _ O VI — v> 2 » 13 c c c/3 .t^ 1— I 3 3 II ^ E + cn •' 3 c3 x: CJ c c u 3 U Q 60

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98 When the susceptible cultivar received nematode inoculum, root galling, number of egg masses, and individual eggs were less in microwaved, air-dried, and in untreated soil than in autoclaved soil (P < 0.05) (Table 5-3). In experiment 1, the number of eggs was further suppressed in the air-dried soil compared with other treatments {P < 0.05). However, this effect was not observed in experiment 2, in which the microwave treated soil, with few fungi present (Table 5-1), was just as suppressive as the air-dried and untreated soil (Table 5-3). In comparison with the uninoculated treatments, nematode reproduction on susceptible Coker 371 -Gold was lower in the autoclaved, microwaved, and air-dried soil, but not in the untreated soil {P < 0.05). Meloidogyne incognita race 1 reproduced slightly on the resistant tobacco cultivar (Table 5-4). With the exception of the air-drying treatment in experiment 2, nematode reproduction on resistant Northrop King K-326 did not differ between inoculation levels {P < 0.05). When no nematode inoculum was added, and generally when inoculum was added, root-knot damage in the untreated soils was higher than in the other treatments (P < 0.05). Plant performance did not differ among treatments in the case of the inoculated susceptible tobacco cultivar (Table 5-3). However, when no inoculum was added, the total weight, shoot weight, and shoot height but not root weight, were generally greater in the autoclaved soil than that in the other treatments (P < 0.05). In experiment 1 , the performance of the uninoculated resistant cultivar was generally favored in autoclaved and untreated soils (Table 5-4). The weight of inoculated roots of either cultivar, and of uninoculated roots of the susceptible cultivar was not affected by soil treatments (Tables 53 and 5-4).

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99 Discussion Nematode reproduction was suppressed consistently in the presence of viable P. penetrans endospores. The reduction most likely resulted from the infection of M. incognita by P. penetrans, since the numbers of egg masses and eggs were reduced more than the number of root galls. This supports earlier observations that infection by P. penetrans causes sterility or reduced fecundity in females (Mankau, 1980a; Minton and Sayre, 1989). Although nematode suppression by soilbome fungi and P. penetrans could not be evaluated separately, an additive nematode suppression would be expected to cause differences in nematode reproduction between the microwaved and air-dried soils. In one experiment, a significantly lower number of eggs was observed in air-dried soil. However, since this effect could not be reproduced in experiment 2, the roles of fiingi in nematode suppression could not be elucidated. These results support the hypothesis that P. penetrans is the main contributor to the suppressiveness of the soil. The reproduction of Meloidogyne incognita race 1 on the resistant cultivar is not unusual in that it has been reported in several other studies with plants carrying the same Mi 1 resistance gene (Huang, 1986; Schneider, 1991; Veech and Endo, 1970). Root-knot damage on the resistant cultivar in the untreated soil probably resulted from infection by the natural population of M.javanica. Nematode reproduction was not increased by additional nematode inoculum. The generally improved plant performance (except root weight) observed in autoclaved soil was probably induced by an increased release of nutrients into the soil solution (Ferriss, 1984). In experiment 2, inoculated plants did not benefit from the autoclave treatment. In this case water and nutrient absorption may have been limited by the heavy nematode infection (Hussey, 1985). Severe stunting and yield reductions caused

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100 by M. incognita on resistant cultivars was previously described as being caused by the hypersensitive response of the resistant roots to the nematode infection (Barker et al., 1981). The suppressive nature of soils containing P. penetrans has been reported previously (Bird and Brisbane, 1988; Dickson et al., 1994; Mankau, 1980a). In all cited cases, continuous replanting of a nematode-susceptible crop apparently caused the nematode antagonist to increase to suppressive levels. Bird and Brisbane (1988) demonstrated qualitatively the biological origin of nematode suppression caused by P. penetrans in field soil. In the present study, the lower number of developing Meloidogyne spp. in the microwaved soil with viable P. penetrans endospores suggests that P. penetrans was important in nematode suppression and the reduction of plant damage. Further improvement of the bioassay for nematophagous fiingi is required in order to make conclusions about their contribution to the suppressiveness of a soil to pantparasitic nematodes.

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CHAPTER 6 POPULATION DENSITY DEVELOPMENT OF MELOIDOGYNE ARENARIA AND PASTEURIA PENETRANS IN METHYL BROMIDE TREATED SOIL Introduction The annual yield suppression of the world's major crops caused by plant-parasitic nematodes was estimated to be approximately 12% (Sasser and Freckman, 1987). The increasing limitation in number and applicability of registered nematicides has stimulated increased research on biological alternatives. Among many naturally occurring organisms antagonistic to root-knot nematodes, Pasteiiria penetrans has the potential to contribute to natural and induced nematode control (Bird and Brisbane, 1988; Brown et al., 1985; Channer and Gowen, 1988; Dickson et al., 1994; Kerry, 1990; Mankau, 1980; Minton and Sayre, 1989; Oostendoip et al., 1991a; Sayre and Starr, 1988; Stirling and White, 1982). Unfortunately, this obligate bacterial parasite cannot be mass-reared in vitro, which limits economic application to potting soil and small-plot agriculture. Until a procedure of mass cultivation is developed, and P. penetrans is successfully commercialized as a biological control agent, the emphasis has to be on amplification of the pathogen in soil. Levels of P. penetrans that are suppressive to populations of Meloidogyne arenaria can be inoculated into root-knot nematode infested soil (Chen et al. 1996b; 1997c) or can built up in soil over time in the presence of a nematode host and a susceptible crop (Dickson et al., 1994; Minton and Sayre, 1989; Oostendorp et al., 1991a; Chapters 2 and 4). Nematode suppressive levels in peanut (Arachis hypogaea L.) took 3 years of cropping 101

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102 to develop, with crop losses in the first 2 years of a microplot study (Oostendorp et al., 1991a; Chapter 2). Increasing the suppressiveness of a soil to disease on a susceptible host with the simultaneous cultivation of a resistant or tolerant crop in an intercropping system with the susceptible crop may limit economic losses. Intercropping is defined as growing two or more crops simultaneously on the same land (Lewandowski, 1987). This cropping technique is practiced frequently in developing countries, but is rarely used in developed countries like the United States (Francis and Decoteau, 1993). Intercropping can reduce management input (Lewandowski, 1987) and increase yields, especially when cereals are intercropped with nitrogen fixing legumes (Elmore and Jacobs, 1986). The cropping of sweet com (Zea mays L.) and pole bean (Phaseolus vulgaris L.) together simultaneously would be a possible example of such an intercropping system. The growth of sweet com has been shown to be unaffected by a bean intercrop (Willey and Osiru, 1972), and many cultivars of sweet com can support high population densities of root-knot nematodes without suffering yield reductions (McSorley, personal communication). Bean is generally susceptible to root-knot nematodes (Sikora and Greco, 1990); therefore, this crop can be used to amplify the nematode and its biological antagonist, P. penetrans. The objectives of this experiment were to monitor the population development of Meloidogyne arenaria and P. penetrans on a bean-sweet com intercrop in rotation with peanut, and to determine the population density of P. penetrans that provided suppression of root-knot nematodes.

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103 Materials and Methods Microplot experiment. At the University of Florida Green Acres Agronomy Farm, Alachua County, 90 microplots (each 76-cm in diameter), were arranged in 10 rows of nine-plots with a distance of 1.5 m between plots in a loamy, siliceous, hypothermic Grossarenic Paleudults with 90% sand, 4% silt, 6% clay, and 1.8% organic matter. In April of 1984, the microplots were fumigated with methyl bromide (98% methyl bromide plus 2% chloropicrin) at 977 kg/ha under 3 mm polyethylene mulch before inoculation with M. javanica (Neal) Chitwood and Paecilomyces lilacinus (Thom) Samson, and planted to crops supporting high levels of Meloidogyne spp. (Hewlett et al., 1988). In 1987, the plots were again treated with methyl bromide at 977 kg/ha under 3 mm polyethylene mulch and either inoculated with M. arenaria alone, with M. arenaria plus P. penetrans, or maintained free of nematodes and bacteria by periodic fumigation with methyl bromide. For the following 7 years, plots were planted to peanut during the summers, and maintained under rye, wheat or vetch, and bare fallow during the winters. For the present study, all 90 microplots were fumigated with 977 kg/ha of methyl bromide as described above, on 10 March 1994. Thirty days later, the plots where uncovered. After aeration for 19 days, six 5-cm-deep holes and six 10-cm-deep holes (2.5-cm diameter) were evenly distributed throughout each plot. Inoculum of 10* eggs of Meloidogyne arenaria race 1 per microplot was applied uniformly in 2 liters of water over the soil. A potato rake was used to further incorporate the nematodes and leave the soil surface level. A 5-cm-deep furrow was made in a north-south direction through the middle of each plot. For intercropping system 1 (Fig. 6-1), sweet com {Zea mays var. saccharata cv. Silver Queen) was planted on 28 April in the furrows. Two weeks later the com plants were thinned to six plants per plot, and pole beans (Phaseolus vulgaris cv.

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Fig. 61 . Cut away illustration of a sweet com (middle plant) and pole bean intercropping system in field microplots (76 cm diameter)

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105 Kentucky Wonder) were planted to each side of the com. After germination, the pole bean plants were thinned to a stand of three plants on each side of the com. Plots were irrigated by a microjet sprinkler system at the rate of 2 ml/cm^ per day. The plots were fertilized with ammonium sulfate (18-0-0) and potash (0-0-10), with a total of 385 kg N/ha and 320 kg K/ha, respectively, provided in one at-planting and three post-planting applications. Over a period of 1 week in midJuly, plant shoots were cut off at ground level and removed from the microplots. Three root systems of both sweet com and pole bean were harvested from each plot, and galling indices were recorded according to the scale: 0 = no galls, 1 = 1-10, 2 = 1 1-20, 3 = 21-55, 4 = 56-80, and 5 = 81-100% of roots galled (Barker et al., 1986). The root systems were retumed to their original plots and tumed into the soil. After 3 weeks of bare fallow, the soil in each plot was leveled and the same crops were planted in an east-west direction (intercropping system 2). Plots were maintained as described above. After 86 days, the root systems were harvested. Three, 2.5-cm-diameter, 15-cm-deep soil cores were taken for nematode extraction with a cone-shaped auger from each plot. Nematodes were extracted from 100-cm^ subsamples using centrifugal flotation (Jenkins, 1964). The numbers of second-stage juveniles (J2), endospores per juvenile, and the rate of attachment by P. penetrans (as averages of 20 randomly selected individuals) were determined with the aid of an inverted microscope at X400 magnification. For the winter, plots were planted in mid-November to a cover crop of hairy vetch {Vicia villosa Roth). On 19 April 1995, the plant shoots of the winter cover crop were removed, and their fresh and dry weights were determined. Soil samples were collected, nematodes were extracted, and data were recorded as above. On 2 May 1995, peanuts cv. Florunner, were germinated on filter paper in petri dishes and six seeds were planted 5 days thereafter in each microplot. Two weeks later, the peanut plants were thinned to three plants per microplot (Fig. 6-2). Plots were irrigated as

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106 Fig. 6-2. Cut away illustration of a peanut crop in field microplots (76 cm diameter).

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107 needed using the system described above. After 123 days, peanut plants were lifted out of the ground, and the galling indices were determined and recorded for each root system, and for pods and pegs. A five-core soil sample was taken with a cone-shaped auger from the center of each plot. Data on nematode and P. penetrans population densities were collected as described above. After 5 days, peanut pods were picked, dried at 60 °C until their moisture level was reduced to about 10%, and weighed. Insect and disease control were maintained based on recommendations for Florida (Whitty et al., 1975). Laboratory bioassays. Throughout the two cropping seasons, the population density of P. penetrans was estimated by bioassay, using J2 of M. arenaria race 1 as a host. Three, 2.5-cm-diameter, 15-cm-deep soil core samples were collected with a coneshaped auger from each microplot on 21 April 1994, 21 April 1995, and 7 September 1995 (referred to as spring 1994, spring 1995, and fall 1995, respectively). The soil was screened through a sieve with 0.8-mm openings, and the moisture content was determined by drying 100 g of soil at 60 °C until weight was consistent. The soil moisture content was 6.2%, 6.5%, and 5.7% in the spring of 1994, spring of 1995, and fall of 1995, respectively. A 50-g sample of soil from each microplot was placed in a petri dish. The soil then was subjected to microwave treatment equivalent to 4 minutes/kg of soil at full power in the 18liter cavity of a microwave oven (1,500 Watts, 2,450 MHz) (Tappan Appliance, Mansfield, OH), which caused the temperature to rise to about 75 °C. About 500, 1to 4-day-old J2 were suspended in 10 ml of water and applied uniformly to the soil using a pipette. Petri plates were incubated at room temperature for 48 hours, and stored at 4 "C until further processed. Within the following 2 days, J2 were extracted by centrifugal flotation (Jenkins, 1964). The number of endospores per juvenile and the rate of attachment by P. penetrans (as averages of 20 randomly selected individuals) were determined with the aid of an inverted microscope at X400 magnification. In the summer of 1996, tomato seedlings were transplanted into 5-cm-diameter plastic pots filled with 250-g samples of

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108 soil from each microplot, which was microwave-treated at the rate of 3 minutes/kg of soil. After two days, 1,000 ± 46, 1to 4-day-old J2 of M. arenaria race 1 were suspended in 5 ml of water and added to the pots. Pots were maintained in the greenhouse for 600 degree days (18 °C base temperature); 20 female nematodes were dissected per root system, and the presence of P. penetrans endospores was determined with the aid of a microscope at XIOOO magnification. Nematode inoculum. Meloidogyne arenaria race 1 was produced on tomato Lycopersicon esculentiim Mill. cv. Rutgers in a greenhouse. For the microplot inoculation, nematode eggs were extracted with 1.05% sodium hypoclorite (Hussey, 1971) from the infected root systems (Hussey, 1973), and stored for a maximum of 3 days at 15 °C. The egg suspension then was adjusted to 4,000 eggs/ml of water, and further diluted to 500 eggs/per ml of water, resulting in 2 liters of inoculum per microplot. To test the viability of eggs used for microplot inoculation, 4,000 eggs were placed on a modified Baermann funnel and incubated at 22 °C. The hatching rates of the M arenaria race 1 were 5.5%, 18.5%, and 39% after 48, 72, and 120 hours, respectively. For the bioassays, nematode eggs were collected as described above and hatched on a modified Baermann funnel at 28 °C up to 4 days before inoculation. Statistical analysis. The density of nematodes and the numbers of endospores per juvenile were transformed with log,o(x + 1) before analysis. Data that were calculated as percentages were transformed with arcsin (Vx) before analysis. Plots were ranked and divided into 10 groups of 9 microplots each, based on the number of endospores per juvenile of M arenaria race 1 as determined by a bioassay in the spring of 1994. The change over time was determined by subtraction of the first or second data set from the final data set. The grouped data were subjected to analysis of variance (ANOVA), and group means were compared with Duncan's multiple-range test. Spearman correlation coefficients (Sokal and Rohlf, 1969) were performed on the ranks of the ungrouped data of

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109 all 90 microplots. Linear regressions were performed to determine the relationship between the initial P. penetrans density and the final nematode and endospore population densities. The linear regressions of the changes in population densities between spring and fall data of 1995 on the initial P. penetrans density were used to estimate the rate at which these changes were influenced by the initial endospore density. Results Bioassay experiments. Grouping of the ranked 90 microplots resulted in 10 groups with mean numbers of 0, 0, 1, 3, 4, 5, 7, 9, 12, and 18 endospores attached per juvenile (Table 6-1). In the spring of 1994, the mean numbers of endospores per juvenile for groups 2 to 10 were different {P < 0.001) (Tables 6-2 and 6-3), which was expected since the groups were formed by ranked data. In the following spring, the numbers of attached endospores generally decreased, especially in plots that previously yielded the highest numbers of endospores per juvenile. In the final bioassay of fall 1995, the estimated P. penetrans population density increased in all groups except group number 1, resulting in three different density levels {P < 0.05) with ranges of means of 1-6 endospores/juvenile, 23 endospores/juvenile, and 53-68 endospores/juvenile. Two groups with mean endospore counts of 45 an 40 did not differ from the intermediate group and the high group. Over two seasons, an increase between 1.1 and 5.1 endospores/juvenile for groups 1 to 3 was less than the increase between 35.1 and 55.8 endospores/juvenile in groups 5 to 10 (P < 0.05) (Table 6-3). With every endospore attached per juvenile in the spring of 1994, an average addition of 3.7 endospores/juvenile was observed over the two seasons (7= 2.4 + 3Jx) (Fig. 6-3 A). Similarly, when the attachment rate in the spring of 1994 was compared to the difference in attachment between the spring and fall of 1995, the rate of change (slope) was 2.9, which indicates that with every endospore attached in the

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110 Table 6-1. Grouping of 90 microplots based on the mean number of Pasteuria penetrans endospores attached per second-stage juvenile of Meloidogyne arenaria race 1 in a bioassay on the microplot soil conducted in the spring of 1994. Group Replicate 1 2 3 4 5 6 7 8 9 10 1 0 0 1.9 3.3 4.2 5.9 7.8 10.7 14 2 0 0 2 3.4 4.4 5.9 8 10.8 14.4 3 0 0 2 3.4 4.8 6.2 8 10.9 14.9 4 0 0 2.3 3.4 4.8 6.4 8.8 10.9 15.3 5 0 0 2.6 3.5 4.9 6.6 9 11.5 15.4 6 0 0 2.6 3.6 5 6.7 9 11.8 18.3 7 0 0 3 3.7 5.1 7 10.3 12 19.3 8 0 0 1.3 3 3.8 5.2 7 10.5 12.8 23.2 9 0 0 1.7 3.3 4.1 5.8 7.1 10.7 13.6 24.7 Group mean 0 0 1.1 2.5 3.6 4.9 6.5 9.1 11.7 17.5 Twenty randomly selected juveniles were observed per sample.

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Ill c/3 ^ I fa ^ «^ S "Si 52 C S c/3 >< -a Si 13 c « o & -2 o 111 c o i ^ O :z! Si ^ o .ts 3 «3 t) S 5 P .ti o °.2 § U c3 C <= S " a C . '-t ~ , '4^ C 6 03 4— » c/3 C/3 oa od IS c« 0 -a c 1) o .3 W CM * s * * * * * * * * * * * c/3 Z * * * * ON ON bx) c * 4ec/3 * * * * * •XON 0\ U-ON gON = UhOn" O-On 3 0\ DhON 03 C/3^C/3'-itLiC/3^P-i On C/3 z c/3 * z J CJ •I c/3 * Z * > o c 03 x: ^ CO J Z J * * * * Xn o d VI c c« o u o c 03 > O c« c/3 >% > 3 2 o C CX c/3 1) &0 c o o 03 »n o d VI a, 4-* CO 00 (L> O C 2i 4—* c 03 O c £>0 O c CO Z c _o '4— » 03 > (D cn Xi O O c c c/3 H Cl,

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112 E > o u x: U ON 15 ON On D. On On 'C o. 00 on c ID C 1) n D. 3 O o ab C3 o O) OCJ d CN >n CO no' NO in V } in o C3 X 00 NO o m CO CN d d n O CN o •n CN NO CN CN d NO NO cn NO CN O CN o O NO CN O d d d d d o d o d o bed •a T3 o bed abc abc X) CvJ abc p 00 CN NO rCN o d d d CN CN CN o o o o O o o o O 0.04 x: -a o X) a o o lO NO CTn in CN CO no" CN NO 00 CJN o a a 3 2 60 C 'oj X) (U X) 03 •a C3n ON C5J0 c c ex I/! c o •o u (A en X) -a u c u 13 C O a u e Ui O *-( oo C C3 & >^ 03 c/i IZl 173 O o •=5 > C3 O ^^ 3 T3 C C3 C o CJ u in o d VI a, C3 C -3 o c > X5 •a o o CJ c a C/2 n c 3 i 03 -4—' T3 -o !/3 u u c l-l I V c« O c 3 Q S o •a s ^ o ^ m d S,c?N O u. XI o H C/3 JJ 'c > 3 *— 1 T3 «J CJ _u s o T3 C 03 o CN CJ C S. x: V3 g D O O % ON ^ o O 60 1) "r X) S 3 N(7N OS •a — ' n o c (4-1 o -o O o 3 O Ui 60 x: CJ 03 w ^ 6 c in .3 ON On > o o 60. S C ^ 03 O, x: t/3 " dJ u x; xi ^ S
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113 144 100 64 ^ in •S ^ 36 16i' 6,400 • in OS 0^ 4,900 • 3,600 • s M VI e 2,500 • i *m e M u e (» 1,600 i i «) o soil 900 1 o 400 ; e •m 100 i M T 0-A Y = 2.37+ 3.65 X :0.43 P <0.05 1 4 9 16 Endospores/juvenfle in bioassay of spring 1994 25 Endospores/juvenile in bioassay of spring 1994 12.25 Endospores/juvenfle in bioassay of spring 1994 Fig. 6-3. Relationship between tiie initial endospore population density (estimated by the number of endospores of Pasteuria penetrans per second-stage juvenile of Meloidogyne arenaria in the spring 1994 bioassay) and A) the final number of endospores per juvenile as determined by bioassay, B) the final number of second-stage juvenile(s) (J2) per 100 cm' soil, and C) the final number of endospores per juvenile as determined by soil sample extraction.

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114 10 .5 ? «M O e 8 • 6 2i, i « 5 e e 8 : -40 S M y -50 -60 -70 Y = 0.82 -K 1.7 X r =0.55 P < 0.05 1 0.5 A At A-T , , , , , 1^ 1.5 2 2.5 3 3.5 4 45 1 Square root of endospores/juvenile ki bioassay of spring 1994 B 1 A Y = 23.37 439 X r ^ = 0.12 P < 0.05 « 1 1 1 . 1 1 1 1 0.5 ^ A 1.5 ^ 2 2.5 3 3.5 4 45 i ^ ^^TT^^^ — A ^ A A A A— 1 Square root of endospores/juvenOe n bioassay of spring 1994 50 Y= 23.31 6.12 X 0.35 P < 0.05 Square root of endospores/juvenile ii bioassay of spring 1994 Fig. 6-4. Rate of change for A) the number of endospores of Pasteuria penetrans per juvenile of Meloidogyne arenaria as determined by bioassay, B) the number of second-stage juveniles (J2)/100 cm soil, and C) the number of endospores/juvenile as determined in soil sample extraction based on the initial endospore population density (estimated by the number of endospores per juvenile in the spring 1994 bioassay) and the change over the 1995 season.

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115 bioassay of spring 1994, the change in the final estimated population density increased over the 1995 season by 2.9 endospores (Y= 0.67 + 2.9x or 7 = 0.82 + IJx after transformation with square root of x) (Fig. 6-4 A). If compared with the numbers of endospores per juvenile, the trends observed for the percentages of J2 with attached endospores were similar (Table 6-4). In the spring of 1994, the attachment by P. penetrans increased through group 5, and leveled off at 87-99% in groups 5 to 10. In the spring of 1995, 29% and 24% of the J2 in groups 1 and 2 had endospores attached, whereas only healthy J2 were recovered from this group in the previous spring. The highest rates of attachment leveled off in groups 5 to 10 with 5776% of the juveniles attached. In the final bioassay (Fall, 1995), an overall increase raised the attachment rates of the three lowest groups to 34-48%; the intermediate group four to 88%; and the six highest groups to 98-100% (P < 0.05). The three lowest groups had lesser increases in attachment over one season than the seven highest groups. However, no difference was observed when the change over both seasons was considered. Microplot soil populations. Under the com-bean intercropping systems in rotation with peanut, the M. arenaria densities and P. penetrans endospore densities, as well as the galling indices were affected by the pretreatment endospore densities in the microplot soil at each sampling date {P < 0.05, P < 0.001) (Table 6-2). No differences were noted in the nematode population densities in spring 1995, as well in the population developments of the nematode and the bacterium and change in galhng indices over one season. Over the two seasons, the endospore attachment, nematode population densities and galling indices were affected by the different endospore densities in the microplots {P < 0.001). During the first season of two consecutive com and bean intercropping systems, the nematode population density increased in the fall of 1994 to an average (across all groups) of 1,838 J2/100 cm^ of soil (Table 6-5). In the spring of 1995, the nematode

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116 •So ^ ^ (j_ o o 5 o M _, o C/3 — _, — * I c St: 2 CI I § u .Si ^ C. Oh U. I « S " ^ -3 <^ On I ^ ^ ON , 2i 3 _ -C| ^ ^ O >l i-i u a c o o (U 1/3 o "3 -n -C w iH c -J t; ca c3 (u < ^ X) H c -o P c i: ^ c K S ^ O > o u jC u in 0\ On 0\ .. ON O C — I C CO ON On 60^ 'C 00 oo c CM 3 O o c3 03 03 C3 03 00 m ON X) XI X) 03 ca 00 CO 00 r~CN in cn m CN CN m CO W CN r o o O o CN CO m o o o x> X) c3 03 03 o3 a 03 00 00 00 00 o Q m 00 On o O O o o 00 O o NO o o o X) X) 03 03 OS ON r~CN NO CN m CN (N cn >n NO NO NO o O CN (N 00 NO CN CN CO CN o Xi ab 03 o3 03 o o ro ON NO On On ON On CO NO 00 ON ON ON ON On ^ CN >n NO 00 ON O oo C/3 ^ o 2^ C C ON

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117 si u o o 2 o 5i .a s u <*o St 8 e/! — C C i E .« 8 a o •2 jc o\ 2 H IVI ^ B o > c x: C/3 3 1/3 n 1/3 C/3 ^ C D C U O O 1/3 (yl > IS ^ C D 1) o. O 3 els ^ c St ^ as VO 3 < 1"^ 13 S3 1) > o D c U IT) as as C3 o. 00 CN O C/2 CM 3 O 173 a C3 03 ,799 ,995 ,682 ,252 ,618 -1,702 ,516 .555 OO CN q_ c o X3 CJ o o CJ >n o C<1 m 'It Ov Tl<3v ON "t CN VO c<^ 00 CO ON CN .348 00 00^ <7v WO vq_ 00 00 O VO CN WO VO wo 00 cn c^ CN CN p_ CN CN CN oo' CN o' CN fo' CN CN d" CO CN o' wo wo" VO c3 a a ca C3 ,692 ,426 ,583 VO CN r~; CN vq_ VO 00 CO uo CN CO CN CN CN CO CN CN ,806 ,835 ,865 CO CN 00 o WO o VO^ CO 00^ CO vq^ VO c^ C<0 cn as VO' CN rn CO CN CN CN oo" ab XI d X) X) XJ X) X3 X) X) 2,312 2,425 2,627 1,446 1,709 1,726 1,552 1,593 2,047 1,544 CN wo VO 00 ON o < > o z < o o 3 U3 •a c 03 •a D Oh 3 O Ml e (U X) 3 CL, 1/3 E o c« o Id c Ul U X) S 3 C (U X! C O u c« 03 X) •o c Ul x: & 0*3' _o u Ul U c c« 'C Oi S o o c/3 u^Os 1bO "*H x: ° O bO 03 C W u -o Dc/3 U x: u c • ^ •4— t 03 o c/3 2 o c o u c/3 o o o o <2 o ^ CliWO u On C <4-C u o , iS u :S > 6 o o bo 03 On X! On O ^ D '4-1 J= O H bo u c • ^ Ul CL 00 lU Xi

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118 population density further increased to an average of 2,750 J2/100 cm^ of soil, and did not differ among the groups (P < 0.05). In the fall of 1995, the nematode population density declined drastically, ranging from the low levels of 12 to 194 J2/100 cm' of soil in groups 4 to 10, to the high levels of 430-945 J2/100 cm' of soil in groups 1-3. The nematode reduction over one season in groups 6, 7, and 9 was greater than in groups 1 and 5 (P < 0.05). There were no differences in the change over two seasons. With every endospore attached per juvenile in the spring 1994 bioassay, a decrease of 31.5 J2/100 cm' of soil was observed over the two seasons {Y= 547.1 31. 5x) (Fig. 6-3 B). Similarly, when the attachment rate in the spring of 1994 was compared to the difference in number of J2 between spring and fall of 1995, the rate of change (slope) was -19.3, which indicates that with every additional endospore attached in the bioassay of spring 1994, the suppression in the nematode population density over the 1995 season increased by 19.3 J2 (Y= -546.2 19.3x or y = -23.4 4.4x after transformation with square root of x) (Fig. 6-4 B). In the fall of 1994, the estimated number of endospores per juvenile in the microplot soil ranged from the lowest level of 0-1 in groups 1 to 4 to the highest level of 8.6 in group 10 (P < 0.05) (Table 6-6). Over one season, the change in the number of endospores was not different among groups, and in the final sample only group 10, with the highest attachment, showed a greater decrease in the number of endospores than the other nine groups. With every additional endospore attached per juvenile in the bioassay from the spring of 1994, an increase of 0.1 endospores/juvenile was observed in the soil analysis over the two seasons (Y= 0.3 + OAx) (Fig. 6-3 C). However, when the attachment rate in the spring of 1994 was compared to the difference in attachment between spring and fall of 1995, the rate of change was -37.5, which indicates that with every endospore attached in the bioassay from the spring of 1994, the decrease in the final estimated endospore population density in the soil increased over the 1995 season by 37.5

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119 (/2 1) C XI u > .2 x; ^ 2 o ^ :^ rS c -a u c '7? S lyj o •a c O cj 1) O OS "PC o E (u Si f. St VI >-' « C 3 i: U C ^ s ^ § ?^ O = " c 5 z ^ 4 ^ -s <^ ^ c a —• o m S u H o p U J3:S o > o C x: U in OS OS a on c/: C O U c o 1/3 c« U 00 c C3 C3 CO C3 C3 X) m OS m so d d d d d d 1 d d d SO c3 C3 CO cd n 00 d d d o d d d d CN 1 o so CN CN so >n m 00 o 00 o d d d d d d d o d O X> o x> OS cd cd (N m OS OS sq OS sq CN d d d CN CN OS ^1 V N o m 00 SO CN d d d d d d O d o o o u X2 X) X3 C3 cd cd o o o SO (N d oi CN 4 ^ ID O CN o c cd O oo — u u X o < > O Z < -o 2 c/; .Si, >,-° X' 3 S " a S c =^ Cd T3 (U O Cl. (N 3 , O (A l-H (U bO E.S cd
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120 endospores/juvenile (Y= 543.4 31. 5x or 7= 23.31 6.12j: after transformation with square root of ;c) (Fig. 6-4 C). Soil samples taken in the fall of 1994 revealed attachment rates ranging from 310% at the lowest levels in groups 1 to 3, and 62-89% at the highest levels in groups 4 to 10 (P < 0.05) (Table 6-7). The attachment rates were similar throughout the remainder of the experiment. However, the change over two seasons was different between groups 1, 2 and 3, for which attachment increased by 2-15%, and group 10, which had a 18% decrease in the attachment of endospores to J2 (P < 0.05). In both consecutive com and bean intercropping systems of the 1994 season, a variable degree of root galling was observed on both plants (Table 6-8). Root galling was generally higher in groups 1, 2, and 3 than in the other groups (P < 0.05). Pods and pegs of the peanut plants were equally well protected from galling; galling indices between 0 and 1.2 for groups 4 to 10 were lower than the indices of 2.8-3.1 recorded for groups 1 to 3. No differences were observed among galling indices between the two com and bean intercrops. Over two seasons of com and bean and then peanut, the galling in groups 6 to 10 was reduced by 2.1 to 2.6 units, which was higher than the reduction by 0.3 to 1.1 units of groups 1, 3, 4, and 5. The rank of the numbers of endospores per juvenile, and the rates of attachment recorded from the bioassays were positively correlated to the rank of the same observations collected by soil sample extraction (Table 6-9) (P < 0.05). Thus, the estimated endospore density determined by the bioassay was representative for the estimated density in the microplot soil. However, the ranks of all endospore counts, as well as attachment rates, were negatively correlated to the ranks of the nematode population densities and galling indices, showing that the nematode population density was negatively correlated with the endospore density. Only the ranks of the nematode population density of spring 1995

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121 'n o •S V» ON s o\ s ° u X) o 5 o 6 o o —I o o o 2 ^ •c :§ C u ^ ^ .S ^ D C E 3 I/; O _ 00 c _ (U D. o o o s " 's =r a. -a ^ box; o\ C go f2 2 > o U in On 60 C 'C CM 00 ON On C3 tin (U C/3 03 CM 3 O O o o n in in in m m o X3 X) 00 (N ca NO ea On NO NO CN in NO
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122 C/3 _o "5. o On b ^ c 2 c ^ 4—* '4— » O (A C 1) I -§ O 5 8 .s «5 1/2 U o c a ^ g 5 C3 00 U c 00 I U -J •3 •a a, s 2 > O (U M C a jc U (4-. C o c o u '-' C „ «-> C3 S > s ^ o -a 03 C a ON , 03 04 o ^ 03 C ^ 8 00 On is o U 03 -a o (X 03 U OQ o U O O ab be 03 ab f 1 w o o OCJ m ON NO 1 d 1 d d CN CN 1 (N CN 1 03 03 03 03 o! 03 o! NO CN CN o ON 00 q d d d d d d 03 03 03 03 03 03 03 0! 03 o 00 ON NO d d d d d d d d d 03 03 o3 X X X) X) X X 00 On CN o CN o o CN CN f^ X) T3 o 03 Vi 03 O o :3 := c "2 i 03 00 3 o u c X. *-» 3 O 00 (D CX, C/3 4— » b 03 .^ "-I U I C U O 3 DS 2 ^ 03 O |2 aj two t« .s O O O CJ c3 aJ O O O o o 00 II m d" oo NO m CN d" CN II CN •S O r;2 -C VI o cx, X) (U IJ ' o c 03 x; CJ 'x. 03 CJ CJ c •a flj aS o o 2^' 00 On pq 00 " .S CJ c J=> CS + D ^ X •o c 03 ^ O On CJ OS Ui '~' ,0 "^^ O V —I S 13 «j ^ > c O 03 00 S «J o X H e o o c o • — H -4—) o Ui Xi 3

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123 t3 ^ c ^ u, O C« y Uh 4) ^ (U .S E E U 3 U > j=; >> • ' C C3 JO v: p (D Oh 6 o .is T3 C C/3 03 O IS s o i3 •a 3 5 2 >^ c ^ > V bC C o c B cj --2 U C — ^ c o> as S O ON O -J CO < a, c3 o u X3 •a c 03 S g E ° S c; o I o o c a 60 O o o C/2 c U P3 o U 0) .QS SirSis ON L» in c On 0 s x: 0 < o 10 I O^ loo ON I I ON l\0 10 I On in >n , o\ m 10 ON (N O Id VI c 03 O o o — (N vO o" o c o r(N (N CN m NO o d d — r~ ts ro (Nl NO (N CN i-S (N o Al o _ <^ ^ ^ ^. c/3 00 o o cn en d d d d tN CS 00 0\ O O IN 00 (30 — cn "n m — VO Tl»0 NO d d o o o o .5 -t-l u o o c o 00 m fs (N CS r~m NO >n in d d d o d c5 d d o — inooinr~oooNooNONO f^^f^df^NO-^j-mNONO o o 000000 NOfNON — oooooNONr~in NDvqp-^cscNinm — -^i-in dddddddddd NO NO NO rd °o d o o >n CNi m in ON T}CO NO d d d d d 03 T3 U •4— » |<4-l o — ren r\0 VO '^^0'~^f~^'^t~^NOTl-TtVONO|lJ 00 o d d d d d d d d \s: D 03 s o o ^-;NOlnr-;NO\oc<^ — vomcNNOvo c>dddddddddddd o o ^ d 00 NO NO >n ^o p~ d d d 00 cN] in NO n r~nt~;ininNOT:l-in d d d d d d d d oor-o\NOcNioovo — 00 inj-joovoNOr^r-Nor^ d d d d d d d d r— r~00 d d (N On NO d d NO NO v£j d d On vo in o\ tN r\0 — ^ NO rr~ d d d d d d d fo ON o\ in in 00 <^ O >n ^ n O — CI m vo in I d d d d d d c V 'o IS u o o c _o s o o 1) » o &, in ON ON -a c ON ON W) c 'C D T3 £U3 C I 03 00 •a c 03 Ml 6 03 to c ex «5 E — csc
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124 showed a random relationship with those of the attachment rates and the endospore densities of the spring data of 1994, as determined by the bioassays. Thus, the initial endospore density had no effect on the nematode population of the following spring. Discussion The nematode population became well established under the com and bean intercropping systems through 1994, and remained at equally high levels in all microplots under the susceptible winter cover crop. No conclusions can be drawn about the effect of the intercropping system on the population development of P. penetrans in the soil, since no altemative treatment was included. During the microplot sampling in the spring of 1995, 14% to 97% of the J2 escaped attachment by P. penetrans endospores, leaving at least 343 J2 free of P. penetrans! \Q0 cm^ of soil for each group. This density of nematodes would be capable of causing 90% yield loss in peanut (Porter et al., 1984). However, nematode populations of the five groups with > 4.9 endospores per juvenile and an attachment rate of > 96% in the standard bioassay showed a decline in the incidence of galling, and the peanut crop was nearly free of galls. In contrast to the two fall samples, the high numbers of J2 in the spring of 1995 were not related to endospore counts and attachment rates in the bioassays. Therefore, in winter of 1994-95, the nematode population did not seem to be affected by the P. penetrans population density in the soil. This observation contradicts the results of other scientists (Oostendorp et al., 1991a), in which, despite the high temperature requirement of P. penetrans (Hatz, et al., 1992 ; Serracin et al., 1997; Stirling, 1981), endospore attachment increased, and nematode populations decreased during the winter

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125 season (Oostendorp et al., 1991a; Chapter 2). The P. penetrans population in the microplot soil may have been unable to reproduce on the predominant nematode host. In this experiment, the endospore population development determined by bioassays was not consistent with the endospore population development determined by soil sample extraction. A 3.7and a 0. 1 -fold increase in the number of endospores per juvenile was estimated with the aid of bioassays or soil sample extraction, respectively. A possible explanation might be given by a population shift to Meloidogyne spp. other than M arenaria that occurred by cultivating the root-knot nematode susceptible pole bean (Mullin et al., 1991). It is possible that M. javanica re-established in the microplots from residual populations of previously conducted experiments (Dickson, personal communication). The P. penetrans population present in the microplots attached rather poorly to M. javanica (Oostendorp et al., 1990). This could explain the good endospore attachment toM arenaria in the bioassays, the poor attachment rates observed in soil samples during the com and bean intercrop (if M. javanica predominated on bean), and the suppression of Meloidogyne spp. other than M. arenaria in the course of the following peanut crop. Unfortunately, the specification of the nematode population was not determined. It is also possible that in unperturbated soil, endospore attachment was affected by other soilbome microorganisms. In related studies, as these microorganisms were selectively eliminated or reduced by the microwave treatment (Chen, et al., 1995; Ferriss, 1984; WeibelzahlFulton et al., 1996; Chapter 3), the number of endospores per juvenile increased. After 2 years, the soils of seven groups with an initial mean number of 2.5 and more endospores per juvenile and a rate of attachment of 69% or higher developed to highly suppressive soils in which 88% or more of J2 were encumbered with 23.5 or more endospores. Pasteuria penetrans is known to suppress root-knot nematodes by affecting juvenile mobility and reducing or inhibiting female fecundity (Stirling, 1991). Mobility

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126 and infectivity of J2 is reduced when between 7 and 50 endospores are attached (Brown and Smart, 1985; Davies et al., 1988; 1990; Sell and Hansen, 1987; Stirling, 1984), and juveniles are prevented from invading roots when they are encumbered with 25 to 30 endospores (Stirling, 1984; Stirling et al., 1990). Previous experiments have shown that root systems can be protected from juvenile invasion by high numbers of P. penetrans endospores in the soil (Chen et al., 1996b; 1997; Weibelzahl-Fulton et al., 1996; Chapter 3). Endospores of P. penetrans are relatively resistant to heat, desiccation, nematicides, and other adverse environmental conditions (Stirling, 1991). The effect of methyl bromide (98% methyl bromide plus 2% chloropicrin) on the survival of the bacterial endospores is unknown at this time. Although the fumigation of soil containing P. penetrans with methyl bromide had little or no effect on the attachment of endospores to the J2 in a greenhouse bioassay, P. penetrans did not complete its life cycle with the formation of mature endospores (personal observation). This observation is in agreement with Freitas (1997), who reported that chloropicrin alone or in combination with methyl bromide was highly detrimental to P. penetrans because endospore formation was inhibited. Hence, the observed increase in endospore attachment in the present study may have resulted from the continuous release and dispersion of endospores from decomposing root material rather than from an endospore build-up following soil fumigation. The nematode population was generally lower in groups with an initial number of 2.5 or more endospores of P. penetrans per juvenile. Since methyl bromide treated endospores of P. penetrans appeared to be unable to reproduce (Freitas 1997), the nematode suppression observed in this experiment most likely resulted from the prevention of root invasion by endospore encumbered juveniles. Endospore attachment is not necessarily followed by nematode infection (De Silva and Gowen, 1994; Freitas,

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127 1997). In this experiment, the host-parasite development was limited to cuticular attachment only, similar to several previous observations (S tiding, 1991). Future scientists must exercise caution and actually determine parasitism before making conclusions about the host-parasite relationship. • It can be concluded that microplots with a certain P. penetrans population density remained in the original population density group through the course of the 2-year experiment. This observation may have been influenced by the effect of methyl bromide on the survival of the endospores in the soil; nonetheless, these density classes remained remarkably intact.

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CHAPTER 7 SUMMARY Since environmental problems have been associated with the application of nematicides (Thomason, 1986), their use has led to great controversy, and increasing attention is being given to alternative management tactics, such as cultural practices, plant resistance and biological control of nematodes with microbial agents such as Pasteuria penetrans (Thome) Sayre & Starr. The potential of P. penetrans in combination with cultural practices to suppress Meloidogyne arenaria on peanut (Arachis hypogaea L.) and Meloidogyne spp. on tobacco {Nicotiana tobacum L.) was investigated in laboratory, greenhouse, microplot, and field experiments. A suppressive-soil test was developed to determine the role of P. penetrans in suppressive soils. In 1987, before the initiation of the present study in 1990, 90 microplots were either infested with M. arenaria, or with M. arenaria and P. penetrans, or maintained nematode free by periodic fumigation (Oostendorp, 1991a). Rye, vetch or wheat, or bare fallow was maintained for winter cover. The population development of P. penetrans and M. arenaria was monitored for the last 3.5 years (1990-1993) of the 6.5-year study. In the fifth, sixth, and seventh cropping season, nematodes were nearly eliminated from plots with the initial P. penetrans inoculum. The peanut yields did not differ from those in fumigated control plots. By the end of the 7-year study, P. penetrans had spread to all plots containing nematodes, which caused an increasing portion of the population of second-stage juveniles (J2) to become encumbered with endospores, and, hence. 128

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129 detrimentally affected. The increase in the estimated number of endospores in the soil was enhanced under susceptible vetch or wheat cover crops. The effect of microwave treatment on soil fungi and bacterial endospores was determined using dilution plating and soil bioassay. An increase in microwave radiation timeand soil water content revealed an increasing detrimental effect on P. penetrans and other soil organisms. A treatment time of 3 minutes/kg of soil at field capacity of 7% soil water reduced fungal populations but left the attachment by P. penetrans nearly unaffected. This microwave treatment was helpful because it reduced fungi that might also be a factor in suppressive soils. In a suppressive-soil test, microplot soil was autoclaved, microwaved, or air dried and bioassayed in the greenhouse using M. arenaria on peanut as a host for potential biological control organisms. The suppressive component was inhibited by autoclaving soil, and this treatment resulted in heavy galling and egg mass formation on the root system. Microwaving or air-drying soil did not affect the suppressiveness of the soils inoculated or contaminated with P. penetrans. Endospores in both soils existed in densities at which infectivity of J2 was reduced and the root systems were protected from severe root-knot infection. Pasteuria penetrans was identified as the main contributor to the suppressiveness of the microplot soils. The suppressive-soil test was a useful tool in the determination of the suppressive component in the soil. However, the effects of the microwave treatment on soil organisms fluctuated with different soil characteristics, and requires additional experimentation. A mixed population of root-knot nematodes was observed to infest a tobacco field from which Pasteuria penetrans and several species of potential nematophagous fungi were isolated (Chen et al., 1994). This field site was planted to two tobacco cultivars (Coker 371, which is susceptible to M. incognita, M. javanica, and M. arenaria; and

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130 Northrup King-326, which is resistant to M. incognita but susceptible to M. javanica and M. arenaria), followed by three autumn cover crops (hairy indigo, forage sorghum, and weeds). It was fertilized with two rates of inorganic nitrogen fertilizers (89 and 158 kg of ammonium nitrate/ha). The nematode suppression increased over 2 years, and P. penetrans was identified to be suppressive to the root-knot nematodes. The endospore build-up in the soil was slightly favored by a cropping history with the susceptible tobacco cultivar. The preferred host for the two isolated P. penetrans populations appeared to be M. incognita over M. javanica. Inorganic nitrogen and winter cover crops had no consistent effects on the endospore build-up in the soil. When the tobacco field soil was subjected to a suppressive-soil test using M. incognita and tomato as hosts for potential nematode antagonists, P. penetrans was identified as the main contributor to the suppressiveness of the microplot soils. However, when the experiment was replicated, soilbome fungi contributed to the nematode suppression. Thus, the role of fungi in the nematode suppression could not be elucidated. Ninety microplots were ranked and grouped based on the number of endospores per juvenile of M arenaria as determined by a bioassay at the beginning of the experiment. The population development of Meloidogyne spp. and P. penetrans was monitored over 2 years, planted with two cycles of a com and bean intercrop in rotation with peanut. The soil of groups with an initial mean number of 3 and more endospores per juvenile and a rate of attachment of 69% or higher developed highly suppressive soil in which 88% or more of J2 were encumbered with 23 or more endospores. In a greenhouse bioassay, P. penetrans was not able to complete its life after soil with endospores was treatment with methyl bromide (98% methyl bromide plus 2% chloropicrin). The current studies revealed that the continuous cultivation of crops susceptible to root-knot nematode for 3 to 4 consecutive years increases levels of P. penetrans

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131 endospores in the soil from relatively low to highly suppressive levels. In the microplot environment, the suppressiveness was maintained in the absence of nematodes for a period of 3 years before the experiment was terminated. Efforts should be directed toward the amplification of endospores in namre and in artificial mass production systems.

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133 Barker, K. R. 1993. Resistance/tolerance and related concepts/terminology in plant nematology. Plant Disease 77:1 1 1-1 13. Barker, K. R., F. A. Todd, W. W. Shane, and L. A. Nelson. 1981. Interrelationship of Meloidogyne species with flue-cured tobacco. Journal of Nematology 13:67-79. Barker, K. R., J. L. Townshend, G. W. Bird, I. J. Thomason, and D. W. Dickson. 1986. Determining nematode population responses to control agents. Pp. 283-296 in K. D. Hickey, ed. Methods for evaluating pesticides for control of plant pathogens. St. Paul, MN: APS Press. Barooti, S. 1989. Distribution of Pasteiiria penetrans, a parasite of nematodes in Iran. Iranian Joumal of Plant Pathology 25:9-10. Barron, G. L. 1992. Lignolytic and cellullolytic fungi as predators and parasites. Pp. 31 1326 in G. C. Carroll and D. T. Wicklow, eds. The fungal community: Its organization and role in the ecosystem. New York: Marcel Dekker. Baxter, R. I., and C. D. Blake. 1969. Oxygen and hatch of eggs and migration of larvae of Meloidogyne javanica. Annals of Applied Biology 63: 191-203. Bennett, F. D. 1974. Criteria for determination of candidate hosts and for selection of biotic agents. Pp. 87-96 in F. G. Maxwell and F. A. Harris, eds. Proceedings of the Summer Institute on biological control of plant insect disease. Jackson, MS: University Press of Mississippi. Bernard, E. C, and T. L. Niblack. 1982. Review of Hoplotylus S. Jacob (Nematoda: Pratylenchidae). Nematologica 28:101-109. Bhattacharya, D., and G. Swamp. 1988. Pasteuria penetrans, a pathogen of the genus Heterodera, its effect on nematode biology and control. Indian Joumal of Nematology 18:61-70. Birchfield, W., and A. A. Antonopoulus. 1976. Scanning electron microscopic observations of Dubosqia penetrans parasitizing root-knot larvae. Joumal of Nematology 8:272-273. Bird, A. F. 1986. The influence of the actinomycete, Pasteuria penetrans, on the hostparasite relationship of the plant-parasitic nematode, Meloidogyne javanica. Parasitoloev 93:571-580. Bird, A. F. 1988. A technique for staining the endospores of Pa^reifna ;7^«e/ra/z.y. Revue de Nematologie 11:364-365. Bird, A. F., I. Bonig, and A. Bacic. 1989. Factors affecting adhesion of micro-organisms to the surface of plant-parasitic nematodes. Parasitology 98: 155-164. Bird, A. F., and P. G. Brisbane. 1988. The influence of Pasteuria penetrans in field soils on the reproduction of root-knot nematodes. Revue de Nematologie 1 1:75-81.

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134 Bird, A. F., P. G. Brisbane, S. G. McClure, and W. L. Kimber. 1990. Studies on the properties of the spores of some populations of Pasteuria penetrans. Journal of Invertebrate Pathology 55:169-178. Bishop, A. H., and D. J. Hilar. 1991. Attempts to culture Pasteuria penetrans in vitro. Biological Science Technology 1:101-114. Brown, S. M., J. L. Kepner, and G. C. Smart, Jr. 1985. Increased crop yields following application of Bacillus penetrans to field plots infested with Melodogyne incognita. Soil Biology and Biochemistry 17 :483-486. Brown, S. M., and D. Nordmeyer. 1985. Synergistic reduction in root galling by Meloidogyne javanica with Pasteuria penetrans and nematicides. Revue de Nematologie 8:285-286. Brown, S. M., and G. C. Smart, Jr. 1984. Attachment of Bacillus penetrans to Meloidogyne incognita. Nematropica 14:171-172. Brown, S. M., and G. C. Smart, Jr. 1985. Root penetration by Meloidogyne incognita juveniles infected with Bacillus penetrans. Journal of Nematology 17: 123-126. Bunt, J. A. 1987. Mode of action of nematicides. Pp. 461-468 m J. A. Veech and D. W. Dickson, eds. Vistas on nematology: A comemorandum of the twenty-fifth anniversary of the Society of Nematologists. Hyattsville, MD: Society of Nematologists. Cappuccino, J. C, and N. Sherman. 1986. Microbiology, a laboratory manual. Menlo Park, CA: Benjamin/Cummings Publishing. Channer, A. G., and S. R. Gowen. 1988. Preliminary studies on the potential of Pasteuria penetrans to control Meloidogyne species. Proceedings of Brighton Crop Conference, Pest and Disease. Surrey, UK: The British Crop Protection Council. Channer, A. G., and S. R. Gowen. 1992. Selection for increased host resistance and increased pathogen specificity in the Meloidogyne-Pasteuria penetrans interaction. Fundamental and Apphed Nematology 15:331-339. Chamecki, J. H. 1997. Pasteuria penetrans spore proteins: Potential function in attachment to Meloidogyne spp. M.S. thesis. University of Florida, Gainesville. Chamecki, J. H., J. F. Preston, and D. W. Dickson. 1996. Fluorescent labeled endospores of Pasteuria penetrans for the characterization of the proteins involved in attachment to root-knot nematodes. Abstracts of the 96th general meeting of the society of microbiology. Washington, DC: American Society of Microbiology. (Abstr.). Chen, S., J. Charnecki, J. F. Preston, D. W. Dickson, and J. D. Rice. 1997a. Antibodies from chicken eggs as probe for antigens from Pasteuria penetrans endospores. Joumal of Nematology 29:268-275.

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135 Chen, S., D. W. Dickson, and D. J. Mitchell. 1995. Effect of soil treatment on the survival of soil microorganisms. Journal of Nematology 27:661-663. Chen, S., D. W. Dickson, and E. B. Whitty. 1994. Response of Meloidogyne spp. to Pasteuria penetrans, fungi, and cultural practices in tobacco. Supplement to the Journal of Nematology 26:620-625. Chen, Z. X. 1996. Biological control potential of Pasteuria penetrans. Ph.D dissertation. University of Florida, Gainesville. Chen, Z. X., and D. W. Dickson. 1997. Estimating incidence of attachment of Pasteuria penetrans endospores to Meloidogyne spp. with tally threshold. Journal of Nematology 29:289-295. Chen, Z. X., D. W. Dickson, L. G. Freitas, and J. F. Preston. 1997b. Ultrastructure, morphology, and sporogenesis of Pasteuria penetrans. Phytopathology 87:273-283. Chen, Z. X., D. W. Dickson, and T. E. Hewlett. 1996a. Quantification of endospore concentration of Pasteuria penetrans in tomato root material. Journal of Nematology 28:50-55. Chen, Z. X., D. W. Dickson, R. McSorley, D. J. Mitchell, and T. E. Hewlett. 1996b. Suppression oi Meloidogyne arenaria race 1 by soil applications of Pasteuria penetrans. Journal of Nematology 28:159-168. Chen, Z. X., D. W. Dickson, D. J. Mitchell, R. McSorley, and T. E. Hewlett. 1997c. Suppression mechanisms of Meloidogyne arenaria race 1 by Pasteuria penetrans. Journal of Nematology 29: 1-8. Cho, M. R., D. W. Dickson, and T. E. Hewlett. 1997. Comparison of inoculation methods and different host plants for production of Pasteuria. Journal of Nematology (in press) (Abstr.). Ciancio, A. 1995. Density dependent parasitism of Xiphinema diversicaudatum by Pasteuria penetrans in a naturally infested field. Phytopathology 85:144-149. Ciancio, A., R. Bonsignore, N. Volvas, and F. Lamberti. 1994. Host record and spore morphometries of Pasteuria penetrans group parasites of nematodes. Journal of Invertebrate Pathology 63:260-267. Ciancio, A., and M. Bourijate. 1995. Relationship between Pasteuria penetrans infection levels and density of Meloidogyne javanica. Nematologia Mediterranea 23:43-49. Ciancio, A., S. Landriscina, and L. Scrano. 1987. Indagini sulla nematofauna degli agrumi nell'Italia Meridionale. Informatore Fitopatologico 37:55-57. Ciancio, A., and R. Mankau. 1989a. Note on Pasteuria sp. parasitic to longidorid nematodes. Nematropica 19:105-109.

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137 Davies, K. G., V. Laird, and B. R. Kerry. 1991. The motility, development, and infection of Meloidogyne incognita encumbered with spores of the obligate hyperparasite Pasteuria penetrans. Revue de Nematologie 14:611-618. Davies, K. G., M. Redden, and T. K. Pearson. 1994. Endospore heterogeneity in Pasteuria penetrans related to adhesion in plant-parasitic nematodes. Letters in Applied Microbiology 19:370-373. Davies, K. G., M. P. Robinson, and V. Laird. 1992. Proteins involved in the attachment of a hyperparasite, Pasteuria penetrans, to its plant-parasitic nematode host, Meloidogyne incognita. Journal of Invertebrate Pathology 59: 18-23. Davis, E. L., and J. R. Rich. 1987. Nicotine content of tobacco roots and toxicity to Meloidogyne incognita. Journal of Nematology 19:23-29. De Leij, P., K. G. Davies, and B. R. Kerry. 1992. The use of Verticillium chlamydosporium Goddard and Pasteuria penetrans (Thorne) Sayre and Starr alone and in combination to control Meloidogyne incognita on tomato plants. Fundamental and Applied Nematology 15:235-242. De Silva, M. P., and S. R. Gowen. 1994. Attempts to adapt a population of Pasteuria penetrans originating from Meloidogyne javanica to its less susceptible host M. arenaria. Afro Asian Journal of Nematology 4:40-43. Dickson, D. W., M. Oostendorp, R. M. Giblin-Davis, and D. J. Mitchell. 1994. Control of plant-parasitic nematodes by biological antagonists. Pp. 575-601 in D. Rosen, F. D. Bennett, and J. L. Capinera, eds. Pest management in the subtropics. Andover, UK: Intercept. Dickson, D. W., and F. B. Struble. 1965. A sieving-staining technique for extraction of egg masses of Meloidogyne incognita from soil. Phytopathology 55:497. (Abstr.). Domingues, H. E., D. D. Baltensperger, P. E. Reith, and R. A. Dunn. 1985. Genotypic variation in Indigofera hirsuta L. reaction to Meloidogyne spp. Soil and Crop Science Society of Florida Proceedings 45:189-192. Dos Santos, J. M. 198 1 . Occurrence of Bacillus penetrans parasitizing Meloidogyne javanica in Brazil. Fitopatologia Brasileira 6:519-522. Dropkin, V. H. 1969. The necrotic reaction of tomatoes and other hosts resistant to Meloidogyne: Reversal by temperature. Phytopathology 59:1632-1637. Dube, B., and G. C. Smart, Jr. 1987. Biological control of Meloidogyne incognita by Paecilomyces lilacinus and Pasteuria penetrans. Journal of Nematology 19:222-227. Dutky, E. M., and R. M. Sayre. 1978. Some factors affecting infection of nematodes by the bacterial spore parasite Bacillus penetrans. Journal of Nematology 10:285.

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APPENDIX RING NEMATODE DATA FOR CHAPTER 2 Table A-1) Effect of Meloidogyne arenaria (RKN) alone and in combination with Pasteuria penetrans (RKN + Pp), and of a rye, vetch, or bare fallow winter cover crop on the ring nematode population density in the falls of 1991 to 1993 of a 6.5-year microplot experiment. Ring nematodes in 100 cm soil Treatment 1991 1992 1993 Main effect means Nematode Control 0.33 b 14.2 0.6 b RKN 76.6 a 9.8 251.2 a RKN + Pp 95.8 a 21.2 205.8 a Cover Crop Rye 33.4 b 3.4 b 47.1 b Vetch 50.5 b 4.7 b 91.6b Fallow 88.9 a 37 a 318.6 a Interactions Rye Control 0.2 2.1 0.5 b RKN 28.1 4.9 90.4 a RKN + Pp 71.8 3.3 50.4 a Vetch Control 0.1 7.5 0.4 b RKN 91 2.7 74.5 a RKN + Pp 60.3 4 200.9 a Fallow Control 0.7 33 0.8 c RKN 110.7 21.7 588.8 a RKN + Pp 155.3 56.2 366.2 b Means followed by no letter or the same letter within a group of three observations are not different according to Duncan's multiple-range test {P < 0.05). Means of 10 or 30 replicates (for main effects), with 20 second-stage juveniles (J2) per replicate. 151

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BIOGRAPHICAL SKETCH I was born on September 4, 1963, to Mrs. Anna Heyl and Mr. Arniin Willi Paul Weibelzahl in the district of Steglitz, Berlin, Germany. I received my elementary and middle school education from the local public school. In 1981, 1 completed a 2-year apprenticeship in horticulture and entered the College of Horticulture in Berlin-Zehlendorf. In 1982, 1 continued my education at the Polytechnics in Berlin-Dahlem, where I obtained the bachelor's degree of horticultural engineering in January 1987. After a period of voluntary work in the Department of Plant Physiology at the Volcani Center in Bet Degan, and the Department of Agriculture in Tel Aviv, Israel, I was attracted to the international approaches of the Department of Agriculture at the University of Reading, UK. Intensive studies in Reading and a nematological field project in Barbados, West Indies, helped me to obtain a Master of Science degree in December 1990. I came to the University of Florida in August of 1991, and have since worked in the laboratories and fields of Don W. Dickson mainly on the biological control of root-knot nematodes. In 1994, my advances were recognized by the Gamma Sigma Delta Honor Society of Agriculmre. In July 1993, I was fortunate to wed my husband, Michael Rory Fulton. Since then I became mother of two wonderful children. 152

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality as a dissertation for the degree of Doctor of Philosophy. D. W. Dickson, Chair Professor of Entomology and Nematology I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality as a dissertation for the degree of Doctor of Philosophy. R. McSorley Professor of Entomology and Nematology I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality as a dissertation for the degree of Doctor of Philosophy. D. J.^itchell Professor of Plant Pathology

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality as a dissertation for the degree of Doctor of Philosophy. Professor of Agronomy This dissertation was submitted to the Graduate Faculty of the College of Agriculture and to the Graduate School and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy. May 1998 ^Oean, College of Agriculture Dean, Graduate School