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Biochemical events during the development of Pasteuria penetrans within the pseudocoelum of Meloidogyne arenaria

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Biochemical events during the development of Pasteuria penetrans within the pseudocoelum of Meloidogyne arenaria
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Brito, Janete Andrade, 1957-
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English
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xv, 148 leaves : ill. ; 29 cm.

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Subjects / Keywords:
Antibodies ( jstor )
Endospores ( jstor )
Epitopes ( jstor )
Nematology ( jstor )
Parasites ( jstor )
Pasteuria ( jstor )
Root knot nematodes ( jstor )
Roundworms ( jstor )
Species ( jstor )
Sporulation ( jstor )
Dissertations, Academic -- Entomology and Nematology -- UF ( lcsh )
Entomology and Nematology thesis, Ph.D ( lcsh )
Peanut root-knot nematode -- Biological control ( lcsh )
Sporeforming bacteria ( lcsh )
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bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

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Thesis:
Thesis (Ph. D.)--University of Florida, 2002.
Bibliography:
Includes bibliographical references (leaves 125-147).
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Also available online.
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Printout.
General Note:
Vita.
Statement of Responsibility:
by Janete Andrade Brito.

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University of Florida
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BIOCHEMICAL EVENTS DURING THE DEVELOPMENT OF Pasteuria penetrans
WITHIN THE PSEUDOCOELOM OF Meloidogyne arenaria











By

JANETE ANDRADE BRITO











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 2002




























To my parents, Alaide Andrade de Brito and Urbano Pinheiro de Brito, for their unconditional
love and support to follow my dreams.














ACKNOWLEDGMENTS

I am indebted to the Coordenagdo de Aperfeigoamento de Pessoal de Nivel Superior (CAPES), Brasilia, DF, Brasil, for financial support.

I express my deep felt appreciation and thanks to Drs. Robin M. Giblin-Davis, chairman, and James F. Preston, cochairman of my supervisory committee, for their guidance, support, encouragement, suggestions, and friendship they gave me throughout this study. Thanks also are expressed to the other members of my committee, Drs. Henry C. Aldrich, Donald W. Dickson, and Grover C. Smart, Jr for their support, suggestions, encouragement, and friendship.

My sincere thanks and appreciation go to Donna S. Williams and John D. Rice for assistance in the laboratory, encouragement, and friendship.

Sincere thanks go to Mrs. Debbie Hall who guided me to follow the University of Florida rules throughout my program, and made sure that I did not miss any deadlines. I also thank Dr. Khuong B. Nguyen for friendship, kindness, and help with portions of my program, and also to Onaur Ruano for his unconditional support, encouragement, and friendship at the beginning of my career as nematologist at the Fundago Institute Agronomico do Parand, Londrina PR, Brazil.

Special thanks are extended to Drs. Waine Dixon, Paul Lehman, and Renato Inserra for their support and friendship.

Thanks to my labmates Claudia Riegel, Fahiem K. El-Borai Kora, Billy W. Crow, iii














Hye Rim (Helena) Han, Zhongxiao Chen, and Ramazan Cetintas. They gave me great friendship and inspiration. Also thanks go to Drs. Hermes Peixoto Santos Filho, Maria de Lourdes Mendes, Rui P. Leite, Alfredo O. A. de Carvalho, Luis G. E. Vieira, and Rui G. Carneiro; Marinalva Pereira Santos, Solange Colavoupe, Lorain M. McDowell, Marisol D~ivila, Heather L. Smith, and Susana B. Carrasco for their friendship, support and sense of humor.

Many thanks go to my dear Brazilian friends for their support and friendship.

Special thanks go to my husband, Don Dickson, and also to my parents, Alaide A. de Brito and Urbano P. de Brito, my sister Maria Augusta, and to my brothers Urbano Filho, Elisiario Neto, and Antonio Marival for their love, patience, and encouragement throughout my life.

Thanks go to my nieces, Rayane, Suian, and Heleninha, for their love.


















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TABLE OF CONTENTS

Page


ACKNOWLEDGMENTS.............................................1ii1

LIST OF ABBREVIATIONS ......................................... viii

LIST OF TABLES................................................... x

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

ABSTRACT ...................................................... xiv

CHAPTERS

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

Host: Root-Knot Nematodes (Meloidogyne spp.) ........................1I
Historical Background.....................................1I
Life Cycle............................................... 2
Symptoms.............................................. 4
Distribution and Economic Importance......................... 5
Management............................................. 6
Parasite: Pasteuria pen etrans..................................... 15
Historical Background .................................... 15
The Genus Pasteuria ..................................... 18
Members of Pasteuria ................................... 18
Systematic and Phylogeny of Pasteuria........................ 21
Distribution............................................ 24
Biological Control Potential................................ 24
The Effect of Other Microorganisms and Pesticides on Pasteuria ..... 26 Life Cycle.............................................. 28
Host Specificity ......................................... 33
Cultivation............................................. 35
Interaction: Host-Parasite........................................ 35
The Role of Adhesin Proteins in the Relationship Host-Parasite ...... 35


V









O bjectives ...................................................... 36

2. SYNTHESIS AND IMMUNOLOCALIZATION OF AN ADHESINASSOCIATED EPITOPE IN Pasteuria penetrans ............................. 38

Introduction ..................................................... 38
M aterials and M ethods ............................................. 40
Nem atode Source ........................................... 40
Pasteuriapenetrans Source ................................... 41
Experimental Design ........................................ 42
Extraction and Determination of Proteins ........................ 44
M onoclonal Antibody ....................................... 45
Epitope Quantification by ELISA .............................. 45
SDS-PAGE Analysis ........................................ 46
Imm unoblotting ............................................ 47
Immunofluorescence of Whole Endospores ...................... 47
Tissue Preparation for Sectioning .............................. 49
Immunogold Labeling ....................................... 50
R esults ......................................................... 51
M icroscopic Examination .................................... 51
Epitope Quantification by ELISA .............................. 51
SDS-PAGE Analysis and Immunoblotting ....................... 53
Immunfluorescence ......................................... 53
Inmunogold Labeling ....................................... 57
D iscussion ...................................................... 68

3. DETECTION OF ADHESIN PROTEINS AND IMMUNOLOGICAL
DIFFERENTIATION OF Pasteuria spp. USING A
MONOCLONAL ANTIBODY ......................................... 73

Introduction ..................................................... 73
M aterials and M ethods ............................................. 75
Origin of Pasteuria Species and Isolates ......................... 75
Propagation of Bacterial Species and Isolates ..................... 76
Extraction and Determination of Proteins ........................ 79
Preparation of Infected Nematodes for TEM ...................... 80
Immunocytochemistry ....................................... 81
SDS-PAGE Analysis ........................................ 82
Im m unoblotting ............................................ 83
R esults ......................................................... 84
Immunocytochemistry ....................................... 84
SDS-PAGE and Immunoblotting Analysis ....................... 84
D iscussion ...................................................... 85


vi











4. SYNTHESIS OF SMALL, ACID-SOLUBLE SPORE PROTEINS IN
Pasteuria penetrans ............................................... 99

Introduction ................................................. 9
Materials and Methods ........................................ 101
Pasteuria penetrans Endospores Source.......................101
Bacillus subtilis Spore Source.............................. 101
Extraction and Determination of SASPs from P. penetrans and
B. subtilis............................................. 101
Conjugation of SASPs Peptide Carrier Proteins ..................102
Purification of the Conjugates.............................. 102
Immunization of Hens for Production of Polyclonal. Antibodies .....103 Determination of IgY Activities in Yolk Extracts ................104
Extraction of IgY from Egg Yolk Extracts ..................... 104
Determination of Activities of Purified IgY ....................105
Concentration of Purified IgY using Centripep .................. 105
Affinity of Anti-Peptide IgY for SASPs .......................105
Results .................................................... 106
Purification of the Conjugates.............................. 106
Determination of IgY Activities in Yolk Extracts ................106
Extraction of IgY from Egg Yolk Extracts .....................11ll
Determination of Activities of Purified IgY ....................111
Affinity of Anti-Peptide IgY for SASPs ......................111Il
Discussion..................................................111l

5. SUMMARY.................................................... 117

APPENDIX A EXTRACTION OF SMALL, ACID SOLUBLE SPORE
PROTEINS FROM SPORES.................................... 122

APPENDIX B ISOLATION OF IgY ANTIB3ODY FROM CHICKEN
EGG YOLKS ............................................... 123

LIST OF REFERENCES............................................. 125

BIOGRAPHICAL SKETCH .......................................... 148









vii














LIST OF ABBREVIATIONS ELISA Enzyme linked immunosorbent assay

BSA Bovine serum albumin

kDa Kilodalton

FITC Fluorescein isothiocyanate

KLH Keyhole Limpet Hemocyanin

M Molar

pl Microliter (s)

11g Microgram (s)

mM Millimolar (s)

gm Micrometer (s)

mg Milligram (s)

ml Millileter (s)

ng Nanogram (s)

nm Nanometer (s)

PAGE Polyacrylamide gel electrophoresis

PBS Sodium phosphate buffer

PBST Sodium phosphate buffer plus Tween





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PBST-BSA 10mM phosphate buffer pH 7.4,150 mM NaC1, 0.05 % Tween, 2%

bovine serum albumin SDS Sodium dodecyl sulfate

UDC 6.0 M urea, 0.03 M dithiothreitol, 0.005 M CHES buffer pH 9.8

1.33x UJDC 8.0 M urea, 0.04 M dithiothreitol, 0.00665 M CHES buffer pH 9.8 WGA Wheat-germ agglutinin






























xix














LIST OF TABLES

Table Page

1.1. Described genera of endospore-forming bacteria and their DNA
base com position .................................................. 23

2.1. Percentage of different developmental stages of Pasteuria penetrans
in Meloidogyne arenaria race 1 ...................................... 52

3.1. Species and isolates of Pasteuria ....................................... 77































x















LIST OF FIGURES

Fiure Page

2.1. Adhesin-associated epitope and total nematode protein per infected
nematode as a function of the development ofPasteuria penetrans .......... 54

2.2. Blots of sodium dodecyl sulfate-polyacrylamide gels of Meloidogyne
arenaria protein extracts after electrophoresis .......................... 55

2.3. Detection of Pasteuria penetrans adhesin-associated epitope as a function
of its development within the pseudocoelom of Melodogyne arenaria
racel ..... .....................................................56

2.4. Differential interference contrast (DIC) and fluorescence microscopy
microphotographs of whole endospores of Pasteuria penetrans P-20 strain ... 58

2.5. Longitudinal section of uninfected second-stage Meloidogyne arenaria
(1-day-old) probed with anti-IgM Mab at 1:10,000 dilution ................ 59

2.6. Immunocytochemical localization of an adhesin-associated epitope
during the development of Pasteuria penetrans ......................... 61

2.7. Labeling of sporogenous stages ofPasteuria penetrans ..................... 63

2.8. Sporogenous stages ofPasteuria penetrans .............................. 65

2.9. Late sporogenous stage of Pasteuria penetrans ............................ 67

3.1. Transmission electron micrographs of Pasteuria endospore sections,
probed with anti-P-20 IgM Mab at 10,000 .............................87

3.2. Gold labeling of endospores of different isolates and species of Pasteuria ...... 89 3.3. Immunoelectron microscopy of endospores ofPasteuria spp ................. 91

3.4. Labeling of endospores of two isolates of Pasteuria spp .................... 93



xi









3.5. Thin section of Pasteuria sp. NA used as a control ......................... 94

3.6. Thin section of an endospore of Pasteuria sp. NA ......................... 95

3.7. Detection of an adhesin-associated epitope in different strains ............... 96

4.1. Activities of antibodies in egg yolk extracts collected from hen 134-5,
34 days after injection of 100 jal KLH-peptide as immunogen
(80 lag per 100 jal) into the wing and 100 lI into the foot pad. A boost
injection was performed at 14 days, 75 lal was injected into the
wing and 75 jal into the footpad. Egg yolk extracts were used at 100 and
1,000 dilution in PBST, whereas the antigen (KLH-peptide) .............. 107

4.2. Activities of antibodies in egg yolk extracts collected from hen 135-1,
34 days after injection of 100 lal KLH-peptide as immunogen
(80 lag per 100 lal) into the wing and 100 lal into the foot pad. A boost injection was performed as in Fig. 4.1. Egg yolk extracts were used at
100 and 1,000 dilution in PBST, whereas the antigen (K.LH-peptide) ....... 108

4.3. Activities of antibodies in egg yolk extracts collected from hen 134-5,
34 days after injection of 100 lal KLH-peptide as immunogen
(80 jig per 100 jil) into the wing and 100 jil into the foot pad. A boost injection was performed as in Fig. 4.1. Egg yolk extracts were used at
100 and 1,000 dilution in PBST, whereas the antigen (BSA-peptide) ....... 109

4.4. Activities of antibodies in egg yolk extracts collected from hen 135-1,
34 days after injection of 100 jil KLH-peptide as immunogen
(80 jig per 100 jil) into the wing and 100 jil into the foot pad. A boost injection was performed as in Fig. 4.1. Egg yolk extracts were used at
100 and 1,000 dilution in PBST, whereas the antigen (BSA-peptide) ....... 110

4.5. Activities of anti-KLH-peptide IgY antibodies extracted from egg yolk
extracts laid by hen 134-5 at 20 to 28 days after injection with
KLH-peptide. Antibodies were used at 1,000 dilution in PBST, pH 7.6.
KLH-peptide and BSA-peptide ..................................... 112

4.6. Activities of purified IgY antibodies (pool 2) from egg yolk extracts
laid by the hen 134-5. Antibodies were dilute to 100; 1,000; and 10,000 in PBST, pH 7.6 whereas the antigens, KLH- peptide and BSA-peptide,
were dilute to 10,000; 100,000; and 1000,000 .......................... 113

4.7. Activities of anti-KLH-peptide IgY antibodies extracted from egg yolk
extracts (pool 2) laid by hen 134-5 Antibody was used at 100; 1,000 and


xii










10,000 dilution in PBST, pH 7.6. SASP-Bacillus subtilis at 100 and
1,000 dilution in coating buffer ....................................114

4.8. Activities of ant-KLH-peptide IgY antibody extracted from egg yolk
extracts (pool 2) laid by the hen 134-5. Antibody was used at 100;
1,000 and 10,000 dilution in PBST, pH 7.6. SASP-Pasteuriapenetrans
at 100 and 1,000 dilution in coating buffer ...........................115







































xiii














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

BIOCHEMICAL EVENTS DURING THE DEVELOPMENT OF Pasteuria penetrans WITHIN THE PSEUDOCOELOM OF Meloidogyne arenaria By

Janete Andrade Brito

May, 2002


Chairperson: Dr. Robin M. Giblin-Davis Major Department: Entomology and Nematology

Pasteuria penetrans is a naturally-occurring bacterial parasite of root-knot

nematodes and a promising biocontrol agent. The endospores of this bacterium attach to the cuticle of second-stage juveniles and complete their life cycle within the infected female. Sequential steps required for the bacterium's propagation include: attachment, infection and germination, vegetative growth, sporulation and release. The hypothesis to be tested in these studies considers that molecular entities present on the surface of mature endospores, designated as spore adhesins, are synthesized at a certain time during the growth and sporulation of P. penetrans, and these allow the bacteria to attach to the nematode host. The objectives of this study were to: 1) determine the temporal relationship between adhesin epitope formation and sporulation of P. penetrans; 2) determine adhesin epitope distribution during spore development and association with xiv









nematode host and; 3) determine if the adhesin epitope is shared by different species of Pasteuria with different host specificities. ELISA and immunoblotting showed that only proteins extracted from P. penetrans-infected root-knot nematodes harvested 24 days after inoculation and growth at 35 oC were recognized by the anti-P-20 IgM Mab that recognizes an adhesin epitope. Labeling, which was first observed in stage II of sporogenesis, identified the epitope distributed over the parasporal fibers, and over other structures, such as sporangium and exosporium, as the bacteria proceeded with the sporogenesis process. However, labeling was not observed on the basal rings, cortex, inner spore coat, outer spore coat, or protoplasm. Immunofluorescence revealed that the epitope does not occur uniformly on the surface of mature endospores. Immunocytochemistry and immunoblot analysis showed that the adhesin epitope is shared by other species of Pasteuria. The uniform distribution of the epitope over the thin sections of mature endospores of strains and species of Pasteuria support a role for the epitope in recognition of the nematode host as an early event in the attachment process.

















xv














CHAPTER 1
INTRODUCTION

Host: Root-knot nematodes (Meloidogyne spp.) Historical Background

Berkeley (1855), in England, reported a plant disease in a greenhouse as "vibrios forming excrescences on cucumber roots". Muller (1884) described the nematode pathogen of the disease as Heterodera radicicola. This pathogen, root-knot nematode, was considered as a single large group for 65 years; nevertheless, it was reclassified several times during that period as follows: Anguillula marioni Cornu, 1879, A. arenaria Neal, 1889, A. vialae Lavergne, 1901, H. javanica Treub, 1885, Tylenchulus arenarius Cobb, 1890, Meloidogyne exigua G61di, 1887, Oxyurus incognita Kofoid and White, 1919, Caconema radicicola Cobb, 1924, and Heterodera marioni (Cornu, 1879) Marcinowski, 1909 (Thorne, 1961). The nematodes gained much attention. There was obvious physiological and biological variability noted among different field populations (Christie, 1946, Christie and Albin, 1944). This led to the classical work by Chitwood in 1949. He re-erected the genus Meloidogyne G6ldi, 1887 to receive all root-knot nematodes. He not only redescribed the type species, M. exigua (G61di, 1887), but also redescribed M.javanica (Treub, 1885), M. arenaria (Neal, 1889), and M. incognita (Kofoid &White, 1919), and described M. hapla, and a new variety, M incognita var. acrita (Hirschmann, 1985). A host-range study conducted by Sasser (1954) showed that


I









2

the host response to root-knot nematode infection was widely variable, not only among species, but also within species of this nematodes. This was the first report calling for the use of differential host plant bioassays to aid with the identification of Meloidogyne species. Taylor and Sasser (1978) modified the original list of host differentials to include the following six differential host plants: cotton (Gossypium hirsutum cv. Deltapine 16), peanut (Arachis hypogea cv. Florunner), pepper (Capsicum annum cv. California Wonder), strawberry (Fragaria x ananassa Duchesne), tobacco (Nicotiana tabacum cv. NC 95), tomato (Lycopersicon esculentum cv. Rutgers), and watermelon (Citrullus lanatus cv. Charleston Gray). Based on these differentials, four races of M. incognita and two races ofM. arenaria were identified (Taylor and Sasser, 1978). In addition, two biological races in M hapla, based on chromosome numbers, have been reported (Triantaphyllou, 1966). Also, some physiological (Carneiro et al., 1990) and genetic variability (Camrneiro et al., 1998, Janati et al., 1982, Triantaphyllou, 1985) has been reported within M javanica and M arenaria (Esbenshade and Triantaphyllou, 1985). Up to 1981, about 80 species of Meloidogyne had been described (Eisenback et al., 1981).

Life Cycle

The life cycle of root-knot nematodes starts with the production of eggs. After the embryogenesis process is completed the first-stage juvenile molts within the egg. The second-stage juvenile (J2) hatches, and migrates freely in the soil. The J2 is the major survival stage and only infective stage. It enters susceptible plant roots to continue its life cycle. The J2 are attracted to plant roots. They migrate to a root of a susceptible plant 25









3

cm vertically in 10 days (Prot, 1978). The J2 generally penetrate roots directly behind the root cap; however, penetration may occur at points where lateral roots emerge (Hussey, 1985). Cellulase, derived from esophageal gland cells, may play a role in the penetration and migration in roots (Bird et al., 1975). The subventral gland cells are the most active in J2 (Bird, 1967). Nonetheless, following the onset of parasitism, the dorsal gland cell increases the production of secretory granules and becomes the predominate gland cell in females (Bird, 1968; 1983; Hussey and Mims, 1991). After penetrating a root, the J2 migrates intercellularly in the cortex toward the region of cell differentiation. When its head reaches the periphery of the vascular tissue, it establishes a feeding site (Hussey, 1985). Secretions injected through the stylet into the vascular tissue of the cells near the head cause morphological and physiological changes in these cells, which enlarge and are called giant cells (Hussey et al., 1998). The roots enlarge at those sites producing galls (Loewenberg, et al., 1960). Five to six multinucleate giant cells develop. These are highly specialized cells on which the nematode feeds (Hussey, 1985). After establishing the feeding site, the J2 becomes sedentary and undergoes morphological changes including increase in its body width but not its length (Taylor and Sasser, 1978). The nematode molts three more times during development to form the third and fourth stage juveniles and the adult stage (male and female). The males are vermiform whereas the females are globose-pyriformn in shape. The rate at which these nematodes develop is influenced by several factors such as temperature, host suitability, and vigor of the host. Tyler (1933) reported that at 27.5 0C to 30 0 C females developed from J2 to the egglaying female in about 17 days, at 24.5 'C in 21 to 30 days, at 20 'C in 31 days, and at









4

15.4 'C in 75 days. Females reproduce mainly by parthenogenesis (Triantaphyllou, 1985). Some species are amphimitic or reproduce both by parthenogenesis or amphimixis. Females lay eggs into a gelatinous matrix that forms an egg mass. The number of eggs per egg mass is highly variable, but may range from almost 200 to 1,000 eggs. The egg masses are generally found outside the galled tissue, but in some host plants the egg mass will lie within the galled tissue. SvM12tms

Plants infected with root-knot nematodes exhibit above-ground and below-ground symptoms. The first below-ground symptoms are the formation of root galls and a poorly developed root-system. The galls result from cell enlargement (hyperplasia), and an increase in cell number (hypertrophy) surrounding the giant cells. Galls usually start to develop in 1 to 2 days after root penetration by a J2. The gall size, which can be small and discrete or large, and in some cases coalesced, is related to the number of nematodes inside the plant tissue (Dropkin, 1954). The size of the galls varies among plant species and nematode species. Generally, egg masses may be observed easily on a galled root, but in some plant species the egg masses are covered by plant tissue. Galls caused by root-knot nematodes can be diagnosed erroneously as nitrogen nodules. Nematode galls are an integral part of root tissue and can not be detached without severely damaging the roots, whereas nitrogen nodules, caused by Bradyrhizobium spp., are round swellings that appear to be attached to the root and are detached easily. Nodules may be infected by root-knot nematodes, and galls and egg masses can be found on the nodules (Minton and Baujard, 1990; Porter et al., 1984).









5

The above-ground symptoms usually depend on the initial nematode density in the soil as well as environmental conditions (Minton and Baujard, 1990). Infected plants have reduced uptake of nutrients and water, which produces yellowing, wilting, and stunting of leaves (Nestscher and Sikora, 1990). Distribution and Economical Imnportance

Meloidogyne spp. are among the most widespread and important plant pathogens limiting crop productivity (Sasser and Carter, 1985). Root-knot nematodes can establish in several soil types; however, suppression of crop yields caused by these nematodes are more severe in sandy soils than in clay soils (Taylor and Sasser, 1978). Heavily infected plants may die when there is severe stress caused by hot, dry conditions. Yield losses caused by plant-parasitic nematodes are approximately $8 billion a year to producers in the United States and nearly $78 billion worldwide (Society of Nematologists, Committee on National Needs and Priorities in Nematology, 1994). However, the damage caused by root-knot nematodes alone is very difficult to determine, and sometimes it is overlooked or underestimated because of the interaction with soilborne fungi, bacteria, viruses, insects, and other nematodes (Nestscher and Sikora, 1990).

Meloidogyne spp. cause damage and are associated with many plants, including economic crops and weeds in all areas of the world (Taylor and Sasser, 1978), but they are considered to be most important in tropical regions (Johnson and Fassuliotis, 1984; Mai 1985). This is mainly due to i) high temperatures and a longer growing season that favors more generations of the nematode per year, resulting in higher nematode densities in the soil; ii) the presence of highly virulent species, such as M incognita, M arenaria,









6

and M. javanica, which are well-adapted to warmer areas, and iii) prevalence of more disease complexes involving root-knot nematodes and soilbome fungi (Mai, 1985). Meloidogyne incognita has the widest geographic distribution of all species described, followed closely by M. javanica, and M. arenaria. Those species are very common in tropical regions, whereas M. hapla is more common in temperate regions of the world (Taylor and Sasser, 1978). The optimum monthly temperature for development of M. incognita is 27 'C; nonetheless it can be found in areas that have an average temperature of 24-30 'C (Eisenback and Triantaphyllou, 1991). In contrast, M. hapla can survive in frozen soil and it can reproduce at temperatures as low as 15 C (Taylor and Sasser, 1978).

Management

Chemical nematicides. In the 1940s, discovery of the nematicidal properties of 1,2-dichloropropane, l,3-dichloropropene (DD) made it possible to demonstrate to producers the damage caused by root-knot nematodes. It marked the beginning of the soil fumigation industry (Johnson and Feldmesser, 1987). After World War II, ethylene dibromide (EDB), 1,2-dibromo-3-chloropropane (DBCP), and bromomethane (methyl bromide, MBr) were formulated as soil fumigants. Each was offered at prices economical for use in the production of moderate to high-value crops (Johnson and Feldmesser, 1987). Later DD, EDB and DBCP were found in ground water, and were withdrawn from the market (Heald, 1987).

Since 1960, different methyl bromide formulations have been used for high-value crops. Methyl bromide has became one of the most popular fumigants because of its









7

broad-spectrum activity and its relatively low cost (Noling and Becker, 1994). It is not only highly efficient in the control of nematodes, but also provides control of fungi, bacteria, insects, rodents, and weeds (Thomas, 1996). Methyl bromide has been used as an agricultural soil fumigant, structural and commodity fumigant, and for quarantine and regulatory purposes (USDA, 1993a; 1993b; Watson, et al., 1992). About 79,000 tons have been used annually on a global basis by agricultural users, mainly as a soil fumigant (75%), but also as a post-harvest fumigant (22%) and for structural (3%) pest control (UNEP, 1995). Worldwide more than half of the production of methyl bromide is used on four crops: tomato, tobacco, strawberries, and melons (Ferguson and Padula, 1994; Stephens, 1 996a; 1 996b).

In Florida and in other states, methyl bromide is used mainly under plastic mulch as a preplant soil fumigant in the production of tomato, pepper, strawberry, other fruits, turfgrass, and nursery crops; however, most methyl bromide is consumed in the tomato, pepper, and strawberry industries (Ferguson and Padula, 1994; Johnson et al., 1962; McSorley et al., 1986; Overman and Jones, 1984).

The emission of methyl bromide into the atmosphere became a major

environmental concern in the late 1980s. The Montreal Protocol Treaty, an international agreement signed by more than 150 countries, governs the world-wide production and trade of ozone-depleting substances. In 1992, the signatories of the Montreal Protocol identified methyl bromide as an ozone depleter (Watson et al., 1992). In 1993, the Montreal Protocol treaty was amended to require that developed countries freeze the production of methyl bromide at 1991 levels by 1995 (USEPA, 1993), and at the 1995









8
meeting, a global methyl bromide production phase-out was approved (Thomas, 1996). Industrial nations were to have a 25% reduction by 2001, a 50% reduction by 2005, and a complete phase-out in 2010, whereas developing nation should freeze the production of methyl bromide in 2002 based upon an average of the years 1995-98 (UNEP, 1995).

In the last several years, studies have been carried out to develop alternative

biocides and to implement new strategies for methyl bromide replacement. Materials that have been identified to have broad spectrum activity in soils include 1,3-dichloropropene (1,3-D) products (Riegel, 2001), dazomet, trichloronitromethane (chloropicrin), dithiocarbamate (metham sodium), sodium tetrathiocarbonate, formalin or formaldehyde, and nonfumigants nematicide-insecticides (Anonymous, 1995). However, none of the materials provide the same level of broad spectrum activity as that provided by methyl bromide. Chloropicrin alone is very efficient for the control of many soilborne fungi, but it does not control plant-parasitic nematodes efficiently. 1,3-D provides control of cyst, root-knot, stubby root, lesion, ring, and dagger nematodes, but it is not effective against fungi (Locascio et al., 1997, Stephens, 1996b). 1,3-D can be mixed with chloropicrin to enhance activity against soilborne fungi. Such products are registered for more than 120 vegetable, field, and nursery crops in the United States (Melicher, 1994).

Crop rotation. Nonchemical alternatives for suppressing nematode populations include the use of crop rotation, resistant varieties, cover crops, soil amendments, flooding, solarization, bare fallowing, and biological control (Christie, 1959; Netscher and Sikora, 1990; Mai, 1985). Some of those techniques have been used for many years, and can be effective against some plant-parasitic nematodes under specific situations, but









9

they do not provide the same broad spectrum of control as methyl bromide.

Crop rotation is one of the oldest ways to manage Meloidogyne spp. However, due to their broad host range, choosing the appropriate crop can be difficult (Potter and Olthof, 1983), and in many cases the best crop choice to manage the nematode densities in the soil is not a suitable choice for the growers. The principle of this method is based on the use of resistant, susceptible, or tolerant crops for the predominant species of rootknot nematode for a specific area (Johnson, 1982). Currently, crop rotation remains an option to reduce the damage caused by root-knot nematodes in the southeastern United States (Johnson, 1982). Rodriguez-Kibana et al. (1988, 1989) showed that castor (Ricinus communis L.), American jointvetch (Aeschynomene americana L.), partridge pea (Cassiafasiculata Michx.), and sesame (Sesamum indicum L.) did not support M. arenaria populations in the field. McSorley et al. (1994) studied the effects of 12 summer crops on M. arenaria race 1 and on the yield of vegetables in microplots. Castor, cotton (Gossypium hirsutum L.), velvetbean (Mucuna deeringiana [Bort.] Merr.), crotalaria (Crotalaria spectablis Roth.), and hairy indigo (Indigofera hirsuta L.) reduced nematode numbers. Yields of vegetable crops were higher following castor than other summer crops, and yields of vegetable crops following castor as a cover crop were approximately double the yields of the same vegetable crop following peanut, a host of M. arenaria race 1.

Resistance. Nematode-resistant cultivars can be an option to manage root-knot nematodes, and they might be used alone or in crop rotation schemes as part of an integrated root-knot nematode control program. Attempts have been carried out to









10

develop cultivars resistant to one or more species of root-knot nematodes. Currently, there are nematode-resistant cultivars of tomato, southern pea, pepper, bean, and sweet potato (Noling and Becker, 1994). However, due to the occurrence of genetic variability within species of root-knot nematodes, it is difficult to develop a cultivar that is resistant to more than one race. In addition, the occurrence of mixtures of races and species of root-knot nematodes within a given area, as well as resistance being broken at high soil temperatures, often limits their usefulness. Even though the tomato resistant gene "Mi" typically confers resistance to M. javanica, M. incognita, and M. arenaria, virulent populations of these nematodes have completely overcome the Mi gene resistance (Castagnone-Sereno 1999; Xu et al., 2001). A greater problem to overcome is the loss of host resistance in tomato that occurs when soil temperatures heat up to over 28 'C (Abdul-Baki et al., 1996; Tzortzakakis, 1997). A loss of resistance to M. incognita in Phaseolus vulgaris was observed at 24 'C and above (Mullin et al., 1991).

Integrated pest management. The integration of different tactics have been

implemented in attempts to manage plant-parasitic nematodes. In the southern United States, M. incognita is a major pathogen of sweet potato (Hall et al.; 1988). A combination of crop rotation, resistant cultivars, nonhost, and nematicides seems to be the most economical method of nematode control on sweet potato (Jatala and Bridge, 1990). Meloidogyne arenaria race 1 is one of the most serious pathogens of peanut in the southern United States. For many years peanut growers have relied on crop rotation, winter cover crops, post harvest crop destruction, and nematicides for managing root-knot nematodes (Dickson, 1998). Recently, the peanut germplasm has been released from









11

Texas A&M University that is resistant to race I of M. arenaria (Simpson and Starr, 1999). With the development of suitable cultivars incorporating this resistance will greatly improve nematode management for peanut producers.

Biological control agents. Root-knot nematodes, their antagonists and parasites, share the same soil habitat. Interactions of these organisms are affected by a number of factors such as the physical and chemical environment of the soil as well as the soil microflora which might play a role in the use of antagonists and parasites in root-knot nematode management (Stirling, 1991). Although several organisms such as fungi, bacteria, viruses, nematodes, mites, insects, protozoans, turbellarians, oligochaetes, and tardigrades have been shown to have some affect on nematode population densities under laboratory and greenhouse conditions, field results have been contradictory (Jairajpuri et al., 1990; Stirling, 1991). Particular attention has been given to effects of soil-inhabiting fungi on the population densities and activities of plant parasitic-nematodes. The known fungal antagonists (Gray, 1988) of nematodes are grouped as i) endoparasites of vermiform nematodes; ii) nematode-trapping fungi, and iii) female and egg parasites and cyst colonizers.

Endoparasitic fungi are classified into three categories based on their mechanism of infection and their taxonomic position: i) group I, encysting species of Chytridiomycetes and Oomycetes such as Catenaria anguillulae, Lagenidium caudatum, Aphanomyces sp. and Leptolegnia sp. which have a flagellated zoospore as their infective propagule; ii) group II, Deuteromycetes producing adhesive conidia and conidia which are ingested; and iii) group III, Basidiomycetes producing adhesive conidia. Fungi of









12

groups II and III initiate the infection process when the conidia either adhere to the nematode's cuticle (Drechmeria coniospora, Hirsutella rhossiliensis, Macrobiophthora vermicola, Myzocytium humicola, Nematoctonus leiosporus, N. concurrens, N. haptocladus, and Verticillium balanoides), or when conidia lodge in the buccal cavity or the gut of the host (all species of Harposporium but one) (Stirling, 1991). This latter group would not likely be efficient for biocontrol of plant-parasitic nematodes because they would be unable to ingest the conidia (Stirling, 1991).

Nematode-trapping fungi or predatory fungi have sparse mycelia that have been modified to form organs capable of capturing nematodes. They are the best known nematophagous fungi, and they have been studied for over a century (Stirling, 1991). There are six mechanisms by which these types of fungi can capture a nematode: i) Adhesive hyphae, produced by Zygomycetes (Stylapage and Cystopage) and a few species of Hyphomycetes. A yellowish adhesive secretion is produced by the fungi. These are considered to be the least sophisticated trapping mechanisms. ii) Adhesive branches produced by a few species of fungi, such as Monacrosporium cionopagum. Erect branches of one or two cells produced on the hyphae may anastomose to form loops or two dimensional networks, which may trap nematodes as they move around. iii) Adhesive mycelial network, the most common type of trap, found in almost all soil types. It forms from the lateral branch growing from the vegetative hypha and curving to fuse with the parent hypha. More loops are produced on this loop or on the parent hypha, until a complex, three-dimensional, adhesive-covered network of anastomosed loops is produced (Arthrobotrys oligospora). iv) Adhesive knobs, formed of distinct adhesive-









13

globose cells that are either sessile on the hypha or borne aloft on a short, erect stalk. These cells occur along the hypha, so that nematodes are often restrained by several knobs. Nematodes may struggle to escape the knobs, which may cause the knobs to detach from their stalks in some species but the knobs remain firmly attached to the nematode and germination occurs quickly. This type of trap mechanism is most common among Hyphomycetes, but it is found also in the Basidiomycetes. Nematoctonus produces non-detachable, hourglass-shaped knobs that are engulfed in a larger, spherical ball of viscous substance (Barron, 1997). v) Non-constricting rings, the most frequent device in nematophagous fungi. Three-celled rings are formed when erect lateral branches from vegetative hyphae thicken and curve, which then fuse to the support stalks. Nematodes are captured when rings become wedged around their bodies. vi) Constricting rings, similar to non-constricting rings. The rings are attached to hypha by short stalks. Nematodes entering these rings trigger them to swell rapidly inward, thereby capturing the nematode. The ring closes in about 0. 1 second once initiated; however there is a lag period of 2 to 3 seconds from the time the nematodes first touch the ring cells until it closes. The nematodes can escape during this short period, which makes this type of mechanism an inefficient trap (Stirling 1991).

Female and egg parasites, and cyst colonizers, are a taxonomically and

ecologically diverse group, ranging from host specific zoosporic fungi to opportunistic species that live largely as soil saprophytes. Over the years many fungi have been isolated from females, cysts, eggs, and egg masses of plant-parasitic nematodes, but the









14

majority have proved to be saprophytes rather than parasites (Chen et al., 1996; MorganJones and Rodriguez-Kibana, 1988; Stirling, 1988).

Rodriguez-Kibana and Morgan-Jones (1988 ) listed 12 genera of fungi that are isolated frequently from females, eggs, and cysts of Heteroderidae in Australia, Europe, and North and South America: Acremonium, Alternaria, Catenaria, Cylindrocarpon, Exophiala, Fusarium, Gliocladium, Nemathophora, Paecilomyces, Penicillium, Phoma, and Verticillium. Among these V. chlamydosporium has been the most widely studied (Stirling, 1988), and proven pathogenic to Meloidogyne, Globodera, and Heterodera. The fungus Paecilomyces lilacinus was found parasitizing eggs of Meloidogyne incognita (Jatala et al., 1979) in Peru. After its discovery, it became the principal organism of interest (Dube and Smart, 1987; Jatala et al., 1979; 1980; 1981). Although it has been found in many geographical areas (Gintis et al., 1983; Godoy et al., 1983; Morgan-Jones et al., 1984; Dackman and Nordbring-Hertz, 1985) it is more common in warmer areas of the world (Domsch et al., 1980). Paecilomyces lilacinus has been shown to be a biocontrol agent of several species of nematodes (Jatala 1985; 1986). However, there are mixed reports on the efficacy of this fungus (Hewlett et al., 1988; Rodriguez-Kibana et al., 1984).

The bacterium, Pasteuria penetrans (Chen et al., 1997b; Eddaoudi and Bourijate, 1998; Freitas, 1997; Trudgill et al., 2000, Tzortzakis and Gowen, 1994; Spiegel et al., 1996), has become the most studied biocontrol agent in the last several years, and is reported to be one of the most promising biological control agents of root-knot nematodes (Chen et al., 1996; Duponnois et al., 1999; Oostendorp et al., 1991; Zaki and Maqbool,









15

1992). Once the problem with its cultivation and mass-production is overcome it may be a very useful biological agent in an integrated root-knot nematode management program.


Parasite: Pasteuria penetrans

Historical Background

The history of Pasteuria spp. has a rather unusual start in that the organism was first reported as a parasite of the water flea Daphnia magna Strauss. This discovery was made in 1887 by Elie Metchnikoff, soon after he accepted a research position offered by Louis Pasteur at the newly formed Pasteur Institute, Paris (Sayre, 1993). In 1888 Metchnikoff erected a new genus, Pasteuria, which he named in honor of Louis Pasteur, to contain the new species, P. ramosa. He emphasized the unique mode of division of this bacterium when he wrote, "Pasteuria sp. was able to undergo as many as five longitudinal divisions at the same time, giving it a characteristic fan shape" (Sayre, 1993 pl01). All attempts made by Metchnikoff to culture the bacterium failed, and thus the type strain was not established (Sayre, 1993).

For many years the description of Pasteuria ramosa enticed researchers around the world to seek the bacterial parasite of water fleas (Henrici and Johnson, 1935; Hirsch ,1972; Staley, 1973). A budding bacterial species of the Blastobacter group, found occasionally on the exterior surfaces of Daphnia sp., was classified erroneously as Metchnikoffts unique bacterium, even though it did not form either endospores, mycelium or branches, was not a parasite of cladocerans, and showed a Gram-negative









16

reaction. This budding bacterium (strain ATCC 27377) was cultivated in vitro, and then assigned erroneously as the type species of the genus Pasteuria (Staley, 1973).

Eighty-nine years after Metchnikoff discovered P. ramosa, it was rediscovered infecting Moina rectirostris, a member of the family Daphnidae (Sayre, 1977). The similarity between the newly discovered bacterial strain and Metchnikoff's bacterium was very clear despite the lack of evidence of longitudinal division. Primary colonies branched and formed a cauliflower-like shape. Daughter colonies were formed by the fragmentation of mother colonies. Quartets, doublets, and single sporangia were produced from the daughter colonies. A sporangium consisted of a conical stem, swollen middle cell, and an endogenous endospore (Sayre et al., 1979; 1983).

Ten years after Pasteuria had been assigned erroneously as strain ATCC 27377, that strain was reclassified as Plactomyces staleyi Starr, Sayre, and Schmidt, 1983 (Starr et al., 1983). Starr et al. (1983) requested that the original description of P. ramosa Metchnikoff, 1888 be conserved and that ATCC 27377 be rejected as the type strain of P. ramosa. Later that request was supported by the Judicial Commission for the Code of Nomenclature of Bacteria (Judicial Commission, 1986), and further studies supported that decision (Sayre et al., 1988; 1989).

Cobb (1906) was the first to report an organism resembling Pasteuria sp.

(numerous highly refractile spores) as a parasite of a nematode, Dorylaimus bulbiferous. He erroneously classified the parasite as a sporozoan. Later Micoletzky (1925) found an organism whose shape and spore size were similar to those reported in 1906 by Cobb. Micoletzky suggested that those spores belonged to the genus of a sporozoan, Duboscqia









17

Perez. Thorne (1940) described in detail an organism parasitizing Pratylenchus pratensis (de Man) Filipjev (later identified as P. brachyurus by Thorne), and on the assumption that the organism was similar to the parasite described by Micoletzky, assigned it to the genus Duboscqia as D. penetrans. However, over the years the taxonomic position of the nematode parasite has been questioned (Canning, 1973; Williams, 1960). The misplacement of the organism, now known to be a bacterial parasite of nematodes as a protozoan, persisted for almost 70 years. Mankau (1975a) reexamined the nematode parasite using electron microscopy and showed for the first time that it is a bacterium rather than a protozoan; he reassigned it to the genus Bacillus as B. penetrans (Thome, 1940, Mankau, 1975). Nonetheless, neither flagella nor active motility were observed in Bacillus penetrans (Sayre and Starr, 1985). Soon more studies on the procaryotic affinities (Mankau, 1975b), biology (Mankau and Imbriani, 1975), ultrastructure (Imbriani and Mankau, 1977), and host (Mankau and Prasad, 1977) of B. penetrans were carried out. B. penetrans was never included in the "Approved Lists of Bacterial Names" (Skerman et al., 1980), thus the confusion on the classification of the bacterial nematode parasite continued.

Sayre and Wergin (1977) observed the similarity between the developmental

stages of a bacterial parasite of Meloidogyne incognita with the original descriptions and drawings of the life cycle of P. ramosa. Later morphological and taxonomic reevaluations of P. ramosa and B. penetrans were provided (Sayre et al., 1983). Finally Sayre and Starr (1985) placed the bacterial parasite of nematodes in the genus Pasteuria,









18

as P. penetrans, due to its similarity with Pasteuria rather than Bacillus, and presented an emended description of the genus Pasteuria Metchnikoff. The Genus Pasteuria

Species of Pasteuria are Gram-positive, endospore-forming bacteria. The genetic and biochemical aspects of the formation of the virulent endospores of Pasteuria spp. are not well understood, but the morphological aspects are (Chen et al., 1977a; Giblin-Davis et al., 1995; Sayre and Starr, 1985; Sayre 1993). These bacteria form a dichotomously branched septate mycelium. The terminal hyphae of a mycelium elongates, and then segments to form the sporangia, and eventually endospores. (Sayre and Starr, 1985). Mother colonies, which resemble a cauliflower or elongate grapes in clusters, fragment to form daughter colonies. Daughter colonies form quartets, doublets, and finally a single sporangia which enclose a single endospore (Chen et al., 1997a; Sayre and Starr, 1985). Endospores are nonmotile and resistant to desiccation and elevated temperatures (Dutky and Sayre, 1978; Stirling, 1985; Williams et al., 1989). Endospores of P. penetrans are cup-shaped and measure, on average 3.4 Am 0.2 by 2.5 kzm + 0.2 using transmission electron microscopy (Sayre 1993).

Members of Pasteuria

There is still considerable confusion about the taxonomy of Pasteuria. Over the years the criteria used to assign species to the genus have been host specificity, developmental characteristics, and size and shape of sporangia and endospores (Sayre and Starr, 1989). However, host specificity overlaps in several cases. Although sizes of









19

endospores and sporangia are considered to be host specific (Ciancio, 1995), endospore diameters of P. penetrans vary from 3.6 to 7.0 /m ( Sayre and Starr, 1985).

Cross-genera hosts have been reported. For example, one isolate of P. penetrans reported from the United States (Mankau, 1975a; Oostendorp et al., 1990), Puerto Rico (Vargas and Acosta, 1990) and China (Pan et al., 1993) parasitizes both Meloidogyne and Pratylenchus spp. An isolate of Pasteuria sp. from India parasitizes Heterodera sp. and M. incognita (Bhattacharya and Swarup, 1988), whereas another strain reported from India parasitizes Heterodera spp., and Rotylenchulus reniformis (Sharma and Davies, 1996). Davies et al. (1990) reported that endospores of a Pasteuria sp. extracted from H. avenae, cereal-cyst nematode, attached to the cuticle of H. shachtii, H. glycines, Globodera rostochiensis, G. pallida, and M. javanica. On the other hand, Pasteuria sp. S-1 showed a high a level of host specificity. S-1 strain attached to B. longicaudatus but did not attach to any of the other nematodes, including J2 of M. arenaria, M. incognita, M javanica, H. galeatus, and Pratylenchus penetrans (Giblin-Daves et al., 1995). These results were confirmed by Bekal et al. (2001). They showed that S-1 did not attached to H. schachtii, Longidorus africanus, M. hapla, M. incognita, M. javanica, P. brachyurus, P. scribneri, P. neglectus. P. penetrans, P. thornei, P. vulnus, or Xiphinema spp.

Some isolates of Pasteuria have been reported to attach to and develop in

different life stages of the nematode host (Abrantes and Vovlas, 1988; Davies et al., 1990). Mature endospores of P. penetrans were observed in the peseudocoelom of J2 and males of Meloidogyne sp. and J2 of H.fici (Abrantes and Vovias, 1988). Davies et al.









20

(1990) found that a Pasteuria sp. isolated from the cereal-cyst nematode, H. avenae Wollenweber, completed its life cycle in the J2 but not in females and cysts.

Different genera of nematodes have been reported to be parasitized by Pasteuria spp. at the same site and in the same growing season. Giblin-Davis, during a survey in South Florida, found that B. longicaudatus, Meloidogyne spp. and Helicotylenchus microlobus were parasitized by Pasteuria spp. in Collier County; B. longicaudatus, Hoplolaimus galeatus, Tylenchorhynchus annulatus, and Meloidogyne spp. in Broward County; and H. microlobus and Meloidogyne spp. in Palm Beach County.

Currently four species of Pasteuria have been described so far: i) P. ramosa, a parasite of the cladocerans (water fleas) Daphnia pulex Leyding and D. magna Strauss (Sayre et al., 1977); ii) P. penetrans, a parasite of root-knot nematodes (Sayre and Starr, 1985), iii) Pasteuria thornei, isolated from Pratylenchus spp. (Starr and Sayre, 1988), and iv) Pasteuria nishizawae (Sayre et al., 1991), a parasite of cyst nematodes (Heterodera and Globodera).

Recently, at least three new species of Pasteuria have been proposed, Pasteuria sp. designated as S-1 (Bekal et al., 2001) from Belonolaimus longicaudatus Rau; North American Pasteuria (Heterodera glycines-infecting Pasteuria) from the soybean cyst nematode, Heterodera glycines Ichinohe, in Urbana, IL, USA (Atibalentja et al., 2000) and one strain from the pea cyst nematode, Heterodera goettingiana Liebscher in Minster, Germany (Sturhan et al., 1994).

Over the years unique isolates of Pasteuria have been reported. For example, a large- and a small-spored isolate of Pasteuria spp., each from Hoplolaimus galeatus








21

(Cobb) Thomrne (Giblin-Davis et al., 1990), and another isolate from Rhabditis sp. (GiblinDavis pers. comm.) were collected from a bermudagrass turf in Fort. Lauderdale, Fl. Two isolates of Pasteuria sp.infecting different ring nematode species have been found: C-1 (Han et al., 1999), and ring nematode Pasteuria (Dickson, pers. comm.). A Helicotylenchus sp.-infecting Pasteuria was isolated from bermudagrass turf shipped from CA (Crow, pers. comm.). Also, three other isolates of Pasteuria that attach and complete their life-cycle in Heterodera spp. have been reported: one isolate from H avenae (Davies et al., 1990); another (HCP) from Heterodera cajani Koshy, the pigeon pea cyst nematode (Walia et al., 1990); and another, HMP, from Heterodera mothi, Khan & Husain (Bajaj et al., 1997).

It is clear that there is a need to use other criteria, in addition to those already

used, to determine species of Pasteuria. The 16S rDNA has been used to determine more precisely the taxonomic position of Pasteuria (Anderson et al., 1999; Atibalentja et al., 2000; Bekal, 2001; Ebert et al., 1996). Once the conditions necessary to mass produce Pasteuria in vitro are known, it will be possible to establish species through genetic and biochemical studies.

Systematics and Phylogeny of Pasteuria

In 1992 13 genera of endospore-forming bacteria were known (Table 1.1). The basis for separating them was morphology, physiology, and genetic diversity (Berkelwy and Ali, 1994). Currently, bacteria are differentiated based on the generally accepted rule that bacteria with DNA base compositions differing by more than 10 mol %GC (G+C) should not be considered as members of the same genus. Strains differing by more than









22

5%GC values should not be regarded as the same species (Bull et al., 1992). The genera Bacillus, Clostridium, and Desulfotomaculum are very heterogenous (Table 1.1). The genera Oscillospira and Pasteuria (four species) have not yet been grown successfully in pure culture. The description of Oscillospira species, O. guillermondii (Berkely and Ali, 1994), was based on morphological characters, whereas the species of Pasteuria were described based on morphological characters, morphometrics, ultrastructure, and host specificity. Otherwise their DNA base composition are unknown.

In the summer of 1992 and throughout 1993 and 1994, P. ramosa was rediscovered parasitizing D. magna collected from several ponds in London, UK (Stimadel and Ebert, 1997). Ebert et al. (1995) used these spores of P. ramosa collected from D. magna, D. pulex, and D. longispina in the previous three summers from England as well as Russia to establish the culture of P. ramosa by co-cultivation in D. magna. These authors ended the uncertainty about the phylogenetic position of Pasteuria Metchnikoff by sequencing the 16S rDNA of the bacterium. They provided strong evidence that P. ramosa belongs to the low G+C Gram-positive endospore-forming bacteria and resides within a clade containing B. tusciae, Alicyclobacillus cycloheptanicus, and A. acidocaldarius, as the closest neighbors. They rejected the placement of P. ramosa in the Actinomycetales. Anderson et al., (1999) provided the first 16S rDNA gene sequence analysis of P. penetrans, and showed that it is correctly placed in the genus Pasteuria. The authors found that P. ramosa is the closest neighbor of P. penetrans, and it is within a clade that includes A. acidocaldarius, A. cycloheptanicus, Sulfobacillus sp., B. tusciae,









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Table. 1. 1. Described genera of endospore-forming bacteria and their DNA base composition.


Genus Mol% GC

Alicyclobacillus 5 2-60

Amphibacillus 36-3 8

Bacillus 32-69

Clostridium 22-54

Desulfotomaculum 3 8-52

Oscillospira

Pasteuria

Sporohalobacter 3 0-32

Sporolactobacillus 3 8-40

Sporosarcina 40-42

Sulfobacillus 54

Syntrophspora 38

Thermoactinomyces 52-55

Source: Berkeley and Ali, 1994.









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B. schlegelii, and P. ramosa. Also Atibalentja et al. (2000), using a sequence of the 16S rDNA, showed that a Heterodera.glycines-infecting Pasteuria (Pasteuria sp. NA) and P. ramosa form a distinct line of descent within the Alicyclobacillus group of the Bacillaceae.

Distribution

Pasteuria spp. have been reported in 51 countries and in various islands in the Atlantic, Pacific, and Indian oceans associated with 205 nematodes species belonging to 96 genera (Sayre and Starr, 1988; Sturhan, 1985). An updated host record list is reported by Chen and Dickson (1998).

Biological Control Potential

There are certain attributes that make P. penetrans a desirable biological control agent: 1) endospores are resistant to desiccation, high temperature, and most nematicides (Dutky and Sayre, Freitas 1997; 1978; Stirling, 1985; Williams et al., 1989); 2) encumbered nematode juveniles have reduced activity and ability to infect roots (Sturhan, 1985); and 3) infected juveniles complete their life cycle, but females have low or no fecundity (Bird, 1986; Bird and Brisbane, 1988).

Pasteuria penetrans has been shown to control root-knot nematodes in

greenhouse tests (Brown and Smart, 1985; De Leij et al., 1992; Stirling 1984) and in field microplots (Brown et al., 1985, Chen et al., 1997b; Dube and Smart, 1987; Oostendorp et al., 1991; Stirling, 1984; Tzortzakakis and Gowen, 1994; Trudgill et al., 2000). Suppression of root-knot nematodes by P. penetrans has been observed in vineyards more than 10 years old in Australia (Stirling and White, 1982) as well as India (Mani et al.,









25

1999), and also in peanut and tobacco fields infected by root-knot nematodes in Florida (Dickson, pers. comm.). Also, suppression of B. longicaudatus by Pasteuria sp. S-I in a bermudagrass turf field in Florida has been reported (Giblin-Davis et al., 1995; 2000).

Studies have been carried out to determine the optimum endospore densities to suppress root-knot nematodes (Chen et al., 1996; Melki et al., 1998; Oostendorp et al., 1991). Chen et al. (1996) found that 10,000 endospores/g of soil was necessary for suppression of M. arenaria race 1 on peanut in plots on a fine sand soil. Melki et al.(1998) reported that the cultivation of a susceptible host for more than one season was needed for P. penetrans to build up its densities to suppressive levels. Oostendorp et al., 1991 showed that endospore attachment to M. arenaria race I increased from 0.11 to 8.6 spores/J2 in plots over a 2-year cropping sequence with peanut (summer) and rye, vetch or fallow (winter)

The use of air-dried soils infested with P. penetrans was one of the first attempts to show the biological control potential of this bacterium. Mankau (1973) used air-dried soil infested with the bacterial spores in greenhouse studies. He reported that after 70 days, plants in the endospore-infested soil had more leaves, greater dry weight, and lower numbers of root galls than in those soil-free of endospores. However, the use of infested soil as a source of endospores is time consuming and inconvenient to transport and handle. Stirling and Wachtel (1980) reported for the first time the use of infested root powder as a source of endospores and as a method for their mass production. The authors showed that when they used 100 mg/kg of soil of air-dried and finely ground roots









26

containing 2x 10' spores/g, that within 24 hours, 99% of the J2 of M. javanica in the pot had endospores attached to their cuticles. Stirling (1984) used tomato roots containing P. penetrans-infected females of M javanica to produce infested, air-dried root powder. Significant control was obtained when at least 80% of the bioassayed J2 were encumbered with 10 or more spores per J2. When the infested root powder was incorporated into root-knot nematode-infested field soil at the rate of 212-600 mg per kilogram of soil, the number of galls and nematodes in the soil at harvest was significantly reduced. Also, the application of P. penetrans in air-dried powdered roots at 55 000 spores/cm3 soil in pots infested with 420 J2 significantly suppressed root galling and egg production of M. javanica through two successive tomato growing seasons. At planting, there was an average of 14 spores per J2 in the soil (Gowen et al., 1998). The application of air-dried root powder infested with P. penetrans strains P-20 and P-100 has been used at Disney World at The Land, Lake Buena Vista, Florida to effectively control M. arenaria, and M. incognita over the several years on sandy plots (Dickson, pers. comm. and Brito, pers. observation).

The Effect of Other Microorganisms and Pesticides on Pasteuira

Duponnois and Ba (1998) studied the influence of soil microflora on the

antagonistic relationship between P. penetrans and M. javanica. The authors showed that the attachment of P. penetrans to J2 ofM. javanica was higher in the presence of larger soil microbial populations, such as fluorescent strains of Pseudomonas, nematophagous and mycorrhizal fungi. One of the explanations given by those authors was that those soil microorganisms may change the soil ionic environment, which favored the attachment of









27

endospores to the nematode cuticle, which is negatively charged (Himmelhoch et al,. 1979). Duponnois et al. (1999) studied the interaction of Enterobacter cloacae and Pseudomonas mendocina, which had been isolated previously from the rhizosphere of tomato cv Roman growing in a field infested by both M. javanica and P. penetrans. Those authors found that P. mendocina and E. cloacae stimulated plant growth, inhibited the reproduction of M. incognita, and increased the attachment of P. penetrans in vitro. Enterobacter cloacae increased significantly the reproduction of P. penetrans. They suggested that the introduction of E. cloacae in soils could enhance the efficacy of P. penetrans.

The compatibility of P. penetrans with some pesticides increased its potential to be used in an integrated management of root-knot nematodes (Brown and Nordmeyer, 1985; Freitas, 1977; Singh and Dhawan, 1998). Carbofuran had no effect on the reproduction of P. penetrans (Brown and Nordmeyer, 1985; Singh and Dhawan, 1998). Freitas (1977) found that treatment with 1,3-dichloropropene (1,3-D) + 17% chloropicrin, 1,3- D + 25% chloropicrin and 1,3-D + 35% chloropicrin reduced significantly the percentage of female nematodes with P. penetrans, whereas metham sodium did not have any effect. However, the author reported that the percentage of nematode females infected by P. penetrans was significantly lower (1.67%) in the soil treated with methyl bromide + 33% chloropicrin than in the untreated control (27.50%) under greenhouse conditions. Under field conditions, the percentage of females infected with P. penetrans from a plot treated with methyl bromide + 33% chloropicrin was 5% compared to the untreated control plot, which had 58% of the females infected (Freitas 1997). The









28

exposure of endospores to the fungicides, hymexazol, fosetyl-Al, and carbendazin had no effect on the attachment or development of endospores (Melki et al., 1998). Life Cycle

Attachment of endospores to the nematode host. Endospores of the P. penetrans attach to second-stage juveniles (J2) of root-knot nematodes as they move through soil pore spaces. After attachment, the sporangial wall and exosporium of the majority of endospores slough off (Sayre and Starr, 1985). The bacterium is reported to attach to J2 and produce virulent endospores only within the pseudocoelom of a mature female. However, one isolate of P. penetrans attached to and developed within the pseudocoelom ofjuveniles, males, and females of M. acronea isolated originally from cotton (Page and Bridge, 1985). Also, an endospore-filled J2 ofMeloidogyne sp. was isolated from a suppressive soil infested with P. penetrans in Florida (Dickson, pers. comm.). Stirling et al. (1990) showed that the number of endospores attached to the cuticle of J2 increased in proportion to both endospore-concentration and time. Davies et al. (1988) found that the number of J2 entering the plant host root was reduced when they were encumbered with 15 or more spores. Ahmed and Gowen (1991) reported that 11 or more endospores per J2 reduced the capability of M. incognita, M. javanica, and M. graminicola to enter the host roots.

Attachment is one of the major steps toward successful development of P.

penetrans within its host, and it has been studied in several laboratories (Afolabi et al., 1995; Bird 1989; Charnecki, 1997, Davies et al., 1996). Persidis et al. (1991) used polyclonal antibodies selected against whole endopsores and wheat germ agglutinin as a









29

probe, and suggested that proteins glycosylated with N-acetylglucosamine are involved in the attachment. Similar results were obtained using a monoclonal antibody raised to whole endospores of P-20 isolate of P. penetrans and wheat germ agglutinin (Chamecki, 1997, Charnecki et al., 1998). Mohan et al. (2001) found that fibronectin-like residues on the cuticle of M. javanica is involved in the attachment of endospores. Other forces such as hydrophobic interactions may be involved in the attachment of endospores to its host (Afolabi et al., 1995; Davies et al., 1996; Esnard et al., 1997)

Germination. Unknown factors trigger the germination of the endospore and the formation of a germ tube. A germ tube emerges through a central opening in the basal attachment layer after an endospore-encumbered juvenile enters a host root and begins feeding (Sayre and Wergin, 1997; Sayre and Starr, 1985; 1988; Serracin et al., 1997). The germ tube penetrates the nematode cuticle and hypodermal tissue, and then enters the pseudocoelom (Sayre, 1993) where unknown growth factors promote its development into a vegetative, spherical colony, containing a dichotomously branched and septate mycelium (Satrr and Sayre 1988). The peripheral fibers of the endospores are closely associated with the cuticle of the nematode (Sayre and Starr, 1985) and are involved in the attachment of endospores to the cuticle.

Vegetative stage. When intercalary cells in the microcolony disperse, many

daughter colonies are formed. Eventually quartets of developing sporangia predominate the pseudocoelom, and then the quartets separate into doublets of sporangia, which separate into single sporangia that will eventually form the endospores (Sayre 1993).









30

Endospore formation. The formation of bacterial endospores is a regulated and complex process. The initiation of sporulation is triggered by several genes, spoO genes, in response to nutrient deprivation (Foster, 1994). It is hypothesized that molecular functions that control sporulation are the same across all genera of endopsore-forming bacteria. Small acid-soluble proteins (SASPs) have been shown to be synthesized by spores of species of Bacillus, Clostridium, and Thermoactynomycetes during sporulation (Setlow, 1988; Setlow and Waites, 1976). The main types of SASPs found in B. subtilis are termed the a/3 type (Connors et al., 1986) and y type (Hackett and Setlow, 1984), which are synthesized during the first 3-4 hours of sporulation, and are found only in spores (Setlow et al., 1992). Previous studies indicated that a/P type-SASPs are DNAbinding proteins, and their binding to the DNA cause UV resistance by modifying spore DNA's UV photochemistry (Manson and Setlow, 1986; Setlow and Setlow, 1987). Another molecule that is found in spores but not in vegetative cells is the dipicolinic acid, which is located in the core of the endospores (Madigan et al, 1997). Studies have shown that calcium, which is present in high concentration in spores, forms a complex with dipicolonic acid in the core, and confers the heat resistance found in endospores (Madigan et al., 1997).

The factors that trigger the sporulation of P. penetrans within the pseudocoelom of the nematode host are not known. However, the sequence of morphological events during the endogenous spore formation of P. penetrans is similar to other Gram-positive endospore-forming bacteria (Chen et al., 1997a; Sayre 1993) as follows: i) formation of a transverse septum within the endospore mother cell; ii) condensation of a forespore from









31

the anterior protoplast; iii) formation of a multilayered wall about the forespore; iv) lysis of the old sporangial wall; and v) release of an endospore (Sayre 1993).

Chen et al. (1997a) found that the sporogenesis process of P. penetrans generally matched stages II through VII following vegetative growth reported for Bacillus thuringiensis. Stage I is unique for Pasteuria sp. The stages are as follows: 1) stage I, mycelium dichotomously branched and microcolonies fully septate; terminal cells elongate to form a sporogenous cell; 2) stage II, the terminal cells increase in size and become oval, 1.2 to 1.7 pm by 0.6 to 1.0 gm, bounded by a 0.002 gm-thick wall; a membrane is formed about 0.4 gm from the anterior end and separates the forespore from the parasporium; 3) stage III, parasporium increases in size and engulfs the forespore. Parasporal fibers are formed and attach to the lower part of the forespore. An inner membrane defines the forespore protoplast and an outer membrane defines the mother cell's protoplast; 4) stage IV, lamella, which rises from the cortex, and inner and outer spore coats start to form; 5) stage V cortex with formation of inner and outer spore coats; the inner spore coat is a narrow multilaminar band whereas the outer spore coat is a wide electron-dense wall; 6) stage VI, formation of exosporium, a delicate membrane that delimits the outermost layer of a typical Gram-positive bacterium; 7) stage VII, complete maturation with formation of endospore, the basal ring surrounding the germinal pore. An epicortical layer, which is a discontinuous, electron-dense band was observed between the cortex and the inner spore coat. Endospores of P. penetrans measure an average of

3.4 im 0.2 by 2.5 um 0.2 (Sayre 1993).









32

The life cycle of this bacterium is not completely synchronized with the life cycle of the nematode since it is possible to observe different developmental stages simultaneously at a given time within the pseudocoelom of a single root-knot nematode female (Chen et al., 1997a). The rate of development is highly temperature-dependent (Hatz and Dickson, 1992; Serracin et al., 1997; Stirling 1981). The optimum temperature for the development of the P. penetrans was 35 'C, at which the bacterium completed its life cycle in 35 days after inoculation (Hatz and Dickson, 1992). An average of 2x 106 endospores have been found within one single female of P. penetrans-infected Meloidogyne sp. (Sturhan, 1985) and P. penetrans-infected M javanica (Stirling 1991).

Soil phase. Endospores are released into soil upon host disintegration.

Endospores are not actively motile in soil; therefore, its contact with the nematode host must rely on the motility of J2, as well as physical factors affecting endospore distribution (Sayre 1993). The factors that mediate the movement and survival of endospores of P. penetrans in soil are not well understood. However, soil water percolation, sizes of soil pore openings, surface charge of soil particles, tillage practices, and soil microflora may play important roles in the distribution of endospores (Sayre, 1993). Kamra and Dhawan (1998) found that at pH 8.0 to 10.0, the average number of endospores encumbered on the bioassayed J2 of H. cajani was 36 and 26 compared to 10 and 7.0 at pH 6.0 and pH 4.0, respectively. Those authors also showed that the movement and distribution of endospores in soil increased with greater pore size, and decreased with an increase in the silt and clay contents of the soil.









33

Host Specificity

Host specificity has been used for many years to determine species of Pasteuria. According to Sayre and Starr (1985; 1989) host range of P. penetrans is limited to species of Meloidogyne.

Host specificity of P. penetrans has been reported in most cases by observing only the attachment of endopores to its host rather than by establishing infection and production of mature endospores. Attachment can occur, but endospores might fail to germinate and propagate within the nematode. Duponnois et al. (2000) tested 25 isolates of P. penetrans, and found that only six attached, developed, and produced mature endospores in M incognita.

Attachment of endospores was greater to the nematode species from which

endospores were originally cultured (Oostendorp et al., 1990; Somasekhar and Metha, 2000). When labeling endospores using monoclonal antibodies, larger areas of the endospores were labeled when Pasteuria were reared on the same nematode population from which endospores were taken than from other populations of root-knot nematodes (Davies et al., 1994). However, Davies et al. (1988) found that a particular isolate of Pasteuria sp. adapted and shifted from one nematode host to another by continually culturing the bacterium on a given nematode host. Stirling (1985) reported that attachment was not always related to the species from which the endospores were isolated, or to the species of the recipient nematode.

Davies et al. (2001) using only the PPI strain of P. penetrans and several field populations of root-knot nematodes collected from Burkino Faso, Ecuador, Greece,









34

Malawi, Senegal, Trinidad, and Tobago showed that the extent of attachment differed between countries. Also, those authors found similar results when endospores of P. penetrans collected from those countries were assayed against M. arenaria and M incognita.

Endospores of P. penetrans did not attach to the entomopathogenic nematodes Steinernema glaseri Steiner, Heterorhaditis zealandica Poinar and H. bacteriophora Poinar and seven new isolates each of Steinernema sp. and Heterorhaditis sp. when juveniles were exposed to I x 105 spores/ml for 24, 48, and 72 hours at 25 oC (Somasekhar and Metha, 2000). Similar results were observed by Mendoza de Gives et al. (1999), who reported that P. penetrans did not attach to animal-parasitic nematodes, free-living nematodes, including wild type Caenorhabditis elegans (Maupas) Dougherty and three of its surface (srf) mutants. Oostendorp (1990) also showed that endospores of P. penetrans did not attach to the free-living nematodes, Panagrelus redivivus (L.) Goodey and C elegans, but attached to different species of plant-parasitic nematodes.

The nature and the amount of protein on the surface of endopores may explain host specificity (Davies et al;. 1992; Persidis et al., 1991). Monoclonal antibodies have shown that the surface of endospores of P. penetrans isolated from M. incognita race 1 is highly heterogenous (Davies et al., 1994). These and other studies (Davies and Redden, 1997) have suggested that the virulence of the bacterium to a certain species of root-knot nematodes is dictated by the surface properties of endospores, and suggested that similar heterogeneity will be present in the nematode cuticle. Differences in cuticle characteristics of J2 of root-knot nematodes have been reported (Davies and Danks









35

(1992). Charnecki et al., 1998 showed that the anti-P-20 IgM MAb recognized differences in the protein extracts from B4, P-20, and P120 isolates of P. penetrans, which have different host specificities.

Cultivation

Pasteuria spp. have not been grown successfully in pure culture (Reise et al., 1988; Williams et al. 1989; Bishop and Eller, 1991). Currently P. penetrans produces virulent endospores only within the pseudocoelom of females of Meloidogyne spp., which in turn must be reared on the roots of a plant host or on excised-root systems (Verdejo and Jaffee, 1988). The mass production of this bacterium relies, currently in the use of dried, powdered roots obtained from infected root systems grown in a greenhouse (Stirling and Wachtel, 1980).



Interaction: Host-Parasite

The Role of Adhesin Proteins in the Host-Parasite Relationship

The surface of Gram-positive bacteria has adhesin proteins, also known as

virulence factors, that allow the bacteria to adhere, invade, and colonize tissues (Salyers and Whitt, 1994). Studies on the composition of the surface proteins have focused mainly on pathogenic bacteria (Kehoe, 1994).

Virulence factors are classified into two major categories: i) promoters of bacterial colonization and invasion of the host; and ii) those that cause disease in the host. Among the virulence factors that promote bacterial colonization are pili, or fimbriae (RobinsBrowne, 1994; Salyers and Whitt, 1994; Suoniemi et al., 1995), and afimbrial adhesins









36

(Salyers and Whitt, 1994) that adhere to mucosal surfaces and bind tightly to the host cells, respectively. Streptococcus pyogenes has a nonfibrillar adhesin (protein F) that mediates its attachment to fibronectin, a protein found on many host cell surfaces, including the mucosa of the human throat (Salyers and Whitt, 1994).

Several environmental signals may affect virulence. These may include

temperature, carbon source, osmolarity, starvation, stress, pH, growth phase; and the levels of specific nutrients including iron, calcium, sulfate, nicotinic acid, and specific amino acids (Mekalanos, 1992). Bacteria use different sigma factors to control different set of genes under specific conditions (Salyers and Whitt, 1994). Similar mechanisms might be used by P. penetrans to produce endospore adhesins involved in recognition and attachment to the nematode host.



Objectives

The biochemical events that occur during the development of P. penetrans within the root-knot nematodes' pseudocoelom are poorly understood and may provide valuable insight into the conditions necessary for the formation of virulent endospores.

The objectives of this research project were to 1) determine the sequence of events required for the formation of P. penetrans spore-associated proteins (adhesins) that are required for the attachment of endospores, as a function of the development of P. penetrans within its nematode host, M. arenaria race 1; 2) determine the distribution of an adhesin-related epitope on the surface of virulent endospores; 3) detect and localize antigens bearing the epitope during the sporogenesis process; and 4) determine whether or









37

not different species or isolates of Pasteuria share the same adhesin-related epitope, which is recognized by the anti-P20 IgM MAb. In addition, a polyclonal antibody against a synthetic polypeptide, which was designed according to the conserved regions of small, acid-soluble proteins (SASPs) of Bacillus spp. was prepared for use as a probe to detect SASPs as a development marker in the sporulation process in P. penetrans.














CHAPTER 2
SYNTHESIS AND IMMUNOLOCALIZATION OF AN ADHESIN-ASSOCIATED EPITOPE IN Pasteuria penetrans

Introduction

Pasteuria penetrans (Thomrne) Sayre & Starr is a Gram-positive, endosporeforming bacterial parasite of Meloidogyne spp. Endospores attach to second-stage juveniles (J2) as they move through soil pore spaces. Unknown factors trigger infection of the nematode host and germination of the endospore. The germination of the endospore occurs after the endospore-encumbered juvenile enters host roots and begins feeding (Sayre and Wergin, 1997; Sayre and Starr, 1985, 1988; Serracin et al., 1997) at some point in development, presumably before the J2 molts to the third-stage juvenile. A germ tube penetrates the nematode cuticle and hypodermal tissue, and then enters the pseudocoelom (Sayre and Starr, 1988), where unknown growth factors promote vegetative growth, differentiation, sporulation, and maturation of endospores. Endospores are released into soil upon host disintegration, and more than 2 million endospores have been found within one single P. penetrans-infected Meloidogyne sp. female (Sturhan, 1985).

There are certain attributes that make P. penetrans a desirable biological control agent: 1) endospores are resistant to desiccation, high temperature, and most nematicides (Dutky and Sayre, 1978; Stirling, 1985; Williams et al., 1989); 2) encumbered juveniles



38









39

have reduced activity and ability to infect roots (Sturhan, 1985); and 3) infected juveniles complete their life cycle, but females have low or no fecundity (Bird, 1986; Bird and Brisbane, 1988). These bacteria complete their life cycle and produce virulent endospores only within the pseudocoelom of Meloidogyne spp., which in turn must be reared on a plant host either in pots or on excised-root systems (Verdejo and Jaffee, 1988). Attempts to culture P. penetrans in vitro have failed to produce virulent endospores (Reise et al., 1988; William et al., 1989; Bishop and Ellar, 1991). The biochemical events that occur during the development of P. penetrans, leading to the formation of virulent endospores within the pseudocoelom, are poorly understood.

The molecular basis for the recognition and attachment has been the subject of investigation in several laboratories. Lectin-carbohydrate interactions have been suggested to be involved in the attachment of P. penetrans to its nematode host. Previous studies have shown that wheat-germ agglutinin (WGA) inhibited the attachment of endospores (Bird et al., 1989; Charnecki 1997; Charnecki et al., 1998; Davies and Danks, 1993). Also, proteins extracted from endospores of P. penetrans were recognized, not only by monoclonal antibodies (Charnecki 1997; Charnecki et al., 1998; Davies and Redden, 1997) and polyclonal antibodies selected against whole endospores of P. penetrans (Charnecki et al., 1998; Chen, S. Y et al., 1997; Davies et al., 1992; Persidis et al., 1991), but also by wheat-germ agglutinin (WGA) (Bird et al., 1989; Charnecki, 1997; Persidis et al., 1991). These results indicate that one or more epitopes detected by the antibodies may be glycosylated with 13-1-4 linked-acetylglucosamine.









40

Understanding the processes that lead to the growth, differentiation, sporulation, and maturation of P. penetrans within the pseudocoelom will likely provide a basis to establish the conditions required for its mass production in vitro. The objectives of this study were to (1) determine the synthesis of spore-associated proteins (adhesins) as a function of P. penetrans development within the pseudocoelom of the nematode host, M. arenaria race 1; (2) determine the distribution of an adhesin-associated epitope on the surface of virulent endospores; and (3) detect and localize an adhesin-associated epitope during the sporogenesis process.



Materials and Methods

Nematode Source

Meloidogyne arenaria (Neal) Chitwood race 1 used in this experiment was isolated originally from peanut (Arachis hypogea L.), Green Acres Research Farm, University of Florida, Alachua County, Florida. The nematode was reared on tomato (Lycopersicon esculentum Mill. cv. Rutgers) maintained in a greenhouse. Eggs of the nematodes were extracted from galled roots by dissolving the gelatinous matrix with

0.5% NaOCl for 20 seconds and collecting the eggs on a sieve with 75 pm-pore openings (200 mesh) nested in a sieve with 25-tm-pore openings (500 mesh) (Hussey and Barker, 1973). Second-stage juveniles were obtained by hatching the eggs in a modified Baermann funnel (Pitcher and Flegg, 1968). Juveniles (up to 3-day-old) were collected on an autoclaved 500-mesh sieve.








41

Pasteuria penetrans Source

Pasteuria penetrans strain P-20 (Oostendorp et al., 1990) used in this study was collected originally from females of M. arenaria race 1 parasitizing peanut in Levy County, FL and reared on M. arenaria race 1 growing on tomato in a greenhouse. One to three-day-old juveniles (J2), with endospores attached to their cuticles were obtained by incubating them with a suspension containing I x 10' endospores/ml overnight, with constant aeration at room temperature. Endospores were exposed to a mild sonification (FS14, Fisher Scientific, Suwanee, GA) for 5 minutes before attachment. Twenty sporeencumbered J2 were chosen randomly from a glass-slide mount, and the number of endospores attached per J2 was estimated with an inverted compound microscope at 400x. The percentage of endospores attached was 100% with an average of 7 3 endospores per juvenile. Tomato plants (45-day-old seedlings) growing in 15-cm-diam. clay pots, were inoculated with endospore-attached J2 (3,000 J2/plant). Three days later, the plants were inoculated again as before. Plants were fertilized twice a week by watering them with a solution containing 0.63 g of 20-20-20 (N-P-K) (Peters Professional, general purpose fertilizer, Division, United Industries Corp., St. Louis, MO) per liter. Water and insecticide applications were provided as needed. At 45 to 60 days after inoculation, the root systems were harvested, washed with tap water and weighed. Roots were cut into pieces 2 to 5 cm long and subjected to digestion in a 1-liter Erlenmeyer containing Rapidase Pomaliq 2F at 1:5 (g/v) (Gist Brocades Pomaliq product number 7003-A/DSM Food Specialities USA Inc., Menominee, WI), previously optimized with a buffer system (Chamecki, 1997), and agitated on a shaker at 120








42

oscillations per minute for approximately 24 hours at room temperature. Softened roots were placed in a sieve with 600 jim-pore openings (30 mesh) nested in a sieve with 150jim-pore openings (100 mesh) and sprayed with a heavy stream of tap water according to Hussey (1971), with modifications. Females and root debris were collected in a beaker by washing the sieve with a jet of deionized H20, and the contents centrifuged through 20% sucrose (w/v) at 1,500 x g for 5 minutes; the pellet fraction was centrifuged again through 47% sucrose (w/v) (Chen et al., 2000). The supernatant containing the females was collected in a beaker and the females were examined for P. penetrans infection with an inverted microscope at 100x. Endospore-filled females were hand-picked with forceps under a dissecting microscope at 40x (Nikon, Marietta, GA ), and placed in a 1.5 ml siliconized microtube containing 300 jll of deionized H20. Infected females were washed three times in deionized water by centrifugation at 10,000 x g for 2 minutes. Endospores were collected by grinding the females with a sterile pestle, and the suspension filtered through a nylon filter either with 21 gim or 18 jim openings (Spectra/Mesh). The concentration of endospores was determined by counting three 10 jtl aliquots using a hemocytometer (Fisher Scientific) at a magnification of 450x. Endospores retained on a sieve with 21 jim openings were stored at 4 'C, and used as inoculum for further production of the bacterium, whereas the endospores retained on a sieve with 18 im openings were stored at 20 'C and used for protein extraction. Experimental Design

Two sets of J2 of M. arenaria, one exposed and the other unexposed to P.

penetrans endospores, were compared with respect to development. These were arranged








43

randomly, with four replications per treatment per each designated "window of P. penetrans development" (harvest time: 12, 16, 24, and 38 days after inoculation). The windows of development were based on those reported by Hatz and Dickson (1992) and Serracin et al. (1997). 'Rutgers' tomato seedlings growing in a clay pot (10-cm-diam.) containing autoclaved sand were inoculated with 3,500 J2/plant ( 2 days old) with and without endospores attached. Plants were maintained in a growth chamber at 25 C for 48 hours to allow the nematodes to enter roots. After 48 hours the plants were removed from pots, and the roots washed thoroughly with tap water to remove any juveniles that had not penetrated. The seedlings were replanted in clay pots (1 5-cm-diam.), placed in a growth chamber at 35 C, and exposed to a 12-hour-day photoperiod. Plants were harvested at 12, 16, 24, and 38 days after inoculation. The root systems harvested from plants were washed in tap water, dried with a paper towel, weighed, cut into pieces 2 to 5 cm long, and incubated in an aqueous solution of commercial Rapidase Pomaliq 2F (Charnecki, 1997). Nematodes and softened roots were collected on a sieve with 600pam-pore openings (30 mesh) nested in a sieve with 25-ptm-pore openings (500 mesh), and washed as before. The nematodes were transferred to a sterile beaker, and twenty nematodes were hand-picked from each root system. To determine the percentage of nematodes infected by P. penetrans, and the stage of development of the bacterium from those nematodes, these were crushed individually in a 2.5 [LI drop of lactophenol and 1% methyl blue (w/v) (Sigma, St. Louis, MO) (Serracin et al., 1997) under a cover glass on a glass slide, and examined with an inverted microscope (Nikon) at 400x magnification. The remaining uninfected and infected nematodes from each harvest time were hand-








44

picked, washed, and stored in 1.5 ml siliconized microtubes containing 10 p.1 PBS (10 mM sodium phosphate buffer, 0.15 M sodium phosphate), pH 7.2 at -20 'C. Extraction and Determination of Proteins

Uninfected and P. penetrans-infected nematodes harvested at each interval after inoculation, and mature endospores (2 x 106 spores/ 10 .l PBS, pH 7.2) used as a control, were obtained as described before. Nematodes in 10 .l PBS, pH 7.2 were disrupted with a pestle, and then 30 lpl of the extraction solution containing 1.33x UDC (8M urea, 0.04 M dithiothreitol, 0.00665 M CHES buffer, pH 10) was added to each microfuge tube containing the samples. Microfuge tubes were placed into a water bath for 2 hours at 37 'C, and treated with 20 seconds of sonication (Brankson Cleaning Equipment Company, Shelton, CN) every 15 minutes. Extracts were centrifuged at 10,000 x g for 5 minutes at room temperature, and aliquots of the supernatant were collected for storage at -20 'C to carry out ELISA and SDS-PAGE analyses. Protein estimation was performed by a microprotein assay, based on the Bradford's method (Bradford, 1976) according to the manufacturer's instructions (BioRad, Hercules, CA). Standard curves were generated using bovine serum albumin (BSA) (Sigma), and colorimetric measurement was performed at 595 rum (Hewlett Packard 8451A Diode Array spectrophotometer, Palo Alto, CA). The extraction solution containing only urea and CHES buffer pH 9.8 was made previously, divided in 0.5 ml aliquots, and stored at -20 'C in 1.5 ml microtubes (Fisher Scientific), and then dithiothreitol was added to it just before the extraction of proteins.








45

Monoclonal Antibody

The anti-P-20 IgM monoclonal antibody (IgM MAb) used in this study was raised in mice against whole endospores ofP. penetrans P-20 strain and purified on a Sephacryl S-300 column (J. F. Preston and J. D. Rice, unpubl.). This monoclonal antibody showed the ability to block attachment of P. penetrans (P-20 strain) to the cuticle of M. arenaria race 1, and the IC50 is 1.3 x 10-0 M. It recognized an epitope shared on several polypeptides separated by SDS-PAGE (Brito et al., 1998; 2000 Charnecki, 1997; Charnecki et al., 1998).

Epitope Quantification by ELISA

Proteins (100 ng/well) extracted from P. penetrans-infected nematodes (either 13 infected nematodes harvested at 12 and 16 DAI or 5 infected nematodes harvested or 24 and 38 DAI) at each harvest interval, or from P-20 strain endospores alone as a positive control (2 x 106 endospores/pl), were applied to appropriate wells of a multi-well plate with 100 pl/well of coating buffer (15.00 mM Na2C03, 33.40 mM NaHC03, and 0.2% NaN3) added, and incubated overnight at 4 'C. After washing the wells four times with PBST (0.2% Tween 20 in 10 mM sodium phosphate buffer, pH 7.6; 154 mM NaC1), the first antibody, anti-P-20 IgM MAb diluted to 1:100,000 in PBST, was added to the appropriate wells (100 ll/well) and incubated for 1.5 hours at room temperature. Wells were washed with PBST again, and the secondary antibody, anti-mouse IgM-alkaline phosphatase conjugated (Sigma) diluted at 1:4000 in PBST was added to all wells, and incubated for another 1.5 hours at room temperature, and the wells were washed with PBST as before. Alkaline phosphatase substrate, 0.1% p-nitrophenol phosphate (w/v)








46
(Sigma) in alkaline phosphatase substrate buffer (0.05 M Na2C03, 0.05 M NaHC03,

0.0005 mM MgCI2) was added to all wells, and color development was measured with an automated microplate reader at 405 nm (BioRad model 2550, Hercules, CA). SDS-PAGE Analysis

Proteins (600 ng of total healthy or infected nematode protein) in an appropriate volume of 10 mM PBS, pH 7.2, were combined with an equal volume of sample buffer (50 mM Tris/HC1, pH 6.8, 2% SDS w/v, 10% glycerol, 0.05% bromophenol blue w/v, 2% f-mercaptoethanol), and boiled for 5 minutes at 100 oC, and then centrifuged for 5 minutes at 10,000 x g. Endospore protein that was extracted from P-20 isolate (2 x 106 endospores/pl) alone was used as a control. Twenty microliters of the supernatants were transferred into appropriate wells of a polyacrylamide gel of 4% stacking gel (pH 6.8) and 12% separating gel (pH 8.8) with Tris-glycine buffer (Laemmli, 1970). A prestained molecular weight marker (SeeBlue TM Prestained Standards, Novel Experimental Technology, San Diego, CA) was loaded onto the same gel. Electrophoresis was carried out at 100 V for 10 minutes, and then was set at 200 V until the dye marker moved to the bottom of the gel. Gels were electro-blotted onto nitrocellulose membranes in blotting buffer (192 mM glycine, 25 mM Tris, 20% methanol) using a Mini Transfer-blot Cell (BioRad, Hercules, CA) at a constant voltage, 50 V for 2 hours. Proteins either were stained with AuroDye according to the manufacturer's instructions (Amersham, Piscataway, NJ) or with anti-P-20 IgM Mab.








47
Immunoblotting

The nitrocellulose membranes were blocked with 0.5% non-fat dry milk (w/v) in PBST (10 mM sodium phosphate buffer, pH 7.2, 150 mM NaCl, 0.2% Tween 20) overnight at 4 oC. Polypeptides containing the epitope recognized by anti-P-20 MAb were detected as follows: incubation of the membranes with anti-P-20 IgM MAb at 1: 2,000 in PBST, pH 7.2 for 1.5 hours at room temperature on a shaker, washed three times for 5 minutes each with PBST; incubated with goat anti-mouse IgM MAb conjugated to alkaline phosphatase (Sigma) diluted to 1:1,000 in PBST, pH 7.2 as secondary antibody for 1.5 hours at room temperature on a rotatory shaker, followed by three washes with PBST as above; incubation with substrate buffer (100 mM Tris-HCI pH 9.5, 100 mM NaC1, 5 mM MgCl2) three times, five minutes each; incubated with alkaline phosphatase substrate (0.1 mg/ml nitrotetrazolium blue, 0.05 mg/ml 5-bromo-4-chloro-3-indolyl phosphate) (Promega, Madison, WI) in substrate buffer on a shaker at room temperature until color development. The blots were washed with deionized water and dried at room temperature.

Immunofluorescence of Whole Endospores

The immunofluorescence staining was performed as described by Pogliano et al. (1985) with modifications. Fresh endospores were washed and purified as before, and then filtered through a woven polyester filter with 18 pim openings. Twenty microliters of the endospore suspension (2 x 106 endospore/tl) were transferred to a 1.5 ml siliconized microtube, and fixed in 230 gl of the primary fixative containing 2.7% formaldehyde and 0.008% glutaraldehyde in 10 mM PBS (10 mM sodium phosphate








48
buffer, pH 7.4, 150 mM NaCI) for 35 minutes on ice. Endospores were placed in 250 tl 10 mM PBS, pH 7.4 and then centrifuged at 6,000 x g three times for 6 minutes each. After resuspending the endospores in 150 jtl PBS, 10 tl of the suspension was transferred into each of three wells of a microscope slide which had been treated previously with

0.1% poly-L-lysine (Sigma). Each slide was incubated for 30 seconds at room temperature and then the suspension was aspirated from the wells with a sterile-transfer pipette (Fisher Scientific). After air drying at room temperature for 30 minutes, the endospores were incubated in a 10 .l/well with in PBST-BSA (2% BSA (w/v) and 0.05% Tween 20 (v/v) in 10 mM PBS, pH 7.4 ) for 15 minutes at room temperature to block nonspecific antibody-binding sites. Primary antibody, anti-P-20 IgM MAb diluted to 1:1,000 in PBST-BSA, was added to the wells and incubated overnight at 4 'C. Wells containing the endospores were washed in PBST, pH 7.4, five times for 5 minutes each, and incubated for 2 hours in the dark at room temperature with micron chain-specific, anti-mouse IgM conjugated with fluorescein isothiocyanate (FITC, Sigma, 1:100 diluted in PBST-BSA). Anti-P-20 IgM MAb was substituted with non-immune ascites fluid at 1:1,000 dilution as negative control. After washing the wells with 10 mM PBS, pH 7.4, 10 times for 5 minutes each, the slides were mounted in Slow Fade in a PBS-glycerol solution (Molecular Probes Inc., Eugene, OR). Preparations were examined with differential-interference contrast and fluorescence microscopy using a Nikon Episcopic Fluorescence attachment with an excitation filter at 495 nm.









49
Tissue Prearation for Sectioning

Uninfected and P. penetrans-infected M arenaria race I harvested at 20 days

after inoculation at 35 'C were obtained as described above. The procedure used to carry out this study was a modification of the work by Aldrich et al. (1995); Chen et al. (I1997a); and Zeikus and Aldrich (1975). Fresh nematodes were ruptured with a surgical knife (Fisher Scientific No. 15) into 40 gl of fixative (1% glutaraldehyde, 4% formaldehyde, 5 % dimethyl sulfoxide in 0. 1 M sodium cacodylate buffer, pH 7.2) to facilitate the penetration of reagents, and then embedded in 2.5% low temperature gelling agarose (Fisher Scientific) at 45 'C and congealed in the refigerator (4 'Q). The gel was sliced into square blocks containing individual nematodes and transferred into 12 x 75 mm culture tubes (Fisher Scientific) containing 1.5 ml of the above-mentioned fixative, and incubated overnight at 4 'C. Agar blocks containing nematodes were washed four times with cold 0. 1 M cacodylate buffer on ice for 30 minutes each and dehydrated in a cold ethanol series containing the following percentages: 12, 25, 38, 50, 65 for 20 minutes each, and then 75 overnight at 4 'C. This was followed by 85, 95 and two changes of 100% ethanol for 20 minutes each. The specimens were embedded in LR White Resin (London Resin White, Electron Microscopy Science, Fort Washington, PA) series (25 and 50% for 3 and 6 hours, respectively, and 75%, 100%, and 100%, overnight each time). Agar blocks containing nematodes were transferred into a 1 -ml gelatin capsule containing LR White, and allowed to polymerize at 50 'C for 4 days. Ultrathin sections (50-70 nm thick) were cut from the resin block with a diamond knife on a LKB








50
8800 Ultratome III microtome (Sweden). Sections were collected on Formvar-coated nickel grids (100 mesh), and processed for immunogold labeling. Immunogold Labeling

Nickel grids with sections of uninfected and P. penetrans-infected nematodes, and with endospore-attached juveniles were floated, section-side down, on 20-tl drops of 1% non-fat dry milk in PBS, pH 7.2 (0.01M sodium phosphate buffer, 0.15 M sodium chloride, pH 7.2) on a piece of Parafilm (American National Can TM, Menasha, WI) for 15 minutes at room temperature to block nonspecific antibody-binding sites (modified from Aldrich et al. 1992, 1995; Dykstra, 1993). Grids were floated on 20-pl drops of primary antibody, anti-P-20 IgM MAb at 1:10,000 dilution in PBS, pH 7.2, and incubated overnight in a closed petri dish inside a moist chamber at 4 'C. Control grids were floated on non-immune ascites fluid at 1:10,000 dilution instead of anti-P-20 IgM MAb. Grids were removed, and floated on 20-jtl drops of high salt-Tween buffer, pH 7.2 (0.1% Tween 20 in 0.02 M Tris-HC1, pH 7.2, 0.5 M Na Cl), two times for 10 minutes each, and then PBS, pH 7.2, two times for 10 minutes each. Sections were incubated with the secondary antibody, goat anti-mouse IgM conjugated to 12-nm colloidal gold particle, jtchain specific (Jackson Immuno Research, West Grove, Pennsylvania), diluted 1:30 in PBS, pH 7.2, at room temperature for 1 hour. After washing as above in high salt-Tween buffer and PBS, the grids were floated in Trumps buffer, pH 7.2 (McDowell and Trump, 1976) for 10 minutes at room temperature in order to stabilize the antigen-antibody complex, and then washed with deionized water. Sections were stained with 0.5% uranyl








51
acetate for 7 minutes, and aqueous lead citrate solution for 2.5 minutes and observed on a Zeiss EM-10 transmission electron microscope at 80 kV. All reagents used to carry out this study were ultrapure-TEM grade.


Results

Microscopic Examination

The vegetative growth stage of P. penetrans was observed only in nematodes harvested at 12 and 16 days after inoculation (Table 2.1). At 24 days after inoculation, mixed developmental stages of thalli showed advanced differentiation, including quintets, quartets, triplets, doublets; sporulation, oval-shaped immature sporangium; and mature endospores with visible exosporium were first observed. At 38 days only various phases of sporulation and mature endospores were present in the pseudocoelom of M. arenaria race 1.

Epitope Ouantification by ELISA

The anti-P-20 IgM MAb did not recognize proteins extracted from infected nematodes harvested at 12 and 16 days after inoculation (Fig. 2.1A). However, the monoclonal antibody reacted with proteins extracted from infected nematodes harvested at 24 and 38 days after inoculation (Fig. 2.1A). The protein per infected nematode was 0.453 jig at 12; 0.466 jig at 16, 1.175 jig at 24, and 2.049 jig/nematode at 38 days after inoculation (Fig. 2.1 B). The total protein per infected nematode increased with developmental time (Fig. 2.1 B), and was correlated with the increase in the signal detected by the anti-P-20 IgM MAb (Fig. 2.1A). At 24 and 38 days after inoculation, the ELISA-based absorbance at 405 jim per infected nematode was 1.50 and 3.20,








52
Table 2.1. Percentage of different developmental stages of Pasteuria penetrans in Meloidogyne arenaria race 1 on tomato 'Rutgers' at 12, 16, 24, and 38 days after inoculation at 35 oCa.

Days Postinoculation


Developmental stage 12 16 24 38

Vegetative growth 90 90 0 0

Differentiation 0 0 15 0

Sporulation 0 0 85 5

Mature endospores 0 0 65 95

aTwenty nematodes were observed at each harvest date, and percentage of
nematodes at 12, 16, 24, and 38 days after inoculation. Nematodes were hand-picked, placed on a glass slides, and crushed separately in 2.5 ptl of lactophenol plus 1% methyl blue (w/v) under a cover glass. Infected nematodes were examined with the use of an inverted microscope (x400) to determine the percentage of the different developmental stages of P. penetrans within the pseudocoelom of Meloidogyne arenaria race 1. Note that at 24 days after inoculation more than one developmental stage was observed within the pseudocoelom of a single nematode. The developmental stages observed were: vegetative growth including mycelial colonies only within the pseudocoelom; differentiation stage, with presence of thalli differentiation, including quintets, quartets, triplets, doublets; sporulation stage, with many doublets and developing endospore with distal swollen ends connected by intercalary ends; and mature endospores; with free endospores with exosporium clearly visible.








53
respectively, which was proportional to the amount of adhesin-associated epitope increased as P. penetrans reached its maturation stage (Fig. 2.1 A). These results suggest that the antigens bearing the epitope, which was recognized by anti-P-20 IgM MAb, were synthesized at later stages of development associated with sporulation of P. penetrans within the pseudocoelom of M. arenaria race 1. SDS-PAGE Analysis and Immunoblotting

Analysis of individual proteins extracted from uninfected and P. penetransinfected nematodes at each window of development showed some differences in the protein profiles related to the infection of the nematode by the bacterium (Figs. 2.2A-B;

2.3A-B). The immunoblot showed that Anti-P-20 IgM MAb did not recognize any protein extracted from uninfected nematodes harvested at 12, 16, 24, and 38 days after inoculation (Lanes 2, 3, 4, and 5) (Fig. 2.2B); nor were proteins extracted from infected nematodes harvested at 12 and 16 days detected in the immunoblot (Lanes 2, and 3) (Fig.

2.3B). However the immunoblot revealed that the monoclonal antibody reacted with protein extracts of infected nematodes harvested at 24 and 38 days after inoculation (Lanes 4, 5) (Fig. 2.3B) and with endospore protein of the P-20 strain used as the control (Lanes 6) (Figs. 2.2B; 2.3B).

Immunofluorescence

Note the general shape of P-20 strain just before it was examined with the

fluorescence microscope (Fig. 2.4A). Labeling of whole endospores of P. penetrans








54







S4
-o
0
E 3




2 1 2... .1 6....2 4....3 8.
2


0



12 16 24 38




=-3
0


"a
0







12 16 24 38

1 Days after inoculation


Fig. 2.1. Adhesin-associated epitope and total nematode protein per infected
nematode as a function of the development of Pasteuria penetrans. A) Levels of adhesinassociated epitope determined by ELISA using anti-P-20 IgM MAb at 1:100,000 dilution in PBST, pH 7.6. Infected nematode total proteins (100 ng/well) was applied in 100 gl/well at the final treatment. Alkaline phosphatase substrate, 0.1% p-nitrophenol phosphate (w/v) was added to all wells, and color development was measured at 405 nm. B) Total nematode protein of infected nematodes. Data shown are 40 minutes readings. Lines above the bars indicate SE of the mean for six replicates per treatment.








55




kDa
1 2 3 4 5 6
250
A
98
64-
50-- .
36-
30-






1 2 3 4 5 6
250-
B 98-64-

50-
36-






Fig. 2.2. Blots of sodium dodecyl sulfate-polyacrylamide gels of uninfected Meloidogyne arenaria protein extracts after electrophoresis. Proteins of uninfected nematodes, harvested at each window of development. Extracts in the appropriate volume of sample buffer were boiled for 5 minutes at 100 'C, and 20 pt of the appropriate extract containing 600 ng of total protein was applied per lane. A) Proteins were detected by staining with AuroDye according to manufacturer's instructions. B) Immunodetection of blotted antigens with anti P-20 IgM MAb at 1: 2,000 dilution in PBST, pH 7.2. Lane 1 Molecular weight markers, See Blue pre-stained proteins; Lanes 2, 3, 4, and 5 Total proteins extracted from uninfected nematodes at 12, 16, 24, and 38 days after inoculation; Lane 6 Proteins extracted from P. penetrans P-20 endospores.








56



kDa
1 2 3 4 5 6
250-
A 98-64-
50-
36-

30-



1 2 3 4 5 6
250-- m
B 98-64-

50-
36-








Fig. 2.3. Detection of Pasteuria penetrans adhesin-associated epitope as a
function of its development within the pseudocoelom of Melodogyne arenaria racel. Nematode total proteins and endospore proteins were extracted as for Fig 2. Proteins, 600 ng in 20 pla of the appropriate extract plus sample buffer was loaded into each lane. A) Detection of blotted proteins with AuroDye. B) Western blot of P. penetrans infected nematodes probed with anti-P-20 IgM MAb at 1:2,000 dilution in PBST, pH 7.2. Lane
1 Molecular weight markers, See Blue pre-stained proteins; Lanes 2, 3, 4, and 5 Epitope bearing proteins extracted from P. penetrans infected nematodes at 12, 16, 24, and 38 days after inoculation; Lane 6 Proteins extracted from P. penetrans P-20 endospores.








57
isolate P-20 by anti-P-20 IgM MAb was not uniform (Fig. 2.4B), which suggests that the adhesin-associated epitope is not uniformly distributed on the surface of mature endospores.

Immunogold Labeling

Anti-P-20 IgM MAb did not recognize any nematode tissue and there were no gold particles observed over the thin section of either uninfected females or J2 with associated endospores (Fig 2.5). The adhesin-associated epitope was not present in the ultrathin sections of vegetative cells (vc) or stage I (Fig. 2.6A) or in stage II of sporogenesis of P. penetrans isolate P-20 (Fig. 2.6B). Note a membrane (arrow head) is forming at 1/3 from the anterior, which occurs at this stage of sporogenesis (Chen et al., 1997b). Labeling of the adhesin-associated epitope was first observed over an ultrathin section of the stage III sporogenesis, mainly on the parasporal fibers (pf) (Fig. 2.7A). The antigens bearing the epitope were detected not only over the parasporal fiber (pf) (Figs.

2.7B-2.9A) but also over the sporangium(s) as P. penetrans continues to sporulate (Figs.

2.8A-B; 2.9A). The mature endospore was heavily labeled, and the epitope was localized in the sporangium (s), exosporium (ex), and parasporal fibers (pf) (Fig. 2.9A). The outer spore coat (oc), inner spore coat (ic), cortex (c), protoplasm (p), and basal ring (br) were not labeled (Fig. 2.9A). No labeling was observed over any structure of the mature endospore when non-immune ascites fluid was used (Fig. 2.9B).








58




e

Ao


















B











Fig. 2.4. Differential interference contrast (DIC) and fluorescence microscopy photomicrographs of whole endospores of Pasteuria penetrans P-20 isolate (100x magnification). A) Overall shape of whole endospores using DIC. B) Labeling of an adhesin-associated epitope on the surface of whole endospores using anti-P-20 IgM MAb at 1:1000 dilution in PBST-BSA, overnight at 4 oC, as primary antibody, and anti IgM Mab-FITC labeled as secondary antibody diluted 1:1000 in PBST-BSA. Arrows heads identify regions of nonuniform labeling.






59




































Fig 2.5. Longitudinal section of uninfected second-stage juvenile of Meloidogyne arenaria (1 -day-old) probed with anti-P-20 IgM MAb at 1:10,000 dilution, and anti-IgM, gold-conjugated at 1:30 dilution. No gold particles are visible over the nematode tissues. Scale Bar = 0.5 um.


























Fig. 2.6. Immunocytochemical localization of an adhesin-associated epitope
during the development of Pasteuria penetrans within the pseudocoelom of M. arenaria. Thin sections of all stages of development of P. penetrans were probed with anti- P-20 IgM MAb at 1:10,000 dilution, and anti-IgM MAb gold-conjugated diluted to 1:30 dilution as secondary antibody and examined by transmission electron microscopy. Scale Bars = 0.5 gm. A) Stage I of sporogenesis. A longitudinal ultrathin section of mycelial colony (arrow) of P. penetrans P-20 isolate. No labeling is visible over the mycelium. B) Stage II sporogenesis of P. penetrans. Note that a membrane is forming at 1/3 distance from the anterior end (arrow read), which is characteristic of this stage. No labeling occurs over any structure of this stage of development of the bacterium.









61







































2! 1. lwv





























Fig 2.7. Labeling of sporogenous stages of Pasteuriapenetrans. Scale bars = 0.5 gim. A) Stage II sporogenesis showing labeling of the adhesin-associated epitope (arrow head) mainly over the parasporal fibers (pf). B) Stage IV sporogenesis, gold particles (arrow head) are concentrate in the parasporal fibers (pf). Note that the vegetative cell
(vc) was not labeled.











63



















k Pf



4r, -ip f






























Fig 2.8. Sporogenous stages of Pasteuria penetrans. Scale bars = 0.5 gm. A) Stage V of sporogenesis. Gold label (arrow head) indicating antibody binding is present over the parasporal fibers (pf) and exosporium (e). B) Stage VI of sporogenesis, labeling of the adhesin-associated epitope is observed over the parasporal fibers (pf) and exosporium (ex).










65











Lp




tv t4









; 7,%A








Pf .










Pf




























Fig. 2.9. Late sporogenous stage of Pasteuria penetrans. Scale Bars = 0.5 gm. A) Stage VII of sporogenesis, a mature endospore showing the sporangium (s) exosporium(ex), and parasporal fibers (pf) heavily labeled, whereas the outer spore coat
(oc), inner spore coat (ic), epicortex (ep), cortex (c), protoplasm (p), and basal ring (br) are not labeled. Note that the parasporal fibers (pf) were not uniformly labeled (arrow head). B) A mature endospore of the Pasteuria penetrans, stage VII used as control. No label is observed over the thin section of the endospore.











67























ex.
41









68
Discussion

Pasteuria penetrans completes its life cycle within the pseudocoelom of female of root-knot nematodes. The physiological aspects of its life cycle have been studied and are reasonably well understood (Chen and Dickson, 1997; Freitas et al., 1997; Hatz and Dickson, 1992; Serracin et al., 1997; Nakasono et al. 1993, Stirling, 1981, Giannakou et al., 1999). However the biochemical aspects are poorly understood. Seven morphological stages of development through sporulation have been determined as I, II, II, IV, VI, and VII (Chen and Dickson, 1997). The initial step in the life cycle of Pasteuria is the recognition/attachment of the endospores to the cuticle of a free living J2 root knot-nematode host. Infection of the host and germination of the endospores occur once the J2 enters the root tissue of a plant host, and establishes a permanent feeding site (Sayre and Starr, 1985, 1988). Vegetative growth, differentiation, and formation of Imbriani, 1975; Sayre, 1993; Sayre and Wergin, 1997; Serracin et al., 1997). The mechanisms involved in the attachment have been the subject of study in several laboratories. The results of these studies have led to the establishment of a model where glycoproteins, designated as adhesins and lectin are involved in the interaction of P. penetrans and the nematode host (Persidis et al., 1991; Davies and Danks, 1993). Previous studies have shown that microbial adhesins, or bacterial surface proteins, known as virulence factors such as pili, or fimbriae (Robins-Browne et al., 1994; Salyers and Whitt, 1994; Suoniemi et al., 1995), and afimbrial adhesins (Salyers and Whitt, 1994), allow bacteria to attach, colonize, and invade their hosts. For instance, Streptococcus pyogenes, a gram-positive pathogen has a nonfibrillar adhesin (protein F) that mediates its









69
attachment to fibronectin, a protein found on many host cell surfaces, including the mucosa of the human throat (Salyers and Whitt, 1994). However, the mechanisms used by Pasteuria spp. to produce virulent endospores within the pseudocoelom of the nematode host is not well understood. Mohan et al. (2001) found that fibronectin-like proteins extracted from M. javanica are involved in the attachment of endospores. In this study, we determined the relative time of the synthesis of an adhesin-associated epitope during the development of P. penetrans within the pseudocoelom of M. arenaria race 2; detected and localized this epitope during endospore development, and also determined the distribution of the epitope on the surface of mature endospores using a monoclonal antibody directly selected against whole mature enendospores of P. penetrans P-20 isolate.

ELISA and immunoblot analysis revealed that only proteins extracted from P. penetrans-infected nematodes at 24 and 38 days after inoculation were recognized by anti- P-20 IgM MAb and the amount of the epitope was highest at the height of sporulation (38 days after inoculation) than at any other developmental stage (12, 16, and 24 days after inoculation). The Western blot showed a higher degree of similarity in the protein profile ofP. penetrans-infected nematodes at 38 days after inoculation to the mature P-20 spore protein, used as a control, than with P. penetrans-infected nematodes from any other window of development. Examination of the infected nematodes harvested at 12 and 16 days by light microscopy revealed that only the vegetative growth stage, including clusters of mycelial colonies and thalli, were found throughout the pseudocoelom of nematodes. At 24 and 38 days, sporulation and maturation stages were









70
observed within the pseudocoelom. Therefore, the synthesis of the adhesin-associated epitope occurred at a certain developmental stage relative to the sporogenesis process, and it was absent in vegetative growth and differentiation stages.

The synthesis of specific molecules at specific times during the germination,

growth, and sporulation of the endospore-forming bacterium, Bacillus subtilis, has been rigorously established. For instance dipicolinic acid (pyridine-2, 6-dicarbonate) is formed during the first 5 hours of sporulation (Schlegel, 1986), whereas the small, acid-soluble spore proteins (SASPs), a group DNA-binding proteins (at neutral to slightly alkaline pH), are synthesized after 3-4 hours into sporulation (Johnson and Tipper, 1981; Setlow 1985). Both molecules are found only in endospores (Fliss et al., 1985; Schlegel, 1986). Even though some molecules are synthesized at specific stages of sporulation, it is possible that they are degraded and used to carry out a certain function at another stage. For instance, during the first 5 hours of sporulation in B. subtilis much of the vegetative cell protein is degraded (Schlegel, 1986).

Immunofluorescence labeling showed that the adhesin-associated epitope is not uniformly distributed on the surface of virulent endospores. The heterogeneity of endospore surface has been observed not only within populations but also between populations of P. penetrans (Davies and Redden, 1997). Previous studies have shown that differences in the amount and nature of spore-surface proteins, as recognized by several monoclonal antibodies, may account for surface heterogeneity of endospores as well as host specificity (Davies et al., 1992). Davies et al. (1994) using monoclonal antibodies showed that the surface of endospores of the PP 1 strain of P. penetrans is









71
highly heterogenous. These and subsequent studies (Davies and Redden, 1997) have suggested that endospore surface properties are responsible for the virulence of P. penetrans.

Antigens bearing the epitope were synthesized during the sporogenesis process. Labeling was first observed at stage E of the sporogenesis, mainly in the parasporal fibers. In contrast to stage III sporogenesis, mature endospores were heavily labeled and the adhesin-associated epitope was localized in the parasporal fibers, sporangium, and exosporium.

The general pattern of the labeling of the adhesin-associated epitope over thin

sections of a mature endospore was similar to a previous study, where mature endospores were probed with a polyclonal antibody (Persidis et al., 1991). These authors concluded that the labeling did not show any preference to a certain structure of the endospore and suggested that a nonspecific binding of the antibodies could have occurred. These observations may reflect a heterogeneity in the polyclonal antibody preparation and/or selection of a single stage of development. In our immunocytochemistry work, it was shown that the adhesin-associated epitope is synthesized at a certain stage of development related to endospore formation and it is localized initially in the parasporal fibers early in stage I, becoming widespread throughout the sporangium and exosporium, but not in the central body of the stages IV, V, VI, and VI of sporogenesis. Label was not uniformly distributed in the parasporal fibers. Also no labeling was observed in the outer or inner spore coat, epicortex, cortex, protoplasm, and basal ring.









72
These observations establish a window of development in which the adhesinassociated epitope is formed, and where further studies concerning the formation of this epitope should be directed. The fact that the epitope is distributed over several structures of the mature endospores suggests its involvement in the recognition of the nematode host as an early event in the attachment process. It may increase the chances for a cooperative interaction between the adhesin epitope with receptors on the cuticle of the nematode host, such as carbohydrate binding proteins (Bird et al., 1989; Davies and Danks, 1993; Persidis et al., 1991) and fibronectin-like residues. (Mohan et al., 2001) as well as other forces, that may be involved in the attachment, such as hydrophobic interactions (Afolabi et al., 1995; Davies et al., 1996; Esnard et al., 1997).














CHAPTER 3
DETECTION OF ADHESIN PROTEINS AND IMMUNOLOGICAL
DIFFERENTIATION OF Pasteuria spp. USING A MONOCLONAL ANTIBODY Introduction

Pasteuria penetrans (Thorne) Sayre & Starr, the first species of Pasteuria

described as a parasite of plant-parasitic nematodes, is a widespread endospore-forming bacterial parasite of root-knot nematodes (Meloidogyne spp.) (Sayre and Starr, 1985). Over the years, several more species of nematodes in other genera have been reported as hosts of species of Pasteuria. (Chen and Dickson, 1998). To date, three species of Pasteuria have been described in addition to Pasteuria penetrans (Sayre and Starr, 1985). These are Pasteuria ramosa, a parasite of water fleas, Daphnia spp. (Sayre et al., 1983) which is the type species of the genus; Pasteuria thornei isolated from Pratylenchus spp. (Starr and Sayre, 1988), and Pasteuria nishizawae a parasite of cyst nematodes of the genera Heterodera and Globodera (Sayre et al.,1991). In recent years more species of Pasteuria have been proposed: i) Pasteuria sp., designated as S-1 strain (Bekal et al., 2000) from Belonolaimus longicaudatus Rau; ii) a large- and a small-spored isolate of Pasteuria spp. each from Hoplolaimus galeatus (Cobb) Thorne (Giblin-Davis et al., 1990); and iii) three isolates which attach and complete their life-cycles in Heterodera spp.; one isolate was from cereal cyst nematode, Heterodera avenae Wollenweber (Davies et al., 1990), a second strain from pea cyst nematode, Heterodera goettingiana



73









74

Liebscher in Mtinster, Germany (Sturhan et al., 1994), and a third isolate that infects soybean cyst nematode, Heterodera glycines Ichinohe, Pasteuria sp. NA (Heterodera glycines-infecting Pasteuria), Urbana, IL, USA (Atibalentja et al., 2000).

Traditionally species of Pasteuria are identified based on morphometrics,

morphology, ultrastructural characteristics, and host specificity (Davies et al., 1990; Giblin-Davis et al., 1995; Sayre and Starr, 1985; Sayre et al., 1983; 1991; Starr and Sayre, 1988; Sturhan et al., 1994). More recently, 16S rDNA has been used to carry out systematics studies of P. ramosa (Ebert et al., 1996), P. penetrans (Anderson et al., 1999), Heterodera glycines-infecting Pasteuria (Atibalentja et al., 2000), and Pasteuria sp. S-1 strain (Bekal et. al., 2000). Also, the use of serology through hybridoma technology might be a useful probe for the identification of Pasteuria spp. The anti-P-20 IgM monoclonal antibody (MAb) raised against whole mature endospores of P-20 isolate of P. penetrans was used as a probe in this study. This MAb was selected on the basis of its ability to block attachment of P. penetrans isolate P-20 to M. arenaria race 1 (Charnecki et al., 1998) (Chapter 2). Previous studies have shown that this MAb recognized an epitope shared on several polypeptides separated by SDS-PAGE (Brito et al., 1998; Charnecki 1997; Chamrnecki et al., 1998). The appearance of an adhesinassociated epitope was tracked during development and localized during sporogenesis of the P-20 within its nematode host (Brito et al., 1998; 1999). The objectives of this study were to determine whether different strains and species of Pasteuria share this adhesin-









75

associated epitope which is involved in the attachment of P. penetrans P-20 strain to M arenaria race 1, and to use anti-P-20 IgM MAb as a probe to separate strains and species of Pasteuria.



Material and Methods

Origin of Pasteuria Species and Isolates

The designations and origins of the species and isolates of Pasteuria spp. (Table 3.1) were as follows: two isolates of P. penetrans; one designated P-20 (Oostendorp et al., 1990) originally collected from M arenaria race 1 (Neal) Chitwood, from peanut (Arachis hypogea cv. Florunner) roots growing in a naturally infested field in Levy County, FL, and the other one designated P1 -UFLA (Souza and Campos, 1997), originally isolated from a mixed population of M javanica and M incognita, Lavras, Minas Gerais, Brazil; H. glycines-infecting Pasteuria, (Pasteuria sp. NA) (Atibalentja et al., 2000) from cysts of H. glycines collected from the rhizosphere of soybean plants (Glycines max (L). Mirril), Urbana, IL. Pasteuria sp. strain S-1 (Bekal, et al., 2001; Giblin-Davis et al., 2001) isolated from the sting nematode B. longicaudatus, L-1 (largespored strain), LS-1 (small- spored strain) from the lance nematode, H. galeatus (GiblinDavis et al., 1990), and Pasteuria from Rhabditis sp. (Giblin-Davis pers. comm.) were all originally collected from bermudagrass (Cynodon spp.) turf growing in a naturally infested field, at the Ft. Lauderdale Research and Education Center, University of Florida, Broward County, Fort Lauderdale, FL. Pasteuria sp. C-1 isolate (Han et al., 1999) was originally collected from Criconemoides sp. in a naturally infested soil where peanut









76

(Arachis hypogea L. cv. Florunner ) was growing at the Green Acres Agronomy Farm, University of Florida, Alachua County, Gainesville. A ring nematode isolate of Pasteuria also isolated from Criconemoides sp. collected in a peanut field (Williston), FL (Dickson per. comm.), and spiral nematode isolate of Pasteuria isolated from Helicotylenchus sp. extracted from the rhizosphera of bermudagrass turf from California (Crow, pers. comm.).

Propagation of Bacterial Species and Isolates

Pasteuria penetrans P-20 and P 1-UFLA isolates were propagated on M. arenaria race 1 and M. javanica respectively, growing on 'Rutgers' tomato. Endospores of each strain were attached separately to second-stage juveniles (J2) (up to 2 day old) of root-knot nematodes using a centrifugation method (Hewlett and Dickson, 1993). Juveniles (3,000 J2 per plant) with approximately six endospores attached per J2 were inoculated on 55-day-old tomato plants growing in 15-cm-diameter clay pots in a greenhouse. Endospore-filled females were harvested from the root systems 45 to 60 days after inoculation. Root systems were placed in a 1-liter Erlenmeyer flask containing Rapidase Pomaliq 2F (Gist Brocadest Pomaliq, 7003-A/DSM Food Specialities USA Inc., Menominee, WI) optimized previously with a buffer system at 1:5 (g/v) (Charnecki, 1997), and placed on a shaker at 120 oscillations/minute for approximately 24 hours at room temperature. The softened roots and nematodes were poured onto a sieve with 600 gm-pore openings nested in a sieve with 150 pm-pore openings, and subjected to a heavy stream of tap water to dislodge the nematodes (Hussey, 1971), with modifications. Nematodes and root debris were collected in a beaker, and the contents centrifuged in









77

Table 3.1. Species and isolates of Pasteuria. Species or isolates Reference

P. penetrans P-20 Meloidogyne. arenaria race 1 (Oostendorp et al.,

1990)

P. penetrans P I-UFLA Meloidogyne spp. (Souza and Campos, 1997)

Hg Pasteuria sp. NA Heterodera glycines (Atibalentja et al., 2000)

Pasteuria sp. S-1 Belonolaimus longicaudatus (Bakel et al., 2001)

C-1 isolate Criconemoides sp. (Han et al., 1999)

L-1 isolate Hoplolaimus galeatus (Gibli-Davis, 1990)

LS-1 isolate Hoplolaimus galeatus (Gibli-Davis, 1990)

Rhabditis infecting-Pasteuria Rhabditis sp. (Giblin-Davis, pers. comm.) Ring nematode-infecting Pasteuria Criconemoides (Dickson, pers. comm.) Spiral nematode-infecting Pasteuria Helicotylenchus sp. (Crow, pers. comm.)









78

20% sucrose (w/v) at 1,500 x g for 5 minutes, and the resulting pellet was again centrifuged in 47% (w/v) sucrose (Chen et al., 2000). Female nematodes were collected and examined for Pasteuria infection with an inverted microscope at 100 x magnification (Leica, Davie, FL). Pasteuria-infected females were hand-picked using a dissecting microscope at 40 x magnification (Nikon, Marietta, GA), and placed in 1.5 ml siliconized microtubes containing 900 pl of deionized water. Nematodes were centrifuged in deionized water three times at 10,000 x g for 2 minutes, and then stored in 500 pl deionized water at 4 oC until used. Pasteuria sp. S-1, L-1, LS-1 isolates, and the Rhabditis sp. infecting-Pasteuira, and spiral nematode-infecting Pasteuria were isolated from their nematode hosts growing in bermudagrass (Cynodon dactylon (L) X C transvaalensis Burt-Davy cv. Tifway or C. magenissii Hurcombe cv. Tifgreen) turf in a naturally-infested field. The C-1 isolate and ring nematode-infecting Pasteuria were obtained from Criconemoides sp. extracted from the rhizosphere of peanut (Arachis hypogea L. cv. Florunner) grown in a naturally-infested soil in a greenhouse, and peanut field, respectively. All nematodes were extracted from the soil using a centrifugalflotation method (Jenkins, 1964). Pasteuria-infected nematodes were hand-picked under a dissecting microscope, and placed in deionized water. After washing the nematodes with deionized water as above, they were stored in 900 pl deionized water at 40 C until used. Pasteuria sp. NA was propagated on H. glycines race 3 reared on soybean cv. Lee growing in a naturally-infected soil in a greenhouse. Pasteuria-infected cysts and females were extracted from the rhizosphere of 3-month old soybean plants by washing the soil through a sieve with 850 g-pore openings nested over a sieve with 180 u-pore openings;









79

and nematodes were collected in a sterile beaker. Nematodes were transferred into 200ml centrifuge tubes containing 150 ml of deionized water, and centrifuged at 2,000 x g for 4 minutes. The resulting pellets were re-suspended with 50% sucrose solution, and again centrifuged for 35 to 45 seconds. The supernatant was poured through a sieve with 180 g-pore openings (Atibalentja et al., 2000), and collected in a sterile beaker. Infected females and cysts were hand-picked based on their opaque appearance, washed three times with deionized water by centrifugation at 10,000 x g for 2 minutes, placed in a 1.5 ml siliconized microtube containing 100 jtl deionized water, and stored at 4 'C until used.

Extraction and Determination of Proteins

Nematodes infected by species or isolates P-20, P1 -UFLA, S-l, C-l, ring

nematode and spiral nematode isolates of Pasteuria, and cysts infected with the Pasteuria sp. NA strain were obtained as described before. Infected nematodes and cysts in the appropriate 1.5 ml siliconized microtube containing deionized water were crushed with a pestle, filtered with 18-1am-pore membrane, and the endospore concentration of the suspension was determined with a hemocytometer (Fisher, Suwanee, GA) under a compound microscope (Leica, Davie, FL) at a magnification of 40x. Ten microliters of endospore suspension was transferred to a 1.5 ml siliconized microtube, and 30 jil of the extraction solution containing 1.33x UDC (8 M urea, 0.04 M dithiothreitol,0.00665 M CHES buffer, pH 9.8) was added. Microtubes were placed into a water bath for 2 hours at 37 'C, with 20 seconds of mild sonication (Brankson Cleaning Equipment Company, Shelton, CN), every 15 minutes. Extracts were centrifuged at 10,000 x g for 5 minutes at









80

room temperature, and aliquots of the supernatant were collected and stored at -20 C until used. Protein estimation was performed by a micro-protein assay, according to the manufacturer's instructions (BioRad, Hercules, CA). Standard curves were generated using bovine serum albumin (BSA) (Sigma, St. Louis, MO), and colorimetric measurement was performed at 595 nm (Hewlett Packard 8451A Diode Array spectrophotometer, Palo Alto, CA). The UDC stock solution was made previously using only urea and CHES buffer, pH 9.8, divided in 0.5 ml aliquots, and stored at -20 C in 1.5 ml microtubes. Dithiothreitol was added to the microtubes just before the extraction of proteins.

Preparation of Infected Nematodes for TEM

All Pasteuria-infected nematodes were obtained as described above except for the NA Pasteuria which was obtained as follows: infested dry soil (50 g) was placed in a I 00x 15 ml petri dishes, and the soil water was adjusted to 100% field capacity to increase the rate of endospore attachment. The dish was left uncovered at room temperature (Brown et al., 1985). After 3 days 1,000 juveniles (2) of H. glycines race 3 were added, and the moisture level was adjusted to 50% of field capacity. Dishes were incubated for 7 days at room temperature (Oostendorp et al., 1990), and the J2 were extracted by the centrifugal-flotation method (Jenkins, 1964). J2 with endospore attached were handpicked, and placed in a 1. 5 ml microtube, washed three times with deionized water by centrifugation at 10,000 x g for 2 minutes, and stored at 4 'C until used.

A modified protocol was used to carry out the TEM part of this study (Aldrich et al., 1995; Chen et al., 1997a; Zeikus and Aldrich, 1975). Nematodes were hand-picked









81

into a 40 tl-drop of fixative (1% glutaraldehyde, 4% formaldehyde, 5% dimethyl sulfoxide in 0.1 M sodium cacodylate buffer, pH 7.2), and cut into 2 to 4 pieces with a surgical knife (Fisher Scientific No. 15) to aid penetration of the reagents. Nematodes were transferred into a 50 jil-drop of 2.5% agarose (Fisher) at 450C and then cooled in a refrigerator. After cutting the gel into square blocks, they were placed in 12x75 millimeter culture tubes (Fisher) containing 1.5 ml of the fixative, and incubated overnight at 4 'C. After rinsing the nematodes four times with 0.2 M cadodylate buffer, pH 7.2 on ice for 30 minutes each, they were dehydrated in a cold ethanol series: 12, 25, 38, 50, 65, 75, 85, 95, and two changes of 100% for 20 minutes each, except for 75%, which was kept overnight at 4 'C. Specimens were infiltrated with LR White resins (London Resins White, Electron Microscopy Science, Fort Washington, PA) series: 25% and 50% for 3 and 6 hours, respectively, 75% and two changes in 100% overnight each time). Blocks were placed in lml-gelatin capsule containing LR White resin, and allowed to polymerize for 4 days at 50 'C. Thin sections, 50-70 nm thick were cut with a diamond knife on a LKB 8800 Ultratome III microtome (Sweden). Sections were collected and mounted on Formvar-coated nickel grids (100 mesh) and processed for immunocytochemistry.

Immunocytochemistry

Nickel grids containing section were placed, face down, on 201il-drops of 1% nonfat dry milk in PBS, pH 7.2 (0.01 M sodium phosphate buffer, 0.15 M sodium chloride) on a piece of Parafilm (American National Can, Menasha, WI) for 15 minutes at room temperature, to block nonspecific antibody-binding sites (Aldrich et al., 1992; 1995;









82

Dykstra, 1993) with modifications. Grids were transferred to 20 [tl-drops of the first antibody, anti-P-20 IgM MAb at 1:10,000 or 1:40,000 dilution in PBS, pH 7.2, and incubated overnight in a closed petri dish inside of a moist chamber at 4 C. Grids were floated in 20 lil-drops of high salt tween buffer, pH 7.2 (0.1% Tween 20 in 0.02M TrisHC1, pH 7.2, 0.5 M NaCl), and in PBS, pH 7.2 twice in each buffer for 10 minutes each, before incubation with goat anti-mouse IgM conjugated to colloidal gold (1:30 dilution in PBS, pH 7.2, 12 nm gold) (Jackson Immuno Research, West Grove, PA) for 1 hour at room temperature. Grids were washed again in high salt tween buffer, and PBS, and were incubated for 10 minutes in Trumps buffer, pH 7.2 (McDowell, and Trump, 1976) at room temperature in order to stabilize the antigen-antibody complex. Sections were washed with deionized water, and stained with 0.5% uranyl acetate for 7 minutes, and aqueous lead citrate solution for 2.5 minutes. Controls were probed with non-immune ascites fluid and goat-anti mouse IgM conjugated to gold to ensure that the results were not due to non-specific binding. Sections were examined on a Zeiss EM-10 transmission electron microscope at 80kV. All reagents used in this study were ultra pure-TEM grade. SDS-PAGE Analysis

Proteins extracted from endospores of Pasteuria NA, S-1, C-I, P1-UFLA, P-20, ring nematode and spiral nematode isolates of Pasteuria were individually combined with equal volume of sample buffer (62.5 mM Tris-HCl, pH 6.8, 2% SDS w/v, 10% glycerol, 0.05% bromophenol blue w/v, 2% P3-mercaptoethanol) (BioRad), boiled for 5 minutes at 100 'C, and centrifuged for 5 minutes at 10,000 x g. Twenty microliters (600 ng of protein) of the supernatant was loaded into the wells of a Tris-glycine polyacrylamide 4%









83

stacking gel (pH 6.80) and 12% separating gel (pH 8.8) (BioRad). Electrophoresis was carried out at 100 V for 10 minutes, and then it was set for 200V until the bromophenol blue dye had migrated to the bottom of the gel. Proteins were transferred onto nitrocellulose membranes in blotting buffer (192 mM glycine, 25 mM Tris, 20% methanol) using a Mini Transfer-blot Cell (BioRad) at a constant voltage, 50 V for 2 hours. Protein bands were visualized either by Aurodye (Amersham, Piscataway, NJ) according to manufacturer's instructions or anti-P-20 IgM MAb (Chapter 2). Standard ladders for molecular mass were loaded in the same gels ( SeeBlue TM Prestained Standards, Novel Experimental Technology, San Diego, CA). Immunoblotting

Blots were first blocked overnight with 0.5% skimmed milk (w/v) in PBST (10 mM sodium phosphate buffer, pH 7.2, 150 mM NaC1, 0.2% [Iv/v] Tween 20 at 4 'C. Blots then were incubated with anti-P-20 IgM MAb diluted 1:2,000 in PBST, pH 7.2 for

1.5 hours on a rotatory shaker at room temperature, and washed with PBST, three times, 5 minutes each. Blots were incubated with goat-anti mouse IgM conjugated to alkaline phosphatase (Sigma) diluted 1:1,000 in PBST, pH 7.2 for 1.5 hours at room temperature, and washed as before with PBST, pH 7.2. After washing blots with substrate buffer (100 mM Tris-HC1, pH 9.5, 100 mM NaCI, 5 mM MgC12) three times, 5 minutes each at room temperature, blots were incubated with alkaline phosphatase substrate (0.1 mg/ml nitrotetrazolium blue, 0.05 mg/ml 5-bromo-4-chloro-3-indolyl phosphate) (Promega, Madison, WI) in substrate buffer on a rotatory shaker at room temperature until color development. Blots were washed with deionized water and dried at room temperature.









84

Results

Immunocytochemistry

Intense gold labeling was specifically associated with sporangium (s), exosporium

(ex), and parasporal fibers (pf) of P-20, P1 -UFLA, Rhabditis-infecting Pasteuria, S-1, LS-1, L-1, and C- 1 (Figs. 3.1-4). Labeling was not observed over the outer spore coat

(oc), inner spore coat (ic), cortex (c) (Figs. 3.1-4), and basal ring (br) (Figs. 3.1A, B) of the endospores of P-20 and P 1-UFLA, collected in USA and Brazil. No labeling was observed over any structure of Pasteuria sp. NA used as a control (Fig. 3.5). Gold particles were not observed on the germ tube (gt) of Pasteuria sp. NA, nor over the cuticle of the cyst nematode, H. glycines (Fig. 3.6), however the parasporal fibers (pt) were labeled heavily (Fig. 3.6).

SDS-PAGE and Immunoblotting Analysis

AuroDye staining showed that at least three bands of proteins (arrow head) are common among the Pasteuria sp. NA, S-1, C-1, P1-UFLA, ring nematode and spiral nematode isolate of Pasteuria and P-20, used as control (Fig 3.7A). Immunoblotting showed qualitative and quantitative differences among all the those isolates and species of Pasteuira (Fig 3.7B). All species and isolates share the same epitope because it was recognized by anti-P-20 IgM MAb (Lanes 2-8) (Fig.3.7B). Isolates P1-UFLA and P-20 showed similar bands of proteins with equal intensity (Lane 5 and 6) (Fig. 3.7B). Similarities in bands of proteins also were observed between spiral nematode isolate of Pasteuria and ring nematode isolate of Pasteuria (Lanes 7 and 8) (Fig. 3.7B). Also the same degree of similarity in the protein profiles was observed among the Pasteuria sp.









85

NA, P1-UFLA, and P-20 extracts (Lanes 2, 5,and 6) (Fig. 3.7B). The strongest bands were observed in proteins extracts from Pasteuria sp. NA, PI-UFLA, and P-20 (Lanes 2, 5, and 6) (Fig. 3.7B). Pasteuria sp. S-1 showed one band of protein (arrow) (Lane 3) (Fig. 3.7B) that is shared among all other strains (Lanes 2, 4, 5, 6, 7, and 8) whereas C-1 strain showed one band of protein (arrow) (Lane 4) (Fig. 3.7B) that is also observed from the protein extract of Pasteuria sp. NA, P-20, and ring nematode-infecting Pasteuria (Lanes 2, 6, and 8). The isolate C-1 showed one strong band of protein with molecular weight between 50 and 36 kDa (Lane 4), which appeared similar to a band of less intensity from the extract of the spiral nematode infecting-Pasteuria (Lane 7) and ring nematode-infecting Pasteuria extract (Lane 8) (Fig. 3.7B).



Discussion

The immunocytochemistry indicated that the adhesin-associated epitope as recognized by anti-P-20 IgM MAb is shared among P-20, P-1 UFLA, NA Pasteuria strain, Rhabditis sp.-infecting Pasteuria, Pasteuria sp. S-1 strain, C-1, LS-1, and L-1. The immuno-gold labeling patterns were similar for all the species and isolates examined. The broad distribution of the adhesin epitope over several structures of endospores of different species and isolates of Pasteuria may increase their capabilities to attach to their host due to cooperative interactions between the adhesin epitope with receptors on the cuticle of the nematode host, such as carbohydrate binding proteins (Bird et al., 1989; Davies and Danks, 1993; Persidis et al., 1991) and fibronectin-like residues (Mohan et al.,




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BIOCHEMICAL EVENTS DURING THE DEVELOPMENT OF Pasteuria penetrans
WITHIN THE PSEUDOCOELOM OF Meloidogyne arenaria
JANETE ANDRADE BRITO
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
2002

To my parents, Alaide Andrade de Brito and
Urbano Pinheiro de Brito, for their unconditional
love and support to follow my dreams.

ACKNOWLEDGMENTS
I am indebted to the Coordenado de AperfeÍ9oamento de Pessoal de Nivel
Superior (CAPES), Brasilia, DF, Brasil, for financial support.
I express my deep felt appreciation and thanks to Drs. Robin M. Giblin-Davis,
chairman, and James F. Preston, cochairman of my supervisory committee, for their
guidance, support, encouragement, suggestions, and friendship they gave me throughout
this study. Thanks also are expressed to the other members of my committee, Drs. Henry
C. Aldrich, Donald W. Dickson, and Grover C. Smart, Jr for their support, suggestions,
encouragement, and friendship.
My sincere thanks and appreciation go to Donna S. Williams and John D. Rice for
assistance in the laboratory, encouragement, and friendship.
Sincere thanks go to Mrs. Debbie Hall who guided me to follow the University of
Florida rules throughout my program, and made sure that I did not miss any deadlines. I
also thank Dr. Khuong B. Nguyen for friendship, kindness, and help with portions of my
program, and also to Onaur Ruano for his unconditional support, encouragement, and
friendship at the beginning of my career as nematologist at the Fundado Institute
Agronómico do Paraná, Londrina PR, Brazil.
Special thanks are extended to Drs. Waine Dixon, Paul Lehman, and Renato
Inserra for their support and friendship.
Thanks to my labmates Claudia Riegel, Fahiem K. El-Borai Kora, Billy W. Crow,
iii

Hye Rim (Helena) Han, Zhongxiao Chen, and Ramazan Cetintas. They gave me great
friendship and inspiration. Also thanks go to Drs. Hermes Peixoto Santos Filho, Maria de
Lourdes Mendes, Rui P. Leite, Alfredo O. A. de Carvalho, Luis G. E. Vieira, and Rui G.
Cameiro; Marinalva Pereira Santos, Solange Colavoupe, Lorain M. McDowell, Marisol
Dávila, Heather L. Smith, and Susana B. Carrasco for their friendship, support and sense
of humor.
Many thanks go to my dear Brazilian friends for their support and friendship.
Special thanks go to my husband, Don Dickson, and also to my parents, Alaide A.
de Brito and Urbano P. de Brito, my sister Maria Augusta, and to my brothers Urbano
Filho, Elisiario Neto, and Antonio Marival for their love, patience, and encouragement
throughout my life.
Thanks go to my nieces, Rayane, Suian, and Heleninha, for their love.
IV

TABLE OF CONTENTS
Page
ACKNOWLEDGMENTS iii
LIST OF ABBREVIATIONS viii
LIST OF TABLES x
LIST OF FIGURES xi
ABSTRACT xiv
CHAPTERS
1. INTRODUCTION 1
Host: Root-Knot Nematodes {Meloidogyne spp.) 1
Historical Background 1
Life Cycle 2
Symptoms 4
Distribution and Economic Importance 5
Management 6
Parasite: Pasteuria penetrans 15
Historical Background 15
The Genus Pasteuria 18
Members of Pasteuria 18
Systematic and Phylogeny of Pasteuria 21
Distribution 24
Biological Control Potential 24
The Effect of Other Microorganisms and Pesticides on Pasteuria 26
Life Cycle 28
Host Specificity 33
Cultivation 35
Interaction: Host-Parasite 35
The Role of Adhesin Proteins in the Relationship Host-Parasite 35
v

Objectives
36
2. SYNTHESIS AND IMMUNOLOCALIZATION OF AN ADHESIN-
ASSOCLATED EPITOPE IN Pasteuria penetrans 38
Introduction 38
Materials and Methods 40
Nematode Source 40
Pasteuria penetrans Source 41
Experimental Design 42
Extraction and Determination of Proteins 44
Monoclonal Antibody 45
Epitope Quantification by ELISA 45
SDS-PAGE Analysis 46
Immunoblotting 47
Immunofluorescence of Whole Endospores 47
Tissue Preparation for Sectioning 49
Immunogold Labeling 50
Results 51
Microscopic Examination 51
Epitope Quantification by ELISA 51
SDS-PAGE Analysis and Immunoblotting 53
Immunfluorescence 53
Immunogold Labeling 57
Discussion 68
3. DETECTION OF ADHESIN PROTEINS AND IMMUNOLOGICAL
DIFFERENTIATION OF Pasteuria spp. USING A
MONOCLONAL ANTIBODY 73
Introduction 73
Materials and Methods 75
Origin of Pasteuria Species and Isolates 75
Propagation of Bacterial Species and Isolates 76
Extraction and Determination of Proteins 79
Preparation of Infected Nematodes for TEM 80
Immunocytochemistry 81
SDS-PAGE Analysis 82
Immunoblotting 83
Results 84
Immunocytochemistry 84
SDS-PAGE and Immunoblotting Analysis 84
Discussion 85
vi

4. SYNTHESIS OF SMALL, ACID-SOLUBLE SPORE PROTEINS IN
Pasteuria penetrans 99
Introduction 99
Materials and Methods 101
Pasteuria penetrans Endospores Source 101
Bacillus subtilis Spore Source 101
Extraction and Determination of SASPs from P. penetrans and
B. subtilis 101
Conjugation of SASPs Peptide Carrier Proteins 102
Purification of the Conjugates 102
Immunization of Hens for Production of Polyclonal Antibodies 103
Determination of IgY Activities in Yolk Extracts 104
Extraction of IgY from Egg Yolk Extracts 104
Determination of Activities of Purified IgY 105
Concentration of Purified IgY using Centripep 105
Affinity of Anti-Peptide IgY for SASPs 105
Results 106
Purification of the Conjugates 106
Determination of IgY Activities in Yolk Extracts 106
Extraction of IgY from Egg Yolk Extracts Ill
Determination of Activities of Purified IgY Ill
Affinity of Anti-Peptide IgY for SASPs Ill
Discussion Ill
5. SUMMARY 117
APPENDIX A EXTRACTION OF SMALL, ACID SOLUBLE SPORE
PROTEINS FROM SPORES 122
APPENDIX B ISOLATION OF IgY ANTIBODY FROM CHICKEN
EGG YOLKS 123
LIST OF REFERENCES 125
vii
BIOGRAPHICAL SKETCH
148

LIST OF ABBREVIATIONS
ELISA
BSA
kDa
FITC
KLH
M
pi
Fg
mM
pm
mg
ml
ng
nm
PAGE
PBS
PBST
Enzyme linked immunosorbent assay
Bovine serum albumin
Kilodalton
Fluorescein isothiocyanate
Keyhole Limpet Hemocyanin
Molar
Microliter (s)
Microgram (s)
Millimolar (s)
Micrometer (s)
Milligram (s)
Millileter (s)
Nano gram (s)
Nanometer (s)
Polyacrylamide gel electrophoresis
Sodium phosphate buffer
Sodium phosphate buffer plus Tween

PBST-BSA
lOmM phosphate buffer pH 7.4,150 mM NaCl, 0.05 % Tween, 2%
bovine serum albumin
SDS
Sodium dodecyl sulfate
UDC
6.0 M urea, 0.03 M dithiothreitol, 0.005 M CHES buffer pH 9.8
1.33x UDC
8.0 M urea, 0.04 M dithiothreitol, 0.00665 M CHES buffer pH 9.8
WGA
Wheat-germ agglutinin
XIX

LIST OF TABLES
Table Page
1.1. Described genera of endospore-forming bacteria and their DNA
base composition 23
2.1. Percentage of different developmental stages of Pasteuria penetrans
in Meloidogyne arenaria race 1 52
3.1. Species and isolates of Pasteuria 77
x

LIST OF FIGURES
Figure Page
2.1. Adhesin-associated epitope and total nematode protein per infected
nematode as a function of the development of Pasteuria penetrans 54
2.2. Blots of sodium dodecyl sulfate-polyacrylamide gels of Meloidogyne
arenaria protein extracts after electrophoresis 55
2.3. Detection of Pasteuria penetrans adhesin-associated epitope as a function
of its development within the pseudocoelom of Melodogyne arenaria
racel 56
2.4. Differential interference contrast (DIC) and fluorescence microscopy
microphotographs of whole endospores of Pasteuria penetrans P-20 strain ... 58
2.5. Longitudinal section of uninfected second-stage Meloidogyne arenaria
(1-day-old) probed with anti-IgM Mab at 1:10,000 dilution 59
2.6. Immunocytochemical localization of an adhesin-associated epitope
during the development of Pasteuria penetrans 61
2.7. Labeling of sporogenous stages oí Pasteuria penetrans 63
2.8. Sporogenous stages of Pasteuria penetrans 65
2.9. Late sporogenous stage of Pasteuria penetrans 67
3.1. Transmission electron micrographs of Pasteuria endospore sections,
probed with anti-P-20 IgM Mab at 10,000 87
3.2. Gold labeling of endospores of different isolates and species of Pasteuria 89
3.3. Immunoelectron microscopy of endospores of Pasteuria spp 91
3.4. Labeling of endospores of two isolates of Pasteuria spp 93
xi

3.5. Thin section of Pasteuria sp. NA used as a control 94
3.6. Thin section of an endospore of Pasteuria sp. NA 95
3.7. Detection of an adhesin-associated epitope in different strains 96
4.1. Activities of antibodies in egg yolk extracts collected from hen 134-5,
34 days after injection of 100 pi KLH-peptide as immunogen
(80 pg per 100 pi) into the wing and 100 pi into the foot pad. A boost
injection was performed at 14 days, 75 pi was injected into the
wing and 75 pi into the footpad. Egg yolk extracts were used at 100 and
1,000 dilution in PBST, whereas the antigen (KLH-peptide) 107
4.2. Activities of antibodies in egg yolk extracts collected from hen 135-1,
34 days after injection of 100 pi KLH-peptide as immunogen
(80 pg per 100 pi) into the wing and 100 pi into the foot pad. A boost
injection was performed as in Fig. 4.1. Egg yolk extracts were used at
100 and 1,000 dilution in PBST, whereas the antigen (KLH-peptide) 108
4.3. Activities of antibodies in egg yolk extracts collected from hen 134-5,
34 days after injection of 100 pi KLH-peptide as immunogen
(80 pg per 100 pi) into the wing and 100 pi into the foot pad. A boost
injection was performed as in Fig. 4.1. Egg yolk extracts were used at
100 and 1,000 dilution in PBST, whereas the antigen (BSA-peptide) 109
4.4. Activities of antibodies in egg yolk extracts collected from hen 135-1,
34 days after injection of 100 pi KLH-peptide as immunogen
(80 pg per 100 pi) into the wing and 100 pi into the foot pad. A boost
injection was performed as in Fig. 4.1. Egg yolk extracts were used at
100 and 1,000 dilution in PBST, whereas the antigen (BSA-peptide) 110
4.5. Activities of anti-KLH-peptide IgY antibodies extracted from egg yolk
extracts laid by hen 134-5 at 20 to 28 days after injection with
KLH-peptide. Antibodies were used at 1,000 dilution in PBST, pH 7.6.
KLH-peptide and BSA-peptide 112
4.6. Activities of purified IgY antibodies (pool 2) from egg yolk extracts
laid by the hen 134-5. Antibodies were dilute to 100; 1,000; and 10,000
in PBST, pH 7.6 whereas the antigens, KLH- peptide and BSA-peptide,
were dilute to 10,000; 100,000; and 1000,000 113
4.7. Activities of anti-KLH-peptide IgY antibodies extracted from egg yolk
extracts (pool 2) laid by hen 134-5 Antibody was used at 100; 1,000 and
xii

10,000 dilution in PBST, pH 7.6. SASP-Bacillus subtilis at 100 and
1,000 dilution in coating buffer 114
4.8. Activities of ant-KLH-peptide IgY antibody extracted from egg yolk
extracts (pool 2) laid by the hen 134-5. Antibody was used at 100;
1,000 and 10,000 dilution in PBST, pH 7.6. SASP-Pasteuria penetrans
at 100 and 1,000 dilution in coating buffer 115

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
BIOCHEMICAL EVENTS DURING THE DEVELOPMENT OF Pasteuria penetrans
WITHIN THE PSEUDOCOELOM OF Meloidogyne arenaria
By
Janete Andrade Brito
May, 2002
Chairperson: Dr. Robin M. Giblin-Davis
Major Department: Entomology and Nematology
Pasteuria penetrans is a naturally-occurring bacterial parasite of root-knot
nematodes and a promising biocontrol agent. The endospores of this bacterium attach to
the cuticle of second-stage juveniles and complete their life cycle within the infected
female. Sequential steps required for the bacterium’s propagation include: attachment,
infection and germination, vegetative growth, sporulation and release. The hypothesis to
be tested in these studies considers that molecular entities present on the surface of
mature endospores, designated as spore adhesins, are synthesized at a certain time during
the growth and sporulation of P. penetrans, and these allow the bacteria to attach to the
nematode host. The objectives of this study were to: 1) determine the temporal
relationship between adhesin epitope formation and sporulation of P. penetrans-, 2)
determine adhesin epitope distribution during spore development and association with
xiv

nematode host and; 3) determine if the adhesin epitope is shared by different species of
Pasteuria with different host specificities. ELISA and immunoblotting showed that only
proteins extracted from P. penetrans-infected root-knot nematodes harvested 24 days
after inoculation and growth at 35 °C were recognized by the anti-P-20 IgM Mab that
recognizes an adhesin epitope. Labeling, which was first observed in stage III of
sporogenesis, identified the epitope distributed over the parasporal fibers, and over other
structures, such as sporangium and exosporium, as the bacteria proceeded with the
sporogenesis process. However, labeling was not observed on the basal rings, cortex,
inner spore coat, outer spore coat, or protoplasm. Immunofluorescence revealed that the
epitope does not occur uniformly on the surface of mature endospores.
Immunocytochemistry and immunoblot analysis showed that the adhesin epitope is
shared by other species of Pasteuria. The uniform distribution of the epitope over the
thin sections of mature endospores of strains and species of Pasteuria support a role for
the epitope in recognition of the nematode host as an early event in the attachment
process.
xv

CHAPTER 1
INTRODUCTION
Host: Root-knot nematodes (Meloidosvne spp.)
Historical Background
Berkeley (1855), in England, reported a plant disease in a greenhouse as “vibrios
forming excrescences on cucumber roots”. Muller (1884) described the nematode
pathogen of the disease as Heterodera radicicola. This pathogen, root-knot nematode,
was considered as a single large group for 65 years; nevertheless, it was reclassified
several times during that period as follows: Anguillula marioni Cornu, 1879, A. arenaria
Neal, 1889, A. vialae Lavergne, 1901, H. javanica Treub, 1885, Tylenchulus arenarais
Cobb, 1890, Meloidogyne exigua Goldi, 1887, Oxyurus incognita Kofoid and White,
1919, Caconema radicicola Cobb, 1924, and Heterodera marioni (Cornu, 1879)
Marcinowski, 1909 (Thome, 1961). The nematodes gained much attention. There was
obvious physiological and biological variability noted among different field populations
(Christie, 1946, Christie and Albin, 1944). This led to the classical work by Chitwood in
1949. He re-erected the genus Meloidogyne Goldi, 1887 to receive all root-knot
nematodes. He not only redescribed the type species, M. exigua (Góldi, 1887), but also
redescribed M. javanica (Treub, 1885), M. arenaria (Neal, 1889), and M incognita
(Kofoid &White, 1919), and described M. hapla, and a new variety, M. incognita var.
acrita (Hirschmann, 1985). A host-range study conducted by Sasser (1954) showed that
1

2
the host response to root-knot nematode infection was widely variable, not only among
species, but also within species of this nematodes. This was the first report calling for the
use of differential host plant bioassays to aid with the identification of Meloidogyne
species. Taylor and Sasser (1978) modified the original list of host differentials to
include the following six differential host plants: cotton (Gossypium hirsutum cv.
Deltapine 16), peanut (Arachis hypogea cv. Florunner), pepper (Capsicum annum cv.
California Wonder), strawberry (Fragaria x ananassa Duchesne), tobacco (Nicotiana
tabacum cv. NC 95), tomato (Lycopersicon esculentum cv. Rutgers), and watermelon
(Citrullus lanatus cv. Charleston Gray). Based on these differentials, four races of M.
incognita and two races of M. arenaria were identified (Taylor and Sasser, 1978). In
addition, two biological races in M. hapla, based on chromosome numbers, have been
reported (Triantaphyllou, 1966). Also, some physiological (Cameiro et al., 1990) and
genetic variability (Cameiro et al., 1998, Janati et al., 1982, Triantaphyllou, 1985) has
been reported within M. javanica and M. arenaria (Esbenshade and Triantaphyllou,
1985). Up to 1981, about 80 species of Meloidogyne had been described (Eisenback et
al., 1981).
Life Cycle
The life cycle of root-knot nematodes starts with the production of eggs. After the
embryogenesis process is completed the first-stage juvenile molts within the egg. The
second-stage juvenile (J2) hatches, and migrates freely in the soil. The J2 is the major
survival stage and only infective stage. It enters susceptible plant roots to continue its life
cycle. The J2 are attracted to plant roots. They migrate to a root of a susceptible plant 25

3
cm vertically in 10 days (Prot, 1978). The J2 generally penetrate roots directly behind the
root cap; however, penetration may occur at points where lateral roots emerge (Hussey,
1985). Cellulase, derived from esophageal gland cells, may play a role in the penetration
and migration in roots (Bird et al., 1975). The subventral gland cells are the most active
in J2 (Bird, 1967). Nonetheless, following the onset of parasitism, the dorsal gland cell
increases the production of secretory granules and becomes the predominate gland cell in
females (Bird, 1968; 1983; Hussey and Mims, 1991). After penetrating a root, the J2
migrates intercellularly in the cortex toward the region of cell differentiation. When its
head reaches the periphery of the vascular tissue, it establishes a feeding site (Hussey,
1985). Secretions injected through the stylet into the vascular tissue of the cells near the
head cause morphological and physiological changes in these cells, which enlarge and are
called giant cells (Hussey et al., 1998). The roots enlarge at those sites producing galls
(Loewenberg, et al., 1960). Five to six multinucleate giant cells develop. These are
highly specialized cells on which the nematode feeds (Hussey, 1985). After establishing
the feeding site, the J2 becomes sedentary and undergoes morphological changes
including increase in its body width but not its length (Taylor and Sasser, 1978). The
nematode molts three more times during development to form the third and fourth stage
juveniles and the adult stage (male and female). The males are vermiform whereas the
females are globose-pyriform in shape. The rate at which these nematodes develop is
influenced by several factors such as temperature, host suitability, and vigor of the host.
Tyler (1933) reported that at 27.5 °C to 30 °C females developed from J2 to the egg-
laying female in about 17 days, at 24.5 °C in 21 to 30 days, at 20 °C in 31 days, and at

4
15.4 °C in 75 days. Females reproduce mainly by parthenogenesis (Triantaphyllou,
1985). Some species are amphimitic or reproduce both by parthenogenesis or
amphimixis. Females lay eggs into a gelatinous matrix that forms an egg mass. The
number of eggs per egg mass is highly variable, but may range from almost 200 to 1,000
eggs. The egg masses are generally found outside the galled tissue, but in some host
plants the egg mass will lie within the galled tissue.
Symptoms
Plants infected with root-knot nematodes exhibit above-ground and below-ground
symptoms. The first below-ground symptoms are the formation of root galls and a poorly
developed root-system. The galls result from cell enlargement (hyperplasia), and an
increase in cell number (hypertrophy) surrounding the giant cells. Galls usually start to
develop in 1 to 2 days after root penetration by a J2. The gall size, which can be small
and discrete or large, and in some cases coalesced, is related to the number of nematodes
inside the plant tissue (Dropkin, 1954). The size of the galls varies among plant species
and nematode species. Generally, egg masses may be observed easily on a galled root,
but in some plant species the egg masses are covered by plant tissue. Galls caused by
root-knot nematodes can be diagnosed erroneously as nitrogen nodules. Nematode galls
are an integral part of root tissue and can not be detached without severely damaging the
roots, whereas nitrogen nodules, caused by Bradyrhiiobium spp., are round swellings that
appear to be attached to the root and are detached easily. Nodules may be infected by
root-knot nematodes, and galls and egg masses can be found on the nodules (Minton and
Baujard, 1990; Porter et al., 1984).

5
The above-ground symptoms usually depend on the initial nematode density in the
soil as well as environmental conditions (Minton and Baujard, 1990). Infected plants
have reduced uptake of nutrients and water, which produces yellowing, wilting, and
stunting of leaves (Nestscher and Sikora, 1990).
Distribution and Economical Importance
Meloidogyne spp. are among the most widespread and important plant pathogens
limiting crop productivity (Sasser and Carter, 1985). Root-knot nematodes can establish
in several soil types; however, suppression of crop yields caused by these nematodes are
more severe in sandy soils than in clay soils (Taylor and Sasser, 1978). Heavily infected
plants may die when there is severe stress caused by hot, dry conditions. Yield losses
caused by plant-parasitic nematodes are approximately $8 billion a year to producers in
the United States and nearly $78 billion worldwide (Society of Nematologists,
Committee on National Needs and Priorities in Nematology, 1994). However, the
damage caused by root-knot nematodes alone is very difficult to determine, and
sometimes it is overlooked or underestimated because of the interaction with soilbome
fungi, bacteria, viruses, insects, and other nematodes (Nestscher and Sikora, 1990).
Meloidogyne spp. cause damage and are associated with many plants, including
economic crops and weeds in all areas of the world (Taylor and Sasser, 1978), but they
are considered to be most important in tropical regions (Johnson and Fassuliotis, 1984;
Mai 1985). This is mainly due to i) high temperatures and a longer growing season that
favors more generations of the nematode per year, resulting in higher nematode densities
in the soil; ii) the presence of highly virulent species, such as M. incognita, M. arenaria,

6
and M. javanica, which are well-adapted to warmer areas, and iii) prevalence of more
disease complexes involving root-knot nematodes and soilbome fungi (Mai, 1985).
Meloidogyme incognita has the widest geographic distribution of all species described,
followed closely by M. javanica, and M. arenaria. Those species are very common in
tropical regions, whereas M. hapla is more common in temperate regions of the world
(Taylor and Sasser, 1978). The optimum monthly temperature for development of M.
incognita is 27 °C; nonetheless it can be found in areas that have an average temperature
of 24-30 °C (Eisenback and Triantaphyllou, 1991). In contrast, M. hapla can survive in
frozen soil and it can reproduce at temperatures as low as 15 °C (Taylor and Sasser,
1978).
Management
Chemical nematicides. In the 1940s, discovery of the nematicidal properties of
l,2-dichloropropane,l,3-dichloropropene (DD) made it possible to demonstrate to
producers the damage caused by root-knot nematodes. It marked the beginning of the soil
fumigation industry (Johnson and Feldmesser, 1987). After World War II, ethylene
dibromide (EDB), l,2-dibromo-3-chloropropane (DBCP), and bromomethane (methyl
bromide, MBr) were formulated as soil fumigants. Each was offered at prices economical
for use in the production of moderate to high-value crops (Johnson and Feldmesser,
1987). Later DD, EDB and DBCP were found in ground water, and were withdrawn
from the market (Heald, 1987).
Since 1960, different methyl bromide formulations have been used for high-value
crops. Methyl bromide has became one of the most popular fumigants because of its

7
broad-spectrum activity and its relatively low cost (Noling and Becker, 1994). It is not
only highly efficient in the control of nematodes, but also provides control of fungi,
bacteria, insects, rodents, and weeds (Thomas, 1996). Methyl bromide has been used as
an agricultural soil fumigant, structural and commodity fumigant, and for quarantine and
regulatory purposes (USDA, 1993a; 1993b; Watson, et al., 1992). About 79,000 tons
have been used annually on a global basis by agricultural users, mainly as a soil fumigant
(75%), but also as a post-harvest fumigant (22%) and for structural (3%) pest control
(UNEP, 1995). Worldwide more than half of the production of methyl bromide is used
on four crops: tomato, tobacco, strawberries, and melons (Ferguson and Padula, 1994;
Stephens, 1996a; 1996b).
In Florida and in other states, methyl bromide is used mainly under plastic mulch
as a preplant soil fumigant in the production of tomato, pepper, strawberry, other fruits,
turfgrass, and nursery crops; however, most methyl bromide is consumed in the tomato,
pepper, and strawberry industries (Ferguson and Padula, 1994; Johnson et al., 1962;
McSorley et al., 1986; Overman and Jones, 1984).
The emission of methyl bromide into the atmosphere became a major
environmental concern in the late 1980s. The Montreal Protocol Treaty, an international
agreement signed by more than 150 countries, governs the world-wide production and
trade of ozone-depleting substances. In 1992, the signatories of the Montreal Protocol
identified methyl bromide as an ozone depleter (Watson et al., 1992). In 1993, the
Montreal Protocol treaty was amended to require that developed countries freeze the
production of methyl bromide at 1991 levels by 1995 (USEPA, 1993), and at the 1995

8
meeting, a global methyl bromide production phase-out was approved (Thomas, 1996).
Industrial nations were to have a 25% reduction by 2001, a 50% reduction by 2005, and a
complete phase-out in 2010, whereas developing nation should freeze the production of
methyl bromide in 2002 based upon an average of the years 1995-98 (UNEP, 1995).
In the last several years, studies have been carried out to develop alternative
biocides and to implement new strategies for methyl bromide replacement. Materials that
have been identified to have broad spectrum activity in soils include 1,3-dichloropropene
(1,3-D) products (Riegel, 2001), dazomet, trichloronitromethane (chloropicrin),
dithiocarbamate (metham sodium), sodium tetrathiocarbonate, formalin or formaldehyde,
and nonfumigants nematicide-insecticides (Anonymous, 1995). However, none of the
materials provide the same level of broad spectrum activity as that provided by methyl
bromide. Chloropicrin alone is very efficient for the control of many soilbome fungi, but
it does not control plant-parasitic nematodes efficiently. 1,3-D provides control of cyst,
root-knot, stubby root, lesion, ring, and dagger nematodes, but it is not effective against
fungi (Locascio et al., 1997, Stephens, 1996b). 1,3-D can be mixed with chloropicrin to
enhance activity against soilbome fungi. Such products are registered for more than 120
vegetable, field, and nursery crops in the United States (Melicher, 1994).
Crop rotation. Nonchemical alternatives for suppressing nematode populations
include the use of crop rotation, resistant varieties, cover crops, soil amendments,
flooding, solarization, bare fallowing, and biological control (Christie, 1959; Netscher
and Sikora, 1990; Mai, 1985). Some of those techniques have been used for many years,
and can be effective against some plant-parasitic nematodes under specific situations, but

9
they do not provide the same broad spectrum of control as methyl bromide.
Crop rotation is one of the oldest ways to manage Meloidogyne spp. However,
due to their broad host range, choosing the appropriate crop can be difficult (Potter and
Olthof, 1983), and in many cases the best crop choice to manage the nematode densities
in the soil is not a suitable choice for the growers. The principle of this method is based
on the use of resistant, susceptible, or tolerant crops for the predominant species of root-
knot nematode for a specific area (Johnson, 1982). Currently, crop rotation remains an
option to reduce the damage caused by root-knot nematodes in the southeastern United
States (Johnson, 1982). Rodríguez-Kábana et al. (1988, 1989) showed that castor
(Ricinus communis L.), American jointvetch (Aeschynomene americana L.), partridge pea
(Cassia fasiculata Michx.), and sesame (Sesamum indicum L.) did not support M.
arenaria populations in the field. McSorley et al. (1994) studied the effects of 12
summer crops on M. arenaria race 1 and on the yield of vegetables in microplots.
Castor, cotton (Gossypium hirsutum L.), velvetbean (Mucuna deeringiana [Bort.] Merr.),
crotalaria (Crotalaria spectablis Roth.), and hairy indigo (Indigofera hirsuta L.) reduced
nematode numbers. Yields of vegetable crops were higher following castor than other
summer crops, and yields of vegetable crops following castor as a cover crop were
approximately double the yields of the same vegetable crop following peanut, a host of
M. arenaria race 1.
Resistance. Nematode-resistant cultivars can be an option to manage root-knot
nematodes, and they might be used alone or in crop rotation schemes as part of an
integrated root-knot nematode control program. Attempts have been carried out to

10
develop cultivars resistant to one or more species of root-knot nematodes. Currently,
there are nematode-resistant cultivars of tomato, southern pea, pepper, bean, and sweet
potato (Noling and Becker, 1994). However, due to the occurrence of genetic variability
within species of root-knot nematodes, it is difficult to develop a cultivar that is resistant
to more than one race. In addition, the occurrence of mixtures of races and species of
root-knot nematodes within a given area, as well as resistance being broken at high soil
temperatures, often limits their usefulness. Even though the tomato resistant gene “Mi”
typically confers resistance to M. javanica, M. incognita, and M. arenaria, virulent
populations of these nematodes have completely overcome the Mi gene resistance
(Castagnone-Sereno 1999; Xu et al., 2001). A greater problem to overcome is the loss of
host resistance in tomato that occurs when soil temperatures heat up to over 28 °C
(Abdul-Baki et al., 1996; Tzortzakakis, 1997). A loss of resistance to M. incognita in
Phaseolus vulgaris was observed at 24 °C and above (Mullin et al., 1991).
Integrated pest management. The integration of different tactics have been
implemented in attempts to manage plant-parasitic nematodes. In the southern United
States, M. incognita is a major pathogen of sweet potato (Hall et al.; 1988). A
combination of crop rotation, resistant cultivars, nonhost, and nematicides seems to be the
most economical method of nematode control on sweet potato (Jatala and Bridge, 1990).
Meloidogyne arenaria race 1 is one of the most serious pathogens of peanut in the
southern United States. For many years peanut growers have relied on crop rotation,
winter cover crops, post harvest crop destruction, and nematicides for managing root-knot
nematodes (Dickson, 1998). Recently, the peanut germplasm has been released from

11
Texas A&M University that is resistant to race 1 of M. arenaria (Simpson and Starr,
1999). With the development of suitable cultivars incorporating this resistance will
greatly improve nematode management for peanut producers.
Biological control agents. Root-knot nematodes, their antagonists and parasites,
share the same soil habitat. Interactions of these organisms are affected by a number of
factors such as the physical and chemical environment of the soil as well as the soil
microflora which might play a role in the use of antagonists and parasites in root-knot
nematode management (Stirling, 1991). Although several organisms such as fungi,
bacteria, viruses, nematodes, mites, insects, protozoans, turbellarians, oligochaetes, and
tardigrades have been shown to have some affect on nematode population densities under
laboratory and greenhouse conditions, field results have been contradictory (Jairajpuri et
al., 1990; Stirling, 1991). Particular attention has been given to effects of soil-inhabiting
fungi on the population densities and activities of plant parasitic-nematodes. The known
fungal antagonists (Gray, 1988) of nematodes are grouped as i) endoparasites of
vermiform nematodes; ii) nematode-trapping fungi, and iii) female and egg parasites and
cyst colonizers.
Endoparasitic fungi are classified into three categories based on their mechanism
of infection and their taxonomic position: i) group I, encysting species of
Chytridiomycetes and Oomycetes such as Catenaria anguillulae, Lagenidium caudatum,
Aphanomyces sp. and Leptolegnia sp. which have a flagellated zoospore as their infective
propagule; ii) group II, Deuteromycetes producing adhesive conidia and conidia which
are ingested; and iii) group III, Basidiomycetes producing adhesive conidia. Fungi of

12
groups II and III initiate the infection process when the conidia either adhere to the
nematode’s cuticle (Drechmeria coniospora, Hirsutella rhossiliensis, Macrobiophthora
vermicola, Myzocytium humicola, Nematoctonus leiosporus, N. concurrens, N.
haptocladus, and Verticillium balanoides), or when conidia lodge in the buccal cavity or
the gut of the host (all species of Harposporium but one) (Stirling, 1991). This latter
group would not likely be efficient for biocontrol of plant-parasitic nematodes because
they would be unable to ingest the conidia (Stirling, 1991).
Nematode-trapping fungi or predatory fungi have sparse mycelia that have been
modified to form organs capable of capturing nematodes. They are the best known
nematophagous fungi, and they have been studied for over a century (Stirling, 1991).
There are six mechanisms by which these types of fungi can capture a nematode: i)
Adhesive hyphae, produced by Zygomycetes (Stylapage and Cystopage) and a few
species of Hyphomycetes. A yellowish adhesive secretion is produced by the fungi.
These are considered to be the least sophisticated trapping mechanisms, ii) Adhesive
#
branches produced by a few species of fungi, such as Monacrosporium cionopagum.
Erect branches of one or two cells produced on the hyphae may anastomose to form loops
or two dimensional networks, which may trap nematodes as they move around, iii)
Adhesive mycelial network, the most common type of trap, found in almost all soil types.
It forms from the lateral branch growing from the vegetative hypha and curving to fuse
with the parent hypha. More loops are produced on this loop or on the parent hypha, until
a complex, three-dimensional, adhesive-covered network of anastomosed loops is
produced (Arthrobotrys oligospora). iv) Adhesive knobs, formed of distinct adhesive-

13
globose cells that are either sessile on the hypha or borne aloft on a short, erect stalk.
These cells occur along the hypha, so that nematodes are often restrained by several
knobs. Nematodes may struggle to escape the knobs, which may cause the knobs to
detach from their stalks in some species but the knobs remain firmly attached to the
nematode and germination occurs quickly. This type of trap mechanism is most common
among Hyphomycetes, but it is found also in the Basidiomycetes. Nematoctonus
produces non-detachable, hourglass-shaped knobs that are engulfed in a larger, spherical
ball of viscous substance (Barron, 1997). v) Non-constricting rings, the most frequent
device in nematophagous fungi. Three-celled rings are formed when erect lateral
branches from vegetative hyphae thicken and curve, which then fuse to the support stalks.
Nematodes are captured when rings become wedged around their bodies, vi) Constricting
rings, similar to non-constricting rings. The rings are attached to hypha by short stalks.
Nematodes entering these rings trigger them to swell rapidly inward, thereby capturing
the nematode. The ring closes in about 0.1 second once initiated; however there is a lag
period of 2 to 3 seconds from the time the nematodes first touch the ring cells until it
closes. The nematodes can escape during this short period, which makes this type of
mechanism an inefficient trap (Stirling 1991).
Female and egg parasites, and cyst colonizers, are a taxonomically and
ecologically diverse group, ranging from host specific zoosporic fungi to opportunistic
species that live largely as soil saprophytes. Over the years many fungi have been
isolated from females, cysts, eggs, and egg masses of plant-parasitic nematodes, but the

14
majority have proved to be saprophytes rather than parasites (Chen et al., 1996; Morgan-
Jones and Rodríguez-Kábana, 1988; Stirling, 1988).
Rodríguez-Kábana and Morgan-Jones (1988 ) listed 12 genera of fungi that are
isolated frequently from females, eggs, and cysts of Heteroderidae in Australia, Europe,
and North and South America: Acremonium, Alternaría, Catenaria, Cylindrocarpon,
Exophiala, Fusarium, Gliocladium, Nemathophora, Paecilomyces, Penicillium, Phoma,
and Verticillium. Among these V. chlamydosporium has been the most widely studied
(Stirling, 1988), and proven pathogenic to Meloidogyne, Globodera, and Heterodera.
The fungus Paecilomyces lilacinus was found parasitizing eggs of Meloidogyne incognita
(Jatala et al., 1979) in Peru. After its discovery, it became the principal organism of
interest (Dube and Smart, 1987; Jatala et al., 1979; 1980; 1981). Although it has been
found in many geographical areas (Gintis et al., 1983; Godoy et al., 1983; Morgan-Jones
et al., 1984; Dackman and Nordbring-Hertz, 1985) it is more common in warmer areas of
the world (Domsch et al., 1980). Paecilomyces lilacinus has been shown to be a
biocontrol agent of several species of nematodes (Jatala 1985; 1986). However, there are
mixed reports on the efficacy of this fungus (Hewlett et al., 1988; Rodríguez-Kábana et
al., 1984).
The bacterium, Pasteuria penetrans (Chen et al., 1997b; Eddaoudi and Bourijate,
1998; Freitas, 1997; Trudgill et al., 2000, Tzortzakis and Gowen, 1994; Spiegel et al.,
1996), has become the most studied biocontrol agent in the last several years, and is
reported to be one of the most promising biological control agents of root-knot nematodes
(Chen et al., 1996; Duponnois et al., 1999; Oostendorp et al., 1991; Zaki and Maqbool,

15
1992). Once the problem with its cultivation and mass-production is overcome it may be
a very useful biological agent in an integrated root-knot nematode management program.
Parasite: Pasteuria penetrans
Historical Background
The history of Pasteuria spp. has a rather unusual start in that the organism was
first reported as a parasite of the water flea Daphnia magna Strauss. This discovery was
made in 1887 by Elie Metchnikoff, soon after he accepted a research position offered by
Louis Pasteur at the newly formed Pasteur Institute, Paris (Sayre, 1993). In 1888
Metchnikoff erected a new genus, Pasteuria, which he named in honor of Louis Pasteur,
to contain the new species, P. ramosa. He emphasized the unique mode of division of
this bacterium when he wrote, “Pasteuria sp. was able to undergo as many as five
longitudinal divisions at the same time, giving it a characteristic fan shape” (Sayre, 1993
plOl). All attempts made by Metchnikoff to culture the bacterium failed, and thus the
type strain was not established (Sayre, 1993).
For many years the description of Pasteuria ramosa enticed researchers around
the world to seek the bacterial parasite of water fleas (Henrici and Johnson, 1935; Hirsch
, 1972; Staley, 1973). A budding bacterial species of the Blastobacter group, found
occasionally on the exterior surfaces of Daphnia sp., was classified erroneously as
Metchnikoff s unique bacterium, even though it did not form either endospores,
mycelium or branches, was not a parasite of cladocerans, and showed a Gram-negative

16
reaction. This budding bacterium (strain ATCC 27377) was cultivated in vitro, and then
assigned erroneously as the type species of the genus Pasteuria (Staley, 1973).
Eighty-nine years after Metchnikoff discovered P. ramosa, it was rediscovered
infecting Moina rectirostris, a member of the family Daphnidae (Sayre, 1977). The
similarity between the newly discovered bacterial strain and Metchnikoff s bacterium was
very clear despite the lack of evidence of longitudinal division. Primary colonies
branched and formed a cauliflower-like shape. Daughter colonies were formed by the
fragmentation of mother colonies. Quartets, doublets, and single sporangia were
produced from the daughter colonies. A sporangium consisted of a conical stem, swollen
middle cell, and an endogenous endospore (Sayre et al., 1979; 1983).
Ten years after Pasteuria had been assigned erroneously as strain ATCC 27377,
that strain was reclassified as Plactomyces staleyi Starr, Sayre, and Schmidt, 1983 (Starr
et al., 1983). Starr et al. (1983) requested that the original description of P. ramosa
Metchnikoff, 1888 be conserved and that ATCC 27377 be rejected as the type strain of P.
ramosa. Later that request was supported by the Judicial Commission for the Code of
Nomenclature of Bacteria (Judicial Commission, 1986), and further studies supported
that decision (Sayre et al., 1988; 1989).
Cobb (1906) was the first to report an organism resembling Pasteuria sp.
(numerous highly refractile spores) as a parasite of a nematode, Dorylaimus bulbiferous.
He erroneously classified the parasite as a sporozoan. Later Micoletzky (1925) found an
organism whose shape and spore size were similar to those reported in 1906 by Cobb.
Micoletzky suggested that those spores belonged to the genus of a sporozoan, Duboscqia

17
Perez. Thome (1940) described in detail an organism parasitizing Pratylenchus pratensis
(de Man) Filipjev (later identified as P. brachyurus by Thome), and on the assumption
that the organism was similar to the parasite described by Micoletzky, assigned it to the
genus Duboscqia as D. penetrans. However, over the years the taxonomic position of the
nematode parasite has been questioned (Canning, 1973; Williams, 1960). The
misplacement of the organism, now known to be a bacterial parasite of nematodes as a
protozoan, persisted for almost 70 years. Mankau (1975a) reexamined the nematode
parasite using electron microscopy and showed for the first time that it is a bacterium
rather than a protozoan; he reassigned it to the genus Bacillus as B. penetrans (Thome,
1940, Mankau, 1975). Nonetheless, neither flagella nor active motility were observed in
Bacillus penetrans (Sayre and Starr, 1985). Soon more studies on the procaryotic
affinities (Mankau, 1975b), biology (Mankau and Imbriani, 1975), ultrastructure
(Imbriani and Mankau, 1977), and host (Mankau and Prasad, 1977) of B. penetrans were
carried out. B. penetrans was never included in the “Approved Lists of Bacterial Names”
(Skerman et al., 1980), thus the confusion on the classification of the bacterial nematode
parasite continued.
Sayre and Wergin (1977) observed the similarity between the developmental
stages of a bacterial parasite of Meloidogyne incognita with the original descriptions and
drawings of the life cycle of P. ramosa. Later morphological and taxonomic
reevaluations of P. ramosa and B. penetrans were provided (Sayre et ah, 1983). Finally
Sayre and Starr (1985) placed the bacterial parasite of nematodes in the genus Pasteuria,

18
as P. penetrans, due to its similarity with Pasteuria rather than Bacillus, and presented an
emended description of the genus Pasteuria Metchnikoff.
The Genus Pasteuria
Species of Pasteuria are Gram-positive, endospore-forming bacteria. The genetic
and biochemical aspects of the formation of the virulent endospores of Pasteuria spp. are
not well understood, but the morphological aspects are (Chen et al., 1977a; Giblin-Davis
et al., 1995; Sayre and Starr, 1985; Sayre 1993). These bacteria form a dichotomously
branched septate mycelium. The terminal hyphae of a mycelium elongates, and then
segments to form the sporangia, and eventually endospores. (Sayre and Starr, 1985).
Mother colonies, which resemble a cauliflower or elongate grapes in clusters, fragment
to form daughter colonies. Daughter colonies form quartets, doublets, and finally a single
sporangia which enclose a single endospore (Chen et al., 1997a; Sayre and Starr, 1985).
Endospores are nonmotile and resistant to desiccation and elevated temperatures (Dutky
and Sayre, 1978; Stirling, 1985; Williams et al., 1989). Endospores of P. penetrans are
cup-shaped and measure, on average 3.4 pm ± 0.2 by 2.5 pm ± 0.2 using transmission
electron microscopy (Sayre 1993).
Members of Pasteuria
There is still considerable confusion about the taxonomy of Pasteuria. Over the
years the criteria used to assign species to the genus have been host specificity,
developmental characteristics, and size and shape of sporangia and endospores (Sayre and
Starr, 1989). However, host specificity overlaps in several cases. Although sizes of

19
endospores and sporangia are considered to be host specific (Ciancio, 1995), endospore
diameters of P. penetrans vary from 3.6 to 7.0 /am ( Sayre and Starr, 1985).
Cross-genera hosts have been reported. For example, one isolate of P. penetrans
reported from the United States (Mankau, 1975a; Oostendorp et al., 1990), Puerto Rico
(Vargas and Acosta, 1990) and China (Pan et al., 1993) parasitizes both Meloidogyne and
Pratylenchus spp. An isolate of Pasteuria sp. from India parasitizes Heterodera sp. and
M. incognita (Bhattacharya and Swarup, 1988), whereas another strain reported from
India parasitizes Heterodera spp., and Rotylenchulus reniformis (Sharma and Davies,
1996). Davies et al. (1990) reported that endospores of a Pasteuria sp. extracted from H.
avenae, cereal-cyst nematode, attached to the cuticle of H. shachtii, H. glycines,
Globodera rostochiensis, G. pallida, and M. javanica. On the other hand, Pasteuria sp.
S-l showed a high a level of host specificity. S-l strain attached to B. longicaudatus but
did not attach to any of the other nematodes, including J2 of M. arenaria, M. incognita,
M. javanica, H. galeatus, and Pratylenchus penetrans (Giblin-Daves et al., 1995). These
results were confirmed by Bekal et al. (2001). They showed that S-l did not attached to
H. schachtii, Longidorus africanus, M. hapla, M. incognita, M. javanica, P. brachyurus,
P. scribneri, P. neglectus. P. penetrans, P. thornei, P. vulnus, or Xiphinema spp.
Some isolates of Pasteuria have been reported to attach to and develop in
different life stages of the nematode host (Abrantes and Vovlas, 1988; Davies et al.,
1990). Mature endospores of P. penetrans were observed in the peseudocoelom of J2 and
males oí Meloidogyne sp. and J2 of H.fici (Abrantes and Vovlas, 1988). Davies et al.

20
(1990) found that a Pasteuria sp. isolated from the cereal-cyst nematode, H. avenae
Wollenweber, completed its life cycle in the J2 but not in females and cysts.
Different genera of nematodes have been reported to be parasitized by Pasteuria
spp. at the same site and in the same growing season. Giblin-Davis, during a survey in
South Florida, found that B. longicaudatus, Meloidogyne spp. and Helicotylenchus
microlobus were parasitized by Pasteuria spp. in Collier County; B. longicaudatus,
Hoplolaimus galeatus, Tylenchorhynchus annulatus, and Meloidogyne spp. in Broward
County; and H. microlobus and Meloidogyne spp. in Palm Beach County.
Currently four species of Pasteuria have been described so far: i) P. ramosa, a
parasite of the cladocerans (water fleas) Daphnia pulex Leyding and D. magna Strauss
(Sayre et al., 1977); ii) P. penetrans, a parasite of root-knot nematodes (Sayre and Starr,
1985), iii) Pasteuria thornei, isolated from Pratylenchus spp. (Starr and Sayre, 1988), and
iv) Pasteuria nishizawae (Sayre et ah, 1991), a parasite of cyst nematodes (Heterodera
and Globodera).
Recently, at least three new species of Pasteuria have been proposed, Pasteuria
sp. designated as S-l (Bekal et ah, 2001) from Belonolaimus longicaudatus Rau; North
American Pasteuria {Heterodera glycines-infecting Pasteuria) from the soybean cyst
nematode, Heterodera glycines Ichinohe, in Urbana, IL, USA (Atibalentja et ah, 2000)
and one strain from the pea cyst nematode, Heterodera goettingiana Liebscher in
Münster, Germany (Sturhan et ah, 1994).
Over the years unique isolates of Pasteuria have been reported. For example, a
large- and a small-spored isolate of Pasteuria spp., each from Hoplolaimus galeatus

21
(Cobb) Thome (Giblin-Davis et al., 1990), and another isolate from Rhabditis sp. (Giblin-
Davis pers. comm.) were collected from a bermudagrass turf in Fort. Lauderdale, FI.
Two isolates of Pasteuria sp.infecting different ring nematode species have been found:
C-l (Han et al., 1999), and ring nematode Pasteuria (Dickson, pers. comm.). A
Helicotylenchus sp.-infecting Pasteuria was isolated from bermudagrass turf shipped
from CA (Crow, pers. comm.). Also, three other isolates of Pasteuria that attach and
complete their life-cycle in Heterodera spp. have been reported: one isolate from H.
avenae (Davies et al., 1990); another (HCP) from Heterodera cajani Koshy, the pigeon
pea cyst nematode (Walia et al., 1990); and another, HMP, from Heterodera mothi, Khan
& Husain (Bajaj et al., 1997).
It is clear that there is a need to use other criteria, in addition to those already
used, to determine species of Pasteuria. The 16S rDNA has been used to determine more
precisely the taxonomic position of Pasteuria (Anderson et al., 1999; Atibalentja et al.,
2000; Bekal, 2001; Ebert et al., 1996). Once the conditions necessary to mass produce
Pasteuria in vitro are known, it will be possible to establish species through genetic and
biochemical studies.
Svstematics and Phvlogenv of Pasteuria
In 1992 13 genera of endospore-forming bacteria were known (Table 1.1). The
basis for separating them was morphology, physiology, and genetic diversity (Berkelwy
and Ali, 1994). Currently, bacteria are differentiated based on the generally accepted rule
that bacteria with DNA base compositions differing by more than 10 mol %GC (G+C)
should not be considered as members of the same genus. Strains differing by more than

22
5%GC values should not be regarded as the same species (Bull et al., 1992). The genera
Bacillus, Clostridium, and Desulfotomaculum are very heterogenous (Table 1.1). The
genera Oscillospira and Pasteuria (four species) have not yet been grown successfully in
pure culture. The description of Oscillospira species, O. guillermondii (Berkely and Ali,
1994), was based on morphological characters, whereas the species of Pasteuria were
described based on morphological characters, morphometries, ultrastructure, and host
specificity. Otherwise their DNA base composition are unknown.
In the summer of 1992 and throughout 1993 and 1994, P. ramosa was re¬
discovered parasitizing D. magna collected from several ponds in London, UK (Stimadel
and Ebert, 1997). Ebert et al. (1995) used these spores of P. ramosa collected from D.
magna, D. pulex, and D. longispina in the previous three summers from England as well
as Russia to establish the culture of P. ramosa by co-cultivation in D. magna. These
authors ended the uncertainty about the phylogenetic position of Pasteuria Metchnikoff
by sequencing the 16S rDNA of the bacterium. They provided strong evidence that P.
ramosa belongs to the low G+C Gram-positive endospore-forming bacteria and resides
within a clade containing B. tusciae, Alicyclobacillus cycloheptanicus, and A.
acidocaldarius, as the closest neighbors. They rejected the placement of P. ramosa in the
Actinomycetales. Anderson et al., (1999) provided the first 16S rDNA gene sequence
analysis of P. penetrans, and showed that it is correctly placed in the genus Pasteuria.
The authors found that P. ramosa is the closest neighbor of P. penetrans, and it is within
a clade that includes A. acidocaldarius, A. cycloheptanicus, Sulfobacillus sp., B. tusciae,

23
Table. 1.1. Described genera of endospore-forming bacteria and their DNA base
composition.
Genus
Mol% GC
Alicyclobacillus
52-60
Amphibacillus
36-38
Bacillus
32-69
Clostridium
22-54
Desulfotomaculum
38-52
Oscillospira
—
Pasteuria
—
Sporohalobacter
30-32
Sporolactobacillus
38-40
Sporosarcina
40-42
Sulfobacillus
54
Syntrophspora
38
Thermoactinomyces
52-55
Source: Berkeley and Ali, 1994.

24
B. schlegelii, and P. ramosa. Also Atibalentja et al. (2000), using a sequence of the 16S
rDNA, showed that a Heterodera.glycines-infecting Pastearía (Pasteuria sp. NA) and P.
ramosa form a distinct line of descent within the Alicyclobacillus group of the
Bacillaceae.
Distribution
Pasteuria spp. have been reported in 51 countries and in various islands in the
Atlantic, Pacific, and Indian oceans associated with 205 nematodes species belonging to
96 genera (Sayre and Starr, 1988; Sturhan, 1985). An updated host record list is reported
by Chen and Dickson (1998).
Biological Control Potential
There are certain attributes that make P. penetrans a desirable biological control
agent: 1) endospores are resistant to desiccation, high temperature, and most nematicides
(Dutky and Sayre, Freitas 1997; 1978; Stirling, 1985; Williams et al., 1989); 2)
encumbered nematode juveniles have reduced activity and ability to infect roots (Sturhan,
1985); and 3) infected juveniles complete their life cycle, but females have low or no
fecundity (Bird, 1986; Bird and Brisbane, 1988).
Pasteuria penetrans has been shown to control root-knot nematodes in
greenhouse tests (Brown and Smart, 1985; De Leij et al., 1992; Stirling 1984) and in field
microplots (Brown et al., 1985, Chen et al., 1997b; Dube and Smart, 1987; Oostendorp et
al., 1991; Stirling, 1984; Tzortzakakis and Gowen, 1994; Trudgill et al., 2000).
Suppression of root-knot nematodes by P. penetrans has been observed in vineyards more
than 10 years old in Australia (Stirling and White, 1982) as well as India (Mani et al.,

25
1999), and also in peanut and tobacco fields infected by root-knot nematodes in Florida
(Dickson, pers. comm.). Also, suppression of B. longicaudatus by Pasteuria sp.
S-l in a bermudagrass turf field in Florida has been reported (Giblin-Davis et al., 1995;
2000).
Studies have been carried out to determine the optimum endospore densities to
suppress root-knot nematodes (Chen et ah, 1996; Melki et ah, 1998; Oostendorp et ah,
1991). Chen et ah (1996) found that 10,000 endospores/g of soil was necessary for
suppression of M arenaria race 1 on peanut in plots on a fine sand soil. Melki et
al.(1998) reported that the cultivation of a susceptible host for more than one season was
needed for P. penetrans to build up its densities to suppressive levels. Oostendorp et ah,
1991 showed that endospore attachment to M. arenaria race 1 increased from 0.11 to 8.6
spores/J2 in plots over a 2-year cropping sequence with peanut (summer) and rye, vetch
or fallow (winter)
The use of air-dried soils infested with P. penetrans was one of the first attempts
to show the biological control potential of this bacterium. Mankau (1973) used air-dried
soil infested with the bacterial spores in greenhouse studies. He reported that after 70
days, plants in the endospore-infested soil had more leaves, greater dry weight, and lower
numbers of root galls than in those soil-free of endospores. However, the use of infested
soil as a source of endospores is time consuming and inconvenient to transport and
handle. Stirling and Wachtel (1980) reported for the first time the use of infested root
powder as a source of endospores and as a method for their mass production. The authors
showed that when they used 100 mg/kg of soil of air-dried and finely ground roots

26
containing 2><109spores/g, that within 24 hours, 99% of the J2 of M.javanica in the pot
had endospores attached to their cuticles. Stirling (1984) used tomato roots containing P.
penetrans-infected females of M. javanica to produce infested, air-dried root powder.
Significant control was obtained when at least 80% of the bioassayed J2 were
encumbered with 10 or more spores per J2. When the infested root powder was
incorporated into root-knot nematode-infested field soil at the rate of 212-600 mg per
kilogram of soil, the number of galls and nematodes in the soil at harvest was
significantly reduced. Also, the application of P. penetrans in air-dried powdered roots at
55 000 spores/cm3 soil in pots infested with 420 J2 significantly suppressed root galling
and egg production of M. javanica through two successive tomato growing seasons. At
planting, there was an average of 14 spores per J2 in the soil (Gowen et al., 1998). The
application of air-dried root powder infested with P. penetrans strains P-20 and P-100 has
been used at Disney World at The Land, Lake Buena Vista, Florida to effectively control
M. arenaria, and M. incognita over the several years on sandy plots (Dickson, pers.
comm, and Brito, pers. observation).
The Effect of Other Microorganisms and Pesticides on Pasteuira
Duponnois and Ba (1998) studied the influence of soil micro flora on the
antagonistic relationship between P. penetrans and M. javanica. The authors showed that
the attachment of P. penetrans to J2 of M. javanica was higher in the presence of larger
soil microbial populations, such as fluorescent strains of Pseudomonas, nematophagous
and mycorrhizal fungi. One of the explanations given by those authors was that those soil
microorganisms may change the soil ionic environment, which favored the attachment of

27
endospores to the nematode cuticle, which is negatively charged (Himmelhoch et al,.
1979). Duponnois et al. (1999) studied the interaction of Enterobacter cloacae and
Pseudomonas mendocina, which had been isolated previously from the rhizosphere of
tomato cv Roman growing in a field infested by both M. javanica and P. penetrans .
Those authors found that P. mendocina and E. cloacae stimulated plant growth, inhibited
the reproduction of M. incognita, and increased the attachment of P. penetrans in vitro.
Enterobacter cloacae increased significantly the reproduction of P. penetrans. They
suggested that the introduction of E. cloacae in soils could enhance the efficacy of P.
penetrans.
The compatibility of P. penetrans with some pesticides increased its potential to
be used in an integrated management of root-knot nematodes (Brown and Nordmeyer,
1985; Freitas, 1977; Singh and Dhawan, 1998). Carbofuran had no effect on the
reproduction of P. penetrans (Brown and Nordmeyer, 1985; Singh and Dhawan, 1998).
Freitas (1977) found that treatment with 1,3-dichloropropene (1,3-D) + 17% chloropicrin,
1,3- D + 25% chloropicrin and 1,3-D + 35% chloropicrin reduced significantly the
percentage of female nematodes with P. penetrans, whereas metham sodium did not have
any effect. However, the author reported that the percentage of nematode females
infected by P. penetrans was significantly lower (1.67%) in the soil treated with methyl
bromide + 33% chloropicrin than in the untreated control (27.50%) under greenhouse
conditions. Under field conditions, the percentage of females infected with P. penetrans
from a plot treated with methyl bromide + 33% chloropicrin was 5% compared to the
untreated control plot, which had 58% of the females infected (Freitas 1997). The

28
exposure of endospores to the fungicides, hymexazol, fosetyl-Al, and carbendazin had no
effect on the attachment or development of endospores (Melki et al., 1998).
Life Cycle
Attachment of endospores to the nematode host. Endospores of the P. penetrans
attach to second-stage juveniles (J2) of root-knot nematodes as they move through soil
pore spaces. After attachment, the sporangial wall and exosporium of the majority of
endospores slough off (Sayre and Starr, 1985). The bacterium is reported to attach to J2
and produce virulent endospores only within the pseudocoelom of a mature female.
However, one isolate of P. penetrans attached to and developed within the pseudocoelom
of juveniles, males, and females of M. acronea isolated originally from cotton (Page and
Bridge, 1985). Also, an endospore-filled J2 of Meloidogyne sp. was isolated from a
suppressive soil infested with P. penetrans in Florida (Dickson, pers. comm.). Stirling et
al. (1990) showed that the number of endospores attached to the cuticle of J2 increased in
proportion to both endospore-concentration and time. Davies et al. (1988) found that the
number of J2 entering the plant host root was reduced when they were encumbered with
15 or more spores . Ahmed and Gowen (1991) reported that 11 or more endospores per
J2 reduced the capability of M. incognita, M. javanica, and M. graminicola to enter the
host roots.
Attachment is one of the major steps toward successful development of P.
penetrans within its host, and it has been studied in several laboratories (Afolabi et al.,
1995; Bird 1989; Chamecki, 1997, Davies et al., 1996). Persidis et al. (1991) used
polyclonal antibodies selected against whole endopsores and wheat germ agglutinin as a

29
probe, and suggested that proteins glycosylated with N-acetylglucosamine are involved in
the attachment. Similar results were obtained using a monoclonal antibody raised to
whole endospores of P-20 isolate of P. penetrans and wheat germ agglutinin (Chamecki,
1997, Chamecki et al., 1998). Mohan et al. (2001) found that fibronectin-like residues on
the cuticle of M. javanica is involved in the attachment of endospores. Other forces such
as hydrophobic interactions may be involved in the attachment of endospores to its host
(Afolabi et al., 1995; Davies et al., 1996; Esnard et al., 1997)
Germination. Unknown factors trigger the germination of the endospore and the
formation of a germ tube. A germ tube emerges through a central opening in the basal
attachment layer after an endospore-encumbered juvenile enters a host root and begins
feeding (Sayre and Wergin, 1997; Sayre and Starr, 1985; 1988; Serracin et al., 1997).
The germ tube penetrates the nematode cuticle and hypodermal tissue, and then enters the
pseudocoelom (Sayre, 1993) where unknown growth factors promote its development
into a vegetative, spherical colony, containing a dichotomously branched and septate
mycelium (Satrr and Sayre 1988). The peripheral fibers of the endospores are closely
associated with the cuticle of the nematode (Sayre and Starr, 1985) and are involved in
the attachment of endospores to the cuticle.
Vegetative stage. When intercalary cells in the microcolony disperse, many
daughter colonies are formed. Eventually quartets of developing sporangia predominate
the pseudocoelom, and then the quartets separate into doublets of sporangia, which
separate into single sporangia that will eventually form the endospores (Sayre 1993).

30
Endospore formation. The formation of bacterial endospores is a regulated and
complex process. The initiation of sporulation is triggered by several genes, spoO genes,
in response to nutrient deprivation (Foster, 1994). It is hypothesized that molecular
functions that control sporulation are the same across all genera of endopsore-forming
bacteria. Small acid-soluble proteins (SASPs) have been shown to be synthesized by
spores of species of Bacillus, Clostridium, and Thermoactynomycetes during sporulation
(Setlow, 1988; Setlow and Waites, 1976). The main types of SASPs found in B. subtilis
are termed the a/p type (Connors et al., 1986) and y type (Hackett and Setlow, 1984),
which are synthesized during the first 3-4 hours of sporulation, and are found only in
spores (Setlow et al., 1992). Previous studies indicated that a/p type-SASPs are DNA-
binding proteins, and their binding to the DNA cause UV resistance by modifying spore
DNA’s UV photochemistry (Manson and Setlow, 1986; Setlow and Setlow, 1987).
Another molecule that is found in spores but not in vegetative cells is the dipicolinic acid,
which is located in the core of the endospores (Madigan et al, 1997). Studies have shown
that calcium, which is present in high concentration in spores, forms a complex with
dipicolonic acid in the core, and confers the heat resistance found in endospores
(Madigan et al., 1997).
The factors that trigger the sporulation of P. penetrans within the pseudocoelom
of the nematode host are not known. However, the sequence of morphological events
during the endogenous spore formation of P. penetrans is similar to other Gram-positive
endospore-forming bacteria (Chen et al., 1997a; Sayre 1993) as follows: i) formation of a
transverse septum within the endospore mother cell; ii) condensation of a forespore from

31
the anterior protoplast; iii) formation of a multilayered wall about the forespore; iv) lysis
of the old sporangial wall; and v) release of an endospore (Sayre 1993).
Chen et al. (1997a) found that the sporogenesis process of P. penetrans generally
matched stages II through VII following vegetative growth reported for Bacillus
thuringiensis. Stage I is unique for Pasteuria sp. The stages are as follows: 1) stage I,
mycelium dichotomously branched and microcolonies fully septate; terminal cells
elongate to form a sporogenous cell; 2) stage II, the terminal cells increase in size and
become oval, 1.2 to 1.7 pm by 0.6 to 1.0 pm, bounded by a 0.002 pm-thick wall; a
membrane is formed about 0.4 pm from the anterior end and separates the forespore from
the parasporium; 3) stage III, parasporium increases in size and engulfs the forespore.
Parasporal fibers are formed and attach to the lower part of the forespore. An inner
membrane defines the forespore protoplast and an outer membrane defines the mother
cell’s protoplast; 4) stage IV, lamella, which rises from the cortex, and inner and outer
spore coats start to form; 5) stage V cortex with formation of inner and outer spore coats;
the inner spore coat is a narrow multilaminar band whereas the outer spore coat is a wide
electron-dense wall; 6) stage VI, formation of exosporium, a delicate membrane that
delimits the outermost layer of a typical Gram-positive bacterium; 7 ) stage VII, complete
maturation with formation of endospore, the basal ring surrounding the germinal pore.
An epicortical layer, which is a discontinuous, electron-dense band was observed between
the cortex and the inner spore coat. Endospores of P. penetrans measure an average of
3.4 pm ± 0.2 by 2.5 pm ± 0.2 (Sayre 1993).

32
The life cycle of this bacterium is not completely synchronized with the life cycle
of the nematode since it is possible to observe different developmental stages
simultaneously at a given time within the pseudocoelom of a single root-knot nematode
female (Chen et al., 1997a). The rate of development is highly temperature-dependent
(Hatz and Dickson, 1992; Serracin et al., 1997; Stirling 1981). The optimum temperature
for the development of the P. penetrans was 35 °C, at which the bacterium completed its
life cycle in 35 days after inoculation (Hatz and Dickson, 1992). An average of2><106
endospores have been found within one single female of P. penetrans-infected
Meloidogyne sp. (Sturhan, 1985) and P. penetrans-infected M. javanica (Stirling 1991).
Soil phase. Endospores are released into soil upon host disintegration.
Endospores are not actively motile in soil; therefore, its contact with the nematode host
must rely on the motility of J2, as well as physical factors affecting endospore distribution
(Sayre 1993). The factors that mediate the movement and survival of endospores of P.
penetrans in soil are not well understood. However, soil water percolation, sizes of soil
pore openings, surface charge of soil particles, tillage practices, and soil microflora may
play important roles in the distribution of endospores (Sayre, 1993). Kamra and Dhawan
(1998) found that at pH 8.0 to 10.0, the average number of endospores encumbered on the
bioassayed J2 of H. cajani was 36 and 26 compared to 10 and 7.0 at pH 6.0 and pH 4.0,
respectively. Those authors also showed that the movement and distribution of
endospores in soil increased with greater pore size, and decreased with an increase in the
silt and clay contents of the soil.

33
Host Specificity
Host specificity has been used for many years to determine species of Pasteuria.
According to Sayre and Starr (1985; 1989) host range of P. penetrans is limited to species
of Meloidogyne.
Host specificity of P. penetrans has been reported in most cases by observing only
the attachment of endopores to its host rather than by establishing infection and
production of mature endospores. Attachment can occur, but endospores might fail to
germinate and propagate within the nematode. Duponnois et al. (2000) tested 25 isolates
of P. penetrans, and found that only six attached, developed, and produced mature
endospores in M. incognita.
Attachment of endospores was greater to the nematode species from which
endospores were originally cultured (Oostendorp et al., 1990; Somasekhar and Metha,
2000). When labeling endospores using monoclonal antibodies, larger areas of the
endospores were labeled when Pasteuria were reared on the same nematode population
from which endospores were taken than from other populations of root-knot nematodes
(Davies et al., 1994). However, Davies et al. (1988) found that a particular isolate of
Pasteuria sp. adapted and shifted from one nematode host to another by continually
culturing the bacterium on a given nematode host. Stirling (1985) reported that
attachment was not always related to the species from which the endospores were
isolated, or to the species of the recipient nematode.
Davies et al. (2001) using only the PP1 strain of P. penetrans and several field
populations of root-knot nematodes collected from Burkino Faso, Ecuador, Greece,

34
Malawi, Senegal, Trinidad, and Tobago showed that the extent of attachment differed
between countries. Also, those authors found similar results when endospores of P.
penetrans collected from those countries were assayed against M. arenaria and M.
incognita.
Endospores of P. penetrans did not attach to the entomopathogenic nematodes
Steinernema glaseri Steiner, Heterorhaditis zealandica Poinar and H. bacteriophora
Poinar and seven new isolates each of Steinernema sp. and Heterorhaditis sp. when
juveniles were exposed to lx 105 spores/ml for 24, 48, and 72 hours at 25 °C
(Somasekhar and Metha, 2000). Similar results were observed by Mendoza de Gives et
al. (1999), who reported that P. penetrans did not attach to animal-parasitic nematodes,
free-living nematodes, including wild type Caenorhabditis elegans (Maupas) Dougherty
and three of its surface (srf) mutants. Oostendorp (1990) also showed that endospores of
P. penetrans did not attach to the free-living nematodes, Panagrelus redivivus (L.)
Goodey and C. elegans, but attached to different species of plant-parasitic nematodes.
The nature and the amount of protein on the surface of endopores may explain
host specificity (Davies et al;. 1992; Persidis et al., 1991). Monoclonal antibodies have
shown that the surface of endospores of P. penetrans isolated from M. incognita race 1 is
highly heterogenous (Davies et al., 1994). These and other studies (Davies and Redden,
1997) have suggested that the virulence of the bacterium to a certain species of root-knot
nematodes is dictated by the surface properties of endospores, and suggested that similar
heterogeneity will be present in the nematode cuticle. Differences in cuticle
characteristics of J2 of root-knot nematodes have been reported (Davies and Danks

35
(1992). Chamecki et al., 1998 showed that the anti-P-20 IgM MAb recognized
differences in the protein extracts from B4, P-20, and PI20 isolates of P. penetrans,
which have different host specificities.
Cultivation
Pasteuria spp. have not been grown successfully in pure culture (Reise et al.,
1988; Williams et al. 1989; Bishop and Eller, 1991). Currently P. penetrans produces
virulent endospores only within the pseudocoelom of females of Meloidogyne spp., which
in turn must be reared on the roots of a plant host or on excised-root systems (Verdejo
and Jaffee, 1988). The mass production of this bacterium relies, currently in the use of
dried, powdered roots obtained from infected root systems grown in a greenhouse
(Stirling and Wachtel, 1980).
Interaction: Host-Parasite
The Role of Adhesin Proteins in the Host-Parasite Relationship
The surface of Gram-positive bacteria has adhesin proteins, also known as
virulence factors, that allow the bacteria to adhere, invade, and colonize tissues (Salyers
and Whitt, 1994). Studies on the composition of the surface proteins have focused
mainly on pathogenic bacteria (Kehoe, 1994).
Virulence factors are classified into two major categories: i) promoters of bacterial
colonization and invasion of the host; and ii) those that cause disease in the host. Among
the virulence factors that promote bacterial colonization are pili, or fimbriae (Robins-
Browne, 1994; Salyers and Whitt, 1994; Suoniemi et al., 1995), and afimbrial adhesins

36
(Salyers and Whitt, 1994) that adhere to mucosal surfaces and bind tightly to the host
cells, respectively. Streptococcus pyogenes has a nonfibrillar adhesin (protein F) that
mediates its attachment to fibronectin, a protein found on many host cell surfaces,
including the mucosa of the human throat (Salyers and Whitt, 1994).
Several environmental signals may affect virulence. These may include
temperature, carbon source, osmolarity, starvation, stress, pH, growth phase; and the
levels of specific nutrients including iron, calcium, sulfate, nicotinic acid, and specific
amino acids (Mekalanos, 1992). Bacteria use different sigma factors to control different
set of genes under specific conditions (Salyers and Whitt, 1994). Similar mechanisms
might be used by P. penetrans to produce endospore adhesins involved in recognition and
attachment to the nematode host.
Objectives
The biochemical events that occur during the development of P. penetrans within
the root-knot nematodes’ pseudocoelom are poorly understood and may provide valuable
insight into the conditions necessary for the formation of virulent endospores.
The objectives of this research project were to 1) determine the sequence of events
required for the formation of P. penetrans spore-associated proteins (adhesins) that are
required for the attachment of endospores, as a function of the development of P.
penetrans within its nematode host, M. arenaria race 1; 2) determine the distribution of
an adhesin-related epitope on the surface of virulent endospores; 3) detect and localize
antigens bearing the epitope during the sporogenesis process; and 4) determine whether or

37
not different species or isolates of Pasteuria share the same adhesin-related epitope,
which is recognized by the anti-P20 IgM MAb. In addition, a polyclonal antibody against
a synthetic polypeptide, which was designed according to the conserved regions of small,
acid-soluble proteins (SASPs) of Bacillus spp. was prepared for use as a probe to detect
SASPs as a development marker in the sporulation process in P. penetrans.

CHAPTER 2
SYNTHESIS AND IMMUNOLOCALIZATION OF AN ADHESIN-ASSOCIATED
EPITOPE IN Pasteuria penetrans
Introduction
Pasteuria penetrans (Thome) Sayre & Starr is a Gram-positive, endospore-
forming bacterial parasite of Meloidogyne spp. Endospores attach to second-stage
juveniles (J2) as they move through soil pore spaces. Unknown factors trigger infection
of the nematode host and germination of the endospore. The germination of the
endospore occurs after the endospore-encumbered juvenile enters host roots and begins
feeding (Sayre and Wergin, 1997; Sayre and Starr, 1985, 1988; Serracin et al., 1997) at
some point in development, presumably before the J2 molts to the third-stage juvenile. A
germ tube penetrates the nematode cuticle and hypodermal tissue, and then enters the
pseudocoelom (Sayre and Starr, 1988), where unknown growth factors promote
vegetative growth, differentiation, sporulation, and maturation of endospores.
Endospores are released into soil upon host disintegration, and more than 2 million
endospores have been found within one single P. penetrans-infected Meloidogyne sp.
female (Sturhan, 1985).
There are certain attributes that make P. penetrans a desirable biological control
agent: 1) endospores are resistant to desiccation, high temperature, and most nematicides
(Dutky and Sayre, 1978; Stirling, 1985; Williams et al., 1989); 2) encumbered juveniles
38

39
have reduced activity and ability to infect roots (Sturhan, 1985); and 3) infected juveniles
complete their life cycle, but females have low or no fecundity (Bird, 1986; Bird and
Brisbane, 1988). These bacteria complete their life cycle and produce virulent
endospores only within the pseudocoelom of Meloidogyne spp., which in turn must be
reared on a plant host either in pots or on excised-root systems (Verdejo and Jaffee,
1988). Attempts to culture P. penetrans in vitro have failed to produce virulent
endospores (Reise et al., 1988; William et al., 1989; Bishop and Ellar, 1991). The
biochemical events that occur during the development of P. penetrans, leading to the
formation of virulent endospores within the pseudocoelom, are poorly understood.
The molecular basis for the recognition and attachment has been the subject of
investigation in several laboratories. Lectin-carbohydrate interactions have been
suggested to be involved in the attachment of P. penetrans to its nematode host. Previous
studies have shown that wheat-germ agglutinin (WGA) inhibited the attachment of
endospores (Bird et al., 1989; Chamecki 1997; Chamecki et al., 1998; Davies and Danks,
1993). Also, proteins extracted from endospores of P. penetrans were recognized, not
only by monoclonal antibodies (Chamecki 1997; Chamecki et al., 1998; Davies and
Redden, 1997) and polyclonal antibodies selected against whole endospores of P.
penetrans (Chamecki et al., 1998; Chen, S. Y et al., 1997; Davies et al., 1992; Persidis et
al., 1991), but also by wheat-germ agglutinin (WGA) (Bird et al., 1989; Chamecki, 1997;
Persidis et al., 1991). These results indicate that one or more epitopes detected by the
antibodies may be glycosylated with P-1-4 linked-acetylglucosamine.

40
Understanding the processes that lead to the growth, differentiation, sporulation,
and maturation of P. penetrans within the pseudocoelom will likely provide a basis to
establish the conditions required for its mass production in vitro. The objectives of this
study were to (1) determine the synthesis of spore-associated proteins (adhesins) as a
function of P. penetrans development within the pseudocoelom of the nematode host, M.
arenaria race 1; (2) determine the distribution of an adhesin-associated epitope on the
surface of virulent endospores; and (3) detect and localize an adhesin-associated epitope
during the sporogenesis process.
Materials and Methods
Nematode Source
Meloidogyne arenaria (Neal) Chitwood race 1 used in this experiment was
isolated originally from peanut (Arachis hypogea L.), Green Acres Research Farm,
University of Florida, Alachua County, Florida. The nematode was reared on tomato
(Lycopersicon esculentum Mill. cv. Rutgers) maintained in a greenhouse. Eggs of the
nematodes were extracted from galled roots by dissolving the gelatinous matrix with
0.5% NaOCl for 20 seconds and collecting the eggs on a sieve with 75 gm-pore openings
(200 mesh) nested in a sieve with 25-pm-pore openings (500 mesh) (Hussey and Barker,
1973). Second-stage juveniles were obtained by hatching the eggs in a modified
Baermann funnel (Pitcher and Flegg, 1968). Juveniles (up to 3-day-old) were collected
on an autoclaved 500-mesh sieve.

41
Pasteuria penetrans Source
Pasteuria penetrans strain P-20 (Oostendorp et al., 1990) used in this study was
collected originally from females of M. arenaria race 1 parasitizing peanut in Levy
County, FL and reared on M. arenaria race 1 growing on tomato in a greenhouse. One to
three-day-old juveniles (J2), with endospores attached to their cuticles were obtained by
incubating them with a suspension containing 1 x 105 endospores/ml overnight, with
constant aeration at room temperature. Endospores were exposed to a mild sonification
(FS14, Fisher Scientific, Suwanee, GA) for 5 minutes before attachment. Twenty spore-
encumbered J2 were chosen randomly from a glass-slide mount, and the number of
endospores attached per J2 was estimated with an inverted compound microscope at
400*. The percentage of endospores attached was 100% with an average of 7 ± 3
endospores per juvenile. Tomato plants (45-day-old seedlings) growing in 15-cm-diam.
clay pots, were inoculated with endospore-attached J2 (3,000 J2/plant). Three days later,
the plants were inoculated again as before. Plants were fertilized twice a week by
watering them with a solution containing 0.63 g of 20-20-20 (N-P-K) (Peters
Professional, general purpose fertilizer, Division, United Industries Corp., St. Louis, MO)
per liter. Water and insecticide applications were provided as needed. At 45 to 60 days
after inoculation, the root systems were harvested, washed with tap water and weighed.
Roots were cut into pieces 2 to 5 cm long and subjected to digestion in a 1-liter
Erlenmeyer containing Rapidase Pomaliq 2F at 1:5 (g/v) (Gist Brocades Pomaliq product
number 7003-A/DSM Food Specialities USA Inc., Menominee, WI), previously
optimized with a buffer system (Chamecki, 1997), and agitated on a shaker at 120

42
oscillations per minute for approximately 24 hours at room temperature. Softened roots
were placed in a sieve with 600 pm-pore openings (30 mesh) nested in a sieve with 150-
pm-pore openings (100 mesh) and sprayed with a heavy stream of tap water according to
Hussey (1971), with modifications. Females and root debris were collected in a beaker
by washing the sieve with a jet of deionized H20, and the contents centrifuged through
20% sucrose (w/v) at 1,500 x g for 5 minutes; the pellet fraction was centrifuged again
through 47% sucrose (w/v) (Chen et al., 2000). The supernatant containing the females
was collected in a beaker and the females were examined for P. penetrans infection with
an inverted microscope at 100*. Endospore-filled females were hand-picked with forceps
under a dissecting microscope at 40* (Nikon, Marietta, GA ), and placed in a 1.5 ml
siliconized microtube containing 300 pi of deionized H20. Infected females were washed
three times in deionized water by centrifugation at 10,000 x g for 2 minutes. Endospores
were collected by grinding the females with a sterile pestle, and the suspension filtered
through a nylon filter either with 21 pm or 18 pm openings (Spectra/Mesh). The
concentration of endospores was determined by counting three 10 pi aliquots using a
hemocytometer (Fisher Scientific) at a magnification of 450x. Endospores retained on a
sieve with 21 pm openings were stored at 4 °C, and used as inoculum for further
production of the bacterium, whereas the endospores retained on a sieve with 18 pm
openings were stored at -20 °C and used for protein extraction.
Experimental Design
Two sets of J2 of M. arenaria, one exposed and the other unexposed to P.
penetrans endospores, were compared with respect to development. These were arranged

43
randomly, with four replications per treatment per each designated “window of P.
penetrans development” (harvest time: 12, 16, 24, and 38 days after inoculation). The
windows of development were based on those reported by Hatz and Dickson (1992) and
Serracin et al. (1997). ‘Rutgers’ tomato seedlings growing in a clay pot (10-cm-diam.)
containing autoclaved sand were inoculated with 3,500 J2/plant (< 2 days old) with and
without endospores attached. Plants were maintained in a growth chamber at 25 °C for
48 hours to allow the nematodes to enter roots. After 48 hours the plants were removed
from pots, and the roots washed thoroughly with tap water to remove any juveniles that
had not penetrated. The seedlings were replanted in clay pots (15-cm-diam.), placed in a
growth chamber at 35 °C, and exposed to a 12-hour-day photoperiod. Plants were
harvested at 12, 16, 24, and 38 days after inoculation. The root systems harvested from
plants were washed in tap water, dried with a paper towel, weighed, cut into pieces 2 to 5
cm long, and incubated in an aqueous solution of commercial Rapidase Pomaliq 2F
(Chamecki, 1997). Nematodes and softened roots were collected on a sieve with 600-
pm-pore openings (30 mesh) nested in a sieve with 25-pm-pore openings (500 mesh),
and washed as before. The nematodes were transferred to a sterile beaker, and twenty
nematodes were hand-picked from each root system. To determine the percentage of
nematodes infected by P. penetrans, and the stage of development of the bacterium from
those nematodes, these were crushed individually in a 2.5 pi drop of lactophenol and 1%
methyl blue (w/v) (Sigma, St. Louis, MO) (Serracin et al., 1997) under a cover glass on a
glass slide, and examined with an inverted microscope (Nikon) at 400* magnification.
The remaining uninfected and infected nematodes from each harvest time were hand-

44
picked, washed, and stored in 1.5 ml siliconized microtubes containing 10 pi PBS (10
mM sodium phosphate buffer, 0.15 M sodium phosphate), pH 7.2 at -20 °C.
Extraction and Determination of Proteins
Uninfected and P. penetrans-infected nematodes harvested at each interval after
inoculation, and mature endospores (2 x 106 spores/10 pi PBS, pH 7.2) used as a control,
were obtained as described before. Nematodes in 10 pi PBS, pH 7.2 were disrupted with
a pestle, and then 30 pi of the extraction solution containing 1.33* UDC (8M urea, 0.04
M dithiothreitol, 0.00665 M CHES buffer, pH 10) was added to each microfuge tube
containing the samples. Microfuge tubes were placed into a water bath for 2 hours at 37
°C, and treated with 20 seconds of sonication (Brankson Cleaning Equipment Company,
Shelton, CN) every 15 minutes. Extracts were centrifuged at 10,000 x g for 5 minutes at
room temperature, and aliquots of the supernatant were collected for storage at -20 °C to
carry out ELISA and SDS-PAGE analyses. Protein estimation was performed by a micro¬
protein assay, based on the Bradford’s method (Bradford, 1976) according to the
manufacturer’s instructions (BioRad, Hercules, CA). Standard curves were generated
using bovine serum albumin (BSA) (Sigma), and colorimetric measurement was
performed at 595 run (Hewlett Packard 8451A Diode Array spectrophotometer, Palo
Alto, CA). The extraction solution containing only urea and CHES buffer pH 9.8 was
made previously, divided in 0.5 ml aliquots, and stored at -20 °C in 1.5 ml microtubes
(Fisher Scientific), and then dithiothreitol was added to it just before the extraction of
proteins.

45
Monoclonal Antibody
The anti-P-20 IgM monoclonal antibody (IgM MAb) used in this study was raised
in mice against whole endospores of P. penetrans P-20 strain and purified on a Sephacryl
S-300 column (J. F. Preston and J. D. Rice, unpubl.). This monoclonal antibody showed
the ability to block attachment of P. penetrans (P-20 strain) to the cuticle of M. arenaria
race 1, and the IC50 is 1.3 * 10'loM. It recognized an epitope shared on several
polypeptides separated by SDS-PAGE (Brito et al., 1998; 2000 Chamecki, 1997;
Chamecki et al., 1998).
Epitope Quantification by ELISA
Proteins (100 ng/well) extracted from P. penetrans-infected nematodes (either 13
infected nematodes harvested at 12 and 16 DAI or 5 infected nematodes harvested or 24
and 38 DAI) at each harvest interval, or from P-20 strain endospores alone as a positive
control (2 x 106 endospores/pl), were applied to appropriate wells of a multi-well plate
with 100 pl/well of coating buffer (15.00 mM Na2C03, 33.40 mM NaHC03, and 0.2%
NaN3) added, and incubated overnight at 4 °C. After washing the wells four times with
PBST (0.2% Tween 20 in 10 mM sodium phosphate buffer, pH 7.6; 154 mM NaCl), the
first antibody, anti-P-20 IgM MAb diluted to 1:100,000 in PBST, was added to the
appropriate wells (100 pl/well) and incubated for 1.5 hours at room temperature. Wells
were washed with PBST again, and the secondary antibody, anti-mouse IgM-alkaline
phosphatase conjugated (Sigma) diluted at 1:4000 in PBST was added to all wells, and
incubated for another 1.5 hours at room temperature, and the wells were washed with
PBST as before. Alkaline phosphatase substrate, 0.1% p-nitrophenol phosphate (w/v)

46
(Sigma) in alkaline phosphatase substrate buffer (0.05 M Na2C03, 0.05 M NaHC03,
0.0005 mM MgCl2) was added to all wells, and color development was measured with an
automated microplate reader at 405 nm (BioRad model 2550, Hercules, CA).
SDS-PAGE Analysis
Proteins (600 ng of total healthy or infected nematode protein) in an appropriate
volume of 10 mM PBS, pH 7.2, were combined with an equal volume of sample buffer
(50 mM Tris/HCl, pH 6.8, 2% SDS w/v, 10% glycerol, 0.05% bromophenol blue w/v, 2%
(3-mercaptoethanol), and boiled for 5 minutes at 100 °C, and then centrifuged for 5
minutes at 10,000 * g. Endospore protein that was extracted from P-20 isolate (2 x 106
endospores/pl) alone was used as a control. Twenty microliters of the supernatants were
transferred into appropriate wells of a polyacrylamide gel of 4% stacking gel (pH 6.8) and
12% separating gel (pH 8.8) with Tris-glycine buffer (Laemmli, 1970). A prestained
molecular weight marker (SeeBlue â„¢ Prestained Standards, Novel Experimental
Technology, San Diego, CA) was loaded onto the same gel. Electrophoresis was carried
out at 100 V for 10 minutes, and then was set at 200 V until the dye marker moved to the
bottom of the gel. Gels were electro-blotted onto nitrocellulose membranes in blotting
buffer (192 mM glycine, 25 mM Tris, 20% methanol) using a Mini Transfer-blot Cell
(BioRad, Hercules, CA) at a constant voltage, 50 V for 2 hours. Proteins either were
stained with AuroDye according to the manufacturer’s instructions (Amersham,
Piscataway, NJ) or with anti-P-20 IgM Mab.

47
Immunoblotting
The nitrocellulose membranes were blocked with 0.5% non-fat dry milk (w/v) in
PBST (10 mM sodium phosphate buffer, pH 7.2, 150 mM NaCl, 0.2% Tween 20)
overnight at 4 °C. Polypeptides containing the epitope recognized by anti-P-20 MAb
were detected as follows: incubation of the membranes with anti-P-20 IgM MAb at 1:
2,000 in PBST, pH 7.2 for 1.5 hours at room temperature on a shaker, washed three times
for 5 minutes each with PBST; incubated with goat anti-mouse IgM MAb conjugated to
alkaline phosphatase (Sigma) diluted to 1:1,000 in PBST, pH 7.2 as secondary antibody
for 1.5 hours at room temperature on a rotatory shaker, followed by three washes with
PBST as above; incubation with substrate buffer (100 mM Tris-HCl pH 9.5, 100 mM
NaCl, 5 mM MgCl2) three times, five minutes each; incubated with alkaline phosphatase
substrate (0.1 mg/ml nitrotetrazolium blue, 0.05 mg/ml 5-bromo-4-chloro-3-indolyl
phosphate) (Promega, Madison, WI) in substrate buffer on a shaker at room temperature
until color development. The blots were washed with deionized water and dried at room
temperature.
Immunofluorescence of Whole Endospores
The immunofluorescence staining was performed as described by Pogliano et al.
(1985) with modifications. Fresh endospores were washed and purified as before, and
then filtered through a woven polyester filter with 18 pm openings. Twenty microliters
of the endospore suspension (2 x 106 endospore/pl) were transferred to a 1.5 ml
siliconized microtube, and fixed in 230 pi of the primary fixative containing 2.7%
formaldehyde and 0.008% glutaraldehyde in 10 mM PBS (10 mM sodium phosphate

48
buffer, pH 7.4, 150 mM NaCl) for 35 minutes on ice. Endospores were placed in 250 pi
10 mM PBS, pH 7.4 and then centrifuged at 6,000 x g three times for 6 minutes each.
After resuspending the endospores in 150 pi PBS, 10 pi of the suspension was transferred
into each of three wells of a microscope slide which had been treated previously with
0.1% poly-L-lysine (Sigma). Each slide was incubated for 30 seconds at room
temperature and then the suspension was aspirated from the wells with a sterile-transfer
pipette (Fisher Scientific). After air drying at room temperature for 30 minutes, the
endospores were incubated in a 10 pl/well with in PBST-BSA (2% BSA (w/v) and 0.05%
Tween 20 (v/v) in 10 mM PBS, pH 7.4 ) for 15 minutes at room temperature to block
nonspecific antibody-binding sites. Primary antibody, anti-P-20 IgM MAb diluted to
1:1,000 in PBST-BSA, was added to the wells and incubated overnight at 4 °C. Wells
containing the endospores were washed in PBST, pH 7.4, five times for 5 minutes each,
and incubated for 2 hours in the dark at room temperature with micron chain-specific,
anti-mouse IgM conjugated with fluorescein isothiocyanate (FITC, Sigma, 1:100 diluted
in PBST-BSA). Anti-P-20 IgM MAb was substituted with non-immune ascites fluid at
1:1,000 dilution as negative control. After washing the wells with 10 mM PBS, pH 7.4,
10 times for 5 minutes each, the slides were mounted in Slow Fade in a PBS-glycerol
solution (Molecular Probes Inc., Eugene, OR). Preparations were examined with
differential-interference contrast and fluorescence microscopy using a Nikon Episcopic
Fluorescence attachment with an excitation filter at 495 nm.

49
Tissue Preparation for Sectioning
Uninfected and P. penetrans-infected M. arenaria race 1 harvested at 20 days
after inoculation at 35 °C were obtained as described above. The procedure used to carry
out this study was a modification of the work by Aldrich et al. (1995); Chen et al.
(1997a); and Zeikus and Aldrich (1975). Fresh nematodes were ruptured with a surgical
knife (Fisher Scientific No. 15) into 40 pi of fixative (1% glutaraldehyde, 4%
formaldehyde, 5% dimethyl sulfoxide in 0.1 M sodium cacodylate buffer, pH 7.2) to
facilitate the penetration of reagents, and then embedded in 2.5% low temperature gelling
agarose (Fisher Scientific) at 45 °C and congealed in the refrigerator (4 °C). The gel was
sliced into square blocks containing individual nematodes and transferred into 12 x 75
mm culture tubes (Fisher Scientific) containing 1.5 ml of the above-mentioned fixative,
and incubated overnight at 4 °C. Agar blocks containing nematodes were washed four
times with cold 0.1 M cacodylate buffer on ice for 30 minutes each and dehydrated in a
cold ethanol series containing the following percentages: 12, 25, 38, 50, 65 for 20
minutes each, and then 75 overnight at 4 °C. This was followed by 85, 95 and two
changes of 100% ethanol for 20 minutes each. The specimens were embedded in LR
White Resin (London Resin White, Electron Microscopy Science, Fort Washington, PA)
series (25 and 50% for 3 and 6 hours, respectively, and 75%, 100%, and 100%, overnight
each time). Agar blocks containing nematodes were transferred into a 1-ml gelatin
capsule containing LR White, and allowed to polymerize at 50 °C for 4 days. Ultrathin
sections (50-70 nm thick) were cut from the resin block with a diamond knife on a LKB

50
8800 Ultratome III microtome (Sweden). Sections were collected on Formvar-coated
nickel grids (100 mesh), and processed for immunogold labeling.
Immunogold Labeling
Nickel grids with sections of uninfected and P. penetrans-infected nematodes, and
with endospore-attached juveniles were floated, section-side down, on 20-pl drops of 1%
non-fat dry milk in PBS, pH 7.2 (0.01M sodium phosphate buffer, 0.15 M sodium
chloride, pH 7.2) on a piece of Parafilm (American National Can â„¢, Menasha, WI) for
15 minutes at room temperature to block nonspecific antibody-binding sites (modified
from Aldrich et al. 1992, 1995; Dykstra, 1993). Grids were floated on 20-pl drops of
primary antibody, anti-P-20 IgM MAb at 1:10,000 dilution in PBS, pH 7.2, and incubated
overnight in a closed petri dish inside a moist chamber at 4 °C. Control grids were
floated on non-immune ascites fluid at 1:10,000 dilution instead of anti-P-20 IgM MAb.
Grids were removed, and floated on 20-pl drops of high salt-Tween buffer, pH 7.2 (0.1%
Tween 20 in 0.02 M Tris-HCl, pH 7.2, 0.5 M Na Cl), two times for 10 minutes each, and
then PBS, pH 7.2, two times for 10 minutes each. Sections were incubated with the
secondary antibody, goat anti-mouse IgM conjugated to 12-nm colloidal gold particle, p-
chain specific (Jackson Immuno Research, West Grove, Pennsylvania), diluted 1:30 in
PBS, pH 7.2, at room temperature for 1 hour. After washing as above in high salt-Tween
buffer and PBS, the grids were floated in Trumps buffer, pH 7.2 (McDowell and Trump,
1976) for 10 minutes at room temperature in order to stabilize the antigen-antibody
complex, and then washed with deionized water. Sections were stained with 0.5% uranyl

51
acetate for 7 minutes, and aqueous lead citrate solution for 2.5 minutes and observed on a
Zeiss EM-10 transmission electron microscope at 80 kV. All reagents used to carry out
this study were ultrapure-TEM grade.
Results
Microscopic Examination
The vegetative growth stage of P. penetrans was observed only in nematodes
harvested at 12 and 16 days after inoculation (Table 2.1). At 24 days after inoculation,
mixed developmental stages of thalli showed advanced differentiation, including quintets,
quartets, triplets, doublets; sporulation, oval-shaped immature sporangium; and mature
endospores with visible exosporium were first observed. At 38 days only various phases
of sporulation and mature endospores were present in the pseudocoelom of M. arenaria
race 1.
Epitope Quantification by ELISA
The anti-P-20 IgM MAb did not recognize proteins extracted from infected
nematodes harvested at 12 and 16 days after inoculation (Fig. 2.1 A). However, the
monoclonal antibody reacted with proteins extracted from infected nematodes harvested
at 24 and 38 days after inoculation (Fig. 2.1 A). The protein per infected nematode was
0.453 pg at 12; 0.466 pg at 16, 1.175 pg at 24, and 2.049 pg/nematode at 38 days after
inoculation (Fig. 2.IB). The total protein per infected nematode increased with
developmental time (Fig. 2.IB), and was correlated with the increase in the signal
detected by the anti-P-20 IgM MAb (Fig. 2.1 A). At 24 and 38 days after inoculation, the
ELISA-based absorbance at 405 pm per infected nematode was 1.50 and 3.20,

52
Table 2.1. Percentage of different developmental stages of Pasteuria penetrans in
Meloidogyne arenaria race 1 on tomato ‘Rutgers’ at 12, 16, 24, and 38 days after
inoculation at 35 °Ca.
Days Postinoculation
Developmental stage
12
16
24
38
Vegetative growth
90
90
0
0
Differentiation
0
0
15
0
Sporulation
0
0
85
5
Mature endospores
0
0
65
95
Twenty nematodes were observed at each harvest date, and percentage of
nematodes at 12, 16, 24, and 38 days after inoculation. Nematodes were hand-picked,
placed on a glass slides, and crushed separately in 2.5 pi of lactophenol plus 1% methyl
blue (w/v) under a cover glass. Infected nematodes were examined with the use of an
inverted microscope (*400) to determine the percentage of the different developmental
stages of P. penetrans within the pseudocoelom of Meloidogyne arenaria race 1. Note
that at 24 days after inoculation more than one developmental stage was observed within
the pseudocoelom of a single nematode. The developmental stages observed were:
vegetative growth including mycelial colonies only within the pseudocoelom;
differentiation stage, with presence of thalli differentiation, including quintets, quartets,
triplets, doublets; sporulation stage, with many doublets and developing endospore with
distal swollen ends connected by intercalary ends; and mature endospores; with free
endospores with exosporium clearly visible.

53
respectively, which was proportional to the amount of adhesin-associated epitope
increased as P. penetrans reached its maturation stage (Fig. 2.1 A). These results suggest
that the antigens bearing the epitope, which was recognized by anti-P-20 IgM MAb, were
synthesized at later stages of development associated with sporulation of P. penetrans
within the pseudocoelom of M. arenaria race 1.
SDS-PAGE Analysis and Immunoblotting
Analysis of individual proteins extracted from uninfected and P. penetrans-
infected nematodes at each window of development showed some differences in the
protein profiles related to the infection of the nematode by the bacterium (Figs. 2.2A-B;
2.3A-B). The immunoblot showed that Anti-P-20 IgM MAb did not recognize any
protein extracted from uninfected nematodes harvested at 12, 16, 24, and 38 days after
inoculation (Lanes 2, 3, 4, and 5) (Fig. 2.2B); nor were proteins extracted from infected
nematodes harvested at 12 and 16 days detected in the immunoblot (Lanes 2, and 3) (Fig.
2.3B). However the immunoblot revealed that the monoclonal antibody reacted with
protein extracts of infected nematodes harvested at 24 and 38 days after inoculation
(Lanes 4, 5) (Fig. 2.3B) and with endospore protein of the P-20 strain used as the control
(Lanes 6) (Figs. 2.2B; 2.3B).
Immunofluorescence
Note the general shape of P-20 strain just before it was examined with the
fluorescence microscope (Fig. 2.4A). Labeling of whole endospores of P. penetrans

54
Fig. 2.1. Adhesin-associated epitope and total nematode protein per infected
nematode as a function of the development of Pasteuria penetrans. A) Levels of adhesin-
associated epitope determined by ELISA using anti-P-20 IgM MAb at 1:100,000 dilution
in PBST, pH 7.6. Infected nematode total proteins (100 ng/well) was applied in 100
pl/well at the final treatment. Alkaline phosphatase substrate, 0.1% p-nitrophenol
phosphate (w/v) was added to all wells, and color development was measured at 405 nm.
B) Total nematode protein of infected nematodes. Data shown are 40 minutes readings.
Lines above the bars indicate SE of the mean for six replicates per treatment.

55
Fig. 2.2. Blots of sodium dodecyl sulfate-polyacrylamide gels of uninfected
Meloidogyne arenaria protein extracts after electrophoresis. Proteins of uninfected
nematodes, harvested at each window of development. Extracts in the appropriate
volume of sample buffer were boiled for 5 minutes at 100 °C, and 20 pi of the
appropriate extract containing 600 ng of total protein was applied per lane. A) Proteins
were detected by staining with AuroDye according to manufacturer’s instructions. B)
Immunodetection of blotted antigens with anti P-20 IgM MAb at 1: 2,000 dilution in
PBST, pH 7.2. Lane 1 - Molecular weight markers, See Blue pre-stained proteins; Lanes
2, 3, 4, and 5 - Total proteins extracted from uninfected nematodes at 12, 16, 24, and 38
days after inoculation; Lane 6 - Proteins extracted from P. penetrans P-20 endospores.

56
kDa
1 2 3 4 5 6
Fig. 2.3. Detection of Pasteuria penetrans adhesin-associated epitope as a
function of its development within the pseudocoelom of Melodogyne arenaria racel.
Nematode total proteins and endospore proteins were extracted as for Fig 2. Proteins,
600 ng in 20 pi of the appropriate extract plus sample buffer was loaded into each lane.
A) Detection of blotted proteins with AuroDye. B) Western blot of P. penetrans infected
nematodes probed with anti-P-20 IgM MAb at 1:2,000 dilution in PBST, pH 7.2. Lane
1 - Molecular weight markers, See Blue pre-stained proteins; Lanes 2, 3, 4, and 5 -
Epitope bearing proteins extracted from P. penetrans infected nematodes at 12, 16, 24,
and 38 days after inoculation; Lane 6 - Proteins extracted from P. penetrans P-20
endospores.

57
isolate P-20 by anti-P-20 IgM MAb was not uniform (Fig. 2.4B), which suggests that the
adhesin-associated epitope is not uniformly distributed on the surface of mature
endospores.
Immunogold Labeling
Anti-P-20 IgM MAb did not recognize any nematode tissue and there were no
gold particles observed over the thin section of either uninfected females or J2 with
associated endospores (Fig 2.5). The adhesin-associated epitope was not present in the
ultrathin sections of vegetative cells (vc) or stage I (Fig. 2.6A) or in stage II of
sporogenesis of P. penetrans isolate P-20 (Fig. 2.6B). Note a membrane (arrow head) is
forming at 1/3 from the anterior, which occurs at this stage of sporogenesis (Chen et al.,
1997b). Labeling of the adhesin-associated epitope was first observed over an ultrathin
section of the stage III sporogenesis, mainly on the parasporal fibers (pf) (Fig. 2.7A). The
antigens bearing the epitope were detected not only over the parasporal fiber (pf) (Figs.
2.7B-2.9A) but also over the sporangium(s) as P. penetrans continues to sporulate (Figs.
2.8A-B; 2.9A). The mature endospore was heavily labeled, and the epitope was localized
in the sporangium (s), exosporium (ex), and parasporal fibers (pf) (Fig. 2.9A). The outer
spore coat (oc), inner spore coat (ic), cortex (c), protoplasm (p), and basal ring (br) were
not labeled (Fig. 2.9A). No labeling was observed over any structure of the mature
endospore when non-immune ascites fluid was used (Fig. 2.9B).

58
I
B
t
Fig. 2.4. Differential interference contrast (DIC) and fluorescence microscopy
photomicrographs of whole endospores of Pasteuria penetrans P-20 isolate (100*
magnification). A) Overall shape of whole endospores using DIC. B) Labeling of an
adhesin-associated epitope on the surface of whole endospores using anti-P-20 IgM MAb
at 1:1000 dilution in PBST-BSA, overnight at 4 °C, as primary antibody, and anti IgM
Mab-FITC labeled as secondary antibody diluted 1:1000 in PBST-BSA. Arrows heads
identify regions of nonuniform labeling.

59
Fig 2.5. Longitudinal section of uninfected second-stage juvenile of Meloidogyne
arenaria (1-day-old) probed with anti-P-20 IgM MAb at 1:10,000 dilution, and anti-IgM,
gold-conjugated at 1:30 dilution. No gold particles are visible over the nematode tissues.
Scale Bar = 0.5 pm.

Fig. 2.6. Immunocytochemical localization of an adhesin-associated epitope
during the development of Pasteuria penetrans within the pseudocoelom of M. arenaria.
Thin sections of all stages of development of P. penetrans were probed with anti- P-20
IgM MAb at 1:10,000 dilution, and anti-IgM MAh gold-conjugated diluted to 1:30
dilution as secondary antibody and examined by transmission electron microscopy. Scale
Bars = 0.5 pm. A) Stage I of sporogenesis. A longitudinal ultrathin section of mycelial
colony (arrow) of P. penetrans P-20 isolate. No labeling is visible over the mycelium.
B) Stage II sporogenesis of P. penetrans. Note that a membrane is forming at 1/3
distance from the anterior end (arrow read), which is characteristic of this stage. No
labeling occurs over any structure of this stage of development of the bacterium.


Fig 2.7. Labeling of sporogenous stages of Pasteuria penetrans. Scale bars = 0.5
pm. A) Stage III sporogenesis showing labeling of the adhesin-associated epitope (arrow
head) mainly over the parasporal fibers (pf). B) Stage IV sporogenesis, gold particles
(arrow head) are concentrate in the parasporal fibers (pf). Note that the vegetative cell
(vc) was not labeled.

63

Fig 2.8. Sporogenous stages of Pasteuria penetrans. Scale bars = 0.5 pm. A)
Stage V of sporogenesis. Gold label (arrow head) indicating antibody binding is present
over the parasporal fibers (pf) and exosporium (e). B) Stage VI of sporogenesis, labeling
of the adhesin-associated epitope is observed over the parasporal fibers (pf) and
exosporium (ex).

65
WK

Fig. 2.9. Late sporogenous stage of Pasteuria penetrans. Scale Bars = 0.5 pm.
A) Stage VII of sporogenesis, a mature endospore showing the sporangium (s)
exosporium(ex), and parasporal fibers (pf) heavily labeled, whereas the outer spore coat
(oc), inner spore coat (ic ), epicortex (ep), cortex (c), protoplasm (p), and basal ring (br)
are not labeled. Note that the parasporal fibers (pf) were not uniformly labeled (arrow
head). B) A mature endospore of the Pasteuria penetrans, stage VII used as control. No
label is observed over the thin section of the endospore.

67

68
Discussion
Pasteuria penetrans completes its life cycle within the pseudocoelom of female of
root-knot nematodes. The physiological aspects of its life cycle have been studied and are
reasonably well understood (Chen and Dickson, 1997; Freitas et al., 1997; Hatz and
Dickson, 1992; Serracin et al., 1997; Nakasono et al. 1993, Stirling, 1981, Giannakou et
al., 1999). However the biochemical aspects are poorly understood. Seven
morphological stages of development through sporulation have been determined as I, II,
II, IV, VI, and VII (Chen and Dickson, 1997). The initial step in the life cycle of
Pasteuria is the recognition/attachment of the endospores to the cuticle of a free living J2
root knot-nematode host. Infection of the host and germination of the endospores occur
once the J2 enters the root tissue of a plant host, and establishes a permanent feeding site
(Sayre and Starr, 1985, 1988). Vegetative growth, differentiation, and formation of
Imbriani, 1975; Sayre, 1993; Sayre and Wergin, 1997; Serracin et al., 1997). The
mechanisms involved in the attachment have been the subject of study in several
laboratories. The results of these studies have led to the establishment of a model where
glycoproteins, designated as adhesins and lectin are involved in the interaction of P.
penetrans and the nematode host (Persidis et al., 1991; Davies and Danks, 1993).
Previous studies have shown that microbial adhesins, or bacterial surface proteins, known
as virulence factors such as pili, or fimbriae (Robins-Browne et al., 1994; Salyers and
Whitt, 1994; Suoniemi et al., 1995), and afimbrial adhesins (Salyers and Whitt, 1994),
allow bacteria to attach, colonize, and invade their hosts. For instance, Streptococcus
pyogenes, a gram-positive pathogen has a nonfibrillar adhesin (protein F) that mediates its

69
attachment to fíbronectin, a protein found on many host cell surfaces, including the
mucosa of the human throat (Salyers and Whitt, 1994). However, the mechanisms used
by Pasteuria spp. to produce virulent endospores within the pseudocoelom of the
nematode host is not well understood. Mohan et al. (2001) found that fibronectin-like
proteins extracted from M. javanica are involved in the attachment of endospores. In this
study, we determined the relative time of the synthesis of an adhesin-associated epitope
during the development of P. penetrans within the pseudocoelom of M. arenaria race 2;
detected and localized this epitope during endospore development, and also determined
the distribution of the epitope on the surface of mature endospores using a monoclonal
antibody directly selected against whole mature enendospores of P. penetrans P-20
isolate.
ELISA and immunoblot analysis revealed that only proteins extracted from P.
penetrans-infected nematodes at 24 and 38 days after inoculation were recognized by
anti- P-20 IgM MAb and the amount of the epitope was highest at the height of
sporulation (38 days after inoculation) than at any other developmental stage (12, 16, and
24 days after inoculation). The Western blot showed a higher degree of similarity in the
protein profile of P. penetrans-infected nematodes at 38 days after inoculation to the
mature P-20 spore protein, used as a control, than with P. penetrans-infected nematodes
from any other window of development. Examination of the infected nematodes
harvested at 12 and 16 days by light microscopy revealed that only the vegetative growth
stage, including clusters of mycelial colonies and thalli, were found throughout the
pseudocoelom of nematodes. At 24 and 38 days, sporulation and maturation stages were

70
observed within the pseudocoelom. Therefore, the synthesis of the adhesin-associated
epitope occurred at a certain developmental stage relative to the sporogenesis process,
and it was absent in vegetative growth and differentiation stages.
The synthesis of specific molecules at specific times during the germination,
growth, and sporulation of the endospore-forming bacterium, Bacillus subtilis, has been
rigorously established. For instance dipicolinic acid (pyridine-2, 6-dicarbonate) is formed
during the first 5 hours of sporulation (Schlegel, 1986), whereas the small, acid-soluble
spore proteins (SASPs), a group DNA-binding proteins (at neutral to slightly alkaline
pH), are synthesized after 3-4 hours into sporulation (Johnson and Tipper, 1981; Setlow
1985). Both molecules are found only in endospores (Fliss et al., 1985; Schlegel, 1986).
Even though some molecules are synthesized at specific stages of sporulation, it is
possible that they are degraded and used to carry out a certain function at another stage.
For instance, during the first 5 hours of sporulation in B. subtilis much of the vegetative
cell protein is degraded (Schlegel, 1986).
Immunofluorescence labeling showed that the adhesin-associated epitope is not
uniformly distributed on the surface of virulent endospores. The heterogeneity of
endospore surface has been observed not only within populations but also between
populations of P. penetrans (Davies and Redden, 1997). Previous studies have shown
that differences in the amount and nature of spore-surface proteins, as recognized by
several monoclonal antibodies, may account for surface heterogeneity of endospores as
well as host specificity (Davies et al., 1992). Davies et al. (1994) using monoclonal
antibodies showed that the surface of endospores of the PP1 strain of P. penetrans is

71
highly heterogenous. These and subsequent studies (Davies and Redden, 1997) have
suggested that endospore surface properties are responsible for the virulence of P.
penetrans.
Antigens bearing the epitope were synthesized during the sporogenesis process.
Labeling was first observed at stage III of the sporogenesis, mainly in the parasporal
fibers. In contrast to stage III sporogenesis, mature endospores were heavily labeled and
the adhesin-associated epitope was localized in the parasporal fibers, sporangium, and
exosporium.
The general pattern of the labeling of the adhesin-associated epitope over thin
sections of a mature endospore was similar to a previous study, where mature endospores
were probed with a polyclonal antibody (Persidis et al., 1991). These authors concluded
that the labeling did not show any preference to a certain structure of the endospore and
suggested that a nonspecific binding of the antibodies could have occurred. These
observations may reflect a heterogeneity in the polyclonal antibody preparation and/or
selection of a single stage of development. In our immunocytochemistry work, it was
shown that the adhesin-associated epitope is synthesized at a certain stage of development
related to endospore formation and it is localized initially in the parasporal fibers early in
stage III, becoming widespread throughout the sporangium and exosporium, but not in the
central body of the stages IV, V, VI, and VI of sporogenesis. Label was not uniformly
distributed in the parasporal fibers. Also no labeling was observed in the outer or inner
spore coat, epicortex, cortex, protoplasm, and basal ring.

72
These observations establish a window of development in which the adhesin-
associated epitope is formed, and where further studies concerning the formation of this
epitope should be directed. The fact that the epitope is distributed over several structures
of the mature endospores suggests its involvement in the recognition of the nematode
host as an early event in the attachment process. It may increase the chances for a
cooperative interaction between the adhesin epitope with receptors on the cuticle of the
nematode host, such as carbohydrate binding proteins (Bird et al., 1989; Davies and
Danks, 1993; Persidis et al., 1991) and fibronectin-like residues. (Mohan et al., 2001) as
well as other forces, that may be involved in the attachment, such as hydrophobic
interactions (Afolabi et al., 1995; Davies et al., 1996; Esnard et al., 1997).

CHAPTER 3
DETECTION OF ADHESIN PROTEINS AND IMMUNOLOGICAL
DIFFERENTIATION OF Pasteuria spp. USING A MONOCLONAL ANTIBODY
Introduction
Pasteuria penetrans (Thome) Sayre & Starr, the first species of Pasteuria
described as a parasite of plant-parasitic nematodes, is a widespread endospore-forming
bacterial parasite of root-knot nematodes (Meloidogyne spp.) (Sayre and Starr, 1985).
Over the years, several more species of nematodes in other genera have been reported as
hosts of species of Pasteuria. (Chen and Dickson, 1998). To date, three species of
Pasteuria have been described in addition to Pasteuria penetrans (Sayre and Starr, 1985).
These are Pasteuria ramosa, a parasite of water fleas, Daphnia spp. (Sayre et al., 1983)
which is the type species of the genus; Pasteuria thornei isolated from Pratylenchus spp.
(Starr and Sayre, 1988), and Pasteuria nishizawae a parasite of cyst nematodes of the
genera Heterodera and Globodera (Sayre et al.,1991). In recent years more species of
Pasteuria have been proposed: i) Pasteuria sp., designated as S-l strain (Bekal et al.,
2000) from Belonolaimus longicaudatus Rau; ii) a large- and a small-spored isolate of
Pasteuria spp. each from Hoplolaimus galeatus (Cobb) Thome (Giblin-Davis et al.,
1990); and iii) three isolates which attach and complete their life-cycles in Heterodera
spp.; one isolate was from cereal cyst nematode, Heterodera avenae Wollenweber
(Davies et al., 1990), a second strain from pea cyst nematode, Heterodera goettingiana
73

74
Liebscher in Münster, Germany (Sturhan et al., 1994), and a third isolate that infects
soybean cyst nematode, Heterodera glycines Ichinohe, Pasteuria sp. NA (Heterodera
glycines-infecting Pasteuria), Urbana, IL, USA (Atibalentja et al., 2000).
Traditionally species of Pasteuria are identified based on morphometries,
morphology, ultrastructural characteristics, and host specificity (Davies et al., 1990;
Giblin-Davis et al., 1995; Sayre and Starr, 1985; Sayre et al., 1983; 1991; Starr and Sayre,
1988; Sturhan et al., 1994). More recently, 16S rDNA has been used to carry out
systematics studies of P. ramosa (Ebert et al., 1996), P. penetrans (Anderson et al.,
1999), Heterodera glycines-infecting Pasteuria (Atibalentja et al., 2000), and Pasteuria
sp. S-l strain (Bekal et. al., 2000). Also, the use of serology through hybridoma
technology might be a useful probe for the identification of Pasteuria spp. The anti-P-20
IgM monoclonal antibody (MAb) raised against whole mature endospores of P-20 isolate
of P. penetrans was used as a probe in this study. This MAb was selected on the basis of
its ability to block attachment of P. penetrans isolate P-20 to M. arenaria race 1
(Chamecki et al., 1998) (Chapter 2). Previous studies have shown that this MAb
recognized an epitope shared on several polypeptides separated by SDS-PAGE (Brito et
al., 1998; Chamecki 1997; Chamecki et al., 1998). The appearance of an adhesin-
associated epitope was tracked during development and localized during sporogenesis of
the P-20 within its nematode host (Brito et al., 1998; 1999). The objectives of this study
were to determine whether different strains and species of Pasteuria share this adhesin-

75
associated epitope which is involved in the attachment of P. penetrans P-20 strain to M.
arenaria race 1, and to use anti-P-20 IgM MAb as a probe to separate strains and species
of Pasteuria.
Material and Methods
Origin of Pasteuria Species and Isolates
The designations and origins of the species and isolates of Pasteuria spp.
(Table 3.1) were as follows: two isolates of P. penetrans; one designated P-20
(Oostendorp et al., 1990) originally collected from M. arenaria race 1 (Neal) Chitwood,
from peanut (Arachis hypogea cv. Florunner) roots growing in a naturally infested field in
Levy County, FL, and the other one designated Pl-UFLA (Souza and Campos, 1997),
originally isolated from a mixed population of M. javanica and M. incognita, Lavras,
Minas Gerais, Brazil; H. glycines-infecting Pasteuria, (Pasteuria sp. NA) (Atibalentja et
al., 2000) from cysts of H. glycines collected from the rhizosphere of soybean plants
{Glycines max (L). Mirril), Urbana, IL. Pasteuria sp. strain S-l (Bekal, et al., 2001;
Giblin-Davis et al., 2001) isolated from the sting nematode B. longicaudatus, L-l (large-
spored strain), LS-1 (small- spored strain) from the lance nematode, H. galeatus (Giblin-
Davis et al., 1990), and Pasteuria from Rhabditis sp. (Giblin-Davis pers. comm.) were all
originally collected from bermudagrass (Cynodon spp.) turf growing in a naturally
infested field, at the Ft. Lauderdale Research and Education Center, University of Florida,
Broward County, Fort Lauderdale, FL. Pasteuria sp. C-l isolate (Han et al., 1999) was
originally collected from Criconemoides sp. in a naturally infested soil where peanut

76
(Arachis hypogea L. cv. Florunner) was growing at the Green Acres Agronomy Farm,
University of Florida, Alachua County, Gainesville. A ring nematode isolate of Pasteuria
also isolated from Criconemoides sp. collected in a peanut field (Williston), FL (Dickson
per. comm.), and spiral nematode isolate of Pasteuria isolated from Helicotylenchus sp.
extracted from the rhizosphera of bermudagrass turf from California (Crow, pers.
comm.).
Propagation of Bacterial Species and Isolates
Pasteuria penetrans P-20 and Pl-UFLA isolates were propagated on M.
arenaria race 1 and M. javanica respectively, growing on ‘Rutgers’ tomato. Endospores
of each strain were attached separately to second-stage juveniles (J2) (up to 2 day old) of
root-knot nematodes using a centrifugation method (Hewlett and Dickson, 1993).
Juveniles (3,000 J2 per plant) with approximately six endospores attached per J2 were
inoculated on 55-day-old tomato plants growing in 15-cm-diameter clay pots in a
greenhouse. Endospore-filled females were harvested from the root systems 45 to 60
days after inoculation. Root systems were placed in a 1-liter Erlenmeyer flask containing
Rapidase Pomaliq 2F (Gist Brocadest Pomaliq, 7003-A/DSM Food Specialities USA
Inc., Menominee, WI) optimized previously with a buffer system at 1:5 (g/v) (Chamecki,
1997), and placed on a shaker at 120 oscillations/minute for approximately 24 hours at
room temperature. The softened roots and nematodes were poured onto a sieve with 600
pm-pore openings nested in a sieve with 150 pm-pore openings, and subjected to a heavy
stream of tap water to dislodge the nematodes (Hussey, 1971), with modifications.
Nematodes and root debris were collected in a beaker, and the contents centrifuged in

77
Table 3.1. Species and isolates of Pasteuria.
Species or isolates
Reference
P. penetrans P-20
Meloidogyne. arenaria race 1 (Oostendorp et al.,
1990)
P. penetrans Pl-UFLA
Meloidogyne spp. (Souza and Campos, 1997)
Hg Pasteuria sp. NA
Heterodera glycines (Atibalentja et al., 2000)
Pasteuria sp. S-l
Belonolaimus longicaudatus (Bakel et al., 2001)
C-l isolate
Criconemoides sp. (Han et al., 1999)
L-l isolate
Hoplolaimus galeatus (Gibli-Davis, 1990)
LS-1 isolate
Hoplolaimus galeatus (Gibli-Davis, 1990)
Rhabditis infecting-Pasteuria
Rhabditis sp. (Giblin-Davis, pers. comm.)
Ring nematode-infecting Pasteuria
Criconemoides (Dickson, pers. comm.)
Spiral nematode-infecting Pasteuria Helicotylenchus sp. (Crow, pers. comm.)

78
20% sucrose (w/v) at 1,500 * g for 5 minutes, and the resulting pellet was again
centrifuged in 47% (w/v) sucrose (Chen et al., 2000). Female nematodes were collected
and examined for Pasteuria infection with an inverted microscope at 100 * magnification
(Leica, Davie, FL). Pasteuria-iniected females were hand-picked using a dissecting
microscope at 40 * magnification (Nikon, Marietta, GA), and placed in 1.5 ml siliconized
microtubes containing 900 pi of deionized water. Nematodes were centrifuged in
deionized water three times at 10,000 x g for 2 minutes, and then stored in 500 pi
deionized water at 4 °C until used. Pasteuria sp. S-l, L-l, LS-1 isolates, and the
Rhabditis sp. infecting-Pasteuira, and spiral nematode-infecting Pasteuria were isolated
from their nematode hosts growing in bermudagrass (Cynodon dactylon (L) X C.
transvaalensis Burt-Davy cv. Tifway or C. magenissii Hurcombe cv. Tifgreen) turf in a
naturally-infested field. The C-l isolate and ring nematode-infecting Pasteuria were
obtained from Criconemoides sp. extracted from the rhizosphere of peanut (Arachis
hypogea L. cv. Florunner) grown in a naturally-infested soil in a greenhouse, and peanut
field, respectively. All nematodes were extracted from the soil using a centrifugal-
flotation method (Jenkins, 1964). Pasteuria-'mÍQCied nematodes were hand-picked under
a dissecting microscope, and placed in deionized water. After washing the nematodes
with deionized water as above, they were stored in 900 pi deionized water at 4° C until
used. Pasteuria sp. NA was propagated on H. glycines race 3 reared on soybean cv. Lee
growing in a naturally-infected soil in a greenhouse. Pasteuria-'mkcXQd cysts and females
were extracted from the rhizosphere of 3-month old soybean plants by washing the soil
through a sieve with 850 p-pore openings nested over a sieve with 180 p-pore openings;

79
and nematodes were collected in a sterile beaker. Nematodes were transferred into 200-
ml centrifuge tubes containing 150 ml of deionized water, and centrifuged at 2,000 x g
for 4 minutes. The resulting pellets were re-suspended with 50% sucrose solution, and
again centrifuged for 35 to 45 seconds. The supernatant was poured through a sieve with
180 p-pore openings (Atibalentja et al., 2000), and collected in a sterile beaker. Infected
females and cysts were hand-picked based on their opaque appearance, washed three
times with deionized water by centrifugation at 10,000 x g for 2 minutes, placed in a 1.5
ml siliconized microtube containing 100 pi deionized water, and stored at 4 °C until
used.
Extraction and Determination of Proteins
Nematodes infected by species or isolates P-20, Pl-UFLA, S-l, C-l, ring
nematode and spiral nematode isolates of Pasteuria, and cysts infected with the Pasteuria
sp. NA strain were obtained as described before. Infected nematodes and cysts in the
appropriate 1.5 ml siliconized microtube containing deionized water were crushed with a
pestle, filtered with 18-pm-pore membrane, and the endospore concentration of the
suspension was determined with a hemocytometer (Fisher, Suwanee, GA) under a
compound microscope (Leica, Davie, FL) at a magnification of 40*. Ten microliters of
endospore suspension was transferred to a 1.5 ml siliconized microtube, and 30 pi of the
extraction solution containing 1.33* UDC (8 M urea, 0.04 M dithiothreitol,0.00665 M
CHES buffer, pH 9.8) was added. Microtubes were placed into a water bath for 2 hours
at 37 °C, with 20 seconds of mild sonication (Brankson Cleaning Equipment Company,
Shelton, CN), every 15 minutes. Extracts were centrifuged at 10,000 x g for 5 minutes at

80
room temperature, and aliquots of the supernatant were collected and stored at -20 °C
until used. Protein estimation was performed by a micro-protein assay, according to the
manufacturer’s instructions (BioRad, Hercules, CA). Standard curves were generated
using bovine serum albumin (BSA) (Sigma, St. Louis, MO), and colorimetric
measurement was performed at 595 nm (Hewlett Packard 8451A Diode Array
spectrophotometer, Palo Alto, CA). The UDC stock solution was made previously using
only urea and CHES buffer, pH 9.8, divided in 0.5 ml aliquots, and stored at -20 °C in 1.5
ml microtubes. Dithiothreitol was added to the microtubes just before the extraction of
proteins.
Preparation of Infected Nematodes for TEM
All Pasteuria-infecied nematodes were obtained as described above except for the
NA Pasteuria which was obtained as follows: infested dry soil (50 g) was placed in a
100x15 ml petri dishes, and the soil water was adjusted to 100% field capacity to increase
the rate of endospore attachment. The dish was left uncovered at room temperature
(Brown et al., 1985). After 3 days 1,000 juveniles (J2) of H. glycines race 3 were added,
and the moisture level was adjusted to 50% of field capacity. Dishes were incubated for 7
days at room temperature (Oostendorp et al., 1990), and the J2 were extracted by the
centrifugal-flotation method (Jenkins, 1964). J2 with endospore attached were hand¬
picked, and placed in a 1. 5 ml microtube, washed three times with deionized water by
centrifugation at 10,000 x g for 2 minutes, and stored at 4 °C until used.
A modified protocol was used to carry out the TEM part of this study (Aldrich et
al., 1995; Chen et al., 1997a; Zeikus and Aldrich, 1975). Nematodes were hand-picked

81
into a 40 pi-drop of fixative (1% glutaraldehyde, 4% formaldehyde, 5% dimethyl
sulfoxide in 0.1 M sodium cacodylate buffer, pH 7.2), and cut into 2 to 4 pieces with a
surgical knife (Fisher Scientific No. 15) to aid penetration of the reagents. Nematodes
were transferred into a 50 pl-drop of 2.5% agarose (Fisher) at 45 °C , and then cooled in a
refrigerator. After cutting the gel into square blocks, they were placed in 12x75
millimeter culture tubes (Fisher) containing 1.5 ml of the fixative, and incubated
overnight at 4 °C. After rinsing the nematodes four times with 0.2 M cadodylate buffer,
pH 7.2 on ice for 30 minutes each, they were dehydrated in a cold ethanol series: 12, 25,
38, 50, 65, 75, 85, 95, and two changes of 100% for 20 minutes each, except for 75%,
which was kept overnight at 4 °C. Specimens were infiltrated with LR White resins
(London Resins White, Electron Microscopy Science, Fort Washington, PA) series: 25%
and 50% for 3 and 6 hours, respectively, 75% and two changes in 100% overnight each
time). Blocks were placed in lml-gelatin capsule containing LR White resin, and allowed
to polymerize for 4 days at 50 °C. Thin sections, 50-70 nm thick were cut with a
diamond knife on a LKB 8800 Ultratome III microtome (Sweden). Sections were
collected and mounted on Formvar-coated nickel grids (100 mesh) and processed for
immunocytochemistry.
Immunocvtochemistrv
Nickel grids containing section were placed, face down, on 20pl-drops of 1% non¬
fat dry milk in PBS, pH 7.2 (0.01M sodium phosphate buffer, 0.15 M sodium chloride)
on a piece of Parafilm (American National Can, Menasha, WI) for 15 minutes at room
temperature, to block nonspecific antibody-binding sites (Aldrich et al., 1992; 1995;

82
Dykstra, 1993) with modifications. Grids were transferred to 20 (a 1-drops of the first
antibody, anti-P-20 IgM MAb at 1:10,000 or 1:40,000 dilution in PBS, pH 7.2, and
incubated overnight in a closed petri dish inside of a moist chamber at 4 °C. Grids were
floated in 20 pl-drops of high salt tween buffer, pH 7.2 (0.1% Tween 20 in 0.02M Tris-
HC1, pH 7.2, 0.5 M NaCl), and in PBS, pH 7.2 twice in each buffer for 10 minutes each,
before incubation with goat anti-mouse IgM conjugated to colloidal gold (1:30 dilution in
PBS, pH 7.2, 12 nm gold) (Jackson Immuno Research, West Grove, PA) for 1 hour at
room temperature. Grids were washed again in high salt tween buffer, and PBS, and were
incubated for 10 minutes in Trumps buffer, pH 7.2 (McDowell, and Trump, 1976) at
room temperature in order to stabilize the antigen-antibody complex. Sections were
washed with deionized water, and stained with 0.5% uranyl acetate for 7 minutes, and
aqueous lead citrate solution for 2.5 minutes. Controls were probed with non-immune
ascites fluid and goat-anti mouse IgM conjugated to gold to ensure that the results were
not due to non-specific binding. Sections were examined on a Zeiss EM-10 transmission
electron microscope at 80kV. All reagents used in this study were ultra pure-TEM grade.
SDS-PAGE Analysis
Proteins extracted from endospores ofPasteuria NA, S-l, C-l, Pl-UFLA, P-20,
ring nematode and spiral nematode isolates of Pasteuria were individually combined with
equal volume of sample buffer (62.5 mM Tris-HCl, pH 6.8, 2% SDS w/v, 10% glycerol,
0.05% bromophenol blue w/v, 2% P-mercaptoethanol) (BioRad), boiled for 5 minutes at
100 °C, and centrifuged for 5 minutes at 10,000 * g. Twenty microliters (600 ng of
protein) of the supernatant was loaded into the wells of a Tris-glycine polyacrylamide 4%

83
stacking gel (pH 6.80) and 12% separating gel (pH 8.8) (BioRad). Electrophoresis was
carried out at 100 V for 10 minutes, and then it was set for 200V until the bromophenol
blue dye had migrated to the bottom of the gel. Proteins were transferred onto
nitrocellulose membranes in blotting buffer (192 mM glycine, 25 mM Tris, 20%
methanol) using a Mini Transfer-blot Cell (BioRad) at a constant voltage, 50 V for 2
hours. Protein bands were visualized either by Aurodye (Amersham, Piscataway, NJ)
according to manufacturer’s instructions or anti-P-20 IgM MAb (Chapter 2). Standard
ladders for molecular mass were loaded in the same gels ( SeeBlue â„¢ Prestained
Standards, Novel Experimental Technology, San Diego, CA).
Immunoblotting
Blots were first blocked overnight with 0.5% skimmed milk (w/v) in PBST (10
mM sodium phosphate buffer, pH 7.2, 150 mM NaCl, 0.2% [v/v] Tween 20 at 4 °C.
Blots then were incubated with anti-P-20 IgM MAb diluted 1:2,000 in PBST, pH 7.2 for
1.5 hours on a rotatory shaker at room temperature, and washed with PBST, three times, 5
minutes each. Blots were incubated with goat-anti mouse IgM conjugated to alkaline
phosphatase (Sigma) diluted 1:1,000 in PBST, pH 7.2 for 1.5 hours at room temperature,
and washed as before with PBST, pH 7.2. After washing blots with substrate buffer (100
mM Tris-HCl, pH 9.5, 100 mM NaCl, 5 mM MgCl2) three times, 5 minutes each at room
temperature, blots were incubated with alkaline phosphatase substrate (0.1 mg/ml
nitrotetrazolium blue, 0.05 mg/ml 5-bromo-4-chloro-3-indolyl phosphate) (Promega,
Madison, WI) in substrate buffer on a rotatory shaker at room temperature until color
development. Blots were washed with deionized water and dried at room temperature.

84
Results
Immunocvtochemistrv
Intense gold labeling was specifically associated with sporangium (s), exosporium
(ex), and parasporal fibers (pf) of P-20, Pl-UFLA, Rhabditis-'mfectmg Pasteuria, S-l,
LS-1, L-l, and C-1 (Figs. 3.1-4). Labeling was not observed over the outer spore coat
(oc), inner spore coat (ic), cortex (c) (Figs. 3.1-4), and basal ring (br) (Figs. 3.1 A, B) of
the endospores of P-20 and Pl-UFLA, collected in USA and Brazil. No labeling was
observed over any structure of Pasteuria sp. NA used as a control (Fig. 3.5). Gold
particles were not observed on the germ tube (gt) of Pasteuria sp. NA, nor over the
cuticle of the cyst nematode, H. glycines (Fig. 3.6), however the parasporal fibers (pf)
were labeled heavily (Fig. 3.6).
SDS-PAGE and Immunoblotting Analysis
AuroDye staining showed that at least three bands of proteins (arrow head) are
common among the Pasteuria sp. NA, S-l, C-l, Pl-UFLA, ring nematode and spiral
nematode isolate of Pasteuria and P-20, used as control (Fig 3.7A ). Immunoblotting
showed qualitative and quantitative differences among all the those isolates and species
of Pasteuira (Fig 3.7B). All species and isolates share the same epitope because it was
recognized by anti-P-20 IgM MAb (Lanes 2-8) (Fig.3.7B). Isolates Pl-UFLA and P-20
showed similar bands of proteins with equal intensity (Lane 5 and 6) (Fig. 3.7B).
Similarities in bands of proteins also were observed between spiral nematode isolate of
Pasteuria and ring nematode isolate of Pasteuria (Lanes 7 and 8) (Fig. 3.7B). Also the
same degree of similarity in the protein profiles was observed among the Pasteuria sp.

85
NA, Pl-UFLA, and P-20 extracts (Lanes 2, 5,and 6) (Fig. 3.7B). The strongest bands
were observed in proteins extracts from Pasteuria sp. NA, Pl-UFLA, and P-20 (Lanes 2,
5, and 6) (Fig. 3.7B). Pasteuria sp. S-l showed one band of protein (arrow) (Lane 3)
(Fig. 3.7B) that is shared among all other strains (Lanes 2, 4, 5, 6, 7, and 8) whereas C-l
strain showed one band of protein (arrow) (Lane 4) (Fig. 3.7B) that is also observed from
the protein extract of Pasteuria sp. NA, P-20, and ring nematode-infecting Pasteuria
(Lanes 2, 6, and 8). The isolate C-l showed one strong band of protein with molecular
weight between 50 and 36 kDa (Lane 4), which appeared similar to a band of less
intensity from the extract of the spiral nematode 'mÍQC\\n%-Pasteuria (Lane 7) and ring
nematode-infecting Pasteuria extract (Lane 8) (Fig. 3.7B).
Discussion
The immunocytochemistry indicated that the adhesin-associated epitope as
recognized by anti-P-20 IgM MAb is shared among P-20, P-1 UFLA, NA Pasteuria
strain, Rhabditis sp.-infecting Pasteuria, Pasteuria sp. S-l strain, C-l, LS-1, and L-l.
The immuno-gold labeling patterns were similar for all the species and isolates examined.
The broad distribution of the adhesin epitope over several structures of endospores of
different species and isolates of Pasteuria may increase their capabilities to attach to their
host due to cooperative interactions between the adhesin epitope with receptors on the
cuticle of the nematode host, such as carbohydrate binding proteins (Bird et al., 1989;
Davies and Danks, 1993; Persidis et al., 1991) and fibronectin-like residues (Mohan et al.,

Fig.3.1. Transmission electron micrographs of Pasteuria endospore sections,
probed with anti-P-20 IgM MAb at 10,000 dilution. Scale bars = 1 pm. A) Thin section
of an mature endospore of P-20 isolate. Labeling was observed over the sporangium (s)
exosporium (ex), and parasporal fibers (pf), whereas the outer spore coat (oc), inner spore
coat (ic), cortex (c), and basal ring (br) were not labeled. Note that the parasporal fibers
(pf) were not uniformly labeled (arrow head). B) Section of a mature endospore of Pl-
UFLA. Sporangium (s), exosporium(e), and parasporal fibers (pf) are heavily labeled.
Labeling is absent over the outer spore coat (o), inner spore coat (i), and basal ring (br).

87

Fig. 3.2. Gold labeling of endospores of different isolates and species of
Pasteuria. Scale bars = 1 pm. A) Thin section of a mature endospore of the Pasteuria
sp. NA attached to a second-stage juvenile showing the labeling of the adhesin-associated
epitope over parasporal fibers (pf), but not on the outer spore coat (oc), inner spore coat
(ic), protoplasm (p), and nematode cuticle (nc). B) Endospore of Rhabditis sp.-infecting
Pasteuria showing the labeling the adhesin-associated epitope over the sporangium (s),
and parasporal fibers (pf). No labeling is observed over the vegetative cell (vc).

89

Fig. 3.3. Immunoelectron microscopy of endospores of Pasteuria spp. A)
Labeling of an adhesin-associated epitope over a thin section of a endospore of S-l strain
Gold particles (gp) are observed on the sporangium (s), exosporium (ex), and parasporal
fibers (pf), but not over the outer spore coat (oc) and, inner spore coat (ic). Scale bar = 1
pm. B) Thin section of a endospore of the C-l isolate showing gold particles (gp) over
the sporangium (s), exosporium(ex), and parasporal fibers (pf)- Scale bar = 0.5 pm.


Fig. 3.4. Labeling of endospores of two isolates of Pasteuria spp. Scale
bars = 0.5 pm. A) LS-1 isolate endospore showing labeling over the sporangium (s),
exosporium(ex), and parasporal fibers (pf), whereas the outer spore coat (oc), inner spore
coat (ic) were not labeled. B) Endospore of the L-l isolate labeling of the adhesin-
associated epitope over the sporangium (s), exosporium (ex), and parasporal fibers (pf).
Gold particles (g).

93

94
Fig 3.5. Thin section of the Pasteuria sp. NA used as a control. Section was
treated as the experimental ones, but the first antibody was replaced with non-immune
ascites fluid. No gold particles were observed over any structure of the mature endospore
(me), parasporal fibers (pf), and nematode cuticle (n). Scale bar = 1 pm.

95
Fig. 3.6. Thin section of a mature endospore (me) of the Pasteuria sp. NA.
Parasporal fibers (pf) were heavily labeled whereas the genn tube (gt), and the central
body (arrow head) were not labeled. Note that the nematode cuticle (n) was not labeled.
Scale bar = 1 pm.

96
1 2 3 4 5 6 7 8
Fig. 3.7. Detection of an adhesin-associated epitope in different strains of
Pasteuria. Endospore proteins, 600 ng in 20 pi of the appropriate extract plus sample
buffer was loaded into each lane. A) Detection of blotted proteins with AuroDye. B)
Western blot of blotted proteins probed with anti-P-20 IgM MAb at 1:2,000 dilution in
PBST, pH 7.2. Lane 1 - Molecular weight markers, See Blue pre-stained proteins ( 10
gl/well); Lane 2 - Pasteuria sp. NA; Lane 3 - Pasteuria sp. S-l; Lane, 4 - C-l; Lane 5 -
Pl-UFLA; Lane 6 - P-20 isolate of P. penetrans; Lane 7 - Spiral nematode-infecting
Pasteuria; and Lane 8 - Ring nematode-infecting Pasteuria.

97
2001), as well as other forces that may be involved in the attachment, such as
hydrophobic interactions (Afolabi et al., 1995; Davies et al., 1996; Esnard et al., 1997;
Spiegel et al., 1996).
Immumoblot analyses showed that all isolates of Pasteuria share the same adhesin
-associated epitope. This epitope also was recognized previously by anti-P-20 IgM Mab
in protein extracts from P-120, and B4 isolates of Pasteuria penetrans (Chamecki, 1997;
Chamecki et al., 1998). Several antigens bearing the same epoitope were recognized by
the MAb; however quantitative and qualitative differences were observed among all
species and isolates. Pasteuria sp. NA, a parasite of Heterodera glycines, and Pl-UFLA
and P-20 isolates from root-knot nematodes have similar profiles of protein bands to
those found for P. penetrans isolates, which are clearly different from those of protein
profiles of extracts from Pasteuia sp. S-l, spiral nematode-infecting Pasteuria, and ring
nematode-infecting Pasteuria. Pasteuria sp. S-l and C-l isolate both showed a certain
degree of uniqueness in their protein profiles, which may differentiate them from the
other isolates and species. Similar results were obtained by Davies et al. working with
polyclonals (1992) and Chamecki (1997) and Chamecki et al, (1998). Davies et al.
(1992) found that proteins extracted from three isolates of P. penetrans: PP1, PNG, and
PCal, probed with a polyclonal antibody previously raised against whole spores of P.
penetrans PP1, showed qualitative and quantitative differences in the protein profiles, but
most of the proteins were conserved. Differences detected with the anti-P-20 IgM MAb
confirm not only previous studies with P-120, P-100 and P-20 isolates by Chamecki

98
(1997) and Chamecki et al., (1998) but also the potential use of this MAb as a probe to
detect different species of Pasteuria.
The detection of the adhesin epitope over thin sections of endospores of the
different species and isolates of Pasteuria indicates that this epitope does not confer host
specificity, but it may confer virulence. Since this epitope has been proven to be involved
in the attachment of P. penetrans P-20 endospores to its nematode host, M. arenaria race
1, its broad distribution on thin sections of endospores of other species and isolates of
Pasteuria with different host specificity indicates that it is only one component of the
attachment process. The results of these studies show that the adhesin associated-epitope,
which was not found associated with any of several species of Bacillus previously
examined (J. Harrison and J. F. Preston, unpubl.; Schmidt et al., 2001) is unique for
strains and species of Pasteuria.

CHAPTER 4
SYNTHESIS OF SMALL, ACID-SOLUBLE SPORE PROTEINS
IN Pasteuria penetrans
Introduction
Pasteuria penetrans (Thome) Sayre & Starr is a Gram-positive mycelial-and
endospore-forming bacterium known from around the world. It has shown great potential
as a biological control agent of root-knot nematodes (Meloidogyne spp.) (Chen and
Dickson, 1998; Dickson et al., 1994; Trudgill et al., 2000). For instance the application
of 10,000 endospores per gram of soil effectively suppressed the peanut root-knot
nematodes on peanut (Chen et ah, 1997b).
Phylogenetic studies using 16S rDNA of Pasteuria ramosa (Ebert et ah, 1996), P.
penetrans (Anderson et ah, 1999) and Heterodera glycines-infecting Pasteuria
(Atibalentja et ah, 2000) have placed Pasteuria spp. in a clade with some species of
Bacillus. Bacillus spp. have had many aspects of their life cycle, such as growth factors,
sporulation genes, and proteins, well characterized, therefore they can provide an
appropriate model to study species of Pasteuria.
Spore of various species of Bacillus have a number of small, acid-soluble spore
proteins (SASPs), which are synthesized during the first 3-4 hours sporulation (Setlow et
ah, 1992). SASPs comprise approximately 5 to 10% of the protein in dormant spores of
Bacillus and Clostridium species (Cabrera-Martinez and Setlow, 1991; Setlow, 1988).
99

100
The major function of these proteins is to bind to and protect the DNA from
environmental trauma. The main types of SASPs found in B. subtilis are termed the a/p
type (Connors et al., 1986) and y type (Hackett and Setlow, 1984). Previous studies
indicated that a/p type-SASPs are DNA-binding proteins, and their binding to the DNA
protect it from UV irradiation (Manson and Setlow, 1986; Setlow and Setlow, 1987).
Studies in vivo and in vitro have shown that a/p-type SASP are non-specific DNA-
binding proteins, which trigger a conformational change in DNA from a B-like to an A-
like structure (Mohr et al., 1991; Setlow et al., 1992). The a, P and y from make up about
18, 18, and 36% of the total SASPs of B. subtilis strain 168 with molecular weights of
5,900, 5,900, and 11,000 kDa (Johnson and Tipper, 1981). Another funtion of these
proteins is to be a source amino-acids for new protein synthesis during spore germination
and outgrowth (Setlow, 1981). Setlow and Primus (1975) showed that spores lack
several important pools of amino acids and germinating spores lack several key amino
acid synthetic enzymes.
Amino acid sequences of a/p type-SASP are highly conserved within and across
species of a variety of endospore-forming bacteria (Cabrera-Martinez, 1991; Setlow,
1988). Setlow (1988) analyzed the SASPs of 19 species of Bacillus and found that 27
residues were conserved exactly and 11 were similar. He also showed that this
conservation exists between the tested Bacillus species and various species of
Clostridium. Therefore the SASPs may be an useful model to study SASPs in P.
penetrans. The objective of this study was to select an antibody against a synthetic

101
peptide (sequence: Cys-Ser-Val-Gly-Gly-Glu-Ile-Thr-Lys-Arg-Leu-Val), a conserved
sequence of amino acid in Bacillus SASPs, and use it as a probe to detect SASPs in P.
penetrans.
Materials and Methods
Pasteuria penetrans Endospores Source
The source and the procedure to obtain the endospore of P. penetrans, P-20 strain,
used in this study was the same as for Chapter 2.
Bacillus subtilis Spore Source
Purified and lyophilized spore of B. subtilis, ATCC strain 6051 was kindly
provided by Dr. James F. Preston and Josh Loomis, Microbiology and Cell Science
Department, University of Florida, Gainesville, Florida.
Extraction and Determination of SASPs from P. penetrans and B. subtilis
The extraction of SASPs from spores of B. subtilis and P. penetrans was
performed according to Johnson and Tipper (1981). Clean and dry spores of B. subtilis
(10 mg/ml) and endospore suspension of P. penetrans (1 x 108/p.l) in 2N HC1 were used to
perform the acid rupture of spores and extraction of SASPs (APPENDIX A). Protein
estimation was performed by a micro-protein assay based on the Bradford method
(Bradford, 1976), according to the manufacturer’s instructions (BioRad, Hercules, CA).
Standard curves were generated using bovine serum albumin (BSA) (Sigma, St. Louis,
MO), and colorimetric measurement was performed at 595 nm (Hewlett Packard 8451A
Diode Array spectrophotometer, Palo Alto, CA).

102
Conjugation of SASP Peptide to Carrier Proteins
A synthetic peptide (sequence: Cys-Ser-Val-Gly-Gly-Glu-Ile-Thr-Lys-Arg-Leu-
Val), which was designed based on a conserved region of Bacillus SASPs, was
synthesized by the Protein Chemistry Core Facility, Interdisciplinary Center for
Biotechnology Research, University of Florida, Gainesville, FL. Two milligrams of the
peptide were added to each of two 1.5 ml microfuge tubes containing 300 pi of
conjugation buffer (83 mM Sodium phosphate buffer, pH 7.2, 0.9 M NaCl, 0.1 M EDTA,
0.02% azide) according to the manufacturer’s instructions (44895 sulfoLink" Kit, Pierce
Chemical Company, Rockford, IL). Twenty microliters of the peptide mixture was added
to a 1.5 ml microfuge tube and stored at 4 °C until use. In separate 1.5 ml microfuge
tubes, pre-activated keyhole limpet hemocyanin (KLH) or bovine serum albumin (BSA)
were mixed with conjugation buffer, pH 7.2 at a concentration of 2 mg per 200 pi. The
peptide mixture (280 pi) was combined with each protein mixture (200 pi) in 1.5 ml
microfuge tube and incubated at room temperature for 2 hours prior to purification.
Purification of the Conjugates
Each of the conjugate mixtures was added to a gel filtration column, which was
washed previously three times with the purification buffer (0.083 M sodium phosphate
buffer, pH 7.2, 0.9 M NaCl) according to the manufacturer’s instructions (44895
sulfoLink" Kit, Pierce Chemical Company, Rockford, IL). Using the purification buffer
as diluent, 15 fractions of 1 ml each were collected and proteins were determined by
measuring the absorbance at 280 nm, to verify which of the fractions had the most
conjugate (Hewlett Packard 8451A Diode Array spectrophotometer, Palo Alto, CA). The

103
peak fraction for each conjugate was retained and concentrated using a Centricon â„¢ 3
device according to the manufacturer s’ instructions (Amicon Inc., Beverly, MA ). The
final concentration was estimated for each of the conjugate based on Abs 280 nm.
Fractions were stored at 4 °C overnight and used to inject in hens for polyclonal antibody
production.
Immunization of Hens for Production of Polyclonal Antibodies
The polyconal antibodies were raised in White Leghorn hens (134-4 and 135-1)
against KLH-peptide and (135-2 and 135-3) against BSA-peptide. Approximately 100 pi
of KLH-peptide was injected in the wing (subcutaneous), and 100 pi in the footpad of
each hen. Two hens were used for each immunogen. Hens were boosted 14 days after
the initial injection as follows: approximately 75 pi of KLH-peptide was injected in the
wing, and 75 pi in the footpad of each two hens (134-4 and 135-1), whereas 50 pi of
BSA-peptide was injected in the wing and 60 pi in the footpad of each two hens ((135-2
and 135-3). Eggs were collected every day and stored at 4 °C. To monitor the formation
of the antibody, two consecutively-laid eggs were combined, extracted, and analyzed for
titers against the KLH-peptide and BSA-peptide. Intact egg yolks (~ 15 ml yolk per egg)
(APPENDIX B) were removed and rinsed three times with 0.1 M sodium phosphate
buffer, pH 7.6. Every two yolks (30 ml/two eggs) were combined and lysed in 120 ml
(four volumes) of 0.1 M sodium phosphate buffer, pH 7.6, and stirred (slowly) at room
temperature for 30 minutes. Yolk extracts were stored at 4 °C overnight. Activities of
the antibodies in each of the egg yolk extracts were determined by ELISA.

104
Determination of IgY Activities in Yolk Extracts
A general procedure for ELISA was done as described in Chapter 2. Each
antigen, KLH-peptide and BSA-peptide diluted at 10,000 in coating buffer (15.00 mM
Na2C03, 33.40 mM NaHC03, 0.0.2% NaN3) was added to the appropriate wells of the
microtiter plate in 75 pi aliquots. Plates containing the antigens were incubated at 4 °C
overnight. Extracts from every two egg yolks were screened for antibody activity at 100-
and 1,000-dilutions in 10 mM PBST, pH 7.6 (10 mM sodium phosphate buffer, pH 7.6 ,
0.9% NaCl, 0.2% Tween 20). After transferring 75 pi of the egg yolk extract to the
appropriate wells, plates were incubated at room temperature for 1.5 hours, and washed
with PBST four times. One milliliter of each yolk suspension was centrifuged at 10,000
x g for 2 minutes prior to dilutions. The secondary antibody, anti-chicken IgG conjugated
to alkaline phosphatase (Sigma) at 1:2,000 in PBST was added to each well (75 pi per
well), and incubated for another 1.5 hours at room temperature. The plates were washed
with PBST four times again as before. To each well, 75 pi of alkaline phosphatase
substrate, 0.1% /7-nitrophenol phosphate (w/v) (Sigma) in alkaline phosphatase substrate
buffer (0.05 M Na2C03, 0.05 M NaHC03, 0.0005 mM MgCl2) was added. The reaction
was monitored by reading the absorbance at 405 nm with an automated microplate reader
(Model 2550 ELA, BioRad, Hercules, CA).
Extraction of IgY from Egg Yolk Extracts
Egg fractions that showed high IgY antibody activities were combined in group of
two, and the IgY antibodies were purified using polyethylene glycol (Sigma) following

105
the method described by Poison et al. (1985) (APPENDIX B). The activities of IgY
antibodies were determined by ELISA as follows.
Determination of Activities of Purified IgY
IgY fractions extracted from each pool were centrifuged as described before, and
each supernatant was diluted to 100, 1,000, and 10,000 in PBST, pH 7.6, and applied to
the appropriate well (100 pl/well). The antigens, KLH-peptide, and BSA-peptide were
diluted to 10,000 in coating buffer (15.00 mM Na2C03, 33.40 mM NaHC03, and 0.0 2%
NaN3), and applied to the wells (100 pl/well). The activities of IgY antibodies in the
purified fractions were determined by monitoring the absorbance at 405nm as above.
Concentration of Purified IuY using Centripep
The pool (pool 2) that exhibited the highest activity (as shown by ELISA) was
concentrated using a Centripep 10 device, following the manufacturers’ instructions
(Amicon Inc., Beverly, MA). The volume of the anti-peptide IgY obtained after the
concentration was 4.5 ml. The affinity of the anti-peptide IgY for SASPs of B. subtilis
and P. penetrans was evaluated by ELISA.
Affinity of Anti-Peptide IgY for SASPs
ELISA procedure was the same as described above. The SASP of B. subtilis and
P. penetrans were diluted to 100 and 1,000 whereas KLH-peptide and BSA-peptide, used
as positive controls, were diluted to 10,000 and 100,000 in coating buffer. Anti-peptide
IgY (pool 2) was diluted to 100, 1000, and 10,000 in PBST.

106
Results
Purification of the Conjugates
Fractions number 3 of each purified conjugate had the highest activity. Fraction 3
containing the KLH-peptide or BSA-peptide was used to inject into the appropriate hen.
A final concentration of 0.8 mg/ml was estimated for each of the conjugates that was used
for immunization.
Determination of IgY Activities in Yolk Extracts
Two White Leghorn hens (134-5 and 135-1) injected with KLH-peptide yielded
levels of anti-KLH-peptide antibody (Figs. 4.1-4.4), whereas hens (135-2 and 135-3)
injected with BSA-peptide produced very low levels of anti-BSA-peptide antibody (data
not shown). These results indicate that KLH-peptide was a better immunogen than BSA-
peptide. The highest antibody activities were observed after the antigenic boost of the
hens. The egg yolk extracts diluted at 100 showed higher antibody activities than at a
dilution of 1,000 dilution regardless of the antigen used in the ELISA, and the hens used
to raise the antibodies. The highest level of anti-peptide activity was observed in yolk
from eggs laid by hen 134-5 at 16 to 26 days after injection (Fig 4.1), using KLH-peptide
as antigen in ELISA. However this high level of activity might be a background due the
presence of KLH, in the antigen used. Using BSA-peptide as antigen, hen 134-5, at 16 to
26 days after injection produced some level of activity (Fig. 4.3). This indicated that the
activity observed using BSA-peptide as antigen was not due to the antigen’s background.
Therefore, egg yolk extracts laid by hen 134-5 between 20 and 28 days after injection
were used for IgY extraction (Fig. 4.3).

107
Days after initial injection
Fig. 4.1. Activities of antibodies in egg yolk extracts collected from hen
134-5, 34 days after injection of 100 pi KLH-peptide as immunogen (80 pg per 100 pi)
into the wing and 100 pi into the foot pad. A boost injection was performed at 14 days,
75 pi was injected into the wing and 75 pi into the footpad. Egg yolk extracts were used
at 100 and 1,000 dilution in PBST, whereas the antigen (KLH-peptide) was diluted at
10,000 in coating buffer. Absorbance (405 nm) represent readings recorded at 45 minutes
after the addition of the substrate in ELISA.

108
Days after initial injection
Fig.4.2. Activities of antibodies in egg yolk extracts collected from hen 135-1, 34
days after injection of 100 pi KLH-peptide as immunogen (80 pg per 100 pi) into the
wing and 100 pi into the foot pad. A boost injection was performed as in Fig. 4.1. Egg
yolk extracts were used at 100 and 1,000 dilution in PBST, whereas the antigen (KLH-
peptide) was diluted at 10,000 in coating buffer. Absorbance (405 nm) represent readings
recorded at 45 minutes after the addition of the substrate in ELISA.

109
Days after initial injection
Fig.4.3. Activities of antibodies in egg yolk extracts collected from hen 134-5, 34
days after injection of 100 pi KLH-peptide as immunogen (80 pg per 100 pi) into the
wing and 100 pi into the foot pad. A boost injection was performed as in Fig. 4.1. Egg
yolk extracts were used at 100 and 1,000 dilution in PBST, whereas the antigen (BSA-
peptide) was diluted at 10,000 in coating buffer. Absorbance (405 nm) represent readings
recorded at 45 minutes after the addition of the substrate in ELISA.

110
Days after initial injection
Fig. 4.4. Activities of antibodies in egg yolk extracts collected from hen 135-1
34 days after injection of 100 pi KLH-peptide as immunogen (80 pg/100 pi) into the
wing and 100 pi into the pad. A boost injection was performed as in Fig. 4.1. Egg yolk
extracts were used at 100 and 1,000 dilution in PBST, whereas the antigen (BSA-peptide)
was diluted at 10,000 in coating buffer. Absorbance (405 nm) represent readings
recorded at 45 minutes after the addition of the substrate in ELISA.

Ill
Extraction IgY from Egg Yolk Extracts
The five pools of egg yolk extracts eggs laid between 20 and 28 days after
injection) from hen 134-5 showing high levels of IgY activities were combined by every
two polls just before the extraction of IgY antibodies was performed. Each new poll
yielded a total of 20, 18, and 16 ml of purified anti-KLH-peptide IgY obtained from a
total of 10 eggs.
Determination of Activities of Purified IgY
Antibodies purified from pool 2 showed the highest activity regardless of the
planting antigens (Fig. 4.5). The titer of anti-KLH-peptide IgY purified from pool 2 was
determined using 100; 1,000; and 10,000 dilutions in PBST, and KLH-peptide and BSA-
peptide antigens at 10,000; 100,000; 1,000,000 dilution in coating buffer (Fig. 4.6).
Affinity of Anti-Peptide IgY for SASPs
Anti-peptide KLH IgY isolated from pool 2 recognized SASPs of B. subtilis as
well as SASP of P. penetrans (Figs. 4.7; 4.8), respectively.
Discussion
Hens successfully raised antibodies against the synthetic peptide (sequence: Cys-
Ser-Val-Gly-Gly-Glu-Ile-Thr-Lys-Arg-Leu-Val), a conserved sequence of amino acid in
SASPs of Bacillus spp. KLH was more efficient as a carrier protein than BSA. The anti¬
peptide KLH IgY purified from yolks extracts laid by hen 134-4 between days 20 to 28
was reactive with SASPs of B. subtilis and cross reactive with SASPs of P. penetrans.
These results indicate that ELISA can be used as a method to detect SASPs of B. subtilis

112
t
Antigen dilutions
Fig. 4.5. Activities of anti-KLH-peptide IgY antibodies extracted from egg yolk
extracts laid by hen 134-5 at 20 to 28 days after injection with KLH-peptide. Antibodies
were used at 1,000 dilution in PBST. KLH-peptide and BSA-peptide at 10,000 dilution
in coating buffer were used as antigens. Readings were recorded at 45 minutes after the
addition of alkaline phosphatese substrate, p-nitrophenyl phosphate in ELISA. Lines
above bars indicate SE for the average of replicates per assay at P = 0.05.

113
Fig. 4.6. Activities of purified IgY antibodies (pool 2) from egg yolk extracts laid
by the hen 134-5. Antibodies were dilute to 100; 1,000; and 10,000 in PBST, pH 7.6
whereas the antigens, KLH- peptide and BSA-peptide, were dilute to 10,000; 100,000;
and 1,000,000 in coating buffer. Readings were recorded at 15 minutes after the addition
of alkaline phosphate phosphatese substrate, /?-nitrophenyl phosphate.

114
Antibody dilutions
Fig. 4.7. Activities of anti-KLH-peptide IgY antibodies extracted from egg yolk
extracts (pool 2) laid by hen 134-5. Antibody was used at 100; 1,000 and 10,000 dilution
in PBST. SASP-Bacillus subtilis at 100 and 1,000 dilution in coating buffer were used as
an antigen. Readings were recorded at 45 minutes after the addition of alkaline
phosphatese substrate, p-nitrophenyl phosphate in ELISA. Lines above bars indicate SE
for the average of replicates per assay at P = 0.05.

115
Antibody dilutions
Fig.4.8. Activities of anti-KLH-peptide IgY antibody extracted from egg yolk
extracts (pool 2) laid by hen 134-5. Antibody was used at 100; 1,000; and 10,000
dilution in PBST. SASP-Pasteuria penetrans at 100 and 1,000 dilution in coating buffer
were used as an antigen. Reading were recorded at 45 minutes after the addition of
alkaline phosphatase substrate, />-nitrophenyl phosphate in ELISA. Lines above bars
indicate SE for the average of replicates per assay at P = 0.05.

116
and P. penetrans. It also indicated that there was some level of similarity between the
SASPs of these two species of endospore-forming bacteria. The anti-peptide KLH IgY
will be purified further, and used as a probe to detect SASPs of B. snbtilis and P.
penetrans. Further studies are needed to characterize the SASPs of P. penetrans, and to
compare them to those of Bacillus spp. and Clostridium spp.
Studies have shown that endospores of P. penetrans are resistant to high
temperature and desiccation (Stirling, 1984), some nematicides (Freitas, 1997), and
microwave oven treatment (Weibelzahl-Fulton, 1996). However, the factors that trigger
resistance of endospores to those conditions are yet to be studied. Since SASPs have been
implicated, as a source of amino acid during spore germination in B. subtilis, they may
serve a similar role in Pasteuria. Thus they may be key nutrients that trigger germination
and allow subsequent vegetative growth. It also will be desirable to determine sequences
of the genes encoding SASPs of Pasteuria to compare isolates and species using specific
probes.

CHAPTER 5
SUMMARY
Plant-parasitic nematodes reduce crop yields by approximately $8 billion a year to
producers in the United States and nearly $78 billion worldwide (Society of
Nematologists. Committee on National Needs and Priorities in Nematology, 1994).
Plant-parasitic nematodes are controlled mainly by crop rotation and chemical
nematicides. Rational schemes are oftentimes very difficult to achieve and environmental
concerns plus high cost of using nematicides limit their application. The impending ban
on methyl bromide has forced the loss of one of the most effective chemical nematicides
in use for the past decade.
Pasteuria penetrans (Thome) Sayre & Starr is one of the most promising
biological agents to replace methyl bromide for the control Meloidogyne spp. (Chen and
Dickson, 1998; Dickson et al., 1994; Trudgill et al., 2000), and has the potential to be
used as part of a management program to control root-knot nematodes (Freitas, 1997;
Stapleton and Heald, 1991). This bacterium has been reported in root-knot nematode-
suppressive soils (Dickson et al., 1994), and also has suppressed root-knot nematodes in
greenhouse tests (Brown and Smart, 1985; De Leij et al., 1992; Stirling 1984) as well as
in field microplots (Brown et al., 1985, Chen et al., 1997c; Dube and Smart, 1987;
Oostendorp et al., 1991; Stirling, 1984; Tzortzakakis and Gowen, 1994; Trudgill et al.,
117

118
2000). Pasteuria-infected nematodes have significantly reduced egg production
capability (Bird, 1986; Bird and Brisbane, 1988).
Pasteuria penetrans completes its life cycle within the pseudocoelom of its
nematode host, and unknown signals trigger its germination, growth, and sporulation.
The stages of development of P. penetrans include: i) recognition and attachment; ii)
germination/infection/; iii) vegetative growth; iv) sporulation; and v) release. To be used
successfully as a biological control agent, endospores of P. penetrans must recognize
and bind to molecules on the surface of the nematode-host cuticle. Protein extracts from
P. penetrans spores have been shown to react with wheat-germ agglutinin, indicating the
presence of a carbohydrate ligand on the surface of the spores. These results have
provided the basis for a model in which glycoproteins bearing (3-l-4-linked N-acetyl
glucosamine residues on the surface of the spores, designated as spores adhesins, are
recognized by lectins on the cuticle of the nematode (Bird et al., 1989; Chamecki, 1997;
Chamecki et ah, 1998; Davis and Danks, 1993). Bird et ah (1989) and Chamecki (1997)
showed that wheat-germ agglutinin also inhibited endospore attachment to the nematode
host.
Previous studies using anti-P-20 IgM monoclonal antibody (MAb),
which was selected directly against whole endospores of a P. penetrans strain from
Meloidogyne arenaria race 1 (designated as P-20) demonstrated that this monoclonal
antibody blocked the attachment of P-20 to M. arenaria race 1 (Chamecki, 1997;
Chamecki et ah, 1998). The MAb also recognized an epitope that was shared among
several polypeptides. In my research, the anti-P-20 IgM MAb was used to characterize an

119
adhesin-related epitope, as well as to determine whether other species and strains of
Pasteuria share the same epitope.
The anti-P-20 MAb was used to follow the appearance of an adhesin-associated
protein during the development of P. penetrans within the pseudocoelom of Meloidogyne
arenaria (Neal) Chitwood race 1. Tomato, cv. Rutgers, that were inoculated with either
second-stage juveniles alone or with endospore-attached second stage juveniles of M.
arenaria race 1. Nematodes, uninfected or infected, were harvested at 12, 16, 24, and 38
days after inoculation (DAI), and were examined to determine the developmental stage of
the bacterium at each window of development. Vegetative growth of P. penetrans was
observed only in infected nematodes harvested at 12 and 16 DAI, whereas cells at
different stages of sporulation and mature endospores were observed at 24 and 38 DAI.
Levels of the adhesin-associated proteins were detected by ELISA using the anti-P-20
IgM MAb as the primary antibody. Proteins extracted from uninfected and Pasteuria-
infected nematodes were resolved by SDS-PAGE, electroblotted, and visualized by Auro
Dye or anti-P-20 IgM MAb. Only proteins extracted from infected nematodes harvested
at 24 and 38 DAI were recognized by the MAb. ELISA and immunoblot revealed that the
amount of the adhesin-associated epitope increased as P. penetrans proceeded through
sporogenesis. These results indicate that the synthesis of adhesin-related proteins
occurred at a certain developmental stage relative to the sporulation process, and are
present after mature spores are formed.
Immunocytochemistry techniques were used to investigate where and when the
adhesin-associated proteins were formed. Labeling was first observed in stage III of

120
sporogenesis. Gold particles were clearly observed over the parasporal fibers. Labeling
was observed over other structures such as sporangium and exosporium during the
sporogenesis process, but labeling was not observed on the cortex, epicortex, inner-spore
coat, outer-spore coat, and protoplasm. Immunogold labeling showed that the adhesin-
related epitope, which is recognized by anti P-20 IgM MAb on P-20 is equally distributed
over endospores of other species and isolates of Pasteuria, including Pl-UFLA, North
American Pasteuria, S-l, C-1, LS-1, L-l, and Rhabditis-iniecting Pasteuria. The
distribution of the adhesins over several structures of the endospores of the different
isolates and species of Pasteuria indicates that this epitope does not confer host
preference or specificity. Since the epitope has been implicated in recognition and
attachment (Chamecki et al., 1998) its broad distribution on Pasteuria spp. with different
host specificity indicates it is only one component of the attachment process. Other
forces associated with surface properties (Afolabi et al., 1995; Kamra and Dhawan, 1998;
Mohan et al., 2001) of both endospore and the nematode cuticle may contribute to a
cooperative process resulting in recognition, initial interaction, and finally irreversible
attachment.
Quantitative and qualitative differences were observed in the protein bands from
extracts of Pasteuria sp. NA, Pasteuria sp. S-l, as well as C-l, Pl-UFLA, ring nematode
and spiral nematode isolates of Pasteuria, compared to P-20 used as control. These
results indicate that the epitope that was recognized by the MAb in the P-20 isolate is
shared among other isolates and species of Pasteuria. This epitope, which was not found
associated with any of several Bacillus spp. examined previously, is considered to be a

121
component of a recognition system shared by different isolates and species of Pasteuria.
(J. Harrison and J. F. Preston, unpubl.; Schmidt et al., 2001) Indirect
immunofluorescence was used to visualize the distribution of the adhesin-associated
epitope on the surface of whole mature endospores. It revealed that the antigen bearing
the epitope does not occur uniformly on the surface of mature endospores of P-20 strain
of P. penetrans.
This study is the first to report a temporal synthesis of an adhesin-associated
epitope during the sporogenesis process of P. penetrans and that this epitope is also
synthesized by other isolates and species of Pasteuria.

APPENDIX A
EXTRACTION OF SMALL, ACID SOLUBLE SPORE PROTEINS FROM SPORES
(Johnson and Tipper, 1981)
1. Add 10 mg of dry and clean spores of B. subtilis (10 mg/ml) or 10 p\ of an endospore
suspension of P. penetrans (1><108//ul) to a 1.5 ml siliconized microtube containing 1 ml 2N HC1.
2. Vortex the suspension until it is homogenous.
3. Incubate at 20 °C for 30 minutes.
4. Centrifuge the suspension at 10,000 x g for 10 minutes.
5. Collect the supernatant (SI) in a microtube, and suspend the pellet in 1 ml 2N HC1.
6. Vortex the pellet until it is homogenous.
7. Incubate at 20 °C for 30 minutes.
8. Centrifuge as above.
9. Collect the supernatant (S2) in a microtube.
10. Combine the supernatants (SI plus S2) and bring it to pH = 5 (use 10N NaOH).
11. Storage at-20 °C.
122

APPENDIX B
ISOLATION OF IgY ANTIBODY FROM CHICKEN EGG YOLKS
(Modified from Poison et al., 1985)
1. Separate the egg white from the yolk, keeping the yolk.
2. Carefully rinse the egg yolk with 0.1 M NaP04, pH 7.6, and then place it in a vessel suitable
for stirring.
3. Add 4 volumes of 0.1 M P04 buffer, pH 7.6 and stir for 30 minutes.
4. At this point the extract can be screened for antibody content.
5. The antibody of interest can be further isolated by using the procedure by Poison et al.,
1985.
6. Add polyethylene glycol (PEG) (P-2139, Sigma, St. Louis, MO) at 0.035g/ml extract
while stirring at room temperature until the PEG is dissolved.
7. Centrifuge at 5000g for 20 minutes at 20 °C.
8. Remove the middle phase, taking care to disrupt the liquid phase as little as possible, and
record the volume.
9. Add PEG at 0.085g/ml extract with slow stirring until dissolved, then let stand for 60 minutes.
10. Centrifuge as above for 25 minutes.
11. Decant the supernatant and resuspend the pellet in 0.1 M P04 buffer, pH 7.6, recording
the volume.
123

124
12. Slowly add PEG at 0.12 g/ml extract with slow stirring. About 10 minutes after
addition of the last of the PEG, centrifuge as per step 10.
13. Dissolve the pellet in 0.1M P04 buffer, pH 7.6, cooled at 4 °C, then add an equal
volume of 50% ethanol (Fisher Scientific, Suwanee, GA), precooled at -20 °C.
14. Stir on ice for 15 minutes, then centrifuge at 10,000 x g for 25 minutes at 5 °C.
15. Dissolve the pellet in 0.1 M P04 buffer, pH 7.6, and dialyze (10000 MWCO tubing,
Pierce Company, Rockford, IL) against 2 to 4 litters of the 0.1M P04 buffer, pH 7.6 in
cold room overnight.
16. Record the volume after dialysis, and add 1% Na N3 for preserving.
17. The extract may be stored indefinitely at 4 °C. Do not freeze it.
18. Lastly, a precipitate may form during storage. As long as the precipitate is not
disturbed, there is apparently no effect on the titer of the preparation.

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225.

BIOGRAPHICAL SKETCH
Janete Andrade Brito was bom on July 25, 1957, to Mrs. Alaide Andrade de Brito
and Mr. Urbano Pinheiro de Brito, Jequié, Bahia, Brazil. She began her undergraduate
program in the University Federal of Bahia, Salvador, Bahia, Brazil, and received a
bachelor’s degree in Agronomy in January, 1981. She did her postbaccalaureate in Plant
Pathology at The Brazilian National Research Institute (EMBRAPA), Cruz das Almas,
Bahia, Brazil, April 1981 to March 1982. In March of 1982, she entered the Graduate
School in the Department of Pathology, Universidade Federal de Vifosa, and obtained a
Master of Science degree in December, 1985. In the next seven and a half years she was
employed by the Funda?áo Instituto Agronómico do Paraná, Paraná, Brazil. In August
1996, she continued her graduate education under the supervision of Drs. Robin M.
Giblin-Davis and James F. Preston. Her research project emphasized the biochemical
aspects of the development of Pasteuria penetrans.
148

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 Phjl
Robin M. Giblin-Davis, 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.
/
J>tfhes F. Preston, Cochair
Professor of Microbiology and Cell Science
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.
Donald W. Dickson
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.
fwAlAyl 0.
Henry C. Aldrich
Professor of Microbiology and Cell Science
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.
Grover C. Smart, Jr
Professor of Entomology and Nematology

This dissertation was submitted to the Graduate Faculty of the College of
Agricultural and Life Sciences and to the Graduate School and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.
May, 2002
Dean, College of Agricul
Sciences
Dean, Graduate School

LD
1780
20 £2_
UNIVERSITY OF FLORIDA
3 1262 08555 3690