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.
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.
TABLE OF CONTENTS
LIST OF ABBREVIATIONS ......................................... viii
LIST OF TABLES................................................... x
LIST OF FIGURES.................................................. xi
ABSTRACT ...................................................... xiv
1. INTRODUCTION ................................................. 1
Host: Root-Knot Nematodes (Meloidogyne spp.) ........................1I
Life Cycle............................................... 2
Distribution and Economic Importance......................... 5
Parasite: Pasteuria pen etrans..................................... 15
Historical Background .................................... 15
The Genus Pasteuria ..................................... 18
Members of Pasteuria ................................... 18
Systematic and Phylogeny of Pasteuria........................ 21
Biological Control Potential................................ 24
The Effect of Other Microorganisms and Pesticides on Pasteuria ..... 26 Life Cycle.............................................. 28
Host Specificity ......................................... 33
Interaction: Host-Parasite........................................ 35
The Role of Adhesin Proteins in the Relationship Host-Parasite ...... 35
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
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
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
LIST OF ABBREVIATIONS ELISA Enzyme linked immunosorbent assay
BSA Bovine serum albumin
FITC Fluorescein isothiocyanate
KLH Keyhole Limpet Hemocyanin
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
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
LIST OF TABLES
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
LIST OF FIGURES
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
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
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
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
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.
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
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).
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
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
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).
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,
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).
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
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
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
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
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
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
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-
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
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,
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
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
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
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,
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
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.
(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
(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
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,
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
Desulfotomaculum 3 8-52
Sporohalobacter 3 0-32
Sporolactobacillus 3 8-40
Source: Berkeley and Ali, 1994.
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.
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.,
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
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
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
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
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).
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
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).
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.
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,
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
(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.
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).
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
(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.
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
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.
SYNTHESIS AND IMMUNOLOCALIZATION OF AN ADHESIN-ASSOCIATED EPITOPE IN Pasteuria penetrans
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
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.
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
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.
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
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
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-
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.
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)
(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.
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
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.
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
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
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.
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,
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.
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.
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).
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
2 1 2... .1 6....2 4....3 8.
12 16 24 38
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.
1 2 3 4 5 6
1 2 3 4 5 6
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.
1 2 3 4 5 6
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 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.
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.
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).
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.
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.
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.
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).
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.
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
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
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
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.
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).
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
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-
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
(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
Table 3.1. Species and isolates of Pasteuria. Species or isolates Reference
P. penetrans P-20 Meloidogyne. arenaria race 1 (Oostendorp et al.,
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.)
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;
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
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
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.
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;
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%
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.
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.
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).
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.,