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SYSTEMATICS OF THE ENTOMOPATHOGENIC BACTERIA BACILLUS POPILLIAE, BACILLUS LENTIMORBUS, AND BACILLUS SPHAERICUS

Karen Rippere Lampe

Dissertation submitted to the Faculty of the Virginia Polytechnic Institute and State University in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY in Biology

Allan A. Yousten, Chair Noel R. Krieg Khidir W. Hilu Eric A. Wong David L. Popham

September 11, 1998 Blacksburg, Virginia

Keywords:

Bacillus popilliae, Bacillus lentimorbus,

Bacillus sphaericus, DNA reassociation, RAPD, vancomycin resistance

SYSTEMATICS OF THE ENTOMOPATHOGENIC BACTERIA BACILLUS POPILLIAE, BACILLUS LENTIMORBUS, AND BACILLUS SPHAERICUS

Karen Rippere Lampe A. A. Yousten, Chairman Department of Biology (ABSTRACT)

Bacillus popilliae and B. lentimorbus, causative agents of milky disease in Japanese beetles and related scarab larvae, have been differentiated based upon a small number of phenotypic characteristics, but they have not previously been examined at the molecular level. Thirtyfour isolates of these bacteria were examined for DNA similarity.

Three distinct but related similarity groups

were identified; the first contained strains of B. popilliae, the second contained strains of B. lentimorbus, and the third contained two strains distinct from but related to B. popilliae.

Some strains received as B.

popilliae were found to be most closely related to B. lentimorbus and some received as B. lentimorbus were found to be most closely related to B. popilliae.

Geographically distinct strains of B. popilliae and B. lentimorbus were analyzed using RAPD.

Eight decamer

primers were tested against nineteen new and seventeen isolates previously described by randomly amplified polymorphic DNA (RAPD) analysis (M. Tran).

Of the new

isolates, ten were found to be B. popilliae while nine

isolates were more related to the B. lentimorbus species. Paraspore formation, believed to be a characteristic unique to B. popilliae, was found to occur among a subgroup of B. lentimorbus strains.

Using a combination of two PCR primer pairs, the cry18Aa1 gene was detected in 31 of 35 B. popilliae isolates and in 1 of 18 B. lentimorbus isolates.

When

hemolymph smears were examined microscopically, a parasporal crystal was seen in three of the four B. popilliae strains where the PCR primers could not amplify the paraspore gene.

The fourth strain was not tested due

to the unavailability of infected hemolymph.

A paraspore

was also detected by microscopic examination in a subgroup of 14 B. lentimorbus strains.

In combination, the primer

pairs CryBp1 and CryBp2 are effective at detecting the paraspore gene in B. popilliae isolates, but not in the B. lentimorbus isolates.

Growth in media supplemented with 2%

NaCl was found to be less reliable in distinguishing the species than was vancomycin resistance, the latter present only in B. popilliae.

The basis for vancomycin resistance in all isolates was investigated using a polymerase chain reaction assay designed to amplify the vanB gene in enterococci. amplicon was identified and sequenced.

An

The amplified

portion of the putative ligase gene in B. popilliae had 77% and 68-69% nucleotide identity to the sequences of the vanA gene and the vanB genes, respectively.

There was 75% and

69-70% identity between the deduced amino acid sequence of the putative ligase gene in B. popilliae and the deduced amino acid sequence of the vanA gene and the vanB genes,

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respectively. It has been determined that the vanE gene is located either on a plasmid greater than 16 kb in size or on the chromosome.

The gene in B. popilliae may have had

an ancestral gene in common with vancomycin resistance genes in enterococci. Bacillus sphaericus strains isolated on the basis of pathogenicity for mosquito larvae and strains isolated on the basis of a reaction with a B. sphaericus DNA homology group IIA 16S rRNA probe were analyzed for DNA similarity. All of the pathogens belonged to homology group IIA, but this group also contained nonpathogens.

It appears

inappropriate to designate this homology group a species based solely upon pathogenicity.

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ACKNOWLEDGEMENTS First and foremost, I would like to thank my advisor Dr. Allan Yousten for his guidance and wisdom throughout this project.

Without his input, I would never have made

it though this program.

I would also like to thank my

committee members, Dr. N. R. Krieg, Dr. K. W. Hilu, Dr. E. Wong and Dr. D. L. Popham for their help, advice and support.

The late Dr. John L. Johnson enabled me to get

started on this project and we miss him very much.

To my

family, thanks for believing in me and supporting me both emotionally and financially.

Finally, to all the other

graduate students on the hall; without you it wouldn't have been as much fun.

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TABLE OF CONTENTS

Page ABSTRACT......................................................ii ACKNOWLEDGEMENTS...............................................v LIST OF FIGURES...............................................ix LIST OF TABLES............................................... xi INTRODUCTION...................................................1 I.

REVIEW OF THE LITERATURE Use of Bacillus popilliae and Bacillus lentimorbus as biological control agents............................4 Pathology of Bacillus popilliae and Bacillus lentimorbus..........................................6 Physiology of Bacillus popilliae and Bacillus lentimorbus..........................................7 Genetics of Bacillus popilliae.........................10 Taxonomy of Bacillus popilliae and Bacillus lentimorbus.........................................12 DNA-DNA similarities...................................15 RAPD analysis..........................................16 Vancomycin resistance..................................18 References.............................................24

II.

MATERIALS AND METHODS Media and Reagents.....................................33 Bacterial strains and growth conditions................35 Isolation of bacteria from dried beetle hemolymph......38 DNA isolation for DNA-DNA reassociation................38 DNA sample preparation.................................39 DNA labeling...........................................40 S1 Nuclease assay......................................42 DNA isolation for RAPD experiments.....................43

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Determination of DNA concentration....................44 RAPD analysis.........................................45 Isolation, amplification and digoxygenin labeling of individual RAPD bands..............................47 Estimation of probe yield.............................49 Southern transfer and hybridization...................50 RAPD band analysis....................................52 Data analysis.........................................52 Multiplex PCR-RFLP for detection of the van ligase....53 Paraspore gene detection using PCR....................54 PCR product sequencing................................55 Labeling of the vanE PCR product......................56 Determination of vanE location in B. popilliae........56 References............................................56 III.

BACILLUS POPILLIAE AND BACILLUS LENTIMORBUS, BACTERIA

CAUSING

MILKY DISEASE IN JAPANESE BEETLES AND RELATED SCARAB

LARVAE Abstract..............................................58 Results DNA similarity.....................................60 Growth in 2% NaCl or vancomycin....................63 Discussion............................................63 References............................................65 IV.

RANDOMLY AMPLIFIED POLYMORPHIC DNA ANALYSIS OF

GEOGRAPHICALLY DISTINCT ISOLATES OF BACILLUS POPILLIAE AND BACILLUS LENTIMORBUS Abstract..............................................68 Results RAPD analysis......................................68 Growth in 2% NaCl or vancomycin....................75 Discussion............................................78 References............................................80

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V.

IDENTIFICATION AND DETECTION OF THE CRY GENE IN STRAINS OF

BACILLUS POPILLIAE AND BACILLUS LENTIMORBUS Abstract..............................................82 Results Detection of the cry operon.......................82 Discussion............................................90 References............................................92 VI.

DNA SEQUENCE RESEMBLING VANA AND VANB IN THE VANCOMYCIN-

RESISTANT BIOPESTICIDE BACILLUS POPILLIAE Abstract.............................................93 Results..............................................95 Discussion..........................................101 References..........................................102 VII.

DNA SIMILARITIES AMONG MOSQUITO-PATHOGENIC AND

NONPATHOGENIC STRAINS OF BACILLUS SPHAERICUS Abstract............................................105 Bacteria and DNA isolation..........................106 DNA similarities....................................107 Results and Discussion..............................108 References..........................................109 SUMMARY.....................................................111 CONCLUSIONS.................................................114 CURRICULUM VITAE............................................116

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LIST OF FIGURES Page CHAPTER THREE Figure 1. Distance dendogram of B. popilliae and B. lentimorbus strains generated from DNA similarity analysis.......................................62 CHAPTER FOUR Figure 1. RAPD banding patterns of B. popilliae and B. lentimorbus isolates using primer OPA-03.........................................70 Figure 2. RAPD banding patterns of B. popilliae and B. lentimorbus isolates using primer OPA-03.........................................71 Figure 3. RAPD banding patterns of B. popilliae and B. lentimorbus isolates using primer OPA-15........................................72 Figure 4. RAPD banding patterns of B. popilliae and B. lentimorbus isolates using primer OPA-15........................................73 Figure 5. Dendogram showing the relationships between strains of B. popilliae and B. lentimorbus generated from RAPD analysis...................74 CHAPTER FIVE Figure 1. Structure of the Bacillus popilliae cry operon.....................................83 Figure 2. ATCC 14706 and NRRL B-4081 PCR products using primer pair CryBp2......................86 Figure 3. B. popilliae cry18Aa1 gene sequences......87 Figure 4. Deduced amino acid sequence comparison of B. popilliae cry genes.........................89 CHAPTER SIX Figure 1. Multiplex PCR-RFLP of enterococcal isolates carrying the vanA and vanB ligase genes and B. popilliae ATCC 14706........................96

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Figure 2. Sequence comparisons of the putative ligase genes in B. popilliae isolates.........97 Figure 3. Comparison of the translation of the putative ligase gene in B. popilliae ATCC 14706 to the translations of four previously characterized vanB genes(isolates 55, 94, 45, and 91) and one vanA gene (isolate)..........98 Figure 4. Southern blot of digested and undigested B. popilliae chromosomal DNA probed with the vanE PCR product....................100

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LIST OF TABLES Page CHAPTER TWO Table 1. B. popilliae and B. lentimorbus strains used in DNA-DNA reassociation.................36 Table 2. B. popilliae and B. lentimorbus strains used in RAPD analysis from diverse host insects and geographic regions.........................36 Table 3. RAPD primer sequences......................45 Table 4. Dilution series for probe estimation.......49 Table 5. Primer sequences used in multiplex PCR-RFLP reaction for detection of van ligase genes in enterococci....................................53 Table 6. Primer sequences used for detection of cry genes in B. popilliae and B. lentimorbus.......55 CHAPTER THREE Table 1. Levels of DNA similarity between B. popilliae and B. lentimorbus as determined by the S1 nuclease method.........................61 Table 2. Characteristics of B. popilliae and B. lentimorbus strains used in DNA similarity studies........................................63 CHAPTER FOUR Table 1. Characteristics of B. popilliae and B. lentimorbus isolates from diverse host insects and geographical regions...........................77 CHAPTER FIVE Table 1. Detection of the paraspore crystal in strains of B. popilliae and B. lentimorbus by visualization and PCR..........................84 CHAPTER SEVEN Table 1.

Bacillus sphaericus strains studied using

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DNA-DNA similarity analysis..................107

Table 2. Levels of DNA similarity among strains of B. sphaericus................................109

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INTRODUCTION

Classification schemes illustrating the relationships among living organisms have been documented since the writings of Aristotle, and the classification of bacteria since the different morphological forms were described by Muller in 1773. Bacterial classification began by organization into taxonomic units based solely on the morphological characteristics of the organisms and has progressed to the use of a wide variety of characteristics including the physiology, biochemistry and genetic material of the bacteria.

Today, the sheer number of

bacterial species that have been identified and the wide diversity among them make classification of these organisms into discreet arrangements both difficult and necessary.

Classification can be described as having three major purposes.

The arrangement of organisms into discrete groups

provides a way to summarize and catalog information about them. The classification takes the form of a database in which information about an organism can be stored and retrieved by the use of a particular name.

The classification can be used to

predict the properties of a group of organisms so that members may be recognized by their defining characteristics.

The

organization of organisms into groups by classifying them must be accomplished before an identification system can be created which will recognize new isolates.

Finally, classification

systems can provide insights into the evolutionary origins and relationships among organisms.

To fulfill these purposes,

classifications should contain as much information as possible, be stable and should be based on empirical evidence.

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The species concept is less rigorously defined for bacteria than for other organisms.

Bergey's Manual defines the bacterial

species as "a collection of strains that share many features in common and differ considerably from other strains."

It goes on

to say that "a species consists of the type strain and all other strains that are considered to be sufficiently similar to it as to warrant inclusion with it in the species."

A more uniform

definition of the bacterial species is desireable and can possibly be obtained through the use of genetic relatedness among bacteria.

Microbiologists typically use two different types of classifications, phenetic classifications and special purpose classifications.

Phylogenetic classifications are beginning to

be developed with the information provided by macromolecule sequencing but have only been applied to select bacterial groups.

Phenetic classifications encompass all bacteria and are

useful to all microbiologists, regardless of their specific discipline.

They are organized using affinities based on the

phenotype and genotype of organisms as they exist in the present, with no regard for evolutionary context.

Special

purpose classifications are designed for a particular discipline.

These systems are often based on a single feature

which is thought to be sufficient and necessary for the placement of an organism within a group.

A disadvantage of

these systems is that they are based on very little information and therefore tend to be unstable.

Due to the lack of

information, an unknown organism that is lacking the single essential feature of the classification would be assigned to the wrong taxon.

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Classification of bacteria allows microbiologists to associate certain characteristics with groups of bacteria.

This

ability to define discrete groups allows for identification of new isolates and the rapid association of certain properties to them.

In addition, classification of bacteria into orderly

groups eliminates confusion that could be caused by the large numbers of bacterial species and the diversity among them.

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CHAPTER ONE Review of the Literature

Use of Bacillus popilliae and Bacillus lentimorbus as biological control agents. Bacillus popilliae and B. lentimorbus are the causative agents of types A and B (respectively) milky disease, a fatal infection of Japanese beetle larvae as well as other members of the family Scarabaeidae (35).

Japanese beetles and other

scarabaeids feed on more than 257 different plants and cause economic loses through damage to turfgrasses and crops, making their control important to various industries (86).

Biological

control of these pests using B. popilliae may be easier, less expensive and ecologically safer than use of synthetic chemicals (41).

The bacteria are also very specific, targeting only the

insect of choice while leaving beneficial insects unharmed (72).

Bacillus popilliae has been used as a biopesticide since 1937 when Dutky artificially added diseased larvae to field plots (32).

He successfully established the disease in one

location and showed that B. popilliae populations built up and spread in the field.

Due to the inability to produce spores in

vitro, a process involving the injection of spores into healthy larvae was developed by White and Dutky in order to mass produce milky disease spores (101).

A standardized spore powder was

developed and used to establish B. popilliae at new field sites (33).

Establishment of milky disease in the field appears to be

dependent on achievement of larval densities between 180 and 480 larvae per square meter (10).

Milky disease organisms may be

spread in the environment by birds, insects, skunks, moles, and mice (100).

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Other methods of producing milky disease spores have been explored, including the use of tissue culture, in vitro culture and sporulation and the use of vegetative cells as disease agents.

Limited sporulation of B. popilliae has been achieved

in vitro using both solid media and chemostat cultures (19, 81). Spores of B. popilliae produced in these ways are less infective than spores produced in the larvae (48).

Bacillus popilliae

spores germinate poorly, requiring injection of 105-107 spores into larvae to cause 50-80% infection.

In contrast, injection

of 102-103 viable vegetative cells causes comparable infection rates in Japanese beetle larvae (88).

Splittstoesser et al.

(85) reported that germination and outgrowth of B. popilliae spores in cabbage looper hemolymph reached 90% in one hour.

The

spores had to be heated at 37oC under alkaline conditions with the addition of tyrosine in order to achieve such rates of outgrowth (85).

Sharpe et al. (82) developed a microscope slide

culture system used to track the germination and outgrowth of B. popilliae B-2309 spores.

They found that the vegetative cells

emerged in 23-24 hours and 5% of total spores showed outgrowth after 48 hours.

However, only 1% of the spores produced visible

colonies on a plate, indicating that 80% of germinating spores fail to develop visible colonies.

Sharpe (82) suggested that

the low rate of germination and outgrowth in vitro may indicate the reason for a low infectivity rate in vivo.

In tissue

culture consisting of hemocytes of Phyllophaga anxia, Luthy (53) reported growth and sporulation of B. popilliae var. melolonthae and growth without sporulation of B. popilliae var. popilliae. Lyophilized vegetative cells pelleted using tung oil polymer coated with paraffin have been shown to have 93% infectivity when injected into larvae (48).

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Milky disease bacteria have been reported to persist in the environment for extended periods of time, often longer than twenty-five years, negating the need for reapplication of spore powders to field plots (48).

Resistance of the insects to B.

popilliae has not been shown to occur in these areas. Investigation of

the taxonomy of B. popilliae and related

species will assist in the development of this element in an integrated approach to pest management.

Pathology of Bacillus popilliae and B. lentimorbus Bacillus popilliae spores are ingested by the beetle larvae during feeding, and once ingested, enter the larval midgut where the spores germinate.

The vegetative cells proliferate and

enter the hemocoel where they continue to multiply. disease can be said to occur in four stages.

Milky

An initial

incubation stage (2 days) where few bacterial cells are found in the hemolymph is followed by rapid proliferation of vegetative cells (day 3 to day 5).

Stage three is characterized by a

change from predominantly vegetative growth to sporulation (days 5-10).

Stage four is a sporulation phase terminating in the

death of the larvae (day 14 to day 21) (19, 86). Infections caused by B. popilliae var. melolonthae do not follow this pattern, instead increase in vegetative cell numbers and sporulation occur simultaneously (48).

Eventually, the number of spores in the hemolymph reaches numbers as high as 5 × 1010 per milliliter of hemolymph.

The

normally clear insect hemolymph becomes turbid, leading to the name “milky disease”.

The B. popilliae spores are released into

the soil from the larval cadaver, thus beginning the process again.

This accounts for the extended persistence of B.

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popilliae in the environment.

In larvae infected by B.

lentimorbus for extended periods of time, there is a build up of blood clots, causing the hemolymph to look brownish in color instead of milky white (19).

Physiology of Bacillus popilliae and B. lentimorbus Gordon, Haynes and Pang (39) provided phenotypic information on 12 strains of B. popilliae and 5 strains of B. lentimorbus.

They reported the vegetative cells to be gram

negative and the prespores and sporangia to be gram positive. Dutky originally reported the vegetative cells to be gram positive (35).

When examined by electron microscopy the cells

exhibit a gram positive cell wall structure (15, 16).

Gordon et

al. (39) reported that B. popilliae was motile by peritrichous flagella while all the strains of B. lentimorbus tested were nonmotile.

Splittstoesser (85) also reported that B. popilliae

cells were extremely motile upon germination and outgrowth in hemolymph slide mounts.

Bacillus popilliae and B. lentimorbus are nutritionally fastidious and only grow well on a rich medium containing yeast extract and digests of casein (19, 78).

Cells reach stationary

phase after 16–20 hours of growth and the maximum number of viable cells at this time is about 6 × 108 for B. lentimorbus and 1.2 × 109 for B. popilliae (86).

After the cultures reach

stationary phase there is a rapid decrease in viability.

The

cause of cell death is not fully understood, but both organisms lack the enzymes peroxidase and catalase, leaving them sensitive to hydrogen peroxide damage (67). It has been hypothesized that the lack of these enzymes may play a role in culture death (26,

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67, 92). Pepper et al. (67) tested for oxygen evolution when hydrogen peroxide was added to a Warburg flask containing B. popilliae cells and were unable to detect any evolution of oxygen.

They also tested for the breakdown of hydrogen peroxide

by B. popilliae by iodometric titration and were unable to detect any breakdown of peroxide.

Bacillus popilliae was

examined for the presence of peroxidase and it was found that while cell extracts rapidly oxidized NADH2, the rate was not enhanced by the addition of hydrogen peroxide (67). St. Julian et al. (86) suggested that hydrogen peroxide toxicity is not the cause of death because its build up in vegetative cells is slight.

They also state that death caused by exposure to the

superoxide radical is unlikely because of the high levels of superoxide dismutase found in B. popilliae cells (86).

Thiamine and tryptophan have been found to be essential nutrients for B. popilliae, while biotin, myoinositol and niacin are stimulatory for growth (86, 90).

Many of the amino acids

must be supplied to B. popilliae and B. lentimorbus in some form, including any amino acids in the serine or aromatic families (93).

Bacillus popilliae metabolizes sugars including

glucose, fructose, mannose, galactose, maltose, sucrose and trehalose, the latter sugar found in the larval hemolymph (19). Products formed by glucose catabolism are lactic acid, acetic acid and carbon dioxide (68).

The decrease in culture medium pH

has a slight effect on the viability of the culture once it reaches stationary phase.

When the culture medium was

appropriately buffered, the amount of growth increased and the survival of the cells was slightly enhanced.

The Embden-

Meyerhof-Parnas pathway and the pentose phosphate pathways are the preferred routes of carbohydrate catabolism in B. popilliae and B. lentimorbus (68, 87).

The EMP pathway is the major route

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of glucose assimilation while the PP pathway functions mostly for formation of biosynthetic intermediates (18).

Pepper et al.

(68) found no evidence for the presence of either the EntnerDudoroff or the phosphoketolase pathways in B. popilliae.

Using

inhibitors specific for the enzymes glyceraldehyde-3-phosphate dehydrogenase and enolase they found that they could inhibit glucose oxidation by 100%.

This provided preliminary evidence

for the lack of the ED and phosphoketolase pathways and further enzyme assays showed no KDPG (2-keto-3-deoxy-6-P-gluconate) aldolase or phosphoketolase activity (68).

The enzymes that

breakdown trehalose are expressed constitutively and both respiration and growth rates are higher when the bacteria are grown with trehalose than with glucose.

Trehalose is

transported into the cell by the PEP phosphotransferase system and the trehalose-6-phosphate is cleaved by a phosphotrehalase into glucose and glucose-6-phosphate (12).

B. popilliae lacks a complete tricarboxylic acid cycle, suggested by some to be the cause of the poor sporulation in vitro (18, 68).

McKay et al. (56) were unable to detect α-

ketoglutarate dehydrogenase activity in B. popilliae strain NRRL B-2309 and its derivatives.

St. Julian et al. (86) suggested

that lack of sporulation in vitro is caused by a decrease in protein synthesis and lipid metabolism once the cells reach the stationary phase of growth.

B. popilliae and B. lentimorbus do

contain cytochromes and are capable of oxygen dependent growth (86).

These characteristics make it difficult to grow and maintain the bacteria in the laboratory.

In addition, strains

such as RM9 are unable to be grown in the laboratory and can be

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maintained only in the insects themselves.

This makes it

difficult to rapidly identify and classify which species of bacteria are present in natural populations in any given area (84).

Strains of B. popilliae were shown to vary in virulence

and for some strains the virulence and host preference could be modified by repeated passage through the insect host (19). Strains have also been noted to vary widely in their growth characteristics in vitro.

Milky disease infections may exhibit

some degree of insect host specificity.

Bacillus popilliae var.

melolonthae was isolated only from the common cockchafer (Melolontha melolontha).

Two distinct B. popilliae isolates

were identified in New Zealand Costelytra zealandica populations (37, 38).

An atypical strain of B. popilliae has been reported

to be associated with the northern masked chafer (Cyclocephala borealis) (34).

The spores from the diseased larvae had an

unusually large paraspore and virtually no cross-infectivity was found between spores from C. borealis and Japanese beetles (Popillia japonica) (19).

Klein (48) stated that this lack of

cross-infectivity stressed the need for commercial spore preparations intended for use against the Japanese beetle to be produced in Japanese beetle larvae.

Milner (60) also found a

lack of cross-infectivity for an isolate he called B. popilliae var. rhopaea.

He showed that this isolate had virtually no

ability to infect Rhopaea morbillosa and Othnonius batesi grubs in Australia but could effectively infect Rhopaea verreauxi larvae.

Due to the possible lack of cross infectivity, it is

necessary to be able to properly identify which species is needed to control a population of insects so that effective control of the insect is achieved.

An understanding of the

classification of these bacteria could lead to a means of distinguishing varieties with specific host infectivity ranges.

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Genetics of B. popilliae Very little is known about the genetics of B. popilliae and B. lentimorbus. and number.

Both species contain plasmids of various sizes

Valyasevi and Kyle (96) reported that an isolate of

B. popilliae collected from infected larvae in New York contained three plasmids denoted pBP149, pBP082 and pBP043. Plasmid pPB149 showed no homology to pBP082.

pBP149 was

estimated to be 12 kb, pBP082 was 7.4 kb, and pBP043 was 4.9 kb in size (96).

Dingman (29) performed a study on interrelated

plasmids in B. popilliae strain KLN4 and B. lentimorbus strain NRRL B-2522, also finding that these isolates contained three plasmids.

However, these plasmids differed from those found by

Valasevi, at 6.8 kb, 8.8 kb and 9.4 kb in size.

These three

plasmids were named pBP68, pBP88 and pBP94, respectively ((29). All three plasmids showed contiguous regions of similarity to each other as tested by hybridization of segments of each plasmid to the others, indicating that they are an interrelated family of plasmids.

A plasmid designated pBP614 has been

characterized from B. popilliae and found to replicate by the rolling circle mechanism (51).

This plasmid is 5.6 kb in size

and the coding strand of the plasmid is deficient in cytosine (16.1% of the total base composition).

Two open reading frames

were found on this plasmid, one corresponding to the rep gene and the other to a protein of unknown function (51).

Bacillus popilliae and B. lentimorbus have been shown to contain N6-methyladenine in GATC sequences distinguishing them from all other Bacillus species tested for this characteristic (30).

The paraspore gene (cry) has been cloned and sequenced

from B. popilliae strain H1, isolated near Heidelberg, Germany. Two open reading frames of a putative operon were sequenced, the

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first codes for a protein of 175 amino acids while the second has been designated cry18Aa1 and codes for the paraspore protein (706 amino acids) (108).

The name cry18Aa1 is in accordance

with the nomenclature of the Bacillus thuringiensis toxin genes as revised and summarized by (27).

The first open reading

frame, designated orf1, shows significant similarity to orf1 of the cry2Aa-cry2Ac operon, orf1 of the cry9Ca operon and p19 of the cry11Aa operon of B. thuringiensis.

orf1 and cry18Aa1 are

transcribed as an operon and EσE and EσK acting at the same site in the promoter can drive transcription of the operon (109). Cry18Aa1 has significant amino acid similarity to the Cry proteins of B. thuringiensis and hydrophobicity distribution throughout the protein seems to be similar to that found in Cry3A and Cry1A toxins of B. thuringiensis (108).

Zhang et al.

(108) suggested that the strong similarity between the B. popilliae cry gene and the cry genes of B. thuringiensis indicates a possible role of the paraspore protein in the pathogenesis of the milky disease organism.

Taxonomy of B. popilliae and B. lentimorbus Isolated and described by Dutky in 1940, B. popilliae and B. lentimorbus were defined as two separate species based on the presence of a refractile parasporal body in B. popilliae and its absence in B. lentimorbus (35).

In addition, there are

differences in the color of the hemolymph from larvae infected by the two bacteria (18).

The mol% G+C of B. popilliae is

listed by Bergey’s manual as 41.3% while that for B. lentimorbus is 37.7% (23).

Generally, greater than two percent difference

in the mol% G+C is considered to be indicative of speciation (23).

Serological differences between the two species have been

demonstrated, as well as minor differences in the fatty acid 12

composition of the bacteria (47, 50, 52).

Both species readily

form spores in vivo, but sporulate poorly or not at all in vitro.

Spore morphology has been examined by scanning electron microscopy and it was found that the spores of B. popilliae and B. lentimorbus share a common ridged surface (20, 64, 88).

The

most widely used characteristic in differentiating B. popilliae from B. lentimorbus is the parasporal inclusion found in B. popilliae.

This inclusion has been considered to be absent in

B. lentimorbus. However, it has been suggested that the parasporal body is not a stable characteristic and should not be used for species identification (107).

The parasporal body is

formed at the time of sporulation as it is for other insect pathogens, such as Bacillus thuringiensis and Bacillus sphaericus.

In contrast to these latter bacteria, the

parasporal body in B. popilliae has not been conclusively shown to play a role in pathogenesis, although recent evidence suggests that it may have a function similar to the Cry toxins of B. thuringiensis (108).

Weiner

(99) found that solubilized

parasporal protein was capable of killing 58 % of larvae in 48 hours when injected into the grubs.

Intact parasporal

inclusions were able to kill 25 % of larvae. fed orally to the larvae was nontoxic (99).

Parasporal protein Zhang et al. (108)

proposed a role for the B. popilliae parasporal protein in milky disease, suggesting that once the spores germinate in the larval gut the paraspore protein is activated.

Once activated, the

protein binds to the brush border membrane and damages the gut wall in some fashion, allowing the vegetative cells to enter the hemolymph and the disease to progress.

The shape of the

parasporal crystal as well as its size and position in the sporangium differs among strains of B. popilliae (59).

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Gordon

et al. (39) reported that B. popilliae was able to grow in broth supplemented with 2% NaCl while B. lentimorbus was unable to grow under this condition.

This finding has been disputed by

Milner (59), as all of the B. lentimorbus strains used by Gordon (39) were from the same source and the same insect.

They may

not have been representative of the non-paraspore forming strains.

Distinct strains of B. popilliae have been isolated

from several different scarabaeids.

These isolates show very

little cross infectivity between insect species, suggesting a fundamental difference between the isolates (49, 59).

Presently, the major criteria for establishing two species of milky disease bacteria are the presence or absence of a parasporal body in sporulated cells, the physical appearance of infected larvae, and the ability to grow in broth supplemented with 2% NaCl.

Several classification schemes have been suggested for the milky disease bacteria.

After their original isolation and

characterization by Dutky (35), many more strains of milky disease bacteria were isolated from different insect species.

A

strain causing milky disease was isolated from the European cockchafer and named B. melolonthae. A similar strain was isolated in Europe but named B. fribourgensis.

These two

strains were later shown to be identical, but the individual names carried on for some time.

Luthy and Krywienczyc (50, 52)

demonstrated that B. popilliae, B. melolonthae and B. lentimorbus shared common antigens and suggested the classification of the milky disease bacteria into two species, B. popilliae (containing three varieties, popilliae, melolonthae and lentimorbus) and B. euloomarahae, an Australian isolate that has not been grown in vitro (11).

14

Milner has described a fourth variety of B. popilliae (var. rhopeae) (61-63).

This isolate produces parasporal inclusions,

like var. popilliae and melolonthae, but has a larger paraspore than these varieties.

This isolate and var. melolonthae will

not grow in vitro at 37oC.

Milner (59) has suggested classifying

the milky disease bacteria on the basis of formation of the parasporal crystal during sporulation and its size and position in the sporangium.

His categories include:

A1 - large spore, produces parasporal body that is often small and overlaps the spore.

Example: B. popilliae var. popilliae.

A2 - large spore, produces parasporal body that is often large and separated from the spore.

Example: RM12, the only example

of this type B1 - large central spore, no paraspore.

Example: B. popilliae

var. lentimorbus B2 - small spore in a small sporangium, spore often eccentric, no paraspore.

Example:

B. popilliae var euloomarahae.

This method of classification has the advantage of being purely morphological in nature, and milky disease bacteria can be identified when viewed under a microscope.

This also allows the

identification of isolates unable to be grown in vitro.

A

disadvantage to this classification scheme is the necessity for infecting larvae to produce the spores.

DNA-DNA Similarities One method of determining phenetic relationships between bacteria is the study of deoxyribonucleic acid similarity.

DNA

similarity has been used to differentiate between bacteria at the species level.

It has been recently proposed that DNA

similarity should be used to examine relationship between

15

closely related strains, while rRNA gene sequence analysis should be used to determine more distant relationships (89). The primary structure of the rRNA gene is highly conserved, and species with more than 70% DNA similarity usually have more than 97% rRNA sequence similarity (94).

DNA similarity values of 70%

or more are generally considered to be indicative of identical species (46).

This demonstrates that rRNA sequence analysis

will not differentiate between closely related members of a species because of the high conservation of the sequences.

DNA similarity studies are based on the fact that deoxyribonucleic acid can be denatured and then renatured back into the native molecule.

If competitor DNA is introduced after

the denaturation of the DNA molecule, to some extent the competitor DNA will renature or hybridize with the original molecule.

The amount that it renatures correlates with the

amount of similarity between the sequences of the two molecules.

Similarity experiments are performed using a small amount of labeled DNA and a large amount of unlabeled competitor DNA. The labeled DNA does not reassociate appreciably with itself because it is a small amount and the strands are outcompeted by the competitor DNA in solution.

Instead, the labeled DNA

reassociates to the extent possible with the unlabeled DNA.

The

reassociated DNA is then treated with S1 nuclease to degrade any single stranded DNA left in the mixture (28).

This eliminates

the radioactive count from any labeled DNA that did not reassociate with another strand. Sheared native salmon sperm DNA is used as a control to determine the amount of reassociation of the labeled DNA (45).

The salmon DNA is highly unrelated to the

bacterial DNA and therefore will not reassociate appreciably with the labeled DNA.

After S1 nuclease treatment, only the

16

rehybridized labeled DNA will be detected.

This allows a

determination of the amount of radioactive background caused by reassociation of labeled DNA molecules to be made (45).

RAPD Analysis A method used to differentiate bacteria, including members of the genus Bacillus, at the strain level is the technique called randomly amplified polymorphic DNA, or RAPD (102). RAPD’s are performed using genomic DNA as a template and arbitrarily chosen PCR primers.

The primers are short in length

(10 base pairs) and may prime the DNA at none, one or many locations.

Polymorphisms in the size of the PCR fragments

result from loss or addition of a primer site through point mutations or through deletions and insertions in the chromosome between primer sites (58).

This differentiates between strains

because any given strain may or may not contain the same site where the primer binds or the same amount of DNA between primer sites.

PCR conditions are optimized in order to facilitate the

binding of an arbitrary primer (annealing temperature 36oC).

The

low annealing temperature allows for a certain amount of base pair mismatching between the primer and the template, thereby increasing the number of PCR fragments received from the primer. The bands created by the use of the random primers could produce a unique fingerprint when electrophoresed. This fingerprint is then compared to that of other strains, and each band is considered to be one characteristic.

It can then be decided

which strains share more characteristics, and their relatedness evaluated based on shared bands.

Originally used as a genetic mapping tool, RAPD analysis has been used extensively to distinguish among strains of

17

bacteria, fungi, plants and animals (7, 58, 102).

RAPD strain

typing has been shown to be much more sensitive than typing using multi-locus enzyme electrophoresis (MLEE).

Wang et al.

(98) found that by using RAPD analysis, they could distinguish 74 out of 75 isolates of Escherichia coli, compared to the identification of 15 groups of the same isolates by MLEE.

RAPD has been correlated to restriction enzyme analysis of PCR amplified small-subunit DNA coding for rRNA.

This

correlation illustrated that RAPD analysis is useful for providing taxonomic information at the species level (8).

In a

later study, Baleiras Couto et al. (7) compared the usefulness of RAPD analysis in discriminating organisms at the strain level to that of restriction enzyme analysis of the internal transcribed spacer (ITS) and nontranscribed spacer (NTS) regions of Saccharomyces cerevisiae.

This study proved that RAPD

primers could give rise to recognizable intraspecies patterns, thereby distinguishing between strains of S. cerevisiae isolated from spoiled beer and wine.

Both RAPD analysis and restriction

enzyme analysis of the ITS and NTS spacer regions of S. cerevisiae were shown to be useful in yeast identification (7). Renders et al. (75) compared RAPD analysis with pulsed field gel electrophoresis (PFGE) of Pseudomonas aeruginosa, showing that RAPD results were very comparable to those obtained from PFGE.

RAPD analysis is technically easier and more straightforward than most of these other molecular typing methods, making it a strain typing method of choice in bacterial systematics and epidemiology (40, 75).

Because RAPD’s are PCR

based, they require only nanogram amounts of DNA, which does not need to be highly purified or double stranded.

This allows

RAPD’s to be used in many situations where isolation of DNA is

18

difficult (98).

It has been shown to be useful both at

identifying bacterial species and bacterial strains with the use of properly selected primers. The formation of the RAPD fingerprint requires no prior genetic knowledge of the organism and is unaffected by DNA modifications such as methylation, making this technique particularly useful for taxonomic purposes (98).

Vancomycin Resistance Stahly et al. (91) showed that certain strains of Bacillus popilliae and B. lentimorbus are resistant to the antibiotic vancomycin.

Vancomycin is a glycopeptide antibiotic that was

isolated from Streptomyces orientalis in 1956 (55).

The

molecular structure of vancomycin is based upon a linear heptapeptide molecule substituted with five aromatic rings. Vancomycin inhibits bacterial growth by halting peptidoglycan synthesis (9). The antibiotic is readily adsorbed onto the cell wall of gram positive bacteria and the UDP-Nacetylmuramylpentapeptide precursors (Chatterjee 1966). Vancomycin binds to the pentapeptide side chain at the terminal D-alanyl-D-alanine residues (70).

This binding is accomplished

through hydrogen bonds formed between the D-alanyl-D-alanine terminus of the precursor and the heptapeptide backbone of the antibiotic molecule (83).

These bonds are strengthened by

hydrophobic interactions between the peptide methyl groups and the hydrocarbons of the antibiotic (Williams 1983). Binding of vancomycin to the terminus of the pentapeptide side chain inhibits transglycosylation of the sugar backbone and transpeptidation of the pentapeptide side chain (9).

19

Vancomycin is only effective against Gram positive organisms as it is unable to cross the outer membrane of Gram negative cells (9).

Vancomycin is unusual in that it never

actually enters the bacterial cell, but is active at the cell surface.

This means that cells are unable to use efflux

mechanisms or metabolism of the antibiotic to protect themselves, relying mainly on changing the antibiotic target to become resistant.

Resistance to this antibiotic has emerged among several clinically important bacteria, including Enterococcus, Staphylococcus epidermidis, Leuconostoc and Pediococcus (Rubin) (21, 24, 80, 95).

Prevalence of vancomycin resistant

enterococci (VRE) in the United States has risen from 0.3% of hospital acquired infections in 1989 to 7.9% of hospital acquired infections in 1993 (25).

Clonally related isolates of

VRE have been obtained from different patients in the same hospital as well as in different cities (22, 65).

It is thought

that the increase in the number of VRE may be due to increased use of glycopeptide antibiotics as prophylactics and their use in patients sensitive to penicillin.

Markopulos et al. (54)

showed that glycopeptide resistance could not be developed in a step-wise fashion in enterococci, however, Staphylococcus epidermidis was able to develop increased resistance to glycopeptides due to selection pressure (54).

These findings

support the idea that increased use of vancomycin and related glycopeptide antibiotics has contributed to the increase in bacterial resistance.

Vancomycin resistance appears to be present in four distinguishable types; A, B, C and D.

Type A resistance (VANA

phenotype) is characterized by a very high minimum inhibitory

20

concentration (MIC) for vancomycin, as well as a high MIC for the related antibiotic teicoplanin (1).

Type A resistance is

encoded by a transposon, Tn1546, a member of the Tn3 family, and is usually found on a plasmid (4, 6, 17).

Like Tn3, the

transposase and resolvase genes are transcribed in opposite directions and the genes for vancomycin resistance are located downstream from the resolvase gene (6).

This transposon, when

placed in a host deficient in general recombination, is replicative and leads to formation of a conjugative plasmid.

The VANA operon consists of seven genes, five of which are necessary for resistance to vancomycin, and two of which are accessory genes (4).

The first two genes in the operon, vanS

and vanR, encode a two component regulatory system analogous to the CheY/CheA and OmpR/EnvZ systems (5).

VanS shows sequence

similarity to the membrane bound histidine kinase sensor proteins while VanR shows response regulator similarity (104). Arthur et al. (5) showed that expression of the downstream genes vanH, vanA and vanX were transcriptionally regulated by VanS and VanR.

Wright et al. (104) proved that the cytosolic domain of

VanS is phosphorylated at His194 and that phosphorylated VanS readily transferred the phosphate to VanR at Asp53. Phosphorylated VanR binds to DNA at the vanH and putative vanR promoter regions, activating transcription of vanH, vanA and vanX in response to vancomycin or related antibiotics teicoplanin and moenomycin (5, 44).

Binding of phosphorylated

VanR to the vanR putative promoter region represses transcription of VanR (44).

VanS was shown to negatively

control promoter activation by VanR in the absence of glycopeptides due to dephosphorylation of VanR by VanS (2).

21

vanH encodes a dehydrogenase which converts pyruvate to Dlactate, providing the substrate for the VanA protein (1, 13, 17).

vanA codes for a ligase of altered specificity.

The

normal cellular ligase (ddl gene product) ligates two D-alanines to provide the D-alanyl-D-alanine precursor used in the synthesis of many bacterial cell walls (97). alanine to the D-lactate produced by VanH.

VanA ligates D-

When this is

incorporated into the pentapeptide precursor and eventually the cell wall, it prevents binding of vancomycin to the peptidoglycan (17).

The final required gene product is VanX, a

d,d-dipeptidase which hydrolyzes the vancomycin sensitive precursor D-alanyl-D-alanine (106).

Digestion of this molecule

ensures that only resistant peptidoglycan will be manufactured by the cell (76).

These five genes and protein products are

required for a cell to exhibit resistance to vancomycin.

The

accessory proteins VanY and VanZ are also encoded by the VANA operon.

VanY is a d,d-carboxypeptidase that cleaves the

terminal D-lactate from side chains that have not participated in crosslinking (4, 105).

VanZ confers resistance to

teicoplanin, a glycopeptide antibiotic structurally related to vancomycin, in an unknown fashion (3).

VANB type resistance is characterized by a variable MIC for vancomycin and sensitivity to teicoplanin (1).

The VANB operon

consists of seven genes and is located on either a large conjugative chromosomal element or on a plasmid (73, 74, 103). The VANB element has been transferred naturally from enterococci to Streptococcus bovis, giving weight to the fear that vancomycin resistance will be eventually transferred to Staphylococcus aureus (43, 71).

VanRB and VanSB comprise a two

component regulatory system that operates in a similar manner to that found in the VANA operon.

VanRB and VanSB have a low amino 22

acid similarity to VanR and VanS, 34 and 23 % respectively. However, VanRB and VanSB do show structural similarity to other two component regulatory system proteins.

The C terminal region

of VanSB contains conserved amino acid residues characteristic of histidine kinase sensor proteins.

The N terminal domain of VanRB

has conserved lysine and aspartate residues characteristic of response regulators (36).

Constitutively expressed, VanRB and

VanSB together trans-activate transcription of downstream genes vanYB, vanW and vanHB.

Preexposing the cells to vancomycin can

induce resistance to teicoplanin.

Activation of VanRB and VanSB

seems to be due to functional activation of VanSB by vancomycin.

The VANB operon contains five additional genes; vanHB, vanB, vanXB, vanW and vanYB.

VanHB, VanB and VanXB show very high

structural and functional similarity to VanH, VanA and VanX (67, 76 and 74 % respectively) (57).

VanY and VanYB share only 30 %

amino acid similarity, although both proteins are d,dcarboxypeptidases (36).

VanW does not show similarity to any

sequence in the databases and the VANB operon does not contain a VanZ homolog, explaining the sensitivity to teicoplanin exhibited by VANB organisms (36).

Type C resistance is considered natural resistance (VANA and VANB are acquired) and is found in organisms such as Leuconostoc, Lactobacillus, and Enterococcus spp.

This

resistance can be either constitutive, found in Leuconostoc and Lactobacillus, or inducible (found in enterococci) (31, 79).

In

E. gallinarum a ligase gene responsible for vancomycin resistance was found and designated vanC-1.

The protein VanC-1

shows 29 % similarity with VanA and 38 % similarity with the Dalanyl-D-alanine ligases of E. coli (31).

However, as opposed

to VanA which ligates D-alanine and D-lactate, VanC-1 ligates D-

23

alanine with D-serine, resulting in peptidoglycan with lowered affinity for vancomycin (14, 77).

Two organisms related to E. gallinarum, E. casseliflavus and E. flavescens were examined and shown to posses different vanC ligases designated vanC-2 and vanC-3 respectively.

vanC-2

shows high nucleotide and amino acid similarity with vanC-1, 66 and 69 % respectively.

vanC-3 differs from vanC-2 by 10

nucleotides, equivalent to 4 amino acid changes (66).

Both E.

casseliflavus and E. flavescens contain an additional ligase gene, designated ddlE.

Cass.

and ddlE.

flav.

These gene products

ligate D-alanine with D-alanine and are related to the ddl genes found in E. coli.

The deduced amino acid sequences for the two

genes found in E. casseliflavus and E. flavescens are identical (66).

These organisms make peptidoglycan that has D-alanyl-D-

lactate at the end of the pentapeptide side chain, rather than the sensitive D-alanyl-D-alanine even though the ddl genes are present.

Lactobacillus and Leuconostoc have also been shown to

synthesize peptidoglycan precursors that terminate in D-lactate in a constitutive manner (14, 42).

VAND has been recently described in Enterococcus faecium by Perichon et al. (69).

It is characterized by constitutive, low

level resistance to both vancomycin and teicoplanin.

The ligase

responsible for this phenotype was identified and designated vanD.

The deduced amino acid sequence of this gene has 69 %

similarity with VanA and VanB and 43 % similarity with VanC. This E. faecium isolate was found to synthesize peptidoglycan precursors that terminate in D-lactate (69).

24

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33

CHAPTER TWO Materials and Methods Media and Reagents

MYPGP broth

1.5% yeast extract, 1.0% MuellerHinton broth, 0.3% K2HPO4, 0.1% sodium pyruvate, 0.2% glucose MYPGP agar MYPGP broth plus 2.0% agar Cell Suspension buffer 10 mM Tris-HCl (pH 8.0), 1 mM disodium EDTA, 0.35 M sucrose 2X Lysing buffer 100 mM Tris-HCl (pH 8.0), 20 mM disodium EDTA, 0.3 M NaCl, 2% (w/v) SDS, 2% (v/v)

β-mercaptoethanol, 100 µg/ml proteinaseK RNase

1 mg/ml RNase A dissolved in 0.15 M NaCl (pH 5.0), 4,000 U/ml T1 RNase TE buffer 10 mM Tris-HCL (pH 8.0), 1 mM EDTA Iodination buffer 7.2 M NaClO4, 0.02 mM KI in 80 mM glacial acetic acid (pH 4.8) TlCl3 catalyst 1.0 mg/ml TlCl3 dissolved in 100 mM acetic acid (pH 4.8) Sodium phosphate buffer 0.5 M NaH2PO4.H2O, 0.5 M Na2HPO4 (pH 6.8) Stop reaction buffer 0.5 M sodium phosphate buffer (pH 7.0) HA buffer 0.14 M sodium phosphate buffer, 0.5% SDS Tris buffer 1.0 M Tris-HCl (pH 8.0) TE-SDS buffer TE buffer plus 0.1% SDS Salmon sperm DNA salmon sperm DNA dissolved in TE, then sheared to 400-600bp sodium acetate buffer 3.0 M sodium acetate S1 Nuclease buffer 0.3 M NaCl, 0.05 M acetic acid, 0.5mM ZnCl2, pH 4.6 HCl buffer 1 M HCl, 1% Na4P2O7.10H2O, 1% NaH2PO4.H2O Acid wash buffer S1 storage buffer High Salt buffer 50X TAE

1:5 dilution of HCl buffer 20 mM Tris, pH 7.5, 50 mM NaCl, 0.1 mM ZnCl2, 50% glycerol 13.2X SSC, 5 mM HEPES, pH 7.0 2.0 M Tris base, 57.1 ml/L glacial acetic acid, 100 ml/L 0.5 M EDTA (pH 8.0)

34

Gel loading dye 10X TNE 10X buffer (Promega) dNTP mixture MgCl2

30% sucrose in TE with bromophenol blue 100 mM Tris-HCl, 10 mM EDTA, 2.0 M NaCl, pH 7.4 500 mM KCl, 100mM Tris-HCl (pH 9.0), 1% Triton X-100 2.5 mM each dATP, dCTP, dTTP, dGTP 25 mM MgCl2

Depurinating solution 250 mM HCl Denaturing solution 0.5 M NaOH, 1.5 M NaCl Neutralization solution 1.0 M Tris-HCl (pH 8.0), 1.5 M NaCl 20X SSC 3.0 M NaCl, 300 mM sodium citrate (pH 7.5) Prehybridization buffer 5X SSC, 1% (w/v) Blocking reagent (Boehringer Mannheim), 0.1% Nlauroylsarcosine, 0.2% SDS 2X wash 2X SSC, 0.1% SDS 0.5X wash 0.5X SSC, 0.1% SDS 10X Maleic acid buffer 100 mM maleic acid, 150 mM NaCl, pH 7.5 Blocking solution 1% Blocking reagent (Boehringer Mannheim) dissolved in 1X maleic acid buffer Detection buffer 100 mM Tris-HCl, 100 mM NaCl, pH 9.5 Color developing solution

45 µl 4-nitroblue tetrazolium chloride (NBT, Boehringer Mannheim) 35µl 5-bromo-4-chloro-3-indoyl-phosphate (Xphosphate, Boehringer Mannheim) dissolved in 10 ml detection buffer

Bacterial strains and growth conditions

Bacterial strains used in this study are listed in Tables 1 and 2. All strains were grown in MYPGP broth or on MYPGP plates (1). Bacteria used to inoculate flasks for DNA isolation were grown overnight in 5 ml of MYPGP broth with shaking at 30oC. Two, two-liter erlenmeyer flasks containing 500 ml of MYPGP broth each were inoculated with 5 ml of culture and incubated for approximately 16 h in a New Brunswick G25 shaker at 30oC with shaking (175 rpm). Oneliter erlenmeyer flasks containing 250 ml of media were inoculated

35

for RAPD DNA isolation. Cells were harvested by centrifugation (12,000 x g for 15 min) and the cell pellet was stored at -20oC. Phenotypic testing was performed on MYPGP plates containing either 150 µg/ml vancomycin (Sigma) or 2% NaCl.

The plates were

streaked from an MYPGP plate grown overnight and then incubated for 1 to 2 days at 30oC. Growth was determined by visual examination of the plates. Bacterial tolerance of 2% NaCl was also tested using MYPGP broth supplemented with 2% NaCl. Klett tubes containing 5 ml media were inoculated and incubated at 30oC on a New Brunswick model TC-5 roller drum shaker (23 rpm). Growth was determined as greater than a doubling in absorbance. Table 1.

B. popilliae and B. lentimorbus strains used in DNA-DNA

reassociation Refer to Rippere et al. 1998.

Molecular systematics of Bacillus popilliae

and Bacillus lentimorbus, bacteria causing milky disease in Japanese beetles and related scarab larvae.

Int. J. Syst. Bacteriol. 48:395-

402. Table 2.

B. popilliae and B. lentimorbus strains from diverse host

insects and geographical regions used in RAPD analysis Strain

Host Insect

Source

ATCC 14706+ ATCC 14707*

Popillia japonica

USA1 USA1

BlPj1+ Bp1*

Popillia japonica

Bp6+ Bp9+ Bp10+ Bp11* Bp12+ Bp13+ Bp14+

Popillia japonica

Popillia japonica Papuana woodlarkiana

Ataenius spretulatus Anomala flavipennis Holotichia oblita Popillia japonica Cyclocephala hirta

36

USA6 Papua New Guinea2 USA2 USA, NY2 USA, NC2 USA2 China2 Australia2 USA, CA2

Bp15* Bp16* Bp17* Bp18* Bp19* Bp21* Bp22+ Bp23++ Bp25* Bp26* BpF+ BpCb1* BpPa1* BpPj1+

Cyclocephala lurida

Phyllophaga anxia

USA, TX2 USA, NC2 USA, TX2 Japan2 Australia2 USA, TN2 Panama2 USA2 USA, NY2 USA, FL2 Europe5 USA6 USA6

Popillia japonica

USA6

DNG 2+

Popillia japonica

USA6

DNG 11+

Anomala orientalis

USA6

DNG 4+

Anomala orientalis

USA6

KLN1+

Popillia japonica

USA6

KLN3+

Popillia japonica

USA6

NRRL B-2524+ NRRL B-4081+

Popillia japonica

USA4 Europe4 USA4

Polyphyla comes Phyllophaga crinita Anomala diversa Rhopaea morbillosa Phyllophaga sp. Popillia japonica Cyclocephala hirta Cyclocephala parallela Cyclocephala borealis

Melolontha melolonthae

NRRL B-4145+ NRRL B-4154+

Odontria

RM23+

Anoplognathus porosus

Australia3

RM29+

Lepidiota picticollis

Australia3

(strain Odontria)

USA4

ATCC, 2Klein, 3Milner, 4Nakamura 5Schnetter, 6Stahley

1 *

B. lentimorbus +B. popilliae ++ B. popilliae Dutky

Stock cultures were made by adding 900 µl of an overnight culture grown in MYPGP broth to 100 µl sterile glycerol to make a final glycerol concentration of 10%.

The cultures were mixed and

stored at -80oC.

Isolation of bacteria from dried beetle hemolymph

37

Hemolymph samples microscope slide. Ten added to a spot on the spores. The water was

were received as a dried film coating a microliters of sterile distilled water were slide to allow resuspension of the dried lifted off of the slide using an Eppendorf

pipettor and added to 90 µl sterile distilled water.

The spore

suspension was mixed and incubated in a 60oC waterbath for 20 min. dilution series was performed and the 10-6 and 10-7 dilutions were plated on MYPGP agar. The plates were incubated at 30oC for approximately one week, during which time any possible B. popilliae

A

colonies were restreaked on MYPGP agar. These cultures were checked for purity by restreaking, microscopic examination and the catalase test (B. popilliae and B. lentimorbus are catalase negative). DNA isolation for DNA-DNA reassociation

The cell pellet was taken out of the freezer and fully thawed. DNA was isolated following a variation of the Marmur procedure (2). Five milliliters of cell suspension buffer were added to the pellet, and the pellet was resuspended using a sterile 5 ml glass pipet. Fifteen ml suspension buffer were added to the cells along with 1

µg/ml lysozyme.

The suspension was transferred to a 125 ml glass

stoppered erlenmeyer flask and incubated at 37oC for 3 h, followed by the addition of 20 ml 2X lysing solution and 10 ml 5 M sodium perchlorate. Following incubation at 55oC for 2 h, 15 ml of phenol:chloroform:isoamyl alcohol (25:24:1) were added to the cells, which were briefly shaken vigorously by hand to homogenize the mixture, followed by vigorous shaking for 20 min on a platform shaker. The mixture was centrifuged at 17,000 x g for 10 min to separate the aqueous layer from the phenol layer. The aqueous layer was removed from the centrifuge tube with an inverted 5 ml glass pipet and placed in the erlenmeyer flask. The phenol:chloroform extractions were repeated until the aqueous layer was clear. After the final phenol:chloroform extraction, the aqueous layer was transferred to a clean erlenmeyer flask and 0.6 volume isopropanol was added to precipitate the nucleic acids. The nucleic acids were clotted by gentle swirling of the flask, and the clot held back with a sterile 5 ml glass pipet while the alcohol was poured off. The nucleic acids were washed with cold 80% ethanol for 15 min with

38

occasional swirling. The ethanol was poured off in the same manner as the isopropanol, and the nucleic acids were allowed to air dry. Once dry, the DNA was resuspended in sterile TE buffer and refrigerated at 4oC overnight.

The next morning, 250 µl RNase mix

were added to the nucleic acids and incubated at 37oC for 1 h to degrade any RNA present. Five milliliters chloroform:isoamyl alcohol (24:1) were added to the DNA, shaken vigorously to homogenize the mixture, and then shaken for 20 min. The DNA was centrifuged at 17,000 x g for 10 min. The aqueous layer was removed and placed in a sterile 100 ml beaker, to which was added 0.1 volume of 3 M sodium acetate (2 ml). The DNA was precipitated by the addition of 2 volumes 95% ethanol. The precipitated DNA was spooled on a glass rod, washed with cold 80% ethanol and allowed to air dry. The dry DNA was resuspended in 3 ml TE buffer, quantified at 260 nm and stored at -20oC. DNA sample preparation

The samples to be used for DNA-DNA reassociation experiments were diluted to a concentration of 0.4 mg/ml in a final volume of 4-5 ml. The samples were passed through a French Pressure Cell (American Instrument Co.) at 16,000 lb/in2 and fragment sizes were determined by electrophoresis on a 0.7% agarose gel. Any sample that had fragment sizes larger than 800 bp was passed through the pressure cell again. After shearing, the DNA samples were heated in a boiling water bath for 5 min, cooled rapidly on ice for 5 min, and centrifuged at 17,000 x g for 10 min at 4oC (2). The samples were stored at -20oC. DNA labeling

Five micrograms (12.5 µl) of the DNA to be labeled were placed in a glass autoinjection vial (Chemical Research Suppliers) and 0.1 volume of 3.0 M sodium acetate (pH 6.0) was added. The samples were mixed well, followed by the addition of 2.0 volumes of cold 95% ethanol. The samples were again mixed well and incubated at -20oC for 1 h, followed by centrifugation at 12,000 x g for 15 min. The supernatant was decanted, cold 80% ethanol added to desalt the pellet, and centrifuged again for 15 min. The supernatant was decanted and the pellet dried at 37oC. The vials were covered with

39

parafilm and stored at -20oC until the labeling reaction could be performed. Fifteen minutes prior to the beginning of the labeling reaction, 23 µl of reaction buffer were added to each dried sample.

Once the

samples were resuspended, 1.0 µl (100 mCi) of sodium iodide (125I, Dupont New England Nuclear) was placed on the side of the vial, followed by 6.0 µl of TlCl3 catalyst on the opposite side of the vial. I was used because it can be chemically linked to cytosine residues in the presence of thallium chloride, thereby eliminating the need to grow the bacteria with a radioactive isotope. A serum bottle cap was crimped onto each reaction vial, the contents mixed and incubated in a 70oC waterbath for 20 min. While the samples were incubating, NAP25 sepharose (Pharmacia) columns were equilibrated by washing three times with HA buffer, and a tuberculin syringe was loaded with stop reaction buffer. The reaction tubes were removed from the waterbath, allowed to cool for 2 min and 0.1 ml of stop reaction buffer was injected into each vial. The contents of the vials were mixed and incubated in a 70oC waterbath for 20 min. During this incubation period, hydroxyapatite was added to Pasteur pipets plugged with glass wool and kept moist by plugging the bottom of the pipet. The columns were placed in a glass culture tube in the 70oC waterbath. For each DNA sample, one tuberculin syringe was loaded with 0.15 ml HA buffer and 125

50 µl salmon sperm DNA (denatured, 0.4 mg/ml) were added to a screw cap tube. The NAP-25 columns were placed in the fume hood and allowed to drain and air dry. The vials were removed from the waterbath and cooled for 2 min. Using the prepared syringes, HA buffer (0.15 ml) was injected into the bottom of each vial, and the contents were drawn back up into the syringe. The reactions were loaded directly onto the top of the NAP25 columns and allowed to drain into the columns. HA buffer (2.2 ml) was added to the column, moving the DNA into the bottom portion of the column. The collection tube containing the salmon sperm DNA was placed under the column, 1.8 ml HA buffer added to the column, and

40

the eluent containing labeled DNA was collected. The DNA was denatured again by heating for 5 min in a boiling water bath. The labeled DNA samples were loaded onto dried HA columns, and movement of the DNA through the columns was monitored with a survey meter. Once the DNA moved into the bottom of the columns and started to elute from the bottom, the columns were moved to new collection tubes to begin collecting the labeled samples. The HA columns were washed with 0.5 ml HA buffer and the wash eluent was collected in the same tubes as the labeled DNA. NAP-25 columns equilibrated with three changes of TE + 0.1% SDS were drained until the surface was dry. A disposable serological pipet was used to draw up the labeled DNA recovered from the HA column and the volume recovered was recorded. The labeled DNA was loaded onto the NAP-25 column and the eluent was allowed to drain. An additional volume of TE + 0.1% SDS was added to the column to make the total volume of DNA up to 2.5 ml and allowed to drain. A screw capped culture tube was placed under the column, 3.5 ml TE + 0.1% SDS were added to the column, and the eluent was collected in the tube. Ten microliters of the labeled DNA were transferred to a scintillation vial for gamma counting to determine the strength of the label. Once counted,the labeled DNA was diluted to an activity of 30,000 cpm/ml and stored at -20oC. S1 Nuclease assay

The labeled and unlabeled DNA samples were thawed and then heated in a 65oC waterbath for five minutes. Using an Eppendorf repeating pipettor, 10 µl of labeled DNA were added to the bottom of each reaction tube (200µl polypropylene tubes, Robbins Scientific). Fifty microliters of test DNA were added to each tube. Four tubes contained sheared, native salmon sperm DNA (0.4 mg/ml), four tubes contained DNA homologous to the labeled DNA, and each heterologous DNA was done in duplicate. Following addition of the DNA samples to the reaction tubes, 50 µl high salt buffer were added, the tubes were closed and vortexed eight times. The tubes were transferred to stainless steel racks, a cover placed over the rack, and the entire rack incubated in a 65oC waterbath for 24 h. Following incubation the

41

reactions were stored at -20oC until the rest of the experiment could be performed. The reaction contents were thawed and allowed to come to room temperature. For each reaction, 1 ml of S1 buffer was added to a plastic digestion tube, followed by 50 µl of denatured salmon sperm DNA (0.4 mg/ml).

The contents of each 200 µl reaction tube were

quantitatively transferred to the digestion tube, and the reaction tube was washed twice with 100 µl S1 buffer. added to the digestion tube.

The washes were also

Ten microliters S1 nuclease (10 U/µl)

were added to each digestion tube, the contents vortexed three times, and incubated for 1 h in a 50oC water bath. Following the incubation period, 50 µl 1.2 mg/ml native salmon sperm DNA were added to each tube to serve as a precipitation matrix for hybridized DNA.

To each

tube, 500 µl cold 1 M HCl were added, followed by an incubation at 4oC for 1 h. After the precipitation was complete, the reactions were filtered through Whatman glass filter strips ( No. 1825 915 GF/F). Each reaction tube was rinsed twice with HCl wash buffer and the rinses were filtered on the same strips. The filter strips were dried under a heat lamp for at least 1 h and once dry, the circles where the DNA was collected were removed with forceps. The circles were placed in the bottom of scintillation vials and counted for 2 min each with a Beckman gamma counter (2).

DNA isolation for RAPD experiments

Thawed cell pellets were resuspended in 8 ml cell suspension buffer and transferred to a 125 ml glass stoppered erlenmeyer flask. Dry lysozyme (final concentration 1 mg/ml) was added to the contents of the flask, mixed and incubated at 37oC for 3 h. After the incubation, 8 ml of 2X lysing solution (55oC) and 4 ml 5 M NaClO4 were added to the mixture. The flasks were incubated at 55oC for 2 h to lyse the cells. Following this incubation, 8 ml phenol:chloroform:isoamyl alcohol (25:24:1) were added to each flask,

42

shaken vigorously to homogenize the mixture and placed on the shaker for 20 min. The mixtures were poured into centrifuge tubes and centrifuged at 17,000 x g and 4oC for 10 min. The aqueous layer was removed from the tube, placed in the flask and the extraction was repeated until no protein layer was present in the centrifuged sample. After the last extraction, the aqueous layer was placed in a clean flask and 0.6 volumes 100% isopropanol were added to precipitate the nucleic acids. The flask was swirled to clot the nucleic acids, and the alcohol was poured off. Cold 80% ethanol was added to the flask and incubated for 10 to 15 min with occasional swirling to wash the samples. The ethanol was poured off, the nucleic acids stuck to bottom of the flask, and the flask was turned upside down to dry. The nucleic acids were rehydrated in 8 ml TE buffer and 125 µl RNase mix were added to the flask. Following an incubation at 37oC for 1 h, 2 ml chloroform:isoamyl alcohol were added to each flask. The flasks were shaken vigorously to homogenize the mixture and placed on the shaker for 20 min. The contents of each flask were poured into centrifuge tubes and centrifuged at 17,000 x g and 4oC for 10 min. The aqueous layer was removed, placed in a 100-ml beaker and 800 µl of 3 M sodium acetate were added.

The sample was overlayed

with two volumes of 95% ethanol and the DNA was collected on a glass rod. The DNA was washed in cold 80% ethanol, the glass rod was inverted and placed in the beaker to let the DNA dry. Once dry, the DNA was resuspended in 1 ml warm TE and stored at -20οC (2). Determination of DNA concentration

The DNA was quantified using a fluorometer (Hoefer TKO-100). a reference, 830 µg/ml standard λ strain CI85757 DNA (USB) was diluted to 250 ng/µl in sterile TE buffer.

The fluorometer was

allowed to warm up and blanked using 2 ml assay solution in a glass cuvette.

The assay solution contained 1X TNE and 0.1 µg/ml Hoechst

33258 (Hoefer Scientific).

The fluorometer was standardized by

43

As

adding 4 µl of λ DNA standard to the cuvette and adjusting the machine to read 250 ng/µl.

Measurements of DNA concentration were

made by adding 4 µl of sample to 2 ml assay solution.

Samples were

diluted to give a final working concentration of 5 ng/µl and stored at -20oC. RAPD analysis

The sequences of the primers (Operon Technologies) used in this study are given in Table 3. The primers were rehydrated in sterile, milli-Q filtered water to a final concentration of 0.125 µg/µl and stored at -20oC. The working solution of dNTP’s was prepared by diluting 100mM dTTP, dATP, dCTP and dGTP together in sterile, milli-Q filtered water and stored at -20oC.

Table 3.

RAPD primer sequences

Primer Name

Sequence

OPA-03

5’-AGTCAGCCAC-3’

OPA-04 OPA-05 OPA-07 OPA-08 OPA-10 OPA-11 OPA-15

5’-AATCGGGCTG-3’ 5’-AGGGGTCTTG-3’ 5’-GAACGGGGTG-3’ 5’-GTGACGTAGG-3’ 5’-GTGATCGCAG-3’ 5’-CAATCGCCGT-3’ 5’-TTCCCGACCC-3’

For at least thirty minutes prior to use, milli-Q filtered water, 50% glycerol, mineral oil, microcentrifuge tubes and rack, gloves, aerosol resistant pipet tips and pipettors were exposed to UV light in a laminar flow hood. Primers, Promega 10X buffer, dNTP’s, MgCl2 and the DNA samples were thawed at room temperature. The Taq DNA polymerase (5000 U/ml, Promega) was stored in Buffer A (Promega) at -20oC until used.

44

All RAPD reactions were prepared in a laminar flow hood after exposure of the contents of the hood to UV light for 30 minutes. The amount of primer used to obtain a final concentration of 0.6 µM varied due to the different molecular weights of the primers. The reagents were added together to make a “master mix” and aliquots were dispensed into the reaction tubes. Each reaction tube contained 0.5

µl 50% glycerol, 2.5 µl 10X buffer, 1.0 µl dNTP’s (100 µM), 3.0 µl MgCl2 (3 mM), 0.3 µl Taq polymerase (1.5 U), 0.6 µM primer, 3.0 µl DNA template (15 ng) and the appropriate amount of milli-Q filtered water to make up a final volume of 25 µl. drops sterile mineral oil.

Samples were overlayed with two

Negative controls in which template DNA

was replaced with 3.0 µl milli-Q filtered water were also prepared for each primer. The RAPD reaction tubes were placed in a PTC-100 thermalcycler (MJ Research) with 1 drop of mineral oil per well. The following temperature profile was programmed: 95oC for 5 min followed by 75 cycles of 94oC for 20 sec, 36oC for 20 sec, and 72oC for 2 min. Upon cycle completion, samples were maintained at 4oC until electrophoresis (6). A 1.7% (w/v) gel composed of 1.0% Synergel (Diversified Biotech) and 0.7% agarose was poured in preparation for electrophoresis. Synergel and agarose were mixed in a slurry with 15 ml 95% ethanol. TAE buffer (1X) was slowly added to the slurry to a final volume of 300 ml and the flask was weighed. The mixture was heated to melt the Synergel-agarose mixture, the ethanol was evaporated off, and water was added (by weight) to the flask to replace the amount which had evaporated during heating. The mixture allowed to cool slightly before pouring the gel. The PCR amplification product was removed from the tube by inserting a pipet tip below the mineral oil layer, expelling an air bubble from the tip, and immediate withdrawal of a 10 µl volume.

The sample was mixed with 3 µl loading buffer on

parafilm and loaded onto the gel.

The gel was electrophoresed at 3.2

45

V/cm in recirculating 1X TAE buffer, stained in 0.5 µg/µl ethidium bromide for 2 h and photographed. Isolation, amplification and digoxygenin labeling of individual RAPD bands

RAPD reactions were prepared as before with the desired primer to isolate single RAPD bands, which were labeled with digoxygenin for use as probes. Reactions were subjected to the thermal cycling conditions described above. Ten microliters of the RAPD reaction were loaded onto a 1.7% low-melt agarose gel prepared with 1X TAE and electrophoresed at 4oC. The gel was stained and photographed as previously described. Using a sterile razor blade, the RAPD band of interest was cut out of the gel and placed in a microcentrifuge tube. The microcentrifuge tube was placed in a 65oC waterbath for 10 min to melt the agarose. The DNA was purified from the agarose using Wizard PCR Preps (Promega). One milliliter of PCR preps resin was added to the gel slice, vortexed briefly and then incubated for 1 min with occasional vortexing. The DNA/resin mixture was added to a 3 ml disposable syringe attached to a PCR preps minicolumn and then dispensed into the column. The column was washed with 2 ml 80% isopropanol, centrifuged for 20 sec at 12,000 x g and placed on a new microcentrifuge tube.

50 µl warm (50-60oC) TE were added directly to

the column and incubated for 1 min, followed by centrifugation for 20 sec at 12,000 x g. The eluted purified DNA was stored at -20oC. To generate a higher concentration of DNA for storage and labeling, the purified DNA product was amplified by PCR using the following reagents. Fifty microliter reactions were prepared containing 1.0 µl 50% glycerol, 5.0 µl 10X buffer, 4.0 µl dNTP’s (200µM), 6.0 µl MgCl2 (3mM), 0.8 µl Taq polymerase (2 U), 2 µM primer, 1.0 µl DNA template and the appropriate amount of sterile milli-Q filtered water to make up the final volume. overlaid with 2 drops sterile mineral oil.

Each reaction was

The reactions were placed in the thermalcycler with 1 drop mineral oil in each well. The temperature profile was as follows:

46

95oC for 5 min followed by 30 cycles of 94oC for 30 sec, 36oC for 30 sec, and 72oC for 2 min. Upon completion, the reaction tubes were held at 4oC. The PCR product was removed from the tube and purified using Wizard PCR Preps (Promega).

The reaction product was added to 100 µl

direct purification buffer and vortexed briefly to mix. One milliliter of PCR preps resin was added to the sample and vortexed. After a 1 min incubation with occasional mixing, the DNA/resin mixture was added to a 3 ml disposable syringe attached to a PCR preps minicolumn and dispensed into the column. The column was washed with 2 ml 80% isopropanol, centrifuged for 20 sec at 12,000 x g and placed on a new microcentrifuge tube. Fifty microliters of warm TE (50-60oC) were added to the column and after a 1 min incubation, the column was centrifuged for 20 sec at 12,000 x g. The eluted product was stored at -20oC. To label the RAPD band, 25 µl RAPD reactions were prepared as previously described. The working solution of dNTP’s was replaced with a 10X concentrated Boehringer Mannheim dig-DNA labeling mixture (1 mM dATP, 1 mM dCTP, 1 mM dGTP, 0.65 mM dTTP, 0.35 mM dig-dUTP). Each reaction contained sterile water to a final volume of 25 µl, 0.5

µl 50% glycerol, 2.5 µl 10X buffer, 2.5 µl dig-dNTP mix (100 µM), 3.0 µl MgCl2 (3 mM), 0.4 µl Taq polymerase (2 U) and 1.0 µl template DNA. The reactions were overlaid with 2 drops sterile mineral oil. The amplification reactions were placed in the thermalcycler with 1 drop of mineral oil per well. The following temperature profile was entered: 95oC for 5 min followed by 30 cycles of 94oC for 30 sec, 36oC for 30 sec and 72oC for 2 min. Upon completion of the reaction, the products were held at 4oC. The PCR product was removed and purified as previously described. The purified probe was stored at -20oC (6). Estimation of probe yield

47

To estimate the probe yield, serial dilutions of the purified PCR products were prepared in DNA dilution buffer (Boehringer Mannheim) and compared to control DNA (Boehringer Mannheim). The dilutions are described in Table 4. Table 4.

Dilution series for probe estimation

Dilution

Final concentration

Total dilution

A.

2 µl DNA/8 µl buffer

1 ng/µl

1:5

B.

2 µl A/18 µl buffer

100 pg/µl

1:50

C.

2 µl B/18 µl buffer

10 pg/µl

1:500

D.

2 µl C/18 µl buffer

1 pg/µl

1:5,000

E.

2 µl D/18 µl buffer

0.1 pg/µl

1:50,000

One microliter of each dilution of the PCR labeled probe and 1

µl of labeled control DNA were spotted onto a positively charged nylon membrane (Boehringer Mannheim). The membrane was placed on a damp paper towel and UV crosslinked (FB UVXL-1000, Fisher) using the optimal setting. The membrane was placed in a glass petri dish, wetted with 1 ml maleic acid buffer and incubated at room temperature for 5 min with enough Blocking solution to cover the membrane. The Blocking solution was discarded and new Blocking solution containing a 1:5000 dilution of anti-DIG-alkaline phosphatase was added to the membrane. The membrane was incubated with gentle shaking at room temperature for 10 min. The membrane was then washed twice with maleic acid buffer, 5 min per wash, and incubated in detection buffer for 2 min. The detection buffer was discarded and the membrane was placed in a heat sealable bag. Color developing solution was added to the bag, the bag sealed and placed in the dark for 30-60 min until adequate color was developed. The reaction was stopped by the addition of TE. Estimation of yield was done by visual comparison of probe intensity to that of the controls. Southern transfer and DNA hybridization

48

The protocol given by Boehringer Mannheim for Southern transfer, prehybridization, hybridization and colorimetric detection of hybridized probe was followed with a few modifications. RAPD DNA to be transferred was electrophoresed, stained and photographed as previously described. The gel was shaken gently at room temperature for 10 min each in depurinating solution and denaturing solution, then soaked twice at room temperature for 20 min each in neutralization solution. The DNA was transferred overnight to a positively charged nylon membrane (Boehringer Mannheim) by capillary action in 10X SSC. After transfer the membrane was placed on a damp paper towel and UV crosslinked using the optimal setting. The membrane was placed in a heat-sealable bag and incubated in 20 ml/100 cm2 standard prehybridization buffer for 2 h in a 65oC water bath. After prehybridization, the solution in the bag was replaced with an equal amount of prehybridization buffer. One and one-half nanogram/100 cm2 of labeled probe was also added to the bag, the bag sealed and incubated overnight in a 65oC waterbath. After hybridization, the membrane was removed from the bag, placed in a glass baking dish and washed twice in 2X wash solution for 5 min each. Then the membrane was washed twice in 0.5X wash solution for 15 min each. To start color development, the membrane was incubated at room temperature in Blocking solution with gentle shaking for 1 h. After the intial incubation, the blocking solution was discarded and antiDIG alkaline phosphatase diluted 1:5000 in blocking solution (20 ml) was added to the membrane. The membrane was incubated with the antibody at room temperature with gentle agitation for 30 min. After the antibody had bound, the membrane was washed twice in 15 ml 1X maleic acid buffer for 15 min each and washed once in 20 ml detection buffer for 2 min. The membrane was placed in a heat sealable bag and the color developing solution was added. Color development was allowed to proceed in the dark at room temperature until sufficient color had been deposited on the membrane. The color development was

49

stopped by the addition of TE buffer to the membrane and the membrane was stored at 4oC in the dark until photographed. RAPD band analysis

The presence (1) or absence (0) of each RAPD band among the strains was determined for each primer by visual examination of the gel photographs. The results for each strain were recorded in an ASCII format as a rectangular matrix consisting of total bands. All detectable RAPD bands present in the strains were analyzed and scored. Data analysis

The percent DNA similarity data were analyzed with the average taxonomic distance algorithm (3, 5). The distance coefficient was utilized in this case because the data were all quantitative real variables without a range of variation, and as such could be treated as points in space. The coefficient calculates the distance between the points and this value is converted into a dissimilarity value. The matrix obtained was subjected to clustering by the unweighted pair group method with arithmetric averages (UPGMA)(5). Cophenetic matrices for the clusters were computed and the correlation between these coefficients and their corresponding rectangular matrix was computed by using normalized Mantel statistics z (5). This determined how much distortion was present in the phenetic tree. The RAPD data were analyzed using either the Jaccard or Dice similarity coefficients (3, 5). The Jaccard coefficient uses only positive matches in the calculation of similarity. This allows characters which are missing in two or more isolates to be ignored, resulting in a similarity value calculated from characters which are present. The Dice coefficient is also a measure of similarity between OTU's, and does not include characters which are absent in each of the isolates being compared. The matrix of coefficients obtained was subjected to clustering by UPGMA. The NTSYS-pc computer program (version 1.8) was used in the analysis of the data (5). Distance coefficient: dij =√1/n∑k(xki + xkj)2

50

Jaccard coefficient: a/(n-d) where a = all positive matches, n = total sample size and d = all negative matches Dice coefficient: 2a/(2a + b + c) where a = all positive matches, b and c = unmatches

Multiplex PCR-RFLP for detection of the van ligase All PCR reactions were set up in a laminar flow hood. The pipettors, tips, gloves, racks and tubes were exposed to ultra-violet light for at least 30 min prior to use. Each reaction contained 10 pmol of each of six primers designed for detection of the van ligase in the enterococci. Table 5.

Primer sequences are shown in Table 5.

Primer sequences used in multiplex PCR-RFLP reaction for

detection of van ligase genes in enterococci (4). Refrer to Patel et al. 1997. Multiplex polymerase chain reaction detection of vanA, vanB, vanC-1, and vanC-2/3 genes in enterococci. J. Clin. Microbiol. 35:703-707. In addition to the primers, each reaction consisted of 1.25 U Taq polymerase, 200 µM each dNTP, 50 mM KCL, 10 mM Tris-HCl pH 8.3, 1.5 mM MgCl2 and 5 % glycerol.

For the Enterococcus strains, a single

24 h colony was picked off a BHI plate supplemented with 4 µg/ml vancomycin and suspended in a 50 µl PCR reaction.

Five nanograms of

DNA isolated as for RAPD reactions was used as a PCR template for Bacillus popilliae strain ATCC 14706 and Bacillus lentimorbus strain ATCC 14707. The reactions were overlaid with two drops sterile mineral oil and placed in the thermalcycler. The reaction profile was as follows: 95oC for 10 min to lyse the enterococci and denature the template DNA, followed by 60 cycles of 94oC for 1 min, 56oC for 1 min and 74oC for 1 min. 4oC.

Upon cycle completion the reactions were held at

One microliter of MspI (10 U/µl) and 5 µl of restriction buffer

were added to each tube, followed by centrifugation at 13,200 × g for 20 sec to drive the enzyme down into the reaction.

51

The tubes were

then incubated overnight at 37oC and the restriction products were electrophoresed on a 4% MetaPhor (FMC Corp. MA) gel in 1X TAE. Primers specific for the ligase gene found in B. popilliae strain ATCC 14706 were designed from the sequenced ATCC 14706 PCR product. The primer sequences are 5’-GCTGCTTGTTATGCGGAATA-3’ (BPOP-FOR) and 5’-AATTGCTTTCGCCGTCTC-3’ (BPOP-REV). The B. popilliae and B. lentimorbus strains were screened for the presence of the ligase gene using these primers and the above PCR conditions.

Paraspore gene detection using PCR The paraspore gene (cry18Aa1) sequence from a European B. popilliae isolate has been previously described (7). Using the published nucleotide sequence, two sets of PCR primers were designed to cover both the open reading frame found just prior to the gene and the gene itself. The primer sequences are shown in Table 6. PCR reactions were set up in the laminar flow hood with all tools subjected to 30 minutes UV exposure prior to use. Reaction mixtures contained 25 ng template DNA (isolated as for RAPD reactions), 5% glycerol, 1 X buffer, 200 µM each dNTP, 3 mM MgCl2, 25 pmol each primer and 0.5 U Taq polymerase. Reaction mixtures were overlaid with two drops sterile mineral oil and placed in the thermalcycler. Cycling parameters were 95oC for 2 min, followed by 35 cycles of 94oC for 1 min, 54oC for 1 min and 72oC for 2 min. Upon completion of the programmed cycles, the tubes were held at 4oC until electrophoresis on a 1 % agarose gel.

Table 6.

Primer sequences used for detection of cry genes in B.

popilliae and B. lentimorbus.

Primer

Sequence

Location

52

Expected

size cryBP1-F cryBP1-R cryBP2-F cryBP2-R

5’-AGGGAATGGACAGAATGG-3’ 5’-GAAAGCTGAACGCCAATC-3’ 5’-AGGATGTTCCTCCGATCCCCATCAC-3’ 5’-GTTCCGTGGCTCGTAAAATCTCTTC-3’

1058 2020 441 1247

962 bp 806 bp

PCR conditions for the second set of primers, cryBP2-F and cryBP2-R were identical except for a primer annealing temperature of 56oC.

PCR product sequencing All DNA sequencing was performed at the Mayo Clinic (Rochester, MN).

Six microliters of PCR product, 1 µl of 1 U/µl shrimp alkaline

phosphatase and 1 µl of 10 U/µl exonuclease I (United States Biochemicals) were incubated at 37oC for 30 min followed by incubation at 80oC for 15 min.

One microliter of sequencing primer (3.2 µM) and

1 µl of dimethyl sulfoxide were added to the mixtures and the DNA sequence was determined in both directions using a Taq dideoxy terminator cycle sequencing kit and a 373 A DNA Sequencer (Applied Biosystems, CA). The sequence data were analyzed using version 8 of the Genetics Computer Group Sequence Analysis software (4). Labeling of vanE PCR product

A portion of the vanE gene was amplified from ATCC 14706 using PCR conditions identical to those used for screening the Bacillus strains for the presence of the ligase gene.

The

amplified product was cleaned using the Wizard PCR Preps system as described for RAPD band labeling.

The PCR reaction for

digoxygenin labeling was set up as described for the detection of the vanE gene in B. popilliae, with the replacement of the dNTP mix with a digoxygenin labeled dNTP mix.

Upon completion

of the cycling program, the product was cleaned as detailed before.

53

The probe concentration was determined following the procedure used for RAPD probe determination. The probe was stored at –20oC.

Determination of vanE location in B. popilliae

DNA from both B. popilliae strain ATCC 14706 and B. lentimorbus strain ATCC 14707 was digested with MboII in a 20 µl reaction.

The

reaction included 1X enzyme buffer, 3 U enzyme and 2 µg BSA, 1 µg DNA and Milli-Q water to the final volume. The digestions were incubated at 37oC for 2 h and then electrophoresed on a 1 % agarose gel. As controls, undigested DNA as well as unlabeled vanE PCR product were also run on the gel. The gel was stained with ethidium bromide and photographed under UV light, followed by a Southern transfer to a positively charged nylon membrane as described for RAPD’s. Hybridization of the probe and colorimetric detection were performed as previously described for RAPD bands. References

1.

Billot-Klein, D., L. Gutmann, S. Sable, E. Guittet, and J. van Heijenoort.

1994. Modification of peptidoglycan precursors is a common feature of the low-level vancomycin-resistant species Lactobacillus casei, Pediococcus pentosaceus, Leuconostoc mesenteroides, and Enterococcus gallinarum. J. Bacteriol. 176(8):2398-2405. 2.

Dingman, D. W., and D. P. Stahly. 1983. Medium promoting sporulation

of Bacillus larvae and metabolism of medium components. Appl. Environ. Microbiol. 46(4):860-869. 3.

Handwerger, S. 1994. Alterations in peptidoglycan precursors and

vancomycin susceptibility in Tn917 insertion mutants of Enterococcus faecalis 221. Antimicrob. Agent. Chemother. 38(3):473-

475. 4.

Johnson, J. L. 1994. Similarity analysis of DNA's, p. 655-682. In

P. Gerhardt, R. G. E. Murray, W. A. Wood, and N. R. Krieg (ed.), Methods for General and Molecular Bacteriology, 1st ed. American Society for Microbiology, Washington, D. C.

54

5.

Patel, R., J. R. Uhl, P. Kohner, M. K. Hopkins, and F. R. Cockerill, III.

1997. Multiplex polymerase chain reaction detection of vanA, vanB, vanC-1, and vanC-2/3 genes in enterococci. J. Clin.

Microbiol. 35:703-707. 6.

Rohlf, F. J. 1994. NTSYS-pc numerical taxonomy and multi-variate

7.

analysis system version 1.80. Exeter Publishing, Setauket, NY. Woodburn, M. A., A. A. Yousten, and K. H. Hilu. 1995. Random amplified polymorphic DNA fingerprinting of mosquito pathogenic and nonpathogenic strains of Bacillus sphaericus. Int. J. Syst. Bacteriol. 45(2):212-217.

8.

Zhang, J., T. C. Hodgman, L. Krieger, W. Schnetter, and H. U. Schairer.

1997. Cloning and analysis of the first cry gene from Bacillus popilliae. J. Bacteriol. 179(13):4336-4341.

55

CHAPTER THREE Bacillus popilliae and Bacillus lentimorbus, Bacteria Causing Milky Disease in Japanese Beetles and Related Scarab Larvae

Abstract Bacillus popilliae and B. lentimorbus, causative agents of milky disease in Japanese beetles and related scarab larvae, have been differentiated based upon a small number of phenotypic characteristics, but they have not previously been examined at the molecular level.

In this study thirty-four isolates of

these bacteria were examined for similarity by DNA reassociation (henceforth referred to as DNA similarity).

Three distinct but

related similarity groups were identified; the first contained strains of B. popilliae, the second contained strains of B. lentimorbus, and the third contained two strains distinct from but related to B. popilliae.

Some strains received as B.

popilliae were found to be most closely related to B. lentimorbus and some received as B. lentimorbus were found to be most closely related to B. popilliae.

Paraspore formation,

believed to be a characteristic unique to B. popilliae, was found to occur among a subgroup of B. lentimorbus strains. Growth in media supplemented with 2% NaCl was found to be less reliable in distinguishing the species than was vancomycin resistance, the latter present only in B. popilliae.

Bacillus popilliae and B. lentimorbus are pathogens of Japanese beetles (Popillia japonica) and related scarab larvae. Larvae feeding in the soil consume spores of these bacteria and following spore germination in the larval gut, vegetative cells penetrate into the hemocoel. A period of vegetative growth is

57

followed by asynchronous sporulation and death of the larvae. At the time of larval death, the hemolymph may contain up to 5 x 1010 spores/ml (1). The milky color of the larval hemolymph at the time of death has given the condition the name “milky disease” (8).

Because of its action against economically

important insect pests, efforts have been made to develop B. popilliae as a biological control agent. However, the inability to mass produce spores in vitro has prevented large scale manufacture and utilization (5).

Dutky (2) reported a difference in color of the hemolymph in insects infected by either B. popilliae (type A disease) or B. lentimorbus (type B disease).

In addition, Dutky (2), Gordon

et al. (3) and St. Julian and Bulla (9) suggested that a primary distinguishing characteristic between the two named species is the production of a parasporal body by B. popilliae but the absence of this structure in B. lentimorbus. studies prompted

Serological

Krywienczyk and Luthy (6) to propose a single

species, B. popilliae, with three varieties, B. popilliae var. popilliae, B. popilliae var. lentimorbus and B. popilliae var. melolonthae (the last variety based on a European isolate also known as fribourgensis).

This approach was similar to that

proposed by Wyss (14) who emphasized physiological and morphological characteristics in his taxonomic arrangement. Milner (7) utilized the position and size of the spore and paraspore in the sporangium to group the bacteria.

In this

system all milky disease isolates were considered varieties of B. popilliae. The A1 group contained strains with small parasporal bodies overlapping the spore.

Group A2 had a large

parasporal body separated from the spore. Group B1 had a large central spore and lacked a paraspore and group B2 had a small spore and no paraspore.

The utilization of these morphological

58

characteristics in species determination is limited because the paraspore is produced at the time of sporulation which only occurs in living larvae.

Therefore, only those laboratories

capable of collecting and infecting the larvae are able to identify the species (11).

It has been reported that B.

popilliae will grow in media containing 2% NaCl whereas B. lentimorbus will not grow under these conditions (11).

The genetic relationship between B. popilliae, B. lentimorbus, and less well-known bacteria producing milky disease is unknown.

In this study we utilized DNA reassociation

to define relationships between these species.

Our results

validate the existence of the two species and identified the presence of subgroups within the species.

Phenotypic

characteristics presented by the species and subgroups were investigated to facilitate identification.

RESULTS DNA similarity.

DNA was prepared from 34 strains of bacteria

that had been originally isolated from scarab larvae suffering from milky disease.

This DNA was compared to labeled reference

DNA from the type strains of both B. popilliae and B. lentimorbus as well as to three additional strains, one of which was a European isolate sometimes referred to as B. popilliae var. melolonthae (NRRL B-4081), shown in Table 1.

The clusters

obtained from the distance and correlation matrices were almost identical in their topology.

The cophenetic correlations for

both clusters were r=0.98 to 0.99, underscoring the extremely high fit between the original matrices and the phenograms. distance-based phenogram will be used here because it showed higher resolution within the groups. revealed the existence of two groups

59

The similarity study

The

Table 1.

Levels of DNA similarity between Bacillus popilliae

and Bacillus lentimorbus as determined by the S1 nuclease method

Refer to Rippere et al. 1998.

Molecular systematics of Bacillus

popilliae and Bacillus lentimorbus, bacteria causing milky disease in Japanese beetles and related scarab larvae.

Int. J.

Syst. Bacteriol. 48:395-402.

of strains (Fig. 1). The first group showed 84 to 97% similarity to the type strain, B. popilliae ATCC 14706T, and a high similarity to BpPj5, another B. popilliae isolate. These strains were primarily North American in origin and most were isolated from diseased Popillia japonica except for a few from Anomola orientalis (northern masked chafer).

Two strains, NRRL B-4081

and Bp3, showing markedly lower similarity (77% and 73% respectively) to the ATCC 14706T reference strain than did the other strains of B. popilliae.

Bp3 displayed 82% similarity to

NRRL B-4081 whereas the other strains of the B. popilliae group showed only 59% to 67% similarity to that reference strain. Two strains, BlPj1 and NRRL B-2522, were received as B. lentimorbus but showed 95% and 86% similarity to the B. popilliae reference strain and 59% and 62% similarity respectively, to the B. lentimorbus type strain.

Following growth and sporulation in

Japanese beetle larvae, paraspores were detected in NRRL B-2522 but not in BlPj1.

Eight strains showed higher similarity to the B. lentimorbus reference than to B. popilliae. Only one of these was received as B. lentimorbus, while the other seven were received as B. popilliae.

60

Figure 1. UPGMA dendogram of B. popilliae and B. lentimorbus strains based on a distance matrix generated from DNA similarity analysis.

Refer to Rippere et al. 1998.

Molecular systematics of Bacillus

popilliae and Bacillus lentimorbus, bacteria causing milky disease in Japanese beetles and related scarab larvae.

Int. J.

Syst. Bacteriol. 48:395-402.

However, these latter seven strains had lower similarity to B. lentimorbus than one strain, KLN2, received as B. lentimorbus (Table 1).

Microscopic examination of hemolymph from Japanese

beetles or masked chafers infected with six of these strains, Bp7, DGB1, BpCb1, BpCb2, BpPa1, and BpCp1, revealed the presence of parasporal bodies in the sporangia.

Strain Bp1 has not yet

been retested.

Growth in 2% NaCl or vancomycin.

Growth in media supplemented

with 2% NaCl has been used as a characteristic to separate B. popilliae from B. lentimorbus.

Although we found this to be an

accurate indicator of species for most strains tested (Table 2), there were a few exceptions on both solid and liquid media.

Stahly et al. (12) described a selective medium designed for the recovery of B. popilliae spores from soil.

This medium

utilized vancomycin to select for B. popilliae while suppressing growth of B. lentimorbus and many other soil microorganisms. Although they reported that B. popilliae was generally resistant to vancomycin, there were several isolates that appeared to be susceptible. When we examined the response to vancomycin of the strains studied by DNA similarity, it was found that strains

61

identified as B. popilliae were resistant to vancomycin and all strains identified as B. lentimorbus were sensitive (Table 4). The strains of B. popilliae that Stahly et al. (12) reported as being sensitive to the antibiotic were found to be B. lentimorbus by DNA similarity and one B. lentimorbus strain, BlPj1, that Stahly et al. reported to be resistant, we have found to be B. popilliae.

Table 2.

Phenotypic characteristics of Bacillus popilliae and

Bacillus lentimorbus strains used in DNA similarity studies

Refer to Rippere et al. 1998.

Molecular systematics of Bacillus

popilliae and Bacillus lentimorbus, bacteria causing milky disease in Japanese beetles and related scarab larvae.

Int. J.

Syst. Bacteriol. 48:395-402.

When the MIC’s were determined for the type strains, B. popilliae was found to be highly resistant, MIC’s ranging from 400 to 800 µg/ml, whereas B. lentimorbus was sensitive to