SYSTEMATICS OF THE ENTOMOPATHOGENIC BACTERIA BACILLUS
129
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
Transcript of SYSTEMATICS OF THE ENTOMOPATHOGENIC BACTERIA BACILLUS
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. Thirty-
four 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
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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. An
amplicon was identified and sequenced. 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
PageCHAPTER THREE
Figure 1. Distance dendogram of B. popilliae andB. lentimorbus strains generated from DNAsimilarityanalysis.......................................62
CHAPTER FOURFigure 1. RAPD banding patterns of B. popilliae
and B. lentimorbus isolates using primerOPA-03.........................................70
Figure 2. RAPD banding patterns of B. popilliaeand B. lentimorbus isolates using primerOPA-03.........................................71
Figure 3. RAPD banding patterns of B. popilliaeand B. lentimorbus isolates using primer OPA-15........................................72
Figure 4. RAPD banding patterns of B. popilliaeand B. lentimorbus isolates using primer OPA-15........................................73
Figure 5. Dendogram showing the relationships betweenstrains of B. popilliae and B. lentimorbusgenerated from RAPD analysis...................74
CHAPTER FIVEFigure 1. Structure of the Bacillus popilliae
cry operon.....................................83Figure 2. ATCC 14706 and NRRL B-4081 PCR products
using primer pair CryBp2......................86
Figure 3. B. popilliae cry18Aa1 genesequences......87
Figure 4. Deduced amino acid sequence comparison ofB. popilliae cry genes.........................89
CHAPTER SIXFigure 1. Multiplex PCR-RFLP of enterococcal isolates
carrying the vanA and vanB ligase genes andB. popilliae ATCC 14706........................96
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Figure 2. Sequence comparisons of the putativeligase genes in B. popilliae isolates.........97
Figure 3. Comparison of the translation of theputative ligase gene in B. popilliae ATCC14706 to the translations of four previouslycharacterized vanB genes(isolates 55, 94, 45, and 91) and one vanA gene (isolate)..........98
Figure 4. Southern blot of digested andundigested B. popilliae chromosomal DNA probedwith the vanE PCR product....................100
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LIST OF TABLES
Page
CHAPTER TWOTable 1. B. popilliae and B. lentimorbus strains
used in DNA-DNA reassociation.................36
Table 2. B. popilliae and B. lentimorbus strainsused in RAPD analysis from diverse host insectsand geographic regions.........................36
Table 3. RAPD primersequences......................45
Table 4. Dilution series for probeestimation.......49
Table 5. Primer sequences used in multiplex PCR-RFLPreaction for detection of van ligase genes inenterococci....................................53
Table 6. Primer sequences used for detection of crygenes in B. popilliae and B. lentimorbus.......55
CHAPTER THREETable 1. Levels of DNA similarity between B.
popilliae and B. lentimorbus as determined bythe S1 nuclease method.........................61
Table 2. Characteristics of B. popilliae and B.lentimorbus strains used in DNA similaritystudies........................................63
CHAPTER FOURTable 1. Characteristics of B. popilliae and B.
lentimorbus isolates from diverse host insectsand geographicalregions...........................77
CHAPTER FIVETable 1. Detection of the paraspore crystal in
strains of B. popilliae and B. lentimorbus byvisualization and PCR..........................84
CHAPTER SEVENTable 1. Bacillus sphaericus strains studied using
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DNA-DNA similarity analysis..................107
Table 2. Levels of DNA similarity among strains ofB. sphaericus................................109
1
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.
3
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. Milky
disease can be said to occur in four stages. 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,
8
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 Entner-
Dudoroff 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
10
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. Both species contain plasmids of various sizes
and number. 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
12
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
13
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. Parasporal protein
fed orally to the larvae was nontoxic (99). 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). Gordon
14
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).
15
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
16
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
17
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
18
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
19
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-N-
acetylmuramylpentapeptide 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).
20
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
21
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).
22
vanH encodes a dehydrogenase which converts pyruvate to D-
lactate, 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). VanA ligates D-
alanine to the D-lactate produced by VanH. 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
23
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,d-
carboxypeptidases (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 D-
alanyl-D-alanine ligases of E. coli (31). However, as opposed
to VanA which ligates D-alanine and D-lactate, VanC-1 ligates D-
24
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).
25
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34
CHAPTER TWO
Materials and Methods
Media and Reagents
MYPGP broth 1.5% yeast extract, 1.0% Mueller-
Hinton 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 1:5 dilution of HCl buffer
S1 storage buffer 20 mM Tris, pH 7.5, 50 mM NaCl, 0.1 mM
ZnCl2, 50% glycerol
High Salt buffer 13.2X SSC, 5 mM HEPES, pH 7.0
50X TAE 2.0 M Tris base, 57.1 ml/L glacial acetic
acid, 100 ml/L 0.5 M EDTA (pH 8.0)
35
Gel loading dye 30% sucrose in TE with bromophenol blue
10X TNE 100 mM Tris-HCl, 10 mM EDTA, 2.0 M NaCl,
pH 7.4
10X buffer (Promega) 500 mM KCl, 100mM Tris-HCl (pH 9.0), 1%
Triton X-100
dNTP mixture 2.5 mM each dATP, dCTP, dTTP, dGTP
MgCl2 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% N-
lauroylsarcosine, 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 (X-
phosphate, 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). One-
liter erlenmeyer flasks containing 250 ml of media were inoculated
36
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+ Popillia japonica USA1
ATCC 14707* Popillia japonica USA1
BlPj1+ Popillia japonica USA6
Bp1* Papuana woodlarkiana Papua New
Guinea2
Bp6+ Popillia japonica USA2
Bp9+ Ataenius spretulatus USA, NY2
Bp10+ Anomala flavipennis USA, NC2
Bp11* USA2
Bp12+ Holotichia oblita China2
Bp13+ Popillia japonica Australia2
Bp14+ Cyclocephala hirta USA, CA2
37
Bp15* Cyclocephala lurida USA, TX2
Bp16* Polyphyla comes USA, NC2
Bp17* Phyllophaga crinita USA, TX2
Bp18* Anomala diversa Japan2
Bp19* Rhopaea morbillosa Australia2
Bp21* USA, TN2
Bp22+ Phyllophaga sp. Panama2
Bp23++ Popillia japonica USA2
Bp25* Cyclocephala hirta USA, NY2
Bp26* Cyclocephala parallela USA, FL2
BpF+ Europe5
BpCb1* Cyclocephala borealis USA6
BpPa1* Phyllophaga anxia USA6
BpPj1+ 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+ Popillia japonica USA4
NRRL B-4081+ Melolontha melolonthae Europe4
NRRL B-4145+ USA4
NRRL B-4154+ Odontria (strain Odontria) USA4
RM23+ Anoplognathus porosus Australia3
RM29+ Lepidiota picticollis Australia3
1ATCC, 2Klein, 3Milner, 4Nakamura 5Schnetter, 6Stahley *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
38
Hemolymph samples were received as a dried film coating a
microscope slide. Ten microliters of sterile distilled water were
added to a spot on the slide to allow resuspension of the dried
spores. The water was 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. A
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
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
39
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
40
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.125I 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, NAP-
25 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
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 NAP-
25 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
41
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
42
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. The washes were also
added to the digestion tube. 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,
43
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). As
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
44
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 5’-AATCGGGCTG-3’
OPA-05 5’-AGGGGTCTTG-3’
OPA-07 5’-GAACGGGGTG-3’
OPA-08 5’-GTGACGTAGG-3’
OPA-10 5’-GTGATCGCAG-3’
OPA-11 5’-CAATCGCCGT-3’
OPA-15 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.
45
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. Samples were overlayed with two
drops sterile mineral oil. 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
46
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. Each reaction was
overlaid with 2 drops sterile mineral oil.
The reactions were placed in the thermalcycler with 1 drop
mineral oil in each well. The temperature profile was as follows:
47
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
48
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
49
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 anti-
DIG 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
50
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
51
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. Primer sequences are shown in Table 5.
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. Upon cycle completion the reactions were held at
4oC. 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. The tubes were
52
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 Expected
53
size
cryBP1-F 5’-AGGGAATGGACAGAATGG-3’ 1058 962 bp
cryBP1-R 5’-GAAAGCTGAACGCCAATC-3’ 2020
cryBP2-F 5’-AGGATGTTCCTCCGATCCCCATCAC-3’ 441 806 bp
cryBP2-R 5’-GTTCCGTGGCTCGTAAAATCTCTTC-3’ 1247
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.
54
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
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1994. Modification of peptidoglycan precursors is a common
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Lactobacillus casei, Pediococcus pentosaceus, Leuconostoc mesenteroides,
and Enterococcus gallinarum. J. Bacteriol. 176(8):2398-2405.
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3. Handwerger, S. 1994. Alterations in peptidoglycan precursors and
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Enterococcus faecalis 221. Antimicrob. Agent. Chemother. 38(3):473-
475.
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5. Patel, R., J. R. Uhl, P. Kohner, M. K. Hopkins, and F. R. Cockerill, III.
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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
analysis system version 1.80. Exeter Publishing, Setauket, NY.
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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.
57
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
58
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. Serological
studies prompted 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
59
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. The
distance-based phenogram will be used here because it showed
higher resolution within the groups. The similarity study
revealed the existence of two groups
60
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.
61
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
62
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 <1
µg/ml. All three strains were sensitive to the related
glycopeptide antibiotic, teicoplanin.
Discussion
DNA similarity analysis was used to elucidate the genetic
relationship between 34 isolates of bacteria causing milky
disease in scarab larvae. The strains separated into two species
based on greater than 70% similarity to the type strains of the
species (4, 10, 13). Twenty four were shown to be B. popilliae
by their relatedness to the type strain, ATCC 14706T, and eight
were shown to be B. lentimorbus by their relatedness to ATCC
14707T. Strains NRRL B-2522 and BlPj1 were received as B.
63
lentimorbus but were found to be most closely related to B.
popilliae. Both strains grew with 2% NaCl in the medium (NRRL B-
2522 only in broth) and both were resistant to vancomycin,
traits that are associated with B. popilliae. The European
isolate referred to as B. popilliae var. melolonthae (NRRL B-
4081) and the North American isolate Bp3 had lower DNA
similarity to the B. popilliae type strain than the remaining
isolates of this species. The main body of B. popilliae
isolates showed less than 70% similarity to NRRL B-4081,
suggesting that these strains may constitute a subspecies of B.
popilliae. The vancomycin resistance of these two strains
points to their relationship to B. popilliae, however, DNA
similarity clearly indicates their uniqueness.
Of the eight strains showing greater than 70% simlarity to
the B. lentimorbus type strain, seven had been received as B.
popilliae. Only one of these, BpPa1, grew with 2% NaCl in the
medium (and then only on plates), and all of them were sensitive
to 150 µg/ml vancomycin. Six of these strains displayed
parasporal bodies when retested by infecting Japanese beetle or
masked chafer larvae. Although the presence of a parasporal
body has been used as a distinguishing characteristic of B.
popilliae, I have shown that paraspores may also be formed by B.
lentimorbus. It appears that the paraspore-forming isolates may
constitute a distinct subgroup of this species. The strains that
were received as B. popilliae but that are now known to be B.
lentimorbus had lower similarity to the type strain (73 to 78 %)
than KLN2 (90%) received as B. lentimorbus. It is noteworthy
that all of the isolates of the second subgroup were isolated
from insects other than Popillia japonica.
64
The observation that vancomycin resistance is a uniform
trait among strains of B. popilliae, as that species is defined
by the DNA similarity, offers a simple phenotypic test for
identifying the species. This test appears to be more reliable
than growth in the presence of 2% NaCl.
This study focused mainly on North American isolates that
were available in pure culture or that I was able to recover
from larval material supplied to me. I have not examined A2 or
B2 isolates, and these may reveal further diversity among the
milky disease bacteria. There are also some strains that have
been described in the literature solely on their appearance in
infected larval hemolymph but which have not been grown in
vitro. It would be of value to be able to examine their
relationships to the better known strains. An understanding of
the genetic relationships among these bacteria and the discovery
of subgroups within the species may provide insight into the
specificity which these bacteria exhibit in their infection of
various species of scarab larvae.
References
1. Bulla, L. A., Jr., R. N. Costilow, and E. S. Sharpe. 1978.
Biology of Bacillus popilliae. Adv. Appl. Microbiol. 2:1-
18.
2. Dutky, S. R. 1940. Two new spore-forming bacteria causing
milky diseases of Japanese beetle larvae. J. Agri. Res.
61(1):57-68.
3. Gordon, R. E., W. C. Haynes, and C. H.-P. Pang. 1973. The
Genus Bacillus, vol. 427. U. S. Department of Agriculture,
Washington, D. C.
65
4. Johnson, J. L. 1973. Use of nucleic-acid homologies in the
taxonomy of anaerobic bacteria. Int. J. Syst. Bacteriol.
23:308-315.
5. Klein, M. G. 1981. Advances in the Use of Bacillus
popilliae for Pest Control, p. 183-192. In H. D. Burges
(ed.), Microbial Control of Pests and Plant Diseases 1970-
1980, 1 ed. Academic Press, London.
6. Krywienczyk, J., and P. Luthy. 1974. Serological
relationship between three varieties of Bacillus popilliae.
J. Invertebr. Pathol. 23:275-279.
7. Milner, R. J. 1981. Identification of the Bacillus
popilliae group of Insect Pathogens, p. 45-59. In H. D.
Burges (ed.), Microbial Control of Pests and Plant Diseases
1970-1980, 1 ed. Academic Press, London.
8. Splittstoesser, C. M., and C. Y. Kawanishi. 1981. Insect
diseases caused by bacilli without toxin mediated
pathologies, p. 190-199. In E. W. Davidson (ed.),
Pathogenesis of Invertebrate Microbial Diseases. Allanheld,
Osmun and Company.
9. St. Julian, G., and L. A. Bulla, Jr. 1973. Milky disease,
p. 57-87. In T. C. Cheng (ed.), Current Topics in
Comparative Pathobiology. Academic Press, New York.
10. Stackebrandt, E., and B. M. Goebel. 1994. Taxonomic Note: A
Place for DNA-DNA Reassociation and 16S rRNA Sequence
Analysis in the Present Species Definition in Bacteriology.
Int. J. Syst. Bacteriol. 44(4):846-849.
11. Stahly, D. P., R. E. Andrews, and A. A. Yousten. 1992. The
genus Bacillus-Insect pathogens, p. 1029-2140. In A.
Balows, H. G. Truper, M. Dworkin, W. Harder, and K.-H.
Schleifer (ed.), The Prokaryotes, 2 ed, vol. 2. Springer-
Verlag, New York.
66
12. Stahly, D. P., D. M. Takefman, C. A. Livasy, and D. W.
Dingman. 1992. Selective medium for quantitation of
Bacillus popilliae in soil and in commercial spore powders.
Appl. Environ. Microbiol. 58(2):740-743.
13. Ursing, J. B., R. A. Rossello-Mora, E. Garcia-Valdes, and
J. Lalucat. 1995. Taxonomic note: A pragmatic approach to
the nomenclature of phenotypically similar genomic groups.
Int. J. Syst. Bacteriol. 45(3):604.
14. Wyss, C. 1971. Sporulationsversuche mit drei varietaten von
Bacillus popilliae Dutky. Zentralbl. Bakteriol. Parasitenk.
Infektionskr. Hyg. II. 126:461-492.
67
CHAPTER FOUR
Randomly Amplified Polymorphic DNA Analysis of Geographically
Distinct Isolates of Bacillus popilliae and Bacillus lentimorbus
Abstract
Geographically distinct strains of Bacillus popilliae and
Bacillus lentimorbus were analyzed using randomly amplified
polymorphic DNA (RAPD). Eight decamer primers were tested
against nineteen new and seventeen previously described
isolates. Of the new isolates, eight were found to belong to the
B. popilliae group containing the type strain ATCC 14706T while
two Australian strains grouped with the subgroup of B. popilliae
containing isolate NRRL B-4081. Nine isolates belong to the B.
lentimorbus species with two isolates in the non-crystal forming
subgroup and seven contained within the subgroup of crystal
forming B. lentimorbus. Vancomycin resistance and 2% NaCl
tolerance were tested for all new isolates.
Thirty-four isolates of B. popilliae and B. lentimorbus
were previously studied by both DNA-DNA similarity and RAPD
analysis, however all but two of those strains were obtained
from the northeastern United States (4). I was interested in
determining whether isolates from a wider range of geographical
regions would reveal the existence of strains of milky disease
bacteria with greater diversity. I have investigated nineteen
geographically diverse isolates using RAPD analysis.
Results
RAPD Analysis. Nineteen geographically diverse milky disease
isolates were tested using eight decamer RAPD primers. Also
68
included in the analysis were seventeen strains that had been
included in a RAPD study performed by M. Tran (4). These 17
isolates were chosen to include representatives from each
possible subgroup suggested by Tran's RAPD analysis. All of the
new isolates fell within the previously described species B.
popilliae and B. lentimorbus. Examples of RAPD banding patterns
are shown for primers OPA-03 and OPA-15 in Figures 1-4.
Primer OPA-03. Negative control reactions containing no
template DNA were run for each primer. Bands appearing in the
negative control reactions were compared to the test reactions
and any band equal in size to a negative control band was not
included in the analysis. Primer OPA-03 generated 15 bands of
different size. All B. popilliae strains except NRRL B-4081,
RM23 and RM29 (Fig. 2, Lanes 12, 14, and 15) had an intense band
of approximately 750 bp that was absent in the B. lentimorbus
strains. RM23 and RM29 had identical banding patterns (OPA-03)
with the exception of one small band found in RM29. However,
the banding patterns obtained from these strains using the other
primers indicated that they are not identical. Primer OPA-03
generated two major bands with strain NRRL B-4081 (Figure 2,
lane 12) which were shared with some B. popilliae (Fig. 1) and
some B. lentimorbus (Fig. 2) isolates. Bacillus lentimorbus
strains ATCC 14707, Bp11 and Bp21 all non-crystal forming
isolates shared a band of approximately 1.2 kb (Figure 2, lanes
3,7, and 10) which was not found in any other B. lentimorbus
isolate.
Primer OPA-15. Primer OPA-15 generated a total of 20 bands
of different sizes. All of the B. popilliae (Fig. 3) isolates
with the exception of Bp22, NRRL B-4081 and RM29 (Fig. 4, Lanes
12, 14, and 15) shared a distinct band of approximately 1.4 kb
69
which was not found in the B. lentimorbus (Fig. 2) strains.
Isolate Bp22 (Fig. 3, Lane 19)
Figure 1. RAPD banding patterns of B. popilliae isolates using
primer OPA-03. Lane 1, 1 kb ladder; 2, ATCC 14706; 3, DNG2; 4,
DNG11; 5, DNG4; 6, NRRL B-2524; 7, KLN3; 8, BlPj1; 9, KLN1; 10,
NRRL B-4145; 11, BpPj1; 12, NRRL B-4154; 13, Bp6; 14, Bp9; 15,
Bp10; 16, Bp12; 17, Bp13; 18, Bp14; 19, Bp22; 20, Bp23; 21, 1 kb
ladder.
70
Figure 2. RAPD banding patterns of B. popilliae (lanes 2, 12,
14 and 15)and B. lentimorbus isolates (lanes 3-11, 13, and 16-
18) using primer OPA-03. Lane 1, 1 kb ladder; 2, BpF; 3, ATCC
14707; 4, Bp1; 5, BpCb1; 6, BpPa1; 7, Bp11; 8, Bp15; 9, Bp19;
10, Bp21; 11, Bp26; 12, NRRL B-4081; 13, Bp25; 14, RM23; 15,
RM29; 16, Bp16; 17, Bp17; 18, Bp18; 19, negative control; 20, 1
kb ladder.
71
Figure 3. RAPD banding patterns of B. popilliae isolates using
primer OPA-15. Lane 1, 1 kb ladder; 2, ATCC 14706; 3, DNG2; 4,
DNG11; 5, DNG4; 6, NRRL B-2524; 7, KLN3; 8, BlPj1; 9, KLN1; 10,
NRRL B-4145; 11, BpPj1; 12, NRRL B-4154; 13, Bp6; 14, Bp9; 15,
Bp10; 16, Bp12; 17, Bp13; 18, Bp14; 19, Bp22; 20, Bp23; 21, 1 kb
ladder.
72
Figure 4. RAPD banding patterns of B. popilliae (lanes 2, 12,
14 and 15) and B. lentimorbus isolates (lanes 3-11, 13, and 16-
18) using primer OPA-15. Lane 1, 1 kb ladder; 2, BpF; 3, ATCC
14707; 4, Bp1; 5, BpCb1; 6, BpPa1; 7, Bp11; 8, Bp15; 9, Bp19;
10, Bp21; 11, Bp26; 12, NRRL B-4081; 13, Bp25; 14, RM23; 15,
RM29; 16, Bp16; 17, Bp17; 18, Bp18; 19, 1 kb ladder; 20,
negative control.
73
gave a unique banding pattern not shared by any other strain
tested with this primer. This isolate did share a band of 450
bp with all of the B. lentimorbus (Fig. 4) strains. Primer OPA-
15 was not able to distinguish among the crystal forming (Fig.
4, Lanes 4-6, 8, 9, 11, 13, 16-18) and the non-crystal forming
(Fig. 4, Lanes 3, 7, 10) B. lentimorbus isolates.
Figure 5. Dendogram illustrating the relationships between
strains of B. popilliae and B. lentimorbus generated from RAPD
analysis.
74
Analysis. The bands derived from each primer were scored
as present or absent for each isolate and combined into a
matrix. The matrix was analyzed using the NTSYS-pc computer
program (5). A dendogram was generated using the Jaccard
coefficient (Figure 5). The analysis of the RAPD bands for each
strain identified nine B. lentimorbus isolates and ten B.
popilliae strains among the 19 new isolates. The two Australian
isolates, RM23 and RM29 were most closely related to the
European isolate, NRRL B-4081, described in the previous
chapter. There are no apparent groupings according to host
insect or the geographic origin of the isolates.
Growth in 2% NaCl or vancomycin. Each isolate was tested
for growth on MYPGP plates containing 2% NaCl and MYPGP plates
containing 150 µg/ml vancomycin. In addition, each isolate was
screened in a PCR based assay for the presence of the vanE
ligase gene (refer to Chapter 6). The vanE gene is related to
the vanA and vanB ligases found in enterococci. The van
ligases, along with several other van gene products confer
resistance to vancomycin. These results are shown in Table 1.
Table 1. Characteristics of B. popilliae and B. lentimorbus
isolates from diverse host insects and geographical regions.
Strain 2% NaCl3 VmR4 vanE5
ATCC 147061 + + +
Bp231 + + +
BlPj11 + + +
NRRL B-41451 +/- + +
KLN11 + + +
BpPj11 + + +
NRRL B-41541 +/- + +
75
Strain 2% NaCl3 VmR vanE5
DNG21 + + +
Bp121 - + +
Bp131 + + +
Bp141 + + +
DNG111 + + +
DNG41 + + +
NRRL B-25241 + + +
Bp91 + + +
Bp101 + + +
BpF1 - + +
KLN31 + + +
Bp61 + + +
Bp221 + - ND6
NRRL B-40811 - + +
RM231 - - -
RM291 - - -
ATCC 147072 - - -
Bp112 - - -
Bp212 - - -
Bp12 - - -
BpCb12 - - -
BpPa12 V7 - -
Bp152 - - -
Bp262 - - -
Bp192 - - -
Bp252 - + +
Bp162 - + +
Bp172 - + +
Bp182 - - -
1 B. popilliae
76
2 B. lentimorbus
3 Growth on MYPGP plates supplemented with 2% NaCl
4 Growth on MYPGP plates supplemented with 150 µg/ml
vancomycin
5 vanE gene detected using PCR assay
6 ND = not determined
7 V = variable reactions
When tested for growth on 2% NaCl eighteen out of twenty-
three B. popilliae strains were capable of growing on MYPGP
plates supplemented with 2% NaCl. Strains Bp12, BpF, RM23, RM29
and NRRL B-4081 were unable to tolerate the NaCl. Three of the
five strains (RM23, RM29, NRRL B-4081) are found on a branch of
the dendogram that is distinct from the majority of the B.
popilliae species. These three isolates form a subgroup of B.
popilliae that is almost as different from B. popilliae as B.
lentimorbus is different from B. popilliae. Twelve out of
thirteen B. lentimorbus strains were negative for growth in 2 %
NaCl. Strain BpPa1 had variable reactions to the NaCl
concentration.
Twenty of twenty-three B. popilliae isolates (Bp22, RM23
and RM29 were negative) tested positive for growth on MYPGP
supplemented with 150 µg/ml vancomycin. The vanE ligase gene was
undetectable in these three strains using the PCR assay
developed for that purpose. RM23, RM29 and Bp22 are distant
from the majority of the B. popilliae strains in the dendogram
generated from the RAPD data. Ten out of thirteen B.
lentimorbus strains were vancomycin sensitive; Bp16, Bp17 and
Bp25 were able to grow on MYPGP plates containing 150 µg/ml
77
vancomycin and the vanE ligase gene was detected in these
strains using the PCR.
Discussion
The inclusion of nineteen new B. popilliae and B.
lentimorbus strains in a RAPD analysis with seventeen strains
that had been previously analyzed by DNA simlarity and RAPD
analysis resulted in the identification of ten B. popilliae and
nine B. lentimorbus isolates. Two isolates formed a cluster on
the dendogram with the B. popilliae NRRL B-4081 subgroup while
eight isolates were found in a cluster with the major B.
popilliae group. Seven strains (Bp15 through Bp18 in Figure 5)
analyzed for the first time were most closely related to the
crystal-forming strains of B. lentimorbus (Bp1, BpCb1, and
BpPa1) that had previously been studied by DNA similarity and by
RAPD. Two (Bp11 and Bp21) were most closely related to the B.
lentimorbus type strain. The strains analyzed in this study are
diverse both with respect to geographic origin and host insect.
There were no apparent grouping patterns relating to either
geographic origin or host insect in either species. At present,
the data supports the classification of milky disease organisms
into two species differentiated only by their reactions to 2 %
NaCl and vancomycin (4). Combining strains that had been
previously analyzed by the RAPD technique with new strains also
being tested by RAPD analysis appears to be of use in
integrating two separate studies. One strain representing each
terminal group of the previous RAPD study was retested in this
study. These seventeen strains retained their original
placement in the dendogram generated by this study, indicating
that the placement of the nineteen new strains is most likely
accurate within the context of the original study (4).
78
Results for two of the eight primers used in the RAPD
analysis were shown in Figures 1-4. These primers were chosen
as representative of the type of banding patterns obtained from
these bacteria. It may be possible to use a RAPD band generated
by one of these primers to distinguish groups within these
species. For example, the non crystal-forming isolates of B.
lentimorbus share a band of 1.2 kb (OPA-03) which could be used
to identify them. It is possible that a RAPD band common to all
the milky disease bacteria could be used to produce a probe for
their rapid identification or that a probe could be produced to
distinguish between species.
Bacillus popilliae has been distinguished from B.
lentimorbus on the basis of the ability to grow in the presence
of 2% NaCl and formation of a parasporal crystal during
sporulation (1). Resistance to vancomycin also appears to be
useful in distinguishing between the two species (4). The
nineteen geographically distinct strains were tested for these
characteristics. All of the newly identified B. lentimorbus
strains were negative for growth in the presence of 2 % NaCl.
However, B. popilliae strains Bp12, BpF, NRRL B-4081, RM23 and
RM29 were also negative. According to the RAPD results, NRRL B-
4081, RM23 and RM29 are more closely related to each other than
to the other B. popilliae strains but Bp12 and BpF are closely
related to other B. popilliae strains that test positive for
growth in 2% NaCl. Rm23 and RM29 were isolated from
Anoplognathus porosus and Lepidiota picticollis, respectively,
in Australia, Bp12 was isolated from Holotrichia oblita in
China, NRRL B-4081 was isolated from Melolontha melolontha in
Europe and BpF was isolated in Europe (insect unknown). The
ability to grow in 2% NaCl appears to be useful in
79
differentiating the species since a high percentage of strains
possess this characteristic while it is lacking in the majority
of B. lentimorbus strains.
When tested for vancomycin resistance, all of the B.
popilliae strains were positive for growth on plates
supplemented with the antibiotic and positive for the presence
of the vanE gene except isolates Bp22, RM23 and RM29. Bp22 was
isolated from a Phyllophaga sp. in Panama, and appears on the
RAPD dendogram as a separate cluster. It is almost as
dissimilar to the majority of the B. popilliae isolates as
strains NRRL B-4081, RM23 and RM29. It is possible that these
four represent distinct varieties of the species although
additional isolates would be required to verify this suggestion.
Six of the nine B. lentimorbus isolates studied by RAPD tested
negative for resistance to vancomycin. Bp16, Bp17 and Bp25 were
able to grow on plates supplemented with vancomycin and all
contain the vanE gene within their genome. These three strains
form a single cluster of the dendogram within the B. lentimorbus
species and appear to be fairly similar to one another. Bp16
was isolated from Polyphyla comes in North Carolina, Bp17 was
isolated from Phyllophaga crinita in Texas and Bp25 was isolated
from Cyclocephala parallela in Florida, showing no pattern with
regard to either geography or host insect. When both growth in
2% NaCl and vancomycin resistance are considered,
differentiation of all of the strains except Rm23 and RM29 is
possible. These two characteristics in combination appear to be
sufficient for identification of milky disease bacteria as
either B. popilliae or B. lentimorbus.
REFERENCES
80
1. Dutky, S. R. 1940. Two new spore-forming bacteria causing
milky diseases of Japanese beetle larvae. J. Agri. Res.
61(1):57-68.
2. Krywienczyk, J., and P. Luthy. 1974. Serological
relationship between three varieties of Bacillus popilliae.
J. Invertebr. Pathol. 23:275-279.
3. Milner, R. J. 1981. Identification of the Bacillus
popilliae group of Insect Pathogens, p. 45-59. In H. D.
Burges (ed.), Microbial Control of Pests and Plant Diseases
1970-1980, 1st ed. Academic Press, London.
4. Rippere, K. E., M. T. Tran, A. A. Yousten, K. H. Hilu, and
M. G. Klein. 1998. Bacillus popilliae and Bacillus
lentimorbus, bacteria causing milky disease in Japanese
beetles and related scarab larvae. Int. J. Syst. Bacteriol.
48:395-402.
5. Rohlf, F. J. 1994. NTSYS-pc numerical taxonomy and multi-
variate analysis system version 1.80. Exeter Publishing,
Setauket, NY.
6. White, R. T., and S. R. Dutky. 1940. Effect of the
introduction of milky diseases on populations of Japanese
beetle larvae. J. Econ. Entomol. 33(2):306-309.
7. Wyss, C. 1971. Sporulationsversuche mit drei varietaten von
Bacillus popilliae Dutky. Zentralbl. Bakteriol. Parasitenk.
Infektionskr. Hyg. II. 126:461-492.
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.
81
CHAPTER FIVE
Identification and detection of the cry gene in strains of
Bacillus popilliae and Bacillus lentimorbus
Abstract
An assay for detection of the cry18Aa1 gene was developed 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. Primer CryBp1 products were of the
expected size, however they were not identified by sequencing.
The ATCC 14706T CryBp2 PCR product was sequenced, compared to the
published cry gene sequence and found to vary from the published
sequence. In combination, the primer pairs CryBp1 and CryBp2
are effective at detecting the paraspore gene in most B.
popilliae isolates, but are unable to detect the B. lentimorbus
paraspore gene.
Results
Detection of the cry operon. Two sets of PCR primer pairs were
designed for the cry18Aa1 gene in B. popilliae. The first pair
(CryBp1) amplifies a DNA fragment from position 1058 to position
2020 of the published cry18Aa1 sequence. The second pair
(CryBp2) amplifies from bp 441 to bp 1247 of the published
sequence(1). This fragment begins in the orf1 that precedes the
82
cry18Aa1 gene and ends in the 5’ end of the cry18Aa1 gene
(Figure 1). The PCR products produced by the CryBp2
Figure 1. Structure of the Bacillus popilliae cry18Aa1 operon.
primer pair had fewer non-specific amplification products than
the products gained from the CryBp1 pair. As a result, the ATCC
14706 and NRRL B-4081 PCR products obtained from primer pair
CryBp2 were sequenced and compared with the published sequence
(Figure 3). Primer pair CryBp1 was homologous enough to detect
the cry gene in 31 of 35 B. popilliae isolates (the gene was
undetected in isolates NRRL B-4154, NRRL B-2522, RM23 and RM29),
but was able to identify the gene in only one B. lentimorbus
isolate, DGB1. Primer pair CryBp2 was designed as a result of
the nonspecific amplification resulting in multiple bands that
occurred with the CryBp1 primers. CryBp2 primers were able to
detect the cry operon in 28 of 35 B. popilliae strains. Isolates
BpPj5, Bp3, Bp22, RM23, RM29, NRRL B-2522, and NRRL B-4145
showed no amplification of the operon under the conditions
tested. CryBp2 primers failed to amplify the cry gene of all B.
lentimorbus isolates examined in this study (Table 1).
CryBp1
CryBp2
83
Table 1. Detection of the paraspore crystal in strains of B.
popilliae and B. lentimorbus by visualization and PCR.
Strain Paraspore1 CryBp22
B. popilliae
ATCC 14706T ND3 +
A8 ND +
BlPj1 - +
Bp3 ND -
Bp6 ND +
Bp9 - +
Bp10 + +
Bp12 - +
Bp13 - +
Bp14 + +
Bp22 + -
Bp23 + +
BpCh1 ND +
BpF ND +
BpPj1 ND +
BpPj2 ND +
BpPj3 ND +
BpPj4 ND +
BpPj5 ND -
DNG1 ND +
DNG2 ND +
DNG4 ND +
DNG10 ND +
DNG11 ND +
DNG12 ND +
KLN1 ND +
84
Strain Paraspore1 CryBp2
KLN3 ND +
NRRL B-2309 ND +
NRRL B-2522 + -
NRRL B-2524 ND +
NRRL B-4081 ND +
NRRL B-4145 ND +
NRRL B-4154 ND -
RM23 + -
RM29 + -
B.
lentimorbus
ATCC 14707T ND -
Bp1 + -
Bp7 + -
Bp11 - -
Bp15 + -
Bp16 + -
Bp17 + -
Bp18 + -
Bp19 + -
Bp21 - -
Bp25 + -
Bp26 + -
BpCb1 + -
BpCb2 + -
BpCp1 + -
BpPa1 + -
DGB1 + -
KLN2 ND -
1Paraspore seen in phase contrast microscopic examination of hemolymph
smear
85
2Gene encoding paraspore detected using PCR primer pair CryBp23ND = not determined
The PCR products for strains ATCC 14706 and NRRL B-4081
were chosen for sequencing. The NRRL B-4081 product obtained
from CryBp2 primers was of the expected size (806 bp), however
the product from ATCC 14706 was approximately 140 bp shorter
than expected (Figure 2).
Figure 2. ATCC 14706 and NRRL B-4081 PCR products using primer
pair CryBp2. Lane 1, 1 Kb DNA ladder; Lane 2, ATCC 14706; Lane
3, NRRL B-4081
86
In addition, PCR products obtained from all other isolates using
primer pair CryBp2 were approximately 140 bp smaller than
expected (Figure 2). The sequences obtained from strains ATCC
14706 and NRRL B-4081 using primer pair CryBp2 were compared to
the published sequence(1). The sequence obtained from NRRL B-
4081 was identical to the published sequence, while the sequence
found in ATCC 14706 was somewhat different. The nucleic acid
sequence comparison is shown in Figure 3 and the amino acid
sequence comparison is shown in Figure 4.
453
cry18Aa1 CGATCCCCAT CACAAAGAAA TTTCTATTTG CTGCACAGAA AGTATCTGTA
ATCC 14706
503
cry18Aa1 TAGATCATGT ACTGAAATGC AGTGTGGAAA CCAGCCCCCA TCATCATGTG
ATCC 14706 -------- ---C------ ---------- -------T--
553
cry18Aa1 GACTGCCATC ATGTGGTAGT TCATGATTTG AAAGCAATCC CAATCCGTGA
ATCC 14706 ---G----C- ----A----- -TG------- ---------- ----------
603
cry18Aa1 AGATCATTGC CGGTTCGTTA AAGTTACGGG GAACTTTCAA TTTCATTATG
ATCC 14706 ---G-----T -A---T---- ------T--- ---------- -------G--
653
cry18Aa1 TAAAGGATTT G1TAAACCGAA ACACAGGCTG GAATGACCCG AGGCGATTTG
ATCC 14706 --------C- -1A-T------ -A---T---- -----CT-T- -------GG-
703 RBS
cry18Aa1 GATAGATTTG AATGCTCATC ATATGAAGGA GGCTATTGGT ATG2AACAATA
ATCC 14706 ---C------ -------C-A ----T-G--- ----C--A-- ---2------T
87
753
cry18Aa1 ATTTTAATGG TGGAAATAAT ACAGGAAATA ACTTTACTGG AAATACTCTA
ATCC 14706 -C----T--- :AA--G-::: :::::::::: :TC-A-GC-- -C--CA-A--
803
cry18Aa1 AGCAACGGAA TTTGTACGAA AAAAAATATG AAAGGAACCC TAAGCAGAAC
ATCC 14706 -A---TAACG :::---:::- TGG-----C- :::::::::- ----------
853
cry18Aa1 TGCTATATTT TCAGATGGGA TTAGTGATGA TTTAATTTGT TGTCTAGATC
ATCC 14706 G---:::::: :::::::::: :::::::::: :----C-::: :::::::::-
903
cry18Aa1 CTATATATAA CAATAACGAT AACAATAACG ATGCTATTTG TGATGAGTTA
ATCC 14706 ---C-A---- ----:::-T- --TCG-GGT- --::::--A- -T-C--A-::
953
cry18Aa1 GGTTTAACTC CAATAGATAA CAATACGATA TGCAGTACTG ATTTTACTCC
ATCC 14706 ---------- ---------- ----TTT--- G-T----A-- G-----T---
1003
cry18Aa1 CATAAATGTA ATGAGAACAG ATCCTTTTCG CAAGAAATCA ACACAAGAAC
ATCC 14706 --G------- -C-----A-- ---------- -----G-A-- ---------T
1053
cry18Aa1 TCACAAGGGA ATGGACAGAA TGGAAAGAAA ATAGTCCTTC TTTGTTTACA
ATCC 14706 ---T------ --------- ----------- -A---G---- ----------
1103
cry18Aa1 CCGGCAATTG TAGGTGTCGT TACCAGTTTT CTTCTTCAAT CATTAAAAAA
ATCC 14706 G-AC------ -------TA- -------AC- ------G--G ----------
1153
cry18Aa1 ACAAGCAACT AGCTTTCTTT TAAAAACTTT GACAGACCTA TTATTTCCTA
ATCC 14706 --T--T-G-G G--AGAG--- ---TGT-A-- ----A----T ----------
88
1203
cry18Aa1 ATAACAGT
ATCC 14706 --C-----
Figure 3. B. popilliae cry18Aa1 gene sequence.
- = identical base
: = missing base1End of orf1 (1)2Start codon of cry gene (1)
1
Cry18Aa1 MNNNFNGGNN TGNNFTGNTL SNGICTKKNM KGTLSRTAIF SDGISDDLIC
ATCC 14706 ---Y-I-KVL S-HHINN-GN GN:::::::: ::------:: ::::::::::
51
Cry18Aa1 CLDPIYNNND NNNDAICDEL GLTPIDNNTI CSTDFTPINV MRTDPFRKKS
ATCC 14706 :-T-T:---V -RG-LVTN:: --------F- G-NG-I-R-- T-K-----RT
101
Cry18Aa1 TQELTREWTE WKENSPSLFT PAIVGVVTSF LLQSLKKQAT SFLLKTLTDL
ATCC 14706 ---FI----- ---K-A---- AP----I--T --EA---LVA GRV-MS--N-
151
Cry18Aa1 LFPNNS
ATCC 14706 ------
Figure 4. Deduced amino acid sequence comparison of cry genes.
- = identical amino acid
: = missing amino acid
89
Discussion
Each strain in this study was examined for the presence of
a gene encoding a paraspore protein using two PCR primer pairs.
A hemolymph sample was available for 11 B. popilliae isolates
and these were examined microscopically for the presence of a
parasporal crystal. Four strains, Bp9, Bp12, Bp13, and BlPj1,
lacked a parasporal crystal when examined by phase contrast
microscopy. These four strains were isolated from different
host insects in different geographical regions. The B.
lentimorbus strains tested microscopically for the presence of a
paraspore revealed a distinctive pattern. A subgroup of B.
lentimorbus strains with the ability to produce a paraspore was
identified in the original DNA similarity study (Chapter 3).
This paraspore-forming subgroup of strains did not group with
the type strain of the species, ATCC 14707. The B. lentimorbus
strains able to make a parasporal body were also identified
through use of RAPD (Chapter 4). They formed a subgroup of the
species along with the representative strains from that group
chosen from the RAPD study by M. Tran. These strains included
Bp15 (Cyclocephala lurida, Texas), Bp26 (Cyclocephala parallela,
Florida), Bp19 (Rhopaea morbillosa, Australia), Bp25
(Cyclocephala hirta, New York), Bp16 (Polyphyla comes, North
Carolina), Bp17 (Phyllophaga crinita, Texas) and Bp18 (Anomala
diversa, Japan) representing a wide range of host insects and
geographical regions.
The cry operon was detected in the isolates by use of two
PCR primer pairs, CryBp1 and CryBp2. CryBp1 primers amplify a
fragment that is internal to the cry18Aa1 gene, beginning near
the N-terminal end of the protein. This primer pair gave
several bands in addition to the expected product, and as a
result, this PCR product was not sequenced. This primer pair
90
could still be used to detect the presence of the parasporal
gene in the majority of the B. popilliae strains. These
reactions contained fragments of the expected size, indicating
that they are most likely the correct amplification product,
although this is not certain because the product was not
sequenced. Four strains, NRRL B-2522, NRRL B-4154, RM23 and
RM29 did not produce a PCR product when tested with this primer
pair. This primer pair was also unable to detect the presence of
the parasporal gene in any of the B. lentimorbus strains
identified in this study. This could be due to a change in one
or both of the primer regions causing the primers to be unable
to anneal to the template under the conditions tested. This
could also occur if these strains produce a paraspore protein
with a different amino acid sequence than that found in the B.
popilliae strains in which the paraspore gene was detected.
Primer pair CryBp2 was designed and used to amplify a portion of
orf1, the spacer region and the 5' region of the cry18Aa1 gene.
Primer pair CryBp2 identified the cry18Aa1 gene in 28 of 35
B. popilliae strains and was unable to detect the gene in any of
the B. lentimorbus strains tested. This primer pair failed to
detect the paraspore gene in B. popilliae strains NRRL B-4154,
NRRL B-2522, Bp3, BpPj5, Bp22, RM23 and RM29. Of these strains,
Bp22, BpPj5 and Bp3 tested positive with the other primer pair,
CryBp1, indicating that the paraspore gene may be similar to the
genes in the other B. popilliae strains. The open reading frame
that precedes the cry18Aa1 gene in these strains may vary enough
from the forward primer sequence that the primer was unable to
anneal to the template and produce a reaction product. In
addition, since the gene sequence of only one strain has been
deposited in the databanks, the reverse primer may be in a
region of the paraspore gene that is somewhat more variable than
91
other regions. The four B. popilliae strains RM23, RM29, NRRL
B-2522 and NRRL B-4154, which tested negative with both primer
pairs could contain paraspore genes that vary widely from the
published sequence, preventing the primer from annealing and
detecting the gene. Strains RM23 and RM29 do contain a
paraspore, as determined microscopically, but this was not
determined for NRRL B-4154. It is possible that this strain
does not produce a paraspore. Primer pair CryBp2, did not
produce a product with any of the paraspore-forming B.
lentimorbus strains tested in this study, supporting the idea
that these isolates may have a protein that varies widely or is
entirely different from the protein that Zhang et al. studied
(1).
References
1. 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.
82
CHAPTER SIX
DNA Sequence Resembles vanA and vanB in the Vancomycin-Resistant
Biopesticide Bacillus popilliae.
Abstract
Biopesticidal powders containing spores of vancomycin
resistant Bacillus popilliae have been used for more than 50
years in the United States for suppression of Japanese beetle
populations. The basis for vancomycin resistance in these
bacteria was investigated using a polymerase chain reaction
assay designed to amplify the vanB ligase genes in enterococci.
An amplicon was identified and sequenced. 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 translation of the putative ligase gene in B.
popilliae and the deduced amino acid sequence of the vanA gene
and the vanB genes, respectively. We have identified a gene
resembling vanA and vanB in B. popilliae and determined that it
is located either on a plasmid greater than 16 kb in size or on
the chromosome. Based on sequence similarity, the gene in B.
popilliae may have had an ancestral gene in common with
vancomycin resistance genes in enterococci.
Biopesticidal powders containing spores of Bacillus
popilliae have been used for more than 50 years in the United
States for the suppression of Japanese beetle (Popillia japonica
Newman) populations (2, 5). The Japanese beetle feeds on more
93
than 257 different plants and annually destroys turf, field
crops, fruits and ornamentals worth millions of US dollars.
Bacillus popilliae was the first microorganism registered in the
US as a pesticide (15).
Pridham et al. (11) first observed that B. popilliae NRRL
B-2309 was vancomycin resistant in the 1960’s. Subsequently,
Stahly et al. (14) described several vancomycin-resistant
isolates of B. popilliae and described a selective medium
containing vancomycin, for quantitation of B. popilliae in soil
and in commercial spore powders.
High level vancomycin resistance was first described in
enterococci in isolates from 1986 (6). The genes associated
with high level vancomycin resistance, vanA and vanB, encode a
ligase responsible for the synthesis of the depsipeptide D-
alanyl-D-lactate which is incorporated into a pentapeptide
peptidoglycan cell wall precursor (which terminates in D-alanyl-
D-lactate) to which vancomycin binds poorly. In contrast, in
vancomycin-susceptible cells, vancomycin complexes with the D-
alanyl-D-alanine termini of normal pentapeptide peptidoglycan
cell wall precursor thereby inhibiting cell wall synthesis.
Enterococci cause about five percent of cases of infective
endocarditis in humans (a uniformly fatal illness if untreated)
and are now the second most common pathogens isolated from
hospital acquired infections. Vancomycin-resistant enterococci
(VRE) are increasingly isolated from clinical specimens, and
infections caused by VRE can be untreatable by any currently
available antimicrobial or antimicrobial combination. With the
increasing presence of VRE in clinical specimens, there is
concern regarding the possibility that vancomycin-resistance
94
genes present in VRE will be transferred to other more virulent
gram-positive bacteria. It has been demonstrated, for example,
that in the laboratory, vancomycin resistance is readily
transferred from enterococci to other gram-positive organisms,
including Staphylococcus aureus (7).
The origin of vancomycin resistance genes in enterococci is
unknown. One hypothesis as to their origin is that vancomycin
resistance present in environmental organisms has been
transferred to enterococci, and these transcipients have been
selected under the pressure of increased oral and parenteral
vancomycin usage in clinical practice. Environmental organisms
carrying genes resembling vanA and vanB have not, however, been
identified to date.
I hypothesized that vancomycin resistance in B. popilliae
might be conferred by a gene resembling the vancomycin-
resistance genes in enterococci. Herein we describe the use of a
PCR assay originally designed for use in enterococci to detect a
gene resembling vanA and vanB, by nucleic acid and amino acid
homology studies, in B. popilliae (9).
Results
Previously described multiplex vanA and vanB ligase gene
primers were used to amplify the putative ligase gene of B.
popilliae ATCC 14706 using PCR (9). The PCR product showed a
restriction pattern that was different from those obtained from
enterococcal isolates carrying the vanA and vanB genes (Figure
1). The sequence of the amplicon obtained was compared to that
of four previously characterized enterococcal isolates carrying
the vanB genes (isolates 55, 94, 45, and 91), one previously
95
Figure 1. Multiplex PCR-RFLP of enterococcal isolates carrying
the vanA and vanB ligase genes and B. popilliae ATCC 14706.
Lane 1, 50 bp ladder; lane 2, empty; lane 3, vanA isolate; lane
4, vanB isolate; lane 5, vanC-1 isolate (negative control); lane
6, vanC-2/3 (negative control); lane 7, ATCC 14706; lane 8, B.
lentimorbus ATCC 14707.
96
characterized enterococcal isolate carrying the vanA gene
(isolate 1), five previously characterized enterococcal isolates
carrying the vanC-1 genes, and six previously characterized
isolates carrying the genes vanC-2/3 (10). The 708 bp fragment
amplified from B. popilliae ATCC 14706T (figure 1) had 77%
nucleotide identity to the sequence of the vanA gene (9), and
68-69% nucleotide identity to the sequences of the vanB genes of
isolates 45 and 91 (9). Comparisons of the putative amino acid
sequence of the B. popilliae ATCC 14706 ligase gene to that of
four previously characterized vanB genes and one previously
characterized vanA gene (10) are shown in figure 2. There was
75% identity between the deduced amino acid sequence of the
putative ligase gene in B. popilliae ATCC 14706 and that of the
deduced amino acid sequence of the previously described vanA
gene, and 69-70% identity between the deduced amino acid
sequence of the putative ligase gene in B. popilliae and that of
the translation of the vanB genes in isolates 45 and 91 (9).
When conservatively substituted, non-identical amino acids were
considered, the homology increased to 82% (vanA gene) and 78%
(vanB gene isolate 45). For comparison, there is 73% nucleic
acid and 75% amino acid identity between the vanA gene and the
vanB gene of isolate 91 (10). Notably, there was 44-50%
nucleotide identity between base pairs 135-490 (Figure 2) of the
B. popilliae putative ligase gene and the vanC-1 or vanC-2/3
genes in enterococci or the D-alanine:D-alanine ligase (ddl)
genes in Lactobacillus spp. and Leuconostoc mesenteroides (data
not shown) (9, 1). I have designated the putative ligase gene
in B. popilliae ATCC 14706 “vanE”.
Figure 2. Sequence of the putative ligase gene in B. popilliae
ATCC 14706. Eleven other isolates of B. popilliae had identical
sequences from position 34 through 572 including Bp6, NRRL B-
97
2309, NRRL B-4145, Bp23, BpPj2, BlPj1, BpPj1, BpPj3, BpPj4, KLN1
and BpCh1. The sequences in Bp3 (12 bp different from ATCC
14706) and NRRL B-4081 (11 bp different from ATCC 14706 are
shown). The sequence in BpF (shown) was identical to that in
KLN3, DNG1, DNG2, DNG4, DNG10, DNG11, DNG12, Bp9, Bp10, Bp12,
Bp14, NRRL B-2524, and Bp13 and differed from that in ATCC 14706
by 7 bp. The sequence in NRRL B-2522 was identical to that in
NRRL B-4154 and differed from that in ATCC 14706 by 2 bp. The
sequence in Bp17 (shown) was identical to that in Bp25 and Bp16
and differed from that in ATCC 14706 by 10 bp.
Refer to Rippere et al. 1998. A gene resembling vanA and vanB
in the vancomycin-resistant biopesticide Bacillus popilliae. J.
Infect. Dis. 178:584-588.
Figure 3. Comparison of the deduced amino acid sequence of the
putative ligase gene in B. popilliae ATCC 14706 to the deduced
amino acid sequences of four previously characterized vanB genes
(isolates 55, 94, 45, and 91) and one previously characterized
vanA gene (isolate 1) (10).
Refer to Rippere et al. 1998. A gene resembling vanA and vanB
in the vancomycin-resistant biopesticide Bacillus popilliae. J.
Infect. Dis. 178:584-588.
Patel et al. have previously identified sequence
variability in the van genes of clinical isolates of enterococci
(10). Given this, I hypothesized that there might be sequence
variability in the putative ligase genes of B. popilliae, and
that possibly, in some isolates of B. popilliae, I might find a
ligase gene with a sequence even more similar to the vanA and
98
vanB genes than was found in B. popilliae ATCC 14706. In order
to test this hypothesis, a segment of the putative ligase genes
of a collection of B. popilliae isolates were sequenced as
follows. Based on the sequence of the ligase gene in B.
popilliae ATCC 14706, a set of PCR primers was designed and used
to amplify the putative ligase gene in a collection of 33 B.
popilliae isolates. Six distinct classes of sequences were
found amongst these isolates and are shown in Figure 2. Members
of each class are described in the legend of Figure 2. All had
homology to the vanA and vanB sequences in enterococci, although
none was markedly more similar to the enterococcal van genes
than that found in B. popilliae ATCC 14706.
I attempted to determine the location of the vanE ligase
gene within the B. popilliae genome by probing both isolated
plasmids and isolated large chromosomal fragments. The vanE PCR
product was labeled with digoxygenin and then used to probe
Southern blots of isolated plasmids. There was no apparent
hybridization of the probe with any of the plasmids, but
isolated chromosomal DNA showed a strong positive hybridization
to the probe. The chromosomal DNA was digested with MboII,
cutting at either end of the sequence obtained from the vanE
ligase gene. When this digest was run on a gel and probed, the
positive signal moved from the top of the gel (undigested DNA)
to two bands of approximately 1 kb and 500 bp in size (Figure
4). The 500 bp band is about the expected size of the digested
piece determined from the known sequence. The 1 kb band most
likely contains the part of the gene adjoining the sequenced
portion.
99
Figure 4. Southern blot of digested and undigested B. popilliae
chromosomal DNA probed with the vanE PCR product. Lane 1, 1 kb
DNA ladder; Lane 2, undigested ATCC 14706 chromosomal DNA; Lane
3, ATCC 14706 chromosomal DNA digested with MboII; Lane 4, vanE
PCR product, Lane 5, undigested ATCC 14707 chromosomal DNA; Lane
6, ATCC 14707 chromosomal DNA digested with MboII.
100
Discussion
I have identified a gene resembling vanA and vanB in B.
popilliae. This represents the first detection of vanA- and
vanB-like genes in an organism other than an enterococcus, where
transmission of the gene from an enterococcus was not suspected.
Bacillus popilliae ATCC 14706 is an ATCC type strain which was
isolated from commercial insecticidal spore dust, and first
described in the medical literature in 1961 (4). Furthermore, I
was able to amplify the putative ligase gene from an isolate
(Bp23) held in dried hemolymph since 1945. There is therefore
compelling evidence that the ligase gene present in B. popilliae
was not transferred to this organism from an enterococcus (high
level vancomycin resistance in enterococci was only described in
the late 1980’s). The putative ligase gene present in B.
popilliae has homology to both the vanA and vanB genes raising
the possibility that it may have been an ancestor to the vanA
and vanB genes found in modern clinical isolates of enterococci.
Alternatively, the van genes in enterococci and the putative
ligase gene in B. popilliae may have had a common ancestor or
ancestors. The ligase gene is most likely located on the
chromosome of B. popilliae, possibly on a conjugative
chromosomal element like that found in enterococci with the VanB
phenotype. In addition, the mechanism of resistance in B.
popilliae may be similar to that found in the enterococci,
involving a change from D-alanyl-D-alanine to D-alanyl-D-lactate
in the peptidoglycan. This can be predicted from the similarity
found between the vanE ligase and the vanA and vanB ligases.
B. popilliae spores have been introduced into turf in the
Eastern United States as a biopesticidal powder since the late
101
1930’s. As an example, in a 14-year period, between 1939 and
1952, approximately 83,600 kg of B. popilliae spore powder,
containing a concentration of 1 × 108 spores per g, was applied
to 194,000 different sites in 14 eastern states (US) and the
District of Columbia and to a total of more than 42,000 ha (8).
Commercial production of spore powder began in the mid-1940’s,
and continues today (8). It has been suggested that spread of
B. popilliae spores may have been increased by birds, insects,
skunks, moles and mice (13). Such widespread distribution of
this organism may have provided the opportunity for its contact
with enterococci. In the presence of the increasing use of oral
and parenteral vancomycin in humans since the late 1970’s for
the treatment of Clostridium difficile and methicillin-resistant
staphylococcal infections, respectively, this transfer would
potentially have been facilitated. The use of B. popilliae
biopesticidal preparations in agricultural practice may have had
an impact on bacterial resistance in human pathogens.
References
1. Elisha, B. G. and P. Courvalin. 1995. Analysis of genes
encoding D-alanine:D-alanine ligase-related enzymes in
Leuconostoc mesenteroides and Lactobacillus spp. Gene 152:79-
83.
2. Fleming, W. E. 1968. Biological control of the Japanese
beetle. U. S. Department of Agriculture Technical Bulletin,
Vol. 1383. U. S. Department of Agriculture, Washington, D. C.
3. Gerhardt, P., R. G. E. Murray, W. A. Wood and N. R. Krieg.
1994. Methods for general and molecular microbiology.
American Society for Microbiology, Washington, D. C.
102
4. Haynes, W., G. St. Julian, M. Shekelton, H. Hall and H.
Tashiro. 1961. Preservation of infectious milky disease
bacteria by lyophilization. J. Insect Pathol. 3:55-61.
5. Klein, M. G. 1988. Pest management of soil-inhabiting insects
with microorganisms. Agric. Ecosyst. Environ. 24:337-349.
6. Leclercq, R., E. Derlot, J. Duval and P. Courvalin. 1988.
Plasmid-mediated resistance to vancomycin and teicoplanin in
Enterococcus faecium. New Engl. J. Med. 319:157-161.
7. Noble, W. C., Z. Virani and R. G. Cree. 1992. Co-transfer of
vancomycin and other resistance genes from Enterococcus
faecalis NCTC 12201 to Staphylococcus aureus. FEMS Microbial.
Lett. 72:195-198.
8. Obenchain, F. D. and B. J. Ellis. 1990. Safety
considerations in the use of Bacillus popilliae, the milky
disease pathogen of Scarabaeidae, pp. 189-201. In M. Laird,
E. Lacey and E. Davidson (ed), Safety of microbial
insecticides. CRC Press, Boca Raton, FL.
9. Patel, R., J. R. Uhl, P. Kohner, M. K. Hopkins and F. R.
Cockerill. 1997. Multiplex polymerase chain reaction
detection of vanA, vanB, vanC-1 and vanC-2/3 genes in
enterococci. J. Clin. Microbiol. 35:703-707.
10. Patel, R., J. R. Uhl, P. Kohner, M. K. Hopkins, J. M.
Steckelberg, B. Kline and F. R. Cockerill. 1998. DNA
sequence variation within vanA, vanB, vanC-1 and vanC-2/3
genes of clinical Enterococcus spp. isolates. Antimicrob.
Agent. Chemother. 42:202-205.
11. Pridham, T. G., H. H. Hall and R. W. Jackson. 1965. Effects
of antimicrobial agents on the milky disease bacteria
Bacillus popilliae and Bacillus lentimorbus. Appl.
Microbiol. 13:1000-1004.
12. Rippere, K. E., M. T. Tran, A. A. Yousten, K. Hilu and M.
Klein. Bacillus popilliae and Bacillus lentimorbus,
103
bacteria causing milky disease in Japanese beetles and
related scarab larvae. Int. J. Syst. Bacteriol. In press.
13. St. Julian, G. and L. A. Bulla. 1973. Milky Disease, pp.
57-87. In T. C. Cheng (ed) Current topics in comparative
pathobiology. Academic Press Inc.
14. Stahly, D. P., D. M. Takeman, C. A. Livasy and D. W.
Dingman. Selective medium for quantitation of Bacillus
popilliae in soil and in commercial powders. Appl.
Environ. Microbiol. 58:740-743.
15. 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. 179:4336-4341.
104
CHAPTER SEVEN
DNA Similarities among Mosquito-Pathogenic and Nonpathogenic
Strains of Bacillus sphaericus.
Abstract
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.
Aerobic bacilli that form shperical endospores are common
in soil and water and are usually classified as Bacillus
sphaericus. There are few useful phenotypic tests for
identification of these bacteria. Spore morphology combined
with negative reactions in tests for fermentation products and
extracellular enzymes have been the basis for taxonomic
placement. The species was found to be comprised of at least
five distinct homology groups, each sufficiently separated from
the others to merit species status (5). Representative strains
of the homology groups have also been examined by rRNA gene
restriction fragment length polymorphisms analyses (ribotyping),
and these analyses confirmed that there are distinct groups
within the B. sphaericus complex (2). Recently, randomly
amplified polymorphic DNA analysis has also clearly
distinguished the groups originally identified by DNA similarity
analysis (9). These five groups have not been designated
105
separate species because of the lack of readily utilizable
phenotypic tests to distinguish them.
In the original study of Krych et al. (5), group II was
divided into two subgroups based on levels of DNA similarity and
DNA heteroduplex stability. It was of considerable interest
that all of the isolates in group IIA were pathogenic for
mosquito larvae. No mosquito pathogens were found in any other
group. These bacteria are pathogenic because they produce one
or more of four toxins, a binary toxin composed of two distinct
proteins and three additional toxins designated Mtx, Mtx2, and
Mtx3 (6-8), Strains that produce the binary toxin are highly
toxic (50% lethal concentrations, around 102 to 103 cells ml-1),
and strains that produce only toxins Mtx, Mtx2, and Mtx3 have
low toxicity (50% lethal concentrations, about 105 to 107 cells
ml-1). It appeared that the group IIA mosquito pathogens might
be designated a separate species. However, only seven
pathogenic isolates were available at the time of the original
DNA similarity study. Now, many more pathogenic isolates from
many geographic locations are available, and although they have
been referred to as group IIA strains on the basis of ribotyping
data, DNA similarity studies have never actually been performed
with them. In this paper we report DNA similarity results for a
large number of strains from diverse geographic locations.
Bacteria and DNA isolation.
The strains of B. sphaericus used in this study are listed
in Table 1 . The bacteria were grown in NY broth (Difco
nutrient broth supplemented with 0.05% yeast extract) at 30oC
with shaking at 150 rpm. Cells were recovered by
centrifugation, suspended in 20 ml of pH 8.0 buffer (10 mM Tris,
106
1.0 mM EDTA, 0.35 M sucrose, 0.1 mg of lysozyme per ml), and
incubated at 37oC for 30 min. A 20-ml portion of lysing solution
(100 mM Tris, 20 mM EDTA, 0.3 M NaCl, 2% [wt/vol] sodium dodecyl
sulfate, 2% [vol/vol] β-mercaptoethanol, 100 µg of proteinase K
Table 1 . Bacillus sphaericus isolates examined by DNA
reassociation.
Refer to Rippere et al. 1997. DNA similarities among mosquito-
pathogenic and nonpathogenic strains of Bacillus sphaericus.
Int. J. Syst. Bacteriol. 47:214-216.
per ml) was added to each preparation, and the mixture was
incubated at 55oC for 1 h. Protein was removed by multiple
phenol-chloroform extractions, and DNA was precipitated with 0.6
volume of isopropanol. The DNA was dried and suspended in 20 ml
of TE, 250 µl of an RNase solution (1 mg of RNase A per ml, 4,000
U of RNase T1 per ml) was added, and the preparation was
incubated 1 h at 37oC. The DNA was chloroform extracted and
precipitated with ethanol. The precipitated DNA was dissolved
in 3 ml of TE and frozen.
DNA similarities
DNA was sheared in a French pressure cell and labeled with125I, and a hybridization analysis was performed by using the S1
nuclease method (4). DNA samples were heated for 5 min at 60oC
before they were used. Reaction tubes containing 10 µl of
labeled DNA (0.4 mg/ml), 50 µl of unlabeled DNA (0.4 mg/ml), and
50 µl of buffer (13.2× SSC, 5 mM HEPES; pH 7.0 [1× SSC is 0.15 M
NaCl plus 0.015 M sodium citrate]) were incubated at 60oC for 24
h to allow reassociation. Following this incubation, 1 ml of
107
buffer (0.3 M NaCl, 0.05 M acetic acid, 0.5 mM ZnCl2), 100 U S1
nuclease, and 50 ml of denatured salmon sperm DNA (0.4 mg/ml)
were added to each reaction mixture, and the mixture was
incubated for 1 h at 50oC. Then 0.5 ml of HCl buffer (1 M HCl,
1% Na4P2O7, 1% NaH2PO4) and 50 µl of native salmon sperm DNA (1.2
mg/ml) were added to the reaction mixture, and the preparation
was incubated for 1 h at 4oC to precipitate the DNA. The
precipitated DNA was collected on Whatman glass fiber filters
and counted with a gamma counter.
Results and Discussion
The strains used as reference strains for the homology
groups were the same as those used in the study of Krych et al.
(5). An additional strain, strain Gt1-a, was labeled and also
used as a reference. Also, six of the seven pathogenic strains
included in group IIA in the original study were analyzed again.
A total of 27 additional mosquito pathogen from a variety of
geographic locations were included in the study. These
pathogens had been isolated on the basis of their ability to
kill mosquito larvae. Each of these isolates, regardless of its
level of toxicity, was found to be a member of homology group
IIA (Table 2). This suggests that the four genes that have been
identified as being responsible for toxicity in these bacteria
have not been transferred beyond this genetically defined group.
As long as isolations were made on the basis of mosquito
pathogenicity, it appeared that homology group IIA might contain
only these distinctive pathogenic bacteria.
Jahnz et al. (3) utilized an oligonucleotide probe based on
a specific region of 16S rRNA from group IIA strains (1) to
108
isolate group IIA strains not on the basis of pathogenicity but
on the basis of membership in homology group IIA. These authors
recovered 20 strains from Brazilian soil that produced ribotype
and isozyme patterns typical of group IIA. However, these
strains lacked mosquito pathogenicity, and probes for the binary
toxin and Mtx toxin revealed that the genes for these toxins
were absent. We included five of these strains in this study
(strains G4a, Gt1-a, Gt1-d, R1e, and R4a) and utilized Gt1-a as
a labeled reference strain. The high levels of homology of
these strains to 1593, the group IIA reference strain, leaves no
doubt that they are in fact members of homology group IIA. In
addition, the group IIA pathogens exhibited high levels of
homology to Gt1-a. Therefore, it appears that although all of
the pathogens belong to homology group IIA, this homology group
also contains nonpathogens. It is interesting that Jahnz et al.
(3) recovered only nonpathogens when they used their probe.
These authors suggested that the nonpathogens may, in fact be
more common in soil than the homology group IIA pathogens.
Whether a pathogen or a nonpathogen is isolated may simply
depend on the method used for selection (i.e., pathogenicity or
response to the group IIA probe).
Table 2. Levels of DNA similarity among strains of B.
sphaericus.
Refer to Rippere et al. 1997. DNA similarities among mosquito-
pathogenic and nonpathogenic strains of Bacillus sphaericus.
Int. J. Syst. Bacteriol. 47:214-216.
In view of this, it does not seem appropriate to utilize
mosquito pathogenicity as the sole characteristic for defining a
new species based on homology group IIA.
109
References
1. Aquino de Muro, M., and F. Priest. 1994. A colony
hybridization procedure for the identification of mosquitocidal
strains of Bacillus sphaericus on isolation plates. J.
Invertebr. Pathol. 63:310-313.
2. Aquino de Muro, M., W. Mitchell, and F. Priest. 1992.
Differentiation of mosquito-pathogenic strains of Bacillus
sphaericus from non-toxic varieties by ribosomal rRNA gene
restriction patterns. J. Gen. Microbiol. 138:1159-1166.
3. Jahnz, U., A. Fitch, and F. Priest. 1996. Evaluation of an
rRNA-targeted oligonucleotide probe for the detection of
mosquitocidal strains of Bacillus sphaericus in soils;
characterization of novel strains lacking toxin genes. FEMS
Microbiol. Ecol. 20:91-99.
4. Johnson, J. L. 1994. Similarity analysis of DNAs, p. 655-
682. In P. Gerhardt, R. G. E. Murray, W. A. Wood, and N. R.
Krieg (ed.), Methods for general and molecular bacteriology.
ASM Press, Washington, D. C.
5. Krych, V., J. Johnson, and A. Yousten. 1980.
Deoxyribonucleotide acid homologies among strains of Bacillus
sphaericus. Int. J. Syst. Bacteriol. 30:476-484.
6. Liu, J.-W., A. Porter, B. Y. Wee, and T. Thanabalu. 1996.
New gene from nine Bacillus sphaericus strains encoding highly
conserved 35.8 kilodalton mosquitocidal toxins. Appl. Environ.
Microbiol. 62:2174-2176.
7. Porter, A., E. Davidson, and J.-W. Liu. 1993. Mosquito
toxins of bacilli and their genetic manipulation for effective
biological control of mosquitoes. Microbiol. Rev. 57:838-861.
8. Thanabalu, T., and A. Porter. 1996. A Bacillus sphaericus
gene encoding a novel type of mosquitocidal toxin of 31.8 kDa.
Gene 170:85-89.
110
9. Woodburn, M. A., A. Yousten, and K. Hilu. 1995. Random
amplified polymorphic DNA fingerprinting of mosquito-pathogenic
and nonpathogenic strains of Bacillus sphaericus. Int. J. Syst.
Bacteriol. 45:212-217.
110
SUMMARY
Bacillus popilliae and B. lentimorbus, pathogens of the
Japanese beetle, have been differentiated by the production of a
parasporal crystal at the time of sporulation and the ability to
grow in the presence of 2% NaCl in B. popilliae and the lack of
these characteristics in B. lentimorbus. Many different
classification systems have been proposed for these bacteria,
but the classification has not been studied using molecular
techniques. Bacillus popilliae and B. lentimorbus were examined
using both DNA similarity studies and randomly amplified
polymorphic DNA (RAPD) analysis. Thirty-four isolates of B.
popilliae and B. lentimorbus were examined for DNA similarity
using the S1 nuclease method. 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 (NRRL B-4081 and Bp3)
distinct from but related to B. popilliae. Twenty-five isolates
were identified as B. popilliae, while 19 isolates were
identified as B. lentimorbus. Two B. popilliae isolates (NRRL
B-2522 and BlPj1) were originally received as B. lentimorbus
while seven B. lentimorbus isolates (DGB1, Bp1, Bp7, BpCb1,
BpCb2, BpPa1 and BpCp1) were originally received as B.
popilliae. These seven strains of B. lentimorbus produce a
paraspore during sporulation, possibly leading to their original
misidentification. All B. popilliae isolates with the exception
of NRRL B-4081 were positive for growth when tested using a
combination of broth and plates supplemented with 2 % NaCl. The
B. lentimorbus isolates (except KLN2, which was positive) were
all negative for growth in 2 % NaCl. When tested for vancomycin
111
resistance, all of the B. popilliae isolates were positive and
all of the B. lentimorbus isolates were negative.
Nineteen milky disease isolates from various geographic
regions were subjected to RAPD analysis. It was hypothesized
that due to their diversity, these strains might reveal new
subgroups of B. popilliae and B. lentimorbus. Included in this
analysis were seventeen strains that had been previously
analyzed by M. Tran. Ten new B. popilliae and nine new B.
lentimorbus isolates were identified, but there were no new
subgroups identified for either species. Patterns relating
groups to either the geographic region or host insect were not
identified. All of the B. lentimorbus strains were negative for
growth in 2 % NaCl, however four B. popilliae strains were also
negative (Bp12, BpF, RM23 and RM 29). When tested for
vancomycin resistance, 16 (Bp22, RM23 and RM29) B. popilliae
strains were positive and 3 (Bp16, Bp17 and Bp25) B. lentimorbus
strains were also positive. Seven B. lentimorbus isolates
(Bp15, Bp16, Bp17, Bp18, Bp19, Bp25 and Bp26) were most closely
related to the crystal-forming subgroup identified in the DNA
similarity study. These isolates did form a crystal during
sporulation as detected through microscopic examination of a
hemolymph smear. DNA similarity and RAPD analysis of B.
popilliae and B. lentimorbus has validated the existence of the
two species originally identified by Dutky (1940).
Bacillus popilliae and Bacillus lentimorbus isolates were
examined for the presence of the cry and van genes. The
paraspore is only detectable in sporulated cells of B. popilliae
and B. lentimorbus. This limits the use of this characteristic
in identification to those laboratories capable of infecting
insect larvae with the bacteria. The design of a rapid assay to
112
detect the cry gene enables all laboratories with access to a
thermalcycler to use paraspore production in the identification
of these species. A PCR assay designed to amplify the cry gene
detected the gene in 31 of 35 B. popilliae isolates and in only
1 of 18 B. lentimorbus isolates. This assay is effective at
detecting the cry gene in B. popilliae but not in B.
lentimorbus. Transferable vancomycin resistance is an emerging
problem in clinical strains of enterococci. B. popilliae was
shown to be vancomycin resistant by Stahly et. al. (1992), but
the mechanism of resistance was unknown. A PCR-RFLP assay
designed to detect the van ligase genes in enterococci was used
to detect a gene in B. popilliae that is related to the
enterococcal van ligase genes. The sequence of the "vanE" gene
in B. popilliae had 76.8 % and 68.4 % nucleotide identity to the
vanA and vanB genes. The vanE gene is located either on a large
plasmid or on the chromosome of B. popilliae. This gene is
predicted to be part of an operon responsible for vancomycin
resistance in B. popilliae. It is yet to be determined if
vancomycin resistance in B. popilliae is transferable, but this
is likely due to the similarity of the vanE gene to both the
vanA and vanB genes.
DNA similarity analysis was used to examine the
classification of 34 B. sphaericus isolates pathogenic for
mosquitoes and 5 non-pathogenic B. sphaericus isolates
identified by a 16S rRNA probe. All of the isolates were
members of the B. sphaericus homology group IIA. As the non-
pathogens were also included in group IIA, it appears to be
inappropriate to designate group IIA a species based only upon
pathogenicity.
113
CONCLUSIONS
1. Most strains of B. popilliae will grow in the presence of 2
% NaCl. In contrast, most strains of B. lentimorbus will not
grow in the presence of 2 % NaCl.
2. Parasporal bodies are present in both B. popilliae and in a
subgroup of B. lentimorbus. Therefore, paraspore formation can
no longer be used as a reliable means to distinguish between the
species.
3. Most strains of B. popilliae are resistant to the antibiotic
vancomycin while most strains of B. lentimorbus are sensitive to
vancomycin.
4. Subgroups of strains were identified among the B. popilliae
isolates studied. There were no apparent relationships between
these strains and the insect from which they were isolated or
between the strains and their geographic origin.
5. PCR primers based upon the published cry18Aa1 nucleotide
sequence (Zhang et al. 1997) detected the presence of the gene
in most strains of B. popilliae known to produce parasporal
inclusions. These primers did not detect the gene in B.
lentimorbus isolates known to produce paraspores. The amount by
114
which the B. lentimorbus parasporal gene differs from the B.
popilliae paraspore gene is unknown.
6. A gene was identified and sequenced in B. popilliae that is
related to the vanA and vanB ligase genes in enterococci. This
gene has been designated vanE and encodes a ligase putatively
involved in vancomycin resistance in B. popilliae.
7. The vanE gene in B. popilliae has been localized to either
the chromosome or a large plasmid.
8. All B. sphaericus mosquito pathogens examined to date belong
to DNA similarity group IIA. A few non-pathogens isolated on
the basis of 16S rRNA similarity also belong to this group. It
may be premature to give species status to this DNA similarity
group based solely upon mosquito pathogenicity.
115
Karen Elaine Rippere Lampe
20712 Crystal Hill Circle #GGermantown, MD 20874
301-972-2708
EDUCATION
Doctor of Philosophy, Microbiology, September, 1998Virginia Polytechnic Institute and State University, Blacksburg, VADissertation: Systematics of the entomopathogenic bacteria Bacillus popilliae,Bacillus lentimorbus and Bacillus sphaericus.Project includes: DNA:DNA similarity analysis, phenotypic analysis, plasmidcuring, Southern blots, hybridizations, nonradioactive detection of probes, RAPDanalysis, DNA sequencing, high performance liquid chromatography, polymerasechain reaction, multiplex PCR-RFLP, SDS PAGE, Sephadex gel filtrationcolumn chromatographyMajor Advisor: Dr. Allan Yousten, Professor of Microbiology
Bachelor of Science,Biology, December 1993Virginia Polytechnic Institute and State University, Blacksburg, VABasic biological principles and microbiological techniques
Secondary EducationSouth River High School, June 1990
PROFESSIONALEXPERIENCE
TeachingGraduate Teaching Assistant: Laboratory Instructor, Department of BiologyVirginia Polytechnic Institute and State University, Blacksburg, VAJanuary 1995-presentTaught laboratory sections in General Biology, General Microbiology, AquaticMicrobiology, Microbial Physiology, Pathogenic Bacteriology and MolecularPlant Systematics
Teaching Experience:Spring 1998 Molecular Plant Systematics laboratoryFall 1996 & 1997 Pathogenic Bacteriology laboratorySpring 1997 General Microbiology laboratorySpring 1996 Aquatic Microbiology laboratoryFall 1995 Microbial Physiology laboratorySpring 1995 General Biology laboratory
116
Related ExperienceGraduate Research Assistant, Biology DepartmentVirginia Polytechnic Institute and State University, Blacksburg, VASpring 1995, Summer 1995, 1996 & 1997Designed and performed experiments related to dissertation
Laboratory TechnicianReichardt Animal Hospital, Edgewater, MDJanuary 1994-August 1994Performed clinical diagnostic tests
Data Entry Clerk, Drug Listing BranchFood and Drug Administration, Rockville, MDData entry into computers, streamlined drug listing process
PUBLICATIONS
Rippere, K. E., R. Patel, J. R. Uhl, K. Piper, J. M. Steckelberg, B. Kline, F.R. Cockerill and A. A. Yousten. A Gene Resembling vanA and vanB in theVancomycin-Resistant Biopesticide Bacillus popilliae. J. Infect. Dis. 178:584-588.
Alban, P. S., D. L. Popham, K. E. Rippere and N. R. Krieg. Identification ofa Gene for a Rubrerythrin/Nigerythrin-Like Protein in Spirillum volutans byUsing Amino Acid Sequence Data from Mass Spectrometry and NH2-terminalSequencing. J. Appl. Microbiol. In press.
Rippere, K. E., M. T. Tran, A. A. Yousten, K. H. Hilu and M. G. Klein.Molecular Systematics of Bacillus popilliae and Bacillus lentimorbus, Bacteriacausing Milky Disease in Japanese Beetles and Related Scarab Larvae. Int. J.Syst. Bacteriol. 48:395-402.
Yousten, A. A. and K. E. Rippere. 1997. DNA Similarity Analysis of a PutativeAncient Bacterial Isolate Obtained From Amber. FEMS Lett. 152:345-347.
Rippere, K. E., J. L. Johnson and A. A. Yousten. 1997. DNA Similaritiesamong Mosquito-pathogenic and Nonpathogenic Strains of Bacillus sphaericus.Int. J. Syst. Bacteriol. 47:214-216.
Pettersson, B., K. E. Rippere, A. A. Yousten and F. G. Priest. Transfer ofBacillus lentimorbus and Bacillus popilliae to the Genus Paenibacillus withDescriptions of Paenibacillus lentimorbus comb. nov. and Paenibacilluspopilliae comb. nov. In review.
117
ABSTRACTS &
PRESENTATIONS
Rippere, K. E. and A. A. Yousten. Studies of the Beetle Pathogens, Bacilluspopilliae and Bacillus lentimorbus. Presented at 6o Simposio de ControleBiologico, May 1998.
Rippere, K. E, R. Patel and A. A. Yousten. A Gene Resembling vanA andvanB in the Biopesticide Bacillus popilliae. Presented at the national AmericanSociety for Microbiology meeting, May 1998.
Rippere, K. E., M. T. Tran, K. Hilu, M. Klein and A. A. Yousten. MolecularSystematics of Milky Disease Bacteria. Presented at the Society for InvertebratePathology meeting, Aug. 1997.
Rippere, K. E., J. L. Johnson and A. A. Yousten. DNA Homologies AmongStrains of Milky Disease Bacteria. Presented at the national American Society forMicrobiology meeting, May 1996
Lampe, R. C., K. E. Rippere, J. L. Johnson, T. Phelps and R. E. Benoit.Characterization of a Deep Subsurface Microaerophile Using 16S rRNASequencing and DNA DNA Reassociation. Presented at the national AmericanSociety for Microbiology meeting, May 1996
Rippere, K. E. DNA Homologies Among Strains of Milky Disease Bacteria.Presented at theVirginia Branch ASM meeting Dec., 1995
HONORS &AWARDS
1998 Sigma Xi Research Grant awarded1997 Sigma Xi Research Grant awarded1996 Phi Kappa Phi Honor Society
PROFESSIONALMEMBERSHIPS
American Society for MicrobiologyAmerican Society for Microbiology (Virginia Chapter)Society for Invertebrate PathologySigma Xi
REFERENCES
Available upon request
