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Order Number 8726807

Characterization of plasmids from Bacillus thuringienais var. israelensis

Clark, Burton David, Ph.D.

The Ohio State University, 1987

UMI SOON.ZeebRA Ann Arbor, MI 48106

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CHARACTERIZATION OF PLASMIDS FROM BACILLUS THURINGIENSIS VAR. ISRAELENSIS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Doctor of Philosophy in the Graduate School of the Ohio State University

By

Burton David Clark, B.Sc.

*****

The Ohio State University 1987

Dissertation Committee: Approved by

D. H. Dean B. R. Oakley W. R. Strohl Adviser Department of Microbiology To My Wife

%% ACKNOWLEDGEMENTS

I would like to thank my advisor, Prof. Donald H. Dean, for his support and encouragement. Don gave me the freedom to make my own mistakes (a privilege I never hesitated to use), but was always available with advice and support when I needed it. I admire his caring for the people in his group beyond caring about their science. I would also like to thank the members of my thesis committee. Prof. Berl R. Oakley and Prof. William R. Strohl for their suggestions and advice.

I am grateful to Tom Boyle, Bill Chu, Steve Cohen, Dan Ellis, John Hopper, Inara Ladzins, John Lohr, Phyllis and Bill Martin, Jim McLinden, Penny Parker, John Perkins, Fred Perlak, Laura Pettine, Josanne Sabourin, Wes Workman, Dan Zeigler, and other members of the lab whose ideas and encouragement were invaluable, and whose tolerance of my humor was admirable.

I am also grateful to Philip Auron at MIT who has been patient and supportive of my completing this thesis.

I want to thank my mother for her interest in my work and for instilling in me a desire to always do my best. Most of all, I want to thank my wife and best friend, Molly, for her love, devotion, and encouragement, and for dragging me away from the laboratory when I needed it most.

%%% VITA

February 6,1957 ...... Born - Beverly, Massachusetts 1975-1979 ...... B.S., Microbiology, University of Massachusetts, Amherst, Massachusetts 1977-1979 ...... Undergraduate research in molecular genetics Department of Microbiology University of Massachusetts, Amherst, MA 1979-present ...... Graduate Research Associate, Department of Microbiology The Ohio State University, Columbus, Ohio 1985-present ...... Research Associate, Massachusetts Institute of Technology, Cambridge, Massachusetts.

PUBLICATIONS & ABSTRACTS

• Clark, B.D., Lohr, J.R., and Dean, D.H. (1980). Genetic manipulation and phenotypic properties of plasmids of the insect pathogen. Bacillus thuringiensis. Second American Society for Microbiology Conference on Genetics and Molecular Biology of Industrial Microbiology. Bloomington, ID. October 7, 1980. (abstract)

* Clark, B.D., Perlak, F.J., Chu, C.-Y., and Dean, D.H. (1982). The Bacillus thuringiensis genetic systems. In: Comparative Pathobiology, edited by LA. Bulla, Jr. and T.C. Cheng, pp. 155-173. Vol. 5. Plenum Press, New York.,

%v • Clark, B.D., Chu, C.-Y., Cohen, S.B., and Dean, D.H. (1982). Plasmid associated toxicity in Bacillus thuringiensis var. israelensis. American Society for Microbiology Annual Meeting. Atlanta, G A. March 7, 1982. (abstract)

• Dean, D.H., Clark, BJ)., Lohr, J.R., and Chu, C.-Y. (1982). Recent advances in the genetics of research in Bacillus thuringiensis. Invertebrate Pathology and Microbial Control: Proceedings of the Third International Colloquium on Invertebrate Pathology. University of Sussex, Brighton, United Kingdom, (abstract)

• Clark, B.D., and Dean, D.H. (1983). A high molecular weight plasmid is associated with toxicity in Bacillus thuringiensis var. israelensis. American Society for Microbiology Annual Meeting. New Orleans, LA. March 9, 1983. (abstract)

• Clark, B.D., and Dean, D.H. (1984). The antimicrobial susceptibility of various Bacillus thuringiensis var. israelensis strains. American Society of Microbiology Annual Meeting. St. Louis, MI. March 7, 1984. (abstract)

• Boyle, T.M., Clark, B.D., and Dean, D.H. (1984). Restriction endonuclease mapping of three plasmids from Bacillus thuringiensis var. israelensis. Annual Meeting of the Society for Invertebrate Pathology. Davis, California. August 9,1984. (abstract)

• Clark, B.D., Boyle, T.M., Chu, C.-Y., and Dean, D.H. (1985). Restriction endonuclease mapping of three plasmids from Bacillus thuringiensis var. israelensis. Gene 36, 169-171.

• McLinden, J.H., Sabourin, J.R., Clark, B.D., Gensler, D.R., and Dean, D.H. (1985). Cloning and expression of an insecticidal K-73 type crystal protein from Bacillus thuringiensis var. kurstaki into Escherichia coli. Applied and Environmental Microbiology 50, 623-628.

• Clark, B.D., Rosenwasser, L.J., Webb, A.C., Irie, S., Gehrke, L., and Auron, P.E. (1986). Expression of biologically active human interleukin

V 1 subpeptides by transfected COS cells. Sixth International Congress of Immunology. Toronto, Canada. July 7, 1986. (abstract)

• Fenton, M.J., Clark, B.D., Collins, K., Rich, A., Webb, A., and Auron, P.E. (1986). Expression of the human IL-1 beta gene in THPl cells following LPS stimulation. Sixth International . Congress of Immunology. Toronto, Canada. July 7, 1986. (abstract)

• Rosenwasser, L.J., Webb, A.C., Clark, B.D., Irie, S., Chang, L., Dinarello, C.A., Gehrke, L., Wolff, S.M., Rich, A., and Auron, P.E. (1986). Expression of biologically active human interleukin 1 subpeptides by transfected simian COS cells. Proc. Natl. Acad. Sci. USA 83, 5243-5246.

• Clark, B.D., Collins, K.L., Gandy, M.S., Webb, A.C., Auron, P.E. (1986). Organization and structure of the human prointerleukin 1 beta gene. Congress on Research in Lymphokines and Other Cytokines. Boston, MA. August 12, 1986. (abstract)

• Fenton, M.J., Clark, B.D., Collins, K.L., Webb, A.C., and Auron, P.E. (1986). Transcriptional regulation of the human prointerleukin 1 beta gene. Congress on Research in Lymphokines and Other Cytokines. Boston, MA. August 12, 1986. (abstract)

• Clark, B.D., Collins, K.L., Gandy, M.S., Webb, A.C., and Auron, P.E. (1986). Genomic sequence for human prointerleukin 1 beta: possible evolution from a reverse transcribed prointerleukin 1 alpha gene. Nucleic Acids Res. 14, 7897-7914.

• Rosenwasser, L.J., Auron, P., Gehrke, L., Clark, B., McDonald, B., Bradley, B., Epstein, E., Collins, K., Webb, A. (1986) Interleukin 1 - Review of structure and function, definition of an active site for T cells, and production by cultured vascular endotheliuni. In Biologically based immunomodulators in the therapy of rheumatic diseases, Pisetsky, P.S. and Rosenwasser, L.J., eds. Elsevier Scientific Publishers, NY.

m • Fenton, M.J., Clark, B.D., Collins, K.L., Webb, A.C., and Auron, P.E. (1987). Transcriptional regulation of the human prointerleukin 1 beta gene. J. Immunol. 138, 3972-3979.

Fenton, M.J., Clark, B.D., Collins, K.L., Webb, A.C., and Auron, P.E. (1987). The human proIL-l;9 gene is regulated by two distinct receptor, pathways. Fifth International Lymphokine Workshop, January 1987, Clearwater, FL. (abstract)

• Fenton, M.J., Clark, B.D., Alexander, S.J., Webb, A.C.j Auron, P.E. (1987). Differential expression of the human prointerleukin 1 beta gene. In: Mechanisms of Control of Gene Expression, UCLA Symposia on Molecular and Cellular Biology, Vol. 67. Collen, B., Gage, L., Siddigui, M., Skalka, A.-M., and Weissback, H., eds. Alan R. Liss, New York.

• Webb, A.C., Collins, K.L., Snyder, S.E., Rosenwasser, L.J., Clark, B.D., Eddy, R.L., Shows, T.B., Auron, P.E. (1987). Molecular cloning, expression and chromosomal assignment of a potential inhibitor of human prointerleukin 1^ processing. Lymphokine Research, Vol 6, No. 1. (abstract)

• Fenton, M.J., Clark, B.D., Collins, A.C., Rich, A., Webb, A.C., Auron, P.E. (1987) Two pathways of human prointerleukin 1)9 gene regulation by PMA and LPS. Lymphokine Research, Vol.6, No. 1. (abstract)

• Clark, B.D., Fenton, M.J., A., Webb, A.C., Auron, PJE. (1987) Characterization of cis and trans acting elements involved in human proIL-l)9 gene expression. International Workshop on Monokines and other Non-lymphocytic Cytokines, December 1987, Hilton Head Island, S.C. (abstract)

• Clark, B.D., Fenton, M.J., A., Webb, A.C., Auron, PÆ. (1987) Expression of the CAT gene in mammalian cells under the control of human prointerleukin 1 beta regulatory sequences. Arthritis Fellows Conference, December 1987, Amelia Island, FL. (abstract)

V ll • Fenton, M.J., Vermeulin, M.W., Clark, B.D., Webb, A.C., and Auron, P.E. (1987) Human prointerleukin-1 beta gene expression is regulated by two distinct pathways, (submitted)

• Clark, B.D., and Dean, D.H. (1987). The antimicrobial susceptibility of Bacillus thuringiensis var. israelensis. (in preparation)

FIELD OF STUDY

Molecular Genetics. Professor Donald H. Dean

m il TABLE OF CONTENTS

Dedication ii Acknowledgements Hi Vita iv List of Tables xii List of Figures ivx

Introduction ' 1

C hapter 1: Literature Reviéw 3

Discovery of Bacillus thuringiensis . 3 Bacillus thuringiensis crystals 5 Additional entomopathogenic properties of Bacillus thuringiensis 11 Use of Bacillus thuringiensis as a microbial insecticide 12 Bacillus thuringiensis var. israelensis 14 Plasmids 15 Plasmids in Bacillus thuringiensis 16 References 18

C hapter 2: Isolation of Plasmid DNA from Bacillus thuringiensis 25 Strains and the. Curing of these Plasmids from Strain 4Q2-WT .

Introduction 25 Materials and Methods 26 Bacterial strains 26 Plasmid curing 28 Isolation of plasmid DNA 28 Agarose gel electrophoresis 30 Serotyping of strains 31 Storage of strains 32 Results 33 Discussion 47

IX References 50

C hapter 3: Characterization of the Three Smallest Bacillus 52 thuringiensis var. israelensis Plasmids and Construction of Shuttle Vectors

Introduction 52 Materials and Methods 54 Bacterial strains 54 Media and growth conditions 55 Plasmid DNA isolation and purification 55 Restriction endonuclease digestion and gel electrophoresis 56 Measurement of restriction fragments 57 DNA-DNA filter hybridization 58 Radioactive labeling of DNA probe 58 Pre-hybridization and hybridization of membranes 59 Results 61 Restriction digestion analysis of pTX14-l 61 Restriction digestion analysis of pTX14-2 66 Restriction digestion analysis of pTXl4-3 66 DNA-DNA filter hybridization 75 Construction of shuttle vectors 75 Discussion 86 References 90

C hapter 4: Association of the Crystal Toxin with a 108 Kb 93 Plasmid

Introduction 93 Crystal isolation . 94 Crystal solubilization 94 Bioassay 95 Materials and Methods 96 Bacterial strains 96 Isolation of crystals for bioassay 96 Bioassays 97 Isolation and purification of crystals 97

X Solubilization of crystal protein and SDS-polyacrylamide 98 gel electrophoresis Results 101 Bioassay of plasmid cured strains 101 Isolation and characterization of crystal proteins 101 Discussion 118 References 123

C hapter 5: The Antimicrobial Susceptibility of Various Bacillus 127 thuringiensis var. israelensis Strains

Introduction 127 Minimal inhibitory concentration 128 Materials and Methods 130 Bacterial strains 130 Antibiotics 130 Antibiotic susceptibility testing 130 agar dilution method 130 broth dilution method 131 Results 132 Discussion 142 References 145

Summary and General Conclusions 147

Appendix 150

List of References . 155 LIST OF TABLES

1.1 List of representative strains for each of the nineteen serotypes 6

1.2 Comparison of the potency of HD-1 powder with the potency of 14 superior B. thuringiensis powders

2.1 List of j B . ihunnjfienszs var. isroe/ensis strains used 27

2.2 Plasmids present in B. thuringiensis var. israelensis 4Q2 mutants 46

2.3 Plasmids present in B. thuringiensis var. israelensis 46 other than 4Q2

3.1 Restriction endonuclease cleavage sites in B. thuringiensis 67 var. israelensis plasmids

3.2 Restriction fragments produced by single and double 68 endonuclease digestions of pTXl4-l

3.3 Restriction fragments produced by single and double 69 endonuclease digestions of pTX14-2

3.4 Restriction fragments produced by single and double 70 endonuclease digestions of pTX14-3

3.5 Fragments produced by partial digestion of pTX14-3 with Hind HI 71

4.1 Conditions for solubilization of B. thuringiensis var. 100 israelensis crystal

4.2 Results of bioassays on crude crystal preparations 103

X ll 4.3 Plasmids present in B. thuringiensis var. israelensis 4Q2 104 mutants

5.1 List of Bacillus thuringiensis var. israelensis strains 133 used in antibiotic studies

5.2 Antibiotics and the concentrations used in culture media during 134 preliminary antibiotic-susceptibility testing using the agar dilution method

5.3 Minimal inhibitory concentrations of eighteen antibiotics for 135 B. thuringiensis var. israelensis and B. thuringiensis var. kurstaki using the agar dilution method

5.4 Minimal inhibitory concentrations of antibiotics tested with 136 B. thuringiensis var. israelensis 4Q2-WT using the broth dilution method

X l l l LIST OF FIGURES

2.1 Plasmid pattern of B. thuringiensis yar. israelensis 4Q2-WT 34

2.2 Pedigree of plasmid cured mutants derived from B. thuringiensis 36 var. israelensis strain 4Q2-WT

2.3a + 2.3b Modified Eckhardt lysate electrophoresis of B. thuringiensis 39 var. israelensis plasmid cured mutants

2.4a - 2.4d Electrophoresis of B. thuringiensis var. israelensis plasmid 42 DNA isolated by the modified PEG method of Kronstad et al.

3.1 Electrophoretic analysis of purified plasmid DNA used for restriction 63 endonuclease studies

3.2 Representative electrophoretic gels of restriction endonuclease 65

3.3 Restriction endonuclease map of pTX14-l 72

3.4 Restriction endonuclease map of pTXl4-2 73

3.5 Restriction endonuclease map of pTX14-3 74

3.6 DNA-DNA filter hybridization of pTX14-3 probe to 4Q2-WT 78 plasmids and DNA controls

3.7 Circular maps of pBDC-3 and pBDC-4 79

3.8 Circular map of pBDC-5 80

xiv 3.9 Electrophoretic analysis of pBDC-3, pBDC-4, and pBDC-5 plasmid 82 DNA

3.10 Circular maps of pBDC-6 and pBDC-7 83

3.11 Electrophoretic analysis of pBDC-6 and pBDC-7 plasmid DNA 85

4.1 Purification of crystals from strain 4Q2-WT with continuous 106 50-80% Renografin-76 gradients

4.2 Polyacrylamide gel electrophoresis analysis of crystal proteins 108 solubilized with different buffers

4.3 Polyacrylamide gel electrophoresis analysis of crystal proteins 110 solubilized with phosphate buffer

4.4 Purification of an inclusion with Renografin-76 gradients 113

4.5 Polyacrylamide gel electrophoresis analysis of crystal and 115 inclusion proteins

4.6 Analysis of crude protein and purified ciTstals from strains 117 4Q2-WT, 4Q2-72, and 4Q2-81

5.1a - 5.Id Susceptibility of B. thuringiensis var. israelensis to 138 antibiotics using the agar dilution method

XV INTRODUCTION

Every year, insects destroy large portions of the world food supply and are vectors for many diseases, both to humans and their livestock. Last year alone, thirty million people were infected with malaria (carried by mosquitoes), and of these, over one million (mostly children) died. For several decades, petroleum based chemical pesticides have been effective in controlling insects, however, they have their drawbacks. Pollution to the environment, increasing expense due to changing oil prices, and insect resistance have necessitated the search for alternative controls.

As an alternative, the bacterium. Bacillus thuringiensis, has been used successfully in pest control. This bacterium produces a parasporal crystal that is toxic to many Lepidoptera and Diptera insects. Recombinant DNA technology has been applied with hopes to further improve the efficacy of this bacterium at controlling insects.

This dissertation describes the characterization of plasmids from Bacillus thuringiensis var. israelensis, one of the more than twenty subspecies. More specifically, this work concentrates on isolating plasmid cured mutants, restriction enzyme analysis, construction of shuttle vectors, and the association of phenotypes to plasmids.

Strains of Bacillus thuringiensis var. israelensis were cured of one to nine plasmids. A strong correlation was observed between the presence of a 108 kilobase pair plasmid and cr^tal production. Strains that did not posses this plasmid did not produce any of the proteins that compose the crystal.

In addition, detailed physical maps positioning restriction endonuclease sites were prepared for the three smallest Bacillus thuringiensis var. israelensis plasmids. They were also studied by Southern hybridization, as well as being used in the construction of shuttle vectors. Finally, the minimal inhibitory concentration for eighteen different antibiotics was determined in hopes of associating antibiotic resistance with one of the plasmids, but no association could be made. Nonetheless, the wealth of information will be valuable in future research with this microorganism. CHAPTER ONE Literature Review

Discovery of Bacillus thuringiensis

The earliest documented record of the bacterium known today as Bacillus thuringiensis, was in Japan in 1901. Ishiwata (1901, 1902) isolated the sporeforining bacteria, Bacillus sotto, now known as B. thuringiensis var. sotto, from diseased silkworms. Bombyx mori . He described "sotto bacillus" disease, which was an epidemic in the sericulture industry. A decade later, while working at a flour mill, Berliner isolated an insect pathogen from diseased Mediterranean flour moths, Anagasta kuehniella, and identiHed a bacterium as the causative agent (Berliner, 1911). In 1915, he named this Gram positive, motile, sporeforming insect pathogen B. thuringiensis after Thueringen, the region in southern Germany where it was discovered (Berliner, 1915). He was the first to describe an irregular, rhomboid crystalline inclusion. This crystal inclusion, called the restkorper by Berliner, was also referred to as the parasporal body, due to its location near the spore, and later as the g-endotoxin, due to its association with insecticidal toxicity. Mattes (1927) verified the presence of the parasporal inclusion, and Husz (1927) and Chorine (1930) used B. thuringiensis for the biological control of corn borer larvae, Pÿrausta nubilalis, in small field trials. However, its use could not compete economically or effectively with petroleum based insecticides and research on B. thuringiensis was inactive for the next twenty years.

Steinhaus (1941) at The Ohio State University studied bacterial pathogens and the normal flora associated with various insects. He isolated B. entomocidus, later to be called B. thuringiensis var. entomocidus, from diseased larvae of Paralipsa gularis and used this bacterium in the control of the alfalfa caterpillar, Colias eurytheme (Steinhaus, 1951). He recognized the potential of B. thuringiensis as a biological insecticide and conceived the term "microbial control", the use of microorganisms to control insects. He is credited with the renaissance of research on -B. thuringiensis. Hannÿ (1953) rediscovered the parasporal crystal and speculated that it may be associated with the entomo­ pathogenic properties of the bacterium. Angus (1954, 1956) , working with the strain originally isolated by Ishiwata, demonstrated that the parasporal crystal was the toxic moiety in silkworms, causing first paralysis and then death after ingestion.

As more and more insects became resistant to the petroleum based organic insecticides used in agriculture, a greater number of investigators became interested in B. thuringiensis and they started isolating sporeforming, crystalliferous bacilli pathogenic to Lepidoptérans (caterpillar forming insects). For example, Delaporte and Beguin (1955) isolated B. anduze from silkworms {Bombyx mori) in 1952. Talalaev (1956) isolated B. dendrolimus from diseased Sibirian silkworms, Dendrolimus sibiricus, which were destroying the pine forests of Siberia, and Isakova (1958) described B. cereus var. galleriae, which he isolated from the greater wax moth. Galleria mellonella. As more crystalliferous bacteria were isolated and reported, rapid and precise identification of these new isolates became a problem.

In 1962, deBarjac and Bonnefoi conducted indepth biochemical and serological tests on 24 crystalliferous strains (deBarjac and Bonnefoi, 1962). The results of this study enabled them to organize what had become a confusing collection of different species names in the literature. They divided the bacteria into six serotypes or varieties of B. thuringiensis (in the literature, serotype and variety are used synonymously). Their divisions were based primarily according to the H antigen of the flagella. Some of the serotypes were divided into sub-serotypes, based on H antigen subfactors or biochemical differences such as acid production from the fermentation of salicin or sucrose. The number of serotypes steadily increased to 19 because of the discovery of new isolates (see Table 1.1) (deBarjac et al., 1973, 1977, 1978, DeLucca et al., 1979, Ohba et al., 1981b, 1981c). Most of the serotypes are pathogenic to Lepidoptera. Noticeable exceptions are B. thuringiensis var. israelensis (serotype 14), which is toxic to mosquitoes and black fly (Dlptera) and B. thuringiensis var. tohokuensis (serotype 17) and B. thuringiensis var. tochigiensis (serotype 19) which are crystalliferons, but non-toxic. By the mid 1970% the search for new isolates of B. thuringiensis declined and the phase of characterizing the bacterium intensified.

Taxonomically, it was difficult to separate this organism from other closely related groups. J3. thuringiensis var. thuringiensis has 81-99% DMA homology with other varieties of B. thuringiensis, 82% DNA homology with B. cereus, 91% DNA homology with B. anthracis, 8% DNA homology with B. subtilis Marburg, and 6% DNA homology with Escherichia coli (Kaneko et al., 1978, Seki et al., 1978). B. thuringiensis and B. cereus are morphologically and histologically similar (Gordon et al., 1973). They share homologous flagellar proteins (Krieg, 1969) and have a similar fatty acid composition (Kaneda, 1968). The key discerning feature of B. thuringiensis which separates this species from others is the parasporal crystal. The other species lack this crystal and therefore are unable to show pathogenicity to insects. B. thuringiensis mutants that have lost the ability to produce parasporal crystal are indistinguishable from B. cereus.

Bacillus thuringiensis crystals

Temporally, crystal formation coincides with sporulation, but its synthesis appears to develop independently of any association with the plasma membranes or with the membranes of the spore (Bechtel and Bulla, 1976, Nishimura and Nishiitsutsuji-Uwo, 1980). Spore and crystal formation is followed by cell lysis and they are released from the cell physically separate from each other. An exception is B. thuringiensis var. finitim us, in which the crystal forms inside the exosporium and is released in association with the spore.

Because of the . economic importance associated with its insect pathogenicity, the crystal of B. thuringiensis has evoked the interest of many people. The first extensive characterization of the crystal by Hanny and Fitz-James (1955) revealed that it composes 30% of the dry weight of sporulated bacteria and is constructed from one protein containing at least 17 different amino acids. They TABLE 1.1. List of representative strains for each of the nineteen serotypes with their H antigen type and biochemical characteristics to aid in their differentiation.

Strain H-serotype* crystal sucrose mannose sallcln starch chltln

thuringiensis 1 + + + + ++ _ finitimus 2 - + - + -• + alesti'* 3a + - -— + kurstaki 3a, 3b + - - + ++ + sotto 4a, 4b + + -— + • dendrolimus 4a,4b + - -- + ++ kenyae 4a,4c + -- ■ + + - galleriae 5a, 5b + - - + ++ + canadensis** 5a,5c + + - + + + subtoxicus 6 + + + - ++ — entomocidus 6 + + + - + — aizavai 7 . + + -- d ++ morrisoni 8a, 8b +. + -- d + ostriniae 8a,8c + - + + d d . tolworthi® 9 + + - + + + darmstadiensls 10 + - -- ++ — toumanoffl 11a,lib + - + - ■ ++ - kyushuensls lia,lie + - + + + - thompsoni 12 ■ + + + + + - Pakistani 13 • + + - + + d israelensis'* 14 + - - - + - dakota 15 + ND ND ND ND ND Indiana 16 + + + + - - tohokuensis 17 + + - - + + kunamotoensls 18 + ■ + - ND + + tochigiensis 19 + + - ND + +

*The a,b, and c associated with the H-serotype refer to subfactors present with the H-antigen. Subfactor "a" is usually common to all B. thuringierms serotypes.

^Can use the pellicle utilization assay to differentiate alesti (-) from israelensis (+).

% an use the pellicle utilization assay to differentiate canadensis (-) from tolworthi (+).

d=weakly active

ND = not determined observed the crystals by light and electron microscopy and found them to be very regular in shape. They are bi-pyramidal, with serrated edges, that at higher resolution, are constructed of parallel rows of dumbbell-shaped dimers. Each dimer is composed of two spherical proteins, each approximately 8.7 nm in diameter (Labaw, 1964). Holmes and Manro (1965) have calculated from x-ray diffraction data that the dimer has a molecular weight of 230,000 daltons. To facilitate further study of the biochemistry and mode of action of the crystal, technologies were developed to purify crystals from the spores and cell debris, taking advantage of the different surface , properties or relative densities of these cell components. For example, Milne et al. (1977) used 50-80% Renografin-76 gradients that very nicely separated a sporulated culture of B.. thuringiensis into three components; debris, crystals, and spores. Ang and Nickerson (1978) and Nickerson and Swanson (1981) achieved similar results using sodium bromide gradients. The amount of material that can be separated by these gradients is limited and overloading can cause distinct bands of pure components to overlap. Overloading can be reduced by first removing some of the spores. Sharpe et al. (1979) took advantage of the hydrophobicity of spores of B. thuringiensis. In a sporulated culture of B. thuringiensis, 50-60% of the spores are present in foam that forms on the surface of a shaking culture. The addition of gelatin can increase the foaming and greater than 90% of the spores can be removed by aspirating off the foam. Using the above purification methods, preparations of crystals with less than 0.03% of spores are achieved, which provide grams of crystal for further study.

In vitro, the crystal is resistant to most standard protein solvents. The crystal protein has intra-molecular diSulHde bonds which are rare in proteins. In addition, there are inter-molecular disulfide bonds which are important for forming dimers, the basic building block of the crystal. Each dimer has approximately six sulfhydryl groups on its external surface which form disulfide bonds with other dimers. These bonds provide the stability for the three dimensional crystal. At physiological pH, these bonds can only be broken and the crystals solubilized if strong reducing agents are present. In vitro, a phosphate or carbonate buffer at a pH of 9.5 or greater will cleave the disulfide 8 bonds, mimicing the high pH of the Leperdopteran gut. Huber et al. (1981) solubilized crystal from ten varieties of B. thuringiensis and separated the protein using polyacrylamide gel electrophoresis (PAGE) with and without the detergent, sodium dodecyl sulfate (SDS). When SDS was not used, they were all, with the exception of crystals from B. thuringiensis var. israelensis, composed of dimers of MW 231,000 daltons. When the crystal was separated on PAGE with SDS, the molecular weight was 130,000 daltons. Both Huber et al. (1981) and Bulla et al. (1977) suggest that the native (non-denatured) protein is a dimer. The difference between what would be the dimeric molecular weight obtained by SDS-PAGE in comparison to PAGE without SDS, is within observed error. Molecular weights ranging from 80,000 - 150,000 daltons have been reported for SDS-PAGE, depending on the method by which the crystal was solubilized, the porosity of the gel, and the amount of binding of SDS to the crystal polypeptide. Molecular weights of crystal by ultra-centrifugation in guanidine hydrochloride, provided molecular weights of 177,000 and 84,000 daltons, the larger molecular weight being the dimer of lower molecular weight proteins. The native dimer, and sometimes the 130,000 dalton monomer, which is also referred to as the protoxin, is later activated by the insect.

Some strains of B. thuringiensis have an additional protein • associated with the crystal and insect toxicity. This protein (P2) was first described by Yamamoto and McLaughlin (1981) as a 65,000 dalton protein. It is toxic to the cabbage looper, Trichoplusia ni, as is the major 130,000 dalton protein component of the crystal. The 130,000 dalton protein is referred to by Yamamoto as P i to distinguish it from P2. The P2 is unique from the PI in its tryptic peptide map (Yamamoto, 1983a) and more importantly, in its toxicity to the mosquito, Aedes taeniorhynchus. The P2 protein is found in the large PI crystal or as small, cuboidal inclusion bodies which have been studied by electron microscopy. Purification of these bodies has proven that they are composed of the 65,000 dalton P2 protein.

In vivo, the crystal protoxin is solubilized by the insect gut juices to smaller, activated, toxic fragments. This activation is rapid, taking less than 20 seconds (Huber, 1977) and symptoms due to the toxicity take place in several minutes depending on the insect species. Most studies on the mode of action have been done using the insect silkworm, Bombyx mari, or large white butterfly, Pieris brassicae. The target site of the toxic moiety is the gut epithelium. The activated crystal, also referred as the 5-endotoxin, causes feeding inhibition, and then total paralysis. Feeding inhibition takes place in 5-10 minutes. This is caused by a degradation of the gut epithelium and a mixing of the hemolymph with the gut juices. The epithelial cells swell and burst as seen by phase contrast microscopy (Fast and Morrison, 1972). Later, Griego et al. (1980) used scanning electron microscopy to see in more detail the swelling and destruction of epithelial cells in the, tobacco hornworm, Manduca sexta. These tissue changes are related to the effect that 5-endotoxin has on cell organelles. It causes the mitochondria to swell, vacuoles to appear in the rough endoplasmic reticulum, and lysosomes to increase in number (Endo and Nishiitsutsuji-Uwo, 1980).

In susceptible Lepidopteran and Dipteran insects, two factors are involved in the activation of the protoxin; proteases, and to a lesser extent, the alkaline environment of the gut. Lecadet and Dedonder (1966) purified two serine proteases, similar to trypsin and chymotrypsin, from the gut juice of the large white butterfly, Pieris brassicae, and found them to cleave the protoxin into fragments that were toxic. to Lepidopterans. If these proteases were heat inactivated, the crystals were no longer solubilized, suggesting that they are essential in crystal activation. The gut juices of other insects, such as the cabbage looper, Trichoplusia ni, bollworm, Heliothis zea, and silkworm. Bombyx mori, also have proteases which are necessary for the activation of the crystal. These enzymes, similar to the two proteases from IHeris brassicae, have optimum activity in the alkali environment of the insect gut (pH 7.5 - 10.0). The alkali environment contributes to the toxicity of B. thuringiensis by partially dissolving the crystal before enzymatic hydrolysis occurs. For example, the crystals oî B. thuringiensis var. aizarvai axe not activated by proteases until partially solubilized by alkali gut juices (Faust et al., 1967). Atypically, the crystals of B. thuringiensis var. tolworthi can be activated solely using alkali buffer. This would suggest that proteases are not always essential for activation. However, this statement has to be made with some caution because it is very 10 difficult to remove all proteases. It has been reported by Cbestukbina et al. (1980) that there are many metalloproteases and serine proteases on the crystal surface.

The varieties of B. thuringiensis have different host toxicity spectrums which are due to either insect species specific proteases required to activate the protoxin or different crystal types. Lecadet and Martouret (1964) first reported that the host toxicity spectrum can be attributed to insect species proteases. They found that crystals of B. thuringiensis var. thuringiensis are more toxic, and are dissolved faster by Pieris brassicae proteases than crystals from B. thuringiensis var. alesti. When equal amounts of pre-dissolved crystal from these two varieties are bioassayed, they have the same toxicity. This suggests that the difference in toxicity of the crystals is due to their proteolysis by the insect enzymes. Another example is the insensitivity of E. uehniella to the action of intact crystal. However, if the insect is fed crystal which has been digested and activated by proteases of Pieris brassicae, it becomes sensitive to the crystal. This difference in susceptibility is due to the specificity of the host proteases (Lecadet and Martouret, 1964). Examples of different host toxicity spectrums due to different crystal types are seen with B. thuringiensis varieties. These differences are seen after digesting their crystals with protease from IHeris brassicae. Even though the protein composing the crystals from the different varieties has a molecular weight of 130,000, the size of the fragments generated by protease differ. Treatment of B. thuringiensis var. thuringiensis crystal gives intermediate fragments of 80,000 and 60,000 daltons and a final fragment of 40,000 daltons. The 80,000 and 60,000 intermediates are not seen during the digestion of B. thuringiensis var. kurstaki crystal, suggesting a different amino acid sequence, resulting in changes in the sites normally recognized by the Pieties brossicoe proteases. These proteases will eventually degrade the crystal into smaller non-toxic fragments. However, the toxic moiety is associated with the larger fragments (Somerville and Pockett, 1975, Travers et al., 1976). In vivo, there is time for the large toxic fragment of the crystal polypeptide to yield entomopathogenic properties before it is further degraded to a non-toxic state. 11

Additional entomopathogenic properties of Bacillus thuringiensis

In addition to the crystal, B. thuringiensis synthesizes three types of metabolites that contribute to its entomopathogenicity. These are ;9-exotoxins, a-exotoxins, and opportunistic enzymes. The )8-exotoxin was first identified in B. thuringiensis var. thuringiensis by McConnel and Richards (1959) and found to be toxic to the greater wax moth, Galleria mellonella, as well as the housefly, Musca domestica (Hall and Arakawa, 1959). Not all varieties of B. thuringiensis synthesize this toxin. It is produced only in serotypes 1, 8, 9, 10, and 11, but it is a strain-specific property, and not a serotype-specific property. Ohba et al. (1981a) tested for its presence in 839 strains of. B. thuringiensis. For example, of 32 strains of serotype 10 tested, 29 strains produced the toxin. The ^S-exotoxin is produced during the vegetative growth phase of B. thuringiensis, and is a nucleotide composed of adenine, ribose, glucose, and phosphorylated allomuciic acid (Farkas et al.j 1969). It is heat stable and can withstand standard autoclaving conditions (Prystas et al., 1976).

The j@-exotoxin inhibits DNA-dependent RNA polymerase and therefore, the polymerization of RNA. It has no direct effect on protein and DNA synthesis, which proceed normally. ;3-exotoxin competes with ATP in vivo for the ATP- specific binding site of the DNA-RNA polymerase complex. It acts as an analogue of ATP and slows down polymerization, often resulting in termination before completion of transcription. Addition of excess ATP, but hot the other ribonucleic acid triphosphates, will relieve the ^-exotoxm effect. Due to the fundamental level at which this toxin works, it can inhibit the RNA polymerases of bacteria, invertebrates, and vertebrates (Beebee et al., 1972, Sebesta and Horska, 1970, Kim et al., 1972, Johnson, 1976). Because of this toxicity to invertebrates and vertebrates, only strains that do not produce the exotoxin are used in commercial preparations in the United States.

The role of another exotoxin, a-exotoxin,. is not as well understood. It has been characterized as a 50,000 dalton protein that is heat labile (Krieg, 1971, Krieg and Lysenko, 1979) . It has activity against insects, and high doses injected intraperitoneally or intravenously can kill mice. However, the a-exotoxin 12

does not hinder the use of B, thuringiensis as a commercial insecticide since it is very unstable and disappears during sporulation. It has not been detected in any commercial crystal preparations because they are allowed to sporulate. It is during this developmental phase that crystals are also produced.

In addition, B. thuringiensis varieties produce several enzymes including phospholipase C, chitinase, and miscellaneous proteases. These enzymes are virulance factors that contribute to the pathogenicity of B. thuringiensis, helping the 6-endotoxin from the crystal destroy the first line of defense of the host, the epithelial cells. One of the exoenzymes, phospholipase C, is able to destroy cell membranes. Chitinases degrade the exoskeletons and connective tissues of the insects that contain chitin. B. thuringiensis produces only small amounts of chitinase and this enzyme pro^des only a minor contribution to the overall pathogenicity. In an attempt to improve commercial preparations, ■ Smirnoff - (1977) reported that the addition of chitinase increased • the pathogenicity of marketed B. thuringiensis products.

Use of Bacillus thuringiensis as a microbial insecticide

Along with several other very effective entomopathogenic toxins, - B. thuringiensis has been widely used as a microbial insecticide in agriculture. Krieg (1981) has reported a list of over 290 different species of Lepidoptera susceptible to the bacterium. In the United States, B. thuringiensis was permitted by the Food and Drug Administration to be used on vegetables and forage crops starting in 1960. B. thuringiensis has been proven to be non-toxic to chicks, lajdng hens, hogs, honey bees or guinea pigs (Fisher and Rosner, 1959), nor is it harmful to birds, pets, or insects other than Lepidoptera (Ignoffo and Anderson, 1979). Inhalation or ingestion of B. thuringiensis by 18 human volunteers showed no toxic effects (Heimpel, 1971). However, in 1982, Ellar presented data that carbonate dissolved B. thuringiensis var. israelensis crystals are cytotoxic to insect and mammalian cells. In addition, some c£^es of diseases associated with B. thuringiensis (secondary infections) have occurred. 13

Commercially, B. thuringiensis Is produced by large scale deep tank fermentation. There is little information in the literature on media formulations and operating conditions, as most of this information is held as trade secrets. In the late 1950's and during the 1960’s, several companies in the United States produced B. thuringiensis var. thuringiensis as a liquid, wettable powder, or dust. There were four major. brands, but they were not consistent in potency from lot to lot. This resulted in poor product acceptance and created a negative attitude to the usefulness of B. thuringiensis. As more studies were done, new isolates of the serotypes were found which exhibited significant increases in toxicity to the same insect species. In 1969, Dulmage (1970) isolated HD-1 (of the B. thuringiensis variety, kurstaki) which was 16 times more toxic than B. thuringiensis var. thuringiensis being used in commercial preparations at this time. In collaboration with industry and the Environmental Protection Agency, he also initiated the International Unit (lU) and prepared the bacterial standard, HD-l-S-1971. All commercial preparations are compared to this standard in their ability to kill insects. This standardization was responsible for improved field results without variation from lot to lot, and in turn, resulted in the increased agricultural use of B. thuringiensis. Before this, potency was determined by spore count, which was not necessarily a good measure of toxicity.

Dulmage, and some twenty scientists in the International Cooperative Program on the Spectrum of the Activities of B. thuringiensis, have compared toxicity of the serotypes against 23 different insects and discovered great variety in the insecticidal activity spectrum. In Table 1.2 are B. thuringiensis powders (and their strains) that are superior to HD-1 in controlling insects.

Today, B. thuringiensis is used as the exclusive insecticide to control the cabbage looper, Trichoplusia ni, on cole crops. It is also used on grape, pecan, tobacco, soybean, and vegetable crops and forests. In Southeast Asia, the Chinese and neighboring countries are actively pursuing its use on the rice leaf roller, Cnaphalocrods medinalis, an insect that destroys rice, a crop that is grown on one-third of the world’s irrigable land. 14

TABLE 1.2. Comparison of the potency of HD-1 powder with the potency of superior B. thuringiensis powders to different insect species.

superior powders

Insect Potency* Potency* Strain Serotype Crystal % Increase of HD-1 of superior type over HD-1 strain

T. nl 39.800 52.400 HD-231 Kurstaki k-1 32 H. vlrescens 16.400 70.600 HD-244 kurstaki k-73 358 H. cunea 47.500 143.000 HD-203 kurstaki k-1 201 B. m o n 50.500 92.800 HD-137 alzawal alz 84 0. nubllalls 29.700 70.900 HD-168 kurstaki k-1 139 S. lltura 22.900 96.000 HD-137 alzawal alz 319

‘Potencies expressed in lU/mg crystal protein,

(after Dulmage, 1981)

Bacillus thuringiensis vw. israelensis

A new strain of Bacillus thuringiensis was isolated by Goldberg and Margalit (1977), that is unique from other B. thuringiensis strains because of its toxicity to mosquitoes. The strain, B. thuringiensis var. israelensis, was isolated from dead larvae of the mosquito, Culex pipiens, taken from a small pond in a dried out river bed in the North Central Negev Desert, near the Zeelim Kibbutz in Israel. Two cultures that originated from an individual colony, called 60A, were sent to deBarjac for characterization (for a review, see Margalit and Dean, 1985). She found 60A (later to be called ONR60A) to have some characteristics similar to other varieties of B. thuringiensis. The strain is a Gram positive, facultative anaerobic bacillus that forms spores and crystals. It has many metabolic activities in common with other strains, such as fermentation of ribose, glucose, and maltose and the production of RNase, phosphatase, and catalase. B. thuringiensis var. israelensis differs from other varieties in the production of 15 flagellar antigen H-14, which does not agglutinate with antisera to other serotypes. Of great commercial importance, B. thuringiensis var. israelensis produce crystals that are highly toxic to mosquito larvae. These crystals are of different shapes and sizes, unlike the organized bipyramidal crystals of other B. thuringiensis varieties that are not toxic to mosquitoes.

The research in this dissertation investigates B. thuringiensis var. israelensis, and was initiated several years after the description of the strain by deBarjac (1978) . More specifically, this dissertation describes the characterization of the plasmids of B. thuringiensis var. israelensis, notably concentrating on restriction enzyme analysis and association of phenotypes to plasmids. However, before discussing the plasmids of B. thuringiensis var. israelensis, it is appropriate to review some basic properties of plasmids.

Plasmids

Plasmids are autonomous extrachromosomal elements that are stably inherited. They are usually composed of double stranded DNA and found in both prokaryotic and eukaryotic organisms. Their sizes range from 1.5 kilobase (kb) pairs to greater than 300 kb. The monomeric unit size of each plasmid is constant, but plasmids may be found as dimeric, trimeric, and even larger multimeric sizes in some organisms. Plasmids carry no genetic information which is essential for cell growth. They carry opportunistic genes that encode for phenotypes such as transfer fertility, degradative enzymes, catabolic enzymes, toxins, .invasiveness, hemolytic activity, restriction and modification systems, heavy metal resistance, plant tumor induction, and bacteriocins.

The term plasmid was coined in 1952 by Joshua Lederberg, who first discovered the F factor (plasmid) in Exoli. Since that time, hundreds of other plasmids have been isolated and studied from E.coli. as well as other bacteria. With the advent of recombinant DNA techniques, thousands of variant plasmids have been constructed. Examples of well studied plasmids in Exoli are the F fertility plasmids, the R antibiotic resistance plasmids, the colicin producing. Col E l plasmid, and the pBR322 series, used extensively in genetic engineering as vectors. 16

Plasmid^ have also been studied in B. subtilis. This organism is second only to E.coli in terms of our understanding the organism’s genetics. Lovett and Bramucci (1975) were the first to describe naturally occurring plasmids in B. subtilis. They found that only two of eighteen strains examined had plasmid DNA. These plasmids were classified as cryptic, because they had no biological function. Four other cryptic plasmids were isolated by Tanaka et al. (1977), as well as four unique plasmids discovered by LeHegarat and Anagnostopoulos (1977) after an extensive search of 83 B. subtilis strains.

The ultimate aim in the search for plasmids in B. subtilis is to develop a genetic engineering system. This organism would be safer than E. coli since it is a non-pathogen, grows aerobically, and is not part of the normal flora of man. In developing a good cloning vector, selectable markers are beneficial. None of the plasmids from strains of B. subtilis encode resistance to antibiotics. Furthermore, plasmids were not isolated from B. subtilis 168, the most extensively studied member of the subtilis species. To overcome these problems, plasmids were transformed into B. subtilis from other bacterial species. The antibiotic resistant plasmids pCl94, pS194, pEl94, and pSC194 from Staphylococcus aureus have been transformed into competent B. subtilis 168 conferring antibiotic resistance to chloramphenicol, streptomycin, erythromycin, and both streptomycin and chloramphenicol, respectively.

Plasmids in Bacillus thuringiensis

Plasmids were first, isolated in B. thuringiensis by Zakharyan et al., (197.6) . They first purported that plasmids play a role in coding for the biosynthesis of crystal and isolated three plasmids from B. thuringiensis var. caucasicus having sizes of 9, 15, and 135 kilobase pairs. Stably et al. (1978b) examined 12 plasmids in B. thuringiensis var. alesti by electron microscopy. Soon afterward, Miteva (1978) presented the first extensive study detecting plasmids in serotypes 1 -11. Most of the plasmids isolated in these serotypes have sizes ranging from. 4.5-13.5 kilobase paire and they are present as covalently closed circular (CGC) forms as well as open circular (00) forms, In a study by lizuka et al. (1981) , clear lysates 17 of cultures from 17 serotypes were examined for the presence of plasmids. They observed that the number of plasmid bands based on agarose gel electrophoresis profiles ranged from one, for B. thuringiensis var. soUo and var. thompsoni, to 16, for B. thuringiensis var. kurstaki. The sizes of the CCC form range from less than 1 kilobase pair to greater than 180 kilobase pairs. A representative of every serotype possessed at least one plasmid. Such diversity is not seen in B. subtilis or any other member of the genus Bacillus . 18

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activity against Anopheles sergentii, Uranotaenia. unguiculata, Culex univitattus, Aedes aegypti, and Culex pipiens. Mosquito News, 1977, S7, 355-358. Gordon, R.E., Haynes, W.C., and Pang, C.H.-N. The genus Bacillus. In Agricultural Handbook, Vol. 4^7, United States Department of Agriculture, 1973. Griego, V.M., Fancher, L.J., and Spence, K.D. Scanning electron microscopy of the disruption of tobacco hornworm, Manduca sexta, midgut by Bacillus thuringiensis endotoxin. J. Invertebr. Pathol., 1980, 35, 186-189. Hall, I.M. and Arakawa, K.Y. The susceptibility of the housefly, Musca domestica Linnaeus, to Bacillus thuringiensis var. thuringiensis Berliner.. J. Insect Pathol., 1959,1, 351-355. Hanny, C.L. Crystalline inclusions in aerobic sporeforming bacteria. Nature, 1953, 173, 1004. Hanny, C.L., and Fitz-James, P. The protein crystals of Bacillus thuringiensis Berliner. Canadian J. Microbiol., 1955, 1, 694-710. Heimpel, A.M. Safety of insect pathogens for man and vertebrates. In Burges, H.D. and Hussey, N.W. (Ed.), Microbial Control of Insects and Mites, Academic Press, New York, pp. 469-487, 1971. Holmes, K.C., and Monro, R.E. Studies on the structure of parasporal inclusions from Bacillus thuringiensis. J. Mol. Biol., 1965, 14, 572-581. Huber, H. E. Zur spezifitat des delta-endotoxins von Bacillus thuringiensis. Diplorha thesis. Faculty Nat. Sd., ETH, 1977, Zurich, Switzerland, . Huber, H.E., Luthy, P., Ebersold, H.R., and Cordier, J.L. The subunits of parasporal crystal of Bacillus thuringiensis: size, linkage, and toxicity. Arch. Microbiol., 1981, ISO, 14-18. Husz, B. Bacillus thuringiensis Berl., a bacterium pathogenic to corn borer larvae. A preliminary report. International Com Borer Invest. Sd. Rpts., 1927, 1987- 1928, 191-193. Ignoffo, C.M. and Anderson, R.F. Bioinsecticides. In Peppier, H.J. and Perlman, D. (Ed.), Microbial Technology 2nd ed.. Academic Press, New York, pp 1-28, 1979. lizuka. T., Faust, R.M., Travers, R.S. Comparative profiles of extrachromosomal 21

DNA in single and multiple crystalliferons strains of Bacillus thuringiensis var. kurstaki. J. Fac. o f AgricuL, 1981, 60, 143-151. Isakova, N.P. Une nouvelle variété dé bacterie du type cereus pathogène pour les insectes. Dokl. Akad. Sd. Navk. Selsk., 1958, 23, 26-27. Ishiwata, S. Examinations of sotto disease Bacillus. Kyoto Sangyo Koshujo Sanji Hokoku, 1902, 2, 346-347. Ishiwata, S. One kind of severe flacherie (sotto disease). Dainihon Sanshi Kaiho, 1901, 9, 1-5. Johnson, D.E. Bacterial membrane transport of )$-exotoxin, an anti-metabolite of RNA synthesis. Nature, 1976, 260, 333-335. Kaneda, T. Fatty acids in the genus Bacillus. II. Similarity in the fatty acid compositions of Bacillus thuringiensis, Bacillus anthracis, and Bacillus cereus. J. Bacteriol., 1968, 95, 2210-2216. Kaneko, T., Nozaki, R., and Aizawa, K. Deoxyribonucleic acid relatedness between BoctMus anthracis, Bacillus cereus, Bacillus thuringiensis. Microbiol. Immunol., 1978, 22, 639-641. Kim, Y.T., Gregory, B.C., and Ignoffa, C.M. The beta-exotoxins of Bacillus . thuringiensis. HI. Effects on in vivo synthesis of macromolecules in an • insect system. J. Invertebr. Pathol., 1972, 20, 46-50. Krieg, A. In vitro determination of Bacillus thuringiensis. Bacillus cereus, and related bacilli. J. Invertebr. Pathol., 1969,15, 313-320. Krieg, A. Concerning a-exotoxin produced by vegetative cells of Bacillus thuringiensis and Bacillus cereus. J. Invertebr. Pathol., 1971, 17, 134-135. Krieg, A. and Langenbruch, G A. Susceptibility of arthropod species to Bacillus thuringiensis. In Burges, HJ). (Ed.), Microbial Control o f Pests and Plant Diseases, Academic Press, London, pp. 851-874, 1981. Krieg, A. and Lysenko, O. Toxine und enzyme bei einigen Bacillus-Artcn unter besonderer berucksichtigung der Bacillus cereus-thuringiensis gruppe. Zbl. Bakt. II Abt., 1979, 134, 70-88. Labaw, L.W. The structure of Bacillus thuringiensis Berliner crystals. J. Ultrastruct. Res., 1964, 10, 66-75. Lecadet, M.M., and Dedonder, R. . . Bull, Soc. Chim. Biol., 1966, 48, 631-:659. 22

Lecadet, M.M., and Martouret, D. . Entomophaga,Mem. Hors. Ser., 1964, 2, 205-212. LeHegarat, J-C, and Anagnostopoulos, C. Detection and characterization of naturally occurring plasmids in Bacillus subtilis. Molec. Gen. Genet., 1977, i57, 167-174. Lovett, P.S., and Bramucci, M.G. Plasmid deoxyribonucleic acid in Bacillus subtilis and Bacillus pumilus. J. Bacteriol., 1975, 124, 484-490. Margalit, J. and Dean, D.H. The story of Bacillus thuringiensis var. israelensis. J. Amer. Mosquito Control Assoc., 1985, 1, 1-7. Mattes, O. Parasitare krankheiten der meulmottenlarven und versuche uber ihre verwendbarkeit als biologisches bekampfungsmittel. Sitzber. Ges. Befoerder. Ges. Naturw. Marburg, 1927, 62, 381-417. McConnell, E. and Richards, A.G. The production by Bacillus thuringiensis Berliner of a heat stable substance toxic for insects. Canadian J. MicrofcioL, 1959, 5,161-168. Milne, R., Murphy, D., and Fast, P.G. Bacillus thuringiensis delta-endotoxin: An improved technique for the separation of crystals from spores. J. Invertebr. Pathol., 1977, 29, 230-231. Miteva, V.I. Isolation of plasmid DNA from various strains of Bacillus thuringienèis Q,nd. Bacillus cereus. Comptes, rendus de I’Acad Burgare de Sd., 1978, SI, 913-916. Nickerson, K.W., and Swanson, J.D. Removal of contaminating proteases from Bacillus thuringiensis parasporal . crystals by density gradient centrifugation in NaBr. European. J. Appl. Microbiol. Biotechnol., 1981, IS, 213-215. Nishimura, M.S. and Nishiitsutsuji-Uwo, J. Sporeless mutants of Bacillus thuringiensis. m . The process of crystal formation. Tissue and Cell, 1980, 233-242. Ohba, M., Tantichodok, A., and Aizawa, K. Production of heat-stable exotoxin by Bacillus thuringiensis and related bacteria. J. Invertebr. Pathol., 1981a, S8, 26-32. Ohba, M., Aizawa, K., and Shimizu, S. A new subspecies of Bacillus thuringiensis isolated in Japan: Badllus thuringiensis subsp. tohokuensis (serotype 17). J. Invertebr. Pathol., 1981b, S8, 307-309. 23

Ohba, M., Ono, K., Aizawa, K., and Iwanami, S.- Two new subspecies of Bacillus thuringiensis isolated in Japan: Bacillus thuringiensis subsp. kumamotoensis (serotype 18) and Bacillus thuringiensis subsp. tochigiensis (serotype 19). J. Invertebr. Pathol., 1981c, 38,184-190. Prystas, M., Kalvoda, L., and Sorm, F. Alternative synthesis of exotoxin from Bacillus thuringiensis. Coll. Czech. Chem. Commun., 1976, 1426-1447. Sebesta, K. and Horska, K. Mechanism of inhibition of DNA-dependent RNA polymerase by exotoxin of Bacillus thuringiensis. Biochim. Biophys. Acta, 1970, 209, 357-376. Seki, T., Chung, C.K., Mikami, H., and Oshima, Y. Deoxyribonucleic acid homology and taxonomy of the genus Bacillus. Interriation. J. Systematic Bacteriol., 1978, 28, 182-189. Sharpe, E.S., Herman, A.I., and Toolan, S.C. Separation of spores and parasporal crystals of Bacillus thuringiensis by flotation. J. Invertebr. Pathol., 1979, 34, 315-316. Smirnoff, W.A. Confirmations expérimentales du potentiel du complexe Bacillus thuringiensis et chitinase pour la repression de la tordeuse des bourgeons de l’epinette, Choristoneura fumiferana (Lepidoptera: tortricidae). Canadian Entomol., 1977, 1Ô9, 351-358. Sommerville, H.J. and Pockett, H.V. An insect toxin from spores of Bacillus thuringiensis and Bacillus cereus. J. Gen. Microbiol., 1975, 87, 359-369. Stahly, D.P., Dingman, D.W., Irgens, R.L., Field, C.C., Feiss, M.G., and Smith, G.L. Multiple extrachromosomal deoxyribonucleic acid molecules in Bacillus thuringiensis. FEMS Microbiol. Letters, 1978b, 3, 139-141. Steinhaus, E. A. A study of the bacteria associated with thirty species of insects. J. Bacteriol., 1941, 757-790. Steinhaus, EA. Possible use of Bacillus thuringiensis Berliner as an aid in the biological control of the alfalfa caterpillar. Hilgardia, 1951, 20, 359-381. Talalaev, E.V. Septikamie de Raupen des sibirischen Awenspinners. Microbiol. Moskva, 1956, 25, 99-102. Tanaka, T., Kuroda, M., and Sakaguchi, K. Isolation and characterization of four plasmids from Bacillus subtilis. J. Bacteriol., 1977, 129, 1487-1494. 24

Travers, R.S., Faust, R.M., and Reichelderfer, C.F. Effects of Bacillus thuringiensis var. kurstaki on isolated Lepidoptera mitochondria. J. Invertebr. Pathol., 1976, .28, 351-356. Yamamoto, T. IdentiOcation of entomocidal toxins of Bacillus thuringiensis by high performance liquid chromatography. J. Gen. Microbiol., 1983a, 129, 2595-2603. Yamamoto, T., and McLaughlin, R.E. Isolation of a protein from the parasporal crystal of Bacillus thuringiensis var. kurstaki toxic to the mosquito larva, Aedes taeniorhynchus. Biochem. Biophys.. Res. Commun., 1981, lOS, 414-421. Zakharian, R.A., Agabalian, A.S., Chil-Hakobian, LA.., Gasparian, N.S., Bakunz, KA., Tatevosian, P.E., and Afrikian, E.K. On the possible role of the extrachromosomal DNA in the biosynthesis of entomocide endotoxin of Bacillus thuringiensis. Dokl. Akad. Nauk. ArmSSR, 1976, 6S, 42-47. CHAPTER TWO Isolation Of Plasmid DNA From B. thuringiensis var. israelensis Strains And The Curing Of These Plasmids From Strain 4Q2-WT

Introduction

As reviewed in Chapter One, B. thuringiensis, upon sporulation, is capable of producing an intracellular crystal which is highly toxic to Lepidopteran or Dipteran larvae, depending on the varietal type. Many of these varieties have been shown to have a large number of plasmids (Miteva, 1978, lizuka et al., 1981). At the time this study was initiated, it was not known if B. thuringiensis var. israelensis possessed plasmids. The first few experiments described in this chapter confirmed that B. thuringiensis var. israelensis h^boured plasmids. This led to one of the main research objectives of these studies; to characterize and gain a greater knowledge about these cryptic plasmids.

As a start to characterize the cryptic plasmids, it may be possible to associate crystal production with a plasmid as shown for other B. thuringiensis varieties. It was suggested by Stahly et al. (1978a, 1978b) that the very high frequency of loss of crystal production in the variety alesti, could be attributed to plasmid instability, although other hypotheses exist. Once the crystal phenotype was lost, it was never regained. This is suggestive of loosing the gene(s) and possibly a plasmid which carries the gene(s), and not a point mutation within the DNA which could later revert. Several years later, the crystal gene was isolated from a large plasmid of the variety kurstaki. The gene was ligated into pBR322, and then expressed in Escherichia coli (Schnepf and Whiteley, 1981), proving that B. thuringiensis plasmids can encode the crystal phenotype.

25 26

Other phenotypes that may be encoded by B. thuringiensis var. israelensis plasmids are resistance to antibiotics and heavy metals. At present, there are few examples of plasmids that are responsible for these phenotypes in Bacillus species. Two naturally occurring examples are pBC16, a tetracycline resistance plasmid from B. cereus (Bernhard et al., 1978), and pPLlO, a plasmid of B. subtilis which codes a bacteriocin-like activity (Lovett and Bramucci, 1976). Other plasmids that code for antibiotic resistance have been introduced into B. subtilis and B. thuringiensis from Staphylococcus aureus. There are no known Bacillus plasmids coding for heavy metal resistance. However, plasmids with this phenotype have been found in Staphylococcus, Escherichia, Salmonella and Pseudomonas.

In this chapter, several experiments are described which were performed to gain greater knowledge about the extrachromosomal elements in B. thuringiensis var. israelensis strains. The initial experiments developed methods to isolate plasmids present in B. thuringiensis var. israelensis, and later experiments focused on preparing mutants of the B. thuringiensis var. israelensis strain, 4Q2, to facilitate the study of the individual plasmids. Additionally, the plasmid profiles of B. thuringiensis var. israelensis strains used by other academicians and industry are compared to the wild type and plasmid cured mutants of strain 4Q2.

MATERIALS and METHODS

Bacterial strains. The strains of B. thuringiensis var. israelensis and their sources are given in Table 2.1. Strain W4Q47 was isolated from a sample of the commercial insecticide, Teknar (Sandoz, Inc., San Diego, CA.). Escherichia coli strain V517 was a kind gift of Dr. F. Macrina (Macrina et al., 1980), and strains J53(RA1), J53(Rl), C600(Rtsl), and C600(pDAC2) were kindly provided by Dr. D. Coplin, The Ohio State University, Columbus, Ohio. These E. coli strains harboured plasmids that were used as size markers to compare their relative mobility to the B. thuringiensis var. israelensis plasmids (Meyers et al., 1976). 27

Table 2.1. List of B. thuringiensis var. israelensis strains used in these studies.

strain Source

4Q2-WT BGSC^ 4Q2-22 Clark, B. 4Q2-32 Clark, B. 4Q2-50 Clark, B. 4Q2-52 Clark, B. 4Q2-S6 Clark, B. 4Q2-58 Clark, B. 4Q2-61 Clark, B. 4Q2-63 Clark, B. 4Q2-64 Clark, B. 4Q2-66 Clark, B. 4Q2-69 Clark, B. 4Q2-72 Clark, B. 4Q2-73 Clark, B. 4Q2-74 Clark, B. 4Q2-75 Clark, B. 4Q2-76 Clark, B. 4Q2-78 Clark, B. 4Q2-79 Clark, B. 4Q2-80 Clark, B. 4Q2-81 Clark, B. 4Q1 BGSC* Btl U.S. 8-1982 Beegle, C. IPS 78 deBarjac, H. IPS 80 Beegle, C. (lot 91413) IPS 82 Beegle, C. WHO 1897 Yousten, A. WHO 1884 Yousten, A. WHO 1887 Yousten, A. WHO 1897 M24 spo- Yousten, A. BactlDos Primary Powder Ross, 0.; Biochem Products Teknar (W4Q47) Sandoz, Inc. B.t. 82161 II Uelln, B. 795/11 1 CHD648) Couch, T. 795/11 2 (HD649) Couch, T. 57-499 CD Vectobae Technical Dewisetty, B.; Abbott Labs WHO 2013-1 Singer, S. WHO 2013-2 Singer, S. WHO 2013-3 Singer, S. WHO 2013-5 Singer, S. WHO 2013-7 Singer, S. WHO 2013-9 Singer, S. WHO 2013-10 Singer. S. WHO 2013-11 Singer, S. ^ Bacillus Genetic Stock Center, The Ohio State University, 28

Plasm id curing. Three methods -were used to cure plasmids from B. thuringiensis var. israelensis. Two methods involved the growth of the vegetative cells either at an elevated temperature, or in the presence of coumermycin. The third method involved the heat shocking of spores. To cure plasmids by growing bacteria at an elevated temperature, a modification of the technique to isolate asporogenous mutant strains of B. subtilis was used (Yousten, 1978). The B. thuringiensis var. israelensis strain 4Q2-WT or derived mutants were routinely inoculated in NSM broth (8g/l nutrient broth [Difco], 7x10'^ M CaClg, 5x10-® M MnClg, and 1x10"^ M MgClg) (Li and Yousten, 1975), and incubated at 41°C - 44°C for 24 h., followed by streaking for isolated colonies on NSM agar (NSM broth with 1.5% agar) plates and incubating again at 41°C - 44°C for 24 h. Distinct colonies that were sporogenous, but more translucent than surrounding colonies, were chosen to isolate their plasmid DNA.

B. thuringiensis var. israelensis grown in L broth (tryptone, 1.0%; yeast extract, 0.5%; and NaCl, 0.5%) with courmermycin (5 /ig/ml) was also used to cure plasmids. This antibiotic has been shown to inhibit gyrase in E. coli (Danilevskaya and Gragerov, 1980). Cultures were incubated at 30°C for 24 h. before streaking for isolated colonies on NSM plates and incubating at the same temperature for 24-48 h. The last method used to cure plasmids involved heat shocking wet spores at 90°C for 1 h. prior to spreading on NSM agar plates for isolated colonies and then incubating at 30°C for 24-48 h.

Isolation of plasm id DNA. Several methods used to isolate plasmid DNA from B. thuringiensis var. israe/ensfs include a modification of the procedure described by Eckhardt (1978) , an SDS lysis procedure, and a polyethylene glycol (PEG) procedure, which is a modification of the method described by Kronstad et al. (1983) . In the modified Eckhardt procedure, an isolated colony putatively cured of plasmids was inoculated into 5 ml SCGYlO broth ( (NH^)gSO^, 0.2%; KgHPO^, 1.4%; KHgPO^, 0.6%; sodium citrate'7HgO, 0.1%; casamino acids [Difco], 0.1%; MgSO^ 7HgO, 0.02%; glucose, 0.5%). The glucose was autoclaved separately as a 50% stock solution and added to the above broth after it had been autoclaved and cooled to room temperature. Cultures were incubated overnight at 30°C with shaking at 150 rpm on an orbital shaker. An equal 29

VQlume of L broth was added to the SCGYlO broth culture and Incubated an additional 2 h. as before. One ml of this culture was transferred into 30 ml SCGYlO broth in a 100 ml Erlenmyer flask and incubated at 30°C with shaking as before to an O.D. (660 nm) = 0.35. The cells were pelleted by centrifugation at 4300 xg and resuspended in 0.5 ml SET (0.25 M Tris, pH 8.0; 0.25 M EDTA, 0.92 M sucrose) with 2-4 mg/ml lysozyme (Sigma Chemical Co., egg white, lyophilized 3X) and incubated at 37°C to remove the bacterial cell wall and create protoplasts. The conversion of cells to protoplast was followed by observing samples using phase contrast microscopy. When greater than 90% of the cells existed as protoplasts, 3-7 /il was placed under a 20 /d detergent layer (SDS, 2%; ■ sucrose, 5%; bromophenol blue, 0.05%) in the wells of a vertical agarose gel to separate the plasmids by electrophoresis as described below (Gonzalez et al., 1981).

The second plasmid isolation method used SDS to lyse the cells. The steps for this procedure consisted of inoculating 5 ml SCGYlO broth as a starter culture and incubating overnight at 30°C with shaking at 200 rpm. One ml of the starter culture was then added to 100 ml SCGYlO broth in a 250 Erlenmyer flask followed with shaking of the culture at 150 rpm for 5 h. at 30°C. The cells were collected by centrifugation at 4300 xg for 15 minutes at 4°C.. The pellet was resuspended in 1.5 ml SET with 10 mg/ml lysozyme and incubated at 37°C with agitation (75 rpm) until grea,ter than 80% protoplast formation was observed. The protoplasts were lysed by adding an equal volume of 8% SDS and stored at 4°C overnight. A precipitate formed with the SDS and was removed by centrifugation at 43000 xg in a Beckman JA-20 rotor for 35 minutes at 0°C. The supernatant was gently decanted and 20-30 fi\ of this cell lysate was separated by agarose gel electrophoresis.

The final method used to isolate plasmids from B. thuringiensis var. israelensis was the PEG method, modified from the procedure described by Kronstad et al. (1983) . Cells to be used as a starter culture are prepared by inoculating 30 ml SPY media (Spizizen medium supplemented with 0.1% yeast extract and 0.1% glucose) and incubating overnight (14 h.) at 30°C and 150 rpm. Five hundred ml SPY media in a 2 liter Erlenmyer flask was inoculated with 15 30 ml of the starter culture and then incubated at 30°C with agitation at 175 rpm until an OD. (660 nm) of 0.7-0.8 was reached. The cells were then centrifuged at 4300 xg at 4°C and the pellet resuspended in 5 ml of IX TE buffer (0.05 M Tris, 0.02 M EDTA, pH 7.9). Cell lysates were prepared by adding 95 ml of IX TE with 1% SDS and 0.085 M NaOH, pH 12.1-12.3. After 30 minutes at room temperature, 10 ml of 10% SDS was added and cell lysis continued by gentle mixing. Finally, 10 ml of 2 M Tris (pH 7.0), was added and gently mixed, and then 30 ml of 5 M NaCl was added and the sample stored in an ice-water bath overnight. The precipitate formed was removed by centrifugation at 12000 xg for 15 minutes at 4°C. Thirty-six ml of 50% PEG 8000 was added to the supernatant, mixed thoroughly by gentle inversion of the sample in a centrifuge bottle, and stored in an ice-water bath for 6-8 h. The PEG-plasmid precipitate was then centrifuged at 5000 xg for 5 minutes at 4°C. The supernatant was poured off and any remaining supernatant removed with a pasteur pipette connected to a vacuum. The pellet (plasmid DNA and PEG) was resuspended in 2-4 ml IX TE buffer. Samples of 10-20 ftl were separated by agarose gel electrophoresis.

Agarose Gel Electrophoresis. Both vertical and horizontal agarose gels" were routinely used to separate and visualize plasmid DNA. For plasmids' isolated by the Eckhardt procedure, 1.5 mm thick, 0.5% vertical agarose (Sigma high-strength Type V agarose) gels were prepared using Tris-borate buffer (0.089 M Tris, 0.89 M boric acid, pH 8.3), the same buffer as used in the reservoirs. Electrophoresis was carried out at 15 volts (V), 3. milliamperes (mA) for 2 h., then 40 V, 8 mA for 0.8 h., and finally 110 V, 27 mA for 4.5 h. using an ISCO model 493 power supply. This was done to slowly pull the detergent through the protoplasts (15 V), then to slowly allow the DNÂ to enter the gel (40 V), and finally to resolve the DNA (110 V). To visualize the DNA separated on the agarose gel, ethidium bromide (0.5 /ig/inl) in HgO was used to stain the DNA bands. Ethidium bromide intercalates between the bases of DNA or RNA molecules and is fluorescent when stimulated with ultraviolet light (Sharp, 1973). The gels were stained for 30 minutes and then destained with HgO for 4-24 h. Plasmid bands were visualized using a Fotodyne ultraviolet (300 nm) 31 transilluminator (Fotodyne Inc.., New Berlin, ,WI.) and photographed using a Poloroid MP-4 Land camera with a Vivitar #25 red filter and Poloroid type 667 film. Film was exposed for 0.5-1 second at f=4.5.

Horizontal agarose gel electrophoresis was used to visualize the plasmids isolated by SDS lysis or PEG, as described above. A 0.7% agarose gel (Seakem agarose, type LE, FMC Corp., Maine Colloids Div., Rockland, ME.) was prepared in IX TAE buffer (0.05 M Tris, pH 8.05; 0.02 M sodium acetate, 0.002 M EDTA). DNA was transferred into wells of a 150 ml, 10 cm by 14 cm horizontal gel. Wells were 8 mm long by 3 mm wide. Electrophoresis was preformed at 7 V/cm for 5 h. Agarose gels were stained.in water with 0.5 Aig/ml ethidium bromide for 30 minutes and then destained in HgO for 24 h. The results were photographed in the same manner as described above for vertical agarose gels.

Serotyping of Strains. Motile cells for serotyping were isolated by first removing the outer 10 mm of agar of a 100 mm diameter nutrient agar plate (nutrient broth [Difco], 0.8%, pH 7.0; agar, 0.2%). This was done with a 5 ml sterile pipet attached to a vacuum. The pipet was held steady while the petri . dish was rotated on a turntable. This helped in removing the same amount of. agar all around the plate. Five ml of nutrient broth (0.8%, pH 7.0) was placed in the trough formed. The center of the plate was then inoculated with motile cells taken from the outer edge of a colony grown on nutrient agar for 12-14 h., and the plate incubated at 30°C. Motile cells were allowed to migrate from the center, across the 0.2% agar, and into the 5 ml nutrient broth. The broth was then removed and the motile cells concentrated by centrifugation at 4300 xg for 10 minutes. The pellet of motile cells (antigen) was resuspended in 5 ml 0.6% formalized saline (NaCl, 8.7 g/1; and formaline, 6 ml/1) and stored at 4°C. The flagella antigen of the motile cells is stable in this solution at 2-8°C for at least 4 months. Antisera (kind gift of G. Donaldson, USDA, Brownsville, TX) was diluted 1:20, 1:40, and 1:80 with saline. One hundred fA of antisera was mixed with 900 ft\ antigen (motile cells in formalized saline) and incubated at 37°C for 1 h. before looking for agglutination with low power microscopy. These assays were done in triplicate to provide better statistical results. 32

Storage of Strains. For short term storage (2-3 weeks), bacterial strains were inoculated on NSM plates and incubated at 30°C for 48 h. to allow sporulation and then saved as isolated colonies at 4°C. Plates were wrapped with parafilm and stacked inverted to avoid condensation from dripping on the bacterial colonies and causing spreading of spores and vegetative cells. For long term storage of sporogenous strains, an isolated colony was used to inoculate 100 ml NSM broth in a 150 ml centrifuge bottle and the culture incubated for 72 h. at 30°C with agitation at 225 rpm. The sporulated culture was then centrifuged in the same bottle at 4300 xg for 15 minutes at 4°C and the pellet resuspended in 3 ml freezer media (0.8% nutrient broth [Difco] and 20% glycerol). One ml aliquots were placed in freezing vials for storage in liquid nitrogen. Liquid nitrogen frozen stocks were used to initiate all new experiments, to decrease the possibility of contamination or strain differentiation. The above protocol was also used for asporogenous strains, except cultures were grown only for 24 h. before storing the vegetative cells. 33

Results

Plasmid DNA was isolated from B. thuringiensis var. israelensis 4Q2 wild type using the modified Eckhardt procedure and separated on a vertical 0.5% agarose gel. The results are seen in Figure 2.1. The sizes of the plasmids were calculated by comparing their migration to plasmids from E. coli V517 and E. coli plasmids pRal, pRl, pRstl, and pDAC2 (data not shown). They were 5.4, 6.5, 7.5, 14.7, 15.8, 98, 108, 165, and 195 Kb. The 14.7 kb band is linear DNA and was not always seen after separating plasmid DNA on an agarose gel, especially when the isolation method had an alkali treatment step used to reduce the amount of sheared linear chromosomal DNA. All the other bands separated on the agarose gel that were less than approximately 90 kb were observed in both their closed covalently circular (CCC) form and open circular (OC) form. The conformation of the plasmid bands was determined by irradiation of CCC DNA in the presence of ethidium bromide. Ultraviolet light generates nicks and converts the DNA into the OC form. The two forms of plasmid DNA were identified after the electrophoretic separation of irradiated plasmids juxtaposed to lanes with non-irradiated plasmids. The four largest plasmids were not seen in their OC forms, even when treated with ethidium bromide and ultraviolet light before being resolved by agarose gel electrophoresis. However, there was an increase of ethidium bromide stained DNA near the top of the gel suggesting the OC. form never entered the gel.

The pedigree of plasmid cured mutants derived from the 4Q2 wild type strain is shown in Figure 2.2. The mutants of the first group (4Q2-22, 4Q2-32, 4Q2-50, 4Q2-52, 4Q2-56, 4Q2-58, 4Q2-61, and 4Q2-66) were isolated after growing 4Q2- WT at 42°C for 24 h., diluting and spreading the cells on NSM plates, and picking cells from 56 colonies that had the same colony morphology but were more translucent than the other colonies. The cells from the majority of these translucent colonies did not produce crystals after observing them by phase microscopy, unlike the normal, opaque colonies which were crystalliferous. Strain 4Q2-66 was a spontaneous mutant that was isolated after growing and isolating DNA from 4Q2-WT 103 times. 34

$

199% 169" I#:':: - s i

(/I «/) I 14.74

7.SH

6 J -

5.4-

Figure 2.1. Plasmid.pattern of B. thuringiensis var. israelensis strain 4Q2- WT (wild type). The vertical bar indicates where linear chromosomal DNA migrates on this 0.7% agarose gel. 35

The second group of mutants (4Q2-62, 4Q2-63, and 4Q2-64) were isolated after growing 4Q2-32 at 44°C for 24 h. and isolating plasmids from 20 randomly selected colonies. Mutants 4Q2-73 and 4Q2-74 were isolated from translucent colonies of 4Q2-32 on NSM agar plates that were stored at 4°C for two months and had become very dry. Mutant 4Q2-69 was isolated after screening only 18 colonies of heat treated (44°C) 4Q2-63. Additional plasmids were cured from 4Q2-69 by growing the strain in L-broth with 5 /ig/ml courmermycin, ten times more than required to inhibit gyrase in E. coli (Danilevskaya and Gragerov, 1980). After analyzing the plasmid DNA from 126 randomly selected colonies of courmermycin treated 4Q2-69, mutants 4Q2-72, 4Q2-75, 4Q2-76, 4Q2-78, and 4Q2-79 were isolated. Finally, 4Q2-80 and 4Q2-81, derived from 4Q2-79 and 4Q2-72 respectively, were each isolated after screening 5 colonies grown from spores that had been heated at 90°C for 1 h. (Stahly et al., 1978a).

Using the modified Eckhardt procedure for screening, plasmid profiles of the cured mutants are seen in Figures 2.3a and 2.3b. Comparison agarose gels of plasmids isolated from the same mutants, but using the PEG method are seen in Figures 2.4a and 2.4b. Plasmid DNA of 4Q2-WT was run on each gel as a standard to compare it to the plasmid DNA= in each mutant (data not shown). The plasmids present in these mutants are listed in Table 2.2. They are denoted pTXl4-l through pTXl4-9, with pTXl4-l being the smallest in size. The nomenclature "TX" is derived from two and three letter codes used to name plasmids that was started by Esther Lederberg. The two letter code for B. thuringiensis is TX, and 14 represents the variety (in this case, for israelensis).

All the strains derived from 4Q2-WT, with the exception of 4Q2-81, contain. 1-8 extrachromosomal elements, as detected using the plasmid isolation methods described in Materials and Methods. Strain 4Q2-81 is a plasmid-less variant of 4Q2. Furthermore, all the mutants are motile and react with H-14 antibody but do not react with anti-flagella antibody from B. thuringiensis var. kurstaki.

The plasmid proGles of B. thuringiensis var. israelensis strains other than mutants derived from strain 4Q2-WT are shown in Figure 2.4c and 2.4d and their plasmids listed in Table 2.3. As with 4Q2-WT, the plasmids fall into two 36

4Q2-WT

4Q2-58 4Q2-22 4Q2-32 4Q2-50 4Q2-52 4Q2-61 4Q2-664Q2-56

4Q2-62 4Q2-644Q2-63 4Q2-73 4Q2-74

4Q2-69

4Q2-72 4Q2-764Q2-75 4Q2-78 4Q2-79

4Q2-81 4Q2-80

Figure 2.2. Pedigree of plasmid cured mutants derived from B. thuringiensis VBT. israelensis strain 4Q2-WT. 37 categories, large plasmids greater than 75 kb, and small plasmids less than 75 kb. Although not always seen in Figure 2.4a through 2.4b, all the strains, with the exception of 4Q2-81, posses a 14.7 kb linear plasmid DNA (data not shown).

The plasmid profiles of strains used as standards by industry (IPS78, IPS80, IPS82, and S-1982) are seen in Figure 2.4c. IPS78 (Institute Pasteur Standard 1978) has all nine plasmids present, but IPS80, IPS82, and the United States B. thuringiensis var. israelensis standard, S-1082, are all missing the 7.5 kb, pTXl4-3 plasmid. The plasmid profiles from strains of B. thuringiensis var. israelensis used in commercial products in comparison to 4Q2 wild type show that both 57-499 ÇD and Bti 82161 II have lost the 7.5 kb, pTXl4-3 plasmid and the 15.8 kb pTX14-4 plasmid (Figure 2.4d). These two strains are used by Abbott Laboratories, North Chicago, Illinois in producing Vectobac. Teknar, a product of Sandoz, Inc., San Diego, CA, has all the plasmids, whereas Bactimos, manufactured by Biochem Products, Montchanin, DE, • has lost the 7.5 kb pTXl4-3 plasmid. Figures 2.3a and 2.3b. ModiHed Eckhardt lysate electrophoresis of B. thuringiensis var. israelensis plasmid cured mutants. The name of each mutant is written above its lane. Adjacent to each gel lane, the letters a - i correspond to the plasmids pTX14-l through pTXl4-9. These lysates were separated on different agarose gels, and due to time variations in electrophoresis, the migration distances of the plasmids are not the same. Lysate from 4Q2-WT was separated on each gel to use as a standard.

38 0 0>n e. • 4Q2-WT

û IT 4 0 2 -5 0

4 0 2 -5 2

4 0 2 -5 6

li 4 0 2 -2 2 ï* I

4 0 2 -3 2

4 0 2 -5 8

4 0 2 -6 4

4 0 2 -6 3

6S 4 0 2 -6 9

m 3 b 4 0 2 -8 0

4 0 2 -8 1

402-W T mfmë i i»*Vi

Ol' Figures 2.4à - 2.4d.Electrophoresis of plasmid DNA isolated by the modlHed PEG method of Kronstad et al. The name of each mutant or strain of B. thuringiensis var. israelensis is written above each lane. Letters a - i adjacent to each lane indicate the position of plasmids pTXl4-l through pTX14-9. Some of these plasmid preparations were separated on different gels and therefore the migration distances are not identical because of variations in the duration of electrophoresis.

41 1402-50

■ r 402-52

402-56

I 402-22 3i mm ™ am i

1402-32

402-58

402-64

4 0 2 -6 3

Zf 402-69

402-78

î 402-79 M a* 402-72

4 0 2 -8 0

402-81

402-WT

SI' 1884

1897

I 1887 3 ta IPS78 k'

IPS80

IPS82

S-1982

I___ 57-499CD

82161 n

795/n 1

A e y m to 7 9 5 /5 2 &

Taknar

Vjiijfe

kkeMm##

2013-2

Sî' 46

Table 2.2. Plasmids present in B. thuringienaia v s l t . .iaraelenaia strain 4Q2 mutants. 1 2 3 4 5 6 7 8 9

4Q2-WT + + + + + + • + + + 4Q2-22 + - — - + -- + +

4Q2-32 - + - + + ■ + - , , + + 4Q2-40 + - - - + -- + + 4Q2-44 + + - - + + - + + 4Q2-50 + + - - + + - ■ + + 4Q2-62 + + - - + + - . + + 4Q2-56 + + - - + + - + + 4Q2-58 -- + - + + - + ■ - 4Q2-63 - - + - + + + + + 4Q2-64 - - + + + + + + + 4Q2-66 - - + - + - + -- 4Q2-69 - - + - + - + + - 4Q2-72 - - -, - + + - - 4Q2-78 - - + - + + - ■ - 4Q2-79 -- + - + - + - - 4Q2-80 - - + - + -- -- 4Q2-81

Table 2.3. Plasmids present in B. thuringienaia var. iaraelenaia strains other than 4Q2 mutants. 1 2 3 4 5 6 7 8 9

57-499CD + + —— + + + + + 82161 II + + — - + + + + + 795/11 1 + + + + + + + + + 795/11 2 + + + + + + - + + Teknar + + + + + + + + ■ + Bactimos + + - + + + + + + 2013-2 + + + + + + + + + 1884 ' + + - - + + - + - 1897 + + - + + + - + + 1887 + + - + + ■ + - + + IPS-78 + + + + + + + + +

IPS-80 + + - + . + + + + + IPS-82 + + - + + + + + + S-1982 + + — + + + + + + ■47

Discussion . ’

The plasmid profiles of various strains of B. thuringiensis var. israelensis are all closely related to 4Q2 wild type, the strain stored in the Bacillus Genetic Stock Center (BGSC), The Ohio State University. This strain was donated to the BGSC by Dulmage from the U.S.D.A. laboratories. Hall originally donated the strain to Dulmage after receiving it from Goldberg. The plasmid profile of 4Q2 is the same as 4Q1, another B. thuringiensis var. israelensis strain in the BGSC. It is one of the 12 original cultures grown on agar slants, each prepared from a surface agar plate, inoculated from a single colony of ONR60A by Goldberg. The 4Q1 culture slant was sent by Goldberg to Dean to be deposited at the BGSC.

. Strain 4Q2 wild type maintains the full complement of plasmids. It has been characterized by Dulmage as being unstable for crystal production (personal communication). In our hands, it easily looses its ability to produce crystals after storage on nutrient agar slants at 4°C for extended periods of time (9-12 months), but is very stable when stored in liquid nitrogen. Strains WHO 1884 and WHO 1897 were also from the original 12 vials. It is interesting that in the passage of these strains among investigators, plasmids have been lost spontaneously. Plasmids pTXl4-3 and pTXl4-4 are missing in WHO 1884, and pTXl4-3 and pTXl4-9 are missing in WHO 1897. These two strains had been sent by Goldberg to deBarjac for serotyping, and then to Briggs, who donated them for these studies. Other B. thuringiensis var. israelensis strains have also lost plasmids spontaneously. Plasmids have been lost from strains 57-499 CD and B.t. 82161 II used by Abbott Laboratories (missing pTX14-3 and pTXl4-4), whereas another strain (795/n 1) used at Abbott Laboratories by Couch has all the plasmids representative of ONR60A. Abbott strain 795/n 2 is missing pTXl4-7. Couch obtained these cultures from Singer, a recipient of one of the original vials of ONR60A from Goldberg. Finally, the international standard, IPS-78, has all the plasmids present, whereas, IPS-80, IPS-82, and the , U.S. standard, U,S. S-1982, have each lost the 7.5 kb pTXi4-3 plasmid. This plasmid was lost at the highest frequency when plasmids were cured from 4Q2 wild type. 48

Strain 4Q2 •wild type has nine extraçhromosomal elements. To better understand these plasmids, it is essential to characterize and associate phenotypes with them. So that these plasmids could be studied individually, plasmid curing experiments were designed to isolate mutant strains cured of various plasmids. Several methods to cure plasmids from B, thuringiensis var. israelensis 4Q2 included the use of heat, which had been used to cure plasmids from Agrobacterium tumerfacians and Staphylococcus aureus (Watson et al., 1975, May et al., 1964, Asheshov, 1966), and the use of coumermycin, which was first used to inhibit gyrase and cure plasmids from Escherichia coli (Danilevskaya and Gragerov, 1980). All the strains derived from 4Q2, with the exception of 4Q2-81, contained extrachromosomal elements. Strain 4Q2-81, producing H-14 reacting antigen, is a plasmid-less variant of 4Q2 wild type. The fact that 4Q2-81 reacts with antibody to B. thuringiensis var. israelensis flagella antigen strongly suggests that this protein, which is used to serotype B. thuringiensis var. israelensis, and thus differentiate it from other B. thuringiensis varieties, is not a plasmid encoded protein.

In general, the frequency of plasmids lost during the curing of strain 4Q2 wild type is similar to those lost spontaneously in other strains used by industry. The plasmids in the smaller size category are lost at a higher frequency than the larger plasmids. Plasmid pTXl4-3 is lost at the highest frequency, followed by pTXl4-4 and pTX14-l. The frequency of plasmid loss was determined from the number of heat or coumermycin cured 4Q2 colonies that had to be screened before a mutant was found with a specific plasmid missing. A possible explaination for the specificity of curing the small plasmids faster than the large plasmids may have something to do with the factors involved with their replication. In E. coli, the large, stringent plasmids are strongly associated with the chromosome and share the same factors used by the cell for chromosomal replication. The small, relaxed plasmids often use a different DNA polymerase and are not associated with the chromosome. In B. thuringiensis var. israelensis, it could be that the factors needed by the small plasmids for their replication are different from those for the large plasmids, and that they are more expendable. 49

In future research, the 4Q2 plasmid cured mutants will be helpful in associating phenotypes to the plasmids. For each plasmid, there is a representative mutant strain cured of that plasmid. The following chapters will describe the characterization of some of these plasmids using many of these mutants. 50

References Asheshov, E.H. Loss of antibiotic resistance in Staphylococcus aureus resulting from growth at high temperature. J. Gen. Microbiol., 1966, 4^, 403-410. •Bernhard, K., Schrempf, H., and Goebel, W. Bacteriocin and antibiotic resistance plasmids in Bacillus cereus and Bacillus subtilis. J. Bacterial., 1978, ISS, 897-903. Danilevskaya, O.N. and Gragerov, A.I. Curing of Escherichia coli K12 plasmids by coumermycin. Molec. Gen. Genet., 1980, 178, 233-235. Eckhardt, T. A rapid method for identification of plasmid deoxyribonucleic acid in bacteria. Plasmid, 1978, 1, 584-588. Gonzalez, J.M., Dulmage, H.T., and Carlton, B.C. Correlation between specific plasmids and delta-endotoxin production in Bacillus thuringiensis. Plasmid, 1981, 5, 351-365. lizuka. T., Faust, R.M., and Travers, R.S. Isolation and partial characterization of extrachromosomal DNA from serotypes of Bacillus thuringiensis pathogenic to lepidopteran and dipteran larvae by agarose gel electrophoresis. J. Sericult. Sci. Japan, 1981, 50, 1-44. Kronstad, J.W., Schnepf, H.E., and Whiteley, H.R. Diversity of locations for Bacillus thuringiensis crystal protein genes. J. Bacterial., 1983, 154, 419-428. Li, E., and Yousten, A. Metalloprotease from Bacillus thuringiensis. Appl. Microbiol., 1975, SO, 354-361. Lovett, P.S. and Bramucci, M.G. Plasmid DNA in Bacilli. In iSchlessinger, D. (Ed.), Microbiology, 1976, American Society for Microbiology, Washington D.C., pp. 388-393, 1976. Macrina, F.L., Kopecko, D.J., Jones, K.R., Ayers, D.J., McCowen, S.M. A multiple plasmid containing Escherichia coli strain: Convenient source of size reference plasmid molecules. Plasmid, 1980, 1, 417-420. May, J.W., Houghton, R.H., Perret, C.J. The effect of growth at elevated temperatures on some inheritable properties of Staphylococcus aureus. J. Gen. Microbiol., 1964, 57, 157-169. Meyers, JA ., Sanchey, D., Elwell, L.P., and Falkow, S. Simple agarose gel electrophoresis method for the identification and characterization of plasmid deoxyribose nucleic acid . J. Bacterial., 1976, 127, 1529-1537. 51

Miteva, V.I. Isolation of plasmid DNA from various strains of Bacillus thuringiensis and Bacillus cereus. Comptes, rendus de I’Acad Burgare de Sci., 1978, 31, 913-916. Schnepf, H.E., and Whiteley, H.R. Cloning and expression of the Bacillus thuringiensis crystal protein m Escherichia coli. Proc. Natl. Acad. Sci. USA, 1981, 78, 2893-2897. Sharp, P.A., Sugden, B., and Sambrook, J. Detection of two restriction endonuclease activities in H. parainfluenzas using analytical agarose- ethidium bromide electrophoresis. Biochem., 1973, IS, 3055-3063. Stahly, D.P., Dingman, D.W., Bulla, LA. Jr., and Aronson, A.I. Possible origin and function of the parasporal crystals, in Bacillus thuringiensis. Biochem. Biophys. Res. Commun., 1978a, 84, 581-588. Stahly, D.P., Dingman, D.W., Irgens, R.L., Field, C.C., Feiss, M.G., and Smith, G.L. Multiple extrachromosomal deoxyribonucleic acid molecules in Bacillus thuringiensis. FEMS Microbiol. Letters, 1978b, 8, 139-141. Watson, B., Currier, T.C., Gordon, M.P., Chilton, MJ)., and Nester, E.W. Plasmid required for virulence of Agrobacterium tumefaciens. J. Bacterial., 1975, 123, 255-264. Yousten;: A.A. A method for the isolation of asporogenic mutants of Bacillus thuringiensis. Can. J. Microbiol., 1978, 24, 492-494. CHAPTER THREE Characterization of the Three Smallest Bacillus thuringiensis var. israelensis Plasmids and Construction of Shuttle Vectors

Introduction

Bacillus thuringiensis is one of the few Bacillus species which have naturally occurring plasmid DNA. A study of plasmid profiles of 24 varieties of B. thuringiensis has shown that while all have plasmid DNA, the number and size of the plasmids differ in the many varieties (lizuka et al., 1981). For example, var. sotto and var. thompsoni each have only one plasmid, while var. kurstaki has sixteen. The smallest plasmid, which is only 1.1 kb, is found in the var. thuringiensis BA-068, and the largest plasmid, which has a size of 443 kb, is found in the var. vmhanensis. Few biochemical functions have been assigned to B. thuringiensis plasmids, Gonzalez et al. (1081) reported that there is a correlation between the ability to produce crystal toxin and the presence of a specific plasmid in four varieties: thuringiensis, kurstaki, alesti, and galleriae. In addition, Schnepf and Whiteley (1981) were able to clone plasmid DNA fragments of B. thuringiensis var. kurstaki into E. coli using the cloning vector p6R322, and demonstrated the presence of toxin polypeptides in the cell extracts by means of radio-immunoassay and insect toxicity bioassay. , Their results provided more direct evidence for the plasmid borne nature of the crystal toxin gene. There have since been several reports of cloning crystal toxin genes that were located on B. thuringiensis plasmids (Ward et al., 1984, Adang et al., 1985, McLinden et al., 1985, Shibano et al., 1985, Waalwijk et al., 1985, Ward and Ellar, 1986).

52 63

Even though the function for the vast majority of B. thuringiensis plasmids is unknown, they offer a rich supply of DNA for the development of vectors for gene cloning within this species. This is especially practical since PEG-induced plasmid transformation of protoplasts, developed for Bacillus subtilis, is possible with the varieties kurstaki, galleriae, and israelensis (Martin et al., 1981, Alikhanian et al., 1981, hfiteva et al., 1981, Lopraser et al., 1986). This is very important because DNA can now be introduced into B. thuringiensis after it has been manipulated in vitro. It should also be possible to test whether a plasmid is compatible with plasmids of another variety. Two plasmids that are compatible can coexist autonomously within the same host. Incompatible plasmids compete with each other, resulting in the loss of one of the two plasmids. In the literature, plasmids are often organized into incompatibility groups. Related plasmids in the same group can not coexist in the same bacteria. Some examples of incompatibility groups are the P and N groups of Ènterobacteria (Datta, 1974, Grant et al.,- 1978) and the Inc groups of Staphylococcus (Novick et al., 1976).

One drawback of using S. thuringiensis var. israelensis plasmids as cloning vectors is the lack of selectable genetic or biochemical markers. However, this problem can be overcome if the plasmid of interest is first ligated with a foreign DNA fragment conferring antibiotic resistance. Restriction fragment maps of all plasmids used in the cloning experiments would be essential for such constructions. Through in vitro manipulation of these plasmids, a suitable cloning vector can be constructed, and possibly the crystal toxin gene can be mobilized from one variety to another. This may eventually lead to the construction of a new strain with either higher toxicity, a broader host spectrum, or both.

Shuttle vectors could also be constructed with plasmid DNA from B. thuringiensis var. israelensis. This type of vector is prepared by combining plasmids from two different species resulting in a chimeric plasmid that is able to replicate in both species. The vector is then useful for cloning and investigating gene expression in the two different hosts. Some examples include pHV33 and pHVl4 (Ehrlich, 1978). These are hybrid plasmids formed by joining . 5 4

Escherichia coli plasmid pBR322 with Staphylococcus aureusjB. subtilis plasmid pdl94. If one of the two plasmids in the hybrid has a Col origin of replication, as do the above examples, then the vector offers the investigator the convenience to prepare large, amounts of DNA m E. coh' using chloramphenicol mediated amplification (Clewell, 1972, Clewell and Helinski, 1972) . In the presence of chloramphenicol, chromosomal DNA replication is terminated but not plasmid replication. The plasmid copy number per cell can reach to greater than 3000, representing a 125 fold increase over the normal cell copy number of 24. Furthermore, shuttle vectors constructed with B. thuringiensis var. israelensis and E. coli plasmids would offer the investigator the well characterized genetic system of E. coli. One could also take advantage of the ease of transforming CaClg treated E. coli.

With the above thoughts in mind, three plasmids from B. thuringiensis var. israelensis, pTXl4-l, pTXl4-2, and pTXl4-3, were isolated, purified, and then subjected to restriction endonuclease studies. Highly purified plasmid DNA was cleaved with a dozen type II restriction endonucleases. By analyzing the restriction fragments obtained from single, double, and partial enzyme digestions, individual restriction maps were constructed. The purified pl^mid DNA was also analyzed by DNA-DNA filter hybridization to see if they had homologous DNA sequences, and in the construction of shuttle vectors.

Materials and Methods

Bacterial strains. Partially cured plasmid mutants of B. thuringiensis var. israelensis (see Chapter Two) were used for plasmid isolation. Mutant strain 4Q2-22, which was cured of plasmid pTXl4-2, pTXl4-3, pTX14-4, and plXl4-7, was used as the source of pTXl4-l. Mutant strain 4Q2-73, which was cured of pTX14-l, pTXl4-3, pTX14-4, and pTXl4-7, was used as a source of pTXl4-2. Mutant strain 4Q2-80, ' cured of all plasmids except pTXl4-3 and pTXl4-5, was used as a source of pTXl4-3. 55

Media and G row th Conditions. The minimal media, SCGYIO, was composed . of the following in grams per liter: (NH^)g80^, 2 g/1; KHgPO^, 6 g/1; KgHPO^, 14 g/1; sodium citrate, 1 g/1; glucose, 5 g/1; yeast extract, 1 g/1; casamino acids, 1 g/1; MgS04, 0.1 g/1. The pH was not adjusted.. The glucose was sterilized separately. Yeast extract and casamino acids were purchased from Difco Laboratories, Detroit, MI. For solid media, 15 grams of agar was added per liter. Bacteria grown on solid media were incubated at 30°C. Bacteria grown in liquid cultures were routinely, incubated with gentle shaking at 150 rpm at 30°C. Cultures were grown in 500 ml SCGYIO broth in 2 liter Erlenmyer flasks, or in 2500 ml SCGYIO broth in 6 liter Erlenmyer flasks with shaking at 30°C. These were inoculated with a volume of starter culture equal to 1/33 that of the volume of the media used. For a routine plasmid isolation, 12 liters of cell culture were grown. These cells were harvested at an optical density (600 nm) of 0.6 - 0.7.

Plasm id DNA Isolation and Purification. Plasmid DNA was isolated using the method of Birnboim and Doly (1979) and scaled-up as described by Ish- Horowicz el al. (Maniatis et al., 1982). The method was modified for B. thuringiensis var. israelensis as described below. Cells were grown as described above in Media and Growth Conditions and harvested by centrifugation at 4500 xg for 10 minutes at 4°C. The cell pellet was resuspended in iO ml of solution I (0.05 M glucose; 0.025 M Tris, pH 8.0; 0.01 M EDTA) with 10 mg/ml lysozyme (Sigma Chemical Company, St. Louis, MO, Grade I) and incubated at 37°C for 30 minutes with gentle shaking. The cells were lysed by the addition of 20 ml of cold (4°C) solution II ( 0.2 N NaOH, 1% SDS) with a pH of 12.3 - 12.5 and the clear lysate was kept at 0°C for 10 minutes and then 15 ml of cold (4°C) solution HI (5 M potassium acetate, pH 4.8) was added to neutralize the alkali. The chromosomal DNA and SDS - protein precipitate was removed by centrifugation at 12,000 x g for 30 minutes at 4°C. Isopropyl alcohol (0.6 volumes) was added to the supernatant and the plasmid DNA was precipitated at 0°C for 60 minutes. The plasmid DNA was then obtained by centrifugation at 12,000 xg for 20 minutes and resuspended in 3 ml Ix TE buffer ( 0.01 M Tris, pH 8.0; 0.001 M EDTA). The resuspended DNA was purified on CsCl-EtBr dye buoyant density gradients by adjusting the solution to a density of 1.648 g/cm^ 56 with CsCl and centrifuging for 40 h. at 40,000 rpm and 4°C in a Beckman. Ti75 rotor. Plasmid DNA was extracted two to four times with 2x volumes of CsCl saturated TES ( 0.03 M Tris, pH 8.0; 0.05 M NaCl; 0.005 M EDTA) buffered isopropyl alcohol to remove the EtBr. The CsCl was removed by dialysis against 2 liters of O.lx SSC ( 0.15 M NaCl, 0.015 M sodium citrate for Ix SBC) for 12 h. with three changes of the buffer at 4°C. After adding 3 M sodium acetate to a final concentration of 0.3 M, the plasmid DNA was precipitated with two volumes of 95% ethanol for 8 h. at -20°C. The precipitated DNA was then centrifuged at 18,000 xg, dried in vacuo, and resuspended in O.lx SSC. The plasmid DNA from strain 4Q2-22 was then cleaved with Psf I in order to convert the large plasmids of this strain into linear fragments while leaving pTX14-l uncut and in its CCC form. Typical digestion mixtures had 8-25 )il (32 ;ig) of plasmid DNA, 8 /il of lOx reaction buffer, 8 /il (80 units) of restriction endonuclease, and the remainder being sterile, demineralized, double distilled HgO to adjust the total volume to 80 /il. Reaction mixtures were inoculated at 37°C for 12 h. A 1 /il sample from each reaction was then separated on 0.7% agarose mini-gels to confirm proper digestion. The reaction mixtures were then combined so that approximately 160 /ig DNA was added to each CsCl-EtBr gradient. The gradients were adjusted to a density of 1.643 g/cm® and the linear fragments separated from the uncut pTXl4-l by centrifugation at 45,000 rpm and 20°C for 18 h. using a Beckman VT165 rotor. The same process was repeated with plasmid DNA from strain 4Q2-73 using Bgl II to yield only pTXl4-2. The plasmid DNA from 4Q2-80 was purified by two successive CsCl- EtBr gradients to provide pTX14-3, the only CCC plasmid in this strain.

Restriction Endonuclease Digestion and Gel Electrophoresis. Twelve restriction endonucleases were used in these studies (Table 3.1). In some studies, Sst I was replaced with its isoschizomer. Sac I. The reaction buffers used were as suggested by the manufacturer, Bethesda Research Laboratories Inc., Gaithersburg, Maryland. In a typical single digestion reaction, the mixture (total volume, 20 /il) contained 2 pi of lOx reaction buffer, G.5-2.0 pi of plasmid DNA (0.5 - 0.7 pg) and 3-4 pi (3-4 units) of restriction endonuclease with the remainder of the volume being made up with sterile, demineralized, double distilled HgO. 57

Reaction mixtures were inoculated at 37°C for 12 h. When double digestions were performed, the first digestion was with the restriction endonuclease requiring the lower salt concentration. After 12 h. incubation at 37°C, the reaction was stopped by heating at 65°C for 10 minutes, and then the salts were adjusted for the second restriction endonuclease digestion. For partial digestion, a series of two fold dilutions of the restriction endonuclease were incubated with a constant amount of plasmid DNA at 37°C for 1 h. Reactions were terminated by the addition of 5 pi loading buffer (100 mM EDTA, 0.25% bromophenol blue, 0.25% xylene cyanol, 30% sucrose in HgO) to the 20 pi reaction mixture. Cleaved plasmid DNA from the same digestion reaction was separated on both a 0.7% horizontal agarose gel and a vertical 10% polyacrylamide gel to resolve large and small restriction fragments respectively. In some instances, a 1.2% horizontal agarose gel was also used to resolve small fragments. The agarose gels (agarose, LE, FMC Corp., Maine Colloids Div., Rockland, ME.) were prepared in TEA electrophoresis buffer (0.05 M Tris, pH 8.05; 0.002 M EDTA; 0.02 M sodium acetate). Electrophoresis was for 6 h. at 4.5 volts/cm (constant voltage). The polyacrylamide gels were prepared as described by Maniatis et al. (1975) . Tris borate electrophoresis buffer (0.089 M Tris, pH 8.4; 0.0025 M EDTA; and 0.089 M boric acid) was used for these gels and electrophoresis was at 5.3 V/cm (constant voltage) for 7.5 h. After electrophoresis, polyacrylamide and agarose gels were stained in HgO with 0.5 pg/ml of EtBr for 30 minutes and destained with HgO for 4-6 h. and 12-18 h. for the polyacrylamide and agarose gels, respectively. The stained gels were visualized with a U.V. transilluminator (Model U.V. Fotodyne, New Berlin, WI) and photographed using a MP-4 Land . Camera with a Vivitar #25 red filter.

If a plasmid was not cleaved with the restriction endonuclease, a control was prepared by mixing lambda DNA with the plasmid DNA in the reaction mixture to assure activity of the enzyme. Proper enzyme activity was confirmed by comparing the size of lambda DNA fragments generated, to their sizes expected from the nucleotide sequence.

M easurem ent of R estriction Fragm ents. A Numonics 1224 electronic digitizer (Numonic Corp., Landsdale, PA.) was used to measure the relative - , 58 mobilities of the restriction fragments from photographs of gels. The restriction fragments of a iKnd HI digest of lambda DNA on agarose gels, or a HaeTSL digest of ^X174 RF DNA on polyacrylamide gels, were used as molecular weight standards. The computer program of Schaffer and Sederoff (1981) , which was modified in these studies by the addition of prompts, was utilized to determine the molecular sizes of the plasmid DNA restriction fragments. The FORTRAN program and directions for its use can be found in the Appendix. This program does least-squares analysis and plots the reciprocal of relative mobility vs. fragment molecular weight or size. The molecular weight of the plasmid DNA restriction fragments was also determined by manually plotting the logarithm of the molecular weight vs. the relative mobility of standard restriction fragments using semi-logarithmic paper. Molecular weights (daltons) were converted to sizes (kilobase pairs) assuming 1 megadalton of DNA = 1 .5 kilobase pairs.

DNA-DNA filter hybridization. Plasmid DNA was transferred from the agarose gel to Zeta-probe membrane sheets (Bio-Rad Laboratories, Richmond, CA.) by electrophoretic blotting. Prior to transfer, the agarose gel was stained with EtBr (0.5 /jg/ml) for 10 minutes and destained in IX TAE (10 mM Tris, pH 7.8; 5 mM sodium acetate; 0.5 mM EDTA) for 3-4 h. before photographing the results. The gel was then placed on a shortwave (256 nm) transilluminator for 10 minutes to nick the DNA. After immersing the gel in 0.2 N NaOH and 0.5 M NaCl for 30 minutes to denature the DNA, it was neutralized by soaking twice in 5X TAE buffer and once in IX TAE (10 min. each time). A sheet of Zeta-probe membrane, which had been pre-soaked in IX TAE for 15 minutes, was carefully placed on the bottom side of the agarose gel to avoid air bubbles that would otherwise form around the wells. The gel and membrane were sandwiched between pre-soaked 3 MM Whatman filter paper and pre-soaked fiber pads on the outside. This sandwich was held firmly in a gel holder which was then placed in the Bio-Rad trans-blot cell. The DNA was transferred to the membrane at 40 y ., 30 mA. for 15 h. and then at 70 V., 75 mA. for 2 h. before being removed and dried at room temperature.

Radioactive labeling of DNA probe. B. thuringiensis var. israelensis plasmid DNA was radiolabeled using nick translation. One /ig of purified • 59 plasmid DNA in iX TE buffer was placed in a sterile 1.5 ml Eppendorf tube sitting on ice. To this tube was added the following: 5 /il of a solution composed of 0.2 mM each of dATP, dGTP, and dTTP in 500 mM Tris-HCl (pH 7.8), 50 mM MgClg, 100 mM 2-ME, and 100 ng/ml nuclease free BSA; 5 /il of a solution of DNA polymerase I (0.4 units//il) and DNAse I (40 pg//il) in 50 mM Tris-HCl (pH 7.5), 5 mM magnesium acetate, 1 mM 2-ME, 0.1 mM PMSF, 50% glycerol, and 100 /ig/ml nuclease free BSA; and 12.5 /il (125 uCi) deoxycytidine 5’-[a-®^P] triphosphate, triethylammonium salt in stabilized aqueous solution with the specific activity of 3000 Ci/mmol (Amersham International). The total volume was adjusted to 45 /il with sterile distilled HgO, mixed by centrifuging for a second or two, and incubating at 15°C for 1 h. The nick translation reaction was terminated by the addition of 5 /il of 300 mM Na^EDTA (pH 8.0). Unincorporated ®^P-dCTP was removed by chromatography using a Sephadex G50 column (Pharmacia, Fine Grade, particle size, 20-80 /im). The Spehadex G50 was autoclaved in IX STE (10 mM Tris, pH 8.0; 100 mM NaCl; 1 mM EDTA) to swell the beads. The column was constructed in a 1 ml tuberculin syringe plugged at one end with glass wool. The syringe was packed to 0.9 ml with Sephadex G50 by repeated centrifugation at 1600 xg for 4 minutes and then equilibrated twice with 0.1 ml IX STÉ with centrifugation at the same speed and for the same amount of time. The nick translated DNA probe was adjusted to 0.1 ml with IX STE and applied to the column. It was centrifuged at the same speed and time as before, collecting the effluent in an Eppendorf tube. One /il of the effluent was added to 1 ml HgO in an Eppendorf tube and counted by Cerenkov counting (Peng, 1977). No corrections were made for the plastic Eppendorf tube, the vial, or the efficiency of Cerenkov counting.

Pre-hybridization . and hybridization of membranes. Prior to hybridization, the membrane was prehybridized with a solution of 5X SSC (IX SSC: 0.15 M NaCl, 0.015 M sodium citrate); 50% formamide; 5X Denhardt’s solution (Denhardt, 1966) (IX Denhardt’s solution: 0.02% bovine serum albumin, 0.02% polyvinylpyrridone, and 0.02% ficoll); 0.1% SDS; 1 mg/ml sheared and denatured salmon sperm DNA; and 25 mM sodium phosphate buffer, pH 6.5. Salmon sperm DNA was sheared by forcefully ejecting it through an 18 gauge 60 needle and then denatured by boiling for 10 minutes. The membrane vrss prehybridized in a polypropylene Seal-n-Save (Sears Roebuck Co.) bag using 0.1 ml prehybridization buffer/cm^ membrane. All air bubbles were removed and the bag was sealed prior to incubation at 42°C for 18-24 h.

After this time, the prehybridization buffer was replaced with an equal volume of hybridization buffer. Hybridization buffer is similar to prehybridization buffer but differs by having a final concentration of 200 /ig/ml salmon sperm DNA (sheared and denatured) and 10% dextran sulfate. The prehybridization buffer has no dextran sulfate but 5X the amount of salmon sperm DNA. The DNA probe was denatured by boiling for 10 minutes and added to the hybridization buffer at 1x10® cpm/cm^ membrane. Air bubbles were removed and the bag sealed and incubated at 42°C for 24 h.

Following hybridization, the membrane was washed to remove non-speciOcally bound probe. The membrane was .first washed in 2X SSC, 0.1% SDS with agitation for 90 minutes at room temperature, then in O.IX SSC, 0.1% SDS for 30 minutes at room temperature, and finally in O.lX SSC, 0.1% SDS for 30 minutes at 55°C. The membrane was dried at room temperature before exposing XR-1 Kodak x-ray film with a Dupont Cronex Lightning-plus intensifying screen at-80°C for 24-72 h. 61

R esults

Figure 3.1 shows samples of the three B. thuringiensis var. israelensis plasmids used in this study at several steps of their purification. Few large molecular weight plasmids are seen because the DNA was isolated by the alkaline lysis method of Birnboim and Doly (1979) which works best for small plasmids. Even though it is difficult to see large plasmids, fragments from these plasmids are visible after cleavage with restriction endonucleases and concentration by NaOAc - ethanol precipitation. These linear fragments are efficiently separated from the CCC form of the desired plasmid during centrifugation in CsCl-EtBr. After centrifugation, some of the CCC plasmid DNA is converted to the OC form. The degree of this conversion varies with each preparation. The variation can be due to physical nicking when the plasmid band is removed from the gradient using a needle, nicking when the DNA and EtBr are exposed to light, or nicking during pipetting when the ethidium bromide is extracted. Also seen in Figure 3.1 is some contamination of pTXl4-3 with linear DNA that has a molecular size of approximately 15 kb. This is most likely chromosomal DNA.

Restriction Digestion Ajialysis of pTX14-l. Plasmid pTXl4-;l was the smallest plasmid investigated. When the plasmid was treated with 12 restriction endonucleases, only certain enzymes could cleave the plasmid (see Table 3.1). Fragments of less than 0.45 kb were not detected clearly or were in a non-linear region for measuring relative mobilities on 0.7% or 1.2% agarose gels. These smaller restriction fragments were easily resolved on 10% polyacrylamide gels. A typical result after resolving the pTX14-l DNA fragments is seen in Figure 3.2.

The positioning of the fragments for each of the restriction endonucleases in the map for pTXl4-l was accomplished by comparing the size of the DNA fragments, shown in Table 3.2, from single and double digests. By choosing a unique restriction endonuclease site as a reference point and knowing the size of all the DNA fragments, it was possible to deduce the relative locations of the other sites. The best arrangement of these fragments is shown in Figure 3.3. Figure 3.1. Electrophoretic analysis of purified plasmid DNA used for restriction endonuclease studies. This DNA is compared to the plasmid DNA profile of B. thuringienaia var. iaraelenaia 4Q2-WT. The DNA samples in each lane are: 1) total plasmid DNA from strain 4Q2>22; 2) plasmid DNA from 4Q2-22 after restriction endonuclease digestion with Pat I; 3) CsCl-EtBr purified pTXl4-l after Pat I cleavage of 4Q2-22 plasmid DNA; 4) total plasmid DNA from 4Q2-73; 5) plasmid DNA from 4Q2-73 after restriction endonuclease digestion with Bgl II; 6) CsCl-EtBr purified pTXl4-2 after Bgl II cleavage of 4Q2-73 plasmid DNA; 7) CsCl-EtBr purified pTXl4-3 from 4Q2-80; 8) purified pTXl4-2 adjacent to plasmid DNA from 4Q2-WT in lane 10; 9) purified pTXl4-l adjacent to plasmid DNA from 4Q2-WT; 10) plasmid DNA of 4Q2-WT isolated by the PEG method of Kronstad et al. (1983) . White open circles indicate the migration position of open circular plasmid DNA. The bracket indicates the position of linear chromosomal DNA fragments. The linear plasmid, pTXl4-5, is not seen on this gel. The plasmid DNA was separated on a horizontal 0.7% agarose gel at 4.5 V/cm for 6 h.

62 63

sm vd 3S 8011)1 NI 3ZIS in moo in in in

Figure 3.1 Figure 3.2. These two gels, representative of the many gels used for restriction endonuclease analysis of plasmid DNA, show larger size restriction fragments on 0.7% agarose and smaller restriction fragments on 10% polyacrylamide. The DNA samples in the agarose gel are: A) lambda DNA cleaved with Hinà IE; B) pTXl4-l cleaved with Af&o I; C) pTXl4-l cleaved with Ptm H; D) lambda DNA cleaved with Hinà IH; E) pTXl4-l cleaved with S a il

Identical samples of pTXl4-l from restriction endonuclease reaction mixtures were separated on the agarose gel, and on the polyacrylamide gel. The samples in the polyacrylamide gel are: A) ÿXl74 RF cleaved with Hae III; B) pTXl4-l cleaved with Mbo I; C) pTXl4-l cleaved with Pvu H; D) ^X174 RF cleaved with Hae ni; E) pTXl4-l cleaved with Sal I.

The size of the standard DNA fragments are in basepairs.

64 65

2 3 1 3 0

A B C D E

Figure 3.2 66

Restriction Digestion Analysis of pTXl4-2. . The results of single digestions with twelve restriction endonucleases shows that pTXl4-2 was cleaved by the enzymes as listed in Table 3.1. By comparing the digestion products of the enzymes JBam HI, Jïae III, HindTSl, Hpa II, Pstl, Pvu ÏÏ, and Sst I, and then the pair-wise double digestion products using these enzymes as listed in Table 3.3, the physical map of pTX14-2 was constructed (Figure 3.4).

Restriction Digestion Analysis of pTXl4-3. The results of single digestions with twelve restriction endonucleases has shown that pTXl4-3 was cleaved with the following enzymes: Bgl I, Bgl II, Eco RI, Hae HI, Hind III, fljpa n. Mho I, Pst I, Pvu II, and Sst I (Table 3.1). The enzymes Bam HI and Sal I had no cleavage sites in pTX14-3. The restriction fragments were ordered on a circular map by comparing the digestion products of the enzymes Bgl I, Bgl II, Eco RI, Hind HI, Pst I, Pvu II, and Sst I, with the pair-wise double digestion products using these enzymes, as shown in Table 3.4. The results of the double digest involving Hind HI with other restriction endonucleases allowed the placement of the three largest Hind HI fragments (A, B, and C). The orientation of Hind III fragments D and E were determined by partial digestion analysis. The .sizes of partial Hind HI fragments (indicated by an *) are shown in Table 3.5'.,.Partial fragments of 1.4, 1.58, 4.83, and 7.22 kb can only be accounted for by single combinations (see proposed order in Table 3.5). The proposed order of the five Hind HI fragments generated in a complete digestion of pTXl4-3 is -B-A-D-E-C- with the largest fragment denoted A and the smallest being E. This order is supported by the positioning of three Hind HI fragments satisfied by the union of C+B-hA from the results of pairwise double digestions with HindJH and other restriction endonucleases and the ordering of A+DH-E from the results of partial digestion. Partial fragments for the possible combinations of B-(-C4-E, B-hD+E, or A+C were not among the detected partial fragments. The 1.58 kb partial fragment can best be satisfied by the union of C+E. It is unlikely that the order is C+D as a partial fragment of 2.33 kb was not detected after partial digestion of pTX14-3. The ordering of restriction sites for pTX14-3 into a circular physical map is seen in Figure 3.5. 67

Table 3.1. Restriction endonuclease cleavage sites ^ pTX14-l PTX14-2 pTX14- 5.40 Kb 6.51 Kb 7.53 ]

Bam HI 1 1 0 Bel I 1 0 2 Bgl II 3 0 1 Eg o RI 2 0 2 Hae III 4 4 9 Hind III 1 2 5 Hpa II 3 5 7 Mbo I 8 8 15 Pst I 0 1 1 Pvu II ••• 3 3 1 Sal I 0 0 0 Sst I 0 1 1

^ fragments were generated after incubation for 12 h. at 37°C with restriction endonucleases 68

Table 3.2» Restriction fragments produced by single and double endonuclease digestions of pTXl4-l

Restriction Size of Fragment (Kb) Total Kb Enzyme AB C DE F G H

Bam HI 6.39 6.39 Bel I 6.48 6.48 Bgl II 3.72 1.63 0.18 6.43 Eco RI 2.80 2.61 6.41 Hae III 3.08 2.04 0.72 0.33 6.17 Hlnd III 6.34 6.34 Hpa I 2.09 1.73 1.66 6.37 Mbo I 1.88 1.04 0.83 0.66 0.41 0.30 0.18 0.14 6.40 Pvu II 2.79 2.40 0.29 6.48 Bam HI-Bcl I 3.09 2.16 6.26 Bam HI-Bgl II 3.02 1.66 0.69 0.17 6.43 Bam HI-Eco RI 2.79 2.26 0.46 6.49 Bam HI-Hae III 2.34 1.40 0.72 0.66 0.33 6.46 Bam HI-Hlnd III 3.32 2.16 6.48 Bam HI-Hpa II 2.10 1.63 1.53 0.23 6.39 Bam HI-Pvu II 2.76 2.340.290.26 6.37 Bel I-Bgl II 2.82 1.62 0.83 0.18 6.34 Bel I-Hae III 2.03 1.63 0.87 0.74 0.33 6.49 Bel I-Hlnd III 6.16 0.17 6.33 Bgl II-Hae III 1:68 1.29 0.72 0.71 0.60 0.33 0.18 6.61 Bgl IlrHpa II 1.62 1.29 0.89 0.89 0.71 0.18 6.46 Bgl II-Pvu II 2.76 1.60 0.65 0.29 0.180.08 6.43 Eco RI-Bgl II 2.67 1.66 0.92 0.30 0.18 6.62 Eco RI-Hae III 1.60 1.02 1.02 0.86 0.72 0.33 6.46 Eco RI-Hpa II 1.67 1.49 1.11 0.62 0.44 6.31 Eco RI-Pvu II 1.98 1.98 0.92 0.41 0.29 6.57 Hae III-Pvu II 1.65 1.37 0.72 0.66 0.38 0.33 0.29 6.39 Hlnd III-Bcl I 5.16 0.17 6.33 Hlnd III-Bgl XI 2.73 1.53 1.04 0.18 6.48 Hlnd III-Bco RI 2.79 2.49 0.15 6.43 Hlnd III-Hae III 2.01 1.82 0.77 0.71 0.33 5.43 Hlnd III-Hpa II 2.06 1.73 1.40 0.32 5.49 Hlnd III-Pvu II 2.37 1.88 1.04 0.29 5.57 69

Table 3.3. Restriction fragments produced by single and double endonuclease digestions of pTXl4-2

Restriction Size of Fragment (Kb) Total Enzyme (Kb) A BC D E F G H

Bam HI 6.61 6.51 Hae III 4.98 1.10 0.33 0.10 6.60 Hlnd III 6.46 1.04 6.60 Hpa II 2.16 1.60 1.43 1.06 0.30 6.64 Mbo I 1.29 1.14 1.01 1.01 0.87 0.76 0.48 0.47 6.69 Pst I 6.48 6.48 Pvu II 6.39 0.101 0.039 6.63 Sst I 6.63 6.63 Hae Ill-Bam HI 4.98 1.10 0.24 0.107 0.096 6.62 Hae III-Hlnd III 3.68 1.37 1.10 1.07 0.32 0.096 6.62 Hae Ill-Pst I 4.98 1.10 0.33 0.101 6.61 Hae III-Pvu II 4.98 0.99 0.33 0.102 0.098 0.039 6.64 Hpa II-Bam II 2.16 1.61 1.44 1.06 0.18 0.12 6.64 Hpa II-Hlnd III 1.62 1.41 1.07 1.06 0.74 0.38 0.30 6.66 Hpa Il-Pst I 2.16 1.62 1.41 1.06 0.27 0.03* 6.64 Hpa Il-Sst I 3.16 1.61 1.44 0.69 0.36 0.30 6.66 Hlnd III-Bam HI 3.92 1.66 1.04 6.61 Hlnd Ill-Pst I 3.64 1.61 1.08 6.63 Hlnd III-Pvu II 3.67 1.80 1.04 0.101 0.039 6.64 Pst I-Pvu II 6:24 0.12 0.101 0.039 6.60 Pvu II-Bam HI 6.08 0.26 0.102 0.039 6.47 Sst I-Bam HI 3.99 2.67 6.56 Sst l-Hae III 3.78 1.23 1.10 0.33 0.099 6.44 Sst l-Hlnd III 4.17 1.31 1.05 6.63 Sst I-Hpa II 2.16 1.61 1.41 0.71 0.36 0.30 6.64 Sst l-Pst I 4.23 2.37 6.69 Sst I-Pvu II 4.26 2.19 0.1010.039 6.69

' This small fragment was not detected on a 10% polyacrylamide gel. The fragment consists of approximately 30 base pairs and does not migrate within the effective range of separation for this gel. 70

Table 3.4. Restriction fragments produced by single and double endoduclease digestions of pTXl4-3

Restriction Size of Fragment (Kb) Total Enzyme (Kb) A B C D EFG H

Bel I 4.68 2.93 7.51 Bgl II 7.62 7.62 Eco RI 7.11 0.47 7.58 Hae III 3.12 1.49 1.23 1.11 0.26 0.11 0.062 0.044 0..033 7.46 Hlnd III 2.48 2.37 1.23 1.01 0.42 7.50 Hpa II 2.24 2.03 1.86 1.02 0.123 0.106 0.063 0.044 7.46 Mbo I 1.10 0.99 0.80 0.68 0.66 0.63 0.67 0.33 0..29 0.29 0.096 0.081 0.067 0.06 7.50 Pat I 7.60 7.50 Pvu II 7.46 7.46 Sst I 7.66 7.55 Bel I-Bgl II 4.69 2.620.38 7.49 Bel I-Hlnd III 2.39 1.68 1.22 0.99 0.72 0.42 0.17 7.58 Bel I-Pst I 2.93 2.33 2.24 7.49 Bel I-Pvu II 4.69 1.63 1.37 7,49 Bgl Il-Eeo RI 6.27 0.93 0.46 7.65 Hlnd III-Bgl II 2.48 2.00 1.23 1.01 0.42 0.36 7.49 Hlnd III-Eeo RI 2.39 1.43 1.19 1.01 0.71 0.46 0.42 7.68 Hlnd Ill-Pst I 2.49 2.37 ■ 1.01 0.86 0.63 0.41 7.66 Hlnd III-Pvu II 2.37 1.66 1.23 1.01 0.83 0.42 7.50 Hlnd Ill-Sst I 2.40 1.60 1.19 1.04 0.96 0.42 7.58 Pst I-Bgl II 4.88 2.70 7.56 Pst I-Eco RI 3.66 3.32 0.47 7.44 Pvu II-Bgl II 6.38 1.10 7.48 Pvu II-Eco RI 7.07 0.21 0.18 7.46 Pvu Il-Pst I 3.86 3.67 7.43 Sst I-Bgl XI 6.33 1.26 7.69 Sst I-Hlad III 2.34 1.60 1.39 1.16 0.93 0.43 7.60 Sst I-Pst X 3.83 3.63 7.44 Sst X-Pvu XI 7.37 0.18 7.66 71

Table 3.5. Fragments produced by partial digestion of pTX14-3 with Hind n i Fragment Size (Kb) Possible Combinations Proposed Combinations

7.22* . A-B-C-D A-B-C-D 6.06* A—B—C or A—B —D A-B-C 4.83* A-BA-B 3.90* A—D—E , B—C—E , or B—D“E A-D-E 3.59* B-C or A-C B-C 2.45 A . 2.37 B 1.58* C-É C-E 1.40* D—E D-E 1.26 C 1.07 D 0.42 E

partially digested fragment 72 Bom HI

PTXI4-I 5.40 kb

Hpo g. Hot SC Eeo RI Hind H I Bgl S B e ll

Figure 3.3. Restriction endonuclease map of pTXl4-l. The relative distances of these sites are based on sizes of restriction fragments resolved on agarose or polyacrylamide gels. This map is to the same scale as the restriction endonuclease maps seen in Figures 3.4 and 3.5. 73

PTXI4-2 6.51 kb

Sst I HoaSE

Figure 3.4. Restriction endonuclease map of pTX14-2. The relative distances of these sites are based on sizes of restriction fragments resolved on agarose or polyacrylamide gels. The exact orientation of the Pvu II fragments B and C are not known. They may be reversed. This map is to the same scale as the restriction endonuclease maps seen in Figures 3.3 and 3.5. Pstl 74

753 kb

Ind Eco m

Figure 3.5. Restriction endonuclease map of pTX14-3. The relative distances of these sites are based on sizes of restriction fragments resolved on agarose and polyacrylamide gels. This map is to the same scale as the restriction endonuclease maps seen in Figures 3.3 and 3.4. 75

DNA-DNA niter hybridization. The autoradiogram in Figure 3.6 used pTXl4-3 DNA as probe to determine if it has any homology with other B. thuringiensis var. israelensis plasmids. The results indicate that pTXl4-2 and pTXl4-3 have related DNA sequences since there is hybridization of pTXl4-3 to pTXl4-2. There is also hybridization between. pTXl4-3 and pTXl4-4. There is some weak hybridization between pTXl4-3 and pTXl4-l but none between pTX14-3 and pTXl4-6 - pTXl4-9. These results were obtained after washing the filter under stringent conditions for Zeta-probe membrane.

Construction of shuttle vectors. The B. thuringiensis var. israelensis plasmids pTX14-2, and pTXl4-3 were each inserted at unique restriction endonuclease sites in E. coli plasmid, pUCl9 to create chimeric plasmids. B. thuringiensis var. israelensis plasmid pTXl4-2 was linearized with Bam HI, Pst I, or Sac I, and ligated to linear pUC19 with the corresponding cohesive ends resulting in chimeric plasmids designated pBDC-3, pBDC-4, and pBDC-5, respectively. Circular maps of these constructs are shown in Figures 3.7 and 3.8. Restriction enzyme analysis (Figure 3.9) confirms that each plasmid has the expected size of 9.2 kb. Cleavage of each construct with the unique restriction endonuclease used to create the chimeric plasmid releases pUCl9 and the B. thuringiensis var. israelensis plasmid. The restriction enzyme analysis also determined the orientation of pTXl4-2 respective to pUC19 (Figure 3.9). Cleavage of pBDC-3 with Hinà. HI generated DNA fragments of 6600, 1550, and 1050 bp, thus confirming the orientation of the B. thuringiensis var. israelensis relative to pUCl9 as shown in the pBDC-3 map (Figure 3.7). The same strategy was used for the other chimeric plasmids. Cleavage of pBDC-4 with Hind IH generated DNA fragments of 6300, 1800, and 1050 bp and cleavage of pBDC-5 with Hind HI generated fragments of 4150, 4000, and 1050 bp., again confirming the orientation of the B. thuringiensis var. israelensis plasmid relative to pUCl9 (Figures 3.7 and 3.8).

Two constructs, pBDC-6 and pBDC-7 were the fusion of pTXl4-3 with pUC19 at their unique Pst l and Sac I sites, respectively. The size of both these chimeric plasmids was 10.2 kb. Figure 3.10 shows the physical maps for these two plasmids and Figure 3.11 shows the electrophoretic analysis of DNA fragments» 76 after cleaving the plasmids with specific restriction enzymes. For pBDC-6 and pBDC-7, cleavage with Pst I and Sac I, respectively, releases pTXl4-3 and pUCl9 (Figure 3.11). A double enzyme digestion of pBDC-6 with Eco RI and Bgl n generated DNA fragments of 5350, 3500, 930, and 420 bp., thus, orientating pTXl4-3 relative to pUC19 (Figure 3.10). Information from a double enzyme dig,estion of pBDC-7 with Eco RI and Pst I, generating fragments of 3600, 3500, 3000, and 100 bp, was used to orientate pTXl4-3 relative to pUC19 in this clone (Figure 3.10). The 100 bp DNA fragment is not seen in this figure. Figure 3.6. DNA-DNA filter hybridization of pTXl4-3 probe to 4Q2-WT plasmids and DNA controls. DNA was electrophoretically resolved through a 0.7% agarose gel (top) before being electroblotted onto Zeta-probe membrane for Southern hybridization (bottom). The DNA in each lane is: A) all pl^mids from B. thuringiensie var. tsraelenata; B) pTXl4-l (lOOng); C) pTX14-2 (lOOng); D) blank; - E) 1:100 dilution of the sample in lane A; F) pUBllO (lOOng); G) pTXl4-3 (500 ng); H) pUC8 (100 ng).

77 78 G H

Y-S,? /,

Figure 3.6 Bm NI sai EcoRI hll 79 hw2 \ 9910

,1100 8000

2000 7000 PBDC-3 9200 bp

(000

5000 I 0000 HindS

Hindi

Ml

lOOO 8000

Hindi

2000

Hindi

900

(ON

(OOO

Figure 3.7. Circular maps of pBDC-3 and pBDC-4. 80

EcoRl Pvu2 \ S a d 9000 ) 9200

1000

Hindî

2000 7000 PBDC-5 9200 bp Pvu2 — HindS P s ll Hindi / BanHI J S a d 30 0 0 6000

BaaHl 5000 4000 PsLI

’vu2 Pvu2

Figure 3.8. Circular map of pBDC-5. Figure 3.9. Electrophoretic analysis of pBDC-3, pBDC-4, and pBDC-5 plasmid DNA. The DNA samples resolved in each lane and the enzymes they were cleaved with are: A) uncut pBDC-3; B) pBDC-3 with Bam HI; C) pBDC-3 w ithiïtndni; D) uncut pBDC-4; E) pBDC-4 with Fat I; F) pBDC-4 with Find ni; G) uncut pBDC-5; ti) pBDC-5 with Sac I; I) pBDC-5 with Find III; J) X DNA with F ind m as molecular size standards (kilobase pairs).

81 82

-23J3

-4.36

Figure 3.9 SKl 83 Ecoll I

Hinfl

HlnO

Hindi kll

Ninfi

Sk i

6IH

k l l Hindi Igl2

. ICON

k l l

10200 bp

Eoell \

Hindi N il

M l k l l

Figure 3.10. Circular maps of pBDOG and pBDC-7. Figure 3.11. Electrophoretic analysis of pBDC-6 and pBDC-7 DNA. The enzyme digestions of the DNA samples in each lane are: A) X, Hinà HI; B) uncut pBDC-6; C) pBDC-6, Pat I; D) pBDC-6, Pco RI; E) pBDC-6, Eco RI, Pgf n double digest; F) uncut pBDC-7; G) pBDC-7, Sac I; H) pBDC-7, Pst I; I) pBDC-7, Eco RI, Pst I double digest; J) X Hind III, Eco Rl double digest.

84 85

23.13-

Figure 3.11 86

D iscussion

The experiments reported here have generated precise restriction cleavage maps for pTXl4-l, pTX14-2, and pTX14-3 from B. thuringiensis var. israelensis. The maps reveal several common restriction endonuclease sites which can be useful in later investigations with these plasmids. For example, pTX14-l contains single restriction cleavage sites for Bam HI, Bel I, and Hind ni. The second smallest plasmid, pTXl4-2, contains single sites for Bam HI, Pst I, and Sst I. Both pTXl4-l and pTX14-2 have a single Bam HI site in common. The 7.5 kb pTXl4-3 contains unique Bgl II, Pst I, Pvu II, and Sst I sites. This plasmid and pTXl4r2 both have unique sites for Pst I and Sst I. Foreign DNA with the appropriate cohesive or blunt ends could be ligated at these single sites if these plasmids were used as cloning vectors. Their small size would benefit their usefulness for several reasons. First, it is much easier to . isolate the smaller plasmids from B. thuringiensis var. israelensis than the larger plasmids. This is because one is less likely to physically damage the small plasmids. Furthermore, small plasmids generally replicate under relaxed control and hence, are present at high copy number in each cell. The stringent replication control of large plasmids is coupled to chromosome replication and their copy number is very low, with only one or two copies per cell. Second, the probability of having unique restriction endonuclease sites is greater in a small plasmid than a larger one. Unique sites simplify the preparation of a cloning vector for the insertion of foreign DNA. Finally, the high copy number of small plasmids, often 20-30 copies per cell, increases the gene dosage of a cloned foreign gene and therefore, may increase the amount of gene product in the cell.

During the construction of the restriction endonuclease cleavage site maps of the three plasmids, the fragment sizes were determined by two methods; plotting the logarithm of the plasmid size against the relative mobilities of standard fragments using semilogarithmic graphs, and plotting the reciprocal of the mobility against fragment size. The sizes determined from these two types of plots were very similar, and the same map is produced using either method. There was only a 1-2% difference in the size of the fragments when comparing 87 these two types of plots. The latter method was preferred, after modifying a computer program (see Appendix), as it generated plasmid fragment sizes much faster.

Comparison of the three restriction maps reveals no apparent homology of large nucleotide sequences and therefore suggests no common origin. If they do have a common origin, it took place so long ago that it is masked by DNA rearrangement and base changes at the majority of the restriction endonuclease sites. DNA-DNA filter hybridization analysis shows that there is homology between pTXl4-2, pTXl4-3, and pTXl4-4. Plasmid pTXl4-3 does not have any homology with the other plasmids of B. thuringiensis var. israelensis. It is not possible to make any conclusions about what DNA sequences pTX14-2, pTXl4-3, and pTXl4-4 share in common. It is not overly surprising to observe DNA homology among the plasmids in B. thuringiensis var. israelensis. With the large number of plasmids in a single strain, there is the possibility that DNA of one plasmid had its origin from another plasmid, possibly due a recombination or transposition event. The fact that there is not a large amount of DNA sequence homology suggests that the plasmids are from different incompatibility groups. Typically, only plasmids within the same incompatibility group share strong DNA homology. In Enterobacteria, the sequence homology is from genes the plasmids share that are involved in replication control, copy number, incompatibility, and stable maintenance (Grant et al., 1978). Since plasmids in B. thuringiensis var. israelensis coexist, it can be concluded that they are not in the same incompatibility group and would not be expected to share these genes. Therefore, DNA sequence homology between the B. thuringiensis var. israelensis plasmids should be relatively weak.

Many attempts to transform J3. thuringiensis with plasmids from B. subtilis or Bacillus cereus have failed. When investigators first reported they were able to transform B. thuringiensis with plasmids from these species, the frequency was very low. The efficiency of plasmid transformation was less than 1 x 10'® transformants//ig DNA (Miteva et al., 1981, Ryabchenko et al., 1980). Recently, the frequency of transformation has improved with higher values of transformation (Lopraser et al., 1986). The construction of shuttle vectors with 88

B. thuringiensis var. israelensis and E. coli may transform at a veîy high frequency. At present though, the putative shuttle vectors have only been propagated in E. coli. A very important experiment will be to transform B. thuringiensis var. israelensis with these vectors. Protoplast transformation will be the best method to introduce these plasmids into B. thuringiensis var. israelensis since positive results have been obtained (Martin et al., 1981, Miteva et al., 1981, Tsenin et al., 1983, Lopraser et al., 1986). The. addition of liposomes, which enhance the frequency of transformation in Streptomyces lividans 100 fold (Rodicio and Chater, 1982), may enhance the protoplast transformation of B. thuringiensis var. israelensis.

The chimeric plasmids have a very good chance of replicating in B. thuringiensis var. israelensis since they have the proper origin of replication and any other cis acting sequences that may be necessary for autonomous replication. Attempts should be made to. transform with all. the constructs, pBDC-3 through pBDC-7. It may be discovered that some of the constructs replicate in B. thuringiensis var. israelensis whereas others do not. This could be due to the splitting of an important plasmid regulatory element when pUCl9 was inserted into the B. thuringiensis var. israelensis plasmick.

After transformation of B. thuringiensis protoplasts with these putative shuttle vectors, it is unlikely that the plasmids will confer ampicillin resistance. It has become clear that when shuttle vectors with B. subtilis and E. coli plasmids are studied, genes expressed in B. subtilis are also expressed in E. coli, but that E. coli genes are not expressed in B. subtilis (Ehrlich, 1978, Kreft et al., 1978). This is due to a difference in the specificity of the RNA polymerases from . these two organisms. The E. coli RNA polymerase is more relaxed in recognizing DNA promoter sequences which the enzyme interacts with prior to initiating transcription. If the ampicillin gene is not expressed in B. thuringiensis var. israelensis, it may be necessary to screen regenerated protoplasts by DNA- DNA filter hybridization using pUCl9 or one of the small B. thuringiensis var. israelensis plasmids as a DNA probe. In the future, shuttle vectors could be prepared between B. thuringiensis var. israelensis plasmids and B. subtilis or S. cereus plasmids that can express their genes in B. thuringiensis 89 var. israelensis. Two candidates would be pBC16 and pCl94 since Lopraser et al. (1986) have reported that these two plasmids can be transformed into B. thuringiensis var. israelensis protoplasts . Plasmid pBCl6 confers tetracycline resistance and pC194 confers chloramphenicol resistance. These antibiotic resistance genes are good choices since B. thuringiensis var. israelensis is relatively sensitive to these two antibiotics (see Chapter Five).

Nonetheless, plasmid clones pBDC-3 - pBDC-7 will be good sources of pTX14-2 and pTX14-3 DNA. The B. thuringiensis var. israelensis plasmid can be easily released from the shuttle vector after cleavage at the unique restriction endonuclease site used in its construction. An advantage will be isolating plasmid DNA from E. coli. Routinely, we have isolated 5 -5 0 fold more plasmid DNA from a one liter culture of E. coli than from B. thuringiensis var. israelensis. 90

References Adang, M.J., Staver, M.J., Rocheleau, TA., Leighton, J., Barker, R.F., and Thompson, D.V. Characterized full-length and truncated plasmid clones of the crystal protein of Bacillus thuringiensis subsp. kurstaki HD-73 and their toxicity to Manduca sexta. Gene, 1985, 36, 289-300. Alikhanian, S.I., Ryabchenko, N.F., Bukanov, N.O., and Sakanyan, V.A. Transformation of Bacillus thuringiensis subsp. gallariae protoplasts by plasmid pBCl6. J. BacterioL, 1981, 146, 7-9. Birnboim, H.C. and Doly, J. A rapid alkaline extraction procedure for screening recombinant plasmid DNA. Nucl. Acids Res., 1979, 7, 1513-1523. Clewell, D.B. Nature of Col plasmid replication in Escherichia coli in the presence of chloramphenicol. J. Bacterial., 1972a, 110, 667-676. Clewell, D.B. and Helinski, D.R. Effects of growth conditions on the formation of the relaxation complex on supercoiled Col Ej deoxyribonucleic acid and protein in Escherichia coli. J. Bacterial., 1972b, 110, 1135-1146. Datta, N. Epidemiology and classification of plasmids. In Schlessinger, D. (Ed.), Microbiology - 1974, American Society for Microbiology, Washington, D.C. pp. 9-15., 1974. Denhardt, D.T. A membrane-filter technique for detecting complementary DNA. Biochem. Biophys. Res. Commun., 1966, 23, 641-646. Ehrlich, S.D. DNA cloning in Bacillus subtilis. Proc. Natl. Acad. Sci. USA, 1978, 75, 1433-1436. Gonzalez, J.M., Dulmage, H.T., and Carlton, B.C. Correlation between specific plasmids and delta-endotoxin production in Bacillus thuringiensis. Plasmid, 1981, 5, 351-365. Grant, R.B., Whiteley, M.H., and Shapley, A.J. Plasmids of incompatibility group P code for the capacity to propagate bacteriophage IKe. J. Bacterial., 1978, 136, 808-811. lizuka. T., Faust, R.M., and Travers, R.S. Isolation and partial characterization of extrachromosomal DNA from serotypes of Bacillus thuringiensis pathogenic to lepidopteran and dipteran larvae by agarose gel electrophoresis. J. Sericult. Sci. Japan, 1981, 50, 1-44. Kreft, J., Bernhard, K., and Goebel, W. Recombinant plasmids capable of replication in B. subtilis and E. coli. Molec. Gen. Genet., 1978, 162, 59-67. 91

Lopraser, S., Pantuwat, S., and Bhumirat, A. Transfer of plasmid pBCl6 and plasmid pC194 into Bacillus thuringiensis subsp. israelensis. J. Invertebr. Pathol., 1986, 48, 325-334. Maniatis, T., Jeffery, A., and Kleid, D.G. Nucleotide sequence of rightward operator of phage X. Proc. Natl. Acad. Sd. USA, 1975, 72, 1184-1188. Ish Horowicz, D. Alkaline Lysis Method. In Maniatis, T., Fritsch, E.F., and Sambrook, J. (Ed.), Molecular Cloning, A Laboratory Manual, Gold Spring Harbor Laboratory, Cold Spring Harbor, New York, pp. 90-91,1982. Martin, P.A.W., Lohr, J.R., and Dean, D.H. Transformation of Bacillus thuringiensis by plasmid deoxyribonucleic acid. J. BacterioL, 1981, 145, 980-983. McLinden, J.H., Sabourin, J.R., Clark, B.D., Gensler, D.R., and Dean, D.H. Cloning and expression of an insecticidal K-73 type crystal protein from Bacillus thuringiensis var. kurstaki into Escherichia coli. Appl. Enmronm. Microbiol., 1985, 50, 623-628. Miteva, V.I., Shivarova, N.I., and Grigorova, R.T. Transformation of Bacillus thuringiensis protoplasts by plasmid DNA. FEMS Microbiol. Letters, 1981, 12, 253-256. Novick, R.P., Clowes, R.C., Cohen, S.N., Curti^, R. Ill, Datta, N., and Falkow, S. Uniform nomenclature for bacterial plasmids: A proposal. BacterioL Rev., 1976, 40, 168rl89. Peng, C:T. Cerenkov counting. In Peng, C.T. (Ed.), Sample preparation in liquid scintilation counting, Amersham Corporation, Arlington Heights, IL., pp. 80-86, 1977. Rodicio, R.M. and Cahter, K.F. Small DNA-free liposomes stimulate transfection of Streptomyces protoplasts. J. BacterioL, 1982, 151, 1078-1085. Ryabchenko, N.F., Alikhanian, S.I., Bukanov, N.O., and Sakanyan, VA. Transformation of protoplasts of Bacillus thuringierisis var. galleria 69-6 by plasmid pBClB. Akad. Nauk. SSSR Doklady, 1980, 253, 729-732. Schaffer, H.E. and Sederoff, R.R. Improved estimation of DNA fragment length from agarose gels. Anal. Biochem., 1981, 115, 113-122. Schnepf, H.E., and Whiteley, H.R. Cloning and expression of the Bacillus thuringiensis crystal protein in Escherichia coli. Proc. Natl. Acad. Sci. U&A, 1981, 78, 2893-2897. 92

Shibano, Y., Yamagata, A., Nakamura, N., lizuka, T., Sugisakl, H., and Takanami, M. Nucleotide sequence coding for the insecticidal fragment of the Bacillus thuringiensis crystal protein. Gene, 1985, S4, 243-251. Tsenin, A.N., Nesterenko, A.V., Rybchin, V.N., Potokon, I.L., and Pisarevskii, Y.S. Experiments with protoplasts of Bacillus thuringiensis. I. Obtaining protoplasts and their reversion to the bacillary form. Genetika, 1983, 19, 517-524. Waalwijk, C., Dullemans, A.M., van Workum, M.E.S., and Visser, B. Molecular cloning and the nucleotide sequence of the M^ 28,000 crystal protein gene of Bacillus thuringiensis subsp. israelensis. Nucl. Acids Res., 1985, 13, 8207-8217. Ward, E.S., Ellar, D.J., and Todd, JA . Cloning and expression in Escherichia coli of the insecticidal delta-endotoxin gene of Bacillus thuringiensis var. israelensis. FEBS Lett., 1984, 175, 377-382. Ward, E.S. and Ellar, D.J. Bacillus thuringiensis var. israelensis 5-endotoxin; nucleotide sequence and characterization of the transcripts in Bacillus thuringiensis and Escherichia coli. J. Mol. Biol., 1986, 191, 1-11. CHAPTER FOUR Association Of The Crystal Toxin With A 108 Kb Plasmid

Introduction

One of the most outstanding phenotypic characteristics of Bacillus thuringiensis var. israelensis is its synthesis of a crystal that is highly toxic to mosquitoes. These crystals are unlike the structurally ordered, bipyramidal crystals found in other varieties of B. thuringiensis. They are irregular in shape and made up of multiple inclusions of different sizes, shapes, and electron densities. They are very well developed by stage VI of sporulation and are 0.6 - 1.2 /im in diameter, three quarters the size of the spore (Charles and deBarjac, 1982).

During the experiments to cure plasmids from B. thuringiensis var. israelensis as described in Chapter Two, it was observed that acrystalliferous strains were generated at a relatively high frequency. It was also observed that these strains never reverted back to producing crystal. In this chapter, experiments were done to see if there was a correlation between the loss of a plasmid, and the inability of the strain to produce crystal. Crude protein preparations from plasmid cured strains were bioassayed for toxicity to the common mosquito, Aedes aegypti, to determine if the toxic component was present. In addition, procedures were modified and developed to isolate crystals and these puriHed crystals were characterized by SDS-PAGE for their protein composition. The final results of these experiments strongly suggest that the different proteins that compose the toxic crystal of B. thuringiensis var. israelensis are encoded by a single 108 Kb plasmid. Prior to describing these

93 94 experiments, it is appropriate at this time to provide some background on isolation and solubilization of crystals, and the bioassay of their toxicity to insects.

Crystal Isolation

Before the discovery of B. thuringiensis var. israelensis, several techniques had been developed for the isolation of crystals from other B. thuringiensis varieties. Some of these methods used rate zonal gradient centrifugation, polyethylene glycol (PEG) or dextran sulfate separation, floatation, or solvents such as carbon tetrachloride and trifluorotrichloroethane to purify crystals. The most popular method is the separation of crystals from spores and cell debris on Renografin-76 rate zonal gradients (Milne et al., 1977). A sporulated B. thuringiensis culture is first'sonicated and then layered on top of a 50-80% Renografin-76 gradient. The spores pellet and the crystals form a tight band at approximately the midpoint of the gradient. Nickerson and Swanson (1981) have substituted NaBr for Renografin-76 and have used large hollow rate zonal rotors instead of tubes in swinging bucket rotors. Another method, floatation, removes greater than 90% of the spores (Sharpe et al., 1979). This method can be combined with Renografin-76 gradient purification resulting in greater than 99.99% of the spores being removed. The purification methods that use. solvents' such as carbon tetrachloride and chloroform separate crystals from spores based on their hydrophobic properties (Murray and Spencer, 1966, Pendleton and Morrison, 1966). Purification using solvents is not used extensively because it must be repeated many times to achieve acceptable purity and because of the possible modifications of the crystal protein by the strong solvents. The purification of crystals using polyethylene glycol or dextran sulfate works because the spores preferentially enter the phase with the most PEG or dextran sulfate (Goodman et al., 1967). This method, however, gives a very poor yield of crystals.

Crystal Solubilization

To study the biochemical properties, of crj^tals, it is important to solubilize them into their protein subunits. However, the crystals of B. thuringiensis that 95 have been extensively studied (those toxic to Lepidopterans) are resistant to solubilization in comparison to most proteins. Only strong alkali (pH 12 or greater), strong denaturing agents, or strong reducing agents at normal pH are capable of solubilizing these crystals. Some of the many published crystal solubilization buffers are carbonate buffer with or without ditbiotbreitol (DTT), pH 10.5; NaOH or KOH with or without DTT; and urea, 2-[N- cyclobexylamino]ethane-sulfonic acid (CHES) solubilization buffer. These solubilization buffers inactivate the toxicity of crystals. The major factor in maintaining toxicity is preserving the internal disulfide bonds of the subunits. There are both inter and intra-molecular disulfîde bonds holding the crystal subunits together. Fast and Milne (1979) have been able to retain the toxicity of solubilized crystal by dissolving with 3-[N-morpholino]propanesulfonic acid (MOPS) buffer, pH 7.8 with 0.05M DTT and IM potassium thiocyanate. There are several reasons why the other solubilization buffers do not retain the toxicity of crystals. For example, strong alkali can solubilize the crystal by hydrolyzing the peptide bonds resulting in non-toxic crystal fragments as described by Glatron et al. (1972) . This hydrolysis, which bypasses the stabilization rendered by disulHde bonds, increases with exposure. Strong reducing agents convert the disulfide bonds into sulfhydryl groups, thus solubilizing the crystal. Urea, SDS, or guanidine-HCl denature the crystal subunits, destroying any ionic interaction and tertiary structure of the polypeptide.

Bioassay

In addition to the biochemical properties of crystals, the insecticidal properties are also very important. Bioassays involve the feeding of crystal to insects and assessing their action on the insect. The type of insect one should use in the bioassay is dependent on the variety of B. thuringiensis being investigated. Dulmage et al. (1971) attempted to standardize the bioassay of the B. thuringiensis varieties that are toxic to Lepidopterans. Basically, their bioassay involves the use of the cabbage looper, Trichoplusia m, an insect that is easy to rear and readily available. A standard agar based diet is prepared to which different concentrations of a crystal standard or the test sample are added 96 before being fed to the larvae. They proposed a standard unit of insect toxicity called the International Unit (lU). It can be calculated by using the equation, lU/mg = LCjq standard / LGgg test sample X lU/mg standard. The LGgg is defined as the minimum concentration of crystal required to kill 50% of the larvae. B. thuringiensis strain E-61 from the Pasteur Institute was provided as the standard with an arbitrary toxicity of 1000 lU/mg. A similar standard has been prepared for B. thuringiensis var. israelensis and is known as IPS78 (Institute Pasteur Standard, 1978). It too, is arbitra,rily given a potency of 1000 lU/mg; New batches, standardized relative to the original IPS78, have since been prepared every two years. The primary insect used for B. thuringiensis var. israelensis bioassays is the mosquito, Aedes aegypti. The World Health Organization has set the standard procedure for the B. thuringiensis var. israelensis bioassay (deBarjac and Large, 1979). Some major factors that must be controlled in this standard procedure are temperature, humidity, age of the mosquito instar, and exposure time of the larvae to the crystal dilution.

Material and Methods

Bacterial strains. All of the B. thuringiensis var. israelensis plasmid mutant strains were derived from 4Q2-WT and are listed in Table 4.2. The plasmids present in these strains (Table 4.3) are described in more detail in Ghapter Two.

Isolation of Crystals for Bioassay. From a 5 ml overnight GYS tube culture grown at 30°G, 1 ml was inoculated into 100 ml of the same media and incubated at 30°G with shaking. This liquor was then centrifuged at 17,000 xg, and 4°G for 20 minutes to pellet spores and crystals. The pellet was resuspended in 1/10 volume of 1 M NaGl with 0.01% Triton X-100 and diluted samples plated on nutrient agar (Difco) to titer the spores. Grude crystal preparations were then diluted with IM NaGl and 0.01% Triton X-100 so that all samples had the same number of colony forming units (GFU)/ml. This material was used in bioassays and the proteins analyzed by SDS-PAGE as described below. 97

Bioassays. Crystals from various strains were assayed for their toxicity to second instar Aedes aegypti larvae. These larvae were hatched from eggs taken from a single mosquito colony. They were kindly provided by Dr. W.A. Foster, Department of Entomology, The Ohio State University. Routinely, 1000-2000 eggs were hatched by placing them in 75 ml HgO. The water (tap) was boiled and then cooled to room temperature three times to remove oxygen and chlorine prior to hatching the eggs. Tap HgO was used because it has trace minerals required by the larvae. The eggs hatched within 3 h. and were reared for 48 h. (second instar) before being used in bioassays. In each bioassay, twenty-four ml of the treated tap water was placed in a 30 ml plastic medicine cup. The bioassay was set up in triplicate. For each strain, four dilutions of the crude protein sample were assayed: 10®, 10"^, 10'^, and 10'®. Dilutions were prepared in sterile

16 X 150 ml tubes with the same source of water as the cups. To each cup was added: 0.5 ml of 20 mg/ml mosquito food (yeast extract, lactalbumin, and pulverized rat chow in equal parts, v/v), 20 second instar larvae, counted as they were pulled into a pasteur pipette, and finally, 0.5 ml of each dilution of crude protein containing crystals and spores. The final concentrations of CFU/ml were

1.6 X 10^, 1.6 X 10®, 1.6 X 10®, and 1.6 x 10^ CFU/ml in the 10® - 10 ® dilutions respectively. Controls were prepared with water, food and larvae but minus crude protein. All cüps were incubated at 26°C and 65% humidity for up to 72 h.

Isolation an d Furifilcation of Crystals. Crystals were routinely isolated from a bacterial strain by first inoculating a 20 ml GYS broth ( yeast extract, 0.2%; (NH^)2SO^, 0.2%; KgHPO^, 0.05%; glucose, 0.1%; MgSO^, 0.02%; CaClg'HgO, 0.008%; and MnSO^'HgO, 0.005%). After incubating Overnight, 3 ml was added to 1.5 liters of the same media and the culture grown for 72 h. at 30°C with shaking. To assure no contamination, the culture was checked by phase contrast microscopy for no bacteria that had a different morphology than B. thuringiensis. Then 30 ml of 25% sterile gelatin (Sigma, swine skin, cat. no. G-2500) was added and the flask was shaken vigorously at 200 rpm to produce foam. The foam which contained the hydrophobic spores was removed with a pipette connected to a vacuum. The addition of gelatin and the shaking to 98 produce foam was repeated 3 times. The remaining liquor was centrifuged at 12,000 xg and 4°C for 20 minutes, using a Sorval GSA rotor. Pellets were resuspended in l/lO volume of the original culture using 1 M NaCl and 0.01% Triton X-100. Each sample was then sonicated for twenty, 30 second busts using a Bonifier Cell Disrupter (Heat Systems Ultrasonic Inc. Model W140) at a setting of 7. The samples were then frozen at -80°C prior to lyophilization. The crystal- spore pellet was resuspended to 100 mg/ml with HgO. Sonifica,tion was repeated, and then the crystals were separated from spores and cellular debris using 50-80% Renografin-76 gradients (Milne et al., 1977), Gradients were prepared in Beckman SW 27.1 nitrocellulose or ultra-clear tubes and 75 mg (0.75 ml) of crystal-spore material was layered on top of each gradient. The components were separated by centrifugation at 12,000 xg and 4°C for 2 h. The crystal fraction was then removed with a syringe, diluted 1:2 with water, and centrifuged to pellet the crystals at 17,000 xg, and 4°C for 20 minutes. The pellet was resuspended in 1 M NaCl with 0.01% triton X-100, sonicated, and then purified 3 more times through 50-80% Renografin-76 gradients.

Solubilization of Crystal Protein and SDS-Polyacrylamide Gel Electrophoresis. Seven different buffers were used to solubilize crystals. These were carbonate buffer (0.05 M sodium carbonate, pH ' 1.0.5) (Gonzalez et al., 1982), carbonate buffer with DTT (0.05 M sodium carbonate, pH 9.5, and 0.01 M DTT) (Huber et aU, 1981), phosphate buffer with DTT and SDS (0.01 M sodium phosphate, pH 7.1, 0.01 M DTT, and 1% SDS), KOH with DTT (0.1 M KOH, pH 12.3, and 0.05 M DTT) (Krywienczyk and Fast, 1980), and urea, CHES, SDS, and dithioerythritol (DTE) buffer (8 M urea, 0.005 M CHES, 1% SDS, and 0.07 M DTE) (Aronson and Pandey, 1978). The two other buffers used were phosphate buffered saline adjusted to pH 11 and pH 12. The conditions for solubilizing crystals with each buffer are listed below in Table 4.1.

Proteins were separated on 12.5% polyacrylamide gels using the method of Laemmli et al. (1970) . The electrode buffer was a Tris-glycine buffer (0.025 M Tris, pH 8.3, 0.192 M glycine, and 0.1% SDS). The pH of the resolving gel was 8.8 and the stacking gel was 6.8. Routinely, crystal to be resolved by PAGE was first solubilized by boiling the protein sample for 3 minutes in phosphate buffer 99

(pH 7.1) with DTT and SDS (composition described above). Typically, 10 /d of a 10 mg/ml crystal stock was solubilized in 40 /d of solubilized buffer in an Eppendorf tube. After solubilization, the sample was centrifuged in a microfuge for 5 minutés at room temperature to pellet any non-solubilized crystals. Twenty- five /il of this supernatant (50 /ig protein) was combined with 50 /A of loading buffer (0.0625 M Tris, pH 6.8, 2% SDS, 10% glycerol, 5% 2-mercaptoethanol, and 1% bromophenol blue) (Laemmli, 1970) and separated on a single lane of the gel.

Electrophoresis was for 6 h. at 0.5 mA/cm. Gels were stained in 1% coomassie brilliant blue, 45% methanol, and 10% glacial acetic acid for 9-12 h. and destained for 12 h., changing the destaining solution (20% methanol and 7.5% glacial acetic acid) every 4 h. Stained gels were photographed using a Polaroid MP-4 Land camera with 667 film exposed through a Tiffen high trans #12 yellow filter at f = l l for 1/125 sec. Protein molecular weight standards were purchased from Bio Rad Inc., Richmond, CA. or Sigma Chemical Co., St. Louis, MO. 100

Table 4.1. Conditions for solubilization of B. thuringiensis var. israelensis crystal.

S o lu b iliz a tio n ^ Amount of'* Tem perature Time buffer crystal

carbonate 100 ng 37°C 120 mln.

carbonate w ith DTT 100 ng 37°C 30 mln. phosphate with DTT 100 ng 100°C 3 mln. and SDS

KOH w ith DTT 100 ng 37°C 30 mln.

U rea, CHES, SDS, 100 ng 37°C 90 mln. and DTE phosphate b u ffere d 100 ng 37°C 30 mln. saline, pH 11 phosphate buffered bufd 100 ftg 37°C 30 mln. saline, pH 12

^ 40ffl buffer was used in each solubilization. ^ The crystal stock concentration was 10 mg/ml. 101

Results

The results in this chapter address the important question ^king if there is a correlation between a plasmid and crystal production. Plasmid cured mutants (Chapter Two) were bioassayed for their toxicity to mosquitoes. After making a positive correlation between the 108 kb plasmid and crystal production, crystals were isolated and characterized from the mutants and compared to crystal from wild type B. thuringiensis var. israelensis.

Bioassay of plasm id cured strains. The results of bioassays on crude crystal preparations are seen in Table 4.2. All samples were titered for colony forming units (CFU)/ml prior to testing their toxicity. The majority of the CFUs were due to the germination of spores. The samples were diluted to a final concentration of 1 x 10® CFU/ml before being used in the bioassays. Two of the plasmid cured strains, 4Q2-40 and 4Q2-44 were asporogenous. The CFU’s/ml present in these samples were due to the presence of vegetative cells, which were at low titers after 72 h. growth, especially for 4Q2-44. The bioassay results were recorded after 1, 12, and 24 h., with the results being the same at all three readings. In general, if a crude protein preparation was going to be toxic to the mosquito larvae, death occurred within the first 15-60 minutes of the assay. The 10® and 10"^ dilutions (1.67 x 10^ and 1.67 x 10® CFU/ml) killed the larvae within 15 minutes whereas the 10"® dilution (1.67 x 10^ CFU/ml) took 1 h. Mortality of larvae was easy to detect because, the dead larvae floated on the surface of the bioassay cup and showed no movement, even in the presence of a bright light from above or the movement of an object over the cups. Larvae that were alive instinctually dive to the bottom of the assay cup as they try to swim away from a bright light source or an object moving over the water.

Isolation and Characterization of Crystal Proteins. Photographs of . RenograTin-76 gradients (50-80%) are seen in Figure 4.1. Examination of samples from 52 fractions (0.5 ml each) of the gradient (tube A) using phase contrast microscopy showed that the spores pelleted and that crystals formed several bands, present in fractions 22-33. The majority of the crystals banded as single 102 particles or in clumps of two - four particles in fractions 29-31. The percentage of Renografin-76 in these three pooled fractions was 66.8% and the refractive index was 1.4051. Closer to the bottom of the tube A as seen in Figure 4.1, crystals were found in association with spores, probably held together by a net-like structure described by Dr. J. DesRosier (personal communications). Increased Bonification decreased this association (data not shown). Using several different solubilization buffers, solubilized crystals (50 ng) were resolved on a 12.5% SDS- PAGE gel, as seen in Figure 4.2. After solubilizing the samples, they were centrifuged briefly to remove non-solubilized material. A pellet was seen only in samples of lanes B, D, F, and G, suggesting that these buffers do not solubilize as efficiently as those buffers used for lanes A, C, and E. The sizes of the proteins were not calculated, however, the relative position of the protein bands can be compared to the crystal proteins which where resolved by PAGE as shown in Figure 4.3. In this figure, the crystal proteins were solubilized using phosphate buffer with SDS and DTT. By comparing the relative migration of the B. thuringiensis var. israelensis crystal to standard proteins of known molecular weight, the molecular weight of the crystal proteins were calculated to be 135,000, 65,000, 48,000, 47,000 and 25,000 daltons from the top of the gel to the bottoDd (standard curve not shown).

When B. thuringiensis var. israelensis are observed by phase contrast microscopy, in addition to seeing spores and crystals, inclusion bodies are seen. These inclusions form bands on Renografin-76 gradients at a position similar to the crystals. Figure 4.4 shows successive Renografin-76 gradient purification of these inclusions. Tube A has a large spore pellet. What appears to be a pellet in tubes B, C, and D is a reflection from a flood light used during photography. The particles of the inclusion band, when observed by phase contrast microscopy had a size (0.25 nm) about 1/3 that of the crystals. When concentrated, they had a pink color, possibly contributing to the creamy, very light pink color of a B. thuringiensis var. israelensis bacterial colony. They showed no toxicity to second instar larvae of Aedes aegypti when substituted for crystal in a bioassay. When material from this inclusion band was solubilized and resolved on SDS- PAGE adjacent to solubilized crystal protein, it can be shown to be a contaminant of purified crystal. 103

Table 4.2. Results of bioassays on crude crystal preparations.

s tr a in spore^ crystal^ % m o rta lity

4Q2-WT + + 100 4Q2-22 + - 0 4Q2-32 + - 0 4Q2—40 - - 0 4Q2-44 -- 0 4Q2-50 + - 0 4Q2-52 + - 0 4Q2-56 + - 0 4Q2-58 + - 0 4Q2-63 + + 100 4Q2-64 ■ + + 100 4Q2-66 + + 100 4Q2-69 + + 100 4Q2-72 + + 100 4Q2-78 + + 100 4Q2-79 + + 100 4Q2-80 + - 0 4Q2-81 + - 0

^ Spores and crystals were verified by phase contrast microscopy. 104

T able 4.3. ^ Plasmids present in B. thuringiensis y&r. israelensis strain 4Q2 mutants

1 2 3 4 5 6 7 8 9

4Q2-WT + + + + + + + + + 4Q2-22 + - - + - - + + 4Q2-32 - ' + - + - + - + + 4Q2-40 + - - - + - - + + 4Q2-44 + + - - + + - + + 4Q2-S0 + + - - + + - + + 4Q2-52 + + - - + + - + + 4Q2-56 + + - - + + - + + 4Q2-58 — ■— + - + + - + - 4Q2-63 - - + - - + + + + 4Q2-64 -- + + + + + + + 4Q2-66 - - + - + - + -- 4Q2-69 + + + 4Q2-72 — • - --- + - 4Q2-78 4Q2-79 - - + - -- + - - 4Q2-80 + 4Q2-81

The plasmid profiles of these mutants can be seen in Chapter Two. Figure 4.1. Purification of crystals from strain 4Q2-WT with continuous 50-80% Renografin-76 ^ gradients. Gradient A-D are successive purifications of the crystals. The crystals from six "A" tubes were combined before successive purifications on single tube gradients B-D. The white spot at the bottom of tubes B, C, and D are not spores but only a reflection of a flood light used for illumination during photography. There were no spores visible at the bottom of these gradients. The abbreviations to the left of tube A are: veg = vegetative cells; cry = crystals; cry -H spo = crystals and spores; spo = spores.

^ trademark of E Jl. Squibb and Sons.

105 J06

..■•\W -«>'»;vi f t

Figure 4.1 Figure 4.2. Polyacrylamide gel electrophoresis analysis of B. thuringieneia var. iaraelensis crystal proteins solubilized with different buffers. The buffers used to solubilize the crystal in each lane are: A) urea, CHES, pH 9.6, SDS, and DTE; B) KOH, pH 12.3, DTT; C) PBS, pH 12.0 with NaOH; D) PBS, pH 11.0 with NaOH; E) phosphate buffer, pH 7.1, SDS, and DTT; F) NaHCOg, pH 10.5; G) NaHCOg, pH 9.5, DTT. The proteins were separated on a 1.5 x 140 x 170 min gel at 20 mA until the dye front reached the bottom of the gel (7 h.)

107 108

A B

VMtlsÆ

laaiae

Figure 4.2 Figure 4.3. Polyacrylamide gel electrophoresis. analysis of purified crystal solubilized with phosphate buffer with DTT and SDS. The protein samples separated in each lane are: A) low molecular weight protein standards; B) purified B. thuringienaia var. iaraelenaia crystal; C) high molecular weight protein standards; D) crude 4Q2-WT protein.

109 » g o> P M UIM o O o«n o ? 1 1 1

Pkirified Cryifal 1 w

Crude Profein Ill

Solubilization of the inclusions with phosphate buffer, SDS and DTT and separation by SDS-PAGE (Figure 4.5) identified two proteins. They have molecular weights of 34,000 and 37,000 daltons (standard curve not shown). Solubilization appeared to be 100% with this buffer since there was no pellet after solubilization and centrifugation. Furthermore, no particulate matter could be seen by microscopy.

Finally, crystals were purified from 4Q2-72 and their protein composition compared to 4Q2-WT and 4Q2-81. Strain 4Q2-72 has only one plasmid, pTXl4-7, which is associated with crystal production. Loss of this plasmid from this strain led to the development of the mutant, 4Q2-81, an acrystalliferous, non-toxic strain. The solubilization of crystals isolated from 4Q2-72 and 4Q2- WT and resolved by SDS-PAGE are shown in Figure 4.6. The proteins that were present from purified crystals were also easily identified in solubilized crude protein preparations from these strains. There are no additional bands found in the purified crystals that are not found in the crude preparations. Strain 4Q2-72 purified crystal has the same protein banding pattern as seen for solubilized crystal from 4Q2-WT, the wild type strain. Figure 4.4. Purification of an inclusion with Renografin-76 gradients. Gradients A-D are successive purifications of an inclusion. The white spot at the bottom of tubes B, C, and D are a reflection of a flood light used for illumination during photography. Indicated to the left of tube A is the banding position of vegetative cells and debris (veg), inclusion particles (incl), and spores (spo). •

112 Figure 4.4 eo Figure 4.5. Polyacrylamide gel electrophoresis analysis of crystal proteins and inclusion proteins. The samples in each lane are: A) /3-lactoglobulin; B+C) trypsinogen; D) bovine albumin; E) phosphorylase b; F) /3-galactosidase from rabbit muscle; G) inclusion; H) inclusion; I) purified crystal. Protein standards were purchased from Sigma Chemical Co., St. Louis, MO.

114 116000 97400-

66000 —

24000 -

18400-

Figure 4.5 Cn Figure 4.6. Analysis of crude protein and purified crystals from strains 4Q2^ WT, 4Q2-72, and 4Q2-81. The name of the strains from which purified and crude protein was isolated is listed above each lane. On the left are the sizes in daltons adjacent to the protein molecular weight standards separated in the first two lanes.

116 ^ a

ÏÎÎP»*-»’

. GQ i h< 402-72 hi 4Q2-WT

402-81

402-72

02-WT

J-‘ 118

Discussion

The data suggests that the genetic information necessary for the production of crystal in B. thuringiensis var. isradensis is associated with a 108 kb plasmid. The absence of this plasmid is linked with the loss of crystal production as observed by phase contrast microscopy and by the lack of toxicity to mosquitoes during bioassay experiments. The association of the 108 kb plasmid with crystal production was first reported in 1982 at the annual meeting of the American Society for Microbiology (Clark et al., 1982). Similar observations were later reported by several other investigators (Ward and Ellar, 1983, Kamdar and Jayaraman, 1983, Gonzalez and Carlton, 1984).

None of the other plasmids appear to be necessary for crystal production as strains missing all the plasmids with the exception of the 108 kb plasmid still produce normal, toxic crystals. However, the bioassays are limited in their ability to detect slight differences in toxicity. As shown in Table 4.2, if crystals were present, there was 100% mortality and if they were absent, there was zero mortality. The bioassay does not detect altered toxicities that may be contributed by other plasmid encoded proteins. If the other plasmids do contribute to or subtract from the overall toxicity of the crystal, it is slight and is overwhelmed by the crystal products associated with the 108 kb plasmid.

In plasmid cured strains missing the 108 kb plasmid, other plasmid products may have low levels of toxicity but may not be detected because the product by itself may be soluble or too small to be ingested by mosquito larvae. Schnell et at. (1984) reported that solubilized crystal proteins are only toxic to larvae when they are bound to a latex bead so that they may be ingested. The latex beads are similar in size to bacteria, a common food of mosquito larvae. Future experiments may be to test the cell lysates of plasmid cured strains missing the 108 kb plasmid but having other plasmids to detect soluble proteins (bound to latex beads). In a wild type B. thuringiensis var. israelensis strain, these proteins may be associated with the crystal matrix. It would be necessary to standardize the crude protein preparations (adjusting the protein concentrations 119 so that all the samples are the same) used in the hloassays so that differences in toxicity are not simply due to differences in the overall protein concentration.

In the assays discussed in this chapter, each crude crystal preparation was standardized for colony forming units (CFU)/ml. An assumption was made that the spore to crystal ratio for each strain was 1:1. This was based on microscopic observations of the different plasmid cured B. thuringiensis var. israelensis strains. Exceptions were 4Q2-40 and 4Q2-44 which have spo', cry' phenotypes and no toxicity to Aedes aegypti. More protein was used in these bioassays for the following reason. It was the vegetative cells of these strains which were responsible for forming colonies on plates (not spores) and since the cultures were old, few vegetative cells were still alive. Therefore, to produce the same number of CFU/ml as the samples with spores, more of the culture pellet had to be used.

Summarizing the results in Table 4.2 and 4.3, there is a strong correlation between the presence of the 108 kb plasmid and the production of a crystal that is toxic to mosquitoes. From these results, however, the exact involvement of the 108 kb plasmid is unknown. One possibility is that the plasmid encodes a irons acting regulatory factor that is responsible for initiating crystal genes located on the chromosome. A more likely possibility is that the plasmid contains the genes for the structural proteins of the crystal. Theoretically, the plasmid has more than enough DNA to encode the major proteins which make up the crystal (proteins to be discussed below). The association of the crystal phenotype with, the 108 kb plasmid and not smaller B. thuringiensis var. israelensis plasmids, is similar to the association of the phenotype with large plasmids in other B. thuringiensis varieties. In B. thuringiensis vars. kurstaki, alesti, and sotto, the gene has been located within large molecular weight plasmids (Schnepf and Whiteley, 1981, Shibano et al., 1985, McLinden et al., 1985, Adang et al., 1985). More recently, several investigators have cloned crystal genes from B. thuringiensis var. israelensis (Ward et al., 1984, Sekar and Carlton, 1985, Waalwijk et al., 1985, McLean and Whiteley, 1987). The DNA for these genes is from the 108 kb plasmid. 120

We have also investigated methods for the isolation of B. thuringiensia var. israelensis crystal to characterize their biochemical properties. Purification of crystals using spore floatation and Renografin-76 gradients provided crystals of acceptable purity for these studies. Even though it was relatively easy to obtain large quantities of crystal, yields were decreased by the aggregation of crystals with spores. This can be seen in tube A of Figure 4.1. This clumping may be due to the net like structure that surrounds the crystal and spore within the cell (J. DesRosier, personal communications). Additional sonification does not significantly reduce the amount of clumping, even when the detergent, triton-X 100, was present to reduce hydrophobic interactions between spores and crystals. Stronger detergents such as SDS were not used for fear they would denature the crystal and reduce the toxicity.

Purified crystals can be solubilized by several different buffers with some being more efficient than others. It is interesting that there is a large variation in efficiency (Figure 4.2). In all solubilization experiments, the same amount of crystal (100/

Establishing that the crystals were most efficiently solubilized with phosphate buffer (pH 7.1) with SDS and DTT (Figure 4.2), the protein subunits of the crystal were further characterized. B. thuringiensis crystals that are toxic to Lepidopterans (i.e. var. kurstaki) produce two proteins on SDS-PAGE after solubilization. They are 135,000 and 68,000 daltons in molecular weight, the later being the toxin processed from the 135,000 dalton protoxin (Tyrell et al., 1981). B. thuringiensis var. israelensis also has proteins of 135,000 and 68,000 daltons. These two proteins most likely are different from the two similar sized proteins in the strains toxic to Lepidopterans since B. thuringiensis var. israelensis is not toxic to this order of insects. Furthermore, antibodies raised against the kurstaki crystal do not react with the B. thuringiensis var. israelensis crystal and vis-versa (data not shown). The most predominant protein that composes the B. thuringiensis var. israelenéis crystal is the 25,000 dalton protein. The two proteins of 34,000 and 37,000 daltons are inclusion proteins. This was determined serendipitously when the inclusions were mistakenly isolated as crystals from a cry strain that was inoculated instead of a crj/"*" strain. This inclusion, which forms a band at a similar position on 50-80% Renografin-76 gradients, is smaller in size than crystal. The overall composition of the inclusion is not known. It has been reported that B. thuringiensis var. kurstai^ and aizawai are good sources of poly-)8-hydrôxybutarate and that this is found in the cells as granules (Nickerson et al., 1981, Wakisaka et al., 1982). The inclusions in B. thuringiensis var. israelensis may be poly-jff-hydroxybutarate granules which have two associated proteins.

In conclusion, experiments and their results have been described in this chapter which show a strong association of the crystal phenotype with the 108 kb plasmid. Methods that have been developed for isolating crystal from other B. thuringiensis varieties have been applied to B. thuringiensis var. israelensis and highly purified crystal has been ii^lated. The protein composition of these crystals have been investigated and five proteins of 135 kd, 65 kd, 48 kd, 47 kd, and 25 kd are proininent with some other minor protein bands present. In 122 addition, two proteins of 37 kd and 34 kd which contaminate crystal are associated with an inclusion body. 123

References Adang, M.J., Staver, M.J., Rocheleau, TA.., Leighton, J., Barker, R.F., and Thompson, D.V. Characterized full-length and truncated plasmid clones of the crystal protein of Bacillus thuringiensis subsp. kurstaki HD-73 and their toxicity to Manduca sexta. Gene, 1985, S6, 289-300. Armstrong, J.L., Rohrmann, G.F., and Beaudreau, G.S. . Delta endotoxin of Bacillus thuringiensis subsp. israelensis. J. BacterioL, 1985, 161, 39-46. Aronson, A.I. and Pandey, N.K. Comparative structural and functional aspects of spore coats. In Chambliss, G. and Vary, J.C. (Ed.), Spores VII, American Society of Aerobiology, Washington D.C., pp. 54-61, 1978. Charles, J.-F. and deBarjac, H. Sporulation et cristallogenese de Bacillus thuringiensis var. israelensis en microscopie électronique. Ann. Microbiol. (Inst. Pasteur), 1982, ISSA, 425-442. Chilcott, C.N., Kalmakoff, J., and Pillai, J.S. Characterization of proteolytic activity associated with Bacillus thuringiensis var. israelensis crystals. FEMS Microbiol. Letters, 1983, 18, 37-41. Clark, B.D., Chu, C.-Y., Cohen, S.B., and Dean, D.H. Plasmid associated toxicity in Bacillus thuringiensis var. israelensis. American Society for Aerobiology, Atlanta, GA, March, 1982. deBarjac, H. and Large, I. Proposal for the adoption of a standardized bioassay method for the evaluation of insecticidal formulations derived from serotype H.14 of Bacillus thuringiensis. WHO, 1979, 79.744, 1-15. Dulmage, H.T., Boening, O.P., Rehnborg, C.S., Hansen, G.D. A proposed standardized bioassay for formulations of Bacillus thuringiensis based on the international unit. J. Invertebr. Pathol., 1971, 18, 240-245. Fast, P.G. and Alne, R. Bacillus thuringiensis parasporal toxin: dissolution of crystals with retention of toxicity. J. Invertebr. Pathol., 1979, S4, 319. Glatron, M.-F., Lecadet, M.-M., and Dedonder, R. Structure of the parasporal inclusion of Bacillus thuringiensis Berliner: characterization of a repetitive subunit. Eur. J. Biochem, 1972, SO, 330-338. Gonzalez, J., Brown, B., Carlton, B. Transfer of Bacillus thuringiensis plasmids coding for 6-endotoxin among strains of B. thuringiensis and B. cereus. Proc. Natl. Acad. Sci. USA, 1982, 79, 6951-6955. 124

Gonzalez, Jr., J.M., and Carlton, B.C. A large transmissible plasmid is required for crystal toxin production in Bacillus thuringiensis variety israelensis. Plasmid, 1984, 11, 28-38. Goodman, N.S., Gottfried, R.J., and Rogoff, M.H. Diphasic system for separation of spores and crystals of Bacillus thuringiensis. J. BacterioL, 1967, 94, 485. Huber, H.E., Luthy, P., Ebersold, H.R., and Cordier, J.L. The subunits of parasporal crystal of Bacillus thuringiensis: size, linkage, and toxicity. Arch. Microbiol., 1981, 129, 14-18. Kamdar, H., and Jayaraman, K. Spontaneous loss of a high molecular weight plasmid and the biocide of Bacillus thuringiensis var. israelensis. Biochem. Biophys. Res. Comm., 1983, 110, 477-482. Krywienczyk, J. and Fast, P.G. Serological relationships of the crystals of Bacillus thuringiensis var. israelensis. J. Invertebr. Pathol., 1980, 86, 139-140. Laemmli, U.K. Cleavage of structural proteins during the assembly of the head of the bacteriophage T4. Nature, 1970, 227, 680-685. - McLean, K.M. and Whiteley, H.R. Expression in Escherichia coli of a cloned crystal protein gene of Bacillus thuringiensis subsp. israelensis. J. BacterioL.,'VàS7, 169, 1017-1023. McLinden, J.H., Sabourin, J.R., Clark, B.D., Gensler, D.R., and Dean, D.H. Cloning and expression of an insecticidal K-73 type crystal protein from Bacillus thuringiensis var. kurstaki into Escherichia coli. Appl. Environm. Microbiol., 1985, 50, 623-628. Milne, R., Murphy, D., and Fast, P.G. Bacillus thuringiensis delta-endotoxin: An improved technique for the separation of crystals from spores. J. Invertebr. Pathol., 1977, 29, 230-231. Murray, E.D. and Spencer, E.Y. A simplified purification technique for parasporal inclusions from certain varieties of Bacillus thuringiensis. J. Invertebr. Pathol., 1966, 8, 418-420. Nickerson, K.W., Zarnick, W.J., and Kramer, V.C. Poly-)3-hydroxybutyrate parasporal bodies in Bacillus thuringiensis. FEMS Microbiol. Letters, 1981, 12, 327-331. Nickerson, K.W., and Swanson, J.D. Removal of contaminating proteases from .125

Badllits thuringiensis parasporal crystals by density gradient centrifugation in NaBr. European. J. Appl. Microbiol. BiotechnoL, 1981, 15,213-215. Pendleton, I.R. and Morrison, R.B. Separation of spores and crystals of Bacillus thuringiensis. Nature, 1966, 212, 728-729. Schnell, D.J., Pfannenstiel, MA., and Nickerson, K.W. Bioassay of solubilized Bacillus thuringiensis var. israelensis crystals by attachment to latex beads. Science, 1984, 223, 1191-1193. Schnepf, H.E., and Whiteley, H.R. Cloning and expression of the Bacillus thuringiensis crystal protein in Escherichia coli. Proc. Natl. Acad. Sd. USA, 1981, 78, 2893-2897. Sekar, V. and Carlton B.C. Molecular cloning of the delta-endotoxin gene of Bacillus thuringiensis var. israelensis. Gene, 1985, 55, 151-158. Sharpe, E.S., Herman, A.I., and Toolan, S.C. Separation of spores and parasporal crystals of Bacillus thuringiensis by flotation. J. Invertebr. Pathol., 1979, 34, 315-316. Shibano, Y., Yamagata, A., Nakamura, N., lizuka, T., Sugisaki, H., and Takanami, M. Nucleotide sequence coding for the insecticidal fragment of the Bacillus thuringiensis crystal protein. Gene, 1985, 34, 243-251. Sriram, R., Kamdar, H., and Jayaraman, K. IdentiHcation of the peptides of the crystal of Badllus thuringiensis var. israelensis involved in the mosquito larvicidal activity. Biochem. Biophys. Res. Comm., 1985, 132, 19-27. Tyrell, D.J., Bulla, LA., Jr., Andrews, R.E., Jr., Kramer, K.J., Davidson, L.I., and Nordin, P. Comparative biochemistry of entomocidal parasporal crystals of selected Bacillus thuringiensis strains. J. BacterioL, 1981, 145, 1052-1062. Waalwijk, C., Dullemans, A.M., van Workum, M.E.S., and Visser, B. Molecular cloning and the nucleotide sequence of the M^ 28,000 crystal protein gene of • Bacillus thuringiensis subsp. israelensis. Nucl. Acids Res., 1985, 13, 8207-8217. Wakisaka, Y., Masaki, E., and Nishimoto, Y. Formation of crystalline 5-endotoxin or poly-;3-hydroxybutyric acid granules by asporogenous mutants of Badllus thuringiensis. Appl. Environm.. Microbiol., 1982, 43, 1473-1480. 126

Ward, E.S., Ellar, D.J., and Todd, JjV. Cloning and expression in Escherichia coli of the insecticidal delta-endotoxin gene of Bacillus thuringiensis. israelensis. FEES Letters, 1984, 175, 377-382. Ward, E.S., and Ellar, D.J. Assignment of the 5-endotoxin gene of Bacillus thuringiensis var. israelensis to a specific plasmid by curing analysis. FEES Letters, 1983, 158, 45-49. Yamamoto, T, lizuka. T., and Aronson, J.N. Mosquitocidal protein of Bacillus thuringiensis subsp. israelensis: identification and partial isolation of the protein. Current Microbiol., 1983b, 9, 279-284. CHAPTER FIVE The Antimicrobial Susceptibility of Various Bacillus thuringiensis var. israelensis Strains

Introduction

In nature, many microorganisms produce antibiotics that inhibit the growth of other microorganisms allowing the antibiotic producing species to carve a larger ecological niche. To counteract these antibiotics, some microorganisms have evolved elaborate mechanisms to detoxify them. Resistance to antibiotics is due in most cases to either degradative enzymes, use of alternative biochemical pathways, permeability of the cell wall, or changes in the target site of the agent.

Sometimes plasmids encode for the antibiotic resistance. The first association of plasmids and antibiotic resistance was in 1961 when Watanabe and Fukasawa reported a R factor (related to the Escherichia coli F fertility factor) involved in transferring streptomycin, sulfonamide, chloramphenicol, and tetracycline resistance among Shigella strains in a single event (Watanabe and Fukasawa, 1961). Since this time, plasmids encoding resistance to antibiotics have been discovered in many other microorganisms including members of Enterobacteriaceae, Staphylococcus, Streptococcus, and Bacillus (Datta and Kontomichalou, 1965, Lacey, 1975, Bernhard et al., 1978, Clewell, 1981). In general, plasmids carry opportunistic genes that encode for phenotypes such as transfer mobility, induction of plant tumors, catabolic enzymes, and degradative enzymes. In some cases, the degradative enzymes detoxify antibiotics. j9-lactamase is an example of a common plasmid encoded enzyme found in both Gram positive and Gram negative bacteria. Plasmids can also convey antibiotic resistance by altering membrane permeability as observed for tetracycline 127 128 resistance (Franklin, 1973, Chopra and Howe, 1978) and by altering the antibiotic target site as reported for erythromycin resistance (Lai and Weisblum, 1971, Lai et al., 1973).

In the course of creating plasmid cured mutants in B. thuringiensis var. israelensis and attempting to discover phenotypes associated with these plasmids, it was thought that one of these plasmids may encode an antibiotic resistance for the strain. In this chapter, experiments are described to determine the minimal inhibitory concentration of several antibiotics for various strains of B. thuringiensis var. israelensis. These strains include B. thuringiensis var. israelensis used in commercial products, international standards, isolates from Romania, and plasmid cured derivatives of the original isolate from Israel, ONR60A (4Q2-WT). However, before proceeding to describing these experiments, it is important to provide a brief definition of the minimal inhibitory concentration (MIC) of an antibiotic and discuss how it relates to bacteriostatic and bacteriocidal antibiotics.

. Minimal Inhibitory Concentration

To determine the efficacy of an antibiotic to inhibit the growth of a microorganism, it is necessary to test different concentrations of the . agent against the bacteria. Most tests compare the susceptibility of the bacteria to serial dilutions of the antibiotic. In a typical test, a series of tubes with culture broth and two-fold dilutions of the antibiotic are each inoculated with the same number of bacteria. This is known as the broth dilution method. After incubation, the minimal inhibitory concentration (MIC) is determined by noting the tube with the lowest concentration of antibiotic in the series showing no visible growth. However, this type of testing does not determine if the antibiotic is bacteriostatic or bacteriocidal to the organism. To test which of these two categories an antibiotic falls into, samples are taken from the tubes that show no visible growth and placed in culture broth without antibiotic. Most likely, growth will be seen after further incubation. This is because the activity of the antibiotic at the MIC concentration is often bacteriostatic and does not cause irreversible damage. As the antibiotic concentration increases, a tube will be 129 found that has no cells capable of recovery after remo^dng the inhibitory agent. This is the bacteriocidal concentration. Normally, bacteriocidal antibiotics are those that show no viable cells in tubes that are one or two tubes apart (2 -4 fold difference in concentration) from the bacteriostatic (MIC) concentration. Antibiotics whose bacteriocidal concentrations are many tubes apart from the MIC are termed bacteriostatic.

There are two other commonly used methods to test the antibiotic susceptibility of a microorganism. These are the agar dilution method and the disk diffusion test, that in principal, are modiHcations of the broth dilution method described above. The main advantage of the agar dilution method over the broth dilution method is that many bacterial strains can be tested at the same time on a single agar based media plate. Several dilutions of antibiotic are added to molten agar, thoroughly mixed, and then poured into petri dishes. Microorganisms are transferred to the plate by an inoculum replicating device so that each inoculum is the same volume. The MIC is the concentration that causes complete inhibition of growth after 24 h.

The disk diffusion test uses standard paper disks impregnated with known concentrations of antibiotic. They are placed on top of a confluent "lawn" of bacteria growing on the surface of agar based media. After incubation, the zone of growth inhibition around each disk is measured. The zones can be described as susceptible, intermediate, and resistant. Tables defining the breakpoints for these three categories of susceptibility relative to the diameters of inhibition zones are often published by the manufacturers of the antibiotic disks (Difco Bulletin 0344, 1080). One disadvantage of this method is that the tables defining the breakpoints varies among bacteria. Tables have been developed for many human pathogens but not for B. thuringiensis. 130

Material and Methods

B acterial Strains. The strains used and their derivation are listed in Table 5.1. Their plasmid component has been previously described in Chapter Two (summarized in Tables 2.2 and 2.3). B. thuringimaia var. Icurataki (HD-l) was from the Bacillus Genetics Stock Center, The Ohio State University. Its plasmids have been described elsewhere (McLinden et al., 1985, Kronstad et al., 1983).

Antibiotics. The antibiotics that were used are listed in Table 5.2. All antibiotics were purchased from Sigma Chemical Co., St. Louis, MO. They were initially dissolved with culture broth except for those antibiotics that were poorly soluble in HgO. Chloramphenicol, chlorotetracycline, erythromycin, and novobiocin were first dissolved in a small volume (2-5 ml) of ethanol (95%) before being diluted with culture broth. Gentamycin was first dissolved in 2-5 ml of methanol, and nalidixic acid was dissolved in 2 ml of 0.1 N NaOH before being diluted with culture broth. Controls were prepared without antibiotic to test is the solvents affected the growth of the cells.

A ntibiotic susceptibility testing. The minimal inhibitory concentration for each of the antibiotics listed in Table 5.2 was determined for each B. thuringienaia var. iaraelenaia strain listed in Table 5.1, utilizing the agar dilution method. The MIC values were also determined for B. thuringienaia var. kurataki for comparing to the B. thuringienaia var. iaraelenaia MIC values. Strain 4Q2-WT was also tested using the broth dilution method.

Agar dilution method. The test culture plates were prepared with Difco Penassay base agar (Antibiotic Media #3, Difco Laboratories, Detroit, hfichigan). The medium was sterilized in flasks with a magnetic stirbar, and cooled to 55°C before addition of the antibiotic so there would be limited heat inactivation of the drug. The antibiotic was then thoroughly mixed into the media by stirring for 3 minutes before pouring 25 ml media into each 100 mm diameter petri plate. The plates were dried at room temperature for 24 h. before being inoculated with cells. 131

Bacterial strains to be tested were first grown on Difco Penassay base media for 24 h. at 30°C. A single isolated colony for each strain was then inoculated into 4 ml Penassay broth and incubated at 30°C with shaking (225 rpm) for 4 h. Cells (0.3 ml) were transferred to alternate wells of a 96 well microtiter plate to avoid cross contamination. An inoculum replicating device, consisting of a brass plate with 96 inoculating pins, was used to transfer cells from the microtiter wells to the antibiotic culture media. The inoculum replicating device was sterilized with ethanol and flaming between transfers. The 5 /A droplet of each transferred culture ( 5-7 x 10® cells) was absorbed by the culture media within 5 minutes and the plates were then inverted and incubated at 30°C. Results were recorded after 12 h. and 24 h. with the MIC being determined at 24 h.

The testing as described above was conducted in two phases; preliminary and defined. The preliminary phase tested a broad range of antibiotic concentrations as listed in Table 5.2. In the defined phase, a narrow range of antibiotic concentrations focused on bracketing the MIC. These tests were repeated 3-4 times to rule out any ambiguities.

Broth dilution method. The broth dilution method was utilized for testing the antibiotic susceptibility of strain 4Q2-WT. Dilutions of antibiotics were prepared in a microtiter plate. Eight dilutions of twelve different antibiotics could be tested in each 96 well plate. Initially, 50 /il of Penassay broth was placed in each well. Then 50 /il of antibiotic stock solution at 4X the highest concentration to be tested was placed in a well of the first row. This was done for twelve different antibiotics. An automatic medimixer (Flow Laboratories, Inc.) that could mix and transfer 50 /il was used to make eight, two-fold serial dilutions of the antibiotic. Finally, 150 /il of broth with 1000 cells was added to each well. The cells added to these plates were grown in Penassay broth to an O.D. of 0.65 (660 nm), directly counted using an Hausser Hy-Lite ultra plane counting chamber, and then diluted to 8.6 x 10^ cells/ml (1000 cells/150 /il). The number of viable vs. dead cells was not determined, howevere, the cells were counted during exponetial growth and assumed to be healthy. The minimal inhibitory concentration for each antibiotic was recorded after 18 h. 132

Results

The preliminary antibiotic susceptibility testing of the antibiotics in the concentrations listed in Table 5.2 provided important information on the overall sensitivities of the B. thuringiensis var. iaraelenaia strains. These preliminary results (data not shown) allowed for more specific testing of each antibiotic against B. thuringienaia var. iaraelenaia. Figure 5.1a-d show the final results of antibiotic susceptibility testing of more than 40 strains of B. thuringienaia var. iaraelenaia. The antibiotics in Figure S .la rd are divided into several groups based on their mode of inhibitory action. These groups are: inhibition of protein synthesis, inhibition of nucleic acid synthesis, and inhibition of cell wall synthesis (see legend of Figure 5.1).

The inhibitory action of these different antibiotics was recorded after 12 and 24 h. The breakpoints between resistant, intermediate, or susceptible were easier to discern at 24 h. than 12 h. and for this reason, the MIC was determined at 24 h. Growth was sparse at 12 h. The criteria used to place the bacteria in these three groups of antibiotic sensitivity are as follows: resistant bacteria grew at the same level with the antibiotic as without it, bacterial growth in the intermediate range showed poor growth, and bacteria susceptible to the antibiotic did not grow at all. All B. thuringienaia var. iaraelenaia strains, whether missing plasmids or not, responded to the antibiotics in the same manner. Therefore, each bar graph (Figure 5.1) is representative for all the strains tested with the exception of B. thuringienaia var. kurataki. The breakpoints defining resistance, intermediate, and susceptibility to an antibiotic are shown in the bar graphs of Figure 5.1. Control agar plates that lacked only the antibiotics showed excellent growth of all strmns tested. Table 5.3 lists the comparison of the MIC values of each antibiotic tested on B. thuringienaia var. iaraelenaia and B. thuringienaia var. kurataki.

The MIC’s for 18 antibiotics tested using the broth dilution method with B. thuringienaia var. iaraelenaia 4Q2-WT are listed in Table 5.4. As with the agar diffusion method, the MIC breakpoint was defined as the minimal antibiotic concentration required to inhibit growth. 133

Table 5.1. List of Bdcillua thuringienaia var. iaraelenaia strains used in these studies.

Strain Source

4Q2-WT BGSC* 4Q2-22 Clark, B. 4Q2-32 Clark, B. 4Q2-60 Clark, B. 4Q2-62 Clark, B. 4Q2-56 Clark, B. 4Q2-G8 Clark, B. 4Q2-61 Clark, B. 4Q2-63 Clark, B. 4Q2-64 Clark, B. 4Q2-66 Clark, B. 4Q2-69 Clark, B. 4Q2-72 Clark, B. 4Q2-78 Clark, B. 4Q2-79 Clark, B. 4Q2-80 Clark, B. 4Q2-81 Clark, B. 4Q1 BGSC* Btl U.S. S-1982 Beegle, C. IPS 78 deBarjac, H. WHO 1897 Yousten, A. WHO 1884 Yousten, A. WHO 1887 Yousten, A. WHO 1897 M24.spo- Yousten, A. 795/11 1 (HD648) Couch, T. 795/11 2 (HD649) Couch, T. 67-499 CD Vectobac Technical Devlsetty, B.; Abbott Labs WHO 2013-2 Singer, S. WHO 2013-5 Singer, S. WHO 2013-7 Singer, S. WHO 2013-9 Singer, S. WHO 2013-10 Singer, S. WHO 2013-11 Singer, 8. HD 667-1 Gonzalez, J. HD 667-19 Gonzalez, J. HD 567-26 Gonzalez, J. HD 667-39 Gonzalez, J. HD 667-40 Gonzalez, J. HD 667-63 Gonzalez, J. T4 . Tam, A. HD-1 (4D1) BGSC*

* Bacillus Genetic Stock Center, The Ohio State University, Columbus, OH 43210 134

Table 5.2. Antibiotics and the concentrations^ used in culture media during preliminary antibiotic-susceptibility testing using the agar dilution method.

acrlflavlii 30, 60, 90, 120, 150 amplclllln 25, 50, 75, 100, 125 bacitracin 50, 75, 100, 125, 150 chloramphenicol 0.1, 0.5, 1, 5, 10

chlorotetracycline 0.01, 0.05, 0.10, 1

cycloserine 70, 80, 90, 110, 130, 150

dihydrostreptomycin 0.5 2.5, 5, 10, 15

erythromycin 0.01, 0.05, 0.1, 0.2, 0.3

gentamycin 2, 3, 4, 5, 6

kanamycin 5, 7.5, 10, 15,: 20

kasugamycin 200, 225, 250, 275, 300

nalidixic acid 0.625, 1.25, 2.5, 5, 7.5

neomycin 4, 6, 8, iO, 12

novobiocin 0.3, 0.6, 0.9, i.25, 2.5

penicillin 1.25, 2.5, 5, 10, 15

rifampicin 0.125, 0.25, 0.50, 0.75, 1.0

streptomycin 1, 2, 3, 4, 5

tetracycline 0.625, i.25, 2.5, 5, 10

^ All antibiotic concentrations are listed as /ig/ml 135

Table 5.3. Minimal inhibitory concentrations* of eighteen antibiotics to B. thuringiensis var. israelensis (Bti) and B. thuringiensis var. kurstaki (Btk). Btl Btk acriflavln 60 40 amplclllln 2 0.5 bacitracin 90 170 chloramphenicol 3 3 chlorotetracycline 0.16 0.30 cycloserine 60 60 dlhydrostreptomycln 5 5 erythromycin 0.30 0.30 gentamycin 1.4 1.4 kanamycin 6 6 kasugamycin 225 300 nalidixic acid 0.6 0.6 neomycin 3.0 3.0 novobiocin 1.0 0.5 penicillin 2 0.5 rifampicin 0.10 0.10 streptomycin 4.0 4.0 tetracycline 0.20 0.70

All minimal inhibitory concentrations are expressed in units of ggfval 136

Table 5.4. Minimal inhibitory concentrations^ of antibiotics tested with B. thuringienais var. israelensis 4Q2-WT using the broth dilution method.

acriflavln 50

amplclllln 1

bacitracin 80

chloramphenicol 12.5

chlorotetracycline 1

cycloserine 50

dlhydrostreptomycln 5

erythromycin 0.4

gentamycin 4

kanamycin 10 kasugamycin ND^ nalidixic acid 5

neomycin 12.5

novobiocin 0.625 penicillin ' ' 2 rifampicin 1.0

streptomycin 20

tetracycline 40

* All minimal inhibitory concentrations are expressed in units of pg/ml ^ ND = not determined Figure 5.1a-d. Susceptibility of B. thuringienaia var. iaraelenaia to antibiotics using the agar diffusion method. Each bar graph is for a single antibiotic and is representative for all B. thuringienaia var. iaraelenaia strains tested. Each tick on the abscissa represents an actual antibiotic concentration tested. A key, defining the three ranges of susceptibility, is shown below.

The mode of action of the antibiotics are divided into three groups. The antibiotics in Figure 5.1 a+b inhibit protein synthesis. The antibiotics in figure 5.1c inhibit nucleic acid synthesis, and those in Figure 5.1d inhibit cell wall synthesis.

(666ôj resistant intermediate 1 1 'susceptible

137 chloramphenicol

0.1 erythromycin

0.01 0.1 0.2 0.3 0.4

chlorotetracycline

0.02 0.04 0.06 0.08 0.10 0.12 0.14 0.16 0.18 0.20 0.22 0.24 0.26 tetracycline

0.01 0.1 0.2 0.3 0.4 0.5 0.6 dihydrostreptomycin

H------1------1------1------1------h H------1 9 10 streptomycin

------1------1------1------1------1

Figure 5.1a gentamycin

kanamycin

kasugamycin

150 1 7 5 200 225 250 275 300 neomycin

h------+ ------H------H------1------1------1------1------

Figure 5.1b COO acriflavin

nalidixic acid

0.0 0.2 0.4 0.6 0.8 1.0 1.2 novobiocin ...... 1 1 I I ' I — .30.4 0.5 0.6 0.7 0.8 0.9 10 1.1 1 rifampicin

H------1------1------1------:------1------1------1------1 0.02 0.04 0.06 0.08 0.10 0.12 0.14 0.16 0.18

Figure 5.1c S ampicillin

■H 10

bacitracin -H so 60 70 SO 90 100 110 120 130 140 150 t 160 cyclosprine

penicillin H 10

Figure 5.Id 142

Discussion

The MIC for 18 antibiotics on various strains of B. thuringiensis var. israelensis were determined. The uniformity of the antibiotic resistance among the strains, even mutants missing from one to nine plasmids suggests that the resistance is not encoded by plasmids and therefore, the genes responsible for resistance must be chromosomally located. There is considerable precedence for chromosomally encoded antibiotic resistance in both Gram positive and Gram negative bacteria. Some examples are jS-lactamase and cephalosporinase that form a single operon in B. cereus (Kuwahara et al., 1970), and streptomycin resistance in E. coli, where a mutation in the ribosomal protein, SIC, alters the target site for the antibiotic. Another example is rifampicin resistance in E. coli, which is attributed to a mutation in the chromosomal gene for RNA polymerase, the target for this antibiotic.

Of interest, are the similarities in the MIC for B. thuringiensis var. israelensis strains in comparison to HD-1 {B. thuringiensis var. kurstaki), as seen in Table 5.3. These results appear to be specific for B. thuringiensis. They differ extensively from the MIC values, reported for Staphylococcus aureus and E. coli (Degener. et al., 1981) which were tested in a manner similar to the method described above in Material and Methods. The similarities between the two different B. thuringiensis varieties exemplifies the relatedness of these bacteria. The target sites for many of the antibiotics that inhibit protein synthesis are ribosomal proteins. The two B. thuringiensis species probably share similar ribosomal proteins (target sites) that respond, to the antibiotics with the same degree of resistance/sensitivity. Along with their related physiology and similar biochemical properties, the only major differences between B. thuringiensis var. israelensis and B. thuringiensis var. kurstaki are their plasmids and the type of crystal they encode. The HD-1 crystal is highly toxic to Lepidopterans, whereas the B. thuringiensis var; israelensis crystal is toxic to Diptera. 143

However, there are some differences in the MIC values between B. thuringiensis var. israelensis and B. thuringiensis var. kurstaki. These differences are for ampicillin, penicillin, bacitracin, tetracycline, novobiocin, and kasugamycin. Penicillin inhibits the linking of peptidoglycan for cell wall synthesis. Ampicillin with a similar ^9-lactam structure, also inhibits cell wall synthesis. For these agents, the difference between B. thuringiensis var. kurstaki and B. thuringiensis var. israelensis may be due to differences in permeability for these agents, different penicillin binding proteins (PBP), or different lipoteichoic acids. It is necessary for penicillin or ampicillin to bind to PBPs for the antibiotic to inhibit cell wall synthesis. E. coli mutants defective in producing PBP are resistant to many )9-lactam antibiotics (Spratt, 1977). It could be that B. thuringiensis var. israelensis is less sensitive to ampicillin and penicillin than B. thuringiensis var. kurstaki because it produces fewer PBP.

One observation made when testing the susceptibility of B. thuringiensis var. israelensis to ampicillin and penicillin was that colonies would occasionally appear that were resistant to 100-150 ftg/ml. Furthermore, when an ampicillin or penicillin G disk was placed on a confluent lawn of 4Q2-WT, several resistant colonies grew within the zone of inhibition (data not shown). However, this resistance was not due to )3-lactamase production since the enzyme was not detected by poly-vinyl alcohol-penicillin testing (data not shown). The resistance may be attributed to a mutation that decreases the production of penicillin binding proteins so the antibiotic does not bind to the cells.

An alternative to explain the difference of /3-lactam susceptibility between B. thuringiensis var. israelensis and B. thuringiensis var. kurstain may be the type and concentration of lipoteichoic acids present in the bacterial membranes. High concentrations of lipoteichoic acid in the cell membrane inhibit autolysins such as )3-N-acetylglucosaminidase as studied in B. suhtilis. These autolysins are strongly involved in the bacteriocidal action of /3-lactam antibiotics. It may be that the difference in the MIC values between B. thuringiensis var. israelensis and kurstaki may be attributed to different lipoteichoic acid concentrations. This may also be true for the B. thuringiensis var. israelensis colonies that grow in the presence of 100-150 /ig/ml ampicillin or penicillin. 144

Bacitracin inhibits peptidoglycan synthesis. The difference in MIC values between B. thuringiensis var. israelensis and B. thuringiensis var. kurstaki could be due to differences in cellular permeability. The same could be true to explain the differences for tetracycline, an antibiotic that inhibits protein synthesis. For example, the mechanism for tetracycline resistance in E. coli (Tait and Boyer, 1978) is attributed to the insertion of three proteins into the membrane that exclude tetracycline from entering the cell and reaching its target. In summary, it is interesting that the antibiotics that have different MIC values between B. thuringiensis var. israelensis and B. thuringiensis var. kurstaki either inhibit cell wall synthesis, or in the case of tetracycline, resistance may be due to changes in the composition of the cell membrane.

The antibiotic resistance data presented in this chapter will be helpful in the future use of B. thuringiensis var. israelensis in several disciplines. In the field of genetic engineering, plasmid vectors used in cloning often have selectable antibiotic resistance markers. The antibiotic susceptibility data will help in deciding the vector to use and the proper selective antibiotic concentration. The data can also be used in identifying B. thuringiensis var. israelensis from other B. thuringiensis varieties. This can be in addition to the different flagella antigens and variations in carbohydrates utilization between B. thuringiensis varieties. Finally, the data could be used in cell fusion and plasmid transfer experiments were one may want to select for one bacterial strain. 145

References Anonymous. Bacto-sensitivity discs for use in the antimicrobiotic susceptibility test. , 1980, Publication 0344, , . Bernhard, K., Schrempf, H., and Goebel, W. Bacteriocin and antibiotic resistance plasmids in Bacillus cereus and Bacillus subtilis. J. Bacterial., 1978, 133, 897-903. Chopra, I., and Howe, T.G.B. Bacterial resistance to . the tetracyclines. Microbiol. Rev., 1978, 4^, 707-724. Clewell, D.B. Plasmids, drug resistance, and gene transfer in the genus Streptococcus. Microbiol. Rev., 1981, .^5, 409-436. Datta, N. and Kontomichalou, P. Penicillinase synthesis controlled by infectious R-factors in Enterobacteriaceae. Nature, 1965, 208, 239-241. Degener, J.E., Thonus, I.P., and Michel, M.F. The antimicrobial susceptibility test: a comparison of the results of four methods. J. Applied M 'crobiol., 1981, 50, 505-517. Franklin, T.J. Antibiotic transport in bacteria. Crit. Rev. Microbiol., 1973, 2, 253-272. Kronstad, J.W., Schnepf, H.E., and Whiteley, H.R. Diversity of locations for Bacillus thuringiensis crystal protein genes. J. Bacterial., 1983, 154, 419-428. Kuwahara, R., Adams, E.P., and Abraham, E.P. The composition of ^S-lactamase I and ;3-lactamase H form from Bacillus cereus 569/H. Biochem., 1970, 118, 475-480. Lacey, R.W. Antibiotic resistance plasmids of Staphylococcus aureus and their clinical importance. Bacterial. Rev., 1975, 39, 1-32. Lai, C.J., Dahlberg, J.E., and Weisblum, B. Structure of an inducibly methylatable nucleotide sequence in 23S ribosomal ribonucleic acid from erythromycin resistant Staphylococcus aureus. Biochemistry, 1973, 12, 457-460. Lai, C.J. and Weisblum. Altered méthylation of ribosomal RNA in an erythromycin resistant strain of Staphylococcus aureus. Proc. Natl. Acad. Sci. USA, 1971, 68, 856-860. 146

McLinden, J.H., Sabourin, J.R., Clark, B.D., Gensler, D.R., and Dean, D.H. Cloning and expression of an insecticidal K-73 type crystal protein from Bacillus thuringiensis var. kurstaki into Escherichia coli. Appl. Environm. Microbiol., 1985, 50, 623-628. Spratt, B.C. Properties of the penicillin-binding proteins of Escherichia coli K12. Eur. J. Biochem., 1977, 72, 341-352. Tait, R.C. and Boyer, H.W. On the nature of tetracycline resistance controlled by the plasmid pSClOl. Cell, 1978, IS, 73-81. Watanabe, T. and Fukasawa, T. Episome media,ted transfer of drug resistance in Enterobacteriacaea. I. Transfer of resistance factors by conjugation. J. BacterioL, 1961, 81, 669-678. SUMMARY AND GENERAL CONCLUSIONS

The use of Bacillus thuringiensis successfully in pest control holds great promise as an alternative to petroleum based, chemical pesticides. There are a large number of B. thuringiensis strains that can be employed as microbial insecticides and they exhibit a broad range of properties including different insecticidal activities, flagella antigens, and carbohydrate utilization. The organism is related to B. cereus with the only major differences between the two organisms being the production of 6-endotoxin and the vast array of plasmids in B. thuringiensis.

This dissertation describes the characterization of the plasmids from B. thuringiensis var. israelensis, one of the more than twenty B. thuringiensis varieties. After a brief review of the relevant literature of B. thuringiensis and their plasmids (Chapter One), experiments are described aimed at creating B. thuringiensis var. israelensis strains cured of plasmids (Chapter Two) from the wild type strain, 4Q2-WT. As described in this chapter, various plasmids were cured after treatment of vegetative cells or sporœ with heat or courmermycin. The final result is that for each of the nine B. thuringiensis var. israelensis plasmids, there is a representative strain cured of that plasmid. These strains were then used as a resource for detailed studies of the B. thuringiensis var. israelensis plasmids and to aid in the association of phenotypes to the plasmids. They will also be beneficial for these types of studies in future research.

Furthermore, the plasmids present in S. thuringiensis var. israelensis strains used by industry and academicians were investigated. Some of these strains had all nine plasmids as did our wild type strain (4Q2-WT) while others were missing one or several plasmids. The plasmids that were lost spontaneously were the same as those lost at high frequency during curing experiments, suggesting plasmid instability. It is interesting that the two plasmids that were cured from 4Q2-WT at the highest frequency, pTX14-3 and pTXl4-4, also have sequence homology as shown by Southern hybridization in Chapter Three. They could be 147 148 competing for factors important for their replication. Another possibility is that recombination resulting in deletions is responsible for plasmid loss. Stable recombination between these two plasmids resulting in the formation of a new plasmid of a different size was not observed.

In Chapter Three, plasmid cured strains 4Q2-22, 4Q2-73, and 4Q2-80 were used as sources of pTXl4-l, pTX14-2, and pTXl4-3, respectively. Each of these plasmids was studied free of all other plasmids. Experiments were designed that resulted in the detailed positioning of restriction endonuclease sites on circular maps. Based on the size comparison of the restriction DNA fragments and the number of sites for each enzyme, there were no large regions of conserved sequence between the three plasmids. Southern hybridization using pTXl4-3 as a DNA, nick translated probe showed hybridization with pTXl4-l, pTXl4-2, and pTXl4-4 but no other plasmid in 4Q2-WT. The hybridization was weak relative to pTXl4-3 hybridizing to itself.

Other experiments described in Chapter Three resulted in the construction of several putative shuttle vectors with pTX14-2, pTX14-3, and pUC19. These chimeric pl^mids are stably replicated in E. coli and are a resource for copious amounts of pTX14-2 and pTX14-3 which can easily be excised from the chimeric plasmids by cleavage with a single restriction endonuclease. A very important set of experiments will be the transformation of B. thuringiensis var. israelensis. These plasmids should replicate within the bacteria since they have the proper origin of replication. They were constructed because many previous attempts to transform JB. thuringiensis var. israelensis with plasmids from B. subtilis and B. cereus were unsuccessful.

Construction of several shuttle vectors between pTX14-l and pUCl9 were attempted but after ligation and transformation of these into E. coli strain HBlOl, deletions of approximately 200 bp occurred within pTX14-l. This happened repeatedly during five trials. These deletions may be due to recombination between pTXl4-l and pUC19. Better results may be obtained if the constructs are transformed into Rec A", Rec BC* E. coli strains since less recombination takes place in these strains. 149

A strong correlation was made between crystal production and the presence of the 108 kb plasmid as described in Chapter Four. Strains that did not possess this plasmid did not produce any of the proteins that compose the crystal. After determining the best conditions for solubilizing the crystal, it was discovered that purified crystal from strain 4Q2-72 (has only the 108 kb plasmid) was composed of five proteins as was the crystal from 4Q2-WT. Strain 4Q2-81 which was cured of all nine B. thuringiensis var. israelensis plasmids did not produce any of these proteins. This suggests that the presence of the 108 kb plasmid is necessary for the production of these proteins. Conclusive evidence that the 108 kb plasmid contains the structural genes for crystal proteins is not provide. However, recent reports on the cloning of the B. thuringiensis var. israelensis crystal genes have used the 108 kb plasmid as a source of DNA. After exhaustive attempts, I was unable to express the crystal gene within E. coli after cloning partially cleaved DNA fragments (using several different restriction enzymes) of the 108 kb plasmid into pBR322 and pUCl9 or in pPL608, an expression vector for B. subtilis.

Finally, in Chapter Five, the minimal inhibitory concentration of eighteen antibiotics to B..thuringiensis var. israelensis were determined. These studies were initiated to answer the question if any of the plasmids conferred antibiotic resistance to B. thuringiensis var. israelensis. There are many examples of other plasmids encoding resistance to an antibiotic. B. thuringiensis is a soil microorganism and it would not be surprising that one of its plasmids encoded an antibiotic resistance so that it could better survive among all the. soil microorganisms. However, no correlation between a plasmid and bacterial resistance to an antibiotic could be made. In fact, all B. thuringiensis var. israelensis strains, whether missing plasmids or not, responded to each antibiotic in the same manner. APPENDIX

The FORTRAN program of Schaffer and Sederoff (1981) for calculating the molecular weight or sizes of DNA fragments was modified to instruct the user when and what information must be entered. Following this is a typical log-in session using the program

EDIT L 00010 DIMENSION WT(50), DISTCSO), DWTC50), DDISTC50), 00020 XDPRODCEO), C(50), D(50) 00030 REAL*4 MO.LO.MWT.MDIST.MPROD 00040 1000 CONTINUE 00045 WRITE (6,10) 00046 10 FORMAT(////' ENTER THE NUMBER OF STANDARDS YOU HAVE RIGHT " 00047 X*JUSTIFIED IN SPACES 1 THROUGH 5.V* EXAMPLE: 00048 X'SSS21 IF YOU HAVE 21 STANDARDS. S MEANS SPACE.'////) 00050 READ (5.1) (WTCI), I = l.N) 00060 1 FORMAT (15) 00070 IF (N .LE. 0) GO TO 99 00075 WRITE (6,20) 00076 20 FORMAT(////• ENTER EITHER THE MOLECULAR WEIGHT ", 00077 X*OR THE LENGTH IN BASES FOR EACH STANDARD RIGHT ", 00078 X/' JUSTIFIED IN 10 SPACES AND 8 VALUES TO A LINE.*. 00079 X / ‘ EXAMPLE: SSSSSS2250SSSSSS1960...'////) 00080 READ (5.2) (WT(I), I = l.N) 00090 2 FORMAT (8F10.0) 00100 READ (5.2) (DIST(I). I = l.N) 00110 SWT = 0. 00120 SDIST = 0. 00130 SPROD = 0. 00140 DO 11 I = l.N 00150 SWT = SWT + WT(I) 00160 SDIST = SDIST + DIST(I) 00170 PROD(I) = WT(I)*DIST(I) 00180 11 SPROD = SPROD + PROD(I) 00190 MWT = SWT/N 00200 MDIST = SDIST/N 00210 MPROD = SPROD/N 00220 DO 12 I = l.N

150 151 00230 DWT(I) = WT(I) - MWT 00240 DDIST(I) = DISTCI) - MDIST 00250 12 DPROD(I) = PROD(I) - MPROD 00260 CSSL = 0. 00270 CSSM = 0. 00280 CSCPML = 0. 00290 CSPMLL = 0. 00300 CSPMLM = 0. 00310 DO 13 I = l.N 00320 CSSL = CSSL + DWT(I)**2 00330 CSSM = CSSM + DDIST(I)**2 00340 CSCPML = CSCPML + DWT(I)*DDIST(I) 00350 CSPMLL = CSPMLL + DPROD(I)*DWT(I) 00360 13 CSPMLM = CSPMLM + DPROD(I)*DDIST(I) 00370 DET = CSSL*CSSM - CSCPML**2 00380 MO = (CSSM*CSPMLL - CSCPML*CSPMLM)/DET 00390 LO = (-CSCPML*CSPMLL + CSSL*CSPMLM)/DET 00400 SC — 0 • 00410 SSC = 0. 00420 DO 14 I = l.N 00430 C(I) = (WT(I) - LO)*(DIST(I) - MO) 00440 SC = SC + (CCI) - CCD) 00450 14 SSC = SSC + ceci) - C(l))**2 00460 CBAR = SC/N + C(l) 00470 SDC = SORT((SSC - SC**2 /N)/(N-1)) 00480 WRITE (6.101) 00490 101 FORMAT(' STD LEN DIST PRED LEN DEV 00500 X ' »D C(D') 00610 SD = 0. 00520 SSD = 0. 00530 DO 15 I = l.N 00540 PREDWT = CBAR/(DIST(I) - MO) + LO . 00550 WTDEV = WT(I) - PREDWT 00560 PERC = 100.*WTDEV/WT(I) 00570 SD = SD + WTDEV/WTd) 00580 SSD = SSD + WTDEV**2 00590 15 WRITE (6,3) WT(I). DIST(I). PREDWT. WTDEV. PERC. C(I) 00600 3 FORMAT (F8.1, 5F11.3) 00610 SDWT = SORT((SSD - SD**2/N)/ (N-3)) 00620 WRITE (6.4) MO. LO. CBAR 00630 4 FORMAT (' MO = '. F12.5, * LO = F12.5.* CBAR = F12.5) 00640 WRITE (6.6) SDC. SDWT 00650 6 FORMAT (' SC = '. F12.5. SD = F12.5) 00660 READ (5.1) NU 00670 IF (NU.LE. 0) GO TO 199 00680 READ (5.2) (D(I), I = l.NU) 00690 DO 16 I = l.NU 00700 PREDWT = CBAR/(D(I) - MO) + LO 00710 16 WRITE (6.5) D(I). PREDWT 00720 5 FORMAT (• FOR A DIST OF '.F12.4.' PREDICT A LEN OF F12.4) 00730 199 GO TO 1000 00740 99 CALL EXIT 00750 END END OF DATA 152

Typical log-in session dial 2-1111

The Ohio State University Hosts are TSO, WYLBUR, CMS, DEC20 Host name?

TSO

GO

LOGON (space) TS1186 P(FORTUSER) typing in this will alleviate the need to type Allocate Dataset... PASSWORD? XXXXX UNIVERSITY I.D. XXXXXXXX

TS1186 LOGON IN PROGRESS AT 00:00:00 ON AAAN 00, 1984. there is tso new about SPSS-X. enter: news to view it.

READY ALLOCATE (space) DATASET (space) •(*) (space) FILE (space) (FT06F001) (return) RUN (space) BURTMAP.FORT

G1 COMPILER ENTERED SOURCE ANALYZED PROGRAM NAME = MAIN * NO DIAGNOSTICS GENERATED

(when you right justify, your numbers go farthest right to the 10 space block when entering mol. wt. or lengths)

ENTER THE NUMBER OF STANDARDS YOU HAVE RIGHT JUSTIFIED IN SPACES 1 THROUGH 5 EXAMPLE: SSSS21 IF YOU HAVE 21 STANDARDS. S MEANS SPACE>

SSSS7 (return) (This is the number of standards if you are using lambda cleaved with Hind HI}) 153

(It has been right justified for 5 spaces.) (You must enter at least 3 and no more than 8)

ENTER EITHER THE MOLECULAR WEIGHT OR THE LENGTH IN BASES FOR EACH STANDARD RIGHT JUSTIFIED IN 10 SPACES AND 8 Vj^LUES TO A LINE EXAMPLE: SSSSSS2250SSSSSS1960.... (LEN and mokcular weight are inter changeable) bases bases

SSSSSS15.0SSSSSS6.12SSSSSS4.26SSSSSS2.84SSSSSS1.51SSSSSS1.32SSSSSS0.37 SSSSSS1.36SSSSSS1.92SSSSSS2.30SSSSSS2.93SSSSSS4.36SSSSSS4.72SSSSSS7.44

LEN DIST PRED LEN DEV %D C(I)STD 15.0 1.360 14.738 0.262 1.747 7.378 6.1 1.920 6.355 -0.235 -3.842 7.013 4.3 2.300 4.458 -0.198 -4.647 6.976 2.8 2.930 2.870 -0.030 -1.043 7.195 1.5 4.360 1.404 0.106 6.991 7.622 1.3 4.720 1.208 0.112 8.489 7.684 0.4 7.440 0.421 -0.051 -13.899 6.918

M 0 = 0.88962 L 0 = -0.68617 CBAR= &.25520 SC= 0.31289 SD= 0.21806

SSSS4 (return) (this is the number of unknowns)

(now type in unknowns with Right justified of 10 and only 8 values to a line)

SSSSSS4.38SSSSSS4.68SSSSSS4.88SSSSSS6.20 (return) (You can type in up to 50 unknown values)

FOR A DIST OF 4.3800 PREDICT A LEN OF 1.3925 FOR A DIST OF 4.6400 PREDICT A LEN OF 1.2484 FOR A DIST OF 4.8800 PREDICT A LEN OF 1.1320 FOR A DIST OF 6.2000 PREDICT A LEN OF 0.6801

(LEN = MOL. WT.)

(SPACE) (return) (this is called a évacuons return and 154 it will start the program over again with a new set of standards. One would enter those new standards at this point)

EDIT

END (return)

READY PROFILE (space) PREFIX (space) (TS1186) (return) [IF NECESSARY] READY LOGOFF (return)

(computer printout of time used, price charged, balance remaining etc., date logged off).

The Ohio State University Hosts are TSO, WYLBUR, CMS, DEC20 HOSTNAME? DISCONNECTED LIST OF REFERENCES

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155 156

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