I 3 1293 00590 2535 w H 5 ‘6 0 ‘3 f Wit1Wlil'lfliflflniiliilllfll This is to certify that the dissertation entitled CLONING AND CHARACTERIZATION OF GENETIC MARKERS FROM THE AFLATOXIN-PRODUCING FUNGUS ASPERGILLUS PARASITICUS presented by Jyh-Song Horng has been accepted towards fulfillment of the requirements for Ph . D. degm in Food Science/ Microbiology Major professor Date 12/07/89 MSUi: an Affirmative Action/Equal Opportunity Institution 042771 LIBRARY Michigan State University PLACE IN RETURN BOX to remove this checkout from your record. To AVOID FINES return on or before dete due. DATE DUE DATE DUE DATE DUE MT? 100-: e» - W93, MSU Is An Affirmetive ActiorVEquel Opportu‘nlty Inetttution CLONING AND CHARACTERIZATION OF GENETIC MARKERS FROM THE AFLATOXIN-PRODUCING FUNGUS ASPERGILLUS PARASITIC US By Jyh-Song Horng A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Food Science and Human Nutrition and Department of Microbiology and Public Health 1989 ABSTRACT CLONING AND CHARACTERIZATION OF GENETIC MARKERS FROM THE AF LATOXIN-PRODUCING FUNGUS ASPERGILLUS PARASITICUS By Jyh-Song Homg Three selectable markers have been isolated from Aspergillus parasiticus as the initial step in developing genetic transformation systems for cloning aflatoxin biosynthetic genes. First, the 0120 gene in the tryptophan biosynthetic pathway was isolated by complementation of an Escherichia coli trpC mutant lacking phospho- ribosylanthranilate isomerase (PRAI) activity. The cloned gene complemented an E. call trpC mutant deficient in indoleglycerolphosphate synthase (IGPS) activity and an Aspergillus nidulans mutant strain that was defective in all three enzymatic activities of the rrpC’ gene (glutamine amidotransferase, IGPS and PRAI), thus indicating the presence of a complete and functional trpC+ gene. The location and organization of the A. parasiticus an’ gene on the cloned DNA fragment were determined by deletion mapping and by hybridization to heterologous DNA probes prepared from cloned rrpC‘ genes of A. nidulans and Aspergillus niger. These experiments suggested that the A. parasiticus rrpC‘ gene encoded a trifunctional polypeptide with functional domains organized identically to those of analogous genes from other filamentous fungi. The A. parasiticus rrpC“ gene was expressed constitutively regardless of the nutritional status of the culture medium. Second. the nitrate reductase structural gene (niaD‘) was isolated by screening an A. parasiticus genomic DNA library with DNA probes prepared from cloned niaD‘ genes of A. niger and Aspergillus oryzae. Three types of nitrate-nonutilizing mutants deficient in niaD', genes coding for a molybdenum cofactor (cruc’), and a regulatory element in nitrogen assimilation (nirA’), respectively, were also isolated based on their resistance to chlorate, a toxic analogue to nitrate. A homologous transformation system has been developed with nt'aD+ as the selectable marker. Third, a gene with homologies to the Neurospora crassa B-tubulin gene was isolated from a genomic DNA library of a benomyl-resistant mutant of A. parasiticus. The possibility that this gene represents a mutated version of the B—tubulin gene and confers resistance to benomyl remains to be determined. The cloned apC‘, niaD‘, and B-tubulin genes should be useful as general markers for fundamental genetic analyses, and as selectable markers in developing homologous transformation systems to analyze the aflatoxin biosynthetic pathway of A. parasiticus. Dedicated to my parents, for their love and encouragement; to my wife Kitty, for her understanding and sharing excitement and frustration with me throughout my graduate career, and to our two clones, Cindy and Ivy, obtained by old-fashioned recombinant DNA. iv ACKNOWLEDGMENTS The author would like to express sincere gratitude to his academic advisor, Dr. James I. Pestka, for encouragement and support throughout the five-year odyssey of graduate study. Special thanks are expressed to Dr. John E. Linz for constant technical guidance regarding recombinant DNA methodology and for helpful suggestions in preparation of this dissertation, and to Dr. C. A. Reddy for pointing out the opportunity of receiving a joint Ph.D. degree in Microbiology and Food Science. He also wishes to thank the rest of his guidance committee members, Dr. E. S. Beneke and Dr. C. -N. Shih, for their valuable advice. The author is indebted to Mei-Ling Liu for her assistance in isolation of plasmid pLH23; to Dr. Perng-Kuang Chang for his assistance in cloning the nitrate reductase gene, and to Chris Skory for the benomyl-resistant mutants of A. parasiticus. TABLE OF CONTENTS Page LIST OF TABLES ....................................... x LIST OF FIGURES ...................................... xi RATIONALE .......................................... 14 CHAPTER 1. LITERATURE REVIEW Genetics of Aflatoxin Biosynthesis ........................... 19 Aflatoxins as Secondary Metabolites ......................... 20 Limited taxonomic distribution ........................... 20 Produced in chemical families ........................... 20 Produced after active cellular growth ....................... 23 Synthesized from simple building blocks .................... 23 The Biosynthesis of Aflatoxins ............................ 23 Overview of aflatoxin biosynthesis ........................ 24 Approaches to elucidating the aflatoxin biosynthetic pathway ....... 33 Chemical approach ................................ 33 Biochemical approach .............................. 34 Intraspecific Variation in Toxin Production .................... 38 Interspecific Variation in Toxin Production .................... 39 Mutational Analysis of Aflatoxigenic Fungi .................... 41 A. parasiticus ..................................... 41 A. flavus ........................................ 44 Viral Association with Aflatoxin Production .................... 45 The Parasexual Analysis ................................ 47 Genetic Transformation in Aflatoxigenic Fungi .................. 49 Genetic Transformation in Filamentous Fungi ..................... 52 Transformation Techniques .............................. 55 Preparation of competent cells ........................... 55 Protoplasts ..................................... 56 Alternatives to protoplasts ............................ 59 Uptake of DNA .................................... 61 Calcium and PEG ................................. 61 Liposome-mediated uptake ........................... 62 Heat shock treatment ............................... 62 Electroporation ................................... 63 Other procedures ................................. 64 Selection of Transformants ............................... 64 Selectable markers .................................. 64 Auxotrophic selectable markers ........................ 64 Dominant selectable markers .......................... 72 Visual screening of n'ansformants ...................... 78 Cou'ansformation ................................... 81 Fates of Transforming DNA .............................. 84 Integration into chromosomes ........................... 84 Modes of integration ............................... 85 Analysis of integration events ........................ 86 Stability of integrated sequences ........................ 89 Autonomous replication ............................... 89 Features of autonomously replicating plasmids ............... 9O Origins of replication ............................... 9O ARP and abortive transformants ........................ 94 Recovery of Transforming DNA ........................... 95 Recovery of plasmids. ................................ 96 Recovery of cosmids. ................................ 96 Sib selection. ..................................... 97 Applications of Transformation ............................ 97 Cloning genes by complementation ........................ 98 Heterologous complementation ......................... 98 Homologous complementation ......................... 99 Gene disruption/replacement ........................... 100 Approaches to gene disruption ........................ 100 The RIP effect: a unique gene disruption ................. 106 Cloning genes by insertional inactivations .................. 108 Gene fusions ..................................... 110 Titration of regulatory gene products ...................... 111 Biotechnology .................................... 111 CHAPTER II. CLONING AND CHARACTERIZATION OF THE TRPC’ GENE FROM AN AFLATOXIGENIC STRAIN OF ASPERGILLUS PARASITICUS INTRODUCTION ..................................... 116 MATERIALS AND METHODS ............................ 120 Bacterial and fungal strains ............................. 120 Growth media ...................................... 122 Preservation of cultures ................................ 122 Vectors .......................................... 123 General Procedures .................................. 126 Isolation and manipulation of genomic DNA of A. parasiticus ....... 127 Preparation of RNA from A. parasiticus ..................... 128 Construction of genomic DNA libraries ...................... 129 Lyric complementation of E. coli trpC mutants ................. 130 Transformation ..................................... 131 Laboratory safety .................................... 131 RESULTS .......................................... 132 Isolation of the A. parasiticus rrpC‘ gene from recombinant phage libraries .................................... 132 Localization and organization of the A. parasiticus trpC‘ gene ....... 135 Complementation of an A. nidulans trpC mutant by the cloned A. parasiticus trpC’ gene ............................. 142 Isolation, complementation analysis, and deletion mapping of plasmid pLHZ3 ......................................... 152 Analysis of the A. parasiticus an‘ transcript .................. 156 DISCUSSION ....................................... 159 CHAPTER III. ISOLATION OF NITRATE-NONUTILIZING MUTANT S AND CLONING OF THE NITRATE REDUCT ASE STRUCTURAL GENE (NIAD‘) FROM ASPERGILLUS PARASITIC US INTRODUCTION ..................................... 165 MATERIALS AND METHODS ............................ 169 Fungal and bacterial strains ............................. 169 Media ........................................... 169 Vectors .......................................... 170 General procedures ................................... 173 Isolation and analysis of nitrate-nonutilizing mutants ............. 173 Assay of nitrate reductase .............................. 174 Preparation of protoplasts from A. parasiticus .................. 175 Transformation of fungi ............................... 176 RESULTS .......................................... 177 Chlorate-resistant mutants of A. parasiticus ................... 177 Cloning the A. parasiticus niaD’ gene ...................... 179 DISCUSSION ....................................... 190 CHAPTER IV. ISOLATION OF A DNA FRAGMENT FROM A BENOMYL- RESISTANT MUTANT OF ASPERGILLUS PARASIT 1C US WITH HOMOLOGY TO THE NEUROSPORA CRASSA TUB-2+ GENE INTRODUCTION ..................................... 194 MATERIALS AND METHODS ............................ 196 General procedures ................................... 196 Isolation of benomyl-resistant mutants ....................... 196 Construction of a genomic DNA library ..................... 196 viii Conditions for Southern analysis and plaque hybridization .......... 197 Transformation of A. parasiticus ......................... 200 RESULTS AND DISCUSSION ............................ 201 CONCLUSIONS ....................................... 207 LIST OF REFERENCES .................................. 210 .U'PP’ 10. 11. 12. LIST OF TABLES flag Incorporation of labeled precursors into aflatoxin B1 ............. 36 Aflatoxin intermediates accumulated by blocked mutants of A. parasiticus ...................................... 37 Quick screening media for analysis of aflatoxin-producing ability ..... 42 Filamentous fungi in which transformation has been achieved ....... 53 Selectable markers used across species lines in filamentous fungi. ..... 66 Filamentous fungi as hosts for heterologous gene expression and product secretion ................................... 113 Complementation of E. coli tryptophan auxotmphs by plasmid pLH23 . 121 Characteristics of the A. parasiticus wild-type genomic DNA libraries . 133 Lytic complementation of E. coli trpC mutants with recombinant lambda EMBL3 phages from the A. parasiticus genomic DNA libraries ..... 134 Transformation of an A. nidulans trpC mutant with the A. parasiticus th‘ gene ....................................... 145 Identification of nitrate non-utilizing mutants from A. parasiticus on the basis of growth on different nitrogen sources ............ 178 Comparison of chlorate-resistant mutant types recovered from three strains of A. parasiticus .............................. 180 10. 11. 12. LIST OF FIGURES nge A basic scheme for cloning genes involved in aflatoxin biosynthesis by complementation of blocked mutants. .................... 16 Structures of the major aflatoxins. ........................ 21 The origin of the carbon atoms and the arrangement of intact acetate units in aflatoxin Bl ............................ 25 The hypothetical scheme for the assembly of anthraquinones by a "polyketide synthase" enzyme complex in Aspergillus species ....... 27 The proposed pathway for the biosynthesis of aflatoxin B, ......... 29 The metabolic scheme proposed for the late stages of aflatoxin biosynthesis ................................ 31 Three patterns of integration of transforming DNA into the fungal genome. .................................. 87 Examples of gene disruption/replacement. ................... 102 The biosynthetic pathway of tryptophan from chorismic acid. ...... 118 Restriction maps of arc-containing plasmids pHY 201 and pAB2-l. ....................................... 124 Restriction endonuclease digestion of DN As from 10 randomly selected recombinant lambda EMBL3 clones with DNA inserts containing the A. parasiticus rrpC’ gene. ................... 136 Restriction endonuclease maps of three A. parasiticus DNA inserts (open bars) in EMBL3 which complement the E. coli arC9830 mutation. ............................ 138 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. Southern hybridization of lambda clone MptrpW9 with A. nidulans trpC’ probes. ............................. Restriction endonuclease maps of plasmids containing the A. parasiticus trpC’ gene. ............................ Southern hybridization analysis of five A. nidulans Trp” transformants. .................................... Hybridization of plasmid pJH34 to A. parasiticus chromosomal DNA fragments. .................................. Localization of the Trp functions in pLH23 by deletion mapping. Nprthem analysis of the A. parasiticus rrpC‘ transcript .......... The nitrate utilization pathway in Aspergillus spp ............... Restriction endonuclease maps of flaw-containing plasmids pSTAlO and pSTA14. ............................... Growth of wild-type and nitrate-nonutilizing mutant strains of A. parasiticus ATCC 36537 on media with various nitrogen sources. Identification of the A. parasiticus genomic DNA fragments containing the niaD’ gene by Southern hybridization. ........... Southern analysis of lambda EMBL3 clones containing the A. parasiticus niaD+ gene ............................. Restriction endonuclease map of plasmid pSL82 .............. Restriction endonuclease map of plasmid pBT3. .............. Identification of DNA restriction fragments from the A. parasiticus chromosome with homologies to the N. crassa [i-tubulin gene. ..... 140 157 166 171 181 184 186 188 202 RATIONALE l4 Aspergillus parasiticus is an imperfect fungus which produces aflatoxins. The ubiquitous existence of this mold and the potent toxicity and carcinogenicity of aflatoxins represent a serious threat to food safety and public health. To date, protection against contamination of the food supply with these toxins is limited to preliminary detection, followed by elimination of contaminated items. By investigating expression and regulation of genes involved in aflatoxin biosynthesis it should be possible to design strategies to efficiently control formation of these toxins. It is thus desirable to develop efficient DNA-mediated trans- formation systems for A. parasiticus to provide a mechanism to clone the genes involved in the aflatoxin biosynthetic pathway in this organism. However, the major obstacle in deveIOping such a n'ansformation system for A. parasiticus has been the lack of suitable selectable markers. Auxotrophic mutants of this imperfect fungus are not easily obtainable, presumably due to the presence of multiple nuclei in the conidia (Y uill, 1950) and the coenocytic nature of the fungal cytoplasm. Moreover, preliminary data indicate that A. parasiticus is highly resistant to various antibiotics such as hygromycin B and kanamycin, which are commonly used in transformation of eukaryotic cells. Utilization of cloned genes from other organisms as selectable markers to develop a heterologous transformation system for A. parasiticus is a feasible approach. However, available evidence suggests that heterologous genes will not always be expressed (Woloshuk et al., 1989), or expressed less efficiently in A. parasiticus, resulting in a lower transformation frequency. Together, these factors significantly impede development of homologous or heterologous transformation systems based on auxotrophic or dominant resistance genes as selectable markers. 15 A general approach to deve10ping transformation systems for cloning genes in the aflatoxin biosynthetic pathway of A. parasiticus is depicted in Figure l. The immediate goal of this investigation was to isolate and characterize selectable markers for use in developing transformation systems for A. parasiticus in suitable recipient strains. Three selectable markers from A. parasiticus were isolated and characterized. These three markers include the trifunctional trpC‘. gene in the tryptophan biosynthetic pathway, which has been widely used in transformation of filamentous fungi; the nitrate reductase structural gene (niaD‘) for which the corresponding mutants can be readily induced on the basis of a positive selection protocol (chlorate resistance), and a mutant allele of the B—tubulin gene which confers resistance to the fungicide benomyl, and is generally used as a dominant selectable marker in transformation of filamentous fungi. 16 Figure l. A basic scheme for cloning genes involved in aflatoxin biosyn- thesis by complementation of blocked mutants. A vector containing a suit- able selectable marker (sel‘) from heterologous or homologous sources is used to construct a genomic DNA library from an aflatoxigenic strain of A. parasiticus. Homologous selectable markers to be incorporated into the cloning vector are obtained by complementation of suitable E. coli mutant strains, or by screening an A. parasiticus wild-type (wt) genomic DNA library by hybridization with heterologous genes with sufficient nucleotide sequence similarity to the marker of interest. DNA purified from the library is used to transform protoplasts of appropriate blocked mutants that are defective in the aflatoxin biosynthesis. Transformants are first selected on the basis of selectable markers (re?) and then screened for their ability to produce aflatoxin (afl’). Genetic markers are: amdS’, coding for acetamidase; bml, conferring resistance to the fungicide benomyl; hygB, conferring resistance to the antibiotic hygromycin B; M, conferring resistance to the antibiotic kanamycin; pyrG”, coding for orotidine 5’-phosphate decarbo- xylase; trpC‘, a trifunctional gene in the tryptophan biosynthetic pathway of filamentous fungi; niaD‘, the structural gene for nitrate reductase. 17 HETEROLOGOUS SYSTEM HOMOLOGOUS SYSTEM markers from other vector A- rasiticusiwt) organisms( 33115: lA-pUCfl-DRKOl I ED! ’ M 9 J‘_ITI_' ) ' DNA. >500“ L.._.._T ..... .l Genomic Library 1 r """" 1 contme Heterologous tenant-«3.1! probing Ieteete ) -------- L.._--r-......J . Cloned Genes (95.93.19? 2129') J . t . . AWEY‘OSRQ'cfl / veCtOr DNA Blocked Mutants L (fir-1'1.) ...... T , Genomic Library Protoplasts L ___________ 1 __________ .J l Transformation Selection (as. .s_-_n::) Characterization l . Cloned Genes CHAPTER I LITERATURE REVIEW I. Genetics of Aflatoxin Biosynthesis The aflatoxins are a group of carcinogenic, toxigenic, mutagenic, and teratogenic compounds of considerable health and economic importance. They are produced by certain strains of the closely related imperfect fungi Aspergillus flavus, A. parasiticus, and A. nomius, a strain which has recently evolved from A. flavus (Kurtzman et al., 1987). Aflatoxins are the most potent naturally occurring carcinogens known and are frequently found in agricultural commodities and dietary staples such as corn, peanuts, treenuts and cottonseed (Jelinek et al., 1989). The potential hazard of aflatoxins to human health was initially realized following their association with acute hepatotoxicity in poultry (Turkey X disease) in 1960 (Blount, 1961; Goldblatt, 1969) and subsequently with fatal toxicoses in India and West Africa (N gindu et al., 1982). Aflatoxins in association with hepatitis B have been implicated as contributory epidemiological factors for primary human liver cancer in areas of Africa, China, and Southeast Asia where there is an extremely high inci- dence of this disease (Hsieh, 1986, 1989; Groopman et al., 1988; Yeh et al., 1989). The mode of action of this class of mycotoxins involves metabolic activation to a reactive epoxide and its subsequent covalent binding to cellular macromolecules such as DNA and proteins (Busby and Wogan, 1981; Swenson, 1981; Kiessling, 1986). Since their discovery in 1960, aflatoxins have been the subject of intense research by agricultural scientists including mycologists, chemists, and toxicologists (Goldblatt, 1969; Heathcote and Hibbert, 1978; Betina, 1984; Smith and Moss, 1985). 19 20 A. Aflatoxins as Secondary Metabolites Aflatoxins are secondary metabolites because they are not essential for growth. Several additional criteria, as summarized by Bennett and Christensen (1983), which distinguish aflatoxins and other secondary metabolites from the primary metabolites, which are ubiquitous and necessary for growth are described below. 1. Limited taxonomic distribution Aflatoxins is only produced by certain strains of a few fungal species: A. flavus, A. nomius, and A. parasiticus (Bennett, 1982). Moreover, the amount of aflatoxin produced by aflatoxigenic strains varies widely. In general, isolates of A. parasiticus consistently produce both B and G aflatoxins, while A. flavus isolates contain a greater proportion of nontoxigenic strains and tend to produce only B aflatoxins (Hesseltine et al., 1970; Klinch and Pitt, 1985). 2. Produced in chemical families The name "aflatoxins" describes a group of closely related compounds. The aflatoxins are substituted coumarins containing the reactive difuran moiety. The major aflatoxin families are called BL B; G,, and G, (Figure 2) based on their respective blue and green fluorescence under long-wave ultraviolet (uv) light 21 Figure 2. Structures of the major aflatoxins (adapted from Zaika and Buchanan, 1989). 22 .3 2:02:33; S 53:32 ...... 4.0%. _ x s... \ am 53:32 ....o- ..z .... a a: 2:83: .... ..m ...o. a 3 23853: .2. .m- z _ \ .o 2.3.2.73 ao/jo _ / can... \ .2 2:053“; .10»: .m z_xo»<...._< £1. 0 / con: 23 and their relative chromatographic mobilities. Typically, aflatoxin B, is the major metabolite produced by all aflatoxigenic strains, with the other aflatoxins produced to a lesser extent. The hydroxylated aflatoxins (aflatoxin B2, and G“) (Figure 2) occasionally occur as minor metabolites (Heathcote and Hibbert, 1978). 3. Produced after active cellular growth Like other secondary metabolites, aflatoxins are synthesized after aetive growth of the mycelia has stopped. Morphological differentiation frequently occurs during this synthesis period (Turner, 1971). 4. Synthesized from simple building blocks Aflatoxins are synthesized from a few simple precursors that are derived from primary metabolism, e.g., acetate, malonate, and the methyl group of methio- nine (Steyn et al., 1980; Applebaum and Marth, 1981; Bennett and Christensen, 1983). B. The Biosynthesis of Aflatoxins While great strides have been made in elucidating the chemistry, toxicity and biological activities of aflatoxins, information on their biochemistry, specifically their biosynthesis, has accumulated more slowly. However, due to improved experimental techniques and analytical instrumentation (as described below), aflatoxin biosynthesis 24 has attracted considerable research attention in recent years. The subject of _ aflatoxin biosynthesis has been extensively reviewed (Maggon et al., 1977; Bennett and Lee, 1979; Steyn, 1980; Bennett and Christensen, 1983; Turner and Aldridge, 1983; McCormick et al., 1986). 1. Overview of aflatoxin biosynthesis The basic skeleton of aflatoxin B, [a CD polyketide (decaketide)] is derived entirely from acetate units via the polyketide pathway, and methionine contributes the methoxy-methyl group (Figure 3) (Donkersloot et al., 1968; Biollaz et al., 1970; Hsieh and Mateles, 1970, 1971). The polyketide pathway can be divided into two distinct phases. "Phase one" is common to all compounds of polyacetate origin, while "phase two" is unique to the particular polyketide being synthesized (Bu’Lock, 1961). During "phase one" ten acetate units are assembled into a polyketide chain, i.e., anthrone (Figure 4). This assembly reaction is presumably catalyzed by the bfiefidfiMfi an enzyme complex very similar to the eukaryotic fatty acid synthase in structural organization, function and mechanism of catalysis (Bennett and Christensen, 1983).. .The anthrone intermediate thus formed is further converted into aflatoxin B, in "phase two" by a series of steps involving several precursors and reactions including oxidation, reduction, condensation and molecular rearrangements (Figure 5). To date the known precursors to aflatoxin B, include seven anthraqui- nones (norsolorinic acid, averantin, averufanin, averufin, hydroxyversicolorone, versiconal hemiacetal acetate, and versicolorin A) and two xanthones [sterigma- tocystin (ST), and O-methylsterigmatocystin (OMST)]. 25 Figure 3. The origin of the carbon atoms and the arrangement of intact acetate units in aflatoxin B, (adapted from Biollaz et al., 1968, 1970). 26 o cu,-c’<- 0" cn,s- cu, cam (my) coon I O A 27 Figure 4. The hypothetical scheme for the assembly of anthraquinones by a "polyketide synthase" enzyme complex in Aspergillus species (adapted from Bennett and Christensen, 1983). 28 29 Figure 5. The proposed pathway for the biosynthesis of aflatoxin B,. Arrows interrupted by a short double line indicate steps that are blocked in the pathway. The corresponding blocked mutants of A. parasiticus are shown. An arrow interrupted by a single dash line indicates the conversion step which is inhibited by the insecticide dichlorvos. Arrows may indicate more than one step. 30 n _A_- erasittcu: a L“ m ATCC 24690 —e l ' . ’I an Noreolor-lnle Acid \ n e Averantln \/ 1" CD. on; i l. hm Averulenln O ’ 1:1'WJ' \/ ’\ a 2 itl . a , Tmcmssim amtrnrslcolom. \’ I] _A_. par-lumen: \ i mic: Vulcan: Hemtacetel mut- ‘chhlorvos \, .1 so ma’ltlcus ,\ m: / Sterlgmatocylun O-methyleterlgmetocyeun A. 35. mitic\g ”\ 02% union 3, 31 Figure 6. The metabolic scheme proposed for the late stages of aflatoxin biosynthesis (modified from Yabe et al., 1988, 1989). __ , confimd reactions; ----, hypothetical reactions. Abbreviations: AFB,, aflatoxin B,; AFB,, aflatoxin B,; AFG,, aflatoxin G,; AFG,, aflatoxin G1; DMST, dimethyl-sterigmatocystin; DHDMST, dihydro-dimethyl-sterigmatocystin; ST, sterigmatocystin; DHST, dihydro-sterigmatocystin; OMST, O~methyl- sterigmatocystin; DHOMST, dihydro-O—methylsterigmatocystin; MT-I, G methyltransferase I; MT-II, O-methyltransferase II; OR, oxidoreductase. No"? l .. .. «92 ll 5.20.3 lpmzo lemzozo l. .. ... p . ’ 32 a mo ....E. 2.2 «it x x 33 The biosynthetic pathway of the other aflatoxin families has not been well understood. Generally it has been assumed that aflatoxins 8,, G,, and G, are synthesized by direct interconversions from aflatoxin B, (Bennett and Christensen, 1983). However, studies of Floyd and Bennett (1981), and Dutton et a1. (1985) provided evidence that aflatoxin B, and B, could arise independently via a branched pathway. Recent findings (Cleveland et al., 1987; Floyd et al., 1987; Henderberg et al., 1988; Yabe et al., 1988, 1989; Cleveland, 1989) provide further support for this branched pathway hypothesis for the late stages of aflatoxin biogenesis. The currently-favored scheme is depicted in Figure 6. 2. Approaches to elucidating the aflatoxin biosynthetic pathway Tluee approaches have been utilized for the elucidation of the aflatoxin biosynthetic pathway. In addition to the genetic approach, which is the major focus of this review, chemical and biochemical approaches have contributed tremendously to our understanding of this pathway. a. Chemical approach. By marking aflatoxin B, with differentially labeled radio- active acetates and subjecting the compounds to selective degradation, Biollaz et a1. (1968, 1970) established unambiguously the polyketide origins of aflatoxin B,. The origin of the 16 skeletal carbon atoms was determined: 9 carbon atoms were derived from 01 of acetate and 7 carbon atoms were derived from C-2 of acetate (Figure 3). This powerful technique, however, is tedious to perform, and therefore is no longer widely applied to the aflatoxin biosynthesis. Nuclear magnetic resonance . 34 (NMR), in particular 13C-NMR (Sequin and Scott, 1974), represents another powerful technique that has been widely used for exploring biosynthetic pathways in recent years. Studies using this technique have provided information concerning bonds broken and formed during the aflatoxin biosynthetic process, and have revealed some mechanistic details concerning formation and subsequent conversion of each individual aflatoxin precursors (Steyn et al., 1980; Townsend, 1986). b. Biochemical approach. Our current knowledge about aflatoxin biosynthesis was derived mostly from the utilization of versatile biochemical techniques. Several techniques have contributed significantly to the elucidation of the aflatoxin biosyn- thetic pathway. 1) Bioconversion with blocked mutants. Aflatoxigenic Aspergillus spp. can be mutagenized to yield mutants that do not produce or produce a limited amount of aflatoxins. These blocked mutants (Figure 5 and Table 2) are presumably impaired in one or more enzymatic steps involved in the aflatoxin biosynthesis. Singh and Hsieh (1977) pioneered the use of biotransformations by blocked mutants to esta- blish a tentative sequence for the intermediates of the aflatoxin pathway. When a mutant blocked early in the pathway is incubated with a putative late intermediate, the recovery of aflatoxins simultaneously supports the role of the putative interme- diate as an aflatoxin precursor and places the compound past the site of the mutant block. Radioactive aflatoxin precursors used in biofeeding studies are normally prepared by fading a radioactive acetate to the blocked mutant that has accumu- lated the desired precursor. 35 2) Radiolabeling. As summarized in Table 1, metabolism of “C-labeled precursor molecules by blocked mutants has been routinely used in radiotracing experiments to confirm and place suspect precursors in the aflatoxin biosynthetic pathway. Radio- acrive compounds can also be used in kinetic pulse-labeling experiments to follow the metabolism of radioactive precursors as a function of time, thus revealing an ordered sequence of metabolite appearance (Zamir and Ginsburg, 1979; Zamir and Hufford, 1981). This technique is also advantageous because transient aflatoxin precursors can be distinguished from "dead-end" metabolites by the rapid formation and disappearance of the former. 3) Metabolic inhibitors. A number of chemicals are able to inhibit aflatoxin production (Zaika and Buchanan, 1987), but only dichlorvos (dimethyl-2,2-dichlo- rovinyl phosphate, an insecticide) has been frequently used for the elucidation of the aflatoxin biosynthetic pathway. Dichlorvos specifically inhibits the enzymatic conversion of versiconal hemiacetal acetate to versicolorin A in the aflatoxin biosynthetic pathway (Rao and Harein, 1972; Hsieh, 1973; Bennett et al., 1976; Singh and Hsieh, 1977). The effect of dichlorvos on intermediate accumulation by A. parasiticus blocked mutants is summarized in Table 1. 4) Enzymology. Only limited information is available with respect to the enzymes associated with the aflatoxin biosynthesis (Dutton, 1988). A few cell-free systems have been developed for conversion of certain precursors into aflatoxin B, or other 36 Table l. Incorporation of labeled precursors into aflatoxin B,. Labeled Blocked “C incor- Dichlorvos inhibition of compound mutant poration (%)l label into aflatoxin B, Norsolorinic NOR-1 2.2 n.a.z Acid Averantin AVN-l 10.2-28.5 Yes Averufanin n.a. 23.0 n.a. Averufin AVR-l 7.4-49.4 Yes Hydroxyversi- HVN-l n.a. n.a. colorone Versiconal hemi- n.a. 8.0-13.7 Yes acetal acetate Versicolorin A VER-l 34.5-50.5 No Sterigmatocystin n.a. 17.0-65.0 No O-Methyl-sterig- SRRC 72.5 No matocystin 2043 1: Data are adapted from Bennett and Lee (1979), and Bennett and Christensen (1983) except for averufanin (McCormick et al., 1987), hydroxyversicolorone (Townsend et al., 1988) and O-methylsterigmatocystin (Bhatrragar et al., 1987). 2: Data not available. 37 Table 2. Aflatoxin intermediates accumulated by blocked mutants of A. parasiticus Blocked mutant Genotype Intermediate accumulated ATCC 24690 (NOR-1) nor-1 N orsolorinic Acid ATCC 56774 (AVN-l) avn-I, nor-I Averantin, Versicolorin A ATCC 24551 (AVR-l) avr-I Averufin HVN-l hvn-I Hydroxyversicolorone ATCC 365 37 (VER- l) ver-I Versicolorin A SRRC 2043 n.a.3 O-Methyl-sterigmatocystin ‘: Mutants NOR-1 (Lee et al., 1970), AVN-l (Bennett et al., 1980a), AVR-l (Donkersloot et al., 1972) and VER-l (Lee et al., 1975) were derived from the same wild-type strain A. parasiticus NRRL 5862 (SU-l or ATCC 56775) by mutagenesis with UV-light or nitrosoguanidine. Mutant HVN-l (Townsend et al., 1988) was derived from A. parasiticus SU-7. SRRC 2043 (Bhatnagar et al., 1987) was originated from an A. parasiticus isolate CP461 (Dorner et al., 1984). 2: Data net available. 38 intermediates in the pathway (Singh and Hsieh, 1976; Anderson and Dutton, 1979; Anderson and Dutton, 1980; Wan and Hsieh, 1980; Jeenah and Dutton, 1983; Bhatrragar et al., 1989), but most of the nature of the enzymatic activities involved remain unclear to date. Purification and characterization of some of the relevant enzymes has been reported recently (Cleveland and Bhatnagar, 1987; Cleveland et al., 1987; Bhatrragar et al., 1988, Yabe et al., 1989). For example, Yabe et al. (1989) have identified two distinct O-methyltransferase activities in the aflatoxin biosynthetic pathway (Figure 6). One of these is responsible for the conversion of dimethyl sterigmatocystin (DMST) and dihydro-DMST to ST and dihydro-ST, resp- ectively. The other is involved in the conversion of ST and dihydro-ST to OMST and dihydro-OMST, respectively. The latter enzyme has been purified to homoge- neity by Bhatnagar et al. (1988). An attempt to clone the gene coding for this enzyme using a cDNA expression library was also initiated by Cleveland and Bhatnagar (1988). C. Intraspecific Variation in Toxin Production The aflatoxin-producing ability is strain-specific among natural populations of A. flaws and A. parasiticus. Not all wild-type strains of these fungi produce afla- toxins. For example, Diener and Davis (1969) screened more than one thousand natural isolates of A. flavus collected from six countries and found that only approximately 60% of them produced detectable aflatoxins. Bennett (1982) surveyed published literature studies encompassing 3343 isolates and found that a total of 1847 (56%) were aflatoxigenic. Recently Wei and long (1986) surveyed 39 the aflatoxin-producin g ability of 169 strains of the Aspergillus reference cultures in the A. flavus group maintained in the American Type Culture Collection (ATCC). A relatively low percentage (33%) of A. flow: strains were found aflatoxigenic, when compared to A. parasiticus strains (85%). Aflatoxin-producing ability can be lost or decreased after serial transfer in the laboratory (Mayne et al., 1971). The basis of this phenomenon is presumably genetic, but little is known about inheritance of aflatoxin-producing ability. D. Interspecific Variation in Toxin Production Aflatoxin production has been reported only from the fungal species A. flavus, A. parasiticus and A. nomius. Early reports that other fungal species also produced aflatoxin have never been confirmed (Detroy et al., 1971; Bennett and Papa, 1988). A. flavus and A. parasiticus are classified in the A. flavus group by Raper and Fennell (1965) along with the koji molds (A. oryzae and A. sojae) which have been used extensively in Asia for the production of various types of fermented foods. The A. flavus group is taxonomically complex. The morphological char- acteristics of strains within this group are not distinct enough to allow unambiguous differentiation among strains. Because of the possibility that koji molds might be aflatoxigenic, hundreds of Japanese industrial and domestic strains of A. oryzae and A. sojae have been screened. No aflatoxin has been detected in early studies (Murakami et al., 1967, 1968a, 1968b; Murakami, 1971), or in a more recent study by Wei and Jong (1986). Wicklow (1983) has recently reexamined various koji molds, and, based on their morphological and cultural characteristics, has concluded 40 that A. oryzae has originated as a "domesticated" strain of A. flavus, with a similar relationship existing between A. sojae and A. parasiticus. In support of this theory, homology studies based on DNA hybridization (Kurtzman et al., 1986) showed no detectable differences between A. flavus and A. oryzae, and 91% relatedness between A. parasiticus and A. sojae. . Several other fungal species have been reported to contain parts of the aflatoxin biosynthetic pathway (Moss, 1977; Steyn et al., 1980). Moreover, as summarized by Bennett and Deutsch (1985), a number of mold species also produce sterigmatocystin, one of the known intermediates in the aflatoxin pathway (Davies et al., 1960; Dean, 1963; Holzapfel et al., 1966; Mislivec et al., 1975; Schroeder and Kelton, 1975; Harnasaki et al., 1977; Rabie et al., 1977; Udagawa et al., 1979; Ayer et al., 1981; Davis, 1981; Sekita et al., 1981). These data suggest that genes encoding enzymes in the aflatoxin biosynthetic pathway up to sterigmatocystin are widely distributed in nature, while genes encoding enzymes for the last few steps are limited to A. flow: and A. parasiticus. From a genetic point of view, the reason that certain isolates of A. flavus, A. parasiticus, or the koji molds do not produce detectable aflatoxins could be that they lack of genes for the aflatoxin pathway, or that they are simply not expressing these genes. Similar reasoning could be used to explain fungal strains which share part of the aflatoxin pathway but produce no aflatoxins. To date, there are not sufficient data to distinguish between these possibilities. Attempts have been inititated in this lab and others (e.g., Woloshuk et al., 1989) to clone the structural genes of the aflatoxin pathway. These cloned genes can then be used as probes to unambiguously answer these questions. 41 E. Mutational Analysis of Aflatoxigenic Fungi A number of mutants have been isolated from A. flavus and A. parasiticus, and are summarized in a recent review by Bennett and Papa (1988). Some of these have been useful as markers for the elucidation of the parasexual cycle (see below); others have been used to shed light on aflatoxin biosynthesis (Steyn, 1980; Bennett and Christensen, 1983). l. A. parasiticus Three types of mutants have been described in A. parasiticus: Q) mutants blocked in the aflatoxin pathway and accumulate aflatoxin intermediates; (_2_) spore color and auxouophic mutants and (_3) the so-called fan and fluff variants which have lost or reduced aflatoxigenicity. Intermediate-accumulating mutants (Table 2) have brightly-colored mycelia which have been used as a visual screen in parasexual studies. Several rapid plate assays for the aflatoxin-producing ability have been developed to facilitate isolation of blocked mutants (Table 3). In these assays, fluorescence of the aflatoxin- producing colonies under the long-wave uv light enables one to differentiate bet- ween aflatoxin-producing and -nonproducing isolates. Spore color and auxotrophic mutants are generally used as markers in elucidating the parasexual cycle and gene- rating linkage maps. 42 Table 3. Quick screening media for analysis of aflatoxin-producing ability'. Name of medium Reference Aspergillus Differential Medium (ADM) Hyflo—Supercel Peanut Agar (HSPA) Aflatoxin Producing Ability Medium (APA) Coconut Agar Medium (CAM) Silica Gel Medium (SGM) Fluorescence Agar Medium (FAM) Bothast and Fennell (1974) deVogel et al. (1965) Hara et al. (1974) Lin and Dianese (1976) Davis et al. (1987) Lemke et al. (1988, 1989) Torrey and Marth (1976) Lennox and Davis (1983) * Modified from Bennett and Deutsch (1985). After incubation at 30°C for 2 to 4 days, cultures in Petri plates were inverted and viewed under illumination with a long-wave ultraviolet light. Aflatoxin-producing ability was indicated by a bright blue fluorescence zone around the producing colonies in all cases except for ADM in which formation of a yellow-orange pigment was a positive evidence for aflatoxi- genesis. 43 Strains of A. parasiticus which produce no detectable or less aflatoxins are of particular interest. They can be induced by successive transfers of mycelial macerates in defined media. Two such variants (fan and flufl‘) were isolated by Bennett et al. (1981a). These strains share the following common features: (1) Both arose spontaneously by positive selection under conditions that favor rapid vegetative growth; (2.) Attenuation or loss of the aflatoxin-producing ability is accompanied by loss of non-vegetative functions. For example, haploid and diploid isolates of the fan and fllgfi' phenotype exhibited less sporulation and altered colony characteristics. The fan variants yielded flat growth with heavy sporulation in the center of the colony but only scant sporulation at the edges. The flufi' variants produced abundant fluffy, aerial mycelium, and few spores. The fan variants produced no detectable aflatoxins, but the flufl' variants were unstable with respect to aflatoxigenicity (Bennett et al., 1986); (3) Both fan and flufi’ were recessive traits. Complementation occurred resulting in production of aflatoxin in heterozygous diploids derived from blocked mutants and the fan variants (Bennett et al., 1981b, 1986); (4) Both are chromosomal mutations but are not linked to any structural genes for aflatoxin synthesis (Bennett et al., 1981b). It is interesting to note that the "lazy" mutants of Fusan‘um graminearwn isolated by Duncan and Bu’Lock (1985) showed the similar phenotypic and geno- typic characteristics to the fan and fllgfl' variants of Aspergillus (Bu’Lock, 1986; Bu’Lock et al., 1986a, 1986b). These mutants arose by a chromosomal change and were most readily obtained through repeated serial subculture under non-limiting conditions on rich media. They were characterized by a radically different colony morphology and pigmentation as well as by the substantial or complete loss of the 44 zearalenone-producing ability. Bu’Lock (1986) and Bu’Lock et al. (1986a, 1986b) suggested that the lazy strains contained the full complement of structural genes for zearalenone production but were unable to express them because the lazy mutation affected a regulatory function that governed the expression of the structural genes for zearalenone synthesis. Whether or not this is also the case for the fan and 1w variants remains unclear. As suggested by Bennett (1982), the high frequency with which fan and first? variants are isolated, the pleiotropic nature of their respective phenotypes, and the anomalous behavior of genes nor-1 and ver-I in crosses involving fan and fllgfl’ imply that genetic transpositions are involved. However, direct evidence for this proposition is lacking. 2. A. flavus Three major classes of mutants have been isolated in A. flavus, including Q) high aflatoxin B,-accumulating strains; L2) spore color and auxotrophic mutants, and (3) aflatoxin-negative mutants. Among these mutants, of particular interest are those which accumulate a high level of aflatoxin B,. A number of these mutants have been isolated from natural environments (Van Walbeek et al., 1968; Schroeder and Carlton, 1973; Gunasekaran, 1981). The strain isolated by Schroeder and Carlton has been used to elucidate the late stages of aflatoxin biosynthesis (Dutton et al., 1985). A mutant producing a high level of aflatoxin B, over B, was induced by nitrosoguanidine treatment (Papa, 1977a). This mutant (afl-BZ) has been linked to the histidine locus on linkage group VIII (Papa, 1977a, 1977b, 1977c, 1981). 45 Other classes of mutants were isolated and characterized by Bennett and Papa (1988). These mutants have been used in parasexual analysis and genetic mapping. F. Viral Association with Aflatoxin Production The possibility that aflatoxin production is influenced by extrachromosomal genetic elements in a non-toxigenic isolate of A. flavus (NRRL 5565) was inves- tigated by Schmidt et al., (1983). This strain harbored unusual proteinaceous virus- like particles (Wood et al., 1974) consisting of double-stranded RNA (ds-RNA) components that shared characteristic features with a virus that is associated with Penicillin»: strains, particularly P. Chrysogenum. (Schmidt et al., 1986). Since the viral particles in A. flavus NRRL 5565 showed unusual properties, such as an extremely low replication rate and a high proportion of smaller, empty capsids (Wood et al., 1974; Schmidt et al., 1986) that could be attributed only to host influences, Schmidt et al., (1986) suggested that the virus in A. flavus NRRL 5565 normally resides in a foreign host. Elimination of this virus by viral antimetabolites (cycloheximide and 5- fluorouracil) resulted in the simultaneous losses of its ds-RNA trait and initiation of stable aflatoxin formation by A. flavus (Schmidt et al., 1983), which, in turn could be switched off by artificial infection with a ds-RNA virus from P. chrysogenwn (Schmidt and Esser, 1985; Schmidt et al., 1986). This finding suggested that the virus from P. chrysogenum was able to repress aflatoxin production of A. flavus and supported the hypothesis that this virus may have been transmitted from a foreign host to the originally toxigenic strain of A. flavus NRRL 5565. These data also 45 suggested that the viral ds-RNA present in NRRL 5565 somehow repressed aflatoxin synthesis. When removed or reduced in titer by treatment with antiviral drugs, the fungus regained a native ability to produce aflatoxin. Thus, the nonproduction of aflatoxins was due, in this case, to the failure of gene expression rather than the absence of structural genes of aflatoxin biosynthesis. The mechanism of this viral control of aflatoxin formation remains unknown at this moment. However, it should be noted that a yellow-spored mutant of A. flavus NRRL 5565 consistently produced no detectable aflatoxin even after repeated exposure to cycloheximide (Lemke et al., 1989), suggesting that previous results of aflatoxin production in this strain following cycloheximide treatment could be due to contamination by a toxigenic strain. Further investigation on this phenomenon may shed light on genetic regula- tion of aflatoxin biosynthesis in this particular A. flavus strain. G. The Parasexual Analysis Although neither A. flavus nor A. parasiticus possess a sexual stage (teleo- morph), there is evidence that a parasexual cycle functions in both species (Bennett, 1979; Gussack et al., 1977; Papa, 1973, 1978). Parasexual analysis involves forma- tion of heterokaryons followed by isolation of heterozygous diploids fiom these heterokaryons, and the subsequent recovery of recombinant cells from the hetero- zygous diploid (Roper, 1966). Unlinked genes segregate independently during haploidization, and recombination between linked genes occurs during mitotic crossing over. This process provides an "alternative sex" for many imperfect 47 (anamorphic) fungi and has made formal genetic analysis possible for the aflatoxi- genic molds. The genetics of A. flavus is more deve10ped than that of A. parasiticus. Over Thirty genes have been mapped to 8 linkage groups via parasexual analysis (Papa, 1976, 1977a, 1977b, 1977c, 1979, 1980, 1982, 1984). A complete list of these genetic loci of A. flavus, including known linkage assignments, has been compiled by Bennett and Papa (1988). It appears that the genes encoding proteins responsible for aflatoxin synthesis and regulation of this metabolic pathway have been mapped to different chromosomes with some clustering of loci on particular chromosomes. Fewer genetic loci have been studied in A. parasiticus. Papa (1978) assigned seven loci to four linkage groups (I through IV), and ten loci were assigned to six linkage groups by Bradshaw et al. (1983). As pointed out by Bennett and Papa (1988), different laboratories used different parent strains, different genetic markers and nomenclatures, as well as different blocked aflatoxin mutants because virtually no collaborative studies have been initiated among laboratories. Therefore, until genetic nomenclature and linkage groups are standardized, the mapping data from each laboratory must be evaluated with caution. The conventional designations for fungal gene symbols and pheno- types as described by Bennett and Lasure (1985b) and Yoder et a1. (1986) are followed throughout this thesis. Although parasexual analysis allows genetic analysis of aflatoxigenic molds, it is somewhat unreliable and laborious to perform due to the following reasons as summarized by Bennett (1982), which significantly impede the development of the genetic maps for the aflatoxigenic molds. 48 a. Cell ploidy. In parasexual analysis, a haploidization agent such as p-fluoro- phenylalanine (FPA) or benlate (containing 50% benomyl) is usually used to incr- ease the number of haploid segregants from heterozygous diploids. In species with uninucleate conidia (e.g., A. nidulans), ploidy is readily determined by conidial diameter, with diploid spores derived from nuclear fusions being larger than haploid spores. However, determination of cell ploidy is nearly impossible in wild-type A. flavus and A. parasiticus which have multinucleate conidia (Y uill, 1950). First, microscopical examinations are not able to detect differences in conidial diameters in haploid, heterokaryotic and diploid spores of these fungi. Second, total DNA content is nearly equal in these spores because spores in diploids have fewer nuclei than do haploids or heterokaryons (Leaich and Papa, 1975; Bennett et al., 1980b). The inability to directly detect cell ploidy by spore diameter entails the utilization of indirect methods which are laborious, and often can not be used to distinguish from strain instability arising from aneuploidy such as has been detected in A. nidulans (Birkett and Raper, 1977). b. Genetic segregation. During parasexual cycle, a number of auxotrophic markers are recovered in extremely low yields on media containing haploidization agents (FPA and benlate). Furthermore, spore and mycelial color markers have been found to segregate nonrandomly (Bennett, 1979; Bennett et al., 1980b). c. Heterokaryon incompatibility. Strains belonging to the same heterokaryon (vegetative) incompatibility (or compatibility) group generally do not mate or allow 49 hyphal anastomosis to form a heterokaryon. Th phenomenon is widespread in the A. flavus group. Formation of heterokaryons has been reported only among comple- menting auxotrophs derived from the same ancestral wild-type strain (Gussack et al., 1977; Bennett, 1982). Papa (1986) detected 22 different heterokaryon compatibility groups among 32 A. flavus strains collected from 15 Georgia counties. Strains within the same heterokaryon compatibility group were not restricted to the same geographical area. Six intraspccific crossing of auxotrophs from A. flavus PC-7 and A. flavus NRRL 5565 failed to grow, as did 12 interspecific combinations of auxo- trophs of A. flavus and A. parasiticus (Gussack et al., 1977). Attempted protoplast fusions between A. flavus and A. parasiticus auxotrophs were also unsuccessful (Bennett, 1982). H. Genetic Transformation in Aflatoxigenic Fungi As discussed above, the development of aflatoxin genetics is still at a very early stage. The difficulties arising from the lack of a sexual reproduction cycle of the aflatoxigenic molds make traditional Mendelian genetic analysis of aflatoxin biosynthesis nearly impossible. In addition, as summarized by Bennett (1982), a number of technical problems impede research on the genetic basis of biosynthesis of other fungal secondary metabolites as well. First, aflatoxins are not direct gene products. They are produced by the enzymatic conversion of small precursor mole- cules. Second, cell-free systems are poorly developed and only a few of the enzymes involved in aflatoxin biosynthesis have been isolawd (as discussed in section B). Third, strain instability in originally high aflatoxin-producing strains 50 occurs at a higher frequency than background mutation rates. Fourth, mutants aff- ecting the aflatoxin biosynthetic pathway are difficult to obtain because conditional lethal protocols cannot be employed, because with all secondary metabolites, afla- toxins are not essential for growth of the producing organism. Recent advancements in the molecular biology of filamentous fungi (as reviewed below) offer innovative and more effective approaches for studying afla- toxin biosynthesis. Transformation is a gene transfer process in which a naked DNA molecule is introduced into cells. It is generally considered an essential step in modern genetic engineering techniques. Transformation systems developed for a variety of filamentous fungi have resulted in cloning of a growing number of genes by functional complementation of mutations (Fincham, 1989). Such a transforma- tion system has recently been developed for A. flavus (Woloshuk et al., 1989). This system is based on complementation of a pyrimidine auxotrophic mutation with vectors containing the heterologous pyr-4‘ gene of N. crassa as a selectable marker. The pyrimidine mutation in the recipient strain was obtained by screening muta- genized cells for resistance to the toxic metabolite 5-fluoro-orotic acid (S-FOA) (Boeke et al., 1984). The pyrimidine mutation in one strain could be transferred into different strains through parasexual recombination, thus making mutant isolation somewhat easier to perform. The transformation frequency of A. flavus with this system is generally low (about 20 transformants/pg DNA) but, in theory, is suffi- cient to screen a cosrnid genomic DNA library. Coupled with aflatoxin-nonpro- ducing mutants, this system may allow one to clone genes involved in aflatoxin synthesis in A. flavus. A more versatile transformation system based on the homologous benomyl resistance gene (a mutated version of the B-tubulin gene) as a 51 selectable marker is also being developed for A. flavus (Seip et al., 1988, 1989). This type of dominant selectable marker is highly desirable because any fungal strain sensitive to the inhibitors (benomyl) can be used as a potential recipient in transformation. A. parasiticus offers a distinct advantage over A. flavus in studying afla- toxin biosynthesis at the molecular level since there exist several blocked mutants of this fungus which will allow cloning of genes involved in the aflatoxin biosynthetic pathway by functional complementation. Development and characterization of selec- table markers for genetic transformation systems in the aflatoxigenic A. parasiticus is the major goal of this investigation. 11. Genetic Transformation in Filamentous Fungi The importance of filamentous fungi in medicine, industry, and agriculture has led to intensive investigations into their physiology and metabolic regulation (Bennett and Lasure, 1985a; Tirnberlake, 1985). However, due to the lack of a “sexual stage, fundamental genetic analysis in many of these important groups of fungi has been impossible or difficult to perform. Therefore, it was desirable to develop a deoxyribonucleic acid (DNA)-mediated transformation systems to conduct genetic studies. The first confirmed DNA-mediated transformation in filamentous fungi was reported by Case et al. (1979) who used the cloned N. crassa qa-Z+ (catabolic dehydroquinase) gene as a selectable marker. Transformants were shown by Southern hybridization analysis to contain chromosomally integrated plasmid molecules. Subsequently, many Neurospora genes have been used as selectable markers for transformation, including pyr-4” (Ballance et al., 1983), trp-I+ (Schechtrnan and Yanofsky, 1983), am’ (Kinnaird et al., 1982), and bml (benomyl resistance) (Orbach et al., 1986). Transformation of A. nidulans, another well- studied filamentous fungus, was first reported by Ballance et al. (1983), who used the N. crassa pyr-4+ gene to complement an A. nidulans pyrG mutation. Shortly after, Tilburn et al. (1983) used the A. nidulans amdS‘ gene, encoding acetamidase ,Hynes et al., 1983), and Yelton et al. (1984) used the A. nidulans t‘rpC+ gene for ransformation. In each case, transforming DNA became integrated into the genome. lubsequently, other A. nidulans genes were used as transformation markers, inclu- ing argB* (Berse et al., 1983; John and Peberdy, 1984), pm+ (Arst et al., 52 ‘ ' ... Table 4. Filamentous fungi in which transformation has been achieved'. Type of fungus Oomyceres Achlya ambisexualis Ascomycetes Ascobolus immersus Aspergillus nidulans Claviceps purpurea Cochliobolus heterostrophus’ Endothia parasitica’ Gaeumannomyces graminis Gibberella pulicaris’ Glomerella cingulara‘ Hypoxylon marmnarum Leprosphaeria maculans‘ Magnaporthe grisea‘ Necrrl'a haematococca’ Neurospora crassa Podospora anserina Sordaria macrospora Related Fungi Imperfecti Aspergillus awamori Aspergillus flavus Aspergillus ficuum Aspergillus niger Aspergillus oryzae Aspergillus terrcus Borryris squamosa Cephalosporium acremonium Colletotrichwn capsici Colletotrichum graminicola Colletotrichum gloeosporioides Colletom‘chum lindemuthianum Colletotrichum trifolii F ulvia fulva' Penicillium caseicolum Penicillium chrysogenwn Penicillin»: nalgiovense Reference Manavathu et a1. (1987) Faugeron et al. (1989) Tilburn et al. (1983) van Engelenburg et al. (1989) Turgeon et al. (1985) Van Alfen et a1. (1988) Henson et al. (1988) Salch & Beremand (1989) Rodriguez & Yoder (1987) Griffin et al. (1989) Farman & Oliver (1988) Parsons et al. (1987) Van Etten & Kistler (1988) Case et al. (1979) Brygoo & Debuchy (1985) Le Chevanton et al. (1989) Cullen & Leong (1986) Woloshuk et al. (1989) Mullaney et al. (1988) Goosen et a1. (1987) Mattem et a1. (1987) Katz & Hynes (1989b) Huang et al. (1989) Skatrud & Queener (1984) Soliday et al. (1989) Panaccione et al. (1988) TeBeest & Dickman (1989) Daboussi et al. (1989) Dickman (1988) Oliver et al. (1987) Daboussi et al. (1989) Diez et a1. (1987) Geisen & Leismer (1989) 54 Table 4 (cont’d.). Basidiomycetes Coprinus cinereus Phanerochaete Chrysosporium Schizophyllwn commune Ustilago maydis Ustilago hordei Ustilago nigra Ustilaga violacea Zygomycetes Absidia glauca Mucar circinelloides Phycomyces blakesleeanus Other Fungi Imperfecti Aphanocladium album Beauveria bassiana Curvularia lunata Fusarium culmorum Fusarium graminearum Fusarium monilifonne Fusarium oxysporium Gliocladium virens Gliocladium rosewn Metarhizium anisopliae Pseudocercosporella herpom'choides Scyralidium flavo-brunneum Septoria nodorum’ Trichoderma reesei Binninger et al. (1987) Alic et al. (1989) Munoz-Rivas et al. (1986b) Banks (1983a) Holden et al. (1988) Holden et a1. (1988) Bej & Perlin (1989) Wbstemeyer et al. (1987) Van Heeswijck & Roncero (1984) Revuelta & Jayaram (1986) Daboussi et al. (1989) Daboussi et al. (1989) Osiewacz & Weber (1989) Madhosingh & Orr (1985) Dickman & Leslie (1989) Leslie & Dickman (1989) Kistler & Benny (1988) Thomas & Kenerley (1989) Thomas & Kenerley (1989) Berrrier et al. (1989) Blakemore et al. (1989) Caprioglio & Parks (1989) Cooley et al. (1988) Penttilil et al. (1987) Adapted from Fincham (1989). ‘: Anamorph: Helminthosporium maydis = Bipolaris maydis = Dreschlera maydis. ~30“. ’: Synonym: Cryphanecrria parasitica, the causal agent of chestnut blight. ’: Anamorph: Pusarium sambucinum, causing dry rot on potato tubers. : Anamorph: Colletotrichurn lindemuthianum, a pathogen of bean. : Synonym: Phoma Iingam, causing the blackleg disease of brassicas. : Anamorph: Pyricularia oryzae or P. grisea, the causal agent of rice blast disease. : Anamorph: Fusarium solani f. sp. pisi, a pathogen of pea. : Synonym: Cladosporium fulvum, a major foliar pathogen of tomato. : Teleomorph: Leprosphaeria nodorum, causing a leaf spot disease of wheat. 55 1985), pyrG* (Oakley et al., 1987), pabaA‘ (Timberer and Marshall, 1989), and BenA‘ (Dunne and Oakley, 1988). Following the success with N. crassa and A. nidulans, transformation systems were rapidly developed in many other fungal species of medical, industrial, and agricultural significance (Fincham, 1989; Table 4). Transformation of filamentous fungi has been extensively reviewed (Bennett and Lasure, 1985a; Johnstone, 1985; Mishra, 1985; Timberlake, 1985; Cullen and Leong, 1986; Esser and Mohr, 1986; Hynes, 1986; Saunders et al., 1986; Rambosek and Leach, 1987; Leong, 1988; Schwab, 1988; Fincham, 1989; Timberlake and Marshall, 1989). A. Transformation Techniques Although the original transformation protocols developed for N. crassa and A. nidulans have been modified and improved, they remain fundamentally unch- anged. These include preparation of competent cells and DNA uptake. 1. Preparation of competent cells DNA-mediawd transformation requires that an organism take up and express exogenous DNA molecules that can confer a selectable advantage on those cells which receive them. Cells that are capable of taking up exogenous DNA are refe- rred to as "competent." In filamentous fungi, competence can be achieved in the following ways: 56 a. Protoplast. The fungal cell wall is a multilayer network structure of complicated organization which serves as an efficient barrier to DNA entry (Bartnicki-Garcia, 1968; Rosenberger, 1976; Farka§, 1985; Wessels, 1986; Cabib et al., 1988). Proto- plasts are typically generated by digesting away the fungal cell walls with cell wall- degrading enzymes. Novozym 234, a commercially available hydrolytic enzyme mixture (notably 1,3-glucanases and chitinase) secreted by the filamentous fungus Trichoderma harzianum, has been commonly used for this purpose (Hamlyn et al., 1981). In some fungal species (Turgeon et al., 1985; Binninger et al., 1987), it is necessary to use other enzymes in conjunction with N ovozym 234. These include enzymes from snail gut (Glusulase, helicase and B-glucuronidase), Zymolyase, cellulase, chitinase, driselase or Others (Peberdy, 1985). Experimental data suggests that the particular enzyme used for digestion of the fungal cell wall was often of great importance. For example, using Glusulase Kinsey et al. (1984) obtain a high yield of u'ansformants (IO‘Iug DNA) when sele- cting for the am‘ (glutamate dehydrogenase) gene. With different lot of Glusulase, however, the frequency decreased 100-fold. Akins and Lambowitz (1985) similarly found that one particular lot of N ovozym 234 was most suitable for preparation of competent Neurospora protoplasts. Because empirical observation suggests that there is significant variation in efficiency of transformation between batches of enzymes, Rambosek and Leach (1987) suggeswd "When in doubt, try another batch of enzymes to generate protoplasts." Protoplasts can be prepared from various cell types of filamentous fungi. In For example, young mycelium has been a major source of protoplasts for Neuro- spora (Buxton and Radford, 1984), Aspergillus (Y elton et al., 1984), and Penicillium 57 species (Sénchez ct al., 1987), as well as for Padospora anserina (Brygoo and Debuchy, 1985) and Ascobolus immersus (Faugeron et al., 1989) which do not pro- duce conidia. Protoplasts released can be easily separated fiom the hyphal debris by filtration. For Basidiomycetes, the sexual basidiospores (Munoz-Rivas et al., 1986b), the dikaryotic mycelium, or the asexual oidia (Binninger et al., 1987) has been used for protoplast production. One of the main advantages of preparing protoplasts from spores is obtaining a homogeneous suspension of pretoplasts with only one nucleus. Mononucleate pro- toplasts are most suitable for recombination and transformation experiments (Bos et al., 1983). Moreover, protoplasts from fungal spores are homogeneous in terms of size distribution and physiological condition, whereas protoplasts derived from mycelium are heterogeneous because of age differences of the hyphal cells from which they are originated (Peberdy and Gibson, 1971; Anne et al., 1974). Thus, protoplasts fiom fungal spores offer certain advantages over those from mycelium for genetic and physiological experiments. Several reports describe production of protoplasts from fungal spores (Chu and Alexander, 1972; Emerson and Emerson, 1958; Bachmann and Bonner, 1959; Weiss, 1965; Garcia Acha and Villanueva, 1963, 1964; Garcia Acha et al., 1966; Laborda et al., 1974; Moore and Peberdy, 1976; Res and Slakhorst, 1981; B03, 1985). Freezing has been used to preserve fungal protoplasts for successive transfor- mation experiments. However, the effect(s) of freezing on competence and survival of protoplasts has not been fully investigated. Competent Neurospora protoplasts could be stored frozen in the transformation bufi'er at -70°C for more than one month without loss in transformation frequency (Aldus and Lambowitz, 1985). 58 Neurospora protoplasts stabilized with sorbitol remained viable indefinitely at -70°c (Vollmer and Yanofsky, 1986). Frozen protoplasts of A. nidulans prepared accor- ding to this procedure yielded more reproducible transformation results than freshly prepared protoplasts (Cullen et al., 1987). However, storage of protoplasts of Trichoderma at -70°C resulted in 50% reduction in transformation frequency (Penttila et al., 1987). A similar result was observed for protoplasts of Schizo- phyllum (Specht et al., 1988). However, they could be stored on ice for 20h before use without affeCting the transformation frequency. Addition of glycerol to 15% (v/v) had no effect on protoplast viability, but completely inhibited transformation in Schizophyllum. Protoplasts are fragile and must be protected by osmotic stabilizers in the suspending medium. The type and concentration of stabilizer may influence the yield and the stability of protoplasts, and depending on the organism used, a wide range of osmotic stabilizers including inorganic salts, sugars and sugar alcohols have been used to stabilize protoplasts released from fungi (Davis, 1985). There is no single stabilizer suitable for all fungi. Some osmotic stabilizers such as KCl, MgSO,, and the sugar alcohol sorbitol and mannitol are used more often than others (Musilkova and Fencl, 1968; Sietsma and de Boer, 1973; Peberdy et al., 1976). Magnesium sulfate (1.2 M) has promoted the release of highly vacuolated protoplasts from S. commune (de Vries and Wessels, 1972) and A. nidulans (Peberdy and Isaac, 1976; Tilburn et al. 1983; Yelton et al., 1984) which could be easily separated from mycelial debris because they floated on the supernatant after cennifugation. However, protoplasts isolated from A. fumigatus in the presence of magnesium sulfate are osmotically very fragile (Hearn et al., 1980). 59 An essential requirement for obtaining growing colonies from protoplasts is the maintenance of the osmotic stabilizer in the growth medium until the cell wall has been regenerated. The same stabilizer is generally used for regeneration as well as for protoplast preparation, but the choice of plating conditions may be determined by the selectable marker in use. For example, cesium chloride is commonly added into media to reduce background nitrogen utilization by nontransformed cells. In this case, sucrose is the stabilizer of choice instead of potassium chloride (Tilburn et al., 1983). In contrast, sucrose or sorbitol prevents direct selection for carbon utilization genes. In general, spreading of the transformed cells embedded in an agar overlay of selective medium tends to improve the transformation yield by increasing the regeneration rate of protoplasts (Ballance and Turner, 1985; Borges and Barreau, 1989). However, there were reported cases in which directly spreading the transformd cells or embedding them in the agar overlay were indifferent with respect to transfonnants recovered. Oakley et al. (1987) were able to achieve a high transformation frequency (2400/ug DNA) in A. nidulans by eliminating top agar and plating transformed cells directly on the selective medium. b. Alternatives to protoplast. Several alternative methods have been developed that allow transformation of fungal cells without the need for tedious protoplast preparation. One approach is to use a mutant recipient strain that may have more perme- able membranes or cell walls. For example, the inl mutant of N. crassa was thou- ght to be particularly competent to take up DNA because of the greater membrane porosity when it was starved for inositol (Mishra and Tatum, 1973; Mishra et al., 60 1973). Germlings of this strain were transformable, although the frequency was low when compared to methods involving protoplasts (Mishra, 1977, 1979; Wootton et al., 1980). Cell wall-deficient mutants of Neurospora have also been used to facilitate the uptake of DNA without protoplast formation (Mishra et al., 1973). A cell-wall-less strain of Neurospora called slime could be used without glusulase treatment. However, genetic analysis of the slime strain of Neurospora is. difficult, making the use of this strain less attractive in transformation experiments (Mishra, 1985). The use of high concentrations of alkali metal ions (Iimura et al., 1983; Ito et al., 1983) represents another effective way to induce permeability of DNA into intact fungal cells. Lithium acetate at 0.1 M has been widely used for this purpose (Ito et al., 1983). This procedure has been successfully applied to N. crassa (Dhawale et al., 1984), Coprinus cinereus (Binninger et al., 1987), Colletorrichum mfolii (Dickman, 1988) and Fusarium solani f. sp. pisi (Nectria hematococa) (Soliday et al., 1989, Marek et al., 1989). Frequencies of transformation are generally lower than those obtained using protoplast methods but considerable time can be saved since generation of protoplasts is not necessary. This method, however, could not be applied in Usrilago maydis due to the toxic effect of the lithium ion to this fungus (Leong, 1988). In contrast, whole sporidial cells from U. violacea could be transformed at a high frequency when treated with lithium acetate and polyethylene glycol (PEG) (Bej and Perlin, 1989). How alkali metal cations assist the passage of DNA into cells is not well understood. 61 2. Uptake of DNA How cells become competent to take up exogenous DNA molecules remains unclear, and is a subject being intensively investigated. Nevertheless, several procedures are available to enhance entry of DNA into fungal cells. 8. Calcium and PEG. Calcium chloride (10 or 50 mM) is typically used in trans- formation protocols for Aspergillus species and other filamentous fungi. A treating time of 15 to 30 min at room temperature or on ice is generally sufficient for DNA uptake. Dimethyl sulfoxide (1%) has been added to the transformation mixtures for Neurospora, and some protocols have included heparin (Kinsey and Rambosek, 1984) and spermidine as well (V ollmer and Yanofsky, 1986). However, these addi- tives is not commonly used in transformation of fungi other than Neurospora species. PEG is one of the key components in virtually all fungal transformation protocols. It brings the treamd cells together and thus allows fusion to occur. DNA molecules are presumably internalized during this process because no transfor- mation occurs when PEG is omitted (Timberlake and Marshall, 1989). The effect(s) on the transformation frequency of varying the molecular weight and concentration of PEG has not been carefully investigated. Concentrations of PEG commonly used range from 20% to 70% with PEG 4000 (from various sources). PEG 6000 and PEG 8000 at 25% or 50% are also common. Tilburn et a1. (1983) found that incr- easing the concentration of PEG from 25 to 60% increased the frequency of 62 transformation in A. nidulans, but Turner and Ballance (1985) did not find the same effect. Increasing the concentration of PEG 8000 from 25% to 50% also increased the u'ansformation frequency 3 to 4-fold in Penicillium (Cantoral et al., 1987). b. Liposome-mediated uptake. An alternative way of introducing DNA into cells that could in principle be more efficient was devised by Fraley and coworkers (Fraley and Papahadjopoulos, 1981). They showed that DNA molecules can be encapsulated within the aqueous interior of phospholipid vesicles (liposomes). Subsequent fusion of liposomes with the plasma membrane results in intracellular delivery of the encapsulated nucleic acid. Whereas transformation by naked DNA is DNase-sensitive, that by liposomes is presumably not. Radford et al. (1981) first applied this technique in transformation of Neurospora. This procedure is effective in protecting transforming DNA but is not widely adopted because it appears to offer no advantages over the routine procedures with naked DNA to justify the extra work of liposome preparation. c. Heat shock treatment. During the transformation protocol, bacterial cells are generally treated with a short pulse of heat to enhance their competence in taking up DNA (Hanahan, 1983). However, the effect of heat shock treatment on the transformation frequency of filamentous fungi has not been well documented except in Podospora. Borges and Barreau (1989) observed that heat shock at an elevated temperature (48°C) for 5 min improved the transformation efficiency 5- to 10-fold in P. anserina protoplasts only if the heat shock was applied before the addition of transforming DNA. An increase in competence was observed immediately after the 63 heat shock. Heat-shocked cells remained competent for 20 to 30 min. The impro- vement in transformation efficiency after heat shock did not depend on the selec- table markers used for transformation. The mechanism by which heat shock impro- ves competence remains unclear, perhaps explaining why heat shock has not been widely used in fungal transformation. (1. Electromration. DNA can also be introduced into protoplasts by exposing cells to a brief electrical pulse or pulses of high field strength (Zimmermann, 1982; Zimmermann and Vienken, 1982). This technique, called electroporation, renders cell membranes temporarily permeable to macromolecules such as DNA. Electro- poration has been used successfully to study the transformation and expression of gene products in various cell types where other methods failed (Fromm et al., 1987; Knight and Scrutton, 1986; Neumann and Bierth, 1986). Ward et al. (1988) first transformed protoplasts of A. niger to oligomycin resistance by electroporation. Subsequently protoplasts from several other filam- entous fungi have been successfully transformed by this technique, including A. nidulans, F. solani (Richey et al. 1989), A. awamori, A. niger (Ward et al., 1989), and Gliocladium spp. (Thomas and Kenerley, 1989). In certain cases, transfor- mation frequencies were comparable to those obtained by the traditional protocol involving treatment of protoplasts with PEG and calcium ion. Electroporation may not be an appropriate method of transformation for species such as A. nidulans or N. crassa for which highly efficient PEG-mediated transformation protocols are already available. However, this method may provide an alternative of introducing foreign DNA into fungi that cannot be transformed by traditional methods. e. Other procedures. In addition to procedures described above, transformation protocols involving particle bombardment (Klein et al., 1987; Fox et al., 1988; Johnston et al., 1988) or partial cell breakage by blending with glass beads (Costanzo and Fox, 1988a, 1988b) have been proven feasible in yeasts, and could also be useful with filamentous fungi. B. Selection of Transformants Transformation systems require the presence of a marker on the trans- forming vector which allows growth of transformed colonies under selective conditions. 1. Selectable markers A wide variety of selection systems are now available in the filamentous fungi (Schwab, 1988) which fall into two broad classes: auxouophic markers and dominant resistance markers. Occasionally certain specific selectable (or screenable) markers such as those based on spore color or colonial morphology are also useful. a. Auxotrophic selectable markers The use of auxou'ophic selectable markers involves complementation of an auxotrophic mutation with the wild-type gene. Selection of transformants with this 65 system is usually straightforward. Several wild-type genes from filamentous fungi (Schwab, 1988; Fincham, 1989) have been particularly useful as general-purpose selectable markers and have frequently been incorporated into cloning vectors. A number of them are functional in several different fungal species (Table 5). In order for this type of selection system to work, it is necessary to obtain both a cloned wild-type gene (either from the same or different fungal strains) as well as the corresponding mutation in the recipient strain. This latter requirement is not always easy to meet, especially for genetically undeveloped fungi. One way to overcome these limitations is to utilize positive selection protocols which have been successful in generating a number of mutants in filamentous fungi. 1) Mutants defective in orotidine 5’-monophosphate (OMP) pyrophosphorylase and/or 0MP decarboxylase. The biosynthesis of uridine monophosphate (UMP) in filamentous fungi proceeds from aspartate and carbamoyl phosphate through the intermediate or0tic acid, to 0MP which is finally decarboxylated to UMP (Radford et al., 1985). The genes involved in the conversion of orotic acid to UMP have been named pyrF’ (OMP pyrophosphorylase) and pyrG‘ (OMP decarboxylase) in Aspergillus (Palmer and Cove, 1975), whereas the same genes in Neurospora are known as pyr-Z’ and pyr-4‘ respectively (Caroline, 1969; Perkins et al., 1982). Boeke et a1. (1984) found that wild-type strains of S. cerevisiae were sensitive to the pyrimidine analog S-fluoroorotic acid (5-FOA), presumably due to its conversion into the toxic 5-fluoro-UMP. Mutants resistant to 5-FOA were frequently uracil auxotrophs. Thus, this toxic pyrimidine analog is potentially useful in isolating mutants defective in genes encoding enzymes responsible for conversion of orotic Anna: .3505 d 83mph Ag: .7 8 83:: 33%» Sgou 853 .v. ES: 88,. d 3:5— bm? .< gen. 63: .1 B 480.0 .2333: .< 3.3% 32%.an 3.83m 35 gm: do 8 8.55 .3533: .v. new? .< $53 .... 8 :3 .38.. Segue: 8mm: .83 d 33m 5> 3883.53 3&er gas ..a 8 29.3..— oauuu Egan: 55: i a .88 85.. .< 3855 fig: .1 8 3855 some. .< .2333: .< 0555 Base E5: .1 8 :38.— Svt Seasons 88: i 8 380 3:88.. 35%“. 88: .283 a. ease 8:29.33 caesium the: 8.5:. d can saunas“. gauze: g: 83> d 3.6.53— SoBuSo 83.8.35 53v .1 8 8095:. aiqoufiog 3:33.30 A25: .1 .0 883mg g Samba? Ammo: 81: d 33. bu? 335‘? 5:53.. 88: i a 3.5:: 53% 3:883. 35.5.. ... £583. .38 . 33:33 $.85; 88c ... a a? caresses acetates: seabeseeu been? .33 8a»— . 883 8e»: 333...: 358an ...-obs...» 33:53 98:82 A on 8538. can. 83 his... Sate go £58»?— all: 623 ... 38% team .emegSéEoéénuggeoggoZaBo—oadozeh 8...... 3.8.0.. a ......m 8.3.. ... .o 558 8...... .53.. ... 8.2... 8.3.. 55.5 a 3.3. 8...... .a ... 9.5 3.5.. ... .o 8.2 ....> 8...... 303 ... game 8...... .- 3 358 8.8.. 8.....05 Ea: ... .... 80...: 5...... ... a 3.5m 8.53 ... .o 9.5.: $8.. .a u .25.... £5: .... .9 ...... Ex... ... ... 8....o .82. .- .o 38. 8...... .. a .3505 ...... 3%.... 8...... a as. Ea: .. u .53: .88.. 8...... a as. .82. .. ... g: 8.3.. .- ... 955...... 8...... ......m ... .23.; 8.5: 3.2.... ... 8.8.. 8...... 32.5. d .859... 88.. ... .o .35.. 8...... .... u ..E..o 8...... ... .o :88: 8...... .... .o 8.... ....> 88.. 5.55 .82. ... ... 8389.... .34.... 3.2.2... ...... ..w .. ......a 5.3:... E33 Eat—ask 8.5.339: £3.33... 9...: 8...... 835...... .... 39.3 8.5.530 33.3... gutsuaou .3... .... 3.85.283. U 5.3.95.3 23.93.3330 8.9.31... .893 233% .< sum... .< 3333.. .< ...:auufiaéu .m 3% Rug... .32.». 33.3.3. 85.. .v. .6»... .< £33.. .< gag»: Sfiiuousugnk BEE. 883?. .....euwfiaiu .k 33mg: 5.3.3.3»: 5.35:3: Sigma: has.» uuuafiggueb 82.55.. 3.2%.... n.a.... 59:33.8 3%.. 55.8.3.8 ...8 .... a £9.35... a»... ...... £9.53. :23}: v... .3583 n 033—. again-35a: $8:qu amusing 88:qu 55: 3.55 G8: 1 .o 6.5.: 6,5: .553 a 5.33: Ex: ... 8 :85 Ex: .33 a x835 $3: 533 a =3 8.8: i u in: 58: £88 $3: $8.. a «8222 Ex: 1 8 3923: Ex: .1 8 .2653? Ex: .1 .o :95. 68: i u 38. 88: i 8 308293: 8 88: .1 8 225 $8: .1 u 823. £815.. a .3 88: .1 8 82o: 88: i u 5...: Ag: .1 8 933 38: d a 55m 83: ..a .0 >280 $8: ... u 8833: _ 38: use d 8:5". Es: 38> d 8.55m 88: 3.23. a. 3.2.. 533.5%. .k 33.3.5 §z§8 953.3 S§§m I .55.. .....385? 5.9... 5 335:3 .u. 3.333.353 awasgk signage .L gags... .L 38h. .2 33. .k 23:928.»: .0 so»? .< 3:335 22% gain $3 3.38 .h gufiah‘u .k g .< a»! .< 33...- .< 383: gases 5%.! 954.55 323 aqua: 9.39:. euuafib Eu 3539.25 5:38: 333$. éfig .m 3338.: utuimu§3 Seaman 6 .383 33985 §.< :8 .M /£§ 9g 2 Q3: 3533. £03.82 .3: 831 3 N95 .3583 m 930,—. awe. 88: 8.2... a 5.880 are .1 8 89829:“: 8 58588.53: 58188.86 8.85.1833 6.5: .83 a 8:5 Ex: 8 8 85 $8: ... 8 8.5.8.. $8: ... 8 5985...: ....» 88: ... 8 ...-.863 68: ...-.3 8 8:5 £85 3.8.. a 5.85 Ex: 8.5. a 8.25 $8: ... 8 8.8... 388: ... 8 83.5 328: ... 8 83.5 328: i 8 82.5 3.8: ... 8 83.5 was: ... 8 83.5 was: .... 8 82.5 8.8: .... 8 .238 .23: .... 8 329...? .28: .- 8 .8895 $8: ... 8 .888: 853.3% .m. 35°... .88: .< 83.3... .2 3.8 ... 803: .< 8.19.5. .3: 88......» 8.35....— .65. 82...». 8.35:... .13 .8852 8...... 60.348 m 033-. 7O .3? ....- ..35.» 83.5. 2.3.8— u. .88: 58.... E5 858: no $3: ... 8 38¢ $8: ..a 8 8:5 $5: .1 3 338m 55: .1 u 8.2% 88: 88: 9:8". a 8.3.5 6:: 1 u :83: 88: ... u «.5: Anna: .1 8 m8— 23 .m 3&3 .< Luna. .< 33.3.. .< 39.3.8 .9 333%.. .< .3334: .< .2533: .< 85!. =8 .m 658.3 .13.. 395.83... .k 333.283 .9 3933: .< $85; sum? .v. 5% .9? .8358 m ...—as ‘71 acid to UMP. Using this protocol, pyrimidine auxotrophs have been readily isolated and used as recipients in transformation of a variety of filamentous fungi including A. flavus (Woloshuk et al., 1989), A. nidulans (Dunne and Oakley, 1988), A. niger (van Hardngsveldt et al., 1987; Goosen et al., 1987). A. oryzae (Mattern et al., 1987), C. graminicola (Rasmussen et al., 1989), P. anserina (Razanamparany and Bégueret, 1986), P. chrysogenum (Dfez et al., 1987), and U. maydis (Kronstad et al., 1989). 2) Mutants defective in nitrate reductase. Mutants in the nitrate reductase struc- tural gene of filamentous fungi [niaD’ in Aspergillus (Cove, 1976a, 1979); nit-3‘ in Neurospora ('I‘omsett and Garrett, 1980)] can be isolated readily by selection for chlorate resistance. Chlorate toxicity arises presumably from its mimicking nitrate’s effect on the control of nitrogen catabolism (Cove, 197 9). Although mutation in several genes could result in chlorate resistance in filamentous fungi (Cove, 1976b), the niaD/nit-3 mutants could be easily distinguished from Other chlorate resistant mutants by a series of simple phenotypic tests (Cove, 1979; Birkett and Rowlands. 1981; Correll et al., 1987; Klittich and Leslie, 1988; Newton and Caten, 1988). Nitrate reductase genes have recently been cloned from A. nidulans (Malardier et al., 1989), N. crassa (Fu and Marzluf, 1987), A. oryzae (Unkles et al., 1989a) as well as A. niger (Unkles et al., 1989b), which should facilitate usage of these genes as selectable markers in transformation of other filamentous fungi. Whitehead et a1. (1989) has successfully transformed a niaD mutant of Penicillin»: with the cloned niaD‘ genes from A. nidulans and A. niger. A niaD mutant of F. oxysporum has also been transformed successfully with the cloned niaD” gene of A. nidulans 72 (Malardier et al., 1989). Furthermore, Unkles et al. (1989a) has successfully expressed the A. oryzae niaD’ gene in A. nidulans, A. niger and P. chrysogenum. The versatility of this transformation system was further demonstrated by Daboussi et al. (1989) who transformed seven filamentous fungi with the A. nidulans niaD‘ gene (Table 5). Thus, the nitrate system can potentially be used as a universal selection system for transformation of nitrate-assimilating fungi. In addition to S-FOA and chlorate, other toxic metabolic analogs are avail- able for selection of mutations in specific nutritional genes, particularly those asso- ciated with amino acid metabolism. For example, in S. cerevisiae, a-amino—adipate can be used to select mutations at the LYSZ’ and LYSS‘ loci (Chattoo et al., 1979) and methyl mercury selects for met2 and metIS mutations (Singh and Sherman, 1974, 1975). In E. coli (Zurawski et al., 1978) and plants (Last and Pink, 1988), tryptophan-requiring mutants could be isolawd by selecting for resistance to 5- methylanthranilic acid. Selection of the corresponding mutants in filamentous fungi with these chemicals may also be feasible. b. Dominant selectable markers Dominant selectable markers (e.g., those encoding resistance to antibiotics) are routinely used in transformation of prokaryotic cells. Any fungal species should be transformable with this type of selectable markers as long as it is sensitive to the applied selective pressure, and that the cloned gene encoding the resistance pheno- type can be expressed in the recipient under selection. Therefore, dominant selec- 73 tion of transformants offers the distinct advantage of not requiring the presence of a particular mutation in the recipient. A number of mutations conferring drug resis- tance have been proven to be suitable for transformation in a variety of filamentous fungi. Some of these dominant resistance markers are also listed in Table 5 (Fincham, 1989). l) Benomyl resistance. Most mutations conferring benomyl [methyl l-(butylcar- bamoyl)-2-benzimidazole carbamate], a fungicide) resistance in fungi are due to an alternation in the B-tubulin structural gene which prevents binding of benomyl to the fl-tubulin molecules (Davidse, 1986; Morris, 1986). A cloned B-tubulin gene from a benomyl-resistant mutant, therefore, should be usable as a dominant selectable marker in transformation experiments. A mutated B—tubulin gene has been cloned from N. crassa (Orbach et al., 1986) and successfully used as a dominant selectable marker in transforming this fungi. Subsequently several fungal species have also been transformed successfully using this marker (Table 5). Fungal species in which a transformation system is being developed using a homologous B-tubulin gene as a dominant selectable marker include A. flavus (Seip et al., 1988, 1989) and Erysiphe graminis (Sherwood and Sommerville, 1988). 2) Hygromycin B resistance. The aminocyclitol antibiotic hygromycin B binds to 708 or 808 ribosomes, inhibiting protein synthesis and growth of prokaryotic or eukaryotic organisms. Hygromycin B is produced by the prokaryote Streptomyces hygroscopicus, which protects its own ribosome formation by synthesizing the enzyme hygromycin B phosphotransferase. This enzyme phosphorylates the drug 74 making it inactive in the host (Pardo et al., 1985). Genes (hygB or hph) encoding hygromycin B phosphotransferase have been cloned from both S. hygroscopius (Malpartida et al., 1983) and E. coli (Rao et al., 1983; Kaster et al., 1983). Resis- tance to hygromycin B has been used as the basis of transformation systems for a number of eukaryotes, including mammalian cells (Santerre et al., 1984), plant cells (Waldron et al., 1985), and S. cerevisiae (Kaster et al., 1984). In filamentous fungi, the hygB gene is the most widely utilized selectable marker in transformation of genetically undeveloped strains which are sensitive to this drug (Table 5). The list will most likely be extended as the hygromycin B resistance marker has been tried on more and more fungal species. It is immrtant to nate that in all these cases, a eukaryotic promoter has been required to obtain adequate levels of expression of the prokaryotic‘ hygB or hph genes. 3) Oligomycin resistance. Oligomycin is a hydrophobic antibiotic which inhibits the subunit 9 oligomer in the membrane-bound (Fo) portion of the mitochondrial ATP synthase complex (Enns and Criddle, 1977). Mutants resistant to this anti- biotic can be readily obtained. In A. nidulans nuclear and extranuclear mutations have been isolated which confer oligomycin resistance (Rowlands and Turner, 1973). Despite a variety of mutant phenotypes, all nuclear resistance mutants mapped to a single locus (oliC) (Rowlands and Turner, 1977) which encoded an altered allele of the subunit 9 structural gene (Sebald and Hoppe, 1981). The nuclear resistance mutants showed incomplete dominance in heterozygous diploids and were hypersen- sitive to triethyltin. The semi-dominant nature of the 011C gene presumably resulted from a mixture of resistant and sensitive subunits in the subunit 9 oligomer. The 75 extranuclear mutation possibly resides in the mitochondrial gene for subunit 6 (Sebald and Hoppe, 1981). A cloned oligomycin-resistance allele (oliC3I) of A. nidulans was shown to be useful as a semi-dominant selectable marker for transformation of this fungus (Ward et al., 1986). However, the A. nidulans oliC31 gene was not able to act as a marker in A. niger (Ward et al., 1988). Oligomycin-resistance alleles isolated from resistant mutants of A. niger (Ward et al., 1988) and Penicillin»: (Bull et al., 1988) have also been used successfully in transformation of these fungi. Particular features of this system include the fact that homologous integration of the plasmid at the 011C locus can be phenOtypically distinguished fi'om non-homologous integra- tion. Transformants having a plasmid integrated at the oliC locus are fully oligomycin resistant or initially semi-resistant but frequently give rise to fully resistant sectors on oligomycin media. Transformants having a plasmid integrated elsewhere in the genome show stable semiresistant phenotype (Ward et al., 1986, 1988). In addition, triethyltin could be used for selection against the oliC allele which may find use in gene replacement experiments. 4) Kanarnyein resistance. The gene encoding the aminoglycoside 3’-phosphotrans- ferase is responsible for resistance to antibiotics neomycin, kanamycin and its analog G-418. The enzyme phosphorylates these antibiotics and renders them inactive. The kanamycin—resistance gene, derived from the bacterial transposon TnS (Jorgensen et al., 1979) or Tn903 (Oka et al., 1981), has been successfully expressed in a slime mold (I-Iirth et al., 1982), in yeasts (Jimenez and Davies, 1980; Webster and Dickson, 1983; Das et al., 1984; Sakai et al., 1984; Sakai and 76 Yamamoto, 1986) and in higher eukaryotes (Colbére-Garapin et al., 1981). Fila- mentous fungi which have been successfully transformed to G-418 resistance are listed in Table 5. In a few cases, the kanamycin-resistance gene could be driven by its native bacterial promoter. However, in the majority of cases, expression of this resistance gene has been placed under control of the promoter fi'om the recipient fungus itself or from the SV 40 virus. 5) Phleomycin resistance. Phleomycin belongs to the metallo-glycopeptide group of antibiodcs of the bleomycin family (Berdy, 1980). Phleomycin and bleomycin are very closely related classes of compounds produced by different strains of Streptomyces verticillus (U mezawa, 1974). Each class of these water-soluble anti- biotics is composed of a mixture of several components differing in the structure of the basic group (Grigg et al., 1985). At low concentrations the phleomycins are effective in killing prokaryotic or eukaryotic organisms. Phleomycin and bleomycin cause scission of DNA in vivo and in vitro (Kross et al., 1982) preferentially at inverted repeat sequences in single stranded DNA (Ueda et al., 1985) and at various unmethylated sites in double stranded DNA (Hertzberg et al., 1985). This reaction is mdiated by oxygen in the presence of iron salts (Pratviel et al., 1986). Two bleomycin resistance genes (ble) have been characterized from Tn5 (Collis and Hall, 1985) and from Staphylococcus aureus (Semon et al., 1987). Driven by the promo- ter from the S. cerevisiae CYC'I+ (iso-l cytochrome C) gene, the ble gene from Tn5 has been used as a dominant selectable marker in transformation of this fungi (Gatignol et al., 1987). Filamentous fungi which have been transformed by the ble 77 gene into phleomycin resistance are listed in Table 5. In each of these cases, expression of the ble gene was driven by a fungal promoter. 6) Utilization of acetarnide. In filamentous fungi. acetamidase (encoded by the amdS‘ gene) converts acetarnide into acetate and ammonium, thereby allowing growth on acetamide as a sole nitrogen or carbon source (Hynes and Davis, 1986). The amdS‘ gene has been cloned from A. nidulans (Hynes et al., 1983) and subse- quently used in development of a transformation system for this fungus (Tilburn et al., 1983). This cloned gene has been proven a valuable heterologous selectable marker for the transformation of several commercially important fungal species (Table 5). Selection depends on poor utilization of acetarnide as a sole nitrogen source by the recipient strain. It therefore has the potential for use with any fungal species which can be grown on nitrogen-limiting defined medium and which has low levels of acetamidase activity. Moreover, in fungal species which are capable of utilizing acetarnide, mutant strains lacking acetamidase activity can be isolated by treatment with fluoro-acetamide, an amide analogue (Hynes, 1979). This compound is converted to the toxic fluoro-acetate by the acetamidase. Mutations preventing or reducing amdS’ expression cause resistance to the analogue. An acetarnide uptake system has not been found in organisms studied to date. Therefore, selection for mutations affecting utilization of acetarnide will most likely isolate mutants altered in the expression of the amdS‘ structural gene (Hynes and Davis, 1986). 7) Sulfonamide. Sulfonamides are structural analogues of para-aminobenzoic acid. They act as competitive growth inhibitors by reducing the amount of synthesis of 78 functional dihydrofolate by dihydropteroate synthetase (DHPS). Recently, a sulfo- namide resistance gene encoding DHPS has been used as a dominant selectable marker in transformation of P. chrysogenum (Carramolino et al., 1989). Expression of the sulfonamide resistance marker in this fungus was made possible by replacing the native bacterial promoter with the trpC’ promoter from P. chrysogenwn. 8) Others. A gene conferring resistance to the herbicide glyphosate [N-(phos- phonomethyl)-glycine] has also been used as a dominant selectable marker in trans- formation of yeast (Kunze et al., 1989). Glyphosate inhibits EPSP synthase, an enzyme which catalyzes the conversion of shikimate-S-phosphate into 5-enol-shiki- mate-B-phosphate. The gene encoding EPSP synthase has been cloned (as part of the pentafunctional arom’ polypeptide) from S. cerevisiae (Larimer et al., 1983) and from E. coli (aroA’) (Duncan and Coggins, 1984). Application of these markers in the selection for transformants of filamentous fungi remains to be explored. c. Visual screening of transformants Expression of transforming genes can be directly visualized in certain cases on plates with or without a visual indicator. 1) Reporter genes. Vectors containing a fusion of the E. coli lacZ‘ (B-galacto- sidase) gene inserted in frame into the coding region of a fungal structural gene allow visual detection of the transformants. Fungal colonies transformed with this construction turn blue on media containing the chromogenic B-galactoside derivative 79 X-Gal (S-bromo-4-chloro-3-indolyl-B-D-galactopyranoside). In filamentous fungi, structural genes of A. nidulans are commonly used for construction of such trans- lational fusions. These include trpC’ (van Gorcom et al., 1985, 1986; Wemars et al., 1987), gpd’ (van Gorcom et al., 1986), and (1de (Davis et al., 1988). In addition to A. nidulans, similar gene fusions have also been shown to function in A. niger (van Gorcom et al., 1986; Davis et al., 1988) and P. chrysogenum (Kolar et al., 1988). The level of expression of the lacZ’ gene under control of the gpd’ gene promoter was approximately tenfold higher than that under control of the an‘ promOter (van Gorcom et al., 1986). The E. coli uidA’ gene, encoding B-glucuronidase, has recently been deve- loped (Roberts et al., 1989) as an alternative reporter gene to the Idol". A chimeric B-glucuronidase gene was created by ligating the A. nidulans gpd' gene (encoding glyceraldehyde 3-phosphate dehydrogenase) promoter to the coding sequence of the E. coli uidA’ gene. Con'ansformation of this vector into A. nidulans, A. niger and a tomato pathogen Fulvia fulva (syn. Cladosporium fulvum) resulted in the expression of B—glucuronidase. Transformants turned blue on plates containing the enzyme substrate X-gluc (5-bromo-4-chloro-3-indolyl glucuronide). A. oryzae was able to express the E. coli uidA+ gene when cotransformed with the homologous niaD* gene (Unkles et al., 1989a). This system is suitable for those fungi (e.g., wood rot fungi and some plant pathogens) in which a gene expression system based on the lacZ‘ gene is not feasible (Roberts et al., 1989). These fungi normally produce abundant endogenous B—galactosidase even in the presence of glucose as the carbon source to repress production of this enzyme. In order to apply this system, the endogenous, B- glucuronidase activity in these fungi should not be detectable. 80 2) Spore color. Genes responsible for the formation of fungal spore color can be used as visual screenable markers. In Aspergillus, the yA* gene encodes a p-diphe- nol oxidase, or laccase, which is needed to convert the yellow pigment intermediate to the mature green pigment of the conidia (Clutterbuck, 1972). yA+ is one of only a few genes selectively activated during asexual reproduction in A. nidulans that has a well defined physiological function in conidiophore development or spore differen- tiation (Glutterbuck, 1977; Timberlake, 1987, Timberlake and Marshall, 1988). A mutant defective in the yA’ locus produced yellow spores, and when successfully transformed with the wild-type yA‘ gene, was capable of producing green spores (Yelton et al., 1985; O’Hara and Timberlake, 1989). Thus, in this particular case the transformants could be easily identified by the change in spore color. 3) Pigment production. The imperfect filamentous fungus Scytalidium flavo- brunneum produces a potent antifungal metabolite, 15-aza-sterol, and a reddish brown pigment at the end of the exponential phase of growth (Rodriguez and Parks, 1980). A plasmid (pSFB-l) was found (Caprioglio and Parks, 1988) in the wild- type strain which conferred the ability to synthesize both the pigment and the aza- sterol simultaneously when transformed into a mutant strain lacking this plasmid (Caprioglio and Parks, 1989). In this case, successful transformants were identified visually by production of the reddish brown pigment that is characteristic of the plasmid-containing wild type. 81 4) Morphological changes. Changes in size, shape and other morphological fea- tures resulting from genetic transformation represent another type of visual selection. For example, in the ascomycete P. anserina, senescence (vegetative death) through premature strain aging is under nucleo-cytoplasmic control and is inducible in juvenile mycelia by a movable, circular mitochondrial plasmid DNA (plDNA or at senDNA), which originates from an intron of the subunit 1 gene for the mitochon- drial cytochrome oxidase and is highly amplified in aging mycelia (Kilck, 1989). Juvenile protoplasts could be transformed into senescence by using purified plDNA (Tudzynski and Esser, 1980). Unequivocal selection of aging transformants was made possible by using a long-living mutant for which spontaneous occurrence of aging was not observed (Tudzynski et al., 1982). 2. Cotransformation Cotransformation is a technique in which a transforming gene that cannot be directly selected is assimilated along with a more readily selectable marker. It appears that when recipient cells are simultaneously exposed to different transform- ing DNAs, the most competent cells tend to take up more than one type of DNA molecules (Fincham, 1989). The efficiency of cotransformation depends somewhat on the fungal species, vectors and selectable markers used in transformation. In the majority of cases, it has been proven to be a high-efficiency process in filamentous fungi. For example, A. niger was transformed with two plasmids containing, respectively, the amdS‘ and the aciA+ genes of A. nidulans. The seven transformants selected for acetarnide 82 utilization also contained aciA’ sequences derived from the unselected plasmid (Kelly and Hynes, 1985). In A. nidulans, Wernars et al. (1987) has shown that approximately 85% of the AmdS+ transformants, obtained with the cloned amdS’ gene and the trpC-lacZ hybird gene on separate vectors, exhibited expression of a functional B-galactosidase fusion protein. This percentage of B—galactosidase-expre— ssing transformants was as high as that obtained with both genes on one composite vector. About 80% and 60% of the hygromycin B resistant transformants of A. niger and A. nidulans, respectively, were cotransformed with the second marker (amdS’ or pyrG’) (Punt et al., 1987). When P. chrysogenum was cotransformed with amdS’ and the gpd-lacZ hybrid gene, 90% of the AmdS+ transformants formed blue colonies when transferred to XGal plates (Kolar et al., 1988). In Trichodemta, 86% of the Arg’ transformants were also amdS’ when cotransformed with argB+ and amdS‘ of A. nidulans (Penttila et al., 1987). Up to 47.5% of the Trp’ transformants were also acu’ when a double mutant of Coprinus cinereus was transformed with a mixture of two plasmids carrying, respectively, tip-1* and the putative acre-7* gene (Mellon et al., 1987). Cotransformation of A. nidulans, A. niger and F. fulva (Roberts et al., 1989) using selected and unselected DNAs resulted in efficiencies varying from 20% with a phleomycin vector to 44% with the B—glucuronidase vec- tor. When transformed with a mixture of plasmids carrying the hygromycin B resis- tance marker and the A. nidulans gpd-E. coli lacZ hybrid gene, 10 of the 13 hygro- mycin B-resistant transformants of A. flcuum produced a blue color in media conta- ining X-Gal (Mullaney et al., 1988). In all these cases, approximately equal con- centrations of the ectransforming DNAs were used. 83 There are also cases in which cotransformation is less efficient. For exam- ple, when A. nidulans was cotransformed with a mixture of pyr—rl+ (carried on a plasmid) and aliC3l (carried on a lambda), selection for pyr-4’ resulted in oliC31 cotransformation at a frequency of only 4% (Turner and Ballance, 1985). A pyrG mutant of A. niger was transformed with a 1:1 mixture of plasmids containing the A. niger pyrG+ and the A. nidulans gpd-E. coli lacZ fusion. About 10% of the Pyr+ transformants showed a blue color on X-gal medium (van Hartingsveldt et al., 1987). Cotransformation of an A. oryzae niaD mutant with the native pyrG‘, the E. coli uidA’, as well as a mutant allele of the N. crassa tub-2+ gene (conferring benomyl resistance) resulted in cotransformation frequencies of only 4, 9, and 6%, respeCtively (Unkles et al., 1989). It appears that the maximum cotransformation frequency which can be achieved depends on the nature of the cotransforming DNA and the order of selection for marker genes. For example, in one cotransformation experiment conducted by Wernars et al. (1987), an amdS, 012C double mutant of A. nidulans was transformed with a mixture of both plasmids containing the corres- ponding wild-type genes, and either AmdS’ or Trp’ transformants were selected. When the AmdS' transformants were selecwd first, 95% of them were also 0720. On the other hand, if Trp’ transformants were selected first, only 50-60% of the colonies also possessed the AmdS’ phenotypes. Catransformation is a useful technique for testing expression of unselected DNA sequences. It avoids the need to make technically difficult and often complex plasmid constructs containing more than one gene. Genes encoding the isocitrate lyase of A. nidulans (acuD‘) (Ballance and Turner, 1986) and C. cinereus (acct-7*) (Mellon et al., 1987) as well as acetate utilization genes of N. crassa (Thomas et 34 al., 1988) and A. nidulans (Sandeman and Hynes, 1989; Katz and Hynes, 1989c) have been cloned and tested for their functions by this technique. A test for the heterologous expression of the C. cinereus acu-7’ gene in an acuD mutant of A. nidulans was also made possible by cotransformation (Hynes, 1989). It is important to note that the unselected gene could be carried in a lambda vector, although the cotransformation was less efficient in this case (Turner and Ballance, 1985; Mellon et al., 1987). Thus, genes carried on lambda or cosmid vectors could be tested directly for function without the need to subclone them (Timberlake et al., 1985). C. Fates of Transforming DNA Without a fungal replication origin, transforming DNA is frequently inte- grated into the host chromosome. Alternatively, transforming DNA can replicate autonomously in the recipient species. 1. Integration into chromosomes The data presently available (Fincham, 1989; Timberlake and Marshall, 1989; Rambosek and Leach, 1987; Hynes, 1986) clearly indicate that incorporation of the transforming DNA into the genome of the recipient cells is the most frequent events in transformation of filamentous fungi. 85 a. Modes of integration. Three types of integration (Figure 7), originally proposed by Hinnen et al. (1978) for transformation of S. cerevisiae, also apply to filamen- tous fungi. The three types of integration are thought to differ in the nature of the recombination event(s) leading to integration. The frequency with which the various types of integration events occur varies with the plasmid and the recipient strain or ‘ species used. 1) Type I (Homologous integration). Transformants of this type result from a single cross-over between cloned and chromosomal fungal sequences resulting in the integration of vector sequences, which are flanked by the cloned (transforming) and the chromosomal fungal sequences. Homologous integration is a common event in many fungal transformation systems (Case et al., 1979; Tilburn et al., 1983; Yelton et al., 1984, Upshall, 1986). Several copies of transforming DNA have been detected in transformants in A. nidulans (Kelly and Hynes, 1985). 2) Type II (Nonhomologous integration). Transformants of this type arise from integration at sites other than the resident chromosomal locus. It occurs frequently in heterologous transformation systems where there is no detectable homology bet- ween introduced sequences and the recipient genome. For example, the N. crassa pyr-4* gene has low homology with A. nidulans genome, however, integrative trans- formants were readily obtained (Ballance et al., 1983; Ballance and Turner, 1985). Integration at nonhomologous sites occurs at a reasonable frequency even where a homologous sequence exists (Case et al., 1979; Tilburn et al., 1983; Yelton et al., 1984). 86 3) Type 111 (Gene conversion or replacement). Transformants of this type are attributed to either gene conversion or a double cross-over event in which the cloned fungal sequences replace their chromosomal counterparts with no integration of the vector sequences (Case et al., 1979; Yelton et al., 1984). Apparent gene conversion events have been observed between oliC'l and oliC genes located in different parts of the A. nidulans genome (Ward et al., 1986). b. Analysis of integration events. Traditional genetic linkage analysis as well as modern molecular probing techniques are usually employed to distinguish among the three integration events described above. Whether or not the integrated transforming DNA is linked to the homologous chromosomal locus can be determined by analyz- ing progenies of a cross between the transformant and the wild type. Absence or extremely low frequency of the untransformed phenotype among the products of the cross indicates close linkage of the integrated DNA to the resident locus. An unlinked transformant is expected to yield 25% of progeny with the untransformed phenotype from such a cross (Case et al., 1979; Kinsey and Rambosek, 1984; Brygoo and Debuchy, 1985; Dhawale and Marzluf, 1985; Wernars et al., 1985; Picard et al., 1987; Kim and Marzluf, 1988). More conclusive results can be obtained by Southern hybridization analysis of restriction fragments of genomic DNA from transformants. Molecular analysis of transformants has led to the identification of all three types of integration events in virtually all filamentous fungi investigated (Kim and Marzluf, 1988; Grant et al., 1984; de Graaff et al., 1988; Faugeron et al., 1989; Tilburn et al., 1983; Binninger 87 Figure 7. Three patterns of integration of transforming DNA into the fungal genome (adapted from Peberdy, 1989). Type 1: integration at a homologous locus by a single recombination event. Type II: integration at random sites of weaker homology by a single recombination event. Type III: integration, at a homologous locus involving a double recombination resulting in a gene conversion. DNA sequences: solid bars, fungal genes used for selection; striped bars, flanking regions of fungal DNA; open bars, bacterial plasmid DNA. 88 es xv. . .4. ... n. ...... 0...! . $14.... and :3, have ... bml ...... 6.5.... ... 25 = 25 _ on»... 89 et al., 1987; Mellon et al., 1987; Alic et al., 1989; Wang et al., 1989; Bull et al., 1988; Van Hartingsveldt et al., 1987). However, the distribution of these three events varies from species to species. There is not sufficient data to accurately estimate relative frequencies of the three types of integration events in filamentous fungi. The mechanism of integration remains virtually unknown (Razanamparany and Bégueret, 1988; Fincham, 1989). There are several examples of integration of transforming DNA sequences in tandemly repeated c0pies in filamentous fungi (Yelton et al., 1984; Wernars et al., 1985, 1987; Goyon and Faugeron, 1989). c. Stability of integrated sequences. In general, integrated transforming DNA sequences are mitotically stable (Boylan et al., 1986). This high degree of mitotic stability is most desirable for genetic engineering of industrial strains. In contrast, integrated DNA sequences are often meiotically unstable (Tilburn et al., 1983; Yelton et al., 1985). In N. crassa (Selker et al., 1987) and A. immersus (Goyon and Faugeron, 1989; Faugeron et al., 1989), integrated DNA sequences are often eliminated at a high frequency during the sexual phase by a process called repeat- induced point mutation (RIP) (see below). 2. Autonomous replication Transformation vectors capable of autonomous replication have been reported in several filamentous fungi including Mucor (van Heeswijck and Roncero, 1984; van Heeswijck, 1986), Phycomyces (Revuelta and Jayaram, 1986; Suarez and Eslava, 90 1988), Absidia (Wbstemeyer et al., 1987), Podospora (Tudzynski et al., 1980; Stahl et al., 1982), Ustilago (T sukuda et al., 1988) and Neurospora (Stohl and Lambowitz, 1983; Stohl et al., 1984; Grant et al., 1984). There is also limited evidence to suggest their existence in A. nidulans (Johnstone, 1985), A. niger (Rambosek and Leach, 1987) and the koji mold A. oryzae (Iimura et al., 1987). a. Features of autonomously replicating plasmids. In general, autonomously rep- licating plasmids (ARPs) can be distinguished from integrating vectors by the following criteria (Rambosek and Leach, 1987): L1) ARPs normally give a high frequency of transformation; (g) Transformants containing ARP are mitotically and meiotically unstable under nonselective growth conditions; (3) DNA isolated from transformants with ARP readily transforms E. coli. However, genomic DNA from integrated transformants doesn’t; (A) Southern analysis of undigested DNA from transformants with ARPs reveals the presence of monomer-sized ARPs, while the integrated plasmid DNA comigrates with undigested chromosomal DNA in a similar analysis. b. Origins of replication. In filamentous fungi, the origin that allows autonomous DNA replication is generally derived from mitochondrial plasmids or chromosomal sequences capable of autonomous replication in yeast cells. I) Mitochondrial plasmids. Although nuclear plasmids are not known in filamen- tous fungi, several mitochondrial plasmids have been discovered in various species (Tudzynski and Esser, 1985; Garber et al., 1986). 91 It is interesting to note that a restriction DNA fragment containing the terminal inverted repeat of a linear plasmid isolated from the mitochondria of N. haematococca (anamorph F. solam) (Samac and Leong, 1988) conferred autonomous replication when incorporated into an integrative transformation vector of U. maydis (Samac and Leong, 1989). Whether this plasmid DNA fragment will also support autonomous replication of vectors in other fungi remains to be investigated. 2) Autonomous replication sequences in filamentous fungi. Autonomous replica- tion sequences (ARS s) refer to DNA sequences isolated from other organisms that promote autonomous replication in yeast. They often do not represent authentic origins of DNA replication in the homologous system. ARSs were first identified as sequences linked to genetic markers that transformed yeast cells at high frequencies (Hsiao and Carbon, 1979; Struhl et al., 1979). Plasmid DNA lacking an ARS can u'ansform yeast cells only by rare integrative recombination into the cellular genome (Hinnen et al., 197 8). In contrast, the transformation frequency may be 10’ to 10‘ times higher when the plasmid carrys an ARS. This differential transformation frequency has been useful for defining ARS sequences operationally (Stinchcomb et al., 1980). Putative ARSs have been isolated from a variety of filamentous fungi, including U. maydis (Banks, 1983b), C. acremonium (Skatrud and Queener, 1984), N. crassa (Suzci and Radford, 1983; Buxton and Radford, 1984), Phycomyces (Revuelta and Jayaram, 1986), P. anserina (Lazdins and Cummings, 1982), and C. heterostrophus (Yoder and Turgeon, 1985). ARS sequences exhibit only a limited function across species lines of fila- mentous fungi. For example, the A. nidulans ans] sequence identified by Ballance 92 and Turner (1985) is capable of increasing the transformation frequencies of vectors containing the N. crassa pyr-4‘, and the A. nidulans argB* as well as trpC’ genes (Cullen et al., 1987). The efficiency of P. chrysogenum transformation was also improved by employing ansI-bearing plasmids (Cantoral et al., 1987). However, vector constructs carrying the am] sequence and the A. niger niaD+ gene did not show increased transformation frequencies (Whitehead et al., 1989). Genetic mapping and DNA sequencing data (Cullen et al., 1987) suggested that ansI may be related to the centromere sequence. Whether this is the basis of the effect of ans] on the efficiency of transformation needs further investigation. An A. nidulans genomic DNA sequence of unknown biological function (de- signated um) has been cloned by Johnstone (1985). Like ans], an: is reiterated several times in the Aspergillus genome, and is capable of enhancing the frequency of transformation, but its mechanism of activity appears to be different. Aspergillus, transformed with a vector containing nut, is very unstable and usually contains considerable amounts of free circular plasmid molecules as well as multiple inte- grated copies (Johnstone and Clutterbuck, 1986). The nature of this sequence remains unknown to date. Despite failures in the search for an ARP from the Ascomycetes, evidence for ARP was obtained from the Zygomycetes and Basidiomycetes. In the Zygomycetes such as Mucor (van Heeswijck and Roncero, 1984; van Heeswijck, 1986), Phyco- myces (Suarez and Eslava, 1988; Revuelta and Jayaram, 1986), and Absidia (Wostemeyer et al., 1987), Southern analysis has shown the presence of plasmids capable of replicating indefinitely in the transformants under selective conditions. 93 An ARS sequence was presumably responsible for the autonomous replication in the cases of Phycomyces and Mucor. Recently, a more detailed study on ARS elements in a basidiomycete, U. maydis, was reported by Tsukuda et al. (1988). A selection scheme similar to that used in yeast which exploited the high transformation frequency of ARP was adopted to isolate°DNA segments from U. maydis capable of autonomous replica- tion. DNA fragments 2-10 kb in size were cloned into an integrative transforming vector (pl-1L1) containing the hygromycin B resistance marker fused to heat shock gene promoter of Ustilago. DNA enriched in plasmid molecules was extracted fi'om the pooled hygromycin-resistant transformants and used to transform competent E. coli cells. A high percentage of plasmids recovered from E. coli transformants was found to contain U. maydis DNA inserts which increased the frequency of U. maydis transformation several-thousandfold. Evidence of transmission of these ARS- containing plasmids as extrachromosomal elements through replication was obtained by monitoring the sensitivity of methylated plasmid DNA to restriction endonuclease digestion after transformation. ARS-containing plasmid prepared from the dam’ E. coli strain DHS, which methylates adenine in the sequence GATC, was cut by Dpnl but not by its isoschizomer Mbol. However, after transformation of U. maydis and 20 generations of growth under antibiotic selection, plasmid DNA extracted from the transformants had become resistant to digestion by Dpnl but sensitive to MboI, indicating a loss of methylation. This loss in the pattern of DNA methy- lation is in agreement with a mode of transmission in which the unmethylated sites arise from new synthesis of plasmid DNA during autonomous replication. These plasmids were maintained at a level of 25 copies per cell but were mitotically 94 unstable as expected. One 1.7-kb ARS (designated UARSl) could be reduced fur- ther into three separate fragments, each of which retained ARS activity. The smallest fragment was 383 base pairs long, which, although not active itself in yeast, contained seven 8-bp direct repeats, two contiguous 30-bp direct repeats, and five ll-bp units in both orientations with sequences similar but not identical to the consensus sequence found to be crucial for ARS activity in S. cerevisiae. Whether any of these sequences plays a functional role in conferring autonomous replication activity remains to be determined. c. ARP and abortive transformants. A common observation in fungal transforma- tion systems is the presence of "abortive" transformants (Buxton and Radford, 1984; Kinnaird et al., 1982), that is, colonies that initially grow on selective media but not after subculturing. One theory suggests that in abortive colonies the transforming DNA has not been stabilized by integration or by autonomous replication and is only transiently expressed. The transformant grows initially, but as the gene product that relieves the selective pressure is diluted out, growth is halted. In support of this theory, Tilburn et a1. (1983) found that transforming DNA is present in abortive transformants after cessation of growth and noted that sectors of vigorous growth could develop from abortive colonies. Southern analysis was used to show that these sectors contained transforming DNA that was stabilized by integration into the recipient genome. In summary, autonomously replicating plasmids have been reported but have net been widely used as molecular cloning vectors in filamentous fungi. Construc- 95 tion of vectors lacking genomic homology is not useful in preventing integration since transforrrring plasmids readily integrate at either homologous or heterologous sites regardless of the homology (Tilburn et al., 1983; Russell et al., 1989). The development of mutant strains completely defective in integrative recombination represents a feasible approach. On the Other hand, centromeric sequences may be used to develop autonomously replicating vectors with increased mitotic stability during segregation. However, such sequences are not yet available from filamentous fungi. The cloned S. cerevisiae centromeres provide an alternative source, but they do not display detectable stabilizing activity in A. nidulans (Boylan et al., 1986). Telomeric sequences from T etrahymena thermophila are able to maintain an integra- tive linear plasmid of P. anserina extrachromosomally (Perrot et al., 1987). A telomere DNA has been isolated from N. crassa (Schechtrnan, 1987). Vectors cons- tructed based on this telomeric sequence may allow development of autonomous replicating linear plasmids in filamentous fungi. D. Recovery of Transforming DNA Transformation is frequently used to directly clone genes by selection for function. The ability to recover the cloned sequences from the transformants is one of the most desirable features of a transformation system which has led to efforts to isolate the autonomously replicating shuttle vectors as discussed above. However, the frequencies of integrative fungal transformation can be sufficiently high for direct cloning and the integrated transforming DNA can be recovered in several ways. 96 1. Recovery of plasmids Although transforming plasmids frequently integrated into the host chromo- some, they can be recovered intact by direct transformation of E. coli with undi- gested genomic DNA from transformants (Stohl and Lambowitz, 1983; Ballance and Turner, 1985; Johnstone et al., 1985). It appears that excision of an integrated plasmid can occur, resulting in free plasmid molecules in transformants (Johnstone et al., 1985; Diallinas and Scazzocchio, 1989). Rearrangement or partial deletion of transforming plasmid DNA can occur during the recovery process (Paietta and Marzluf, 1985; Barnes and MacDonald, 1986). However, they often still carry the selectable marker. Alternatively, an integrated transforming DNA sequence can be recovered by cleaving the transformant genomic DNA with a restriction enzyme that cuts once within the transforming sequence. The resulting DNA fragments are circularized and used to transform E. coli (Yelton et al., 1984; Ballance and Turner, 1985). For example, the genes encoding isocitrate lyase (Turner and Ballance, 1985) and orotidine 5’-phosphate decarboxylase (Oakley et al., 1987) have been cloned in this way. 2. Recovery of cosmids Cosmids are plasmids with bacteriophage lambda cos (packaging) sequences. The genomic DNA isolated from transformants carrying cosmid vectors can be sub- 97 jected to an in vitro lambda packaging system to recover an intact copy of the transforming DNA. This cosmid rescue procedure has been successfully used to clone yA’ (encoding laccase) and other developmentally regulated Aspergillus genes (Yelton et al., 1985). 3. Sib selection This strategy is originally developed by Akins and Lambowitz (1985) for N. crassa, and represents a very efficient procedure for marker rescue especially when coupled with cosmid genomic banks (Vollmer and Yanofsky, 1986). In this appr- oach, a transforming marker is recovered by successive rounds of transformation of - a suitable recipient strain, using pools of DNAs from a genomic library which are progressively reduced in complexity until a single candidate clone is identified. A similar procedure was used to identify plasmid clones containing the A. nidulans pyrG’ gene (Oakley et al., 1987), and to identify the mating-type b1 allele from a cosmid library of U. maydis (Kronstad and Leong, 1989). E. Applications of Transformation Transformation represents a new approach to altering the biological charac- teristics of filamentous fungi. It has been used extensively in studying control mechanisms for growth, metabolism, and development as well as for isolation and manipulation of genes of patential application in medicine, industry and agriculture. 98 1. Cloning genes by complementation Techniques used for isolating genes from other organisms have been used in filamentous fungi with varying degrees of success. Basic cloning strategies such as differential hybridization (Hynes et al., 1983), hybridization with synthetic DNA probes (Kinnaird et al., 1982), and cDNAs prepared from abundant messenger RNAs (Nunberg et al., 1984) have been employed successfully. Cloning genes by functional complementation of mutation can be heterolo- gous or homologous in nature. In heterologous systems, appropriate mutant strains of E. coli, yeast or Other fungal species than the recipient strain are often used as recipients, whereas for homologous systems mutants of filamentous fungi from which the desired wild-type gene will be isolated from is the recipient. a. Heterologous complementation. Several fungal genes, for example, the N. crassa qa-Z+ gene (Vapnek et al., 1977) and the A. nidulans trpC’ gene (Yelton et al., 1984) have been successfully cloned by complementation of E. coli auxotrophic mutants. However, this approach is not generally applicable since regulation and GXpression signals of fungal genes may not be recognized in E. coli. Genes isolated by complementation of E. coli mutants are presumed not to have introns in the functional domain. Yeast cells, in general, are not suitable hosts for heterologous complemen- tation since they can not efficiently process introns of genes from filamentous fungi (Petunia et al., 1984; McKnight et al., 1935; Woudt et al., 1985; Innis et al., 1985). I‘IOWever, cDNA libraries constructed on a yeast promoter have been successfully 99 used to isolate several genes from filamentous fungi (McKnight et al., 1985; Kronstad et al., 1989). The close relatedness of certain fungi to the well-characterized perfect fungus A. nidulans has made cloning genes by complementation of the latter feasible, in particular for those genes that are reasonably conserved through evolution. For example, the A. niger argB‘ gene (encoding omithine carbamoyl transferase) has been cloned by transforming an argB mutant of A. nidulans with a genomic DNA library of A. niger (Buxton et al., 1987). Alternatively, a gene can be cloned from one species by detecting its expression in a different species. For example, a gene encoding pisatin demethylase from the plant pathogen N. haematacocca was cloned by screening its enzymatic activity in A. nidulans transformed with DNAs from a cosmid library (Weltring et al., 1988). b. Homologous complementation. Virtually any gene can be cloned by homolog- ous complementation providing that an appropriate selection protocol and a mutant recipient with a low reversion frequency are available. The rapid advancement in molecular genetics of A. nidulans and N. crassa has allowed a number of structural genes to be cloned (Rambosek and Leach, 1987; Schwab, 1988). In addition, a strategy to cloning promOter sequences has been developed in Cochliobolus (Turgeon et al., 1987). This technique should be of general applicability for cloning regulatory sequences in other less well characterized f urr gal species. 100 2. Gene disruption/replacement Gene disruption or replacement refers to a procedure whereby an altered cloned gene is used to disrupt or replace its chromosomal counterpart by homo- logous recombination after transformation. Insertion of plasmids at specific chromosomal sites is important for analysis of mutations generated in vitro and for making targeted in vivo mutations. a. Approaches to gene disruption Several transformation techniques are available for selection of plasmid integration events at specific sites (Fincham, 1989; Leong, 1988) (Figure 8). 1) Transformation with a truncated cloned gene. In this approach (Figure 8A) a homologous recombination by a single crossover between the transforming plasmid , which carries an internal fragment of the target gene, and the resident locus of the recipient chromosome results in two incomplete genes, one lacking sequences at the 5 ’ end of the gene and the other lacking sequences at the 3’ end. For example, mutants of the A. nidulans yA+ gene produce yellow instead of the normal green sPores (Pontecorvo, 1953). Homologous integration with transforming plasmids car- 1‘.)’ing a truncated yA+ gene results in formation of readily selectable yellow-spored mumts (O’Hara and Timberlake, 1989). 101 2) One-step gene replacement. In this approach (Figure SE) a plasrrrid (linear or circular) containing a selectable marker inserted into (and thus disrupting) the cloned gene of interest is used to transform a mutant defective in the selectable gene. Selection is made for the disrupting marker. Type III integration will result in replacement of the target gene by the nonfunctional disrupted allele. Undesirable transformants arising from type I integration of the disrupting marker into the chromosomal locus can be largely avoided by using a deletion mutant as the recipient. Alternatively, transformants derived from type III integration should contain no vector DNA sequences, and thus can be distinguished from those derived from type I event by Southern analysis of the chromosomal DNA. Gene disruption seems to be a low-frequency event in A. nidulans and N. crassa (Paietta and Marzluf, 1985; Miller et al., 1985). However, it may occur at a higher frequency in U. maydis. To disrupt the genomic copy of the U. maydis pyr6’ gene, Kronstad et al. (1989) inserted the hygromycin B resistance marker into a DNA fragment containing the cloned pyr6‘. The resulting linear construct was used to transform U. maydis to hygromycin B resistance. Approximate 70% of the transformants required uracil for growth. Southern blot analysis revealed that allthese transformants resulted from direct gene replacement by homologous recom- bination. Similar results were also obtained in a gene replacement experiment using a DNA fragment carrying an allele of the mating-type b locus of U. maydis (Kronstad and Leong, 1989). By inserting the hygromycin B resistance marker into the cloned leu‘ gene, Fotheringham and Holloman (1989) have shown that the one- step disruption procedure in U. maydis could be as effective as in yeast. Among 102 Figure 8. Examples of gene disruption/replacement (modified fi'om Fincham, 1989). (A) Transformation with a truncated cloned gene. Integration of the transforming plasmid carrying a deleted copy of the yA“ gene by homologous recombination splits the chromosomal yA+ gene into two nonfunctional pieces. (B) One-step gene disruption (Miller et al., 1985). The transforming plasmid carries a copy of argB’ disrupted by the selectable marker trpC‘. Type III integration of the th‘ selectable marker at the chromosomal argB“ locus results in its disruption. (C) Two-step gene replacement. A plasmid containing a partially deleted spoCI-C‘ gene of the A. nidulans spoCI’ gene cluster (thicker lines with open arrows) and the trpC" gene is used to ‘transform a an mutant (Miller et al., 1985). A Trp” transformant contain- ing a tandem duplication of the spoCI-C’ region is self-fertilized. TrpC mutants are obtained which have lost trpC’ but retained the partially deleted spoCI -C* gene by intrachromosomal recombination. (D) Gene replacement by cotransformation (Wernars et al., 1987). Two plasmids, one carrying the A. nidulans trpC/lacZ fusion gene and one carrying amdS’, are used in co- transformation. LacZ‘ gene activity is detected among a large number of AmdS+ transformants screened. 103 .Ilf' w 4. argB Transform + | Transform 1 Selection '0! 1’PC 3' 7 Selection tor apC+ 5' f——‘ M #’f W Disrupted )4 90M Disrupted 038 (me ‘ .sX —-— 0006 car transform - selection for "DC ' Ce-trenstermetron at ”43' mutant - selection tor and? a." + l O IODMCOMOM [’11: ’1. {”11 g I L 1"“. i I lipC/ldtz "43‘ 104 hygromycin B resistant transformants, one quarter to one half were simultaneously Leu'. 3) Two-step gene replacement. In this approach (Figure 8C) type I integrative transformants are isolated which contain tandem c0pies of the target gene under investigation and the modified replacement version. Then segregants are identified which .have lost sequences associated with the selectable marker through differential double reciprocal recombinations, resulting in restoration of the native gene or its replacement. In general, these two recombination events can be distinguished by Southern hybridization analysis. Alternatively, they can be readily differentiated without resort to Southern analysis in cases where a simple assay is available to detect the replacement gene. For example, a simple color assay for B—galactosidase was used to detect meiotic progenies of A. nidulans expressing the amdS-lacZ fusion which replaced the resident amdS’ gene during a two-step gene replacement experi- ment (Davis et al., 1988). Selectable markers which can be selected (positively selective) and counter- selected (negatively selective) are extremely useful in gene replacement experiments. For example, in filamentous fungi, mutations resulting in loss of orotidine 5’-phos- phate decarboxylase (encoded by pyrG’, pyr-4’ or pyr6’, depending on fungal spec- ies) confer resistance to the normally inhibitory analog 5-FOA (Diez et al., 1987; Dunne and Oakley, 1988; Kronstad et al., 1989). This two-way selection has been used in A. nidulans for a two-step gene replacement of a mutant benA allele ([3- tubulin) with the wild-type allele (Dunne and Oakley, 1988). A benomyl-resistant (benA), pyrimidine-auxotrophic (pyrG) strain of A. nidulans was transformed with a 105 plasmid carrying the wild-type benA allele and the pyr-4+ gene Of N. crassa. The pyr-4* gene complemented pyrG but had insufficient homology to direct integration at the pyrG’ locus. Consequently, the majority Of the primary transformants carried a duplication Of benA alleles (one wild type and one benomyl resistant) interrupted by plasmid sequences due to integration at the benA locus. Such transformants (Pyr', Ben‘) showed intermediate degree Of benomyl resistance and were not able to grow in the presence Of high levels Of benomyl. The cells from the transformant colony were then incubated on medium containing 5-FOA which killed cells carry- ing a functional pyr-4‘ gene and thus selected for cells that had lost the functional pyr-4+ by reciprocal recombination. Of the twenty-two such Pyr’ colonies isolated, 7 were benomyl sensitive (Ben') and 15 were Ben'. Southern hybridization Of DNA from 3 Of the Ben', Pyr colonies revealed that the plasmid sequences had been lost and only a single c0py of benA was present in each case, demonstrating the replace- ment Of the resident mutant benA allele by the transforming wild-type allele. Gene replacement can also be achieved in transformants Of A. immersus by a two-step procedure (Goyon and Faugeron, 1989) similar to that described for A. nidulans and Other filamentous fungi. 4) Gene replacement by cotransformation. Cotransformation can be used in gene replacement (Figure 8D). For example, about 75% Of the A. nidulans AmdS‘ trans- formants express ligalactosidase activity when cotransformed with the homologous amdS‘ and the a'pC-lacZ hybrid gene. Some Of these transformants have an‘ replaced by the trpC-lacZ fusion (Wernars et al., 1987). 106 In summary, direct and indirect gene disruption and replacement have been demonstrated in several filamentous fungi. The efficiency with which these events occur is sufficiently high that the desired transformants can readily be found among the background of nonspecific integrations. These powerful techniques can be used (1) to return an in vitro-altered gene to its resident site, and thus create a mutant which could Otherwise difficult to Obtain (Wernars et al., 1987); (i_i) to prove the identity Of newly cloned genes (Osmani et al., 1988a; Boylan et al., 1987; Turner et al., 1985), and map their chromosomal location (May et al., 1985); ('3') to demo- nstrate the physiological functions Of cloned genes (Aramayo et al., 1989), and Q!) to analyze regulatory genetic elements (Frederick et al., 1989). b. The RIP effect: a unique gene disruption Any cloned N. crassa gene can be potentially disrupted by the premeiotic disruption procedure discovered by Selker et al. (1987). They found that crossing a strain containing duplicated DNA sequences generated by transformation with any Other strain resulted in heavy methylations and numerous nucleotide sequence changes in both duplicated COpies in part Of the meiotic products (ascospores). The effect presumably will destroy the function Of any gene present within the dupli- cated region. The phenomenon was initially called the RIP (rearrangement induced premeiotically) effect because it occurs immediately before karyogamy and meiosis but later renamed to "repeat-induced point mutation" because the changes were found usually within the duplicated sequences without rearrangement Of their positions in the genome (Fincham et al., 1989; Cambareri et al., 1989). A detailed 107 characterization Of this phenomenon by DNA sequencing and heteroduplex analyses (Cambareri et al., 1989) indicated that the process produces an extremely high frequency Of G-C to AT transition point mutations (of the order Of 50% Of G-C base pairs may be affected). The changes occur principally at sites where adenine is 3’ Of the changed cytosine. The mechanism and function Of the RIP effect remain unclear. The RIP effect also exists in Ascobolus (Goyon and Faugeron, 1989). As pointed out by Fincham (1988, 1989), the RIP effect probably accounts for L1) a reported case Of two-copy lethality in P. anserina (Debuchy et al., 1988); (g) the fact that many transformants fail to transmit the transformed character through crosses in A. nidulans (Tilburn et al., 1983; Yelton et al., 1984), P. anserina (Picard et al., 1987; Razanamparany and Begueret, 1986), as well as in N. crassa (SzabO and Schablik, 1982), and (3) the Observation Of Case (1986) that transforming qa- 2* sequences whose activity had been lost during outcrossing appeared tO remain present (but nonfunctional) in the genome. The RIP effect may explain the meiotic instability Of selectable markers in transformants with closely linked duplicated genes. Unlinked duplicate gene OOpies Of the selectable marker would show less meiotic instability. Furthermore, the RIP effect may play a role in limiting the distribution Of the Tad transposon in strains Of Neurospora. Tire Tad transposable element was detected as a 7-kb insertion in two independently isolated spontaneous forward mutants Of the N. crassa am’ gene (Kinsey and Helber, 1989). DNA hybri- dization analysis (Kinsey, 1989) indicated that Tad was present only in the Neurospora strain Of origin from AdiOpOdume, Ivory Coast; laboratory strains do not contain this element. This has led Kinsey (1989) to suggest that the RIP process 108 might play a role in protecting Neurospora against transposable elements. Tad DNA sequences would become extensively altered and failed to hybridize the cloned Tad probe if the RIP process occurred during repeated crosses. NOt all Ascomycetes with a dikaryotic phase in their life history are subject to the RIP effect. For example, duplications created by transformation in Sordaria macrospora, an ascomycete closely related to N. crassa, are not inactivated during meiosis (Le Chevanton et al., 1989). As the inactivation Of duplicated sequences takes place just before meiosis, it was suggested that such inactivation may act in heterothallic species (e.g., N. crassa) as a regulatory mechanism associated with the coordinated expression Of the two nuclei Of different mating type. Such a mecha- nism would not be required in a homothallic species such as S. macrospora in which the two nuclei are most likely derived from the same parental nucleus. 3. Cloning genes by insertional inactivation As discussed previously, plasmids lacking homology with the host genome frequently integrate at random chromosomal sites in transformation. This feature is similar to a transposable element, and Offers a convenient method for cloning genes by selection for loss Of function due to insertional inactivation (Timberlake and Marshall, 1989). This approach does not require previous isolation Of mutants as does the functional complementation technique. Using this inactivational cloning technique, Diallinas and Scazzocchio (1989) were able to clone the A. nidulans uapA’ gene encoding the uric acid/xanthine permease. Cloning uapA+ by direct complementation is laborious because uapA 109 mutants are leaky, due to the presence Of Other perrneases that transport uric acid into the cell (Arst and Scazzocchio, 1975). In their procedure, a yellow-spored ' strain of A. nidulans carrying a complete deletion Of the amdS‘ gene was trans- formed with a plasmid carrying an intact amdS‘ gene and bearing nO sequences homologous to the genomic DNA of the recipient strain. Selection Of the uapA mutants resulting from inactivational integration at the uapA’ locus was made simple by an Observation by Alderson and Scazzocchio (1967): in the presence Of the xanthine analog 2-thioxanthine, wild-type strains Of A. nidulans produce yellow rather than normal green conidia. This effect requires uptake Of the analog and its subsequent Oxidation to 2-thiouric acid which presumably acts by chelating the copper Of laccase (polyphenol oxidase) which is involved in the conversion Of yellow tO green pigment (Alderson and Scazzocchio, 1967). Mutations at any locus necessary for this process result in resistance to 2-thioxanthine and produce green conidia even in the presence Of 2-thioxanthine. Thus, transformants were selected on acetarnide as nitrogen source in the presence of 2-thioxanthine. Among appro- ximately 12,000 A. nidulans AmdS’ transformants, 34 were 2-thioxanthine-resistant, i.e., showed green conidia. In theory, integration events at nine genes could result in resistance to 2-thioxanthine. However, these possibilities can be easily distinguished by growth and complementation tests (Darlington and Scazzocchio, 1967; Alderson and Scazzocchio, 1967; Scazzocchio and Arst, 1978). Genetic mapping indicated that all 34 green-spored transformants were uapA mutants due to integration Of the amt? gene at the uapA’ locus. A plasmid containing an intact amdS" gene but an incomplete uapA’ gene was recovered by transforming E. coli tO ampicillin resistance with genomic DNA prepared from transformants. A functional 110 copy of the uapA‘ gene was finally isolated from a gene library by using this plasmid as a probe. 4. Gene fusions The E. coli B—galactosidase is easily detected in plates or protein extracts, and therefore its structural gene (lacZ‘) is frequently used for transcriptional or translational fusions tO measure the activity Of a regulatory DNA sequence from filamentous fungi (Davis et al., 1988; Hamer and Timberlake, 1987; van den Hondel et al., 1985, 1986; van Gorcom et al., 1985). Gene replacement can be used to target the fused hybrid gene at specific chromosomal sites (see above) to avoid position effects on gene expression (Miller et al., 1987). Similarly, regulatable promoters can be used to study expression patterns of structural genes. For example, several A. nidulans structural genes have been fused to the A. nidulans ach’ promoter to investigate the functions Of their products (Osmani et al., 1988b; Adams et al., 1988; Mirabito et al., 1989; Waring et al., 1989). The ach’ gene encodes catabolic alcohol dehydrogenase and the promoter is subject to substrate induction and carbon catabolite repression (Gwynne et al., 1987). Expression Of genes fused to this promoter can be manipulated by growth condi- tions. Ill 5. Titration Of regulatory gene products Expression of the A. nidulans amdS‘ gene is controlled by the trans-acting gene products Of areA’ and ade’ under different growth conditions (Hynes and Davis, 1986). Transformants with multiple copies of amdS‘ grow poorly on nine. gen sources due to titration Of the areA’ and ade+ gene products by binding sites upstream Of each Of the multiple copies Of amdS* (Kelly and Hynes, 1987). The activities regulated by ade‘ were depressed upon introduction Of multiple copies Of an amdS‘ upstream DNA fragment, but could be restored by transformation with multiple copies Of ade’ (Andrianopoulos and Hynes, 1988). This titration approach has allowed identification and mapping Of cis-acting regulatory sequences (Hynes et al., 1988) as well as cloning of genes that are controlled by common trans-acting factors (Katz and Hynes, 1989a; Richardson et al., 1989). 6. Biotechnology Filamentous fungi have a long history Of use in fermentations for production Of a variety of industrial enzymes and pharmaceuticals. In general, these organisms are more versatile hosts for the development Of heterologous gene expression systems than Other organisms such as E. coli and S. cerevisiae (van Brunt, 1986; Cullen and Leong, 1986; Upshall, 1986) because many Of them have the ability to properly process and secrete high level Of heterologous proteins Of eukaryotic origin. For example, A. niger has been reported to secrete up to 20 grams per liter Of the 112 glycoprotein amyloglucosidase (van Brunt, 1986). An additional advantage is the ability to generate transformants containing multiple copies Of integrated plasmid sequences of sufficient mitotic stability. With the development Of molecular methods for manipulating filamentous fungi, the exploitation Of their secretion and processing capability to produce heterologous proteins, especially those with therapeutic and commercial value is being actively investigated (Saunders et al., 1989). Results from such attempts are summarized in Table 6. In general, the rapid developments in the molecular biology of A. nidulans make this fungus suitable for initial development Of a secretion system, from which technology may be transferred to the important industrial species such as A. niger, A. awarnori, and A. oryzae. Cotransformation, which is a very efficient procedure in Aspergillus (see previous discussion), is usually used to introduce the desired gene into the expression host. Both consti- tutive (Upshall et al., 1987) and controllable (Gwynne et al., 1987a, 1989; Christensen et al., 1988; Tumbull et al., 1989; Harkki et al., 1989; Cullen et al., 1987) promoter elements have been used in construction Of heterologous expression systems, but the latter is most desirable. Being able to control production Of the desired protein allows one to direct product synthesis at particular times during the fermentation process. Thus, the secreted products are only exposed briefly to the extracellular environment prior to harvest. For example, the expression Of the A. nidulans ach+ is normally repressed by glucose but can be induced in the absence Of it (Sealy-Iewis and Lockington, 1984; Lockington et al., 1985). This allows the construction Of strains which can be rapidly grown in the presence of glucose tO accumulate biomass and which can then be induced to initiate the production (and 433625888893. fineness-8858588888,: .5.m26882§&8§=u82=8=§ gig—.3595. .3385; .5geqzég68m. .%:85 53338.»... 3:83:503888.25Eguueggfiagéguezwbiggggue 9.5.2.9. 88: ..e a as: 3... 91: 88:8 8:8 :38 8.8. .2 :28 e82 .s 858.:— 9288 853k. 25.—8 533: $3: a e 8.33 as... a.-. 23... .z 36.2 .2 2.3.2.. 5.5 .3 03.353.» 2M2 .< 58: .a e 883?: e ... use. 23... .2 .8353. :85 .< 5.2:... 658.:— 888 man—:88.» 5M2 .< 85 ... 8 e885 .5... 2-2 is... 882822 his: 5:.Ne .< 88% 88 .< 8585. 668.8 858-. 8.88 68: ..e 3 55 .e ... is... .3 33: .z «5.22.. 2333.. a. 8338 5:58..— B Ea: ... e 8.8: S: 8. 2.3. 5.5... ..ss 33...: .... 2.2.. 2332 ... 1 $3: a a 5:5 8: 8.8 ‘ 88:8 8:8 09.588.» a»... e 55.. .53.? .< m .233 5.9235 38: a 8 :35; we: a 88. as. =8 .m .25 23.8.. a. v... 2.3.8.. .... as: a e 8.55 S a. 85588.» 8.2 .< .83 33.: a. 25.2.. .532 ... .98.. 532» A8: a e 855 3:. 8 .2538 8.5.. .83 .53.... a. 2.5.2.. .538 .< gag—wan 38: a 3 825 a ... .2: 39835 .83 33...: .< 25.2; 933.... .< 88.8 Sea: a e 8&5 .e ... .2: 3.9835 08.588.» 8.... .... 8.5.. 2:32 .< 28: a a 8&5 E... _ 88.95-... 8.5.. ..Se 3.32 .< 25.7..— 2532 ... sea: a e 8&5 S: 3 8.8.95.6 5...... 08.588.» 8.... .... mice 33...: .... 8.5.5 .8888 8588 89.8.8. .350 58> 8: .85 .88..— ..e 8&5 gigéfiflggégagggfiegeh 114 secretion) of the recombinant protein (Gwynne et al., 1987b, 1989). By using a recombinant A. nidulans host with multiple integrated copies of the alcR+ regulatory gene whose gene product is a positive transcriptional regulatory factor for the 0ch+ gene (Doy et al., 1985; Gwynne et al., 1987a; Felenbok et al., 1988), Gwynne et al. (1989) was able to increase the secretion efficiency of A. niger glucoamylase up to 1.2 g/liter complex medium. Sufficient amounts of the alcR‘ gene product derived from the multiple copies of the integrated alcR+ were presumably able to relieve the titration effect caused by the multiple cis-acting regulatory sites of £1ch+ in the expression cassette. CHAPTER II CLONING AND CHARACTERIZATION OF THE TRPC‘ GENE FROM AN AFLATOXIGENIC STRAIN OF ASPERGILLUS PARASITICUS I. INTRODUCTION The five biosynthetic steps from chorismate to. tryptophan, catalyzed by seven enzymatic activities (Figure 9), appear to be the same in bacteria, yeasts, and filamentous fungi (Hiitter et al., 1986). The seven enzymatic functions of Escherichia coli are encoded by 5 genes (trpA’ through an‘) in a single operon and are designated domain A through G (l-liltter et al., 1986). In the filamentous fungi Neurospora crassa and Aspergillus nidulans, four unlinked genes encode four polypeptides, of which two are monofunctional, one is bifunctional, and one is trifunctional. The activities of the fungal trifunctional gene correspond to the domains G, C, and F of E. coli, which encode glutamine amidotransferase (GA'I'), indoleglycerolphosphate synthase (IGPS), and phosphoribosylanthranilate isomerase (PRAI), respectively. In E. coli, the trpC‘ gene product is a bifunctional poly- peptide with IGPS and PRAI activities. The trpC‘ gene has been used previously as a selectable marker for several molecular studies in filamentous fungi. There is considerable conservation of domain structure of this gene throughout the prokaryotic (Crawford, 1975, 1989) and eukaryotic worlds (Htmer et al., 1986). The genes that encode PRAI activity of different filamentous fungi including N. crassa (Schechtrnan and Yanofsky, 1983), A. nidulans (Yelton et al., 1983), Aspergillus niger (Kos et al., 1985), Cochliobolus heterostrophus (Turgeon et al., 1986), Schizophyllwn commune (Munoz-Rivas, et al., 1986a), Penicillium chrysogenum (sanchez et al., 1986), and Phycomyccs blakes- Ieeanus (Revuelta and Jayaram, 1987) have been cloned by complementation of the 116 117 PRAI deficiency in E. coli trpC mutants. Therefore, the A. parasiticus th‘ gene could be selected in suitable E. coli mutant strains based on the same strategy. 118 Figure 9. The biosynthetic pathway of tryptophan from chorismic acid. 119 coon (CAT) coon GI H GI \ (“II-i3 {‘3 3 /u "“3 Phosphoribosyl-PP Pyruvate ' OH Anthranilate Anthranilic Phosphoribasyl- Chorismie acid ' synthetase acid transferasa Coon NH (FD-CFC": 0 HO OH Phosphoribosyt enthunihte 1"°m¢"“(PHA I) coon NH éu—c—cn—cu—caaofi) H H 5H ' End-Hmboaty-phenyhmlnol- 14MNW10W'5'P lndolo glycerol-P CO synthetase : 3-Phospho- (lGPS) glyceraldohyde NH; \ EH OH I Set I -eH,—'-cn—coou J / H—CH—CH3—0® Tryptophan I synthetase N H Twptephan . H lndole glycerol-P 11. MATERIALS AND METHODS A. Bacterial and fungal strains The E. coli K-12 strain DHS (F endAI hstI7 supE44 thi-I recAI gyrA96 reIAI rk'mf) (Hanahan, 1983) was used for plasmid library construction. DHS and E. coli H3101 [hsdszd (13.-mg) recA13 araI4 proAZ lacYI galKZ rpsl20 smr xyLs mtll supE44] were used to propagate plasmids. E. coli JA209 (twp/136 argH metE xyl recA56 str' glyH) (Clarke and Carbon, 1978), JA300 (trpC1117 thr leuB6 thyA W 11st )2de str) (Tschumper and Carbon, 1980), MC1066 [trpC9830 Alac(IPOZYA)X74 galU galK strA )2st leuB6 pyrF74::Tn5 (km’)] (Casadaban et al., 1983) as well as mutants of strain W3110 (Schechtrnan and Yanofsky, 1983), each carrying a different mutation in the rrp operon (Table 7), were used for complemen- tation assays. E. coli NM539 [supF hst (rgmf) (P,)] (Frischauf et al., 1983) and strain 1.13392 (F hst514 supE44 supF58 lacYI galKZ gaITZZ metBI rrpR55 2;) (Murray et al., 1977) were the host for amplification and propagation of the lambda phage library, respectively. The wild-type aflatoxigenic strain A. parasiticus NRRL 5862 (SU-l) (Bennett and Papa, 1988) was the source of DNA for genomic library construction. A. nidul- ans mutant strain FGSC 237 (pabaAI yA2 rrpC 801 , deficient in all trifunctional trpC“ activities) (Kafer, 1977) was the host for studies of heterologous gene expression. 120 121 Table 7. Complementation of E. coli tryptophan auxotrophs by plasmid pLH23. Recipient Deficient Growth on minimal medium° strain' domain(s)" with TRP without TRP MC1066 trpC9830 F + + JABOO trpC1117 F + + W3110 trpC9830 F + + W3110 trpC9941 C + + W3110 AtrpC10-16 C, F + + W3110 AtrpES E + - W3110 AtrpLDIOZ B, D + - W3110 01789578 B + - JA209 trpA36 A + - ‘: Strain JA300 rmC1117 and JA209 tip/136 were purchased from American Type Culture Collection (ATCC); strain MC1066 trpC9830 was obtained from W. E. Timberlake (University of Georgia, Athens, GA); all the W3110 strains were obtained from C. Yanofsky (Stanford University, Stanford, CA). ”: Enzymatic activity encoded by each domain: A, subunit of tryptophan synthase, which catalyzes the conversion of indole-3-glycerophosphate to indole; B, subunit B of tryptophan synthase, which catalyzes the conversion of indole to tryptophan; C, IGPS; D, phosphoribosyl transferase; E, anthranilate synthase (with ammonia as the amino group donor); F, PRAI; G, GAT, interacts with the E domain in the glutamine-dependent anthranilate synthase. ': Cells of each strain were transformed with pLH23, plated on M9 minimal agar, and incubated overnight at 37°C. Colonies which appeared were transferred to M9 minimal medium with or without tryptophan (TRP). 122 B. Growth media For growing A. parasiticus, Czapek Dox broth (agar) and a synthetic low salt medium (Reddy et al., 1971, 1979) were used as minimal media; potato dextrose broth (agar) supplemented with 0.5% yeast extract (PDY) (Bennett and Papa, 1988) was used as a complete medium. Standard complete and minimal media for E. coli (Maniatis et al., 1982; Miller, 1972) and for A. nidulans (Pontecorvo, 1953; Barratt et al., 1965) were used where appropriate. C. Preservation of cultures E. coli strains were routinely stored as frozen (-70°C) stock with 50% glycerol. For storage of fungal strains, young cultures germinated from single spores (conidia) that were isolated by a serial dilution and plating technique (Raper and Fennell, 1965) were transferred onto the center of Petri plates with appropriate media. They were grown for 7 to 10 days at 28-30°C, until the surface of the media was covered with conidia. Suspensions of conidia were prepared by adding sterile solution of 0.01% (v/v) Triton X-100 and lightly scraping plates with an inoculating loop. Suspensions were filtered through a sterile cheese cloth, glass wool or Miracloth (Calbiochem, San Diego, CA) and washed 2 to 4 times with dis- tilled water by centrifugation at about 800 xg. For long-term preservation, stock cultures were generally maintained as frozen (-70°C) spore suspensions in 15-20% glycerol or on silica gel granules as described for Neurospora (Perkins, 1962; Davis and de Serres, 1970). In brief, conidia were suspended in sterile, reconstituted dry 123 skim milk (Difco Laboratories, Detroit, MI). Small, screw-cap vials were half filled with silica gel granules (6-12 mesh, grade 40, desiccant, activated, without indicator, Davison Chemical Corp., Baltimore, MA) and sterilized in a ISO-200°C oven for 2 hr. Samples of the spore suspension were transferred to the vials, cooled with ice, and the vials were sealed. Silica gel stocks were stored at room temp. They were reactivated by sprinkling granules onto the surface of agar-solidified growth medium. For shorter term storage, cultures grown on Petri dishes or slants for one week were sealed with tape or Parafilm and stored at 4°C. Alternatively, short slants were covered with sterile mineral oil (heavy white oil, Sigma Chemical Co., St. Louis, MO) and stored at 4°C. D. Vectors Plasmids pHY201 (Figure 10) and pHYlOl (Yelton et al., 1984), both con- taining a complete trifunctional A. nidulans trpC‘ gene, as well as plasmid pABZ-l (Figure 10; Kos et al., 1985), containing an intact A. niger trpC‘ gene, were sources of probes in heterologous hybridization study. Plasmid pHYlOl was constructed by ligating a 4.3-kb XhoI fragment, which contains the entire A. nidulans an“ gene, with the XhoI-digested pACYC177 (Chang and Cohen, 1978). 124 Figure 10. Restriction maps of trpO-containing plasmids pHY201 and pAB2-1. Plasmid pI-IY201 (Yelton et al., 1984) was constructed by ligating a 4.3-kb XhoI fragment containing a complete wild-type copy of the A. nidulans trpC* gene to Sail-digested pBR329 DNA, thereby interrupting the tetracycline-resistance ('I‘c“) gene. The plasmid confers resistance to ampicillin (Ap‘) and chloramphenicol (Cm‘) to E. coli. Plasmid pAB2-1 (Kos et al., 1985) was isolated from an A. niger gene bank by complemen- tation of an E. coli trpC mutant. It consists of the yeast vector YIpS with a. 7.2-kb A. niger DNA inserted at the BamHI site. The 3.3-kb PvuII-HindlII fragment contains the complete A. niger trpC‘ gene. Small solid triangles indicate restriction sites used to generate probes for heterologous hybri- dization analyses. Thin lines, vector DNA sequences; heavy lines, fungal DNA sequences. Abbreviations are: E, EcoRI; H, HindIII; C, ClaI; Pv, Pqu; X, 20101; B, BamHI; Bg, Bng; S, 5011; P, PstI; Xb, XbaI; Sm, Smal. 125 Slll ".“mtc'onl ' Hind 80m Ml HYZO! be at P “auxin 8.2 Kiloboses Solltxuon 126 Plasmid pRK9 (Schechtrnan and Yanofsky, 1983) was used for the construction of the A. parasiticus genomic DNA library. pRK9 is a pBR322 (Balbas et al., 1986) derivative with a unique BamI-II site created by replacing the 380-base-pair (bp) EcoRI-BamHI fragment of pBR322 with a 96-bp Serratia marcescens trp operon promoter. The bacterial plasmid pUC19 (Yanisch-Perron et al., 1985) and the yeast Saccharomyces cerevisiae vector YIpS (Botstein et al., 1979) were used for sub- cloning. E. General Procedures Preparation of bacteria and plasmid DNA, restriction enzyme digestions, agarose gel electrophoresis, and hybridization analyses were performed by standard procedures (Maniatis et al., 1982; Ausubel et al., 1987). Phage DNA was purified according to the procedure of Carlock (1986). Using a kit purchased from Boehringer Mannheim Biochemicals (Indianapolis, IN), DNA probes were radio- labeled with 3“-P (New England Nuclear Corp., Wilmington, DE; or Amersham Corp., Arlington Heights, IL) by the random primer technique (Feinberg and Vogelstein, 1983, 1984) to a specific activity of greater than 10' cpm/ug of DNA. Unincorporated radioactive precursors were separated from the probes by Sephadex G 5080 (Sigma) exclusion column chromatography as described by Maniatis et al. (1982). Filters with DNAs or RNAs were hybridized in 6x SSC (1x SSC is 0.15 M sodium chloride and 0.015 M sodium citrate), 5x Denhart solution (1x Denhart solution contains 1% Ficoll, 1% bovine serum albumin and 1% polyvinylpyrro- lidone), 40% formamide, 0.1% sodium dodecyl sulfate (SDS), 5 mM 127 ethylenediamine tetraacetic acid (EDTA) and 100 ug/ml denatured salmon sperm DNA at 65°C for higher stringency, and at 37 or 42°C for lower stringency. Filters were washed twice in 0.1% SDS and 2x SSC at room temperature for 40 min followed by a final wash in 0.1% SDS and 0.1% SSC at 65°C for one hour. Auto- radiograms were obtained by exposing filters to Kodak X-OMAT XAR5 diagnostic X-ray film (Eastman Kodak Company, Rochester, NY) with or without Cronex Lightning plus intensifying screens (E. I. Du Pont Nemours & Co., Wilmington, DE) at -70°C for variable time. Restriction enzymes and Other enzymes for manipulation of recombinant DNA were purchased from Bethesda Research Laboratories Inc. (Gaithersburg, MD), Boehringer Mannheim Biochemicals, New England Biolabs (Bevery, MA), New England Nuclear Corp. or Promega Biotec, and were used according to the instruc- tions of the suppliers. Novozym 234 used for production of fungal protoplasts was purchased from Novo Laboratories, Inc. (Wilton, CT). Nitrocellulose and Nytran nylon membranes were purchased from Schleicher & Schuell Inc., (Keene, NH); Gene Screen Plus membrane was supplied by New England Nuclear Corp. All chemicals were of analytical grade and were purchased from Sigma Chemical Co., Aldrich Chemical Co. (Milwaukee, WI), or Fluka Chemical Corp. (Ronkokoma, NY). F. Isolation and manipulation of genomic DNA of A. parasiticus High-molecular-weight chromosomal DNA of A. parasiticus was prepared by _ a procedure modified from that of Cihlar and Sypherd (1980). Mycelia of A. 128 parasiticus NRRL 5862 were harvested fiom YES medium (2% yeast extract and 6% sucrose), lyophilized and blended into a powder with a Waring blender and then ground in a mortar and pestle with glass powder. The resulting mycelial powder was suspended in TSE buffer (100 mM Tris hydrochloride [pH 8.0], 150 mM NaCl, 100 mM EDTA) and incubated with 1% SDS at 37°C for 1 hr with occasional gentle shaking. The suspension was centrifuged (5,000 x g, 10 min) to remove cell debris. The supernatant was extracted with equal volume of TSE-saturated phenol for 1 hr at room temp with gentle shaking. After centrifugation (7,500 x g, 10 min), the aqueous layer was removed and extracted with equal volume of TSE- saturated phenol-chloroform-isoarnylalcohol (25:24: 1). The nucleic acids were then precipitated from the aqueous phase with ethanol, dried under vacuum, and resus- pended in TE (10 mM Tris hydrochloride [pH 8.0], 1 mM EDTA). Contaminating RNA and proteins were removed by treatment with RN ase A and proteinase K (37°C, 1 h each). The enzymes were removed by phenol extraction and the genomic DNA was recovered by ethanol precipitation. With this protocol yields were normally greater than 1 mg of A. parasiticus DNA from 10 g wet mycelia. The majority of the genomic DNA migrated more slowly than phage lambda DNA in agarose gel electrophoresis (larger than 50 kilobase [kb]) and was readily digested with restriction enzymes. G. Preparation of RNA from A. parasia'cus Total RNA was isolated by the hot phenol method of Maramatsu (1973). Frozen mycelia of A. parasiticus NRRL 5862 were ground into powder with a 129 mortar and pestle under liquid nitrogen. The mycelial powder was suspended in RNA extraction buffer (50 mM NaOAc, 1.0 mM EDTA, 1% SDS, pH 5.0) pre- warmed to 65°C. An equal volume of phenol saturated with extraction buffer and prewarmed to 65°C was added immediately and the mixture was incubated at 65°C for 30 min with gentle shaking. The aqueous phase was recovered by centri- fugation (5,000 rpm, 10 min), extracted with an equal volume of 65°C, buffer- saturated phenol, followed by an equal volume of phenol:chloroformzisoamylalcohol (25:24:1) mixture and finally with water-saturated ether. Total RNA was then precipitated from the aqueous phase by adding 1/6 volume of 3 M NaOAc (pH 5.2) and 2.5 volume of absolute ethanol at -20°C. The RNA was collected by centri- fugation (20,000 rpm, 20 min), dried under vacuum and resuspended in RNase-free water. Poly(A)° messenger RNAs were separated from total RNA by oligo(dT) cellulose (Boehringer Mannheim) affinity column chromatography, done by using standard procedures (Maniatis et al., 1982). H. Construction of genomic DNA libraries For construction of the phage lambda library, high-molecular-weight genomic DNA from A. parasiticus was partially digested with restriction endonuclease Sau3A1. DNA restriction fragments (15 to 20 kb) were isolated by fractionation twice on 5-25% sodium chloride gradients (Kaiser and Murray, 1985) and ligated to phage AEMBLB BamHl arms (Promega Biotec). Recombinant phage DNAs were packaged according to the protocol of the supplier (Promega Biotec). Libraries 130 were amplified by the procedure of Maniatis et al. (1982), distributed into 1.5-ml tubes with screw cap, and stored at 4°C with a drop of chloroform. For construction of the plasmid library, genomic DNA fragments with an average size of 5 to 10 kb were generated by partial digestion with Sau3A1 followed by fractionation twice on 5-24% sodium chloride gradients, and then ligated with BamHl-digested, phosphatase-treated plasmid pRK9 (Schechtrnan and Yanofsky, 1983). A. parasiticus DNA fragments were inserted into the unique BamHI site in the S. marcescens twp leader region of pRK9 in order to maximize expression of the gene cloned downsueam from the promoter. The recombinant plasmids were used to transform competent cells of E. coli DH5 (Bethesda Research Laboratories) to ampicillin resistance. The primary plasmid library was amplified as described by Vogeli et al. (1985) and was stored as l-ml aliquots at -70°C with 15% glycerol. I. Lytic complementation of E. coli an mutants Lytic complementation was performed by the procedure of Davis et al. (1980). E. coli rrpC mutants MC1066 and W3110 trpC9830, defective in PRAI activity, were infected with 107 recombinant phages from the A. parasiticus genomic DNA library (multiplicity of infection, < 0.001) and plated on M9 medium lacking tryptophan. Plaques appeared after overnight incubation at 37°C. Phage clones were further purified by reinfecting the susceptible host under selective conditions to isolate single plaques. 131 J. Transformation E. coli Strains were transformed by the hexamine cobalt chloride method (Hanahan, 1983). For transformation of A. nidulans, the procedure of Oakley et al. (1987) was used with the following slight modifications: young germlings for production of protoplasts were prepared from frozen conidia stocks. Protoplasting solution was filtered-sterilized through a 0.45 um Millex-HA Millipore filter (Millipore Products Division, Bedford, MA) and the final protoplast preparation was resuspended in 0.6 M KCl, 0.05 M CaCl, and 10 mM Tris hydrochloride, pH 7.5. K. Laboratory safety Guidelines for the recombinant DNA researches as established by the National Institutes of Health (1986) were observed throughout the experiments. Glasswares, equipments, waste media and cultures that were suspect of conta- mination with aflatoxins or their toxic precursors were routinely soaked in commercially available 10% NaOCl solution for at least 30 min followed by treat- ment with acetone and autoclaving to destroy these toxins (Castegnaro et al., 1980; Yang, 1972; Fischbach and Campbell, 1965; Stoloff and Trager, 1965). III. RESULTS A. Isolation of A. parasiticus trpC’ gene from recombinant phage libraries Two primary phage libraries were constructed (Table 8). Library I contained 1.2 x 10’ recombinant phage clones. It was generated during the preliminary test process to determine the optimal packaging condition for the phage particles. Library 11 contained 2.1 x 10’ recombinant phage clones, and was generated using the Optimal packaging condition. Restriction digestion analysis of DNA from 20 randomly selected clones from both libraries indicated that 99% of them contained A. parasiticus DNA inserts averaging 15 kb. Assuming that the genomic size of A. parasiticus is comparable to that of the A. nidulans (Timberlake, 1978) and N. crassa (Krumlauf and Marzluf, 1979) genomes, the probability of finding any given DNA sequence in the library was greater than 99.9% (Clarke and Carbon, 1976). The primary library was amplified in E. coli NM539, which supports only the growth of recombinant phage particles (Frischauf et al., 1983). The amplified libraries consisted of 5.1 x 10’ (library I) and 6.7 x 10’ (library II) plaque-forming units (PFU) in total. Their titers decreased only 2 to 3-fold after storage at 4°C for approximately three years. E. coli MC1066 and W3110 carrying the trpC9830 missense mutation were used as recipients for lytic complementation. Plaques appeared after infection followed by overnight incubation (Table 9). Twenty phage isolates were purified and replated on both host strains under selective conditions, i.e., without 132 .68: 8:58. Eoeflaapa 9.. 2 8333838832 u a." buggy Sausages .< 053gagnggvlggsagugggagofiigagueggcofi.n. 5.85.5 Eaémcfiiguggggmaugg3§§§3u§§8§~§3u. 2 u 44. 2.2. 2.2. .2 3 c6 2 2 3e :85 < a >< v. xum < >w m .28 .m we 5323803800 9: 156 E. Analysis of A. parasiticus th’ transcript To determine the transcriptional pattern and direction of the cloned trpC‘ gene, filters with poly(A)° RNAs purified from A. parasiticus mycelia grown in minimal and rich medium were hybridized to ”P-labeled RNA probes generated with an SP6 transcription system (Riboprobe System II, Promega Biotec). Probe l but not probe 2 hybridized to a single species of poly(A)* RNA, 2.7 kb in length, which was approximately the same size as trpC‘ transcripts from other filamentous fungi (Figure 18) (Choi et al., 1988; Kos et al., 1988; Mullaney et al., 1985; Revuelta and Jayaram, 1987; sanchez et al., 1986; Schechtrnan and Yanofsky, 1983). The direc- tion of transcription of the A. parasiticus trpC‘ gene (Figure 14, longer arrow) must therefore have been the same as that of the template which gave rise to probe 2. These data were fully consistent with those obtained by heterologous hybridization analysis of phage clones and by deletion mapping of pLH23. Transcriptional ana- lysis did not show significant differences in the level of the trpC‘ mRNA in cell grown in minimal medium or in rich medium, indicating a substantial constitutive expression of the A. parasiticus trpC‘ gene. 157 Figure 18. Northern analysis of the A. parasiticus trpC' transcript. Poly (A)+ RNA isolated from A. parasiticus NRRL 5862 grown in minimal medium (Czapek Dox broth; lane 1, 10 ug) or rich medium (Czapek Dox broth supplemented with 100 ug of L-tryptophan per ml; lane 2, 10 ug) was fractionated on 1.2 % agarose gels containing 2.2 M formaldehyde (Foumey et al., 1988), blotted onto a Gene Screen Plus membrane and hybridized to anP-labeled probe 1 (A) or probe 2 (B) generated with an SP6 transcription system (Riboprobe System 11; Promega Biotec). The templates for SP6 RNA polymerase were prepared by inserting a 0.5-kb BgIII fragment purified from the trpC’-coding region in plasmid pJH34 in the two possible orientations into BamHI-digested Riboprobe plasmid vector pSP64. The resulting recom- binant plasmids were linearized with AMI and subjected to in vitro transcrip- tion with SP6 RNA polymerase by the procedure of Melton et al. (1984). Specific activities were >10' chug of RNA. A solid bar in the diagram indicates the SP6 promoter region. The size of the RNA transcript was estimated with a 0.24. to 9.5-kb RNA ladder from Bethesda Research Laboratories. The 2.7-kb A. parasiticus trpC‘ transcript is indicated by an arrow. For abbreviations of restriction enzymes, see legend to Figure 14. 158 Prob. 2 Probe 1 IV. DISCUSSION Several lines of evidence demonstrate that the complete A. parasiticus trpC‘ gene is present on the plasmid pJI-134. First, pJH34 was capable of complementing different strains of E. coli trpC mutant lacking PRAI activity. Hybridization experiments revealed that A. parasiticus DNA was inserted in pJH34 (Figure 16). A truncated version of this plasmid (pLH23) obtained independently not only complemented the PRAI mutation but also an IGPS mutation (Table 7). Southern blot analysis showed that the 3.4-kb insert of pJH34 was homologous with the trpC° gene of A. nidulans (Figure 13) and A. niger (data not shown). Finally, transfor- mation of an A. nidulans trpC mutant deficient in GAT, IGPS, and PRAI activities with pJH34 resulted in tryptophan prototrophy (Table 10). Plasmid pJH34 was present in DNA prepared from these Trp° transformants. The ability of pJI-134 to restore prototrophic growth of an A. nidulans trpC mutant lacking the GAT, IGPS and PRAI enzymatic activities further indicated that the A. parasiticus trpC° gene codes for a polypeptide with the same enzymatic activities as the A. nidulans trpC* gene product. On the basis of these observations, it is concluded that pJH34 carries a functional A. parasiticus trpC’ gene which can be expressed in both E. coli as well as homologous and heterologous fungal cells. Deletion mapping and Southern hybridization analyses using cloned trpC° genes from other filamentous fungi as probes allow prediction of the direCtion of transcription of the A. parasiticus trpC‘ gene and its functional organization. The data suggested that organization of the A. parasiticus trpC” gene was identical to that of the trpC° (trp-I’) genes of other filamentous fungi in which the gene product 159 160 is a trifunctional polypeptide harboring trpG+ (GAT), trpC‘ (IGPS), and trpF’ (PRAI) activities arranged in the order of NIL-GAT. IGPS. PRAI-COOH. Plasmids carrying the 3.4-kb HindIII insert of A. parasiticus were also capable of complementing different E. coli mutants lacking PRAI activity. The observation that complementation of a an mutation in E. coli was not dependent on the cloning vector or the orientation of the insert in the vector (Table 10) suggested that expression of the gene in E. coli was independent of vector DNA sequences. This implies that the sequences required for transcription and translation initiation in E. coli occur fortuitously within the coding sequence of the A. parasit- icus trpC‘ gene. The PRAI activity encoded by the A. parasiticus th’ was appa- rently initiated in E. coli from within the coding region and not from the A. para- siticus promoter, since the 5’ end of the gene could be eliminated (plasmid pIV 238) and the PRAI activity was still retained. In support of this inference clone Mptrle (Figure 12), which contained only the C-terminal portion of the A. para- siticus trpC’ gene, was able to complement the PRAI deficiency in E. coli. The fact that pJH34 encoded a complete A. parasiticus trpC’ gene but that only PRAI activity could be expressed in E. coli also supported this inference. This same pattern was observed for expression of the A. nidulans th' gene (Y elton et al., 1984) and the N. crassa tip-1° gene (Schechtrnan and Yanofsky, 1983). Expression of both PRAI and IGPS activities encoded by pLH23 in E. coli suggested that trans- cription from this construct was initiated at the Serratia tip promoter. However, the A. parasiticus DNA insert must have provided a Shine-Dalgarno sequence (ribosome binding site) for translation because both pRK9 and the Serratia trp promoter lack this sequence (Schechtrnan and Yanofsky, 1983). 161 Complementation of E. coli mutations is a common practice to isolate eukar- yotic genes. However, since E. coli cells can not process eukaryotic intervening sequences (introns), in principle, only eukaryotic genes lacking introns can be cloned in this way. Consistent with this inference, available DNA sequences of the fila- mentous fungal trpC‘ genes isolated by complementation of E. coli mutations do not contain introns (Choi et al., 1988; Kos et al., 1988; Mullaney et al., 1985; Revuelta and Jayaram, 1987; Schechtman and Yanofsky, 1983). Moreover, attempts to clone the A. parasiticus pyrG* gene by complementation of E. coli pyrF mutants (ATCC 35673, DB6656 [Bach et al., 1979] and MC1066) were not successful, presumably due to the existence of inuon(s). In support of this view, genes in the pyrimidine biosynthetic pathway of N. crassa (Newbury et al., 1986) and P. ansen'na (Béguert et al., 1984) isolated by complementation of E. coli mutations had no introns, whereas A. nidulans pyrG° gene which had one intron could not complement the E. coli pyrF mutation (Oakley et al., 1987). Expression of the trifunctional trpC‘ genes is regulated differently among fungi. For instance, in A. nidulans, the level of the 0226” transcript from cultures grown in a minimal medium is considerably higher than those grown in tryptophan- rich medium (Yelton et al., 1983). In contrast, cultures of C. heterostrophus (Turgeon et al., 1986) and P. blakesleeanus (Revuelta and Jayaram, 1987) grown in minimal or rich media show no significant differences in the amount of TRPI” mRNA. In A. parasiticus, the amount of the trpC* mRNA did not appear to be affected by the presence or absence of tryptophan in the culture medium. There is increasing interest in studies of heterologous gene expression. Apart from the practical applications of developing new gene transfer systems, 162 heterologous gene expression studies are of interest from the evolutionary standpoint in order to see how conserved transcription and regulatory signals are between the major taxonomic groups of fungi. Studies with ascomycete fungi (see Table 5 for summary) have shown that it is not only possible to transfer genes between different fungal species but that these genes can still be metabolically regulated, thus there is conservation of both transcriptional and regulatory signals. The demonstration in this study that the A. parasiticus trpC' gene was able to complement the A. nidulans trpC mutant in transformation supported this generalization. It will be interesting to see if trpC‘ genes from different fungi can be expressed in other heterologous fungal hosts. Casselton and de la Fuente Herce (1989) had shown that a trp-Z mutant of the basidiomycete C. cinereus (corresponding to the trpC mutant in Aspergillus) could be complemented by trpC“ genes from two other basidiomycete species, S. commune and P. chrysosporium, but not by the trpC’ gene from the ascomycete A. nidulans. Cotransformation experiments indicated that the A. nidulans trpC’ gene had integrated into the Coprinus genome but was not expressed. Too little sequence data are available at present to understand the restraints which may limit gene expression across different taxonomic groups of fungi. It should be noted that the ascomycete gene lacks the CAAT and TATAAA motifs characteristic of the promoters of higher eukaryotic genes (Hamer and Timberlake, 1987), whereas these motifs are found in the promoter of the phycomycete gene (Revuelta and Jayaram, 1987). To my knowledge, this report represents the first successful cloning of a gene from an aflatoxin-producing strain of A. parasiticus. This is particularly significant in the task of analyzing the aflatoxin biosynthetic pathway of this fungus 163 at the molecular level. For example, there exist several A. parasiticus mutants blocked at various stages of aflatoxin biosynthesis (Bennett and Papa, 1988). The 0770 gene vector could be used in complementation studies to isolate aflatoxin biosynthetic genes. This cloned trpC” gene will offer a useful selectable marker in parasexual analysis and for development of a DNA-mediated transformation system for A. parasiticus. CHAPTER III ISOLATION OF NITRATE-NONUTILIZING MUTANT S AND CLONING OF THE NITRATE REDUCT ASE STRUCTURAL GENE (NIAD’) FROM ASPERGILLUS PARASITICUS I. INTRODUCTION The assimilation of nitrate has been extensively studied at the biochemical and genetic levels in a variety of eukaryotic and prokaryotic organisms (Wray and Kinghom, 1989). In filamentous fungi, the nitrate reduction pathway has been well- characterized in N. crassa (Marzluf, 1981; Marzluf et al., 1985) and A. nidulans (Cove, 1979). Extracellular nitrate is taken up by a permease (the product of crnA° in A. nidulans). Two enzymes are then induced to reduce nitrate to ammonium: nitrate reductase (A. nidulans m‘aD° and N. crassa nit-3+ gene products), which converts nitrate to nitrite, and nitrite reductase (A. nidulans niiA° and N. crassa nit- t5° gene products), which reduces nitrite to ammonium (Figure 19). Numerous genes, however, control the nitrate assimilation process in these fungi (at least 11 genes in A. nidulans and eight in N. crassa), and their regulation is complex (Cove, 1979; Marzluf et al., 1985). In general, the genes necessary for nitrate assimilation in Aspergillas and Neurospora appear to be similar in function. For example, both the nitrate and nitrite reductase structural polypeptides are encoded by single genes in these fungi. Both fungi also require several genes (at least five in A. nidulans and four in N. crassa) for the synthesis of a molybdenum-containing cofactor that is part of the nitrate reductase complex. This cofactor is also essential for purine dehydrogenase (Pateman et al., 1964) activity in the purine catabolism pathway. In addition to these structural genes, two regulatory genes are known in A. nidulans and N. crassa. The product of the nitrate-assimilation pathway-specific regulatory gene (the A. nidulans nirA° and the N. crassa nit-4° gene) controls the induction 165 166 Figure 19. The nitrate utilization pathway in Aspergillus spp. (modified from Correll et al., 1987). Extracellular nitrate is transported into the cell by a permease encoded by the crnA+ gene, converted into nitrite by the action of nitrate reductase encoded by niaD° gene, and finally converted into ammo- nium ion by nitrite reductase, the m’t‘A° gene product. Nitrate reductase shares a common molybdenum cofactor (encoded by a number of cnx“ genes) with the hypoxanthine and xanthine (purine) dehydrogenase. 167 c— «:0 3.3 S: ...-2.55:5. AI A V 03.52 AIAQ . v 33:2 A «Stow 32.22 35935.. 83260.. 3.35.8.— . 93.52 3952 2.38. .8858 5:53:32 unaccueeemsoe 05.....— / Eon 3.5 111.. \ 055:3” .AT «555.8%: 168 of nitrate reductase and nitrite reductase. The gene product of a second locus, the major nitrogen regulatory gene (the A. nidulans areA+ and the N. crassa nit-2* gene), represses the synthesis of both nitrate reductase and nitrite reductase whenever a preferred nitrogen source such as ammonium or glutamine is present (Cove, 1979; Marzluf et al., 1985). An advantage of the nitrate system for genetic transformation is the ease with which recipient mutants can be generated on the basis of resistance to chlorate (Cove, 1976a; Tomsett and Garrett, 1980; Birkett and Rowlands, 1981; Correll et al., 1987; Newton and Caten, 1988). The positive screening of mutants deficient in nitrate reductase activity through the resistance to chlorate has been successfully used to develop a transformation system in a number of filamentous fungi (summa- rized in Table 5). This procedure is of general applicability and is particularly suitable for fungi without uninucleate spores such as A. parasiticus. In addition, the nitrate-nonutilizing, chlorate-resistant mutants can be easily characterized by their ability to grow on various nitrogen sources (Table 12). In filamentous fungi that have been investigated (Cove, 1979; Tomsett and Garrett, 1980; Birkett and Rowlands, 1981; Newton and Caten, 1988; Klittich and Lesile, 1988; Unkles et al., 1989a, 1989b), the nitrate reductase is encoded by a single gene, and mutants having a lesion in this gene can be recovered at a high frequency. Furthermore, the conserved homology among the nitrate reductase genes (Kinghorn and Campbell, 1989) has allowed identification of this gene from filamentous fungi using cloned heterologous genes as probes (U nkles et al., l989a,b). Thus, it seems feasible that the nitrate reductase gene and appropriate mutants can be obtained and used to develop a transformation system for A. parasiticus. 11. MATERIALS AND METHODS A. Fungal and bacterial strains Escherichia coli strains HBlOl or DHSa (Bethesda Research laboratories, Gaithersburg, MD) were used to propagate plasmids; strain LB392 was used for propagation of phage. The following parental strains of A. parasiticus were used for selection of nitrate-nonutilizing mutants: A. parasiticus NRRL 5862, a wild-type aflatoxigenic strain (Bennett and Papa, 1988); A. parasiticus ATCC 24690, a brown- spored mutant which accumulates a brick-red pigment, norsolorinic acid (Lee et al., 1970); A. parasiticus ATCC 36537, a white-spored mutant which accumulates a yellow pigment, versicolorin A (Lee et al., 1975). Both A. parasiticus ATCC 24690 and 36537 were derived from A. parasiticus NRRL 5862 and produced no detect- able aflatoxins. A. nidulans FGSC A691 (biAI; niaDIS), a nitrate reductase mutant obtained from the Fungal Genetics Stock Center ( University of Kansas Medical Center, Kansas), was used in experiments for heterologous gene complementation. B. Media Media for growth of E. coli, A. nidulans and A. parasiticus were previously described (Chapter II, Horng et al., 1989). Biotin (10 ug/ml) was added to media for growth of A. nidulans FGSC A691. The synthetic low salt medium (SLS) (Reddy et al., 1971, 1979) and potato dextrose broth supplemented with 0.5% yeast exu’aCt (PDY) were used to grow A. parasiticus for protoplast preparation. A 169 170 modified minimal medium for A. nidulans (Pontecorvo, 1953; Barratt et a1, 1965) was used to select and propagate A. parasiticus mutants impaired in nitrate assimilation. This medium (MMG) consists of sodium L-glutamate (1.69 g), pota- ssium chloride (0.52 g), magnesium sulfate.7 H,O (0.52 g), potassium dihydrogen phosphate (1.52 g), glucose (10 g), zinc sulfate (250 mg), ferric sulfate (50 mg), agar (1.5%) and distilled water (1000 ml). The pH was adjusted to 6.5 with sodium hydroxide before sterilization. For growth tests of the chlorate-resistant mutants, the sodium L-glutamate was replaced with one of the following nitrogen sources: sodium nitrate (0.85 g/l); sodium nitrite (0.69 M): ammonium tartrate (1.84 g/l); hypoxanthine (0.1 g/l); uric acid (0.1 g/l). When specified, potassium chlorate was added to selective media at a concentration of 470 mM before autoclaving. C. Vectors Plasmids pSTAlO and pSTA14 (Figure 20; kindly supplied by J. R. Kinghorn, Plant Molecular Genetics Unit, U.K.) containing the niaD“ gene of A. niger (Unkles et al., 1989b) and A. oryzae (Unkles et al., 1989a), respectively, were sources of probes used in heterologous hybridization experiments to isolate the analogous gene from A. parasiticus. 171 Figure 20. Resuiction maps of mum-containing plasmids pSTAlO and pSTA14. Plasmid pSTAlO (Unkles et al., 198%) consists of a 7.5-kb A. niger genomic DNA fragment, which harbors the entire niaD" gene, inserted at the BamI-II site of plasmid pUC8. Plasmid pSTA14 (Unkles et al., 1989a) was constructed by ligating an 8.2-kb SaII fragment, which contained the complete A. oryzae niaD* gene, with the Sail-digested pUC18. One of the BamI-II sites on the polylinker in pSTAlO was disrupted during the cloning process. No cutting sites were found for enzymes Ach, HindIII, PstI, SmaI or XmaI on plasmid pSTA10, and for enzymes BscI, Pstl, Smal, XbaI, or XmaI on plasmid pSTAl4. The approximate start position for niaD’ of A. niger and A. oryzae are determined by hybridization of restriction endo- nuclease-digested, Southem-blotted plasmids to an extreme 5’ end of the A. nidulans m’aD+ fragment. The arrows indicate the direction of transcription as judged by hybridization to a 3’ end of the A. nidulans m’aD+ probe. The approximate positions of the niaD‘ genes within both plasmids are indicated by solid bars; vector DNA sequences are represented by single lines. Small solid triangles indicate restriction sites used to generate DNA probes in heterologous hybridization analyses. Abbreviations are: B, BamI-II; Bg, BgIII; Bs, BssI-III; E, EcoRI; H, HindIII; Hc, HincII; K, KpnI; N, NruI; S, SalI; X, XbaI; Xh, XhoI. 172 ’7 psrmo '1 1 10.2 kb ; . \ 397/ PSTA14 \ Bgfl 10.9 kb 173 D. General procedures Standard recombinant DNA techniques were performed as previously des- cribed (Chapter II; Horng et al., 1989; Maniatis et al., 1982). Where specified, nonradioactive DNA probes generated with the ECL gene detection system (Amersham Corp.) were also used in Southern hybridization analyses according to the procedures of the supplier. E. Isolation and analysis of nitrate-nonutilizing mutants A. parasiticus spontaneous mutants defective in nitrogen assimilation were isolated by positive selection for resistance to chlorate on a minimal medium with sodium L—glutamate as the sole nitrogen source (Cove, 1979). Approximately 107 spores from frozen stocks were spread with a glass rod onto a single 85-mm Petri dish containing MMG supplemented with 470 mM potassium chlorate. Plates were incubated at 30°C. Chlorate-resistant colonies appeared were transferred with toothpicks onto fresh MMG plates supplemented with chlorate to confirm the chlo- rate resistance. Stable resistant mutants were subject to single spore purification (Roper and Fennel, 1965) on selective medium for three rounds before preservation as frozen spore stocks. For classification of mutants, chlorate-resistant colonies were tested for growth on media with chlorate and various nitrogen sources. 174 F. Assay of nitrate reductase Mycelium for enzyme extraction was obtained from cultures inoculated with approximately 10‘ spores and grown overnight at 30°C in PDY. The mycelium was washed and then u'ansferred to fresh minimal medium containing 10 mM nitrate as the sole nitrogen source and shaken for 6 more hours to induce the nitrate reductase activity. Cell-free extracts were prepared by grinding frozen mycelium in liquid nitrogen and resuspending the powder (1 g) in cold 0.1 M phosphate buffer (pH 7.0). The resulting slurry was centrifuged at 4°C for 30 min (15,000 x g). The supernatant was used as a crude enzyme extract. Nitrate reductase activity was assayed with the colorimetric procedure described by Cove (1966). An assay mixture was made as follows: 200 [11 sodium nitrate (337.5 mg/ml); 100 11.1 NADPH (tetrasodium salt) (16 mg/ml); 10 pl FAD (disodium salt) (1.05 mg/ml); 100 pl phosphate buffer (pH 7.75) (0.3 M); 100 [.11 cell extract and distilled water to give a final volume of 3 ml. After 20 min at room temperature, 0.5 ml of a 1% solution of sulfanilamide (dissolved in 25% HCl) was added followed by 0.5 ml of a 0.02% aqueous solution of N -(1-naphthyl)ethylenediamine hydrochloride. The resultant pink color was proportional to the amount of nitrite present, and was estimated by determining the absorbance of the assay mixture specuophmometrically (540 um). Conuol tubes lacking NADPH were used to correct for nitrite present in the extracts, and any turbidity caused by the samples. 175 G. Preparation of protoplasts from A. parasiticus Pr0toplasts of A. parasiticus were prepared by a modification of the procedure of Yelton et al. (1984). Approximately 10' spores from frozen stocks were inoculated into a one-liter Erlenmeyer flask with 400 ml of 81.8 or PDY broth. Mycelia were harvested onto a disc of Miracloth (Calbiochem, San Diego) after overnight growth (16-20 h) at 37°C with shaking. The mycelia were washed with sterile distilled water followed by 0.6 M MgSO., and resuspended in an osmotic buffer (0.8-1.2 M MgSO. in 0.1 M sodium phosphate buffer, pH 5.8; 5 ml/g wet mycelium). A filter-sterilized hydrolytic enzyme solution consisting of Novozym 234 (5 mg/ml), B-glucuronidase (Sigma, 0.2 ml/g wet mycelium) and bovine serum album (12 mg/g wet mycelium) was added. The mixture was incu- bated at 30°C with gentle shaking for 2-3 h. Samples were withdrawn at intervals to check the progress of digestion and possible contamination. The digestion mixture was transferred into sterile centrifuge tubes and carefully overlaid with equal volume of ST buffer (1.2 M sorbitol in 10 mM Tris hydrochloride, pH 7.5). After centrifugation (4,000 x g for 15 min at 4°C using Sorval SS-34 fixed-angle r0tor), the promplasts which banded at the buffer interface were withdrawn with a sterile Pasteur pipette, washed three times by suspending in STC buffer (ST plus 10 mM CaCl,) and centrifugation (2000-3000 rpm at room temperature for 5 min each) in a table-top centrifuge. Protoplasts were resuspended in a suitable volume of STC buffer to give a final concenu'ation of approximately 10'/m1 as determined with a hemacytometer. The regeneration frequency of the pr0toplasts was determined by calculating the portion of the protoplasts that were capable of forming colonies in 176 MG with 1.2 M sorbitol or 0.6 M KCl as osmotic stabilizers. This was generally found to be in the range of 5-15%, depending on the fungal strains used. H. Transformation of fungi For transformation of A. parasiticus a suitable amount (typically 5-10 ug in TE buffer) of DNA was mixed with approximately 101 protoplasts in a volume of 100-200 ul of STC buffer, followed by 50 ul of PEG solution [50% PEG 4000 (Fluka Chemical Corp. or Sigma Chemical Co.), 10 mM CaCl,, 10 mM Tris hydro- chloride, pH 7 .5]. The mixture was allowed to stand on ice for 20 min. One ml of PEG solution was added, mixed thoroughly and incubated at room temperature for 30 min. The transformation mixture was then diluted with ten volumes of STC buffer, added to 400 ml of melted (precooled to 45 to 50°C) selection medium (MMG supplemented with 1.2 M sorbitol and 10 mM nitrate instead of glutamate as the sole nitrogen source). The mixture was poured into 15 to 20 Petri plates, and incubated at 30°C. Control protoplasts were treated as above but with vector DNA only (no insert) or without addition of DNA. A. nidulans was transformed by a modified procedure of Oakley et al. (1987) as described previously (Chapter II). III. RESULTS A. Chlorate-resistant mutants of A. parasiticus Spontaneous chlorate-resistant mutants of various colony sizes were readily recovered at a frequency of approximately one in 10‘ viable spores from all three suains of A. parasiticus tested after incubation for 7 to 10 days at 30°C (Table 11). Growth of wild-type strains was restricted on 470 mM chlorate. However, all nitrate-nonutilizing mutants were resistant to chlorate and showed wild-type growth on MMG. After transfer onto a fresh chlorate medium containing nitrate as the sole nitrogen source, those that grew as thin, expansive, nitrogen-starved colonies with little or no aerial mycelium were considered nitrate—nonutilizing mutants. Selected mutants derived from all three A. parasiticus strains were isolated and purified on chlorate-containing minimal medium with glutamate as the sole source of nitrogen. The ability of these mutants to grow on nitrate, nitrite, ammonium, hypoxanthine, uric acid or glutamate as the sole nitrogen source was assessed. Chlorate resistant mutants could be divided into three phenotypic classes (Figure 21 and Table 12). These classes presumably represent a mutation at a nitrate reductase structural gene (niaD‘), a niu'ate-assimilation pathway-specific regulatory gene (nirA‘), or genes (cnx‘) that affect the assembly of a molybdenum- containing cofactor necessary for niu'ate reductase activity. The phenotype designations used for A. nidulans have been adopted for comparable phenotypes in A. parasiticus. On MMG medium a high frequency of the niaD° 177 178 585-83”. §u8§o§>§ Seawaasaeefiaeeegggigeegegeeg55833233883353 .53”. 9.255.»..— ggncngagsgoefiifiéggfifi"-.gafibéaaznbu+u588_3§§a3§§ fizizafissagcéofigégzaozoggaiggust—">555?33.85320". .ggbugggzgxoéiggggafiuogzr 08: ...—.3 98» gm». 058%.EEE «Se 05» gm». couch! 8.32 310.3 389. 85.. 8.. econ 83:5.— 23:... 85a 8888 sag—e: :58 so» 1583 gap. 932 :52 98» .2383 8332 olez .3: 0m: <3 x: £2 52 .Oz 838::— 9685 259.06 hop-8335:3556 «8:8 325... 298% so 532» 3 $8.. 2. 8 9.33.3 .v. 89¢ 8:52.. 9.3.3-8.. 8.5:. 3 guaca— .= «...-h 179 phenotype was found among the chlorate-resistant mutants (Table 12). All three parental strains were able to overcome the inhibitory effect of chlorate when grown on media with uric acid or ammonium as a sole source of nitrogen, but not with the other nitrogen sources tested (Figure 21). To confirm that the niaD mutants were deficient in nitrate reductase, enzyme activities were measured in crude extracts of selected mutants and wild-type strains. Only the wild-type strains, when induced, displayed nitrate reductase activity. No attempt was made to further characterize the nirA or the cm: mutants. On testing selected niaD mutant strains for reversion to nitrate utilization, it was found that the frequency was generally less than one in 101 viable spores. Thus, the niaD mutants were suitable recipient strains for transformation, because transformation frequencies could be expected to exceed the spontaneous reversion frequency. B. Cloning the A. parasiticus niaD’ gene To test if heterologous niaD° genes could be used as probes to identify the corresponding gene from A. parasiticus, internal restriction fragments from the niaD+ genes of A. niger and A. aryzae (Figure 20) were purified, radiolabeled, and used to probe Southern blots of genomic DNA digests from two strains of A. parasiticus. Homology among niaD+ genes of these three fungi were detected when low strin- gency hybridization and washing conditions were used (Figure 22), 180 Table 12. Comparison of chlorate-resistant mutant types recovered from three strains of A. parasiticus' Types of mutant Su'ain Number“ niaD nirA cnx NRRL 5 862 50 25 15 10 ATCC 24690 62 34 25 3 ATCC 36537 154 70 75 9 ‘: Chlorate-resistant mutants were recovered from basal media with 10 mM L- glutamate as the sole niu'ogen source and supplemented with 470 mM potassium chlorate. ': Numbers of chlorate-resistant mutants analyzed. Each number represents an average of results fiom two independent isolations. 181 Figure 21. Growth of wild-type and nitrate-nonutilizing mutant strains of A. parasiticus ATCC 36537 on media with various nitrogen sources. The arrangement of colonies on each plate is: upper left, wild-type strain ATCC 36537; upper right, niaD mutant; lower left, nirA mutant; lower right, car mutant. A, Nitrate medium (although difficult to see, all mutant strains on this plate showed thin, spidery growth with colonial diameter similar to the wild-type su'ain). B, Nitrite medium; note thin growth of the nirA mutant. C, Ammonium medium; note less growth of the wild-type strain. D, Hypo- xanthine medium; note thin growth of the cnx mutant. E, Uric acid medium; note less growth of the wild-type strain. F, Glutamate medium; note thin growth of the wild-type strain. All but plate A contain 470 mM potassium chlorate. Pictures were taken after incubation for 7 days at 30°C. 182 183 indicating that it was feasible toclone the A. parasiticus niaD° gene by hetero- logous hybridization. An amplified A. parasiticus genomic DNA library in phage lambda ( Chapter II; Horng et al., 1989) was screened by in situ plaque hybridization for the nitrate reductase structural gene using probes prepared from rtiaD° genes of A. niger and A. oryzae. Three phage clones which hybridized strongly to these probes were identified and plaque-purified. Restriction digestion of DNAs purified from these clones with SalI (which releases the DNA insert from the recombinant phage genome) indicated that they contained unique DNA inserts, approximately 17 kb in length, with overlapping regions. These clones were further analyzed by restriction digestion and Southern hybridization using internal DNA fragments from the niaD+ gene of A. oryzae or A. niger as probes. The following unique fragments were identified which hybridized strongly to the A. niger niaD° probe (Figure 23): a 5.2- kb. EcoRI (from clone #1), a 6.2-kb HindIII (from clones #2 and #3), a 5.4-kb SalI (from clone #1), and an 8.2-kb $011 (from clone #2 and #3) fragments. The same fragments also hybridized strongly to the A. oryzae niaD+ probe (data not shown) and were detected in the genomic DNA blots (Figure 22, data for $011 digests were not shown), suggesting that DNA inserts in these three phage clones were not rearranged during the cloning process. The A. niger niaD+ gene probe hybridized strongly to the middle region of the 8.2-kb Sail fragment (Figure 24), suggesting that this fragment is large enough to encode a complete copy of the A. parasiticus niaD° gene. This fragment was subcloned into vector pUCl9 to yield plasmid pSL82 (Figure 24). 184 Figure 22. Identification of the A. parasiticus genomic DNA fragments containing the niaD+ gene by Southern hybridization. Genomic DNAs from (1) A. parasiticus ATCC 36537 and (2) A. parasiticus NRRL 5862 were digested with restriction enzymes BamHI (B), EcoRI (E) and HindIII (H). Restriction fragments were separated by electrophoresis through a 0.7% agarose gel, transferred onto a nitrocellulose membrane, and probed with a 1.7-kb 32P-labeled SalI-BglII fragment purified from plasmid pSTAlO which contains the A. niger niaD° gene (panel a), or with a 2.0—kb a“P-labelled BamHI fragment purified from plasmid pSTAl4 containing the A. oryzae niaD° gene (panel b). Hybridization (in 6x SSC, 5x Denhardt solution, 40% formamide, 0.1% SDS, 5 mM EDTA, 100 pg salmon sperm DNA per ml, 37°C) and washings (twice in 2X SSC-0.1% SDS at room temp for 40 min, followed by a final wash in 2x SSC-0.1% SDS at 42°C for l h) of blots were done under less stringent conditions. Molecular size markers in kilobases are indicated. 185 a E H BE H ! '121'2"12'12121'2' 186 Figure 23. Southern analysis of lambda EMBL3 clones containing the A. parasiticus niaD* gene. Purified DNAs from three phage clones (l, 2 and 3) were digested with restriction enzymes BamI-II (B), EcoRI (E), HindIII (H), and Still (S), fractionated by electrophoresis in a 0.9% agarose gel, and transferred onto a nitrocellulose membrane. The blot was probed with a 1.7- kb Sall-BgIII fragment excised from plasmid pSTAlO which contained major- ity of the coding sequence for the A. niger niaD° gene. The probe was non- radioactively labeled by the ECL gene detection system from Amersham Corp. Due to a nonspecific binding region between top portions of lanes E2 and H2, an approximately 20-kb fragment on lane H1 which hybridized strongly to the probe was invisible. Light bands on lanes 81, 82 and S3 were caused by incomplete digestions. Molecular size markers in kb are indicated. 2.3- 2.0- ‘ 1.4- . 1.1- 187 188 Figure 24. Restriction endonuclease map of plasmid pSL82. The double lines represent the 8.2-kb 8011 DNA insert from A. parasiticus. Plasmid pUC19 is represented by a single line. A striped block represents the minimal coding region of the A. parasiticus niaD+ gene which hybridizes strongly to the A. niger niaD° gene. Abbreviations for restriction enzymes are: E, EcoRI; H, HindIII; K, KpnI; P, Pstl; S, SaII; X, XbaI; Xh, XhoI. 189 IV. DISCUSSION The wild-type strain of A. parasiticus is capable of utilizing nitrate as the sole nitrogen source. Nitrate-nonutilizing mutants of this fungus were readily obtainable by selecting spontaneous mutants that were resistant to chlorate. The genotype of these mutants could be easily characterized by growth tests on various nitrogen sources. Only three types of mutants (niaD, nirA and cm) were isolated when glutamate was used as the sole nitrogen source in the chlorate selection medium. A significant portion (approximately 50%) of these chlorate-resistant mutants were niaD mutants. This high proportion of niaD mutants was consistent with results from Cove (1976b) who found that nitrogen sources had significant effects on relative frequencies and types of chlorate-resistant mutants recovered. Glutamate favored the selection of niaD mutants. Using L-arginine as the sole nitrogen source for selection, Papa (1986) also recovered three classes of chlorate- resistant mutants with similar phenotypes from A. flavus, an aflatoxigenic fungus closely-related to A. parasiticus. Therefore, nitrate assimilation in A. parasiticus appears to be very similar to that in A. nidulans in that both species are sensitive to chlorate and produce spontaneous mutants with similar physiological characteristics. The cm:+ loci encoding molybdenum cofactor in A. parasiticus were not further analyzed in this study, nor was the nirA° locus. Depending on species, there are 4 to 7 distinct cm:° loci in filamentous fungi (Unkles, 1989). These are generally distinguished from each other by complementation tests. The structural gene for the nitrate reductase of A. parasiticus was isolated on the basis of its homology with analogous genes from A. niger and A. oryzae. The, 190 191 identity of this gene is being characterized by subcloning and transformation of heterologous and homologous niaD mutants. Preliminary experiments indicated that plasmid pSL82 or DNA fragments carrying the cloned A. parasiticus niaD” gene were able to transform a putative niaD mutant derived from A. parasiticus ATCC 36537 at a frequency of approximately 100 transformants/11g DNA. However, att- empted transformations of A. nidulans FGSC A691 with pSL82 were not successful. The reason for these failures remains unclear. Chlorate resistance provides a system for the positive selection of mutations with an auxotrophic phenotype in several different genes and has wide potential applications. The procedure is economical in terms of labor and materials and, furthermore, does not require the use of mutagens, thereby avoiding problems resulting from the mutagenic induction of chromosomal aberrations or multiple mutations. The procedure is particularly suitable for isolation of mutants from genetically-undeveloped fungal strains such as A. parasiticus. This study has demonstrated that chlorate resistant mutants can be easily isolated from three strains of A. parasiticus, including two blocked mutants that are impaired in aflatoxin biosynthesis. These mutants are being used as recipient mains to develop a genetic transformation system for cloning genes associated with aflatoxin biosynthesis. In addition, chlorate resistance should be useful as a general marker in genetic mapping analyses to construct linkage maps. Furthermore, the niaD+ gene represents a selectable marker which can be selected and counterselected. This feature will be extremely useful in designing gene replacement and disruption experiments (see Chapter I for discussion) for A. parasiticus. Moreover, chlorate resistant mutants of 192 A. flavus have been successfully used in studies of vegetative (heterokaryon) compatibility (Papa, 1986; Bayman and Cotty, 1989). CHAPTER IV ISOLATION OF A DNA FRAGMENT FROM A BENOMYL—RESIST ANT MUTANT OF ASPERGILLUS PARASITICUS WITH HOMOLOGY TO THE NE UROSPORA CRASSA TUB-2’ GENE I. INTRODUCTION The tubulins are a family of the principal protein subunits of microtubules in the eukaryotic cell cytoplasm. Tubulins polymerize into a diverse number of microtubule arrays whose assembly and functional properties are defined both by specific programs of cellular differentiation and by cell cycle determinants. Microtubules represent the principal structtual components of mitotic and meiotic spindles. They participate in several aspects of intracellular transport, in maintenance of various cell surface properties, and establish overall cell shape and internal cytoplasmic architecture (Roberts and Hyams, 1979; Borisy et al., 1984). Tubulin amino acid sequences are highly conserved among evolutionarily diverse organisms (Cleveland and Sullivan, 1985). This has allowed tubulin genes to be cloned and characterized from many different organisms. Multiple genes encoding different tubulin proteins are common in eukaryotes. For example, in the filamentous fungus A. nidulans, tubA° encodes two a-tubulin polypeptides and tubB° encodes the third a—tubulin. The B—tubulins parallel the Ot-tubulins in that the ban° locus codes for two Botubulin polypeptides, and the tubC’ locus codes for another (Morris, 1986). Recently, the y—tubulin polypeptide, a new member of the tuban super-family encoded by the mipA° locus, was discovered in A. nidulans by Oakley and Oakley (1989). Fewer tubulin genes have been found in yeasts and other filamentous fungi. For example, two a-tubulin genes and one B-tubulin gene have been identified in the fission yeast Schizosaccharomyces pombe (Hiraoka et al., 1984; Toda et al., 1984), and in the budding yeast Saccharomyces cerevisiae (Neff et al., 1983; Schatz et al., 1986). Neurospora crassa contains only one B—tubulin 194 195 gene (Orbach et al., 1986). The dimorphic, pathogenic fungus Histoplasma capsulatum contains a single a— and a single B-tubulin gene (Harris et al., 1989). Polymerization of tubulin monomers into microtubules is inhibited by the antimitotic fungicide benomyl [methyl 1-(butylcarbamoyl)-2-benzimidazole carba- mate]. Benomyl resistance is caused by alternations in tubulins which apparently inhibit or reduce binding of benomyl to the tuban protein (Davidse, 1986). In fungi, most mutations conferring resistance to benomyl map in the B-tubulin structural genes (Hiraoka et al., 1984; Neff et al., 1983; Sheir-Neiss et al., 1978; Thomas et al., 1985; Orbach et al., 1986). A mutant allele of the B-tubulin gene has been cloned from a benomyl-resistant strain of N. crassa, and was successfully used as a dominant selectable marker to transform this fungus (Orbach et al., 1986) and several others (summarized in Table 5). Preliminary investigations have demonstrated that growth of A. parasiticus is completely inhibited by benomyl at a concentration of 5 pig/ml. Thus, this fungus potentially can be transformed to benomyl resistance. However, repeated transfor- mations of A. parasiticus using the N. crassa benomyl resistance gene as a selec- table marker were not successful, indicating that this marker either was not expressed or was insufficiently expressed in A. parasiticus. An attempt was thus initiated to clone the native B-tubulin gene from a benomyl-resistant strain of A. parasiticus, and to use it as a dominant selectable marker for transformation of this fungus. A similar strategy was used to develop a transformation system for A. flavus (Seip et al., 1989). II. MATERIALS AND METHODS A. General procedures Purification of DNAs from fungi, phage particles and plasmids were pre- viously described (Homg et al., 1989; Chapter II). Standard recombinant DNA procedures were performed as described by Maniatis et al. (1982). B. Isolation of benomyl-resistant mutants Approximately 10' spores of A. parasiticus ATCC 36537 were irradiated with ultraviolet (uv) light (254 nm) resulting in a 10% survival level. The treated spore suspension was spread onto a single Petri dish containing Czapek-Dox agar medium supplemented with benomyl (5 uglml) (DuPont Co, technical grade, 98% pure). Resistant colonies appeared after incubation at 30°C and were purified through single spore isolation. The level of resistance of these isolates to benomyl was determined on Czapek-Dox agar plates supplemented with various concentrations of benomyl. C. Construction of a genomic DNA library Chromosomal DNA was purified from a benomyl resistant isolate (20 ug/ml) of A. parasiticus ATCC 36537, and used to construct a genomic DNA library in the lambda vector EMBL3 following the procedures as described previously (Homg et al., 1989; Chapter II). 196 197 D. Conditions for Southern analysis and plaque hybridization A 2.6—kb SalI fragment excised from plasmid pBT3 (Figure 25) containing the cloned mutant allele of the N. crassa B-tubulin gene (Orbach et al., 1986) was used as a heterologous probe to identify the analogous B-tubulin gene of A. parasi- ticus. Southern hybridization analysis was conducted with the radiolabeled N. crassa B-tubulin gene probe and A. parasiticus genomic DNA on a nitrocellulose filter (BA 85, Schleicher & Schuell). The hybridization reaction was performed at 42°C in 6x SSC-5x Denhardt solution-40% formamide-0.l% SDS-100 ug of dena- tured salmon sperm DNA per ml. Filters were washed twice for 20 min at room temp in 2x SSC-0.1% SDS and then twice for 30 min at 63°C in 0.1x SSC-0.1% SDS. The A. parasiticus lambda DNA library was screened by in situ plaque hybridization with the N. crassa B-tubulin probe as described previously (Chapter III). Filters with phage DNAs were hybridized at 45°C and washed at 52°C in the same solutions as described for genomic DNA blots. 198 Figure 25. Resuiction endonuclease map of plasmid pBT3. The double lines represent the 3.1-kb N. crassa DNA insert. Plasmid pUC12 is represented by a single line. The pUC12 polylinker region is expanded on the left side of the plasmid. The line with an arrow, above the N. crassa DNA, repre- sents the B-tubulin transcript. The dashes at the 5’ end indicate that this end has not been determined precisely. Restriction sites indicated by solid triangles are used to prepare the DNA probe for use in heterologous hybridi- zation. Restriction sites are abbreviated as follows: E, EcoRI; Ss, SstI; Sm, SmaI; B, BamI-II; Xb, XbaI; Sa, 5011; P, PstI; H, HindIII; K, KpnI; N, NcoI; Xh, XhoI. The map is complete for these enzymes. 1° represents 16 bp (Orbach et al., 1986). 199 5x a“ moan w e. we. whma mm: 2x ‘ xm s nxx>z x x 5224‘ 200 E. Transformation of A. parasiticus Protoplasting and transformation of A. parasiticus were performed essentially as described previously (Chapter III). Benomyl-resistant uansformants were selected by two methods. First, portions of the transformation mix were added to Czapek- Dox top agar (1%) containing 1.2 M sorbitol, and plated onto agar (1.5%) plates of similar composition, excluding the sorbitol and including benomyl at 5 rig/ml. Alternatively, portions of the transformation mixture were spread onto Czapek-Dox agar plates (1.5% agar) containing 1.2 M sorbitol. After incubation at 30°C overnight, top agar (1%) containing 5 jig/ml benomyl was gently overlaid on the surface of the medium. Benomyl was made fresh as a 0.5 mg/ml stock solution in 100% ethanol, and was added without further sterilization after the medium had been autoclaved. III. RESULTS AND DISCUSSION Although the N. crassa benomyl resistance gene has been successfully used as a dominant selectable marker in transformation of several filamentous fungi (see Table 5), preliminary transformation studies on A. parasiticus with plasmid pBT3 and cosmid pSV50 (V ollmer and Yanofsky, 1986), both containing the mutant allele of the N. crassa tub-2* gene conferring resistance to benomyl (Orbach et al., 1986), suggested that this marker was not sufficiently expressed in A. parasiticus for selection of transformants. Similar results were also found (Woloshuk et al., 1989) in attempts at transformation of A. flavus with the benomyl resistance genes fiom N. crassa and A. nidulans. DNA restriction fragments with strong homology to the cloned N. crassa rub- 2° gene were identified in the genome of A. parasiticus using Southern analysis under high-suingency hybridization and washing conditions (Figure 26). These data indicated that it was feasible to identify the analogous B-tubulin gene in an A. parasiticus genomic DNA library using this heterologous probe. A strain of A. parasiticus resistant to 20 ug/ml of benomyl was isolated by repeated mutagenesis with uv irradiation. Hybridization analysis failed to detect any difference in distribution of restriction fragments between the benomyl-resistant mutants and the wild-type strain of A. parasiticus. A genomic DNA library (1.4 x 10’ PFU) was constructed in the vector lambda EMBL3 using genomic DNA puri- fied from this strain. A total of 12,000 plaques from the library were screened at a density of 2,400 plaques per 82-mm Petri plate using a 2.6-kb SaII restriction fragment excised from plasmid pBT3 as probe. 201 202 Figure 26. Identification of restriction DNA fragments from the A. parasi- ticus chromosome with homologies to the N. crassa B-tubulin gene. Purified genomic DN As from A. parasiticus ATCC 36537 (lane 1), and its benomyl- resistant isolates (lane 2, resistant to 10 ug/ml benomyl; lane 3, resistant to 20 rig/ml benomyl) were digested with various restriction enzymes, fractio- nated by electrophoresis through a 0.8% agarose gel, and transferred onto a nitrocellulose membrane. The blot was hybridized with a radiolabeled 2.6- kb Sail fragment excised from plasmid pBT3. The aberration in lane E3 was caused by an incomplete digestion of the genomic DNA by EcoRI. Abbreviations are: B, Band-II; E, EcoRI; H, HindIII; P, PstI; S, SalI; X, XbaI. Molecular size markers in kb are indicated. 203 — 3 2 9.4‘ 4.4- 2.3- 2.0- 1 .4- 204 Three plaques that hybridized strongly were recovered and purified. Restriction enzyme analyses of DNAs purified from these clones indicated that two of them were identical and shared few common restriction fragments with the, third clone for most of the resuiction enzymes tested (data not shown). The same restriction fragments present in the two identical phage clones, but not in the third clone, were also deteCted in the A. parasiticus chromosome, suggesting that the two identical phage clones probably contain a complete copy of the putative mutant allele of the A. parasiticus B—tubulin gene (designated ban"), with the copy in the third clone being incomplete or rearranged. However, a 7.4-kb Nsfl fragment and a 12-kb SmaI fragment that hybridized strongly to the N. crassa fl-tubulin probe were present in all three phage clones (data not shown). These fragments are being subcloned into plasmid pUCl9. Further analyses of these plasmids by transformation of A. parasiticus and by DNA sequencing will provide insight into the identity and su'ucture of the cloned fi-tubulin gene. Attempted transformations of A. parasiticus with DNAs purified from the three phage clones were not successful. The reasons for this failure remain unclear. However, there are several possible explanations. First, with the high-molecular- weight phage DNA, the transformation frequency may be too low to detect. Second, it is possible that the cloned gene did not confer resistance to benomyl, or conferred insufficient resistance for selection of transformants. Mutated [S-tubulin genes are not responsible for all cases of benomyl resistance. For example, in the fungus Sporobolomyces roseus resistance to benomyl can be caused by the differ- ential uptake of this fungicide by the sensitive and resistant strains (Nachmias and Barash, 1976). Alternatively, the cloned gene may not represent a mutated B- 205 tubulin gene, but an a- or a 'y-tubulin gene which may not confer resistance to benomyl. This is reasonable since all the a- and B-tubulins reported to date share 36-42% identity with each other, and the y-tubulin from A. nidulans is 33.3% identical to the corresponding B-tubulins (Oakley and Oakley, 1989). However, the hybridization reaction used to identify this gene was conducted under high- stringency conditions which tended to rule out this possibility. The screening procedure, in theory, should allow only fl-tubulin gene to be isolated, unless the homology between the N. crassa B-tubulin gene and the A. parasiticus a- or y- tubulin gene is stronger than that between the N. crassa B-tubulin gene and the A. parasiticus B-tubulin gene. Finally, the method and time of application of selective pressure in the current transformation protocol may not be appropriate for selection of benomyl-resistant transformants in A. parasiticus. Orbach et al. (1986) nated that if benomyl was included in the overlaying top agar, the transformation frequency for benomyl resistance of N. crassa was reduced 4- to 10-fold, suggesting that transformed cells require a growth period to allow expression of the benomyl resistance gene before selection is applied. In conclusion, a systematic analysis is required to characterize the nature of the mutation in the benomyl-resistant mutants and the putatively cloned B-tubulin gene before they can be used in developing a transformation system for A. parasi- ticus. CONCLUSIONS 207 Selectable markers are necessary for development of genetic transformation systems for cloning genes involved in the aflatoxin biosynthesis from A. parasiticus. Three such genetic markers, trpC“, niaD’, and ben’, have been isolated and charac- terized from this fungus in this study, while a fourth marker, pyrG° was also isolated in the lab. A transformation system based on the cloned trpC‘ gene as a selectable marker can be developed once suitable trpC mutants of A. parasiticus are available to serve as recipient strains. Attempts to isolate a trpC auxotroph from A. parasiticus by traditional mutagenesis and screening techniques were not successful, presumably due to the existence of multiple nuclei in its conidia and the coenocytic nature of the fungal cytoplasm. Gene disruption and replacement techniques as previously described offer an alternative strategy to obtain such a mutant. Plasmid pLH23 (Figure 14) and its deletion derivatives (Figure 17) that carry different truncated versions of the A. parasiticus trpC° gene are well suited for this purpose. This approach requires an established transformation system with an appropriate selectable marker for the primary selection. Successful development of transformation systems based on niaD° or the benomyl resistance marker should make this approach feasible. The gene coding for nitrate reductase (niaD’) provides another potential selectable marker to develop a transformation system for A. parasiticus. The niaD” gene has been cloned and mutants deficient in nitrate reductase activity have been isolated from A. parasiticus. A homologous transformation system has been deveIOped based on the niaD‘ as the selectable marker. This is a transformation system of general applicability since this investigation has demonstrated that the 208 recipient mutants can be easily selected from wild-type and blocked mutant strains of A. parasiticus by virtue of their resistance to chlorate. A gene (ben') has been cloned from a benomyl-resistant strain of A. para- siticus by virtue of its homology to the N. crassa B-tubulin gene. The ben’ gene potentially encodes a mutant allele of the B-tubulin gene which confers resistance to the fungicide benomyl, and can be used as a dominant selectable marker in transfor- mation of A. parasiticus since this fungus is reasonably sensitive to benomyl. Comparison of DNA sequences between B-tubulin genes isolated from wild-type and the benomyl-resistant strains of A. parasiticus will shed light on the nature of the cloned benomyl resistance marker. This work represents the first systematic effort to explore the aflatoxigenic A. parasiticus by the modern molecular genetic techniques. The wild-type A. para- siticus genomic DNA libraries consumed and preserved in this study are of suffi- cient quality to allow successful isolation of three genes (trpC’, niaD’, and pyrG‘), and should provide stable sources for obtaining more genetic markers from this fungus. Genes isolated in this study can be used as general markers for routine genetic analyses. Further characterization of these cloned genes by DNA sequence analysis will provide insight into the structure, organization, and nature of the regulatory elements for genes of this fungus, which should help in designing cloning and expression vectors to analyze the aflatoxin biosynthetic pathway. LIST OF REFERENCES 10. 11. REFERENCES . Adams, T. H., M. T. Boylan, and W. E. Timberlake. 1988. (MA is nece- ssary and sufficient to direct conidiophore development in Aspergillus nidulans. Cell 54:353-362. Adye, J., and R. I. Mateles. 1964. Incorporation of labelled compounds into aflatoxins. Biochirrr. Biophys. Acta 86:418-420., Akins, a. A., and A. M. Lambowitz. 1985. General method for cloning Neurospora crassa nuclear genes by complementation of mutants. Mol. Cell. Biol. 5:2272-2278. Alderson, T., and C. Scazzocchio. 1967. A system for the study of inter- locus specificity for both forward and reverse mutations in at least eight gene loci in Aspergillus nidulans. Mutat. Res. 4:567-577. Alic, M., J. R. Komegay, D. Pribnow, and M. H. Gold. 1989. Transforma- tion by complementation of an adenine auxotroph of the lignin-degrading basidiomycete Phaerochaete chrysosporium. Appl. Environ. Microbiol. 55:406-411. Anderson, M. S., and M. F. Dutton. 1979. The use of cell free extracts derived from fungal protoplasts in the study of aflatoxin biosynthesis. ‘Experientia 35:21-22. Anderson, M. S., and M. F. Dutton. 1980. Biosynthesis of versicolorin A. Appl. Environ. Microbiol. 40:706-709. Andrianopoulos, A., and M. J. Hynes. 1988. Cloning and analysis of the positively acting regulatory gene (2de from Aspergillus nidulans. Mol. Cell. Biol. 8:3532-3541. Anne, 1., H. Eyssen, and P. de Somer. 1974. Formation and regeneration of Penicillium chrysogenum protoplasts. Arch. Microbiol. 98:159-166. Applebaum, R. S., and E. H. Marth. 1981. Biogenesis of the C” polyketide, aflatoxin. Mycopathologia 76:103-114. Aramayo, R., T. H. Adams, and W. E. Timberlake. 1989. A large cluster of highly expressed genes is dispensable for growth and development in Aspergillus nidulans. Genetics 122:65-71. ' 210 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 211 Arnau, J., F. J. Murillo, and S. Torres-Martinez. 1988. Expression of TnS-derived kanamycin resistance in the fungus Phycomyces blakesleeanus. Mol. Gen. Genet. 212:375-377. Arst, H. N., Jr., and C. Scazzocchio. 1975. Initiator constitutive mutation with an "up-promoter" effect in Aspergillus nidulans. Nature 254:31-34. Arst, H. N., Jr., and C. Scazzocchio. 1985. Formal genetics and molecular biology of the control of gene expression in Aspergillus nidulanS. PP. 309-343. In Bennett, J. W., and L. L. Lasure (eds.), Gene manipulations in fungi. Academic Press, Inc., New York, NY. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K, Struhl. 1987. Current Protocols in molecular biology. John Wiley & Sons, New York. Ayer, W. A., L. Pena-Rodriguez, and J. C. Vederas. 1981. Identification of sterigmatocystin as a metabolite of Monocillium nordinii. Can. J. Microbiol. 27:846-847. Bach, M. L., F. Lacroute, and D. Botstein. 1979. Evidence for transcrip- tional regulation of orotidine-5’-phosphate decarboxylase in yeast by hybridization of mRNA to the yeast structural gene cloned in E. coli. Proc. Natl. Acad. Sci. USA 76:386-390. Bachmann, B. J., and D. M. Bonner. 1959. Protoplasts from Neurospora crassa. J. Bacteriol. 78:550-556. Balbas, P., X. Soberon, E. Merino, M. Zurita, H. Lomeli, F. Valle, Flores, N, and F. Bolivar. 1986. Plasmid vector pBR322 and its special purpose derivatives- a review. Gene 50:3-40. Ballance, D. J., and G. Turner. 1985. Development of a high-frequency transforming vector for Aspergillus nidulans. Gene 36:321-331. Ballance, D. J., and G. Turner. 1986. Gene cloning in Aspergillus nidulans: isolation of the isociuate lyase gene (acuD). Mol. Gen. Genet. 202:271-275. Ballance, D. J., F. P. Buxton, and G. Turner. 1983. Transformation of Aspergillus nidulans by the orotidine-5’-phosphate decarboxylase gene of Neurospora crassa. Biochem. BiOphys. Res. Commun. 112:284-289. Banks, G. R. 1983a. Transformation of Ustilago maydis by a plasmid containing yeast 2-micron DNA. Curr. Genet. 7:73-77. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. \35. 36. 212 Banks, G. R. 1983b. Chromosomal DNA sequences from Ustilago maydis promote autonomous replication of plasmids in Saccharomyces cerevisiae. Curr. Genet. 7:79-84. Banks, G. R., and S. Y. Taylor. 1988. Cloning of the PYR3 gene of Usn'lago maydis and its use in DNA transformation. Mol. Cell. Biol. 8:5417-5424. Barnes, D. E., and D. W. MacDonald. 1986. Behavior of recombinant plasmids in Aspergillus nidulans: structure and stability. Curr. Genet. 10:767-775. Barratt, B. W., G. B. Johnson, and W. N. Ogata. 1965. Wild-type and mutant stocks of Aspergillus nidulans. Genetics 52:233-246. Bartnicki-Garcia, S. 1968. Cell wall chemisu'y and taxonomy of fungi. Annu. Rev. Microbiol. 22:87-108. Bayman, P., and P. J. Cotty. 1989. Vegetative compatibility groups in Aspergillus flavus. APS Annu. Meet. Abst. No. 406. Bégueret, J., V. Razanamparany, M. Perrot, and C. Barreau. 1984. Cloning gene ura5 for the orotidylic acid pyrophosphorylase of the filamentous fungus Podospora anserina: transformation of protoplasts. Gene 32:487-492. Bej, A. K., and M. H. Perlin. 1989. A high efficiency transformation system for the basidiomycete Ustilago violacea employing hygromycin resistance and lithium-acetate treatment. Gene 80:171-176. Bennett, J. W. 1979. Aflatoxins and anthraquinones from diploids of Aspergillus parasiticus. J. Gen. Microbiol. 113:127-136. Bennett, J. W. 1981. Loss of norsolorinic acid and aflatoxin production by a mutant of Aspergillus parasiticus. J. Gen. Microbial. 124:429-432. Bennett, J. W. 1982. Genetics of mycotoxin production with emphasis on aflatoxins, pp. 549-561. In Krumphanzl, V., B. Sikyta, and Z. Vanek (eds.), Overproductian of microbial products. Academic Press, Inc., London. Bennett, J. W., andS. B. Christensen. 1983. New perspectives on aflatoxin biosynthesis.lAdv. Appl. Microbiol. 29:53-92. ,. Bennett, J. W., and E. Deutsch. 1986. Genetics of mycotoxin biosynthesis, pp. 51-64. In Steyn, P. 8., and R. Vleggaar (eds.), Mycotoxins and phyco- toxins. Elsevier Science Publishers, Amsterdam. 37. 38. 39. 40. 41. 42. 43. 45. 46. 47. 48. 49. 213 Bennett, J. W., and L. A. Goldblatt. 1973. The isolation of mutants of Aspergillus flavus and A. parasiticus with altered aflatoxin producing ability. Sabouraudia 11:235-241. Bennett, J. W., and L. L. Lasure. 1985a. Gene manipulations in fungi. Academic Press, Inc., New York, NY. Bennett, J. W., and L. L. Lasure. 1985b. Conventions for gene symbols, pp. 537-544. In Bennett, J. W., and L. L. Lasure (eds.), Gene manipulations in fungi. Academic Press, Inc., New York. Bennett, J. W., and L. S. Lee. 1979. Mycotoxins- their biosynthesis in fungi: aflatoxins and other bisfuranoids. J. Food Prat. 42:805-809. Bennett, J. W., and K. E. Papa. 1988. The aflatoxigenic Aspergillus spp. Adv. Plant Pathol. 6:263-280. Bennett, J. W., L. S. Lee, and A. F. Cucullu. 1976. Effect of dichlorvos an aflatoxin and versicolorin A production in Aspergillus parasiticus. Bot. Gaz. 137:318-324. Bennett, J. W., L. S. Lee, S. M. Shoss, and G. H. Boudreaux. 1980a. Iden- tification of averantin as an aflatoxin B, precursor: placement in the biosynthetic pathway. Appl. Environ. Microbiol. 39:835-839. Bennett, J. W., P. -M. Leong, S. Kruger, and D. Keyes. 1986. Sclerotial and law aflatoxigenic morphological variants from haploid and diploid Aspergillus parasiticus. Experientia 42:848-851. Bennett, J. W., R. B. Silverstein, and S. J. Kruger. 1981a. Isolation and characterization of two nonaflataxigenic classes of morphological variants of Aspergillus parasiticus. J. Amer. Oil Chem. Soc. 58:952A-955A. Bennett, J. W., C. H. Vinnett, and W. R., Jr., Gaynes. 1980b. Aspects of parasexual analysis in Aspergillus parasiticus. Can. J. Microbial. 26:706-713. Bennett, J. W., D. G. Wheeler, and J. J. Dunn. 1981b. Genetic analysis of aflatoxin production by Aspergillus parasiticus, pp. 417-422. In Moo-Young, M., C. Vezina, and K. Singh (eds.), Advances in biotechnology. Vol. III. Fermentation products. Pergaman Press, Toronto. Berdy, J. 1980. Bleamycin type antibiotics, pp. 459-491. Handbook of antibiotic compounds, Vol. IV. CRC Press, Boca Raton, Florida. Berges, T., and C. Barreau. 1989. Heat shock at an elevated temperature improves transformation efficiency of protoplasts from Podospora anserina. J. Gen. Microbial. 135:601-604. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 214 Beri, R. K., and G. Turner. 1987. Transformation of Penicillium chrysogenum using the Aspergillus nidulans amdS gene as a dominant selective marker. Curr. Genet. 11:639-641. Bemier, L., R. M. Cooper, A. K. Charnley, and J. M. Clarkson. 1989. Transformation of the entomopathogenic fungus Metarhiziwn anisopliae to benomyl resistance. FEMS Microbiol. Lett. 60:261-266. Berse, B., A. Dmochawska, M. Skyrzypek, P. Weglenski, M. A. Bates, and R. L. Weiss. 1983. Cloning and characterization of the arnithine carbamoyl- transferase gene from Aspergillus nidulans. Gene 25:109-117. Betina, V. 1984. Mycotoxins- production, isolation, separation and purifi- cation. Elsevier Science Publishers, Amsterdam. Bhatnagar, D., and T. E. Cleveland. 1988. Fate of the methyl group during the conversion of sterigmatocystin into O-methylsterigmatocystin and aflatoxin B, by cell-free preparations of Aspergillus parasiticus. Biochimie 70:743-747. Bhatnagar, D., '1‘. E. Cleveland, and A. R. Lax. 1989. Comparison of the enzymatic composition of cell-free extracts of non-aflatoxigenic Aspergillus parasiticus with respect to late stages of aflatoxin biosynthesis. Arch. Environ. Contam. Toxicol. 18:434-438. Bhatnagar, D., S. P. McCormick, L. S. Lee, and R. A. Hill. 1987. Identi- fication of O-methylsterigmatocystin as an aflatoxin B, and G, precursor in Aspergillus parasiticus. Appl. Environ. Microbiol. 53:1028-1033. Bhatnagar, D., A. H. J. Ullah, and T. E. Cleveland. 1988. Purification and characterization of a methyltransferase from Aspergillus parasiticus SRRC 163 involved in aflatoxin biosynthetic pathway. Preparative Biochem. 18:321-349. Binninger, D. M., C. Skrzynia, P. J. Pukkila, and L. A. Casselton. 1987. DNA-mediated transformation of the basidiomycete Coprinus cinereus. EMBO J. 6:835-840. Biollaz, M., G. Biichi, and G. Milne. 1968. Biosynthesis of aflatoxins: the biogenesis of bisfuranoids in the genus Aspergillus. J. Amer. Chem. Soc. 90:5017-5020. Biollaz, M., G. Biichi, and G. Milne. 1970. The biosynthesis of aflatoxins. J. Amer. Chem. Soc. 92:1035-1043. 61. 62. 63. 65. 66. 67. 68. 69. 70. 71. 72. 215 Birkett, J. A., and J. A. Roper. 1977. Chromosome aberrations in Aspergillus nidulans, pp 293-303. In Smith J. E., and J. A. pateman (eds.), Genetics and physiology of filamentous fungi. Academic Press, Inc., London. Birkett, J. A., and R. T. Rowlands. 1981. Chlorate resistance and nitrate assimilation in industrial strains of Penicillium chrysogenum. J. Gen. Microbiol. 123:281-285. Blakemore. E. J. A., M. J. Dobson, M. J. Hocart, J. A. Lucas, and J. F. Peberdy. 1989. Transformation of Pseudocercosporella herpotrichoides using two heterologous genes. Curr. Genet. 16:177-180. Blaunt, W. P. 1961. Turkey "X" disease. J. Brit. Turkey Federation 9:55-58. Boeke, J. D., F. LaCraute, and G. R. Fink. 1984. A positive selection for mutants lacking oratidine-5’-phosphate decarboxylase activity in yeast: 5-fluoro-or0tic acid resistance. Mol. Gen. Genet. 197:345-346. Borisy, G. G., D. W. Cleveland, and D. G. Murphy. 1984. Molecular biology of the cytoskeleton. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Bas, C. J. 1985. Protoplasts from fungal spores, pp. 73-85. In Peberdy, J. F., and L. Ferenczy (eds.), Fungal protoplasts: applications in biochemistry and genetics. Marcel Dekker, Inc., New York, NY. Bas, C. J., and S. M. Slakhorst. 1981. Isolation of protoplasts from Aspergillus nidulans canidiospares. Can. J. Microbiol. 27:400-407. Bas, C. J., A. J. M. Debets, A. W. van Heusden, and H. T. A. M. Schepers. 1983. Fusion of protoplasts from conidiospores of Aspergillus nidulans. Experientia (Suppl.) 45:298-299. Bothast, R. J., and D. I. Fennell. 1974. A medium for rapid identification and enumeration of Aspergillus flaws and related organisms. Mycologia 66:365-369. Botstein, D., S. C. Falco, S. E. Stewart, M. Brennan, S. Scherer, D. T. Stinchcomb, K. Struhl, and R. W. Davies. 1979. Sterile host yeasts (SHY): a eukaryotic system of biological containment for recombinant DNA experiments. Gene 8:17-24. Boylan, M. T., M. J. Holland, and W. E. Timberlake. 1986. Saccharomyces cerevisiae centromere CENI I does not induce chromosome instability when integrated into the Aspergillus nidulans genome. Mol. Cell. Biol. 6:3621-3625. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 216 Boylan, M. T., P. M. Mirabito, C. E. Willett, C. R. Zimmerman, and W. E. Timberlake. 1987. Isolation and physical characterization of three essential conidiation genes from Aspergillus nidulans. Mol. Cell. Biol. 7:3113-3118. Bradshaw, R. E., J. W. Bennett, and J. F. Peberdy. 1983. Parasexual analysis of Aspergillus parasiticus. J. Gen. Microbiol. 129:2117-2123. Brygoo, Y., and R. Debuchy. 1985. Transformation by integration in Podospora anserina. 1. Methodology and phenomenology. Mol. Gen. Genet. 200:128-131. Bull, J. H., and J. C. Wootton. 1984. Heavily methylated amplified DNA in transformants of Neurospora crassa. Nature 310:701-704. Bull, J. H., D. J. Smith, and G. Turner. 1988. Transformation of Penicillium Chrysogenum with a dorrrinant selectable marker. Curr. Genet. 13:377-382. Bu’Lock, J. D. 1961. Intermediary metabolism and antibiotic synthesis. Adv. Appl. Microbiol. 3:293-342. Bu’Lack, J. D. 1986. Genetic aspects of mycotoxin formation, pp. 1-12. In Kleinkauf, H., H. V. Dohren, H. Domauer, and G. Nesemann (eds.), Regulation of secondary metabolite formation. VCH Verlagsgesellschaft mbH, Weinheim. Bu’Lock, J. D., J. P. Mooney, and C. E. Wright. 1986a. Regulation of mycotoxin production by Fusarium graminearwn: complementation effects between two mutant types. Biotechnol. Lett. 8:323-326. Bu’Lock, J. D., C. E. Wright, and J. E. Mooney. 1986b. Use of a proto- plast fusion test to establish the status of mycotoxin genes in an edible Fusarium. Biotechnol. Lett. 8:621-624. Busby, W. F., and G. N. Wogan. 1981. Aflatoxins, pp. 3-27. In Shank, R. C. (ed.), Myc0toxins and N-nitroso compounds: environmental risks. Vol. II. CRC Press, Boca Raton, Florida. Buxton, F. P., and A. Radford. 1983. Cloning of the structural gene for orotidine 5’-phosphate carboxylase of Neurospora crassa by expression in Escherichia coli. Mol. Gen. Genet. 190:403-405. Buxton, F. P., and A. Radford. 1984. The transformation of mycelial sphero- plasts of Neurospora crassa and the attempted isolation of an autonomous replicator. Mol. Gen. Genet. 196:339-344. 85. 86. 87. 88. 89. 91. 92. 93. 94. 95. 96. 217 Buxton, F. P., D. I. Gwynne, and R. W. Davis. 1985. Transforrrration of Aspergillus niger using the argB gene of Aspergillus nidulans. Gene 37:207-214. Buxton, F. P., D. I. Gwynne, S. Garven, S. Sibley, and R. W. Davis. 1987. Cloning and molecular analysis of the omithine carbamoyl transferase gene of Aspergillus niger. Gene 60:255-266. Cabib, E., B. Bowers, A. Sburlati, and S. J. Silverman. 1988. Fungal cell wall synthesis: the construction of a biological structure. Microbial. Sci. 5:370-375. Cambareri, E. B., B. C. Jensen, E. Schabtach, and E. C. Selker. 1989. Repeat-induced G-C to A-T mutations in Neurospora. Science 244:1571-1575. Campbell, E. 1., S. E. Unkles, J. A. Macro, C. van den Handel, R. Contreras, and J. R. Kinghom. 1989. Improved transformation efficiency of Aspergillus niger using the homologous niaD gen for nitrate reductase. Curr. Genet. 16:53-56. Cantoral, J. M., B. Diez, J. L. Barredo, E. Alvarez, and J. F. Martin. 1987. High-frequency transformation of Penicillium chrysogenum. BiofI‘echnol. 5:494-497. Caprioglio, D. R., and L. W. Parks. 1988. Purification and characterization of plasrrrid-like DNA from the antimycotic producing fungus, Scytalidium flavo-brunneum. Plasmid 20:175-181. Caprioglio, D. R., and L. W. Parks. 1989. Temporal expression of trans- cription and relative copy number of plasmid pSFB-l in Scytalidium flavo-brunneum. J. Bacteriol. 171:4876-4880. Carlock, L. R. 1986. Analyzing lambda libraries. Focus 8(2):6-8. Caroline, D. F. 1969. Pyrirnidine synthesis in Neurospora crassa: gene-enzyme relationships. J. Bacteriol. 100:1371-1377. Carramolino, L., M. Lozano, A. Perez-Aranda, V. Rubia, and F. sanchez. 1989. Transformation of Penicillium chlysogenum to sulfonamide resistance. Gene 77:31-38. Casadaban, M., A. Martinez-Arias, S. Shapina, and J. Chow. 1983. Beta-galactosidase gene fusions for analyzing gene expression in Escherichia coli and yeast. Methods Enzymol. 100B:293-308. 97. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 108. 218 Case, M. E. 1986. Genetical and molecular analyses of QA-Z transformants in Neurospora crassa. Genetics 113:569-587. Case, M. E., M. Schweizer, S. R. Kushner, and N. H. Giles. 1979. Efficient transformation of Neurospora crassa by utilizing hybrid plasmid DNA. Proc. Natl. Acad. Sci. USA 76:5259-5263. Casselton, L. A., and A. de la Fuente Herce. 1989. Heterologous gene expression in the basidiomycete fungus Coprinus cinereus. Curr. Genet. 16:35-40. Castegnaro, M., M. Friesen, J. Michelon, and E. A. Walker. 1981. Problems related to the use of sodium hypochlorite in the detoxification of aflatoxin B,. Amer. Ind. Hyg. Assoc. J. 42:398-401. Castegnaro, M., D. C. Hunt, E. B. Sansone, P. L. Schuller, M. G. Siriwardana, G. M. Telling, H. P. van Egmond, and E. A. Walker. 1980. Laboratory decontamination and destruction of aflatoxins B,. B,, G,, G2 in laboratory wastes. International Agency for Research on Cancer, Lyon. Chang, A. C. Y., and S. N. Cohen. 1978. Construcrion and characterization of amplifiable multicopy DNA cloning vehicles derived from the PISA cryptic rniniplasmid. J. Bacteriol. 134:1141-1156. Chattoo, B. B., F. Sherman, D. A. Azubalis, T. A. Fjellstedt, D. Mehvert, and M. Ogur. 1979. Selection of Iys2 mutants of the yeast Saccharomyces cerevisiae by the utilization of a-amino-adipate. Genetics 93:51-65. Choi, H. T., J. L. Revuelta, C. Sadhu, and M. Jayaram. 1988. Structural organization of the TRP] gene of Phycomyces blakesleeanus: implications for evolutionary gene fusion in fungi. Gene 71:85-95. Christensen, T., H. Woeldike, E. Boel, S. B. Mortensen, K. Hjortshoej, L. Thirn, and M. T. Hansen. 1988. High level expression of recombinant genes in Aspergillus nidulans. Bio/Technol. 6:1419-1422. Chu, S. B., and M. Alexander. 1972. Resistance and susceptibility of fungal spores to lysis. Trans. Br. Mycol. Soc. 58:489-497. Cihlar, R., and P. S. Sypherd. 1980. The organization of the ribosomal RNA genes in the fungus Mucor. Nucleic Acids Res. 8:793-804. Clark, L., and J. Carbon. 1976. A colony bank containing synthetic ColEl hybrid plasmids representative of the entire E. coli genome. Cell 9:91-99. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 219 Clarke, L., and J. Carbon. 1978. Functional expression of cloned yeast DNA in Escherichia coli: specific complementation of argininosuccinate lyase (argH) mutations. J. Mol. Biol. 120:517-532. Cleveland, D. W., and K. F. Sullivan. 1985. Molecular biology and genetics of tubulin. Annu. Rev. Biochem. 54:331-365. Cleveland, T. E. 1989. Conversion of dihydro-O-methylsterigmatocystin to aflatoxin B, by Aspergillus parasiticus. Arch. Environ. Contam. Toxicol. 18:429-433. , Cleveland, T. E., and D. Bhatnagar. 1987. Individual reaction requirements of two enzyme activities, isolated from Aspergillus parasiticus, which together catalyze conversion of sterigmatocystin to aflatoxin B,. Can. J. Microbiol. 33:1108-1112. Cleveland, T. E., and D. Bhatnagar. 1988. Construction of a cDNA library from Aspergillus parasiticus mRN A isolated at the time of first expression of enzymes catalyzing aflatoxin biosynthesis. APS Annu. Meet. Abst. No. 608. Cleveland, T. E., D. Bhatnagar, C. J. Foell, and S. P. McCormick. 1987. Conversion of a new metabolite to aflatoxin B, by Aspergillus parasiticus. Appl. Environ. Microbiol. 53:2804-2807. Cleveland, T. E., A. R. Lax, L. S. Lee, and D. Bhatnagar. 1987. Appear- ance of enzyme activities catalyzing conversion of sterigmatocystin to aflatoxin B, in late-growth-phase Aspergillus parasiticus cultures. Appl. Environ. Microbiol. 53:1711-1713. Clutterbuck, A. J. 1972. Absence of laccase from yellow-spared mutants of Aspergillus nidulans. J. Gen. Microbiol. 70:423-435. Clutterbuck, A. J. 1977. The genetics of conidiation in Aspergillus nidulans, pp. 305-317. In Smith, J. E., and J. A. Pateman (eds.), Genetics and physiology of Aspergillus. Academic Press, Inc., London. Colbere-Garapin, F., F. Horodniceanu, P. Kourilsky, and A. -C. Garapin. 1981. A new dorrrinant hybrid selective marker for higher eukaryotic cells. J. Mol. Biol. 150:1-14. Collis, C. M., and R. M. Hall. 1985. Identification of a Tn5 determinant conferring resistance to phleomycins, bleomycins and tallysomycins. Plasmid 14:143-151. 120. 121. 122. 123. 124. 125. 126. 127. 128. 129. 130. 131. 132. 220 Cooley, R. N., R. K. Shaw, F. C. H. Franklin, and C. E. Caten. 1988. Transformation of the phytopatlrogenic fungus Septoria nodorum to hygromycin B resistance. Curr. Genet. 13:383-389. Correll, J. C., C. J. R. Klittich, and J. F. Leslie. 1987. Nitrate nonutilizing mutants of F usarium oxysporum and their use in vegetative compatibility tests. Phytopathol. 77:1640-1646. Costanzo, M. C., and T. D. Fox. 1988a. Transformation of yeast by agita- tion with glass beads. Genetics 120:667-670. Costanzo, M. C., and T. D. Fox. 1988b. Specific translational activation by nuclear gene products occurs in the 5’ untranslated leader of a yeast mitochondrial mRNA. Proc. Natl. Acad. Sci. USA 85:2677-2681. Cove, D. J. 1976a. Chlorate toxicity in Aspergillus nidulans: studies of mutants altered in nitrate assimilation. Mol. Gen. Genet. 146:147-159. Cove, D. J. 1976b. Chlorate toxicity in Aspergillus nidulans: the selection and characterization of chlorate resistant mutants. Heredity 36:191-203. Cove, D. J. 1979. Genetic studies of nitrate assimilation in Aspergillus nidulans. Biol. Rev. 54:291-327. Crawford, 1. P. 1975. Gene rearrangements in the evolution of the tryp- t0phan synthetic pathway. Bacterial. Rev. 39:87-120. Crawford, 1. P. 1989. Evolution of a biosynthetic pathway: the tryptophan paradigm. Annu. Rev. Microbiol. 43: 567-600. Cullen, D., and S. A. Leong. 1986. Recent advances in the molecular genetics of industrial filamentous fungi. Trends Biotechnol. 42285-288. Cullen, D., G. L. Gray, L. J. Wilson, K. J. Hayenga, M. H. Lamsa, M. W. Rey, S. Norton, and R. M. Berka. 1987. Controlled expression and secretion of bovine chymosin in Aspergillus nidulans. Bio/Technol. 5:369-376. Cullen D, S. A. Leong, L. J. Wilson, and D. J. Henner. 1987. Transfor- mation of Aspergillus nidulans with the hygromycin-resistance gene, hph. Gene 57:21-26. Daboussi, M. J., A. Djeballi, C. Gerlinger, P. L. Blaiseau, Bouvier, I, M. Cassan, M. H. Lebrun, D. Parisot, and Y. Brygoo. 1989. Transformation of seven species of filamentous fungi using the nitrate reductase gene of Aspergillus nidulans. Curr. Genet. 15:453-456. 133. 134. 135. 136. 137. 138. 139. 140. 141. 142. 143. 144. 145. 221 Darlington, A. J., and C. Scazzocchio. 1967. Use of analogues and the substrate-sensitivity of mutants in analysis of purine uptake and breakdown in Aspergillus nidulans. J. Bacteriol. 93:937-940. Das, S., E. Kellerrnann, and C. P. Hollenberg. 1984. Transformation of Kluyveromyces fragilis. J. Bacteriol. 158:1165-1167. Davidse, L. C. 1986. Benzimidazole fungicides: mechanism of action and biological impact. Annu. Rev. Phytopathol. 24:43-65. Davies, J. E., D. Kirkaldy, and J. C. Robert. 1960. Studies in mycological chemistry. Part VII. Sterigmatocystin, a metabolite of Aspergillus versicolor (Vuillemin) Tiraboschi. J. Chem. Soc. 1960:2169-2178. Davis, B. 1985. Factors influencing protoplast isolation, pp. 45-71. In Peberdy, J. F., and L. Ferenczy (eds.), Fungal protoplasts: applications in biochemistry and genetics. Marcel Dekker, Inc., New York, NY. Davis, M. A., C. S. Cabbett, and M. J. Hynes. 1988. An amdS-IacZ fusion for studying gene regulation in Aspergillus. Gene 63:199-212. Davis, M. A., and M. J. Hynes. 1987. Complementation of areA'-regu1atory gene mutations of Aspergillus nidulans by the heterologous regulatory gene nit-2 of Neurospora crassa. Proc. Natl. Acad. Sci. USA. 84:3753-3757. Davis, N. D. 1981. Sterigmatocystin and other mycotoxins produced by Aspergillus species. J. Food Prat. 44:711-714. Davis, N. D., S. K. Iyer, and U. L. Diener. 1987. Improved method of screening for aflatoxin with a coconut agar medium. Appl. Environ. Microbiol. 53:1593-1595. Davis, R. H., and F. J. de Serres. 1970. Genetic and microbiological research techniques for Neurospora crassa. Methods Enzymol. 17:79-143. Davis, R. W., D. Botstein, and J. R. Roth. 1980. Advanced bacterial genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. de Graaff, L., H. van den Braeck, and J. Visser. 1988. Isolation and transformation of the pyruvate kinase gene of Aspergillus nidulans. Curr. Genet. 13:315-321. de Ruiter-Jacobs, Y. M. J. T., M. Broekhuijsen, S. E. Unkles, E. 1. Campbell, J. R. Kinghom. R. Contreras, P. H. Pouwels, and C. A. M. J. J. van den Handel. 1989. A gene transfer system based on the homologous pyrG gene and efficient expression of bacterial genes in Aspergillus oryzae. Curr. Genet. 16:159-163. 146. 147. 148. 149. 150. 151. 152. 153. r54. 155. 156. 157. 222 Dean, P. M. 1963. Naturally occurring oxygen ring compounds. Butter- worths, London. Debuchy, R., E. Cappin-Raynel, D. Le Coze, and Y. Brygoo. 1988. Chro- mosome walking towards a centromere in the filamentous fungus Podospara anserina: cloning of a sequence lethal at a low-copy state. Curr. Genet. 13:105-111. Delorme, E. 1989. Transformation of Saccharomyces cerevisiae by electro- poration. Appl. Environ. Microbiol. 55:2242-2246. Detroy, R. W., E. B. Lillehoj, and A. Ciegler. 1971. Aflatoxins and related compounds, pp 4-178. In Ciegler. A., S. Kadis, and S. J. Ajl (eds), Microbial toxins, Vol. VI. Fungal toxins. Academic Press, Inc., New York, NY. De Vogel, P., R. Van Rhee, and W. A. A. Blanche-Kaelensmid. 1965. A rapid screening test for aflatoxin-synthesizing aspergilli of the flavus-oryzae group. J. Appl. Bacterial. 28:213-220. de Vries, O. M. H., and J. G. H. Wessels. 1972. Release of protoplasts from Schizophyllum commune by a lytic preparation from Trichoderma viride. J. Gen. Microbiol. 73:13-22. Dhawale, S. S., and G. A., Marzluf. 1985. Transformation of Neurospora crassa with circular and linear DNA and analysis of the fate of the transforming DNA. Curr. Genet. 10:205-212. Dhawale, S. S., J. V. Paietta, and G. A. Marzluf. 1984. A new, rapid and efficient transformation procedure for Neurospora. Curr. Genet. 8:77-79. Diallinas, G., and C. Scazzocchio. 1989. A gene coding for the uric acid-xanthine permease of Aspergillus nidulans: inactivational cloning, characterization, and sequence of a cis-acting mutation. Genetics 122:341-350. Dickinson, L., M. Harboe, R. van Heeswijck, P. Stroman, and L. P. Jepsen. 1987. Expression of active Mucor rrriehei aspartic protease in Mucor circinelloides. Carlsberg Res. Commun. 52:243-252. Dickman, M. B. 1988. Whole cell transformation of the alfalfa fungal pathogen Colletotrichum trifolii. Curr. Genet. 14:241-246. Dickman, M. B., and J. F. Leslile. 1989. Regulation of niu'ogen metabolism in Fusarium by a homologous Neurospora gene. APS Annu. Meet. Abst. No. 296. 158. 159. 160. 161. 162. 163. 164. 165. 166. 167. 168. 169. 223 Diener, J. S., and N. D. Davis. 1969. Aflatoxin formation by Aspergillus flavus, pp. 13-54. In Goldblatt, L. A. (ed.), Aflatoxin: scientific background, control and implications. Academic Press, Inc., New York, NY. Diez, B., E. Alvarez, J. M. Cantoral, J. L. Barredo, and J. F. Martin. 1987. Selection and characterization of pyrG mutants of Penicillium chrysogenum lacking orotidine-5’-phosphate decarboxylase and complementation by the pyr4 gene of Neurospora crassa. Curr. Genet. 12:277-282. Donkersloot, J. A., D. P. H. Hsieh, and R. I. Mateles. 1968. Incorporation of precursors into aflatoxin B,. J. Amer. Chem. Soc. 90:5020-5021. Donkersloot, J. A., R. I. Mateles, and S. S. Yang. 1972. Isolation of averufin from a mutant of Aspergillus parasiticus impaired in aflatoxin biosynthesis. Biochem. Biophys. Res. Commun. 47:1051-1055. Domer, J.W., R. J. Cole, and U. L. Diener. 1984. The relationship of Aspergillus flavus and Aspergillus parasiticus with reference to production of aflatoxins and cyclopiazonic acid. Mycopathologia 87:13-15. Day, C. H., J. A. Pateman, J. E. Olsen, H. J. Kane, and E. H. Creaser. 1985. Genomic clones of Aspergillus nidulans containing ach, the structural gene for alcohol dehydrogenase and aIcR, a regulatory gene for ethanol metabolism. DNA 4:105-114. Duncan, J. S., and J. D. Bu’Lock. 1985. Degeneration of zearalenone production in Fusarium graminearum. Exp. Mycol. 9:133-140. Duncan, K., and J. R. Coggins. 1984. Subcloning of the Escherichia coli genes ar05 (5-enolpyruvyl shikimate 3-phosphate synthase) and aroB (3-dehydroquinate synthase). Biochem. Soc. Trans. 12:274-275. Duncan, K., and J. R. Coggins. 1986. The serC-aroA operon of Escherichia coli: a mixed function operon encoding enzymes from two different amino acid biosynthetic pathways. Biochem. J. 234:49-57. Dunne, P. W., and B. R. Oakley. 1988. Mitotic gene conversion, reciprocal recombination and gene replacement at the benA, beta-tubulin, locus of Aspergillus nidulans. Mol. Gen. Genet. 213:339-345. Dutton, M. F. 1988. Enzymes and aflatoxin biosynthesis. Microbiol. Rev. 52:274-295. Dutton, M. F., and M. S. Anderson. 1978. The use of fungal protoplasts in the study of aflatoxin biosynthesis. Experientia 34:22-24. 170. 171. 172. 173. 174. 175. 176. 177. 178. 179. 180. 181. 182. 224 Dutton, M. F., and M. S. Anderson. 1982. Role of versicolorin A and its derivatives in aflatoxin biosynthesis. Appl. Environ. Microbiol. 43:548-551. Dutton, M. F., K. Ehrlich, and J. W. Bennett. 1985. Biosynthetic rela- tionship among aflatoxins B,, B,, M,, and M,. Appl. Environ. Microbiol. 49:1392-1395. Emerson, S., and M. R. Emerson. 1958. Production, reproduction and rever- sion of protoplastlike structures in the osmotic strain of Neurospora crassa. Proc. Natl. Acad. Sci. USA 44:668-671. Enns, R. K., and R. S. Criddle. 1977. Affinity labelling of yeast mito- chondrial adenosine triphosphatase by reduction with [’H]borohybride. Arch. Biochem. Biophys. 182:587-600. Esser, K., and G. Mohr. 1986. Integrative transformation of filamentous fungi with respect to biotechnological application. Process Biochem. 21:153-159. Farkas, V. 1985. The fungal cell wall, pp. 3-29. In Peberdy, J. F., and L. Ferenczy (eds.), Fungal pr0toplasts: application in biochemisu'y and genetics. Marcel Dekker, Inc., New York, NY. Farman, M. L., and R. P. Oliver. 1988. The transformation of protoplasts of Leprosphaeria maculans to hygromycin B resistance. Curr. Genet. 13:327-330. Faugeron, G., C. Goyon, and A. Gregoire. 1989. Stable allele replacement and unstable non-homologous integration events during transformation of Ascobolus immersus. Gene 76:109-119. Feinberg, A. P., and B. Vogelstein. 1983. A technique for radiolabeling DNA restriction endonuclease fragments to high specific activity. Anal. Biochem. 132:6-13. Feinberg, A. P., and B. Vogelstein. 1984. A technique for radiolabeling DNA restriction endonuclease fragments to high specific activity. Addendum. Anal. Biochem. 137:266-267. Felenbok, B., D. Sequeval, M. Mathieu, S. Sibley, D. I. Gwynne, and R. W. Davies. 1988. The ethanol regulon in Aspergillus nidulans: characterization and sequence of the positive regulatory gene aIcR. Gene 73:385-396. Fernandez-Lama, J., and U. Stahl. 1989. Transformation of Podospora anserina with a dominant resistance gene. Curr. Genet. 16:57-60. Fincham, J. 1988. Hazards in sexual transformation. Nature 331:207-208. 183. 184. 185. 186. 187. 188. 189. 190. 191. 192. 193. 194. 225 Fincham, J. R. S. 1989. Transformation in fungi. Microbiol. Rev. 53:148-170. Fincham, J. R. S., 1. F. Connerton, E. Notarianni, and K. Harrington. 1989. Premeiotic disruption of duplicated and triplicated copies of the Neurospora crassa am (glutamate dehydrogenase) gene. Curr. Genet. 15:327-334. Fischbach, H., and A. D. Campbell. 1965. Note on detoxification of the aflatoxins. J. Assoc. Off. Anal. Chem. 48:28. Floyd, J. C., and J. W. Bennett. 1981. Preparation of “C-labeled aflatoxins and incorporation of unlabeled aflatoxins in a blocked versicolorin A- accumulating mutant of Aspergillus parasiticus. J. Amer. Oil Chem. Soc. 58:956A-959A. Floyd, J. C., J. C. 111., Mills, and J. W. Bennett. 1987. Biotransformation of sterigmatocystin and absence of aflatoxin biotransformation by blocked mutants of Aspergillus parasiticus. Exp. Mycol. 11:109-114. Fotheringham, S., and W. K. Holloman. 1989. Cloning and disruption of Ustilago maydis genes. Mol. Cell. Biol. 9:4052-4055. Foumey, R. M., J. Miyakoshi, R. S. 111., Day, and M. C. Paterson. 1988. Northern blotting: efficient RNA staining and transfer. Focus 10:5-7. Fox, T. D., J. C. Sanford, and T. W. McMullin. 1988. Plasmids can stably transform yeast mitochondria lacking endogenous mtDNA. Proc. Natl. Acad. Sci. USA 85:7288-7292. Fraley, R., and D. Papahadjopoulos. 1981. New generation liposomes: the engineering of an efficient vehicle for intracellular delivery of nucleic acids. Trends Biochem. Sci. 6:77-80. Frederick, G. D., D. K. Asch, and J. A. Kinsey. 1989. Use of transfor- mation to make targeted sequence alterations at the am (GDH) locus of Neurospora. Mol. Gen. Genet. 217:294-300. Frischauf, A. -M., H. Lehrach, A. Poustka, and N. Murray. 1983. Lambda replacement vectors carrying polylinker sequences. J. Mol. Biol. 170:827-842. Fromm, M., J. Gallis, L. P. Taylor, and V. Walbot. 1987. Electroporation of DNA and RNA into plant prot0plasts. Methods Enzymol. 153:351-366. 195. 196. 197. 198. 199. 200. 201. 202. 203. 204. 205. 206. 226 Fu, Y. -H., and G. A. Marzluf. 1987. Molecular cloning and analysis of the regulation of nit-3, the structural gene for nitrate reductase in Neurospora crassa. Proc. Natl. Acad. Sci. USA 84:8243-8247. Garber, R. C., J. J. Lin, and O. C. Yoder. 1986. Mitochondrial plasmids in Cochliobolus heterostrophus. PP 105-118. In Wickner, R. B., A. Hinnebusch, A. M. Lambowitz, I. C. Gunsalus, and A. Hollander (eds.), Extrachromosomal elements in lower eukaryotes. Plenum Press, New York. Garcia Acha, 1., and J. R. Villanueva. 1963. Differences in the mode of aetion of strepzyme and Helix pomatia enzyme preparations on Tricho- teciwn roseum spores. Nature 200:1231. Garcia Acha, 1., and J. R. Villanueva. 1964. ProtOplasts from conidia of Fusarium culmorum. Can. J. Microbiol. 10:99-101. Garcia Acha, 1., F. Lopez-Belmonte, and J. R. Villanueva. 1966. Preparation of protoplast-like structures from conidia of F usariwn culmorum. Antonie van Leeuwenhoek J. Microbiol. Serol. 32:299-311. Gatignol, A., M. Baron, and G. Trraby. 1987. Phleomycin resistance encoded by the ble gene from transposon Tn5 as a dominant selectable marker in Saccharomyces cerevisiae. Mol. Gen. Genet. 207:342-348. Geisen, R., and L. Leistner. 1989. Transformation of Penicillium nalgiovense with the amdS gene of Aspergillus nidulans. Curr. Genet. 15:307-309. Glazebrook, J. A., K. Mitchell, and A. Radford. 1987. Molecular genetic analysis of the pyr-4 gene of Neurospora crassa. Mol. Gen. Genet. 209:399-402. Gold, S. E., and N. T. Keen. 1989. Cloning and sequence analysis of gene(s) encoding B-tubulin in the Hemiascomycete plant pathogen Geotrichum candidum. APS Annu. Meet. Abst. No. 548. Goldblatt, L. A. 1969. Aflatoxin: scientific background, control and implications. Academic Press, Inc., New York, NY. Gomi, K., Y. Iimura, and S. Hara. 1987. Integrative transformation of Aspergillus oryzae with a plasmid containing the Aspergillus nidulans argB gene. Agr. Biol. Chem. 51:2549-2555. Goosen, T., G. Bloemheuvel, C. Gysler, D. A. de Bie, H. W. J. van den Broek, and K. Swart. 1987. Transformation of Aspergillus niger using the homologous orotidine-5'-phosphate decarboxylase gene. Curr. Genet. 11:499-503. 207. 208. 209. 210. 211. 212. 213. 214. 215. 216. 217. 227 Goyon, C., and G. Faugeron. 1989. Targeted transformation of Ascobolus immersus and de novo methylation of the resulting duplicated DNA sequences. Mol. Cell. Biol. 9:2818-2827. Grant, D. M., A. M. Lambowitz, J. A. Rambosek, and J. A. Kinsey. 1984. Transformation of Neurospora crassa with recombinant plasmids containing the cloned glutamate dehydrogenase (am) gene: evidence for autonomous replication of the transforming plasmid. Mol. Cell. Biol. 4:2041-2051. Gray, G. L., K. Hayenga, D. Cullen, L. J. Wilson, and S. Norton. 1986. Primary structure of Mucor miehei aspartyl protease: evidence for a zymogen intermediate. Gene 48:41-53. Griffin, D. H., M. DeVit, and R. Tuori. 1989. Protoplast formation and transformation of Hypoxylon mammatum. APS Annu. Meet. Abst. No. 549. Grieg. G. W., R. M. Hall, N. K. Hart, D. R. Havulak, J. A. Lamberton, and A. Lane. 1985. Amplification of the antibiotic. Effects of the bleomycins, phleomycins and tallysomycins: its dependence on the nature of the variable basic groups. J. Antibiot. 38:99-110. Groopman, J. D., L. G. Cain, and T. W. Kensler. 1988. Aflatoxin exposure in human populations: measurements and relationship to cancer. CRC Crit. Rev. Toxicol. 19:113-145. Gunasekaran, M. 1981. Optimum culture conditions for aflatoxin B, pro- duction by a human pathogenic strain of Aspergillus flavus. Mycologia 73:697-704. Gussack, G., J. W. Bennett, S. Cavalier, and L. Yatsu. 1977. Evidence for the parasexual cycle in a strain of Aspergillus flavus containing virus-like particles. Mycopathologia 61:159-165. Gwynne, D. I., F. P. Buxton, S. Sibley, R. W. Davies, R. A. Lockington, C. Scazzocchio, and H. M. Sealy-Lewis. 1987a. Comparison of the cis-acting control regions of two coordinately controlled genes involved in ethanol utilization in Aspergillus nidulans. Gene 51:205-216. Gwynne, D. 1., F. P. Buxton, S. A. Williams, S. Garven, and R. W. Davies. 1987b. Genetically engineered secretion of active human interferon and a bacterial endoglucanase from Aspergillus nidulans. Bio/Technol. 5:713-719. Gwynne, D. 1., F. P. Buxton, S. Williams, A. M. Sills, J. A. Johnstone, J. K. Buch, Z. -M. Guo, D. Drake, M. Westphal, and R. W. Davies. 1989. Development of an expression system in Aspergillus nidulans. Biochem. Soc. Trans. 17:338-340. 218. 219. 220. 221. 222. 223. 224. 225. 226. 227. 228. 229. 230. 228 Hahm, Y. T., and C. A. Batt. 1988. Genetic transformation of an argB mutant of Aspergillus oryzae. Appl. Environ. Microbiol. 54:1610-1611. Harnasaki, T., T. Nakagorrri, Y. Hatsuda, K. Fukuykama, and Y. Katsube. 1977. 5,6-Dimethylsterigmatocystin, a new metabolite from Aspergillus multicolor. Tetrahedron Lett. 32:2765-2766. Hamer, J. E., and W. E. Timberlake. 1987. Functional organization of the Aspergillus nidulans trpC promoter. Mol. Cell. Biol. 7:2352-2359. Hamlyn, P. F., R. E. Bradshaw, F. M. Mellon, C. M. Santiago, J. M. Wilson, and J. F. Peberdy. 1981. Efficient protoplast isolation from fungi using commercial enzymes. Enzyme Microb. Technol. 3:321-325. Hanahan, D. 1983. Studies on transformation of Escherichia coli with plasmids. J. Mol. Biol. 166:557-580. Hara, S., D. I. Fennell, and C. W. Hesseltine. 1974. Aflatoxin-producing strains of Aspergillus flavus detected by fluorescence of agar medium under ultraviolet light. Appl. Microbiol. 27:1118-1123. Harkki, A., J. Uusitalo, M. Bailey, M. Penttilit, and J. K. C. Knowles. 1989. A novel fungal expression system: secretion of active calf chymosin from the filamentous fungus Trichoderma reesei. Bio/Technol. 7:596-603. Harris, G. S., E. J. Keath, and J. Medoff. 1989. Characterization of alpha and beta tubulin genes in the dimorphic fungus Histoplasma capsulatum. J. Gen. Microbiol. 135:1817-1832. Hearn, V. M., E. V. Wilson, and D. W. R. Mackenzie. 1980. The prepara- tion of protoplasts from Aspergillus fumigatus mycelium. Sabouraudia 18:75-77. Heathcote, J. G., and J. R. Hibbert. 1978. Aflatoxins: chemical and biological aspects. Elsevier Science Publishers, Amsterdam. Henderberg, A., J. W. Bennett, and L. S. Lee. 1988. Biosynthetic origin of aflatoxin G,: confirmation of sterigmatocystin and lack of confirmation of aflatoxin B, as precursors. J. Gen. Microbiol. 134:661-667. Henson, J. M., N. K. Blake, and A. L. Pilgeram. 1988. Transformation of Gaeumannomyces graminis to benomyl resistance. Curr. Genet. 14:113-117. Hertzberg, R. P., M. J. Caranfa, and S. M. Hecht. 1985. DNA methylation diminishes bleomycins-mediated strand scission. Biochemistry 24:5285-5289. 231. 232. 233. 234. 235. 236. 237. 238. 239. 240. 241. 242. 229 Hesseltine. C. W., W. G. Sorenson, and M. Smith. 1970. Taxonomic studies of the aflatoxin-producing strains in the Aspergillus flavus group. Mycologia 62:123-132. Hinnen, A., J. B. Hicks, and G. R. Fink. 1978. Transformation of yeast chimeric ColEl plasmid carrying LEUZ. Proc. Natl. Acad. Sci. USA 75:1929-1933. Hiraoka, Y., T. Toda, and M. Yanagida. 1984. The NDA3 gene of fission yeast encodes B-tubulin: a cold sensitive nda3 mutation reversibly blocks spindle formation and chromosome movement in mitosis. Cell 39:349-358. Hirth, K. -P., C. A. Edwards, and R. A. Firtel. 1982. A DNA-mediated transformation system for Dictyostelium discoidewn. Proc. Natl. Acad. Sci. USA 79:7356-7360. Holden, D. W., J. Wang, and S. A. Leong. 1988. DNA-mediated transfor- mation of Ustilago hordei and Ustilago nigra. Physial. Mol. Plant Pathol. 33:235-239. Holzapfel, C. W., I. F. H. Purchase, P. S. Steyn, and L. Gouws. 1966. The toxicity and cherrrical assay of sterigmatocystin, a carcinogenic myc0toxin, and its isolation from two new fungal sources. S. A. Medical J. 40:1100-1101. . Homg, J. S., J. E. Linz, and J. J. Pestka. 1989. Cloning and charac- terization of the trpC gene from an aflatoxigenic strain of Aspergillus parasiticus. Appl. Environ. Microbiol. 55:2561-2568. Hsiao, C. -L., and J. Carbon. 1979. High-frequency transformation of yeast by plasmids containing the cloned yeast ARG4 gene. Proc. Natl. Acad. Sci. USA 76:3829-3833. Hsieh, D. P. H. 1973. Inhibition of aflatoxin biosynthesis of dichlorvos. J. Agric. Food Chem. 21:468-470. Hsieh, D. P. H. 1986. The role of aflatoxin in human cancer, pp. 447-456. In Steyn, P. S., and R. Uleggar (eds.), Mycotoxins and phycotoxins. Elsevier Science Publishers, Amsterdam. Hsieh, D. P. H. 1989. Carcinogenic potential of mycotoxins in foods, pp. 11-30. In Taylor, S. L., and Scanlan, R. A. (eds.), Food toxicology: a perspective on the relative risks. Marcel Dekker, Inc., New York. Hsieh, D. P. H., and R. I. Mateles. 1970. The relative contribution of acetate and glucose to aflatoxin biosynthesis. Biochim. Biophys. Acta 208:482-486. 243. 244. 245. 246. 247. 248. 249. 250. 251. 252. 253. 254. 255. 230 Hsieh, D. P. H., and R. I. Mateles. 1971. Preparation of labeled aflatoxins with high specific activities. Appl. Microbiol. 22:79-83. Hsieh, D. P. H., M. T. Lin, and R. C. Yao. 1973. Conversion of sterig- matocystin to aflatoxin B, by Aspergillus parasiticus. Biochem. Biophys. Res. Conunun. 52:992-997. Hsieh, D. P. H., M. T. Lin, R. C. Yao, and R. Singh. 1976. Biosynthesis of aflatoxin: conversion of norsolorinic acid and other hypothetical inter- mediates into aflatoxin B,. J. Agr. Food Chem. 24:1170-1174. Hsieh, D. P. H., R. Singh, R. C. Yao, and J. W. Bennett. 1978. Anthra- quinones in the biosynthesis of sterigmatocystin by Aspergillus versicolor. Appl. Environ. Microbiol. 35:980-982. Huang, D., S. Bhairi, and R. C. Staples. 1989. A transformation procedure for Botryotinia squamosa. Curr. Genet. 15:411-414. Huge-Jensen, B., F. Andreasen, T. Christensen, M. Christensen, L. Thim, and E. Boel. 1989. Rhizomucor miehei triglyceride lipase is processed and secreted from transformed Aspergillus oryzae. Lipids 24:781-785. Hiltter, R., P. Niederberger, and J. A. DeMoss. 1986. Tryptaphan biosyn- thetic genes in eukaryotic microorganisms. Annu. Rev. Microbiol. 40:55-77. Hynes, M. J. 1979. Fine su'ucture mapping of the acetamidase structural gene and its controlling region in Aspergillus nidulans. Genetics 91:381-392. Hynes, M. J. 1986. Transformation of filamentous fungi. Exp. Mycol. 10:1-8. Hynes, M. J. 1989. Complementation of an Aspergillus nidulans mutation by a gene from the basidiomycete Coprinus cinereus. Exp. Mycol. 13:196-198. Hynes, M. J., and M. A. Davis. 1986. The amdS gene of Aspergillus nidulans: control by multiple regulatory signals. BioAssays 5:123-128. Hynes, M. J., C. M. Conick, J. M. Kelly, and T. G. Littlejohn. 1988. Identification of the sites of action for regulatory genes controlling the amdS gene of Aspergillus nidulans. Mol. Cell. Biol. 8:2589-2596. Hynes, M. J., C. M. Conick, and J. A. King. 1983. Isolation of genomic clones containing the amdS gene of Aspergillus nidulans and their use in the analysis of structural and regulatory mutations. Mol. Cell. Biol. 3:1430-1439. 256. 257. 258. 259. 260. 261. 262. 263. 264. 265. 266. 267. 268. 231 Iimura, K., K. Gorrri, and H. Uzu. 1987. Transformation of Aspergillus oryzae through plasmid-mediated complerrrentation of the methionine- auxotrophic mutation. Agr. Biol. Chem. 51:323-328. Iimura, Y., K. Gotoh, K. Ouchi, and T. Nishima. 1983. Transformation of yeast without the spheroplasting process. Agr. Biol. Chem. 47:897-901. Innis, M. A., M. J. Holland, P. C. McCabe, G. E. Cole, V. P. Wittman, R. Tal, W. K. Watt, D. H. Gelfand, J. P. Holland, and J. H. Meade. 1985. Expression, glycosylatian, and secretion of an Aspergillus glucoamylase by Saccharomyces cerevisiae. Science 228:21-26. Ito, H., Y. Fukuda, K. Murata, and A. Kimura. 1983. Transformation of intact yeast cells treated with alkali cations. J. Bacteriol. 153:163-168. Jeenah, M. S., and M. F. Dutton. 1983. The conversion of sterigmatocystin to O-methylsterigmatocystin and aflatoxin B, by a cell-free preparation. Biochem. Biophys. Res. Commun. 116:1114-1118. Jelinek, C. F., A. E. Pohland, and G. E. Wood. 1989. Worldwide occurr- ence of mycotoxins in foods and feeds- an update. J. Assoc. Off. Anal. Chem. 72:223-230. Jimenez, A., and J. Davies. 1980. Expression of a transposable antibiotic resistance element in Saccharomyces. Nature 287:869-871. John, M. A., and J. F. Peberdy. 1984. Transformation of Aspergillus nidulans using the argB gene. Enzyme Microb. Technol. 6:386-389. Johnston, S. A., P. Q. Anziano, K. Shark, J. C. Sanford, and R. A. Butow. 1988. Mitochondrial transformation in yeast by bombardment with rrricro- projectiles. Science 240:1538-1541. Johnstone, I. L. 1985. Transformation of Aspergillus nidulans. Microbial. Sci. 2:307-311. Johnstone, 1. L., and A. J. Clutterbuck. 1986. An Aspergillus DNA sequence giving a high frequency of unstable transformants. Heredity 57:131. Johnstone, I. L., S. G. Hughes, and A. J. Clutterbuck. 1985. Cloning an Aspergillus nidulans developmental gene by transformation. EMBO J. 4:1307-1311. Jones, I. G., and H. M. Sealy-Lewis. 1989. Chromosomal mapping and gene disruption of the ADI-III! gene in Aspergillus nidulans. Curr. Genet. 15:135-142. 269. 270. 271. 272. 273. 274. 275. 276. 277. 278. 279. 280. 281. 232 Jorgensen, R. A., S. J. Rothstein, and W. S. Reznikoff. 1979. ‘A restriction enzyme cleavage map of Tn5 and location of a region encoding neomycin resistance. Mol. Gen. Genet. 177:65-72. Kafer, E. 1977. The anthranilate synthetase enzyme complex and the tri- functional trpC gene of Aspergillus. Can. J. Genet. Cytol. 19:723-738. Kaiser, K., and N. E. Murray. 1985. The use of phage lambda replacement vectors in the construction of representative genomic DNA libraries, pp. 1-47. In Glover, D. M. (ed), DNA cloning, a practical approach, Vol. I. IRL Press, Ltd., Oxford. Kaster, K. R., S. G. Burgett, and T. D. Ingolia. 1984. Hygromycin B resistance as dominant selectable marker in yeast. Curr. Genet. 8:353-358. Kaster, K. R., S. G. Burgett, R. N. Rao, and T. D. Ingolia. 1983. Analysis of a bacterial hygromycin B resistance gene by transcriptional and trans- lational fusions and by DNA sequencing. Nucleic Acids Res. 11:6895-6911. Katz, M. E., and M. J. Hynes. 1989a. Characterization of the ade- controlled lamA and lamB genes of Aspergillus nidulans. Genetics 122:331-339. Katz, M. E., and M. J. Hynes. 1989b. Gene function identified by inter- specific transformation. Gene 78:167-171. Katz, M. E., and M. J. Hynes. 1989c. Isolation and analysis of the acetate regulatory gene, facB, from Aspergillus nidulans. Mol. Cell. Biol. 9:5696- 5701. Keesey, J. K., Jr., and J. A. DeMoss. 1982. Cloning of the tip-1 gene from Neurospora crassa by complementation of a trpC mutation in Escherichia coli. J. Bacteriol. 152:954-958. Kelly, J. M., and M. J. Hynes. 1985. Transformation of Aspergillus niger by the amdS gene of Aspergillus nidulans. EMBO J. 4:475-479. Kelly, J. M., and M. J. Hynes. 1987. Multiple copies of the amdS gene of Aspergillus nidulans cause titration of ‘ trans-acting regulatory proteins. Curr. Genet. 12:21-31. Kiessling, K. -H. 1986. Biochemical mechanism of action of mycotoxins. Pure Appl. Chem. 58:327-338. Kim, S. Y., and G. A. Marzluf. 1988. Transformation of Neurospora crassa with the trp-I gene and the effect of host strain upon the fate of the transfomring DNA. Curr. Genet. 13:65-70. 282. 283. 284. 285. 286. 287. 288. 289. 290. 291. 292. 233 Kinghom. J. R., and E. 1. Campbell. 1989. Amino acid sequence relation- ships between bacterial, fungal, and plant nitrate reductase and nitrite reductase proteins, pp. 385-404. In Wray, J. L., and J. R. Kinghom (eds.), Molecular and genetic aspects of nitrate assimilation. Oxford Science Publications, Oxford. Kinnaird, J. H., M. A. Keighren, J. A. Kinsey, M. Eaton, and J. R. S. Fincham. 1982. Cloning of the am (glutamate dehydrogenase) gene of Neurospora crassa through the use of a synthetic DNA probe. Gene 20:387-396. Kinsey, J. A. 1989. Restricted distribution of the Tad transponson in strains of Neurospora. Curr. Genet. 15:271-275. Kinsey, J. A., and J. Helber. 1989. Isolation of a transposable element from Neurospora crassa. Proc. Natl. Acad. Sci. USA 86:1929-1933. Kinsey, J. A., and J. A. Rambosek. 1984. Transformation of Neurospora crassa with the cloned am (glutamate dehydrogenase) gene. Mol. Cell. Biol. 4:117-122. Kistler, H. C., and U. K. Benny. 1988. Genetic transformation of the fungal plant wilt pathogen, Fusarium oxysporum. Curr. Genet. 13:145-149. Klein, T. M., E. D. Wolf, R. Wu, and J. C. Sanford. 1987. High-velocity microprojectiles for delivering nucleic acids into living cells. Nature 327:70-73. Klinch, M. A., and Pitt, J. I. 1985. The theory and practice of distin- guishing species of the Aspergillus flavus group, pp. 211-220. In Samson, R. A., and J. I. Pitt (eds.), Advances in Penicillium and Aspergillus systematics. Plenum Press, New York. Klittich, C. J. R., and J. F. Leslie. 1988. Nitrate reduction mutants of Fusarium moniliforme (Gibberella fujikuroi). Genetics 118:417-423. Knight, D. E., and M. C. Scrutton. 1986. Gaining access to the cytosol: the technique and some applications of elecuopermeabilization. Biochem. J. 234:496-507. Kolar, M., P. J. Punt, C. A. M. J. J. van den Handel, and H. Schwab. 1988. Transformation of Penicillium chrysogenum using dominant selection markers and expression of an Escherichia coli lacZ fusion gene. Gene 62:127-134. 293. 294. 295. 296. 297. 298. 299. 300. 301. 302. 303. 304. 234 Kos, A., J. Kuijvenhoven, K. Wernars, C. J. Bas, H. W. J. van den Broek, P. H. Pouwels, and C. A. M. J. J. van den Handel. 1985. Isolation and characterization of the Aspergillus niger trpC gene. Gene 39:231-238. Kos, T., A. Kuijvenhoven, H. G. M. Hessing, P. H. Pouwels, and C. A. M. J. J. van den Hondel.1988.Nucle0tide sequence of the Aspergillus niger trpC gene: structural relationship with analogous genes of other organisms. Curr. Genet. 13: 137- 144. Kronstad, J. W., and S. A. Leong. 1989. Isolation of two alleles of the b locus of Ustilago maydis. Proc. Natl. Acad. Sci. USA 86:978-982. Kronstad, J. W., J. Wang, S. F. Covert, D. W. Holden, G. L. McKnight, and S. A. Leong. 1989. Isolation of metabolic genes and demonstration of gene disruption in the phytopathogenic fungus Ustilago maydis. Gene 79:97-106. Kross, J., W. D. Henner. S. M. Hecht. and W. A. Haseltine. 1982. Speci- ficity of deoxyribonucleic acid cleavage by bleomycin, phleomycin and tallysomycin. Biochemistry 21:4310-4318. Krurrrlauf, R., and G. A. Marzluf. 1979. Characterization of the sequence complexity and organization of the Neurospora crassa genome. Bioche- mistry 18:3705-3713. Kilck, U. 1989. Mitochondrial DNA rearrangements in Podospora anserina. Exp. Mycol. 13:111-120. Kunze, G., R. Bode, H. Rintala, and J. Hofemeister. 1989. Heterologous gene expression of the glyphosate resistance marker and its application in yeast transformation. Curr. Genet. 15:91-98. Kurtzman, C. P., B. W. Horn, and C. W. Hesseltine. 1987. Aspergillus nomius, a new aflatoxin-producing species related to Aspergillus flaws and Aspergillus tamarii. Antonie van Leeuwenhoek 53:147-158. Kurtzman, C. P., M. J. Smiley, C. J. Robnett, and D. T. Wicklow. 1986. DNA relatedness among wild and domesticated species in the Aspergillus flavus group. Mycologia 78:955-959. Laborda, F., I. Garcia Acha, and J. R. Villanueva. 1974. Studies on a strepzyme capable of obtaining protoplasts from Fusarium culmorum conidia. Trans. Br. Mycol. Soc. 62:509-518. Larimer, F. W., C. C. Morse, A. K. Beck, K. W. Cole, and F. H. Gaertner. 1983. Isolation of the ARC] cluster gene of Saccharomyces cerevisiae. Mol. Cell. Biol. 3:1609-1614. 305. 306. 307. 308. 309. 310. 311. 312. 313. 314. 315. 316. 235 Last, R. L., and G. R. Fink. 1988. Tryptaphan-requiring mutants of the plant Arabidopsis thaliana. Science 240:305-310. Lazdins, I. B., and D. J. Cummings. 1982. Autonomously replicating sequences in young and senescent mitochondrial DNA from Podospora anserina. Curr. Genet. 6:173-178. Leaich, L. L., and K. E. Papa. 1975. Identification of diploids of Aspergillus flavus by the nuclear condition of conidia. Mycologia 67:674-678. Le Chevanton, L., and G. Leblon. 1989. The ura5 gene of the ascomycete Sordaria macrospora: molecular cloning, characterization and expression in Escherichia coli. Gene 77:39-49. Le Chevanton, L., G. Leblon, and S. Lebilcot. 1989. Duplications created by transformation in Sordaria macrospora are not inactivated during meiosis. Mol. Gen. Genet. 218:390-396. Lee, L. S., J. W. Bennett, A. F. Cucullu. and R. L. Ory. 1976. Biosynthesis of aflatoxin B,: conversion of versicolorin A to aflatoxin B, by Aspergillus parasiticus. J. Agr. Food Chem. 24:1167-1170. Lee, L. S., J. W. Bennett, A. F. Cucullu. and J. B. Stanley. 1975. Synthesis of versicolorin A by a mutant strain of Aspergillus parasiticus deficient in aflatoxin production. J. Agr. Food Chem. 23:1132-1134. Lee, L. S., J. W. Bennett, L. A. Goldblatt, and R. E. Lundin. 1970. Norso- lorinic acid from a mutant main of Aspergillus parasiticus. J. Amer. Oil Chem. Soc. 48:93-94. Lemke, P. L., N. D. Davis, and G. W. Creech. 1989. Direct visual detec- tion of aflatoxin synthesis by rrrinicolonies of Aspergillus species. Appl. Environ. Microbiol. 55:1808-1810. Lemke, P. A., N. D. Davis, S. K. Iyer, G. W. Creech. and U. L. Diener. 1988. Fluorometric analysis of iodinated aflatoxin in minicultures of Aspergillus flavus and Aspergillus parasiticus. J. Indust. Microbial. 3:119- 125. Lennox, J. E., and C. K. Davis. 1983. Selection of and complementation analysis among aflatoxin-deficient mutants of Aspergillus parasiticus. Exp. Mycol. 7:192-195. Leong, S. A. 1988. Recombinant DNA research in phytopathogenic fungi. Adv. Plant Pathol. 6:1-26. 317. 318. 319. 320. 321. 322. 323. 324. 325. 326. 327. 328. 236 Leslie, J. F., and M. B. Dickman. 1989. Heritability of hygromycin resistance in transformed strains of Fusarium moniliforme. APS Annu. Meet. Abst. No. 408. Lin, M. T., and J. C. Dianese. 1976. A coconut-agar medium for rapid detection of aflatoxin production by Aspergillus spp. Phytopath. 66:1466-1469. Lin, M. T., D. P. H. Hsieh, R. C. Yao, and J. A. Donkersloot. 1973. Conversion of averufin into aflatoxins by Aspergillus parasiticus. Biochem. 12:5167-5171. Lockington, R. A., H. M. Sealy-Lewis, C. Scazzocchio, and R. W. Davies. 1985. Cloning and characterization of the ethanol utilization regulon in Aspergillus nidulans. Gene 33:137-149. Madhosingh, C., and W. Orr. 1985. Zearalenone production in Fusariwn culmorum after transformation with DNA of F. graminearum. Plant. Pathol. 34:402-407. Maggon, K. K., S. K. Gupta, and T. A. Venkitasubramanian. 1977. Biosyn- thesis of aflatoxins. Bacterial. Rev. 41:822-855. Malardier, L., M. J. Daboussi, J. Julien, F. Roussel, C. Scazzocchio, and Y. Brygoo. 1989. Cloning of the niu'ate reductase gene (niaD) of Aspergillus nidulans and its use for transformation of Fusarium oxysporum. Gene 78:147-156. Malpartida, F., M. Zalacain, A. Jimenez, and J. Davies. 1983. Molecular cloning and expression in Streptomyces Iividans of a hygromycin B phosphotransferase gene from Streptomyces hygroscopicus. Biochem. Biophys. Res. Commun. 117:6-11. Manavathu, E. K., K. Suryanarayana, S. E. Hasnain, M. Leung, Y. F. Lau, and W. -C. Leung. 1987. Expression of Herpes simplex virus thymidine kinase gene in aquatic filamentous fungus Achlya ambisexualis. Gene 57:53-59. Manavathu, E. K., K. Suryanarayana, S. E. Hasnain, and W. -C. Leung. 1988. DNA-mediated transformation in the aquatic filamentous fungus Achlya ambisexualis. J. Gen. Microbiol. 134:2019-2028. Maniatis, T., E. F. Fritsch, and J. Sambrook. 1982. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Maramatsu, M. 1973. Preparation of RNA from animal cells. Methods Cell Biol. 7:23-51. 329. 330. 331. 332. 333. 334. 335. 336. 337. 338. 339. 237 Marek, E. T., C. L. Schardl, and D. A. Smith. 1989. Molecular transfor- mation of F usarium solani with an antibiotic resistance marker having no fungal DNA homology. Curr. Genet. 15:421-428. Marzluf, G. A. 1981. Regulation of nitrogen metabolism and gene expre- ssion in fungi. Microbiol. Rev. 45:437-461. Marzluf, G. A., K. G. Perrine, and B. H. Hahm. 1985. Genetic regulation of nitrogen metabolism in Neurospora crassa, pp 83-94. In W. E. Timberlake (ed.), Molecular genetics of filamentous fungi. Alan R. Liss, Inc., New York, NY. Mattem, I. E., S. Unkles, J. R. Kinghom. P. H. Pouwels, and C. A. M. J. J. van den Handel, 1987. Transformation of Aspergillus oryzae using the A. niger pyrG gene. Mol. Gen. Genet. 210:460-461. May, G. S., J. Gambino, J. A. Weatherbee, and N. R. Morris. 1985. Identi- fication and functional analysis of beta-tubulin genes by site specific integrative transformation in Aspergillus nidulans. J. Cell. Biol. 101:712-719. Mayne, R. Y., J. W. Bennett, and J. Tallant. 1971. Instability of an aflatoxin-producing strain of Aspergillus parasiticus. Mycologia 63:644-648. McComrick, S. P., D. Bhatnagar, and L. S. Lee. 1986. Averufanin: an aflatoxin B, precursor between averantin and averufin in the biosynthetic pathway. App. Environ. Microbiol. 53214-16. McCormick, S. P., D. Bhatnagar, and L. S. Lee. 1987. Averufanin is an aflatoxin B, precursor between averantin and averufin in the biosynthetic pathway. Appl. Environ. Microbiol. 53:14-16. McKnight, G. L., and B. L. McConaughy. 1983. Selection of functional cDNAs by complementation in yeast. Proc. Natl. Acad. Sci. USA 80:4412-4416. McKnight, G. L., H. Kato, A. Upshall, M. D. Parker, G. Saari, and P. J. O’Hara. 1985. Identification and molecular analysis of a third Aspergillus nidulans alcohol dehydrogenase gene. EMBO J. 4:2093-2099. Mellon, F. M., P. F. R Little, and L. A. Casselton. 1987. Gene cloning and transformation in the basidiomycete fungus Coprinus cinereus: isolation and expression of the isocitrate lyase gene (acu-7). Mol. Gen. Genet. 210:352-357. 340. 341. 342. 343. 344. 345. 346. 347. 348. 349. 350. 351. 352. 238 Melton, D. A., P. A. Krieg,M. R. Rebagliati, T. Maniatis, K. Zinn, and M. R. Green. 1984. Efficient in vitro synthesis of biologically active RNA and RNA hybridization probes from plasmids containing a bacteriophage SP6 promoter. Nucleic Acids Res. 12:7035-7056. Miller, B. L., K. Y. Miller, K. A. Roberti, and W. E. Timberlake. 1987. Position-dependent and -independent mechanisms regulate cell-specific expression of the SpoCI gene cluster of Aspergillus nidulans. Mol. Cell. Biol. 7:427-434. Miller, B. L., K. Y. Miller, and W. E. Timberlake. 1985. Direct and indirect replacements in Aspergillus nidulans. Mol. Cell. Biol. 5:1714-1721. Miller, J. H 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Mirabito, P. M., T. M. Adams, and W. E. Timberlake. 1989. Interactions of three sequentially expressed genes control temporal and spatial specificity in Aspergillus development. Cell 57:859-868. Mishra, N. C. 1977. Characterization of the new osmotic mutants (as) which originated during genetic transformation in Neurospora crassa. Genet. Res. 29:9-19. Mishra, N. C. 1979. DNA-mediated genetic changes in Neurospora crassa. J. Gen. Microbiol. 113:255-259. Mishra, N. C. 1985. Gene transfer in fungi. Adv. Genet. 23:73-178. Mishra, N. C., and E. L. Tatum. 1973. Non-mendelian inheritance of DNA-induced inositol independence in Neurospora. Proc. Natl. Acad. Sci. USA 70:3875-3879. Mishra, N. C., G. Szabo, and E. L. Tatum. 1973. Nucleic acid induced genetic changes in Neurospora, pp. 259-268. In Niu, M. C., and S. J. Segal (eds.), The role of RNA in reproduction and development. Elsevier/North Holland Publishing Co., Amsterdam. Mislivec, P. B., C. T. Dieter, and V. R. Bruce. 1975. Mycotoxin-producing potential of mold flora of dried beans. Appl. Microbiol. 29:522-526. Moore, P. M., and J. F. Peberdy. 1976. Release and regeneration of proto- plasts from the conidia of Aspergillus flavus. Trans. Br. Mycol. Soc. 66:421-425. , Morris, N. R. 1986. The molecular genetics of microtubule proteins in fungi. Exp. Mycol. 10:77-82. 353. 354. 355. 356. 357. 358. 359. 360. 361. 362. 363. 239 Moss, M. O. 1977. Aspergillus mycotoxins, pp. 499-524. In Smith, J. E., and J. A. Pateman (eds.), Genetics and physiology of Aspergillus. Academic Press, Inc., New York, NY. Mullaney, E. J., J. E. Hamer, K. A. Roberti, M. M. Yelton, and W. E. Timberlake. 1985. Primary structure of the trpC gene from Aspergillus nidulans. Mol. Gen. Genet. 199:37-45. Mullaney, E. J., P. J. Punt, and C. A. M. J. J. van den Handel. 1988. DNA mediated transformation of Aspergillus ficuum. Appl. Microbiol. Biotechnol. 28:451-454. Munoz-Rivas, A. M., C. A. Specht, R. C. Ullrich, and C. P. Novotrry. 1986a. Isolation of the DNA sequence coding indole-3-glycerol phosphate synthetase and phosphoribosylanthranilate isomerase of Schizophyllum commune. Curr. Genet. 10:909-913. Mur‘ioz-Rivas, A., C. A. Specht, B. J. Drummond, E. Froeliger, C. P. Novotny, and R. C. Ullrich. 1986b. Transformation of the basidiomycete, Schizophyllum commune. Mol. Gen. Genet. 205:103-106. Murakami, H. 1971. Classification of the koji mold. J. Gen. Appl. Microbiol. 17:281-309. Murakami, H., H. Sagawa, and S. Takase. 1968a. Non-productivity of aflatoxin by Japanese industrial strains of the Aspergillus. 111. Common characteristics of the aflatoxin-producing strains. J. Gen. Appl. Microbiol. 14:251-262. Murakarrri, H., S. Takase, and T. Ishii. 1967. Non-productivity of aflatoxin by Japanese industrial strains of Aspergillus. 1. production of fluorescent substances in agar slant and shaking culture. J. Gen. Appl. Microbiol. 13:323-334. Murakami, H., S. Takase, and K. Kuwabara. 1968b. Non-productivity of aflatoxin by Japanese industrial su'ains of Aspergillus. IL production of fluorescent substances in rice koji, and their identification by absorption spectrum. J. Gen. Appl. Microbiol. 14:97-110. Murray, N. E., W. J. Brarrrmar, and K. Murray. 1977. Lambdoid phages that simplify the recovery of in vitro recombinants. Mol. Gen. Genet. 150:53-61. Musflkova, M., and Z. Fencl. 1968. Some factors affecting the formation of protoplasts in Aspergillus niger. Folia Microbiol. 13:235-239. 364. 365. 366. 367. 368. 369. 370. 371. 372. 373. 374. 375. 240 Nachmias, A., and J. BaraSh. 1976. Decreased permeability as a mechanism of resistance to methyl benzirnidazol-Z-yl-carbamate (MBC) in Sporobolomyces roseus. J. Gen. Microbiol. 94:167-172. National Institute of Health, Department of Health and Human Services. 1986. Guidelines for Research involving recombinant DNA molecules. Federal Register 51:16957-16985. Neff, N. F., J. H. Thomas, P, Grisafi, and D. Botstein. 1983. Isolation of B- tubulin gene from yeast and demonstration of its essential function in viva. Cell 33:211-219. Neumann, F., and P. Bierth. 1986. Gene transfer by electroporation. Amer. Biotechnol. Lab. 4:10-15. Newbury, S. F., J. A. Glazebrook, and A. Radford. 1986. Sequence analysis of the pyr-4 (orotidine 5’-P decarboxylase) gene of Neurospora crassa. Gene 43:54-58. Newton, A. C., and C. E. Caten. 1988. Auxou'ophic mutants of Seproria nodorum isolated by direct screening and by selection for resistance to chlorate. Trans. Br. Mycol. Soc. 90:199-207. N gindu, A., P. R. Kenya, D. M. Ocheng, T. N. Omondi, W. Ngare, D. Gatei, B. K. Johnson, J. A. Ngira, H. Nandwa, A. J. Jansen, J. N. Kaviti, and T. A. Siongok 1982. Outbreak of acute hepatitis caused by aflatoxin poisoning in Kenya. Lancetzl346-1348. Nunberg, J. H., J. H. Meade, G. Cole, F. C. Lawyer, P. McCabe, V. Schweikart, R. Tal, V. P. Wittrrran, J. E. Flatgard, and M. E. Innis. 1984. Molecular cloning and characterization of the glucoamylase gene of Aspergillus awamori. Mol. Cell. Biol. 4:2306-2315. Oakley, B. R., J. E. Rinehart, B. L. Mitchell, C. E. Oakley, C. Carmana, G. L. Gray, and G. S. May. 1987. Cloning, mapping and molecular analysis of the pyrG (orotidine-5’-phosphate decarboxylase) gene of Aspergillus nidulans. Gene 61:385-399. Oakley, C. E., and B. R. Oakley. 1989. Identification of y-tubulin, a new member of the tubulin superfamily encoded by mipA gene of Aspergillus nidulans. Nature 338:662-664. O’Hara, E. B., and W. E. Timberlake. 1989. Molecular characterization of the Aspergillus nidulans yA locus. Genetics 121:249-254. Oka, A., H. Sugisaki, and M. Takanami. 1981. Nucleotide sequence of the kanamycin resistance transposon Tn903. J. Mol. Biol. 147 :217-226. 376. 377. 378. 379. 380. 381. 382. 383. 384. 385. 386. 241 Oliver, R. P., I. N. Roberts, R. Harling, L. Kenyon, P. J. Punt, M. A. Dingemanse, and C. A. M. J. J. van den Handel. 1987. Transformation of F ulvia fulva, a fungal pathogen of tomato, to hygromycin B resistance. Curr. Genet. 12:231-233. Orbach, M. J., E. B. Porro, and C. Yanofsky. 1986. Cloning and charac- terization of the gene for beta-tubulin from a benomyl-resistant mutant of Neurospora crassa and its use as a dominant delectable marker. Mol. Cell. Biol. 6:2452-2461. Orrell, J. C., C. J. R. Klittich, and J. F. Leslie. 1987. Nitrate nonutilizing mutants of F usarium oxysporum and their use in vegetative compatibility tests. Phytopathol. 77:1640-1646. Osiewacz, H. D., and A. Weber. 1989. DNA mediated transformation of the filamentous fungus Curvularia lunata using a dominant selectable marker. Appl. Microbiol. Biotechnol. 30:375-380. Osmani, S. A., D. B. Engle, J. H. Doonan, and N. R. Morris. 1988a. Spindle formation and chromatin condensation in cells blocked at interphase by mutation of a negative cell cycle control gene. Cell 52:241-251. Osmani, S. A., R. T. Pu, and N. R. Morris. 1988b. Mitotic induction and maintenance by overexpression of a G,-specific gene that encodes a potential protein kinase. Cell 53:237-244. Paietta, J. V., and G. A. Marzluf. 1985. Gene disruption by transformation in Neurospora crassa. Mol. Cell. Biol. 5:1554-1559. Palmer, L. M., and D. J. Cove. 1975. Pyrinridine biosynthesis in Aspergillus nidulans: isolation and preliminary characterization of auxotrophic mutants. Mol. Gen. Genet. 138:243-255. Panaccione, D. G., M. McKieman, and R. M. Hanau. 1988. Colletotrichum graminicola transformed with homologous and heterologous benomyl- resistance genes retains expected patlrogenicity to corn. Mal. Plant-Microbe Interact. 1:113-120. Papa, K. E. 1973. The parasexual cycle in Aspergillus flavus. Mycologia 65:1201-1205. Papa, K. E. 1976. Linkage groups in Aspergillus flavus. Mycologia 68:159-165. 387. 388. 389. 390. 391. 392. 393. 394. 395. 396. 397. 398. 399. 242 Papa, K. E. 1977a. Genetics of aflatoxin production in Aspergillus flavus: linkage between a gene for a high B,:B, ratio and the histidine locus on linkage group VIII. Mycologia 69:1185-1190. Papa, K. E. 1977b. Mutants of Aspergillus flavus producing more aflatoxin B, than B,. Appl. Environ. Microbiol. 33:206. Papa, K. E. 1977c. Genetic analysis of a mutant of Aspergillus flavus producing more aflatoxin B, than B,. Mycologia 69:556-562. Papa, K. E. 1978. The parasexual cycle in Aspergillus parasiticus. Mycologia 70:766-773. Papa, K. E. 1979. Genetics of Aspergillus flavus: complementation and mapping of aflatoxin mutants. Genet. Res. 34:1-9. Papa, K. E. 1980. Dominant aflatoxin mutant of Aspergillus flavus. J. Gen. Microbiol. 118:279-282. Papa, K. E. 1982. Norsolorinic acid mutant of Aspergillus flavus. J. Gen. Microbiol. 128:1345-1348. Papa, K. E. 1984. Genetics of Aspergillus flavus: linkage of aflatoxin mutants. Can. J. Microbiol. 30:68-73. Papa, K. E. 1986. Heterokaryon incompatibility in Aspergillus flavus. Mycologia 78:98-101. Pardo, J. M., F. Malpartida, M. Rico, and A. Jimenez. 1985. Biochemical basis of resistance to hygromycin B in Streptomyces hygroscopicus- the producing organism. J. Gen. Microbiol. 131:1289-1298. Parsons, K. A., F. G. Chumley, and B. Valent. 1987. Genetic u'ansfor- mation of the fungal pathogen responsible for rice blast disease. Proc. Natl. Acad. Sci. USA 84:4161-4165. Pateman, J. A., D. J. Cove, B. M. Rever, and D. B. Roberts. 1964. A common cofactor for nitrate reductase and xanthine dehydrogenase which also regulates the synthesis of nitrate reductase. Nature 201:58-60. Peberdy, J. F. 1985. Mycolytic enzymes, pp. 31-44. In Peberdy, J. F., and L. Ferenczy (eds.), Fungal protoplasts: applications in biochemistry and genetics. Marcel Dekker, Inc., New York, NY. Peberdy, J. F. 1989. Fungi without coats- protoplasts as tools for myco- logical research. Mycal. Res. 93:1-20. i. 401. 402. 403. 405. 406. 408. 410. 411. 412. 413. 243 Peberdy, J. F., and L. Ferenczy. 1985. Fungal Protoplasts: applications in biochemistry and genetics. Marcel Dekker, Inc., New York, NY. Peberdy, J. F., and R. K. Gibson. 1971. Regeneration of Aspergillus nidulans protoplasts. J. Gen. Microbiol. 69:325-330. Peberdy, J. F., and S. Isaac. 1976. An improved procedure for protoplast isolation from Aspergillus nidulans. Microbios. Lett. 3:7-9. Peberdy, J. F., C. E. Buckley, D. C. Daltry, and P. M. Moore. 1976. Factors affecting pr0toplast release in some filamentous fungi. Trans. Br. Mycol. Soc. 67:23-26. Penalva, M. A., A. Tourir'ro, C. Patiflo, F. sanchez, J. M. Femandez-Sousa, and V. Rubia. 1985. Studies on transformation of Cephalosporium acremonium, pp. 59-68. In Timberlake, W. E. (ed.), Molecular genetics of filamentous fungi. Alan R. Liss, Inc., New York, NY. Penttilli, M. E., K. M. H. Nevalainen, A. Raynal, and J. K. C. Knowles. 1984. Cloning of Aspergillus niger genes in yeast. Expression of the gene coding Aspergillus B-glucosidase. Mol. Gen. Genet. 194:494-499. Penttila, M., H. Nevalainen, M. Ratta, E. Salminen, and J. Knowles. 1987. A versatile transformation system for the cellulolytic filamentous fungus Trichoderma reesei. Gene 61:155-164. Perkins, D. D. 1962. Preservation of Neurospora stock cultures with anhydrous silica gel. Can. J. Microbiol. 8:591-594. Perkins, D. D., A. Radford, D. Newmeyer, and M. Bjorkman. 1982. Chromosomal loci of Neurospora crassa. Microbiol. Rev. 46:426-570. Perrot, M., C. Barreau, and J. Begueret. 1987. Nonintegrative transformation in the filamentous fungus Podospora anserina: stabilization of a linear vector by the chromosomal ends of Tetrahymena thermophila. Mol. Cell. Biol. 7:1725-1730. Picard, M., R. Debuchy, J. Julien, and Y. Brygoo. 1987. Transformation by integration, in Podospora anserina. 11. targeting to the resident locus with cosmids and instability of the transformants. Mol. Gen. Genet. 210:129-134. Picknett, T. M., and G. Saunders. 1989. Transformation of Penicillium chrysogenum with selection for increased resistance to benomyl. FEMS Microbiol. Lett. 60:165-168. Picknett, T. M., G. Saunders, P. Ford, and G. Holt. 1987. Development of a gene transfer system for Penicillium chrysogenum. Curr. Genet. 12:449-455. 414. 415. 416. 417. 418. 419. 420. 421. 422. 423. 424. 425. 244 Pitt, J. 1., A. D. Hacking, and D. R. Glenn. 1983. An improved medium for the detection of Aspergillus flavus and A. parasiticus. J. Appl. Bacterial. 54:109-114. Pontecorvo, G. 1953. The genetics of Aspergillus nidulans. Adv. Genet. 5:141-238. Pratviel, G., J. Bemadou, and B. Meunier. 1986. DNA breaks generated by the bleomycin-iron III complex in the presence of KHSOS, a single oxygen donor. Biochem. Biophys. Res. Commun. 136:1013-1020. Punt, P. J., R. P. Oliver, M. A. Dingemanse, P. H. Pouwels, and C. A. M. J. J. van den Handel, 1987. Transformation of Aspergillus based on the hygromycin B resistance marker from Escherichia coli. Gene 56:117-124. Rabie, C. J., M. Steyn, and G. C. van Schalkwyk. 1977. New species of Aspergillus producing sterigmatocystin. Appl. Environ. Microbiol. 33:1023-1025. Radford, A., F. P. Buxton, S. F. Newbury, and J. A. Glazebrook. 1985. Regulation of pyrimidine metabolism in Neurospora, pp. 127-143. In Timberlake, W. E. (ed.), Molecular genetics of filamentous fungi. Alan R. Liss, Inc., New York, NY. Radford, A., 8. Pope, A. Sazei, M. J. Fraser, and J. H. Parish. 1981. Liposome-mediated genetic transformation of Neurospora crassa. Mol. Gen. Genet. 184:567-569. Raj, H. G., L. Viswanathan, H. S. R. Murthy, and T. A. Venkitasubramanian. 1969. Biosynthesis of aflatoxins by cell-free preparations from Aspergillus flavus. Experientia 25:1141-1142. Rambosek, J ., and J. Leach. 1987. Recombination DNA in filamentous fungi: progress and prospects. CRC Crit. Rev. Biotechnol. 6:357-393. Randall, T., T. R. Rao, and C. A. Reddy. 1987. Transformation of Phanerochaete chrysosporium, a lignin-degrading white-rot basidiomycete, to G418 resistance. ASM Annu. Meet. Abst. p 142. Rao, H. R. G., and P. K. Harien. 1972. Dichlorvos as an inhibitor of aflatoxin production on wheat, corn, rice, and peanuts. J. Econo. Entom. 65:988-989. Rao, H. R. G., and P. K. Harein. 1973. Inhibition of aflatoxin and zearalenone biosynthesis with dichlorvos. Bull. Environ. Contam. Toxicol. 10:112-115. 426. 427. 428. 429. 430. 431. 432. 433. 434. 435. 436. 437. 245 Rao, R. N., N. E. Allen, J. N. Hobbs, W. E., Jr., Alborn, H. A. Kirst, and J. W. Paschal. 1983. Genetic and enzymatic basis of hygromycin B resis- tance in Escherichia coli. Antirrricrob. Agents Chemother. 24:689-695. Raper, K. B., and D. I. Fennell. 1965. The genus Aspergillus Williams and Wilkins, Baltimore. Rasmussen, J. B., D. G. Panaccione, and R. M. Hanau. 1989. Improved transformation of Colletotrichum graminicola protoplasts and its use in cloning the PYRI gene. APS Annu. Meet. Abst. No. 551. Razanamparany, V., and J. Begueret. 1986. Positive screening and trans- T formation of ura5 mutants in the fungus Podospora anserina: characteri- zation of the transformants. Curr. Genet. 10:811-817. Razanamparany, V., and J. Bégueret. 1988. Non-homologous integration of transforming vectors in the fungus Podospora anserina: sequences of .. junctions at the integration sites. Gene 74:399-409. L Reddy, T. V., L. Viswanathan, and T. A. Venkitasubramanian. 1971. High aflatoxin production on a chemically defined medium. Appl. Microbiol. 22:393-396. Reddy, T. V., L. Viswanathan, and T. A. Venkitasubramanian. 1979. Factors affecting aflatoxin production by Aspergillus parasiticus in a chemically defined medium. J. Gen. Microbiol. 114:409-413. Revuelta, J. L., and M. Jayararrr. 1986. Transformation of Phycomyces blakesleeanus to G-418 resistance by an autonomously replicating plasmid. Proc. Natl. Acad. Sci. USA 83:7344-7347. Revuelta, J. L., and M. Jayaram. 1987 . Phycomyces blakesleeanus TRP] gene: organization and functional complementation in Escherichia coli and Saccharomyces cerevisiae. Mol. Cell. Biol. 7:2664-2670. Richardson, 1. B., S. K. Hurley, and M. J. Hynes. 1989. Cloning and molecular characterization of the ade controlled gatA gene of Aspergillus nidulans. Mol. Gen. Genet. 217:118-125. Richey, M. G., E. T. Marek, C. L. Schardl, and D. A. Smith. 1989a. Transformation of filamentous fungi with plasmid DNA by electroporation. Phytopathol. 79:844-847. Richey, M. G., C. L. Schardl, and D. A. Smith. 1989b. Isolation of the ARGB gene from Fusarium solani f. sp. phaseoli. APS Annu. Meet. Abst. No. 293. 438. 439. 441. 442. 443. 445. 447. 449. 246 Roberts, K., and J. S. Hyams. 1979. Microtubules. Academic Press, Inc., London. Roberts, 1. N., R. P. Oliver, P. J. Punt, and C. A. M. J. J. van den Handel. 1989. Expression of the Escherichia coli B—glucuronidase gene in industrial and phytopathogenic filamentous fungi. Curr. Genet. 15:177-180. Rodriguez, R. J., and L. W. Parks. 1980. Growth and antifungal homoaza- sterol production in Geotrichum flavo-brunneum. Antimicrob. Agents Chemother. 18:822-828. Rodriguez, R. J., and O. C. Yoder. 1987. Selectable genes for transfor- mation of the fungal plant pathogen Glomerella dingulata f. sp. phaseoli (Colletotrichurn lindemuthianum). Gene 54:73-81. Roper, J. A. 1966. Mechanisms of inheritance. 3. the parasexual cycle, pp. 589-617. In Ainsworth, G. C., and A. S. Sussman (eds.), The fungi. Vol. II. The fungal organism. Academic Press, Inc., New York, NY. Rosenberger, R. F. 1976. The cell wall, pp. 328-344. In Smith, J. E., and D. R. Berry (eds.), The filamentous fungi, Vol. 11. Edward Arnold, London. Rowlands, R. T., and G. Turner. 1973. Nuclear and extranuclear inheritance of oligomycin resistance in Aspergillus nidulans. Mol. Gen. Genet. 126:201-216. Rowlands, R. T., and G. Turner. 1977. Nuclear-extranuclear interactions affeCting oligomycin resistance in Aspergillus nidulans. Mol. Gen. Genet. 154:311-318. Russel, P. J., J. A. Welsch, and S. Wagner. 1989. Transformation of Neurospora crassa by an integrative transforming plasmid is not enhanced by ribosomal DNA sequences. Biochim. Biophys. Acta 1008:243-246. Sakai, K., and M. Yamam0to. 1986. Transformation of the yeast, Saccha- romyces carlsbergensis, using an antibiotic resistance marker. Agric. Biol. Chem. 50:1177-1182. Sakai, K., J. Sakaguchi, and M. Yamamoto. 1984. High-frequency cotrans- formation by c0polymerization of plasmids in the fission yeast Schizo- saccharomyces pombe. Mol. Cell. Biol. 4:651-656. Salch, Y. P., and M. N. Beremand. 1989. Identification of sequences with promoter activity from Gibberella pulicaris (Fusarium sambucinum) transformants. APS Annu. Meet. Abst. No. 294. 450. 451. 452. 453. 454. 455. 456. 457. 458. 459. 461. 462. 247 Samac, D. A., and S. A. Leong. 1988. Two linear plasmids in mitochondria of Fusarium solani f. sp. cucurbitae. Plasmid 19:57-67. Samac, D. A., and S. A. Leong. 1989. Characterization of the termini of linear plasmids from Nectria haemarococca and their use in construction of an autonomously replicating uansformation vector. Curr. Genet. 16:187- 194. Sénchez, F., M. Lozano, V. Rubia, and M. A. Penalva. 1987. Transforma- tion in Penicillium chrysogenum. Gene 51:97-102. Sénchez, F., A. Tourir'io, S. Traseira, A. P6rez-Aranda, V. Rubia, and M. A. Pefialva, 1986. Molecular cloning and characterization of the trpC gene from Penicillium chrysogenum. Mol. Gen. Genet. 205:248-252. Sandeman, R. A., and M. J. Hynes. 1989. Isolation of the facA (acetyl- coenzyme A synthetase) and acuE (malate synthase) genes of Aspergillus nidulans. Mol. Gen. Genet. 218:87-92. Santerre, R. F., N. E. Allen, J. N., Jr., Hobbs, R. N. Rao, and R. J. Schmidt. 1984. Expression of prokaryotic genes for hygromycin B and G418 resistance as dominant-selection markers in mouse L cells. Gene 30:147-156. Saunders, G., M. F. Tuite, and G. Holt. 1986. Fungal cloning vectors. Trends. Biatechnol. 4:93-98. Saunders, G., T. M. Picknett, M. F. Tuite, and M. Ward. 1989. Hetero- logous gene expression in filamentous fungi. Trends Biotechnol. 7:283-287. Scazzocchio, C., and H. N., Jr., Arst, 1978. The nature of an initiator constitutive mutation in Aspergillus nidulans. Nature 274:177-179. Schatz, P. J., L. Pillus, P. Grisafi, F. Solomon, and D. Botstein. 1986. Two functional alpha tubulin genes of the yeast Saccharomyces cerevisiae encode divergent proteins. Mol. Cell. Biol. 6:3711-3721. Schechtrnan, M. G. 1987. Isolation of telomere DNA from Neurospora crassa. Mol. Cell. Biol. 7:3168-3177. Schechtrnan, M. G., and C. Yanofsky. 1983. Structure of the uifunctional trp-I gene from Neurospora crassa and its aberrant expression in Esche- richia coli. J. Mol. Appl. Genet. 2:83-99. Schmidt, F. R., and K. Esser. 1985. Aflatoxins: medical, economic impact, and prospects for control. Process Biocherrr. 20:167-174. 463. 464. 465. 466. 467. 468. 469. 470. 471. 472. 473. 474. 248 Schmidt, F. R., N. D. Davis, U. L. Diener, and P. A. Lemke. 1983. Cyclo- heximide induction of aflatoxin synthesis in a nontoxigenic strain of Aspergillus flavus. Bio/Technol. 1:794-795. Schmidt, F. R., P. A. Lemke, and K. Esser. 1986. Viral influences on aflatoxin formation by Aspergillus flavus. Appl. Microbiol. Biotechnol. 24:248-252. Schroeder, H. W., and W. W. Carlton. 1973. Accumulation of only afla- toxin B, by a strain of Aspergillus flavus. Appl. Microbiol. 25:146-148. Schroeder, H. W., and W. H. Kelton. 1975. Production of sterigmatocystin by some species of the genus Aspergillus and its toxicity to chicken embryos. Appl. Environ. Microbiol. 30:589-591. Schroeder, H. W., R. J. Cole, R. D. Grigsby, and H., Jr., Hein. 1974. Inhibition of aflatoxin production and tentative identification of an aflatoxin intermediate "versiconal acetate" from treatment with dichlorvos. Appl. Microbiol. 27:394-399. Schwab, H. 1988. Strain improvement in industrial microorganisms by recombinant DNA techniques. Adv. Biochem. Eng. Biotechnol. 37:129-168. Sealy-Lewis, H. M., and R. A. Lockington. 1984. Regulation of two alcohol dehydrogenases in Aspergillus nidulans. Curr. Genet. 8:253-259. Sebald, W., and J. Hoppe. 1981. On the structure and genetics of the proteolipid subunit of the ATP synthase complex. Curr. Top. Bioenerg. 12:1-64. Seip, E. R., C. P. Woloshuk. G. A. Payne, and C. R. Adkins. 1988. Iso- lation of cloned DNA from a benomyl resistant mutant of Aspergillus flavus with homology to the benA gene of A. nidulans. APS Annu. Meet. Abst. No. 294. Seip, E. R., C. P. Woloshuk, G. A. Payne, and C. R. Adkins. 1989. Expre- ssion of MBC (benomyl) resistance in Aspergillus flavus transformed with a homologous B-tubulin gene. APS Annu. Meet. Abst. No. 552. Sekita, S., K. Yoshihira, S. Natori, S. Udagawa, T. Muroi, Sugiyama, Y, and H. Kurata. 1981. Mycotoxin production by Chaetomium spp. and related fungi. Can. J. Microbiol. 27:766-772. Selker, E. U., E. B. Cambareri, B. C. Jensen, and K. R. Haack. 1987. Rearrangement of duplicated DNA in specialized cells of Neurospora. Cell 51:741-752. 475. 476. 477. 478. 479. 480. 481. 482. 483. 484. 485. 486. 487. 249 Semon, D., N. R. Movva, T. F. Smith, M. El Alama, and J. Davies. 1987. Plasmid-determined bleomycin resistance in Staphylococcus aureus. Plasmid 17:46-53. Sequin, U., and I. Scatt. 1974. Carbon-13 as a label in biosynthetic studies. Science 186:101-107. Sheir-Neiss, G., M. Lai, and N. Morris. 1978. Identification of a gene for [3- tubulin in Aspergillus nidulans. Cell 15:639-647. Sherwood, J. E., and S. C. Sommerville. 1989. Molecular characterization of a B-tubulin gene from Erysiphe graminis. APS Annu. Meet. Abst. No. 560. Sietsma, J. H., and W. R. de Boer. 1973. Formation and regeneration of protoplasts from Pythium PRL 2142. J. Gen. Microbiol. 74:211-217. Singh, A., and F. Sherman. 1974. Characteristics and relationships of mercury-resistant mutants and methionine auxotrophy of yeast. J. Bacteriol. 118:911-918. Singh, A., and F. Sherman. 1975. Genetic and physiological characterization of met15 mutants of Saccharomyces cerevisiae: a selective system for forward and reverse mutations. Genetics 81:75-97. Singh, R., and D. P. H. Hsieh. 1976. Enzymatic conversion of sterigma- tocystin into aflatoxin B, by cell-free extracts of Aspergillus parasiticus. Appl. Environ. Microbiol. 31:743-745. Singh, R., and D. P. H. Hsieh. 1977. Aflatoxin biosynthetic pathway: elucidation by using blocked mutants of Aspergillus parasiticus. Arch. Biochem. Biophys. 178:285-292. Siyan, A., T. E. Stasz, G. E. Harman, and M. Hemmat. 1989. Transfor- mation of T richoderma spp. to hygromycin B resistance. APS Annu. Meet. Abst. No. 298. Skatrud, P. L., and S. W. Queener. 1984. Cloning of a DNA fragment from Cephalosporium which functions as an autonomous replication sequence in yeast. Curr. Genet. 8:155-163. Skatrud, P. L., S. W. Queener, L. G. Carr, and D. L. Fisher. 1987. Efficient integrative transformation of Cephalosporium acremonium. Curr. Genet. 12:337-348. Smith, J. E., and M. 0. Moss. 1985. Mycotoxins: formation, analysis and significance. John Wiley & Sons, New York, NY. 488. 489. 490. 491. 492. 493. 494. 495. 496. 497. 498. 499. 500. 250 Soliday, c. r... M. B. Dickman, and P. E. Kolattukudy. 1989. Structure of the cutinase gene and detection of promoter activity in the 5’-flanking region by fungal transformation. J. Bacteriol. 171:1942-1951. Specht, C. A., A. Munoz-Rivas, C. P. Novotrry, and R. C. Ullrich. 1988. Transformation of Schizophyllum commune: an analysis of parameters for improving transformation frequencies. Exp. Mycol. 12:357-366. Stahl, U., P. Tudzynski, U. Kuck. and re Esser. 1982. Replication and expression of a bacterial-mitochondrial hybrid plasmid in the fungus Podospora anserina. Proc. Natl. Acad. Sci. USA 79:3641-3645. Steyn, P. S. 1980. The Biosynthesis of Mycotoxins: a study in secondary metabolism. Academic Press, Inc., New York, NY. Steyn, P. S., R. Vleggaar, and P. L. Wessels. 1980. The biosynthesis of aflatoxin and its congeners, pp. 105-155. In Steyn, P. S. (ed.), The biosynthesis of mycotoxins: a study in secondary metabolism. Academic Press, Inc., New York, NY. Stinchcomb, D. T., M. Thomas, J. Kelly, E. Selker, and R. W. Davis. 1980. Eukaryotic DNA segments capable of autonomous replication in yeast. Proc. Natl. Acad. Sci. USA 77:4559-4563. Stahl, L. L., and A. M. Lambowitz. 1983. Construction of a shuttle vector for the filamentous fungus Neurospora crassa. Proc. Natl. Acad. Sci. USA 80: 1058- 1062. Stahl, L. L., R. A. Atkins, and A. M. Lambowitz. 1984. Characterization of deletion derivatives of an autonomously replicating Neurospora plasmid. Nucleic Acids Res. 12:6169-6178. Stoloff, L., and W. Trager. 1965. Recommended decontamination proce- dures for aflatoxin. J. Assoc. Off. Anal. Chem. 48:681-682. Struhl, K., D. T. Stinchcomb, S. Scherer, and R. W. Davis. 1979. High- frequency transformation of yeast: autonomous replication of hybrid DNA molecules. Proc. Natl. Acad. Sci. USA 76:1035-1039. Suarez, T., and A. Eslava. 1988. Transformation of Phycomyces with a bacterial gene for kanamycin resistance. Mol. Gen. Genet. 212:120-123. Suzci, A., and A. Radford. 1983. ARS8 sequences in the Neurospora genome. Neurospora Newslett. 30:13. Swenson, D. H. 1981. Metabolic activation and detoxication of aflatoxins. Rev. Biochem. Toxicol. 3:155-192. 501. 502. 503. 504. 505. 506. 507. 508. 509. 510. 511. 512. 251 TeBeest, D. O., and M. B. Dickrnam. 1989. Transformation of Colleto- trichum gloeosporioides f. sp. aeschynomene. APS Annu. Meet Abst. No. 301. Thomas, G. H., I. F. Connerton, and J. R. S. Fincham. 1988. Molecular cloning, identification and transcriptional analysis of genes involved in acetate utilization in Neurospora crassa. Mol. Microbiol. 2:599-606. Thomas, 1. H., N. Neff, and D. Botstein. 1985. Isolation and character- ization of mutations in the B-tubulin gene of Saccharomyces cerevisiae. Genetics 112:715-734. Thomas, M. D., and C. M. Kenerley. 1989. Transformation of the mycopa- rasite Gliocladium. Curr. Genet. 15:415-420. Tilburn, J., C. Scazzocchio, G. G. Taylor, J. H. Zabicky-Zissman, R. A. Lockington, and R. W. Davis. 1983. Transformation by integration in Aspergillus nidulans. Gene 26:205-221. Timberlake, W. E. 1978. Low repetitive DNA content in Aspergillus nidulans. Science 202:773-775. Timberlake, W. E. 1985. Molecular genetics of filamentous fungi. Alan R. Liss, New York, NY. Timberlake, W. E. 1987. Molecular genetic analysis of development in Aspergillus nidulans, pp. 63-82. In Loomis, W. F. (ed), Genetic regulation of development. Alan R. Liss, Inc., New York, NY. Timberlake, W. E., and M. A. Marshall. 1988. Genetic regulation of deve- lopment in Aspergillus nidulans. Trends Genet. 4:162-169. Timberlake, W. E., and M. A. Marshall. 1989. Genetic engineering of fila- mentous fungi. Science 244:1313-1317. Timberlake, W. E., M. T. Boylan, M. B. Cooley, P. M. Mirabito, E. B. O’Hara, and C. E. Willett. 1985. Rapid identification of mutation-comple- menting resuiction fragments from Aspergillus nidulans cosmids. Exp. Mycol. 9:351-355. Toda, T., Y. Adachi, Y. Hiraoka, and M. Yanagida. 1984. Identification of the pleiotropic cell division cycle gene NDA2 as one of two different alpha tubulin genes in Schizosaccharomyces pombe. Cell 37:233-242. 513. 514. 515. 516. 517. 518. 519. 520. 521. 522. 523. 524. 252 Tomsett, A. B., and R. H. Garrett. 1980. The isolation and characterization of mutants defective in nitrate assimilation in Neurospora crassa. Genetics 95:649-660. Torres, J., J. Guarro, G. Suarez, N. Sune, M. A. Calvo, and C. Ramirez. 1980. Morphological changes in suains of Aspergillus flavus Link ex Fries and Aspergillus parasiticus Speare related with aflatoxin production. Myc0pathologia 72: 17 1-174. Torrey, G. S., and E. H. Marth. 1976. Silica gel medium to detect molds that produce aflatoxin. Appl. Environ. Microbiol. 32:376-380. Townsend, C. A., K. A. Plavcan, K. Pal, and S. W. Brobst. 1988. Hydroxy- versicolorone: isolation and characterization of a potential intermediate in aflatoxin biosynthesis. J. Amer. Chem. Soc. 53:2472-2477. Townsend, C. A. 1986. Progress toward a biosynthetic rationale of the aflatoxin pathway. Pure Appl. Chem. 58:227-238. Tschumper, G., and J. Carbon. 1980. Sequence of a yeast DNA fragment containing a chromosomal replicator and the TRP] gene. Gene 10:157-166. Tsukuda, T., S. Carleton, S. Fotheringham, and W. K. Holloman. 1988. Isolation and characterization of an autonomously replicating sequence from Ustilago maydis. Mol. Cell. Biol. 8:3703-3709. Tudzynski, P., and K. Esser. 1980. Transformation to senescence with plasmid like DNA in the ascomycete Podospora anserina. Curr. Genet. 2:181-184. Tudzynski, P., and K. Esser. 1985. Mitochondrial DNA for gene cloning in eukaryotes, pp. 403-416. In Bennett, J. W., and L. L. Lasure (eds.), Gene manipulation in fungi. Academic Press, Inc., New York, NY. Tudzynski, P., U. Stahl, and K. Esser. 1982. Development of a eukaryotic cloning system in Podospora anserina. I. Long-lived mutants as potential recipients. Curr. Genet. 6:219-222. Turcq, B., and J. Begueret. 1987. The ura5 gene of the filamentous fungus Podospora anserina: nucleotide sequence and expression in transformed strains. Gene 53:201-209. Turgeon, B. G., R. C. Garber, and O. C. Yoder. 1985. Transformation of the fungal maize pathogen Cochliobolus heterostrophus using the Aspergillus nidulans amdS gene. Mol. Gen. Genet. 201:450-453. 525. 526. 527. 528. 529. 530. 531. 532. 533. 534. 535. 253 Turgeon, B. G., R. C. Garber, and O. C. Yoder. 1987. Development of a fungal transformation system based on selection of sequences with promoter activity. Mol. Cell. Biol. 7:3297-3305. Turgeon, B. G., W. D. MacRae, R. C. Garber, G. R. Fink, and O. C. Yoder. 1986. A cloned tryptophane-synthesis gene from the ascomycete Cochliobolus heterostrophus functions in Escherichia coli, yeast and Aspergillus nidulans. Gene 42:79-88. Tumbull, I. F., K. Rand, N. S. Willetts, and M. J. Hynes. 1989. Expression of the Escherichia coli enterotoxin subunit B gene in Aspergillus nidulans directed by the amdS promoter. Bio/Technol. 7:169-174. Turner, G., and D. J. Ballance. 1985. Cloning and transformation in Aspergillus, pp. 259-278. In Bennett, J. W., and L. L. Lasure (eds.), Gene manipulations in fungi. Academic Press, Inc., New York, NY. Turner, G., D. J. Ballance, M. Ward. and R. K. Beri. 1985. Development of cloning vectors and a marker for gene replacement techniques in Aspergillus nidulans, pp. 15-28. In Timberlake, W. E. (ed.), Molecular genetics of filamentous fungi. Alan R. Liss, Inc., New York. NY. - Turner, W. B. 1971. Fungal metabolites. Academic Press, Inc., New York, NY. Turner, W. B., and D. C. Aldridge. 1983. Fungal metabolites 11. Academic Press, Inc. New York, NY. Tyagi, J. S., A. K. Tyagi, and T. A. Venitasubrarnanian. 1981. Preparation and properties of spheroplasts from Aspergillus parasiticus with special reference to the de novo synthesis of aflatoxins. J. Appl. Bacterial. 50:481-491. Udagawa, S., T. Muroi, H. Kurata. S. Sekita, K. Yoshihira, and S. Natori. 1979. The production of chaetoglobsins, sterigmatocystin, O-methylsterig- matocystin, and chaetocin by Chaetomium spp. and related fungi. Can. J. Microbiol. 25:170-177. Ueda, K., S. Kobayashi, H. Sakai, and T. Komano. 1985. Cleavage of stem and loop structure DNA by bleomycin. J. Biol. Chem. 260:5804-5807. Ullrich, R. C., C. P. Novotny, C. A. Specht, E. H. Froeliger, and A. M. Munoz-Rivas. 1985. Transforming Basidiomycetes, pp. 39-57. In Timberlake, W. E. (ed.), Molecular genetics of filamentous fungi. Alan R. Liss, Inc., New York, NY. 536. 537. 538. 539. 540. 541. 542. 543. 544. 545. 254 Umezawa, H. 1974. Chemistry and mechanism of action of bleomycin. Fed. Proc. 33:2296—2302. Unkles, S. E. 1989. Fungal biotechnology and the nitrate assimilation pathway, pp 341-363. In Wray, J. L., and J. R. Kinghom (eds.), Molecular and genetic aspects of nitrate assimilation. Oxford Science Publications, Oxford. Unkles, S. E., E. 1. Campbell, Y. M. J. T. de Ruiter-Jacobs, M. Broekhuijsen, J. A. Macro, D. Carrez, R. Contreras, C. A. M. J. J. van den Handel, and J. R. Kinghom. 1989a. The development of a homologous transformation system for Aspergillus oryzae based on the nitrate assimilation pathway: a convenient and general selection system for filamentous fungal transformation. Mol. Gen. Genet. 218:99-104. Unkles, S. E., E. 1. Campbell, D. Carrez, C. Grieve, R. Contreras, W. Fibers, C. A. M. J. J. van den Handel, and J. R. Kinghom. 1989b. Trans- formation of Aspergillus niger with the homologous nitrate reductase gene. Gene 78:157-166. Upshall, A. 1986. Filamentous fungi in biotechnology. BioTechniques 4:158-166. Upshall, A., A. A. Kumar, M. C. Bailey, M. D. Parker, M. A. Favreau, K. P. Lewison, M. L. Joseph, J. M. Maraganore, and G. L. McKnight. 1987. Secretion of active human tissue plasminogen activator from the filamentous fungus Aspergillus nidulans. Bio/Technol. 5:1301-1304. Van Alfen, N. K., A. C. L. Churchill, D. R. Hansen, 1. M. Cuiffetti, and H. D. Van Etten. 1988. Transformation of Cryphonectria (Endothia) parasitica using a variety of fungal promoters. APS Annu. Meet. Abst. No. 290. Van Brunt, J. 1986. Fungi: the perfect hosts? Bio/Technol. 4:1057-1062. van den Handel, C. A. M. J. J., R. F. M. van Gorcom, T. Goosen, H. W. J. van den Broek, W. E. Timberlake, and P. H. Pouwels. 1985. Development of a system for analysis of regulation signals in Aspergillus, pp. 29-38. In Tirrrberlake, W. E. (ed.), Molecular genetics of filamentous fungi. Alan R. Liss, Inc. New York, NY. van den Handel, C. A. M. J. J., P. J. Punt, B. L. M. Jacobs-Meijsing, W. van Hartingsveldt, R. F. M. van Gorcorrr, and P. H. Pouwels. 1986. Analysis of transcription-control signals in Aspergillus, pp. 365-369. In Bailey, J. (ed.), Biology and molecular biology of plant-pathogen interactions. Springer-Verlag, Berlin. 546. 547. 548. 549. 550. 551. 552. 553. 554. 555. 556. 557. 255 van Engelenburg, F., R. Srrrit, T. Goosen, H. van den Brock, and P. Tudzynski. 1989. Transformation of Claviceps purpurea using a bleomycin resistance gene. Appl. Microbiol. Biotechnol. 30:364-370. Van Etten, H. D. and H. C. Kistler. 1988. Necrria haematococca, mating populations 1 and VI. Adv. Plant Pathol. 6:189-206. van Gorcom, R. F. M., P. H. Pouwels, T. Goosen, J. Visser, H. W. J. van den Broek, J. E. Hamcr, W. E. Timberlake, and C. A. M. J. J. van den Handel. 1985. Expression of an Escherichia coli B-galactosidase fusion gene in Aspergillus nidulans. Gene 40:99-106. van Gorcom, R. F. M., P. J. Punt, P. H. Pouwels, and C. A. M. J. J. van den Handel. 1986. A system for the analysis of expression signals in Aspergillus. Gene 48:211-217. van Hartingsveldt, W., I. E. Mattem, C. M. J. van Zeijl, P. H. Pouwels, , and C. A. M. J. J. van den Handel. 1987. Development of a homologous transformation system for Aspergillus niger based on the pyrG gene. Mol. Gen. Genet. 206:71-75. van Heeswijck, R. 1986. Autonomous replication of plasmids in Mucor transformants. Carlsberg Res. Commun. 51:433-443. van Heeswijck, R. V., and M. I. G. Roncero. 1984. High frequency trans- formation of Mucor with recombinant plasmid DNA. Carlsberg Res. Commun. 49:691-702. van Walbeek, W., P. M. Scatt, and F. S. Thatcher. 1968. Mycotoxins from food-borne fungi. Can. J. Microbiol. 14:131-137. Vapnek, D., J. A. Jautala, J. W. Jacobson, N. H. Giles, and S. R. Kushner. 1977. Expression in Escherichia coli K-12 of the structural gene for catabolic dchydroquinase of Neurospora crassa. Proc. Natl. Acad. Sci. USA 74:3508-3512. Vogeli, G., E. Horn, M. Laurent, and P. Nath. 1985. Recombinant DNA techniques: storage and screening of cDNA libraries with large numbers of individual colonies from initial transformations. Anal. Biochem. 151:442-444. Vollmer, S. J., and C. Yanofsky. 1986. Efficient cloning of genes of Neurospora crassa. Proc. Natl. Acad. Sci. USA 83:4869-4873. Waldron, C., E. B. Murphy, J. L. Roberts, G. D. Gustafson, S. L. Armour, and S. K. Malcolm. 1985. Resistance to hygromycin B: a new marker for plant transformation studies. Plant Mol. Biol. 5:103-108. 558. 559. 560. 561. 562. 563. 564. 565. 566. 567. 568. 569. 256 Wan, N. C., and D. P. H. Hsieh. 1980. Enzymatic formation of the bisfuran structure in aflatoxin biosynthesis. Appl. Environ. Microbiol. 39:109-112. Wang, J., D. W. Holden, and S. A. Leong. 1988. Gene transfer system for the phytopathogenic fungus Ustilago maydis. Proc. Natl. Acad. Sci. USA 85:865-869. Ward, M., K. H. Kodama, and L. J. Wilson. 1989. Transformation of Aspergillus awamori and A. niger by electroporation. Exp. Mycol. 13:289-293. Ward, M., B. Wilkinson, and G. Turner. 1986. Transformation of Aspergillus nidulans with a cloned. oligomycin-resistant ATP synthase subunit 9 gene. Mol. Gen. Genet. 202:265-270. Ward, M., L. J. Wilson, C. L. Carmona, and G. Turner. 1988. The oliC gene of Aspergillus niger: isolation, sequence and use as a selectable marker for transformation. Curr. Genet. 14:37-42. Waring, R. B., G. S. May, and N. R. Morris. 1989. Characterization of an inducible expression system in Aspergillus nidulans using ach and tubulin-coding genes. Gene 79:119-130. Webster, T. D., and R. C. Dickson. 1983. Direct selection of Saccharo- myces cerevisiae resistant to the antibiotic G418 following transformation with a DNA vector carrying the kanamycin-resistance gene of Tn903. Gene 26:243-252. Wei, D. -L., and S. -C. Jong. 1986. Production of aflatoxins by strains of the Aspergillus flavus group maintained in ATCC. Mycopathologia 93:19-24. Weiss, B. 1965. An electron microscope and biochemical study of Neurospora crassa during development. J. Gen. Microbiol. 39:85-94. Weiss, R. L. 1985. Expression of Aspergillus genes in Neurospora, pp. 279-292. In Bennett, J. W., and L. L. Lasure (eds.), Gene manipulations in fungi. Academic Press, Inc., New York, NY. Weltring, K. -M., B. G. Turgeon, O. C. Yoder, and H. D. Van Etten. 1988. Isolation of a phytolexin-detoxification gene from the plant pathogenic fungus Nectria haemarococca by detecting its expression in Aspergillus nidulans. Gene 68:335-344. Wernars, K., T. Goosen, B. M. J. chnekes, K. Swart. C. A. M. J. J. van den Handel, and H. W. J. van den Broek. 1987 . Cotransformation of Aspergillus nidulans: a tool for replacing fungal genes. Mol. Gen. Genet. 209:71-77. 570. 571. 572. 573. 574. 575. 576. 577. 578. 579. 580. 257 Wernars, K, T. Goosen, L. M. J. Wennekes, J. Visser, C. J. Bos, H. W. J. van den Broek, R. F. M. van Gorcom, C. A. M. J. J. van den Handel, and P. H. Pouwels. 1985. Gene amplification in Aspergillus nidulans by trans- formation with vectors containing the amdS gene. Curr. Genet. 9:361-368. Wessels, J. G. H. 1986. Cell wall synthesis in apical hyphal growth. Int. Rev. Cytol. 104:37-79. Whitehead, M. P., S. E. Unkles, M. Ramsden, E. 1. Campbell, S. J. Gurr, D. Spence, C. van den Handel, R. Contreras, and J. R. Kinghom. 1989. Transformation of a nitrate reductase deficient mutant of Penicillium chrysogenum with the corresponding Aspergillus niger and A. nidulans niaD genes. Mol. Gen. Genet. 216:408-411. Wicklow, D. T. 1983. Taxonomic features and ecological significance of sclerotia, pp. 6-12. In Diener, U. L., R. L. Asquith, and J. W. Dickens (eds.), Aflatoxin and Aspergillus flaws in com. Auburn University, Craftmaster Printers, Opelika, Alabama. Woloshuk, C. P., E. R. Seip, G. A. Payne, and C. R. Adkins. 1989. Genetic transformation system for the aflatoxin-producing fungus Aspergillus flavus. Appl. Environ. Microbiol. 55:86-90. Wood, H. A., R. F. Bozarth, J. Adler, and D. W. Mackenzie. 1974. Protei- naceous virus-like particles from an isolate of Aspergillus flavus. J. Viral. 13:532-534. Wootton, J. C., M. J. Fraser, and A. J. Baron. 1980. Efficient transfor- mation of germinating Neurospora conidia using total nuclear fragments. Neurospora Newslett. 27:33. Wastemeyer, J ., A. Burrnester, and C. Weigel. 1987. Neomycin resistance as a dominantly selectable marker for transformation of the zygomycete Absidia glauca. Curr. Genet. 12:625-627. Woudt, L. P., J. J. van den Heuvel, M. M. C. van Raamsdonk-Duin, W. H. Mager, and R. J. Planta. 1985. Correct removal by splicing of a Neurospora intron in yeast. Nucleic Acids Res. 13:7729-7739. Wray, J. L., and J. R. Kinghom. 1989. Molecular and genetic aspects of nitrate assimilation. Oxford Science Publications, Oxford. Yabe, K., Y. Ando, and T. Harnasaki. 1988. Biosynthetic relationship among aflatoxins B,, B,, G,, and 0,. Appl. Environ. Microbiol. 54:2101-2106. 581. 582. 583. 584. 585. 586. 587. 588. 589. 590. 591. 592. 593. 258 Yabe, K., Y. Ando, J. Hashimoto, and T. Harnasaki. 1989. Two distinct O-mcthyltransfcrases in aflatoxin biosynthesis. Appl. Environ. Microbiol. 55:2172-2177. Yang, C. Y. 1972. Comparative studies on the detoxification of aflatoxins by sodium hypochlorite and commercial bleachcs. Appl. Microbiol. 24:885-890. Yanisch-Perron, C., J. Vieira, and J. Messing. 1985. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUCl9 vectors. Gene 33:103-119. Yao, R. C., and D. P. H. Hsieh. 1974. Step of dichlorvos inhibition in the pathway of aflatoxin biosynthesis. Appl. Microbiol. 28:52-57. Yeh, F. -S., M. C. Yu, C. -C. Ma, S. Luo, M. J. Tang, and B. E. Henderson. 1989. Hepatitis B virus, aflatoxins, and hepatocellular carcinoma in Southern Guangxi, China. Cancer Res. 49:2506-2509. Yelton, M. M., J. E. Hamcr, E. R. DeSouza, E. J. Mullaney, and W. E. Timberlake. 1983. Developmental regulation of the Aspergillus nidulans trpC gene. Proc. Natl. Acad. Sci. USA 80:7576-7580. Yelton, M. M., J. E. Hamer, and W. E. Timberlake. 1984. Transformation of Aspergillus nidulans by using a trpC plasmid. Proc. Natl. Acad. Sci. USA 81:1470-1474. Yelton, M. M., W. E. Timberlake, and C. A. M. J. J. van den Handel. 1985. A cosmid for selecting genes by complementation in Aspergillus nidulans: selection of the developmentally regulated yA locus. Proc. Natl. Acad. Sci. USA 82:834-838. Yoder, O. C., and B. G. Turgeon. 1985. Molecular bases of fungal patho- genicity to plants, pp. 417-48. In Bennett, J. W., and L. L. Lasure (eds.), Gene manipulations in fungi. Academic Press, Inc., New York, NY. Yoder, O. C., B. Valent, and F. Chumlcy. 1986. Genetic nomenclature and practice for plant pathogenic fungi. Phytopathol. 76:383-385. Yuill, E. 1950. The numbers of nuclei in conidia of Aspergilli. Trans. Br. Mycol. Soc. 33:324-331. Zaika, L. L., and R. L. Buchanan. 1987. Review of compounds affecting the biosynthesis or biorcgulation of aflatoxins. J. Food Prat. 50:691-708. Zamir, L. O., and R. Ginsburg. 1979. Aflatoxin biosynthesis: detection of transient, acetate-dependent intermediates in Aspergillus by kinetic pulse-labeling. J. Bacteriol. 138:684-690. 594. 595. 596. 597. 259 Zamir, L. 0., and K. D. Hufford. 1981. Precursor recognition by kinetic pulse-labeling in a toxigenic aflatoxin B,-producing strain of Aspergillus. Appl. Environ. Microbiol. 42:168-173. Zimmermann, U. 1982. Electric field-mediated fusion and related electrical phenomena. Biochim. Biophys. Acta 694:227-277. Zimmerrnann, U., and J. Vienken. 1982. Electric field-induced cell-to-ccll fusion. J. Membrane Biol. 67:165-182. Zurawski, G., D. Elsevicrs, V. Stauffer, and C. Yanofsky. 1978. Transla- tional control of transcription termination at the attenuator of the Escherichia coli tryptophan operon. Proc. Natl. Acad. Sci. USA 75:5988-5992. "11111111111111.1115